Protein Science Encyclopedia

1 Microcontact Printing of Proteins Emmanuel Delamarche IBM Research, Zürich Research Laboratory, Rüschlikon, Switzer...

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Microcontact Printing of Proteins Emmanuel Delamarche IBM Research, Zürich Research Laboratory, Rüschlikon, Switzerland

Originally published in: Nanobiotechnology. Edited by Christof M. Niemeyer and Chad A. Mirkin. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30658-9

1 Introduction

Biomolecules on surfaces have applications that range from medical diagnostics, analytical chemistry, and culturing and studying cells on surfaces, to synthesizing or engineering DNA, carbohydrates, polypeptides, or proteins. Defining patterns of biomolecules – and of proteins in particular – on surfaces is no simple task considering how complex and fragile these molecules can be. Photolithography – the art of structuring surfaces at lateral scales of less than 1 micrometer – was used recently to create DNA microarrays. Photolithography affords the capability of synthesizing strands of DNA using lithographic masks and photochemistry, but it is unlikely that a similar approach would permit the fabrication of arrays of proteins, which cannot be synthesized block-by-block at present. In photolithography, UV light, organic solvents, photoresists, and resist developers can compromise the structure and function of even a simple protein. For these reasons, novel approaches to patterning proteins include defining regions on surfaces that attract, bind, or repel proteins from solution, or in a more direct manner, by delivering small volumes of a solution of protein to a surface using drop-on-demand systems or microfluidic devices [1–3]. This chapter describes the patterning of proteins on surfaces by means of microcontact printing (µCP), where proteins are applied like ink to the surface of a stamp and transferred to a substrate by printing. Microcontact printing was originally developed by Whitesides and coworkers at Harvard for printing alkanethiols on gold with spatial control [4]. Many variants of µCP were later developed and collectively termed “soft lithography” [5]. The central element in µCP is the stamp. This is a silicone-based elastomer that is microstructured by curing liquid prepolymers of poly(dimethylsiloxane) (PDMS) Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Microcontact Printing of Proteins

on a lithographically fabricated master (or mold). Once cured, the stamp is peeled off the mold by hand; the stamp then bears an inverted pattern of that of the mold. One mold can be used to replicate many stamps. The relative softness of the stamp, compared to that of a lithographic mask, allows it to follow the contours of surfaces onto which it is applied. It is the work of adhesion between the stamp and the substrate that drives the spreading of the initial zones of contact at the expense of an elastic adaptation of the stamp [6]. In µCP – and soft lithography in general – the contact between the elastomer and a substrate occurs at the molecular scale and is termed “conformal”; it ensures the homogeneous transfer of ink from the stamp to the printed areas of the substrate [4]. PDMS materials have the following features. They are transparent to optical light and even UV down to ∼240 nm, resistant to many chemicals and pH environments, good electrical insulators, thermally stable, nontoxic, and can have tailored mechanical properties using various degrees of crosslinking and amounts of resin fillers [7]. A PDMS stamp can be a simple piece cut from a PDMS slab or accurately molded and mounted on a stiff backplane [8, 9]. It can be composed of PDMS layers having different mechanical characteristics, or shaped like a paint roller [10]. Handling stamps with tweezers and printing by hand is sufficient for most needs of experimentalists. Mounting a stamp on a printing tool, however, provides the ability to vary and control the pressure applied during printing, and to align the stamp with the substrate. PDMS stamps have advancing and receding contact angles with water of ∼115◦ and ∼95◦ and thus are hydrophobic and promote the spontaneous deposition of proteins from solution [11]. This deposition is nonspecific and self-limiting to a monolayer of proteins if the stamp is rinsed after the inking step. An important difference between microcontact printing proteins on surfaces and alkanethiols on noble metals is the limited amount of proteins on the stamp. Alkanethiols can diffuse inside a PDMS stamp in sufficient amounts for multiple prints, whereas reinking stamps with proteins is necessary after each print unless hydrogel “stampers” are used [12]. These stamps can carry a large reserve of protein solution used for the contact processing (CP) of substrates. PDMS stamps derivatized with biomolecules provide the basis for selective inking strategies and lead to the affinity-contact printing (αCP) technique [13]. Figure 1 delineates the operations that CP, µCP, and αCP involve. These techniques are described in detail in the following sections.

2 Strategies for Printing Proteins on Surfaces 2.1 Contact Processing with Hydrogel Stamps

Contact processing (left-hand panel in Figure 1) mimics the deposition of proteins from an aqueous environment to a surface by utilizing a hydrogel swollen with a solution of protein [12–14]. The proteins can diffuse through this hydrophilic matrix and adsorb onto the substrate without uncontrolled spreading. The stamp

2 Strategies for Printing Proteins on Surfaces

Fig. 1 Three related methods can pattern proteins from a stamp to a surface. In contact processing (left), a hydrogel stamp mediates the diffusion of proteins from its bulk to a surface. Microcontact printing (center) utilizes an elastomeric stamp inked with

proteins to print the proteins on a substrate without having a liquid. The stamp in affinity contact printing (right) is derivatized with capture proteins, which allows it to be selectively inked with target proteins released to a substrate during printing.

consists of two parts. The first is a reservoir above the hydrogel containing proteins dissolved in a biological buffer. The second is the hydrogel that makes contact with the substrate and mediates the transport of proteins to the substrate. A stamp that has a hydrogel made of poly(6-acryloyl-β-O-methyl-galactopyranoside), for example, embedded in a fine capillary can pattern proteins with a resolution of ∼20 µm [14]. Hydrogels having a refined composition and a greater degree of crosslinking exhibited better mechanical resistance, and were patterned by replication of a mold [15]. The latter approach should allow a protein to be patterned on a surface with a resolution better than 20 µm. CP based on hydrogel stamps has interesting features. First, biomolecules remain in a biological buffer until the stamp is removed and the substrate dried. Denaturation of proteins in this case should be minimal, and may be similar to that of proteins adsorbed from solution onto polystyrene microtiter plates. Second, it is straightforward to reuse such stamps for multiple CP experiments [14]. 2.2 Microcontact Printing

Microcontact printing of proteins uses PDMS stamps replicated from a mold (middle panel in Figure 1). Inking the stamp with proteins is simple, and analogous to depositing a layer of capture antibody (Ab) on polystyrene for conducting a solidphase immunoassay. The duration of inking and the concentration of protein in the

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ink solution determine the coverage of protein obtained on the stamp [16]. Inking a stamp can be local and/or involve multiple types of proteins when the stamp is locally exposed using a microfluidic network (µFN) or microcontainers to one or more solutions of protein [17]. The transfer of proteins can be remarkably homogeneous and effective, depending on the wetting properties of the substrate [18]. The large area of interaction of proteins with substrates and their high molecular weight account for the high-resolution potential of µCP of proteins. At the limit, single protein molecules can be printed as arrays on a surface [16], whereas the diffusion of alkanethiols on noble metals or the reactivity of silanes with themselves limit the practical resolution achieved for microcontact printing self-assembled monolayers on surfaces. Microcontact printing proteins on surfaces appears to be limited by the resolution and mechanical stability of the patterns on the stamp. Stamps made of Sylgard 184 and using masters prepared using rapid prototyping or photolithography can have micrometer-sized patterns on fields even larger than 10 cm2 [19]. Microcontact printing proteins with arbitrary patterns and submicrometer resolution benefits from the use of a PDMS elastomer stiffer than Sylgard 184 and masters patterned using electron-beam lithography [8]. 2.3 Affinity-Contact Printing

Tailoring the surface chemistry of stamps to ink a particular type of biomolecule is crucial for αCP (right-hand panel in Figure 1). The chemical stability of silicone elastomers is both an advantage for preparing chemically resistant stamps and an obstacle to modifying the surface of PDMS stamps. Exposing PDMS to an oxygen-based plasma forms a glassy silica-like surface layer [20]. The oxidized layer is a few nanometers thick and contains silanol groups ( Si OH), which are useful for anchoring organosilanes [21]. Oxidized PDMS can thus be derivatized similarly to glass or SiO2 in a few chemical steps using silane monolayers and with crosslinkers for proteins [22]. Affinity-contact printing is the technique of covalently immobilizing ligand biomolecules onto a PDMS stamp, and using them to ink a stamp selectively with receptor molecules. A stamp for αCP is roughly analogous to a chromatography column due to its ability to extract proteins selectively from a mixture, although releasing them involves printing them onto a surface [13]. Biomolecules that are naturally present in crude solutions and have a function on a surface are ideal candidates for applications of αCP. Cell adhesion molecules is one example that has already been demonstrated, but αCP could well be extended to a large variety of biomolecules for which ligands exist. Stamps in αCP are reusable and may include sites of different affinity to capture and print multiple types of protein in parallel [23]. 3 Microcontact Printing Polypeptides and Proteins

Many different types of proteins can be inked from an aqueous solution onto a hydrophobic silicon rubber such as PDMS [24]. Hydrophobic polymers in general

3 Microcontact Printing Polypeptides and Proteins

promote the deposition of proteins from solution through a variety of interactions, and slight or pronounced conformational changes of the protein structure can accompany this adsorption process. The kinetics of formation of a layer of protein on hydrophobic surfaces is often compared to a Langmuir-type isotherm: the rate of deposition of the protein molecules scales with their concentration in the bulk of the solution and reaches a plateau when all sites on the substrate become occupied [25]. Hydrophobic substrates, as a general rule, have stronger interactions with hydrophobic proteins, and their adsorption process is less influenced by the pH and ionic strength of the solution and by the isoelectric point of the protein than when polar or charged substrates are employed [25]. The size of the protein does not seem to play an important role on their inking behavior. A wide range of proteins in terms of structure and functions has been microcontact printed, which includes cytochrome c (12.5 kDa) [11], streptavidin and bovine serum albumin (BSA; ∼60 kDa) [26–28], protein A and immunoglobulins G (150 kDa) [11], glucose oxidase (160 kDa) [29], laminin (∼210 kDa) [30], and fibronectin (440 kDa) [31]. It is sometimes necessary to employ stamps with a hydrophilic surface to ink hydrophilic polypeptides such as polylysine (with MW ranging from 38 to 135 kDa) [32] or lipid bilayers [33]. In other cases, small biomolecules such as amino-derivatized biotins were inked and printed onto surfaces reactive to amino groups [34, 35]. In general, the derivatization of biomolecules with thiol groups allows the printing of biomolecules on gold substrates [36], where patterning by printing can be complemented by the adsorption of other types of molecules from solution. The chemisorption of small biomolecules on surfaces might be necessary for efficient transfer from the stamp and to prevent rinsing the printing molecules during subsequent steps. 3.1 Printing One Type of Biomolecule

Immunoglobulins G (IgGs) are interesting candidate molecules for µCP: these Abs are useful on surfaces for heterogeneous immunoassays. Their numerous disulfide linkages make them robust, they adsorb from biological buffers to PDMS in a nonreversible manner [24], and they can be conjugated to fluorescent centers, metal particles, enzymes, or ligands such as biotin. Fluorescence microscopy is a versatile method to follow the results of microcontact printing IgGs onto a glass surface (Figure 2). There, TRITC-labeled anti-chicken Abs were inked everywhere on the stamp but transferred to glass only in the regions of contact [11]. The patterns on the glass are accurate and correspond to zones of the stamp where the inked proteins are missing. The contrast of the 1 µm-wide features in the pattern is high and accurate and, as no fluorescence above background is measured in the nonprinted regions, it is clear that no transfer of Ab occurred in the recessed areas of the stamp. This might not always be the case, because small features have limited mechanical stability [6]. Demolding the stamp from the mold, capillary effects during inking and drying the stamp, and the printing itself may compromise the mechanical stability of patterns [37]. Implementing support structures in the design of the pattern, controlling the forces exerted during printing and affixing a

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Fig. 2 Microcontact printing proteins on glass. Fluorescence microscopy images revealing TRITC-labeled chicken Abs on a stamp after inking and accurately transferred in the regions of contact to a glass substrate. Reproduced with permission from Ref [11]. (Copyright 1998 American Chemical Society.)

stiff backplane to the stamp improve the stability of patterns. Stamps can be very large and have features measuring from micrometers to centimeters, making it possible to print proteins of one kind on large substrates to pattern cells indirectly. Examples include microcontact printing fibronectin [31], polylysine [30, 38, 39], laminin [40], and adhesion peptides [41]. 3.2 Substrates

Substrates for biomolecules cover a wide range of materials, from simple glass slides to complex functional microelectronic devices or sensors. Having conformal contact between the stamp and substrate during printing is the first requirement for microcontact printing biomolecules. For this reason, the substrate should not be too rough [6], or have too prominent structures [39]. Polystyrene, poly(styrene terephthalate), glass, amphiphilic comb polymers, Si wafers, and substrates covered with a thin evaporated metal and/or a self-assembled monolayer can be microcontact printed with proteins and stamps made of Sylgard 184 [11, 29, 34, 42]. The printing time does not seem to play a role, and takes the few seconds necessary to propagate the initial contact to the entire substrate. The details of how and why proteins transfer from a stamp to a surface were intriguing until the recent discovery that the difference in wettability by water between the stamp and the surface determines whether transfer occurs [18]. Proteins tend to transfer when the substrate is more wettable, or has a higher work of adhesion for water, than the stamp. In this respect, the chemical composition of the surface does not seem to play a particular role other than defining the wettability of the surface (Figure 3). The surface of the stamp can be derivatized with fluorinated silanes to raise the wettability threshold


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of the substrate below which transfer remains effective, for example. A remarkable incidence of printing proteins occurs with poly(ethylene glycol) (PEG)-derivatized surfaces [18]. Surfaces covered with a sufficient density of PEGs resist the deposition of proteins from solution because in order to interact with the PEG layer, proteins have to remove water solvat-ing EG repeat units and reduce the number of possible conformations of the PEG chains [43]. Both of these requirements are energetically unfavorable to the deposition of proteins from solution onto PEG-treated surfaces [44]. The mechanism accounting for the transfer of protein in µCP might thus involve the dry state of the PEG layer during printing [43], the local pressure exerted by the stamp at the line front propagation of the conformal contact [45], or some contamination of the PEG layer by low-molecular-weight silicone residues from the stamp. The deposition of proteins from solution or by printing exhibits antagonistic behaviors: proteins are more difficult to print on a hydrophobic surface than on a hydrophilic one whereas the opposite situation generally occurs in solution with, as an extreme case, PEG surfaces, which are protein-repellant [46]. Printing proteins is not limited to the patterning of planar substrates but is possible on curved surfaces, structured surfaces, and over large areas [5]. A stamp can be molded directly curved or planar and then curved and rolled over a surface [10]. Large stamps (≥ 10 cm) can be molded with a pattern having an accuracy of better than 1 mm [47]. The mechanical properties of stamps can be varied from 1 MPa (Young’s modulus) to over 30 MPa by adjusting the formulation of the polymer with respect to its average molecular weight between junctions, the junction functionality, and the density and size of filler particles added to the polymer [8]. The hardness of a stamp, its work of adhesion with the substrate, the pressure applied during printing, and the topography and work adhesion of the substrate all determine whether conformal contact will occur. The stability of features on the stamp might be compromised, however, when the stamp is made too soft and pressed too hard during printing [6, 9, 48]. 3.3 Resolution and Contrast of the Patterns

High resolution in lithography refers to patterning features of arbitrary shape at a length scale where it becomes crucial to optimize all parameters of the technique ← Fig. 3 Influence of the wettability of the substrate by water on the degree of transfer of proteins from a PDMS stamp to a Au surface. (A) The substrate is derivatized with SAMs comprising variable mole fractions of two constituents having different end-groups. (B) Proteins are adsorbed from solution onto the stamp. (C) The resulting printed patterns are analyzed using fluorescence microscopy. The fluorescence micrograph corresponds to fluorescently la-

beled proteins printed onto a 100% COOHterminated SAM. The transfer of proteins followed on SAMs having hydrophilic components functionalized with (D) COOH, (E) OH, or (F) EG correlates with the wettability of the mixed SAM (G). Figure kindly provided by J. L. Tan, J. Tien and C.S. Chen, and reprinted with permission from Ref. [18]. (Copyright 2002 American Chemical Society.)


(e. g., conditions for exposing and developing the resist, transfer of the resist pattern onto the substrate). Electron-beam lithography has a high-resolution regime for making features < 100 nm, photolithography for features < 250 nm, and µCP for features < 500 nm. In conventional lithography, shrinking the dimensions of patterns is driven by the necessity to improve the performance of integrated circuits at invariant or lower cost. The resolution of lithographic techniques limits the smallest sizes of components made today. It will be the physics of tomorrow’s devices that will ultimately be the limiting obstacle to further integration. Patterning biomolecules has a different paradigm for the resolution limit than conventional lithography because single functional elements, an enzyme for example, are available but do not have to be constructed. Microcontact printing meets several requirements that are necessary to place single proteins at predefined positions on a surface: (i) it is possible to fabricate Si molds with features as small as 40 nm using electron-beam lithography [16]; (ii) PDMS-replicated structures can be 80 nm and even smaller [9, 47]; (iii) proteins remain in the areas of contact, unlike alkanethiols and monolayerforming molecules which generally diffuse away from the initial printed zones on the substrate when an excess ink is present on the stamp; and (iv) the solution of protein used to ink the stamp can be diluted to limit the number of proteins inked per feature on the stamp [16]. Figure 4 shows high-resolution patterns of Abs on Si and glass and how a highresolution stamp can look. Each feature in the atomic-force microscopy (AFM) image in Figure 4A comprises ∼1000 Abs of the same type that were printed on a Si wafer using a PDMS stamp made of Sylgard 184 [17]. The structures have a width of 500 nm and an edge resolution better than 50 nm. The contrast of the patterns seems perfect because no Ab is present outside of the printed areas. An excellent contrast, together with specific binding events between printed ligands and receptors from solution, are desirable for high-sensitivity biological assays. The photography in Figure 4B shows a 8 × 4 cm2 stamp composed of a 30 µm-thick layer of PDMS attached to a flexible glass backplane 100 µm thick [8]. The PDMS layer of this stamp has numerous fields with 250 nm-wide lines, is about five times harder than Sylgard 184, but is also more brittle. The patterns are consequently more stable against collapse, and the glass backplane contributes significantly to the long-range accuracy of the pattern while making the stamp simple to handle, mount and align on a printer tool [47]. The surface tension of the polymer is an important limiting factor for the resolution of µCP. Features as small as 5 nm can be written by electron-beam lithography in PMMA and developed. PMMA is brittle, however, and hence not soft enough to form a reliable contact over surfaces. Unmolding even relatively hard PDMS stamps (having a Young’s modulus > 12 MPa) from a master structured with 40 nm-wide lines results in their broadening by 20 nm owing to surface tension effects [16]. Such a broadening is relatively less important for stamps with features ≥ 100 µm, and can be compensated in the electron-beam lithography layout. The stability of small features on stamps limits the freedom of design for high-resolution patterns. Dots, lines, and meshes do not have all the same mechanical stability against pressure; some recessed areas may collapse during printing. Incorporating support structures with micrometer dimensions around the high-resolution fields can remedy these problems [6]. Another strategy

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Fig. 4 Microcontact printing proteins at high resolution. (A) AFM images showing that only ∼1000 chicken Abs were printed onto a Si wafer in each element of this pattern. (B) High-resolution :CP is best done using stamps harder than Sylgard 184 and

having a flexible glass backplane. (C) AFM images showing rabbit Abs printed on glass as a mesh comprising 100 nm-wide lines (left) or on regularly spaced areas that have none, one or a few Ab molecules (right).

4 Activity of Printed Biomolecules

is to transfer the resist pattern into the Si master with a reactive ion etching, where the etch rates depend on the geometry of the features. When large structures are made deeper in the master than smaller structures, a larger part of the load during printing is exerted on the larger structures without inducing collapse of the smallest features [16]. The AFM images in Figure 4C correspond to Ab molecules microcontact printed from a PDMS stamp (material B) [8] onto glass using a mesh of 100-nm-wide lines (left image) and 100-nm hemispherical posts [16]. Both the posts and the lines were 60 nm high. The detail of a mesh reveals that two to four Ab molecules define the width of the lines. In the case of posts, one to three Ab molecules occupy each visible printed site, and the statistical analysis of larger printed zones revealed that sites could have none, one or a few printed Ab molecules [16]. There, the resolution limit for microcontact printing a single molecule is reached while still leaving space for improvement to form homogeneous arrays having only one biomolecule per site. A high concentration of protein in the ink, a long inking time, a further reduction of the dimensions of the posts, and a substrate with a high work of adhesion could help printing large arrays of single protein.

4 Activity of Printed Biomolecules

Many studies emphasize that while the adsorption of a protein on a surface is simple to perform, it is nevertheless a complicated phenomenon in which the biological activity of the immobilized biomolecule might be lost or significantly altered [25]. Microcontact printing biomolecules harbors this risk twice: when proteins are inked onto the stamp, and when they are printed. In principle, the deposition of proteins from solution onto PDMS should be analogous to that of proteins on hydrophobic surfaces [24]. The second concern is more difficult to weigh. Transferring a protein by printing implies that the adhesion of the protein with the substrate overcomes that of the protein with the stamp. At the limit, this might create a mechanical stress on the protein and could lead to irreversible conformational changes. It might be interesting to evaluate the yield of transfer as a function of the peeling rate to better characterize the transfer mechanism [49]. Comparing the activity of different types of biomolecules printed or adsorbed onto polystyrene suggests that enzymes are more susceptible to denaturation during printing than during adsorption from solution [11]. A layer of printed proteinase K displayed half the activity of a layer deposited from solution. Abs are more robust against loss of function; the ability of printed polyclonal Abs to capture antigens from solution was similar to that where the captured Abs were adsorbed. Printed monoclonal Abs seemed to have a ∼10% loss of capture efficiency compared to ones adsorbed from solution. The surface activity of microcontact printed biomolecules belonging to three important biological classes is illustrated in Figure 5. Cell adhesion molecules are ideal candidates for printing biomolecules because these molecules are useful on surfaces as they can direct the adhesion and growth of cells to specific regions of a substrate. Moreover, many are “simple” polypeptides. Polylysine [30, 32, 38, 39],

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Fig. 5 Microcontact printed proteins preserve a sufficiently high degree of activity for (A) promoting cell adhesion, (B) performing immunoassays, and (C) performing surface enzymatic catalysis. (A) The adhesion peptide PA22-2, which was printed onto a thiol-reactive surface, was immunostained (left-hand fluorescence micrograph) in two steps using an anti-PA22-2 Ab and a secondary Ab labeled with fluorescein, and is useful for attaching hippocampal neurons with spatial control (phase-contrast micrograph on the right). (B) These AFM images

illustrate three steps of an immunoassay in which chicken Abs were printed on a Si wafer (left), BSA was adsorbed from solution during the blocking step (middle), and the printed Abs were recognized by anti-chicken Abs. (C) This fluorescence image shows the deposition from solution of a fluorescent product made insoluble by printed alkaline phosphatase. The images ¨ in (A) were kindly provided by Offenhausser et al. and reprinted from Ref. [41]. (Copyright 2000 with permission from Elsevier Science.)

laminin [50], polylysine fused with laminin [40], fibronectin [31], specific adhesion peptides [41, 42], and neuron-glial cell adhesion molecules [13] (NgCAM), for example, were microcontact printed to promote or guide the attachment of cells to surfaces. In other examples, Ab–cell interactions were used to pattern cells on patterns of printed Abs [51, 52]. In some instances, the stamp was made hydrophilic and the substrate activated with a crosslinker to increase, respectively, the inking

5 Printing Multiple Types of Proteins

and transfer efficiency. The left-hand fluorescence image in Figure 5A corresponds to the immunostaining of the adhesion peptide PA22-2 that was microcontact printed onto a glass surface activated with amine-reactive crosslinkers [41]. The phase-contrast image (right) shows that the printed pattern of peptide was suitable to grow viable hippocampal neurons in the printed regions specifically These results illustrate well the conservation of the function of printed adhesion molecules. The capability of printing a pattern in registry with structures predefined on a substrate opens the way to placing cells wherever desired on a complex surface to study their function and to form networks of immobilized cells [39, 50, 53]. The next example (Figure 5B) is a printed polyclonal Ab, which serves as antigen to bind polyclonal Abs from solution [11]. AFM reveals the printed regions of a Si surface, each of which comprises ∼1000 molecules of chicken Ab molecules. Blocking the free Si surface with BSA is the next step necessary to prevent nonspecific deposition of proteins during the recognition step. After the blocking step, the Si surface is either covered with BSA or printed Abs; the prints are no longer visible. Recognition of the printed chicken Abs by anti-chicken Abs faithfully reflects the printed pattern. This experiment is an example of a highly miniaturized surface immunoassay in which the printed antigens were recognized by their specific Abs. Enzymes are probably more fragile than Abs and suffer more from a random orientation on a surface with respect to their activity than antigens for polyclonal Abs. The activity of printed enzymes can be evaluated using flat stamps, polystyrene substrates and colorimetric measurements [11]. It can be useful, however, to keep enzymatic products near their sites of production and to assess the activity of the enzymes with high spatial resolution. This is possible by using precipitating fluorescent products that accumulate in the regions of the substrate having enzymatic activity (Figure 5C) [17]. Real-time analysis of the development of fluorescence on the printed sites is even possible with this method. At least a part of the alkaline phosphatases printed on the glass surface in Figure 5C are active. This suggests that enzyme-linked immunosorbent assays can be performed using captured Abs that are printed. Interestingly, the same type of reporter enzyme can unambiguously reveal an ensemble of binding events, which are discernible through their localization. The challenge in this case remains to derivatize a surface with several types of protein.

5 Printing Multiple Types of Proteins 5.1 Additive and Subtractive Printing

An obvious application for microcontact printing proteins is the preparation of protein microarrays that can be used to screen different analytes in parallel while conserving reagents and still obtaining high-quality signals [54–56]. The simplest method to place two types of protein on a surface is to print one and adsorb the other from solution, as has been done with two different types of Abs [11]. This strategy requires that the printed layer of protein be complete enough to limit the

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adsorption of Abs from solution into the printed regions. The fabrication of arrays comprising n types of protein is possible using additive patterning steps: once a substrate is microcontact-printed, more proteins can be printed next to or over the previously printed ones (Figure 6A) [17, 57]. The transfer of proteins from a hydrophobic stamp to a more wettable substrate accounts for this finding because surfaces covered with printed proteins are more hydrophilic than PDMS stamps. This ability to stack proteins on top of each other is peculiar to µCP and might be useful for constructing protein-based architectures. It is also possible to place a variety of proteins on a substrate without the need for precise alignment during printing. Stamps with parallel lines can, for example, be inked and printed with a rotation between each print [17, 57]. Subtractive approaches are also possible (Figure 6B). One strategy is to ink a flat stamp homogeneously and to remove a subset of the proteins by printing them onto a structured surface. The sites on the stamp made free for adsorption are then covered by proteins from solution and the operation can be repeated [17]. An original way to pattern a surface with multiple types of protein is to fabricate a three-dimensional stamp and ink the different layers of the stamp with different types of proteins. Applying increasing pressure to the stamp brings each layer of the stamp successively in contact with the substrate [58]. The different patterns of protein are inherently aligned, and the difficulty of fabricating and inking the stamp might be compensated for by the relatively simple printing operation. 5.2 Parallel Inking and Printing of Multiple Proteins

Serial methods are simple, but probably not suitable, to pattern substrates with a large number of different proteins. A parallel inking approach of a stamp using a µFN can solve the problem of inking a stamp with different types of proteins (Figure 6C) [17, 59]. With such a strategy, a µFN having an ensemble of independent flowing zones is placed on a flat PDMS stamp [60, 61]. Sealing the microchannels results from the conformal contact between the µFN and the stamp. When solutions of proteins are flushed through the microchannels, proteins deposit in the areas of the stamp exposed to the conduits. Filling a µFN can be done serially, or with an array of dispensing heads or tips. The deposition of protein on the PDMS might be as fast as a few seconds when it is not limited by the mass transport of proteins from solution or the depletion of proteins from the channels. There, inking the stamp with or without a µFN could involve pin spotting, drop-on-demand, or microinjection techniques. Prefilling individual wells of a structured surface and applying it to a PDMS surface is also suited to locally derivatize a stamp with different types of proteins [23]. 5.3 Affinity-Contact Printing

The inking of a stamp with a large number of different types of protein before each print can quickly become cumbersome when it is desirable to print a large series

5 Printing Multiple Types of Proteins

Fig. 6 Fluorescence microscopy images illustrating three strategies for printing several types of protein on a surface. (A) Two fluorescently labeled proteins were printed subsequently on a glass surface. Proteins on the stamp transferred during the second print both to glass and to the lines of proteins already patterned. (B) This pattern includes two fluorescently labeled protein and BSA printed simultaneously from a flat stamp. First, BSA was inked homogeneously by adsorption from solution onto

the stamp and patterned by subtractive printing. Proteins labeled with fluoroscein isothiocyanate (FITC) were then adsorbed in the regions complementary to the BSA pattern. This was repeated to remove BSA and the second protein along lines that were filled with TRITC-labeled protein adsorbed from solution. (C) A stamp was inked with different lines of proteins using independent channels of a :FN and printed onto a polystyrene surface in one step. Reproduced from Ref. [17].

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of substrate with the same pattern of protein. The ability to attach biomolecules to PDMS covalently yields the opportunity to define zones on a stamp that can actively bind a target molecule from an ink. Crosslinking capture molecules (e.g., Abs or antigens) to a stamp allows inking the stamp by exposing it to a solution containing targets for the surface-bound capture molecules. This inking strategy, termed αCP, is analogous to capturing a protein on a column for affinity chromatography, although release of the captured species can occur during printing (Figure 7A) [13]. The idealized view of αCP is to have an affinity stamp (α-stamp) with multiple sites for capturing target molecules from solution in parallel. This could be used for many cycles of capture and printing. The fluorescence image in Figure 7B corresponds to the detection of fluorescently labeled Abs that were inked onto an α-stamp and printed on a glass substrate. This stamp had two types of binding sites (antigens) that were crosslinked to PDMS with spatial control by means of subtractive printing and ordinary printing [23]. The α-stamp used to print the Abs in Figure 7C was prepared by attaching capture antigens to a stamp activated with a crosslinker for proteins using a µFN. The yellow line corresponds to the printing of TRITC and FITC-labeled Abs that were simultaneously captured on a line of protein A immobilized on the stamp. Affinity contact printing, in particular when it employs stamps having multiple affinity sites, is both powerful and challenging. It is powerful because the ink can be a complex solution of biomolecules, the stamp is reusable, and patterns can have high resolution, as in µCP. The difficulty of αCP lies in the preparation of the α-stamp because the PDMS surface must be derivatized with crosslinkers and the quality of the patterns may degrade when preparation involves a large number of steps. The capture and release of radioactive or fluorescent proteins using αCP demonstrated the specificity of the capture event and the reusability of the α-stamp for at least 10 cycles [13]. Neuron-glial cell adhesion molecules (NgCAM), which are 200 kDa transmembrane proteins present at a concentration of ∼1 mg mL−1 in membrane homogenates of chicken brain, were captured by monoclonal Abs of an α-stamp and patterned on a polystyrene surface. The patterned surface appeared to be suitable for the attachment and growth of dorsal root glial neurons (Figure 7D and 7E), which was not the case where polystyrene was exposed directly to a nonpurified source of NgCAM [13]. Among all the printing methods reviewed here, αCP might be the most powerful method owing to the selective inking step and the reusability of α-stamps. It also has, in principle, the potential to print biomolecules with a defined orientation on a surface.

6 Methods 6.1 Molds and Stamps

Molds for preparing stamps are most often Si wafers patterned with a combination of photolithography and reactive ion etching. Reactive ion etching Si, instead of

6 Methods

Fig. 7 Affinity contact printing. (A) Stamps for "CP are prederivatized with capture sites and alternatively inked selectively and used for releasing the captured molecules on a substrate during the printing step. The fluorescence microscopy images correspond to (B) fluorescently labeled Abs co-captured and co-printed on glass using an "-stamp that had two types of capture sites; (C)

fluorescently labeled proteins captured on lines of an "-stamp and printed on glass; (D) the immunofluorescent detection of NgCAM that was patterned on polystyrene using an "-stamp decorated with lines of anti-NgCAM mAbs; and (E) the staining by immunofluorescence of neurons which attached to and developed on the printed pattern of NgCAM.

17


using directly the pattern of photoresist as the mold, prolongs the lifetime of molds. Si molds can be washed and cleaned; passivation of the Si surface is necessary before pouring PDMS with a release layer. Passivation can be done in situ after reactive ion etching or using a simple dessicator that can be evacuated. Typical release agents are fluorinated silanes. High-resolution molds are Si wafers patterned using electron-beam lithography. The density and resolution of the high-resolution features determine the price of these molds, which easily reaches $1000 per written cm2 for features having a critical dimension of 100 nm or less. Rapid prototyping is suitable for the preparation of stamps with a resolution of ∼25 µm [62] and only requires access to a high-resolution printer (5000 dpi). Replicating molds to make stamps and other operations such as inking stamps and printing are best done in a clean room or using a laminar flow bench to minimize the contamination of surfaces by dust particles. Sylgard 184 (Dow Corning) is used to prepare stamps in many cases, and comprises two prepolymers which, once mixed at a ratio of 1:10 (catalyst and hydridosiloxanes:vinyl-functionalized siloxanes), are poured on the master and cured at 60◦ C overnight. The formulation of harder, mechanically more stable PDMS is sometimes necessary when stamps have tall, isolated features [8]. Stamps can be cured on a backplane made of a thin steel plate or glass sheet [47], or on another PDMS layer [63]. Backplanes are typically flexible, but nevertheless give dimensional and long-range stability to stamps, which can then be mounted on a mask aligner, a printer, or handled by hand more conveniently. Design rules to make stamps are described jointly with the analytical description of the formation of conformal contact between stamps and substrates [6, 48]. This information is valuable to estimate the mechanical stability of stamps against pressure and to predict whether conformal contact will occur on rough surfaces or surfaces having topography. Hydrogel stamps are composed of a polymer that is crosslinked to the desired value (2–4%) and embedded in a microcapillary [12] or patterned by photocuring the hydrogel precursor sandwiched between a slide and a mold [15]. Both the inking and printing with hydrogel stamps rely on the diffusion of proteins through the hydrogel medium. 6.2 Surface Chemistry of Stamps

PDMS is hydrophobic and promotes the adsorption of proteins from solution in a manner analogous to polystyrene. Modifying the surface chemistry of stamps is necessary in two cases. Polypeptides and homogeneously polar biomolecules require stamps to have a hydrophilic surface for inking [38]. Affinity stamps must be derivatized with a ligand specific for biomolecule targets [13]. Exposing a PDMS stamp to an O2 -based plasma creates a silica-like layer on PDMS in a self-limiting manner. Stamps should be freshly inked (within ∼5 min) after the plasma treatment to prevent the recovery of their hydrophobic character [20, 22, 46]. This hydrophobic recovery originates from the migration of low-MW silicone residues from the bulk to

6 Methods 19

the surface. The plasma-induced scission of some polymer chains might also create mobile residues. It is impractical to extract these residues from the prepolymer components or after polymerization. Instead, plasma-treated stamps can be kept under water for long periods of time (more than days). Anchoring crosslinkers for proteins on plasma-treated stamps in one or more steps permits attaching covalently ligands onto the stamps [23]. Unreacted crosslinkers can be quenched with chemicals or reacted with noninterfering proteins such as BSA. 6.3 Inking Methods

The time to ink a stamp with a full monolayer of protein can be relatively long: up to 30 min at room temperature to obtain a monolayer of Ab using a concentration in phosphate-buffered saline (PBS) of 1 mg mL−1 , and 45 min with a solution of 5 µg mL−1 [11, 16]. The stamp is rinsed and dried after the inking step and then placed in contact with a substrate. The inking of hydrogel stamps with 1 mg mL−1 solutions of Abs in PBS takes similar times, and might even be faster if the gel is initially dry [12]. Shortening the inking time of PDMS stamps and localizing the inking is possible using µFNs [59]. In this case, the adsorption of proteins to the stamp might not be mass transport-limited, and is local in the regions of the stamp exposed to the channels. Localized inking is equally possible using microcontainers. These are small reservoirs microfabricated in Si, for example, and filled with the same, or different, solutions of proteins by hand or using pipetting robots [23]. Subtractive inking corresponds to inking entirely a flat PDMS stamp with proteins and transferring a subset by printing on a structured target. This strategy can remove proteins from areas of the stamp that become free for the inking of other proteins from solution [17, 23, 57]. This method can be repeated to form patterns with different types of protein next to each other but it requires an alignment step. Inking a stamp for αCP is analogous to linking a protein to a chromatography column via NH2 residues. Crosslinking protocols are usually well detailed by chemical suppliers or reviewed elsewhere [64]. 6.4 Treatments of Substrates

Surfaces more wettable by water than PDMS stamps are, in principle, suited for printing proteins [18]. Otherwise, they can be derivatized appropriately using plasma deposition techniques, oxidizing methods, or by grafting ultrathin organic layers. The wettability criteria may not apply when hydrophilic stamps are used to print certain polar biomolecules. In this case, derivatization of the substrate might also be necessary. Polylysine has been printed on glass directly [38] and on glass derivatized with glutaraldehyde [32], biotin on amine-reactive substrates [34, 35, 65, 66], and the adhesion peptide PA22-2 on a thiol-reactive surface [41]. In principle, crosslinkers can be attached to many types of substrates to bind proteins from stamps with high efficiency.


6.5 Printing

Handling a stamp with tweezers is the simplest approach to microcontact print proteins on surfaces both at low and high resolution. Typically, the stamp is brought close to the surface at an angle and set down gradually to ensure that conformal contact propagates from the initial contact areas without trapping air. Occasionally, (dust) particles or topography on the substrate are an obstacle to the propagating contact; applying gentle pressure to the stamp helps spread the contact to the rest of the substrate. Hybrid stamps are convenient to mount on printing tools [50] such as modified mask aligners, or on home-built printers using step motors to print substrates (up to 40 cm in lateral dimensions) with curved stamps while controlling the pressure applied to the stamp during printing [67]. Alignment of the stamp to preexisting structures on the substrate is desirable to print cell adhesion molecules on electrodes or to pattern substrates with multiple types of proteins, for example [39, 50]. Other methods already employed in soft lithography might be applicable for printing proteins. One example is printing surfaces and curved surfaces with cylindrical stamps [5, 10].

6.6 Characterization of the Printed Patterns

Surface-sensitive techniques such as ellipsometry, contact angle microscopy, and X-ray photoelectron spectroscopy can provide chemical information about surfaces printed with proteins. Patterns are best characterized using: (i) AFM, for which no labeling of the proteins is necessary; (ii) fluorescence and scanning confocal fluorescence microscopy; (iii) scanning electron microscopy, provided that the protein layer attenuates the emission of secondary electrons sufficiently to yield fair contrast [68]; or (iv) time-of-flight secondary ion mass spectroscopy [69]. AFM yields rich data concerning the contrast and resolution of the patterns, as well as the appearance and height of the printed layer, but suffers from the difficulty in localizing small printed areas [16]. Fluorescence microscopy is conversely effective in localizing signals from fluorescently labeled proteins, albeit with much less resolution than AFM. Colocalization of fluorophores having different spectral properties allows the detection of successively different types of proteins forming complex patterns. Optimization of the fluorescent signals is greatly facilitated by a` priori knowledge of the geometry of a printed pattern. Detection of unlabeled proteins is possible by immunoassays with detecting Abs either fluorescently labeled or conjugated with a reporter enzyme. In the latter case, the enzymatic conversion of chromogenic precursor into a precipitating fluorescent product keeps the fluorescent signal localized to the printed areas [17]. Staining using electroless deposition also keeps signals local [23]. Microcontact printing proteins onto diffraction gratings [51] or surfaces suited for plasmon resonance [26] offers the exciting capability of following binding events in real time and over sites in parallel.

7 Acknowledgments 21

7 Conclusion

The possibilities of microcontact printing biomolecules on a variety of surfaces with spatial control and resolution down to single protein molecules is unprecedented. Many fields could benefit in principle from these achievements. Surfaces could be decorated with high-quality patterns of microcontact-printed proteins for diagnostic applications. Very small volumes and quantities of reagents would be necessary for this purpose, using inking strategies based on microfluidic networks. In this case, numerous different types of protein could be patterned next to each other and used for surface ligand assays [70]. Printing proteins with tools that control the pressure during printing – hybrid stamps that are accurate and mechanically stable – and in alignment with predefined structures on the substrate has been demonstrated. The next steps are to build on these concepts, and mass fabricate high-quality arrays of proteins on surfaces such as glass slides or polystyrene surfaces for diagnostic applications. Stamps of various types can be devised. Some incorporate proteins in solution in a hydrogel-based reservoir, some have sites capable of binding target molecules from a complex ink, and others display numerous inked sites of micrometer lateral dimensions. Wherever biomolecules can be used on a surface, they might be placed advantageously by means of µCP. This is clearly the case for the growth of cells on surfaces, for which it becomes possible to construct hybrid architectures with cells connected to parts of electronic devices [53]. Positioning biomolecules on a biosensor surface is equally interesting. Microcontact printing can pattern areas of a sensing element with great precision and contrast, enabling the real-time monitoring of binding events on a surface. Possibly, the interaction between a large number of analytes and multiple printed sites could be screened for multianalyte immunoassays or drug screening. Microcontact printing is also well suited for preparing samples to investigate the biophysical properties of single biomolecules. Arrays of single biomolecules provide the advantage of having multiple sites to study each immobilized molecule with easy localization, without suffering from averaging effects, and with minimal signal degradation (photobleaching). It appears that although µCP was originally developed as a tool for applications in lithography [5], it is such a versatile technique that chemists and biologists have diverted it towards many more purposes. Microcontact printing clearly has unique, impressive features to manipulate and pattern biomolecules on surfaces, and it will be interesting to see how these will translate into firmly established applications for diagnostics, and biology in general.

Acknowledgments

I am very grateful to my colleagues Bruno Michel for his indefectible support of the work, to André Bernard and Jean Philippe Renault for having pioneered and carried out challenging experiments on microcontact printing proteins, and to Sergei Amontov, Helen Berney, Hans Biebuyck, Alexander Bietsch, Hans


Rudolf Bosshard, Isabelle Caelen, Dora Fitzli, Matthias Geissler, Bert Hecht, David Juncker, Max Kreiter, Heinz Schmid, Peter Sonderegger, Richard Stutz, Heiko Wolf, and Marc Wolf for their close collaboration on our biopatterning activities.

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1

Bacteriorhodopsin and Its Potential in Technical Applications Norbert Hampp

Philipps-Universität Marburg, Marburg, Germany

Dieter Oesterhelt Max-Planck Institute for Biochemistry, Planegg-Martinsried, Germany


1 Introduction

In biotechnology, the production of biological macromolecules for technical processes is state-of-the art. The biosynthetic capabilities of cells go far beyond those of organic chemistry. Materials such as functional biomolecules, enzymes, antibodies, and hormones are indispensable in the food industry, cleaning, medical diagnosis, pharmacy, and therapy, and consequently many companies supplying systems such as DNA chips and readers, and high-throughput screening platforms were established to meet demands. The biotechnology industry is booming – indeed, it is very likely to become the most important high-tech industry within the next few decades. Why is “bio” the coming technology? The advantages and the need for enduring miniaturizations are an increasing challenge as long as conventional lithographic methods are employed. The utilization of self-organization principles and bioengineering of functional biological structures seems to be a promising alternative approach. Re-engineering of biomolecules in order to realize technically desirable functions has become possible. However, there remains another problem – namely, the communication between classical microsystems (in particular electronic systems) and the nanoscaled biomolecules. This interface remains the major challenge in the realization of “cross-technology” products. Last – but not least – there is the problem of stability, as biomaterials are less stable than organic and semiconductor structures. This is of course a problem

Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Bacteriorhodopsin and Its Potential in Technical Applications

today, but in future technologies, where repair mechanisms like those in living organisms may be implemented, this obstacle may be overcome. Today, we have the need to seek methods for stabilizing the structures of biomaterials before they can be considered for technical applications. Bacteriorhodopsin has been studied over the past two decades as a material for technical applications. Its stability is adequate, it has several technically interesting functions, tools for both its modification and production in technical quantities have been developed, and it offers various interface principles, whether optical, electrical, or chemical. In the following sections, the activities for technical applications of bacteriorhodopsin will be reviewed and future developments will be discussed.

2 Overview: The Molecular Properties of Bacteriorhodopsin

In the first section of this chapter, the organisms in which bacteriorhodopsin is found are described, after which the processes used to modify this protein and to produce it in large quantities will be outlined. Finally, the current technical applications of bacteriorhodopsin will be summarized. 2.1 Haloarchaea and their Retinal Proteins

Archaea form, together with Bacteria and Eucarya, the three domains in life. Archaea are unicellular organisms thriving in a variety of habitats. Most of the archaea so far isolated and cultivated prefer extreme environments, although recent investigations have revealed archaea to be a standard component in biomasses of terrestrial and marine environments [1, 2]. Archaea are split into the crenarchaeotal and the euryarchaeotal branches. Members of the crenarchaeotal family comprise the full temperature range of life, from habitats at −1.8◦ C in the Antarctic to hyperthermal springs where Pyrolobus fumarii holds the world record in optimal temperatures for growth at 113◦ C [3]. Typical of the euryarchaeota are the methanogenic archaea. These occur ubiquitously in locations where organic matter is decomposed under strictly anaerobic conditions, such as in aquatic sediments, in marshes, or in the rumen of herbivores, and they contribute heavily to biogenic methane production. A second branch of the euroarchaeota comprises extreme halophilic organisms, the so-called halobacteria; these are ubiquitous on earth wherever salt occurs in solute concentrations close to saturation (Figure 1). Archaea and their proteins won commercial interest for the unusual physicochemical conditions under which they live and reproduce. The best example is bioleaching (biohydrometallurgy) of ore rubble, which was first carried out during the 1950s for copper and is now used to produce a variety of metals [4]. Heatstable enzymes are valuable molecular biological tools; examples include heat-stable catalysts for the polymerase chain reaction, or use in washing powders. Further

2 Overview: The Molecular Properties of Bacteriorhodopsin 3

Fig. 1 Halobacterium salinarum is found in nature in concentrated salt solutions as they occur in salines. A purple color is caused by bacteriorhodopsin, and this is the key protein of the photosynthetic capabilities

of H. salinarum. The proton pathway with the amino acids involved and the lysinebound retinylidene residue are shown in the structural model of bacteriorhodopsin (foreground).

applications of archaeal proteins which are halophilic or especially resistant to extreme pH values might become apparent in the near future. A high potential for applications in nanotechnology lies in the fabrication of devices on the basis of archaeal surface-layer proteins [5]. These readily form two-dimensional crystals, and actually occur on the cell surface as natural crystals with pores of precisely defined size. To date, six genera of halophilic archaea have been identified [6], and retinal proteins occur ubiquitously among these. In two genera, the extreme halophilicity is connected to a second extreme condition of life, alkalophilism, and these archaea are found in large masses in salty natron lakes. Halophilic archaea in general grow on organic substrates and have optimized their bioenergetics during evolution. With a sufficient supply of organic nutrients and oxygen, they respire in standard fashion. However, it must be noted that oxygen solubility in saturated salt solution is about five times lower than in water. Under the anaerobic conditions that commonly occur in their habitats, these organisms have acquired three alternative means for energy conversion: either they use nitrate, dimethylsulfoxide or trimethylaminoxide as end electron acceptors [7, 8] or they ferment arginine to carbamoyl phosphate for the production of ATP [9–11]. The third choice is a very powerful system, and the second route to photosynthesis in nature. This system does not use chlorophyll-based reaction centers (as do bacteria and plants) but rather


relies on retinal as a photon absorber and retinal-containing proteins as energy transducers [12]. The light-driven proton pump bacteriorhodopsin drives a proton circuit across the cell membrane, and ATP is produced via photophosphorylation as the chemical energy source for cell growth. The process is supported by the chloride pump halorhodopsin, which converts light energy via chloride transport into electrochemical energy used for the maintenance of osmotic balance. Halobacteria in nature grow very slowly, due to the limited supply of organic nutrients in hypersaline lakes. (The reader is referred to Refs. [7, 8] for a detailed description of their ecophysiology and the natural cycle of their “blooms” in salt lakes.) An example of such massive growth density is shown in Figure 1; an indication of heavy growth is the presence of an intense reddish to purple color. These blooms may occur only once each year (or even less frequently), but under optimization of nutrient supply in the laboratory generation times are usually in the order of hours. For biotechnological use, these two facts have to be considered: (i) a salt concentration of 25% is required to prepare peptone media, and this may be detrimental even to stainless steel fermentors; and (ii) peptone media are usually expensive to produce. These two obstacles are respectively overcome by using salt-resistant fermenting units and low-priced yeast extracts to replace the peptone. Another point to consider is the genetic stability of strains. Here, extensive progress has been made in the availability of stable standard strains used in biotechnology. In particular, strains which produce bacteriorhodopsin can be maintained stably by the use of phototrophic selection procedures [13]. Halophilic archaea, among them the genus Halobacteria, are unique in the sense that they are the only group of archaea that contains retinal proteins. While originally thought to occur only in higher animals capable of vision, retinal proteins recently have also been found in unicellular plant organisms [14], in fungi [15], in bacteria [16] and, most diverse in function, in halophilic archaea [12]. While all other groups seem to harbor either one of the two principal functions of retinal proteins – that is, a sensory function (eucaryotes) or a presumed energy-converting function (bacteria) – only halophilic archaea have developed a set of four retinal proteins, two of which serve a sensory function, while two convert light energy to chemical energy. Retinal, or vitamin-A aldehyde, originates from β-carotene by oxidative cleavage in the center of the molecule. The aldehyde in the free state is a chemically labile molecule with five conjugated double bonds. It is oxygen-sensitive and shows lightinduced isomerization around all double bonds. Light and oxygen together (photooxidation) destroy the free retinal easily. All known proteins containing retinal protect the molecule against photooxidation and select specific photoisomerization reactions, e. g., from 11-cis retinal to all-trans retinal in visual pigments and all-trans retinal to 13-cis retinal in the haloarchaeal proteins. All retinal proteins known are intrinsic membrane proteins and possess a transmembrane helical topography. Retinal always binds to the ε-amino group of a lysine residue of the seventh transmembrane helix, and a protonated Schiff base results; this becomes embedded in a cage of amino acids, which in turn drastically modifies the spectroscopic, chemical, and photochemical properties. For example visual pigments cover the color range


Fig. 2 Bioenergetics of Halobacterium salinarum. Bacteriorhodopsin acts as a light-driven, outward-directed proton pump. The generated proton gradient over the cell membrane drives a membrane-bound ATPase. These two proteins together form the simplest photosynthetic system known.

of the entire visible spectrum, and bacteriorhodopsin is chemically more stable than most proteins. Moreover, its light fastness is greater than that of organic dye molecules. Four molecular structures of retinal proteins are presently known: the visual pigment rhodopsin [17]; bacteriorhodopsin [18], halorhodopsin [19]; and sensory rhodopsin II [20]. Halobacterium salinarum, for example, makes extensive physiological use of retinal proteins (Figure 2). Bacteriorhodopsin drives the above-mentioned photosynthetic process for the production of ATP in light, and is also coupled to a highcapacity energy storage system in the form of a molar potassium gradient [21]. In order to maintain osmotic balance during growth – when a volume increase occurs under conditions of isoosmolar conditions, both inside and outside the cell – halorhodopsin is used as a light-driven anion pump [22]. This transports chloride ions into the cells, against the existing electrochemical potential, and allows a net salt accumulation to occur during the volume increase. This is a second way of avoiding the extensive use of respiratory energy at the expense of organic nutrients, by using light energy instead. Finally, two other retinal proteins occur in the cell which monitor the intensity and wavelength of the environmental illumination. These two photoreceptors – sensory rhodopsin I and II – receive orange light as an attractive stimulant, and blue light and near UV-light as repulsive stimulants (for a review, see Ref. [23]). Both photoreceptors signal to a two-component system of the cell, thereby regulating the frequency of the flagellar motor’s changes in rotational sense for directing halobacterial cells into an environment of optimal conditions. These photoreceptors are only two among a total of 18 receptors, all of


which convert chemical and physical signals of the environment to direct the cell into areas of optimal growth. The structures of all four halobacterial retinal proteins are very similar, and the molecular structures of two – bacteriorhodospin and halorhodopsin – have been elucidated to the atomic level. Moreover, based on details of the sequence alignment of the two ion pumps and the two sensors, together with the details of some two dozen other structures of archaeal retinal proteins, a tree has been created for this unique family of proteins [24]. Of particular interest in biophysical and biochemical terms is that the functions of the proton pump, the chloride pump, and the sensors are largely inter-convertible, either by varying the physical conditions under which the molecules operate, or by introducing minor genetic modifications. As an example, a point mutation in bacteriorhodopsin will convert this proton pump into a chloride pump [25, 26]. 2.2 Structure and Function of Bacteriorhodopsin

Bacteriorhodopsin is by far the best-studied archaeal retinal protein. It is naturally overproduced in the cells under conditions of illumination and limited aeration of a growing cell culture, but constitutive overproducers have been isolated. These cell lines produce up to 300 000 copies per cell, covering about 80 % of the cell surface as patches of two-dimensional natural crystals of bacteriorhodopsin with specific lipidic molecular species forming the so-called purple membrane [18]. Being a paradigm of light-driven proton pumps and seven-transmembrane helical proteins, the intense studies on this molecule over the past three decades by dozens of laboratories have produced a large body of knowledge on its biophysics, biochemistry, and molecular biology. Thus, the biotechnology of this “myoglobin” of membrane proteins is well founded on detailed knowledge about this molecule (for a recent review, see Ref. [27]). Like the other archaeal retinal proteins, bacteriorhodopsin is an intrinsic membrane protein with the common seven-transmembrane helix topology (see Figure 1) and an approximate molecular weight of 26 kDa. The seven helices are arranged in two arcs: an inner arc with helices B, C, and D; and an outer arc with helices E, F, G, and A. A transmembrane pore is formed mainly between helices B, C, F, and G. The retinal is bound to Lys216 in helix G as a protonated Schiff base, which interrupts the pore and separates an extracellular (EC) half channel (Figure 1, downward oriented) from a cytoplasmic (CP) half channel (Figure 1, upward oriented). The retinal side chain in the binding pocket is closely packed between four tryptophan residues, and the positively charged Schiff base interacts electrostatically with the protein environment. The chromophore is defined as the retinylidene moiety and the side chains in contact with it. Its color is tuned by electrostatic interactions specifically with the Schiff base and a complex counterion consisting of several amino acids. Light absorption causes photoisomerization of the retinal (all-trans to 13-cis and vice versa) and energy storage which includes ion-affinity shifts (H+


Fig. 3 Photocycle of bacteriorhodopsin. (A) Upon absorption of a photon, the initial $-state of bacteriorhodopsin is converted photochemically to the J-state from where a series of thermal steps leads back to the initial state. The proton transport is intimately coupled to the photocycle, which is observed as a sequence of intermediates which are represented by the common single-letter code with their absorption maxima given as subscripts. In the dark, bacteriorhodopsin relaxes thermally to the D-

state which has 13-cis configuration. The resulting mixture of $- and D-states is called dark-adapted bacteriorhodopsin. From the O-state, a photochemical conversion of alltrans to 9-cis retinal is possible which is not thermally reisomerized to the initial state. (B) The proton transport and related retinal configurations as well as accessibility of the nitrogen in the Schiff-base linkage between retinal and Lys216 are indicated. This sequence represents several of the molecular changes involved in the proton transport.

in bacteriorhodopsin and Cl− in halorhodopsin) of the Schiff base and internal binding sites as well as conformational changes as central elements of the catalytic cycle. This cycle may formally be represented as a sequence of six steps which are indispensable for transport: the all-trans to 13-cis photoisomerization and its thermal reversal (isomerization, I); a reversible change in accessibility (switch, S) of the Schiff base for ions in the EC and CP channel respectively; and ion transfer (T) reactions to and from the Schiff base active center (Figure 3). Specifically in


wild-type bacteriorhodopsin the order of these elementary steps is photoisomerization, proton transfer from the Schiff base to the acceptor aspartic acid (D) 85 in the EC channel, change in accessibility of the Schiff base from EC to CP, proton transfer from the donor aspartic acid (D) 96 in the CP channel to the Schiff base and thermal reisomerization, followed by reset of the Schiff base accessibility from CP to EC. The two proton transfer steps are intimately linked to cooperative changes in the EC and CP channels where hydrogen networks exist [28], and proton conduction over the total distance of about 48 Å is very likely based on a Grotthuss-type mechanism [29]. Structural key players in the EC channel are, besides D85, the arginine residue 82 and the two glutamic acid residues 194 and 204. They are connected by a water bridge to form a dyad which, by all likelihood, is the proton-release unit on the EC surface [18]. Other water molecules in the EC channel also play an important role [30, 31] in addition to amino acid side chains. One water molecule is located between D85 and the Schiff base, and disappears in the structure of the key intermediate M [32]. Three more water molecules below D85 are interconnected with side chains to form an extended hydrogen network as suggested by early experiments [33]. The CP channel from which the Schiff base later in the catalytic cycle receives the proton back is functionally dominated by the protonated D96. The distance between D96 and the nitrogen is 15 Å, and the hydrophobic nature of the space between them could provide an insulating layer against the membrane potential of 280 mV in the ground state. One water molecule occurs in contact with D96, and one water molecule is located between D96 and the Schiff base. Again, changes of this part of the molecule are observed in the key intermediate M [32]. At the proton entrance of the CP channel, aspartic acid D38 plays a role in the refeeding mechanism of protons [34]. The most dramatic effect of mutations on the structure and function of bacteriorhodopsin have the two carboxylates at positions 85 and 96. The removal of the proton acceptor D85 prevents the deprotonation of the Schiff base and almost completely annihilates the proton transport function (for a detailed description, see Ref. [35]). The lack of the proton donor D96 slows the catalytic cycle by a factor of up to several hundred, depending on temperature, humidity, azide concentration, etc. [36]. The reason is that the lack of the internal proton donor renders the rate of reprotonation of the Schiff base depending on external pH. Another important feature of the M-intermediate is its capacity to absorb blue light, and by this to reconvert photochemically into the initial state. Thus, in mutants lacking D96 the life-time of the M-intermediate and therefore the speed of color changes can be regulated either by physico-chemical parameters such as pH, temperature, and humidity, or alternatively by simultaneous application of two photons of different quality, for example green and blue photons. The three most important features, which eventually lead to biotechnological use of bacteriorhodopsin on the basis of its catalytic cycle are: 1. The color changes, which can be used for any type of information processing and storage process.


2. The photoelectric events which are due to the changing geometry of the Schiff base upon photoisomerization and the movement of the proton. Such electric changes occur from the picosecond to the millisecond time regime. 3. The pH change between the inside and the outside of bacteriorhodopsincontaining membrane systems as the net result of proton translocation.

So far, the color changes are under most intensive investigation because of the velocity of light-triggered reactions in bacteriorhodopsin and the possibility of regulating the speed of the color changes over a very wide range. 2.3 Genetic Modification of Bacteriorhodopsin

The molecular biology and genetics of halophilic archaea are developing at a slow, but constant, pace. The development of a transformation system was a major breakthrough, as this opened the door to site-specific mutagenesis and gene deletion or replacement [37, 38]. The basic process consists of a protoplast (spheroblast) preparation by incubation of cells with EDTA to remove magnesium ions. These are necessary for the integrity of the surface layer formed by the glycoprotein in the cell wall of archaea. Spheroblasts are incubated with a mixture of vector DNA carrying an antibiotic resistance and polyethyleneglycol. After a curing period in complex medium the cells are plated with antibiotic to select transformed clones [39]. Not many antibiotics have been reported as efficient agents against halophiles, and only two resistance genes have been cloned and inserted into suitable transformation vectors. Mevinolin inhibits the β-hydroxymethylglutaryl CoA (HMG) reductase and thus prevents isoprenoid synthesis of archaea. As lipids of halophilic archaea are diphytanoyl ethers of substituted glycerol, mevinolin completely blocks growth [40]. Novobiocin prevents DNA replication by inhibiting Gyrase B. The two selection marker genes used are hmg and gyrB; these were isolated from Haloferax volcanii and should have highly homologous genes in most other halophilic archaea [41]. The creation of site-specific retinal protein mutants requires appropriate host strains and vectors. The very stable strain H. (halobium) salinarum L33 [42] was introduced as a host for bacteriorhodopsin muteins [39]. It was found as a spontaneous mutation, carries an insertion element in the bop gene, and does not revert to a wild-type phenotype under any condition tested. Further strains were selected after either spontaneous or induced mutation which lack one or more of the retinal proteins, and many more strains with various phenotypes bearing favorable properties for investigations on the bioenergetics and signal transduction of halophiles have been reported. These are not covered in this chapter. More recently, systematic deletions of retinal protein genes have been described; one such example is the strain SNOB (S9 without bop) which is derived from the bacteriorhodopsinoverproducing strain S9 and has the bop gene deleted [43]. In consecutive rounds of a deletion procedure which uses the same antibiotic for selection repeatedly, all four retinal protein genes have been deleted to reach the strain NAOMI (now all


opsins missing; M. Otsuka and D. Oesterhelt, unpublished results). This strain has the advantage that complementation with opsin genes allows the combination of their physiological function at will. Vectors have been introduced first on the basis of a replicative element and a mevinolin-resistant determinant from a halophilic cell, together with a replicon and an ampicillin resistance from E. coli to serve as shuttle vectors [44]. In many laboratories, a plethora of vectors was subsequently designed with a size smaller than 10 kb, and without halobacterial origin of replication (suicide vectors) to enforce homologous recombination and therefore stable integration into the halobacterial genome. For interruption of a functional bop gene, the resistance gene used for selection is usually inserted into the coding sequence and remains stably associated with it permanently. Site-specific mutated genes can be favorably introduced into wildtype (S9) or mutated (L33) background in the following way. The resistance gene is placed next to the mutant bop gene on the suicide vector DNA. After transformation, antibiotic-resistant clones are selected which must result from a single crossover genetic event leading to the integration of vector DNA into the chromosome. These clones are allowed to grow without selection pressure, and are then replica plated with and without antibiotic. Among several genetic results is one which removes the vector DNA with the resistance gene and the original version of bop gene and leaves the mutated bop gene in the correct place stably integrated. These clones are found at various frequencies (usually one among several hundred) as the phenotype which does not grow on the antibiotic-containing plate but on its antibiotic-free counterpart. Besides genetic stability, the procedure provides the additional advantage of repeated use of the same antibiotic in further rounds of genetic alterations. Mutations of bacteriorhodopsin have been produced by the hundred and, once published, are available from the various laboratories. While the genetic background into which the mutated bop genes are introduced is often different, the mutagenesis procedure presently is usually the overlapping PCR method which replaced the preceding approach of gapped duplex DNA and is easily and quickly applied. Recently, a β-galactosidase gene as a reporter gene has been shown to act as an indicator gene for promoter strength and for blue-white selection procedures [45–47]. In conclusion, the production of bacteriorhodopsin muteins has become a routine method, but the maintenance of stable strains overproducing these muteins requires much care. In addition, the maximal level of mutein production can be very different, is unpredictable, and whether a given mutein will produce the crystalline arrangement of the purple membrane is also in doubt. 2.4 Biotechnological Production of Bacteriorhodopsins

The biotechnology of bacteriorhodopsin is based on simplicity of its isolation and chemical and photochemical stability when in the form of purple membranes. As

3 Overview: Technical Applications of Bacteriorhodopsin 11

mentioned, bacteriorhodopsin forms 2D crystals in vivo, and these can be isolated as purple membranes. The isolation is facilitated by two facts: (i) halobacterial cells are unstable in water and cell constituents are released upon lysis; (ii) the cell membrane is fragmented, and for unknown reasons the crystalline patches of the purple membrane are set free as fragments of largest size and highest buoyant density. These two specific features are used in the isolation procedure, either in a combination of sedimentation and isopycnic gradient centrifugation leading to a product of highest purity, or by a filtration procedure. The purification step yields a product which is 95–100% pure depending on the conditions and the mutein under consideration. Although these methods have not yet been exposed to economic competition, it is expected that the biotechnological production of purple membranes might in the future represent a competitive biomaterial in information technology. Certain limits of this material should be mentioned, however. As a membrane protein, bacteriorhodopsin cannot be produced via inclusion bodies as although the refolding and reconstitution of the active chromoprotein into membranes is possible in principle, it is certainly not feasible on an economic basis. Halobacterial cells do not form invaginations of their cell membrane like phototrophic bacteria (e. g., Rhodopseudomonas), which concentrate their reaction centers for photosynthesis in units called chromatophores. Although the purple membrane may occupy about 80 % of the cell membrane area, the amount of purple membranes obtained from cells is comparably low. It is needless to point out that the secretion of the integral membrane protein bacteriorhodopsin from cells into the medium has no biochemical basis. In practical terms, the current production of bacteriorhodopsin is at a level of 25 g m−3 nutrient broth. A 25-g quantity of bacteriorhodopsin would produce 50 L of suspension with an optical density of 1 (at a layer thickness of 1 cm this would allow the passage of only 10 % of light; i.e. it is an intense color). The molecular biology or production of bacteriorhodopsin variants has been established over the years, and at present the genetics is well known, including the sequence of the entire genome [48]. One problem in the molecular genetics of the halophilic archaea, especially H. salinarum, is that of insertion elements, as this sometimes causes instability of strains. However, with increasing knowledge on transformation procedures, vector construction and the creation of new antibiotics for the cells, these problems may be removed, at least for industrial purposes. It should be mentioned again, however, that cells producing bacteriorhodopsin variants do not always overproduce bacteriorhodopsin as do wild-type cells.

3 Overview: Technical Applications of Bacteriorhodopsin

The remarkable physico-chemical properties of bacteriorhodopsin and its numerous technically attractive molecular functions have led to many potential technical uses for this material [49–51]. In most of these applications, bacteriorhodopsin is used in the purple membrane (PM) form because its deliberation from the


crystalline package reduces both the chemical and thermodynamic stability of bacteriorhodopsin to a significant degree. Bacteriorhodopsin comprises three basic molecular functions which may be used in technical applications: photoelectric, photochromic, and proton transport properties. The molecular mechanisms responsible for each of these properties have been already discussed, and here we will focus briefly on the related applications.

3.1 Photoelectric Applications

The use of bacteriorhodopsin as a molecular level photoelectric conversion element is one of the fields where technical applications of the material have been examined. Upon illumination, a photovoltage up to 250 mV per single bacteriorhodopsin layer is generated, and this may be used either as an indicator or control element for various applications. Triggered by the absorption of a photon, the bacteriorhodopsin molecule undergoes a series of very rapid molecular changes, one of which is the generation of a photovoltage caused by changes in the orientation of molecular dipole moments that are triggered by the isomerization of the retinal on the femtosecond scale (Figure 4A). The proton released through the outer proton half channel may be either transferred to the outer medium or conducted along the surface of the PM [52]. The proton conductivity along the PM in H. salinarum cells supports delocalization of protons on the surface and proton conduction to the membrane-bound ATPase molecules. In most of the photoelectric applications of PM, the water content of the films is reduced, and this in turn causes proton conduction along the membrane surface. A single PM sheet contains several thousands of unidirectionally oriented BR molecules. Upon illumination, a number of bacteriorhodopsin molecules proportional to the intensity of light will be excited to accomplish a proton transport process. As all of the bacteriorhodopsin molecules in a single PM patch are oriented in the same direction, the voltage generated over a single membrane is independent of the number of active molecules (Figure 4B), but the proton current generated is proportional to the light intensity. The photovoltage generated can be easily measured by embedding the PM layer between two transparent electrodes. The light-triggered photovoltage induces a compensating polarization voltage in the outer electrodes, and this in turn may be detected as a high impedance voltage signal (Figure 4C). In a perfectly capacitive coupled system of this type, an induced photovoltage with the characteristics shown in Figure 4C is observed. A voltage is induced only during the light intensity change. The polarity of the signal is different for the OFF =⇒ ON and the ON =⇒ OFF transition. This is called the “differential responsivity” of PM layers. There are two major issues to be solved for an attractive use of bacteriorhodopsin in photoelectric processing. The first is that a high degree of orientation of the PMs is obtained because counter-oriented PMs cancel out each others’ photoelectric effects. The second is that a coupling of the light-dependent proton-motive force


Fig. 4 Photoelectric properties of bacteriorhodopsin. (A) Absorption of a photon by bacteriorhodopsin causes molecular changes which lead on a pico- to nanosecond timescale to the generation of a photovoltage over the molecule. The bacteriorhodopsin molecules in a single purple membrane (PM) patch are unidirectionally oriented. The light-driven vectorial proton transport of all the bacteriorhodopsin molecules is switched in parallel. This means that, over a single PM patch, it is not the voltage but the proton current which is proportional to the light intensity. The protons transported through the bacteriorhodopsin molecules can either be released to the outer medium or move along the surface of the PM patch due to proton conduction. (B) Oriented bacteriorhodopsin molecules, each represented by an arrow, sandwiched in a capacitor structure can be used as a photoelectric indicator cell. Upon illumination, a charge separation over the bacteriorhodopsin layer is generated which

induces a proportional charging of the outer electrode layers. No electric conduction between the bacteriorhodopsin layer and the electrodes is required. The electric field of the charge distribution induced in the electrodes compensates the electric field caused by bacteriorhodopsin. (C) Depending on the type of outer circuitry, either the induced voltage or the induced charge motion can be measured. In the latter case, a signal is recorded which corresponds to the first derivative of the temporal change of the light. (D) In a pixelated structure which is coated with oriented bacteriorhodopsin, photovoltages are measured only in those spots where a change in the light intensity occurs; hence, this is called novelty filtering. (E) In a volume (e. g., in 3-D data storage) the detection of the photovoltage in an outer capacitor structure was considered for readout. If the bacteriorhodopsin in the point of excitation is in the $-state, the absorption of a photon will lead to a photoinduced voltage, but not in the M-state.


of bacteriorhodopsin to the electromotive forces used in conventional electronics needs to be achieved. 3.1.1 Preparation of Oriented PM Layers The photoelectric signal of PM is high enough so that a single layer only of PM is needed for applications where the PM acts as a photoelectric indicator molecule. The Langmuir-Blodgett technique is often used for the preparation of such devices. The main problem is to obtain a high degree of orientation of the PM patches. The PM patches are thin (5 nm), large (up to micrometers), and flexible, and for this reason a mechanical orientation is difficult to obtain. The physical properties of the two sides of the PM are definitively different, but the differences are not so large that orientation (e. g., in an electric field) achieves more than a preorientation. Any preorientation obtained needs to be made permanent, for example, by covalent crosslinking or by polymer embedding of the PM patches. The only reliable method described so far is the coating of a surface with monoclonal antibodies against bacteriorhodopsin [53]. Since the antibodies selectively react with the cytoplasmic or extracellular side of the PM, a highly oriented monolayer of PM can be obtained using this method, although unfortunately it is restricted to a single PM layer. 3.1.2 Interfacing the Proton-Motive Force The other problem in photoelectric applications is that under light exposure bacteriorhodopsin transports protons, but not electrons. The interface between the biological component bacteriorhodopsin and its proton-motive force and the electromotive forces required for conventional electronics needs to be considered in the design of devices. Interfacing the proton-motive force of BR with the electron-conducting outer electrodes requires an electrolyte layer which couples both “worlds”. The balancing between the electronic circuitry and the photoelectric properties of a PM layer is crucial, and is the reason why results from different laboratories are so incomparable. The more than complete review published by Hong [54] is recommended for the reader who seeks much more detail on this subject. 3.1.3 Application Examples Ultrafast photodetection was one of the earliest proposals for the use of bacteriorhodopsin in a technical application [55, 56]. Two-dimensional photoelectric arrays, as artificial retinas or as control elements for a liquid crystal spatial light modulator, were developed later. Last, but not least, the photoelectric properties of bacteriorhodopsin can be used for indicator purposes in 3-D memories. 3.1.3.1 Artificial retinas Most devices which make use of the photoelectric properties of bacteriorhodopsin are called “artificial retinas” [57–59], the reason being that they offer certain preprocessing features known from the retina, including edge detection and novelty filtering. The physical background is called the “differential response” of PM layers. A device (see Figure 4D) comprising an electrode array which is covered with one or more oriented layers of PM may be used for


novelty filtering. Each of the electrodes is connected to an amplifier electronics. First, assume that the dark rectangular structure shown in Figure 4D prevents a set of electrodes from being exposed to light. Then, if this structure is moved over the light-sensitive sensor area (see arrow), it causes a voltage to be induced in each of the pixel electrodes, with a sign proportional to the light change. Because only those electrodes “fire” where a change of the illumination occurs, this type of sensor is called an “artificial retina”, and this type of preprocessing is called “novelty filtering”. Due to the differential response of bacteriorhodopsin, the polarity of the electrode signal carries the information whether a pixel was switched to “ON” or to “OFF”, and in turn the direction of the movement of the object can be derived. 3.1.3.2 Electro-optically controlled spatial light modulators Another use of the artificial retina described above may be in optically addressed spatial light modulators (SLMs). The amplifier electronics which detects the photovoltages induced in the electrode pixels may be connected to a SLM device. In particular, liquid crystal (LC)-based SLMs are state-of-the-art. In this case, the electrodes of the bacteriorhodopsin-based artificial retina are connected one-to-one to the pixels of a LC-SLM. An advanced version omits the wiring and amplifiers. The LC-layer of the SLM is controlled directly by the artificial retina device [60, 61]. 3.1.3.3 Readout in 3-D Memories Another application of the photoelectric properties of PM was investigated for the readout of volume storage units with bacteriorhodopsin. The basis is a cube of oriented PM patches in either the purple initial state or the yellow M state. Upon illumination of a PM patch in the initial state (which may be addressed by actinic light), a photovoltage signal is induced. A PM patch in the M state would not respond to the actinic light, but two electrodes on the outer surfaces of the bacteriorhodopsin cube could detect the photovoltage generated, and by this method the 3-D distribution of the photochemical states of PM patches could be read out. This is a prerequisite for a 3-D memory based on bacteriorhodopsin (Figure 4E). 3.2 Photochromic Applications

Most applications currently investigated utilize the photochromic properties of bacteriorhodopsin [50, 62]. During the photocycle, bacteriorhodopsin cycles through a pair of spectroscopically distinguishable intermediates (see Figure 3A), all of which have an absorption maximum which is different from that of the initial B state. However, in most applications the photochromism of bacteriorhodopsin is used in connection with the purple to yellow absorption change which is related to M state formation. Upon acidification, a blue membrane is formed which has a significantly different photocycle. The formation of a 9-cis retinal-containing state is observed, and this is thermally stable. In contrast, 13-cis retinal is isomerized by the bacteriorhodopsin molecule to all-trans retinal at room temperature, and the isomerization of 9-cis


Fig. 5 Photochromism of bacteriorhodopsin and its application. (A) The retinylidene residues are strongly anisotropic. (B) Upon illumination with polarized light, the retinylidene residues which are in parallel to the electric field vector of the actinic light are preferentially excited and isomerized. (C)

Transient photochromic change of bacteriorhodopsin between the initial purple state and the yellowish M-state (middle). (D) The photochemical formation of 9-cis retinal may be utilized for photochromic long-term storage. (E) Permanent storage of information in bacteriorhodopsin.

retinal is not catalyzed. This pathway opens the route to long-term storage materials based on bacteriorhodopsin. 3.2.1 Photochromic Properties of Bacteriorhodopsin Isomerization from all-trans to 13-cis is the first occurrence after the photochemical excitation of bacteriorhodopsin, and this causes significant transient shifts in the absorption spectrum. In addition to the isomerization change, deprotonation of the chromophoric group is observed. In the L to M transition, a proton from the Schiff base nitrogen group is transferred to Asp85, and this deprotonation causes a drastic blue shift of the absorption to 410 nm. The photochromism of bacteriorhodopsin is dominated by the intermediate which has the longest life-time; hence, this forms a “bottleneck” in the photocycle. The retinylidene residue inside bacteriorhodopsin which forms part of the photochromic group in the molecule is strongly anisotropic. A PM layer may be considered as a crystalline arrangement of chromophoric groups which are oriented angles of 120◦ between them (Figure 5A). However, in most applications where a statistic number of PM patches is used, the angular chromophore distribution appears anisotropic. Due to the anisotropy of the retinylidene groups, excitation of a random distribution of PM patches with polarized light causes the chromophores to become oriented in parallel to the actinic light polarization such that a


preferentially converted (and in turn photoinduced) anisotropy is obtained (Figure 5B). In solutions containing PM this is masked by diffusion, but in bacteriorhodopsin-films where the PM patches are fixed, the photo-induced anistropy can be easily observed and utilized. Today, three types of photochromic changes in bacteriorhodopsin have been described which enable different applications. The first is the photochromic shift between the B and M states (Figure 5C), and this is used mainly for optical processing tasks. The second is photoerasable data storage using 9-cis-containing states of blue membrane or suitably modified BR-variants (Figure 5D). However, the very low quantum efficiency for recording of far below 1 % is a major limitation. And last, but not least, permanent photochromic changes obtained through two-photon absorption in bacteriorhodopsin are suitable for long-term data storage (Figure 5E). 3.2.2 Preparation of Bacteriorhodopsin Films Optical films are prepared from bacteriorhodopsin by polymer embedding. Optically clear, water-soluble polymers are suitable for this purpose (e. g., polyvinylalcohol, gelatin). The film formation is usually carried out by mixing the polymers with PMs and additives in aqueous solution, this being cast on a glass support. The water is generally removed by drying in air, but the films may also be sealed with a second glass plate. 3.2.3 Interfacing the Photochromic Changes The reason why photochromic applications of bacteriorhodopsin are more developed than others is because of the ease with which an interface can be implemented between the bacteriorhodopsin films and any type of optical system. The bacteriorhodopsin film is completely sealed, the only interface being the light which transports energy and information simultaneously. 3.2.4 Application Examples Many applications based on the photochromism of bacteriorhodopsin have been suggested, and some are described briefly here as an indication of the wide range of potential uses. 3.2.4.1 Photochromic color classifier As the different rhodopsins in the eye enable color perception, the use of bacteriorhodopsins with different absorption maxima would allow a biomimetic system for color perception to be set up. The photoelectric response of three sensor elements coated with three different bacteriorhodopsin types (i. e., wild-type and two containing retinal analogues) are combined and coupled to a simple neural network for color recognition properties. This functions quite reliably, though it is unclear whether it has any technical advantages over conventional color sensors. Two limitations can be identified in this system. First, the absorption maxima of the three BR types used today are relatively similar, and they do not span the visual wavelength range as well as the human rhodopsins, notably in the blue region. Second, the conventional systems which typically comprise three different color filters and semiconductor light-sensitive elements reliably


supply the required information. At present, it is difficult to identify any advantages of the bacteriorhodopsin-based systems over conventional systems [63]. 3.2.4.2 Photochromic inks Another development is the preparation of photochromic inks. These inks differ from the polymer films for optical recording by their rheological properties. Depending on the method of application (e. g., screen printing, offset printing), the viscosity and surface tension must be considered. The basic photochromic properties are quite similar to those of the optical films, but auxiliaries in the compositions adjust the required application dependent properties. A major problem is to identify suitable compositions which do not interfere with the photochemical properties of the bacteriorhodopsin embedded [64]. 3.2.4.3 Electrochromic inks The main color shift in bacteriorhodopsin is due not to a primary photochemical reaction but to the protonation change of specific groups, in particular the Schiff base linkage and the Asp85 residue. As protons are charged particles, their removal from the binding position by electric fields should be possible. Indeed, this can be demonstrated, though the speed and efficiency of the decoloration/coloration process is quite low. Nonetheless, the basic principle was successfully demonstrated [65], indicating a potential development of electrochromic paper using bacteriorhodopsin. 3.2.4.4 Photochromic photographic film It is well known that a permanent bleaching of bacteriorhodopsin may be achieved with hydroxylamine. The chemical reaction of hydroxylamine with the retinal binding site occurs in an intermediate state only, and no reaction of BR in the β-state is observed. Due to this finding, which dates back to the early retinal extraction experiments, it is possible to fabricate nonreversible optical films from bacteriorhodopsin [66]. These films behave quite similarly to photographic films, except that the compounds needed for the chemical development are already contained in the film. The reason why this process has not been used technically is that, after image formation, the nonreacted hydroxylamine must be removed from the film, or fading of the contrast will occur when stored in light. 3.2.4.5 Long-term photorewriteable storage of information Much interest has centered on photorewriteable optical storage with bacteriorhodopsin. For this purpose, the conventional all-trans to 13-cis conversion is not suitable because of the retinal reisomerization at room temperature in bacteriorhodopsin. In blue membrane, a photochemical conversion from all-trans to 9-cis retinal, which appears pink in bacteriorhodopsin, may be induced by high light intensities. 9-cis retinal is thermally stable and requires photochemical excitation for reconversion of all-trans. The main disadvantage of blue membrane is that its formation from PM requires either acidification or the removal of divalent cations, as both cause destabilization of the bacteriorhodopsin molecule. Aggregation of the PM patches is also seen. The requirement is for bacteriorhodopsin variants which have a similar photochemistry but at ambient pH value and without removal of cations. BR variants with alterations


in position 85 show such desired properties, and one of the first reported for this purpose was D85N. Another approach is to use wild-type BR at neutral pH values and to switch the material photochemically to the 9-cis state from the so-called O-state [67]. This scheme was named branched photocycle memory [68]. 3.2.4.6 Neural networks Due to the fact that its absorption state may be shifted with blue and yellow light in different directions, bacteriorhodopsin is also a suitable material for neural networks. The output from an absorptive bacteriorhodopsin-cell is used to control the absorption state of another such cell. This has been demonstrated in principle [69], but the bacteriorhodopsin has a “fan-out” of much less than one and for this reason is not really a suitable material for the implementation of optical neural networks. (“Fan-out” is a term from electronics which characterizes the signal power output of a device compared to the required signal power input the same device requires.) A “fan-out” of 10 means that the output terminal of the device supplies enough power that 10 input terminals of identical devices could be supplied with enough energy to signal them the input state reliably. In photochromic devices where no amplification occurs, the “fan-out” is generally less than 1. Without external amplification, it is almost impossible to set up control loops. 3.2.4.7 3-D information storage The use of bacteriorhodopsin in 3-D information storage has been investigated for some time. Bacteriorhodopsin shows an astonishingly high two-photon absorption cross-section of the initial B state. This is used in a two-photon absorption setup to address the absorption state of bacteriorhodopsin in three dimensions [70]. The recording process is quite well handled, but the readout is an intrinsic problem. The advantage of such a memory device is its tolerance towards electromagnetic radiation. 3.2.4.8 Nonlinear optical filtering The strongly nonlinear optical response of bacteriorhodopsin towards the incident light intensity has given rise to many applications which use the nonlinear response for image processing purposes (e. g., edge enhancement, noise reduction). The response curve of the bacteriorhodopsin can be tuned over several orders of magnitude by changing the lifetime of the M-state. This may be accomplished by changing the pH, as well as using modified bacteriorhodopsins. 3.2.4.9 Holographic pattern recognition and interferometry The use of bacteriorhodopsin as a short-term memory has been tested in several applications. The most challenging was the construction of a real-time holographic pattern recognition system which operates at video frame rate. In this system, the holographic comparison of images allows similarities between objects used for identification purposes for complex images to be quantified [71, 72]. In the bacteriorhodopsinfilm, holograms are recorded, read-out and erased at video frame rate. In the early 1990s, when the system was first developed, it was unchallenged by computer systems, but as computing power has advanced the bacteriorhodopsin-film method has been abandoned.


Typical photochromic applications include a holographic real-time correlator (Figure 6A) and a holographic camera for nondestructive testing (Figure 6B). Examples of photochromic inks made from bacteriorhodopsin are shown in Figure 6C, with the initial colored (purple) and bleached, yellowish inks in the foreground and background, respectively. An ID card sample with a bacteriorhodopsin-based optical storage in the purple-colored strip is shown in Figure 6D. 3.3 Applications in Energy Conversion

The use of bacteriorhodopsin as a light energy-converting device seems to be first choice as with regard to technical applications of the molecule [73, 74]. All applications of this type have as the key element in common a bacteriorhodopsin-driven charge separation over the PM. Although this is the basic function of bacteriorhodopsin embedded in PM, until now it has not been possible to prepare artificial PMs with technically relevant dimensions. Methods devised to overcome this problem have been unable to provide PMs capable of producing passive proton transport, and research is continuing in this area.

4 Methods

The methods required for this type of research span from biochemistry and bioengineering to various printing and optical techniques. All have been well documented, with the methods of cultivating halobacteria and isolating PMs being common to all applications. A comprehensive laboratory manual on Archaea was produced in 1995 [40]. For specific conditions of phototrophic growth [13], laboratory protocols of cultivation and media composition, the reader is referred to Ref. [40] (pages 13–21 and 225–230), while for the isolation of PMs from halobacteria, the reader is referred to Ref. [75]. The genetic modification of halobacteria is described in Refs. [47, 76]. The very first publication on the discovery of the bacteriorhodopsin [77], as well as the latest volume on its applications [78], should also be mentioned in this context. Information on specific processing steps for the wide range of applications is taken best from the relevant patent applications, and a summary of patents related to applications of bacteriorhodopsin may be found in Ref. [51].

5 Conclusions

Bacteriorhodopsin is today the biological photochromic material for which technical applications in optical information processing are much more developed than for any other biomaterial. The entire sequence, from an analysis of molecular

5 Conclusions 21

Fig. 6 Photochromic applications of bacteriorhodopsin. (A) Holographic correlator; (B) holographic camera for interferometric testing; (C) photochromic inks for security applications; (D) optical storage.


function to its controlled modification and the development of suitable applications, has been demonstrated with this molecule. Ideas of utilizing the evolutionary optimized functions of biological molecules in technical processes by using genetic engineering to produce tailor-made modified versions of natural molecules with improved technical properties have been demonstrated with bacteriorhodopsin for the first time. Biomaterials as blueprints for technical materials with nanoscale functions form the basis of the concept of nanobionics – and bacteriorhodopsin was the first such example.

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1

Biomolecular Motors Operating in Engineered Environments Stefan Diez, Jonne H. Helenius, and Jonathon Howard Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany


1 Overview

Recent advances in understanding how biomolecular motors work has raised the possibility that they might find applications as nanomachines. For example, they could be used as molecule-sized robots that: Ĺ work in molecular factories where small, but intricate structures are made on tiny assembly lines; Ĺ construct networks of molecular conductors and transistors for use as electrical circuits; Ĺ or that continually patrol inside “adaptive” materials and repair them when necessary.

Thus, biomolecular motors could form the basis of bottom-up approaches for constructing, active structuring and maintenance at the nanometer scale. We will review the current status of the operation of biomolecular motors in engineered environments, and discuss possible strategies aimed at implementing them in nanotechnological applications. We cite reviews whenever possible for the biochemical and biophysical literature, and include primary references to the nanotechnological literature. Biomolecular motors are the active workhorses of cells [1]. They are complexes of two or more proteins that convert chemical energy – usually in the form of the high-energy phosphate bond of ATP – into directed motion. The most familiar motor is the protein myosin which moves along filaments, formed from the protein actin, to drive the contraction of muscle. In fact, all cells – not just specialized


2 Biomolecular Motors Operating in Engineered Environments

muscle cells – contain motors that move cellular components such as proteins, mitochondria, and chromosomes from one part of the cell to another. These motors include relatives of muscle myosin (that also move along actin filaments), as well as members of the kinesin and dynein families of proteins. The latter motors move along another type of filament called the microtubule. The reason that motors are necessary in cells is that diffusion is too slow to transport molecules efficiently from where they are made (which typically is near the nucleus) to where they are used (which is often at the periphery of the cell). For example, the passive diffusion of a small protein to the end of a 1 meter-long neuron would take approximately 1000 years, yet kinesin moves it in a week. This corresponds to a speed of 1–2 µm s−1 , which is typical for biomolecular motors [2]. Actin filaments and microtubules form a network of highways within cells, and localized cues are used to target specific cargoes to specific sites in the cell [3]. By using filaments and motors, cells build highly complex and active structures on the molecular (nanometer) scale. Little imagination is needed to envisage employing biomolecular motors to build molecular robots [4]. Biomolecular motors are unusual machines that do what no man-made machines do: they convert chemical energy to mechanical energy directly rather than via an intermediate such as heat or electrical energy. This is essential because the confinement of heat, for example, on the nanometer scale is not possible because of its high diffusivity in aqueous solutions [2]. As energy converters, biomolecular machines are highly efficient. The chemical energy available from the hydrolysis of ATP is 100 × 10−21 J = 100 pN nm−1 (under physiological conditions, where the ATP concentration is 1 mM and the concentrations of the products ADP and phosphate are 0.01 mM and 1 mM, respectively). With this energy, a kinesin molecule is able to perform an 8-nm step against a load of 6 pN [2]. The energy efficiency is therefore almost 50%. For the rotary motor F1 F0 -ATPase synthase which uses the electrochemical gradient across mitochondrial and bacterial membranes to generate ATP, the efficiency is reported to be between 80 and 100% [5, 6]. This high efficiency demonstrates that, like other biological systems, the operation of biological motors has been optimized through evolution. High efficiency is but one feature that makes biomolecular motors attractive for nanotechnological applications. Other features are: 1. They are small and can therefore operate in a highly parallel manner. 2. They are easy to produce and can be modified through genetic engineering. 3. They are extremely cheap. For example, 20 × 109 kinesin motors can be acquired for 1 US cent from commercial suppliers (1 mg = 3.3 × 1015 motors cost $1500; Cytoskeleton, Inc., Colorado, USA) and the price could be significantly decreased if production were scaled up. 4. A wide array of biochemical tools have been developed to manipulate these proteins outside the cell.

1 Overview

This review focuses on two broad categories of molecular motors: Ĺ Linear motors generate force as they move along intracellular filaments. In addition to myosin and kinesin mentioned above, linear motors also include enzymes that move along DNA and RNA. Ĺ Rotary motors generate torque via the rotation of a central core within a larger protein complex. They include ATP synthase, mentioned above, as well as the motor that drives bacterial motility.

Representatives of both categories have been used to manipulate molecules and nanoparticles. Mechanical and structural properties of relevant filaments are contained in Table 1, and those of several associated motors in Table 2. The general set-ups for studying motor proteins outside cells – the so-called motility assays – are depicted in Figure 1. In the gliding assay, the motors are immobilized on a surface and the filaments glide over the assembly (Figure 1A). In the stepping assay, the filaments are laid out on the surface where they form tracks for the motors to move along (Figure 1B). In both assays, movement is observed under the light microscope using fluorescence markers or highcontrast techniques. Variations on these assays have been used to reconstitute linear motility on the four types of filaments – actin filaments, microtubules, DNA, and RNA. The gliding motility assay has provided detailed data on the directionality, speed, and force generation of purified molecular motors [2, 7]. However, for use in nanotechnological applications, the movement of gliding filaments must be controllable in space and time. For example, a simple application would be to employ a moving filament to pick up cargo at point A, move it along a user-defined path to point B, and then release it. A number of methods for the spatial and temporal control of filament movement have been developed. Spatial control has been achieved using topographical features [8–11], chemical surface modifications [10, 12–14], and a combination of both [15–18]. Electrical fields [19–21] and hydrodynamic flow [22, 23] have also been used to direct the motion of gliding filaments. An example from our laboratory of gliding microtubules that are guided by channels is shown in Figure 2. Temporal control has been achieved by manipulating the ATP concentration [9, 24]. In addition to these basic techniques for controlling motion, some simple applications of the gliding assay have been demonstrated. These include the transport of streptavidin-coated beads [9], the transport and stretching of individual DNA molecules [25], the measurement of forces in the pN range [26], and the imaging of surfaces [27]. The stepping assay opens up additional possibilities. Initially, micrometer-sized beads were coated with motor proteins and visualized as they moved along filaments. The movement of beads can be tracked with nanometer precision to determine the speed and step size [2], and the use of optical tweezers allows forces to

3

13 2

25 nm 2 nm

2 nm

RNA

2

2

6 nm

Actin filament Microtubule DNA

Strands per filament

Diameter

Filament

0.34 nm

8 nm 0.34 nm

5.5 nm

Repeat length

75 nm

5 mm 50 nm

10 µm

Persistence length

1.5 GPa

2 GPa 1 GPa

2 GPa

Young’s modulus

30 µm

10 cm 100 mm

100 µm

Maximum length

Kinesin, Dynein RNA polymerase, DNA helicase, topoisomerase Ribosome

Myosin

Motors

80

78 79

77

Reference

where k is the Boltzmann constant and T is absolute temperature. Young’s modulus (E) is calculated assuming that the filament is homogenous and isotropic. The repeat length describes the periodicity along a strand of the filament

Table 1 Physical attributes of actin filaments, microtubules, DNA, and RNA. The persistence length (Lp) is related to the flexural rigidity (EI) by: Lp = EI/kT,


The sizes refer to the motor domains.

120◦

8 × 14 45

*

0.34 up to 43 nm/turn 0.34

15

DNA DNA DNA pilus NA NA

0.34

5 36 8

16 24 6 24

Actin Actin Microtubule Microtubule DNA

Myosin II Myosin V Conventional kinesin Dynein T7 DNA polymerase (exonuclease activity) RNA polymerase Topoisomerase Bacteriophage portal motor Type IV pilus retraction motor F1 -ATPase Flagellar motor

Step size [nm]

Filament

Motor

Size* [nm]

5 − 100 bps 1000 8 rps 300 rps

30 000 300 800 6400 >100 bps

Maximum speed [nm s−1 ]

57 pN 110 pN 100 pN nm 550 pN nm

25 pN

10 pN 1.5 pN 6 pN 6 pN 34 pN

Maximum force [pN]

80

NA NA

NA

50 50 50

Efficiency [%]

Table 1. The sizes refer to the motor domains. Dynamic parameters were determined by in-vitro experiments at high ATP concentration

87, 88 89, 90 91 92, 93 6 94

2, 81 82 2, 83 84, 85 86

Reference(s)

Table 2 Values characterizing the operation of several important biomolecular motors. The filaments along which the linear motors operate are indicated in

1 Overview 5


Fig. 1 Biomolecular motor systems currently applicable for nanotechnological developments. (A) Linear transport of filaments by surface bound motor molecules (gliding assay). (B) Linear movement of motor proteins along filaments (stepping assay). (C) Rotation generated by a rotary motor.

1 Overview

Fig. 2 (A) Directed movement of gliding microtubules along microstructured polyurethane channels on the surface of a coverslip. The initial positions of the microtubules are shown in orange, while the paths they traveled over the subsequent 12 s are shown in green. (B) Scanning electron microscopy image of the polyurethane channels. The channels are a replica mold of a

Si-master (channel width 500 nm, periodicity 1000 nm, depth 300 nm) produced using a poly(dimethylsiloxane) (PDMS) stamp as an intermediate. Note, that the ridges have been “undercut”. This probably aids the guiding of the microtubules in the channels. (Silicon master provided by T. Pompe, Institute of Polymer Research, Dresden, Germany.)

be measured [28]. In addition to beads, 10 (µm-diameter glass particles [29] and Simicrochips [30] have been transported and membrane tubes have been pulled [32]a along filaments. In another variation, high-sensitivity fluorescence microscopy is used to visualize individual motor molecules as they step along filaments [31, 32]. An example from our laboratory of a single kinesin motor fused to the green fluorescent protein moving along a microtubule is shown in Figure 3. Despite the power of single-molecule techniques, they have yet to be exploited for nanotechnological applications. Rotary motors can be studied in vitro by fixing the stator to a surface and following the movement of the rotor (see Figure 1C). Rotation can be visualized under the light microscope by attaching a fluorescent label or a microscopic marker to the rotor. Both techniques have been used to investigate the stepwise rotation generated by F1 -ATPase, which is a component of the F1 F0 -ATP synthesis machinery [5, 33]. Individual motors have been integrated into nanoengineered environments by arraying them on a nanostructured surface and using them to rotate fluorescent microspheres [34] or to drive Ni-nanopropellers [6].

7


Fig. 3 Movement of a single kinesin molecule (labeled with the green fluorescent protein) along a microtubule (red). Micrographs were acquired at the indicated times using total-internal-reflection fluorescence microscopy.

2 Methods

There are many challenges in applying biomolecular motors to nanotechnology. Motility must be robust, it must be controlled both spatially and temporally, and the motors must be hitched to and unhitched from their cargoes. This section summarizes key techniques towards these ends. 2.1 General Conditions for Motility Assays

Motility assays are performed in aqueous solutions that must fulfill a number of requirements. We will illustrate these requirements with the kinesin/microtubule system. Kinesin uses ATP as its fuel; the maximum speed is reached at ∼0.5 mM, approximately equal to the cellular concentration. Other nucleotides such as GTP, TTP, and CTP can substitute for ATP, but the speed is lower [35]. Motility also

2 Methods 9

requires divalent cations, with magnesium preferred over calcium, and strontium and barium unable to substitute [36]. Optimal motility, assessed by gliding speed, occurs over a range of pH, between 6 and 9 [35, 37], and over a range of ionic strengths, between 50 mM and 300 mM [37]. The speed increases with temperature, doubling for each 10◦ C between 5◦ C and 50◦ C [24, 38]; motility fails at higher temperatures. The force is independent of temperature between 15◦ C and 35◦ C [39]. When assays are performed in the middle of these ranges, motility is robust and only a small drop in the mean velocities is seen after 3 hours [24, 37]. If fluorescent markers are used, then an oxygen-scavenging enzyme system must be present in order to prevent photodamage. Many experimental details, including a discussion of the densities of the motors, can be found in Ref. [7].

2.2 Temporal Control

Motors can be reversibly switched off and on by regulating the concentration of fuel, or by adding and removing inhibitors. The ATP concentration can be rapidly altered by flowing in a new solution. In such a setup, the kinesin-dependent movement of microtubules can be stopped within 1 s and restarted within 10 s (unpublished data from our laboratory). Similarly, inhibitors such as AMP–PNP (a non-hydrolyzable analogue of ATP [40]), adocia-sulfate-2 (a small molecule isolated from sponge [41]) and monastrol [42] can be perfused to stop motility. An alternative method to control energy supply is to use photoactivatable ATP. In this method, a flash of UV light is used to release ATP from a derivatized, nonfunctional precursor; an ATP-consuming enzyme is also present to return the ATP concentration to low levels following release. Using such a system, microtubule movement has been repeatedly started and stopped [9], though the start-up and slow-down times were slow, on the order of minutes. The advantage of this method is that the solution in the flow cell does not have to be exchanged. Fortuitously, many proteins possess natural regulatory mechanisms and, once understood, these might offer additional means to regulate the motors in vitro. Examples include the regulation of myosins by phosphorylation and calcium/calmodulin [43] and the inhibition of kinesin by its cargo-binding “tail” domain [44]. Because such natural controls might not always be applicable in a synthetic environment, there is strong interest in the development of artificial control mechanisms for motor proteins. Towards this end, metalion binding sites have been genetically engineered into the F1 -ATPase motor. The binding of ions at the engineered site immobilizes the moving parts of the motor, thus inhibiting its rotation [45]. ATP-driven rotation can be restored by the addition of metal ion chelators. Clever genetic engineering of motors could provide temporal control mechanisms that may be switched by temperature, light, electrical fields, or buffer composition.


2.3 Spatial Control

In order to control the path along which filaments glide – a process that we call “guiding” – it is necessary to restrict the location of active motors to specific regions of a surface. This can be done by coating a glass or silicon surface with resist polymers such as PMMA, SU–8, or SAL601 and using UV, electron beam or soft lithography to remove resist from defined regions [12–19]. The motor-containing solution is then perfused across the surface. By choosing appropriate properties of this solution [e. g., the concentration of motors, salts, other blocking proteins such as casein and bovine serum albumin (BSA), and detergents such as Triton X-100], motility can be restricted to either the unexposed, resist surface or to the exposed, underlying substrate. For example, it has been found that myosin motility is primarily restricted to the more hydrophobic resist surfaces while kinesin motility is primarily restricted to the more hydrophilic non-resist surfaces. However, the detailed interactions of the motors with these surfaces are not well understood. One limitation of this approach to binding proteins to surfaces is that the motors tend to bind everywhere, so it is difficult to attain good contrast. A proven method to prevent motor binding is to coat a surface with polyethylene oxide (PEO) [10, 46]. Techniques to bind motors and filaments via affinity tags to surfaces are summarized in section 2.4. While chemical patterning can restrict movement of filaments to areas with a high density of active motors, walking off the trails is not prevented. This was demonstrated by Hess et al. [10], who showed that microtubules move straight across a boundary between high motor density (non-PEO) and low motor density (PEO), where they dissociate from the surface. The problem with a purely chemical pattern is that if a rigid filament is propelled by several motors along its length, there is nothing to stop the motors at the rear from pushing the filament across a boundary into an area of low motor density. The behavior of microtubules colliding with the walls of channels imprinted in polyurethane has been studied by Clemmens et al. [11]. They found that the probability of a filament being guided by the walls decreased as the approach angle increased. At high incident angles, guiding was not observed and instead the microtubules climbed the walls. Combining chemical and topographic features – as occurs in the lithographic studies described above – leads to more efficient guiding. For example, in the study of Moorjani et al. [18], filaments remained at the bottom of the channels formed in the SU–8 even when they collided with the walls at angles above 80◦ . When the leading end of the microtubule hits the wall, the motors at the rear force the microtubule to bend into the region of high motor density, and in this way the motion is guided by the boundary (see Figure 4, unpublished results from our laboratory). While it is possible to use chemical and topographical patterning to guide filaments – that is, to restrict their movement to particular paths – it is more difficult to control the direction of movement along the path. The difficulty arises because the orientation in which motors bind to a uniform surface is not controlled. Some motors will be oriented so that they propel filaments in one direction along the path, whereas others will propel filaments in the opposite direction. The reason

2 Methods

Fig. 4 Sequence of fluorescent images showing the kinesin-driven, unidirectional movement of a rhodamine-labeled microtubule (red) along chemically and topographically structured Si-chip. The bottom of the channels (green), the depths of which

are 300 nm, is coated with kinesin. The surrounding regions are blocked by polyethylene glycol. (Research in collaboration with R. M.M. Smeets, M.G.L. van den Heuvel, and C. Dekker, Delft University of Technology, The Netherlands.)

that motors do not counteract each other is that filaments are polar structures: the orientation of the proteins that form up the filaments is maintained all along the length of the filament (see Figure 1). Because the motors bind stereospecifically to the filament, they will exert force in only one direction. Thus, the orientation of the filament determines its direction of motion; one end always leads. The direction of filament gliding can be controlled by the application of external forces. Actin filaments and microtubules both possess negative net charges and, consequently, in the presence of a uniform electric field, will experience a force directed towards the positive electrode. It is possible to apply high enough electric fields to steer motor-driven filaments in a specified direction [19, 21]. Because the refractive index of protein differs from that of water, filaments become electrically polarized in the presence of an electric field, and consequently in a nonuniform field they move in the direction of highest field strength. This so-called dielectrophoretic force has been used to direct the gliding of actin filaments on a myosin-coated substrate [20]. It is even possible to manipulate a microtubule using optical gradients produced by focusing a laser beam (i. e., an optical tweezers) [47]. Directional control of microtubule gliding has also been achieved using hydrodynamic flow fields [23, 29]. An alternative approach to directionality relies on more sophisticated guiding concepts. For example, unidirectional movement of filaments can be achieved if

11


guiding geometries based on arrow and ratchet structures are employed [10, 15]. An example of the unidirectional movement of a microtubule on a topographically and chemically structured silicon chip is depicted in Figure 4 To control the direction of motion in stepping assays, the orientation of the filaments on the surface must be controlled. Towards this end, the generation of isopolar filament arrays has been achieved by binding specific filament ends to a surface, and using hydrodynamic flow to align the filaments along the surface to which they are subsequently adhered to [30, 48–50]. Alternatively, moving filaments can be aligned in a particular orientation by a flow field prior to fixation by glutaraldehyde [23, 29], which has been shown not to interfere with kinesin motility [51]. Fluid flow has also been used to align microtubules binding to patterned silane surfaces, though the orientation of the microtubules was not controlled [52]. 2.4 Connecting to Cargoes and Surfaces

Cargoes can be attached to filaments using several different approaches. The prospective cargo can be coated with an antibody to the filament [53] or to a filament-binding protein such as gelsolin [54]. A clever refinement of this technique is genetically to fuse gelsolin with a cargo protein, thereby generating a dual-functional protein [55]. Alternatively, the cargo can be coated with streptavidin which binds to filaments that have been derivatized with biotin [56]. There are many other possibilities which have not yet been realized. Analogous methods can be used to couple motors to surfaces. For example, the motor can be fused with the bacterial biotin-binding protein [57] and in this way bound to streptavidin-coated cargoes or surfaces. There any many peptide tags that can be fused to proteins to aid their purification [58, 59]. These tags can be used to couple these proteins to surfaces coated with the complementary ligand. A popular tag is the hexahistidine tag which binds Ni2+ and other metals that are chelated to nitriloamines (NTA). A nice approach is to couple the NTA to the terminal ethyleneoxides of triblock copolymers containing PEO. In principle, this provides specific binding of a histagged motor (or another protein) to a surface while the PEO groups block nonspecific binding [46, 60]. Controlled unloading of cargo has not been demonstrated, but ought to be feasible. For example, there are biotins that can be irreversibly cleaved by light and reversibly cleaved by reducing agents, and the histidine-Ni2+ -NTA connection can be broken by sequestering the Ni2+ with EDTA.

3 Conclusions

Although the first steps have been made towards the operation of biomolecular motors in engineered environments, many advances are necessary before these

3 Conclusions

motors can be used in nanotechnological applications such as working in molecular factories and building circuits. An immediate task is to improve the spatial and temporal control over the motors. By combining improved surface techniques with the application of external electric, magnetic, and/or optical fields it should be possible, in the near future, to stretch and collide single molecules, to control cargo loading and unloading, and to sort and pool molecules. Another goal is to control the position and orientation of motors with molecular precision. This means placing motors with an accuracy of ∼10 nm on a surface and controlling their orientation within a few degrees. In this way both the location and the direction of motion of filaments can be controlled. One approach to molecular patterning is to “decorate” filaments with stereospecifically bound motors. Once aligned along the filament matrix, the motors can be transferred to another surface. This approach was taken by Spudich et al. [48, 61] and should be followed up. A further development of this idea is to directly produce (perhaps by stamping a mold made with a filament) surfaces that have structures functionally similar to motor-binding sites. An alternative approach is to use dippen lithography or other AFM techniques [62] to directly pattern motors on surfaces. The robustness of motors must be increased. Motors operate only in aqueous solutions and under a restricted range of solute concentrations and temperatures. While it is inconceivable that protein-based motors could operate in a nonaqueous environment, two approaches to increasing their robustness can be envisaged. First, motors could be purified from thermophilic or halophilic bacteria, some of which grow at temperatures up to 112◦ C and salinities above 5 M. There are also extreme eukaryotes that grow at up to 62◦ C or 5 M NaCl. This approach has already been taken for ATP synthase [63], but not with linear motors because no obvious homologues of myosins or kinesins have been found in bacteria. Second, a genetic screening approach might reveal mutations that allow motors to operate in less restrictive or different conditions. A longer-term goal is to use the design principles learnt from the study of biomolecular motors to build purely artificial nanomotors that can operate in air or vacuum. This is a daunting prospect however, and it is not even clear what fuel(s) might be used. A potential way forward is to use chemical energy from a surface: for example, it was demonstrated that tin particles slide across copper surfaces driven by the formation of bronze alloy [64], this being analogous to paraffin-driven toy boats. Besides the motor systems discussed so far, other biomechanical assemblies are good candidates for nanotechnological applications. In addition to providing paths along which motors move, active biological filaments on their own might find use in nanotechnological applications. The pushing and pulling forces generated by the polymerization and depolymerization of actin filaments and microtubules provide an alternative method of moving molecules [2, 65]. This ability is of particular interest because bacteria possess actin- [66] and microtubule-like [67] filaments and, as mentioned above, the proteins of extremophilic bacteria function in extreme environmental conditions. Filaments and motors can also self-organize under certain conditions [68–71]. On a side note, the flagellar filament in conjunction

13


with the flagellar motors allow the bacteria to move in three-dimensional liquid space [72]. In addition to the motors that we have described so far, cells contain numerous biomolecular machines that can also be thought of as motors (for example, see Ref. [3]). These machines use chemical energy to replicate DNA (DNA polymerases) and process it (recombinases, topoisomerases and endonucleases), to produce RNA (RNA polymerases) and splice it (spliceosomes), to make proteins (ribosomes) and fold them (chaperones) and move them across membranes (translocases), and finally destroy them (proteasomes). The energy is provided by another group of machines that generate the electrochemical gradients (electron transport system, bacteriorhodopsin) used by the F1 F0 -ATP synthase to make ATP or by flagellar motors to propel bacteria. All these machines are candidates for nanotechnological applications, and a recent report of the use of chaperones to maintain nanoparticles in solution [73] is a step in this general direction. We finish up by pointing out that the high order and nanometer-scale periodicity of DNA, actin filaments and microtubules make them ideal scaffolds on which to erect three-dimensional nanostructures. While these features have been exploited to make DNA-based structures [74], the use of DNA motors to address specific sites (based on nucleotide sequence) has not, to our knowledge, been realized. Some years ago it was proposed that the regular lattice of microtubules might serve as substrates for molecular computing and information storage [75, 76]. While these ideas seem crazy in the context of the living organism, they may be realizable for biomolecular motors operating in engineered environments. At the moment, anything is possible!

Acknowledgements

The authors thank U. Queitsch for help with experiments on guiding microtubules, T. Pompe, R. M.M. Smeets, M.G.L. van den Heuvel, and C. Dekker for fabricating microstructured channels, and F. Friedrich for assistance with the illustrations.

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68 F. J. NEDELEC, T. SURREY, A. C. MAGGS, S. LEIBLER, Nature 1997, 389, 305. 69 T. SURREY, M. B. ELOWITZ, P. E. WOLF, F. YANG, F. NEDELEC, K. SHOKAT, S. LEIBLER, Proc. Natl. Acad. Sci. USA 1998, 95, 4293. 70 K. KRUSE, F. JULICHER, Phys. Rev. Lett. 2000, 85, 1778. 71 D. HUMPHREY, C. DUGGAN, D. SAHA, D. SMITH, J. KAS, Nature 2002, 416, 413. 72 W. S. RYU, R. M. BERRY, H. C. BERG, Nature 2000, 403, 444. 73 D. ISHII, K. KINBARA, Y. ISHIDA, N. ISHII, M. OKOCHI, M. YOHDA, T. AIDA, Nature 2003, 423, 628. 74 N. C. SEEMAN, Nature 2003, 421, 427. 75 S. R. HAMEROFF, R. C. WATT, J. Theoret. Biol. 1982, 98, 549. 76 R. PENROSE, Ann. N. Y. Acad. Sci. 2001, 929, 105. 77 P. SHETERLINE, J. CLAYTON, J. SPARROW, Protein Profile 1995, 2, 1. 78 E. NOGALES, Annu. Rev. Biochem. 2000, 69, 277. 79 C. BUSTAMANTE, Z. BRYANT, S. B. SMITH, Nature 2003, 421, 423. 80 P. J. HAGERMAN, Annu. Rev. Biophys. Biomol. Struct. 1997, 26, 139. 81 C. RUEGG, C. VEIGEL, J. E. MOLLOY, S. SCHMITZ, J. C. SPARROW, R. H. FINK, News Physiol. Sci. 2002, 17, 213. 82 A. MEHTA, J. Cell Sci. 2001, 114, 1981. 83 F. J. KULL, Essays Biochem. 2000, 35, 61. 84 C. SHINGYOJI, H. HIGUCHI, M. YOSHIMURA, E. KATAYAMA, T. YANAGIDA, Nature 1998, 393, 711. References 199 85 S. A. BURGESS, M. L. WALKER, H. SAKAKIBARA, P. J. KNIGHT, K. OIWA, Nature 2003, 421, 715. 86 G. J. WUITE, S. B. SMITH, M. YOUNG, D. KELLER, C. BUSTAMANTE, Nature 2000, 404, 103. 87 M. D. WANG, M. J. SCHNITZER, H. YIN, R. LANDICK, J. GELLES, S. M. BLOCK, Science 1998, 282, 902. 88 N. R. FORDE, D. IZHAKY, G. R. WOODCOCK, G. J. WUITE, C. BUSTAMANTE, Proc. Natl. Acad. Sci. USA 2002, 99, 11682. 89 J. J. CHAMPOUX, Annu. Rev. Biochem. 2001, 70, 369. 90 T. R. STRICK, G. CHARVIN, N. H. DEKKER, J. F. ALLEMAND, D. BENSIMON, V. CROQUETTE, Comptes Rendus Physique 2002, 3, 595.

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93 B. MAIER, L. POTTER, M. SO, H. S. SEIFERT, M. P. SHEETZ, Proc. Natl. Acad. Sci. USA 2002, 99, 16012. 94 D. J. DEROSIER, Cell 1998, 93, 17.

17

1

Biocatalysis in Non-conventional Media Andreas Sebastian Bommarius

Georgia Institute of Technology, Atlanta, USA

Bettina R. Riebel Emory University School of Medicine, Atlanta, USA

Originally published in: Biocatalysis. Andreas Sebastian Bommarius and Bettina R. Riebel. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30344-1

Introduction

Contrary to expectations that enzymes are only active in aqueous solution, activity in almost anhydrous organic solvents was already demonstrated in the 1930s and rediscovered in 1977. It was not water-miscible hydrophilic solvents such as methanol or acetone that proved to be the best reaction media, but hydrophobic water-immiscible solvents such as toluene or cyclohexane. Supposedly, the cause is the partitioning of water between the enzyme surface and the bulk phase of the organic solvent. As comparably hydrophilic solvents such as methanol or acetone can take up basically infinite amounts of water, they strip the remaining water molecules off the enzyme surface. As a consequence, the enzyme is no longer active because it requires a small but measurable amount of water for developing its activity. The advantages of organic media for enzyme reactions are: (i) an enhancement of the solubility of reactants; (ii) a shift of equilibria in organic media; (iii) easier separation of organic solvents than water; (iv) enhanced stability of enzymes in organic solvents; and (v) altered selectivity, including substrate specificity, enantioselectivity, prochiral selectivity, regioselectivity, and chemoselectivity, of enzymes in organic solvents. In water (with no organic phase present) the yield of the ester is about 0.01%, whereas in a biphasic water–water-immiscible organic solvent system consisting of porous glass impregnated with aqueous buffer solution it is practically 100%. Upon dehydration, many enzymes become extremely thermostable, up to and beyond 100◦ C. Most categories of enzymes have been found to be active in organic solvents, not just lipases but also proteases, dehydrogenases, peroxidases, and several others. Enzymatic reactions in organic solvents follow saturation kinetics; mechanisms Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Biocatalysis in Non-conventional Media

and active sites have been found to be the same as in aqueous solution. Kinetic constants, however, do not correlate with a few simple solvent parameters such as hydrophobicity, dielectric constant, or dipole moment; instead, case-by-case correlations are found. While enzymes in organic solvents often display activities that are orders of magnitude of lower than in water, there are several activation mechanisms which each yield about an order of magnitude of improvement: (i) sufficient hydration in organic solvent (between 1% v/v and saturation); (ii) lyophilization at the pH of maximum activity in water; (iii) addition of lyoprotectants such as polyols; (iv) addition of hydrophobic binding pocket protectors such as phenols or anilines; and (v) lyophilization in strong salts such as KCl (if present at > 90% w/w). Whereas “protein engineering” covers knowledge of improving the biocatalyst, the complementary “medium engineering” optimizes the reaction medium, resulting in improved prochiral, regio-, and enantioselectivity. Many such changes have been observed, demonstrating that not only the catalyst but also the medium exerts a decisive influence on the outcome of an enzymatic reaction.

1 Enzymes in Organic Solvents

Conventional wisdom says that enzymes are active in water and in water only, just as in nature; according to these ideas, enzymes immediately deactivate in organic solvents. This picture is wrong for several reasons: Ĺ in nature, enzymes are not active in water but in the proximity of cell membranes, so a double layer of lipid molecules or a micelle would be a more suitable environment to emulate than an aqueous bulk phase; Ĺ enzymes do not necessarily deactivate in organic solvents – other phenomena such as inaccessibility owing to insolubility or denaturation during lyophilization can account for lack of activity.

Despite these arguments and classic papers demonstrating activity of enzymes in partially or wholly organic solvents [1], results from the Lomonosov University in Moscow, Russia, at the end of the 1970s came somewhat as a surprise: Klibanov et al. demonstrated esterification of N-acetyl-L-tryptophan with ethanol in chloroform [2]. As porous glass was impregnated with aqueous buffer solution, Klibanov et al. termed this system a biphasic water–water-immiscible organic solvent system. They considered the system to consist of two phases, the organic phase and the aqueous phase on the glass bead, noted the shift of equilibria for water-forming reactions in nearly anhydrous media and provided equations modeling the situation. In water (no organic phase) the yield of the ester is about 0.01%, whereas in the biphasic system it is practically 100%. However, it was not water-miscible hydrophilic solvents such as methanol or acetone that proved to be the best reaction media but, on the contrary, hydrophobic

2 Evidence for the Perceived Advantages of Biocatalysts in Organic Media 3

water-immiscible solvents such as toluene or cyclohexane. Supposedly, the cause is the partitioning of water between the enzyme surface and the bulk phase of the organic solvent. As comparably hydrophilic solvents such as methanol or acetone can take up basically infinite amounts of water, they strip the remaining water molecules off the enzyme surface. As a consequence, the enzyme is no longer active because it requires a small but measurable amount of water for developing its activity. What are the advantages of organic solvents as media for biocatalytic reactions? 1. The main advantage is an enhanced solubility of many, mostly hydrophobic, reactants insoluble or only sparingly soluble in water. Even for reactants with a moderate solubility in water, that solubility can often limit the attainable reaction rate. 2. In nearly anhydrous media, equilibria of hydrolytic reactions can be shifted towards condensation. In this fashion, reactions are possible in organic media whose equilibrium constants favor the reactants to such a degree that they alone are present in water. 3. As the enzyme in most cases is not dissolved in hydrophobic media but only suspended, it can be separated relatively easily after the reaction is finished. As the enzyme is not immobilized, connected disadvantages such as low activity yields or mass transfer limitations are avoided. 4. Enzymes are sometimes more stable in anhydrous organic solvents than in water. Lysozyme in water at 100◦ C was demonstrated to be 50% deactivated after 30 s (pH 8) or after 100 min (pH 4), but after 140 h in cyclohexane or even after 200 h as a dry powder [3]. 5. Owing to the influence of the medium, enzymes should feature different substrate specificities and selectivities than water. This would allow for tuning of these parameters under the control of the operator. 6. Organic solvents can be recycled more easily than water with its known high enthalpy of vaporization.

2 Evidence for the Perceived Advantages of Biocatalysts in Organic Media 2.1 Advantage 1: Enhancement of Solubility of Reactants

The rates of asymmetric sulfoxidation of thioanisole in nearly anhydrous (99.7%) isopropyl alcohol and methanol catalyzed by horseradish peroxidase (HRP) were determined to be tens to hundreds of times faster than in water under otherwise identical conditions [4]. Similar effects were observed with other hemoproteins. This dramatic activation is due to a much higher substrate solubility in organic solvents than in water and occurs even though the intrinsic reactivity of HRP in isopropyl alcohol and in methanol is hundreds of times lower than in water.


In addition, the rates of spontaneous oxidation of the model prochiral substrate thioanisole in several organic solvents was observed to be some 100- to 1000-fold slower than in water. This renders peroxidase-catalyzed asymmetric sulfoxidations synthetically attractive. 2.2 Advantage 2: Shift of Equilibria in Organic Media 2.2.1 Biphasic Reactors Multi-phase reactors are of interest in biocatalytic reactions if one or several components of the reaction are insoluble or insufficiently soluble in aqueous phases but if an aqueous phase has to be kept, if only for the biocatalyst. However, a two-phase system can be utilized advantageously for the shifting of an equilibrium; this is demonstrated below. We analyze the simple reaction A ⇐⇒ B in an organic-aqueous two-phase system with the assumption that reactant A and product B partition between the two phases. The partition coefficients Pw and Porg are defined by Eq. (1).

PX = [X]org /[X]w

(1)

In both phases, equilibrium exists between A and B [Eq. (2)]. K w = [B]w /[A]w and K org = [B]org /[A]org

(2)

A biphasic equilibirum constant also can be defined as in Eq. (3). K biphas = [B]tot /[A]tot

(3)

[X]tot can be linked to [X]org and [X]w via a mass balance [Eq. (4)]. [X]tot Vtot = [X]w Vw + [X]org Vorg

(4)

In addition, the sum V tot = V w + V org is valid, so insertion of Eq. (4) into Eq. (3) yields Eq. (5). K biphas = {[B]w Vw + [B]org Vorg {/{[A]w Vw + [A]org Vorg }

(5)

Upon division by [A]w V w and introduction of the phase ratio α (α = V org /V w ) we finally obtain Eq.: K biphas = K w · {(1 + α PB )/(1 + α PA )}

(6)

It can be seen from this equation that the effective equilibrium constant K bisphas is a function of the phase ratio α. Within limits, the equilibrium can be shifted in the

2 Evidence for the Perceived Advantages of Biocatalysts in Organic Media 5

Fig. 1 Dependence of the effective equilibrium constant K biphas on the phase ratio " (from Chaplin, 1990).

desired direction by controlling and shifting α. If the reaction is of the form A + B ⇐⇒ C + D, Eq. (7) is equivalent to Eq. (6). K biphas = K w · {(1 + α PC )(1 + α PD )/(1 + α PA )(1 + α PB )}

(7)

Figure 1 further elucidates the connection between K biphas and α: 2.3 Advantage 3: Easier Separation

In comparison to the high enthalpy of evaporation of water (57.36 kJ mol−1 ), the respective values are much lower for organic solvents, which are thus much easier to separate than water. Table 1 lists some boiling points and enthalpies of evaporation for common organic solvents. 2.4 Advantage 4: Enhanced Stability of Enzymes in Organic Solvents

Porcine pancreatic lipase catalyzes the transesterification reaction between tributyrin and various primary and secondary alcohols in a 99% organic medium [5]. Upon further dehydration, the enzyme becomes extremely thermostable. Not only

6 Biocatalysis in Non-conventional Media Table 1 Boiling points and enthalpies of evaporation for common organic solvents Solvent MTBE Acetone CH3 OH THF Hexane DIPE B.p. [◦ C] 55.2 29.70 Hvap [kJ mol−1 ]

56.2 32.00

65.0 39.26

67.0 28.76

69.0 31.93

EA

C2 H5 OH C6 H12 Toluene DMF

69.0 77.1 32.56 34.75

78.5 40.50

80.7 32.79

110.6 149 39.23 60.45

MTBE: methyl t-butyl ether; THF: tetrahydrofuran; EA: ethyl acetate; C6 H12: cyclohexane; DIPE: diisopropyl ether; DMF: dimethylformamide.

can the dry lipase withstand heating at 100 ◦ C for many hours, but it exhibits a high catalytic activity at that temperature. Reduction in water content also alters the substrate specificity of the lipase: in contrast to its wet counterpart, the dry enzyme does not react with bulky tertiary alcohols. During transesterification of amino acid esters with alcohols in the presence of chymotrypsin and subtilisin, both thermal and storage stabilities of chymotrypsin in non-aqueous solvents are observed to be greatly enhanced compared to water. 2.5 Advantage 5: Altered Selectivity of Enzymes in Organic Solvents

One of the most profound influences a solvent system can have on a reaction is a change of selectivity. Enzymes in organic solvents have been discovered in many cases to feature altered selectivity, including substrate specificity, enantioselectivity, prochiral selectivity, regioselectivity, and chemoselectivity [6, 7].

3 State of Knowledge of Functioning of Enzymes in Solvents 3.1 Range of Enzymes, Reactions, and Solvents

Not only do enzymes work in anhydrous organic media, but in this unnatural milieu they acquire remarkable properties such as enhanced stability, altered substrate and enantiomeric specificities, molecular memory, and the ability to catalyze unusual reactions [8]. Regarding the latter point, hydrolases, such as lipases, catalyze not only transesterifications in organic media but also other types of reactions, including esterification, aminolysis, thiotransesterification, and oximolysis. As all of these reactions compete with hydrolyses, which tend to dominate in aqueous media, some of them proceed to an appreciable extent only in non-aqueous solvents. Soon after the initial discovery, it became apparent that neither the source of the enzyme, nor the type of enzyme, nor the type of solvent seem to constrain the

3 State of Knowledge of Functioning of Enzymes in Solvents 7 Table 2 A selection of enzymes active in organic solvents

Class

E.C. number Enzyme

Reaction

Reference

yeast ADH horse-liver ADH alcohol oxidase HR peroxidase PP lipase

ethanol oxidation alcohol oxidation ethanol oxidation sulfoxidation transesterification

Deetz [17] Zaks [19] Zaks [19] Dai [4] Zaks [5]

3.2.1.17 3.4.21.1 3.4.21.62 3.4.21.4 3.4.24.27 3.5.1.5 3.5.1.11

lysozyme α-chymotrypsin subtilisin trypsin thermolysin urease Pen G acylase

transesterification transesterification transesterification dipeptide synthesis hydrolysis of urea resolution of β-amino esters

Zaks [3] Zaks [16] many van Unen [36] Bedell [4] Ghatorae [29] Roche [67]

4.1.2.10 4.2.1.84

(R)-oxynitrilase HCN addition to C O nitrile hydratase H2 O addition to nitriles

Oxido-reductases 1.1.1.1 1.1.1.1 1.1.3.13 1.11.1. Hydrolases 3.1.1.3

Lyases

Wajant [80] Thomas [29]

HR peroxidase: horseradish peroxidase; PP lipase: porcine pancreatic lipase.

use of organic solvents [3]. Various types of enzymes, such as lipases, proteases (chymotrypsin, subtilisin), oxidoreductases (alcohol dehydrogenase, oxidases, and peroxidases), and others, react in organic solvents. A selection of enzymes found to be active is listed in Table 2. Lipases from three different sources, porcine pancreatic, yeast, and mold, act vigorously as catalysts in a number of nearly anhydrous organic solvents. The range of solvents encompasses the whole spectrum from hydrophilic ones such as methanol, isopropanol, and acetonitrile via THF to the hydrophobic ones such as dialkyl ethers, toluene, hexane, cyclohexane, and heptane. 3.2 The Importance of Water in Enzyme Reactions in Organic Solvents 3.2.1 Exchange of Water Molecules between Enzyme Surface and Bulk Organic Solvent Early after the (re)discovery of enzyme activity in organic solvents, the higher level of activity in very hydrophobic solvents, such as hydrocarbons, in comparison with rather hydrophilic solvents, such as alcohols, was noted. Soon the hypothesis was raised that water molecules partition between the enzyme surface and the bulk organic solvent, and they are “stripped off” the enzyme surface in dry organic solvents. This idea explained why hydrophilic solvents, which strip water off an enzyme surface more readily than hydrophobic ones, deactivate enzymes much more readily than the latter. The hypothesis was proven correct by exchange of water on the enzyme surface by tritiated water, T2 O: suspension of the tritiated enzyme in the organic solvent, filtration of the enzyme, and measurement of the


amount of tritiated water in the solvent by scintillation counting demonstrated that several different enzymes (α-chymotrypsin, Subtilisin Carlsberg, and horseradish peroxidase) yielded water molecules immediately after suspension in the solvent. Depending on the solubility of water in the solvent, different amounts of T2 O were desorbed, from 0.4% in hexane to 62% in methanol [9]. The amount of water required by chymotrypsin and subtilisin for catalysis in organic solvents is several hundred molecules per protein molecule, less than required to form a monolayer on the surface. While subtilisin and α-chymotrypsin act as catalysts in a variety of dry organic solvents, the vastly different catalytic activities in these organic solvents are partly due to stripping of the essential water from the enzyme by more hydrophilic solvents and partly due to the solvent directly affecting the enzymatic process. Enzymes as different as yeast alcohol oxidase, mushroom polyphenol oxidase, and horse-liver alcohol dehydrogenase demonstrated vastly increased enzymatic activity in several different solvents upon an increase in the water content, which always remained below the solubility limit [10]. While much less water was required for maximal activity in hydrophobic than in hydrophilic solvents, relative saturation seems to be most relevant to determining the level of catalytic activity. Correspondingly, miscibility of a solvent with water is not a decisive criterion: upon transition from a monophasic to a biphasic solvent system, no significant change in activity level was observed [11]. Therefore, the level of water essential for activity depends more on the solvent than on the enzyme.

3.2.2 Relevance of Water Activity While water level itself seems to be of no significance for enzyme activity in organic solvents, the relative amount of water with respect to total water solubility seems to be very influential. Correspondingly, instead of using concentration cw or mole fraction xw of water as a measure of the amount of water, water activity aw should be preferred. [Activity, linked to concentration by the activity coefficient γ w , aw = γ w · cw , written here for the case of water, is the thermodynamically rigorous way to consider amounts per volume in thermodynamic formulae such as mass action laws.] At the solubility limit in any solvent, water activity is always equal to unity, regardless of the mole fraction. The transesterification of vinyl butyrate with n-octanol with Pseudomonas cepacia lipase yielded increasing K M values with rising water activity; however, vmax values demonstrated a maximum at water activity levels aw between 0.11 and 0.38 [7]. Activity maxima at certain aw values have also been found with many other enzymes, such as optima at aw = 0.55 for Mucor miehei lipase-catalyzed reaction in several solvents of different polarity from hexane to 3-pentanone [13]. In comparing five different lipases, considerable variations not just of the optima of aw but also the dependence of activity on aw itself were found [14]. Moreover, in polar solvents, correlations of activity with aw often cannot be found at all, probably owing to solvent effects beyond those controlling hydration of the enzyme [5].

3 State of Knowledge of Functioning of Enzymes in Solvents 9

Summarizing the findings above, control of water activity aw during enzyme reactions in organic solvents is extremely important, as water activity exerts a crucial influence, and enzyme reactivity crucially depends on it. 3.3 Physical Organic Chemistry of Enzymes in Organic Solvents 3.3.1 Active Site and Mechanism In organic solvents, enzymes react at the same active site as in water; covalent modification of the active center of the enzymes by a site-specific reagent renders them catalytically inactive in organic solvents [16]. Upon replacement of water with octane as the reaction medium, the specificity of chymotrypsin towards competitive inhibitors reverses. Early results with enzymes in organic solvents demonstrated that enzymes work best in organic solvents if there are lyophilized from aqueous solution at or near the optimum pH value in water (see Section 3, below). Any specific pH effect of organic solvents, however, can be dismissed: it was found that the pKa of amino, carboxylic, and phenolic compounds does not differ by more than 0.3 units in the aqueous and lyophilized states [13]. Enzymatic transesterifications in organic solvents follow Michaelis-Menten kinetics, and in quite a few cases, the values of v/K M roughly correlate with the solvent’s hydrophobicity [16]. The dependence of the catalytic activity of the enzyme in organic media on the pH of the aqueous solution from which it was recovered is bell-shaped, with the maximum coinciding with the pH optimum of the enzymatic activity in water. Hammett analysis and finding of corresponding LFERs suggests that the mechanism of an enzyme reaction in organic solvents is the same as in aqueous solution [18]. 3.3.2 Flexibility of Enzymes in Organic Solvents From experiments on (i) replacement of water with other hydrogen bond forming additives and (ii) titration of enzyme amino groups in an organic medium, as well as the literature data on dehydrated enzymes, it is concluded that the water required by enzymes in non-aqueous solvents provides them with sufficient conformational flexibility for catalysis [19]. The activity of lyophilized HL-ADH in several solvents increased by orders of magnitude upon addition of small amounts of water to solvents of dielectric constant ε from 1.9 to 36 [20]. Enzyme flexibility, as measured by electron spin resonance (ESR) spectroscopy with a spin label at the active site, did not depend on water content but on ε of the pure solvent: HL-ADH turns less flexible with decreasing ε. Correlating positively with the hydrophobicity of the solvent, different fractions of inactivated active centers were measured in different solvents with solid-state NMR spectroscopy (13 C-cross-polarization/magic angle spinning (MAS) NMR) [9]. Just as with tritiated water (see above), immediate desorption of water molecules from the protein surface was observed after addition to the organic solvent.


In some instances, a relationship between enzyme enantioselectivity and flexibility has been found: while there are some positive examples in the literature [8], other investigations either found a slightly negative correlation or did not find any correlation at all [23]. 3.3.3 Polarity and Hydrophobicity of Transition State and Binding Site As enzyme active sites commonly have to be totally anhydrous to allow chemical catalysis to occur, substrates have to be desolvated when transferring from solvent to active site. The effect of six solvents ranging from water to hexane on the standard test reaction in organic solvents, the transesterification of N-acetyl-L-Phe-OEt with n-PrOH, with the help of Subtilisin Carlsberg, has been interpreted by taking into account substrate desolvation, as measured by relative substrate solubility [24]. Analysis of a thermodynamic cycle between ground (E + S) and transition state (ES= ) in the solvent and in acetone yields Eq. (8). =

=

=

G obs = G intr + G s = G ES −G E

(8)

Equation (8) states that the apparent and intrinsic differential activation energies G= obs and G*intr differ by the differential Gibbs free solvation energy Gs , with Gs relating the saturation solubilities of N-Ac-L-Phe-OEt in a solvent to acetone as the standard state [Eq. (9)], so that, with G = H – T · S, Eq. (10) follows. G s = −RT ln([S]solvent /[S]acetone ) G

=

=

obs

= H

intr −T

(9) =

· S

intr

−RT ln([S]solvent /[S]acetone )

(10)

While G= intr failed to yield a straight-line correlation, an LFER, with log ε (manifesting solvent polarity) and G= obs in fact did feature such a correlation. The results indicate that substrate desolvation is important and that the intrinsic activation energy of subtilisin catalysis is lowest in polar organic solvents, which may be due to transition-state stabilization of the enzyme’s polar transition state for transesterification. Furthermore, the results strongly point towards the same transition state in water and organic solvents across a wider range of solvent polarity ε. The dramatic activation of serine proteases in non-aqueous media owing to prior lyophilization in the presence of KCl could be due in principle to the relaxation of potential substrate diffusional limitations in clumps of heterogeneous, undissolved enzyme in organic solvent. This hypothesis was checked with a classic experiment for diffusional limitation: Subtilisin Carlsberg was lyophilized with KCl and phosphate buffer in varying proportions ranging from 1 to 99% (w/w) enzyme in the final lyophilized solids [4]. The active enzyme content of a given biocatalyst preparation was controlled by mixing native subtilisin with subtilisin preinactivated with the serine protease inhibitor PMSF and lyophilizing the enzyme mixture in the presence of different fractions of KCl and phosphate buffer. Initial catalytic rates, normalized per weight of total enzyme in the catalyst material, were measured as

3 State of Knowledge of Functioning of Enzymes in Solvents 11

a function of active enzyme for biocatalyst preparations containing different ratios of active to inactive enzyme. Plots of initial reaction rates as a function of percentage active subtilisin in the biocatalyst were found to be linear for all biocatalyst preparations. Thus, enzyme activation, as high as 3750-fold in hexane for the transesterification of N-Ac-L-PheOEt with n-PrOH, is a manifestation of intrinsic enzyme activation and not of relaxation of diffusional limitations resulting from diluted enzyme preparations. Thus, the above-mentioned hypothesis was refuted. As similar results were found for the metalloprotease thermolysin, activation due to lyophilization in the presence of KCl may be a general phenomenon. 3.4 Correlation of Enzyme Performance with Solvent Parameters

Given the multitude of potential organic solvents for enzyme reactions, design rules or at least heuristics were sought early on for picking organic solvents without trying a large number of them experimentally. Organic solvents can be described and classified according to several criteria, such as volatility (melting and boiling points, Tm and T boil ), size (molar mass, M [g mol−1 ]), accommodation of charge (dielectric constant ε), hydrophobicity (water/octanol partition coefficient, log P), distribution of charge (dipole moment µ), or polarizability (polarization constant α); for data, see Table 3. The water/octanol partition coefficient log P of a component A is defined by Eq. (11), where A is in dilute solution below the solubility limit. logP = log{[A]water /[A]n−octanol }

(11)

n-Octanol is chosen as a reference because it mimics biological membranes with a hydrophobic tail and a hydrophilic head capable of hydrogen bonding.

Table 3 Some properties of common organic solvents Solvent MTBE Acetone CH3 OH THF Hexane DIPE B.p. [◦ C] M [g/mol] ε[−] log P µ [D]

55.2 88.15 4.5 1.15 1.23

56.2 65.0 67.0 58.08 32.04 72.10 20.7 32.63 7.58 − 0.23 − 0.76 0.49 2.70 1.71 1.74

EA

69.0 69.0 77.1 86.17 102.17 88.10 1.89 3.23 6.02 3.5 1.9 0.68 0.0 1.24 1.83

C2 H5 OH C6 H12 Toluene DMF 78.5 46.07 24.3 − 0.24 1.74

80.7 110.6 153 84.16 92.13 73.09 2.02 2.38 36.7 3.2 2.5 − 1.0 0.0 0.30 3.24

MTBE: methyl t-butyl ether; THF: tetrahydrofuran; EA: ethyl acetate; C6 H12 : cyclohexane; DIPE: diisopropyl ether; DMF: dimethylformamide.


Fig. 2 Correlation of activity data with solvent parameters [27].

3.4.1 Control through Variation of Hydrophobocity: log P Concept In an influential early investigation, correlation of biocatalytic activity data of aerobic and anaerobic whole-cell biocatalysis with solvent properties resulted in the strongest correlation for the partition coefficient log P, whereas both Hildebrand’s solubility parameter δ and the dielectric constant ε showed either a weak correlation with activity data or none at all [26, 27] (Figure 2). This simple picture, unfortunately, did not hold for all enzyme reactions in organic solvents. 3.4.2 Correlation of Enantioselectivity with Solvent Polarity and Hydrophobicity The enantioselectivity of the transesterification of vinyl butyrate with secphenylethanol was influenced greatly by the solvent; however, not log P (hydrophobicity) but both the dielectric constant ε and the dipole moment µ of the solvent correlated best with the E values [27]. Table 4 demonstrates the correlation of µ with E. Table 4 Correlation of the enantioselectivity E of subtilisin in the transtesterification of

racemic 1-phenylethanol in various organic solvents, and the dielectric constants ε of the solvents [28]

Solvent Dioxane Benzene Triethylamine Tetrahydrofuran pyridine

er

E

Solvent

2.2 2.3 2.4 7.6 12.9

61 54 48 40 31

Dimethylformamide Nitromethane Acetonitrile Methylacetamide

er

E

36.7 35.9 35.9 191.3

9 5 3 3

4 Optimal Handling of Enzymes in Organic Solvents 13 Table 5 Effects of the hydrophobicity (log P) of solvents on the prochiral selectivity (S) of

Pseudomonas sp. lipase in the monohydrolysis of 2-(1-naphthoylamino)trimethylene dibutyrate [29])

Solvent

log P

Acetonitrile Nitrobenzene Acetone Cyclohexanone Butanone 2-Pentanone Chloroform Tetrahydrofuran 2-Hexanone Dioxane t-Butyl acetate t-Butyl alcohol t-Amyl alcohol Triethylamine Toluene Benzene Carbon tetrachloride

−0.33 1.8 −0.23 0.96 0.29 0.80 2.0 0.49 1.3 −1.1 1.7 0.80 1.4 1.6 2.5 2.0 3.0

S >30 >30 18 18 16 16 9.9 9.1 8.8 5.4 5.3 4.9 4.8 4.7 3.5 3.2 2.6

In a third example, the prochiral selectivity results of the Pseudomonas sp. lipasecatalyzed monohydrolysis of 2-(1-naphthoylamino)trimethylene dibutyrate correlate inversely with hydrophobicity, as expressed by log P (Table 5; [29]). The three examples provided in this section demonstrate that each case has to be considered individually. None of the solvent parameters has been found to correlate reliably with enzyme activity or selectivity data, except in individual cases; predictions at this point are not possible.

4 Optimal Handling of Enzymes in Organic Solvents

While different enzyme reactions yield different results in different solvents, thus rendering comparison difficult, a procedure akin to a “standard protocol” for conducting enzyme reactions in organic solvents has been developed. It calls for Ĺ lyophilization at optimum pH in aqueous solution; Ĺ meeting the requirement for water for enzyme function; and Ĺ testing the enzyme with a standard reaction under standard conditions.

In the case of subtilisin, the standard reaction is the transesterification of Nacetyl-L-phenylalanine ethyl ester with n-propanol [Eq. (12)]. N−Ac−L−Phe−OEt + n−PrOH → NAc−L−Phe−OPr + EtOH

(12)


More data have been obtained with this reaction on enzymes in organic solvents than with any other. 4.1 Enzyme Memory in Organic Solvents

Repeatedly, researchers have found the phenomenon that enzyme molecules seem to “remember” the aqueous conditions from which they were prepared [30]. As mentioned above, early results with enzymes in organic solvents demonstrated that enzymes work best in organic solvents if there are lyophilized from aqueous solution at or near the optimum pH value in water. No specific pH effect of organic solvents on enzyme reactions was found: the pK a of amino, carboxylic, and phenolic compounds does not differ by more than 0.3 units in the aqueous and lyophilized state [13]. Catalytic activities of α-chymotrypsin and Subtilisin Carlsberg in various hydrous organic solvents were measured as a function of how the enzyme suspension had been prepared [31]. Direct suspension of the lyophilized enzyme in the solvent containing 1% water was compared with precipitation of the same enzyme from its aqueous solution by a 100-fold dilution with anhydrous solvent. The reaction rate in a given non-aqueous enzymatic system was found to depend on the nature of both enzyme and solvent, but to depend strongly on the mode of enzyme preparation. The common mode of suspending lyophilized enzymes in organic solvents containing very little water results in just a low specific activity. Much higher specific activity can be achieved if the enzymes are dissolved in water first and then diluted with anhydrous organic solvent to the same water content [14]; the lower the water content of the medium, the greater the discrepancy. The mechanism of this phenomenon, termed “lyophilization-induced inactivation”, was found to arise from reversible denaturation of the oxidases on lyophilization: because of its conformational rigidity, the denatured enzyme exhibits very limited activity when directly suspended in largely non-aqueous media but renatures and thus yields much higher activity if first redissolved in water. Lyophilizationinduced inactivation could be minimized, at least for the case of four oxidases employed, by at least two strategies, both involving the addition of excipients to the aqueous enzyme solution before lyophilization: Ĺ addition of lyoprotectants such as polyols or polyethylene glycol to preserve the overall enzyme structure during dehydration; Ĺ addition of phenolic and aniline substrates that presumably bind to the hydrophobic pocket of the enzyme active site.

Not only was the effect of these excipients found to extend activity by up to orders of magnitude but the effect of the two excipient groups was found to be additive, affording up to complete protection against lyophilization-induced inactivation when representatives of the two are present together.

4 Optimal Handling of Enzymes in Organic Solvents 15

4.2 Low Activity in Organic Solvents Compared to Water

One important issue in the discussion of the biotechnological opportunities afforded by non-aqueous enzymology and its impact on biocatalysis is the often drastically diminished enzymatic activity in organic solvents compared to that in water. The loss of comparative specific activity can be as high as 103 −105 . Several recent studies have addressed the issue and have made great strides towards both elucidating causes of the phenomenon of activity loss and designing strategies that systematically enhance activity by multiple orders of magnitude [33]. The goal ultimately is to bring the activity level of enzymes in organic solvents to aqueous-like level. Dipeptide synthesis in acetonitrile is found to be enhanced 425-fold in the α-chymotrypsin-catalyzed reaction between the 2-chloroethyl ester of N-acetyl-Lphenylalanine and L-phenylalaninamide upon lyophilization of the enzyme in the presence of 50 equivalents of 18-crown-6 [34]. Acceleration is observed in different solvents and for various peptide precursors. The enhancement of enzyme activity upon addition of 18-crown-6 to the organic solvent can be reconciled with a mechanism in which macrocyclic interactions of 18-crown-6 with the enzyme play an important role [35]. Macrocyclic interactions (e.g., complexation with lysine ammonium groups of the enzyme) can lead to a reduced formation of inter- and intramolecular salt bridges and, consequently, to lowering of the kinetic conformational barriers, enabling the enzyme to refold into thermodynamically stable, catalytically (more) active conformations. This assumption is supported by the observation that the crown-ether-enhanced enzyme activity is retained after removal of the crown by washing with a dry organic solvent. Much stronger crown ether activation is observed when 18-crown-6 is added prior to lyophilization, and this can be explained by a combination of two effects: the beforementioned macrocyclic complexation effect, and a less specific, non-macrocyclic, lyoprotecting effect. The magnitude of the total crown ether effect depends on the polarity and thermodynamic water activity of the solvent, the activation being highest in dry and apolar media, where kinetic conformational barriers are highest. By determination of the specific activity of crown-ether-lyophilized enzyme as a function of the enzyme concentration, the macrocyclic crown ether (linearly dependent on the enzyme concentration) and the non-macrocyclic lyoprotection effect (not dependent on the enzyme concentration) could be separated. These measurements reveal that the contribution of the non-macrocyclic effect is significantly larger than the macrocyclic refolding effect. Immobilization in a sol–gel matrix accelerated the propanolysis of N-acetyl-Lphenylalanine ethyl ester in cyclohexane for several serine proteases compared to the non-immobilized lyophilized enzymes: 31-fold for Subtilisin Carlsberg, 43-fold for α-chymotrypsin, and 437-fold for trypsin [36]. The activity yield upon immobilization was 90% (α-chymotrypsin). The rate enhancement effect of immobilization on the enzyme activity is highest in hydrophobic solvents.

16 Biocatalysis in Non-conventional Media Table 6 Approximate activation factors for enzymes in organic solvents for various strategies

Action

Approx. activation factor

Sufficient hydration in organic solvent (≥ 1%, < saturation) Lyophilization at pH of maximum activity in water Lyoprotectants such as polyols Hydrophobic binding pocket protectors (phenols, anilines) Strong salts such as KCl (if present at > 90% w/w)

10 10 10 10 10

total:

105

The activation mechanisms, with the corresponding activation ratios by which the maximum attainable rate constants of enzymes in organic solvents can be enhanced, are summarized in Table 6. Activation by a factor of 105 closes most of the gaps between specific activity levels in water and organic solvents. The state of affairs regarding the preparation of highly active enzyme formulations for use in non-aqueous media is summarized by Lee and Dordick [37]. Improved mechanistic understanding of enzyme function and activation in dehydrated environments will lead to the development of a broad array of techniques for generating more active, stable, and enantioselective and regioselective tailored enzymes for synthetically relevant transformations. This, in turn, should result in an increase in the opportunities for enzymatic processes to be developed on a commercial scale. 4.3 Enhancement of Selectivity of Enzymes in Organic Solvents

Enzymatic enantioselectivity in organic solvents can be markedly enhanced by temporarily enlarging the substrate via salt formation [38]. In addition to its size, the stereochemistry of the counterion can greatly affect the enantioselectivity enhancement [39]. In the Pseudomonas cepacia lipase-catalyzed propanolysis of phenylalanine methyl ester (Phe-OMe) in anhydrous acetonitrile, the E value of 5.8 doubled when the Phe-OMe/(S)-mandelate salt was used as a substrate instead of the free ester, and rose sevenfold with (R)-mandelic acid as a Brønsted-Lewis acid. Similar effects were observed with other bulky, but not with petite, counterions. The greatest enhancement was afforded by 10-camphorsulfonic acid: the E value increased to 18 ± 2 for a salt with its R-enantiomer and jumped to 53 ± 4 for the S. These effects, also observed in other solvents, were explained by means of structure-based molecular modeling of the lipase-bound transition states of the substrate enantiomers and their diastereomeric salts. The relatively meager stereoselectivity of horseradish peroxidase in the sulfoxidation of thioanisole in 99.8% (v/v) methanol is vastly improved when the enzyme forms a complex with benzohydroxamic acid [16]. The generality of the observed “ligand-induced stereoselectivity enhancement” is demonstrated with other hydrophobic hydroxamic acids, as well as with additional thioether

5 Novel Reaction Media for Biocatalytic Transformations 17

substrates, and rationalized by means of molecular dynamics simulations and energy minimization.

5 Novel Reaction Media for Biocatalytic Transformations 5.1 Substrate as Solvent (Neat Substrates): Acrylamide from Acrylonitrile with Nitrile Hydratase

One of the best examples for discussing biotransformations in neat solvents is the enzymatic hydrolysis of acrylonitrile, a solvent, to acrylamide. For several applications of acrylamide, such as polymerization to polyacrylamide, very pure monomer is required, essentially free from anions and metals, which is difficult to obtain through conventional routes. In Hideaki Yamada’s group (Kyoto University, Kyoto, Japan), an enzymatic process based on a nitrile hydratase was developed which is currently run on a commercial scale at around 30 000–40 000 tpy with resting cells of third-generation biocatalyst from Rhodococcus rhodochrous J1. Characteristic of this process are its extremely high concentration level and space-time-yield. Solid nitrile substrates render precipitating product up to a solid medium at high degrees of conversion; liquid nitriles can be run as neat substrate. Figure 3 illustrates the connection between substrate concentration up to 15 M (!) and achievable degree of conversion for the example of the transformation of 3-cyanopyridine to nicotinamide.

Fig. 3 Relationship of degree of conversion with substrate concentration for the reaction of cyanopyridine to form nicotinamide [64].


5.2 Supercritical Solvents

Supercritical solvents combine extremely low viscosity as well as diffusivity of gases with the high solubility of compounds in solution. In addition, products can be separated by simply depressurizing the reaction vessel and boiling off the solvent. After having been tested for chemical reactions, supercritical solvents were also tested for enzymatic reactions in around 1985. Owing to the expected dependence of the reaction parameters on temperature and pressure, supercritical solvents should allow the investigation of that influence. Supercritical CO2 (SCCO2 ), especially, with its accessible critical parameters (T crit = 31◦ C, pcrit = 50 bar) offers access to this technique. However, results demonstrated that SCCO2 does not show good results for all reactions. It was found for the synthesis of acrylates by transesterification of methylacrylate with lipase from Candida rugosa that owing to its hydrophilic character SCCO2 was inferior to more hydrophobic supercritical solvents such as propane, ethane, or even SCSF6 with respect to reactivity [41]. A comparison between SCCO2 and n-hexane for the esterification of oleic acid showed that in SCCO2 inhibition by ethanol was less important but that vmax in nhexane was about an order of magnitude higher. Enzyme stability was comparable and satisfactory in both cases [27]. For a review of the field of biocatalysis in supercritical solvents, see Mesiano [85]. 5.3 Ionic Liquids

Ionic liquids are composed of organic cations with a compatible anion; examples are [BMIM][PF6 ] and [MMIM][MeSO4 ] (Figure 4). Unlike conventional organic solvents, ionic liquids possess no vapor pressure but good solvent properties, and a wide range of thermal stability over a wide range of liquid state (≤ 300◦ C); also, they can be used to form two-phase systems with many solvents [42].

Fig. 4 Some ionic liquids [42]: a) 1-butyl-3-methylimidazolium hexafluorophosphate, [BMIM][PF6 ]; b) 1-butyl-3-methylimidazolium bis((trifluoromethyl)sulfonyl)amide, [BMIM][CF3 SO2 ]2 N;

c) 1-methyl-3-methylimidazolium methylsulfate, [MMIM][MeSO4 ]; d) 1-butyl-3-methylimidazolium triflate, [BMIM][CF3 SO3 ]; e) 1-butyl-4-methylpyridium tetrafluoroborate, [MBPy][BF4 ].


Given these properties, they can extend the application of solvent engineering to biocatalytic reactions. Instead of just serving as a replacement for organic solvents, they often lead to improved process performance. The first report on thermolysincatalyzed Z-aspartame synthesis from Z-L-aspartate and L-phenylalanine methyl ester · HCl in 1-butyl-3-methylimidazolium hexafluorophosphate [BMIM][PF6 ] achieved an initial rate (1.2 nmol (min−1 (mg protein)−1 ) comparable to rates in organic solvents, and remarkable enzyme stability obviating immobilization [25]. Since then, a range of enzymatic reactions and whole-cell processes have achieved remarkable yields [42, 78], (enantio) selectivity [45, 46], or enzyme stability [42, 47]. 5.4 Emulsions [Manufacture of Phosphatidylglycerol (PG)]

Fatty acids are important raw materials which are gained from naturally occurring triglycerides. However, thermal processes to obtain fatty acids result in discoloration, so alternative processes have been sought. Lipases (E.C. 3.1.1.3.) catalyze the hydrolysis of lipids at an oil/water interface. In a membrane reactor, the enzymes were immobilized both on the side of the water phase of a hydrophobic membrane as well as on the side of the organic phase of a hydrophilic membrane. In both cases, no other means for stabilization of the emulsion at the membrane were required. The synthesis reaction to n-butyl oleate was achieved with lipase from Mucor miehei, which had been immobilized at the wall of a hollow fiber module. The degree of conversion reached 88%, but the substrate butanol decomposed the membrane before the enzyme was deactivated. Phosphatidylglycerol (PG), which is useful as a lung surfactant for the treatment of respiratory illnesses, can be obtained from phosphatidylcholine (PC) in a transphosphatidylation reaction [Eq. (13)] catalyzed by phospholipase D (PLD, E.C. 3.1.4.4.), with the hydrolysis to phosphatidic acid (PA) as a side reaction [Eq. (14)]. PC + glycerol → PG + choline

(13)

PC + H2 O → PA + choline

(14)

Commonly, the reaction is conducted in an emulsion stabilized by surfactants and containing the substrate (PC). In a microporous membrane reactor, PLD stability could be increased sevenfold by the addition of ethers to the chamber side. Additionally, no surfactants were required as the product could be separated in simple fashion; 20% product was formed, compared with 4% in the simple emulsion system. 5.5 Microemulsions

If the objective is to keep the enzyme active and stable in an aqueous phase but otherwise to use as much organic phase as possible, microemulsions are an option as


Fig. 5 Microemulsion droplets and possible average location of enzyme molecules [59].

a reaction medium. In contrast to ordinary emulsions they are thermodynamically stable and, at a particle diameter of 1–20 nm, accommodate most often only one enzyme molecule (Figure 5). The microemulsion droplets communicate rapidly and exchange their contents through elastic collisions. The boundary between microemulsions and reversed micelles is not clearly delineated, and the two notions are often used interchangeably. Enzyme of almost all classes and structures have been solubilized in microemulsion systems and used for reactions [48]. Activity and stability are often comparable to values in aqueous media. Many substrates which cannot be made to react in water or in pure organic solvents such as hexane owing to lack of solubility can be brought to reaction in microemulsions. Whereas enzyme structure and mechanism do not seem to change upon transition from water to the microemulsion phase [6], partitioning effects often are very important. Besides an enhanced or diminished concentration of substrates in the vicinity of microemulsion droplets and thus of enzyme molecules, the effective pH values in the water pool of the droplets can be shifted in the presence of charged surfactants. Frequently, observed acceleration or deceleration effects on enzyme reactions can be explained with such partitioning effects [41]. 5.6 Liquid Crystals

As described above, microemulsions feature the disadvantage that the reaction product can be separated only with difficulty from other components of the system. This disadvantage can be compensated by employing highly viscous to solid liquidcrystalline phases which consist of the same components as microemulsions (oil, surfactant, water). Many enzymes display activity in several phases of a threecomponent mixture (Figure 6) [51]. At the beginning of the investigation a surfactant/water/oil phase diagram has to be prepared in case it has not been done already. Subsequently, one can generate the reaction medium by mixing corresponding amounts of components according to the phase diagram; in most cases, several phases will form. In cubic inverse phases, which are mechanically very stable, a series of enzymatic reactions have been conducted [51]. Often, stability of enzymes has been found to be significantly


Fig. 6 Liquid-crystalline phases and catalytic activity of peroxidase in ternary water/surfactant/organic solvent phases [59]. Phases: a = normal micelles; b = reversed micelles; c = lamellar aggregates; d = hexagonal aggregates.

higher in such liquid-crystalline phases than in pure isotropic continuous phases and media. 5.7 Ice-Water Mixtures

Even more than 30 years ago, it had been found that hydroxylaminolysis of amino acid esters with the help of trypsin catalysis is accelerated in frozen aqueous phases at −23◦ C in comparison to liquid aqueous phases at 1◦ C [34]. For kinetically controlled peptide synthesis with different serine or cysteine proteases, strongly enhanced yields of di- and oligopeptides have been observed [53] (Table 7; Figure 7). The strength of nucleophiles seems to be much higher in ice/water phases than in the purely liquid phase, as even unprotected dipeptides render good yields and even unprotected amino acid render at least moderate yields (Table 7; Figure 7). The reverse action of a trypsin-free elastase isolated from porcine pancreas was studied in frozen aqueous systems and was found to catalyze peptide bond formation more effectively than in solution at room temperature [36]. The acceptance of free amino acids as nucleophilic amino components indicates a changed specificity of the endoprotease in frozen reaction mixtures. In elastase-catalyzed formation of Ser-, Ile-, and Val-X-bonds in frozen aqueous reaction mixtures, peptide yields obtained depended on the P1 amino acid and the acyl donor chain length. The highest yields have been found at temperatures between −20 and −10◦ C and at pH values where more than 90% of the nucleophile was present as free

22 Biocatalysis in Non-conventional Media Table 7 Yields of α-chymotrypsin-catalyzed peptide synthesis in water and ice using maleoyl-

l-tyrosine methyl ester as acyl donor and various peptides, amino acid amides, and amino acids as nucleophiles , [53] Peptide yield [%] Amino component of the reaction H-Gly-Ala-OH H-Gly-Gly-OH H-Gly-Gly-Gly-OH H-D-Leu-NH2 H-L-Leu-NH2 H-Arg-OH H-Lys-OH H-β-Ala-Gly-OH

25◦ C 5.8 2.6 5.1 9.9 79.1 99% e.e. of the S-product (Figure 10) [7, 38].

Suggested Further Reading G. CARREA, G. OTTOLINA, and S. RIVA, Role of solvents in the control of enzyme selectivity in organic media, Trends Biotechnol. 1995, 13, 63–70. G. CARREA and S. RIVA, Properties and synthetic applications of enzymes in

organic solvents, Angew. Chem. Int. Ed. 2000, 39, 2226–2254. A. M. KLIBANOV, Improving enzymes by using them in organic solvents, Nature 2001, 409, 241–246.

References 1 E. A. SYM, The action of esterase in the presence of organic solvents, Biochem. J. 1936, 30, 609–617. 2 A. M. KLIBANOV, G. P. SAMOKHIN, K. MARTINEK, and I. V. BEREZIN, 1977, A new approach to preparative enzymatic synthesis, Biotechnol. Bioeng. 19, 1351–1361, reprinted in Biotechnol. Bioeng 2000, 67, 737–747. 3 A. ZAKS and A. M. KLIBANOV, Enzyme-catalyzed processes in organic solvents, Proc. Natl. Acad. Sci. USA 1986a, 82, 3192–3196. 4 L. DAI and A. M. KLIBANOV, Peroxidase-catalyzed asymmetric

sulfoxidation in organic solvents versus in water, Biotechnol. Bioeng. 2000, 70, 353–357. 5 A. ZAKS and A. M. KLIBANOV, Enzymatic catalysis in organic media at 100 degrees C, Science 1984, 224, 1249–1251. 6 C. R. WESCOTT and A. M. KLIBANOV, The solvent dependence of enzyme specificity, Biochim. Biophys. Acta 1994, 1206, 1–9. 7 Y. HIROSE, K. KARIYA, Y. NAKANISHI, Y. KURONO, and K. ACHIWA, Inversion of enantioselectivity in hydrolysis of 1, 4-di-hydropyridines by point mutation of lipase PS, Tetrahedon Lett. 1995, 36, 1063–1066.

References 8 A. M. KLIBANOV, 1989, Enzymatic catalysis in anhydrous organic solvents, Trends Biochem. Sci. 14, 141–144. 9 L. A. S. GORMAN and J. S. DORDICK, Organic solvents strip water off enzymes, Biotechnol. Bioeng. 1992, 39, 392–397. 10 A. ZAKS and A. M. KLIBANOV, Substrate specificity of enzymes in organic solvents vs. water is reversed, J. Am. Chem. Soc. 1986b, 108, 2767–2768. 11 V. S. NARAYAN and A. M. KLIBANOV, Are water-immiscibility and apolarity of the solvent relevant to enzyme efficiency? Biotechnol. Bioeng. 1993, 41, 390–393. 12 R. BOVARA, G. CARREA, G. OTTOLINA, and S. RIVA, Effects of water activity on V max and K M of lipase catalyzed transesterification in organic media, Biotechnol. Lett. 1993, 15, 937–942. 13 R. H. VALIVETY, P. J. HALLING, and A. R. MACRAE, Reaction rate with suspended lipase catalyst shows similar dependence on water activity in different organic solvents, Biochim. Biophys. Acta 1992a, 1118, 218–222. 14 R. H. VALIVETY, P. J. HALLING, A. D. PEILOW, and A. R. MACRAE, Lipases from different sources vary widely in dependence of catalytic activity on water activity, Biochim. Biophys. Acta 1992b, 1122, 143–146. 15 G. BELL, A. E. M. JANSSEN, and P. J. HALLING, Water activity fails to predict critical hydration level for enzyme activity in polar organic solvents: interconversion of water concentrations and activities, Enzyme Microb. Technol. 1997, 20, 471–477. 16 A. ZAKS and A. M. KLIBANOV, Enzymatic catalysis in nonaqueous solvents, J. Biol. Chem. 1988a, 263, 3194–3201. 17 H. R. COSTANTINO, K. GRIEBENOW, R. LANGER, and A. M. KLIBANOV, On the pH memory of lyophilized compounds containing protein functional groups, Biotechnol. Bioeng. 1997, 53, 345–348. 18 L. T. KANERVA and A. M. KLIBANOV, Hammett analysis of enzyme action in organic solvents, J. Am. Chem. Soc. 1989, 111, 6864–6865. 19 A. ZAKS and A. M. KLIBANOV, The effect of water on enzyme action in organic media, J. Biol. Chem. 1988b, 263, 8017–8021.

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1

Photoproteins as Reporters in Whole-cell Sensing Jessika Feliciano, Patrizia Pasini, Sapna K. Deo, and Sylvia Daunert University of Kentucky, Lexington, USA

Originally published in: Photoproteins in Bioanalysis. Edited by Sylvia Daunert and Sapna K. Deo. Copyright ľ 2006 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31016-6

1 Introduction

Reporter genes code for proteins that produce a detectable signal, which can be differentiated in a mixture of endogenous intra- or extracellular proteins [1, 2]. They have found applications in studies of transcription control, gene expression, cell-signaling mechanisms, drug target discovery, cell-based analyte biosensing, and environmental monitoring of specific microorganisms [3], among others. Photoproteins can serve as reporters in two different ways: via genetically engineered systems that are selective toward specific analytes or by using a constitutively bioluminescent/fluorescent cell line that measures overall toxicity. This chapter will focus on the applications of photoproteins as reporters in genetically engineered whole-cell sensing systems.

1.1 Biosensors Using Intact Cells

A biosensor combines a biological component (the sensing element), which is responsible for the selectivity of the device, with a detection system (the transducer) for measuring the reaction of the biological component with the substance (analyte) being monitored. The biological component can be an enzyme, an antibody, a receptor, or whole cells, while the biological reaction that is monitored can be the activity of an enzyme, the binding of an analyte to an antibody or receptor, the induction of gene expression within cells, or even cell death [4, 5]. A number of detection systems have been employed to monitor these biological events, including electrochemical, optical, and piezoelectric systems [6, 7]. Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Photoproteins as Reporters in Whole-cell Sensing

There are a number of advantages in using intact microorganisms, rather than isolated biological components, in biosensors. Microorganisms are usually more tolerant of assay conditions that would be detrimental to an isolated enzyme or protein. Because microorganisms have mechanisms that help them regulate their internal environment, they are also more tolerant of suboptimal pH and temperatures than purified enzymes and are less likely to be inhibited by solutes in the sample being assayed. Microorganisms are cheaper to use in biosensors because the active biological component does not have to be isolated and unlimited quantities can be prepared relatively inexpensively [8–10]. Because an analyte, such as a pollutant, must be taken up by the microorganism before a response is produced, using intact microorganisms can provide information about the bioavailability of the analyte. Classical analytical methods of detection cannot distinguish between bioavailable pollutants from those that are unavailable. In that regard, sensing systems using intact cells complement the more conventional sensors where isolated biological components are being used. These systems may provide physiologically relevant data and give an indication of the bioavailability of a pollutant, which is especially important when decisions regarding remediation of contaminated environments have to be made [9]. While only the bioavailable amount can be obtained, cell-based biosensors are preferred over other biological biosensors because one can acquire other types of information such as toxicity, mutagenicity, and pharmacological activity. Because of all these characteristics, cell-based biosensing systems have found applications in biotechnology, pharmacology, molecular biology, and clinical and environmental chemistry. Among the advances over the years is the constant wish to improve the systems. Improved high-throughput assays for gene function, automation to enhance screening and capacity, and miniaturization to reduce costs have been developed. Systems for the correction of changes in signal that are due to the physiological state of the cell have also been developed. For a good review on how the bioreporter design can be improved by mutagenizing the sensing and the regulatory proteins involved, readers are encouraged to review Ref. [9]. Several reviews have been written on cell-based biosensors, including by our group in 2000 [10]. This chapter will present a general view of the photoproteins with an emphasis on their application specifically to bacterial whole-cell sensing systems. The most advanced technology from our 2000 review will be highlighted, as well as advances in reporter gene technology in cell-based assays since then. 1.2 Reporter Genes in Genetically Engineered Whole-cell Sensors

Genetically engineered whole-cell sensing systems can be developed by coupling a sensing element, which recognizes the analyte and confers selectivity to the system, with a reporter gene, which produces the detectable signal and determines the sensitivity of the system. In this chapter whole-cell sensors that employ receptors as sensing elements will not be discussed. Such biosensing systems are usually based

1 Introduction 3

Fig. 1 Schematic representation of a whole-cell sensing system based on reporter genes.

on eukaryotic cells, either yeast or mammalian cells, and are routinely used in drug screening assays and for the environmental detection of dioxin- and estrogen-like compounds [11–13]. Instead, we will focus on inducible operon-based whole-cell sensors, in which the sensing element is composed of regulatory proteins that are capable of molecular recognition and promoter regions (O/P) of DNA. In a whole-cell sensing system, the regulatory protein controls the transcription of the fused genes. When the target analyte is present, the regulatory protein is capable of recognizing the analyte and will unbind itself from the operator/promoter region, causing transcription to occur. Thus, when the induction takes place in the presence of the target analyte, the reporter gene is co-expressed along with the other genes of the operon. Consequently, the concentration of the inducer can be quantified by measuring the signal generated by the reporter protein (Fig. 1). When a reporter is being selected, one has to take into consideration which type of application the reporter is needed for, including how sensitive the assay should be, in what concentration range one needs to detect, and the nature of the target analyte. In the case of sensitivity, when the reporter gene codes for an enzyme, internal amplification cascades can be used to increase the sensitivity of the device. The stability of the reporter plays a very important role as well. For instance, green fluorescent protein variants that are unstable have different halflives and are more appropriate for kinetic studies than the red fluorescent protein, which takes days to maturate. A good reporter should be environmentally safe, easy to use, highly sensitive, relatively small in size (for inclusion in the vector), and nontoxic, so that expression is related only to the gene under study and not to overall toxicity (biologically inert and benign). Additionally, the reporter should produce a wide dynamic response, its signal should be distinguishable over background, and the expressed product should be enough so that it can be detected directly or

4 Photoproteins as Reporters in Whole-cell Sensing Table 1 Advantages and disadvantages of photoproteins in whole-cell sensing applications

Reporter protein

Advantages

Disadvantages

Bacterial luciferase

High sensitivity. Does not require addition of a substrate. No endogenous activity in mammalian cells. No light source requirement.

Heat labile and therefore of limited use in mammalian cells. Narrow linear range.

Firefly luciferase

High sensitivity. Broad linear range. No endogenous activity in mammalian cells. High quantum yield. Spectral variants.

Sea pansy luciferase

No endogenous activity in bacterial and mammalian cells. No light source requirement. Its substrate, coelenterazine, is membrane permeable. High sensitivity. No endogenous activity in mammalian cells. No light source requirement. Detection down to subattomole levels. Autofluorescent and thus does not require addition of a substrate or cofactors. Spectral variants. No endogenous homologues in most systems. Stable at biological pH.

Requires addition of a substrate. Requires an aerobic environment and ATP. Requires addition of solubilizers for the substrate to permeate the cell. Flash-type emission. Requires addition of a substrate.

Aequorin

Green fluorescent protein

Requires addition of a substrate and the presence of Ca2+ . Flash-type emission. Moderate sensitivity (no signal amplification). Requires post-translational modification. Background fluorescence from biological systems may interfere. Potential cytotoxicity in some cell types.

indirectly. Table 1 summarizes the advantages and disadvantages of the reporter photoproteins. Details about specifics on each of the reporter proteins have been the subject of reviews elsewhere and will not be discussed here. The following section covers the specific characteristics of each photoprotein when used as a reporter in a cell-based assay.

2 The Luciferases

Bioluminescence is the term used to describe the light produced by chemical excitation within living organisms. In many cases light emission is based on luciferasemediated oxidation reactions. Luciferase is a generic name for an enzyme that catalyzes a light-emitting reaction [14]. Luciferases can be found in both terrestrial and aquatic organisms, including bacteria, algae, fungi, jellyfish, insects, shrimps, squids, and fireflies [15]. The most widely used luciferase genes in reporter gene

2 The Luciferases Table 2 Chemical reactions involving photoproteins

Reporter protein

Catalyzed reaction

Bacterial luciferase

FMNH2 + R-CHO + O2 → FMN + H2 O + RCOOH + hν (490 nm)

Firefly luciferase

Firefly luciferin + O2 + ATP → oxyluciferin + AMP + Pi + hν (550–575 nm)

Renilla luciferase

Renilla luciferase + Coelenterazine + O2 → coelenteramide + CO2 + hν (482 nm)

Aequorin

Coelenterazine + O2 + Ca2+ → coelenteramide + CO2 + hν (469 nm)

Green fluorescent protein

Post-translational formation of an internal chromophore

assays are the bacterial luciferase genes (lux genes of terrestrial Photorhabdus luminescens and marine Vibrio harveyi bacteria) and the eukaryotic luc and ruc genes from firefly (Photinus pyralis) and sea pansy (Renilla reniformis). 2.1 Bacterial Luciferases

Bacterial luciferases have found applications in gene transfer and expression, promoter analysis, gene delivery, imaging of gene expression [16], drug discovery and signaling pathways, etc. However, because they are heat-labile dimeric proteins, their applications in mammalian cell-based systems have been limited [1]. Bacterial luciferase catalyzes the oxidation of a reduced flavin mononucleotide (FMNH2 ) and a long-chain aldehyde to FMN and the corresponding long-chain carboxylic acid, emitting light at 490 nm (Table 2). The blue-green light produced in this reaction has a quantum yield between 0.05 and 0.15 [17]. The lux genes from bacteria luciferases have been isolated and extensively employed in the construction of bioreporters. They are five genes that are organized in the luxCDABE operon: luxA and luxB code for the α and β subunits of bacterial luciferase, while luxC, luxD, and luxE code for the synthesis enzymes for the long-chain aldehyde substrate. These five genes are conserved in all bacterial species identified to date. Depending on the combination of the genes employed, different types of bioluminescent sensing systems can be developed [18]. 2.1.1 luxAB Bioreporters These bioreporters contain only the luxA and luxB genes, which are sufficient for generating the bioluminescent signal; however, because the genes that code for the production of the substrate are not present, the substrate must be supplied to the cell. Typically, decanal is added at some point during the assay. This system has the advantage that one can control the moment when the reaction is triggered and the measurement be taken right after, dramatically reducing the basal

5


bioluminescence resulting from “leaky” promoters. Various luxAB sensing systems have been developed in bacterial, yeast, insect, nematode, plant, and mammalian cell sensing systems. Nevertheless, the addition of a substrate makes it impractical for deployable applications, and platforms for biosensors have utilized the entire luxCDABE cassette instead because expression of all five genes has the advantage of not requiring addition of a substrate. 2.1.2 luxCDABE Bioreporters These bioreporters contain the full lux operon cassette, thereby allowing for a completely independent bioluminescent system that requires no external addition of a substrate. In this biosensing system, the bioreporter is exposed to the target analyte and a quantitative change in bioluminescence is detected, often within an hour. The rapidity of the assay, its ease of use, and the substrate-free characteristic of this system make luxCDABE bioreporters ideal for real-time, online, field, and highthroughput applications. For that reason, luxCDABE-based bacterial luciferases are the most widely used photoproteins in whole-cell sensing applications. They have been incorporated into a diverse range of platforms and have found applications in environmental and clinical applications, among others. 2.1.3 Naturally Luminescent Bioreporters One of the earliest applications of bacterial luciferase as a reporter was in the Microtox assay [19, 20]. This assay relies on the changes in light production of the natural luminescence of Photobacterium phosphoreum and is used to measure overall toxicity. Similarly, toxin-sensitive cells can be used in biosensors for the nonspecific detection of toxins. When the cells are exposed to critical amounts of toxins, either the metabolic activity of the cells decreases or the cells die. In either case, as the concentration of toxins in the environment increases there is a corresponding decrease in the signal produced by a reporter protein expressed constitutively within the cells. It should be noted that this type of biosensor will respond to any type of substance that is toxic to the cells. Based on this principle, biosensors have been developed for determining the biotoxicity of a sample [21, 22]. 2.2 Eukaryotic Luciferases 2.2.1 Firefly Luciferase Firefly luciferase, encoded by the luc gene, was first isolated from the North American firefly Photinus pyralis [23, 24]. The structure of this luciferase is different from that of bacterial luciferase, and the resulting bioluminescent reaction exhibits other characteristics as well. The oxygen-dependent bioluminescent reaction converts the substrate luciferin to oxyluciferin based on energy transfer from ATP to the substrate to yield AMP, carbon dioxide, and light in the 550–575 nm range. The bioluminescence produced by firefly luciferase is characterized by a peak after 300 ms, after which it exhibits decay. This decay is produced by the product oxyluciferin, which associates with the enzyme and inhibits light production. This can be

3 Aequorin

overcome by addition of coenzyme A in the reaction mix, which will render dissociation of oxyluciferin from the enzyme and prolong emission over several minutes [23, 24]. The bioluminescence resulting from firefly luciferase exhibits the highest quantum yield known for a bioluminescent reaction (0.88, which is about 10 times larger than that of bacterial luciferase) [1]. Like bacterial luciferase, mammalian organisms do not exhibit endogenous firefly luciferase activity. However, the genes required to synthesize the substrate (luciferin) are not present; thus, addition of an exogenous substrate is always required for bioluminescence to occur. There has been controversy in this matter because the substrate cannot be permeated into the cell [25]. The addition of solubilizers such as DMSO has been required to facilitate entry into the cell. An interesting feature of some eukaryotic luciferases is that they can exhibit light in different wavelengths, ranging from 547 nm to 593 nm [26]. This characteristic has been employed only in colony detection on the same plate, but it can be further exploited in multi-analyte detection by fusing each mutant to a different regulatory gene. Firefly luciferase has found applications in whole-cell sensing systems for the detection of heavy metals and aromatic organics. Its high sensitivity (subattomole level) and broad dynamic range (eight orders of magnitude) make this reporter a favorite in mammalian cell-based sensing systems. 2.2.2 Sea Pansy Luciferase The sea pansy Renilla reniformis is a coelenterate that, unlike other bioluminescent organisms, emits a bright green light. In vivo, Renilla luciferase oxidizes its substrate coelenterazine, and after energy transfer to the green fluorescent protein, light is emitted at 509 nm. In vitro, it is represented by the Rluc gene and produces a blue light of 482 nm [27]. Like other luciferases, Renilla luciferase has no endogenous activity in bacteria and thus can find applications in bacteria-based sensing systems. Studies have shown a sensitivity and detection range similar to the firefly luciferase. Despite its major advantage over firefly luciferase that coelenterazine is membrane permeable, it has limited applications in analytical chemistry. This is probably due to a flash half-life of several seconds. The main application of this eukaryotic luciferase has been in combination with firefly luciferase, in a dual luciferase-based reporter system [28]. Here, both eukaryotic luciferases show a linear response to the target analyte: five orders of magnitude for Renilla luciferase and seven orders of magnitude for firefly luciferase. In both cases, substrate addition makes it impractical for field applications.

3 Aequorin

Aequorin is a calcium-binding photoprotein isolated from the jellyfish Aequorea victoria. This protein is comprised of three components: apoaequorin (the precursor of the photoprotein aequorin), a coelenterazine chromophore, and molecular oxygen [29]. Within the protein, it contains three Ca2+ -binding sites that, once occupied, cause the protein to undergo a conformational change, which in turn triggers the

7


oxidation of coelenterazine to a highly unstable intermediate, coelenteramide. Like Renilla luciferase, it catalyzes the oxidation of coelenterazine to yield a flash blue light of 469 nm that lasts five seconds. This flash-type light emission is accompanied by the release of CO2 , has a quantum yield between 0.15 and 0.20, and requires a Ca2+ concentration of 0.1–10 µM to induce light emission [30]. To date, aequorin is one of the most extensively studied calcium-binding photoproteins. It can be regenerated by dialyzing with EDTA to remove Ca2+ and adding fresh coelenterazine. Among the advantages of using aequorin in reporter gene assays are its high sensitivity and low background. Aequorin can be detected at subattomole levels and is not toxic to various types of cells [31]. Furthermore, a complete absence of endogenous aequorin in mammalian cells can find applications in this type of cellbased sensing systems. Despite these advantages, aequorin is not a popular reporter in cell-based assays. This is probably due to its flash-type light emission (lasting less than five seconds), which requires a luminometer that can initiate and detect the reaction rapidly. In addition, the exogenous requirement of coelenterazine makes this impractical for out-of-the-laboratory applications. Currently, there are two ways of assessing aequorin bioluminescence: detection of aequorin itself or detection of the luminescence triggered by the addition of Ca2+ as the product of the signaling cascade. The latter is more commonly used. Aequorin is widely used as a label in immunoassays and in nucleic acid probe assays, and, because it is a Ca2+ -binding protein, it is used to monitor intracellular levels of free calcium in both prokaryotes and eukaryotes. However, to our knowledge there is only one application in cell-based sensing assays. Rider et al. developed the CANARY (Cellular Analysis and Notification of Antigen Risks and Yields) assay for the detection of pathogenic bacteria and viruses [32, 33]. In this assay, B cells were genetically engineered to express membrane-form antibodies and apoaequorin. The cells were patterned on glass substrates using photolithography and packed into a flow chamber of a microfabricated device. Once the antigen binds, an intracellular signaling cascade is triggered, causing the release of Ca2+ and the activation of the luminescent reaction of aequorin. Detection of pathogenic bacteria and viruses was achieved in less than 1 min with high specificity and sensitivity. An advantage of this assay is that it can be engineered for any target analyte and can be specific to antigens of different pathogens.

4 Fluorescent Proteins

Fluorescent proteins absorb light at a certain maximum wavelength and emit it at a different maximum wavelength, normally a higher one. The green fluorescent protein (GFP) is by far the most popular fluorescent reporter protein in whole cellbased assays. GFP variants and the red fluorescent protein (DsRed) also have found applications in this field. Spectral properties of fluorescent proteins are reported in Table 3.

4 Fluorescent Proteins Table 3 Excitation and emission maxima of fluorescent photoproteins

Reporter protein

Green fluorescent protein (GFP) Enhanced green fluorescent protein (EGFP) Green fluorescent protein UV (GFPuv) Blue fluorescent protein (BFP/EBFP) Cyan fluorescent protein (CFP/ECFP) Yellow fluorescent protein (YFP/EYFP) Red fluorescent protein (DsRed)

Excitation kmax (nm)

Emission kmax (nm)

395 488 395 380 433 513 558

509 509 509 440 475 527 583

4.1 Green Fluorescent Protein

The green fluorescent protein, encoded by the gfp gene, has been isolated and cloned from the jellyfish Aequorea victoria. In vivo the role of GFP is to shift the blue bioluminescence of aequorin to green fluorescence by means of a radiationless energy transfer from aequorin to GFP [34]. The wild-type GFP has an excitation maximum at 395 nm, with a minor peak at 475 nm, and an emission maximum at 509 nm with a small shoulder at 540 nm [35]. The main advantage of GFP as a reporter protein is that it does not require the addition of exogenous substrates or cofactors to produce light; therefore, its use is not limited by the accessibility of substrates [1]. Detection of GFP requires only irradiation by light, which makes GFP the reporter of choice for in situ detection of ana lytes. Induced bacteria expressing GFP can be detected at a single-cell level by using techniques such as epifluorescence microscopy, flow cytometry, or con focal laser scanning microscopy [36]. GFP is a very stable protein, which is advantageous because long-lasting GFP produced from weak promoters or under conditions of low metabolic activity can accumulate in the biosensor cells over time. GFP can be easily expressed in a wide range of organisms, in particular bacteria. This enabled extensive mutational studies of the native protein and the functions of individual amino acids, with the final goal of altering GFP stability and spectral properties (Fig. 2). Variants with improved fluorescence intensity, thermostability, and chromophore folding have been obtained. Shifts within the excitation and emission spectra have also been achieved, thus making available mutants such as the blue, cyan, and yellow variants that can be used for multi-analyte detection purposes [37]. Different strategies have been used to generate variants with shorter half-lives that are suitable for time-dependent induction studies [38]. Limitations to the use of GFP as a reporter include lower detectability compared to other reporter systems. This is due in part to lack of the amplification step occurring with enzyme reaction-based reporters. Additionally, despite the absence of endogenous GFP homologues in most target organisms, the sensitivity of GFP may be compromised by the presence of other fluorescent molecules found in

9


Fig. 2 Expression of fluorescent reporter proteins in E. coli. Reprinted in part from Ref. [10] with permission from the American Chemical Society.

biological systems, which increase background signal [2]. The need for O2 for folding and maturation also limits application to aerobic conditions. Overall, GFP has unique characteristics that have led to its successful application as a reporter for various analytes [10, 38]. 4.2 Red Fluorescent Protein

The dsRed gene encodes a red fluorescent protein (DsRed) with homology to GFP. The red fluorescent protein has been cloned from coral Discosoma sp. [39]. It has an excitation peak at 558 nm and an emission peak at 583 nm. These spectral properties are suitable for its use as a reporter protein. In fact, cellular autofluorescence is supposed to be lower at longer wavelengths, which should result in lower background signal with DsRed as compared to GFP [40]. However, the use of DsRed as a reporter is somewhat limited by the fact that it has a maturation period of several hours to days [41]. In one study the performance of dsRed was compared to that of other reporter genes (gfp, luc, and luxCDABE) in identical constructs in which the reporter genes were inserted under the control of mercury-responsive regulatory units (mer) [42]. As expected, both fluorescent proteins gave a slow response, with DsRed being much slower than GFP. DsRed was expected to give better sensitivity than GFP because of lower autofluorescence; however, that did not happen with the mer–dsRed construct used in the study.

5 Multiplexing

Many of these reporters have been used in combination. Multiple reporter genes are commonly utilized to provide multi-analyte detection or as viability control. Various combinations have been employed, including using different spectral variants of the same photoprotein and using a combination of a fluorescent photoprotein

6 Applications

and bioluminescent one either within the same construct in the same cell [43, 44], within separate constructs within the same cell, or in different cell lines [45, 46]. For instance, the click beetle has four different luciferases that produce bioluminescence emission at different emission maxima (548 nm, 560 nm, 578 nm, and 590 nm) [26, 47]. If their emission spectra do not overlap, the genes encoding the 548-nm and 590-nm emission could be used in a dual-detection system. Not only would detection of more than one analyte be feasible, but also the assay would provide a much simpler system because only one substrate would be required. When using bioluminescent photoproteins, the signals are simultaneously generated and independently measured because of either different kinetics (half-life) or different emission spectrum. In contrast, the fluorescent photoproteins have to be independently excited and/or measured because of different excitation and emission spectra. We developed a dual-analyte sensing system for the simultaneous detection of β-lactose and L-arabinose as model analytes. Two variants of the green fluorescent protein, BFP2 and GFPuv, were used as the signal-generating reporter proteins. The cells were excited at a given wavelength, and emission for the two reporters was collected at wavelengths characteristic of each reporter. The fluorescence emission thus obtained could be correlated with the amount of sugars present in the sample. The system proved to be highly selective for the two analytes [48]. Dual-reporter reagents using Renilla and firefly luciferase reporters are commercially available. For viability control, we developed a fluorescent dual-reporter system that utilizes two fluorescent proteins with different emission spectra (GFPuv and YFP) within the same genetic construct. One of the fluorescent proteins was used for the analytical signal, while the other was the internal reference of general cell viability. Consequently, any environmental conditions that resulted in a nonspecific effect on the analytical signal could be corrected. Such a system with internal response correction is useful for environmental samples whose complex matrices can affect cell viability [49].

6 Applications

Luciferases are the most widely used of the reporter proteins in whole-cell sensing systems. Unlike fluorescent photoproteins, there is no light scattering from other components in the mixture, as well as no background emission from the media because the media itself is not luminescent. This low background expression along with the enzymatic nature and high quantum efficiency of bioluminescent reactions account for the higher and faster detectability of bioluminescence when compared to fluorescence. Because luminometers do not contain a light source, whole-cell sensing systems based on bioluminescence, specifically bacterial luciferases, have been more widely used in real-sample analysis and field applications. The small number of instrument components required has made possible the construction

11

12 Photoproteins as Reporters in Whole-cell Sensing Table 4 Whole-cell sensing systems using photoproteins as reporters able to respond to dif-

ferent stress factors Stress inducer

Promoter

Stress response

Reporter genes

References

Protein damage Oxidative damage Oxidative damage Membrane synthesis DNA damage

GrpE KatG MicF FabA RecA

Heat shock Hydrogen peroxide Superoxide Fatty acid synthesis SOS

luxCDABE luxCDABE luxCDABE luxCDABE luxCDABE gfpuv

[44, 51–56] [43, 52–54, 56, 57] [52] [51–57] [43, 51–55, 57–60]

of cheaper and more compact devices that are suitable for portability in contrast to fluorescence devices. 6.1 Stress Factors and Genotoxicants

Recombinant bacterial whole-cell sensing systems for the detection of general stress, oxidative stress, and genotoxic compounds have been developed. In these systems, reporter genes are fused to promoters of different stress-response regulons, which are activated by a broad range of compounds. A panel of these bacterial strains is usually employed in environmental monitoring studies [50]. Numerous bacterial strains have been developed that are able to respond to different stressful conditions with the production of a reporter photoprotein. Selected examples of such strains are reported in Table 4. 6.2 Environmental Pollutants

A variety of bacterial whole-cell sensing systems based on the promoter–reporter gene concept have been developed for the specific or selective detection of single pollutants or classes of pollutants. Many microorganisms have evolved the ability to survive in suboptimal conditions, including contaminated environments. Such ability usually relies on genetically encoded resistance systems. In the presence of toxic compounds such as heavy metals and metalloids, particular bacteria can synthesize specific proteins that confer resistance to those substances. The mechanisms of resistance vary. Some microorganisms develop efflux pumps, loss-of-uptake systems, or chemical detoxification systems [61, 62]. Other bacterial strains living in contaminated sites can degrade organic xenobiotics and utilize them as carbon and energy sources [63]. Many of these resistance pathways are inducible, meaning that protein synthesis occurs only as required by the presence of given compounds, which makes their use possible for analytical purposes. The gene sequences of numerous regulatory systems have been cloned and plasmid reporter vectors have been constructed that feature the regulatory sequence in gene fusion with a reporter gene. The pioneering work of Sayler and coworkers, who constructed a whole cell-based

6 Applications Table 5 Whole-cell sensing systems using photoproteins as reporters for the detection of

environmental inorganic and organic analytes Analyte

Reporter genes

Regulatory unit

References

Arsenite, arsenate, antimonite

luc luxAB gfp gfpuv

ars

[67–71]

Cadmium, lead

luc luxAB

cad

[72]

Cadmium, lead, zinc

luc luxCDABE rs-GFP

znt

[73, 74]

Chromate

luxCDABE

chr

[75]

Copper

luxCDABE

copA

[73, 76, 77]

Iron

gfp

pvd

[78]

Mercury, organomercurials

luc

mer

[79–84]

luxAB luxCDABE gfp Nickel

luxCDABE

cnr

[85]

Silver

luxCDABE

copA

[77]

Alkanes

luxAB

alk

[86]

Aromatic compounds

luxCDABE

sep

[87]

Benzene, toluene, ethylbenzene, xylene (BTEX)

luc gfp luxCDABE

xyl tbu tod

[88–90]

Naphtalene, salicylate

luxCDABE

nah

[91]

Phenolic compounds

luxCDABE luxAB

mop dmp

[92] [93]

Polychlorinated biphenyls

luxCDABE

bph

Nitrate

gfp

nar

[94, 95]

Phosphate

luxCDABE

pho

[96]

sensing system for the detection of naphthalene and salicylate [64], prompted the development of a plethora of constructs responsive to organic and inorganic pollutants and including different reporter genes. A number of these constructs, along with the application of whole cell-based biosensors to different environmental samples, have been presented and discussed in recent reviews [65, 66]. Many whole-cell sensing systems for the detection of metals and metalloids have been developed, while smaller numbers of biosensing systems for organic pollutants are available. In Table 5 we list inorganic and organic analytes for which whole cell-based biosensors have been developed, along with the regulatory units and reporter genes used. Whole-cell sensing systems are suitable for the detection of metabolites that are formed along the pollutants degradation pathways. As an example, a whole-cell

13


biosensor for the detection of hydroxylated polychlorinated biphenyls in biological and environmental samples is currently under development in our laboratory. Although whole-cell sensing systems have been mainly developed for environmental monitoring purposes, detection of biologically relevant molecules, such as sugars, has benefited from the analytical performance of reporter gene-based whole-cell biosensors [48, 97, 98]. Whole-cell sensing systems for the detection of quorumsensing signaling molecules and antibiotics have been also developed. These are described in more detail in the following sections of this chapter. 6.3 Quorum-sensing Signaling Molecules

Quorum sensing is a phenomenon that enables certain bacteria to communicate with each other by producing and responding to secreted signaling molecules in proportion to cell density [99]. This process allows a population of bacteria to regulate expression of specialized genes in response to changes in its size. When cell population reaches a critical size, certain genes are expressed, hence the term quorum sensing. This phenomenon is responsible for many processes, including expression and secretion of virulence factors, formation of biofilms, competence, conjugation, antibiotic production, motility, sporulation, symbiosis, and bioluminescence [99]. Several quorum-sensing regulatory systems have been identified so far. The best-characterized are those of the LuxI/LuxR type found in most gramnegative bacteria. These quorum-sensing circuits use N-acyl-homoserine lactones (AHLs) as signaling molecules and contain regulatory proteins with homology to the regulatory proteins LuxI and LuxR, which control bioluminescence in the marine bacterium Vibrio fischeri [100]. The gene sequences of some of these regulatory systems have been cloned and inserted in plasmid reporter vectors, thus allowing the development of whole-cell sensing systems for the detection of quorum-sensing signaling molecules. Whole cell-based sensors that contain the luxCDABE cassette as a reporter in gene fusion with quorum-sensing regulatory sequences have been employed to study structure-activity relationships of AHLs [101]. Such biosensing systems have limits of detection in the nanomolar range. They have been applied successfully to the study of AHLs in sputum of cystic fibrosis patients with bacterial lung infections [102]. Whole-cell sensing systems may also represent a tool to identify AHL-producing bacteria by detecting AHLs in their cell-free culture supernatants. Using this approach, it has been shown that Porphyromonas gingivalis, a gramnegative bacterium implicated in the etiology of human periodontal disease, actually regulates the production of virulence factors by quorum sensing but does not utilize AHL-mediated systems [103]. In another study, it has been reported that plant growth-promoting Pseudomonas putida produces cyclic dipeptides that potentially cross talk with the LuxI/LuxR quorum-sensing system [104]. In general, these biosensors can be used as a preliminary screening tool to identify active samples, which can be then subjected to mass spectrometric and NMR analysis in order to determine the chemical identity of individual molecules.

6 Applications

Recently, the molecular mechanisms of bacterial quorum sensing have been proposed as new drug target. In fact, the pharmacological inhibition of quorum sensing for the treatment of bacterial infections may be an alternative to traditional antibiotic treatments, whose therapeutic efficacy is currently decreasing as a result of the occurrence of multiple antibiotic-resistant pathogenic bacteria [105]. Although physical-chemical methods for the detection of AHLs have been developed [106], detailed investigations of quorum-sensing inhibition by methods such as HPLC, GC-MS, or TLC are not practical for routine screening. Whole-cell sensing systems are capable of easily, rapidly, and inexpensively investigating large numbers of compounds and thus are suitable for high-throughput screening of quorum-sensing inhibitors as potential antimicrobial drugs. A GFP-based whole-cell sensing system was developed to perform single-cell analysis and online studies of AHL-mediated communication among bacteria [107]. In this application, a gfp mutant that encodes a protein with a relatively short half-life was used in order to monitor variations in AHL concentrations in real time. Such a biosensor represents a powerful tool for in vivo studies of cell communication and for in vivo efficacy studies of quorum-sensing inhibitors. Indeed, it has been used to detect the production of AHLs from Pseudomonas aeruginosa in a mouse infection model [108]. A similar biosensor allowed detection of AHL production in laboratorybased P. aeruginosa biofilms and showed the ability of a synthetic compound to penetrate microcolonies and block quorum-sensing signaling in most biofilm cells [109]. GFP-based whole-cell sensing systems combined with flow cytometry analysis have been successfully employed for in situ detection of AHLs in soil [110]. 6.4 Antibiotics

Antibiotics belonging to the tetracycline family are extensively used in the therapy and prophylactic control of bacterial infections in human and veterinary medicine and as food additives for growth promotion in the farming industry. Intensive use of tetracyclines has led to widespread antibiotic resistance in bacterial species. The resistance mechanism genes are located in plasmids that can be efficiently transferred from one strain to another. In addition to the development of drug-resistant pathogens, the use of tetracycline has been associated with problems such as unacceptable levels of drug residues in food products for human consumption and release of drugs into the environment. Control of usage in animal farming is possible by monitoring antibiotic residues in different biological samples. Conventional methods for the detection of tetracycline residues include microbial inhibition tests, immunoassays, and chromatographic methods. Recently, whole-cell sensing systems have been proposed as a sensitive, simple, fast, and inexpensive method for measuring tetracyclines in biological fluids and food samples [111]. A bioluminescent bacterial strain containing the bacterial luciferase operon luxCDABE under the control of the tetracycline-responsive element, which is part of the regulation unit of the tetracycline resistance factor, has been developed. It allows for the specific detection of different clinically relevant tetracyclines, with limits of

15


detection at picomole levels and an induction time of 90 min [112]. Moreover, freeze-dried cells can detect tetracyclines as sensitively as freshly cultivated cells, thus envisioning the possibility of on-site use. The developed whole-cell sensing system has been tested on different biological samples. It has proved to be suitable to screen veterinary serum samples for tetracycline residues in real time [113] and to detect tetracyclines in raw bovine milk samples below the official limits set by the European Union and the U.S. Food and Drug Administration [114]. The biosensing strain was also applied to the detection of traces of tetracyclines in fish, after simple sample preparation. In this case, tetracycline residues were detected below official limits, and the results correlated with those obtained by conventional HPLC [115]. In another study, a GFP-based whole-cell sensor system for in situ detection of tetracyclines was developed. The biosensor was used for qualitative detection of oxytetracycline production by the bacterium Streptomyces rimosus in soil microcosms [116]. The tetracycline-induced GFP-producing biosensor cells were detected by using flow cytometry and fluorescence-activated cell-sorting (FACS) analysis. This study showed the biosensor potential for microbial ecology studies. The same sensor strain was employed in a separate study for in vivo detection and quantification of tetracyclines in rat intestine [117]. Bioavailable tetracycline concentration within the bacterial growth habitat of the intestine proved to be proportional to the intake concentration, but significantly lower. This finding, made possible by the use of the whole-cell sensing system, may help to clarify and optimize antimicrobial therapy in the intestinal environment.

7 Technological Advances

An emerging trend of research in luminescent whole-cell biosensors is moving toward integration of the bacterial reporter strain into the appropriate detection device. This goal is achieved mainly by immobilizing the cells onto optical fiber tips. Different immobilization methods that utilize membranes or alginate, solid agar, and sol-gel matrices have been developed and optimized. The obtained fiber-optic, whole-cell sensors have been proposed as self-contained, disposable biomonitoring devices for environmental analysis [59, 118–121]. When coupled to a portable photon-counting system, they allow performance of measurements in the field. Additionally, the use of multiple fibers enables multiple detection [122]. An interesting application of optical fiber led to the development of a high-density cell assay platform [123]. Here, fluorescent reporter cells were placed into ordered microwell arrays that were fabricated on an optical imaging fiber. Fluorescence signals from individual cells were detected and spatially located using an imaging detector. The main advantage of this cell array technology is the simultaneous analysis of multiple responses from a large number of different strains, thus showing potential for high-throughput screening applications. A different approach to integration is the implementation of chip-based systems. An example of a whole-cell biochip is the bioluminescent bioreporter

References

integrated circuit (BBIC), in which recombinant bioreporter cells are interfaced with an integrated circuit. In this application, the cells are either immobilized or encapsulated to integrate them into the circuit. Alternatively, a flow system is employed to bring the cell bioreporters in proximity with the photodiode-containing chip of a microluminometer optimized for very low-level luminescence detection [124, 125]. Further developments in whole-cell sensing aim toward system miniaturization, with consequent implementation of high-density assay platforms, including 96and 384-well microtiter plates, microarrays, and microfluidics devices. The main advantage of these analytical formats is that they allow for high-throughput screening of samples. Not only can large numbers of samples be analyzed, but also large amounts of information from each sample can be assessed simultaneously and in a relatively short time. Additionally, miniaturized systems allow decreased reagent consumption, sample volume, and analysis time. Several whole cell-based assay systems that use multi-well platforms have been developed. Reporter cells can be suspended in culture media within the wells [126], immobilized in agar matrices at the bottom of single wells [127], or encapsulated in sol-gel matrices that form thick silicate films [128]. The main advantage of immobilized cells is that, while maintaining the analytical characteristics of free cells, they can be used under continuous flow conditions for on-line monitoring and as multiple-use sensing elements. In that regard, the optimization of deposition and immobilization techniques can contribute significantly to the advancement of whole-cell sensing technology [129]. An example of whole-cell microarray is the Lux-Array, which includes hundreds of E. coli reporter strains, high-density printed to membranes on agar plates [130]. The system was developed for genome-wide transcriptional analysis by fusing a collection of E. coli functional promoters to a reporter gene. However, it shows the potential for assembling an array of reportersensing strains to be used for analytical purposes. Recently, we took advantage of microfluidics to develop a miniaturized whole cell-based sensing system [71]. A recombinant reporter strain for the detection of arsenite/antimonite was adapted to a microcentrifugal microfluidics platform in the shape of a compact disk. Centrifugal force controlled valving, reagent flow, and mixing and drove fluids through microfabricated channels and reservoirs. The force was generated by a motor that spun the disk at programmed velocities. Fluorescence from the reporter was detected using a fiber-optic probe. Despite miniaturization, the biosensing system retained its analytical performance and was able to perform quantitative analysis within short response times. Moreover, the developed system has the potential for portability and applicability in the field.

Acknowledgments

This work was supported in part by the National Institute of Environmental Health Sciences (Grant P42 ES 007380) and the National Science Foundation (Grant CHE0416553).

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1

Catalytic Antibody Technology Donald Hilvert Swiss Federal Institute of Technology, Zürich, Switezerland

Originally published in: Catalytic Antibodies. Edited by Ehud Keinan. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30688-6

1 Introduction

Antibody molecules elicited with rationally designed transition-state analogs catalyze numerous reactions, including many that cannot be achieved by standard chemical methods. Although relatively primitive when compared with natural enzymes, these catalysts are valuable tools for probing the origins and evolution of biological catalysis. Mechanistic and structural analyses of representative antibody catalysts, generated with a variety of strategies for several different reaction types, suggest that their modest efficiency is a consequence of imperfect hapten design and indirect selection. Development of improved transition-state analogs, refinements in immunization and screening protocols, and elaboration of general strategies for augmenting the efficiency of first-generation catalytic antibodies are identified as evident, but difficult, challenges for this field. Rising to these challenges and more successfully integrating programmable design with the selective forces of biology will enhance our understanding of enzymatic catalysis. Further, it should yield useful protein catalysts for an enhanced range of practical applications in chemistry and biology.

2 Exploiting Antibodies as Catalysts

Some three decades ago, Jencks suggested that stable molecules resembling the transition state of a reaction might be used as haptens to elicit antibodies with tailored catalytic activities and selectivities [1]. Implementation of this clever idea was made possible by the development of monoclonal antibodies, viable transition-state analogs, and versatile screening assays, and more than 100 reactions have now been Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Catalytic Antibody Technology

successfully accelerated by antibodies [2, 3]. These include pericyclic processes, group transfer reactions, additions and eliminations, oxidations and reductions, aldol condensations, and miscellaneous cofactor-dependent transformations. Because selectivities in these systems generally reflect the structure of the hapten and can rival those of natural enzymes, transformations that cannot be achieved efficiently or selectively via more traditional chemical methods are the subject of much current research [4]. Total synthesis of the natural product epothilone from a chiral intermediate prepared by antibody catalysis [5] and activation of a prodrug in vivo [6] are two recent accomplishments that illustrate the potential of this technology. Nevertheless, practical applications are still the exception rather than the rule. Low catalytic efficiency, in particular, appears to be a significant limitation. While modest rate accelerations are easily achieved, enzyme-like activity remains elusive. The goal of this review is to address this problem in the light of recent structural and mechanistic work. After first considering the intertwined issues of catalytic efficiency and hapten design, a few well-characterized examples of antibodies promoting a diverse set of reactions are used to illustrate how antibody binding energy is exploited for catalysis and to identify factors that limit overall efficiency. Finally, possible strategies for improving these systems are discussed.

3 Catalytic Efficiency

By almost any criterion, natural enzymes are incredibly efficient catalysts. The fastest enzymes are limited by the rate at which they encounter substrate, and even those that have not achieved this level of evolutionary perfection typically have apparent bi-molecular rate constants (kcat /K m ) between 106 and 108 M−1 s−1 , irrespective of the rate of the corresponding uncatalyzed reaction [7]. Rate accelerations over background (kcat /kuncat ) are also very high, in the range 106 to 1012 , and can even reach 1017 in some special cases [7, 8]. These extraordinary effects have been explained by an enzyme’s ability to bind transition states more tightly than ground states [9]. When enzymatic and nonenzymatic reactions occur by the same mechanism and chemistry is rate determining, a simple thermodynamic cycle based on transition-state theory can be used to show that kcat /kuncat = K m /K TS , whereK m andK TS are the dissociation constants for substrate and transition state from the enzyme [10, 11]. The chemical proficiency of an enzyme, as defined by Wolfenden [7], is then given as the ratio (kcat /K m )/kuncat or 1/K TS . This term represents the lower limit of a protein’s affinity for the transition state and varies from 108 to 1023 M−1 for a series of natural enzymes [7, 8]. Although true transition-state affinity may be underestimated if chemistry is not clearly rate limiting or if the enzyme uses a different mechanism than the uncatalyzed reaction, this term provides a useful measure of catalytic power for the purpose of discussion.

3 Catalytic Efficiency

One practical consequence of the application of transition-state theory to enzymes has been the design of potent inhibitor molecules through mimicry of structural and electronic features of otherwise ephemeral transition states (for recent reviews, see [12–14]). Another has been the use of such compounds to generate catalytic antibodies [2, 3]. The latter strategy has been found to be broadly useful, as any chemical transformation compatible with a biological milieu is potentially amenable to antibody catalysis so long as an appropriate transition-state analog can be devised. Broad scope and high, programmable catalyst selectivity certainly make this route to tailored enzymes one of the most promising to emerge in the last two decades. In terms of efficiency, however, catalytic antibodies do not yet match their natural counterparts. Published kcat /K m values for the best catalysts are only in the range 102 to 104 M−1 s−1 , well below the limit for diffusion-controlled processes, and rate accelerations are usually between 103 - to 105 -fold over background [15, 16]. Smaller effects are often found, but higher efficiencies are extremely rare. Differences between antibodies and enzymes are readily apparent in Fig. 1, where rate acceleration is plotted against chemical proficiency for a representative set of reactions. Antibody-catalyzed reactions cluster in the lower left quadrant of the plot, corresponding to the lowest activities, whereas enzymatic reactions lie above and to the right, spanning a much greater range of efficiency. Because K m values are roughly comparable in all of these systems, an excellent correlation between rate acceleration and proficiency is observed. In essence, better transitionstate binding translates directly into higher activity. Although the best antibodies approach the efficiency of the least efficient enzymes, it should be noted that the

Fig. 1 Comparison of chemical proficiency, -log (K Ts ), and rate acceleration, kcat /kuncat , for a series of reactions catalyzed by enzymes (filled symbols) and antibodies (open symbols). Rearrangements (, ), hydrolyses (,), decarboxylations (, ), de-

protonations (•,◦) and retroaldol reactions (♦, ) are included. Enzyme data are from [7, 112, 131, 132] and antibody data were taken from [25, 26, 38, 75, 91, 110, 113, 115, 121, 127, 200–202].

3


corresponding reactions often involve conversions of relatively activated substrates (e.g., the hydrolysis of aryl esters). In contrast, enzymes specialize in accelerating extremely slow reactions like the cleavage of amides (t1/2 = 7.3 years) and phosphate diesters (t1/2 = 130 000 years), few of which have yielded to significant antibody catalysis. In a complementary analysis, Stewart and Benkovic [15] found a weak correlation between rate acceleration and the ratio of the equilibrium binding constants of the reaction substrate and the hapten: kcat /kuncat = K m /K TS ≈ K m /K i . In other words, affinity for the transition-state analog roughly approximates affinity for the true transition state, in accord with the basic premise underlying the production of catalytic antibodies. Practically speaking, this means that the search for highperformance catalysts can often be reduced to a search for the best hapten binders – provided the hapten is a reasonably good transition-state mimic. However, association constants for low molecular weight haptens (1/K i = 106 –1010 M−1 ) are generally much smaller than chemical proficiencies of the most effective enzymes. Given K m values of ca. 10−4 M, these affinities suggest that kcat /kuncat will seldom exceed ca. 106 for first generation catalytic antibodies, as found experimentally. For reactions with modest activation barriers, effects of this magnitude may be useful. If product inhibition and protein production pose no further technical hurdles, kinetic resolutions can be achieved, the fate of reactive intermediates controlled and so on. For truly difficult reactions, however, such effects are almost certainly inadequate.

4 Hapten Design

Before turning to a description of specific antibodies, a few words about hapten design are in order. Transition states themselves have fleeting lifetimes and cannot be isolated. Synthesis of effective analogs must therefore draw on our chemical intuition about the conformational, stereochemical, and electronic properties of the reaction under study. Because no stable molecule can reproduce all the characteristics of an actual transition state, design efforts have tended to focus on salient features distinguishing transition state from ground state [12–14]. For instance, changes in hybridization or charge occurring as a reaction proceeds can be mimicked by incorporating different elements or charged groups at appropriate sites within the analog. Conformational constraint can help approximate the reactive geometry of a flexible substrate, while multisubstrate analogs are used to imitate the relative disposition of reactants in a bimolecular process. For reactions that require catalytic functionality within the antibody pocket, more sophisticated strategies appear to be needed. In the “bait-and-switch” approach, charge complementarity between hapten and antibody is exploited to induce appropriately positioned acids, bases and nucleophiles. Alternatively, catalytic residues can be selected directly by irreversible chemical modification when mechanismbased inhibitors [17, 18] are employed as haptens. The latter strategy, dubbed

5 Representative Catalytic Antibodies

“reactive immunization” [19], has the virtue of allowing rational engineering of covalent catalysis. For each new reaction, hapten design must be optimized to maximize the probability of finding an antibody catalyst. Because subtle differences between even the best transition-state analogs and actual transition states almost certainly contribute to lower efficiencies of antibodies compared with enzymes, it is important to understand how instructions implicit in any given hapten design are realized in the complementary immunoglobulin binding pocket. Characterization of successful antibody catalysts at the atomic level currently provides the most useful insights into how binding energy is exploited for catalysis.

5 Representative Catalytic Antibodies 5.1 Proximity Effects

Utilization of binding energy to constrain flexible molecules into reactive conformations or to preorganize reactants for bimolecular reaction is a potentially powerful strategy for accelerating reactions with unfavorable entropies of activation [20, 21]. To test whether antibodies might serve as entropy traps [21], concerted pericyclic reactions requiring neither nucleophilic nor acid-base catalysis have been investigated. 5.1.1 Sigmatropic Rearrangements The biologically important Claisen rearrangement of chorismate to prephenate (Fig. 2) and the abiological oxy-Cope rearrangement (Fig. 3) are typical [3,3]-sigmatropic processes. They proceed via highly ordered, entropically unfavorable, cyclic transition states involving the simultaneous formation of a carbon–carbon bond and cleavage of either a carbon-oxygen or another carbon-carbon bond. For the chorismate rearrangement, the conformationally locked oxabicyclic dicarboxylic acid 1 [22] proved to be a successful hapten (Fig. 2). It has a chair-like geometry very similar to that of the presumed transition state and is an effective inhibitor of natural chorismate mutases [22, 23]. Antibodies that bind 1 catalyze the chorismate rearrangement enantioselectively with rate accelerations (kcat /kuncat ) of 102 to 104 over background [24–27]. For comparison, enzymes accelerate this reaction by a factor of ca. 106 [28]. Spectroscopic and X-ray studies of 1F7 (which achieves a 200-fold rate enhancement) have provided insights into the origins of catalysis in these systems. Transferred nuclear Overhauser effects show that 1F7 binds the flexible chorismate molecule in the diaxial conformation specified by the transition-state analog [29]. Crystallographic data confirm that the induced binding pocket faithfully reflects hapten design [30]. Compound 1 is deeply buried in the complex, and the active site’s overall shape and charge are complementary to a single hapten

5


Fig. 2 (Top) The Claisen rearrangement of chorismate to prephenate catalyzed by antibody 1F7; the flexible chorismate adopts an extended pseudo-diequatorial conformation in solution and must undergo a conformational change to populate the less

stable pseudodiaxial conformer in order for reaction to proceed. (Bottom) The conformationally restricted dicarboxylic acid 1 [22] mimics the transition state of the Claisen rearrangement and was used to elicit catalytic antibody 1F7 [25].

Fig. 3 Antibody-catalyzed oxy-Cope rearrangement [36]. The aldehyde product is trapped with hydroxylamine to give an oxime to prevent time dependent inactivation of the catalyst. The 2,5-diaryl cyclohexanol derivative 2 was used to imitate the structure of the pericyclic transition state.


Fig. 4 (a) Active site of antibody 1F7 [30], showing interactions with hapten 1. The light and heavy chains are pink and blue, respectively; the hapten is yellow. Note that ArgH95 forms a salt bridge with the ligand’s secondary carboxylate but is too far away to form a hydrogen bond with the ether oxygen. (b) Active site of E. coli chorismate

mutase [203]. Bound 1 is completely inaccessible to solvent and makes numerous contacts with protein residues; hydrogen bonds from Lys39 and Gln88 to the ligand’s ether oxygen are essential for high activity [28]. Graphics were prepared with the programs BobScript [204] and Raster3D [205].

enantiomer (Fig. 4a). Consequently, only the corresponding (–)-isomer of chorismate binds in a conformation appropriate for reaction. The subsequent rearrangement of bound substrate then occurs by the same concerted mechanism as that deduced for the uncatalyzed reaction and for natural chorismate mutases. Nevertheless, 1F7 is likely to be a much poorer entropy trap than mutase enzymes. It exploits many fewer hydrogen bonds and electrostatic interactions for ligand recognition (Fig. 4a,b). It also appears to accommodate charge separation in the transition state less effectively. Isotope effects show that the transition state for the rearrangement is highly polarized, with C–O bond cleavage preceding C–C bond formation [31, 32]. The enzymes stabilize this species electrostatically by placing a cationic residue (either Arg or Lys) near the partially negatively-charged ether oxygen of the breaking C–O bond [28, 33, 34], but 1F7 lacks an analogous feature (Fig. 4a,b). These differences presumably explain the antibody’s 104 -fold lower efficiency. They can be attributed, in large part, to shortcomings in hapten design. While 1 reproduces the geometry of the actual transition state reasonably well, it mimics the polarized character of this high energy species poorly [35]. Shortcomings in hapten design are also likely to account for the modest activity of antibody AZ-28. This antibody was raised against cyclohexanol derivative 2

7


(K d = 17 nM) and catalyzes the oxy-Cope rearrangement of the corresponding 2,5diaryl-3-hydroxy-1,5-hexadiene with a kcat /kuncat of 5300 (Fig. 3) [36]. In this case, disposition of the aryl substituents in the transition state is imitated imperfectly in the stable hapten. Like the chorismate mutase antibody, AZ-28 has been shown by TRNOE measurements to preorganize the normally extended hexadiene substrate into a cyclic conformation so that its termini are in close proximity [37]. In this case, ligand recognition is mediated by extensive van der Waals contacts, π-stacking interactions with the aromatic rings, and hydrogen bonding interactions with the alcohol, all evident in the X-ray structure of the antibody-hapten complex (Fig. 5) [38]. However, the conformation adopted by the substrate at the AZ-28 active site is unlikely to be optimal for reaction. The two aryl substituents at C2 and C5 are key recognition elements (Fig. 5) but are oriented very differently in the hapten, where they are sp3 hybridized and equatorial to the plane of the cyclohexane ring, and the transition state, where they are sp2 hybridized and conjugated with the reacting olefins (Fig. 3). Thus, even though the hapten-induced pocket brings together the ends of the hexadiene substrate, binding energy directed to the peripheral aryl groups

Fig. 5 (a) Stereoview of the AZ-28 active site showing hapten-contacting residues [38]. The 5-phenyl ring of 2 sits deep in the hydrophobic pocket, while the 2-phenyl substituent and linker are near the surface. The hapten’s alcohol forms a hydrogen bond

with HisH96 . Two water molecules in the pocket are depicted as red spheres. (b) GRASP [206] surface representation of AZ28 showing the high degree of shape complementarity between ligand and protein.


almost certainly imposes physical constraints that preclude effective alignment of the reacting [4π + 2σ ] orbitals. Alterations in antibody structure leading to improved orbital overlap should therefore result in significant increases in catalytic efficiency. This inference is supported by studies of AZ-28’s germline precursor. This antibody binds hapten 2 with 40-fold lower affinity than the mature AZ-28 but is a substantially better catalyst, achieving a 163 000-fold rate acceleration over background [38]. Mutagenesis experiments showed that replacement of SerL34 in the germline sequence with Asn is responsible for both effects [38]. In the crystal structure of AZ28, AsnL34 interacts directly with the cyclohexyl ring of the hapten and is therefore in a position to influence the conformation of the substrate at the active site. Although designed as entropy traps, and despite evident restriction in the conformational freedom of their substrates, neither 1F7 nor AZ-28 lowers the entropy of activation for its reaction. Both have S‡ values that are 10–20 cal K−1 mol−1 less favorable than the corresponding uncatalyzed reactions [25, 37]. This contrasts with some natural chorismate mutases which do reduce the entropy barrier to reaction significantly [39]. Mechanistic interpretations of activation parameters are necessarily uncertain [40], but the unfavorable S‡ values are consistent with the need for substantial conformational change in the bound substrate as the reaction proceeds. However, other factors, including changes in solvation or conformation associated with the antibody, cannot be excluded. More generally, these two examples show how the chemical instructions implicit in hapten structure, including deficiencies with respect to transition-state mimicry, are accurately imprinted on an antibody binding site. Improved transition-state analogs should therefore yield much better catalysts. To obtain more efficient chorismate mu-tase antibodies, for example, haptens containing additional negative charges might be used to elicit the catalytically essential cation in the vicinity of the substrate’s ether oxygen. Similarly, haptens in which the aryl substituents are coplanar with the cy-clohexyl ring should increase the probability of identifying faster catalysts for the oxy-Cope rearrangement of Fig. 3. The sensitivity of the latter reaction to anionic substituent effects [41] could also be drawn on. Haptens containing an appropriately positioned ammonium group might induce an antibody residue capable of deproto-nating the substrate alcohol. Fine-tuning of the first-generation antibodies is also likely to yield substantially better catalysts. Identification of a second chorismate mutase antibody possessing significantly higher activity than 1F7 [26] supports the feasibility of such an undertaking. Plausible strategies for optimizing activity include site-directed mutagenesis or random mutagenesis coupled with in vivo selection. The 1F7 Fab fragment’s ability to replace the missing enzyme in a chorismate mutase-deficient yeast cell line [42] is a promising indication that it will be amenable to directed evolution in the laboratory [43]. Overall, there are many ways to bind any given transition-state analog, only some of which will be effective for catalysis. Indeed, a significant fraction of hapten binders in any given experiment is usually found to be inactive. Hapten affinity rather than catalytic activity drives maturation of the immune response, so mutations can arise that favor tighter hapten binding but are deleterious for catalysis,

9


as seen for AZ-28. Broad screening of antibodies raised to each hapten is therefore necessary to guarantee a representative sampling of the immune response. In the present instance, antibodies other than 1F7 and AZ-28 might be obtained that ultimately prove to be better starting points for optimization. 5.1.2 Cycloadditions Loss of both translational and rotational degrees of freedom should make bimolecular reactions particularly sensitive to proximity effects [20]. Diels-Alder reactions between dienes and dienophiles have been used to test this notion. They typically have high activation entropies in the range −30 to −40 cal K−1 mol−1 [44], reflecting the low probability of bringing together two substrates in an orientation optimal for reaction. The transition state for these concerted cycloadditions is highly ordered and resembles the boat form of the cyclohexene product more closely than is does the starting materials. Antibodies raised against bicyclic compounds that mimic the transition-state geometry have displayed a range of useful catalytic effects, including control over reaction pathway and absolute stereochemistry [45–49]. The general approach to catalysis is exemplified by antibody 1E9, which promotes the [4+2] cycloaddition cycloadditionand N-ethylmaleimide (Fig. 6) with multiple turnovers [45]. The initially formed adduct 3 spontaneously eliminates sulfur dioxide to give N-ethyl tetrachlorophthalimide as the final product after oxidation in situ. The endo hexachloronorbornene derivative 4, an excellent mimic of the intermediate and its flanking transition states, served as the hapten (K d = 2 nM). Because the planar product is structurally so different from 4, it binds 105 -fold less tightly to the induced antibody, effectively minimizing product inhibition. In this case, catalytic efficiency can be estimated as an effective molarity (EM) [50]. EM is the ratio of the pseudo-first-order rate constant for the antibody reaction (kcat ) to the second-order rate constant for the uncatalyzed process (kuncat ). This ratio

Fig. 6 Diels-Alder condensation of tetrachlorothiophene dioxide (TCTD) and N-ethyl maleimide (NEM) yields a high-energy intermediate (3) which eliminates sulfur dioxide. The initially formed product is subsequently

oxidized in situ. The transition state for cycloaddition and chelotropic elimination of SO2 closely resemble the hexachloronorbornene derivative 4 used as a hapten to elicit antibody 1E9 [45].


gives the nominal concentration of one reactant needed to convert the spontaneous bimolecular reaction into a pseudo-first-order process with a rate equivalent to that achieved in the antibody ternary complex. It is usually interpreted as the entropic advantage of a unimolecular over a bimolecular process, with an upper limit of about 108 M for 1 M standard states [51]. For 1E9, the EM is ca. 103 M [52]. Although much lower than the theoretical limit, this value is significantly higher than EMs reported for other antibody Diels-Alderases, which rarely exceed 10 M [46–49], making 1E9 the most efficient such catalyst described to date. Recent structural work has shown that the 1E9 active site is exactly what one might expect of an antibody that functions as an entropy trap [52]. Extensive van der Waals contacts, π-stacking interactions, and a strategically-placed hydrogen bond to one of the succinimide carbonyl groups create a pocket that is highly complementary to the hapten (Fig. 7a). When complexed, the ligand (excluding the hexanoate linker) is 86% buried. Its fit to the protein is so snug that no interfacial cavities are detectable, even when a probe of 1.2 Å radius is used. Thus, the 1E9

Fig. 7 The active site of Diels-Alderase 1E9 provides a snug, complementary binding surface for its hexachloronorbornene hapten [52]. With the exception of a hydrogen bond between the deeply buried carbonyl group of the ligand and the side chain of AsnH35 (not shown), most of the contacts are

hydrophobic in nature. (b) Antibody 39A11 (see Fig. 8) binds its hapten much more loosely. Note that the “dienophilelike” N-aryl succinimide is significantly better packed than the bicyclo[2.2.2]octene moiety, which serves as a diene surrogate.

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binding pocket appears ideally suited to the task of preorganizing its diene and dienophile substrates in a reactive complex that closely approximates the transitionstate geometry. Nevertheless, a simple entropy trap mechanism does not appear to be operative. The temperature dependence of kcat and kuncat shows that catalysis by 1E9 is achieved entirely by reducing the enthalpy of activation; the solution and the antibody processes are equally unfavorable entropically, with S‡ values of −22 cal K−1 mol−1 [52]. Nor is the rate acceleration due to a simple medium effect associated with the apolar binding cavity, since the uncatalyzed reaction is 10 times slower in acetonitrile than in water. Catalysis by 1E9 can be explained by enthalpic stabilization of the transition state through an unusually close fit to the apolar binding surface of the antibody active site and a strong hydrogen bond between the side chain of AsnH35 and the maleimide carbonyl. Quantum mechanical calculations indicate that the numerous, energetically favorable van der Waals interactions provide the driving force for binding [52]. Although these interactions distinguish poorly between ground and transition state, they hold the substrates against a relatively unfavorable electrostatic field that becomes substantially more favorable as the transition state is approached because of the increased strength of the hydrogen bond to the dienophile carbonyl. Comparison of 1E9 with another Diels-Alderase, 39-A11 [47], dramatically illustrates the importance of close packing for high efficiency. Antibody 39-A11 was generated with the substituted bicyclo[2.2.2]octene derivative 5, which contains an ethano bridge locking the cyclohexene ring into the requisite boat conformation (Fig. 8). It catalyzes the Diels-Alder reaction between an electron-rich acyclic diene and an N-aryl maleimide to give a cyclohexene derivative, albeit with a relatively low effective molarity of 0.35 M.

Fig. 8 Antibody 39-A11 catalyzes a Diels-Alder reaction between an electron-rich acylic diene and an N-aryl maleimide. It was elicited with the bicyclo[2.2.2]octene hapten 5. The ethano bridge in 5 locks the cyclohexane ring into a boat conformation but has no counterpart in the substrate itself.


Considering the low dissociation constant reported for the antibody-hapten complex (K d = 10 nM) [47], the fit of the bicyclo[2.2.2]octene to 39-A11 is surprisingly loose (Fig. 7b) [53]. Only 66% of the hapten surface area is buried in the complex. Poor complementarity is indicated by the large cavity volume of 117 Å3 detected between ligand and antibody. The portion of the hapten corresponding to the reacting [4+2] system is particularly poorly packed, whereas peripheral substituents of 5, especially the aryl side chain, appear to be important recognition elements. Moreover, the hapten’s ethano bridge, which has no counterpart in the substrates or the transition state, carves out additional unwanted space within the pocket. Consequently, the bound substrates – particularly the diene, which must bind in the least complementary region of the pocket – are likely to retain considerable degrees of freedom. Low catalytic efficiency is therefore unsurprising. Consistent with this idea, introducing large aromatic groups at positions L91 and L96 to improve packing interactions with the kinetically favored endo transition state results in 5 to 10-fold higher kcat values [54]. Relatively non-polar as it is, the environment of the 39-A11 active site may further erode catalytic efficiency. Like 1E9, 39-A11 provides a hydrogen bond (also from AsnH35 ) to the dienophile carbonyl, but its reaction involves a strong donor diene and acceptor dienophile rather than two electron-deficient addends. The corresponding transition state should therefore be more polar and hence more sensitive to transfer from water than that of the 1E9-catalyzed reaction. Unfortunately, mutagenesis experiments to augment activity by providing additional hydrogen bonds to the dienophile have not been successful [54]. Structurally distinct haptens notwithstanding, 1E9 and 39-A11 are unexpectedly closely related in primary sequence and tertiary structure [52, 53, 55]. Both belong to a family of polyspecific antibodies that exhibit extensive cross-reactivity for hydrophobic ligands containing one or two polar groups. For example, 39-A11 and its germline precursor accommodate a range of structurally diverse compounds [53], while 1E9 and the related progesterone-binding antibody DB3 [56, 57] bind each other’s ligands with affinities only 25-fold to 50-fold lower than their own [55]. The side chain of AsnH35 and conserved hydrophobic interactions seem to be particularly important for achieving recognition. However, docking experiments [52] with 1E9 and DB3 suggest that non-cognate molecules bind randomly in the apolar cavity, whereas specific ligands adopt a single, well-defined binding mode similar to that seen in crystal structures of the corresponding antibody-hapten complexes. Specificity in these systems appears to be conferred by a small number of mutations to the shared scaffold. In the case of 39A11, for instance, substituting Val for Ser at position L91 in the germline sequence accounts for almost all the 40-fold increase in hapten affinity achieved during affinity maturation [53]. Similarly, a somatic mutation in the L3 CDR loop (SerL89 → Phe) and a rare mutation in the antibody framework region (TrpH47 → Leu) substantially alter the shape of the 1E9 combining pocket compared to that of 39-A11 or DB3 [52]. These changes are primarily responsible for its virtually perfect shape complementarity to the transition-state analog. This complementarity appears crucial for reactivity, since DB3 does not detectably accelerate the 1E9 reaction despite its affinity for hapten 4.

13


Given the immune system’s enormous combinatorial diversity (more than 108 antibodies are available in the primary response [58]), it is surprising that similar antibodies were generated in separate immunization experiments with 4, 5 and progesterone. Nor, as will be shown below, is this finding unique. To what extent does utilization of a few restricted sets of antibody germline genes limit the catalytic potential of the immune system? Do these frequently selected scaffolds represent local minima from which it will be difficult to evolve more active catalysts? Poor shape complementarity between 39-A11 and the bicyclo[2.2.2]octene is a caseinpoint. However, site-directed mutagenesis experiments demonstrate that the mature antibody is not an evolutionary dead end. The same ValL91 Tyr change that improves catalysis 10-fold also increases hapten affinity 2.5 times [54], and it should be possible to find additional mutations that further tighten the structure. Because only ten antibodies were screened for activity, we cannot know whether analogous mutations were present in the antibody population induced in response to hapten 5. It is conceivable that they never emerged: a 10 nM dissociation constant for the 39-A11-hapten complex is more than adequate for the immune system’s purposes, so there may be little or no selection pressure to increase affinity beyond that point. As shown by 1E9, the combining pocket can be molded remarkably well under favorable circumstances to achieve nearly perfect shape complementarity with a ligand. Superior fit is not necessarily manifest in tighter binding, since 1E9 and 39-A11 have similar hapten affinities (K d = 2 nM versus 10 nM). Compound 4 is more highly optimized than 5 with respect to transition-state mimicry, however, and important binding interactions in 1E9 are concentrated where they are needed for catalysis, rather than loosely dispersed as in 39-A11. A high degree of complementarity at the site of reaction thus appears to pay off in this case in terms of a more efficient catalytic outcome. Even in the case of 1E9, though, much higher efficiency should be attainable. An analogous enzyme is unavailable for direct comparison (see [59] for evidence regarding possible Diels-Alderases in Nature), but 1E9’s chemical proficiency, defined as (kcat /K diene K dienophile )/kuncat = 1.4 × 107 , is far from what is expected of a fully evolved catalyst. Given an already excellent fit between protein and ligand, it is unlikely that mutation of residues lining the binding cavity will substantially improve complementarity. To optimize electrostatic interactions with the transition state and reduce any remaining degrees of freedom available to the bound substrates, residues distant from the active site will have to be modified. Because such mutations will be difficult to identify by inspection of the protein structure, combinatorial mutagenesis and an efficient screening assay [60] or selection protocol will be needed if 1E9 variants with enhanced properties are to be developed. 5.2 Strain

Substrate destabilization through strain has been proposed as another mechanism for achieving rate accelerations with enzymes [20, 61]. Binding energy can be used to strain molecules in various ways. Destabilization can involve geometric


distortion of the substrate, electrostatic repulsion between groups of like charge, or desolvation effects. If substrate destabilization is relieved at the transition state, significant reductions can result in the free energy of activation for reaction. Rate accelerations obtained in this way are potentially quite large, limited only by the amount of binding energy available to force the substrate into the destabilizing environment. As with entropic effects, it was predicted that strain mechanisms might be readily exploited for antibody catalysis. 5.2.1 Ferrochelatase Mimics Ferrochelatase is an example of an enzyme that is believed to exploit geometric distortion for catalysis. As the terminal enzyme in heme biosynthesis, it promotes complexation of Fe2+ by protoporphyrin IX [62, 63]. Early work suggested that fer-rochelatase functions by distorting the substrate porphyrin from its preferred planar conformation into a bent structure [62]. This distortion exposes the nitrogen of one of the pyrroles to solvent, thereby facilitating metal ion complexation. In fact, non-planar N-methylated porphyrins are known to chelate metal ions 3 to 5 orders of magnitude faster than their non-alkylated counterparts [62]. They are also potent inhibitors of ferrochelatase [64]. When used as haptens, they have yielded antibodies [65] that catalyze insertion of divalent metal ions into mesoporphyrins with kcat values approaching those of the enzyme (Fig. 9). Ferrochelatase antibody 7G12 has been characterized in some detail. It promotes incorporation of Zn2+ and Cu2+ into mesoporphyrin IX with kcat values of 0.022 s−1 and 0.0069 s−1 , respectively [65]. By way of comparison, the kcat value for recombinant Bacillus subtilis ferrochelatase is ca. 0.4 s−1 for metalation with Fe2+ , Zn2+ and Cu2+ [66]. K m values for the porphyrin are also comparable (50 µM for the antibody and 8 µM for the enzyme). However, whereas substrate metal ions bind tightly to the enzyme (20–170 µM), no evidence for saturation of the antibody has been observed up to concentrations up to 2.5 mM, suggesting that binding of metal ions to the antibody does not contribute to catalysis. Severe inhibition of the antibodycatalyzed reaction by the metalloporphyrin product is a further point of contrast. Mechanistic and structural studies have clarified how the antibody chelatase functions. The crystal structure of 7G12 complexed with its hapten has been solved [67]. The N-methylated porphyrin binds at the junction of the heavy and light chains

Fig. 9 N-methyl mesoporphyrin and the mesoporphyrin metalation reaction catalyzed by antibody 7G12 [65].

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Fig. 10 Stereoview of the 7G12 binding site [67], showing how bound porphyrin packs against the heavy chain (blue) and exposes one face to solvent. The side chains of heavy chain residues AspH96 and ArgH96 are visible under the surface behind the porphyrin. Interactions with the light chain

(pink) are largely restricted to contacts with the methyl and ethyl substituents of the A and B pyrrole rings of the porphyrin. Although a mixture of N-alkylated porphyrins was used, only the derivative with the A ring methylated appears to bind to the antibody.

(Fig. 10). One face of the ligand makes extensive contacts with VH , while the other is relatively exposed to solvent. Packing interactions from light chain Tyr residues may reinforce the distortion from planarity of pyrrole ring A, which bears the N-methyl group. Replacement of these residues with Ala caused large increases in K m and 10-fold to 40-fold decreases in kcat /K m [67], suggesting that they may play a similar role with the non-methylated substrate. In analogy with the enhanced reaction rates achieved by porphyrin alkylation, a catalytic mechanism involving binding and distortion of the porphyrin by the protein, followed by direct chelation of metal ions from solution, seems plausible. Although its precise role is still unclear, amino acid AspH96 is evidently required for this process [67]. Its carboxylate side chain is directed from the VH domain toward the center of the porphyrin ring with one oxygen roughly equidistant from the porphyrin’s four pyrrole nitrogens (Fig. 10). The other oxygen is fixed in place through a hydrogen bond to ArgH95 . It is conceivable that the carboxylate acts as a base which shuttles protons from the porphyrin during metal ion exchange. This residue is also probably responsible, at least in part, for product inhibition, since axial coordination to the metal ion will anchor the metaloporphyrin to the antibody. Direct experimental evidence for porphyrin distortion by 7G12 has been obtained with resonance Raman spectroscopy [68]. Spectral data show that the antibody


induces an alternating up-and-down tilting of the pyrrole rings very similar to the distortion produced by porphyrin alkylation. In contrast, yeast ferrochelatase apparently causes all four pyrrole rings to tilt in the same direction in a domed fashion. The enzymatic reaction is regulated allosterically by a metal-dependent protein conformational change. Since the antibody has no metal binding site, the distortion it induces must be brought about entirely by binding interactions between porphyrin and protein. An atomic-level explanation of this effect will require elucidation of the antibody-substrate complex structure. The broad lesson to be derived from these experiments is that substrate destabilization can be a very successful approach to antibody catalysis. Although 7G12 and ferrochelatase perturb the porphyrin structure in different ways, both kinds of distortion appear to be effective for metal ion chelation, yielding kcat values within a factor of 10 of each other. That said, destabilization mechanisms are expected to have little to no effect on kcat /K m [69], the steady-state parameter generally optimized through evolution. By this criterion, 7G12 is substantially more primitive than its natural counterpart. Because chelatases catalyze a bimolecular reaction, flux through the catalyst is limited by the least favorably processed substrate. For both enzyme and antibody, this is the metal ion. Hence, rate enhancements are given by [(kcat /K M2+ )/kuncat ]. Since K M2+ values for 7G12 are at least 103 times larger than those for natural ferrochelatases (assuming the metal ion binds at all), the antibody suffers an equivalent rate disadvantage under practical operating conditions, despite its favorable kcat . Similar considerations apply to the chemical proficiencies of the two catalysts [(kcat /K M2+ K porphyrin )/kuncat ]. The antibody’s transition-state affinity is at least four orders of magnitude lower than that of the enzyme. To improve 7G12, a suitable binding site for metal ions should be constructed, perhaps by extending light chain CDR loops that are near the exposed face of the bound porphyrin. The challenge will be to bring the metal ion into close proximity with the porphyrin without further increasing the active site’s affinity for product. As for the chorismate mutase antibody discussed above, an in vivo selection strategy is feasible. Ferrochelatase-deficient yeast auxotrophs have been reported [70]. Complementing this metabolic deficiency with the antibody catalyst would provide a means of identifying more efficient variants. 5.2.2 Other Systems Large rate accelerations are also expected when charged reactants are enthalpically destabilized relative to a charge delocalized transition state by desolvation [20]. Such a mechanism may contribute to the efficacy of enzymes that promote biologically relevant decarboxylations [71–74]. Antibody catalysis of model reactions, such as the solvent-sensitive decarboxylation of 3-carboxy-benzisoxazoles to salicylonitriles [75, 76] and the difficult decarboxylation of orotate to uracil [77], has been used to probe the role of medium effects in a tailored binding pocket. Significant rate enhancements have been achieved, and structural work characterizing the properties of successful active sites will be instructive. Given the well-established and often dramatic sensitivity to solvent change of many reaction types, medium effects are likely to be pervasive in antibody and enzymatic catalysis. Reactions that display

17


large changes in charge localization, including decarboxylations, nucleophilic substitutions, and aldol condensations, should be especially amenable to such effects. 5.3 Electrostatic Catalysis 5.3.1 Acyl Transfer Reactions Enzymes frequently utilize hydrogen bonding and charged groups to stabilize polar transition states electrostatically [69]. When haptens containing positive and negative charges are used, electrostatic interactions can also be exploited for antibody catalysis. For example, anionic phosphonates and phosphonamidates, originally designed as potent inhibitors of hydrolytic enzymes [12, 78], have been useful in the production of antibodies that hydrolyze esters, carbonates and (more rarely) amides [79]. Such analogs resemble the transition state for hydrolysis in a number of ways, including tetrahedral geometry, negative charge, and increased bond lengths. How these features are reflected inthe induced binding pockets has been deduced through structural studies of six different esterase families [80–90]. Antibody 48G7 is typical of this class of catalyst [91]. It was generated against p-nitrophenyl phosphonate (6) and accelerates the hydrolysis of the corresponding activated ester 7a and carbonate 7b by factors of 1.6 × 104 and 4 × 104 , respectively (Fig. 11). Both the antibody and its germline precursor, with and without bound hapten, have been characterized to provide insight into the origins and evolution of its catalytic effects [85–87]. The mature antibody has a deep, well-defined combining site rich in aromatic residues (Fig. 12a). It binds the hapten in an extended conformation with the aryl group at the bottom of a hydrophobic cleft formed between CDRs L1 and H3. The negatively charged phosphonate moiety lies near the pocket entrance, where it forms multiple interactions with charged and neutral antibody residues. The pro-R phosphonyl oxygen hydrogen bonds with the TyrH33 phenolic hydroxyl group and forms a salt bridge with the ArgL96 guanidinium group, while the pro-S

Fig. 11 Hydrolysis of activated aryl esters 7a and carbonates 7b proceeds via an anionic and tetrahedral intermediate (in brackets). Hydrolytic antibody 48G7 was elicited with an aryl phosphonate derivative (6) that mimics this high-energy species and its flanking transition states [91].


oxygen hydrogen bonds to the HisH35 -imino group and the TyrH96 backbone NH. Comparison of the mature antibody with and without the transition-state analog shows no major conformational changes [86], suggesting a simple lock-and-key mechanism for hapten binding (see, however, [92]). Antibody 48G7 has a 30 000-fold higher affinity for the phosphonate hapten and 20-fold greater catalytic efficiency than its germline precursor. Nine somatic mutations, all lying outside the combining site, are responsible for these effects [85, 87, 93]. Although not in direct contact with bound ligand, the mutated residues appear to preorganize the pocket for binding and catalysis. Improved packing and secondary hydrogen-bonding interactions help limit side-chain and backbone flexibility inherent in the germline protein. In fact, germline 48G7 (Fig. 12b) is conformationally much more flexible than the mature antibody and undergoes significant reorganization upon hapten binding [87]. The hapten itself binds quite differently in the two complexes (Fig. 12a,b). The phosphonate moiety occupies essentially the same location in both, but it cannot form a hydrogen bond with TyrH33 in the germline complex because of an altered conformation of the CDR H1 loop. Further, the p-nitrophenyl group is rotated away from the position it adopts in the mature antibody to occupy a hydrophobic cleft constructed from framework residues. This second apolar pocket is present, but empty, in mature 48G7. The alternative binding mode is made possible by removal of an otherwise repulsive interaction with the nitro group of the hapten in the mature antibody by the somatic mutation SerL34 Gly. Concordant with hapten design, the crystallographic data suggest that 48G7 is a relatively simple catalyst which promotes ester hydrolysis by direct attack of hydroxide on the scissile carbonyl. Although the two hapten binding modes seen in the germline and mature 48G7 complexes create some ambiguity about the substrate’s preferred orientation, the side chains of TyrH33 , HisH35 and ArgL96 and the backbone amide of TyrH96 apparently stabilize the tetrahedral and anionic transition states electrostatically through hydrogen-bonding and ionic interactions (Fig. 12). An analogy between this anion binding site and the well-characterized “oxyanion hole” of serine proteases is apparent. Individually, however, the antibody residues are not very effective oxyanion stabilizers. Mutations at positions H33, H35 and L96 cause only 3 to 30-fold reductions in kcat [85]. Loss of the hydrogen bond between TyrH33 and the transition state probably contributes to the germline’s 20fold lower efficiency, as well. In contrast, replacement of a single asparagine in the oxyanion hole of the protease subtilisin results in 102 to 103 -fold losses in specific activity [94, 95]. High solvent accessibility and/or conformational mobility of the antibody residues may account for their limited efficacy. The 48G7 active-site structure and hapten-recognition properties are strikingly similar to those of other, independently-derived anti-phosphonate antibodies [96]. These include three catalysts (CNJ206 [80, 81], 17E8 [89] and 29G11 [90]) for the cleavage of p-nitrophenyl esters and three (D2.3, D2.4 and D2.5 [83, 84]) for the energetically more demanding hydrolysis of p-nitrobenzyl esters. Although many of these esterolytic antibodies derive from different germline sequences, they appear to have in common a deep hydrophobic pocket in the framework region into

19


Fig. 12 Stereoview of the active site of mature 48G7 (a) and its germline precursor (b), showing the different orientation of the hapten in the two complexes [85–87]. The phosphonate moiety of 6 binds in roughly the same location in both, although the network of hydrogen-bonding interactions that

constitute the oxyanion hole is slightly different. The deeply bound aryl group binds in one of two available hydrophobic clefts at the bottom of the pocket depending on whether the residue at position L34 is Ser (germline) or Gly (mature).

which the leaving group’s aryl/benzyl ring binds (Fig. 13). All exploit multiple hydrogen bonds and salt bridges near the mouth of the cavity for recognition of the phosphonate moiety. Consequently, it is likely that all employ the same basic hydrolytic mechanism as 43G7 [96]. Given this, their comparative efficiency, which ranges over two orders of magnitude, must reflect differing abilities to stabilize the hydrolytic transition state relative to the bound ground state. Examination of the hapten complexes suggests that the rate enhancement (kcat /kuncat = 103 – 105 ) roughly parallels the number of hydrogen bonds to the phosphonate. Flexibility in the active site (as seen in the 48G7 germline antibody) also correlates with low efficiency [80, 81, 87]. In some cases, the antibodies may exploit binding energy to


Fig. 13 Overlay of three hydrolytic antibodies, 48G7 (green), 17E8 (blue), and CNJ206 (red), shows remarkable structural convergence in these independently-generated active sites [96]. Only haptencontacting side chains are illustrated; the purple balls indicate the position of the phosphorus of the respective bound hapten.

distort the substrate ester from its thermodynamically favorable Z-conformation, making it easier to reach the tetrahedral transition state during catalysis [82, 85]. For example, this factor may come into play in 48G7, depending on whether the substrate binds like the hapten in the mature or the germline complex (Fig. 12). The remarkable degree of structural convergence observed in antibodies selected for tight binding to aryl phosphonate transition-state analog finds parallels in a number of other systems. For example, the immune responses to phosphorylcholine [97], p-azophenylarsonates [98], and 2-phenyloxazolones [99] are dominated by specific combinations of heavy and light chain variable regions. Interestingly, when a phenyloxazolone-binding antibody was found with a unique VH domain, its structure showed conservation of important antigen binding residues [100]. These results point to the immune system’s utilizing a relatively limited number of mechanisms to recognize any given type of antigen. Apparently, strong selective pressure reduces the broad diversity initially present in the primary repertoire to a small set of “best” solutions that can be further optimized by somatic mutation. As discussed above for the Diels-Alderases, structural convergence is even evident in antibodies raised against unrelated haptens. Rather than possessing an infinite variety of differently configured active sites, the immune system appears to play with a limited deck. Variations within the general theme do arise. Another esterase that has been structurally characterized, antibody 6D6, catalyzes the hydrolysis of a chloramphenical monoester with relatively low efficiency (kcat /kuncat = 900) [101]. It shows specific

21


differences from the other antibodies [82]. In particular, the hapten is bound more shallowly, although the stacked aromatic rings of the leaving group and the acyl side chain are the most deeply buried portions of the molecule. The tetrahedral phos-phonate is highly solvent exposed, forming only one hydrogen bond to the antibody, presumably explaining 6D6’s relatively low efficiency. At this point,itisnot clear howmuch activity can be obtained from antiphosphonate antibodies. Over 50 esterolytic antibodies have been showntohave properties roughly comparable to those discussed here: simple hydrolytic mechanisms, rate accelerations up to 105 over background, and chemical proficiencies of 107 − 108 M−1 [15, 16]. While notable, particularly considering concomitant high and predictable selectivity [3], such effects are still orders of magnitude lower than those achieved by analogous enzymes. The immunological approach appears to have hit a ceiling in catalytic efficiency. It seems likely that these activities reflect what can be achieved with an “oxyanion hole” mechanism alone. Recent attempts to augment activity by rational mutagenesis or affinity selection with phage-displayed antibody fragments have met with only limited success [102–105]. Negative results should not be overinterpreted, but theydo raise concerns that the aryl phosphonate binding pocket common to these catalysts is not an intermediate that can be further refined but an evolutionary dead end. Alternative antigen presentation strategies and in vitro selection methods may provide access to different subsets of the immune repertoire more amenable to optimization. For example, antibodies that bind tetrahedral anions not at the entrance to the combining site but deep within their active site, as seen for enzymes that promote hydrolyses, would be of interest. Of course, highly evolved hydrolytic enzymes are more than simple oxyanion holes. They exploit arrays of catalytic groups (and, often, metal cofactors) to catalyze energetically demanding reactions such as amide hydrolysis. Induction of several precisely aligned functional groups in a single immunization step is extremely improbable, however. Such arrays are unlikely to be present in the primary repertoire of the immune system, and, depending on the basic immunoglobulin scaffold, may not be accessible through somatic hypermutation. Occasionally, though, serendipitous mutations that open up new opportunities for catalysis can and do occur. Although phosphonate transition-state analogs specify a simple hydrolytic mechanism, some anti-phosphonate antibodies have been identified that exploit more complex mechanisms. For example, the lipase-like antibody 21H3, generated against a typical benzyl phosphonate ester [106], unexpectedly accelerates ester hydrolysis by a two-step mechanism involving transient acylation of an amino acid within the binding pocket [107]. Its efficiency at hydrolyzing esters is no greater than that of the esterolytic antibodies discussed above, but use of covalent catalysis makes possible stereoselective transesterification reactions that cannot be carried out in water without the antibody [107, 108]. Similarly, considerable biochemical evidence has been adduced for a two-step sequence involving an acyl-antibody intermediate in hydrolytic reactions catalyzed by antibody 43C9 [109]. The latter was generated against phosphonamidate 8 [110], rather than a phosphonate, and it is


Fig. 14 Phosphonamidate 8, which is a transition-state analog for amide hydrolysis, yielded antibody 43C9 [110]. This antibody cleaves structurally related p-nitroanilides 9 and esters (not shown).

unique in its ability to promote the hydrolysis of activated amides as well as esters (Fig. 14). A 2.5 × 105 -fold rate enhancement (chemical proficiency = 5 × 108 M−1 ) achieved in the cleavage of p-nitroanilide 9 makes 43C9 one of the most efficient hydrolytic antibodies known. The crystal structures of free 43C9 and its complex with p-nitrophenol were recently solved [88]. Although detailed understanding of interactions specific to ligand recognition and catalysis await characterization of the hapten complex, likely participants in the reaction are identifiable upon inspection of the binding pocket (Fig. 15). As with other hydrolytic antibodies, several residues are available for stabilizing the negative charge that develops in the transition state, many of which are shared with 48G7. These include HisH35 and ArgL96 , whose importance for catalysis is supported by the results of site-directed mutagenesis experiments [103]. Residue AsnH33 , like 48G7’s TyrH33 , may also play a role in transition-state stabilization. What distinguishes 43C9 from other esterolytic antibodies, though, is the presence of a second histidine at position L91 . HisL91 is directed into the pocket toward the region where substrate must bind (Fig. 15). Docking experiments have suggested

Fig. 15 Stereoview of the active site of amidase 43C9 with bound p-nitrophenol [88]. The imidazole side chain of HisL91 (green) points into the pocket toward the region normally occupied by the phosphonate in

other hydrolytic antibodies. AsnH33 , HisH35 and ArgL96 are potential oxyanion stabilizers. Two active-site water molecules are shown as red spheres.

23


a plausible orientation of the substrate, placing its scissile carbonyl in an excellent position for nucleophilic attack by this residue’s imidazole side chain [88]. Detailed mechanistic inferences should be considered tentative in the light of likely perturbations to the binding pocket caused by packing interactions between the active sites of adjacent molecules in the crystal [88]. Nevertheless, mutagenesis of HisL91 to Gln decreases catalytic efficiency >50-fold with little effect on ligand binding, supporting its role as the nucleophile that is transiently acylated during catalysis [103]. Consistent with this possibility, the acyl intermediate detected by electrospray mass spectrometry at pH 5.9 is not observed with the HisL91 Gln variant [111]. Covalent catalysis by imidazole is well-established in non-enzymatic reactions of carboxylic acid derivatives [1]. Its efficiency is a consequence of imidazole’s high nucleophilicity and the relative instability of the acyl-imidazole intermediate. The presence of HisL91 can thus explain why 43C9, but not 48G7 or other esterolytic antibodies, cleaves an amide. Context is clearly relevant, since lipase 21H3 lacks amidase activity though it also exploits nucleophilic catalysis [106, 107]. The fact that 43C9’s mechanism is unspecified by its hapten’s design also underscores the importance of serendipity in these experiments. The rarity of such occurrences is reflected in many failed experiments to generate amidases with phosphonamidate haptens. Despite its relative mechanistic sophistication, 43C9 is still a primitive amidase when compared with a typical protease. Subtilisin, which also exploits nucleophilic catalysis, catalyzes the cleavage of succinyl-Ala-Ala-Pro-Phe-p-nitroanilide 104 times more efficiently than 43C9. Its rate acceleration is 3.9 × 109 over background and its chemical proficiency is 2.2 × 1013 M−1 [112]. Moreover, subtilisin can cleave unacti-vated amides, whereas 43C9 is restricted to substrates with leaving groups of pK a < ca. 12 [113]. The high efficiency of covalent catalysis in serine proteases like subtilisin derives from cooperative action of several functional groups in addition to the active site nucleophile. Acid-base chemistry, in particular, is used to activate the serine nucleophile, facilitate proton transfers, and stabilize the amide leaving group. Removal of any single component of the protease’s catalytic triad results in 104 to 106 -fold decreases in activity [112]. Unsurprisingly, 43C9 lacks an analogously complex catalytic machinery. Although it has proven difficult to install additional functional groups by mutagenesis [103], more productive changes may become obvious when the structure of the hapten complex is known. Lessons from the immunological evolution of 48G7 [53, 87] suggest that mutagenesis of residues outside the binding pocket will be required to fine-tune critical synergies between participants in catalysis. 5.4 Functional Groups

Functional groups – acids, bases, and even exogenous cofactors — can clearly extend the capabilities of catalytic antibodies. They will certainly be needed if reactions with large activation barriers are to be significantly accelerated. As we have seen in the case of amidase 43C9, unplanned but useful residues can appear by chance in the


immunoglobulin pocket during affinity maturation. To increase the probability of eliciting such functional groups where they are needed, several strategies have been devised. Charge complementarity between an antibody and its ligand is the easiest principle to exploit in generating functionalized binding pockets. Cationic haptens have been used, for instance, to elicit negatively charged carboxylates that can serve as bases or nucleophiles or as general acids when protonated. Elimination reactions, epoxide ring openings, cationic cyclizations, and hydrolyses of esters, ketals, and enol ethers have been successfully catalyzed by this approach [114]. In favorable cases, very large catalytic effects have been achieved. For example, antibodies raised against a protonated benzimidazolium derivative use an active-site Asp or Glu to deprotonate substituted benzisoxazoles with effective molarities of 40 000 M and rate accelerations in excess of 108 over the acetate-promoted background reaction [115]. Nevertheless, it is unlikely that a single hapten will ever elicit arrays of residues as sophisticated as those present in highly evolved enzymes. Heterologous immunization has been explored as a method of circumventing this limitation [116]. In this procedure, two molecules, each containing different functional groups, are serially used as haptens to elicit the immune response. Ideally, a subset of the resulting antibodies will possess multiple catalytic groups induced in response to both tem-plating molecules and have, as a result, enhanced activities. An important advantage of this strategy is that simplified haptens can be used, reducing the need for laborious synthesis. Although generality must still be established, results from initial studies indicate that antibody esterases generated by heterologous immunization are more efficient than those generated in response to the individual haptens [116, 117]. Mechanism-based enzyme inhibitors [17, 18] are potentially of even greater utility than haptens [19, 118–122]. Such molecules exploit a protein’s ability to initiate a cascade of events, ultimately leading to its own covalent modification. This irreversible chemical reaction thus provides a means of selecting immunoglobulins in vivo and/or in vitro on the basis of their activity. When the selected antibody is challenged with substrate rather than hapten, the same group(s) responsible for protein modification can be used to promote the desired chemical transformation. Covalent catalysis can thus be specified through hapten design. The potential of reactive immunization is perhaps best illustrated by the production of aldolase antibodies [120]. These catalysts not only utilize a complex chemical mechanism but are among the most efficient catalytic antibodies described to date.

5.4.1 Aldolases Aldol condensations are broadly useful carbon-carbon bond-forming reactions in organic synthesis. They pose special difficulties for biocatalysts, however, because they proceed via a series of consecutive transition states, each requiring acid-base catalysis. Class I aldolases solve these problems by using a reactive lysine to activate the ketone donor through Schiff base formation at the active site [123–125]. Deprotonation of the Schiff base yields an enamine that then adds stereoselectively

25


Fig. 16 $-Diketone 10, used as a hapten to raise antibodies 38C2 and 33F12, traps a reactive, active-site amine to form a stable, chromophoric vinylogous amide [120]. These antibodies promote diverse aldol

condensations. In the example shown (bottom), the two products are formed in a ca. 1:1 ratio with the indicated diastereoselectivities.

to the acceptor aldehyde to form a new carbon-carbon bond. Subsequent hydrolysis of the Schiff base releases product and regenerates the active catalyst. β-Diketones inhibit class I adolases by forming a Schiff base with the active-site lysine and then rearranging to a more stable vinylogous amide. When β-diketone 10 was used as a hapten instead of a more conventional transition-state analog [120], two analogously modifiable antibodies (33F12 and 38C2) were obtained (Fig. 16). The reactive group on the antibodies is a lysine with an anomolously low pK a (5.5 for 33F12 and 6.0 for 38C2 [123]). The vinylogous amide it forms with the hapten (λmax 316 nm, 15 000 M−1 cm−1 ) can be irreversibly trapped by reduction with sodium cyanoborohydride. These same antibodies also mimic the activity of class I aldolases [120]. Their reactive lysine reacts with a wide range of ketones to form enamine adducts. These condense with diverse aldehydes to form aldol products. The structure of antibody aldolase 33F12 [123] shows the catalytic lysine (LysH93 ) buried at the bottom of a hydrophobic pocket (Fig. 17). It is not hydrogen bonded with other amino acids and there are no charged residues within 7Å. The absence of such interactions must be responsible for this group’s unusual reactivity: the hydrophobic microenvironment lowers the amine’s pK a by disfavoring the protonated state. In contrast, class I aldolases are believed to increase the acidity of their catalytically essential lysine through proximity to several positively charged residues [126]. Unfortunately, the unliganded antibody provides few clues about the interactions that stabilize the transition states for formation and breakdown of the carbinol amine, deprotonation to afford the enamine, and creation of the new carbon-carbon bond. Sequestered water molecules or the hydroxyl groups of Tyr or Ser residues


Fig. 17 The binding pocket of antibody 33F12 [123] is seen through a slice in the molecular surface calculated with a sphere of 1.4 Å radius [206]. Only the g-amino group of LysH93 contacts the molecular surface at the bottom of the antigen binding site.

that constitute the pocket walls may be involved. For instance, a simple rotation of the Lys side chain from its position in the unliganded antibody would bring its amino group near SerH100 , which lies on the opposite side of the pocket. Structures of enamine adducts will be important for clarifying these points. An unanticipated feature of the aldolase antibodies is their promiscuity. Over 100 different reactions, including aldehyde-aldehyde, ketone-aldehyde and ketoneketone condensations, are subject to catalysis [123]. Because the 33F12 pocket is relatively hydrophobic, polyhydroxylated aldehydes, such as glyceraldehyde, glucose, and ribose, which are good acceptors for natural aldolases, are not substrates for the antibodies. Aside from this restriction, a wide range of donors and acceptors is tolerated. Non-specific van der Waals interactions likely provide the driving force for sequestering the first substrate. Once bound, it encounters the reactive lysine and forms the nucleophilic enamine. Provided there are no steric clashes, similar interactions should allow the aldehyde acceptor to bind and undergo aldol addition. The binding site is 11 Å deep and quite capacious, much larger in fact than the β-diketone that induced it, accounting for this broad specificity. These properties have been rationalized as a consequence of the reactive immunization process itself [123]: capture of the antibody by a covalent chemical event early in the process of affinity maturation may obviate the need for further refinement of the binding pocket by somatic mutation. Although induced with an achiral hapten and despite their broad substrate specificity, these aldolase antibodies are surprisingly stereoselective (Fig. 16). When acetone is the donor substrate, addition preferentially occurs on the si-face of the aldehyde acceptor; with hydroxy acetone, attack occurs on the re-face. In most cases, enantiomeric excesses >95% are found [127]. Enantioselective Robinson annulations [128], resolution of tertiary aldols [129], and preparation of chiral intermediates for the total synthesis of brevicomins [130] and epothilones [5] illustrate the synthetic utility of these catalysts. Kinetic resolution of the epothilone precursor

27


was carried out on a gram scale using 0.06 mol% of antibody 38C2 (corresponding to 0.5 g of IgG). The reaction proceeded with good yield (37%) and high enantiomeric excess (90%). More recently, antibody 38C2 has been used to activate prodrugs [6]. It catalyzes the selective removal of generic drug-masking groups via sequential retro-aldol and retro-Michael reactions. This advance could prove useful in the development of selective chemotherapeutic strategies. How do these remarkable antibody aldolases compare to natural aldolases? To address this question, it is easiest to consider representative retroaldol reactions. For such transformations, the antibodies achieve turnover numbers ranging from 0.0003 to 0.08 s−1 [127]. One of the best antibody substrates is 4-(4 isobutyramidophenyl)-4-hydroxy-2-butanone. In the presence of 38C2, it undergoes retroaldolization with kcat and kcat /K m values of 0.083 min−1 and 3.3 × 103 M−1 s−1 , respectively. The rate constant for the background reaction is 1.4 × 10−9 s−1 (aqueous buffer, pH 7 and 25 ◦ C). The antibody thus accelerates this reaction by a factor of 2 × 107 -fold; its chemical proficiency is 6 × 1010 M−1 . These are impressive effects for a catalytic antibody, and may be contrasted with activity accruing to FDP aldolase and KDPG aldolase, typical class I enzymes involved in sugar metabolism. FDP aldolase catalyzes the interconversion of fructose-1,6-diphosphate (FDP) to give dihydroxyacetone phosphate and glyceraldehyde-3-phosphate. Its steady state kinetic parameters in the cleavage direction are kcat = 48 s−1 and kcat /K m = 1.6 × 107 M−1 s−1 [131]. KDPG aldolase, which cleaves 2-keto-3-deoxy-6-phosphogluconate (KDPG) into pyruvate and glyceraldehyde-3-phosphate, is even more active, having kcat and kcat /K m values of 290 s−1 and 4.0 × 10 M−1 s−1 , respectively [132]. Although rate constants for the corresponding uncatalyzed reactions are unavailable, taking 1.4 × 10−9 s−1 as a conservative estimate [127] yields rate enhancements of ca. 1010 − 1011 and chemical proficiencies of ca. 1015 –1016 M−1 . By either criterion, the natural enzymes are several orders of magnitude more efficient than the antibody aldolases. Natural aldolases are more restrictive in their substrate requirements than the antibodies, though they could lose considerable activity and still be competitive. For example, removal of a single phosphate in FDP causes a 50-fold loss in rate with FDP aldolase [131]. The extensive interactions that confer such specificity are almost certainly coupled to the enzyme’s high efficiency. By further refining the antibody pocket tothe precise steric and electronic demandsofaparticular aldol reaction, while maintaining the active-site lysine’s high reactivity, enzyme levels of activity may be attainable. Narrowed scope may be the unavoidable cost of truly high efficiency, however. 6 Perspectives 6.1 General Lessons from Comparisons of Enzymes and Antibodies

Structural and mechanistic work reviewed here reveals many notable parallels between antibodies and their more highly evolved counterparts. Not only are the

6 Perspectives

sizes and shapes of their active sites comparable, but antibodies and enzymes utilize the same set of molecular interactions to bind their respective ligands and stabilize transition states. Although antibodies tend to be less extensively functionalized than enzymes, the basic mechanistic strategies they employ to lower kinetic barriers are strikingly similar. As more primitive catalysts, however, they provide an alternative vantage point for examining the relationship between binding energy and catalysis. In this regard, simplicity is a virtue. Rather than working backwards from a fully evolved enzyme, uncomplicated, tailored model systems can be constructed to illuminate specific mechanistic questions. As multiple mechanistic strategies are combined to augment efficiency, valuable insight into the evolution of catalytic function can be gained. Functional analysis of the antibody intermediates that arise during affinity maturation [38, 53, 85, 87] also sheds light on these issues. Currently, antibodies appear less successful than enzymes intheir ability to achieve the fine level of recognition required for optimal discrimination between transition states and ground states. Their modest efficiencies appear to be a direct consequence of the simple strategy used to generate them. While the process of natural selection optimizes enzymes on the basis of their catalytic activity, the immune system’s microevolutionary mechanisms select antibodies for increased affinity to an imperfect transition-state analog. It is unrealistic toexpect that proteins engineered to recognize such haptens will provide an ideal steric and electrostatic environment for chemical transformation. Even with a perfect transition-state analog, the chances of obtaining a fully evolved catalyst through immunization would be low. As noted above, there is generally insufficient selection pressure to attain the high binding energies that characterize complexes between true enzymes and their transition states. On both micro and macro levels, mechanistic improvements arise as a function of time, so differences in time scales for the evolution of enzymes and antibodies – millions of years versus weeks or months – also come in to play. Although Nature uses a wide variety of different protein scaffolds to build enzyme active sites [133], she does not seem to have adopted the immunoglobulin fold. It is therefore conceivable that antibody structure itself places intrinsic limitations on the kind of reactions amenable to catalysis and on attainable efficiencies. In general, though, structural studies show excellent shape and chemical complementarity between antibodies and their ligands. Depending on the hapten, deep pockets, clefts, grooves, and flatter, more undulating surfaces can be created [134, 135]. Because certain classes of haptens tend to be recognized in the same way [52, 53, 83, 96], structural diversity must be considerably more restricted than might have been expected given 108 variants available in the primary immune repertoire [136, 137], but whether these consensus sites significantly restrict the catalytic capabilities of antibodies is still unclear. Conformational flexibility is another potential concern. Protein conformational changes in enzymes provide a means of excluding water from the active site and enable the catalyst to adjust tochanges in substrate as the reaction coordinate is traversed [138, 139]. Antibodies are known to undergo a comparable range of ligandinduced conformational changes, including alterations in side chain rotamers, segmental movements of hypervariable loops and changes in the relative disposition

29


of the VH and VL domains [134]. Without direct selection for activity, however, these dynamic effects will be difficult to exploit deliberately for catalysis. In fact, conformational flexibility in catalytic antibodies, when observed [80, 81, 87], usually results in lower rather than higher efficiency. 6.2 How efficient does catalysis need to be?

Enzymes represent an extraordinarily high standard against which to judge new catalysts that are rationally designed from simple principles. From the perspective of the chemist, one exciting aspect of catalytic antibody technology is its ability to deliver tailored catalysts for reactions which are difficult to carry out selectively using existing methods or for which natural enzymes do not exist. Must such systems attain enzyme-like efficiency to be useful? Because antibodies are biocompatible and have long serum half-lives, many in vivo applications would be conceivable if sufficient activity were available. In fact, existing catalytic antibodies have already achieved significant effects in biological systems. When expressed at high levels, they have been shown to be competent (if inefficient) catalysts in metabolism, replacing essential enzymes in amino acid [42] and pyrimidine biosynthesis [77] pathways. Therapeutically relevant concentrations of the aldolase antibodies discussed earlierhave beenusedtoactivate prodrugsand kill colon and prostate cell lines [6]. Similarly, an esterolytic antibody has been employed as a cocaine antagonist, protecting rats from cocaine-induced seizures and sudden death [140, 141]. Many chemical reactions cannot proceed in the absence of catalysts because competing pathways have lower energies. In several instances, antibody binding energy has been successfully utilized to alter the course of such reactions by selectively stabilizing the less favorable transition state. For example, antibody catalysts have been developed for normally disfavored syn-eliminations [142], exo rather than endo Diels-Alder cycloadditions [46, 48], and 6-endo-tet ring closures of epoxy alcohols [143, 144]. Antibodies have also been used to control the fate of high-energy intermediates, allowing them to partition along only one of several possible pathways, as in the case of conversion of an enol ether to a cyclic ketal in water [145]. Formation of a strained cyclopropane derivative in a cationic olefin cyclization is another such example [146]. Binding energies up to 5.5 kcal/mol are typically available for achieving such discrimination, and even more energy may be available in favorable cases [15]. Such selectivities could be of great utility in organic synthesis. Assuming an antibody is available for any given transformation, its turnover and cost will ultimately determine whether it is used in practice. Presently, the steady state parameters of typical catalysts necessitate high antibody concentrations (≥ 10 µM = 1.6 mg/ml) and long reaction times to achieve useful conversions [12]. Preparative applications of several antibodies show that gram-scale reactions are feasible, particularly if antibody selectivity is high and competing reactions are substantially slower than the desired transformation [5, 147, 148]. Costs associated with high-volume antibody production are certainly an issue, but some antibodies

6 Perspectives

are now produced on a large scale for diagnostic and medical applications. They are readily obtained in good yield through ascites production [149] or by fermentation in hollow fiber reactors [150, 151]. Their Fab and Fv fragments can often be produced efficiently in plants [152, 153] or microorganisms [154]. Technical advances in microbial fermentation can be expected to make antibody production even more economically favorable in the future. In short, current levels of activity may be adequate for some laboratory applications, but higher efficiencies would certainly be beneficial. A 103 -fold increase in turnover would mean that 103 -fold less catalyst is needed to achieve useful levels of performance. Enhanced proficiency will certainly be necessary if energetically more demanding reactions are to be tackled. The creation of antibody equivalents of site-specific proteases, glycosidases, and nucleases, for example, remains a significant yet unrealized goal. The use of antibodies to synthesize or modify structurally complex and biologically important macromolecules will depend on solving this basic problem. 6.3 Strategies for Optimizing Efficiency

If imperfect design and indirect selection for binding rather than function are the primary reasons for low catalytic efficiency, creation of substantially better antibody catalysts will be feasible. Conceptually, two approaches can be envisaged: (1) refining methods for producing first generation catalysts, and (2) developing new strategies to optimize existing active sites. Improved transition-state analogs and more effective screening of the immune response address the first point. Rational reengineering and directed evolution methods are relevant to the second. These strategies have already been discussed in the context of specific antibody catalysts but are summarized in more general terms below. 6.3.1 Better Haptens The rewards of good – and the penalties of deficient – design are evident in the properties of catalytic antibodies characterized to date. In general, however, it is not clear which structural features of a transition state are most important to mimic to elicit maximally effective catalysts. Statistically meaningful correlation of different hapten types and the properties of their complementary active sites are needed to optimize analogs for each type of reaction. Incorporating design features that maximize transition-state affinity while minimizing ground-state stabilization remains a major challenge. For example, aryl groups are constituents of many haptens. Binding energy directed toward them, while increasing hapten affinity, may be useless or even harmful for catalysis, since these elements are common to both ground and transition state. Inefficient utilization of intrinsic binding energy in this way may help to explain the modest activities seen in the oxy-Cope [38] and Diels-Alder reactions [53] discussed above. The finding that catalytic activity for a series of polyclonal esterases correlates inversely with the size and hydrophobicity of the haptenic aryl phosphates [155] also illustrates this

31


problem. When binding energy is used to recognize parts of the substrate distant from the site of reaction, product inhibition becomes another concern. Strategies to facilitate product release must therefore be considered integral to hapten design. Antigen presentation is a further issue. Small molecules are not immunogenic and mustbe coupledto carrier proteins. Variation of the tether site maybe a useful means of focusing immunorecognition to a hapten’s catalytically relevant epitopes [141]. Linkage of chemical reactivity with the immune system’s selection and amplification processes may mitigate some limitations in design. For this reason, the reactive immunization strategy [19, 118–122] merits increased attention. Many additional examples will be needed to establish its true scope and limitations. Given their importance in natural enzymes[156], metal ions and exogenous organic cofactors should considerably extend the properties of antibody catalysts, as well. Although versatile hybrid catalysts that combine the intrinsic reactivity of the metal ion or coenzyme with the tailored binding specificity of the antibody can be readily envisaged, this strategy has been surprisingly underutilized. Metal ion binding sites have been engineered into antibody binding sites [157], creating a sensitive Zn(II) sensor in one case [158], but catalysis has not been realized. An alternative, seemingly promising strategy using metal chelates for peptide cleavage has received little attention since first reported [159]. Modest activities have been described for other miscellaneous cofactor-dependent reactions [158, 160–163], but more work is obviously needed. Such strategies will be very important for promoting reactions with high kinetic barriers and reactions that cannot be carried out with protein residues alone. 6.3.2 Screening Because unusual germline sequences or fortuitous mutations may be necessary for high activity, the best antibody catalysts are also potentially the rarest. For this reason, more extensive screening of the immune response may dramatically increase the probability of finding highly active clones. Usually, small panels of antibodies chosen for their ability to bind the transition-state analog are purified and tested individually for catalytic activity. This procedure is necessarily indirect and slow. Sensitive chemical [164], biological [42, 77, 91, 165] and immunological assays [60, 166, 167] can facilitate the screening of thousands of candidates directly for catalysis and thereby accelerate the preliminary evaluation process. One practical problem associated with broad screening is that some antigens yield many hapten binders, others relatively few. To increase the size and diversity of the antibody population available for testing, multiple fusions can be performed and several mouse strains utilized for immunization. Mice prone to autoimmunity have been shown to yield unusually large numbers of esterolytic antibodies and may prove more generally useful for expanding the repertoire of catalytic clones elicited by a single transition-state analog [168]. Significant progress has also been made in copying the combinatorial processes of the immune system in vitro. Libraries of antibody fragments containing more than 106 members can be constructed and produced in microorganisms or

6 Perspectives

displayed on phage particles [169–172]. These systems are attractive vehicles for exploring the catalytic potential of different subsets of the primary immunological repertoire. Binders can be selected from these libraries on the basis of hapten affinity and subsequently screened for catalytic activity [169]. Clever strategies for capturing active clones based on their activity should be even more effective [173– 177]. Alternatively, catalysts can be obtained directly by selection in vivo using yeast or bacterial auxotrophs [42, 77, 91]. In these approaches, iterative rounds of mutagenesis and reselection replace somatic mutation as a means of refining initial hits [170]. Domain swapping [178] and powerful DNA shuffling methods [179, 180] have been developed to speed up this process. 6.3.3 Engineering The upper limit on activity that can be achieved with antibodies is unknown and may be reaction dependent. It is therefore important to push several test cases as far as possible. Site-directed mutagenesis is an attractive tool for improving catalytic power, particularly given the availability of increasing numbers of high-resolution structures. In general, it will probably be easiest to reengineer the poorest catalysts, since changes that improve packing or provide missing but critical interactions may be relatively obvious. The mutational study leading to an order of magnitude increase in activity of the Diels-Alderase antibody 39-A11 is a case in point [54]. The fact that relatively few changes are needed to tailor the properties of germline structure during affinity maturation [53, 85, 93] is also encouraging. Pinpointing subtle changes needed to optimize more active clones is likely to be more difficult, however. The obstacles encountered in augmenting the activity of hydrolytic antibodies sound a cautionary note [103–105]. Ultimately, our understanding of will determine what can be achieved in this way. 6.3.4 Selection Enzymes have been brought to peak efficiency over millions of years by the process of natural selection. An analogous process in the laboratory, involving recursive cycles of mutagenesis and genetic selection for function, may provide the ultimate test of the capabilities of antibody catalysts. Evolution of antibodies can be accomplished perhaps most directly by complementation of auxotrophic yeast or bacterial strains. The chorismate mutase antibody 1F7 [42] and an orotate decarboxylase antibody [77] have been shown to confer a significant growth advantage under selective conditions to host cells lacking the corresponding enzymes. Though the experiments are technically difficult because of poor antibody expression in microorganisms, preliminary results with the chorismate mutase antibody have demonstrated the feasibility of selecting antibodies with novel properties [43]. Recent work showing that cytoplasmic production of antibody fragments is optimizable by selection [181] augers well for these efforts. Selection systems are available for many transformations, including the ferrochelatase and metabolic reactions already mentioned [42, 70, 77]. A generalized selection scheme for hydrolytic reactions has also been reported [91]; analogous assays could be developed to exploit the ability of a catalyst to synthesize or destroy nutrients, drugs, hormones, or toxins. In addition to

33


providing clues about the perfectibility of catalytic antibodies, such experiments may yield fundamental insights into structure-activity relationships and the evolution of molecular function. 6.3.5 Other Scaffolds The immune system was originally tapped as a source of catalysts as a matter of convenience: it is unrivaled in its ability to fashion high affinity protein receptors – the antibodies – to virtually any natural or synthetic molecule, essentially on demand [58, 136, 137]. However, now that the combinatorial processes of the immune system can be mimicked in vitro and libraries of macromolecules can be generated relatively easily using the tools of molecular biology, there is no compelling need to restrict Jencks’s original strategy to a single protein fold or even to a single class of macromolecule. Indeed, in analogy to catalytic antibody experiments, catalytic RNAs and DNAs have been obtained from large libraries of nucleic acids by selection for binding to transition-state analogs. Catalysts for porphyrin metalation obtained in this way [182–184] have activity comparable to that of the ferrochelatase antibody 7G12 discussed above, but an RNA rotamase is 30 times less effective than its antibody counterpart [185]. The lower activity of the latter largely reflects the RNA’s lower affinity for the hapten used (K d = 7 µM, compared with 0.21 µM for the antibody). A similar explanation has been invoked for RNAs that bind the 1E9 hapten (compound 4, Fig. 6) but fail to catalyze the corresponding Diels-Alder reaction [186]. Nucleic acids are likely to be intrinsically more limited than proteins in their capacity for high affinity molecular recognition of structurally diverse ligands as well as for catalysis [187]. Direct selection for function rather than transition-state analog binding has proven to be a much more powerful approach for obtaining nucleic acids with novel catalytic properties (for a recent comprehensive review, see [188]). RNA and have been prepared in this way for a variety of reactions, including a Diels-Alder cycloaddition [189]. Although the resulting catalysts often have relatively modest efficiency, phosphoryl transfers, which are difficult to achieve with antibodies because of the dearth of stable analogs of the pentacoordinate transition state, have been particularly amenable to catalysis. One of the most impressive accomplishments in this regard is the selection of highly efficient ribozymes capable of a self-ligation reaction from large pools of random sequence [190]. The number of starting molecules in these experiments was huge (ca. 1015 ), dwarfing the diversity of the primary immune repertoire (ca. 108 molecules), allowing even extremely rare catalysts to be found. One of the ribozymes obtained by selection was subsequently reengineered to function as a true catalyst; it promotes an intermolecular ligation with multiple turnovers and a rate acceleration approaching 109 [191]. This impressive activity is higher than any seen for most antibody catalysts, and provides an impressive demonstration of the power of direct selection. In principle, in vitro selection of peptides and proteins from vast combinatorial libraries is now possible as well using ribosome display [192–194], mRNAprotein fusion methods [195, 196] and more established phage display formats [172].

References

Coupled with efficient ways of linking genotype with phenotype [174–176], these methods can be expected to facilitate the production of proteins with novel properties and functions. As such, they powerfully complement other efforts to harness the power of evolution to redesign the structures and activities of existing enzymes [197–199, 207, 208].

7 Conclusions

Catalytic antibody technology combines programmable design with the combinatorial diversity of the immune system. This fusion has allowed the field to progress in relatively short order from simple model reactions to complex multistep processes, but much remains to be learned. Early efforts focused largely on defining the scope and limitations of this technology. Now that the approach is well established, attention must be paid to strategies for optimizing catalytic efficiency and for promoting more demanding transformations. In many ways, these are far greater challenges than identifying first-generation catalysts with modestactivity. Continued mechanistic and structural analysis of these systems will inform such endeavors. In addition, learning how to create, manipulate and evolve large combinatorial libraries of proteins outside the immune system should help to automate the processes of catalyst discovery and optimization.

Acknowledgements

The author is indebted to Kinya Hotta for creating the graphics for this article and to the National Institutes of Health, the ETH Zurich, the Swiss National Science Foundation, and Novartis Pharma for generous support.

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41

1

Structure and Function of Catalytic Antibodies Nicholas A. Larsen Harvard Medical School, Boston, USA

Ian A. Wilson The Scripps Research Institute, La Jolla, USA


1 Introduction

An explosion of antibody structures has emerged in the last 15 years [1, 2]. A subset of these antibodies is catalytic, and the latest version of the Protein Data Bank (PDB) (03/2003) contains 56 such structures representing 27 different antibodies (Table 1). Catalytic antibodies are elicited purposely by immunization with a transition state analog. In principle, antibodies that bind such an analog may also stabilize the true transition state of a chemical reaction relative to the ground state, thereby effecting catalysis [3–7]. Remarkably, the immunoglobulin fold possesses the inherent plasticity to harness the usual combination of electrostatic, van der Waals, hydrophobic, hydrogen bond, and cation-π interactions to generate tailor-made catalysts, which, in rare instances, even rival their natural enzyme counterparts. This general plasticity has been exploited to create model systems that explore a number of common and controversial themes in classical enzymology. Thus, substrate strain, approximation, control of the microenvironment, general acid-base chemistry, induced fit, and covalent catalysis have all been systematically examined in the context of the antibody binding pocket. The main objective of structural analysis is to develop hypotheses about key interactions in the binding pocket for binding substrate, transition state (TS), and product, and to propose plausible mechanisms for catalysis. For some antibodies, these hypotheses have been examined rigorously by biochemistry and mutagenesis (Table 1). Mutagenesis studies allow further quantification of the key interactions and provide an avenue for exploring the routes through the energy landscape that confer catalytic potential to the germline antibody precursor. Structural studies of


∗∗)

for diene and dienophile, respectively, equivalent to the so-called effective molarity. ∗∗∗) for substrate and cofactor (periodate), respectively.

∗)

0.28 0.30 0.34 0.08 0.64 0.46 0.05 0.39 0.05 0.004 0.22 3, 10*) 2, 30*) 1, .7* 0.17 0.26 0.27 .04, .25 ***) 0.4 0.36 0.32 0.23 0.21 0.15 0.074 0.051 1.3

3.6 1.0 0.07 0.4 60 220 1500 5.5 0.12 0.11 2.2 0.001 13 40 0.24 1.5 1.4 490 19.2 1.7 0.02 0.02 0.003 0.41 0.02 0.07 0.0016

D2.3 D2.4 D2.5 CNJ206 29G11 17E8 43C9 48G7 6D9 7C8 15A10 13G5 1E9 39-A11 9D9 10F11 33F12 28B4 21D8 5C8 19A4 4C6 1D4 7G12 AZ-28 1F7 4B2

Ester hydrol. pH = 8.3 Ester hydrol. pH = 8.3 Ester hydrol. pH = 8.3 Ester hydrol. pH = 8.0 Ester hydrol. pH = 9.5 Ester hydrol. pH = 9.5 Ester hydrol. pH = 9.3 Ester hydrol. pH = 8.2 Ester hydrol. pH = 8.0 Ester hydrol. pH = 8.0 Ester hydrol. pH = 8.0 Diels-Alder Diels-Alder Diels-Alder Retro-Diels-Alder Retro-Diels-Alder Retro-aldol condensation Sulfide oxidation Decarboxylation Cyclization Cationic cyclization Cationic cyclization Syn-elimination Metal chelatase Oxy-Cope Chorismate mutase Allylic isomerization

kcat (min-1 ) KM(mM)

Antibody Reaction

Table 1 Summary of catalytic antibody structures

2.00 – 2.40 2.15 1.90–2.80 1.61 – 2.10 2.00 – 2.50 2.70 1.30–2.45 1.80–1.85 1.60–2.60 2.00 – 2.80 3.00 1.87

1.85 – 2.10 3.10 2.20 3.20 2.30 2.50 2.20–2.30 1.95–2.70 1.80 2.20 2.35 1.95 1.90 2.10–2.40

x9 1YEC x1 1YED x1 1YEE x1 1KNO x1 1A0Q x1 1EAP x2 43CA x4 1GAF x2 1HYX x1 1CT8 x1 1NJ9 x1 1A3L x1 1C1E x2 1A4K x1 x4 1LO0 x1 1AXT x4 1KEL x2 1C5C x3 25C8 x1 1CF8 x2 1ND0 x2 1JGV x5 3FCT x4 1AXS x1 1FIG x1 1F3D

Yes – – – – Yes Yes Yes – – Yes – – Yes – – Yes Yes Yes – – – – Yes Yes – –

1.3 × 105 3.6 × 104 2.5 × 103 1.6 × 103 2.2 × 103 8.3 × 103 2.7 × 104 1.6 × 104 9.0 × 102 7.1 × 102 2.3 × 104 7 M**) 103M**) 0.4 M**) 4.0 × 102 2.5 × 103 1.7 × 107 9 M**) 2.0 × 104 ND ND ND ND – 5.3 × 103 2.5 × 102 1.5 × 103

1.1 × 105 3.3 × 104 1.3 × 103 0.7 × 102 2.4 × 104 9.0 × 102 6.8 × 104 8.7 × 104 9.0 × 102 1.2 × 10 2.2 × 104 – – – 4.1 × 102 2.5 × 103 – – 7.5 × 103 – – – – – 4.4 × 103 8.5 × 101 4.3 × 102

PDB ID

kcat kuncat -1 KM KDTSA −1 Mutant analysis Res. (Å)

[28] [28] [28] [26] [23] [23, 24] [27] [130] [20, 131] [131] [132, 133] [91] [94] [92, 93] [88, 134] [88, 134] [115–117, 135] [66, 68, 69] [14, 62, 65, 136, 137] [138] [16, 109] [17, 107] [100, 101] [70, 111, 113] [80, 81] [74, 77] [71, 72]

Refs.

2 Structure and Function of Catalytic Antibodies

2 Electrostatic Complementarity

non-catalytic germline precursors further illustrate how the antibodies evolve their catalytic potential by affinity maturation. Such a thorough understanding of these interactions is necessary if improvements are to be made to the antibody catalyst by either mutation or redesign of the immunizing hapten. Importantly, comparison of antibody structures has revealed common structural motifs that are elicited by different haptens during the immune response. Thus, convergent and divergent evolution has been observed directly within the immune system. Most of the catalytic antibody structures found in the are complexes with their respective transition state analog, and provide detailed views of the interactions that contribute to catalysis. In nearly all cases, a high degree of electrostatic and shape complementarity is observed for the corresponding transition state analog (TSA), and, in rarer instances, putative reactive residues that initiate covalent catalysis have been proposed. Consequently, this chapter summarizes antibody-catalyzed reactions in the general context of these three broad headings and concludes with a section on future challenges in the abzyme field for structural biologists and chemists alike.


Many antibody-TSA complexes exhibit a high degree of electrostatic complementarity. Recurring binding motifs for anions (e.g., phosphate, arsenate, sulfate) have been identified, indicative of structural convergence from a variety of germline sequences [8–14]. In contrast, general motifs for binding cations (e.g., primary, secondary, tertiary amines) have not yet been identified. Eliciting an Asp/Glu carboxylate using hapten baiting strategies has met with mixed success. Frequently, cations are buried preferentially in hydrophobic pockets but surrounded by aromatic side chains, which are believed to form stabilizing cation-π interactions [15–17].

2.1 Anionic Binding Motifs and Ester Hydrolysis

Nearly half of the catalytic antibody structures found in the PDB correspond to various hydrolytic antibodies in complex with substrate analogs and products (Table 1). Thus, hydrolytic antibodies are by far the most thoroughly studied catalytic antibodies from both a structural and a biochemical vantage, and have already been reviewed extensively in the literature [18, 19]. These hydrolytic antibodies have been raised generally against either phosphonate or phosphonamidate transition state analogs (Scheme 1). The phosphonate moiety is tetrahedral and carries a negative charge, thereby mimicking the transition state for solvent-mediated hydrolysis that involves nucleophilic attack by a water molecule. With the exception of 6D9 [20] 7C8, and 15A10, these hydrolytic antibodies catalyze the hydrolysis of benzyl or

3


Scheme 1 Hydrolytic Antibodies 48G7, 17E8, 29G11, CNJ206, 43C9, D2.3, D2.4, D2.5

activated 4-nitrophenol or 4-nitrobenzyl esters (Scheme 1), which are particularly convenient model substrates for both synthetic manipulation and sensitivity during hybridoma screening. 2.2 Structural Motifs

The first structural evidence that antibodies recognize antigens by electrostatic complementarity originates from the pioneering study of mAb McPC603, which binds phosphocholine [8, 9]. Further data have indicated that electronically-related pheny-larsonate compounds elicit a conserved anionic binding motif from the immune repertoire [10–13]. Likewise, crystal structures of catalytic antibodies 48G7, 17E8, 29G11, CNJ206, and 43C9 reveal a recurring motif that coordinates the phosphonate moiety of the hapten (Fig. 1a, b) [18, 19]. In general, the constellation of side-chains derived from L96 (CDR3), H33, H35 (CDR1), and the backbone amide of H96 (CDR3) constitute an anionic binding site, arising from structural convergence within the antibody binding pocket (Fig. 1a, b) [18, 19]. For example, antibody 48G7 utilizes ArgL96, TyrH33, HisH35, and the backbone amide of H96 for phosphonate binding (Fig. 1a) [21, 22]. Antibodies 17E8 and 29G11 are related to each other (92% sequence identity [23]) and use the same residues for phosphonate binding [24, 25] as 48G7, except that a hydrogen bond from the LysH93-Nz amine substitutes for the observed TyrH33 hydroxyl in 48G7. Antibody CNJ206 also similarly uses hydrogen bonds from HisH35 and the backbone amides of H96 and H97 to bind the phosphonate [26]. Although a crystal structure of 43C9 in complex with the transition state analog is not available, it is proposed that HisL91, ArgL96, AsnH33, and HisH35 likewise contribute to the anionic binding site (Fig. 1b) [27]. 2.3 Mechanistic Considerations

Clearly, aryl phosphonate/arsonate/sulfonate-like haptens preferentially elicit electrostatic complementarity from the immune response and may explain the commonly observed binding motif observed in 48G7, 17E8, 29G11, CNJ206, 43C9 and other non-catalytic antibodies. For the hydrolytic antibodies, the significance of the

2 Electrostatic Complementarity Fig. 1. Top view of hydrolytic antibodies. The light chain is represented in pink and the heavy chain in blue; side chains are brown and the ligand is yellow. Hydrogen bonds are represented as dashed green lines. A The hapten complex of 48G7 illustrates the prototypic anionic binding motif composed, for the most part, of heavy chain residues. B 43C9 likely has a similar motif as 48G7. The only available structure is a complex with the product p-nitrophenol. HisL91 is proposed to form an acylated intermediate at the N*1 atom. However, the interpretation of the kinetic and structural data is not fully convincing. C D2.3 has a unique anionic binding motif located on the light chain side of the antibody. The Tyr is proposed to form the key interaction responsible for oxyanion stabilization. Catalytic antibodies D2.3–5, 6D9, and 7C8 differ from this general theme. In D2.3, phosphonate binding arises from hydrogen bonds to AsnL34, TrpH95, and TyrH100D (Fig. 1c) [28]. Thus, the anionic binding site is located on the opposite side of the binding pocket and is composed of different residues. Finally, the phosphonate binding motifs in antibody 6D9 and 7C8 are completely different from those seen in any of the other antibodies and consist of a single, solvent-exposed, hydrogen bond donated from a single HisL27D or TyrH95, respectively [20, 29, 30]. Not surprisingly, these antibodies exhibit lower catalytic activity than any other structurallydetermined hydrolytic antibodies (Table 1). Nevertheless, 6D9 has been recently evolved to have 20-fold higher activity with the addition of a second hydrogen bond to the phosphonate [31].

anionic binding site is underscored by the general agreement between transition state stabilization relative to the ground state, K M K DTS −1 , and the observed rate acceleration, kcat kuncat −1 [32] (Table 1). This agreement implies that hapten-binding interactions observed in the crystal structures correspond to the interactions found in the transition state, and that binding energy is converted into catalysis with near 100% efficiency. Thus, the simplest possible mechanism for ester hydrolysis in this family of antibodies is oxyanion stabilization within the anionic binding site. Oxyanion stabilization has a differential impact on catalysis for various natural esterase and protease enzymes. For example, biochemical studies in a bacterial cocaine esterase have demonstrated that the oxyanion hole is critical, contributing at least 5 kcal mol−1 to catalysis [33]. Oxyanion stabilization is estimated to contribute 2–3 kcal mol−1 to catalysis in papain [34] and cutinase [35], 3-4 kcal mol−1

5


in prolyloligopeptidase [36] kcal mol−1 in subtilisin [37–39], and 5–7 kcal mol−1 in acetylcholinesterase [40] and Zn2+ peptidase [41]. These values are within the range ascribed to the oxyanion contribution in the hydrolytic antibodies. For example, a contribution of 5–7 kcal mol−1 from oxyanion stabilization was estimated for the D2.3 hydrolytic antibody [42]. Here, the oxyanion was formally assigned to TyrH100D from pH rate profile and the inactivity of a Tyr → Phe mutant (Fig. 1c) [42]. The alternative candidate, AsnL34, was excluded as the Asn → Gly mutant retained full activity [42]. The activity of this antibody is significantly enhanced by spiking the assay buffer with potent α-nucleophiles, such as peroxide [42]. Thus, the catalytic potential of D2.3 is far from realized, and could be increased by a carefully positioned general acid/base that could more effectively activate water for nucleophilic attack. Even with the more potent nucleophile, the activity of this antibody is apparently limited by product inhibition [42] and conformational isomerization [43]. A largely overlooked ramification of the known anionic binding sites is the asymmetry of the coordinating residues (Fig. 1a–c). This asymmetry means that attack of the substrate carbonyl carbon and oxyanion stabilization has enantiofacial selectivity [44], since the phosphonate oxygens are non-equivalent (prochiral). Thus, the lack of 18 O incorporation in the substrates of hydrolytic antibodies should not be regarded as evidence for an acyl enzyme intermediate [45], but rather that the oxygen atoms are not racemized in the stereoselective binding pocket after hydroxide attack. This enantiofacial selectivity has been noted in other naturally-occurring hydrolytic enzymes [46, 47]. 2.4 Mechanistic Pitfalls

There is an obvious temptation to ascribe complex mechanismsto hydrolytic antibodies to elevate them to the same status as their natural enzyme counterparts. In the case of 17E8, for example, the proximity of a serine and histidine to the phosphonate in the crystal structure led to the proposal that the mechanism involved a catalytic dyad, analogous to the familiar catalytic triad [24]. This structure-based hypothesis was further bolstered by hydroxylamine partitioning data and pH rate profiles that suggested an acyl enzyme intermediate [48]. Initially, this interpretation was criticized because the refined serine rotamer was not within hydrogen-bonding distance to the histidine in the putative dyad [18, 49], but alternative rotamers of a serine have been noted in other serine hydrolases [47, 50, 51]. Eventually, the acyl enzyme mechanism was effectively discounted by a Ser → Ala mutant, which led to twofold improvement in kcat over the wild-type [52]. Likewise, the hydrolytic mechanism for 43C9 was proposed to involve an unprecedented nucleophilic attack by the Nδx atom of HisL91 on the substrate to form an acylated histidine intermediate [53]. In serine hydrolases, the nucleophilicity of histidine N2 atom is enhanced by a buried aspartate that H-bonds to Nδ1 stabilizing the developing positive charge on the imidazole. In the 43C9 pocket, HisL91 N2 H-bonds to TyrL36, but the nearest charged residue is ArgL96 (Fig. 1b). Hence, the available structural data [27] do not


yet provide sufficient evidence for such a mechanism. Therefore, the interpretation of the kinetic, mutagenesis, and structural data [45, 53–56] should be carefully reexamined or, at the very least, regarded with skepticism until a direct observation of an acylated histidine intermediate is demonstrated in a crystal structure. 2.5 Structural and Functional Considerations for Hapten Design

The initial use of a phosphonate analog as the transition state for solvent-mediated hydrolysis (Scheme 1) provided the initial breakthrough in the catalytic antibody field [6, 7]. However, the persistent and ubiquitous use of such analogs to elicit hydrolytic antibodies has now become somewhat of a red herring. Phosphonate compounds are not transition state analog inhibitors for serine hydrolases [57]. Thus, it would be impossible to elicit a serine hydrolase-like antibody using a phosphonate analog. Nevertheless, the elicited antibodies are inevitably compared to serine hydrolases [24], or to hydrolase mutants where the entire catalytic triad has been mutated to alanine [58]. Fluorophosphonate compounds (nerve gas reagents, for example) are potent mechanistic inhibitors of serine esterases because of covalent modification of serine with phosphate and concomitant expulsion of fluoride – a good leaving group. Phosphonate and phosphonamidate compounds do not form the same modification in serine esterases, since O− is a poor leaving-group. In contrast, phosphonate compounds are potent transition state analog inhibitors for Zn2+ proteases [59], which utilize a completely different mechanism from that of serine hydrolases and probably do not involve covalent acylation by the enzyme [60]. Rather, Zn2+ is believed to facilitate water-mediated attack of the ester/amide carbonyl carbon with arginine stabilization of the oxyanion [41, 60]. Unfortunately, no design feature in a phosphonate analog would be likely to elicit Zn2+ binding in an antibody combining site. Thus, the likelihood of eliciting hydrolytic antibodies with either serine hydrolase or Zn2+ peptidase-like features would seem unlikely without some innovative new hapten design. 2.6 Anionic Binding Motifs and Control of the Microenvironment in Decarboxylation

Catalytic antibody 21D8 catalyzes a classic decarboxylation reaction (Scheme 2) with a charge-delocalized transition state. This reaction exhibits high sensitivity to the surrounding medium, where the first-order rate constant in aqueous solution is enhanced 106 -fold in aprotic dipolar solvents [61]. Thus, this reaction is a particularly useful model for studying medium effects and control of the microenvironment in a heterogeneous binding pocket [62, 63]. Antibody 21D8 accelerates the decarboxylation reaction by 2 × 104 (Table 1) [62, 64]. The hapten design incorporated two sulfonate moieties to mimic the carboxylate anion of the substrate. Not surprisingly, this antibody has an anionic binding site consisting of HisH35 and ArgL96 [14]. An additional anionic binding site was observed for the second sulfonate group on the opposite side of the binding pocket. However, assigning which of these anionic

7


Scheme 2 Decarboxylation Antibody 21D8

binding sites was relevant for binding to the substrate carboxylate was hindered because of the unfortunate symmetry of the hapten. Moreover, attempts at cocrystallization with the substrate were unsuccessful [14]. However, mutagenesis studies support the hypothesis that HisH35 and ArgL96 are indeed key substratebinding residues [65]. It is proposed that the antibody shields the substrate from solvent, but the tetrahedral hydrogen-bonding pattern of the anionic binding site is not fully optimized for binding a planar carboxylate [14, 65]. The pocket is polar with apparently non-optimal hydrogen bonds to the transition state, which may be reminiscent of a dipolar aprotic solvent. Such a model would explain, in part, how the antibody and other enzymes could control the microenvironment to facilitate catalysis. 2.7 Anionic Binding Motifs and the Periodate Cofactor in Sulfide Oxygenation

Catalytic antibody 28B4 catalyzes a bimolecular reaction between sodium periodate (NaIO4 ) and a sulfide to form a sulfoxide (Scheme 3, Table 1) [66]. Oxygenation reactions in naturally-occurring enzymes normally require flavin, heme, or other cofactors as well as NADPH to regenerate the cofactor [67]. Here, periodate is exploited as an artificial chemical cofactor, readily available at high excess and obviating the need for cofactor recycling. The oxygenation reaction is believed to involve nucle-

Scheme 3 Sulfide oxygenation Antibody 28B4


ophilic attack of a sulfur lone pair of electrons on a periodate oxygen through a polar transition state (Scheme 3). The amine in the hapten mimics the partial positive charge that develops on the sulfur, while the phosphate mimics the tetrahedral periodate. The crystal structure reveals an anionic binding site similar to that seen in the phosphocholine antibody McPC603 [8, 9] that coordinates periodate (Fig. 2a) [68]. However, the baiting strategy employed to stabilize the developing positive charge on the sulfur was not as effective as the postulated cation-π interaction with a nearby tyrosine does not have optimal geometry (Fig. 2a). Extensive mutagenesis has been performed on this antibody, and the putative germline antibody was also cloned and crystallized in complex with hapten [69]. Here, the p-nitrophenyl portion of the hapten adopts a completely different orientation in the binding pockets of the and affinity-maturated antibodies [69]. In addition, the free and bound forms of the germline antibody structure show significant conformational differences in CDRH3 and CDRL1, while comparison of the free and bound mature antibody shows no significant changes on binding [69]. Thus, the germline is postulated to exhibit wider structural plasticity than the mature antibody [69]. The loss of induced fit binding during the affinity maturation process has been noted in at least two other antibodies [22, 70], but does not rule out the importance of induced fit in affinity-matured antibodies [1].

Fig. 2. Top and side views of sulfide oxygenation and allylic isomerization antibodies, respectively. A Antibody 28B4 has an anion-binding pocket that binds the cofactor periodate in a biomolecular reaction with sulfide to form sulfoxide. This pocket was elicited by a phosphate functionality in the immunizing hapten. B Antibody 4B2 catalyzes an isomerization reaction of a $-( unsaturated ketone. GluL34 acts as general acid/base in the reaction by abstracting the proton from the substrate. Here, GluL34 interacts with the amidinium part of the hapten.

9


Scheme 4 Allylic isomerization Antibody 4B2

2.8 Cation Stabilization and Allylic Isomerization

Several antibodies have been elicited that are believed to have mechanistically important, negatively charged carboxylates in their binding pocket. In principle, such carboxylates may be elicited from a baiting strategy that incorporates positivelycharged substituents in the immunizing hapten. Few structures of this type have been determined, however, and for at least one of these antibodies (1D4), the baiting strategy failed. Antibody 4B2 catalyzes an allylic rearrangement of a β–γ unsaturated ketone (Scheme 4, Table 1). Formally, the reaction would be a [1, 3] sigmatropic migration of hydrogen. An amidinium group was incorporated in the hapten design to elicit a carboxylate general acid/base in the binding pocket (Scheme 4) [71]. The crystal structure of 4B2 shows that this baiting strategy was successful, as the two NH groups of the amidinium form H-bonds to each oxygen of the GluL34 carboxylate (Fig. 2b) [72]. The pH rate profile is bell shaped with optimal activity at pH 4.5, the approximate pK a of glutamic acid. An enol intermediate was proposed, based on lack of any antibody-catalyzed 2 H exchange between α and γ carbons in a deuterated substrate, implying that the reaction is not concerted, but involves exchange with solvent [71]. This is a reasonable hypothesis, although it is unlikely that any antibody residue would be positioned to stabilize this transient enol intermediate [72].

3 Shape Complementarity and Approximation

Shape complementarity simply refers to the similarity between the contours of the binding pocket molecular surface and that of its corresponding ligand. Certainly, it is possible to conclude from most of the catalytic antibody structural studies that exquisite shape complementarity is sufficient to explain the stereoselectivity for the majority of these catalysts. Thus, when the catalysts are challenged with chiral (or prochiral) substrates, chiral products are obtained, typically in high enantiomeric excess (ee). In addition, every reaction requires, to some extent, approximation to bring reactive centers together to either break or make new bonds. The antibody molecule is ideally tailored for this role, as it may be programmed to bind flexible substrates in a particular conformation that would otherwise be disfavored in solution, and can bring two substrates together in bimolecular reactions to increase the effective molarity of the substrates relative to one another. In principle,


approximation lowers the entropy barrier for activation and can effect catalysis. Thus, electrostatic and overall shape complementarity of the binding pocket for the transition state is a key feature for any catalyst and is one of the most important features cited for many catalytic antibody structures. However, approximation alone probably does not fully explain catalysis in all of these systems. Solvent accessibility, control of the microenvironment dielectric, and hydrogen-bonding potential are also important parameters – some reactions are accelerated simply by changing the solvent system. Indeed, not surprisingly, nonspecific interactions with bovine serum albumin (BSA) can accelerate some reactions [73]. In addition, key catalytic residues form specific interactions essential for the reaction to proceed. Catalytic residues have been elicited with mixed success by TSA baiting strategies, reactive immunization, and sometimes by pure chance. Notwithstanding, approximation will be a recurrent theme throughout the remaining sections. 3.1 Unimolecular Rearrangements 3.1.1 Antibody 1F7 Antibody 1F7 catalyzes the [3,3] sigmatropic Claisen rearrangement of chorismate to prephenate (Scheme 5, Table 1) [74]. This reaction is normally catalyzed by chorismate mutase in the biosynthesis of aromatic amino acids. The enzymatic reaction proceeds through an ordered chair-like transition state (S‡ ≈ −13 cal mol−1 K − 1 ) with a rate acceleration of 3 × 106 or four orders of magnitude better than the antibody [75]. Remarkably, even with this modest activity, over-expression of 1F7 complements growth of yeast auxotrophs [76]. The original structure was determined at 3.0 Å resolution from data merged from two crystals collected at room temperature [77]. More recent (unpublished) data at 2.4 Å show some differences in the detailed interactions between the TSA and Fab [77]. However, the general placement of the hapten on the heavy chain side of the antibody is maintained and illustrates a clear and interesting difference in the orientation of the TSA in the antibody pocket as opposed to the enzyme active site [78]. For the antibody, the hapten-binding orientation is determined by the placement of the linker used for covalent attachment to the immunogen, while the enzyme has no such restriction (Scheme 5). The enzyme also utilizes more H-bond interactions than the antibody, which are important for stabilizing the dipolar transition state [77]. Thus, the reaction is probably being catalyzed by preferentially binding the substrate in the

Scheme 5 Chorismate mutase Antibody 1F7

11


requisite reactive conformation, while poorly optimized hydrogen bond interactions may account for the inefficiency of the antibody catalyst. This theme recurs in which the substrate in an antibody is inserted upside down compared to that in the natural enzyme [47, 79]. 3.1.2 Antibody AZ-28 AZ-28 catalyzes a related [3,3]sigmatropic oxy-Cope rearrangement (Scheme 6, Table 1) [80]. The chair-like transition state is highly ordered, requiring overlap of the rearranging 4π+ 2σ orbitals of the diene (S‡ ≈ −15 cal mol−1 K−1 ). Antibodies raised against the appropriate, conformationally-restricted TSA would be expected to catalyze the reaction by acting as an “entropy trap.” In addition, appropriate alignment of the 2- and 5-phenyl rings are believed to contribute stereoelectronically to the reaction through hyperconjugation with the 3-hydroxyl (Scheme 6) [81]. This hyperconjugation may accelerate the oxy-Cope rearrangement by an anionic substituent effect. Indeed, the crystal structure shows that packing interactions with the 2- and 5-phenyl rings constrain the cyclohexyl group in the required chairlike transition state (Fig. 3). However, the cyclohexyl ring of the TSA is rotated ca. 80◦ out of plane of the 2- and 5-phenyl rings, so that the phenyl π-orbitals do not align appreciably with the 3-hydroxyl (Fig. 3) [81]. The orientation of the cyclohexyl group is determined by packing interactions, as well as by hydrogen bonds to the 3-hydroxyl (Fig. 3). Surprisingly, biochemical analysis revealed that six mutations occurred during affinity maturation, one of which dramatically decreases the catalytic efficiency [81]. Thus, the evolution of an antibody binding pocket with increased affinity to the TSA does not always correlate with improved catalysis. The germline-encoded antibody structure was determined and revealed an induced-fit mode of binding, as opposed to a lock-and-key type binding in the mature antibody. This apparent conformational flexibility and corresponding decrease in affinity is interpreted as evidence for possible torsional flexibility in the 2-phenyl substituent [82]. This flexibility, in turn, is postulated to facilitate π-overlap and explain the

Scheme 6 Oxy-Cope Antibody AZ-28

3 Shape Complementarity and Approximation Fig. 3. Side view of oxy-Cope antibody AZ-28. The cyclohexyl ring of the TSA is rotated out of plane with the 2- and 5-phenyl rings because of packing interactions and hydrogen bonds to the 3-hydroxyl. The consequent lack of hyperconjugation in the rearranging B-orbitals may explain the decreased rate acceleration in the affinity mature antibody, as opposed to the germline Fab.

improved catalysis. Studies to characterize the dynamics of the antibody catalyzed reaction would provide further evidence to support this hypothesis. 3.2 Bimolecular Rearrangements and the Diels-Alder Reaction

The D-A reaction is one of the most important carbon-carbon bond-forming reactions that is available for organic syntheses of complex natural products and therapeutic agents [83, 84]. Surprisingly, the D-A reaction has not been exploited widely in nature as only a handful of natural D-A enzymes are known [85]. The reaction is a pericyclic [4π+ 2π] cycloaddition of a diene and dienophile. An optimal D-A reaction occurs between an electron-rich diene (HOMO - highest occupied molecular orbital) and an electron-deficient dienophile (LUMO - lowest unoccupied molecular orbital). These abzyme-catalyzed bimolecular reactions are typically much slower and have much higher K M s than esterolytic antibodies (Table 1). The transition state for the D-A reaction is highly ordered for which S‡ is typically -30 to -40 entropy units [86]. Consequently, the recurring theme for these D-A antibodies is shape complementarity; in general, the antibodies with higher shape complementarity to the transition state are better catalysts. Thus, these antibodies may be considered as model systems for studying the effects of approximation in catalysis. 3.2.1 Retro-Diels-Alder Reaction Antibody 10 F11 catalyzes a unimolecular retro-D-A reaction that expels the dieneophile nitroxyl and diene anthracene (Scheme 7, Table 1). The dihedral angles between the phenyl rings in the substrate (110◦ ) and product (180◦ ) differ by 70◦ [87]. A TSA was designed with an intermediate dihedral angle of 140◦ [87]. Four crystal structures were determined including substrate analog (SA), transition state analog (TSA), and product analog (PA) [88]. The kinetics of the reaction suggest that binding energy is converted to catalysis with high efficiency (Table 1), with a clear hierarchy in the shape complementarity index between the antibody and ligands, ranked as TS > SA > PA [88]. This ranking supports the hypothesis that the antibody exerts strain on the substrate to preferentially stabilize the transition state.

13


Scheme 7 Retro Diels-Alder Antibody 10F11

Antibody 9D9 catalyzes the same reaction with slower kinetics, which could result from poorer overall shape complementarity [88]. The CDRH3 loop also undergoes a slight induced-fit con-formational change from binding the substrate to the transition state [88]. Finally, a water-mediated hydrogen bond was observed between the antibody backbone and the oxygen that is released from the substrate in the nitroxyl dieneophile product [88]. A second hydrogen bond is formed directly between the antibody backbone and the same oxygen of the substrate. These hydrogen bonds are postulated to act as Lewis acids that further stabilize the LUMO energy of the dieneophile and contribute to the regioselectivity of the retro-D-A reaction [88]. 3.2.2 Disfavored exo Diels-Alder Reaction For antibody 13G5, a novel hapten design was examined (Scheme 8, Table 1). Rather than using a conformationally-restricted TSA, an unrestrained ferrocene derivative was used to test whether the immune system would select a conformer that mimics one of the Diels-Alder transition states (Scheme 8) [89]. In principle, a panel of catalysts would be generated, each of which would selectively catalyze the formation of one of the four possible ortho-diastereomers. In fact, of all the antibodies examined, only one enantiomer was formed, indicative of possible structural bias in the immune response [90]. The catalyzed reaction proceeds with high regio-, diastereo-, and enantioselectivity to form the ortho-exo (S,S) product with ee > 95% after the background exo D-A reaction is subtracted [90]. The background endo:exo product distribution is 85:15 in PBS and 66:34 in toluene, so the exo-transition state is slightly disfavored by ca. 1 kcal mol−1 . The antibody rate acceleration is modest with a kcat(exo) kuncat(exo) −1 of 6.9 M and a kcat(exo) kuncat(endo) −1 of only 1.7 M [89, 91].

Scheme 8 Diels-Alder Antibody 13G5


Thus, the background reaction to form unwanted ortho-endo product will dominate the final product distribution of the antibody-catalyzed reaction unless sufficiently high concentrations of antibody catalyst are used [90]. The crystal structure of 13G5 shows the cyclopentadienyl ferrocene deeply buried in the antibody combining site [91]. Three H-bonds are observed between the TSA and TyrL36, AspH50, and AsnL91. TyrL36 was initially hypothesized to act as a Lewis acid, activating the dienophile for nucleophilic attack, while AsnL91 and AspH50 form H-bonds to the carboxylate side chain that substitutes for the diene substrate [91]. However, the limited rate accelerations indicate that catalytic residues in the binding pocket have a modest affect, if any, on the kinetics. Rather, the observed diastereo- and enantioselectivity of the antibody could be explained simply by exclusion of solvent (e.g., note toluene product distribution) and specific van der Waals interactions and hydrogen bonds that anchor the substrates in a particular orientation that favors ortho-exo (S,S) product formation due to approximation [86]. The performance of this catalyst may be explained by the relatively poor shape complementarity between the antibody and the true transition state of the reaction, which differs substantially from the hapten as the distance between the cyclopentadienyl rings in the ferrocene structure is ca. 3.3 Å. But what is truly remarkable is that a specific antibody could be generated against such a flexible hapten. 3.2.3 Endo Diels-Alder Reactions Antibody 39-A11 was elicited from immunization with a constrained bicyclic hapten (Scheme 9), and selectively accelerates formation of the kinetically-favored endo product [92]. The acceleration over the background or effective molarity is modest (Table 1). The X-ray structure of the hapten complex indicates that the antibody would bind the diene and dienophile in a reactive conformation, as preprogrammed in the hapten design strategy. The structure also suggests that the catalyst is enantioselective, although an ee has not been determined [93]. AsnH35 forms a hydrogen bond with the dienophile, which could make it more electron deficient and, thereby, more reactive [93]. Interestingly, any evident shape complementarity is lacking between antibody and the endo transition state; the binding pocket can accommodate either the endo or exo transition states (Fig. 4a). Mutations were designed to improve the packing interactions around the kinetically-favored endo transition state, and three of six mutants led to a 5- to 10-fold improvement in kcat [93]. This successful structure-based engineering is a significant and a rare accomplishment in this field. Also, only two somatic mutations were required to generate this catalytic antibody from the germline precursor. The most relevant of these two mutations is SerL91Val (CDR3), located in the binding pocket. The K DTSA for the germline antibody is 380 nM compared to 130 nM in the mature 39-A11. As expected, the catalytic proficiency of the germline antibody is also slightly decreased [93]. In order to circumvent substrate or product inhibition, more sophisticated hapten design strategies incorporate extraneous chemical groups as a negative design, while still promoting high affinity interactions to the transition state (Scheme 10) [94]. 1E9 was originally developed to evaluate the role of proximity

15


Scheme 9 Diels-Alder Antibody 39-A11

effects in catalysis, and is the most efficient D-A catalytic antibody yet developed (Table 1) with an effective molarity (EM) of 103 M, where the theoretical limit is 108 M [95]. Two structural features can explain the mechanism of the observed antibody catalysis. First is the conserved H-bond formed between AsnH35 and the succinimide carbonyl oxygen of the hapten, which represents the carbonyl oxygen of the dienophile substrate (Scheme 10). Since an H-bond to the carbonyl oxygen should cause the dieneophile to be electron deficient, it is more susceptible to the attack by the diene. Second, the binding pocket features almost perfect van der

Fig. 4 Molecular surface representation of Diels-Alder antibodies. Antibody 39-A11 (A) and 1E9 (B) are derived from the same germline precursor, yet exhibit a large difference in activity. This difference is attributed to the poor shape complementarity of the

antibody for the transition state in 39-A11 compared to 1E9. Mutations were designed in 39-A11 to improve the shape complementarity, and several improved catalysts were identified.

4 Shape Complementarity and Control of the Reaction Coordinate

Scheme 10 Diels-Alder Antibody 1E9

Waals complementarity to the hapten, providing a clear structural demonstration of approximation in bimolecular catalysis (Fig. 4b) [96]. Interestingly, 1E9, antiprogesterone antibody DB3, and the D-A antibody 39A11 are derived from the same germline gene [93, 97–99]. Structures of these antibodies thus represent possible snapshots in the evolution of substrate binding and catalytic efficiency [94]. Subtle differences in the evolution of their binding pockets through somatic mutations have resulted in antibodies that specifically bind progesterone or catalyze a D-A reaction. The exceptional efficiency of 1E9 is hypothesized to arise from a rare somatic mutation that significantly deepens the active site, creating exquisite complementarity to the transition state [94].


In many reactions, several products may be formed, but one product predominates because it is favored kinetically. The energy of activation is much lower for the favored product, which reduces the likelihood of forming the disfavored product. If the activation barrier for the reverse reaction is sufficiently large, then the reaction is essentially irreversible, or under kinetic control. Similarly, for some reactions, many possible products may result (e.g., the D-A reaction or cationic cyclization), and complex mixtures result. The antibody catalyst can steer a reaction through a complex reaction coordinate to alter the final reaction profile, often forming exclusively a single product, and, in some cases, a kinetically-disfavored one. 4.1 Syn-Elimination

Antibody 1D4 is a model system for studying the selective catalysis of a disfavored syn-elimination reaction toform a cis product (Scheme 11, Table 1) [100]. The TSA design incorporated a rigid bicyclic ring structure to constrain the two phenyl groups of the substrate in the eclipsed conformation that characterizes the transition state (Scheme 11). In addition, the α-keto proton that is abstracted during elimination was replaced with a positively-charged amine in the hapten to elicit a complementary general base in the antibody combining site. Although

17


Scheme 11 Syn-elimination Antibody 1D4

the catalyzed reaction proceeds slowly (Table 1) [100], the background elimination reaction to the cis-product is undetectable in the absence of antibody; the competing anti-elimination reaction (k = 2.5 × 10−4 min−1 ) dominates the background reaction, such that the product distribution is entirely trans. Crystal structures of free and bound 1D4 show a conformational change in CDR H2, indicative of induced fit [101]. The pocket exhibits high shape complementarity to the TSA, which accounts for the selectivity for the eclipsed transition-state of the reaction (Fig. 5) [101]. Hence, the antibody combining site likely utilizes the majority of its binding energy toward overcoming the extreme steric barriers, and, hence, stabilizing the eclipsed transition state. The key mechanistic issue for any βelimination process is whether the reaction is E1, E2, or E1cB [102]. Unfortunately, a crystal structure alone cannot resolve such a question. A water molecule was modeled into electron density that is near the hapten amine [101] and within

Fig. 5 Electrostatic potential mapped to the molecular surface of 1D4. The exquisite shape complementarity for the phenyl rings that characterize the eclipsed transition state of the reaction accounts for the selectivity of the catalyst.


hydrogen-bonding distance to a histidine. Thus, HisH58 is proposed to promote catalysis through interaction with the water [101]. 4.2 Disfavored Ring Closure

Antibody 5C8 catalyzes the disfavored endo-tet cyclization reaction inviolation of Baldwin’s rules for ring closure (Scheme 12) [15]. The N-methylpiperidinium hapten was anticipated to elicit negative charge in the binding pocket that would stabilize the developing positive charge at the oxirane carbon of the substrate. Several changes were observed between the bound and unbound structures, indicative of induced fit. The largest rearrangement was observed in the backbone and side chain orientations of CDR H3 with the largest differences seen for the two tyrosine residues at the tip of the loop, which fold into the binding site, creating a solvent-inaccessible cavity. AspH95 and AspH101 bind the positive charge of the quaternary amine like a pair of tweezers (Fig. 6) [15]. In addition, the positive charge appears to be stabilized by a cation-π interaction with TyrL91. AspH95 and HisL89 were hypothesized to effect rudimentary general acid/base catalysis (Fig. 6). In the proposed mechanism, AspH95 forms an H-bond to the epoxide-oxygen, favoring the oxirane ring opening, while HisL89 forms an H-bond with the alcohol to promote nucleophilic SN 2-like attack to form the disfavored six-membered ring [15]. This stabilization of the positive charge that develops along the reaction coordinate appears to be an important factor for the rate enhancement and for directing the reaction along the otherwise disfavored pathway [15]. 4.3 Selective Control of a Reactive Carbocation: Antibodies 4C6 and 19A4

Cationic cyclization proceeds via formation of a reactive carbocation, and may be separated into three steps: initiation, propagation, and termination. A carbocation may be formed either by ionization or by electrophilic addition to an unsaturated olefin, while cyclization is terminated by nucleophilic addition or elimination of a proton. The stereochemistry of such transformations should follow the StorkEschenmoser hypothesis [103, 104]. Natural enzymes catalyze polycyclization of squalene through a reactive carbocation intermediate to form a variety of cycloisoprenoids (e.g., cholesterol). For such enzymes, initiation occurs either by protonation of a double bond or by release of pyrophosphate [105]. Structural studies have shown that the enzyme active sites are typically lined with aromatic residues that

Scheme 12 Disfavored cyclization Antibody 5C8

19


Fig. 6 Disfavored ring closure antibody 5C8. AspH95 is proposed to form an Hbond with the epoxide-oxygen, favoring oxirane ring opening, with HL89 forming an H-bond with the alcohol to promote SN2-

like attack to form the disfavored ring. The piperidinium moiety was used as bait to elicit the desired negative charge in the binding pocket, which should stabilize the developing positive charge in the actual ring closure reaction.

stabilize the carbocation through cation-π interactions. Catalytic antibodies 4C6 and 19A4 also contain a number of aromatic residues in the binding pocket, which are postulated to play a similar role. Thus, the enzymes and antibodies exhibit similar features. Catalytic antibody 4C6 catalyzes a model reaction that had been shown under other conditions (98% formic acid) to proceed through a carbocation [106]. This reactivity was elicited with an N-oxide hapten [107] that is structurally related to the 5C8 hapten. Structurally analogous N-oxide TSAs are known inhibitors for natural squalene cyclases, validating this design strategy [108]. The oxide was included to promote stabilization of the negative charge that develops in the sulfonic acid leaving group, while the nitrogen would elicit residues that increase the electrophilicity at C1 (Scheme 13, Table 1). The hapten also includes a silane moiety that could stabilize the carbocation via hyperconjugation [107]. In addition, conjugation with silane makes the olefin more electron rich, which facilitates attack of C1 . Carbocations are highly reactive species, yet the antibody selectively catalyzes the formation of only one product in mild conditions where no background reaction is detectable (Scheme 13) [107]. The crystal structure shows several aromatic residues in the active site that could stabilize the reactive carbocation and shield it from solution [17]. In the proposed mechanism, TyrL96 and/or TyrH50 initiate carbocation formation at C1 by promoting SN 1-like departure of the acetamidobenzenesulfonic acid leaving group. Electrophilic attack of the C5 -C6 olefin by the developing C1 carbocation closes the ring and shifts the carbocation to C5 , where the reaction terminates by nucleophilic addition of water [17]. This mechanism, however, does not explain why a related compound (Si exchanged for C) has a ca.100-fold reduction in


Scheme 13 Cationic cyclization Antibody 4C6

kcat , but similar K M for the reaction, nor why the related substrate that lacks the C5 C6 double bond exhibits no reaction (kinetics monitor formation of leaving group). Together, these data suggest that anchimeric assistance of the electron-rich double bond is the central feature of the mechanism [107]. Thus, the reaction could be initiated by backside attack of C1 by the electron-rich olefin, followed by termination by nucleophilic addition of water at C5 . This attack is facilitated by approximation due to the shape complementarity of the binding pocket. Additional kinetics and mutagenesis could help resolve the details of how the reaction is initiated. Catalytic antibody 19A4 catalyzes a more complicated, tandem-cationic cyclization to form bridge-methylated decalins (Scheme 14, Table 1) [109]. The hapten design was similar to that for 4C6, with the same N-oxide functionality to elicit residues that would promote initiation of the cyclization cascade by expulsion of the sulfonic acid leaving group. In contrast to 4C6, this reaction is terminated by elimination. The antibody does not possess the exquisite control over the reaction that was observed for 4C6, as three different decalins are formed accounting for only 50% of the overall products [109]. Nevertheless, the catalysis is remarkable, as no bicyclic products are formed in the uncatalyzed reaction. The crystal structure shows a hydrophobic binding pocket lined with several aromatic residues that may stabilize the carbocation [16]. In addition, the binding pocket has high shape complementarity to the productive chair-chair conformation of the substrate that is required for effective cyclization (Scheme 14). Here, a hydrogen bond from

Scheme 14 Cationic cyclization Antibody 19A4

21


AsnH35a to the sulfonic acid leaving group is believed to facilitate SN 1-like departure with concerted anchimeric assistance from the C5-C6 π bond, yielding the first ring with the carbonium ion at the tertiary C5 carbon [16]. In the next step, the C9-C10 π bond undergoes electrophilic attack by the C5 carbonium to form the second ring. Although an epoxide was included in the original hapten to elicit a catalytic residue in the antibody, there is no catalytic base to direct the regiochemistry of the final termination step of the reaction [16]. Thus, the reaction results in a mixture of elimination products. Structure-based engineering would afford a unique opportunity to control the product distribution for this particular reaction.

5 Shape Complementarity and Substrate Strain

Many catalytic antibodies impose varying degrees of strain on the substrate to achieve reactive conformations that would otherwise be disfavored in solution (e.g., 1D4, 4C6, 19A4, 13G5, 10F11, AZ-28). The enzyme ferrochelatase, however, is a particularly attractive model system for studying substrate strain. Ferrochelatase inserts Fe2+ into porphyrin in heme biosynthesis. In this reaction, the enzyme is proposed to distort the porphyrin ring system to expose the pyrrole nitrogen lone pair electrons, facilitating metal ion complexation. Indeed, N-methyl protoporphyrin – a distorted porphyrin analog – is a potent inhibitor of ferrochelatase [110]. Antibodies elicited against this compound catalyze insertion of Cu2+ or Zn2+ into mesoprotoporphyrin IX, supporting the hypothesis that the hapten is a true transition state analog for the enzyme-catalyzed reaction (Scheme 15, Table 1) [111]. Thus, this reaction is a model system for studying the effects of substrate strain in catalysis [4]. In the antibody-catalyzed reaction, no saturation kinetics were observed for the metal ion, suggesting that metal binding to the antibody is not a prerequisite of catalysis, in contrast to ferrochelatase, which has a K M of 32 µM for iron [112]. The crystal structure, however, reveals that an Asp carboxylate is directed toward

Scheme 15 Metal chelation antibody 7G12

6 Reactive Amino Acids and the Possibility of Covalent Catalysis

Fig. 7 Superposition of free (green) and TSA bound (blue) 7G12. (A) The germline 7G12 shows an induced-fit mode of binding, as demonstrated by the conformation change in CDRH3. (B) The affinity mature

antibody, in contrast, has a lock-and-key mode of binding. The maturation of a preformed binding pocket is associated with higher affinity for the transition state and overall improved catalytic proficiency.

the center of the protoporphyrin (Fig. 7) and is proposed to either guide metal complexation or act as a proton shuttle [113]. The residue is clearly important as Asp → Asn and Asp → His mutations are inactive [113]. In addition, the crystal structure shows that packing interactions between the antibody and substrate induce strain in the Michaelis complex [70]. Although accurate modeling and exact refinement of any distortions in the Michaelis complex at 2.6 Å resolution are not reasonable, the electron density does provide clear qualitative indications of deviation from ideal planarity [70]. The free and hapten-bound structures of the germline and mature antibodies were compared and nicely illustrate the evolution from an induced fit mode of binding to lock-and-key (Fig. 7a,b) [70]. The evolution of this lock-and-key binding results in tighter binding to the TSA (due to faster association and slower dissociation rates), and correlates with a ca. 100-fold improvement in kcat K M −1 [70]. Together, these structural and biochemical results demonstrate the importance of substrate strain in antibody-mediated catalysis.


The recently developed reactive immunization strategy [114, 115] has shown that it is possible to create antibodies that catalyze aldol reactions, one of the most important carbon-carbon bond-forming reactions in chemistry and biology. The mechanism for the natural class I aldolase enzyme involves the key formation of a Schiff base between the substrate ketone (aldol donor) and a reactive lysine, followed by condensation with an aldehyde (aldol acceptor) and subsequent hydrolysis to release product (Scheme 16). The binding pocket of aldolase antibody 33F12 is an elongated cleft more than 11 Å deep [116]. The only lysine residue in the active site

23


Scheme 16 Previously proposed retro-aldol mechanism for aldolase antibody 33F12

is located at the bottom of the hydrophobic pocket (Fig. 8a). This environment was hypothesized to perturb the pK a of the lysine, making it amore potent nucleophile [116]. The antibody can then use the reactive -amino group to form an enamine with the corresponding substrates, analogous to the natural class I aldolase. The LysH93 → Ala mutant is inactive, which confirms the importance of this residue [117]. An additional residue that may be of mechanistic interest is TyrL36, which forms a water-mediated hydrogen bond with the -amino group of lysine H93 (Fig. 8a). Importantly, a water molecule was postulated to play a key role shuttling protons in the mechanism of a class I aldolase [118]. Interestingly, the antibody exhibits exceptionally broad substrate specificity, implying that the binding pocket is unrefined for a particular substrate presumably due to the lack of further affinity maturation due to the proposed covalent nature of the hapten complex [116]. Antibodies 40F12, 84G3, 93F3 and others were generated with a sulfone β-diketone hapten [119]. Notably, antibodies 93F3 and 84G3 show reversed enantioselectivity compared to 33F12 [119]. Sequence analysis and our unpublished structural data [120] reveal a lysine at position L89 on the opposite side of the binding pocket, which could explain the reversed selectivity of this antibody. The relatively unrefined binding pocket of 33F12 contrasts with that of class I aldolase enzymes, which catalyze the reversible condensation of aldehydes and ketones with high stereospecificity (121,122). The bacterial 2-deoxyribose-5-phosphate aldolase (DERA) (123,124) catalyzes the reversible aldol reaction of acetaldehyde and D-glyceraldehyde-3-phosphate to form 2-deoxyribose-5-phosphate. The ultra high resolution structure of DERA reveals a β-barrel with two lysines in the active site, one of which forms the covalent Schiff base intermediate (Fig. 8b) [118]. Compared to the hydrophobic antibody binding pocket, a number of charged/polar residues form a sophisticated hydrogen-bonding network within the active site, which is likely important for tuning the reactivity of the lysine and the substrate specificity [118]. The mechanism by which Lys167 is rendered nucleophilic in the enzyme is unclear – the pK a was proposed to be depressed by a neighboring Lys201 (Fig. 8b) [118], but mutation of Lys201 → Leu clearly does not hinder formation of the Schiff


base [118]. Surprisingly, inhibition of natural aldolases by diketones has not been fully characterized. A lingering challenge is to determine the mode of binding of the aldolase family of antibodies to substrate-related inhibitors, as well as the diketone compound originally used as a hapten to elicit the antibody response (Scheme 16). Despite our best efforts, we have obtained no structural evidence demonstrating the formation

Fig. 8 Comparison of aldolase antibody and enzyme active sites. Red spheres represent water molecules. A The aldolase antibody contains a lysine buried at the bottom of hydrophobic gorge, which has been postulated to form covalent intermediates with an aldol donor (ketone) followed by condensation with an aldol acceptor (aldehyde). TyrL36 may form a water-mediated hydrogen bond with the reactive lysine ,amino group, which may be of importance in the reaction mechanism. B The bacte-

rial 2-deoxyribose-5-phosphate aldolase (DERA) likewise contains a reactive Lys167, which was observed to form a covalent intermediate with the substrate in the crystal structure. Lys167 is probably rendered nucleophilic by the sophisticated hydrogenbonding network in the active site, which includes nearby Asp102 and Lys201. A nearby water molecule forms key stabilizing interactions to the substrate, and is believed to play a key role in the catalytic mechanism.

25


of a covalent bond in the antibody binding pocket for the aldolase antibody. This contrasts with the relative ease by which we and others have obtained covalent adducts in enzyme systems [118]. Surprisingly, the Lys167Leu DERA mutant catalyzes the retro aldol reaction for its optimal substrate (3.6 min−1 ) [118] three times faster than the antibody for its optimal substrate (1.4 min−1 ) [125]. Clearly, catalysis of the retro-aldol condensation may occur in the absence of covalent intermediates. Alternative mechanisms for 33F12 and 38C2 were never seriously considered [116], but could involve non-covalent stabilization of an enolate or aldol donor that subsequently attacks the aldol acceptor (analogous to a class II aldolase). Such a mechanism would be more consistent with our structural observations. In any event, the mechanistic supposition that a lysine could be rendered nucleophilic in the hydrophobic environment of the antibody binding pocket led, in part, to the appreciation that secondary amines, e.g., proline, could catalyze the aldol reaction in organic and, to lesser extent, aqueous solvent [126]. Thus, structural and biochemical characterization of aldolase antibodies has led to the development of important new synthetic methodologies.

7 Conclusions

Classically, antibodies are effector molecules that link the recognition elements of the Fab to the destructive elements of the immune system. Recently, this paradigm has been challenged by the discovery that antibodies also possess the intrinsic capability to catalyze the formation of hydrogen peroxide from singlet oxygen and water in the absence of any other metal ions or cofactors [127, 128]. This catalytic capability would potentially link recognition and oxidative killing within the same molecule. This remarkable chemistry is likewise catalyzed by the T-cell receptor (TCR), but not β 2 -microglobulin, which shares the immunoglobulin fold [128]. Very little is understood about where this reaction occurs on the antibody or TCR, and docking studies have not resolved this issue [129]. Assaying light chain dimers and camel antibodies (heavy chain only) would provide some indication of where the chemistry occurs. We have grappled with this problem indirectly by pressurizing numerous antibody crystals under a xenon atmosphere [128]. Our data reveal several hydrophobic pockets in the core of the immunoglobulin fold. Since xenon is larger than oxygen (O2 ), oxygen could potentially diffuse into the same hydrophobic cavities. However, the mechanistic relevance of these hydrophobic cavities, if any, is not yet resolved. Unfortunately, a more direct experimental approach is extremely challenging; distinguishing unambiguously between water, singlet oxygen, hydrogen peroxide, ozone, and other reactive oxygen intermediates (which are believed to be short lived) from an electron density map is not feasible, even at ultra high resolution. Moreover, discriminating between radical chemistry derived from the antibody or synchrotron radiation is non-trivial and requires numerous control experiments. However, we have observed generally that soaking antibody crystals in hydrogen

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1

N-terminal Ubiquitination Aaron Ciechanover

Technion — Israel Institute of Technology, Haifa, Israel

Originally published in: Protein Degradation, Volume 1. Edited by R. John Mayer, Aaron Ciechanover and Martin Rechsteiner. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30837-7 Abstract

The ubiquitin–proteasome system (UPS) is involved in selective targeting of innumerable cellular proteins via a complex pathway that plays important roles in a broad array of processes. An important step in the proteolytic cascade is specific recognition of the substrate by one of many ubiquitin ligases, E3s, that is followed by generation of the polyubiquitin degradation signal. For most substrates, it is believed, though it has not been shown directly, that the first ubiquitin moiety is conjugated, via its C-terminal Gly76 residue, to an ε-NH2 group of an internal lysine residue. Recent findings indicate that for an increasing number of proteins, the first ubiquitin moiety is fused linearly to the α -NH2 group of the N-terminal residue. An important biological question relates to the evolutionary requirement for an alternative mode of ubiquitination.

1 Background

Two distinct structural elements play a role in the ubiquitination of a target protein: (i) the E3 recognition site and (ii) the anchoring residue of the polyubiquitin chain. In most cases, it is believed, though it has been shown for only a few proteins, that the first ubiquitin moiety is transferred to an ε-NH2 group of an internal lysine residue in the substrate. The N-terminal domain of the target protein has attracted attention both as an E3 recognition domain and, recently, as a ubiquitination site. As for specific recognition, in certain rare cases, the stability of a protein is a direct function of its N-terminal residue, which serves as a binding site for the ubiquitin ligase E3α (Ubr1 in yeast; ‘N-end-rule’; [1, 2]). Accordingly, two types of N-terminal residues have been defined, “stabilizing” and “destabilizing”. For the


2 N-terminal Ubiquitination

Mos protein, it was found that its stability is governed primarily by the penultimate proline residue and by a phosphorylation/dephosphorylation cycle of serine3 [3]. A mechanistic explanation for the role of the Pro and Ser residues is still missing. As for the lysine residue targeted, there is no consensus as to its specificity. In some cases distinct lysines are required, while in others there is little or no specificity. Thus signal-induced degradation of IκBα involves two particular lysine residues, 21 and 22 [4]. In the case of Gcn4, lysine residues in the vicinity of a specific PEST degradation signal serve as ubiquitin attachment sites [5]. Mapping of ubiquitination sites of the yeast iso-2-cytochrome c has revealed that the polyubiquitin chain is synthesized almost exclusively on a single lysine [6]. In two other examples, that of Mos (see above; [3]) and the model “N-end rule” substrate X-β-gal (where X is a short fused peptide not encoded by the native molecule [7]), one and two lysines, respectively, that reside in proximity to the degradation signal are required for ubiquitination. In striking contrast, ubiquitination of the ζ chain of the T-cell receptor is independent of any particular lysine residue and proceeds as long as one residue is present in the cytosolic tail of the molecule [8]. Similarly, no single specific lysine residue is required for ubiquitination of either c-Jun [9] or cyclin B [10]: any single lysine residue, even artificially inserted, can serve as a ubiquitin acceptor. Important in this context is that only in a handful of cases it has been shown directly, via chromatographic or mass spectrometric analyses, that ubiquitin is indeed anchored to a lysine residue (see for example Refs. [11, 12]). In most cases studied, and there are not too many, the assumption that an internal lysine serves as the polyubiquitin chain anchor is indirect and based on mutational analyses. One interesting case involves the artificial fusion protein ubiquitin-Pro-X-βgalactosidase. In this chimera, the ubiquitin moiety was fused to the N-terminal Pro residue of the protein. Unlike other ubiquitin-B-X-β-galactosidase species (where B is any of the remaining 19 amino acid residues), here ubiquitin is not removed by isopeptidases and serves as a degradation signal following generation of a polyubiquitin chain that is anchored to Lys48 of the artificially fused ubiquitin moiety [13]. However as noted, in this case the ubiquitin moiety was fused to the N-terminal residue artificially. The first substrate that was identified in which the N-terminal residue serves as a ubiquitination target was MyoD. The basic helix–loop–helix (bHLH) protein MyoD is a tissue-specific transcriptional activator that acts as a master switch for muscle development. MyoD forms heterodimers with other proteins belonging to the bHLH group, such as the ubiquitously expressed E2A, E12 and E47. These dimers are probably the transcriptionally active forms of the factor. Association of MyoD with HLH proteins of the Id family (inhibitors of differentiation that lack the basic domain) inhibits its DNA-binding and biological activities. MyoD is a short-lived protein with a half-life of ∼45 min [14, 15]. Degradation of MyoD is mediated by the ubiquitin system both in vitro and in vivo. Furthermore, the process is inhibited by its consensus DNA-binding site. In contrast, addition of Id1 destabilizes the MyoD–E47–DNA complex and renders the protein susceptible to degradation [15].

2 Results

2 Results

To analyze specific ubiquitination sites in MyoD, we used site-directed mutagenesis to substitute systematically all the lysine residues with arginines [16]. The protein contains nine lysine residues, most of them located within the N-terminal domain of the molecule. The nine residues are in positions 58, 99, 102, 104, 112, 124, 133, 146 and 241. The various proteins were generated either by expression in bacteria followed by purification, or by in vitro translation in reticulocyte lysate in the presence of [35 S]methionine. Conjugation and degradation of the proteins were monitored in a reconstituted cell-free system or in cells. Proteins were detected by either Western blot analysis or PhosphorImaging. Surprisingly, even a MyoD species that lacked all lysine residues was still degraded efficiently in an ATPdependent manner in vitro. To demonstrate involvement of the ubiquitin system in the process, we followed the degradation of wild-type (WT) and lysine-less (LL) MyoD in the absence and presence of ubiquitin. Similar to the degradation of the WT protein, degradation of the LL MyoD was completely dependent upon the addition of exogenous ubiquitin to an extract that does not contain it (Fraction II). Furthermore, addition of methylated ubiquitin, which cannot form polyubiquitin chains and serves as a chain terminator [17], inhibited the degradation of LL MyoD. The inhibition could be alleviated by the addition of excess of free ubiquitin. These results strongly suggested that polyubiquitination of LL MyoD is necessary for degradation of the protein. Furthermore, they implied that the polyubiquitin chain is synthesized on internal lysine residues of ubiquitin. To demonstrate directly polyubiquitinated LL MyoD, we used in-vitro-translated 35 S-labeled protein in a partially reconstituted system. We demonstrated that LL MyoD generates high molecular mass ubiquitinated adducts. It should be noted, however, that these conjugates are of somewhat lower molecular mass than those of the WT MyoD. This can be attributed to the role that the internal lysine residues also play in the process (see also below). To investigate the physiological relevance of the observations in the cell-free system, we followed the fate of the different MyoD lysine-mutated proteins in vivo, using pulse-chase labeling experiments in COS-7 cells that were transiently transfected with the different MyoD cDNAs. In agreement with our in vitro data, the lysine-less MyoD protein is degraded efficiently in cells as well. However, we could observe a progressive increase in the half-life of the proteins of up to ∼2-fold with the gradual substitution of the lysine residues. While the half-life of WT MyoD was ∼50 min, that of LL MyoD was ∼2 h. Interestingly, we found that the stability of MyoD is not affected by the substitution of any specific lysine residue, and it is the total number of these residues that determines the half-life of the protein. To identify the system involved in the destruction of LL MyoD in vivo, transfected cells were incubated in the presence of inhibitors of proteasomal and lysosomal degradation. Chloroquine, a general inhibitor of lysosomal proteolysis, and E-64, a cysteine protease inhibitor that affects lysosomal, but also certain cytosolic proteases, had no effect on the stability of the LL MyoD. In striking contrast, the proteasomal

3


inhibitors MG132 and lactacystin blocked degradation of the LL protein significantly. To demonstrate the intermediacy of ubiquitin conjugates in the degradation of LL MyoD, we incubated COS-7 cells, transiently transfected with either WT or LL MyoD cDNAs, with MG132, and followed generation of ubiquitin-MyoD adducts. Immunoprecipitation with anti-MyoD antibody followed by Western blot analysis with anti-ubiquitin antibody revealed accumulation of high molecular mass compounds in cells transfected with either WT or LL MyoD. A similar analysis of mock-transfected cells clearly demonstrated the specificity of both the anti-MyoD and anti-ubiquitin antibodies. Based on these results, it was clear that polyubiquitination is essential for targeting MyoD for degradation. The lack of internal lysine residues, the only known targets for ubiquitin modification, made it important to identify the functional group that can serve as an attachment site for ubiquitin. Chemically, several groups can generate covalent bonds with ubiquitin. Ser and Thr can participate in ester bond formation, while Cys can generate a thiol ester bond. However, these bonds are unstable and are hydrolyzed in either high pH (Ser and Thr) or high concentration of -SH groups (Cys). The stability of the MyoD-ubiquitin adducts under these conditions made it highly unlikely that any of these modifications is the one we observed. A likely candidate, however, was the free amino group of the N-terminal residue of the protein, which can generate a stable peptide bond with the C-terminal Gly residue of ubiquitin. Edman degradation of the N-terminal residue of bacterially expressed, in-vitro-translated and cellularly expressed MyoDs, has revealed that the ubiquitin attachment site can be the free, unmodified initiator methionine: the proteins were not modified and the N-terminal residue was not acetylated. To demonstrate a role for the free N-terminal amino group in the degradation of MyoD, we chemically modified this group. Initially, we blocked this group in the LL MyoD protein by reductive methylation. While this procedure blocks all amino groups in a protein in a non-discriminatory manner, in this case, it could have been only the α-NH2 group, which is the only free amino group left in the MyoD molecule. The modification stabilized the protein completely. Whereas a free α-NH2 appears to be sufficient for degradation (probably following ubiquitination) of LL MyoD, it is not clear whether it also plays a physiological role in targeting the WT molecule, which has nine available lysine residues. In order to investigate the role and biological relevance of the free α-NH2 group in the targeting of WT MyoD, we selectively blocked it by carbamoylation with potassium cyanate at low pH. This procedure does not modify ε-NH2 groups of internal lysine residues. Automated Edman degradation along with fuorescamine determination of the extent of remaining free NH2 groups confirmed that the modification affected only the N-terminal group. The modified protein was subjected to in vitro degradation and conjugation in cell extract. In contrast to LL MyoD, the N-terminally carbamoylated protein could not be ubiquitinated and was stable. Thus, a free and exposed NH2 terminus of MyoD appears to be an essential site for degradation, most probably because it serves as an attachment site for the first ubiquitin moiety. As an additional control, we selectively modified the internal lysine residues of WT MyoD by guanidination

2 Results

with O-methylisourea. The modification, which does not affect the N-terminal group, generates a protein that is essentially the chemically modified counterpart of the LL MyoD that was generated by site-directed mutagenesis. Similar to the LL protein, this MyoD derivative is degraded efficiently in the cell-free system in a ubiquitin-and an ATP-dependent mode. To analyze the role of the N-terminal residue of MyoD as a ubiquitination site, we fused to WT MyoD, upstream to the N-terminal residue, a 6 × Myc tag, and monitored the stability of the tagged protein. We showed that it is stable both in vitro and in vivo. It should be noted that the two first N-terminal residues of the Myc tag, methionine and glutamate, are identical to the first two N-terminal residues in MyoD. In addition, the Myc tag also contains a lysine residue. Thus, altogether, six additional lysine residues were added to WT MyoD in addition to its own nine native residues. Nevertheless, the tag stabilizes it, probably by blocking access to a specific N-terminal residue, and, as became clear later (see below), to its neighboring domain. Taken together, these findings strongly suggested that MyoD is first ubiquitinated at its N-terminal residue, and the polyubiquitin chain is synthesized on this first conjugated ubiquitin moiety. Internal lysine residues also play a role, probably by serving as additional anchoring sites, whose ubiquitination accelerates degradation. Yet they are not essential for proteolysis to occur. In contrast, ubiquitination of the N-terminal residue plays a critical role in governing the protein’s stability [16] (Figure 1). Using a similar, though not a complete, set of experiments, 12 additional proteins have been identified recently that appear to undergo N-terminal ubiquitination: (i) the human papilloma virus-16 (HPV-16) E7 oncoprotein (18), (ii) the latent membrane protein 1 (LMP1)(19) and (iii) 2A (LMP2A)(20) of the Epstein Barr virus (EBV), involved in viral activation from latency, (iv) the cell cycle-dependent kinase (CDK) inhibitor p21 (21,22), (v) the extracellular signal-regulated kinase 3, ERK3 (22), the inhibitors of differentiation (vi) Id2 (23) and (vii) Id1 (24), two pro-proliferative Helix-Loop-Helix proteins, (viii) hydroxymethyglutaryl-Coenzyme A reductase (HMG-CoA reductase), the first and key regulatory enzyme in the cholesterol biosynthetic pathway (25), (ix) p19ARF , the mouse Mdm2 inhibitor and (x) p14ARF , its human homologue (26), (xi) the HPV-58 E7 oncoprotein, and (xii) the cell cycle regulator p16INK4a (27). As for HMG-CoA reductase, several specific internal lysine residues have also been shown to be important for its targeting, and therefore the essentiality of the N-terminal residue in the process has to be further substantiated. The case of p21 requires further investigation, as one study reported that its degradation by the proteasome does not require ubiquitination [28], while an independent study has demonstrated a role for Mdm2 in targeting p21, also without a requirement for ubiquitination [29]. As we noted for MyoD, substitution of the internal lysines inhibited slightly (up to twofold) both conjugation and degradation of HPV-16, LMP1, and Id2, suggesting that these residues, probably also by serving as ubiquitin anchors, can modulate the stability of these proteins. It is possible to suppose that N-terminal ubiquitination and modification of internal lysines is catalyzed by different ligases that may be even located in different subcellular

5


NH

A

Lys48 - based polyubiquitin chain

K48 G76

C=O NH Ubiquitin K48 G76 C=O NH NH2

X

Kn

COOH

Protein target

B

NH

Lys48 – based polyubiquitin chain

K48 G76

C=O NH NH2

K48 Ubiquitin G76

C NH O

X

Fig. 1 Ubiquitination on an internal lysine and on the N-terminal residue of the target substrate. (A) The first ubiquitin moiety is conjugated, via its C-terminal Gly76 residue, to the g-NH2 group of an internal lysine residue of the target substrate (Kn ). (B) The first ubiquitin moiety is conjugated to a free

Kn

COOH

Protein target "-NH2 group of the N-terminal residue, X. In both cases, successive addition of activated ubiquitin moieties to internal Lys48 on the previously conjugated ubiquitin moiety leads to the synthesis of a polyubiquitin chain which serves as the degradation signal for the 26S proteasome.

compartments (e.g. the nucleus and cytosol). Because of the role that internal lysines play in modulating the stability of these proteins, and in order to better understand the physiological significance of this novel mode of modification, it was important to identify proteins whose degradation is completely dependent on N-terminal ubiquitination. An important group of potential substrates for Nterminal ubiquitination is that of naturally occurring lysine-less proteins – NOLLPs. Since these proteins cannot use the “canonical” lysine conjugation pathway, in order to be targeted by the ubiquitin system they must use, an alternative site for their tagging. Searching the database, we were able to identify 177 eukaryotic NOLLPs, 14 of which occur in humans. In addition, we have identified 111 viral NOLLPs. We have shown that two of the proteins mentioned above, the human tumor supressor p16INK4a and the viral oncoprotein HPV-58 E7 are degraded via

2 Results

the N-terminal ubiquitination pathway [27]. Interestingly, we demonstrated that p16INK4a is ubiquitinated and degraded only in sparse cells, and is stable in dense cells. Similar findings were reported for the NOLLPs p19ARF and p14ARF (26). For E7-16 [18], LMP1 [19], and MyoD (unpublished), it has been shown that truncation of a short N-terminal segment of 10–20 residues stabilized the proteins, suggesting that the entire domain beyond the single N-terminal residue plays a role in governing the stability of these proteins. Such a segment can allow the mobility/flexibility necessary for the N-terminal residue to serve as a ubiquitin acceptor. It can also serve as a recognition domain for the cognate E3. There is no homology between the N-terminal domains of these three proteins, suggesting that if the three N-terminal domains serve as recognition motifs, they recognize different components of the ubiquitin system. Interestingly, the LMP2A E3 was identified as a member of the NEDD4 family of HECT domain ligases, AIP4 and/or WWP2 [20]. A PY motif in LMP2A is recognized by the E3. It resides in the Nterminal domain of the molecule, supporting the hypothesis that in these proteins the E3-binding domain may reside in close proximity to the N-terminal residue ubiquitination site. Is there any direct evidence for N-terminal ubiquitination? All the different and independent lines of evidence in the various studies strongly suggest that ubiquiti-nation occurs on the N-terminal residue, and any other scenario is highly unlikely. Yet, the only direct evidence must be demonstration of a fusion peptide between the C-terminal domain of ubiquitin and the N-terminal domain of the target substrate. The study on p21 [21] and E7-58 [27] brought us a little closer. Bloom and colleagues [21] transfected cells with N-terminally His-tagged ubiquitin and N-terminally HA-tagged p21 that contained a Factor X proteolytic site immediately after the HA tag and upstream of the p21 reading frame. They then immunoprecipitated and resolved the cell-generated ubiquitin conjugates of p21 and treated the mono-ubiquitin–p21 adduct with Factor-X protease. This treatment released a smaller species of p21 (lacking His-ubiquitin and the HA-tag-Factor X site) and His-ubiquitin-HA-Factor X site, thus demonstrating that the HA-tag-Factor X site, which was previously part of p21, had now become part of the Factor X-cleaved ubiquitin. A similar experimental evidence was brought for the NOLLP HPV E7-58 [27]. Here, Ben-Saadon and colleagues generated two species of the protein containing the eight amino acid sequence of the Tobacco Etch Virus (TEV) protease cleavage site inserted either 21 amino acid residues after the iMet [E7-58-TEV(21)] or immediately after the iMet [E7-58-TEV(1)]. The prediction from this experiment was that if ubiquitin is indeed attached to the N-terminal residue of E7-58, TEV protease-catalyzed cleavage will generate an extended ubiquitin molecule that will also contain the respective N-terminal domain of E7-58 [21 residues or 1 residue, respectively, dependent upon whether the substrate of the reaction is E7-58-TEV(21) or E7-58-TEV(1), and the six amino acids derived from the TEV cleavage site]. Such extended ubiquitin moieties were indeed generated following incubation of the substrate with labeled methylated ubiquitin (which generated mostly the mono-ubiquitin adduct of the E7-58 protein), followed by cleavage of the adduct with TEV. The only conclusion that can be derived from these experiments

7


is that the ubiquitin moiety was fused to any of the amino acid residues of the HA tag-Factor X site at the N-terminal domain of p21, or to any of the first 21 amino acids of E7 or the TEV site (part of it; the protease cleaves after the sixth amino acids out of eight in the complete site). Such internal modification is unlikely, however, as it must require a novel chemistry since none of the residues in the tags, the protease sites or the E7 N-terminal fragment, is lysine. Yet, formally, it is still possible that such a modification occurs. The HA tag contains, for example, three Tyr residues. Thus, the evidence provided by these two experiments clearly limits an unlikely non-peptide bond ubiquitination, such as esterification, to a much smaller zone in the N-terminal domain of the tagged p21 or the TEV-containing E7-58, but does not demonstrate directly that the modification occurs indeed on the N-terminal residue. As noted, only identification of a fusion peptide between the C-terminal domain of ubiquitin and the N-terminal domain of the target protein will constitute such an evidence. Ben-Saadon and colleagues have recently isolated the long sought after fusion peptide [27]: mass spectrometric analysis of a tryptic digest of the isolated mono-ubiquitin adduct of HPV-58 E7 revealed a peptide of 11 amino acids, GG-MHGNNPTLR which represents the last two C-terminal amino acids of ubiquitin, GlyGly, and the first nine residues of E7, MetHisGlyAsnAsnProThrLeuArg (MHGNNPTLR). It should be noted that WT E7-58 contains an Arg residue in position 2. It was necessary to substitute this Arg with His, since otherwise the digesting enzyme, trypsin, would have generated a tetrapeptide, GG-MR, that would have been difficult, if not impossible, to identify in the MS analysis. MS/MS analysis of the 11-mer, verified its internal sequence. Coulombe and colleagues were also able to isolate and identify the sequence of a fusion peptide between the C-terminal domain of ubiquitin and the N-terminal domain of HA-tagged p21 that also contained, downstream of the tag, a stretch of residues derived from the N-terminal domain of the native substrate, but without the iMet (which was removed during the construction of the tagged protein) [22].

3 Discussion

N-terminal ubiquitination is a novel pathway, clearly distinct from the N-end rule pathway [30]. In the latter, the N-terminal residue serves as a recognition and binding motif to the ubiquitin ligase, E3α; however, ubiquitination occurs on an internal lysine(s). In contrast, in the N-terminal ubiquitination pathway, modification occurs on the N-terminal residue, whereas recognition probably involves a downstream motif. It should be mentioned that in yeast, using the model fusion protein ubiquitin-Pro-X-β-galactosidase (where X is a short sequence derived from the λ repressor), a new proteolytic pathway has been described, designated the UFD (ubiquitin fusion degradation) pathway [31]. The stably fused ubiquitin moiety (note that in this exceptional case, with Pro and not any other amino acid residue as the linking residue, the ubiquitin moiety is not cleaved off by ubiquitin C-terminal

3 Discussion

hydrolases), functions as a degradation signal, where its Lys48 serves as an anchor for the synthesized polyubiquitin chain. This pathway involves several enzymes, UFD 1–5, some of which appear to be unique and are not part of the “canonical” UPS. It is possible that N-terminal ubiquitination is the most upstream event in the UFD pathway – which was discovered using an artificial chimeric ubiquitin–protein model substrate: the N-terminal ubiquitination pathway can function by providing substrates to the UFD pathway. The physiological significance of N-terminal ubiquitination is still obscure. Naturally occurring lysine-less proteins, NOLLPs, that are degraded by the ubiquitin system must traverse this pathway. Many such proteins, mostly viral, can be found in the database (see above and in Refs. [26, 27]). We believe that many additional lysine-containing proteins, will be discovered to be targeted via this novel mode of modification. Of note is that all the proteins that are N-terminally ubiquitinated must contain a free, unmodified N-terminal residue. Such proteins constitute approximately 25% of all cellular proteins, while the remaining 75% are Nα-acetylated. Whether a protein will be acetylated is dependent on the structure of the N-terminal domain of the mature protein. This is determined by the combined activities of methionine aminopeptidases (MAPs) and N-terminal acetyltransferases (NATs), which are dependent on the specific sequence of up to the first four N-terminal residues of the target protein substrates (reviewed in Ref. [32]). Thus, it is possible to predict which proteins will be potential substrates of the N-terminal ubiquitination pathway. Internal C-terminal fragments of Nα-acetylated proteins can also be modified by ubiquitin at their “new” N-terminal residue following limited processing. Many proteins, such as the NF-κB precursors p105 and p100 or caspase substrates are processed initially in a limited manner, generating a C-terminal fragment with a newly exposed N-terminal residue. For all lysine-containing proteins, the intact free N-termini as well as the products of processing, the assumption is that their internal lysines are not easily accessible, for whatever reason, for ubiquitination, and it is only the N-terminal residue that can be modified. Interestingly, most of the substrates identified thus far have a few lysine residues that might not be accessible to the E3s: For example, MyoD has nine (out of 319), E7 two (out of 97), LMP1 has a single lysine residue (out of 440), LMP2A has three (out of 497), Id2 nine (out of 134) and p21 six (out of 164). From the random discovery of thirteen N-terminally ubiquitinated proteins, it appears that their number could well be larger and that many more will be discovered, which will help in the unraveling of the unique characteristics that distinguish this group of substrates.

Acknowledgments

Research in the laboratory of A.C. is supported by grants from Prostate Cancer Foundation Israel - Centers of Excellence Program, the Israel Science Foundation – Centers of Excellence Program, a Professorship funded by the Israel Cancer Research Fund (ICRF) and the Foundation for Promotion of Research in the Technion administered by the Vice President of the Technion for Research. Infrastructural

9


equipment was purchased with the support of the Wolfson Charitable Fund Center of Excellence for studies on Turnover of Cellular Proteins and its Implications to Human Diseases.

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10 KING, R. W., GLOTZER, M. and KIRSCHNER, M. W. Mutagenic analysis of the destruction signal of mitotic cyclins and structural characterization of ubiquitinated intermediates. Mol. Biol. Cell, 1996, 7, 1343–1357. 11 GOLDKNOPF, I. L. and BUSCH, H. Isopeptide linkage between nonhistone and histone 2A polypeptides of chromosomal conjugate-protein A24. Proc. Natl. Acad. USA, 1977, 74, 864–868. 12 GRONROOS, E., HELLMAN, U., HELDIN, C. H. and ERICSSON, J. Control of Smad7 stability by competition between acetylation and ubiquitination. Mol. Cell, 2002, 10, 483–493. 13 JOHNSON, E. S., BARTEL, B., SEUFERT, W. and VARSHAVSKY, A. Ubiquitin as a degradation signal. EMBO J. 1992, 11, 497–505. 14 THAYER, M. J., TAPSCOTT, S. J., DAVIS, R. L., WRIGHT, W. E., LASSAR, A. B. and WEINTRAUB, H. Positive autoregulation of the myogenic determination gene MyoD1. Cell, 1989, 58, 241–248. 15 ABU-HATOUM, O., GROSS-MESILATY, S., BREITSCHOPF, K., HOFFMAN, A., GONEN, H., CIECHANOVER, A. and BENGAL, E. Degradation of the myogenic transcription factor MyoD by the ubiquitin pathway in vivo and in vitro: regulation by specific DNA binding. Mol. Cell. Biol., 1998, 18, 5670–5677. 16 BREITSCHOPF, K., BENGAL, E., ZIV, T., ADMON, A. and CIECHANOVER, A. A novel site for ubiquitination: The N-terminal residue and not internal lysines of MyoD is essential for conjugation and degradation of the protein. EMBO J., 1998, 17, 5964–5973. 17 HERSHKO, A. and HELLER, H. Occurrence of a polyubiquitin structure in ubiquitin-protein conjugates. Biochem. Biophys. Res. Commun. 1985, 128, 1079–1086.

References 18 REINSTEIN, E., SCHEFFNER, M., OREN, M., SCHWARTZ, A. L. and CIECHANOVER, A. Degradation of the E7 human papillomavirus oncoprotein by the ubiquitin-proteasome system: 2 N-terminal Ubiquitination: No Longer Such a Rare Modification targeting via ubiquitination of the N-terminal residue. Oncogene, 2000, 19, 5944–5950. 19 AVIEL, S., WINBERG, G., MASSUCCI, M. and CIECHANOVER, A. Degradation of the Epstein-Barr virus latent membrane protein 1 (LMP1) by the ubiquitin-proteasome pathway: targeting via ubiquitination of the N-terminal residue. J. Biol Chem., 2000, 275, 23491–23499. 20 IKEDA, M., IKEDA, A. and LONGNECKER, R. Lysine-independent ubiquitination of the Epstein-Barr virus LMP2A. Virology, 2002, 300, 153–159. 21 BLOOM, J., AMADOR, V., BARTOLINI, F., DEMARTINO, G. and PAGANO, M. Proteasome-mediated degradation of p21 via N-terminal ubiquitinylation. Cell, 2003, 115, 1–20. 22 COULOMBE, P., RODIER, G., BONNEIL, E., THIBAULT, P. and MELOCHE, S. N-Terminal ubiquitination of extracellular signal-regulated kinase 3 and p21 directs their degradation by the proteasome. Mol. Cell Biol., 2004, 24, 6140–6150. 23 FAJERMAN, I., SCHAWARTZ, A. L. and CIECHANOVER, A. Degradation of the Id2 developmental regulator: Targeting via N-Terminal Ubiquitination. Biochem. Biophys. Res. Commun., 2004, 314, 505–512. 24 AZAR-TRAUSCH, J. S., LINGBECK, J., CIECHANOVER, A. and SCHWARTZ, A. L. Ubiquitin-proteasome-mediated degradation of Id1 is modulated by MyoD. J. Biol. Chem., 2004, 279, 32614–32619. 25 DOOLMAN, R., LEICHNER, G. S., AVNER, R. and ROITELMAN, J. Ubiquitin is conjugated by membrane ubiquitin ligase to three

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sites, including N-terminus, in transmembrane region of mammalian 3-hydroxy-3- methylglutaryl coenzyme a reductase: Implications for sterol-regulated enzyme degradation. J. Biol. Chem., 2004, 279, 38184–38193. KUO, M. L., DEN BESTEN, W., BERTWISTLE, D., ROUSSEL, M. F., and SHERR, C. J. N-terminal polyubiquitination and degradation of the Arf tumor suppressor. Genes and Dev., 2004, 18, 1862–1874. BEN-SAADON, R., FAJERMAN, I., ZIV, T., HELLMAN, U., SCHWARTZ, A. L., and CIECHANOVER, A. The Tumor Suppressor Protein p16INK4a and the Human Papillomavirus oncoprotein E7-58 are Naturally Occurring Lysine-Less Proteins that are Degraded by the Ubiquitin System: Direct Evidence for Ubiquitination at the N-Terminal Residue. J. Biol. Chem., 2004, 279, 41414–41421. SHEAFF, R. J., SINGER, J. D., SWANGER, J., SMITHERMAN, M., ROBERTS, J. M. and CLURMAN, B. E. Proteasomal turnover of p21Cip1 does not require p21Cip1 ubiquitination. Mol. Cell, 2000, 5, 403–410. JIN, Y., LEE, H., ZENG, S. X., DAI, M. S. and LU, H. Mdm2 promotes p21waf cip1 proteasomal turnover independently of ubiquitylation. EMBO J., 2003, 22, 6365–6377. VARSHAVSKY, A. The N-end rule: Functions, mysteries, uses. Proc. Natl. Acad. Sci. USA, 1996, 93, 12142–12149. JOHNSON, E. S., MA, P. C., OTA, I. M. and VARSHAVSKY, A. A proteolytic pathway that recognizes ubiquitin as a degradation signal. J. Biol. Chem., 1995, 270, 17442–17456. POLEVODA, B. and SHERMAN, F. N-terminal acetyltransferases and sequence requirements for N-terminal acetylation of eukaryotic proteins. J. Mol. Biol., 2003, 325, 595–622.

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1

The COP9 Signalosome and Its Role in the Ubiquitin System Dawadschargal Bech-Otschir

Western General Hospital, Edinburgh, United Kingdom

Barbara Kapelari Max-Planck-Institute for Biochemistry, Martinsried, Germany

Wolfgang Dubiel Charité-Universitätsmedizin Berlin, Berlin, Germany

Originally published in: Protein Degradation, Volume 1. Edited by R. John Mayer, Aaron Ciechanover and Martin Rechsteiner. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30837-8

1 Introduction

The COP9 signalosome (CSN) is a multimeric, highly-conserved protein complex [1]. Just like the ubiquitin system it occurs in all studied eukaryotic cells. Following its 1994 discovery in plant cells the complex was postulated to function in signal transduction [2]. Originally described as a regulator of light-dependent growth in plants [3, 4], identification and characterization of the CSN from mammalian cells led to the discovery of sequence homologies between CSN subunits and subunits of the 26S proteasome lid complex [5, 6] as well as subunits of the translation-initiation complex eIF3 [7]. Significant progress has been made towards understanding its structure and function by analyzing different eukaryotic organisms. The complex is involved in developmental processes of plants [8] and Drosophila [9] and is essential for embryogenesis in mice [10]. It seems to participate in processes such as DNA repair [11], cell-cycle regulation [12] and angiogenesis [13]. At the moment the pleiotropic effects of the CSN can be explained by its regulatory impact on the ubiquitin system. Here we provide a summary of current knowledge of CSN function in the ubiquitin system.


2 The COP9 Signalosome and Its Role in the Ubiquitin System

2 Discovery of the CSN

Deng and co-workers discovered the CSN in Arabidopsis thaliana when they characterized mutants of light-dependent development, and they called it the COP9 complex [2]. Morphogenesis of germinating seedlings is light-dependent. Light triggers a developmental process called photomorphogenesis. A number of mutations in the Arabidopsis COP/DET/FUS loci result in the loss of the COP9 complex accompanied with cop phenotypes in which germinating seedlings exhibit light-independent expression of light-induced genes [3]. Therefore the complex was originally hypothesized to be a repressor of photomorphogenesis [14]. The mammalian CSN complex was independently isolated during preparations of the 26S proteasome and called the JAB1-containing signalosome [15]. The same studies identified proteins such as JAB1 [15] and TRIP15 [16] as components of the complex and revealed homologies between subunits of the CSN and components of the 26S proteasome lid complex. Purification and analysis of the complex from Arabidopsis, pork spleen [6, 17] and human red blood cells [5, 18] led to the conclusion that each subunit of the CSN has its paralog subunit in the 26S proteasome lid complex. These data suggested a common origin for the two complexes during evolution. Because they have similar architectures, the two complexes have been postulated to perform related functions (see below). Unfortunately there is only limited information on the structure or function of the eIF3 complex, and its relationship to the CSN and the lid is not well understood [7]. Studies have revealed that the CSN possesses both intrinsic and extrinsic (associated) activities, which will be reviewed in detail below. Historical gene names of the CSN have been summarized before [1]. In this article we use the unified nomenclature of the CSN [1].

3 Architecture of the CSN 3.1 CSN Subunit–Subunit Interactions

The CSN is composed of eight subunits called CSN1 to CSN8, which are highly conserved in eukaryotes, although only six of them occur in fission yeast. Two hybrid screens and biochemical methods such as far westerns, pull downs and co-precipitation defined a number of CSN subunit–subunit interactions. Figure 1 illustrates known subunit–subunit interactions. Initial insight into the architecture of CSN came from the first 2D electron microscopic analysis of purified CSN from human red blood cells [19] (see also Figure 2 below). The CSN architecture shows similarity to that of the lid. Both complexes have an asymmetric arrangement of their subunits and exhibit a central groove structure [19]. Whether the structural similarity of the two complexes is connected with

3 Architecture of the CSN 3

Fig. 1 Subunit–subunit interactions of the CSN and interactions of CSN subunits with other proteins. Subunits are numbered according to the unified nomenclature [1]. CSN subunit–subunit interactions have been published before [19]. Darker shad-

ing indicates subunits with MPN domains and lighter those with PCI domains. Known phosphorylated subunits are indicated. Details on CSN subunit interactions with other proteins can be found in the text.

similar functions remains unclear. The exact arrangement of CSN and lid subunits within their complexes remains uncertain in the absence of high-resolution crystal structures for the two complexes. Interestingly, the occurrence of smaller CSN sub-complexes apart from the large 500-kDa CSN complex has been described in different species such as Arabidopsis, Drosophila, Schizosaccharomyces pombe and mammalian cells (for a review see Ref. [20]). At the moment the physiological function of CSN sub-complexes is unclear. It can be speculated that a controlled equilibrium exists between the large and small CSN complexes. The small complexes may have a function in shuttling between nucleus and cytoplasma and/or between large multi-subunit complexes such as the 26S proteasome and cullin-based Ub ligases. 3.2 CSN-subunit Interactions With Other Proteins

Apart from subunit–subunit interactions within the CSN, a considerable number of cellular proteins interact with CSN subunits (see Figure 1). Although the


physiological relevance of many of the identified interactions is questionable, most of them might be attributed to a role of the CSN complex in signal transduction and ubiquitin-dependent proteolysis. CSN1 formerly called Gps1 was first described as a signal transduction repressor in Arabidopsis [21]. Over-expression of CSN1 suppresses the activated JNK signaling pathway and also inhibits UV- and serum-induced c-fos expression as well as MEKKactivated AP1-activity in mammalian cells [21, 22]. It remains unclear whether overexpressed CSN1 plays a role as dominant negative regulator when in the CSN complex. Whereas the N-terminal region of CSN1 is sufficient for repression, the C-terminal region is necessary for its integration into the complex and for the stability of the CSN complex [22]. Curiously, the N-terminal region of CSN1 may be not required for the CSN-associated deneddylation of cullin 1 (CUL1) and cullin 3 (CUL3), components of cullin-based E3 Ub ligases in Arabidopsis, although it appears to be one of the binding sites of the CSN for cullin-based complexes [23]. Moreover, CSN1 is the receptor site for the interaction of the CSN with inositol 1,3,4-trisphosphate 5/6 kinase [24]. In addition, CSN1 represents the interactor for a subunit of the translation-initiation factor 3, eIF3c/NIP1 [25], and for the 26S proteasome non-ATPase subunit Rpn6 [26]. Possible functions of these interactions are discussed later. CSN2 also known as alien [27] is perhaps an important regulatory subunit of the CSN. Firstly, CSN2 was identified as Trip15 (thyroid hormone receptor-interacting protein) using a yeast-two-hybrid screen [16]. The binding site of CUL1 and CUL2 is located at the N-terminal region of CSN2. This interaction is important for the CSNmediated deneddylation of cullin-based complexes that regulate their Ub-ligase activity [28]. Additionally, CSN2 binds to the transcription factor, ICSBP (interferon consensus sequence binding protein), which modulates interferon-directed gene expression [29]. Moreover it interacts with the nuclear receptors DAX-1, COUP1TF1 and ecdysone receptor [27, 30]. Interestingly, CSN2 is phos-phorylated by the CSN-associated kinases CK2 and PKD [19, 31]. However, the phosphorylation sites and their physiological function remain unclear. The CSN3 subunit interacts with IKKγ , a component of the IκB-kinase complex controlling NF-κB activity [32]. Additionally, it is the binding site for the CSNassociated kinases CK2 and PKD [31]. The subunit of the translation-initiation factor 3 complex, Int6/eIF3e, and the ubiquitin-conjugating enzyme variant, COP10, have been identified as other cellular interactors [33, 34]. Also the HIV-1 Tat protein interacts with CSN3 (our unpublished data). CSN4 is a poorly studied subunit of the CSN. Only one interactor of CSN4 has been identified, the ubiquitin-conjugating enzyme COP10 [34]. CSN5 appears to be a most important subunit both in terms of interactions with other cellular proteins and because it is a component with intrinsic metalloprotease activity (see below). The binding of CSN5 to cellular proteins including the transcription factors p53 [35] and c-Jun [15], the cell-cycle regulator protein p27 [36], rLHR (lutropin/choriogonadotropin receptor precursor) [37], Smad4 (TGF-β signaling pathway common effector) [38] and HIF1α (hypoxia-inducible factor 1) [39] appears to regulate their metabolic stability. In many cases the CSN5-interacting

3 Architecture of the CSN 5

proteins are phosphorylated by the CSN-associated kinases, which determines the speed of their destruction [40]. In contrast, Id1 and Id3 binding to the CSN complex via CSN5 leads to their stabilization, not to their phosphorylation [41]. The interaction of CSN5 to the member of the IκB multigene family Bcl3, the progesterone receptor PR, and the steroid receptor co-activator SRC-1, leads to stabilization of Bcl3-p50 and PR-SRC-1 complexes and enhances transcriptional activity [42, 43]. Whereas AP-1 activity is stimulated by interaction of CSN5 with the integrin adhesion receptor LFA-1 [44], the opposite effect was reported in the case of the cytokine migration inhibitor factor, MIF [45]. Additionally, there are other published interactors of CSN5 including the membrane-associated RING-finger Ub ligase TRC8 [46], hepatopoietin (HPO) [47], germ-line RNA helicases (GLHs) [48], and the ubiquitin C-terminal hydrolase PGP9.5 [49]. However, the exact role of these interactions remains unclear. Several groups reported the occurrence of a free CSN5 subunit [50] or CSN5 as a component of a smaller complex [51], although the exact physiological function of the different CSN5 forms is so far unclear. It is also unknown whether the occurrence of the different CSN5 forms is regulated. Moreover, little is known about CSN5 interactions in vivo, how they are regulated and under what circumstances they take place. CSN6 like CSN8 exists in eukaryotes except in S. pombe [52]. There are only a few published interactions of CSN6 with other cellular proteins. It binds to the HIV-1 Vpr protein affecting cell-cycle-associated signaling [53] and to the RING-finger protein of the SCF-complex, Rbx1 [54, 55]. In addition, CSN6 is another binding site for Int-6/eIF3e [33]. Interestingly, in mammalian cells two homologs of CSN7, CSN7a and CSN7b, have been found [6]. S. pombe contains only one form of CSN7 whereas Arabidopsis contains two alternative splicing variants, CSN7i and CSN7ii [52, 56]. CSN7 interacts with the polyamine-modulated factor PMF-1 [57]. Interestingly, CSN7 also binds the protein kinase CK2, one of the CSN-associated kinases, which phosphorylates CSN7 [31]. Whether the phosphorylated form of CSN7 is necessary for CSN complex assembly or for other regulatory events is unclear. Little is known of CSN8 interactions. CSN8 like CSN3 and CSN4 binds to COP10 [34]. 3.3 PCI and MPN

Six of the CSN subunits contain PCI (proteasome, COP9 signalosome, initiation factor 3) domains and two contain MPN (Mpr-Pad1-N-terminal) domains [58]. These two characteristic domains have been found in three protein complexes: the CSN, the 26S proteasome lid complex (lid) and the eIF3 complex. The two domains are composed of about 150 to 200 amino acids at the N- or C-terminus of the CSN subunits. Apparently, the PCI domain has been shown to be important for interactions between CSN subunits. Thus, it might have a scaffolding function [22, 59].


The CSN subunit CSN5 has been shown to contain a metalloprotease motif localized on its MPN domain, which is essential for the cleavage of the ubiquitinlike modifier NEDD8 from cullins [60] (see below). Apart from the catalytic activity of the MPN domain of CSN5 it appears to be the receptor for different cellular proteins associated with the CSN complex (see above and Figure 1). Interestingly, an MPN domain similar to that of CSN5 is located in the N-terminal region of CSN6. However, this MPN domain has no deneddylation catalytic center like CSN5. The function of the CSN6 MPN domain remains obscure.

4 Biochemical Activities Associated With the CSN 4.1 Deneddylation Activity

Studies in fission yeast and Arabidopsis have revealed that the CSN has a role in the cleavage of NEDD8 from cullins [54, 55, 61]. The MPN domain of CSN5, like its paralog subunit Rpn11 of the 26S proteasome lid complex, possesses a highly conserved pattern of four charged amino acid residues: one glutamate, two histidines and one aspartate. This pattern represents a new type of metalloprotease motif called the JAMM (Jab1/MPN domain metalloenzyme) or MPN+ motif [62, 63]. In CSN5 the catalytic region is important for the cleavage of the ubiquitinlike modifier NEDD8 from its targets. Mutations in the conserved histidine and asparxstate residues of CSN5 led to suppression of its deneddylation activity [60]. Crystal-structure analysis obtained with bacterial CSN5/MPN+ domain-containing AF2198 protein confirmed the metal-ion-dependent hydrolytic activity of CSN5, although it was inhibited by the alkylating agent NEM, an inhibitor of cysteine proteases [64]. NEDD8 is activated by a heterodimeric complex of APP-B1 and Ubα3 and is conjugated to target proteins by the conjugating enzyme Ubc12. So far, the only known targets are cullin-family proteins (CUL1–5), which are components of the cullin-based E3 ligase complexes. The covalent linkage of NEDD8 to cullins in vivo is thought to activate Ub-ligase complex activity by facilitating ubiquitin-conjugating enzyme E2 recruitment [65]. Deneddylation of cullins inactivates ubiquitination in vitro, but seems to stimulate the Ub E3 ligase complex activity in vivo [66, 67]. In cell lysates only a small fraction of CUL1 is neddylated, but in csn deletion cells 100% of CUL1 is modified by NEDD8. The purified CSN complex is able to deneddylate, although recombinant CSN5 protein cannot. Obviously CSN5 deneddylation activity is dependent on its assembly into the CSN complex [28, 54]. The fact that null mutants in most CSN subunits lack the deneddylation activity in the presence of excess CSN5 supports the fact that CSN5 alone is inactive in deneddylation [61]. So far, the exact role of deneddylation is questionable (see below).

4 Biochemical Activities Associated With the CSN 7

Fig. 2 Association of the CSN complex with enzymes. The Figure shows an electronmicroscopy image of purified CSN complex from human erythrocytes. As indicated by arrows the CSN is associated with the Ubspecific protease Ubp12, the proteasome, presumably with most of the cullin-based

Ub-ligase complexes, with a number of kinases, and with subunits of the translation initiation complex eIF3. In addition, subunit CSN5 has an intrinsic metalloprotease activity, which deneddylates cullins and also removes Ub conjugated to other proteins (for details see text).

4.2 Protein Kinases

The CSN is associated with enzymes such as kinases, proteases and Ub ligases, which perhaps, besides the intrinsic deneddylation activity, determine the specific function of the CSN in the Ub system. Here we summarize the associated (extrinsic) activities of the CSN shown in Figure 2. 4.2.1 Associated Protein Kinases Originally, a protein kinase was the first enzyme identified with the CSN purified from human erythrocytes. The CSN-associated kinase activity phosphorylated several serine and threonine residues in the N-terminal region of c-Jun [5] resulting in stabilization of c-Jun and increased AP-1 transcriptional activity. The pathway responsible for this c-Jun stabilization/activation was called CSN-directed c-Jun signaling [68]. It was subsequently shown that the CSN-directed c-Jun signaling pathway controls most of the VEGF (vascular endothelial growth factor) production in tumor cells [13]. VEGF is essential for tumor angiogenesis (see below).


In contrast to c-Jun, phosphorylation of the tumor suppressor p53 by CSNassociated kinases targets the protein for degradation by the Ub system [35]. For p53 stability, modification on Thr155 is most important as shown by mutational analysis [35] and by using different p53 peptides [31]. Mutation of Thr155 to Val led to stabilization of the transiently expressed p53 mutant in HeLa as well as in HL60 cells [35]. Inhibitors of CSN-associated kinases such as curcumin [18] caused stabilization of cellular p53 followed by massive cell death [35]. In addition to p53 and c-Jun, p27, ICSBP (interferon consensus sequence binding protein) and IκBα were identified as substrates of the CSN-associated kinases (for a review see Ref. [40]). Similar to p53, the phosphorylation of p27 results in accelerated degradation of the cyclin-dependent kinase inhibitor p27 by the Ub system (our unpublished data). In the case of ICSBP and IκBα, it is still unclear whether CSNmediated phosphorylation influences their stability. Interestingly, two of the CSN subunits, CSN2 and CSN7, are phosphorylated by the associated kinases [19, 69]. The physiological relevance of these modifications is currently obscure. 4.2.1.1 Identification of associated protein kinases Based on phosphopeptide analyses it became clear that associated kinases modify principally serine and threonine residues. Moreover, the analysis of putative phosphorylation-specific consensus sequences of p53, c-Jun, p27, ICSBP and IκBα revealed that the protein kinase CK2 and a member of the protein kinase C family might be associated with the CSN. It has been shown by immunoblotting that CK2 and the protein kinase Cµ (also called protein kinase D, PKD) co-purify with the CSN from human erythrocytes [31]. In addition, the two kinases co-immunoprecipitated together with the CSN from HeLa cells. Interaction of CK2 as well as PKD with the CSN is mediated by CSN3, as is the interaction between CK2 and the CSN7 subunit [31]. Interestingly, CSN7 itself is phosphorylated. Majerus and co-workers have published work on the co-purification of inositol 1,3,4-trisphosphate 5/6-kinase (5/6-kinase) with the CSN from bovine brain [24, 70]. Although the 5/6-kinase was not detected in the final preparation of the CSN from human erythrocytes [31], it cannot be excluded that the enzyme is associated with another pool of CSN particles. The enzyme phosphorylates c-Jun, IκBα as well as p53 and is sensitive to curcumin. These characteristics are very similar to those described for CK2 and PKD. It has been shown that the 5/6-kinase interacts with CSN1 and that over-expression of CSN1 inhibits its activity [24]. It might be that it interacts with the N-terminal part of CSN1, which has been shown to suppress activation of an AP-1 promoter [22]. Future studies will show whether additional kinases besides 5/6-kinase, CK2 and PKD can interact with the CSN under certain circumstances. For example, an interaction of CSN3 with IKKγ , a component of the IKK kinase complex, has been published [32]. 4.2.1.2 Functions of associated protein kinases Phosphorylation of a number of Ub-dependent substrates by CSN-associated kinases regulates the stability of the proteins towards the Ub system [40], presumably by promoting substrate ubiquitination. Most of the proteins bind to the CSN via CSN5, are phosphorylated and

4 Biochemical Activities Associated With the CSN 9

subsequently channeled to the associated Ub ligase for ubiquitination (see below). Modification of p53 induces a conformational change of the tumor suppressor, which leads to tighter binding to the Ub ligase [35]. In addition, there is evidence that phosphorylation might directly affect Ub-ligase activity. The transcriptional regulator Id3 interacts with the CSN, but is not phosphorylated. Nevertheless, inhibitors of CSN-associated kinases induce ubiquitination and degradation of the Id3 protein [41]. Because of associated kinases and their function in ubiquitination the CSN has been described as a complex “at the interface between signal transduction and ubiquitin-dependent proteolysis” [40]. This becomes even more significant if upstream regulation of the associated kinases is taken into account. Unfortunately at the moment little is known about the receptors or signal-transduction pathways leading to modification of the CSN and its associated kinase activities. It is also unclear whether there are interactions between the kinases and other associated activities of the CSN. 4.3 Deubiquitinating Enzymes

To date two deubiquitinating activities associated with the CSN have been identified. By mutational analysis one deubiquitinating activity has been mapped to the metalloprotease motif His–X–His–X10–Asp of the JAMM or MPN+ domain of CSN5 [11]. The conserved Asp residue of that motif was mutated and the mutant Flag-CSN5 was integrated into the CSN. The mutated CSN lost its ability to remove ubiquitin from the isopeptide bond of a mono-ubiquitinated conjugate [11]. Obviously the same intrinsic metalloprotease activity is responsible for deneddylation of mono-neddylated cullins [60], which is not surprising because of the homologies between Ub and NEDD8. It would be interesting to test whether the lid subunit Rpn11, which exhibits a deubiquitinating MPN+ domain [62], is able to deneddylate mono-neddylated proteins. In addition, another deubiquitinating activity associated with the CSN disassembles poly-Ub chains [11, 71]. This activity is catalyzed in fission yeast by the Ub-specific protease Ubp12, which is a CSN-associated enzyme [71]. The interaction of Ubp12 with the CSN is required for Ubp12 transport to the nucleus. Presumably in the nucleus, S. pombe Pcu1- and Pcu3-based Ub-ligase activities are inhibited by Ubp12 enzyme, since the deubiquitinating enzyme protects a specific adapter protein, Pop1p, from autocatalytic destruction [71]. Thus it seems that the CSN has dual activity in suppressing cullin-based Ub-ligase reactions: one is the intrinsic deneddylation and the other is deubiquitination via associated Ubp12; both reactions serve to inhibit cullin-based Ub ligases in vitro. Data have accumulated showing that CSN-associated deneddylation and deubiquitination are required for Ub-ligase activity in vivo. It has been hypothesized, therefore, that CSN-mediated inhibition of cullin-based ubiquitination might be necessary for the assembly of new cullin-based Ub-ligase complexes. After release from the CSN the new cullin-based complex would be active. It has to return to the


CSN for re-assembly or is degraded after auto-ubiquitination [71, 72]. For example, p27 has to be degraded at the transition from G1 to S phase. In a first step p27 may bind to the CSN which signals, perhaps by phosphorylation, the assembly of the required SCF complex containing the specific F-box protein Skp2. After formation of the p27-specific SCF complex both p27 and the Ub ligase might be released from the CSN perhaps again by phosphorylation which then results in ubiquitination and complete degradation of p27. Finally the Skp2-containing SCF complex is auto-ubiquitinated and degraded unless additional substrate appears. 4.4 Ubiquitin Ligases

Data have been accumulated demonstrating interactions of the CSN with Ub ligases, in particular with the cullin-based Ub ligases. Cullins 1 to 7 (CUL1–CUL7) form a protein family detected in all eukaryotic cells, which is involved in protein ubiquitination. It is known that CUL1 to CUL5 interact with the RING-domain protein Rbx1, the Ub ligase of the cullin-based complexes. So far it has been shown that the CSN interacts with CUL1 to CUL4 [11, 12, 54, 55, 61, 73]. Binding studies with CUL1 and with CUL2 revealed that the two cullin proteins bind via CSN2 to the complex [28, 54, 55]. In addition, Rbx1 seems to interact with CSN6 [54, 55]. Moreover, CUL1 interacts with Skp1, which makes the connection to a substrate-specific F-box protein. Therefore, CUL1-based Ub ligases are called SCF complexes (Skp1CDC53/CUL1-F-box protein) (for a review see Ref. [74]). CUL2 can be linked to the substrate-adapter protein the von Hippel–Lindau tumor suppressor via elongin C and elongin B forming the so called VCB (von Hippel–Lindau–elongin C–elongin B) complex (for a review see Ref. [74]). BTB/POZ-domain proteins have been identified as possible substrate-specific adaptors of CUL3-based Ub ligases [73, 75, 76]. There are more than 200 putative BTB/POZ-domain proteins expressed in mammalian cells and together with the large number of possible F-box proteins one can estimate that several hundreds of different cullin-based Ub-ligase complexes with different substrate specificities can be formed. The CUL4–Rbx1 complex has been characterized, and seems to be important for checkpoint control [12], DNA repair [11] and ubiquitination of c-Jun [77]. Most likely all cullin-based complexes interact with the CSN. In other words, the CSN is associated with ubiquitinating activity (see Figure 2). There are just a few data on interactions of the CSN with other Ub ligases besides the cullin-based complexes. For example, Mdm2, the RING domain Ub ligase of the tumor suppressor p53, binds to the CSN and is modified by CSN-associated kinases (our unpublished data). Whether Mdm2 is also modified by other CSN-associated activities has to be tested in the future. In addition, COP1, a putative RING-domain Ub ligase, which probably cooperates with the COP1-interacting protein 8 (CIP8) also binds to the CSN (for a review see Ref. [4]). However, some data indicate that COP1 is associated with a CUL4A complex in which it acts together with DET1 as a heterodimeric substrate adapter [77]. In this complex the CSN interacts with both the CUL4A and the COP1.

5 Association of the CSN With Other Protein Complexes 11

5 Association of the CSN With Other Protein Complexes 5.1 The eIF3 Complex

MPN and PCI domains have been also found in subunits of the eIF3 complex. Because MPN and PCI domains are most likely involved in protein–protein interactions (see above), it is not surprising that there are also cross-interactions between subunits of the CSN, the eIF3 and the lid. It has been reported that eIF3e/INT6 possessing a PCI domain interacts with CSN7 [33, 78]. Another eIF3 subunit eIF3c/p105 co-immunoprecipitated with eIF3e/INT6, eIF3b, CSN1 and CSN8 [78]. eIF3e/INT6 was used as bait in a two-hybrid screen that revealed possible interactions with the 26S proteasome ATPase Rpt4, CSN3 and CSN6 but also with CSN7 [33]. Interestingly, the subunit of the CSN-like complex in Saccharomyces cerevisiae Pci8/CSN11 [79] seems also to be a subunit of the budding yeast eIF3 complex and perhaps plays a similar role to eIF3e/INT6 in eukaryotic cells [80]. It has been speculated that these interactions allow the CSN to control translation. Interactions between eIF3e/INT6 or eIF3i with the 26S proteasome have also been described [33, 81]. It has been shown that eIF3e/INT6 interacts with Rpn5 of the lid complex. This has an impact on 26S proteasome activity/localization, presumably affecting cell division and mitotic fidelity [82]. Perhaps there exists a network of “PCI complexes” as suggested [83], which shares polypeptides and communicates via proteins such as eIF3e/INT6. 5.2 The Proteasome

In 1998 it was reported that the CSN co-fractionates with the 26S proteasome from human cells [5]. A yeast two-hybrid screen revealed that the C-terminal domain of the Arabidopsis atCSN1 subunit interacts with atRpn6 of the 26S proteasome lid [26]. Recently gel-filtration size-fractionation of material from Arabidopsis in the presence of ATP and phosphatase inhibitors indicated that the CSN1 and CSN6 subunits co-elute in the same fractions as subunits of the 26S regulatory complex [84]. Based on these data it has been speculated that the CSN might be an alternative lid of the 26S proteasome [85]. The “alternative lid hypothesis”, however, makes little sense if the CSN interacts with the 26S proteasome via the lid component Rpn6 [26]. CSN pull-down experiments and subsequent mass-spectrometry analysis of co-precipitated proteins also revealed the presence of proteasome subunits in the precipitate [73]. However, since proteasome subunits are very abundant in cells, one has to be cautious with this type of data. So far there is no systematic binding study showing physical interaction of the CSN with sub-complexes of the 26S proteasome. Moreover, up to now there is no functional evidence for such a CSN/26S proteasome interaction.


6 Biological Functions of the CSN 6.1 Regulation of Ubiquitin Conjugate Formation

In general, and including all its activities, intrinsic as well as associated, the CSN seems to be a regulator of ubiquitination. Deneddylation, deubiquitination as well as CSN-mediated phosphorylation (at least with c-Jun and Id3 as substrates) cause inhibition of ubiquitination. It is likely that suppression of ligase activity is an essential step in the dynamic process of specific E3 complex assembly/reassembly. According to the model of Wolf et al. [72] cullin-based Ub-ligase complexes might assemble/reassemble in a protected environment produced by the CSN. In the CSN-associated-state, binding of any E2 to the Ub ligase is prevented, perhaps by deneddylation [65], self-ubiquitination is blocked by continuous deubiquitination [71] and substrate binding could be inhibited by phosphorylation [31]. Only under these conditions can the Ub ligase reassemble without itself being destroyed. For example, an SCF complex might associate with another F-box protein, or a CUL3Ub ligase with another BTB/POZ-domain protein, as an adaptation to the next phase of cell cycle or signal transduction upon the appearance of a new substrate, which has to be degraded. Following this argument a major question arises. How does the substrate signal the assembly of the required Ub ligase performing its ubiquitination? Is it by binding to the CSN and subsequent signaling via specific kinases? In the case of the SCF complexes, another protein called CAND1/Tip120A seems to be involved in the dynamic assembly/reassembly process of the E3 [86]. CAND1 binds to the deneddylated CUL1 and inhibits Ub-ligase activity by competing for the Skp1–F-box-protein unit of the SCF complex [87]. After the release of CAND1, a new Skp1–F-box-protein unit can dock to the CUL1-Rbx1 unit to form an SCF complex possessing the necessary substrate specificity. Now the freshly formed Ub ligase has to be released from the CSN to become active. At the moment it is unclear how the Ub ligase might be released from the CSN. The attractive model of CSN-assisted Ub-ligase-complex assembly has to be tested in the future. In this model the CSN would function as a platform for Ub-ligase assembly. Interestingly, there are no reports of interactions between the 26S proteasome lid complex and Ub ligases. Known E3s directly interacting with the 26S proteasome seem to bind via base ATPases [88, 89]. This is an interesting functional difference between the CSN and the lid developed during evolution. In an alternative model the CSN might be the platform for complete proteolysis. It forms supercomplexes consisting of both the ubiquitinating and the proteolytic machineries. According to this model, the substrate first binds to the CSN, is then ubiquitinated by the associated Ub ligase and finally directly channeled into the 26S proteasome. Deneddylation, deubiquitination and phosphorylation are necessary to maintain the supercomplex, to protect the intermediates and to stimulate proteolysis.

6 Biological Functions of the CSN 13

6.1.1 Cell-cycle and Checkpoint Control Initial insight of the role of CSN in cell-cycle control came from the finding that csn1 and csn2 deletion S. pombe strains have an S-phase delay [52]. Interestingly, this effect did not occur in strains missing other CSN subunits. The S-phase delay was caused by the accumulation of the cell-cycle inhibitor Spd1 (S-phase delayed 1), which is involved in the misregulation of the ribonucleotide reductase (RNR). RNR catalyzes the production of deoxyribonucleotides for DNA synthesis and is composed of four subunits including Suc22. Activation of RNR is regulated by nuclear export of Suc22, which is suppressed by Spd1 [12]. Upon DNA damage or during S phase Spd1 is rapidly degraded, presumably leading to the RNRdependent production of dNTPs. However, in csn1 and csn2 deletion mutants, Spd1 accumulates, causing Suc22-dependent suppression of RNR connected with the S-phase delay and DNA-damage sensitivity [12, 15]. In mammalian cells, binding of HIV-1 Vpr-protein to the CSN6 results in cellcycle arrest at the G2/M phase [53]. Additionally, CSN is involved in the cell cycle via the nuclear export of cell-cycle kinase inhibitor p27kip1 (p27). CSN5 binds to p27 and promotes its nuclear export followed by its proteasome-dependent degradation. The over-expression of CSN5 in mouse fibroblasts counteracts cell-cycle arrest induced by serum depletion [36, 51]. Microinjection of the purified CSN complex into synchronized G1 cells blocks the S-phase entry in a deneddylation-dependent manner [28]. Furthermore, the reduction of CSN subunit expression by RNAi in Caenorhabditis elegans causes the failure of Mei-1 degradation by regulation of its specific Ub-ligase CUL3-based complex, which leads to severe effects during mitotic cell division [76]. Moreover, the CSN is involved in checkpoint control. The double deletions of csn1 and csn2 mutants crossed with checkpoint pathway mutants such as rad3, chk, and cds1 are synthetically lethal in S. pombe [52]. Cds1 kinase is constitutively activated in csn1 mutants. Similarly, loss of csn5 in Drosophila results in activation of Mei-41, one of the ATM/ATR family kinases involved in meiotic checkpoint upon DNA damage [90]. 6.1.2 DNA Repair Two papers have assigned the CSN a function in DNA repair. One study reports on the existence of two different complexes containing human CSN and either one of the two nucleotide-excision-repair proteins, DDB2 or CSA. DDB2 is involved in the global genome-repair pathway (GGR) and CSA functions in the transcriptioncoupled repair pathway (TCR). Additionally, these complexes possess Ub-ligase activity and contain cullin-based Ub-ligase components such as CUL4 and Rbx1/Roc1, and DDB1, a UV-damage DNA-binding protein [11]. However, so far their targets remain unclear. CSN differentially regulates the ubiquitin-ligase activity of the DDB2- and CSA-containing complexes in response to UV irradiation. In support of direct involvement of the CSN is the finding that knockdown of CSN5 with RNAi causes a failure in NER mechanisms [11]. Similarly, CSN in combination with the CUL4–Rbx1 complex is involved in Ub-dependent degradation of CDT1, a licensing factor of the pre-replication complex (preRC), after UV- or γ -irradiation.


Knockdown of CSN completely suppresses CDT1 degradation, causing a defect G1 checkpoint in response to DNA damage [91]. 6.1.3 Developmental Processes Although CSN is not essential in yeast, the csn1 and csn2 S. pombe deletion mutants display slow growth and sensitivity to UV- and γ -irradiation [52]. Other csn mutants did not show significant phenotypes apart from the loss of cullin’s deneddylation activity [61]. In mutants of CSN-like complexes in the budding yeast S. cerevisiae the sensitivity to the DNA-damage reagents is not affected [92]. In some S. cerevisiae mutants such as csn5, csn9 and csn12 deletions, increased mating efficiency and enhanced pheromone response has been observed [63]. In Drosophila, mutations of CSN causes lethality in early larval stages and defects during oogenesis or photoreceptor R cell differentiation [9, 90, 93, 94]. More specifically, lack of CSN5 leads to the activation of a DNA double-strand-break-dependent checkpoint mediated by Mei-41. This effect is caused by CSN5-dependent inhibition of gurken (Grk) protein translation [90]. In C. elegans, knockdown of CSN5 by RNAi resulted in a sterile phenotype, which could be explained by CSN interaction with germ-line RNA helicases [48]. The best studied physiological role of the CSN in developmental processes is derived from studies on Arabidopsis. Csn mutants can survive embryogenesis, but they die soon after germination. The csn mutants exhibit a defect in photomorphogenesis, a light-dependent developmental process of germinating seedlings. Even in total darkness the mutants display a light-dependent morphology and signalindependent expression of light-induced genes [3, 14, 95]. One key mechanism is the CSN-dependent regulation of the stability of the transcription factor HY5, a positive regulator of light-induced genes. In the dark it is degraded by the Ub system [8]. It has been suggested that in darkness the RING-finger protein COP1 ubiquitinates HY5 and triggers its degradation by the 26S proteasome (for a review see Ref. [4]). In the light, COP1 is relocated to the cytoplasm allowing expression of genes through HY5. Although the exact mechanism remains unclear, the CSN may be required for relocation of COP1 from cytoplasm to the nucleus in darkness. Identical phenotypes caused by different csn mutants in Arabidopsis could be explained by a role of the CSN as a whole complex (for a review see Ref. [20]). There is accumulating evidence for cooperation of the CSN and cullin-based complexes in specific developmental processes [96]. First insight has been provided by studies on auxin response where the CSN interacts with SCFTIR1 , modulating its activity [55]. Similarly, binding of the CSN to other cullin-based complexes regulates their activity in mediating various developmental processes such as flower development, and plant defense responses [97, 98]. 6.2 Tumor Angiogenesis

Tumor angiogenesis is the vascularization of solid tumors, an essential requirement for tumor growth and metastasis. After a solid tumor has reached a size

6 Biological Functions of the CSN 15

of approximately 2 mm3 , it needs nutrient supply from blood vessels, otherwise it dies from necrosis. Many tumor cells are able to induce angiogenesis. In an initiation phase the tumor cells produce large amounts of pro-angiogenic factors such as vascular endothelial growth factor, VEGF. During proliferation and invasion VEGF stimulates migration of endothelial cells. Finally, after a maturation phase, vascularization of solid tumors is completed. Now the tumor can grow, and some tumor cells penetrate through vessel membranes and spread via the circulation. Therefore, inhibition of tumor angiogenesis has become an important strategy in tumor therapy. There is functional cooperation between the CSN and the Ub system in tumor angiogenesis [13]. It has been known for some time that curcumin is an inhibitor of angiogenesis [99]. However, only in 2001 did it become clear that it acts via inhibition of CSN-associated kinases [13]. It has been demonstrated that over-expression of CSN2 subunit leads to elevated amounts of de novo assembled CSN complex connected with increased c-Jun levels and enhanced AP-1 transactivation activity [68]. This c-Jun activation/stabilization is independent of the JNK and the MAP kinase pathway and is called CSN-directed c-Jun signaling [68]. This process can be inhibited by curcumin or other inhibitors of CSN-associated kinases (Figure

Fig. 3 The CSN-directed c-Jun signaling pathway. (A) The active CSN-directed cJun signaling pathway is shown. In case of active CSN-associated kinases c-Jun is phosphorylated, which stabilizes the transcription factor towards the Ub system. In addition, phosphorylation of the responsible E3 might inactivate the enzyme. In this situation Id1 and Id3 are also stabilized. Stable/active c-Jun causes enhanced AP-1 transactivation connected with an increase of VEGF production by tumor cells (see

text). VEGF is a major pro-angiogenic factor produced by many tumor cells. Id1 and Id3 are transcriptional regulators essential for tumor angiogenesis. (B) In the presence of curcumin or other kinase inhibitors the responsible Ub ligase is most likely active and ubiquitinates both c-Jun and Id3. In addition, unphosphorylated c-Jun might have higher affinity to its Ub ligase. This leads to quick degradation of the proteins by the Ub system.


3). The CSN-directed c-Jun signaling controls up to 75% of VEGF production in tumor cells [13]. In addition, Id1 and Id3 are also essential factors of tumor angiogenesis [100] and are degraded in the presence of CSN-associated kinase inhibitors in an Ub-dependent manner just like c-Jun [41]. Therefore, specific inhibition of CSN-associated kinases might become important for tumor therapy. The application of CSN-associated kinase inhibitors in tumor therapy could be beneficial owing to another effect of curcumin-like compounds, namely they stabilize cellular p53 and, at least in tumors with wild-type p53 protein, massive cell death can be observed [35].

7 Conclusions

The CSN is a regulatory complex of the Ub system. Physically it interacts with the proteasome and with Ub ligases. Although the exact mechanism remains obscure, the CSN regulates ubiquitination of important cell-cycle factors and transcriptional regulators. Its intrinsic deneddylating as well as the associated kinase and deubiquitinating activities seem to be required for determining protein stability towards the Ub system. As a major regulator of the Ub system the CSN is involved in processes such as DNA repair, cell-cycle progression and development. Its role in tumor angiogenesis makes the complex attractive for future tumor therapies.

Acknowledgment

We thank Rasmus Hartmann-Petersen for critical reading of the manuscript. The work was supported by research grants from the Deutsche Forschungsgemeinschaft and from the German-Israeli Foundation to W. D.

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proteolysis. Nat Cell Biol. 2002, 4, 1003–1007. CORN, P. G., MCDONALD, E. R., 3rd, HERMAN, J. G., EL-DEIRY, W. S. Tat-binding protein-1, a component of the 26S proteasome, contributes to the E3 ubiquitin ligase function of the von Hippel-Lindau protein. Nat. Genet. 2003, 35, 229–237. DORONKIN, S., DJAGAEVA, I., BECKENDORF, S. K. CSN5/Jab1 mutations affect axis formation in the Drosophila oocyte by activating a meiotic checkpoint. Development 2002, 129, 5053–5064. HIGA, L. A., MIHAYLOV, I. S., BANKS, D. P., ZHENG, J., ZHANG, H. Radiation-mediated proteolysis of CDT1 by CUL4-ROC1 and CSN complexes constitutes a new checkpoint. Nat Cell Biol. 2003, 5, 1008–1015. WEE, S., HETFELD, B., DUBIEL, W., WOLF, D. A. Conservation of the COP9/signalosome in budding yeast. BMC Genet. 2002, 3, 15. ORON, E., MANNERVIK, M., RENCUS, S., HARARI-STEINBERG, O., NEUMAN-SILBERBERG, S., SEGAL, D., CHAMOVITZ, D. A. COP9 signalosome subunits 4 and 5 regulate multiple pleiotropic pathways in Drosophila melanogaster. Development 2002, 129, 4399–4409. SUH, G. S., POECK, B., CHOUARD, T., ORON, E., SEGAL, D., CHAMOVITZ, D. A., ZIPURSKY, S. L. Drosophila JAB1/CSN5

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acts in photoreceptor cells to induce glial cells. Neuron 2002, 33, 35–46. MA, L., ZHAO, H., DENG, X. W. Analysis of the mutational effects of the COP/DET/FUS loci on genome expression profiles reveals their overlapping yet not identical roles in regulating Arabidopsis seedling development. Development 2003, 130, 969–981. SCHWECHHEIMER, C., SERINO, G., DENG, X. W. Multiple ubiquitin ligase-mediated processes require COP9 signalosome and AXR1 function. Plant Cell 2002, 14, 2553–2563. FENG, S., MA, L., WANG, X., XIE, D., DINESH-KUMAR, S. P., WEI, N., DENG, X. W. The COP9 signalosome interacts physically with SCF COI1 and modulates jasmonate responses. Plant Cell 2003, 15, 1083–1094. WANG, X., FENG, S., NAKAYAMA, N., CROSBY, W. L., IRISH, V., DENG, X. W., WEI, N. The COP9 signalosome interacts with SCF UFO and participates in Arabidopsis flower development. Plant Cell 2003, 15, 1071–1082. ARBISER, J. L., KLAUBER, N., ROHAN, R., van LEEUWEN, R., HUANG, M. T., FISHER, C., FLYNN, E., BYERS, H. R. Curcumin is an in vivo inhibitor of angiogenesis. Mol. Med. 1998, 4, 376–383. BENEZRA, R., RAFII, S., LYDEN, D. The Id proteins and angiogenesis. Oncogene 2001, 20, 8334–8341.

1

Molecular Chaperones and the Ubiquitin–Proteasome System Cam Patterson

University of North Carolina, Chapel Hill, USA

Jörg Höhfeld Rheinische Friedrich-Wilhelms-Universität Bonn, Bonn, Germany


1 Introduction

The biological activity of a protein is defined by its unique three-dimensional structure. Attaining this structure, however, is a delicate process. A recent study suggests that up to 30% of all newly synthesized proteins never reach their native state [1]. As protein misfolding poses a major threat to cell function and viability, molecular mechanisms must have evolved to prevent the accumulation of misfolded proteins and thus aggregate formation. Two protective strategies appear to be followed. Molecular chaperones are employed to stabilize nonnative protein conformations and to promote folding to the native state whenever possible. Alternatively, misfolded proteins are removed by degradation, involving, for example, the ubiquitin–proteasome system. For a long time molecular chaperones and cellular degradation systems were therefore viewed as opposing forces. However, recent evidence suggests that certain chaperones (in particular members of the 70- and 90-kDa heat shock protein families) are able to cooperate with the ubiquitin–proteasome system. Protein fate thus appears to be determined by a tight interplay of cellular protein-folding and protein-degradation systems.

2 A Biomedical Perspective

The aggregation and accumulation of misfolded proteins is now recognized as a common characteristic of a number of degenerative disorders, many Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Molecular Chaperones and the Ubiquitin–Proteasome System

of which have neurological manifestations [2, 3]. These diseases include prionopathies, Alzheimer’s and Parkinson’s diseases, and polyglutamine expansion diseases such as Huntington’s disease and spinocerebellar ataxia. At the cellular level, these diseases are characterized by the accumulation of aberrant proteins either intracellularly or extracellularly in specific groups of cells that subsequently undergo death. The precise association between protein accumulation and cell death remains incompletely understood and may vary from disease to disease. In some cases, misfolded protein accumulations may themselves be toxic or exert spatial constraints on cells that affect their ability to function normally. In other cases, the sequestering of proteins in aggregates may itself be a protective mechanism, and it is the overwhelming of pathways that consolidate aberrant proteins that is the toxic event. In either case, lessons learned from genetically determined neurodegenerative diseases have helped us to understand the inciting events of protein aggregation that ultimately lead to degenerative diseases. Mutations resulting in neurodegenerative diseases fall into two broad classes. The first class comprises mutations that affect proteins, irrespective of their native function, and cause them to misfold. The classic example of this is Huntington’s disease [4, 5]. The protein encoded by the huntingtin gene contains a stretch of glutamine residues (or polyglutamine repeat), and the genomic DNA sequence that codes for this polyglutamine repeat is subject to misreading and expansion. When the length of the polyglutamine repeat in huntingtin reaches a critical threshold of approximately 35 residues, the protein becomes prone to misfolding and aggregation [6]. This appears to be the proximate cause of neurotoxicity in this invariably fatal disease [7, 8]. A number of other neurodegenerative diseases are caused by polyglutamine expansions [9, 10]. For example, spinocerebellar ataxia is caused by polyglutamine expansions in the protein ataxin-1 [11]. In other diseases, protein misfolding occurs due to other mutations that induce misfolding and aggregation; for example, mutations in superoxide dismutase-1 lead to aggregation and neurotoxicity in amyotrophic lateral sclerosis [12, 13]. Other mutations that result in neurodegenerative diseases are instructive in that they directly implicate the ubiquitin–proteasome system in the pathogenesis of these diseases [14]. For example, mutations in the gene encoding the protein parkin are associated with juvenile-onset Parkinson’s disease [15, 16]. Parkin is a RING finger–containing ubiquitin ligase, and mutations in this ubiquitin ligase cause accumulation of target proteins that ultimately result in the neurotoxicity and motor dysfunction associated with Parkinson’s disease [17–20]. Repressor screens of neurodegeneration phenotypes in animal models have also linked the molecular chaperone machinery to neurodegeneration [21–24]. Taken together, the pathophysiology of neurodegenerative diseases provides a compelling demonstration of the importance of the regulated metabolism of misfolded proteins and provides direct evidence of the role of both molecular chaperones and the ubiquitin–proteasome system in guarding against protein misfolding and its consequent toxicity.

3 Molecular Chaperones: Mode of Action and Cellular Functions 3

3 Molecular Chaperones: Mode of Action and Cellular Functions

Molecular chaperones are defined by their ability to bind and stabilize nonnative conformations of other proteins [25, 26]. Although they are an amazingly diverse group of conserved and ubiquitous proteins, they are also among the most abundant intracellular proteins. The classical function of chaperones is to facilitate protein folding, inhibit misfolding, and prevent aggregation. These folding events are regulated by interactions between chaperones and ancillary proteins, the co-chaperones, which in general assist in cycling unfolded substrate proteins on and off the active chaperone complex [25, 27, 28]. In agreement with their essential function under normal growth conditions, chaperones are ubiquitously expressed and are found in all cellular compartments of the eukaryotic cell (except for peroxisomes). In addition, cells greatly increase chaperone concentration as a response to diverse stresses, when proteins become unfolded and require protection and stabilization [29]. Accordingly, many chaperones are heat shock proteins (Hsps). Four main families of cytoplasmic chaperones can be distinguished: the Hsp70 family, the Hsp90 family, the small heat shock proteins, and the chaperonins. 3.1 The Hsp70 Family

The Hsp70 proteins bind to misfolded proteins promiscuously during translation or after stress-mediated protein damage [26, 30]. Members of this family are highly conserved throughout evolution and are found throughout the prokaryotic and eukaryotic phylogeny. It is common for a single cell to contain multiple homologues, even within a single cellular compartment; for example, mammalian cells express two inducible homologues (Hsp70.1 and Hsp70.3) and a constitutive homologue (Hsc70) in the cytoplasm. These homologues have overlapping but not totally redundant cellular functions. Members of this family are typically in the range of 70 kDa in size and contain three functional domains: an amino-terminal ATPase domain, a central peptide-binding cleft, and a carboxyl terminus that seems to form a lid over the peptide-binding cleft [28] (Figure 1). The chaperones recognize short segments of the protein substrate, which are composed of clusters of hydrophobic amino acids flanked by basic residues [31]. Such binding motifs occur frequently within protein sequences and are found exposed on nonnative proteins. In fact, mammalian Hsp70 binds to a wide range of nascent and newly synthesized proteins, comprising about 15–20% of total protein [32]. This percentage is most likely further increased under stress conditions. Hsp70 proteins apparently prevent protein aggregation and promote proper folding by shielding hydrophobic segments of the protein substrate. The hydrophobic segments are recognized by the central peptide-binding domain of Hsp70 proteins (Figure 1). The domain is composed of two sheets of β strands that together with connecting loops form a cleft to accommodate extended peptides of about seven amino acids in length, as revealed


Fig. 1 Schematic presentation of the domain architecture and chaperone cycle of Hsp70. Hsp70 proteins display a characteristic domain structure comprising an amino-terminal ATPase domain (ATP), a peptide-binding domain (P), and a carboxylterminal domain (C) that is supposed to form a lid over the peptide-binding domain. In the ATP-bound conformation, the binding

pocket is open, resulting in a low affinity for the binding of a chaperone substrate. ATP hydrolysis induces stable substrate binding through a closure of the peptide-binding pocket. Substrate release is induced upon nucleotide exchange. ATP hydrolysis and nucleotide exchange are regulated by diverse co-chaperones.

in crystallographic studies of bacterial Hsp70 [33]. In the obtained crystal structure, the adjacent carboxyl-terminal domain of Hsp70 folds back over the β sandwich, suggesting that the domain may function as a lid in permitting entry and release of protein substrates (Figure 1). According to this model, ATP binding and hydrolysis by the amino-terminal ATPase domain of Hsp70 induce conformational changes of the carboxyl terminus, which lead to lid opening and closure [28]. In the ATP-bound conformation of Hsp70, the peptide-binding pocket is open, resulting in rapid binding and release of the substrate and consequently in a low binding affinity (Figure 1). Stable holding of the protein substrate requires closing of the binding pocket, which is induced upon ATP hydrolysis and conversion of Hsp70 to the ADP-bound conformation. The dynamic association of Hsp70 with nonnative polypeptide substrates thus depends on ongoing cycles of ATP binding, hydrolysis, and nucleotide exchange. Importantly, ancillary co-chaperones are employed to regulate the ATPase cycle [27, 30]. Co-chaperones of the Hsp40 family (also termed J proteins due to their founding member bacterial DnaJ) stimulate the ATP hydrolysis step within the Hsp70 reaction cycle and in this way promote substrate binding [34] (Figure 1). In contrast, the carboxyl terminus of Hsp70-interacting protein CHIP attenuates ATP hydrolysis [35]. Similarly, nucleotide exchange on Hsp70 is under the control of stimulating and inhibiting co-chaperones. The Hsp70-interacting protein Hip slows down nucleotide exchange by stabilizing the ADP-bound conformation of the chaperone [36], whereas nucleotide exchange is stimulated by the co-chaperone BAG-1 (Bcl-2-associated athanogene 1), which assists substrate unloading from Hsp70 [37–39]. By altering the AT-Pase cycle, the co-chaperones directly modulate the folding activity of Hsp70. In addition to chaperone-recognition motifs, co-chaperones often possess other functional


Fig. 2 Domain architecture of diverse cochaperones of Hsp70. DnaJ: domain related to the bacterial co-chaperone DnaJ; TPR: tetratricopeptide repeat; Sti1: domain related to the yeast co-chaperone Sti1; CC: coiled-coil domain; U box: E2-interacting domain present in certain ubiquitin ligases;

PG: polyglycine region; ARM: armadillo repeat; TRSEEX: repeat motif found at the amino terminus of BAG-1 isoforms; ubl: ubiquitin-like domain; BAG: Hsp70-binding domain present in BAG proteins; WW: protein interaction domain.

domains and therefore link chaperone activity to distinct cellular processes [27, 40] (Figure 2). Indeed, as discussed below, the co-chaperones BAG-1 and CHIP apparently modulate Hsp70 function during protein degradation. 3.2 The Hsp90 Family

The 90-kDa cytoplasmic chaperones are members of the Hsp90 family, and in mammals two isoforms exist: Hsp90α and Hsp90β. The Hsp70 and Hsp90 families exhibit several common features: both possess ATPase activity and are regulated by ATP binding and hydrolysis, and both are further regulated by ancillary co-chaperones [41–48]. Unlike Hsp70, however, cytoplasmic Hsp90 is not generally involved in the folding of newly synthesized polypeptide chains. Instead it plays a key role in the regulation of signal transduction networks, as most of the known substrates of Hsp90 are signaling proteins, the classical examples being steroid hormone receptors and signaling kinases. On a molecular level, Hsp90 binds


Fig. 3 Cooperation of Hsp70 and Hsp90 during the regulation of signal transduction pathways. The inactive signaling protein, e.g., a steroid hormone receptor, is initially recognized by Hsp40 and delivered to Hsp70. Subsequently, a multi-chaperone complex assembles that contains the Hsp70 co-chaperone Hip and the Hsp70/Hsp90organizing protein Hop. Hop stimulates recruitment of an Hsp90 dimer that accepts the substrate from Hsp70. At the final stage

of the chaperone pathway, Hsp90 associates with p23 and diverse cyclophilins (cycloph.) to mediate conformational changes of the signaling protein necessary to reach an activatable state. Upon activation, i.e., hormone binding in the case of the steroid receptor, the signaling protein is released from Hsp90. In the absence of an activating stimulus, the signaling protein folds back to the inactive state when released and enters a new cycle of chaperone binding.

to substrates at a late stage of the folding pathway, when the substrate is poised for activation by ligand binding or associations with other factors. Consequently, Hsp90 accepts partially folded conformations from Hsp70 for further processing. In the case of the chaperone-assisted activation of the glucocorticoid hormone receptor and also of the progesterone receptor, the sequence of events leading to attaining an active conformation is fairly well understood [49–53]. It appears that the receptors are initially recognized by Hsp40 and are then delivered to Hsp70 [54] (Figure 3). Subsequent transfer onto Hsp90 requires the Hsp70/Hsp90-organizing protein Hop, which possesses non-overlapping binding sites for Hsp70 and Hsp90 and therefore acts as a coupling factor between the two chaperones [55]. In conjunction with p23 and different cyclophilins, Hsp90 eventually mediates conformational changes that enable the receptor to reach a high-affinity state for ligand binding. On other signaling pathways Hsp90 serves as a scaffolding factor to permit interactions between kinases and their substrates, as is the case for Akt kinase and endothelial nitric oxide synthase [56]. Since many of the Hsp90 substrate proteins are involved in regulating cell proliferation and cell death, it is not surprising that the chaperone recently emerged as a drug target in tumor therapy [57–59]. The antibiotics


Fig. 4 Alteration of chaperone action during signal transduction induced by Hsp90 inhibitors such as geldanamycin and radicicol. In the presence of the inhibitors the activation pathway is blocked, and signaling

proteins are targeted to the proteasome for degradation in a process that involves the co-chaperone CHIP and other E3 ubiquitin ligases that remain to be identified.

geldanamycin and radicicol specifically bind to Hsp90 in mammalian cells and inhibit the function of the chaperone by occupying its ATP-binding pocket [60–63]. Drugs based on these compounds are now being developed as anticancer agents, as they potentially inactivate multiple signaling pathways that drive carcinogenesis. Remarkably, drug-induced inhibition of Hsp90 blocks the chaperone-assisted activation of signaling proteins and leads to their rapid degradation via the ubiquitin–proteasome pathway [64–69] (Figure 4). Hsp90 inhibitors therefore have emerged as helpful tools to study chaperone- proteasome cooperation. 3.3 The Small Heat Shock Proteins

The precise functions of small heat shock proteins (sHsps) including Hsp27 and the eye-lens protein αB-crystallin are incompletely understood. However, they seem to play a major role in preventing protein aggregation under conditions of cellular stress [70–73]. All members investigated so far form large oligomeric complexes of spherical or cylindrical appearance [74, 75]. Complex formation is independent of ATP binding and hydrolysis, but appears to be regulated by temperature and phosphorylation. The structural analysis of wheat Hsp16.9 suggested that the oligomeric complex acts as a storage form rather than an enclosure for substrates, as the active chaperone appears to be a dimer [75]. In agreement with this notion, dissociation of


the oligomeric complex formed by yeast Hsp26 was found to be a prerequisite for efficient chaperone activity [76]. Subsequent refolding may occur spontaneously or may involve cooperation with other chaperones such as Hsp70 [77]. 3.4 Chaperonins

The chaperone proteins best understood with regard to their mode of action are certainly the so-called chaperonins, which are defined by a barrel-shaped, doublering structure [25, 28]. Members include bacterial GroEL, Hsp60 of mitochondria and chloroplasts, and the TriC–CCT complex localized in the eukaryotic cytoplasm. Based on their characteristic ring structure, a central cavity is formed, which accommodates nonnative proteins via hydrophobic interactions. Conformational changes of the chaperonin subunits induced through ATP hydrolysis change the inner lining of the cavity from a hydrophobic to a hydrophilic character [78–80]. As a consequence the unfolded polypeptide is released into the central chamber and can proceed on its folding pathway in a protected environment [81]. The chaperonins are therefore capable of folding proteins such as actin that cannot be properly folded via other mechanisms [82].

4 Chaperones: Central Players During Protein Quality Control

Due to their ability to recognize nonnative conformations of other proteins, molecular chaperones are of central importance during protein quality control. This was elegantly revealed in studies on the influence of the Hsp70 chaperone system on polyglutamine diseases using the fruit fly Drosophila melanogaster as a model organism (reviewed in Refs. [23] and [83]). Hallmarks of the polyglutamine disease spinocerebellar ataxia type 3 (SCA3), for example, were recapitulated in transgenic flies that expressed a pathological polyQ tract of the ataxin-3 protein in the eye disc [84]. Transgene expression caused formation of abnormal protein inclusions and progressive neuronal degeneration. Intriguingly, co-expression of human cytoplasmic Hsp70 suppressed polyQ-induced neurotoxicity. In a similar experimental approach, Hsp40 family members protected neuronal cells against toxic polyQ expression [22]. Enhancing the activity of the Hsp70/Hsp40 chaperone system apparently mitigates cytotoxicity caused by the accumulation of aggregationprone proteins. These findings obtained in Drosophila were confirmed in a mouse model of spinocerebellar ataxia type 1 (SCA1) [85, 86]. Unexpectedly, however, the Hsp70 chaperone system was unable to prevent the formation of protein aggregates in these models of polyglutamine diseases and upon polyQ expression in yeast and mammalian cells [84, 85, 87–89]. Elevating the cellular levels of Hsp70 and of some Hsp40 family members affected the number of protein aggregates and their biochemical properties, but did not inhibit the formation of polyQ aggregates. Notably, Hsp70 and Hsp40 profoundly modulated the aggregation process of

5 Chaperones and Protein Degradation

polyQ tracts in biochemical experiments; this led to the formation of amorphous, SDS-soluble aggregates, instead of the ordered, SDS-insoluble amyloid fibrils that form in the absence of the chaperone system [88]. These biochemical data were confirmed in yeast and mammalian cells [88, 90]. Although unable to prevent the formation of protein aggregates, the Hsp70 chaperone system apparently prevents the ordered oligomerization and fibril growth that is characteristic of the disease process. In an alternate but not mutually exclusive model to explain their protective role, the chaperones may cover potentially dangerous surfaces exposed by polyQcontaining proteins during the oligomerization process or by the final oligomers. Intriguingly, elevated expression of Hsp70 also suppresses the toxicity of the nonpolyQ-containing protein α-synuclein in a Drosophila model of Parkinson’s disease without inhibiting aggregate formation [24]. Hsp70 may thus exert a rather general function in protecting cells against toxic protein aggregation. This raises the exciting possibility that treatment of diverse forms of human neurodegenerative diseases may be achieved through upregulation of Hsp70 activity. The mentioned examples illustrate that one does not have to evoke the refolding of an aberrant protein to the native state in order to explain the protective activity of Hsp70 observed in models of amyloid diseases. In some cases it might be sufficient for Hsp70 to modulate the aggregation process or to shield interaction surfaces of the misfolded protein to decrease cytotoxic effects. Another option may involve presentation of the misfolded protein to the ubiquitin–proteasome system for degradation.


Hsp70 and Hsp90 family members as well as small heat shock proteins have all been implicated to participate in protein degradation. For example, the small heat shock protein Hsp27 was recently shown to stimulate the degradation of phosphorylated IκBα via the ubiquitin–proteasome pathway, which may account for the antiapoptotic function of Hsp27 [91]. Similarly, Hsp27 facilitates the proteasomal degradation of phosphorylated tau, a microtubule-binding protein and component of protein deposits in Alzheimer’s disease [92]. Hsp70 participates in the degradation of apolipoprotein B100 (apoB), which is essential for the assembly and secretion of very low-density lipoproteins from the liver [93]. Under conditions of limited availability of core lipids, apoB translocation across the ER membrane is attenuated, resulting in the exposure of some domains of the protein into the cytoplasm and their recognition by Hsp70. This is followed by the degradation of apoB via the ubiquitin–proteasome pathway. Elevating cellular Hsp70 levels stimulated the degradation of the membrane protein, suggesting that the chaperone facilitates sorting to the proteasome. Genetic studies in yeast indicate that cytoplasmic Hsp70 may fulfill a rather general role in the degradation of ER-membrane proteins that display large domains into the cytoplasm [94]. In agreement with this notion, Hsp70 also takes part in the degradation of immaturely glycosylated and aberrantly

9


folded forms of the cystic fibrosis transmembrane conductance regulator (CFTR) [95–98]. CFTR is an ion channel localized at the apical surface of epithelial cells. Its functional absence causes cystic fibrosis, the most common fatal genetic disease in Caucasians [99, 100]. The disease-causing allele, F508, which is expressed in more than 70% of all patients, drastically interferes with the protein’s ability to fold, essentially barring it from functional expression in the plasma membrane. However, wild-type CFTR also folds very inefficiently, and less than 30% of the protein reaches the plasma membrane [99]. While trafficking from the endoplasmic reticulum (ER) to the Golgi apparatus, immature forms of CFTR are recognized by quality-control systems and are eventually directed to the proteasome for degradation [101–104]. A critical step during CFTR biogenesis is the inefficient folding of the first of two cytoplasmically exposed nucleotide-binding domains (NBD1) of the membrane protein [105, 106]. The disease-causing F508 mutation localizes to NBD1 and further decreases the folding propensity of this domain. During the co-translational insertion of CFTR into the ER membrane, cytoplasmic Hsp70 and its co-chaperone Hdj-2 bind to NBD1 and facilitate intramolecular interactions between the domain and another cytoplasmic region of CFTR, the regulatory R-domain [96, 107]. However, Hsp70 is also able to present CFTR to the ubiquitin–proteasome system [97], and heterologous expression of CFTR in yeast revealed an essential role of cytoplasmic Hsp70 in CFTR turnover [98]. Hsp70 is thus a key player in the cellular surveillance system that monitors the folded state of CFTR at the ER membrane. Interestingly, CFTR and the disease form F508 are deposited in distinct pericentriolar structures, termed aggresomes, upon overexpression or proteasome inhibition [108]. Subsequent studies established that aggresomes are induced upon ectopic expression of many different aggregation-prone proteins (reviewed in Refs. [109] and [110]). Aggresomes form near the microtubule-organizing center in a manner dependent on the microtubule-associated motor protein dynein, and are surrounded by a “cage” of filamentous vimentin [108, 111]. Aggresome formation is apparently a specific and active cellular response when production of misfolded proteins exceeds the capacity of the ubiquitin–proteasome system to tag and remove these proteins. They likely serve to protect the cell from toxic “gain-of-function” activities acquired by misfolded proteins. Aggresomes are also of clinical relevance as they share remarkable biochemical and structural features, for example, with Lewy bodies, the cytoplasmic inclusion bodies found in neurons affected by Parkinson’s disease [112]. The pathways that regulate aggresome assembly are only now being explicated. Histone deacetylase 6 (HDAC6) appears to be a key regulator of aggresome assembly [113]. HDAC6 is a microtubule-associated deacetylase that has the capacity to bind both multi-ubiquitinated proteins and dynein motors and is believed to recruit misfolded proteins to the pericentriolar region for aggresome assembly. Deletion of HDAC6 prevents aggresome formation and sensitizes cells to the toxic effects of misfolded proteins, which supports the hypothesis that aggresomes sequester misfolded proteins to protect against their toxic activities. Components of the ubiquitin–proteasome system and chaperones such as Hsp70 are abundantly present in and are actively recruited to aggresomes [114–116]. Furthermore, elevating cellular Hsp70 levels can reduce aggresome formation by


stimulating proteasomal degradation [117]. It appears that these subcellular structures are major sites of chaperone – proteasome cooperation to mediate the metabolism of misfolded proteins. The formation of aggresome-like structures is also observed in dendritic cells that present foreign antigens to other immune cells [118]. Immature dendritic cells are located in tissues throughout the body, including skin and gut. When they encounter invading microbes, the pathogens are endocytosed and processed in a manner that involves the generation of antigenic peptides by the ubiquitin–proteasome system. Upon induction of dendritic cell maturation, ubiquitinated proteins transiently accumulate in large cytosolic structures that resemble aggresomes and were therefore termed DALIS (dendritic cell aggresome-like induced structures). It was speculated that DALIS formation may enable dendritic cells to regulate antigen processing and presentation. DALIS contain components of the ubiquitin–proteasome machinery as well as Hsp70 and the co-chaperone CHIP [118, 119]. Again, an interplay of molecular chaperones and the ubiquitin–proteasome system during regulated protein turnover is suggested. The cellular function of molecular chaperones is apparently not restricted to mediating protein folding; instead, chaperones emerge also as vital components on protein-degradation pathways. Remarkably, the balance between folding and degradation activities of chaperones can be manipulated. In cells treated with Hsp90 inhibitors, for example, with geldanamycin (see above), the chaperone-assisted activation of signaling proteins is abrogated and chaperone substrates such as the protein kinases Raf-1 and ErbB2 are rapidly degraded by the ubiquitin–proteasome system [64–69, 120]. This appears to be due, in part, to transfer of the substrates back to Hsp70 and progression toward the ubiquitin-dependent degradation pathway. Substrate interactions with chaperones – and consequently their commitment either toward the folding pathway or to their degradation via the ubiquitin– proteasome machinery – apparently serve as an essential post-translational protein quality-control mechanism within eukaryotic cells. The partitioning of proteins to either one of these mutually exclusive pathways is referred to as “protein triage” [121]. Although some misfolded proteins may be directly recognized by the proteasome [122], specific pathways within the ubiquitin–proteasome system are probably relied on for the degradation of most misfolded and damaged proteins. For example, E2 enzymes of the Ubc4/5 family selectively mediate the ubiquitylation of abnormal proteins as revealed in genetic studies in Saccharomyces cerevisiae [123]. It is well accepted that chaperones play a central role in the triage decision; however, less well understood are the events that lead to the cessation of efforts to fold a substrate, and the diversion of the substrate to the terminal degradative pathway. It is possible that chaperones and components of the ubiquitin–proteasome pathway exist in a state of competition for these substrates and that repeated cycling of a substrate on and off a chaperone maintains the substrate in a soluble state and increases, in a stochastic fashion, its likelihood of interactions with the ubiquitin machinery (Figure 5A). However, some data argue for a more direct role of the chaperones in the degradation process. Hsp70 plays an active and necessary role in the ubiquitylation of some substrates [124]; this activity of Hsp70

11


Fig. 5 Interplay of molecular chaperones with the ubiquitin–proteasome system. (A) Chaperones and the degradation machinery (i.e., ubiquitylation systems) compete with each other in the recognition of folding intermediates. Interaction with the chaperones directs the substrate towards folding. However, when the protein substrate is unable to attain a folded conformation, the chaperones maintain the folding intermediate in a soluble state that can be recog-

nized by the degradation machinery. (B) The chaperones are actively involved in protein degradation. Through an association with certain components of the ubiquitin conjugation machinery (degrading partner), the chaperones participate in the targeting of protein substrates to the proteasome. A competition between degrading partners and folding partners determines chaperone action and the fate of the protein substrate.

requires its chaperone function, indicating that conformational changes within substrates may facilitate recognition by the ubiquitylation machinery. Plausible hypotheses to explain these observations include direct associations between the chaperone and ubiquitin–proteasome machinery to facilitate transfer of a substrate from one pathway to the other, or conversion of the chaperone itself to a ubiquitylation complex (Figure 5B). It is also entirely possible that several quality-control pathways may exist and that the endogenous triage decision may involve aspects of each of these hypotheses.

6 The CHIP Ubiquitin Ligase: A Link Between Folding and Degradation Systems

Major insights into molecular mechanisms that underlie the cooperation of molecular chaperones with the ubiquitin–proteasome system were obtained through the functional characterization of the co-chaperone CHIP (reviewed in Ref. [40]). CHIP was initially identified in a screen for proteins containing tetratricopeptide repeat (TPR) domains, which are found in several co-chaperones – including Hip, Hop, and the cyclophilins – as chaperone-binding domains [27, 55] (Figure 2). CHIP

6 The CHIP Ubiquitin Ligase: A Link Between Folding and Degradation Systems 13

contains three TPR domains at its amino terminus, which are used for binding to Hsp70 and Hsp90 [35, 125]. Besides the TPR domains, CHIP possesses a Ubox domain at its carboxyl terminus [35] (Figure 2). U-box domains are similar to RING finger domains, but they lack the metal-chelating residues and instead are structured by intramolecular interactions [126]. The predicted structural similarity suggests that U boxes, like RING fingers, may also play a role in targeting proteins for ubiquitylation and subsequent proteasome-dependent degradation, and this possibility is borne out in functional analyses of U box–containing proteins [127, 128]. The TPR and U-box domains in CHIP are separated by a central domain rich in charged residues. The charged domain of CHIP is necessary for TPRdependent interactions with Hsp70 [35] and is also required for homodimerization of CHIP [129]. The tissue distribution of CHIP supports the notion that it participates in protein folding and degradation decisions, as it is most highly expressed in tissues with high metabolic activity and protein turnover: skeletal muscle, heart, and brain. Although it is also present in all other organs, including pancreas, lung, liver, placenta, and kidney, the expression levels are much lower. CHIP is also detectable in most cultured cells, and is particularly abundant in muscle and neuronal cells and in tumor-derived cell lines [35]. Intracellularly, CHIP is primarily localized to the cytoplasm under quiescent conditions [35], although a fraction of CHIP is present in the nucleus [97]. In addition, cytoplasmic CHIP traffics into the nucleus in response to environmental challenge in cultured cells, which may serve as a protective mechanism or to regulate transcriptional responses in the setting of stress [130]. CHIP is distinguished among co-chaperones in that it is a bona fide interaction partner with both of the major cytoplasmic chaperones Hsp90 and Hsp70, based on their interactions with CHIP in the yeast two-hybrid system and in vivo binding assays [35, 125]. CHIP interacts with the terminal-terminal EEVD motifs of Hsp70 and Hsp90, similar to other TPR domain–containing co-chaperones such as Hop [55, 131, 132]. When bound to Hsp70, CHIP inhibits ATP hydrolysis and therefore attenuates substrate binding and refolding, resulting in inhibition of the “forward” Hsp70 substrate folding/refolding pathway, at least in in vitro assays [35]. The cellular consequences of this “anti-chaperone” function are not yet clear, and in fact CHIP may actually facilitate protein folding under conditions of stress, perhaps by slowing the Hsc70 reaction cycle [130, 133]. CHIP interacts with Hsp90 with approximately equivalent affinity to its interaction with Hsp70 [125]. This interaction results in remodeling of Hsp90 chaperone complexes, such that the co-chaperone p23, which is required for the appropriate activation of many, if not all, Hsp90 client proteins, is excluded. The mechanism for this activity is unclear – p23 and CHIP bind Hsp90 through different sites – yet the consequence of this action is predictable: CHIP should inhibit the function of proteins that require Hsp90 for conformational activation. The glucocorticoid receptor is an Hsp90 client that undergoes activation through a well-described sequence of events that depend on interactions of the glucocorticoid receptor with Hsp90 and various Hsp90 co-chaperones, including p23, making it an excellent model to test this prediction. Indeed, CHIP inhibits glucocorticoid receptor substrate binding and


steroid-dependent transactivation ability [125]. Surprisingly, this effect of CHIP is accompanied by decreased steady-state levels of glucocorticoid receptor, and CHIP induces ubiquitylation of the glucocorticoid receptor in vivo and in vitro, as well as subsequent proteasome-dependent degradation. This effect is both U-box- and TPR-domain-dependent, suggesting that CHIP’s effects on GR require direct interaction with Hsp90 and direct ubiquitylation of GR and delivery to the proteasome. These observations are not limited to the glucocorticoid receptor. ErbB2, another Hsp90 client, is also degraded by CHIP in a proteasome-dependent fashion [120]. Nor are they limited to Hsp90 clients. For example, CHIP cooperates with Hsp70 during the degradation of immature forms of the CFTR protein at the ER membrane and during the ubiquitylation of phosphorylated forms of the microtubule-binding protein tau, which is of clinical importance due to its role in the pathology of Alzheimer’s disease [97, 134]. The effects of CHIP are dependent on both the TPR domain, indicating a necessity for interactions with molecular chaperones, and the U box, which suggests that the U box is most likely the “business end” with respect to ubiquitylation. The means by which CHIP-dependent ubiquitylation occurs is not clear. In the case of ErbB2, ubiquitylation depends on a transfer of the client protein from Hsp90 to Hsp70 [120], indicating that the final ubiquitylation complex consists of CHIP, Hsp70 (but not Hsp90), and the client protein. In any event, the studies are consistent in supporting a role for CHIP as a key component of the chaperone-dependent quality-control mechanism. CHIP efficiently targets client proteins, particularly when they are partially unfolded (as is the case for most Hsp90 clients when bound to the chaperone) or frankly misfolded (as is the case for most proteins binding to Hsp70 through exposed hydrophobic residues). Once the ubiquitylation activity of CHIP was recognized, it was logical to speculate that its U box might function in a manner analogous to that of RING fingers, which have recently been appreciated as key components of the largest family of ubiquitin ligases. If CHIP is a ubiquitin ligase, then its ability to ubiquitylate a substrate should be reconstituted in vitro when a substrate is added in the presence of CHIP, E1, an E2, and ubiquitin. Indeed, this is the case [135–137] (Figure 6). CHIP is thus the first described chaperone-associated E3 ligase. The ubiquitin ligase activity of CHIP depends on functional and physical interactions with a specific family of E2 enzymes, the Ubc4/Ubc5 family, which in humans comprises the E2s UbcH5a, UbcH5b, and UbcH5c. Of interest is the fact that the Ubc4/Ubc5 E2s are stress-activated, ubiquitin-conjugating enzymes [123]. CHIP can therefore be seen as a co-chaperone that, in addition to inhibiting traditional chaperone activity, converts chaperone complexes into chaperone-dependent ubiquitin ligases. Indeed, the chaperones themselves seem to act as the main substrate-recognition components of these ubiquitin ligase complexes, as efficient ubiquitylation of chaperone substrates by CHIP depends on the presence of Hsp70 or Hsp90 in reconstituted systems [136, 137] (Figure 6). The chaperones apparently function in a manner analogous to F-box proteins, which are required as substrate recognition modules in many RING finger–containing ubiquitin ligase complexes [138–140]. Recently, another surprising function for CHIP has been identified, that of activation of the stress-responsive transcription factor heat shock factor-1 (HSF1)

7 Other Proteins That May Influence the Balance

Fig. 6 Characterization of CHIP as a chaperone-associated ubiquitin ligase. Purified CHIP, UbcH5b, the ubiquitinactivating enzyme E1, ubiquitin, and the Hsp70–Hsp40 chaperone system were incubated with the bacterially expressed protein kinase Raf-1 (for details, see Ref. [137]). Raf-1 and ubiquitylated forms of the kinase

(ub(n) -Raf-1) were detected by immunoblotting using a specific anti-Raf-1 antibody. Efficient ubiquitylation of Raf-1 through the CHIP conjugation machinery depends on the recognition of the chaperone substrate by Hsp70, which presents the kinase to the conjugation machinery.

[130]. Through this association, CHIP regulates the expression of chaperones such as Hsp70 independently of its ability to modify their function through direct interactions. The mechanisms through which CHIP activates HSF1 are not entirely clear, but they are dependent in part on the induction of HSF1 trimerization, which is required for nuclear import and DNA binding. In addition, activation of HSF1 by CHIP seems to be independent of CHIP’s ubiquitin ligase activity. The consequences of this activation are important for the response to stress, in that cells lacking CHIP are prone to stress-dependent apoptosis and mice deficient in CHIP (through homologous recombination) succumb rapidly to thermal challenge. These data indicate that CHIP plays a heretofore unsuspected role in coordinating the response to stress, not only by serving as a rate-limiting step in the degradation of damaged proteins but also by increasing the buffering capacity of the chaperone system to guard against stress-dependent proteotoxicity.

7 Other Proteins That May Influence the Balance Between Chaperone-assisted Folding and Degradation

CHIP is ideally suited to mediate chaperone-proteasome cooperation, as it combines a chaperone-binding motif and a domain that functions in ubiquitindependent degradation within its protein structure (Figure 2). Some other cochaperones display a similar structural arrangement [40]. For example, BAG-1 contacts Hsp70 through a BAG-domain located at its carboxyl terminus and, in

15


Fig. 7 Schematic presentation of the BAG1–Hsp70–CHIP complex. BAG-1 associates with the ATPase domain of Hsp70, while CHIP is bound to the carboxyl terminus. BAG-1 mediates an association of Hsp70 with the proteasome via its ubiquitin-like

domain (ubl), whereas CHIP acts in conjunction with Ubc4/5 as a chaperoneassociated ubiquitin ligase to mediate the attachment of a polyubiquitin chain to the chaperone substrate.

addition, possesses a central ubiquitin-like domain that is used for binding to the proteasome [141] (Figure 2). The co-chaperone thus belongs to a family of ubiquitin domain proteins (UDPs), many of which were shown to be associated with the proteasome [142]. This domain architecture enables BAG-1 to provide a physical link between Hsp70 and the proteasome, and elevating the cellular levels of BAG-1 results in a recruitment of the chaperone to the proteolytic complex. Notably, BAG-1 and CHIP occupy distinct domains on Hsp70 (Figure 7). Whereas BAG-1 associates with the amino-terminal ATPase domain, CHIP binds to the carboxylterminal EEVD motif of Hsp70 [35, 37]. Ternary complexes that comprise both co-chaperones associated with Hsp70 can be isolated from mammalian cells, suggesting a cooperation of BAG-1 and CHIP in the regulation of Hsp70 activity on certain degradation pathways. In fact, BAG-1 is able to stimulate the CHIPinduced degradation of the glucocorticoid hormone receptor [137]. A cooperation of diverse co-chaperones apparently provides additional levels of regulation to alter chaperone-assisted folding and degradation pathways. Interestingly, BAG-1 and also Hsp70 and Hsp90 are themselves substrates of the CHIP ubiquitin ligase [135, 143] (J.H. unpublished). Yet, CHIP-mediated ubiquitylation of the chaperones and the co-chaperone does not induce their proteasomal degradation. Instead, it seems to provide additional means to regulate the association of the chaperone systems with the proteasome. In the case of BAG-1, ubiquitylation mediated by CHIP indeed stimulates the binding of the co-chaperone to the proteasome [143]. It remains to be elucidated, however, why Hsp70 and BAG-1 are not degraded when sorted to the proteasome through CHIP-induced ubiquitylation, in contrast to chaperone substrates such as the glucocorticoid hormone receptor. Possibly, the folded state of the proteins may serve to distinguish targeting factors and substrates doomed for degradation. Efficient ubiquitylation of BAG-1 mediated by CHIP is dependent on the formation of the ternary BAG-1–Hsp70–CHIP complex [143]. The formed chaperone complex would thus expose multiple signals for sorting to the proteasome, e.g., the integrated ubiquitin-like domain of BAG-1 and polyubiquitin chains attached to BAG-1, Hsp70, and the bound protein substrate. Such a redundancy of sorting information might be considered unnecessary. Intriguingly, however, several

7 Other Proteins That May Influence the Balance

subunits of the regulatory 19S particle of the proteasome are currently thought to act as receptors for polyubiquitin chains and integrated ubiquitin-like domains, including Rpn1, Rpn2, Rpt5, and Rpn10. The Rpn10 subunit was initially identified as a polyubiquitin chain receptor and was later shown to also bind integrated ubiquitin-like domains presented by UDPs [144–146]. Rpn10 possesses two distinct ubiquitin-binding domains, of which only one is used for UDP recognition [145–147]. However, conflicting data exist as to whether the subunit acts as a ubiquitin receptor in the context of the assembled 19S complex [148, 149]. More recently, Rpn1 was identified as a receptor for integrated ubiquitin-like domains [149], and a similar function may be fulfilled by the Rpn1-related subunit Rpn2 [150]. Polyubiquitin chains seem to be recognized by the Rpt5 subunit, one of the AAA ATPases present in the ring-like base of the regulatory 19S complex [151]. Its receptor function was revealed when tetraubiquitin was cross-linked to intact proteasomes [148]. Multiple docking sites for ubiquitin-like domains and polyubiquitin chains are apparently displayed by the regulatory particle of the proteasome. This may provide a structural basis for the recognition of multiple sorting signals exposed by the CHIP–chaperone complex (Figure 8). A similar mechanism involving multiple-site binding at the proteasome was recently proposed based on the

Fig. 8 The co-chaperone network that determines folding and degradation activities of Hsp70. BAG-1 and CHIP associate with Hsp70 to induce the proteasomal degradation of a Hsp70-bound protein substrate. When BAG-1 is displaced by binding of HspBP1 to the ATPase domain of Hsp70, the ubiquitin ligase activity of CHIP is attenuated in the formed complex, enabling

CHIP to modulate Hsp70 activity without inducing degradation. The ATPase domain can also be occupied by Hip, which stimulates the chaperone activity of Hsp70 and participates in the Hsp70/Hsp90-mediated regulation of signal transduction pathways. At the same time, Hop displaces CHIP from the carboxyl terminus of Hsp70 and recruits Hsp90 to the chaperone complex.

17


observation that two unrelated yeast ubiquitin ligases associate with specific subunits of the 19S regulatory complex [152]. In these cases substrate delivery involves interactions of proteasomal subunits with the substrate-bound ubiquitin ligase, with the polyubiquitin chain attached to the substrate, and with the substrate itself. Multiple-site binding may function to slow down dissociation of the substrate from the proteasome and to facilitate transfer into the central proteolytic chamber through ATP-dependent movements of the subunits of the 19S particle. Human cells contain several BAG-1-related proteins: BAG-2, BAG-3 (CAIR-1; Bis), BAG-4 (SODD), BAG-5, and BAG-6 (Scythe, BAT3) [153] (Figure 2). It appears that BAG proteins act as nucleotide-exchange factors to induce substrate unloading from Hsp70 on diverse protein folding, assembly, and degradation pathways. Notably, BAG-6 is another likely candidate for a co-chaperone that regulates protein degradation via the ubiquitin–proteasome pathway. Similar to BAG-1, BAG-6 contains a ubiquitin-like domain that is possibly used for proteasome binding [154]. However, experimental data verifying a role of BAG-6 in protein degradation remain elusive so far. The cooperation of diverse co-chaperones not only may allow promotion of chaperone-associated degradation but also may provide the means to confine the destructive activity of CHIP. The Hsp70-binding protein 1 (HspBP1) seems to fulfill such a regulatory function [155]. HspBP1 was initially identified in a screen for proteins that associate with the ATPase domain of Hsp70 and was shown to stimulate nucleotide release from the chaperone [156, 157]. Notably, association of HspBP1 with the ATPase domain blocks binding of BAG-1 to Hsp70 and at the same time promotes an interaction of CHIP with Hsp70’s carboxyl terminus. In the formed ternary HspBP1–Hsp70–CHIP complex, the ubiquitin ligase activity of CHIP is attenuated and Hsp70 as well as a chaperone substrate are no longer efficiently ubiquitylated [155]. By interfering with CHIP-mediated ubiquitylation, HspBP1 stimulates the maturation of CFTR and promotes the sorting of the membrane protein to the cell surface. HspBP1 apparently functions as an antagonist of the CHIP ubiquitin ligase to regulate Hsp70-assisted folding and degradation pathways (Figure 8). The HspBP1-mediated inhibition of the ubiquitin ligase activity may enable CHIP to modulate the Hsp70 ATPase cycle without inducing degradation. In fact, degradation-independent functions of CHIP have recently emerged [130, 133, 158, 159]. CHIP was shown to regulate the chaperone-assisted folding and sorting of the androgen receptor and of endothelial nitric oxide synthase without inducing degradation [158, 159]. Moreover, CHIP fulfills an essential role in the chaperone-mediated regulation of the heat shock transcription factor, independent of its degradation-inducing activity [130]. It remains to be seen, however, whether HspBP1 cooperates with CHIP in these instances, as HspBP1 displayed a certain specificity with regard to chaperone substrates. The co-chaperone interfered with the degradation of CFTR, but did not influence the CHIP-mediated turnover of the glucocorticoid hormone receptor. Such a client specificity may arise in part from the fact that HspBP1 inhibits the ubiquitin ligase activity of CHIP in a complex with Hsc70, but leaves Hsp90-associated

8 Further Considerations

ubiquitylation unaffected [155]. In addition, direct interactions between HspBP1 and a subset of chaperone substrates may contribute to substrate selection. In any case, the cooperation of CHIP with other co-chaperones apparently provides a means to regulate chaperone-assisted protein degradation. It is likely that there are multiple degradation pathways for misfolded proteins in the eukaryotic cytoplasm. Although CHIP participates in the degradation of chaperone substrates induced by applying Hsp90 inhibitors to cell cultures (see above), drug-induced degradation is not abrogated in cells that lack the CHIP ubiquitin ligase [120]. Furthermore, CHIP cooperates with Hsp70 in the ER-associated degradation of CFTR, but the Hsp70-assisted degradation of apoB at the cytoplasmic face of the ER membrane does not involve CHIP [97]. Taken together, these data strongly argue for the existence of other, yet to be identified, ubiquitin ligases that are able to target chaperone substrates to the proteasome. A likely candidate in this regard is Parkin, a RING finger ubiquitin ligase, whose activity is impaired in juvenile forms of Parkinson’s disease [17]. Hsp70 and CHIP were found to be associated with Parkin in neuronal cells, suggesting an involvement of Parkin in the proteasomal degradation of chaperone substrates [160]. Interestingly, α-synuclein, the main component of protein deposits observed in dopaminergic neurons of Parkinson patients, and synphilin, a protein that binds α-synuclein and induces deposit formation, both associate with yet other ubiquitin ligases: Siah-1 and Siah-2 [161, 162]. In the case of Siah-1, a link to cytoplasmic chaperone systems is suggested by the finding that the Hsp70 co-chaperone BAG-1 is a binding partner of the ubiquitin ligase and suppresses some of the cellular activities of Siah-1 [163]. Taken together, it is tempting to speculate about a role of Parkin and Siah on chaperone-assisted degradation pathways; yet, this remains to be explored in detail.

8 Further Considerations

Although the appreciation of interplay between molecular chaperones and ubiquitin-dependent proteolysis has greatly expanded over the past decade, a number of critical issues remain to be resolved. It is not entirely clear what determines whether a misfolded protein will undergo repeated attempts at misfolding versus diversion to the ubiquitin–proteasome pathway. Recruitment of CHIP into chaperone complexes appears to be a critical component of this reaction, which therefore begs the question as to what regulates this step. Since this step in protein quality control must be both rapidly activated and easily reversible, it is likely that regulation occurs at the post-translational level rather than through changes in steady-state protein levels. The precise sorting mechanisms for ubiquitinated proteins are also unclear. BAG-1 is a player, and it is also likely that overlap exists to some extent for sorting of the cytoplasmic and endoplasmic reticulum quality-control pathways. Nevertheless, much remains to be learned about these steps. From a broader perspective, it is now also imperative to understand the pathophysiological roles of cytoplasmic quality-control mechanisms regulated by

19


chaperone-proteasome interactions. As mentioned previously, there is a strong association between chaperone dysfunction and accumulations of misfolded proteins that characterizes genetic neurodegenerative diseases. An imbalance between protein folding and degradation may also contribute to some features of senescence and organismal aging. The link between chaperone systems and aging is based on increasing appreciation that modified, misfolded, and aggregated proteins accumulate with age [164]. Dysregulation of chaperone expression has been observed with aging and is therefore implicated in aging-related changes [165]; in general, it is accepted that induction of the major chaperones is impaired with aging, a fact confirmed by recent gene-profiling experiments in vivo [166], although given the diversity of chaperones it is probably not surprising that age-related changes in expression are fairly complicated [167]. The mechanism underlying this dysregulation is not entirely clear, but seems to be due in part to impaired activation of the stress-responsive transcription factor HSF1. Overexpression of heat shock proteins in yeast, C. elegans, and Drosophila leads to increased longevity [168–170]. More recently, conclusive genetic evidence from C. elegans indicates that mutation of HSF1 causes a dramatic and significant reduction in lifespan [170, 171], further implicating the accumulation of misfolded proteins in age-related phenotypes.

9 Conclusions

The associations between molecular chaperones and the ubiquitin–proteasome system represent a critical step in the response to proteotoxic damage. Whether attempts should be made to refold damaged proteins (thus conserving cellular resources) or degrade them instead (to prevent the possibility of protein aggregation and concomitant toxicity) requires a consideration of cellular economy. Defects in the quality-control mechanisms may have enormous consequences even if only slight imbalances occur between protein folding and degradation, as these imbalances can cause accumulated toxicity over time. The relationship between chaperone–proteasome interactions and pathophysiological events is only now being unraveled. Modulation of this system may provide a unique therapeutic target for degenerative diseases and pathologies associated with aging. References 1 U. SCHUBERT, L. C. ANTON, J. GIBBS, C. C. NORBURY, J. W. YEWDELL, J. R. BENNINK, Rapid degradation of a large fraction of newly synthesized proteins by proteasomes, Nature 404 (2000) 770–774. 2 J. P. TAYLOR, J. HARDY, K. H. FISCHBECK, Toxic proteins in neurodegenerative disease, Science 296 (2002) 1991–1995.

3 A. HORWICH, Protein aggregation in disease: a role for folding intermediates forming specific multimeric interactions, J. Clin. Invest. 110 (2002) 1221–1232. 4 E. SCHERZINGER, R. LURZ, M. TURMAINE, L. MANGIARINI, B. HOLLENBACH, R. HASENBANK, G. P. BATES, S. W. DAVIES, H. LEHRACH, E. E. WANKER,

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29


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1

Ubiquitin-conjugating Enzymes Michael J. Eddins Johns Hopkins School of Medicine, Baltimore, USA

Cecile M. Pickart Johns Hopkins Bloomberg School of Public Health, Baltimore, USA


1 Introduction

In this chapter we review the biochemical, structural, and biological properties of ubiquitin-conjugating enzymes (also called E2 enzymes). Because length restrictions preclude a comprehensive treatment, we focus on key findings that have revealed important general insights and principles. Throughout the piece we try to point out important unanswered questions concerning the E2 enzyme family. A few words about nomenclature are necessary. The yeast E2 genes were numbered in the order of their discovery, but the situation is more complicated in mammals. There are currently three naming systems in use for human E2s: one based on protein molecular mass (e.g. E225K is a 25-kD E2), one based on temporal order of gene cloning (e.g. UbcH10 is specified by the tenth E2 gene cloned in humans), and one based on relationship to yeast E2s (e.g. HR6A is one of two human homologs of yeast Rad6/Ubc2). The second system is the least ambiguous, but also the least informative. In this chapter, we generally name mammalian E2s according to their relationship to yeast E2s. When this is not possible, we use one of the published names.

2 Historical Background

Ubiquitin’s best-understood function is that of a protein cofactor in an intracellular protein-degradation pathway that terminates with the destruction of Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Ubiquitin-conjugating Enzymes

Fig. 1 The ubiquitin-conjugation pathway. Steps in ubiquitin activation and substrate modification. E1, ubiquitin activating enzyme; E2, ubiquitin-conjugating enzyme; E3, ubiquitin-protein ligase. Atoms involved in the thiol ester and amide bonds are shown.

ubiquitin-tagged substrates by 26S proteasomes [1]. The discovery in 1980 that this 76-residue protein is conjugated to proteolytic substrates through the formation of a peptide-like bond, and in an ATP-dependent manner, suggested that ubiquitin activation would be part of the conjugation process [2, 3]. A ubiquitin activating enzyme (E1) was soon identified and shown to employ an aminoacyl-tRNA-synthetase-like mechanism [4]. E1 first catalyzes the addition of an adenylate moiety to the carboxyl group of ubiquitin’s C-terminal residue, G76. The AMP-bound ubiquitin is then transferred to a cysteine residue in the E1 active site, concomitant with the formation of a new molecule of ubiquitin adenylate. The thiol-linked ubiquitin is the proximal source of activated ubiquitin for downstream steps. From a chemical point of view, the E1/ubiquitin thiol ester should be competent to donate ubiquitin to a substrate amino group. In fact, aminoacyl-enzyme thiol esters are used in exactly this way in non-ribosomal polypeptide synthesis, a process that was discovered around the same time as ubiquitin–protein conjugation [5]. In spite of the attractive simplicity of this model, however, biochemical reconstitution studies showed that besides E1 two additional fractions were required to conjugate ubiquitin to a model substrate. They were called ubiquitin carrier protein (E2) and ubiquitin-protein ligase (E3), respectively, since the respective factors seemed to act sequentially [6]. Interestingly, the E2 factor apparently formed a thiol ester with ubiquitin. Based on these results, Hershko and co-workers proposed the “ubiquitin conjugation cascade” (Figure 1). Multiple thiol ester-forming proteins were present in the E2 fraction [6], but only the smallest of them reconstituted substrate ubiquitination catalyzed by the thenknown E3 [7]. This result suggested that there could be multiple E2s with distinct functional properties. Confirmation of this hypothesis came with the cloning of the first two E2 genes, RAD6/UBC2 and CDC34/UBC3, which indeed encoded homologous yeast proteins with a signature cysteine-containing active-site motif [8, 9]. The two E2s functioned in distinct biological processes – DNA damage tolerance [8, 10] and cell-cycle control [9] – providing the first hint that ubiquitination might regulate a broad range of biological processes. A family of E2 enzymes naturally suggested that there would also be a family of E3 enzymes. This prediction has since been strikingly confirmed. We now know that specific E2/E3 complexes function to modify specific substrates with ubiquitin.

3 What is an E2? 3

3 What is an E2?

A protein can be identified as an E2 enzyme according to several different criteria. Functionally, the E2 occupies an intermediate position in the conjugation cascade – that is, it acts between the E1 and the E3 (Figure 1). This property accounts for the original name of ubiquitin carrier protein, which drew an analogy to the acyl carrier proteins used in fatty acid biosynthesis and non-ribosomal peptide synthesis [6]. Subsequently, with the recognition that E2 enzymes often play an active role in conjugation, the conjugating enzyme name gained favor. Mechanistically, the E2 first participates in a thiol ester transfer reaction, in which the activated ubiquitin is moved from the active-site cysteine of E1 to that of the E2 (Figure 1). The E2/ubiquitin thiol ester intermediate is strictly required for downstream steps, as shown by ablation of substrate ubiquitination following mutation of the active-site cysteine residues of different E2s (see, for example, Refs. [11, 12]). The ubiquitin is then transferred from the E2 active site to the ε-amino group of the substrate’s lysine residue, forming an isopeptide bond. The conjugation site can also be a specific lysine on a previously conjugated ubiquitin, which leads to polyubiquitin chain elongation; chains linked through K48 are the principal signal for targeting substrates to proteasomes [1]. Transfer of ubiquitin to the substrate requires the assistance of the E3 [1, 6]. If this enzyme belongs to the HECT domain family (Homologous to E6AP C-Terminus), the ubiquitin is first transferred to an active-site cysteine residue of the E3; if the E3 belongs to the RING domain family (Really Interesting New Gene), ubiquitin is transferred directly to the substrate’s amino group (Section 6.3). Collectively, these properties constitute the biochemical definition of an E2 enzyme: it is a protein that accepts ubiquitin in thiol ester linkage from E1, and cooperates with an E3 enzyme to deliver this ubiquitin to the substrate. The functional specialization of individual E2s (Section 4) reflects the specificity of interaction of each E2 with its cognate E3(s), in conjunction with the E3’s substrate specificity. Therefore an E2 enzyme can also be defined according to the cognate E3(s) with which it interacts. The E3 partners of many of the eleven ubiquitin-dedicated E2s in Saccharomyces cerevisiae are conserved in higher organisms (Section 4). However, both the E2 and E3 families are much larger in higher organisms than in yeast. Present accounting suggests that there are 50–70 E2s, and hundreds of E3s, in mammals [13, 14]. The amino acids surrounding the thiol ester-forming cysteine residue are particularly highly conserved, but there is sequence similarity throughout the 150-residue E2 core domain (Figure 2). This bioinformatic definition makes it easy to identify E2 genes in sequenced genomes [13, 14]. In fact, many E2s consist of just this core domain (Figure 2). The fact that such E2s are often functionally distinct from one another indicates that modest sequence variation within the core domain can be highly significant. Structural biology has begun to shed light on this structure/function correlation (Section 6). Other E2s display N- and C-terminal extensions to the core


3 What is an E2? 5

Fig. 2 Ubc13 (1JBB). Canonical "/$ E2 fold with the active-site cysteine shown in ball-and-stick.

domain (Figure 2), which may play a role in E3 and/or substrate specificity (see Refs. [15–19]). Structural biology provides a final way to define an E2 enzyme. As expected from the strong sequence conservation, the E2 core domain adopts a conserved fold. At the time this article was being prepared, twelve different E2 structures had been deposited in the Protein Data Bank. The average root-mean-square deviation of the 150 Cα positions of these structures is less than 2Å. E2s are α/β proteins containing a central anti-parallel four-stranded β-sheet (S1–S4), four α-helices (H1, H3, H4, H5), and a small 310 helix (H2) (Figure 3) [20, 21]. The cysteine is located on an extended loop after β-strand 4 and immediately before the short 310 helix H2. The active-site cysteine sits in a shallow groove composed of residues from the H3–H4 and S4–H2 loops. The canonical α/β E2 fold is highly versatile, allowing E2 enzymes to associate with several different proteins in the ubiquitin conjugation cascade without any perturbation of the E2’s tertiary structure (Section 6). Residues occupying the face opposite the active site are less conserved than those surrounding the cysteine [21]. Sequence variation in this region contributes to the functional diversity of the E2 family by permitting specific interactions of individual E2s with cognate E3s and (perhaps) substrates.


4 Functional Diversity of Ubiquitin-conjugating Enzymes

The functional range of the ubiquitin-conjugating enzyme family is easily appreciated by considering the family members in a single organism. Table 1 summarizes key properties of the complete set of E2s in the yeast S. cerevisiae, including notable structural features, known cognate E3(s) and their key substrates, and biological functions (see also [22, 23]). We cannot give a comprehensive review of E2 functions in higher organisms, but we do comment on some notable instances of functional conservation, expansion, and divergence (see also [23]). 4.1 Functions Related to Proteasome Proteolysis

In many cases, the specific function(s) of a given E2 enzyme reflect its role in targeting one or more substrates for degradation by 26S proteasomes. The scope of this function varies considerably between E2 family members, however. At one extreme, the functionally redundant enzymes Ubc4 and Ubc5 are necessary for the turnover of many substrates, as shown by a marked reduction in the rate of turnover of endogenous short-lived and abnormal proteins in a ubc4ubc5 yeast strain [24]. The slow growth and stress sensitivity of this strain [24, 25] can also be ascribed to inhibition of proteasomal proteolysis since these phenotypes are characteristic of proteasome mutants [22, 26]. Despite the important role of Ubc4/5 in proteasome degradation, few E3 partners relevant to this function have been identified. One is Ufd4, a HECT-domain E3 that mediates the degradation of linear ubiquitin fusion proteins [27]. A ufd4 strain grows normally, however, indicating that Ubc4 has other cognate E3s. Rsp5, an essential HECT-domain E3, is one likely candidate since this E3 partners with Ubc4 in other pathways (see below). UBC1 is essential for viability in the ubc4ubc5 strain, suggesting that Ubc1 shares substantial functional overlap with Ubc4/5 in directing substrates to proteasomes for degradation [28]. The Ubc4/5 sub-family of E2s is much larger in mammals, where it includes both constitutively and selectively expressed enzymes. Notable human E2s in this group are UbcH5a/b/c, UbcH7, and UbcH8 (see Ref. [23]). The expansion is likely to reflect the much larger size of the E3 family in higher organisms. However, while the results of in vitro conjugation assays and protein-protein interaction studies suggest that certain E3s partner specifically with individual Ubc4/5 sub-family members, the degree of E3 (hence, functional) selectivity in the cellular setting remains quite uncertain (discussed in Refs. [23, 29]). RNA interference studies and mouse knockout models may be helpful in addressing this question in the future. Ubc3/Cdc34 supports the proteasome-mediated proteolysis of numerous substrates through its role as the specific E2 partner of a large family of multi-subunit RING E3s called SCF E3s (Skp-Cullin-F-box, Section 6.3). This role is preserved in higher organisms [30]. Yeast ubc3 mutants arrest in G1 phase of the cell cycle because they fail to degrade Sic1 [31], an inhibitor of the G1/S transition that is

4 Functional Diversity of Ubiquitin-conjugating Enzymes 7 Table 1 E2 enzymes of the yeast Saccharomyces cerevisiae

Gene

Amino acids

UBC1

215 [28]

Cognate E3

Unknown

Hrd1 [39] UBC2 (RAD6)

172 [8]

Ubr1 [156]

Ubr1 [52]

Rad18 [73], [157] Bre1 [67, 69]

UBC3 (CDC34)

295 [9]

SCF E3s

UBC4

148 [24]

Unknown Doa10 Rsp5

UBC5

148 [24]

UBC6

250 [46]

See UBC4

Unknown

Doa10 [42] Doa10 [42] UBC7

165 [25] Hrd1 [39]

Doa10 [42] UBC8

206 [160]

Unknown

UBC10

165 [75]

Unknown

Functions and substrates Short C-terminal tail harbors ubiquitin-associated (UBA) domain [155] Essential in ubc4ubc5 genetic background, suggesting a redundant role with Ubc4/5 in proteasomal turnover of short-lived and abnormal proteins [28] Role in ERAD that is not fulfilled by Ubc4/5 [39, 40] Proteasomal degradation of N-end rule [158] substrates, including cohesin fragment [51] Proteasomal degradation of Cup9 transcriptional repressor regulates peptide import DNA-damage tolerance [8] via monoubiquitination of PCNA [65] (non-proteolytic function) Ubiquitination of histone H2B [68] regulates gene transcription and silencing [71] (non-proteolytic function) Essential gene; long C-terminal tail; targets diverse substrates for proteasomal degradation [30, 34, 36]; regulation of cell-cycle progression Proteasomal degradation of diverse shortlived proteins [24] Proteasomal degradation of MATα2 transcriptional repressor [49] Endocytosis of membrane proteins [60, 61]; protein trafficking (see Ref. [63]) 92% identical to Ubc4; functionally redundant [24] C-terminal tail provides anchoring to ER membrane [46] Together with Ubc7, proteasomal degradation of some ERAD substrates [47, 48, 159] Proteasomal turnover of Ubc6 is Ubc6-, Ubc7-, and Doa10-dependent [42, 50] In conjunction with Ubc7, proteasomal turnover of MATα2 Localized to ER membrane via Cue1 [37] Role in proteasomal degradation via ERAD [38, 48] confers resistance to cadmium and other ER stresses [25] In conjunction with Ubc7, proteasomal turnover of MATα2 Glucose-induced proteasome degradation of fructose-1,6-bisphosphatase [57] Also called Pas2/Pex10. Peroxisome biogenesis [75]; Pex10 is a candidate E3 [78]

8 Ubiquitin-conjugating Enzymes Table 1 (continued)

Gene

Amino acids

Cognate E3

Functions and substrates

UBC11

156 [84]

Unknown

UBC13

153

Unknown; similar to clam E2-C (E2-C functions in mitotic cyclin turnover [79], but Ubc11 is dispensable for this process in yeast [84]) Heterodimerizes with Mms2 (UEV) [72] DNA-damage tolerance [72] via polyubiquitination of PCNA [65] (non-proteolytic function) Essential gene; E2 dedicated to Smt3 (SUMO) [163] Septin modification E2 dedicated to Rub1 (Nedd8) Modification of specific cullin lysine residue activates cullin-based E3s [165]

Rad5 [73]

UBC9

157 [161]

UBC12

188 [164]

Siz1/2 [162] SCF E3s

recognized and polyubiquitinated by a specific SCF E3 [32, 33]. Although this is the only essential function of yeast Ubc3 [31], this E2 partners with many other SCF E3s to target diverse substrates for degradation by proteasomes (see Refs. [30, 34–36]). Although studies in yeast suggest that Cdc34 is the main E2 partner of SCF E3s, some SCF E3s seem to partner with Ubc4/5-type E3s (see Refs. [23, 36]). Ubc3 has a long C-terminal tail (Table 1), making it the most distinctive yeast E2 in terms of primary structure. A chimeric E2 in which the Ubc3 tail is appended to the core domain of Ubc2 fulfills the essential function of Ubc3 in yeast, suggesting that the tail of Ubc3 is necessary for key interactions with the E3 or Sic1 [15, 16]. Ubc7 is localized to the endoplasmic reticulum (ER) through an interaction with a partner protein, Cue1 [37], and plays a major role in proteasome degradation. Ubc7 acts on misfolded proteins of the ER, which are ejected from this compartment as a prelude to ubiquitination at the cytosolic face of the ER membrane and degradation by cytosolic proteasomes [38]. Ubc7’s role in ERAD (ER Associated Degradation) explains why a ubc7 strain is conditionally sensitive to agents that induce protein misfolding in the ER [25, 39–41]. Ubc7 frequently partners with Hrd1, an ERlocalized RING E3 [39], but some ERAD substrates of Ubc7 seem to be recognized in cooperation with a different ER-localized RING E3, Doa10 [42]. Consistent with Ubc7’s prominent role in ERAD, the yeast UBC7 and CUE1 genes are induced as part of the Unfolded Protein Response (UPR) and there is a synthetic lethal relationship between certain ERAD and UPR genes [40, 41]. Yeast Ubc1 also plays a significant, but poorly-defined, role in ERAD [39, 40]. Mammalian Ubc7 also functions in ERAD [43–45]. Ubc6 localizes to the ER through its own C-terminal membrane anchor [46]. Although Ubc6 plays a role in ERAD, its function in this process is less conspicuous than that of Ubc7 [43, 47, 48]. Interestingly, Ubc6 and Ubc7 both contribute to the Doa10-dependent degradation of a soluble nuclear protein [42, 49], and Ubc6 is

4 Functional Diversity of Ubiquitin-conjugating Enzymes 9

itself rapidly degraded by proteasomes in a manner that depends on its own activesite cysteine, its C-terminal membrane anchor, functional Ubc7, and Doa10 [42, 50]. The purpose of this instability remains mysterious. Ubc2 functions rather selectively in proteasome proteolysis. In yeast, two specific E3 partners are known, leading to proteasome degradation events that regulate chromosome stability [51], peptide import [52], and homing endonuclease stability [53]. Mammals have two closely-related Ubc2 isoforms, each of which complements most of the functions of the yeast ubc2 strain [54]. But the mammalian Ubc2 isoforms also have specialized functions – one of them is required for spermatogenesis in the mouse [55] and at least one of them can be inferred to be necessary for cardiovascular development [56]. So far, Ubc8 has been implicated in the regulated turnover of just one substrate, and its E3 partner(s) remain unknown [57]. Interestingly, the closest mammalian relative of yeast Ubc8 is expressed with a restricted tissue specificity and (in some tissues) in a regulated manner [58, 59]. 4.2 Endocytosis and Trafficking

Just because an E2 functions in proteasome proteolysis does not mean that its functions are limited to this pathway. This is because the E2’s functional range is largely determined by the substrate specificity of its E3 partner(s). For example, yeast Ubc4 and Ubc5 play a prominent role in proteasome degradation, but they also cooperate with a HECT E3, Rsp5, to mono-ubiquitinate certain plasma membrane receptors [60–62]. This modification signals receptor endocytosis, leading to degradation in the vacuole (equivalent to the mammalian lysosome). Ubc1 is partially redundant with Ubc4/5 in endocytosis [61, 62], as seen in ubiquitination reactions leading to proteasome degradation (above). Thus, Ubc1, 4, and 5 may be able to substitute for one another in complexes involving many different E3s, probably reflecting the strong conservation of the core domain between Ubc1 and Ubc4/5. Ubc4 and Ubc5 may also act with Rsp5 to regulate protein trafficking downstream of endocytosis (reviewed in Ref. [63]). 4.3 Non-proteolytic Functions

As mentioned in Section 2, Ubc2 is the defining player in a conserved DNA damagetolerance pathway [8, 10, 64]. Here Ubc2 partners with a RING E3, Rad18, to modify a DNA polymerase processivity factor with a single ubiquitin [65]. This modification signals error-prone bypass of DNA lesions [66]. Ubc2 partners with a different RING E3 (Table 1) to mono-ubiquitinate histone H2B [67–69]. This modification promotes histone methylation, which in turn regulates transcription and silencing [70, 71]. Ubc13 participates in the same DNA damage-tolerance pathway as Ubc2 [72]. Ubc13 collaborates with two enzyme partners (Table 1) to modify the DNA polymerase cofactor (above) with a K63-linked polyubiquitin chain, which signals


error-free lesion bypass [65, 72, 73]. In higher organisms, Ubc13 also helps to synthesize K63-linked polyubiquitin chains in a second non-proteolytic signaling pathway (see Ref. [74]). The function and mechanism of Ubc13 are discussed in more detail below (Section 7). 4.4 E2s of Uncertain Function

Ubc10 is required for the biogenesis of the peroxisome, an oxidative organelle [75]. This E2 plays a role in peroxisomal protein import [76] and is recruited to the peroxisomal membrane through an interaction with a partner protein [77]. Membrane-localized Ubc10 also seems to be spatially proximal to Pex10, which has a RING-like domain [78]. Whether Pex10 is a cognate E3 of Ubc10 remains to be determined, as does the mechanistic role of ubiquitin conjugation in peroxisome biogenesis. The function of the remaining yeast E2, Ubc11, remains uncertain. Ubc11 is very similar to E2-C, a clam E2 that acts with an essential multi-subunit RING E3, the anaphase promoting complex (APC) or cyclosome, to ubiquitinate mitotic cyclins [79]. This reaction leads to cyclin degradation by proteasomes, which drives exit from mitosis [35, 80]. Mitotic cyclin ubiquitination can be reconstituted in vitro with apparent amphibian and fission yeast orthologs of either Ubc11 or Ubc4 [81, 82] and other data implicate both E2s in this process in higher cells [82, 83]. However, mitotic cyclins are efficiently degraded in budding yeast ubc4 and ubc11 strains, indicating that other E2 enzymes can support this essential function in S. cerevisiae [84]. 4.5 E2 Enzymes and Disease

There are now several striking examples of disease-related defects in ubiquitin conjugation, but most of them involve E3s rather than E2s. This is not surprising given the paramount role of E3s in substrate selection and the corresponding intensity of research effort that has been focused on E3s. Still, there are several hints that defects at the E2 level of the conjugation cascade can also contribute to disease. Many viruses subvert the ubiquitin system to evade the host cell’s defenses or modulate the cellular environment so as to promote viral replication (see Refs. [85, 86] and Section 7). The genome of African swine fever virus encodes an E2 enzyme that is somewhat similar to yeast Ubc3 [87, 88]. This enzyme might alter the activity or specificity of the host cell’s conjugation cascade so as to benefit the virus, or it could act on specific viral proteins. Herpes Simplex Virus-1 (HSV-1) encodes an E3 enzyme that specifically binds the host cell’s Ubc3/Cdc34 enzyme and targets this E2 for ubiquitination and (presumably) degradation – events that may help to stabilize specific cyclins and promote viral replication [89]. A different kind of relationship between an E2 enzyme and disease is exemplified by the finding that the Alzheimer’s amyloid-β peptide induces the expression of

6 The Biochemistry of E2 Enzymes

E225K , a mammalian relative of yeast Ubc1 [90]. E225K was found to play a major role in amyloid-β-dependent neuronal cell killing. This effect may be related to the E225K -dependent production of aberrant polyubiquitin chains, leading to the inhibition of proteasomes [90, 91]. Other studies showed that the human homolog of yeast Ubc11 is over-expressed in numerous cancer cell lines and primary tumors and that forced over-expression of this E2 in cultured cells can drive proliferation and transformation [92]. Similarly, transformation and chromosomal abnormalities were observed following over-expression of human Ubc2b [93]. Such disease-related over-expression effects could arise in two different ways. The higher E2 concentration could lead to a relaxation of specificity – that is, pairings with non-cognate E3s – leading to inappropriate ubiquitination events. Alternatively, specificity could be maintained, but an inappropriately high flux through the normal E2/E3 pathway could lead to the excessive ubiquitination of cognate substrates.

5 E2 Enzymes Dedicated to Ubiquitin-like Proteins (UbLs)

Ubiquitin is just one member of a family of protein modifiers that share a common fold and a common mechanism of isopeptide tagging [70, 94, 95]. Like ubiquitin, individual UbLs are activated at a C-terminal glycine residue by a specific E1 enzyme. Often, the next step is transfer to a specific E2 enzyme. Certain UbL-specific E2 enzymes are so similar to ubiquitin-conjugating enzymes that they were initially thought to be members of the ubiquitin-conjugating enzyme family. This was true of yeast Ubc9 and Ubc12, which are dedicated to Smt3/SUMO and Rub1/Nedd8, respectively (Table 1). SUMO modifies numerous cellular proteins and has a broad functional range [94], but the only known target of Nedd8 is a specific lysine residue in one subunit (the cullin) of SCF E3s. Nedd8 modification activates these E3s (see Ref. [70]). The reader should consult earlier reviews [70, 94, 95] for a detailed discussion of UbL biology and biochemistry. There are two important points for the current discussion. First, the conjugation cascades of UbLs differ from that of ubiquitin chiefly in terms of complexity – there is one conjugating enzyme per UbL, and many fewer E3s. Second, because modifier proteins (including ubiquitin) do not interact strongly with their dedicated E2s (Section 6.1), it is believed that E1 enzymes play the major role in matching E2s with the correct modifier protein (see Ref. [96]).

6 The Biochemistry of E2 Enzymes 6.1 E1 Interaction

An E2 needs to associate with several different proteins in the course of the ubiquitin conjugation cascade, with the first being the E1. Mutational studies conducted with

11


the SUMO-specific E2 Ubc9 suggest that free Ubc9 associates with its free cognate E1 through a surface of Ubc9 that includes the C-terminal residues of α-helix H1 and residues in a loop between β-strands S1 and S2 (Figure 3) [97]. This surface of Ubc9 is also important for thiol ester bond formation [97]. Several residues in α-helix H1, particularly the C-terminal residues, are poorly conserved among E2s, and Ubc9 contains a five-residue insertion in the loop between β-strands S1 and S2 (Figure 2). Thus, this region of E2s may contribute to specificity for their cognate E1s. Consistent with this idea, the N-terminal helix (H1) of a ubiquitin E2 was found to be important for E2/ubiquitin thiol ester formation [98]. One cautionary note is that Ubc9 displays a substantial affinity for its free E1 [97], whereas ubiquitin E2s bind tightly to their E1 only after it has been loaded with ubiquitin [6, 99, 100]. The structural basis of this effect remains to be determined. 6.2 Interactions with Thiol-linked Ubiquitin

As a consequence of interacting with ubiquitin-loaded E1, the E2 accepts the activated ubiquitin at its active-site cysteine residue. This thiol ester complex, although biochemically detectable [6], has not been crystallized because it is labile in comparison to the requirements of structural biology. However, NMR chemical shift perturbations have been used to map the binding surface of ubiquitin onto human Ubc2b [101], yeast Ubc1 [102], and human Ubc13 [103]. All three models map the ubiquitin-binding surface of the E2 to a common area that includes parts of α-helix H3, the loop between α-helices H3 and H4, and residues around the active-site cysteine in the extended S4-H2 loop, including part of the 310 helix H2 (Figures 2 and 4). The C-terminus of ubiquitin extends around part of the E2 and is constrained in a cleft [102, 103]. Although this contact surface is detectable in the thiol ester, free E2s display a negligible affinity for free ubiquitin [6]. Thus, the covalent E2/ubiquitin bond enables the formation of these non-covalent contacts. 6.3 E3 Interactions

After E2/ubiquitin thiol ester formation, the ubiquitin must be transferred to the substrate, which is sometimes another ubiquitin. An E3 is usually required for this reaction in vitro, and is always required in vivo. There are three known types of E3s: the RING domain, HECT domain, and U-box (UFD2 homology) families. RING and U-box E3s act as bridging factors for E2s and substrates, but HECT E3s use a different mechanism, adding an extra step to the pathway (Section 6.3.3). 6.3.1 RING E3/E2 Interactions The small RING domain coordinates two zinc ions in a cross-brace arrangement [104]. The domain is defined by the presence of eight zinc-binding groups (cysteines and histidines) with a conserved spacing, such that the distance between the two zincs is conserved at 14 Å [104]. Sequence conservation between RING domains is


Fig. 3 Ubc1/ubiquitin thiol ester complex model (1FXT). The surface of Ubc1 is shown with residues implicated in ubiquitin binding colored purple and the active-site cysteine colored yellow. Ubiquitin is colored green.

otherwise minimal. The RING-domain fold consists of a central α-helix and several small β-strands separated by loops with variable lengths [105]. RING E3s can be either single-subunit or multi-subunit enzymes. The crystal structure of UbcH7 complexed to a single-subunit RING E3, c-Cbl, shows that the RING domain is the main site of contact, although there are a few intermolecular hydrogen bonds to a non-RING helix of the E3 [106] (Figure 5). The structure of the E2 in the c-Cbl/UbcH7 complex is unchanged relative to free E2 structures. The main basis for the interaction is the packing of several hydrophobic residues of UbcH7, notably F63, into a shallow groove on the RING domain surface. These residues come from the S3–S4 and H2–H3 loops (Figure 3). The α-helix H1 of UbcH7 makes the hydrogen-bond contacts to the non-RING α-helix. Interestingly, even though the c-Cbl/UbcH7 structure undoubtedly shows conserved RING/E2 contacts, this complex is catalytically inactive (cited in Ref. [107]). Therefore additional E2/RING contacts may be needed for catalytic competence. The surface of UbcH7 that contacts c-Cbl does not overlap with the E2 surface that contacts ubiquitin (Figures. 4 and 5; see also Ref. [108]), confirming that the

13


Fig. 4 UbcH7/c-Cbl complex (1FBV). The surface of UbcH7 is shown with residues interacting with the c-Cbl RING domain shown in red and the active-site cysteine shown in yellow. c-Cbl is colored green.

E2/ubiquitin thiol ester can associate with a RING E3. The E2 surface that contacts cCbl does, however, overlap the E2 surface implicated in E1/E2 interactions (Section 6.1). Thus, the E1 may have to depart from the E2/ubiquitin complex before E2/E3 interactions can take place. The closest approach of a RING-domain residue to the active-site cysteine of UbcH7 is about 15 Å, arguing against a role for RING E3s in chemical catalysis [106]. Instead, RING E3s have been proposed to facilitate ubiquitination by inducing physical proximity of the E2/ubiquitin thiol ester and the substrate [23, 30, 106, 109]. Catalysis would result from the increased local concentrations of the two reactants (discussed further below). RING/E2 interactions have also been studied with BRCA1. This E3 is unique in that it must heterodimerize with a second RING-domain protein, BARD1, in order to display maximal E3 activity [110]. Even though the heterodimer interface leaves both RING domains available for interaction [110], UbcH5c binds exclusively to the BRCA1 RING [107]. The interacting surface of UbcH5c is homologous to the surface of UbcH7 that contacts the c-Cbl RING domain in that several residues of UbcH5c pack into a cleft on the BRCA1 RING domain. But UbcH5c also makes several


contacts with the C-terminus of the BRCA1 RING domain and with a non-RING region of the heterodimer [107]. These extra contacts are not observed when UbcH7 binds to the BRCA1/BARD1 complex [107]. Because the BRCA1/BARD1/UbcH7 complex is inactive, the extra contacts observed with UbcH5c may help to create a competent E2/E3 complex. Despite the greater complexity of multi-subunit RING E3s, a common theme is evident – all SCF E3s, as well as several other types of cullin-based E3s, utilize a common RING-domain subunit, the small protein Rbx1 (reviewed in Refs. [30, 111]). Four subunits compose the minimal SCF E3 ligase complex: a cullin scaffold, Rbx1, an adaptor protein (Skp1), and a substrate-binding subunit that connects to the adaptor through a conserved domain called the F-box (see Ref. [30]). The cullin acts as a scaffold, with Rbx1 binding to one end to form a cullin/Rbx1 subcomplex that recruits the E2 and, in many cases, displays a substrate-independent ubiquitinligase activity (see Refs. [23, 30]). The crystal structure of the mammalian SCFSkp2 (Skp1/Cul1/F-boxSkp2 /Rbx1) E3 ligase shows a remarkably rigid, elongated complex [112]. The Cul1 scaffold contains three cullin-repeat motifs that span 110Å, with Rbx1 binding to a discrete C-terminal α/β domain. The Skp1/F-boxSkp2 complex binds to the opposite (Nterminal end) of the cullin. Rbx1 displays a hydrophobic groove, as seen previously in the c-Cbl RING domain [106, 112]. In c-Cbl, this groove provides an interaction surface for UbcH7 and it is reasonable to assume a similar mode of interaction in the case of Rbx1. Interestingly, the site where Nedd8 modifies the cullin is close to where the E2 binds, consistent with data which suggest that neddylation modulates E2 binding or activity [113, 114]. A model of the full SCF/E2 complex [112] shows that the end of Skp2 which binds the substrate is pointed toward the Rbx1-bound E2, with a 50-Å gap between the two. Models based on two other SCF structures show similar distances between the F-box protein and the E2 [109, 115]. Whether this gap can be bridged by the bound substrate is currently unclear. It has been suggested that the E2 may bind to Rbx1 somewhat differently than UbcH7 is observed to bind in the c-Cbl RING/UbcH7 complex, but it is not obvious that this can lead to a 20 Å movement of the E2 toward the bound substrate as suggested [109]. One could also imagine that the bound substrate and E2 “meet” through conformational changes of the SCF complex. However, the rigid separation produced by the Cul1 scaffold seems to be important for activity – introducing a flexible linker into the center of Cul1 produced a protein that could still bind an E2, but did not catalyze substrate ubiquitination [112]. An interesting study established a positive correlation between the rate of dissociation of the Ubc3/ubiquitin intermediate from the RING domain and the rate of Sic1 ubiquitination catalyzed by SCFCdc4 [116]. The authors proposed that the role of the RING domain is to bring the charged E2 into the vicinity of the SCF-bound substrate, but that release of the charged E2 is important to bridge the gap and enable multiple substrate lysines to be targeted. However, although these mechanisms may place the substrate’s lysine residue in the general vicinity of the E2’s active site, it is unclear that they can establish an effective orientation of the lysine residue and the thiol ester bond. Since

15


there is no known consensus site for ubiquitination [23, 117, 118], it is unlikely that specific molecular contacts in the vicinity of the substrate’s lysine residue are used to position this attacking group. Overall, it remains unclear how the substrate’s lysine residue approachs the E2 active site. 6.3.2 U-box E3/E2 Interactions The U-box family of E3s bind E2s through the small U-box domain [119]. Some U-box E3s do not seem to have their own cognate substrates, but instead promote polyubiquitination of the substrates of other E3s [120]. Other U-box E3s have defined cognate substrates and behave in a canonical manner [121, 122]. An NMR structure [123] confirmed an earlier prediction [124] that the Ubox domain has a RING-domain-like fold. Remarkably, the U-box domain uses hydrogen-bonding networks in place of zinc coordination to support the characteristic cross-brace arrangement. These interactions stabilize a globular fold consisting of a central α-helix surrounded by several β-strands, which are separated by loops of variable length [123]. There is a shallow groove in the surface located in a position homologous to the E2-interacting surface of RING domains. Mutational studies have linked E3 ligase activity to some of the residues in the surface groove [123, 125]. Since these mutations do not disrupt the U-box fold, they are likely to abrogate E2 binding. Although it is likely that E2s bind similarly to the U-box and RING domains, no E2/U-box structure has been reported so far. 6.3.3 HECT E3/E2 Interactions HECT-domain E3s are defined by the presence of a domain of 350 residues that is homologous to the C-terminus of the founding family member, E6AP (E6 Associated Protein [126]). E6AP is known for its role in binding the E6 protein of oncogenic human papilloma viruses and targeting the p53 tumor suppressor for ubiquitination and degradation [127]. HECT-domain E3s possess an active-site cysteine residue positioned 35 residues upstream of the C-terminus; a thiol ester with ubiquitin is formed at this site and is required for substrate ubiquitination [128]. The crystal structure of an E6AP/UbcH7 complex showed that the HECT domain is L-shaped, with a large, mostly α-helical, N-terminal lobe and a small C-terminal lobe with an α/β structure [108] (Figure 6). UbcH7 binds to the end of the Nterminal lobe and somewhat parallel to the C-terminal lobe, forming an overall U-shaped complex. UbcH7 binds in a large hydrophobic groove in the N-terminal lobe [108]. As seen in other E2/E3 structures, neither the E2 enzyme nor the HECT domain changes its overall fold upon binding. UbcH7 contacts its binding groove with residues from the S3–S4 loop and the H2–H3 loop (Figure 3). A few contacts are also made with the C-terminal portion of α-helix H1. Remarkably, these are the same two loops and helix that bind the RING domain in the c-Cbl/UbcH7 structure [106]. That two different E3s contact a largely similar surface on UbcH7 (Figures 5 and 6) can be explained through the nature of key side-chain contacts. The S3–S4 loop seems to provide most of the specificity, as it contains the F63 residue that is present in all E2s that are known to bind both


Fig. 5 UbcH7/E6AP (1C4Z). The surface of UbcH7 is shown with residues interacting with the E6AP HECT domain shown in blue and the active-site cysteine shown in yellow. E6AP is colored green with the active-site cysteine between the N- and C-lobe shown in yellow.

HECT E3s and c-Cbl [106, 108]. In c-Cbl, the main contacts for F63 are isoleucine and tryptophan residues located in the RING groove [106]. Both F63 and its contact site in c-Cbl are seen to vary in other E2/RING E3 pairs, suggesting that interactions between these three specific residues are needed, but that the nature of the contact can vary [106]. In other words, a different E2 could bind to a different E3 with a similar geometry, but through different types of side-chain contacts. UbcH7-F63 makes specific contacts with six E6AP residues, so this interaction is likely to be important for all HECT/E2 pairs [108]. The H2-H3 loop that makes the other major contacts with the HECT domain is part of the more variable E2 surface (Section 3). The specific contacts made between the H2-H3 loop of UbcH7 and the E6AP HECT domain could be used to correctly predict the E2 preferences of E6AP and Rsp5 [108]. Thus, residues in the S3-S4 and H2-H3 loops play an important role in determining the specificity of E2/E3 interactions. Positioned near the bend between the two lobes of the HECT domain is its activesite cysteine [108]. This residue is 41 Å away from the active-site cysteine of UbcH7 (Figure 6), suggesting that a large conformational change is needed to bring about

17


transfer of ubiquitin from E2 to E3. Such a mechanism was confirmed in the crystal structure of another HECT domain [129]. The WWP1-HECT domain resembles the E6AP HECT domain in having two lobes, but their relative positions differ. In the WWP1-HECT domain the two lobes form an inverted T-shape as opposed to the Lshape seen in the E6AP HECT structure [108, 129]. This conformational change can be brought about by a rotation around three residues in a hinge loop connecting the two lobes. Modeling in the E2 in a position homologous to that seen in the E6AP/UbcH7 structure, the distance between the two active-site cysteines decreases to 16 Å [129]. With additional rotation around the hinge loop the WWP1 activesite cysteine can be brought within 5 Å of the E2’s active-site cysteine. Mutational studies suggested that the flexibility of this hinge loop is indeed important for ligase activity [129]. This flexibility requirement points to possible models for ubiquitin transfer and polyubiquitin chain elongation. One possibility is that the HECT domain adds ubiquitin to target substrates one at a time. This would imply that the E3 changes specificity – from recognizing the substrate to recognizing the ubiquitin – following transfer of the first ubiquitin to the substrate. A different possibility is that the chain is built up by the HECT E3 first, and then transferred as a unit to the substrate. This would require two active sites, one to hold the growing polyubiquitin chain, and the other to hold the next ubiquitin to be added. HECT E3/E2 complexes would satisfy this condition. RING E3/E2 complexes cannot, and thus would have to utilize another mechanism, presumably building on their rigid architecture. In an attractive model [129], the HECT cysteine could hold the first ubiquitin (and later the growing chain), while the C-lobe could rotate to position the first ubiquitin’s target lysine near the thiol ester bond of the bound E2/ubiquitin intermediate. Subsequent rounds of ubiquitin addition to the chain terminus would require the C-lobe to keep rotating, ultimately ending in steric problems for the chain which could favor its transfer to the substrate. A third general model for polyubiquitin chain extension is that the initiation and elongation phases of the reaction involve different E2 enzymes, different E3 enzymes, or different E2/E3 complexes. The modification of a substrate with a noncanonical polyubiquitin chain follows the third model. The Rad6/Rad18 complex ligates the first ubiquitin, while the Mms2/Ubc13/Rad5 complex extends the chain [65, 73]. The extension of K48-linked chains from ubiquitin fusion proteins seems to involve the sequential action of two different E3s with the same E2 [120]. In another possible example, two E2s (orthologs of Ubc11 and Ubc4) have been suggested to act sequentially with the APC in the polyubiquitination of mitotic cyclins in fission yeast [82]. 6.4 E2/Substrate Interactions

With HECT domain E3s, all of the chemistry of isopeptide bond formation occurs at the E3 active site. With RING and U-box E3s, however, the E2 participates directly in this chemical reaction, so the substrate’s lysine must closely approach


the E2’s active site. A crystal structure of the SUMO E2 Ubc9, complexed with a large fragment of RanGAP1 (an efficient sumoylation substrate), reveals the specificity of this interaction [130]. Unlike ubiquitination, sumoylation is site-specific. The target lysine for sumoylation lies within a tetrapeptide sequence motif -K-X-D/E, where is a hydrophobic residue, K is the target lysine, and X is any residue. Ubc9 makes specific interactions with each of these consensus-motif residues in a manner that places the lysine ε-amino group within 3.5 Å of the Ubc9 active-site cysteine [130]. The lysine approaches the cysteine from what is expected to be its unencumbered (by SUMO) side [102]. The interacting surface on Ubc9 involves α-helix H4, the loop preceding it, and the extended S4-H2 loop, including the active-site cysteine [130] (Figure 3). This surface does not overlap with the presumptive binding surface for SUMO. This mode of interaction is unlikely to hold with ubiquitin E2s, since no general consensus site for ubiquitination is known. A model for ubiquitin E2/substrate interactions has also been proposed for the special case in which the substrate is ubiquitin [131]. The crystal structure of the Mms2/Ubc13 complex led to the modeling of an E2/UEV/ubiquitin (donor)/ubiquitin (acceptor) model. As discussed in Section 7, UEV (Ubiquitin E2 Variant) proteins such as Mms2 are homologous to E2s, but lack the active-site cysteine residue. Known UEV/Ubc13 complexes act as E2 enzymes specialized for the synthesis of K63-linked polyubiquitin chains [72, 132]. In the model [131], Ubc13 is bound to the donor ubiquitin through a thiol ester bond in a manner that agrees well with inferences from NMR analysis of the Ubc1/ubiquitin thiol ester [102] (Figure 7). The position of the non-covalently bound acceptor ubiquitin is determined by the orientation of Mms2 on Ubc13 (Figure 7). The acceptor ubiquitin has its K63 side chain placed to enter the active site of Ubc13 to form a diubiquitin conjugate. The model suggests that K63 of ubiquitin is selected as the conjugation site through steric exclusion of other lysines, as determined by an interaction between Mms2 and a region of ubiquitin that is distant from K63 [131]. Recent NMR studies have confirmed and refined this model [103]. Thus, the substrate lysine is presented to the active-site cysteine through an indirect mechanism, in contrast to the Ubc9/RanGAP1 example in which the E2 interacts directly with the lysine residue itself [130]. Unlike most ubiquitination reactions, the modification of ubiquitin itself is often site-specific. The Mms2/Ubc13/ubiquitin model can help to explain this phenomenon. 6.5 E2 Catalysis Mechanism

Chemical catalytic mechanisms in the ubiquitin conjugation cascade have proved difficult to decipher. The reactions leading to E2/ubiquitin thiol ester and isopeptide bond formation would be facilitated by electrostatic stabilization of the oxyanion and deprotonation of the attacking amino group (isopeptide bond formation) by a general base [133, 134]. However, while the sequence conservation around the E2 active site is very high (Figure 2), all E2 structures show a lack of candidate catalytic residues close to the cysteine (see Refs. [23, 135]). Although catalytic residues could

19


Fig. 6 Ubc13 interaction surface. The interacting surfaces have been mapped onto Ubc13. The active-site cysteine is shown in yellow. Colored surfaces contact: covalently bound ubiquitin (purple); RING domains (red); E1 (presumptive, green); acceptor ubiquitin involved in K63-linked polyubiqutin-chain synthesis (blue).

be contributed by other enzymes in the cascade or by the E2 backbone, structural data argue strongly against a chemical catalytic contribution where it may be needed most – in reactions involving RING and U-box domain E3s. Recent studies [136] addressed the role of a strictly conserved asparagine positioned just upstream of the active-site cysteine (N79 in Ubc1 numbering, Figure 2). In existing E2 structures the asparagine is hydrogen-bonded to the backbone or a side chain. It is distant from the E3 contact surface and, as expected, it is dispensable for E2 binding to RING domain E3s. However, the asparagine is critical for E2-catalyzed and RING E3/E2-catalyzed ubiquitin conjugation reactions. The similar effect of asparagine mutation on the two types of conjugation reactions is reasonable given that E2s do not experience structural perturbations upon binding to E3s (above). The data suggest that an intrinsic catalytic role of the asparagine side chain is brought into play through RING-mediated recruitment of the catalytically competent E2/ubiquitin thiol ester. The asparagine is dispensable for upstream and downstream thiol transfer reactions, suggesting that catalytic residues for these reactions may be located in the E1 and HECT E3 active sites. A specific proposal for the role of the asparagine was developed in a model which breaks the hydrogen bonds to the backbone and rotates the asparagine toward the active-site cysteine [136]. Molecular modeling suggested that the asparagine can be positioned to donate a hydrogen bond to the oxyanion (Figure 8) [136]. Many

7 Functional Diversification of the E2 Fold 21

Fig. 7 Model for catalytic role of E2 activesite asparagine. The side chain of the asparagine in the conserved “HPN” motif (Figure 2) stabilizes the oxyanion that

forms when the substrate’s lysine attacks the E2/ubiquitin thiol ester bond. N79 is numbering for Ubc1 (Figure 2).

cysteine proteases, including deubiquitinating enzymes, use an amide side chain in this manner [134, 137–139]. Structural studies of a deubiquitinating enzyme have shown that the entry of ubiquitin into the active site causes a histidine and an asparagine to shift their positions so that the histidine becomes the general base and the asparagine provides the oxyanion hole [137]. Similarly, ubiquitin binding in the E2 active site could be a trigger that repositions the asparagine. So far, no general base is evident in E2s, but this group may not be needed due to the lability of the thiol ester bond.

7 Functional Diversification of the E2 Fold

Increasing evidence suggests that evolution has used (and is using) the E2 fold for new purposes. In one apparent example of functional expansion, E2 core domains have been observed to be embedded within much larger polypeptide chains [140, 141]. The functional properties of these massive E2s remain poorly characterized, and it is likely that more of them will be discovered. But the clearest case of functional diversification is provided by the UEV proteins. UEVs are related to E2s in their primary, secondary, and tertiary structures, but they lack an active-site cysteine residue and therefore cannot function as canonical E2s [142]. Nonetheless they play several different roles in ubiquitin-dependent pathways. Mms2 and its close (mammalian) relative Uev1a form heterodimers with Ubc13 [72, 132]. Each complex plays a key role in the synthesis of K63-linked chains, which act as non-proteolytic signals in different cellular pathways. The Mms2/Ubc13 complex participates in the UBC2/RAD6-dependent DNA damage tolerance pathway by polyubiquitinating the DNA polymerase processivity factor called PCNA (Proliferating Cell Nuclear Antigen) [65, 72]. To be activated for this pathway, PCNA is first mono-ubiquitinated by the Rad6/Rad18 complex, and then modified


with a K63-linked polyubiquitin chain by the Mms2/Ubc13/Rad5 complex (Rad18 and Rad5 are RING E3s) [65, 73]. The Mms2/Ubc13 complex has a core ubiquitin polymerization activity [72]. Rad5 might stimulate this activity [132] or target the Uev/E2 complex to PCNA [73], or both. The related human Uev1a/Ubc13 complex is involved in NFκB signal transduction [132]. It plays an intermediate role in the signaling cascade that starts with a proinflammatory cytokine signal and culminates in the nuclear translocation of the active NFκB transcription factor. In this pathway the Ubc13/Uev1a complex modifies a RING E3, Traf6, with K63-linked polyubiquitin chains [132]. This modification is linked to Traf6 oligomerization. It instigates a cascade of kinase reactions ultimately cause the ubiquitination and degradation of NFκB’s inhibitory partner, IκBα [74, 143]. The crystal structure of Mms2 has been solved alone and in complex with Ubc13 [131, 144]. The overall fold is similar to that in E2s, containing a central fourstranded anti-parallel β-sheet surrounded by α-helices. Differences include the absence of the C-terminal α-helix H5 in the shorter Mms2 protein. The helical Nterminus of Mms2 is also extended compared to Ubc13, and this region plays the major role in heterodimer formation. Two ubiquitins can bind to the heterodimer (Section 6.4) and the surfaces they contact do not overlap with the surface contacted by Rad5 [131, 145]. Ubiquitin plays a crucial role in a protein-trafficking pathway that delivers specific cargo proteins to regions of the late endosome membrane that invaginate into the lumen, thereby targeting these proteins to the vacuole/lysosome (reviewed in Ref. [146]). A different UEV protein, called Tsg101 in humans (Tumor Susceptibility Gene) and Vps23 in yeast (Vacuolar Protein Sorting), is part of a large complex that plays a critical role in the sorting step. Cargo proteins are selected based on their conjugation to mono-ubiquitin; the specific role of the UEV protein is to bind the cargo-linked mono-ubiquitin moiety [147]. HIV-1 and certain other viruses subvert this function of Tsg101 in order to bud from the plasma membrane [148– 150]. Mechanistically, Tsg101 is recruited to the virus budding sites by binding to a tetrapeptide “PTAP” motif in the late domain of viral proteins such as HIV1-GAG. Tsg101 is essential for virus budding from the plasma membrane [148], so it is possible that the endocytic budding machinery is hijacked to the plasma membrane via the Tsg101/GAG interaction [85]. The solution structure of the Tsg101 UEV domain has been solved alone and in complex with a PTAP-containing peptide [151, 152]. Human Tsg101 contains 390 amino acids, with the UEV domain located at the N-terminus [152]. The UEV domain is the minimal region needed to bind HIV-1 GAG, and is also the domain involved in mono-ubiquitin recognition and binding [153]. The overall fold of the UEV domain is similar to that of E2s. One notable difference is the presence of an extra N-terminal α-helix on Tsg101 [152]. The other major difference is the absence in Tsg101 UEV of the two C-terminal α-helices of the E2s – a truncation that was also seen in the Mms2 structures. This truncation appears to be a special trait of UEV proteins [154]. In Tsg101 UEV, the absence of the C-terminal helices helps to create the binding site for the PTAP peptide [151].

8 Conclusions

When aligning the structures of a canonical E2, Mms2, and the Tsg101 UEV domain, the hydrophobic core and the region surrounding the vestigial active site are quite similar, but the Tsg101 UEV domain differs from Mms2 and canonical E2s in the positions of the first two β-strands [152]. In Tsg101 they are elongated and shifted toward the N-terminus, forming a β-hairpin that extends 11 residues outside the main body of the domain [152]. This loop is important for ubiquitin binding by Tsg101 [152]. As determined by chemical shift mapping and mutagenesis studies, the Tsg101/ubiquitin binding interface involves the bottom half of the four-stranded β-sheet, including the β-hairpin (loop S1–S2). The binding interface for ubiquitin on Tsg101 is distinct from the surface that Mms2 uses to position the acceptor ubiquitin within the Ubc13/Mms2 complex [131, 144]. Thus, two different UEVs bind ubiquitin in two different ways. The structural biology and biochemistry of UEVs illustrates how modest changes to an E2-like module can create new, functionally important interaction sites. The UEV domain is just one of a growing set of small domains that can endow other protein domains with ubiquitin-binding capability (reviewed in Ref. [63]). Such binding elements are likely to play important roles in transducing ubiquitin signals in diverse cellular pathways.

8 Conclusions

We have emphasized the biochemical properties of E2s, particularly interactions with other factors in the conjugation cascade, because these properties are central to the biological actions of E2s. We have tried to give a flavor of the “creativity and economy” [103] with which E2s have evolved to maximize the interaction potential of a relatively small and conserved surface (Figure 7). Owing to the large scope of the relevant literature and the limited length of this chapter, we have not done full justice to the biological breadth of the E2 enzyme family. For example, we have focused on yeast and mammalian enzymes, but ubiquitin conjugation is increasingly being studied in other model organisms, including flies, worms, and plants. These systems offer powerful tools to address outstanding questions about ubiquitin-dependent pathways in general and E2 enzymes in particular. What are some of those questions? Significant uncertainties remain concerning E2 catalysis and mechanism, as discussed in Section 6. Another important question has been largely ignored in this review – exactly why are there so many E2s? One appealing model is that the identity of the E2 can modulate the substrate specificity of the E3, but experimental evidence for this model remains sparse. Another possibility is that the E2 has little or no influence on substrate choice, but rather helps to control the flux of activated ubiquitin to its cognate E3. In view of the remarkable developments in ubiquitin biology over the last decade, we should be prepared for both interesting and unexpected answers to these (and other) questions.

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1

The Deubiquitinating Enzymes

Nathaniel S. Russell and Keith D. Wilkinson Emory University, Atlanta, USA


1 Introduction

In the mid-1970s ubiquitin was found to be a covalent modifier of proteins [1]. At the time, it was quite surprising to find a protein that covalently modified another protein. Since then, the reversible covalent modification of proteins by other proteins is known to be commonplace and ubiquitin is used to covalently modify hundreds of proteins, often for the purpose of targeting them to the proteasome for degradation. Protein degradation through the ubiquitin–proteasome system is facilitated by covalently linking ubiquitin to the ε-amino group of a lysine of a substrate protein through its C-terminal glycine [2]. A polyubiquitin (polyUb) chain is formed by linking subsequent ubiquitins to the lysine 48 residue of the preceding ubiquitin in the chain. A chain of four ubiquitins is sufficient for the targeted protein to be recognized and degraded by the proteasome [3]. Conjugation of ubiquitin to other proteins is catalyzed by a three-enzyme cascade [4]. Conjugation begins by activation of ubiquitin by an E1, or Ub-activating enzyme, forming a high-energy thiol ester bond in an ATP-dependent reaction. The ubiquitin is transferred to an E2, or Ub-conjugating enzyme, which then usually pairs with an E3, or Ub-ligase enzyme, to conjugate the ubiquitin to a specific target protein. The usefulness of ubiquitin conjugation is not limited to the ubiquitin– proteasome pathway. Mono- and polyubiquitin are used as signals in various pathways including endocytosis, DNA repair, apoptosis, and transcriptional regulation [5–8]. Polyubiquitin chains can be formed using lysine residues other than K48, the linkage required for proteasomal degradation [9–11]. In addition, there are a number of other ubiquitin-like proteins that also behave as signaling molecules although they are not involved directly in proteasomal degradation. This group Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 The Deubiquitinating Enzymes

includes SUMO (small ubiquitin-related modifier), Nedd8 (neural precursor cell expressed, developmentally down-regulated 8), ISG15 (interferon-stimulated gene 15), and others [12–14]. These proteins are conjugated to substrates in a similar manner to ubiquitin, using an E1, E2, and E3 cascade of enzymes specific for the particular ubiquitin-like protein involved [15–17]. Soon after it was shown that ubiquitin is conjugated to proteins, it was determined that this was a reversible process and deubiquitinating enzymes, or DUBs, could remove ubiquitin from ubiquitinated proteins [18, 19]. As the genes for ubiquitin and ubiquitin-like proteins were identified it became clear that all ubiquitin family members were synthesized as proproteins and processed to reveal the C-terminal glycylglycine of the active proteins [20]. Based on this information, DUBs were defined as proteases that cleave at the C-terminus of ubiquitin or ubiquitin-like proteins to reverse conjugation to target proteins and also process the proproteins. Over 100 DUBs have been identified (see Table 1 for a list of the DUBs whose roles are known or suspected) and they are used to regulate ubiquitin and ubiquitin-like protein metabolism. Since cells utilize a combination of mono-ubiquitin, polyubiquitin, and ubiquitin-like proteins for a multitude of reasons and conjugate them to thousands of proteins, a regulatory system has evolved that is exceedingly complex and must be exquisitely regulated (see Figure 1). DUBs help regulate this system by processing proubiquitin into a mature form, recycling free polyubiquitin chains into monomeric Ub, assisting the degradation of proteasomal substrates, and regulating the ubiquitination levels of proteins in cellular pathways other than proteolysis. Thus, DUBs play crucial roles in determining the cellular fates of many proteins and regulating cellular function.

Fig. 1 DUB families and substrate specificity. DUBs can be classified by genetic relationships (family) or by substrate specificities (activity). Arrows point towards the substrates that members of each family can

process in a physiologically relevant way. Each family is capable of processing multiple substrates and each activity can be catalyzed by members of more than one family.

1 Introduction 3 Table 1 Physiological roles of DUBs revealed by deletion or knockdown experiments

DUB

Organism

Deletion/knockdown phenotype

Ubiquitin C-terminal hydrolases (UCHs) UCH-L1 mouse gracile axonal dystrophy UCH-L3

mouse

no detectable phenotype

UCH37

human

unknown

BAP1 YUH1

human S. cerevisiae

unknown cannot process proRUB1

Functional role

predominant neuronal UCH [112] undetermined neuronal function [113] edits polyubiquitin chains at proteasome [91] tumor suppressor? [22] processes proRUB1, Ub-adducts [83]

Ubiquitin specific processing proteases (UBPs) UBP1 S. cerevisiae no phenotype detected undetermined [97] UBP2 S. cerevisiae no phenotype detected undetermined [96] UBP3/ S. cerevisiae, polyubiquitin accumulation transcriptional silencing inhibitor, USP10 human regulates membrane transport [66, 118] UBP6 S. cerevisiae low levels of free ubiquitin processes polyubiquitin chains at proteasome [94] UBP8 S. cerevisiae increase in ubiquitinated transcriptional regulator [88] histone H2B UBP14/IsoT S. cerevisiae, increased polyubiquitin recycles free polyubiquitin to human levels, proteasome defects mono-ubiquitin [33, 86] UBP16 S. cerevisiae no phenotype detected undetermined function at mitochondria [70] DOA4 S. cerevisiae Ub-depletion, defective recycles Ub and polyubiquitin Ub recycling adducts [71, 84] Unp/USP4 mouse, human unknown undetermined USP7 human indirect p53 activation regulates p53 ubiquitination [36] UBPy/USP8 human increase in protein cell-growth regulation [119] ubiquitination USP14 mouse (UBP6 ataxia regulating synaptic activity plus homolog) proteasome [109] USP21 human unknown process Ub and Nedd8 conjugates, growth regulator? [32] USP25 human unknown over-expression has possible role in Down’s Syndrome [120] UBP41 human unknown promotes apoptosis [6] UBP43 mouse accumulation of Isg15 processes Isg15, regulates Isg15 conjugates conjugate levels [31] CYLD human cylindromatosis regulates K63 polyubiquitination of substrates in NF-κB pathway [7, 103, 104] fat facets Drosophila defective germ cell regulates specific developmental specification, eye formation processes [74, 75] DUB1, 2, 2A mouse unknown cytokine specific growth regulators VDU1 human unknown regulation of Ub-proteasome pathway? [64]

4 The Deubiquitinating Enzymes Table 1 (Continued)

DUB

Organism

Deletion/knockdown phenotype

Functional role

JAMM Isopeptidases Rpn11 Yeast, human Csn5 Yeast, Drosophila

processes polyubiquitin chains at proteasome [46, 47] defects in SCF E3s in yeast, regulation of cullin neddylation lethal in Drosophila levels [45]

OTU DUBs cezanne

human

unknown

A20

mouse

otubain1 otubain2 VCIP 135

human human human

severe inflammation, premature death unknown unknown unknown

lethal

Ubiquitin-like proteases (ULPs) Ulp1 S. cerevisiae lethal Ulp2

S. cerevisiae

Den1/SENP8 mouse SENP6 human

increased SUMO conjugates, DNA repair defective unknown unknown

Others Apg4B

mouse

unknown

ataxin-3

human

unknown

negative regulation of NF-κB pathway [43] negative regulation of NF-κB pathway [44, 121] editing DUB? [42, 122] undetermined [42] membrane fusion after mitosis [72] regulates cell cycle progression [39] desumoylating enzyme [123]

deneddylates cullins [26, 37] involved in reproductive function? [124] processes autophagy-related UbLs [116] processes polyubiquitin chains? [125]

The organism listed for each DUB refers either to where it was discovered or where the work characterizing the deletion strain and function was performed. DUBs with multiple organism identifiers are either highly similar in sequence in each organism or functional homologs. An unknown deletion phenotype indicates that a deletion, knockout, or knockdown of a particular DUB has yet to be generated. No detectable phenotype indicates that a deletion strain has been made, but no phenotypes were observed.

The study of DUBs has moved at a rapid rate since their initial discovery in the early 1980s. Yet despite all the progress, the total number of DUBs and the substrate specificity of most DUBs are still undetermined. The discovery of novel DUB families including the JAMM isopeptidases and OTU DUBs has highlighted that there may be still more unidentified DUBs. Because of the large number of potential DUB substrates and the exquisite specificity that some individual DUBs exhibit (see Figure 1) study of these enzymes has often been challenging. A burgeoning amount of structural data and recent technical advances are being used to address this challenge. The goal of this chapter is to highlight recent developments in the

2 Structure and In Vitro Specificity of DUB Families 5

DUB field by giving an overview of DUB families, including DUBs that act on ubiquitin-like proteins, to discuss how DUBs achieve their specificity, and to show how the physiological roles of DUBs and their substrates are being elucidated.

2 Structure and In Vitro Specificity of DUB Families 2.1 Ubiquitin C-terminal Hydrolases (UCH)

The first class of DUBs discovered, the ubiquitin C-terminal hydrolases (UCHs), is a relatively small class with only four members in humans and one in budding yeast. UCHs are cysteine proteases related to the papain family of cysteine proteases. Most UCHs consist entirely of a catalytic core that has a molecular mass of about 25 kDa, although Bap1 and UCH37 have C-terminal extensions [21, 22]. All UCHs have a highly conserved catalytic triad consisting of the active-site cysteine, histidine, and aspartate residues that are absolutely required for function [23]. In vitro studies have determined that UCHs have significant activity in removing small adducts from the C-terminus of ubiquitin, including short peptides, ethyl ester groups, and amides [24]. They are also very efficient at cotranslationally processing the primary gene products (proubiquitin or Ub-ribosomal subunit fusions) to expose the C-terminal gly–gly motif required for conjugation of ubiquitin and ubiquitin-like proteins to substrates. However, UCHs are unable to cleave the isopeptide bond between ubiquitins in a polyubiquitin chain or to act on ubiquitin conjugated to a folded protein. They are similarly inefficient in acting upon small peptide substrates based on the sequence of the ubiquitin Cterminus [25]. As described below, the binding of ubiquitin is required for optimal UCH activity. Nedd8, a closely related ubiquitin-like protein, is also a substrate for human UCH-L3, albeit with three orders of magnitude less efficiency than ubiquitin [26]. The crystal structures of human and yeast UCHs have been solved, the latter in complex with the inhibitor ubiquitin aldehyde [27, 28]. The UCH fold is closely related to that of the papain family of cysteine proteases. Ubiquitin is bound in a cleft on a surface that is highly conserved in all UCHs. NMR studies on the binding of ubiquitin to human UCH-L3 show a similar mode of interaction and define three regions on the surface of ubiquitin involved in this binding [29]. As noted below and in Figure 2, the same surface of the ubiquitin fold is also involved in binding to USP7 and ULP1. This is remarkable as these DUBs are from different families and not significantly homologous in sequence or structure. A second feature of UCHs is the presence of a “blocking loop” spanning the active site and limiting the size of substrates that can be accommodated. Yuh1 and other UCH DUBs contain a mobile, ∼20-residue loop that is disordered in the unliganded protein, but becomes ordered upon substrate binding. The loop passes directly over the active site and the leaving group attached to the gly–gly at


Fig. 2 Substrate binding by DUBs revealed by X-ray crystal structures. In the ribbon diagrams, the DUB is represented in white and the substrate in color. The ubiquitin (yellow or green) or SUMO (red) substrates are shown in the same orientation to highlight the similarity of substrate binding by different DUB classes. (A) Ubiqui-

tin (yellow) bound to YUH1. (B) Ubiquitin (green) bound to USP7. (C) SUMO (red) bound to ULP1. (D) Superimposition of substrates from A-C. The regions of each substrate that are within 3.5Å of the DUB when bound are highlighted in color to demonstrate the conserved regions that are recognized by the different DUB classes.

the ubiquitin C-terminus has to pass directly through this loop in order to access the catalytic cysteine. The maximum diameter of this loop was calculated to be ∼15Å, too small for any folded substrate save a single helix [27]. The loop thus allows small substrates to be efficiently cleaved, but excludes larger Ub–protein conjugates. This loop explains, at least in part, the preference of UCHs for small or disordered leaving groups.

2 Structure and In Vitro Specificity of DUB Families 7

2.2 Ubiquitin-specific Processing Proteases (UBP/USP)

The ubiquitin specific processing proteases (referred to as UBPs in yeast and USPs in human and mouse) were the second class of DUBs discovered. Catalytically, the UBPs are very similar to the UCHs in that they also utilize the catalytic triad of an active-site cysteine and a conserved histidine and aspartate. The UBP catalytic core of about 400 amino acids contains blocks of conserved sequences (Cys and His boxes) around these catalytic residues [23]. The UBPs are generally larger and more variable in size than the UCH class, ranging from 50 to 300 kDa. Nterminal extensions to the catalytic core account for most of the increased size although a few UBPs have C-terminal extensions. These N-terminal extensions are highly divergent in sequence, unlike the conserved regions of the catalytic core. The sequence and size variations of these extensions are thought to aid in determining UBP localization and substrate specificity. There are 16 UBPs in yeast and more than 50 USPs identified in humans, making the UBP/USP family much larger than the UCH family [30]. UBPs also process a wider variety of substrates than UCH DUBs, including proubiquitin, free polyubiquitin chains of various linkages, and mono-or polyubiquitin conjugated to target proteins in vitro and in vivo. In addition, some family members can act on ubiquitin-like proteins. UBP43 has been demonstrated to act on ISG15 while USP21 cleaves conjugated Nedd8 [31, 32]. The diversity of the UBPs and breadth of substrates they act upon, makes them useful in a wide variety of cellular pathways and locations. UBPs regulate apoptosis, DNA repair, endocytosis, and transcription in addition to the ubiquitin–proteasome pathway (see below). The same diversity presents a challenge in determining the specificity of UBPs and with the exception of Isopeptidase T (UBP14/USP5), there have been few quantitative studies of in vitro specificity [33, 34]. In general, specificity has been described with qualitative “yes or no” assays that are not particularly useful in suggesting in vivo roles. The structure of one UBP catalytic domain has been solved, that of USP7 complexed to ubiquitin aldehyde [35]. The data may be applicable to the way in which other UBPs function because the catalytic core of many UBPs is highly conserved. USP7 (also called HAUSP) is a human ubiquitin-specific protease that regulates the turnover of p53 [36]. USP7 consists of four structural domains; an N-terminal domain known to bind p53 and EBNA1, a catalytic domain, and two C-terminal domains. The 40-kDa catalytic domain exhibits a three-part architecture comprising Fingers, Palm, and Thumb (see Figure 2). The leaving ubiquitin moiety is specifically coordinated by the Fingers, with its C-terminus placed in a deep cleft between the Palm and Thumb where the catalytic residues are located. The domains form a pocket ideal for binding ubiquitin. Residues in the structure important for the above functions are conserved amongst UBPs, indicating that many UBPs may utilize the Fingers, Palm, and Thumb architecture to bind and cleave ubiquitinated substrates.


Another interesting structural observation is that water molecules cushion ubiquitin in the binding pocket. This is necessary because the binding surfaces of ubiquitin are uncharged, and the USP7 binding pocket is made up of predominantly acidic amino acid residues. These water molecules form extensive networks of hydrogen bonds with the bound ubiquitin and USP7. It is possible that they contribute to USP7’s substrate specificity by allowing the protein to provide for relatively weak binding of ubiquitin and forcing itself to interact with the target protein to achieve specificity. This seems to be borne out by the fact that ubiquitin does not form a tightly bound complex with USP7 [35]. 2.3 Ubiquitin-like Specific Proteases (ULP)

The ubiquitin-like specific proteases (ULPs) are a third class of DUB first thought to act only on SUMO-related ubiquitin-like proteins. There are two yeast ULPs and seven human ULPs (also called sentrin specific proteases, or SENPs). Further analysis determined that ULPs have little or no activity on ubiquitin substrates, but one (SENP8) acts on Nedd8 [26, 37, 38]. Despite acting on non-ubiquitin substrates, ULPs are still classified as DUBs because the function and mechanism of catalysis is so similar to those of the DUBs that act on ubiquitin. ULPs lack significant sequence homology to other DUBs and are more closely related to viral proteinprocessing proteases [39]. In addition to the lack of sequence homology, ULPs have little structural homology to other DUB classes except in the active site. The structure of ULP1 (see Figure 2) in complex with the C-terminal aldehyde of yeast SUMO (SMT3) illustrates that, like most other DUBs, ULPs are thiol proteases, utilizing a conserved catalytic triad consisting of an active-site cysteine, histidine, and aspartate [40]. Also, they require a gly–gly motif at the C-terminus of their UbL substrate for tight binding. The SUMO binding pocket of ULP1 recognizes SUMO through a number of polar and charged-residue interactions, including multiple salt bridges that are not present in the USP7 ubiquitin-binding site, and does not utilize water molecules or a “blocking loop”. 2.4 OTU DUBs

A class of DUBs only identified since 2002 is the OTU (ovarian tumor protein) DUB class. The OTU domain was originally identified in an ovarian tumor protein from Drosophila melanogaster, and over 100 proteins from organisms ranging from bacteria to humans are annotated as having an OTU domain. The members of this protein superfamily were annotated as cysteine proteases, but no specific function had been demonstrated for any of these proteins. The first hint of a role for OTU proteins in the ubiquitin pathway was afforded by the observation that an OTUdomain-containing protein, HSPC263, reacted with ubiquitin vinyl sulfone (an active-site-directed irreversible inhibitor of DUBs) [41].

3 DUB Specificity 9

Then two groups almost simultaneously discovered that several OTU-containing proteins have DUB activity. Two human OTU DUBs were identified by purification with Ub-aldehyde (a reversible DUB inhibitor) affinity resin [42]. These proteins, named otubain1 and 2 (OTU-domain Ub-aldehyde binding protein) have a mass of approximately 35 kDa and are able to cleave polyubiquitin chains in vitro. However, the cleavage mechanism and their true substrates in vivo have yet to be determined. The other OTU DUB found was Cezanne, a 100-kDa protein that is similar to the A20 negative regulator of NF-κB [43]. Like A20, Cezanne plays a role in regulating NF-κB signaling pathways and has general DUB activity [44]. These OTU DUBs have highly conserved catalytic cysteine and histidine residues, implying that they utilize a catalytic triad to catalyze cleavage of polyubiquitin. It is unclear if most proteins containing OTU domains are DUBs, as analysis of the OTU family for DUB activity is only just beginning. 2.5 JAMM Isopeptidases

JAMM isopeptidases also constitute a recently identified class of DUBs. The members of this interesting class of DUBs were the first non-cysteine protease DUBs identified. Two JAMM isopeptidases have been confirmed as DUBs: Rpn11, which acts on ubiquitin conjugates, and Csn5, which acts on Nedd8 conjugates [45–47]. A number of other eukaryotic proteins have been annotated as containing the JAMM motif, but whether they have DUB activity has yet to be determined. Instead of cysteine proteases, they are metalloproteases belonging to a family of proteins that contain the Jab1/Csn5 and MPN domains [48]. Their activity depends on the JAMM motif (EXn HS/THX7 SXXD) in the JAMM domain. The two histidines and an aspartic acid act as ligands to bind a metal ion, presumably zinc although this has not been proven, to achieve catalysis through polarization of a bound water molecule. A glutamic acid serves as a general acid-base catalyst. The crystal structure of a JAMM metalloprotease from Archaeoglobolus fulgidus bacteria has been recently been solved, but no structures of a JAMM isopeptidase with DUB activity are yet available [49, 50].

3 DUB Specificity

Why are there so many DUBs and how do they achieve specificity? The numerous DUBs identified to date suggest that DUBs have specifically evolved to act on distinct cellular substrates rather than to have general deubiquitinating activity (see Figure 1). We can ask what common features of these enzymes define them as DUBs and what differences allow specific DUBs to act on mono- vs. polyubiquitin? How have they evolved to cleave only ISG15 or SUMO-modified substrates, for instance? A body of data has been accumulated that at least partially answers these questions.


3.1 Recognition of the Ub-like Domain

All DUBs appear to recognize the body of the ubiquitin fold. UCH-L3, for example, makes contact with three regions of ubiquitin; residues 6–12, 41–48, and 69–74 [29]. These surfaces are highly conserved in Nedd8, but divergent in ISG15 and SUMO. Correspondingly, UCH-L3 can cleave ubiquitin and Nedd8 adducts but not those of the other ubiquitin-like proteins [26]. The same regions appear to be important for interactions of ubiquitin with many other DUBs (see Figure 2) and Ub-binding proteins. Importantly, all ubiquitinbinding domains examined utilize these same surfaces in binding ubiquitin. Recognition of a ubiquitin domain can be accomplished by ubiquitin-associated domains (UBA), which are present in many proteins, including some DUBs, and interact with polyubiquitin up to 1000-fold better than mono-ubiquitin [51]. However, other binding domains such as UIM (ubiquitin-interacting motif) and CUE (coupling of ubiquitin conjugation to ER degradation) domains utilized in endocytic pathways prefer binding mono-ubiquitin [52–54]. Polyubiquitin-binding proteins recognize a subset of this binding surface of ubiquitin, often described as the hydrophobic patch. It is a group of three amino acids, Leu8, Ile44, and Val70, which are oriented in the ubiquitin molecule to form a small hydrophobic patch [55]. Polyubiquitin chains incorporating ubiquitins with mutations at residues 8 and 44 were unable to be disassembled by DUBs present in the 19S subunit of the proteasome [56]. In addition to providing a recognition site for DUBs, the patch is also important in determining the quaternary structure of polyubiquitin, another feature utilized by DUBs in substrate recognition. One UbL protein, ISG15, consists of a fusion of two ubiquitin domains. The crude mimicking of an Ub-dimer could potentially contribute to its specific recognition by deISGylating enzymes. Polyubiquitin chains linked through all seven lysines in ubiquitin have been detected in vivo, and these poorly characterized forms of non-K48-linked polyubiquitin are also likely to have significant roles in the cell [57]. Polyubiquitin chains that are linked through different lysines are expected to be different enough in structure that individual DUBs could distinguish between them. K63-linked polyubiquitin is a well characterized alternative linkage and unlike K48-linked polyubiquitin, is not involved in proteolytic degradation [58, 59]. Structural data confirms the idea that these two types of polyubiquitin can have different structures [60, 61]. The structures of these dimers were solved by NMR analysis and they were found to have quite different conformations. Indicative of this, non-hydrolyzable ubiquitindimer analogs containing different linkages have markedly different effectiveness when used to inhibit the enzymatic activity of Isopeptidase T [62]. Isopeptidase T binds and cleaves polyubiquitin linked through at least four of the seven possible chain linkages found in vivo, although the catalytic efficiency of these activities is not known [10]. It is interesting to speculate that Isopeptidase T utilizes its two UBA domains to regulate binding of different polyubiquitin substrates. Mutational analysis of the UBA domains and structural data are needed to determine if this is the case and whether it is applicable to other DUBs as well.

3 DUB Specificity 11

3.2 Recognition of the gly–gly Linkage

The central feature that defines all DUBs is that they recognize and act at the C-terminus of the ubiquitin or ubiquitin-like domain. All mature ubiquitin and ubiquitin-like proteins have a C-terminal gly–gly motif and DUB cleavage releases leaving groups attached to the carboxyl group of the C-terminal glycine. With the exception of the JAMM metalloproteases, DUB catalysis starts with the nucleophilic attack of the catalytic cysteine on the carbonyl carbon of the scissile bond to form the tetrahedral intermediate. This is converted to an acyl-enzyme intermediate by expelling the C-terminal leaving group. Attack by a water molecule allows regeneration of the free thiol on the catalytic cysteine and releases free ubiquitin. The JAMM isopeptidases appear to use a classical metalloprotease mechanism [50]. DUB structures have evolved to recognize this C-terminal glycylglycine with exquisite specificity. Analysis of ubiquitin-fusion proteins lacking the gly–gly motif has clearly shown that they are not cleaved efficiently by DUBs [63]. All DUBs exclude larger amino acids at the C-terminus of the ubiquitin domain by having a deep cleft in their respective structures that is only large enough to hold two glycines. The narrowest region of USP7’s catalytic cleft sterically excludes amino acids with any type of side chain, enforcing specificity for ubiquitin conjugates [35]. However, the end of the cleft is open, which allows USP7 to act on large ubiquitin conjugates like its substrate, ubiquitinated p53. ULP1 uses a similar type of cleft to recognize the gly–gly motif except that it uses a tryptophan residue to restrict access to the catalytic site when a substrate is bound to the enzyme [40]. UCHs have a similarly constrained cleft and also use the previously described “blocking loop” to assist in specifically recognizing the C-terminus of ubiquitin. 3.3 Recognition of the Leaving Group

In principle, DUBs might also recognize the leaving group to which ubiquitin is attached. In fact, such a mechanism seems likely as several DUBs have little affinity for ubiquitin and several have been shown to bind the un-ubiquitinated target protein (see Table 2). Interactions between DUBs and putative substrates have been shown for the mammalian DUBs VDU1, USP11, and UBPy, as well as UBP3 from yeast and fat facets from Drosophila [64–68]. In other cases, DUBbinding proteins may serve as scaffolds or adaptors that localize DUBs (discussed below). 3.4 Substrate-induced Conformational Changes

DUBs are not general hydrolases for cleaving after a gly–gly sequence even though they recognize the gly–gly motif at the C-terminus of ubiquitin and ubiquitin-like proteins. What is so special about these particular gly–gly sequences that DUBs will only recognize and act on them and not others? The answer comes from the fact

12 The Deubiquitinating Enzymes Table 2 Identification of DUBs and DUB-binding partners through physical and genetic inter-

action screens DUB

Affinity

Characterized by MS

UCHs UCH37 UCH-L3 UCH-L1 UBPs DOA4 UBP3

Yeast two-hybrid

Interaction partner(s) Synthetic lethal

X X X

X

UBP6 UBP8 USP5 X USP7 USP11 CYLD fat facets UBPy OTU DUBs cezanne otubain1 and 2 X VCIP 135 X JAMM Isopeptidases Rpn11 Others ataxin 3

X

S14, UIP1 [21] Nedd8 [126] JAB1, p27 [127] X X

X X

X X X X X

ubiquitin [43] ubiquitin aldehyde [42] VCP/P47 [72] X

X

SLA1, SLA2 [128] SIR4, Bre5, Stu1 [66, 118, 129] 19S proteasome [130] SAGA, SLIK acetyl transferases [88] ubiquitin [131] ataxin [132] RanBPM [67] NEMO [7] Vasa [75] CDC25(Mm) [133]

UBP6 [92] RAD23, HHR23A, HHR23B [134]

that DUBs interact with the rest of the ubiquitin or ubiquitin-like substrate, and this interaction causes conformational changes in the DUB that are necessary to achieve catalysis. These changes result in rapid and efficient cleavage of only the particular substrate that the DUB is equipped to bind. It also explains why peptides with a gly–gly in them are not susceptible to cleavage by DUBs as they are lacking the substrate-binding domains that cause the DUB conformational change required for cleavage. The different DUB classes utilize a number of conformational changes that are induced upon substrate binding to assist in promoting efficient cleavage. UCH DUBs have been the most thoroughly analyzed. Comparison of the ubiquitin—UCH complex with unliganded UCH shows two significant conformational differences that contribute to keeping the unliganded enzyme in an inactive state. First, the previously described “blocking loop” becomes ordered as it interacts with ubiquitin. Invariant residues form hydrogen bonds with the ubiquitin substrate and other UCH residues, indicating that the loop has functional importance during substrate binding [27]. Second, the side chain of L9 in UCH-L3 intrudes into

4 Localization of DUBs 13

the substrate-binding cleft, occluding the catalytic cysteine and preventing binding of peptide substrates [29]. When ubiquitin binds to the UCH-L3, an interaction between ubiquitin and UCH-L3 repositions L9, allowing access to the active site cleft. Thus, the energy of ubiquitin binding is required to activate UCH-L3, allowing its cleavage. This type of selectivity (where ubiquitin binding is required for activity) may be necessary to prevent deleterious cleavage of other protein substrates by UCHs. A similar situation was observed when the crystal structure of USP7 was solved in the absence of substrate [35]. The catalytic cysteine of the unliganded protein is not in an orientation that would allow catalysis to take place. The histidine residue needed to interact with the active-site cysteine is too distant for a catalytic-triad mechanism to function. Binding of ubiquitin aldehyde induces a significant conformational change that realigns the catalytic triad residues so the hydrogen bonding required for catalysis can take place. Thus, like UCH DUBs, the unliganded protease is inactive and only becomes catalytically active when it is binding substrates. ULP1 also uses conformational changes to “clamp down” on the gly–gly motif when a SUMO substrate is bound. Trp448 lies directly above the active site and interacts with the SUMO C-terminus by Van der Waals interactions, sandwiching the gly–gly motif between Trp448 and the active-site cysteine when SUMO binds [40]. Despite the various methods utilized, all DUBs require a conformational change triggered by binding of a specific substrate to catalyze cleavage. These required conformational changes are driven by the energy of interaction between the DUB and the body of the ubiquitin domain.

4 Localization of DUBs

While many DUBs are cytoplasmic, localization of DUBs is also known to be important in regulating DUB specificity. The localization of ULP1, for example, is important in determining its substrates. The N-terminal domain of ULP1 is known to localize the enzyme to the nuclear envelope, and truncation mutations lacking this domain remain in the cytoplasm [69]. When the truncated protein is expressed in ULP1 yeast strains, the cells grow at wild-type levels, and the truncated protein is able to cleave SUMO substrates in vitro. However, analysis of ULP1 cells expressing this truncation shows an accumulation of SUMO conjugates. Apparently, the localization of ULP1 to the nuclear envelope is necessary in order for it to act on specific nuclear-envelope-localized substrates. The localization helps constrain ULP1 isopeptidase activity so ULP1 does not inappropriately act on cytoplasmic substrates. Other examples of DUB activity regulated by localization include UBP6, which is fully active only when bound to the proteasome (see below) and UBP16 residence on the outer membrane of the mitochondria, although its function there is undetermined [70]. Other DUBs have been found to associate with membranes and regulate membrane-associated cellular processes, although they appear not to be membrane


anchored like UBP16. The ability of DOA4 to remove ubiquitin from membranebound endocytic substrates promotes their degradation in the vacuole or lysosome [71]. DUBs are also important for membrane fusion events as shown by the fact that an OTU domain DUB, VCIP135 (VCP/p47 complex-interacting protein of 135kd), is necessary for p97-p47-mediated Golgi cisternae reassembly after mitosis [72]. Also, a neuronal DUB, synUSP, was found to localize to post-synaptic lipid rafts (membrane microdomains involved in membrane trafficking and signal transduction) [73]. However, its function at that location has yet to be characterized. A well-studied example of a tissue-specific DUB activity is fat facets, a UBP originally found in Drosophila[74]. It is important in eye development and germcell specification and is active only in specific cell types during certain stages of development [65, 75]. The lack of fat facets results in defective posterior patterning, germ-cell specification, and eye formation. Fat facets activity is required to prevent the inappropriate degradation of vasa and liquid facets. In this case, the role of the DUB appears to be defined by the restricted expression of its known substrates. Temporal regulation of DUB expression also appears important. D’Andrea and colleagues first described a small family of DUBs that are induced as immediate early gene products of cytokine stimulation [76]. Different cytokines were shown to induce different DUBs and the expression of these enzymes was short-lived [77]. It appears that these DUBs may be involved in down-regulating cytokine receptors, perhaps by removing the ubiquitin involved in sorting of the receptor at the early endosome. Likewise, UBP43, the short-lived processing protein for ISG15, is present at very low levels in normal cells and highly expressed upon interferon induction [78].

5 Probable Physiological Roles for DUBs 5.1 Proprotein Processing

One important function of DUBs is the processing of ubiquitin or ubiquitin-like proteins to their mature forms. Ubiquitin is expressed in cells as either linear polyubiquitin or N-terminally fused to certain ribosomal proteins [79, 80]. These gene products are processed by DUBs to separate the ubiquitin into monomers and expose the gly–gly motif at the C-terminus. Many DUBs process linear polyubiquitin or Ub-fusion proteins in vitro, but this processing appears to take place cotranslationally in vivo and is extremely rapid. This makes analysis difficult and leaves unanswered the question of which DUBs actually perform this function in vivo. Multiple DUBs may be able to perform this processing at a physiologically relevant level since DUB deletions rarely shows processing defects [81]. Ubiquitin-like proteins are also expressed as proproteins with a short C-terminal extension of a few amino acids that must be removed to make the UbL available for conjugation to target proteins. All ULPs have been shown to metabolize their

5 Probable Physiological Roles for DUBs 15

respective proprotein to an active form in vitro although again it is unclear which ULPs are responsible for this activity in vivo. An exception to the confusion is the finding that RUB1 (the yeast homolog of Nedd8) is processed by YUH1 in Saccharomyces cerevisiae. Conjugation of RUB1 to Cdc53 is required for efficient assembly of certain SCF (skp1, cullin, F-box) E3 ubiquitin ligases [82]. Yuh1 deletion strains do not process Rub1 or modify Cdc53 with Rub1 [83]. Modification of Cdc53 by Rub1 could be reconstituted in a Yuh1 strain by expressing a mature Rub1 construct lacking the C-terminal asparagine normally removed by processing. This demonstrated that Yuh1 processes RUB1 proprotein into the mature form in vivo. It is not known which DUB performs the processing of proNedd8. 5.2 Salvage Pathways: Recovering Mono-ubiquitin Adducts and Recycling Polyubiquitin

It has been speculated that without UCH function, all ubiquitin in the cell would be conjugated with glutathione or other cellular amines and therefore unavailable for conjugation. This would quickly result in the cessation of the ubiquitin–proteasome system function and cell death due to lack of active ubiquitin to conjugate to substrates. The effectiveness of UCH DUBs in liberating ubiquitin from other small adducts makes them likely candidates to act on these particular adducts. In addition, the cell must regenerate mono-ubiquitin from polyubiquitin and various mono-ubiquitinated proteins to maintain levels of mono-ubiquitin for conjugation. Doa4 appears to remove small peptides attached to mono- and di-ubiquitin intermediates resulting from proteasomal degradation as well as removing ubiquitin from proteins targeted for endocytosis [84, 85]. Loss of Doa4 function in yeast results in depleted levels of mono-ubiquitin and increased cell death during stationary phase. Another function for DUBs is regenerating free ubiquitin from unanchored polyubiquitin chains removed from proteasome substrates or proteins targeted for other pathways. Polyubiquitin inhibits the proteasome and lowers the amount of free ubiquitin available for conjugation to proteins. Thus, these chains need to be processed to mono-ubiquitin to prevent polyubiquitin accumulation and inhibition of the proteasome. This type of DUB activity has been well characterized in vivo and in vitro and Isopeptidase T appears to be the DUB that is responsible for the majority of this activity. Deletion of UBP14 in yeast is not lethal, although large amounts of polyubiquitin build up in the cell and proteasome function is impaired [86]. Isopeptidase T seems to serve as a general DUB for regenerating mono-ubiquitin as it cleaves polyubiquitin containing various linkages [59]. 5.3 Regulation of Mono-ubiquitination

DUBs have increasingly been found to be important in regulating the ubiquitination level of proteins not targeted to the proteasome for degradation. Some DUBs are active participants in the regulation of mono-ubiquitin (or mono-UbL) conjugation and others can regulate the conjugation of multiple types of ubiquitin or UbLs


to a single substrate. For instance, deneddylating enzymes may regulate the neddylation of cullin proteins both by processing proNedd8 and by removing Nedd8 from neddylated cullins [26, 38]. As a component of the SCF E3 ligase complexes, cullins require neddylation in order for their cognate E3 ligase to be efficiently assembled [82]. Regulation of this modification indirectly regulates the ubiquitination of a subset of proteins. Defects in deneddylation could lead to inappropriate ubiquitination of substrates owing to inappropriate recruitment of E2s to the SCF E3s [87]. Regulating mono-ubiquitination of proteins by DUBs is important in histone modification where ubiquitination is thought to modulate chromatin structure and transcriptional activity. Normally, about 10% of the histone core octomers contain ubiquitinated histones and the ubiquitin is removed at mitosis by DUB activity. UBP8 has been demonstrated to regulate the ubiquitination of histone H2B, which is important in transcriptional activation of many genes [88]. Many cell-surface receptors are ubiquitinated upon internalization and the ubiquitin is removed by DUBs at the early endosome. Properly sorted receptors are then shuttled to the lysosome for degradation. In the absence of Doa4, the ubiquitin is not removed upon sorting and instead is co-degraded in the vacuole, resulting in ubiquitin depletion [84]. Another DUB, UBP3 assists Golgi-ER retrograde transport by deubiquitinating B -COP, thus preventing its degradation [66]. Mono-ubiquitinated B -COP cannot be assembled into the COP1 complex without UBP3/Bre5 complex DUB activity. Disruption of the complex in Bre5 strains reduces the efficiency of Golgi-ER transport and facilitates the polyubiquitination and degradation of B -COP by the proteasome. One fascinating observation is that PCNA (proliferating cell nuclear antigen) can be modified by multiple forms of ubiquitin, demonstrating that DUBs with different specificities can act at the same location on a specific substrate. PCNA can be modified by mono-ubiquitin, 63-linked polyubiquitin, or SUMO at K164 [89]. Modification of PCNA by mono- or polyubiquitin determines whether it is utilized in translesion synthesis or error-free DNA repair, respectively. SUMO modification prevents PCNA function in DNA repair and instead promotes DNA replication. It is probable that multiple DUBs, as yet unidentified, are required to regulate PCNA modification. 5.4 Processing of Proteasome-bound Polyubiquitin

DUBs play a crucial role in regulating the function of the proteasome. For a long time it was unclear what happens to polyubiquitin conjugated to a proteasome substrate when that substrate is at the proteasome ready for degradation. Was the conjugated polyubiquitin processed by the proteasome and degraded or was it removed by a DUB and released from the proteasome? The small 13-Å diameter entrance to the 20S catalytic core of the proteasome requires all substrates to be fed through as unfolded polypeptides [90]. A branched polypeptide such as a ubiquitin–protein conjugate apparently has difficulty fitting through the pore,

6 Finding Substrates and Roles for DUBs 17

greatly reducing proteasome efficiency [47]. Thus, it seemed likely that DUBs must remove polyubiquitin from proteasome substrates before they enter the 20S catalytic core of the proteasome. To date, three DUBs are known to perform this function and all are components of the 19S lid of the mammalian proteasome. The first described was UCH37, although its exact function is still unclear [91]. UCH37 is thought to be an editing DUB that assists in clearing the proteasome of ubiquitinated proteins. UCH37 slowly cleaves one ubiquitin at a time from the distal end of the polyubiquitin chain. If chain trimming is faster than the degradation process, loss of the polyubiquitin signal could result in partial degradation or release of proteins from the proteasome. UCH37 activity could also be necessary to recover proteasomes that are having difficulty degrading ubiquitinated proteins. UCH37 is only found in higher eukaryotes, but the other two proteasome-bound DUBs, RPN11 and UBP6 (USP14), are found in all eukaryotes. RPN11 and UBP6 remove polyubiquitin from substrates that are committed to degradation by the proteasome [92]. The mechanisms for this, and exactly what role each DUB plays in removing ubiquitin, are not fully understood. Interestingly, the Rpn11 DUB activity was first detected over 10 years ago when 26S proteasome DUB activity was inhibited by o-phenanthroline, a metal chelator [93]. The metalloprotease DUB activity was not identified until recently [46]. Rpn11 is thought to remove polyubiquitin chains from proteasome substrates before they are degraded, allowing the unfolded substrate to enter the pore of the 20S subunit of the proteasome. It has been proposed that Rpn11 removes most of the polyubiquitin chain attached to a proteasome substrate and then UBP6 acts to remove the remaining one or two ubiquitin residues. Despite the lack of mechanistic understanding, the DUBs are clearly required for efficient proteasomal degradation to take place. The Rpn11 deletion is lethal in yeast and temperature-sensitive mutants show massive accumulation of polyubiquitin conjugates [46, 47]. UBP6 is approximately 300 times more active when it is associated with the proteasome than in its purified form [94]. The UBP6 deletion is not lethal in yeast, but a large decrease in the cellular pool of mono-ubiquitin occurs, indicating that ubiquitin is fed into the proteasome and degraded rather than being released from the proteasome and recycled [95].

6 Finding Substrates and Roles for DUBs

Surprisingly, little is known about the in vivo substrate specificity of DUBs. Difficulty in defining the substrate specificity of individual DUBs often arises from a lack of observable phenotypes in deletion strains. Deletion studies in yeast where up to 4 of the 17 DUBs have been deleted in a single strain have not produced significant phenotypes [96]. It is unclear if this is due to the subtle nature of the phenotypes or if the remaining DUBs compensate for the missing ones. However,


several tactics have been fruitful in defining the physiological roles of DUBs. First, definition of in vitro specificities can be useful in focusing genetic screens. For example, the first UBPs were cloned and analyzed after it was discovered that they could cleave ubiquitin-fusion proteins [96, 97]. Second, directed screening of deletions or knockdown studies to identify roles for DUBs have also been successful (see Table 1 for DUB-deletion phenotypes). Study of DUB deletions, including UCH-L1, UCH-L3, and USP14 (see below), in the mouse have demonstrated their importance in neuronal function. Third, potential roles for DUBs have also been identified by physical and genetic interaction screens. Table 2 shows in more detail the interaction screens that have been used in discovering DUBs and characterizing their in vivo roles by identifying novel binding partners. For example, Cezanne was suggested to be a DUB after two-hybrid studies demonstrated its interaction with ubiquitin and UBP6 was identified as a component of the 19S subunit of the proteasome by mass spectrometry.

7 Roles of DUBs Revealed in Disease 7.1 NF-jB Pathway

NF-κB is a transcription factor that can be activated by a number of cellular signals, including stress, inflammation (via tumor necrosis factor) and antigen receptors among others [98, 99]. After receptor stimulation, a cascade ensues that results in the release of NF-κB from its inhibitor IκB. Released NF-κB translocates to the nucleus and activates transcription of a number of genes. Ubiquitin metabolism plays a significant regulatory role in the NF-κB pathway. For NF-κB release from IκB and nuclear translocation to take place, IκB is phosphorylated by IκB kinases, resulting in K48-linked polyubiquitination and proteasomal degradation of IκB [100]. A number of other proteins involved in this pathway such as NEMO, IKKγ , and TRAF6 have K63-linked polyubiquitin chains conjugated to them [101, 102]. It is not clear what purpose the K63-linked chains serve, but they appear to be a regulatory component of the NF-κB pathway. Most of the DUB activity characterized in the NF-κB pathway appears to act on K63-linked polyubiquitin, suggesting that modulation of K63-linked polyubiquitination by DUBs is important for control of the NF-κB pathway. CYLD, a tumor suppressor gene, has been confirmed as a DUB [7, 103, 104]. Loss of CYLD function leads to cylindromatosis, a syndrome characterized by large benign tumors on the face and neck. This is one of the few examples where a defective DUB has been defined as the direct cause of a specific disease. Preferred in vivo substrates of CYLD are believed to be 63-linked polyubiquitin-protein conjugates of NEMO, TRAF6, and TRAF2 components of the NF-κB pathway, but the exact in vivo regulatory role of CYLD is still unknown.

7 Roles of DUBs Revealed in Disease

OTU family DUBs such as Cezanne and A20 also play significant roles as negative regulators of the NF-κB pathway [43, 44]. A20 can cleave K48- and K63-linked polyubiquitin chains in vitro while Cezanne has only been tested on K48-linked chains. Although these DUBs are known to be part of the NF-κB pathway, their in vivo substrates are unknown. It is also unclear as to how these DUBs negatively regulate the NF-κB pathway. 7.2 Neural Function

DUBs, specifically UCHs, appear to play significant roles in neurodegenerative diseases such as Parkinson’s, Alzheimer’s, Huntington’s, and others [105]. A mutant form of UCH-L1 with reduced enzymatic activity has been found in a small family of Parkinson’s patients and the S18Y allele of UCH-L1 has been associated with a reduced risk of sporadic Parkinson’s disease [106, 107]. Many of the inclusion bodies found in patients with Parkinson’s are known to contain high amounts of UCH-L1, ubiquitin, and ubiquitinated proteins, as determined by immunostaining [108]. This suggests that defects in some DUBs or their regulation can cause significant harm to the neuronal system, resulting in disease. DUBs have also been implicated in the formation of other neural inclusion bodies. In addition to the case for their involvement in Parkinson’s disease it has been shown that the mutation of USP14 (the mammalian homolog of yeast UBP6) results in Ataxia in the mouse [109]. Many neurological diseases, including Ataxia, result in damaged or mutated proteins aggregating as polyubiquitinated forms at the microtubule organizing center (MTOC) to form inclusions called aggresomes [110]. An adapter, the tubulin deacetylase HDAC6 (histone deacetylase 6), has recently been shown to bind these polyubiquitinated proteins and tether them to the microtubules where they are then transported to the MTOC [111]. The classic Lewy Body of Parkinson’s disease has all the hallmarks of such an aggresome. The formation of an aggresome is thought to be protective and in its absence the aggregated proteins can trigger apoptosis. Thus, the dynamics of ubiquitination and aggregate formation are important responses to this type of cellular stress and several DUBs can modulate this process. Deletion of UCH-L1 and/or UCH-L3 in mice has demonstrated that they are both involved in neuronal regulation, but have separate functions. The GAD (gracile axonal dystrophy) mouse has been shown to lack UCH-L1, the predominant neuronal UCH [112]. These mice show a unique neuronal “dying back” phenotype that results in paralysis of the limbs due to death of nerves originating in the gracile nucleus. Mice lacking UCH-L3 have no obvious abnormalities or defects [113]. However, the double deletion mouse shows more severe defects including reduced weight, a more severe gracile axonal dystrophy than the L1 deletion, and earlier lethality caused by a loss of the ability to swallow resulting in starvation [114]. This demonstrates that the two DUBs are not redundant and have separate neuronal functions.

19


8 New Tools for DUB Analysis

Despite all the DUB structures and substrates previously described, in most cases the in vivo substrate for a particular DUB is unknown. Structural and localization data can provide clues to determine in vivo DUB specificity, especially if one knows what ubiquitin or ubiquitin-like protein it acts upon. Genomic databases have helped, but many annotated DUBs have never been tested for DUB activity and some DUBs thought to act on one type of substrate (based on their homology) are found to act on another when tested. The characterization of hundreds of potential DUBs is a daunting task and in vivo characterization is even more difficult. To make headway, novel tools are needed to conclusively identify potential DUBs and their substrates to help direct appropriate in vivo studies.

8.1 Active-site-directed Irreversible Inhibitors and Substrates

The most promising tools developed for this sort of analysis are active-site-directed irreversible inhibitors of DUBs. These inhibitors are ubiquitin or ubiquitin-like proteins chemically modified at the C-terminus by an electrophilic moiety such as a Michael acceptor or alkyl halide. The modified ubiquitin can be incubated with a purified DUB or a cell lysate containing DUB activity. Ubiquitin vinyl sulfone (UbVS) is one such irreversible inhibitor because the vinyl sulfone moiety reacts with the active-site cysteine of the DUB, forming a thioether linkage. The covalent adduct is stable and can be detected in a variety of ways. Labeling of DUBs is specific, as only a DUB active-site cysteine will efficiently react with the vinyl sulfone moiety. To create these inhibitors, an N-terminally tagged ubiquitin or ubiquitin-like protein (lacking the C-terminal glycine) is expressed using the intein expression system (New England Biolabs). Briefly, in this system a fusion protein consisting of a ubiquitin or a ubiquitin-like protein lacking the C-terminal glycine, an intein linker, and a chitin-binding domain (CBD) is expressed in E. coli. Clarified cell lysate is incubated with chitin beads to bind the ubiquitin-fusion protein. The ubiquitin is then cleaved from the CBD and intein linker by adding mercaptoethanesulfonic acid (MESNA). After MESNA elution, the resulting truncated ubiquitin C-terminal thioester is reacted with glycine vinyl methyl sulfone to create the Ub or UbL vinyl sulfone derivative. The N-terminal tag on the ubiquitin molecule allows analysis of DUB labeling by immunoprecipitation and Western blotting. This labeling has been used with success in yeast where 6 of the 17 known DUBs were labeled with UbVS [115]. Incomplete labeling likely results from DUBs that do not act on mono-ubiquitin or where the UbVS could not access the active site. The labeling has also been used with great success in mammalian cell lysates to identify novel ubiquitin DUBs [41]. A novel deneddylating enzyme and a novel DUB that acts on autophagy-related UbL proteins have also been identified using vinyl sulfone labeled probes [26, 37, 116].

8 New Tools for DUB Analysis 21

This ubiquitin intein system can also be utilized to make a DUB substrate rather than inhibitors by attaching a C-terminal fluorescent tag such as 7amidomethylcoumarin (AMC) instead of vinyl sulfone. DUBs cleave the ubiquitin derivative and release the fluorescent tag, a process that can be followed fluorometrically. Fluorometric assays can then be used to determine a particular DUB’s preferred substrate or to quantitate DUB activity in crude lysates. AMC substrates have turned out to be excellent tools for identifying the substrates of individual DUBs. Den1, for example, was shown to cleave Nedd8-AMC 60 000-fold faster than it cleaves ubiquitin–AMC, and the ratio was even higher when compared to SUMO–AMC [26]. Clearly, these reagents are powerful tools for identifying novel DUBs and identifying potential DUB substrates.

8.2 Non-hydrolyzable Polyubiquitin Analogs

Other modified ubiquitin reagents that are useful in analyzing DUBs are nonhydrolyzable polyubiquitin analogs. These analogs are polyubiquitin chains where the ubiquitins are linked by cross-linking reagents. To synthesize them, one ubiquitin is mutated to cysteine at the C-terminal glycine and another has cysteine introduced at a particular lysine residue. These ubiquitins can then be linked through their cysteine residues with a bifunctional thiol reagent such as dichloroacetone (DCA). As the native ubiquitin sequence contains no cysteines, the ubiquitins will only be linked through the introduced cysteine residues. The result is a ubiquitin dimer analog that mimics physiological dimers. The isopeptide bond is replaced by a DCA linkage, but the ubiquitin subunits retain the appropriate spatial orientation. Thus, DUBs should bind these dimers, but will be unable to cleave them because they cannot hydrolyze the DCA linkage. To make longer polyubiquitin chains, cysteine residues or sulfhydryl groups must be introduced at the desired lysine and the C-terminal glycine on the same ubiquitin molecule. These chain analogs have been used to characterize DUBs in two fashions. First, they can be used as inhibitors of DUBs [62]. Cleavage of Ub–AMC by Isopeptidase T, a polyubiquitin-binding DUB, was inhibited by the addition of differently linked dimer analogs and kinetic inhibition constants were determined. The K i values of dimer analogs were all much lower than the K i for mono-ubiquitin. Further, there was considerable selectivity, as inhibition constants varied depending on the linkage present in the dimer [62]. This demonstrated that the analogs act as faithful mimics of native polyubiquitin. The other way these chain analogs are used is to synthesize them on affinity supports and analyze cell lysates for DUBs that bind a specific type of polyubiquitin chain. These affinity resins have proven useful in identifying a number of binding proteins, including DUBs, from yeast cell lysates [117]. Analogs with different linkages bind a different subset of proteins, perhaps suggesting a way to identify DUBs acting upon polyubiquitin with linkage specificity. Thus, these analogs are excellent tools for characterizing the substrate preferences of known DUBs and discovering novel ones.


9 Conclusion

DUBs are clearly an essential cellular component needed for a variety of pathways including protein degradation, DNA repair, apoptosis, membrane trafficking, stress response, and transcriptional regulation. Not only do they act on various ubiquitin substrates, but they are also needed to process ubiquitin-like substrates. Over 100 DUBs from five major families have been identified and the number is likely to increase. Factors that enhance DUB specificity are the presence of a binding pocket that only accommodates the physiological substrate, the requirement for a substrate-induced conformational change that prevents undesired catalysis, and the recognition of the ubiquitous C-terminal gly–gly motif of all DUB substrates. Subcellular localization to a specific organelle or protein complex and tissue-specific as well as temporal expression are also important components of DUB specificity and function. In spite of all that has been learned about DUBs and their function, much remains to be discovered. Future studies are likely to focus on identifying in vivo DUB substrates, novel DUBs, DUB-binding partners, and phenotypes of DUB deletions. Further development of new reagents, such as the non-hydrolyzable polyubiquitin analogs and active-site-directed inhibitors or substrates will help greatly. Directed proteomics studies should assist in identifying DUBs, loss-offunction phenotypes, and potential binding partners. Despite the major gaps that remain in our understanding of DUBs, our knowledge of their roles and importance has progressed amazingly rapidly. Novel DUB gene families have been identified, new ubiquitin-like DUB substrates have been uncovered, and structural data has been analyzed to elucidate how DUBs perform catalysis and specifically recognize their substrates. In vivo substrates of DUBs are beginning to be identified and the tools and techniques needed to search for novel DUBs and analyze known ones for their specificity are rapidly being created. With so much discovered, and yet so much remaining to be found, deubiquitinating enzymes are a vibrant field of study.

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108 LOWE, J., MCDERMOTT, H., LANDON, M., MAYER, R. J., and WILKINSON, K. D. Ubiquitin carboxyl-terminal hydrolase (PGP 9.5) is selectively present in ubiquitinated inclusion bodies characteristic of human neurodegenerative diseases, J Pathol, 1990, 161, 153–160. 109 WILSON, S. M., BHATTACHARYYA, B., RACHEL, R. A., COPPOLA, V., TESSAROLLO, L., HOUSEHOLDER, D. B., FLETCHER, C. F., MILLER, R. J., COPELAND, N. G., and JENKINS, N. A. Synaptic defects in ataxia mice result from a mutation in Usp14, encoding a ubiquitin-specific protease, Nat Genet 2002, 32, 420–425. 110 GARCIA-MATA, R., GAO, Y. S., and SZTUL, E. Hassles with taking out the garbage: aggravating aggresomes, Traffic, 2002, 3, 388–396. 111 KAWAGUCHI, Y., KOVACS, J. J., MCLAURIN, A., VANCE, J. M., ITO, A., and YAO, T. P. The deacetylase HDAC6 regulates aggresome formation and cell viability in response to misfolded protein stress, Cell, 2003, 115, 727–738. 112 SAIGOH, K., WANG, Y. L., SUH, J. G., YAMANISHI, T., SAKAI, Y., KIYOSAWA, H., HARADA, T., ICHIHARA, N., WAKANA, S., KIKUCHI, T., and WADA, K. Intragenic deletion in the gene encoding ubiquitin carboxy-terminal hydrolase in gad mice, Nat Genet 1999, 23, 47–51. 113 KURIHARA, L. J., SEMENOVA, E., LEVORSE, J. M., and TILGHMAN, S. M. Expression and functional analysis of Uch-L3 during mouse development, Mol Cell Biol, 2000, 20, 2498–2504. 114 KURIHARA, L. J., KIKUCHI, T., WADA, K., and TILGHMAN, S. M. Loss of Uch-L1 and Uch-L3 leads to neuro-degeneration, posterior paralysis and dysphagia, Hum Mol Genet, 2001, 10, 1963–1970. 115 BORODOVSKY, A., KESSLER, B. M., CASAGRANDE, R., OVERKLEEFT, H. S., WILKINSON, K. D., and PLOEGH, H. L. A novel active site-directed probe specific for deubiquitylating enzymes reveals proteasome association of USP14, Embo J, 2001, 20, 5187–5196. 116 HEMELAAR, J., LELYVELD, V. S., KESSLER, B. M., and PLOEGH, H. L. A single protease,

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1

The 26S Proteasome Martin Rechsteiner University of Utah Medical School, Salt Lake City, USA

Originally published in: Protein Degradation, Volume 1. Edited by R. John Mayer, Aaron Ciechanover and Martin Rechsteiner. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30837-8 The 26S proteasome is a large ATP-dependent protease composed of more than 30 different polypeptide chains. Like the ribosome, the 26S proteasome is assembled from two “subunits”, the 19S regulatory complex and the 20S proteasome. The 19S regulatory complex confers the ability to recognize and unfold protein substrates, and the 20S proteasome provides the proteolytic activities needed to degrade the substrates. The 26S proteasome is the only enzyme known to degrade ubiquitylated proteins, and it also degrades intracellular proteins that have not been marked by ubiquitin. The 26S proteasome is located in the nucleus and cytosol of eukaryotic cells, where the enzyme is responsible for the selective degradation of a vast number of important cellular proteins. Because rapid proteolysis is a pervasive regulatory mechanism, the 26S proteasome is essential for the proper functioning of many physiological processes.

1 Introduction

It has become apparent since the mid-1990s that the ubiquitin-proteasome system (UPS) plays a major regulatory role in eukaryotic cells. The UPS helps to control such important physiological processes as the cell cycle [1, 2], circadian rhythms [3], axon guidance [4], synapse formation [5] and transcription [6–8], to name just a few. In view of the growing family of ubiquitin-like proteins [9, 10], it is possible that covalent attachment of ubiquitin and its relatives will even surpass phosphorylation as a regulatory mechanism. Although ubiquitin serves non-proteolytic roles, such as histone modification [11, 12] the activation of cell signaling components [13], endocytosis [14], or viral budding [15], the protein’s principal function appears to be targeting proteins for destruction [16]. To do this, the C-terminus of ubiquitin is activated by an ATP-consuming enzyme (E1) and transferred to one of several


2 The 26S Proteasome

Fig. 1 Schematic representation of the Ubiquitin–Proteasome pathway. Ubiquitin molecules are activated by an E1 enzyme (shown green at 1/3 scale) in an ATPdependent reaction, transferred to a cysteine residue (yellow) on an E2 or Ub carrier protein and subsequently attached to

amino groups (NH2 ) on a substrate protein (lysozyme shown in purple) by an E3 or ubiquitin ligase, (the multicolored SCF complex). Note that chains of Ub are generated on the substrate, and these are recognized by the 26S proteasome depicted in the upper right at 1/20 scale.

dozen or more small carrier proteins (E2s) in the form of a reactive thiol ester. The E2s collaborate with members of several large families of ubiquitin ligases or E3s, and ubiquitin is transferred once again to lysine amino groups on the proteolytic substrates (S) and to itself, thereby generating chains of ubiquitin. The substrate bearing the ubiquitin chains is subsequently recognized and degraded by the 26S proteasome, a large, complex ATP-dependent protease (see Figure 1). Eukaryotic genomes contain information for more than 20 E2s and hundreds of E3s. In contrast to the wealth of components devoted to marking protein substrates for destruction, only one enzyme, the 26S proteasome, has been found to degrade ubiquitylated proteins. However, there is complexity here as well, since the 26S proteasome is an assemblage of at least 30 different subunits. Moreover, there is


a growing list of proteins that act as proteasome activators, adapters, or accessory factors. In this article I focus on basic biochemical and physiological properties of the 26S proteasome, drawing occasionally from findings on structurally similar prokaryotic, ATP-dependent proteases [17].

2 The 20S Proteasome 2.1 Structure

We know the molecular anatomy of archaebacterial, yeast and bovine proteasomes in great detail since high-resolution crystal structures have been determined for all three enzymes [18–20]. The archaebacterial proteasome is composed of two kinds of subunits, called α and β. Each subunit forms heptameric rings that assemble into the 20S proteasome by stacking four deep on top of one another to form a “hollow” cylinder. Catalytically inactive α rings form the ends of the cylinder while proteolytic β subunits occupy the two central rings. The quaternary structure of the 20S proteasome can therefore be described as α7β7β7α7. The active sites of the β-subunits face a large central chamber about the size of serum albumin. The αrings seal off the central proteolytic chamber and two smaller antechambers from the external solvent. Archaebacterial proteasomes, with their fourteen identical β subunits, preferentially hydrolyze peptide bonds following hydrophobic amino acids and are therefore said to have chymotrypsin-like activity [21]. Eukaryotic proteasomes maintain the overall structure of the archaebacterial enzyme, but they exhibit a more complicated subunit composition. There are seven different α-subunits and at least seven distinct β-subunits arranged in a precise order within their respective rings. Although current evidence indicates that only three of its seven β-subunits are catalytically active, the eukaryotic proteasome cleaves a wider range of peptide bonds, containing, as it does, two copies each of trypsin-like, chymotrypsin-like and post-glutamyl-hydrolyzing subunits. For this reason it is capable of cleaving almost any peptide bond, having difficulty only with proline–X, glycine–X and to a lesser extent with glutamine–X bonds [22]. 2.2 Enzyme Mechanism and Proteasome Inhibitors

Whereas standard proteases use serine, cysteine, aspartate, or metals to cleave pep-tide bonds, the proteasome employs an unusual catalytic mechanism. Nterminal threonine residues are generated by self-removal of short peptide extensions from the active β-subunits and act as nucleophiles during peptide-bond hydrolysis [23]. Given its unusual catalytic mechanism, it is not surprising that there are highly specific inhibitors of the proteasome. The fungal metabolite lactacystin

3


and the bacterial product epoxomicin covalently modify the active-site threonines and inhibit the enzyme [24, 25]. Other inhibitors include vinylsulfones [26] and various peptide aldehydes, which are generally less specific. A peptide boronate inhibitor of the proteasome, Velcade, has been approved for the treatment of multiple myeloma [27]. 2.3 Immunoproteasomes

Interferon γ (IFNγ ) is an immune cytokine that increases expression of a number of cellular components involved in Class I antigen presentation [28]. Among the IFNγ -inducible components are three catalytically active β-subunits of the proteasome, called LMP2, LMP7 and MECL1 [29]. Each replaces its corresponding constitutive subunit resulting in altered peptide-bond cleavage preferences of 20S immunoproteasomes. For example, immunoproteasomes exhibit much reduced cleavage after acidic residues and enhanced hydrolysis of peptide bonds following branch-chain amino acids such as isoleucine or valine. Class I molecules preferentially bind peptides with hydrophobic or positive C-termini, and proteasomes generate the vast majority of Class I peptides [28, 30]. Hence, the observed β-subunit exchanges are well suited for producing peptides able to bind Class I molecules.

3 The 26S Proteasome 3.1 The Ubiquitin–Proteasome System

Bacteria express as many as five ATP-dependent proteases, all of which contain nucleotide-binding domains that belong to the AAA+ family of ATPases [31]. By contrast, the 26S proteasome is the only ATP-dependent protease discovered so far in the nuclear and cytosolic compartments of eukaryotic cells. Because the 20S proteasome’s internal cavities are inaccessible to intact proteins, openings must be generated in the enzyme’s outer surface for proteolysis to occur. A number of protein complexes have been found to bind the proteasome and stimulate peptide hydrolysis (see Figure 2). The most important of the proteasome-associated components is the 19S regulatory complex (RC) for it is a major part of the 26S ATP-dependent enzyme that degrades ubiquitin-tagged proteins in eukaryotic cells. In Figure 2 the 20S proteasome is shown binding only one 19S RC although doublycapped 26S proteasomes also exist (see Figure 3 below). The 20S proteasome also binds activators such as PA28 or PA200. Each of these activators can be present in 26S proteasome complexes forming what are called hybrid proteasomes. Finally a protein called Ecm29p has been found associated with the 26S proteasome. Ecm29p is thought to act as an adapter coupling the 26S enzyme to secretory vesicles.


Fig. 2 Interaction of the 20S proteasome with other cellular components.

Fig. 3 Electron-microscopic reconstructions of the 26S proteasome. Three images of a doubly-capped 26S proteasome are presented to illustrate the positions of the lid and base subcomplexes of the 19S RC and to identify the most probable location of the RC ATPases.

5


3.2 Ultrastructure of the 26S Proteasome and Regulatory Complex

Electron micrographs of purified 26S proteasomes by Baumeister and colleagues [32] reveal a dumbbell-shaped particle approximately 40 nm in length in which the central 20S proteasome cylinder is capped at one or both ends by asymmetric RCs looking much like Chinese dragonheads (Figure 3). In doubly capped 26S proteasomes the regulatory complexes face in opposite directions, indicating that contact between the proteasome’s α-rings and the RC is highly specific. However, the contacts may not be especially tight since image analysis of Drosophila 26S proteasomes suggests movement of the RCs relative to the 20S proteasome [33]. Electron microscopy (EM) images of the 26S proteasome from several organisms appear similar, indicating that the overall architecture of the enzyme has been conserved in evolution. This conclusion is also supported by sequence conservation among RC subunits (see below). A yeast mutant lacking the RC subunit Rpn10 contains a salt-labile RC that dissociates into two subcomplexes called the lid and the base [34]. The base contains nine RC subunits, which include six ATPases described below, the two largest RC subunits S1 and S2, and S5a; the lid contains the remaining RC subunits. Thus the RC is composed of two sub-complexes separated on one side by a cavity, i.e. the dragon’s mouth (see Figure 3). Ultrastructural studies have also been performed on the lid and on a related protein complex called the COP9 signalosome [35]. Both particles lack obvious symmetry. Some particles exhibit a negative stainfilled, central groove; other classes of particles exhibit seven or eight lobes in a disclike arrangement. Since both the lid and the COP9 signalosome are composed of eight subunits, the lobes may represent individual subunits. 3.3 The 19S Regulatory Complex

The regulatory complex is also called the 19S cap, PA700, and the µ particle. As its most commonly used name suggests, the 19S RC is roughly the same size as the 20S proteasome. In fact it is a more complicated protein assembly containing 17 different subunits ranging in size from 25 kDa to about 110 kDa. In animal cells the subunits are designated S1 through S15. Homologs for each of these subunits are present in budding yeast where an alternate nomenclature has been adopted (see Table 1). Sequences for the 17 RC subunits permit their classification into a group of 6 ATPases and another group containing the 11 non-ATPases. 3.4 ATPases of the RC

The six ATPases belong to the rather large family of AAA ATPases (for ATPases Associated with a variety of cellular Activities) whose eukaryotic members include the motor protein dynein, the membrane fusion factor NSF, and the chaperone VCP/Cdc48 and whose prokaryotic members include five ATP-dependent pro-

3 The 26S Proteasome Table 1 Subunits of the 19S regulatory complex

Mammalian nomenclature Yeast nomenclature Function

Motifs

S1 S2 S3 p55 S4 S5a S5b S6 S6 S7 S8 S9 S10a S10b S11 S12 S13 S14

Leu-rich repeats, KEKE Leu-rich repeats, KEKE PCI PCI AAA nucleotidase UIM, KEKE

Rpn2 Rpn1 Rpn3 Rpn5 Rpt2 Rpn10 none Rpt3 Rpt5 Rpt1 Rpt6 Rpn6 Rpn7 Rpt4 Rpn9 Rpn8 Rpn11 Rpn12

Ub/UbL binding Ub/UbL binding ? ? ATPase polyubiquitin binding ? ATPase ATPase ATPase ATPase ? ? ATPase ? ? Isopeptidase ?

AAA nucleotidase AAA nucleotidase AAA nucleotidase AAA nucleotidase PCI PCI, KEKE AAA nucleotidase PCI MPN, KEKE MPN PCI

teases [31]. The six ATPases, denoted S4, S6, S6 , S7, S8, and S10b in mammals, are about 400 amino acids in length and homologous to one another. Based on their sequences, one can distinguish three major regions: (1) A central nucleotidebinding domain of about 200 amino acids, which is roughly 60% identical among members of the RC subfamily; (2) the C-terminal region, approximately 100 amino acids in length and with a lesser, though significant, degree of conservation (40%); and (3) a highly divergent N-terminal region (1 indicates the conformation is more stable than coil, Thr > Glu > Ser > Asn > Gly > Gln > Ala > Val. Interestingly, their result conflicted with the statistical survey result that Asn is one of the most frequently found N-caps in proteins [12]. Experimentally Asn destabilized the helices by 1.3 kcal mol−1 relative to Thr. The rank order of amino acids N-cap preferences in T4 lysozyme was found to be Thr > Ser > Asn > Asp > Val = Ala > Gly [21]. They suggested that Asn can be inherently as good an N-cap as Ser or Thr, but it requires a change in backbone dihedral angles of N-cap residues, which might be altered in native proteins as the results of tertiary contacts. Indeed Asn is the most stabilizing residue at N-cap in a peptide model in the absence of tertiary contacts and other side-chain interactions (see below). The Kallenbach group [187] substituted several amino acids at the N-cap position in peptide models in the presence of a capping box. They found that Ser and Arg are the most stabilizing residues with G relative to Ala of −0.74 and −0.58 kcal mol−1 , respectively, whilst Gly and Ala are less stabilizing. The results are in agreement with the results of Forood et al. [23], who found that the trend in α-helix inducing ability at the N-cap is Asp > Asn > Ser > Glu > Gln > Ala. A more comprehensive work to determine the preferences for all 20 amino acids at the N-cap position used peptides with a sequence of NH2 -XAKAAAAKAAAAKAAGYCONH2 [22, 29, 99]. N-Capping free energies ranged from Asn (best) to Gln (worst) (Table 6). We have used a similar approach using peptide models to probe the preferences at N1 [95], N2 [96], and N3 [97] using peptides with sequences of CH3 COXAAAA-QAAAAQAAGY-CONH2 , CH3 CO-AXAAAAKAAAAKAAGY-CONH2 and CH3 CO-AAXAAAAKAAAAKAGY-CONH2 , respectively. The results have given N1, N2 and N3 preferences for most amino acids for these positions (Table 6) and these agree well with preferences seen in protein structures, with the interesting exception of Pro at N1. Petukhov et al. similarly obtained N1, N2, and N3

5 Forces Affecting "-helix Stability 29 Table 6 Amino acid propensities at N- and C-terminal positions of the helix

G relative to Ala for transition from coil to the position (kcal mol−1 )

Residue

A C◦ C− D◦ D− E◦ E− F G H◦ H+ I K+ L M N P Q R+ S T V W Y

N-cap

N1

29

95 164;188

0

0

−1.4 1.0 0.5 −1.6 0 1.0 −0.7 0.1 −0.7 1.4 −1.2 1.0 −0.7 0.7 −0.5 0.1 −0.7 −0.3 −1.7 −0.4 2.5 −0.1 −1.2 −0.7 −0.1 −1.3 −0.9

0.5 0.7 0.4 0.5 0.6 0.5 0.7 0.4 0.5 0.6 0.4

N2

0

96

N3

C3

164;188 97 164;188 189 0

0

0.9 0.7 −0.2 −0.2 −0.4 0.9 0.7

0

0

0

C2 190 0

C1 C-cap 189 189 0

0

29

0.6 1.3 0.4

0 0.2

0

0.2

0.3

−0.4 −0.5

0.3

0.8

2.1

0.6 1.0 0.6

0.4

0.5

0.2

0.2 0.4

0.5

0.5

0.1 0.1 0.7 0.4

−0.1 −0.1 −0.3 0.3 0.1

0.1

0.2 0.1 0.6 0.5 0.3 0.4 0.5

0.5 0.8 0.7 0.5 0.5 0.8

0.1 −1.1

2.6 −0.2

0.6 0.9 0.5 0.7 1.7

251

1.1

0.8 0.4

C

0.5 0.5 0.3 0.7

0.7 0.9 0.8 0.7

0.4 0.4 0.7

0.3

1.2

0.2

0.5 0.5 0.4

1.1 1.2

0.6 0.6 0.4

4.0 1.2

0.2 −0.02 0.2 0.05 −0.5 −0.4 0.6 0.5 0.7 0.5 0.8 0.8 0.6 0.5 0.8 0.3 0.4 0.7 0.6 0.9

−0.9 1.5 −0.1 0.1 −0.4 1.2 −0.1 −0.2 0.3 1.1 1.6 0.7

−2.2

NH2 −XAKAAAAKAAAAKAAGY-CONH2 CH3 CO-XAAAAQAAAAQAAGY-CONH2 c CH3 CO-XAAAAAAARAAARGGY-NH2 d CH3 CO-AXAAAAKAAAAKAAGY-CONH2 e CH3 CO-AXAAAAAARAAARGGY-NH2 f CH3 CO-AAXAAAAKAAAAKAGY-CONH2 g CH3 CO-AAXAAAAARAAARGGY-NH2 h NH2 −YGGSAKEAAARAAAAXAA-CONH2 i Substitution of residue 32 (C2 position) of α-helix of ubiquitin. j NH2 −YGGSAKEAAARAAAAAXA-CONH2 k NH2 −YGGSAKEAAARAAAAAAX-CONH2 l CH3 CO-YGAAKAAAAKAAAAKAX-COOH m Substitution of residue 35 (C position) of α-helix of ubiquitin. a b

preferences for nonpolar and uncharged polar residues by applying AGADIR to experimental helical peptide data, and found almost identical results [164, 188]. The complete sequences of peptides used can be seen in the table footnote. In general, at N1, N2, and N3, Asp and Glu as well as Ala are preferred, presumably because negative side chains interact favorably with the helix dipole or NH groups while Ala has the strongest interior helix preference.

30 Design and Stability of "-Helices

Although it is also unique in terms of the presence of unsatisfied backbone hydrogen bonds, the C-terminal region is less explored experimentally. The Cterminus of the α-helix tends to fray more than the N-terminus, making C-terminal measurements less accurate. Preferences at the C-cap position differ from those at the N-cap. At the N-terminus, the helix geometry favors side chain-to-backbone hydrogen bonding, so polar residues are preferred [14, 19]. At the C-terminus unsatisfied backbone hydrogen bonds are fulfilled by interactions with backbone groups upstream of the helix. Zhou et al. [48] found that Asn is the most favored residue at the C-cap followed by Gln > Ser∼Ala > Gly∼Thr. Forood et al. [23] tested a limited number of amino acids at the C-terminus (C1) finding a rank order of Arg > Lys > Ala. Doig and Baldwin [29] determined the C-capping preferences for all 20 amino acids in α-helical peptides. The thermodynamic propensities of some amino acids at C, C-cap, C1, C2, and C3 are also included in Table 6 [189, 190]. 5.3 Phosphorylation

Phosphoserine is destabilizing compared with serine at interior helix positions [191, 192]. We investigated the effect of placing phosphoserine at the N-cap, N1, N2, N3, and interior position in alanine-based α-helical peptides, studying both the −1 and −2 phosphoserine charge states [193]. Phosphoserine stabilizes at the N-terminal positions by as much as 2.3 kcal mol−1 , while it destabilizes in the helix interior by 1.2 kcal mol−1 , relative to serine. The rank order of free energies relative to serine at each position is N2 > N3 > N1 > N-cap > interior. Moreover, −2 phosphoserine is the most preferred residue known at each of these N-terminal positions. Experimental pK a values for the −1 to −2 phosphoserine transition are in the order N2 < N-Cap < N1 < N3 < interior. Phosphoserine can form stabilizing salt bridges to arginine [192]. 5.4 Noncovalent Side-chain Interactions

Many studies have been performed on the stabilizing effects of interactions between amino acid side chains in α-helices. These studies have identified a number of types of interaction that stabilize the helix including salt bridges [83, 86, 88, 152, 194–198], hydrogen bonds [152, 198–200], hydrophobic interactions [100, 201–203], basic/aromatic interactions [106, 204], and polar/nonpolar interactions [101]. The stabilizing energies of many pairs in these categories have been measured, though some have only been analyzed qualitatively. As described earlier, residue side chains spaced i, i + 3 and i, i + 4 are on the same face of the α-helix, though it is the i, i + 4 spacing that receives most attention in the literature, as these are stronger. A summary of stabilizing energies for side-chain interactions is given in Table 7. We give only those that have been measured in helical peptides with the side-chain interaction energies determined by applying helix/coil theory. Almost all are attractive, with the sole exception of the Lys-Lys repulsion.

5 Forces Affecting "-helix Stability 31 Table 7 Summary of side-chain interaction energies from literature

Interaction Ile-Lys (i, i + 4) Val-Lys (i, i + 4) Ile-Arg (i, i + 4) Phe-Met (i, i + 4) Met-Phe (i, i + 4) Gln-Asn (i, i + 4) Asn-Gln (i, i + 4) Phe-Lys (i, i + 4) Lys-Phe (i, i + 4) Phe-Arg (i, i + 4) Phe-Orn (i, i + 4) Arg-Phe (i, i + 4) Tyr-Lys (i, i + 4) Glu-Phe (i, i + 4) Asp-Lys (i, i + 3) Asp-Lys (i, i + 4) Asp-His (i, i + 3) Asp-His (i, i + 4) Asp-Arg (i, i + 3) Glu-His (i, i + 3) Glu-His (i, i + 4) Glu-Lys (i, i + 3) Glu-Lys (i, i + 4) Phe-His (i, i + 4) Phe-Met (i, i + 4) His-Asp (i, i + 3) His-Asp (i, i + 4) His-Glu (i, i + 3) His-Glu (i, i + 4) Lys-Asp (i, i + 3) Lys-Asp (i, i + 4) Lys-Glu (i, i + 3) Lys-Glu (i, i + 4) Lys-Lys (i, i + 4) Leu-Tyr (i, i + 3) Leu-Tyr (i, i + 4) Met-Phe (i, i + 4) Gln-Asp (i, i + 4) Gln-Glu (i, i + 4) Trp-Arg (i, i + 4) Trp-His (i, i + 4) Tyr-Leu (i, i + 3) Tyr-Leu (i, i + 4) Tyr- Val (i, i + 3) Tyr- Val (i, i + 4) Arg (i, i + 4) Glu (i, i + 4) Arg Arg (i, i + 3) Glu (i, i + 3) Arg Arg (i, i + 3) Glu (i, i + 4) Arg Arg (i, i + 4) Glu (i, i + 3) Arg Phosphoserine-Arg (i, i + 4)

G (kcal mol−1 )

Reference

−0.22 −0.25 −0.22 −0.8 −0.5 −0.5 −0.1 −0.14 −0.10 −0.18 −0.4 −0.1 −0.22 −0.5 −0.12 −0.24 >−0.63 >−0.63 −0.8 −0.23 −0.10 −0.38 −0.44 −1.27 −0.7 −0.53 −2.38 −0.45 −0.54 −0.4 −0.58 −0.38 −0.46 +0.17 −0.44 −0.65 −0.37 −0.97 −0.31 −0.4 −0.8 −0.02 −0.44 −0.13 −0.31 −1.5 −1.0 −0.3 −0.1 −0.45

[101] [101] [101] [100] [100] [200] [200] [106] [106] [106] [204] [106] [106] [252] [227] [227] [253] [253] [254] [227] [227] [152] [152] [198] [203] [198] [241] [227] [227] [227] [227] [227] [227] [196] [153] [153] [203] [199] [152] [252] [161] [153] [153] [153] [153] [255] [255] [255] [255] [192]


5.5 Covalent Side-chain interactions

Lactam (amide) bonds formed between NH3 + and CO2 − side chains can stabilize a helix, acting in a similar way to disulfide bridges in a protein by constraining the side chains to be close, reducing the entropy of nonhelical states [205]. Lactam bridges between Lys-Asp, Lys-Glu, and Glu-Orn spaced i, i + 4 have been introduced into analogs of human growth hormone releasing factor [206], and proved to be stabilizing with Lys-Asp most effective. The same Lys-Asp i, i + 4 lactam was stabilizing in other helical peptide systems [207–210], while Lys-Glu i, i + 4 lactam bridges were less effective [211]. Two overlapping Lys-Asp lactams were even more stabilizing [212]. The effect of the ring size formed by the lactam was investigated by replacing Lys with ornithine or (S)-diaminopropionic acid. A ring size of 21 or 22 atoms was most stabilizing (a Lys-Asp i, i + 4 lactam is 20 atoms) [206]. Lactams between side chains spaced i, i + 7 [213, 214] or i, i + 3 [214]; [215], spanning two or one turns of the helix have also been reported. i, i + 7 disulfide bonds have been introduced into alanine-based peptides, using (D)- and (L)-2-amino-6-mercaptohexanoic acid derivatives [216]. Strongly stabilizing effects were observed. Some interesting recent work has shown that helix formation can be reversibly photoregulated. Two cysteine residues are cross-linked by an azobenzene derivative which can be photoisomerized from trans to cis, causing a large increase or decrease in the helix content of the peptide, depending on its spacing [217–219].

5.6 Capping Motifs

Although the N-terminal capping box sequence stabilizes helices by inhibiting N-terminal fraying, it does not necessarily promote elongation unless accompanied by favorable hydrophobic interactions as in a “hydrophobic staple” motif [220, 221]. The nature of the capping box stabilizing effect thus not only arises from reciprocal hydrogen bonds between compatible residues, but also from local interactions between side chains, helix macrodipole-charged residue interactions and solvation [222]. Despite statistical analyses revealing that Schellman motifs are observed more frequently that expected at the helix C-terminus, this motif populates only transiently in aqueous solution but it is formed in 30% TFE [223]. This might be due to the C-terminus being very frayed and the increase of helical content contributed from this motif is small. Energetically this motif is not very favorable due to the entropic cost of fixing a Gly residue at the position C . The Schellman motif is believed to be a consequence of helix formation and does not involve α-helix nucleation [224]. The α L motif seems to be more stable than the alternative Schellman motif [221].

5 Forces Affecting "-helix Stability 33

5.7 Ionic Strength

Electrostatic interactions between charged side chains and the helix macrodipole can stabilize the helix [92, 102, 225]. The interactions are potentially quite strong, but are alleviated by the screening effects of water, ions, and nearby protein atoms. In theory, increasing ionic strength of the solvent (up to 1.0 M) should stabilize the helix through interactions with α-helix dipole moments by shifting the equilibrium between α-helix and random coil, which has a random orientation of the peptide dipoles [226]. The energetics of the interaction between fully charged ion pairs can be diminished by added salt and completely screened at 2.5 M NaCl [197, 227]. In peptides containing side chain-to-side-chain interactions, the effect of ion pairs and charge/helix-dipole interactions cannot be clearly separated. There are, however, indications that the interactions of charged residues with the helix macrodipole are less affected than those between charged side chains [227, 228]. In coiled-coil peptides, salt also affects hydrophobic residues by strengthening their interactions at the coiled-coil interface. This can be explained through alterations of the peptide-water interactions at high salt concentration. However, this requires a strong kosmotropic anion to accompany the screening cation [229]. 5.8 Temperature

Thermal unfolding experiments show that the helix unfolds with increasing temperature [230–232]. There is no sign of cold denaturation, as seen with proteins. Enthalpy and entropy changes for the helix/coil transition are difficult to determine as the helix/coil transition is very broad, precluding accurate determination of highand low-temperature baselines by calorimetry [230]. Nevertheless, isothermal titration calorimetric studies of a series of peptides that form helix when binding a nucleating La3+ , find H for helix formation to be −1.0 kcal mol−1 [135, 233], in good agreement with the earlier work. 5.9 Trifluoroethanol

Peptides with sequences of helices in proteins usually show low helix contents in water. An answer to this problem is to add TFE (2,2,2-trifluoroethanol) to induce helix formation [234–237]. For many peptides, the concentration of TFE used to increase the helix content is only up to 40% [234–236, 238, 239]. TFE may act by shielding CO and NH groups from the water solvent while leading to hydrogen bond formation between them. The conformational equilibrium thus shifts toward more compact structures, such as the α-helical conformation [184]. The mechanism involves interaction between TFE and water with several interpretations. One view suggests that TFE indirectly disrupts the solvent shell on α-helices [240, 241]. Another view proposes that TFE destabilizes the unfolded species and thereby


indirectly enhances the kinetics and thermodynamics of folding of the coiled coil [242]. A more compromising view suggests that TFE forms clusters in water solution, which at lower concentration pulls the water molecules from the surface of proteins. At higher concentration, TFE clusters associate with appropriate hydrophobic side chains reducing their conformational entropy and switch the conformation at TFE concentration > 40% [243]. The propagation propensities of all amino acids increase variably in 40% TFE relative to water. The propagation propensities of the nonpolar amino acids increase greatly in 40% TFE whilst other amino acids propensity increase uniformly. However, glycine and proline are strong helix breakers in both in water and 40% TFE solvents [30]. In addition, 40% TFE dramatically alters electrostatic (and polar) interactions and increases the dependence of helix propensities on the sequence [244]. 5.10 pK a Values

Evaluation of pK a values of titrable amino acids in a peptide sequence can be used to analyze the strength of the possible interaction they form in water. pK a shifts of charged residues at the helix termini are significant because they can potentially interact with unsatisfied hydrogen bonds of the NH groups and CO group at the N-terminus and C-terminus, respectively, or the helix dipole. The pK a values can be measured accurately from the change in ellipticity across a broad range of pH. The asymptotic values of the ellipticities for the different protonation states are fitted to a Henderson-Hasselbach equation to calculate the pK a . In general, the pK a values of Glu and His at N1–N3 are normal compared with those in model compounds. In contrast, Asp and Cys have shifted pK a to lower values [95, 96, 102, 152, 225, 245–248]. An exception for negatively charged residues at the N-cap is that they have a lower pK a [29]. This may be because side chains at the N-cap can form strong hydrogen bonds to NH groups of N2 and N3, while the bonds formed by side chains at N1, N2, and N3 are much weaker [14, 19]. The negatively charged residues at higher pH destabilize helices when at the Ccap [29]. The increased pK a may result from an unfavorable electrostatic interaction with the C-terminal dipole or partial negative charges on the terminal CO groups. 5.11 Relevance to Proteins

Many of the features studied in peptide helices are also applicable to proteins and can be used to rationally modify protein stability or to design new helical proteins. Helices in proteins are often found on the surface with one face exposed to solvent and the other buried in the protein core. Helix propensities and sidechain interactions measured in peptides are thus directly applicable to the solventexposed face. Substitutions at buried positions are much more complex and tertiary interactions also make major contributions to stability. Tertiary interactions at helix

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We thank all our coworkers in this field, namely Avi Chakrabartty, Carol Rohl, Buzz Baldwin, Tod Klingler, Ben Stapley, Jim Andrew, Eleri Hughes, Simon Penel, Duncan Cochran, Nicoleta Kokkoni, Jia Ke Sun, Jim Warwicker, Gareth Jones, and Jonathan Hirst. Current work in our lab on helices is supported by the Wellcome Trust (grant references 057318 and 065106). TMI thanks the Indonesian government for a scholarship.

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1

Design and Stability of Peptide β-sheets Mark S. Searle University of Nottingham, Nottingham, United Kingdom

Originally published in: Protein Folding Handbook. Part I. Edited by Johannes Buchner and Thomas Kiefhaber. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30784-2

1 Introduction

The pathway by which the polypeptide chain assembles from the unfolded, “disordered” state to the final active folded protein has been the subject of intense investigation. Hierarchical models of protein folding emphasize the importance of local interactions in restricting the conformational space of the polypeptide chain in the search for the native state. These nuclei of structure promote interactions between different parts of the sequence leading ultimately to a cooperative ratelimiting step from which the native state emerges [1–4]. Designed peptides that fold autonomously in water (α-helices and β-sheets) have proved extremely valuable in probing the relationship between local sequence information and folded conformation (the stereochemical code) in the absence of the tertiary interactions found in the native state of proteins. This has allowed intrinsic secondary structure propensities to be investigated in isolation, and enabled the nature and strength of the weak interactions relevant to a wide range of molecular recognition phenomena in chemistry and biology to be put on a quantitative footing. While the literature is rich in studies of α-helical peptides [5–8], water-soluble, nonaggregating monomeric β-sheets have emerged relatively recently [9–11]. For reasons of design and chemical synthesis, these are almost exclusively antiparallel β-sheets, although others have used nonnatural linkers to engineer parallel strand alignments [12–14]. Here we focus on contiguous antiparallel β-sheet systems. Autonomously folding β-hairpin motifs, consisting of two antiparallel β-strands linked by a reverse β-turn (Figure 1), represent the simplest systems for probing weak interactions in β-sheet folding and assembly, although more recently a number of three- and four-stranded β-sheet structures have been described. From this growing body of data, key factors have come into focus that are important in rational Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Design and Stability of Peptide $-sheets

Fig. 1 a) Beta-strand alignment and interstrand hydrogen bonds in an antiparallel $-hairpin peptide; R groups represent amino acid side chains, main chain N and angles are shown. N-terminal $-hairpin of ubiquitin in which the native TLTGK turn se-

quence has been replaced by the sequence NPDG. b) Native strand alignment giving rise to a type I NPDG turn. c) Nonnative strand alignment with a G-bulged type I turn (NPDGT).

design. The following will be considered: the role of the β-turn in promoting and stabilizing antiparallel β-sheet formation; the role of intrinsic backbone φ, ψ propensities in preorganizing the extended conformation of the polypeptide chain; the role of cooperativity in β-sheet folding and stability and in the propagation of multistranded β-sheets, and quantitative approaches to estimating the energetics of β-sheet stability and folding. 2 β-Hairpins Derived from Native Protein Sequences

The early focus on peptides excised from native protein structures provided the first insights into autonomously folding β-hairpins, preceding the more rational

3 Role of $-turns in Nucleating $-hairpin Folding

approach to β-sheet design. Peptides derived from tendamistat [15], B1 domain of protein G [16, 17], ubiquitin [18, 19], and ferredoxin [20] showed that these sequences could exist in the monomeric form without aggregating, but that in most cases they showed a very limited tendency to fold in the absence of tertiary contacts. The use of organic co-solvents appeared to induce native-like conformation [17, 18, 20]. The study by Blanco et al. [20] of a peptide derived from the B1 domain of protein G provided the first example of native-like folding in water of a fully native peptide sequence. In contrast, the studies of hairpins isolated from the proteins ubiquitin and ferredoxin, which are structurally homologous to the G B1 domain (all form an α/β-roll fold) showed these to be unfolded in water [18, 20]. More recent studies of the native ubiquitin peptide have now revealed evidence for a small population of the folded state in water [21], while a β-turn mutation has been shown to significantly enhance folding [22]. The apparent lack of evidence for folding of the peptide derived from residues 15–23 of tendamistat (YQSWRYSQA) [15], and from the N-terminal sequence of ubiquitin (MQIFVKTLTGKTITLEV) [19] led to partial redesign of the sequence to enhance folding through modification to the β-turn sequence by introducing an NPDG type I turn, which is the most common type I turn sequence in proteins. Thus, in the former peptide SWRY was replaced by NPDG [15], and in the latter the G-bulged type I turn TLTGK was replaced by NPDG [19]. By introducing this tight two-residue loop across PD it was envisaged that both hairpins would be stabilized. This was certainly the case, however, the most striking observation was that both peptides folded into nonnative conformations with a three-residue G-bulged type I turn (PDG) reestablished across the turn (Figure 1b and 1c). These initial studies in rational redesign led to strikingly irrational results, prompting a much more systematic approach to β-hairpin design. The above results revealed that the βturn sequence, which dictates the preferred backbone geometry, appears to be an important factor in dictating the β-strand alignment. From the protein folding viewpoint, as demonstrated by the redesigned ubiquitin hairpin sequence, it is evident that one important role of the native turn sequence may be to preclude the formation of nonnative conformations that may be incompatible with formation of the native state.

3 Role of β-turns in Nucleating β-hairpin Folding

The systematic classification of β-turns in proteins reveals a wide variety of geometries and sizes of loop [2, 23, 24]. For the design of β-hairpins, the emphasis has been on incorporation of the smallest turn sequence possible to limit the entropic destabilization effects. While two-residue type I and type II turns are generally common in β-turns, the backbone conformation (φ and torsion angles) of a two-residue type I or type II turn results in a local left-handed twist, which is not compatible with the right-handed twist found in protein β-sheets. Consequently, these turns are less commonly found between contiguous antiparallel β-strands. However, the diastereomeric type I and II turns have the φ and and angles complementary

3


to the right-handed twisted orientation of the β-strands. These conclusions appear to rationalize, at least in part, the above observations with β-turn modifications introduced into the β-hairpins of tendamistat and ubiquitin. The introduced NPDG type I turn is not compatible with the right-handed twist of the β-strands resulting in a refolding to a more flexible G-bulged type I turn. The work of Gellman et al. [25] has shown the importance of backbone φ, angle preferences for the residues in the turn sequence by comparing the stability of a number of β-hairpin peptides derived from the ubiquitin sequence (MQIFVKSXXKTITLVKV) containing either XX = L-Pro-Xaa or D-Pro-Xaa. Replacing L-Pro with D-Pro switches the twist from left-handed (type I or II) to right-handed (type I or II ), making the latter compatible with the right-handed twist of the two β-strands. NMR data (Hα shifts and long-range nuclear Overhauser effects (NOEs)) indicate that each of the D-Pro containing peptides showed a significant degree of folding, whereas the L-Pro analogs appeared to be unfolded. Similar conclusions were drawn from studies of a series of 12-mers containing XG turn sequences, with X = L-Pro or D-Pro [26]. A number of natural L-amino acids are commonly found in the α L region of conformational space and are compatible with the type I or II turn conformation. Statistical analyses from a number of groups have identified Xaa-Gly as a favored type I turn. A number of studies have used the Asn-Gly sequence to design βhairpin motifs and have demonstrated a high population of the folded structure with the required turn conformation and strand alignment [27–32]. The work by de Alba et al. [28] identified two possible conformations of the peptide ITSYNGKTYGR. The NOE data appear to be compatible with rapidly inter-converting conformations involving an YNGK type I turn and a NGKT type II turn, each giving a distinct pattern of cross-strand NOEs. Ramirez-Alvarado et al. [27] also described an NGcontaining 12-mer (RGITVXGKTYGR; X = N), which they subsequently extended to a series of hairpins to examine the correlation between hairpin stability and the database frequency of occurrence of residues in position X [29]. Using nuclear magnetic resonance (NMR) and circular dichroism (CD) measurements they concluded that X = Asn > Asp > Gly > Ala > Ser in promoting hairpin folding, in agreement with the intrinsic φ, preferences of these residues. Despite extensive analysis of NG type I turns in the protein database (PDB), it is still not entirely clear why Asn at the first position is so effective in promoting turn formation. There is no evidence for specific side chain to main chain hydrogen bonds that might stabilise the desired backbone conformation, although specific solvation effects cannot be ruled out [30]. Evidence that the NG turn is able to nucleate folding in the absence of cross-strand interactions was demonstrated in a truncated analog of one designed 16-residue β-hairpin sequence KKYTVSINGKKITVSI (β1 in Figure 2), in which the sequence was shortened to SINGKKITVSI, lacking the N-terminal six residues [30]. Evidence from NOE data showed that the turn was significantly populated with interactions observed between Ser6 Hα ↔ Lys11 Hα and Ile7 NH ↔ Lys10 NH (Figure 2b) that are only compatible with a folded type I turn around NG. Two destabilized β-hairpin mutants (KKYTVSINGKKITKSK with electrostatic repulsion between the N- and

3 Role of $-turns in Nucleating $-hairpin Folding

Fig. 2 Structure of the 16-residue $-hairpin sequence $1, and mutants $2;,$3 and $4 in which cross-strand salt bridges (Lys-Glu) have been introduced at position X2 -X15 and X6 -X11 . In b), hairpin $1 has been truncated by removing the N-terminal hydrophobic

residues (1–5). The resulting peptide still shows the ability to fold around the turn sequence as evident from medium range NOEs (indicated by arrows) that show that the INGK sequence is adopting a type I NG turn.

C-terminal Lys residues, and KKATASINGKKITVSI with the loss of key hydrophobic residues in one strand) showed no evidence from NMR chemical shift data for cross-strand interactions; however, careful examination of NOE data revealed evidence for NG turn nucleation [30]. Titration with organic co-solvent showed both peptides to fold significantly, indicating that the turn sequence probably already predisposes the peptide to form a β-hairpin but that favorable cross-strand interactions are required for stability. The work of de Alba et al. convincingly illustrated this principle in a series of six hairpin sequences (10-mers) where strand residues were conserved but turn sequences varied [28]. Using a number of NMR criteria they were able to show that changes in turn sequence could result in a variety of turn conformations including two residue 2:2 turns, 3:5 turns and 4:4 turns (see earlier nomenclature [22–24]), with different pairings of amino acid side chains. As with the earlier examples cited with the turn modification described for hairpins derived from tendamistat and ubiquitin, the bulged-type I turn (3:5 turn) appears to be an intrinsically stable turn with the necessary right-handed twist. Together these data strongly support a model

5


for hairpin folding in which the turn sequence strongly dictates its preferred conformation, and that strand alignment, cross-strand interactions and subsequently conformational stability are dictated by the specificity of the turn.

4 Intrinsic φ, Propensities of Amino Acids

Statistical analyses of high-resolution structures in the PDB have provided significant insights into residue-specific intrinsic backbone φ; ψ preferences in polypeptide chains. A novel approach presented by Swindells et al., was to determine φ, ψ propensities of different residues in nonregular regions of protein structure where backbone geometry is free of interactions associated with regular hydrogen bonded β-sheet or α-helical secondary structure [33]. The striking observation is that in this context φ and ψ angles (see Figure 3) are far from randomly distributed, and that most occupy regions of Ramachandran space associated with regular secondary structure. The observed φ; ψ distributions for individual residues has been taken as representative of those found in denatured states of proteins providing the basis of a “random coil” state from which residue-specific NMR parameters (3 JNH-H α and NOE intensities) can be derived as a reference state for folding studies [34, 35].

Fig. 3 Ramachandran plots of residue backbone N and angles taken from a database of 512 high-resolution protein Xray structures showing: a) residues in regular $-sheet (N, −120◦ , 120◦ ) and "-helix (N;, −60◦ , 60◦ ), and b) residues in the irregular coil regions of the same structures

(N, 60◦ , 0◦ is the "L region of conformationa space mainly occupied by Gly). The distribution in b) shows that residues have a natura propensity to occupy the " and $ regions of conformational space even when they are not involved in regular protein secondary structure. Taken from Ref [36].

4 Intrinsic N, Propensities of Amino Acids 7

While β-propensity is found to vary significantly from one residue to the next, context-dependent effects also appear to play an important part [36]. While V, I, F, and Y, for example, have a high intrinsic preference to be in the βregion of the Ramachandran plot where steric interactions with flanking residues are minimized, their conformation is relatively insensitive to the nature of the flanking residues. In contrast, small or unbranched side chains have a higher preference for the α-helical conformation, however, this preference can be significantly modulated by its neighbors through a combination of steric and hydrophobic interactions, as well as both repulsive and attractive electrostatic interactions. General effects of flanking residues (grouped as α-like or β-like, reflecting φ propensities) on the central residue of a XXX triplet are illustrated in Figure 4 as an average over all residues at the central position. More specific effects are also shown for

Fig. 4 Effects of neighboring residues on residue $-propensities within the triplets XNX calculated from the data in Figure 3. The $-propensity of residue N is a measure of the number of times a particular residues is found in the $-region of the Ramachandran plot as a fraction of the total distribution between "- and $-space [$/(" + $)]. The context dependence of the $-propensity is estimated by considering the nature of the neighboring residue (X) (X = any residue, " is a residue that prefers the "-helical region – Asp, Glu, Lys, or Ser, $ is

a residue that prefers the $-sheet region – Ile, Val, Phe, or Tyr). The effects of neighboring residues on the averages-propensity is shown in (a), specific effects on Ser (b), Val (c) and Lys (d) are also shown. While Val is relatively insensitive to the nature of the flanking residues, the smaller Ser residue can be forced to adopt a higher $propensity if it has bulky neighbors. Thus, the intrinsic $-propensity of a particular residue is highly context dependent. Taken in part from Ref. [36].


Ser, Val, and Lys (Figure 4). With Ser, for example, having bulky flanking residues either side with high β-propensity (denoted βSβ, where β could be V, I, F or Y), significantly increases the β-propensity of the Ser residue to minimize the steric repulsion between the two bulky neighboring residues [36]. Thus, intrinsic structural propensities appear to be highly context dependent. This statistical framework has been extended to a number of experimental systems to examine the extent to which isolated β-strand sequences (in the absence of secondary structure interactions) are predisposed by the primary sequence to adopt an extended β-like conformation. The isolated 8-mer (GKKITVSI), corresponding to the C-terminal β-strand of the hairpin β1 (Figure 2; KKYTVSINGK-KITVSI), was examined by NMR analysis of 3 JNH-Hα values and backbone NOE intensities. Surprisingly, many of these parameters are similar to those for the folded hairpin despite the monomeric nature of the 8-mer [32, 36]. In an analogous study of the C-terminal strand of the ubiquitin hairpin described above [21, 37], similarly large deviations of coupling constants and NOE intensities from random coil values suggested that the isolated β-strands are partially preorganized into an extended conformation supporting a model for hairpin folding which may not require a significant further organisation of the peptide backbone, a factor that may contribute significantly to hairpin stability. Several studies of denaturated states of proteins have also highlighted the influence of neighboring residues in modulating main chain conformational preferences [38–41], and the importance of residual structure in the unfolded state in guiding the conformational search to the native state [42, 43].

5 Side-chain Interactions and β-hairpin Stability

There is a strong case, at least in the context of isolated peptide fragments, that the origin of the specificity of β-hairpin folding is largely dictated by the conformational preferences of the turn sequence. However, the stability of the folded state has been attributed to interstrand hydrogen bonding and/or hydrophobic interactions, though which dominates is still a matter of debate. Ramirez-Alvarado et al. [27] reported that the population of the folded state of the hairpin RGITVNGKTYGR was significantly diminished by replacing residues on the N-terminal strand, and then the C-terminal strand, by Ala. The loss of stability was attributed firstly to a reduction in hydrophobic surface burial, but also due to the intrinsically lower β-propensity of Ala, the latter contributing through an adverse conformational entropy term. To compensate for this de Alba et al. [44] described a family of hairpins derived from the sequence IYSNSDGTWT. The effects of residue substitutions in the first three positions was examined while maintaining the overall β-character of the two strands. Several favorable cross-strand pair-wise interactions were identified that were apparent in earlier, and more recent, PDB analysis of β-sheet interactions [45, 46]. For example, Thr-Thr and Tyr-Thr cross-strand pairs produced stabilizing interactions, whilst Ile-Thr and Ile-Trp had a destabilizing effect. Other studies of

5 Side-chain Interactions and $-hairpin Stability

Ala substitution in one β-strand have similarly highlighted hydrophobic burial as a key factor in conformational stability [30], while the observation of large numbers of side chain NOEs have been used as evidence for hydrophobic stabilization in water (see below) [19, 25–27, 30]. 5.1 Aromatic Clusters Stabilize β-hairpins

The first example of a natural β-hairpin sequence that folded autonomously in water (residues 41–56 of the B1 domain of protein G) identified an aromatic-rich cluster of residues that appears to impart considerable stability [17]. The interstrand pairing of Trp/Val and Tyr/Phe has subsequently been exploited in the design of a number of model hairpin systems, in particular to examine the relationship between the position of this stabilizing cluster and the β-turn sequence. The separation between the loop sequence and cluster strongly influences stability and the extent of participation in β-sheet forming interactions (Figure 5a). In an isomeric family of 20-mers (peptides 1, 2, and 3), all of which contain exactly the same residue composition, the most stable hairpin is that in which the smallest cost in conformational entropy is paid to bring the cluster together, i.e., where the cluster is closest to the loop sequence [47]. This arrangement results in the largest Hα chemical shift deviations, indicating a well-formed core; however, the terminal residues show a lower propensity to fold. Thus, there exists a strong interplay between the two key stabilizing components of the hairpin. A statistical model was developed to rationalize the experimental observations and estimate the free energy change versus the number of peptide hydrogen bonds formed (Figure 5b). It is a well-known phenomenon that α-helical peptides become more stable as the length increases, reflecting the fact that while helix nucleation is energetically unfavorable, the propagation step has a small net increase in stability. In β-hairpin systems, Stanger et al. [48] suggest that this may not be the case. There is some evidence for an increase in stability as strands lengthen from five to seven residues, however, further extension (to nine) does not lead to a further stability increment, suggesting that there may be an intrinsic limit to strand length. Since the choice of sequence extension in this study was limited to an all Thr extension or an alternating Ser-Thr (ST)n extension the conclusions should be viewed with caution. Since crossstrand Ser/Ser and Thr/Thr pairings do not bury very much hydrophobic surface area, it is not surprising perhaps that cross-strand side-chain interactions may only just compensate for the entropic cost of organizing the peptide backbone. Studies with other more favorable pairings may be enlightening. Undoubtedly the most successfully designed structural motif to date has been the tryptophan zipper (trpzip) motif [49], whose stability exceeds substantially all those already described. The design is based around stabilizing nonhydrogen-bonded cross-strand Trp-Trp pairs (Table 1). The most successful designs involved two such Trp-Trp pairs which NMR structural analysis reveals are interdigitated in a zipper-like manner stabilized through face-face offset π-stacking giving a compact structure (Figure 6). This arrangement

9


Fig. 5 Interplay between hydrophobic cluster and turn position in a family of isomeric $-hairpins. The distance of the WVYF cluster from the turn is shown in (a) for peptides 1, 2, and 3. A statistical mechanical model was used to estimate the free energy of folding for the three peptides according to the number of peptide hydrogen bonds formed and the relative position of the stabilizing hydrophobic cluster (b). Thus, peptide 1, which has the hydrophobic cluster

closest to the turn, has the smallest energetic cost of forming the cluster which then significantly stabilizes this core motif, but leads to fraying of the N- and C-termini. In contrast, in the case of peptide 3, there is a large energy penalty in ordering the peptide backbone to bring the residues of the cluster together. Overall the core hairpin is less stable but more residues are involved in ordered structure. Adapted from Ref. [47].

5 Side-chain Interactions and $-hairpin Stability Table 1 trpzip β-hairpin peptides, sequences and stability

β-hairpin

Sequence

Turn type

Tm

H (kJ mol−1 )

trpzip1 trpzip2 trpzip3

SWTWEGNKWTWK SWTWENGKWTWK SWTWED PNKWTWK

(Type II turn) (Type I turn) (Type II turn)

323 345 352

−45.4 −70.6 −54.8

Fig. 6 NMR structures of trpzips 1 and 2. A) Ensemble of 20 structures of trpzip1 (residues 2–11) showing the relative orientations of the indole rings. B) Overlay of trpzip 1 and 2 aligned to the peptide backbone of residues 2–5 and 8–11 (top),

and rotated by 90◦ for the end-on view of the indole rings. The backbone carbonyl of residue 6 is labeled to illustrate the different turn geometries of the two hairpins (type II and type I ). Taken from Ref [49].

11


of the indole rings results in a pronounced signature in the CD spectrum with intense exciton-coupling bands at 215 and 229 nm indicative of interactions between aromatic chromophores in a highly chiral environment (see Figure 7). The high sensitivity of the CD bands permits thermal denaturation curves to be determined with high sensitivity, in contrast to other β-sheet systems where only small changes in the CD spectrum at 216 nm are evident, resulting in poor signal-to-noise. The thermal unfolding curves are sigmoidal and reversible, fitting to a two-state folding model (Figure 7). The trpzip hairpins 1, 2, and 3 show high T m values (see Table 1) with folding strongly enthalpy driven. Changing the turn sequence (GN versus NG versus DPN) has a significant impact on stability. The unfolding curve for trpzip2 with the NG type I turn gives the most cooperative unfolding transition and appears to be the most stable of the three at room temperature, despite other studies that suggest that the unnatural D-PN (type II ) turn is the most stabilizing [25]. Clearly, context-dependent factors are at work. Also, the strongly enthalpy-driven signature for folding is different to that reported previously for other systems [32], reflecting differences in the nature of the stabilizing interactions which in the case of the trpzip peptides involve π-π stacking interactions. Despite the fact that the trpzip hairpins represent a highly stabilized motif, no such examples have been found in the protein structure database. The authors suggest that on steric grounds it may be difficult to accommodate the trpzip motif within a multistranded β-sheet or through packing against other structural elements. 5.2 Salt Bridges Enhance Hairpin Stability

The high abundance of salt bridges on the surface of hyperthermophilic proteins [50, 51], together with examples of rational enhancement of protein stability through redesign of surface charge, has strongly implicated ionic interactions as a stabilizing force [52, 53]. In model peptides that are only weakly folded in water, the relative importance of ionic versus hydrophobic interactions in stabilizing local secondary structure has been less well investigated in terms of a detailed quantitative description. To this end, peptide β1 (Figure 2) was mutated to introduce Lys-Glu salt bridges at two positions within the hairpin involving substitution of Ser → Glu: one salt bridge is postioned adjacent to the β-turn, while the other involves residues close to the N- and C-termini (see Figure 2) [54]. Using NMR chemical shift data from Ha resonances we have estimated the net contribution to stability from these two interactions. Although the contribution to stability in each case is small, Hα chemical shifts are extremely sensitive to small shifts in the population of the folded state, as evident from the δHα data in Figure 8. On this basis, the individual contributions of these two interactions was estimated (K2-E15, peptide β2, and E6-K11, peptide β3) compared with their K2-S15 and S6-K11 counterparts (peptide β1) and found to be similar (−1.2 and −1.3 kJ mol−1 at 298K). When the two salt bridges are introduced simultaneously into the hairpin sequence (β4), the energetic contribution of the two interactions together (−3.6 kJ mol−1 ) is


Fig. 7 Folding data for trpzips 1 to 3 determined from near- and far-UV CD spectra. a) CD spectrum of trpzip 1, with inset of nearUV CD showing buried aromatic residues (10-fold expansion). b) Reversible thermal

denaturation of trpzip 1 monitored by CD at 229 nm (unfolding and refolding curves overlayed). c) Temperature dependence of folding for trpzips 1 to 3 plotted as fraction folded. Taken from Ref. [49].

13


Fig. 8 Effects of salt bridges (Lys-Glu) on $-hairpin stability. H" chemical shift deviation from random coil values (*H" values) for $-hairpin peptides $2 to $4 compared with those of the reference hairpin

protein $1 containing Lys-Ser cross-strand pairs at the X2 -X15 and X6 -X11 mutation sites (see Figure 2). a) $1 versus $2; b) $1 versus $3, and c) $1 versus $4. All data at 298K and pH 5.5. Taken from Ref. [54].


significantly greater than the sum of the individual interactions. This effect is readily apparent from the large increase in -δ-Hα values in Figure 8, indicating that the contribution of a given interaction appears to depend on the relative stability of the system in which the interaction is being measured. Similar observations have been reported using a disulfide cyclized β-hairpin scaffold [55]. The indication is

Fig. 9 a) Thermodynamic cycle showing the context-dependent energetic contribution to hairpin stability of each electrostatic interaction in the $1 to $4 family of hairpins (Figure 2) by comparing relative hairpin stabilities at 298K determined from NMR chemical shift data; G values are shown for Lys-Glu interactions at pH 5.5. The stabilizing contribution of each salt bridge is context dependent, showing some degree

of cooperativity between the two mutation sites according to the degree of preorganization of the hairpin structure. b) Schematic illustration of the abundance of cross-strand NOEs in hairpin peptide $4. c) Family of five NMR structures of hairpin $4 showing the peptide backbone alignment, with some fraying at the N- and C-termini. Taken from Ref. [54].

15


that the strength of the interaction appears to depend on the degree of preorganization of the β-hairpin template that pays varying degrees of the entropic cost in bringing the pairs of side chains together. This is more clearly illustrated by the schematic representation shown in Figure 9 that shows the thermodynamic cycle indicating the effects on stability of the introduction of each mutation. Thus, the energetic contribution of the S15/E15 mutation could be measured either from β1 ←→ β2 or from β3 ←→ β4. The values obtained are quite different, the latter suggesting that the interaction is more favorable (−1.2 versus −2.3 kJ mol−1 ). This appears to correlate with the fact that the stability of β3 is greater than β1 and that the degree of preorganization of the former determines the energetic contribution of the interaction. The same principle is evident when determining the energetics of the E6-K11 interaction. This can be estimated from β1 ←→ β3 or β2 ←→ β4. Again, the energetics are quite different (−1.3 versus −2.4 kJ mol−1 ) with the larger contribution from the β2 ←→ β4 pair, where the intrinsic stability of the β2 reference state is higher. The NMR data show that the folded conformation of β4 is highly populated (> 70%) giving rise to an abundance of cross-strand NOEs (Figure 9) [55]. On the basis of 173 restraints (NOEs and torsion angle restraints from 3 JNH-Hα values) we have calculated a family of structures compatible with the NMR data (Figure 9). The large number of van der Waals contacts between hydrophobic residues (V, I, and Y)

Fig. 10 CD melting curves for $4 at various concentrations (% v/v) of MeOH, as indicated. The nonlinear least squares fit is shown in each case from which thermodynamic data for folding were determined

(see Ref. [54]). The change in heat capacity on folding in 30% and 50% aqueous methanol is assumed to be zero; there is evidence for cold-denaturation in water and 10% methanol. Taken from Ref. [54].

7 Quantitative Analysis of Peptide Folding 17

evident from the NOE data leads us to conclude that hydrophobic interactions still provide the overall driving force for folding with structure and stability further consolidated by additional Coulombic interactions from the salt bridges. CD melting curves for hairpin β4 in water and various concentrations of methanol (Figure 10) show the same characteristics described above from temperature-dependent NMR data for β1: a shallow melting curve for β4 in water with evidence for cold denaturation is consistent with the hydrophobic interaction again providing the dominant driving force for folding (H = +11:9 kJ mol−1 and S = +38 J K−1 mol−1 ), while folding becomes strongly enthalpy driven in 50% aqueous methanol (H = −39:8 kJ mol−1 and S = −106 J K−1 mol−1 ).

6 Cooperative Interactions in β-sheet Peptides: Kinetic Barriers to Folding

Folding kinetics for a β-hairpin derived from the C-terminus of the B1 domain of protein G have been described from measurements of tryptophan fluorescence following laser-induced temperature-jump [56, 57]. Kinetic analysis of this peptide (and a dansylated analogue) reveals a single exponential relaxation process with time constant 3.7 ± 0:3 µs. The data indicate a single kinetic barrier separating folded and unfolded states, consistent with a two-state model for folding. Subsequently, the authors developed a statistical mechanical model based on these observations, describing the stability in terms of a minimal numbers of parameters: loss of conformational entropy, backbone stabilization by hydrogen bonding and formation of a stabilizing hydrophobic cluster between three key residues. This model seems sufficient to reproduce all of the features observed experimentally, with a rough, funnel-like energy landscape dominated by two global minima representing the folded and unfolded states. The formation of the hydrophobic cluster appears to be a key folding event. Nucleation by the turn seems most likely, consistent with experimental measurements of loop formation on the timescale of ≈1 µs [58]. However, simulation studies by others suggest that folding may proceed by hydrophobic collapse followed by rearrangement to form the hydrophobic cluster, with hydrogen bonds then propagating outward from the cluster in both directions [59]. Such a model does not appear to require a turn-based nucleation event.

7 Quantitative Analysis of Peptide Folding

Quantitative analysis of the population of folded β-sheet structures in solution still presents a challenge, largely as a consequence of uncertainties in limiting spectroscopic parameters for the fully folded state. Far UV-CD has been considered to be unreliable as a consequence of the complicating influence of the β-turn conformation and possibly aromatic residues, where present [10]. Added to this, the CD spectrum of β-sheet is intrinsically weak compared with α-helical secondary structure.


The trpzip peptides [49] represent the exception to the rule, as discussed above. The use of NMR parameters (Hα chemical shifts, 3 JNH-Hα values and NOE intensities) to quantify folded populations has been discussed [9–11, 32, 60]. NMR offers the advantage that several independent parameters can be used in quantitative analysis to provide a consensus picture of the folded state. There still appear to be significant discrepancies between CD analysis and NMR, with peptides that appear to be significantly folded by NMR giving rise to a largely random coil CD spectrum. One interpretation of this observation is that in aqueous solution the peptides fold as a collapsed state with an ill-defined hydrogen bonding network dominated by sidechain interactions. Interestingly, in many cases the addition of organic co-solvents changes the CD spectrum dramatically. The interpretation of co-solvent-induced folding is also subject to some uncertainty. Does trifluorethanol or methanol actually significantly perturb the equilibrium between the folded and unfolded states (induce folding), or do these solvents exert their influence by changing the nature of the folded state such as to stabilize interstrand hydrogen bonding interactions without significantly changing the folded population? The latter hypothesis would appear to more readily account for the observation of solvent-induced effects on the CD spectrum, and finds some support from studies of cyclic β-hairpin analogs where the folded population is fixed, but whose CD spectrum undergoes large solvent-induced changes [60]. The use of cyclic β-hairpin analogs has been exploited in a number of studies to generate a fully folded NMR reference state for comparison with the folding of acyclic analogs [61, 62]. Backbone cyclization through amide bond formation or through a disulfide bridge seem to work equally well. Such an approach has been used effectively to measure the thermodynamics of folding of a short hairpin carrying a motif of aromatic residues [62]. The cyclic analogs show a much higher stability than their acyclic counterparts, including significant protection from amide H/D exchange due to enforced interstrand hydrogen bonding. While peptide cyclization seems a worthwhile approach to defining the fully folded state, there may also be some caveats to this approach that have not been tested. When conformational constraints (β-turns) are imposed at both ends of the structure this may affect the intrinsic twist of the two β-strands, resulting in a more pronounced twist in the cyclic analog than in the acyclic hairpin. Further, the latitude for conformational dynamics in the fully folded state will also be different. Both of these factors are likely to influence Hα shifts chemical shifts to some degree.

8 Thermodynamics of β-hairpin Folding

The number of β-hairpin model systems is expanding rapidly with a greater focus now on quantitative analysis and thermodynamic characterization. The thermodynamic signature for the folding of β1 (Figure 2) presents an insight into the nature of the stabilizing weak interactions in various solvent milieu [30, 32]. β1 exhibits

8 Thermodynamics of $-hairpin Folding

Fig. 11 a) Temperature-dependent stability of the hairpin peptide $1 (see Figure 2) at pH 5.5 in water, 20% methanol and 50% methanol. Temperature is plotted against the RMS value for the deviation of H" chemical shifts from random coil values, assuming a two-state folding model. In water the peptide shows “hot” and “cold” denaturation, but in 50% methanol the folded population increases at low temperature. The best fits to the three sets of data are shown; in water folding is slightly entropydriven with a large negative C◦ p value, while in 50% methanol folding is strongly

enthalpy-driven with C◦ p close to zero. In 20% methanol the values are in between. (Reproduced with permission from the Journal of the American Chemical Society [32]). b) Variable-temperature FTIR spectra of i) 2 mM aqueous solution of the 8-mer peptide GKKITVSI (C-terminal $-strand of hairpin $1); ii) 2 mM solution of $-hairpin peptide $1; iii) 10 mM solution of $1, all in the temperature range 278–330 K and in D2 O solution, pH 5.0 (uncorrected). In iii) the band that appears around 1620 cm−1 is formed irreversibly indicative of peptide aggregation. Reproduced from Ref [64].

the property of cold denaturation, with a maximum in the stability curve occurring at 298K as judged by changes in Hα chemical shift (Figure 11). Such pronounced curvature is clear evidence for entropy-driven folding accompanied by a significant change in heat capacity. Both of these thermodynamic signatures are hallmarks of the hydrophobic effect contributing strongly to hairpin stability. More recent studies by Tatko and Waters [63] have also shown that cold denaturation effects can be observed for simple model peptides, consistent with substantial changes in heat capacity on folding. Further examination of folding in the presence of methanol co-solvent shows that the signature changes such that folding becomes strongly enthalpy driven, and that in 50% aqueous methanol the temperaturestability profile is indicative of the absence of any significant contribution of the hydrophobic effect to folding [32]. The population of the folded state appears to be enhanced by methanol, reflecting similar observations in helical peptides where the phenomenon has been attributed to the effects of the co-solvent destabilizing

19


the unfolded peptide chain so promoting hydrogen bonding interactions in the folded state. It is unlikely that the folded state of a model β-hairpin peptide resembles a β-sheet in a native protein, with the former sampling a much larger number of conformations of similar energy stabilized by a fluctuating ensemble of transient interactions. IR analysis of the amide I band of β1 does not show significant differences in the region expected for β-sheet formation (≈ 1630 cm−1 ) from data on a nonhydrogen-bonded short reference peptide [64]. However, this band does appear under aggregating conditions (Figure 11). In other cases, IR spectral features characteristic of β-hairpins have been shown to form reversibly at relatively high peptide concentrations [65]. Similarly, β-hairpins that appear to be well folded on the basis of various NMR criteria seem to be weakly folded by CD analysis [60]. This discrepancy has been attributed largely to weak interstrand hydrogen bonding interactions in the folded state. Indeed, molecular dynamics simulations using ensemble-averaging approaches or time-averaged NOEs tend to de-emphasize the role of hydrogen bonding between the peptide backbone of the two strands, but emphasize the role of hydrophobic side chains interactions [30, 60, 67]. In all cases described, side chain NOEs across the β-strands support such interactions, however, cross-strand backbone NOEs only imply the possibility of hydrogen bonds since weakly populated folded states lead to only small NH ←→ ND protection factors in water. In studies (both calorimetric and NMR) of the folding of the C-terminal hairpin from the B1 domain of protein G, enthalpy-driven folding is observed in water [68]. In contrast with the above data, where the stabilizing hydrophobic interactions involved aliphatic side chains, here an aromatic cluster is responsible for folding. A strongly enthalpy-driven transition is also apparent for the trpzip motifs described above [49] all of which is consistent with π-π interactions stabilizing β-hairpin structures through fundamentally electrostatic interactions rather than through only solvophobic effects. Studies of a designed β-hairpin system, also carrying the same motif of three aromatic residues, demonstrate qualitatively similiar enthalpydriven folding [62], while the thermodynamics of the N-terminal hairpin component of a designed three-stranded antiparallel β-sheet enables similar conclusions to be drawn (see further below) [69]. The difference in the thermodynamic signature for aliphatic versus aromatic side-chain interactions has been suggested to have its origins in enthalpy-entropy compensation effects, such that enthalpy-driven interactions may be fundamentally a consequence of tighter interfacial interactions, giving rise to stronger electrostatic (enthalpic) interactions [62]. More recent work by Tatko and Waters [70] probing the energetics of aromatic-aliphatic interactions in hairpins suggests that there is a preference for the self-association of aromatics over aromatic-aliphatic interactions, and that the unique nature of this interaction may impart some degree of sequence selectivity. The limited data available from such systems suggest that the thermodynamic driving force for β-hairpin folding is highly dependent on the nature of the side-chain interactions involved. Further quantitative analysis of peptide folding is required to substantiate these hypotheses.

9 Multistranded Antiparallel $-sheet Peptides

9 Multistranded Antiparallel β-sheet Peptides

The natural extension of the earlier studies on β-hairpin peptides was to design three- and four-stranded antiparallel β-sheets using the design principles already discussed, focusing on the importance of turn sequence in defining stability and strand alignment, and employing motifs of interacting side chains already identified to impart stability. An overriding question concerns the extent to which cooperative interactions perpendicular to the strand direction are important in stabilizing these structures (see Figure 12); in other words, how good is a pre-organized βhairpin motif at templating the interaction of a third strand. Several studies have attempted to address this important question. The earliest study described a 24residue peptide incorporating two NG turns (KKFTLSINGKKYTISNGKTYITGR) that showed little evidence for folding in water but was significantly stabilised in aqueous methanol solutions [71, 72]. By comparison with a 16-residue β-hairpin analog consisting of the same C-terminal sequence (GKKYTISNGKTYITGR), it was possible to show that the Hα shift perturbations for the C-terminal β-hairpin were greater in the presence of the interactions of the third strand. Subsequent design strategies, incorporating the D-Pro-Gly loop, together with other Asn-Glycontaining turn sequences have illustrated that it is possible to design structures that fold in water [69, 73–75], some of which show a degree of cooperative stabilization between different strands. Other studies have reported peptides that fold to three- (and four-) stranded β-sheets in organic solvents [76], but a quantitative dissection of stabilizing interactions between β-strands has not been presented. Redesign of the three-stranded Betanova, following earlier work [74], reported the use of an automated approach using the algorithm PERLA (protein engineering rotamer library algorithm) to define stabilizing and destabilizing single and multipleresidue mutations producing incremental increases in stability over the original design of up to ≈4 kJ mol−1 [77]. Schenck and Gellman [73] demonstrated cooperative interactions using their D-ProGly to L-ProGly switch, the latter destabilizing one hairpin component selectively. From chemical shift analysis they showed that the individual β-hairpins are cooperatively stabilized by the presence of the third strand. De Alba et al. [75] also reported a designed β-sheet system, but were unable to find convincing evidence for cooperative stabilization of either hairpin by the third strand. It is clear that folding is not a highly cooperative process in these peptide systems. Thus, the folded three-stranded β-sheet is more than likely in equilibrium with populations of the individual hairpins and the unfolded state (see Figure 12). One quantitative study of cooperative interactions between the strands of a threestranded β-sheet [69] was based on a designed system incorporating a previously studied β-hairpin with the third strand capable of forming a stabilizing motif of aromatic residues (Figure 13), similar to that already described [17, 62]. The earlier study of the isolated C-terminal β-hairpin showed cold denaturation [30], approximating to two-state unfolding. In the designed three-stranded system, the same hairpin component shows the same cold denaturation profile, however, the N-

21


Fig. 12 a) Models for the folding of $hairpin and three-stranded $-sheet peptides illustrating the possibility of cooperative interactions being propagated parallel i) and perpendicular ii) to the $-strand direction. b) Four-state model for the folding of a three-stranded antiparallel $-sheet peptide showing the presence at equilibrium of the

unfolded state and partially folded states in which the C-terminal $-hairpin is formed but the N-terminus is disordered, the Nterminal $-hairpin is formed but the Cterminus is disordered, and the fully folded state. The preformed hairpins can act as templates for the folding of the third strand.

terminal hairpin (sharing a common central strand) showed increased folding at low temperature. While the first process is characterized by entropy-driven folding, the latter is enthalpy-driven (Figure 13). Clearly, two different thermodynamic profiles are not consistent with a single two-state folding model, but the data could be rationalized in terms of a four-state model in which the individual hairpins with an unfolded tail are also populated (Figure 12). Examination of the folding of the isolated C-terminal hairpin, and comparison of the data with that of the three-

9 Multistranded Antiparallel $-sheet Peptides

Fig. 13 a) Mean NMR structure of the fully folded state of the three-stranded antiparallel $-sheet structure of sequence KGEWTFVNG9 KYTVSING17 KKITVSI showing the core cluster of hydrophobic residues (underlined). b) Temperature-dependent stability profiles for the various $-hairpin components of the three-stranded sheet using the H" splitting of Gly9 and Gly17 in the two $-turns: Gly9 (filled circles), Gly17 (open circles). Gly17 (open squares)

in the isolated C-terminal hairpin peptide (G9 KYTVSING17 KKITVSI) is also shown. A larger Gly H" splitting indicates a higher population of the folded hairpin. Different profiles for Gly9 and Gly17 in the threestranded sheet show that the peptide cannot be folding via a simple two-state model involving only random coil and fully folded peptide. The data have been fitted assuming the four-state model shown in Figure 12. Reproduced from Ref. [69].

23


stranded analog, shows good evidence that the C-terminal hairpin is cooperatively stabilized by the interaction of the third strand, even though overall folding is not highly cooperative. The data in Figure 13 show the temperature dependence of the splitting of the Gly Hα resonances in the two NG turns. Larger values indicate a higher folded population. The splitting for G17 is greater in the three-stranded sheet than for the isolated C-terminal hairpin, while many Hα resonances are further downfield shifted than in the isolated hairpin. The temperature dependence of the stability shows the C-terminal hairpin in both cases to undergo cold denaturation. The N-terminal hairpin carrying the aromatic motif of residues increases in population at low temperature; fitting the data shows the former to be entropy driven and the latter enthalpy driven [69]. Entropic factors seem to be the likely explanation for the small cooperative stabilization effect on the C-terminal hairpin (< 2 kJ mol−1 ), with each hairpin providing a possible template against which the third strand can interact. With one strand preorganized, the entropic

Fig. 14 Structure of the four-stranded $-sheet peptide D PD PD P-cc (a), and the cyclic reference peptide c(D P)2 -II (b) used for estimating limiting values for the fully folded state of the C-terminal hairpin of D PD PD P-cc. Adapted from the work of Ref. [78].

10 Conclusions

cost of association of an additional strand is largely confined to the associating strand [73, 78]. The nature of the folded state is unlikely to compare with that of a β-sheet in a native protein, more likely, hydrophobic contacts between side chains stabilize a collapsed conformation where interstrand hydrogen bonds may play a minor stabilizing role. These “loosely” defined interactions between side chains, rather than a native-like “crystalline” array of hydrogen bonds, may explain why cooperative interactions have only a small effect on overall stability because only a small energy barrier separates the folded from partially or fully unfolded states. Several of the above studies have attempted to address the issue of whether cooperative interactions are propagated orthogonal to the strand direction as the number of β-strands increases. Gellman and coworkers have examined the influence of strand number on antiparallel β-sheet stability in designed three- and four-stranded structures (see Figure 14), using the D-Pro-Gly β-turn sequence to define β-turn position and strand length. The results are not dissimilar to those described above; a third strand stabilizes an existing two-stranded β-sheet by ≈2 kJ mol−1 , while a fourth strand seems to confer a similar small stability increment [79].

10 Conclusions

In contrast to the highly cooperative folding behavior characteristic of native globular proteins, simple model β-sheet systems, which lack defining tertiary interactions, do not possess this characteristic. The limited number of three- and fourstranded β-sheet peptides so far described confirms this, although some evidence for cooperative interactions between β-strands have been presented and rationalized on the basis of the entropic benefits associated with adding an additional β-strand to a preformed template. In all cases to date, it seems that designed threeand four-stranded β-sheet structures are in equilibrium with their partially folded β-hairpin components. This is an indication that the energy landscape is relatively flat unlike that for most native proteins where there is a significant energy difference between the single native structure and other conformations. It is interesting to look at examples of the smallest known β-sheet proteins and make comparisons with the designed systems described above. The WW domains [80–82] form a single folded motif consisting of three strands of antiparallel β-sheet but with well-defined tertiary interactions arising from folding back of the N- and C-termini to form a small compact hydrophobic core. This defining feature appears to allow the WW domains to fold co-operatively despite their small size (some as small as 35 residues) (Figure 15). Apart from the desire to be able to design molecules to order with specific tailored properties, the model βsheet systems described have enabled considerable progress to be made in testing our understanding of the basic design principles of β-sheets and fundamental aspects of weak interactions.

25


Fig. 15 NMR structure of the WW domain of the formin binding protein (PDB code: 1EOI), consisting of a three-stranded antiparallel $sheet motif; the side chains of conserved residues are shown with tertiary contacts evident between W8 and P33.

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42 KLEIN-SEETHARAMAN, J., OIKAWA, M., GRIMSHAW, S. B. et al. Science 2002; 295, 1719–1722. 43 SHORTLE, D., ACKERMAN, M. S. Science 2001; 293, 487–489. 44 De ALBA, E., RICO, M., JIMENEZ, M. A. Protein Sci. 1997; 6, 2548–2560. 45 WOUTERS, M. A., CURMI, P. M. G. Protein Struct. Funct. Genet. 1995; 22, 119–131. 46 HUTCHINSON, E. G., SESSIONS, R. B., THORNTON, J. M., WOOLFSON, D. N. Protein Sci. 1998; 7, 2287–2300. 47 ESPINOSA, J. F., MUNOZ, V., GELLMAN, S. H. J. Mol. Biol. 2001; 306, 397–402. 48 STANGER, H. E., SYUD, F. A., ESPINOSA, J. F., GIRIAT, I., MUIR, T., GELLMAN, S. H. Proc. Natl Acad. Sci. USA 2001; 98, 12105–12120. 49 COCHRAN, A. G., SKELTON, N. J., STAROVASNIK, M. A. Proc. Natl Acad. Sci. USA 2001; 98, 5578–5583. 50 KARSHIKOFF, A., LADENSTEIN, R. Trends Biochem. Sci. 2001; 26, 550–556. 51 KUMAR, S., NUSSINOV, R. Cell. Mol. Life Sci. 2001, 58, 1216–1233. 52 TAKANO, K., TSUCHIMORI, K., YAMAGATA, Y., YUTANI, K. Biochemistry 2000, 39, 12375–12381. 53 LOLADZE, V. V., IBARRA-MOLERO, B., SANCHEZ-RUIZ, J., MAKHATADZE, G. I. Biochemistry 1999, 38, 16419–16423. 54 CIANI, B., JOURDAN, M., SEARLE, M. S. J. Am. Chem. Soc. 2003; 125, 9038–9347. 55 RUSSELL, S. J., BLANDL, T., SKELTON, N. J., COCHRAN, A. G. J. Am. Chem. Soc. 2003; 125, 388–395. 56 MUNOZ, V., THOMPSON, P. A., HOFRICHTER, J., EATON, W. A. Nature 1997; 390, 196–199. 57 MUNOZ, V., HENRY, E. R., HOFRICHTER, J., EATON, W. A. Proc. Natl Acad. Sci. USA 1998; 95, 5872–5879. 58 HAGEN, S. J., HOFRICHTER, J., SZABO, A., EATON, W. A. Proc. Natl Acad. Sci. USA 1996; 93, 11615–11617. 59 DINNER, A. R., LAZARIDIS, T., KARPLUS, M. Proc. Natl Acad. Sci. USA 1999; 96, 9068–9073. 60 LACROIX, E., KORTEMME, T., LOPEZ DE LA PAZ, M., SERRANO, L. Curr. Opin. Struct. Biol. 1999; 487–493.

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61 SYND, F. A., ESPINOSA, J. F., GELLMAN, S. H. J. Am. Chem. Soc. 1999; 121, 11577–11578. 62 ESPINOSA, J. F., GELLMAN, S. H. Angew. Chem. 2000; 39, 2330–2333. 63 TATKO, C. D., WATERS, M. L. J. Am. Chem. Soc. 2004; 126, 2018–2034. 64 COLLEY, C. S., GRIFFITHS-JONES, S. R., GEORGE, M. W., SEARLE, M. S. J. Chem. Soc. Chem. Commun. 2000; 593–594. 65 HILARIO, J., KUBELKA, J., KEIDERLING, T. A. J. Am. Chem. Soc. 2003; 125, 7562–7574. 66 MA, B., NUSSINOV, R. J. Mol. Biol. 2000; 296, 1091–1104. 67 WANG, H., SUNG, S.-S. Biopolymers 1999; 50, 763–776. 68 HONDA, S., KOBAYASHI, N., MUNEKATA, E. J. Mol. Biol. 2000; 295, 269–278. 69 GRIFFITHS-JONES SEARLE, M. S. J. Am. Chem. Soc. 2000; 122, 8350–8356. 70 TATKO, C. D., WATERS, M. L. J. Am. Chem. Soc. 2002; 124, 9372–9373. 71 SHARMAN, G. J., SEARLE, M. S. J. Chem. Soc. Chem. Commun. 1997; 1955–1956. 72 SHARMAN, G. J., SEARLE, M. S. J. Am. Chem. Soc. 1998; 120, 5291–5300.

73 SCHENCK, H. L., GELLMAN, S. H. J. Am. Chem. Soc. 1998; 120, 4869–4870. 74 KORTEMME, T., RAMIREZ-ALVARADO, M., SERRANO, L. Science 1998; 281, 253–256. 75 De ALBA, E., SANTORO, J., RICO, M., JIMENEZ, M. A. Protein Sci. 1999; 8, 854–865. 76 DAS, C., RAGHOTHAMA, S., BALARAM, P. J. Am. Chem. Soc. 1998; 120, 5812–5813; DAS, C., RAGHOTHAMA, S., BALARAM, P. Chem. Commun. 1999; 967–968. 77 LOPEZ DE LA PAZ, M., LACROIX, E., RAMIREZ-ALVARADO, M., SERRANO, L. J. Mol. Biol. 2001; 312, 229–246. 78 FINKELSTEIN, A. V. Proteins 1991; 9, 23–27. 79 SYUD, F. A., STANGER, H. E., SCHENCK MORTELL, H. et al. J. Mol. Biol. 2003; 326, 553–568. 80 MACIAS, M. J., GERVAIS, V., CIVERA, C., OSCHKINAT, H. Nat. Struct. Biol. 2000; 7, 375–379. 81 KOEPF, E. K., PETRASSI, H. M., RATNASWAMY, G., HUFF, M. E., SUDOL, M., KELLY, J. W. Biochemistry 1999; 38, 14338–14351. 82 KANELIS, V., ROTIN, D., FORMAN-KAY, J. D. Nat. Struct. Biol. 2001; 8, 407–412.

1

Engineering Proteins for Stability and Efficient Folding Bernhard Schimmele, and Andreas Plückthun Universität Zürich, Zürich, Switzerland

Originally published in: Protein Folding Handbook. Part II. Edited by Johannes Buchner and Thomas Kiefhaber. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30784-2

1 Introduction

The industrial, biotechnological, and medical applications of proteins are often limited by an insufficient protein stability or related problems. Such applications commonly require that proteins be produced on a large scale and remain stable enough to fulfill their functions for a reasonable length of time, often under harsh conditions. However, natural proteins are typically only marginally stable, and it is thus a major challenge for protein engineers to optimize stability and folding efficiency. The approaches that have been successfully employed to achieve this goal are rational design, semi-rational strategies based on sequence comparisons, and the methods of directed protein evolution. Of course, these methods are not mutually exclusive and can be combined to solve practical problems. All studies employing these methods have revealed important rules for protein engineering and at the same time shed light on the principles and mechanisms responsible for the folding and stability of proteins. Recent advances in stability engineering have demonstrated that merely small changes in a given protein sequence can have profound effects on its biophysical properties. The major challenge is therefore to correctly identify and remedy these shortcomings. It is the goal of this chapter to summarize the biophysical principles and technological approaches useful in improving the biophysical properties of proteins through sequence modification. Considering the enormous array of technologies involved in this endeavor, ranging from computer algorithms to selection technologies, it is not possible to give detailed experimental protocols in this chapter; instead, we will guide the reader to the cited literature.


2 Engineering Proteins for Stability and Efficient Folding

2 Kinetic and Thermodynamic Aspects of Natural Proteins 2.1 The Stability of Natural Proteins

Evolution does not per se provide proteins with high stability. In fact, stability is just one of many evolutionary constraints on proteins. Proteins have to fold to a defined structure with adequate yield in a reasonable time and then have to be just stable enough to perform their function over a certain period. There is no evolutionary incentive to make a protein any “better” than what is needed to fulfill its cellular functions. In contrast, the use of a protein in a formulation at high concentration, its prolonged activity at 37 ◦ C, and its large-scale expression and crystallization, just to name a few conditions, may put far higher demands on the protein than its natural environment. Thus, the natural sequence may not be able to provide these properties, but a mutant sequence may. Proteins exist and have evolved in order to fulfill a given function, and evolution drives the structural properties of a protein mainly towards increased functionality [1]. In fact, most proteins are only marginally stable, with Gfolding in the range of −20 to −60 kJ mol−1 . It is still a matter of debate whether this marginal stability is actually a “design feature,” e.g., to allow degradation at a certain rate, whether it is caused by the selection pressure towards higher functionality that may not be compatible with high stability, or whether it is just a side effect of the lack of selection for high stability. Function is often linked to higher structural flexibility in certain regions of a protein. Lower stability as a result of this higher local structural flexibility might therefore simply represent an adaptation to increased functionality [2]. If this were generally true, however, stability engineering would fail in most cases, as it would not be able to reconcile stability with preserved protein function. An alternative, more optimistic view for the protein engineer is that marginal stability can be interpreted as a result of genetic drift [3]. In other words, lower stability is not intended; but it simply does not matter, provided that function is maintained. Random mutations occurring during evolution are more likely to destabilize the structure of a given parental protein sequence than to stabilize it or be neutral. However, as long as this stability decrease is not sufficient to render the protein nonfunctional, these destabilizing mutations are likely to accumulate in the sequence. This tendency has also been referred to as “sequence entropy” [4]. As a consequence, stability engineering could be interpreted as the art of identifying these unfavorable mutations in order to reestablish a more stable sequence. The concept discussed above also sets the basis for the consensus approach to stability engineering, which is discussed in Section 3.1 Based on the physical principles of protein folding and a structure-based analysis of the interactions between amino acid residues, rational design can give hints as to which residues need to be altered to achieve a desired effect, and this is discussed in Section 3.2. The third focus will be set on the methods of directed evolution, which mimic the

2 Kinetic and Thermodynamic Aspects of Natural Proteins

mechanisms of Darwinian evolution to evolve proteins with enhanced folding and stability properties (see Section 4). 2.2 Different Kinds of “Stability”

Before discussing in detail different strategies for rendering a protein more stable, it is worth taking a closer look at some basic features associated with protein stability and folding properties. The term “stability” itself is rather vague, and its precise meaning has often been adapted depending on the problem being addressed. This leads to different definitions of “stability.” We will now briefly analyze the definitions and differences between thermodynamic, kinetic, and thermal stability, as well as folding efficiency, which is also sometimes discussed in this context. Even though these properties are interconnected, they are not equivalent. This has important implications for protein engineering, as it is difficult to predict in advance how a given mutation will influence each of these properties. These influences will in fact be different for any protein under investigation according to the free energies of the native and unfolded states and the folding intermediates, as well as the folding pathway and the respective rate constants. 2.2.1 Thermodynamic Stability Thermodynamics describes the global unfolding behavior of a protein. The corresponding thermodynamic stability G describes the differences between the free energies of the native (N) and the unfolded (U) states. Importantly, G is an equilibrium property for a reaction involving two or more states. The simplest model of an unfolding reaction is the equilibrium of a two-state unfolding reaction:

where kunfolding and kfolding are the rate constants of the respective unfolding and folding reactions. As a consequence, values for G can be deduced only if the described process is fully reversible and no intermediate states are populated to a significant extent, unless they are explicitly known and measured. G describes to what degree the two states are populated at a given temperature according to G unfolding = −RT In K

(1)

where K is the equilibrium constant of the unfolding process. The treatment of experimental data is much simplified in such a model, as stability is affected only by the free energies of the folded and unfolded states. Because G also provides an overall measure of all energetic contributions of interactions occurring upon protein folding, it is a convenient quantity for comparing the energetic effects of single amino acid replacements in a given folded structure. Because of the great scientific interest in deducing the principles of protein folding on a quantitative

3


basis, many studies have been carried out with carefully selected model systems in which these strict requirements that allow the determination of G are fulfilled. Moreover, G represents an intrinsic quantity, at a given temperature in a given buffer, that is independent of the experimental setup and therefore reproducible in any case. The major problem in such studies, however, is confirmation that the observed transition can in fact be described as a fully reversible process, and denaturant-induced transition data alone are often not sufficient to allow reliable judgment. Only if data derived from measurements on the basis of different spectral probes (or better yet, additionally by differential scanning calorimetry, yielding the model-independent enthalpy change of unfolding) agree with each other are the deduced G values likely to be correct. Nevertheless, many useful conclusions can be drawn about the effects of mutations even if these strict requirements cannot be completely fulfilled in all cases. Another fact complicates the use of G as a measure for protein stability. Mostly as a result of the fact that the hydrophobic effect is the major driving force of protein folding [5], G itself is a characteristic, curved function of temperature. It is defined as G(T ) = G U −G N = H(T )−T S(T ) = −RT In K

(2)

emphasizing especially that the enthalpy change H is itself a function of temperature. This change of H with T can be described by the heat capacity change C p =

∂H(T ) ∂T

(3) p

Although often ignored, the temperature dependence of G should thus not be left out of the account if one is dealing with the stability engineering of proteins. The large heat capacity change upon the transfer of nonpolar solutes to water, which is the basis of the hydrophobic effect, results in a curved function of G versus temperature. By using the definition of Cp , G can be approximated as a function of T, the melting temperature T m , the enthalpy change at T m H(T m ), and Cp . G(T ) = (1−

T T )H(Tm ) + (T −Tm )C p −T C p In Tm Tm

(4)

If G(T) is plotted as a function of T, the curve increases at low temperatures and decreases at high temperatures (Figure 1). The temperature at which folded and unfolded states are equally populated, and thus G = 0, is called the melting temperature T m . The respective curvature of G versus temperature is strongly dependent on the change of heat capacity Cp upon unfolding. A mutation may change Cp , H(T m ), and T m in any combination, thereby altering the shape and the position of this curve. Higher thermodynamic stability at a given temperature (G(T)) can


Fig. 1 The complex relationship between the melting temperature and the free energy of folding. Schematic representations of the free energy difference between the folded and unfolded states of a protein, GN , as a function of temperature, T. A typical protein is shown in curve 1 (–––). The shape of this curve changes if a mutation affects C p ; H(T m ), and/or T m of the protein. In the examples shown, not just one but several of these parameters are changed. Higher thermodynamic stability at a given temperature can phenomenologically be the result of a curve upshift (. . .. . .. . .). Right-shifting the

curved function (----) results in a right-shift of the maximum of the GN function as well as in a shift of the melting temperature T m towards higher temperatures (T m ). If the GN function is flattened because of a small C p (. . .), a higher melting temperature (T m ) can result, even though the thermodynamic stability is actually decreased over a broad temperature range. All depicted curves represent extreme cases, and typically a combination of the described alterations will occur upon mutating a protein.

thus be achieved, e.g., by an upshift of this curve with constant maximum. Rightshifting of the curve will decrease G at lower temperatures but increase it at higher temperatures, which goes along with an increase of T m . Flattening of the curve due to a lower change of heat capacity upon unfolding may cause lower thermodynamic stability at most temperatures, even though T m is increased. These considerations contain important implications for the analysis of engineered mutants. First, a measured decrease of G at a certain temperature does not necessarily mean that at higher temperatures the thermodynamic stability might not have been increased. Second, by determining the change of G, no conclusions can be drawn about a potential change of the melting temperature T m . While they are typically related, a low G at a given temperature does not necessarily mean that T m will be decreased. T m and G must therefore be regarded as different properties. It should also be recalled that thermodynamic parameters, while easily reproduced, give no information about how long it will take until equilibrium is reached or, in the case of a nonreversible reaction, until a certain fraction of proteins is inactivated. Thus, thermodynamic stability does not necessarily provide information about whether a protein will meet the stability requirements for an intended application. T m itself often serves as a measure of protein stability, more precisely of thermal stability. In the literature, the expression “thermal stability” is again used with different meanings. In the described case, it represents the melting temperature for a reversible process. However, a complete thermodynamic analysis of the vast majority of proteins is not possible, because either intermediate folding states are

5


populated or folding in the absence of denaturants that solubilize the unfolded state is not a fully reversible process. As will be discussed in the following section, thermal stability can nevertheless serve as a very practical means of describing protein stability, defined either as the transition temperature of an irreversible process or as the half-life of a protein under a given set of conditions. 2.2.2 Kinetic Stability In most cases, outside the biophysical research lab, protein unfolding is an irreversible reaction. Initially, the aggregation of unfolded molecules or of folding intermediates prevent the back-reaction of folding, and this is followed, after prolonged times at high temperature, by chemical inactivation or, in impure samples, proteolysis. In a simple model, kinetic inactivation can be described as

As discussed in the Introduction, one evolutionary constraint on proteins is that their three-dimensional structure remains viable for a certain period of time. In the case of non-equilibrium conditions and irreversible reactions, reaction kinetics becomes the important parameter. The folded state can resist high temperatures for a considerably long time if either the rate constant of inactivation or the rate constant of unfolding is sufficiently low. The rate constants are determined by the free energy of activation, e.g., the difference in energy between the folded state and the transition state of unfolding. Proteins can thus be kinetically stabilized by increasing this activation barrier. This kinetic stability is different in some crucial aspects from the thermodynamic stability mentioned above. Engineering of proteins for enhanced stability has to deal with both aspects. The best mutations for enhancing kinetic stability will not necessarily be the best mutations for enhancing thermodynamic stability, and not every mutation that increases thermodynamic stability will automatically have a positive effect on protein half-life. Kinetic stabilization is a common theme in nature, and there are several indications that many proteins from thermophilic organisms are indeed stabilized kinetically rather than thermodynamically [6, 7]. Another important example is that of proteins from the coats of viruses and phages that have to protect their genetic material under very adverse conditions [8]. In extreme cases the native state of a protein can even be less stable than its denatured state, but the native fold can still be kinetically trapped, and large kinetic unfolding barriers can provide the protein with an extremely long half-life [9]. There are different ways of describing and determining kinetic stability. Even if the reaction proceeds in an irreversible way, a practical “melting curve” can still be determined, and the observed transition is cooperative. The midpoint of this transition can serve as a practical means to compare the thermal resistance of different protein variants. Alternatively, one can use the half-life of the protein at a given temperature as an empirical means of stability. However, one has to be aware


that this will not reflect equilibrium conditions. The observed values are actually kinetic values and thus are not independent of the exact experimental conditions, such as the protein concentration, the heating rate, etc. A more thorough analysis of kinetic stabilization must include kinetic measurements to determine the respective unfolding rates at different temperatures [10, 11]. For medical applications, engineered proteins usually have to fulfill defined stability requirements such as the absence of aggregation in the formulation used, long-term stability, and prolonged activity at 37 ◦ C. The mere analysis of thermodynamic and kinetic parameters under defined reaction conditions in vitro is not always a reliable indicator of protein behavior under in vivo conditions. Mechanisms other than the intrinsic properties of the protein, such as proteolysis or aggregation with other proteins, can affect half-life. For practical utility, the half-life can therefore also be determined by measuring the percentage of molecules that retain their function after incubation under the respective conditions, e.g., in human serum at 37 ◦ C for several days [12]. 2.2.3 Folding Efficiency A different issue of kinetics is related not to unfolding but to the folding of the protein in vivo. More precisely, the question is, which percentage of a protein will actually fold to the native state, as opposed to going to misfolded states, soluble aggregates, or inclusion bodies? Even though the folding efficiency in vivo is not directly related to stability, correlations can often be observed [13]. Additionally, the efficiency of protein folding in vivo is usually the predominant factor influencing the expression yield and is therefore also crucial for large-scale production of functionally intact proteins. Importantly, many mammalian proteins with a high potential for medical applications, especially those secreted or expressed on the cell surface, can rely on the complex folding machinery of the eukaryotic cell to reach their final native state, and the secretory quality-control system of eukaryotic cells allows discrimination of proteins by their folding behavior [14]. Moreover, they usually do not need to be expressed in high amounts in their native physiological context. The resulting lack of selection pressure on their efficiency of folding during evolution is likely to be one of the causes for the difficulties often observed when attempting their overexpression. Unfortunately, these tend to be the proteins of greatest pharmacological utility. Despite the fact that thermodynamic stability underlies folding efficiency, the kinetic partitioning into productive folding or aggregation is influenced by many different factors. For illustrative purposes, we can again describe this by a very simple scheme:

Folding intermediates are often the source of aggregation, and the overall folding efficiency will therefore depend mainly on the nature of these intermediates. This includes the free energy of the folding intermediates themselves, as well as their

7


half-lives and any efficient pathways to aggregation. in vivo, the situation becomes much more complex, as additional parameters such as interactions with cellular components and chaperones or degradation by host cell proteases come into play. The final output of a properly folded protein will therefore depend on all of these kinetic competitions [15]. Protein expression in the bacterial cytoplasm in many cases shows correlations between soluble expression yield and thermodynamic stability of the protein [16, 17]. Additional complications can arise from transport steps. For example, the expression of proteins in the periplasm of E. coli is dependent on the prior transport of the polypeptide chain through the inner membrane, and the folding yield is subsequently influenced by the folding and aggregation reaction in the periplasmic space as well as by interactions with periplasmic factors such as chaperones and proteases. In some cases, mutations that show positive effects on in vivo folding yield have no influence on the overall stability of the protein [18]. Conversely, mutations that strongly increase thermodynamic stability sometimes result in lower folding yields [19]. Nevertheless, many mutations act synergistically on both properties, because they are likely to reduce the free energy of the folded state as well as the free energy of folding intermediates and thereby lower the energetic activation barrier to folding [13].

3 The Engineering Approach 3.1 Consensus Strategies 3.1.1 Principles As discussed in the preceding sections, marginal protein stability is likely to be a side effect of “sequence entropy” occurring during natural evolution, because the major driving force of evolution is positive selection towards an enhanced functional property, while stability has to be maintained at only a minimum level to secure function. Mutations are likely to occur in a random fashion during this process; the probability that a mutation will have a stabilizing effect on the protein is very low, whereas the probability that the mutation will have destabilizing effects is very high. However, as long as the remaining amino acid sequence is still able to fold into a given structure and the overall domain stability does not fall below a certain threshold, the resulting protein sequences will not be eliminated during the course of evolution [20]. Destabilizing mutations are therefore often selectively neutral and thus accumulate in a given parental sequence. The same should also be true for folding efficiencies. Most proteins are not needed at high concentrations or may even become harmful to the organism in such a case. Similar to stability, folding yield is selectively neutral, provided that the minimum level for cellular function is maintained. This sets the basis for a semi-rational approach to protein stabilization, which is called the consensus approach [21] and is based on sequence statistics. Because mutations occur randomly, the distribution of amino acids at a given position in

3 The Engineering Approach

a set of homologous proteins can be described, in a very crude approximation, by Boltzmann’s law. The consensus approach assumes that at a specific position in a sequence alignment of homologous proteins, the contribution of the respective consensus amino acid to the stability of the protein is on average higher than the contribution of any non-consensus amino acid. Replacement of all non-consensus amino acids in a sequence by the respective consensus amino acid should therefore increase the overall stability of the protein. Obvious advantages of the consensus approach are that it is comparatively simple and is not strictly dependent on structural information at high resolution. The prerequisite for building a non-biased consensus is the availability of sequences homologous to the protein under investigation. The number of sequences should be large enough to make the sequence statistics reliable and to exclude bias in the resulting consensus sequences. Figure 2 shows an alignment of homologous sequences of single repeat modules, the smallest structural entity of a class of proteins known as leucine-rich repeat (LRR) proteins. Because the length of these modules varies among the different classes of LRR proteins – influencing their topology – only repeat modules of a length of 24 amino acid residues have been used for this alignment. The probability of each amino acid occurring at a given position is calculated to derive a consensus sequence, representing the most frequently occurring amino acid residue at each position. The distribution of residues at each position can provide information on structurally forbidden residues and allows weighing the consensus with respect to variability. In most cases, the consensus will contain the residues important for defining the structure of the proteins. In the case of an enzyme family, it will also include the “functional” residues, i.e., those of the active site. In the case of binding proteins, such as antibodies or repeat proteins, the “functional” residues (those involved in binding) are not conserved but are different for each individual molecule, which has to adapt to its target. Although at first sight no sophisticated structural analysis seems to be required, this is true in only the simplest of cases, where a single family of related sequences can be represented by a single consensus. Frequently, multiple families have emerged that use mutually incompatible solutions of packing. A good example is that of antibodies for which subgroup-specific consensus building has been very fruitful [22]. Averaging over all families would simply yield the consensus of the most-represented family and, if they are equally represented, may result in mutually incompatible residues. Thus, structural analysis can be very helpful in deciding whether an “averaging” of different sequence families is permissible or not. Because of this problem of interacting residues, a simple averaging may lead to incompatible pairs; therefore, these residues should be changed only as groups. The danger of disrupting these interactions by substitutions with consensus residues is especially high in cases where a very broad set of sequences is used for the alignment. To minimize this risk, the sequence statistics can be extended by analysis of covariance in order to derive probabilities that describe the joint occurrence of amino acid residues at two defined positions [23]. As explained above, before deriving a consensus, the aligned sequences can be divided into subclasses, which are likely to contain interacting pairs or groups of residues. Certain

9


Fig. 2 Example of the analysis for deriving and analyzing a consensus sequence. (a) From a sequence alignment of 3077 sequences of 24 amino acid LRR motifs of the Pfam database (http://www.sanger.ac.uk/Software/Pfam/),

the relative frequencies of amino acid residues at each position are calculated. (b) Based on the relative frequencies calculated from the alignment, a consensus sequence can be derived representing the most frequently occurring residue at each position.


variations of residues involved in salt bridges, distinct hydrogen-bonding patterns, or packing of the hydrophobic core are characterized by complementary changes between these subclasses, with mutations to a certain residue at one position being compensated by a mutation to a complementary residue at another position. The subclasses can be built either by being based on sequence homology alone or by including structural information if available. Even though the definition of these subclasses is always dependent on the homology cutoff set by the investigator, a simple dendrogram analysis can be used to group the complete set of sequences into distinct families. Additionally, by building the consensus sequence of each family separately, followed by a comparison of these consensus sequences, distinct structural features of each group can be recognized in some cases. The consensus concept has been applied successfully to a large variety of proteins and structural motifs to date. Important lessons for an effective application of the consensus approach can be learned from these studies, and we will therefore give a few examples to briefly discuss some of the advantages and limitations of the method. 3.1.2 Examples There is always the concern that the stabilizing and destabilizing effects of introduced mutations will counterbalance each other and that the overall change in protein stability will be small. Steipe et al. [20] applied the method for the first time on immunoglobulin VL domains and predicted 10 potentially stabilizing mutations. Six mutations were indeed stabilizing, three had no effect, and one was destabilizing. When applied to GroEL mini-chaperones, 34 predicted amino acid replacements were individually checked, out of which 13 were stabilizing, five showed no effect, and 16 were destabilizing [24]. Lehmann et al. [25] extended the approach to an entire protein. In a first set of experiments, 13 homologous sequences of

←-----------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------Fig. 2 (Continued) The color code is given on the right hand side of the panel. Residues occurring in more than 80% of all sequences at the respective position are colored in red. Based on these values, consensus sequences with a given homology threshold can be derived (not shown), i.e., with a higher threshold, more positions will be “undefined.” (c) Preferences at a given position are reflected not only by the absolute frequency but also by the total number of different amino acid types occurring at each position (residue variability). As an example, the relative frequencies of each amino acid are shown for the highly conserved position 2 and the highly variable position 4. By plotting the relative frequencies of amino acids at a particular position,

preferences for certain amino acid types as well as “forbidden” residues can be identified. At position 2, a strong preference for leucine can be observed, and the occurrence of other residues is restricted mostly to hydrophobic side chains. At position 4, all residues are “allowed” except for proline, which is “disallowed” due to secondary structure propensity reasons. (d) By normalizing the relative frequency of the consensus amino acid with the number of “allowed” residues at a given position, a modified consensus sequence can be deduced. The variability V is calculated according to V = 100×N/F, where N is the total number of different residue types occurring at each position and F is the frequency of the most frequent residue in percentage.

11


a fungal phytase were used to build the consensus, and the resulting consensus enzyme showed an increase of 15–22 degrees in unfolding temperature and an increase of the temperature optimum for catalysis of 16–26 degrees compared with each of its parents. In a second set of experiments, additional mutations were predicted by simply adding more sequences to the alignment. By checking the effects of the individual mutations on thermal stability and combining mutations with positive effects, the unfolding temperature could be increased by an additional 21 degrees to 90 ◦ C. No loss of catalytic activity of the enzyme was observed in any case. This work showed that the number of sequences used for the alignment is indeed an important factor because it might help to optimize ambiguous positions. In addition, the observed large increase in thermal stability could not be attributed to the effect of one single amino acid substitution but rather to the synergistic effects of many replacements. Even though these and other studies show clearly that the consensus approach allows one to predict stabilizing mutations with a rather high success rate in a rapid way, the effect of each predicted mutation carries some uncertainty, and it is possible that some may contribute destabilizing effects that can counterbalance the stabilizing ones. However, the effects of stabilizing mutations were often found to be additive. Therefore, instead of examining each mutation individually, it is often useful to combine groups of “rather certain” mutations and others that are more speculative. The application of the consensus concept to families of repeat proteins [23, 26–28] represents, in some respects, a special case due to a number of favorable features. It can nevertheless illustrate the importance of some principles of the approach. The non-globular fold of repeat proteins consists of repeated motifs of 20–40 amino acids. Several results indicate that consensus repeat proteins are indeed much more stable than natural repeat proteins [29, 30]. Repeat proteins might represent an extreme case in which the principles of the consensus approach become very apparent. The structural entities (the repeat modules) are small and thus each protein contributes several repeats to the databases; the number of available sequences consequently becomes very large compared with other proteins. Therefore, a consensus sequence for a single repeat module can easily be assigned and ambiguous positions will occur with lower frequency in the statistical output (Figure 2). In addition, interactions that are present within or between several repeats will add to the free energy of folding multiple times, while problem spots would equally be potentiated. Therefore, effects on stability are likely to be consecutively added by introducing additional modules to the array. Even though more-detailed analyses are still needed, initial results pinpoint some of the structural reasons for the stability gains observed upon building a consensus in each of these studies. The regular arrangement of structural motifs gives rise to a more regular H-bonding pattern with a higher number of inter- and intra-repeat H bonds [30]. Loop insertions in natural repeat proteins that are likely to result in more flexible local regions are removed, thereby eliminating local centers of unfolding. In the left-handed helical and disallowed regions of the Ramachandran plot, glycine residues are always present in the consensus proteins, while they are avoided in other places by this design, where their flexibility is not needed and may be harmful to stability.


3.2 Structure-based Engineering

Structure-based engineering relies on a detailed analysis of 3D structures, followed by site-directed mutagenesis. We avoid the term “rational” engineering, as it would elicit expectations of perfect predictability and implicitly suggest that all other approaches are free of logical reasoning. In structure-based engineering, positions have to be identified at which suboptimal amino acids in the original sequences lead to a loss of stability. Subsequently it needs to be specified which amino acids should be introduced as a replacement. The ab initio prediction of protein structure, however, is still not a feasible task due to the multitude of potential interactions within the protein and between protein and solvent, which leads to an extremely high number of possible conformations and intermediates of similar energy [31]. Hence, a prerequisite for structure-based engineering is, next to experimentally determined structures, usually the existence of a large experimental data-set within a group of structural homologues that can be used as a basis for predictions. High-resolution structures are necessary to allow the estimation of possible conformational, energetic, and steric influences upon replacement of particular amino acid side chains and can thus help to avoid unfavorable strain in the resulting mutants. Because of the present efforts in the field of structural genomics, these structure-based approaches are likely to become even more important in the future. With this structural information in hand, the goal of designing more stable variants is then to pinpoint particular regions and positions associated with possible stability defects and to subsequently find a better solution to the problem. In contrast, semi-rational approaches like the previously discussed consensus concept are rather crude methods for introducing stabilizing features. Structural and energetic analysis can be used to reexamine the changes proposed by the consensus approach and to fine-tune the system to reintroduce structural features that might have been lost in the averaging process. Proteins of hyperthermophilic organisms have been of special interest for examining the structural mechanisms of thermostabilization and have been contemplated as guides for the engineering of “problem” proteins for better properties. From a phenomenological point of view, the basis of increased thermostability is frequently set by a flattened G-versus-T curve that is due to a smaller change of heat capacity upon unfolding (Figure 1) or by a kinetic stabilization that is due to a strong decrease in the rate of unfolding. The crucial question is, however, what the molecular differences are that give these proteins their favorable properties. Genome-wide comparisons between hyperthermophilic and mesophilic organisms with respect to amino acid composition did not yield any obvious common rules of how these effects are achieved [32]. Therefore, hope was placed on the increasing number of pairwise high-resolution structure comparisons of thermophilic proteins with their mesophilic counterparts. While they provided a more differentiated picture, a “global” rule still could not be derived. A few highly specific mutations are often enough to provide considerably stabilizing effects, but the additive effect of many small contributions, none of them dramatic by itself, may be the usual case.

13


Moreover, rather than relying on one universal strategy, nature utilizes a variety of strategies for the thermal adaptation of proteins [33]. In fact, the list of stabilizing structural features in hyperthermophilic proteins reflects the diverse principles of protein stability and folding that protein engineers try to exploit and that will be discussed in this section. High-resolution structures of thermophilic proteins can thus provide a detailed view of how nature implements these principles to create proteins of higher stability [34]. However, the lack of a unifying “rule” and the multitude of strategies nature uses provide an important lesson for protein engineers. Depending on the protein under investigation, the strategy of choice can be different, and even for a given protein there may be more than one optimal solution to the problem. Before choosing from the available set of strategies, the focus should therefore be on identifying potential “weak points” responsible for stability defects in a given protein structure. We will now discuss some structural features associated with protein stability as well as strategies for altering these features towards more favorable biophysical properties. The given list of course lays no claim to completeness but should point out some important principles. Any replacement in a given sequence may have multiple effects on protein stability, and destabilizing effects can often outweigh the stabilizing ones. An assessment of potential destabilizing effects is therefore crucial. Wherever possible, references are made to the different forms of “stability” discussed in the Introduction. In the case study provided at the end of this section, an example will be given to demonstrate how consensus approaches and structural analysis can be combined to yield useful results. 3.2.1 Entropic Stabilization An obvious strategy for increasing the free-energy difference between the folded and unfolded states, and thus the thermodynamic stability, is to decrease the entropy of the unfolded state. The underlying concept is to decrease the flexibility of the polypeptide chain, usually by introducing an additional intrachain linkage. Such entropic stabilization has become a common strategy for protein engineers. The prerequisite for success is that the mutations rendering the unfolded protein less flexible do not introduce unfavorable strain in the folded three-dimensional structure or result in any steric incompatibilities [35]. We now discuss several ways to achieve an entropic stabilization. 3.2.1.1 Introduction of Disulfide Bridges The introduction of additional disulfide bridges is a straightforward way of establishing an intrachain linkage to reduce the entropy of the unfolded state [36, 37]. The magnitude of the entropic effect is thought, as a crude approximation, to be proportional to the logarithm of the number of residues between the two bridged cysteines [38]. The spatial distance between the residues to be replaced with cysteines has to be evaluated with care in the model of the folded protein in order to prevent perturbations of the native structure upon formation of the disulfide bridge. It should be noted, however, that the energetic effect of additional disulfide bridges is not only entropic but also of a far more complex nature, giving rise to entropic as well as enthalpic contributions


to the change in the free energy of folding [39, 40]. For example, an additional decrease in the free energy of folding can result from the reduced solvation energy of the unfolded state [40]. In contrast, a reduced solvation energy of the folded state would have the opposite effect, while residual structure in the denatured state would again push the equilibrium to the side of the folded protein. Because the disulfide bond itself is hydrophobic in nature, it is often engineered into the interior of the protein. This is not an easy task, as it can negatively affect core packing. Even though there are several examples of successful protein stabilization by introducing artificial disulfide bridges [41, 42], the complex energetic effects can also cause a destabilization of the protein [43]. Furthermore, the introduction of additional cysteines often results in a rather drastic decrease in folding efficiency, because incorrect and intermolecular disulfide formation can remove large portions of expressed protein by aggregation. Since disulfide formation does not occur in the cytoplasm, secretion to the bacterial periplasm or to the eukaryotic ER is required for functional expression, usually associated with lower yields than for the production of cytoplasmic proteins. Alternatively, if the protein is produced by refolding, redox conditions have to be adjusted, which can be difficult if the native protein also contains free cysteines. 3.2.1.2 Circularization An alternative approach with the same underlying concept is the circularization of proteins by fixing the loose N- and C-termini via a peptide bond. In addition to the entropic effect, the fixing of the loose ends can prevent local unfolding events occurring at the termini and thereby kinetically stabilize the native structure. With the discovery of inteins, which mediate protein-splicing reactions, a tool that allows the directed formation of peptide bonds between ends fused to different parts of the intein became available. Intein-mediated protein ligation has been used to covalently link the termini of β-lactamase, a protein that is especially amenable to this strategy due to the close proximity of its N- and C-termini [44]. In accordance with polymer theory, the thermal stability of the protein was enhanced by about 6 degrees, from 45 ◦ C to 51 ◦ C. For circularized DHFR [45] an increased half-life at elevated temperature was observed. The close proximity of the termini is of course a prerequisite for this procedure, and the stabilizing effect is likely to become marginal if the loose ends are linked via long unstructured loops. Similar to the situation upon introduction of artificial disulfide bridges, destabilizing enthalpic effects may negate the favorable entropic contribution [46]. In addition, low protein ligation efficiencies and difficulties in separating circular from linear forms of the protein often cause additional technical challenges. It remains to be seen whether this technology is robust enough for biotechnological or biomedical applications. 3.2.1.3 Shortening Solvent-exposed Loops Short, solvent-exposed loops are rather fixed in the native state, but a comparably large number of additional conformations become accessible in the unfolded state, while long loops have a large number of conformations also available in the native state. Thus, the shortening of loops should in principle lead to a relative decrease in the loss of conformational entropy upon folding. Conversely, increasing the loop lengths by insertion of glycine residues into

15


the loops of the four-helix bundle protein Rop has indeed resulted in a strong and continuous decrease in thermodynamic stability [47]. In addition, loop shortening can have the effect of abolishing hot spots of local unfolding events and may result in kinetic stabilization. Even though it has become obvious that loop shortening or tying down of loops by external interactions is a common theme in thermostable proteins of thermophilic organisms [48, 49], the strategy is often hard to realize for a given protein target, as the danger of introducing additional strain in the native state is high, and solvent-exposed loops are often important with respect to function. 3.2.1.4 Reduction of Chain Entropies By considering the conformational entropies of amino acid side chains, another strategy for decreasing the entropy of the unfolded state becomes apparent. Because of the five-membered-ring nature of the proline side chain, it not only restricts the possible conformations of the preceding residue but also can adopt only a few conformations itself. It therefore has the lowest conformational entropy of all amino acids [50]. In contrast, glycine, which has no side chain, has the highest conformational entropy. Substitutions of non-glycine residues with proline or the replacement of glycines by other residues should therefore reduce the entropy of the unfolded state. Positions that allow substitutions with proline are, however, very rare. Because proline is poorly compatible with α-helices and incompatible with β-strands, the position of a new proline must not be part of these secondary structure elements. At most positions in the native structure, the respective torsion angles will be incompatible, and the mutations are thus very much restricted to loop and turn regions. Again, care should be taken not to remove any favorable interactions of the replaced amino acid side chain [51]. In order to examine in advance whether the respective site is permissive for a substitution with proline, the dihedral angles of the site can be checked and should lie in the range of φ/ −50 to −80/120 to 180 or, alternatively, −50 to −70/−10 to −50. Similar restrictions apply for the replacement of glycines with any other residue. In many cases this will create steric overlaps, and such negative structural crowding effects can outweigh the positive energetic benefits. 3.2.2 Hydrophobic Core Packing Exposed residues are often directly involved in ligand or substrate binding and therefore often play a functional role. In contrast, the residues of a protein’s interior usually play mostly a structural role, and the associated hydrophobic effect is thought to be the main driving force of protein folding and thermodynamic stability. In known structures the core residues fill almost the entire interior space, provide many favorable van der Waals interactions, and maximize hydrophobic stabilization by exclusion of the solvent. In principle, an increase in thermodynamic stability of 4–8 kJ mol−1 can be achieved for each additional methylene group buried [52]. Paradoxically, the importance of the hydrophobic effect for folding and stability of proteins simultaneously limits its applicability for protein engineering. Because


the hydrophobic core is already densely packed in almost all native proteins, most changes here will create over-packing or packing defects, causing an overall destabilization rather than an improvement in stability. In addition, even subtle changes of core residues can lead to a rearrangement of external residues and thereby alter the functional properties of the protein [53]. Care should also be taken to avoid the introduction of conformational strain by the mutation of core residues, as destabilizing effects from a strained conformation can sometimes compensate the energetic gain of an increase in buried hydrophobic volume [54]. Improvement of core packing must therefore be based on analyses made from high-resolution structural information in combination with sequence comparisons. This allows one to specifically look for cavities in the core that indicate imperfect packing. If the hydrophobic surface area around the cavity is large, additional van der Waals interactions can be provided by the introduction of sterically fitting alkyl or aryl groups from hydrophobic side chains, thereby decreasing the size of the cavity [55, 56]. 3.2.3 Charge Interactions Oppositely charged amino acid residues, if appropriately positioned, have the potential to form salt bridges, whereas like-charged residues lead to repulsions. The magnitude of the effect of charge-charge interactions on overall protein stability is still a matter of discussion [57]. In the case of ionic interactions between side chains buried in the hydrophobic core, the high energetic cost of transferring charged ions from aqueous solution to the low-dielectric interior of the protein also has to be taken into account. If a single charge were buried in a protein, which would be extremely rare in a natural protein, the design of an ion pair would be very attractive. However, if a hydrophobic pair were to be replaced by an ion pair, the resulting energy would have to be higher than the loss of the previous pair plus the cost of burying the charge. Nevertheless, the high contribution of buried salt bridges to the overall stability of the native protein structure underlines their potential for introducing additional stability [58]. The optimal spatial arrangement of the interacting side chains and their respective charges is, however, crucial. Moreover, buried charged side chains are often not only part of interacting charge pairs but also part of complex charge clusters built from many side chains, which are able to magnify the effect. Similar rules apply for the interactions of charged residues on the protein surface. Only the perfect arrangement of charges seems to be able to make up for the desolvation penalty that has to be paid upon formation of a salt bridge. The effect on thermal stability, however, can be drastic. Increasing the free energy of unfolding by the changing of charges includes maximizing the number of salt bridges and, equally important, the removal of repulsive interactions [59], which are not uncommon in natural proteins. Predictions on a structural basis can be difficult due to the often higher flexibility of side chains on the protein surface, but simple models that allow predictions about potential stabilizing and destabilizing surface charges can be used [60]. The key is to consider not only nearest neighbors but also a whole network of charges that have to optimally interact and avoid repulsions.

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Because the surface charge distribution can have a huge impact on stability, but is defined by many residues at different positions, these residues are also a valuable target to be combined with selection techniques as discussed in Section 4. A special case of electrostatic interaction is the “helix dipole.” By reducing the net partial charges at the helical ends through placement of side-chains, which productively interact with the helix dipole, the helical structure is stabilized [61]. Introduction of negatively charged residues at the N-terminal end and positively charged residues at the C-terminal end leads to this stabilizing effect. The provided stability gain is, however, marginal (less than 4 kJ mol−1 ). 3.2.4 Hydrogen Bonding There was no initial reason to believe that intrachain hydrogen bonds in the native state would be more energetic than those of the unfolded chain to water [5]. By including terms of entropy change of the solvent and additional van der Waals interactions upon polar group burial, however, the positive contribution of hydrogen bonding to protein stability has become generally accepted [58, 62]. Despite this ongoing discussion, hydrogen-bonding patterns are a highly valuable target for the stability engineering of proteins. Because engineering deals with improving folded proteins, the major concern has to be how to satisfy the existing hydrogen-bonding network in a structural context. The basis for this endeavor is structural information of high resolution, and the most lucrative goal is to identify potential residues that represent buried but unsatisfied donors of hydrogen bonds. Site-directed mutagenesis of a nearby residue to provide a hydrogen-bonding acceptor can cause a stability gain in the range of 2–10 kJ mol−1 , depending on the geometry and other compensating effects [63, 64]. A special structural context that can provide significant additional stabilization by either hydrogen bonds or ionic interactions is the anchoring of relatively loose structural elements like loop structures or the N- and C-termini, thereby tightening “hot spots” of local unfolding [49, 65]. 3.2.5 Disallowed Phi-Psi Angles The stereochemistry of the polypeptide backbone can be defined by the dihedral angles φ and , and any individual residue in a structure is defined by a single set of φ, values. For conformational analysis of protein structures, the Ramachandran plot representing the dihedral angle space is an excellent starting point [66]. The φ, values of amino acid residues in protein structures usually reside in three preferred or “allowed” regions of the Ramachandran plot, called right-handed helical, extended, and left-handed helical. The right-handed and extended conformations correspond to α-helix and β-strand secondary structures, respectively, and the vast majority of non-glycine residues lie within these two regions. The left-handed region corresponds to structural features at the termini of secondary structure elements and describes regions involved in the reversal of the polypeptide chain. There is a high preference in this region for glycine residues, as the β-carbons of non-glycine residues can sterically interact with the polypeptide backbone, resulting in unfavorable energies. In some cases, the substitution of non-glycine residues


by glycines in the left-handed helical region increases thermodynamic stability (up to 8 kJ mol−1 in RNase H) [67]. Although the strict introduction of glycines in such positions is an important point to consider for de novo protein design, it does not represent a general rule for stabilizing the native states of proteins. In some cases, the energy penalty for the accommodation of unfavorable strain can be offset by lost unfavorable or new favorable local interactions, such as hydrogen bonding or hydrophobic interactions. Some replacements of this kind can therefore even lead to destabilization rather than stabilization [68]. The same rules apply in principle to the disallowed regions of the Ramachandran plot. Steric clashes result in a high energetic cost in the folded structure, especially for non-glycine residues in all disallowed regions. Stabilizing mutations to glycine have been introduced with energy gains of up to 18 kJ mol−1 [69]. It has been noted, however, that certain non-glycine residues also have propensities to occur in the disallowed regions – such as Asn and Ala in the type II turn region– and the energetic cost of their occurrences is often low [70]. Other residues found in the disallowed regions are small polar residues [71] that compensate for the energy cost by making additional hydrogen bonds. Unfavorable conformations often occur in very short loops [72], where the rest of the structure may constrain the loop efficiently, and interactions with the solvent may also offset the energy costs. Conformational analysis by the Ramachandran plot therefore provides a convenient and fast way to assess possible conformational strain in the tertiary structure associated with particular target residues. However, a close inspection of possible side chain interactions is required, and the analysis should be extended to identify potential compensating features of the residues to be replaced. 3.2.6 Local Secondary Structure Propensities Effects on the overall stability of a protein can also be influenced by the respective secondary structure propensities of amino acid residues for the α-helical and βsheet conformations. However, these effects are usually marginal. Nevertheless, even at the expense of other favorable trends, such as avoiding the exposure of hydrophobic side chains to the solvent, a given residue is often favored at a certain position due to its secondary structure propensity [73]. If a particular secondary structure element does not form efficiently, many interactions between this element and other parts of the protein can be lost. The energetic consequences and the influences on folding efficiency are, however, hard to predict at this point. 3.2.7 Exposed Hydrophobic Side Chains The removal of exposed hydrophobic side chains increases the polar surface of a protein. Such mutations are not likely to affect thermodynamic stability but presumably do affect the folding efficiency by influencing the rate of aggregation of intermediates. Interestingly, they also do not affect the solubility per se (the amount of native protein that can be dissolved in buffer) and seem to act mostly on folding intermediates [74]. Because of lateral interactions, e.g., between neighboring loops, partially exposed hydrophobic amino acids can even increase stability. It must be kept in mind however, that not only the aggregation pathway, but also the stabilities

19


of folding intermediates are an important parameter for kinetic stability, which can possibly be influenced by the existence of exposed hydrophobic side chains. Moreover, hydrophobic cavities at the protein surface are often involved in specific binding functions of the protein, and the removal of these “functional hot spots” has to be avoided. Therefore, the hydrophobicity of the protein surface must be carefully balanced. 3.2.8 Inter-domain Interactions In proteins consisting of more than one domain, additional principles come into play. The overall stability not only reflects the intrinsic stabilities of the single domains but is also influenced by the stability of the interface of the domains in cases where they are interacting with each other. Stabilization of this interface can be achieved mainly by increasing and optimizing the hydrophobic surface area of the interface. Because hydrophobic side chains at the interface are usually exposed during folding and transient opening of the domain interface, a tradeoff between interface stability and folding efficiency is often observed. Additional dramatic effects on stability can be observed in cases where the two domains exhibit very different intrinsic stabilities [75]. In such a scenario, the unfolding of the protein and the loss of function are strongly related to the unfolding of the less stable domain [76]. An important aspect of kinetic stabilization can be observed in twodomain proteins, where one domain can slow down the unfolding, and therefore the aggregation, of the other. Thus, the native state becomes kinetically stabilized in the assembly and a stable domain interface reduces the extent of its transient openings and, thereby, the resulting exposure of hydrophobic patches that would favor aggregation. Alternatively, covalent cross-links (e.g., disulfide bonds) between the interfaces of multimeric proteins can be introduced. For example, the introduction of disulfide bonds between the interfaces of Lactobacillus thymidylate synthase not only increases their thermal stability but also leads to reversible thermal unfolding [77]. 3.3 Case Study: Combining Consensus Design and Rational Engineering to Yield Antibodies with Favorable Biophysical Properties

The following example illustrates how consensus approaches, rational design, and experimental data can be combined in a synergistic fashion to iteratively optimize biophysical properties. The smallest form of an antibody able to bind the antigen in the same manner as the whole IgG consists of two domains, the variable domain of the heavy chain (VH ) and the variable domain of the light chain (VL ). Both domains interact with each other via an interfacial region of highly hydrophobic character. In single-chain Fv (scFv) antibody fragments, the two domains are covalently linked by a flexible linker region of typically 15–20 amino acids (Figure 3a). The binding site for the antigen usually involves three loop regions in each of the domains, named complementarity-determining regions (CDRs). Antibodies that are based


Fig. 3 (a) Structure of an antibody Fv fragment consisting of the variable heavy chain (VH ) and the variable light chain (VL ). Each domain VH and VL is characterized by three hydrophobic core regions (upper [green],

central [yellow], and lower [orange] core) and a charge cluster at the base of each domain (red). Even though the residues defining these regions are conserved within the same germ-line family, these sequence

21


on human antibody sequences possess great potential for many medical applications, either directly as an antibody fragment or after reconstructing an IgG [78]. Antibody fragments can be expressed in a convenient manner in E. coli, thereby providing rapid access to these proteins [79]. Nevertheless, there are often drastic differences between individual antibodies concerning their expression yield and their stabilities. Ideally, the recombinant antibodies would all provide favorable biophysical properties. The starting point for such recombinant antibodies is a library. One fully synthetic library of this kind, the human combinatorial antibody library (HuCAL), was designed based on the consensus concept [22]. Based on human antibody germline sequences, several sequence families were created. Importantly, instead of averaging over all possible human antibody sequences, the consensus sequences were built for each family separately, resulting in seven consensus frameworks for VH (VH 1a, VH 1b, VH 2, VH 3, VH 4, VH 5, and VH 6) and seven frameworks for VL (Vκ 1, Vκ 2, Vκ 3, Vκ 4, Vλ 1, Vλ 2, and Vλ 3). This diversity is important, as the use of different frameworks allows a variety of non-CDR contacts to the target, thereby greatly increasing the range of targets being recognized. By using this strategy, each human VH and VL subfamily that is frequently used during an immune response is represented by one consensus framework, and thus the immune response is closely mimicked. The consensus building was further restricted to the framework regions, while the CDRs were diversified in a manner guided by structure. Thereby, functional diversity is maintained in optimized sets of frameworks. Instead of fully relying on the statistical output of the consensus building, structural modeling was employed in order to decrease the risk of disrupting interactions between certain amino acid residues that might be in contact with each other in the three-dimensional structure. Because many structures of antibody domains are available, the modeled structures of each framework family could be compared with the respective natural template structures. The models were checked according to several principles, which have been outlined in the preceding section. At this point ←-----------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------Fig. 3 (Continued) motifs differ between different germ-line families. (b) Arrangement of the residues defining the charge cluster of VH 3 and Vk 3 domains [13]. Importantly, the charge cluster consists of a network of buried charges and hydrogen bonds rather than pairwise interactions between individual residues. (c) Furthermore, subtype-dependent packing differences occur. As an example, the residues defining the upper hydrophobic core region of different human VH subtypes are shown as structural superpositions. Structural alignments are shown of VH 4 (PDB entry 1DHV), VH 1a (1DHA), and VH 5 (1DHW) [22], with the most stable framework, VH 3

(1DHU) (left panel). While the upper core residues of VH 3 are densely packed, cavities occur for the other subtypes. In the least stable subtypes, VH 4 and VH 6, the bulky aromatic residues Phe29 and Phe31 are replaced by smaller residues, and the created space is only partly filled up by compensating residues Trp41 and Val25. The loss of the phenyl ring by replacement with Gly in VH 1a (middle panel) as well as the substitution of Leu89 by Ala are not compensated for by larger side chains at other positions, thereby creating hydrophobic cavities. The same is true for position 89 in VH 5 (right panel). Adapted from Ewert et al. [13].


the models were mostly checked for whether the interactions in natural antibody domains were correctly recreated. The consensus sequence models were inspected for any unfavorable strain in the structures, represented by unfavorable regions of the Ramachandran plot or any obvious cavities in the hydrophobic core. Moreover, the sequences were checked for the existence of residues already known to be involved in conserved intra-domain interaction patterns – such as conserved charge clusters and hydrogen-bonding patterns – as well as for exposed hydrophobic side chains known to decrease expression levels. Nevertheless, differences between the natural subclasses became apparent in the models. Empirical results had suggested that certain natural framework types display more favorable stabilities and expression levels than others. These differences are already existing in the original natural human germ-line sequences. The intrinsic differences in terms of biophysical properties for each subtype were then experimentally explored in a systematic fashion, first on single domains, then on scFv fragments [13]. The experiments confirmed the observed trends, showing that VH 3 displays the highest thermodynamic stability and soluble expression level, when expressed as an individual domain, among all VH subtypes. In contrast, VH 2, VH 4, and VH 6 display the least favorable properties in terms of stability, folding yield, and the tendency to aggregate. For the VL domains, members of the Vκ subtypes showed slightly higher stabilities and expression yields than the Vλ subtypes, but the behavior was much more homogenous overall. In order to trace back these differences to the structural level, the model structures of the different subtypes were compared with each other. Several structural features can be invoked to explain the extraordinary stability of VH 3 domains in comparison with the even-numbered VH subtypes. First, differences in the hydrogen-bonding networks have an influence on the thermodynamic stabilities. Long-range interactions involving several residues are concentrated in a charge cluster at the base of VH domains to establish a complex interaction network. In VH 3 domains the ionic and hydrogen-bonding interactions within this charge cluster are very well satisfied, whereas fewer interactions are observed at the analogous positions in other subtypes (Figure 3b). Corresponding to the subclass, different hydrogen-bonding networks are formed in the charge cluster of VH domains. Some of these networks are less extended and contain a smaller number of interactions than in VH 3. Additionally, based on the residues at three different positions in the first β-strand of the VH domains, the domains can be classified into four different structural subtypes with respect to their conformations in the first framework region [80]. Mutations bringing together incompatible residues and thus “mixing” subtypes have previously been shown to have a large unfavorable effect on the stability of the whole scFv [81]. Also, clear differences can be observed for hydrophobic core packing of the family subtypes. The upper core region of VH 3 is densely packed, whereas cavities can be identified in VH 4, VH 1a, and VH 5 on the basis of structural alignments (Figure 3c). In the lower core, two of the stable domains have an aromatic residue, while the others do not.

23


Finally, a comparison of the Ramachandran plots showed additional non-glycine residues with positive φ-angles and residues with higher secondary structure propensity at certain positions for the even-numbered subtypes compared with the odd-numbered ones. The immediate question was, therefore, whether the results of this structural trouble-shooting could be used to project favorable properties of VH 3 domains onto the less stable subtypes and thereby add another step to the optimization of antibody sequences while maintaining the structural diversity of the immune system. Instead of using the stable VH 3 framework exclusively, resulting in a loss of diversity, some point mutations might be enough to correct some of the shortcomings of the less stable domains. Based on these comparisons, the mutation of six residues in a scFv containing a VH 6 framework led to an overall increase in thermodynamic stability of 20 kJ mol−1 and a fourfold increase in soluble expression yield [73], indeed bringing this framework to the level of VH 3. The effects of the single mutations on stability were almost fully additive, while the effects of folding efficiency (soluble expression yield) were only qualitatively additive. The most dramatic effects on thermodynamic stability were obtained by mutations removing an unsatisfied H-bond donor in the hydrophobic core and introducing glycines at positions with positive -angles. Individually, all mutations, except the one in the hydrophobic core, led to slight increases in soluble expression yield. Interestingly, one mutation that removed a hydrophilic, solvent-exposed glutamine residue by a replacement with hydrophobic valine on the basis of higher secondary structure propensity also significantly increased the expression yield, possibly since valine secures this stretch to be in β-sheet structure. In antibodies, disulfide bond engineering has also been investigated. Optimized disulfide bonds engineered between VH and VL indeed significantly increased the half-life of an Fv fragment at 37 ◦ C [82]. This strategy was subsequently extended to interchain disulfides in generic framework positions [83]. Even more stable proteins can be obtained by combining the single-chain Fv approach with the engineered disulfide [84, 85]. It should be pointed out, however, that the additional inter-domain disulfide significantly reduces the yield of folded protein when produced in the bacterial periplasm, and such proteins have to be prepared by in vitro refolding. Therefore, the additional disulfides are not ideal for antibody libraries; instead, optimized frameworks have shown the greatest promise for combining diversity with stability. In summary, the consensus concept provides a convenient tool for proceeding with large steps in sequence space and a high probability of accumulating features that are favorable for higher stability. In many cases, structural analysis can serve as a trouble-shooting tool to identify shortcomings that might have been created by the consensus-building process or, as in the case of the antibodies described here, that are inherent to the natural sequence. In addition, it serves to rank the potential mutations identified by the consensus approach, keeping the mutational load on the target molecule to a minimum. Experimental data are important not only to validate these results but also to give important hints for future design approaches.

4 The Selection and Evolution Approach 25

Many entries in the table of experimental antibody stability data linked to mutations have come from directed evolution experiments. By combining semi-rational and rational approaches with experimental data, optimization of biophysical properties can be achieved in an iterative fashion. This interplay of experiment and structural analysis can therefore be an effective way to probe the vast sequence space in a systematic manner in order to find the valleys of free energy.

4 The Selection and Evolution Approach 4.1 Principles

The previous section may have led to the impression that mutations enhancing the biophysical properties of proteins can rapidly be identified. In the highlighted case of antibody domains, only the wealth of structural data and empirical measurements available allowed predictions with a high probability of success. Some of the empirical data used successfully for structure-based engineering have come, paradoxically, from directed evolution experiments (see Section 3.3). Despite the rapidly increasing amount of structural data and the better understanding of folding mechanisms of proteins, the effects of introduced mutations still cannot be predicted with a high degree of accuracy in most cases. The main reason is that even slight alterations in the primary sequence can lead to profound conformational changes in tertiary or quaternary structure; consequently, structural predictions have to be very accurate and usually must be backed up by empirical data. Because rational engineering uses site-directed mutagenesis followed by biophysical investigation to probe the effects of specific amino acid substitutions, it becomes very labor-intensive if many mutants have to be checked individually and if no additional hints are available. Additionally, as soon as small synergistic effects of several mutations need to be checked, the combinatorial explosion rapidly exceeds the sample number that can be handled efficiently. More importantly, the restriction to certain target residues automatically excludes alternative solutions to a given problem that may not have been obvious by the initial analysis. For example, affinity improvement of a protein to its binding partner is often achieved more efficiently by slight spatial adjustments of residues directly involved in binding, rather than by substitution of these residues. This kind of spatial adjustment can be caused by mutation of residues whose location in the native structure is further away from the actual binding site (so-called “second-sphere mutations”) [86]. Today, it is almost impossible to predict this kind of mutation by rational means. When it comes to stability engineering of proteins, the problems associated with rational design procedures are even intensified. First, because the energetic contributions of single-site mutations are usually small, the need to sample multiple mutants – in which synergistic effects of several mutations are combined – is more acute. Second, the factors and principles responsible for the overall stability of a


native protein are still far from being completely understood, and a multitude of different forces and interactions contribute to it. A complete analysis will have to consider not only the interaction network of the native protein but also the effects on the denatured state. Additionally, potential aggregation pathways will have to be considered. Even simple substitutions like the ones discussed in the previous sections often make contributions to the entropy as well as the enthalpy of the folded and unfolded states, including the solvent in either state. Third, because rational approaches always rely on the current theoretical knowledge, new principles underlying protein stability will rarely be uncovered. In any case, rational engineering requires a clear definition of the problem by the investigator in order to find a solution. Ironically, in the field of protein engineering, the exact definition of the problem is often the problem itself. Thus, to overcome these limitations, an experimental setup is needed that allows the creation of a vast number of variants of a given protein and that subsequently can identify “superior” molecules that best fulfill a desired property. Nature samples the vast sequence space by the strategy of Darwinian evolution, a cyclic iteration of randomization and selection. Nature thereby adapts proteins to fulfill a function under the given environmental conditions. Recent developments in molecular biology have made it possible to mimic Darwinian evolution in a reasonable time in vitro. Not only has this “evolutionary approach” become the most powerful method to date for engineering proteins towards a desired property, but it also provides new insights into the mechanisms and principles that are responsible for this property. However, as has been illustrated in the case study on antibodies, such experiments can be used not only to solve a particular problem but also to gather information about which residues tend to become enriched in particular positions. This again provides a database for rational engineering. Many different selection and evolution strategies have been developed in recent years, but all of them have several features in common that reflect the principles of Darwinian evolution. The starting point is the generation of a genetic library of mutants derived from the wild-type sequence of the protein under examination. Several methods exist to create sequence diversity. In error-prone PCR, the error rate of polymerases is increased by performing the PCR reaction in the presence of deoxynucleotide analogues or in the presence of other metal ions. By using bacterial mutator strains, which are characterized by deficient DNA repair systems, random mutations are introduced during DNA amplification in the bacterial cell, albeit usually at a lower frequency [87]. In DNA shuffling, which mimics the natural process of sexual recombination, genes are randomly fragmented by nuclease digestion and reassembled by a PCR reaction in which homologous fragments act as primers for each other [88] (Figure 4). The staggered extension process [89] is another possibility to obtain recombined genes in vitro. Here, the polymerase-catalyzed extension of template sequences is extremely abbreviated, and repeated cycles of denaturation and extension lead to several template switches – thereby recombining elements from different genes – before the extension finally yields full-length products. Additionally, techniques are available to focus the diversity to certain regions on the whole gene, such as the use of degenerate primers in PCR or the “doping” of a


Fig. 4 Methods to create genetic diversity by biochemical means. On the left, errorprone PCR (see text) is depicted schematically. Two types of mutations are shown: favorable ones (open squares) and unfavorable ones (closed circles). In successive cycles of PCR, more mutations of each are introduced, and usually molecules will contain some of either type. Thus, the beneficial effect of the favorable mutations can be completely obscured by the presence of unfavorable ones, if the error rate is too high.

Therefore, this method is most successful if it is not used at too high an error rate. On the right, DNA shuffling according to Stemmer [88] is shown. A short DNAse digestion breaks up the DNA into small pieces, and PCR is used to reassemble the gene. Thereby, mutations are “crossed” and genes with largely favorable mutations can be obtained that can be enriched by selection. Nevertheless, successful evolution experiments have been carried out with either method.

shuffling reaction with degenerate primers. Depending on the problem and the sequence under investigation, each of these methods has its advantages and disadvantages. In any case, the generated library should obey two major criteria: it must be diverse enough to contain individual sequences with beneficial mutations, and it should be of high enough quality to reduce the experimental “noise” (such as sequences with stop codons or frameshifts) in the subsequent selection experiment [90]. A primary prerequisite of any selection system is the coupling of the genotype (the gene sequence) and phenotype (the respective protein displaying the properties) of any individual library member. Briefly, this can be done by two strategies. Either the gene and the protein need to be compartmentalized in cells or artificial compartments, such that the “improved” phenotype stays connected to the altered


Fig. 5 Two principal strategies to link phenotype and genotype are depicted. The genetic material must be connected in a unique way to the protein, which defines the phenotype, such that the gene encoding the valuable mutation can be selectively amplified. A collection of two mutant proteins and mutant genes is shown (light gray and dark gray). (a) This connection can be realized through a direct link (e.g., in phage display or ribosome display), as shown on the top. In this case, all assemblies can freely diffuse in the same volume and the selection must filter out those proteins with the desired function, e.g., by binding to a ligand. (b) Alternatively, gene and protein

must be in the same compartment. In nature, this is realized in cells, and microbial cells can be manipulated so that each takes up one variant of a mutant collection. The key is to identify the improved phenotype. This can be done by screening (individual assays on cells) or selection (giving cells with the desired property of the protein a growth advantage). Rather than natural cells, water-in-oil emulsions can be used to create artificial compartments of small water droplets in an oil phase. Usually, the emulsion must be broken and the proteins must be selected by binding to identify the one with the desired phenotype. For details, see text.

gene, or the protein has to be physically coupled to the gene, such that they can be isolated as a particle containing both gene and protein (Figure 5). By selecting the proteins displaying the desired properties, the linked gene sequence can be inherited and amplified subsequently. The various selection technologies differ mainly in the way this physical linkage or compartmentalization is achieved. This will be discussed in more detail in the following section. The library must then be screened or subjected to selection for a certain function, and a defined selection pressure can be applied to direct the selection towards a molecular quantity of interest. It is necessary at this point to discriminate between “screening” and “selection.” Screening methods examine individual members of a library for a given property (e.g., catalytic activity or solubility). For certain properties, screening is often the only way to go. The number of mutants to be screened in a reasonable time depends on the versatility of the screening method. Despite constant progress in automation and miniaturization, even the best screens to date usually do not allow assessment of more than 106 variants in a reasonable time. In contrast, selection methods force the single library members to compete with each other, and members that best fulfill the specified criteria are enriched. Often, the selection


is based on the binding of particular variants to an immobilized ligand. Note that the binding is only a “surrogate quantity” of the real property to be improved. The basis for a successful enrichment is an efficient counter-selection of variants that do not possess properties fulfilling these criteria. Especially during the selection of proteins for higher stability, an efficient counter-selection, in addition to the correct choice of the applied selection pressure, is one of the major experimental challenges [91]. The basic rule of screening and selection technology describes the importance of assigning the correct selection pressure. This has been succinctly phrased: “You only get what you screen for!” [92]. An analogous statement can be made for selection. Even though the rule sounds plausible, it is often difficult to translate this statement into an experimental setup, because an additional complication is introduced by the fact that an explicit selection pressure towards just one property is impossible to realize. Depending on the selection procedure used, more than one molecular property will have an influence on the enrichment process, including, for example, affinity or catalytic activity, thermodynamic stability, folding efficiency, and toxicity of the respective molecule. The outcome of the selection experiment will therefore always be a “compromise solution” with respect to the weighting of many properties in a given experimental setup. Assigning the right selection pressure therefore means biasing the selection towards a certain property, rather than exclusively altering this property. For these reasons we will discuss the application of the various selection technologies with an emphasis on the available selection pressures for stability engineering. Some of the described methods and examples will be based on mere screening of library members, but the main focus will be on the selection for favorable protein variants. Choosing the appropriate selection pressure is just one side of the coin. The proper adjustment of its strength is another important factor. If the selection pressure on the system is too low, molecules with the desired properties will be lost in the background noise of the experiment – i.e., they are not enriched. On the other hand, if the selection pressure is too stringent, even the best variants might not pass the “survival threshold.” The genes of variants that survive the applied selection pressures are subsequently amplified and subjected to another so-called “round of selection.” This either can be performed in the absence of further mutagenesis to simply enrich the best members of the initial library (which is constant) or, to completely mimic the Darwinian principle, the selected members can be subjected to alternating rounds of randomization or recombination before subsequent selection, leading to an adaptation of the library from round to round. However, as most of the mutations will be non-beneficial, care has to be taken that the mutation rate is low enough to allow successful enrichment of improved members and not to extinguish the whole population. We use the terms “combinatorial selection” for a process in the absence of such mutations and “evolutionary selection” for a process that includes such mutations. It has been pointed out that the method of DNA family shuffling is in some respects similar to the consensus approach because it combines gene fragments


Fig. 6 Common display systems used for selection for enhanced biophysical properties. In in vitro display technologies (a), the proteins are produced by in vitro translation, and protein production does not rely on a host organism. The displayed proteins are linked to RNA either by stabilized ternary complexes (ribosome display, upper panel)

or by a covalent puromycin cross-linker (mRNA display, lower panel), thereby establishing the genotype-phenotype linkage. (b) In contrast, partial in vitro display systems rely on cells to produce the displayed protein, but selections can be performed in vitro. In phage display (upper panel), the protein is displayed on the


from homologous sources in a random fashion. For mere statistical reasons, the probability of replacing a residue with a consensus residue by gene shuffling is also higher than replacing it with a non-consensus residue [93]. There is, however, a crucial difference: when recombining genes in a random fashion, a selection or screening step is needed subsequently to identify the “fittest” members of the resulting collection. Rather than being based on theoretical assumptions, gene-shuffling methods mimic the natural process of evolution by identifying the members exhibiting a desired property by explicitly subjecting them to selective pressure for this desired property. In practice, however, even with the largest of libraries, a full “re-equilibration” of residues will not take place. In contrast, the consensus approach is based on the explicit assumption that the statistical preferences in a given (often limited or biased) set of sequences indeed reflect the energetic preferences. However, there may be other reasons that certain sequences are prominent.

4.2 Screening and Selection Technologies Available for Improving Biophysical Properties

As described above, any selection technology relies on four major steps: (1) generation of a genetic library, (2) establishment of the link between genotype and phenotype upon translation into protein, (3) subsequent screening or selection under defined conditions, and (4) re-amplification of selected members. Depending on how these steps are performed, selection technologies can be subdivided into in vitro methods, partial in vitro methods, and in vivo methods. While in vivo systems rely on a host organism to express the protein and to carry the respective genetic information, all in vitro display technologies have in common that the protein production and the selection process are performed entirely in vitro, i.e., that the protein is obtained by cell-free translation. In the partial in vitro methods, the genetic information is introduced in cells, where protein production occurs, while the selection process is performed in vitro. All techniques differ in the way the physical linkage between the protein and its genetic information is established. The principles of linking genotype and phenotype are shown in Figure 6.

←-----------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------------Fig. 6 (Continued) surface of filamentous phages, usually fused to the CT domain of the minor coat protein g3p. The three domains (N1, N2, and CT) of the minor coat protein g3p are depicted. The phage particle also carries the gene encoding the displayed protein. In other partial in vitro technologies, the proteins are displayed on the surface of the expressing host cell itself, which are either bacterial cells (bacterial surface display) or yeast cells (yeast surface display)

(lower panel) that also harbor the respective plasmid DNA. (c) in vivo screening or selection systems use the properties of fused reporter proteins for screening (upper panel) or split proteins for intracellular selection (lower panel) in which cellular growth and therefore amplification of the genetic material occurs only if the two protein halves are reconstituted upon interaction of the fused library protein with its target.


4.2.1 In Vitro Display Technologies in vitro display technologies started off with the selection of peptides [94] and were then made efficient enough to select for functional proteins [95]. A number of technologies have been developed, which will be described briefly: ribosome display [95, 96], RNA-peptide fusions [97, 98], and water-in-oil emulsions [99, 100]. Water-in-oil emulsions constitute artificial compartments, rather than a physical link, and thereby create a phenotype-genotype coupling and can also use a selection step for interaction with a target. In ribosome display, the genetic library, usually in the form of a PCR product, is directly used to produce mRNA by in vitro transcription. This mRNA contains a ribosome-binding site for the subsequent translation of the protein and several features that stabilize it against degradation. The encoded protein variants are expressed in a cell-free translation reaction by a stoichiometric number of ribosomes. The essential linkage of the translated proteins to their respective mRNA molecule is achieved by eliminating the stop codon and stabilizing the ribosomal complexes, which prevents the release of the translated protein from the ribosome. In the related method of mRNA-peptide fusions, additional steps are used after this stoichiometric translation to covalently couple a linker between the end of the mRNA and the protein. The ribosome is then removed and the complexes are purified. These complexes are then used to bind to an immobilized target. In all these methods, after the selection process the re-amplification of genetic material can be performed solely by biochemical means, using reverse transcriptase to first synthesize RNA-DNA hybrids, which are then subsequently amplified in a PCR reaction. The resulting DNA then serves as a template for the production of mRNA used for the next round of selection (Figure 7). In vitro display technologies offer several advantages over in vivo selection systems. First, much larger library sizes are accessible because the creation of large molecular diversity on the genetic level does not pose a great challenge, and diversities up to 1013 are easily achieved. For in vivo systems, the critical step that limits the size of the displayed library is the transformation efficiency of the respective host organism. Only a small fraction of the initially generated pool will actually enter the cells, and, depending on the host organism, library sizes of about 107 variants in yeast or up to 1011 variants in bacteria can be achieved. In contrast, for in vitro selection systems such a transformation step is not necessary and the diversity of the library is defined by the number of different RNA molecules added to the cell-free translation reaction or by the number of functional ribosomes, whichever number is smaller. For a library of 1014 , a 10-mL translation reaction is required. A second advantage is that, because translation and protein folding take place in vitro, reagents can be added during protein synthesis, which can either promote protein folding or minimize aggregation of the displayed proteins (e.g., chaperones), or substances that exert certain selection pressures can be added. in vitro translation encompasses an additional advantage: many proteins are not compatible with in vivo selection systems because the wild type or at least some of the mutants are toxic to the host organism, undergo severe degradation, or cannot be expressed functionally in the respective cellular environment. Last but not least, a major advantage

Fig. 7 Selection cycle of ribosome display (left panel) and phage display (right panel). For ribosome display, all necessary steps are performed in vitro, whereas for phage display protein translation, folding, and phage assembly take place in the bacterial host cell. Other selection criteria can be applied to the protein of interest (see Figure 8) before selecting for binding to a target, thereby introducing selection pressures for higher stability. It has to be guaranteed, however, that the selection conditions are not too harsh to destroy the linkage of genotype and phe-

notype. Importantly, the genetic pool of library members in phage display remains constant in the course of several selection rounds, whereas in ribosome display, the genetic pool undergoes continuous modifications during the in vitro amplification steps by the limited accuracy of the polymerases, resulting in a true Darwinian evolution. In addition, this pool can deliberately and conveniently be altered by introducing additional mutations or recombination events such as DNA shuffling.



results from the amplification process after the selection has been performed. Because all enzymes used to convert and amplify the genetic material possess an intrinsic error rate, the genetic pool is never constant and is continuously modified from round to round. Therefore, even without special measures to increase the error rate, usually some “evolution” is observed. The error rates can be additionally increased by performing error-prone PCR or by recombining favorable mutations by means of DNA shuffling as explained in the previous section. 4.2.2 Partial in vitro Display Technologies We define partial in vitro display technologies as those methods in which host organisms are employed to carry and amplify the genetic information and to produce the proteins from this genetic library, usually encoded on plasmids. The selection step, however, is performed in vitro. Therefore, selection pressures similar to those in the in vitro methods can be applied. The array of available techniques includes phage display [101], bacterial surface display [102], and surface display on yeast [103]. Phage display is still the most popular selection technique, and due to its robustness, most of the studies that aim for enhanced biophysical properties of proteins have utilized this technique. Originally, phage display was applied as a method for the identification and selection of peptides or proteins binding to a specific target [101]. More recently, several developments paved the way for its application as a tool for studying protein folding, and as a result, it is now used for the selection of proteins with improved biophysical properties. We will briefly review the main principles of phage display in its most common format. Variants of the protein of interest are fused to the minor coat protein g3p of the filamentous bacteriophage, which consists of three domains (namely, N1, N2, and CT) that are connected by glycine-rich linkers. The fusion protein is usually encoded on a phagemid vector, expressed in the E. coli host and assembled. Typically, the other proteins necessary to produce an intact phage particle are provided by the helper phage. The assembled phages are secreted by the cell, which does not lyse, and the phages can be collected. While the protein is displayed on the exterior of the phage (typically as an N-terminal fusion to either N1 or CT), the gene of interest encoding a library member is packed into the phage particle upon its assembly in the bacterial host. The phages can then be subjected to the selection process. Phages compete with each other for binding to an immobilized target, using the displayed library members for interaction, and can thereby be captured. Optionally, before capturing, the phages can be subjected to harsh environmental conditions. The captured phages are subsequently amplified by re-infecting bacteria, which then produce phages for a new round of selection (Figure 7). In the ideal case, only phages that specifically recognize the target would be amplified. Due to reasons such as nonspecific “sticking” of phages to the surface, the efficiency of selection is in practice at most a 1000- to 10 000-fold enrichment over nonspecific molecules and can even be very much lower. Several aspects are important when phage display is used as a means of evolving proteins with improved biophysical properties. The correct folding of the fusion


protein during phage morphogenesis is an important parameter determining the frequency of incorporation of correctly folded proteins displayed on the phage. Folding intermediates that have a strong aggregation tendency lead to aggregation of a particular library member during the assembly process, and rather than the g3p fusion, the g3p wild-type protein from the helper phage will be incorporated. Despite this inherent selection pressure for the folding efficiency of displayed proteins, the enrichment of improved variants is slow [104], and the utility of phage display to study and improve folding of a given protein was initially not obvious [105]. Combined with a selection for protein functionality, it nevertheless sets the basis for selecting proteins according to their folding properties, as will be discussed in Section 4.3.2. Since the selection procedure is performed in vitro, a large variety of external selection pressures can be applied to the phage particles and be combined with functional selection. If function is omitted as a direct indicator for the native state, other criteria that directly correlate with the native state have to be translated into a selectable feature. This opens the door to select for proteins, which do not bind a ligand, albeit with the caveat that it is not assured that the native state is being selected. Such general selection approaches for physical properties take advantage of the fact that unfolded proteins are more susceptible to proteolytic cleavage than are compactly folded ones [106]. One variation on this theme makes use of the modular nature of the g3p protein, thereby linking phage infectivity directly to the proteolytic susceptibility of the target protein. This system has been called “Proside” (protein stability increase by directed evolution) [107] and will be discussed in Section 4.4.4. 4.2.3 In Vivo Selection Technologies In vivo selection technologies such as the yeast two-hybrid system [108, 109] or other split protein complementation assays [110, 111] are valuable tools for studying protein-protein interactions in living cells. They commonly employ reporter systems, in which a covalent link between the protein of interest and another so-called reporter protein is established. Cell growth or a colorimetric reaction is dependent on either a specific protein-protein interaction or the solubility of a critical component. The reporter protein transduces certain properties of the host protein, most importantly its native fold and its ability to interact and/or its solubility or its resistance to cellular proteases, to a screenable or selectable feature of the fused reporter protein itself [112]. Examples of reporter proteins are transcription factors [108], critical metabolic enzymes [111], or proteins that can easily be assayed [113]. Several facts, however, limit the applicability of in vivo systems for stability engineering. First, as the viability of the host organism has to be guaranteed, external selection pressures cannot easily be applied. Thermophilic organisms represent in some cases an interesting alternative, but conditions allowing selections are hard to establish. Additionally, the host environment as well as the fused reporter proteins possibly perturb the characteristics of the target protein. Furthermore, control experiments must ensure that growth is really dependent on the interaction


of interest and that mutations in the host have not “short-circuited” the selection strategy. 4.3 Selection for Enhanced Biophysical Properties

As has been explained in the Introduction, desirable biophysical parameters include solubility, stability, and folding efficiency. Even though the various selection methods and conditions usually do not improve one of these properties exclusively, they are often biased towards one of them. We will therefore discuss different ways to exert selection pressure on the system with respect to the property that is likely to be changed. 4.3.1 Selection for Solubility Solubility, correctly defined as the maximal concentration of the native protein that can be kept in solution, is usually not a property to be selected, as most proteins – with the exception of membrane proteins – are sufficiently soluble. The word “solubility” in the context of screening is often inaccurately used to refer to “soluble expression yield,” and thus it usually mirrors the efficiency of folding in the cell. Even though soluble expression yield is not necessarily equivalent to the folding properties and even less to the stability of a protein, a correlation can often be observed, and soluble expression yield is comparatively easy to screen for. In some cases, a function for a given protein cannot be assigned, or it is difficult and laborious to screen for, and then this property becomes especially important. The fluorescence yield of bacterial colonies expressing proteins or protein domains fused to the green fluorescent protein (GFP) correlates over a wide range with the soluble expression yield of the fused domain [114]. Screening for fluorescence intensity is a versatile method, and, combined with directed evolution, has the potential to select variants of proteins that are less aggregation-prone than their progenitors [115, 116]. In a similar setup, reporter proteins can be used as selection markers instead, such as by fusing the protein of interest to chloramphenicol acetyltransferase [117]. Assays exploiting protein-protein interactions can also be used for this purpose and they have the additional advantage that the fused reporter sequence can be much smaller and thereby minimize the risk of a perturbing influence of the reporter itself on the domain of interest [113]. To completely exclude such perturbing influences, reporter systems might possibly be established that do not need any fusion at all and exploit instead the stress response of the expression host cell. As the overexpression of proteins often activates particular stress response genes by the accumulation of insoluble aggregates, the respective gene promoters can be employed to activate reporter genes instead [118]. In special cases, reporter systems can also be combined with other selection methods to select for folding properties. Many intrinsically stable proteins contain permissive sites in loops, into which polypeptide chains of variable length can be inserted without loss of function of the host protein. The higher the number of residues inserted, the larger the entropic cost of ordering these residues will be.


Consequently, the overall stability of the host protein should decrease. However, the entropic cost of inserting a folded sequence will be lower than that of an unfolded sequence of the same length. The probability of the host protein reaching the native (functional) state should thus correlate directly with the ability of the inserted sequence to fold into a compact structure. If host proteins with various inserted sequences are displayed on phages and if the host protein allows a selection for “foldedness” by means of binding to a target, folded sequence insertions are enriched. This system has been termed “loop entropy reduction phage display selection” [119]. While attractive in theory, it was found experimentally that mostly sequences that keep the hybrid protein soluble are enriched. Protein solubility, as used here, can also be interpreted as a lower degree of exposed hydrophobic residues, and folded proteins usually display fewer hydrophobic residues on their surface than do unfolded proteins. Conversely, the exposure of hydrophobic residues is usually a sign of non-native states. Display technologies can thus, for example, use the interaction with hydrophobic surfaces to select against more hydrophobic, i.e., less “folded,” proteins [120]. In summary, protein solubility can reflect in many cases the folding properties of proteins and thus be used as a selection criterion. Moreover, soluble expression in vivo is often a major requirement for the large-scale production and the convenient in vitro handling of proteins. In any case, solubility should never be confused with protein stability. Even though both properties might correlate in some cases, the governing principles are often of a different nature. 4.3.2 Selection for Protein Display Rates The in vivo folding efficiency of proteins represents an intrinsic selection pressure when using phage display. Because correct folding is a prerequisite for both incorporation into the phage coat and binding to the target, the subsequent selection of phages that are able to bind to the target is partly influenced by the folding properties of the displayed molecule. However, this does not automatically imply that the selection is driven towards superior folding properties. As mentioned above, the enrichment factors of proteins with superb folding behavior over the poorly folding members are low [104]. Moreover, even though the binding of a given protein to its target is a very direct way to monitor and screen for its proper folding, the selection criterion “folding” is not decoupled from the selection criterion “binding affinity.” If the enrichment factor for one property (folding) is low, the selection is more likely to be driven towards the other (affinity). As a result, selected members will represent a compromise between folding properties, which only have to be sufficient under the given experimental conditions, and the binding affinity to their target. By randomizing exclusively the residues that build the hydrophobic core of the IgG binding domain of peptostreptococcal protein L, which are not involved in ligand binding but determine the stability and folding kinetics of this small protein, Gu et al. [121] could unambiguously demonstrate the utility of phage display for studying the stability and folding efficiency of proteins. A more extensive randomization effort with subsequent characterization of the selected variants pointed out


some important aspects of selection for folding [122]. First, it showed that folding kinetics is not the critical parameter for the selection but rather the overall stability of the mutants, a tendency which could be confirmed by more advanced selection approaches (see Section 4.4.2). Second, the authors demonstrated the utility of selection methods to identify certain residues that are important for the folding mechanism. Third, the selected pools were highly diverse and the thermodynamic stabilities of all variants were lower compared to the wild type. In fact, most of the selected proteins denatured just above room temperature. These results illustrate the principle that any selection – be it natural or in the test tube – simply continues until the minimum requirements are met, in this case, functionality at the selection temperature. Consequently, this suggests that additional, more stringent selection pressures are necessary to really accomplish a directed evolution for improved properties. 4.3.3 Selection on the Basis of Cellular Quality Control Additional selection pressure can be provided by the host organism itself. For example, the secretory quality-control system of eukaryotic cells discriminates proteins according to their folding behavior in an efficient way. This is based on mechanisms that lead to retention of misfolded proteins in the endoplasmic reticulum (ER), followed by degradation of these proteins. In yeast, the Golgi complex can reroute misfolded proteins, which have escaped ER retention, to the vacuole for degradation, thereby constituting an additional important quality-control pathway [14]. In combination with yeast surface display, these mechanisms can be employed to bias functional selections towards enhanced folding efficiency. Several studies have shown that the surface display rate of proteins strongly correlates with their ther-mostability and their soluble secretion efficiency [123, 124]. As an example, by making use of elevated temperatures during expression as an additional selection pressure, improved T-cell receptor fragments, whose thermostability exceeded by far the expression temperatures, could be obtained [125]. Even though the low transformation efficiency of yeast restricts the accessible library size, one obvious advantage of the method is the applicability to glycosylated eukaryotic proteins, which generally are not amenable to yeast two-hybrid or phage display methodologies [126]. 4.4 Selection for Increased Stability 4.4.1 General Strategies The key to all selections for stability is to introduce a threshold that separates the molecule with desired properties from the starting molecules. If the population initially lacks functionality, while only a few members are above the selection threshold, the selection system can distinguish between these slight energetic differences. This is the starting situation if the target protein is initially of very low stability and should be brought to “average” properties.


If, however, the starting protein is already of considerable stability, but should be brought to even higher stability, it can be advantageous to intentionally destabilize it prior to selection in order to find stabilizing mutations that reconstitute its functionality. This principle can be applied to very different kinds of proteins, provided that mutations are known that destabilize the protein to an extent that will subsequently allow the selection for alternative stabilizing mutations (Figure 8a). Because the effects of independent stabilizing mutations are often additive, the deliberately introduced destabilizing mutations can be reverted in the context of the additional newly selected ones, thereby rendering the molecule far more stable than the original one (Figure 8b). In principle, all reagents and conditions known to destabilize proteins can decrease the number of library members populating the folded state and can therefore be used to exert increasing selection pressure on the system. Increased temperature and denaturing agents are obvious methods that are useful for selecting proteins of higher thermodynamic stability. The major problem is presented by the compatibility of the conditions with the selection method used. 4.4.2 Protein Destabilization An example of the stabilization of a naturally unstable domain by using phage display was a selection performed with the prodomain of the protease subtilisin BPN [127]. At room temperature the prodomain folds into a stable conformation only upon binding to subtilisin. By randomizing positions that are not directly in contact with subtilisin and subsequent selection on subtilisin, a mutant was selected that showed an increase of Gunfolding by 25 kJ mol−1 , from −8 kJ mol−1 to 17 kJ mol−1 , despite the fact that the library size was comparatively small. Intriguingly, the predominant energetic contribution was mediated by a selected disulfide bond. Previously, a similar strategy had been used to select for thermodynamically favored β-turns of the B1 domain of protein G [128]. Based on considerations described above and on the fact that the replacement of amino acids on the surface of proteins would be predicted to have only moderate effects on stability, the authors reasoned that such a selection could be successful only for proteins of marginal stability. Most of the substitutions would then lead to a positive free energy of folding, and thus the molecule would fail to fold into a functional form at all. In fact, selected turn sequences showed clear sequence preferences only if less stable host proteins were used to accept the turn. In this case, the selected sequences either resembled the wild-type turn or reflected the statistical preferences of turn sequences in the databases and stabilized the protein by 12–20 kJ mol−1 , compared to random sequences. Moreover, increased temperature was used as an additional selection pressure during the phage selection. As discussed in the previous section, the inability of the wild-type target protein to fold is the prerequisite for performing an efficient positive selection for functionality by additional mutations. The internal disulfide bond of immunoglobulin domains significantly contributes to the stability of antibodies [129]. The dramatic loss of free energy of folding upon their removal usually renders antibodies nonfunctional. By first destabilizing a scFv antibody fragment through replacement

Fig. 8 Principle of selection for proteins with improved stability. (a) By random mutagenesis of a given sequence, many mutations are obtained. Some of these mutations are favorable for the stability of the native protein and others are unfavorable. As a result, a diverse pool of proteins with a distribution of different energies is created. Upon exposure of the pool to an external selection pressure, only mutants with stabilities exceeding the selection threshold can be recovered and amplied. However, the selection threshold has to be set high enough to allow an efficient selection of “improved”

members. (b) Destabilization of the wild-type sequence prior to selection allows reducing the necessary selection threshold. Thereby, alternative mutations that stabilize the native fold of the protein can be identited and the selection process becomes more efficient. Moreover, additional stability gains can be achieved by removing the deliberately introduced destabilizing mutations after the selection. The initially lost energy is therefore regained, resulting in mutants of higher stability than the wild-type protein. Adapted ¨ et al. [76]. from Worn



of cysteines with other residues in both variable domains separately, a completely disulfide-free antibody was identified after several cycles of functional selection and recombination by DNA shuffling [17]. One globally stabilizing mutation was found to compensate for the initial stability loss. This study illustrates some important features of how several parameters exert influence on different stages of the selection process. Interestingly, the stabilizing mutation was already selected during the first rounds, showing that the loss of thermodynamic stability was the primary problem that had to be overcome. Because the stability gain of this mutation was large enough to shift the free energy of folding above the required threshold, all successive rounds did not affect protein stability but led to a fine-tuning with respect to the improvement of folding yield [17]. To illustrate the principle of additivity, reintroduction of the disulfide bridge into the selected variant yielded a scFv antibody of very high stability and superior expression yields [84]. 4.4.3 Selections Based on Elevated Temperature in vivo selection systems using thermophilic expression hosts have been employed for the stabilization of enzymes [130]. However, a rather large set of requirements must be met to apply such a selection system. In order to use stability of the enzyme at elevated temperatures as the selection criterion, its enzymatic activity must be vital to the thermophilic organism, and the corresponding gene of the thermophilic host has to be deleted. Randomized versions of a mesophilic enzyme can then be screened by means of metabolic selection. Although the approach is very powerful, the utility of these systems is restricted to special cases. To evolve enzymes with altered thermal stability, conventional screening for enzymatic activity in vitro after randomization or recombination of the respective gene and subsequent expression of the enzyme is still the most widely used method. An activity screen rapid and sensitive enough to identify slightly improved members from a vast pool of mutants is the key feature of this directed evolution approach [131]. However, individual screens have to be developed for each specific class of enzymes. Even though screening limits the explorable sequence space considerably compared to selection, mesophilic enzymes such as subtilisin E [132] and p-nitrobenzyl esterase [133] could be converted into mutants functionally equivalent to thermophilic enzymes by only a few rounds of directed evolution. Similar to observations discussed before, only very few mutations were necessary to increase the melting temperatures by more than 14 degrees. An important feature of screening compared to selection is that the much smaller library size is partially compensated for by directly measuring the quantity of interest: enzymatic activity at elevated temperature. In contrast, most selection systems use a surrogate measure where “false positives” can lead to the phenotype by mechanisms different from the ones desired. Elevated temperatures not only can be used in screening but also can be combined with selection technologies. While ribosome display is not an option – because low temperatures are essential to keep the ternary complexes of RNA, ribosome, and nascent polypeptide intact and thus ensure the coupling of genotype and phenotype – phage display has proved to be a very suitable method for harsh conditions


due to the robustness of the phage particles. Nevertheless, future improvements of in vitro technologies may allow their application under more stringent conditions. As of today, in the case of in vitro technologies, it is vital to destabilize the protein first (see Section 4.4.2); then, very significant stability improvements can be selected [134]. The upper temperature limit for selections using filamentous M13 phages is approximately 60 ◦ C [135]. Above this temperature, re-infection titers of the phages are severely decreased, presumably due to the irreversible heat denaturation of phage coat proteins. Phages displaying the protein of interest can be incubated up to this temperature, and proteins still able to function can be selected. It should be noted that the exposure of phages to higher temperature after phage assembly exerts a somewhat different stress on the proteins than in the methods described before. While in the previous examples the functionality of the proteins was influenced mainly by in vivo folding efficiency and the protein stability during the panning procedure, it is the irreversible unfolding reaction at a given temperature that now becomes an additional parameter. The fraction of unfolded molecules will therefore reflect the rate of unfolding and thus kinetic stability. 4.4.4 Selections Based on Destabilizing Agents The use of protein destabilizing agents for biasing the selection pressure towards stability is limited to in vitro and partial in vitro display methods. Like temperature, the concentrations of denaturing agents can be controlled precisely and can be varied from round to round, allowing a gradual increase of stringency. Even though phages are quite resistant to denaturing agents [136], one should be aware of possible general problems when selections are based on the chemical denaturation properties of the displayed proteins. However, if chemical denaturation is combined with a selection for binding, high concentrations of denaturant can prevent binding to the target, even if the protein is not yet unfolded: as the forces governing ligand binding are very similar to those responsible for protein stability, both are disturbed by chemical denaturants. Conversely, because chemical denaturation is in fact often reversible, removal or dilution of the denaturant prior to ligand binding will often result in refolding on the phage and thus release of selection pressure. Moreover, ionic denaturants such as guanidinium chloride weaken electrostatic interactions and strengthen hydrophobic ones. This might impair the selection of mutations that introduce additional ionic interactions on the protein surface [59] and may favor additional hydrophobic interactions, which is not necessarily desired. If the protein is known to be stabilized by disulfide bridges, a strategy similar to the one used by Proba et al. [17] described above may be applicable. To identify globally stabilizing mutations, which compensate for the stability loss upon removal of disulfide bridges, selection can be performed in the presence of reducing agents such as DTT. Thus, the disulfide-forming cysteines do not have to be removed in advance. However, one should be aware of the fact that some disulfide bridges in proteins, once formed, are often hard to reduce, especially if they are buried within the protein core. Thus, it is advantageous to add the reducing agent at a time


when the protein is not yet folded. Jermutus et al. [134] stabilized a scFv antibody fragment by using ribosome display. In contrast to the phage display method, the synthesis of the targeted protein occurs in vitro, which makes the polypeptide chain accessible to reagents during its synthesis on the ribosome and before folding has taken place. DTT was added during translation of the scFv, and its concentration was continuously increased from round to round. When using ribosome display, the population undergoes slight changes due to mutations occurring during PCR, resulting in an iterative adjustment of the selected variants towards tolerating the increasingly stringent conditions. 4.4.5 Selection for Proteolytic Stability In each of the examples cited above, selection for stability was based on a functional selection. Therefore, only those proteins for which a specific function can be assigned could be targeted; in addition, this function has to be screenable or, better yet, selectable. It would be highly advantageous to completely uncouple function from stability in the selection process to extend the range of problems to which this can be applied. A more general approach would thus be an invaluable tool for engineering any given protein and for selecting stable folds from a pool of de novo designed proteins. For functional selections, an additional problem arises from the fact that after a certain stability threshold is reached, allowing enough proteins to populate the folded state, the selection is likely to run towards improved binding properties instead. On the other hand, certain mutations might be stabilizing but might result in slight structural rearrangements that affect functionality in a negative way. Thus, it will not be possible to recover these mutations [106]. A general approach that has proved to be powerful for stability engineering is based on the concept that compactly folded proteins are much more resistant to proteolysis than are partly folded or unfolded proteins [137]. Several variations on this theme have been applied to phage display selections. In principle, it is only necessary that the phages displaying proteins, which can be cleaved by the protease, can be efficiently separated from the phages displaying protease-resistant proteins. This can be achieved in a physical way by providing the displayed protein with an N-terminal tag sequence allowing capture of only phages with non-cleaved proteins [106]. Alternatively, the ability of phages to re-infect bacteria can be directly linked to the protease resistance of the protein of interest [107, 136]. This second alternative takes advantage of the modular structure of the protein domains responsible for phage infection. If the displayed domains are inserted between the carboxy-terminal domain of gp3 and the amino-terminal domains N1 and N2, which are required for phage infectivity, proteolytic cleavage of the displayed domain renders the phage noninfectious. This was inspired by the so-called selectively infective phage method (SIP), where it was shown that the g3p domains could be interrupted by additional domains and even an interacting pair [138]. By employing a phage system derived from the SIP technique, which lacks any wild-type g3p, it is assured that infectivity is completely abolished upon proteolytic cleavage [107] (Figure 9). Alternatively, phages can be engineered to make the remaining wild-type g3p itself susceptible to the proteolytic attack [136]. The principle of protease selection can also be


Fig. 9 Strategies for selecting for improved protein stability and folding by phage display. (a) Methods utilizing the binding to a given target (affinity selection) as a means for selecting members with improved folding behavior and higher stability from a protein library. Several types of selection pressure can be applied in order to recover mutants with enhanced stability. Destabilizing mutations are deliberately introduced (top) to render the protein “nonfunctional,” and alternative stabilizing mutations are identified by selecting variants whose functionality is regained. Alternatively (bottom), elevated temperatures or denaturing agents can be used to render most of the protein variants “nonfunctional,” allowing the selection of simply the “fittest” members. (b) The resistance of the target protein to a protease can be combined with the ability of phages

to re-infect bacteria. The protein library is inserted between the C-terminal domain and the N-terminal domain in all copies of the g3p protein (cloned into the phage genome), which are needed for reinfection of bacterial cells. By proteolytic cleavage of the inserted protein, these domains are cut off and phage infectivity is lost. In alternative approaches, an N-terminal tag sequence is fused to the protein of interest instead of the N-terminal g3p domains. Upon proteolytic cleavage, the tag is lost. In contrast, phages presenting proteins that are resistant to proteolytic cleavage can subsequently be captured on an affinity matrix binding to the tag sequence. In order to further increase the selection threshold, the shown selection strategies can also be combined.

5 Conclusions

applied to in vitro selection techniques in which the displayed protein is freely accessible [120]. Even though proteolysis seems at first glance to be less correlated to the stability of proteins than temperature, it is suitable for optimizing packing of the hydrophobic core [139] as well as for optimizing electrostatic interactions on the protein surface [59]. One reason for the strength of this approach is based on the fact that proteolytic cleavage is an irreversible reaction, which is not necessarily the case for denaturation by temperature or denaturing agents. Furthermore, protease resistance monitors the flexibility of the polypeptide chain rather than complete denaturation. It is therefore capable not only of selecting against completely unfolded variants but also of detecting local unfolding events. Because sites of local unfolding often initiate the global unfolding process, it can be advantageous to remove such sites. Nevertheless, the effective cleavage of flexible parts of the protein restricts the method to proteins that do not have extended flexible regions in the native state. Furthermore, a selection against the primary recognition sequence of the protease is clearly a possible outcome of such experiments. None of the described methods can stand completely on its own. Many methods can easily be combined to increase the stringency of a selection. A combination of temperature stress and increasing amounts of denaturing agents may, for example, lead to higher flexibility in certain regions of the protein, thereby increasing the sensitivity of the subsequent proteolytic attack. Additionally, temperature and the concentration of denaturing agents allow a much tighter control of the selection pressure than do increasing concentrations of protease and thus open the possibility of a well-controlled gradual increase of selection stringency.

5 Conclusions

Several different approaches are now available for engineering proteins for enhanced biophysical properties. In many cases, few and specific mutations are sufficient to provide proteins of marginal stability with considerably stabilizing features. Thus, the challenge for protein engineering is to identify these positions in a given sequence among the vast number of possible changes and to correctly alter them. Each of the applied techniques has its own merits and bottlenecks. Instead of playing them off against each other, the future challenge will be to find ways to synergistically use them to improve a given molecular property. Evolutionary methods have the advantage of being much less biased by theoretical assumptions or working hypotheses. Additionally, proteins stable in new environments may be evolved, e.g., proteins that fulfill a given function in nonaqueous solutions or high concentrations of detergents. Because the biophysical principles in such environments are of a different nature, rational design has to rely on a much smaller empirical dataset. While rational approaches are likely to become more important as more structural and experimental data become available, notably also those from selection experiments, the large number of variations

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that are potentially able to improve the biophysical properties of a protein often still exceeds the experimentally accessible number. Moreover, because rational engineering has to rely on the available dataset, mutations that lie off the beaten track will rarely be identified. Nevertheless, selection experiments often identify the same “key” mutations in proteins. It is then useful to exploit this information and directly introduce such mutations. A “rational” analysis can also help to recombine important “key” mutations in selected clones and to reduce the effects of a selection-neutral genetic drift. Another combination of rational and combinatorial methods is the creation of “smart” libraries of variants. Library design that is based on such structural considerations and principles will therefore allow more accurate focusing of the selection on specific regions of interest and thereby increase the chances for success. Acknowledgements

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1

Protein-based Artificial Enzymes Ben Duckworth and Mark D. Distefano

University of Minnesota, Minneapolis, USA

Originally published in: Artificial Enzymes. Edited by Ronald Breslow. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31165-1

1 Introduction

The design of catalysts that rival the specificity and speed of natural enzymes is a challenging objective. Various approaches have been employed to create such molecules. Polypeptides are a logical choice of possible framework for catalyst design because they provide a simple means for generating complex structures. The catalyst structure can be changed by altering the corresponding gene and expressing the desired protein in bacteria. However, chemical methods can also be incorporated into such an approach to greatly expand the range of possible reactions that can be studied. In such a chemogenetic strategy, a small molecule-based catalyst is coupled to a protein scaffold devoid of catalytic activity. The small molecule provides intrinsic reactivity while the protein component controls specificity; the protein can also be used to tune or augment the reactivity of the small molecule catalyst. Such an approach provides enormous flexibility in the design of new catalytic materials. Early efforts in this field focused on using chemical modification to alter the type of reaction catalyzed by existing enzymes or to change their substrate specificity. Thiosubtilisin [1, 2], selenosubtilisin [3] and flavopapain [4] are all examples of proteins that manifest altered reactivity resulting from chemical modification. Other work with sta-phylococcal nuclease [5] and subtilisin [6, 7] employed chemical modification to modulate substrate selectivity. While that pioneering work served as the foundation for protein-based catalyst design, these efforts were, notably, based on protein frameworks that possessed native catalytic activity. Such proteins already contained highly evolved substrate binding sites and/or catalytic machinery. The present chapter focuses on the design of protein-based catalysts starting with polypeptides that have no intrinsic catalytic activity. While this is an enormous challenge when compared with approaches starting with existing enzymes, it opens Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Protein-based Artificial Enzymes

up a plethora of exciting possibilities; if it could be accomplished, the ability to create catalysts with enzyme-like properties would have dramatic effects that would extend well beyond the field of chemistry. Several recent articles provide a more comprehensive review of the field of protein design, including work with existing enzymes [8–19]. 2 Artificial Nucleases Based on DNA and RNA Binding Proteins 2.1 Introduction

The process of cleaving the phosphodiester bond of RNA and DNA is a vital and widespread phenomenon in nature. Scission of the nucleic acid backbone plays crucial roles in myriad in vivo and in vitro processes, including mutational repair, cell death, cloning, and sequencing. Much attention has been given to the discovery and development of nucleases, both natural and artificial, which might benefit the fields of biotechnology and pharmacology [20]. Several chemical agents possess nuclease activity. However, such compounds have low affinity for oligonucleotides and may not be optimally positioned for attack and scission of the backbone. Therefore, if a chemical nuclease could be incorporated into a DNA or RNA binding protein, substrate binding and nuclease activity would be enhanced. 2.2 Artificial Nucleases from Native Protein Scaffolds

To create an artificial nuclease, Sigman and co-worker enhanced the nuclease specificity of 1,10-phenanthroline(OP)-copper by covalently linking the moiety to the Escherichia coli trp repressor (TrpR) [21]. OP is a robust chemical nuclease that cleaves the phosphodiester bonds of nucleic acids. A copper-oxo species attacks the C-1H of the deoxyribose in the minor grove, yielding 3 - and 5 -phosphomonoesters, free base, and 5-methylene furanone (5-MF) (Figure 1) [22, 23]. This makes it possible to convert DNA-binding proteins into nucleases by conjugation with OPcopper. Since most DNA/RNA binding proteins bind the major groove, it is crucial for the OP-copper group to interact solely with the minor groove to avoid disruption of key binding interactions. To conjugate E. coli trp repressor with OP-copper, the four lysine amino groups were first converted into sulfhydryl groups by reaction with 2-iminothiolane hydrochloride in the presence of the corepressor L-tryptophan (Figure 2) [21, 24, 25]. The thiols were then alkylated with 5-(iodoacetamido)-1,10-phenanthroline (IOP, Figure 2-1), yielding the TrpR-OP conjugate. DNase I footprinting experiments indicated that the DNA-binding ability of TrpR was not jeopardized after modification with OP. Early experiments involved the incubation of trpEDCBA operator with OP-TrpR copper complex for 20 h, which resulted in double-strand scission as well as single-strand nicks in 50% of the labeled strands [21].

2 Artificial Nucleases Based on DNA and RNA Binding Proteins

Fig. 1 Shortened chemical mechanism for DNA cleavage by 1, 10-phenanthroline-copper complexes.

Fig. 2 (A) Procedure for the attachment of 1,10-phenanthroline to proteins via lysine derivatization. (B) 5-(iodoacetamido)-1, 10-phenanthroline (IOP, 1) and 5-(iodoacetylglycylamido)-1, 10-phenanthroline (IAOP, 2).

3


2.3 OP Nuclease Design by Mutagenesis and Chemical Modification

An additional approach to artificial nuclease design is cysteine mutagenesis followed by conjugation with the desired catalytic group. This avoids the need to convert amino acid residues into reactive thiols, while allowing precise control over the placement of the reagent within the DNA-binding protein. Sigman and coworkers applied this strategy to create an artificial nuclease using the bacteriophage λCro protein [26]. When complexed with DNA, the C-terminal arm binds within the minor groove and is close to the C-1 hydrogen of the deoxyribose on either DNA strand [26–30]. Therefore, an alanine close to the C-terminus was mutated to a cysteine and derivatized with IOP. After incubation with the Cro A66C-OP conjugate, 40% of the 17 base pair OR-3 operator site was cleaved within 10 min [11]. Thus, placement of the phenanthroline group close to the DNA substrate ensured efficient nuclease activity. Recently, the carboxy terminal domain of NarL, NarLC , was modified with 1,10phe-nanthroline. NarL is a response regulatory protein of E. coli and binds to a heptameric consensus sequence [31]. Two residues of the C-terminal domain, which were mutated and conjugated with IOP (NarLC K201C-IOP, NarLC K211CIOP), showed high site-specific cleavage efficiency of the top strand (Figure 3). When the mutated NarLC was modified with IAOP (Figure 2-2), a similar DNA

Fig. 3 Model of the NarLC -DNA complex showing the locations of amino acids 201 and 211. Two molecules of NarLC are positioned above the major groove of the DNA. Color scheme: Protein secondary structure (green), DNA (white), residue 211 (pink), residue 201 (orange).

2 Artificial Nucleases Based on DNA and RNA Binding Proteins

cleavage pattern was observed on the bottom strand. This IAOP conjugate contains a 4 Å longer linker arm than does the previous IOP conjugate, which allows the OP-Cu complex to extend and cleave the adjacent bases of the DNA substrate. These artificial cleavage enzymes could ultimately be used to identify the number, position and orientation of NarL monomers on several promoters and may be extended to other regulatory proteins. These results highlight the promise of chemical nucleases as tools for biological investigation. 2.4 Additional Applications for OP Conjugates

Other research groups have also created artificial nucleases by OP conjugation. Johnson and co-workers created an artificial nuclease, which aided in the understanding of DNA conformational changes upon protein binding [32]. E. coli Fis (factor for inversion stimulation) was chosen as the protein scaffold and binds to DNA with a low sequence specificity. X-Ray crystallography and electrophoretic mobility studies indicated significant bending of the DNA upon Fis binding [33, 34]. To aid in the study of DNA-Fis complexes, OP-Cu was conjugated to four separate mutated cysteines. Of these four conjugates, N98C-OP and N78C-OP displayed nuclease activity. Scission patterns from these two nucleases not only confirmed earlier studies of DNA bending, but also revealed that the DNA wrapped around the Fis dimer. The E. coli catabolite gene activator protein (CAP) binds to a 22 base-pair, twofold symmetric DNA recognition site with high affinity (K A = 4 × 1010 M−1 ) [35]. Ebright and co-workers successfully converted CAP into a site-specific DNA cleavage enzyme. In their first attempt, OP was conjugated to the only solvent accessible cysteine (C178 ) of CAP. The catalytic group was placed at this position so that intra-CAP interactions would not be disrupted, CAP-DNA contacts would not be comprised, and OP would be in close proximity for a favorable attack on the phosphodiester bond. The binding constant for this conjugate was 1 × 108 M−1 , indicating that the DNA binding ability of CAP was not significantly lost due to conjugation with OP. The CAP178-OP conjugate cleaved the DNA site at four adjacent nucleotides (Figure 4).

Fig. 4 DNA cleavage sites produced by the CAP178–1, 10-phenanthroline conjugate. Longer arrows indicate sites where greater cleavage efficiency occurred.

5


Earlier work indicated that in the specific CAP-DNA complex the DNA was bent 90◦ away from the protein but was not distorted in the nonspecific complex. (CAP bound to its noncognate DNA site) [3, 36]. This DNA bending phenomenon was exploited to construct a new CAP-OP nuclease capable of single site cleavage [37]. The crystal structure of the specific CAP-DNA complex revealed that amino acids 24–26 and 89–91 of CAP were close to the DNA substrate [38]. With this in mind, residue 26 was mutated to a cysteine and modified with IOP. While the catalytic group neighbors the DNA in the specific complex, OP is too far from the substrate in the nonspecific complex, and thus cannot cleave the DNA. Ultimately, conjugation of OP at position 26 yielded an artificial nuclease that proceeded to ∼90% cleavage with no detectable nonspecific cleavage at distal sites.

2.5 A Fe-EDTA Artificial Nuclease

An alternative DNA-cleaving agent was used to identify the σ subunit DNA contact sites of E. coli RNA polymerase [39]. Iron-EDTA protein conjugates have previously been shown to cleave DNA by producing hydroxyl radicals, which ultimately attack the deoxyribose backbone [40]. To create this artificial nuclease, Minchin and co-workers mutated several residues within the σ subunit to cysteines. These residues were conjugated with (S)-1-[p-(bromoacetamido)benzyl]-EDTA (BABE) (Figure 5). By identifying the cleavage products of several related promoters, this Fe-BABE conjugate confirmed the location of the σ subunit-promoter DNA contact sites.

2.6 Concluding Remarks

Through direct conjugation of chemical nucleases to DNA binding proteins, several groups have successfully created artificial nuclease from proteins with no

Fig. 5 Procedure for the derivatization of proteins with (S)-1[p-(bromoacetamido)benzyl]-EDTA (BABE).

3 Catalysts Based on Hollow Lipid-binding Proteins 7

native nuclease ability. These nucleases can utilize the favorable protein–DNA binding elements and can position the hydrolytic metal close to the DNA backbone for efficient cleavage [41]. These nucleases have provided useful biological insights and a better understanding of the molecular details involved in protein–DNA interactions.

3 Catalysts Based on Hollow Lipid-binding Proteins 3.1 Lipid-binding Proteins

Lipid-binding proteins are a class of molecules found in eukaryotic cells involved in the transport of fatty acids and other types of hydrophobic compounds. The protein structure consists of two orthogonal planes of β-sheets that form a cup-shaped cavity that is capped off with a helix-turn-helix element. Considerable structural data, obtained from X-ray and NMR analysis, exists for this family of proteins [42, 43]. Of particular interest is the presence of a large, solvent sequestered, cavity whose overall volume varies between 500 and 1000 A3 depending on the exact identity of the protein; with the exception of certain membrane-bound channel proteins, there are few other examples of macromolecules that possess such a cavity. This structural feature serves as the ligand binding site in the protein cavity for a diverse range of molecules. Such a large, solvent sequestered, binding site also provides a useful scaffold for the design of catalysts. Formally, this cavity can be viewed as a protein equivalent to the cyclodextrin template used in much of the pioneering work of Breslow and co-workers in their development of enzyme mimics [18]. 3.2 Initial Work

In their initial work Distefano and co-workers used the protein ALBP (adipocyte lipid-binding protein) for their protein scaffold. This protein contains a unique cysteine residue a position 117 that can be selectively modified using reagents that capitalize on the unique reactivity of the thiol side chain. Kuang and co-workers developed a reagent, TP-PX (6-1) that contained an activated disulfide suitable for protein derivatization. This molecule was used to incorporate a PX moiety into ALBP at position 117, resulting in the formation of a construct denoted ALBP-PX (Figure 6, 6-2) [44]. This semisynthetic biocatalyst aminated reductively various α-keto acids (7-2) to ami-no acids (7-4) with 0 to 94% ee (Figure 7). Because these reactions were performed in the absence of any additional amine source, only single turnovers were obtained. The reaction rates were not, however, significantly faster than those involving free pyridoxamine (7-1b). This suggested that the protein


Fig. 6 Derivatization reagents for the preparation of pyridoxamine conjugates of fatty acid binding proteins.

cavity functions as a chiral environment that controls the facial selectivity of the protonation of the aldimine intermediate without forming specific interactions with the bound pyridoxamine cofactor, which could accelerate the reaction, as confirmed by a X-ray crystal structure [45]. Modeling of the Schiff base complexes with several amino acids indicated that one face of the putative aldimine intermediate was protected against the approach of the solvent or buffer molecules that must be the proton source for the reaction given the lack of suitable functional groups within the cavity. This structural data provided a rationale for explaining the enantioselectivity observed in the ALBP-PX system. 3.3 Exploiting the Advantage of a Protein-based Scaffold

The next step in developing these protein-based catalysts was to take advantage of the synthetic flexibility provided by a protein scaffold. Since the protein is encoded by a DNA sequence, changes can be made in the gene that will result in modifications in the protein. Mutagenesis and other genetic methods can produce alterations in the protein framework much more rapidly than the organic synthesis

Fig. 7 Single turnover, reductive amination, reactions promoted by fatty acid binding protein–pyridoxamine conjugates.


required to make changes in synthetic scaffolds. Initially, the position of catalyst attachment within the cavity was varied to probe its effect on reaction rate and selectivity. For these experiments, a different fatty acid binding protein (IFABP) was employed. The wild-type IFABP contains no cysteine residues in its primary sequence and is, hence, a useful template for the introduction of single cysteine residues at different positions [46]. Cysteine residues were introduced at several positions, including V60 , L72 , and A104 , using site-directed mutagenesis. Figure 8 gives the locations of these mutations within the protein structure. The three different mutant proteins described above were purified and the corresponding conjugates were prepared using the TP-PX reagent (6-1). The conjugates were then evaluated in single turnover conditions (Figure 7) with several α-keto acids to produce amino acids in enantiomerically enriched form. This collection of catalysts exhibited various differences in reactivity and selectivity. Compared to ALBP-PX, IFABP-PX60 reacted at least 9.4-fold more rapidly, while IFABP-PX72 displayed opposite enantioselectivity, and IFABP-PX104 showed a clear selectivity preference for unbranched substrates [47]. From these experiments, the position of cofactor attachment is clearly an important parameter in modulating the reactivity and specificity of these biocatalysts. Moreover, these results underscore the utility of site-directed mutagenesis and the power of using a protein-based scaffold for catalyst development.

Fig. 8 Stereo view showing positions 60, 72, 104 and 117 in IFABP that have been used for the attachment of pyridoxamine and other catalytic groups. From top to bottom: A104, Y117, L72 and V60. Color scheme: Protein secondary structure (green), carbon (white), oxygen (red), nitrogen (blue).


Fig. 9 Multiple turnover reactions that produce chiral amino acids from achiral keto acids.

3.4 Catalytic Turnover with Rate Acceleration

Due to its intriguingly more rapid reaction rate than ALBP-PX and free pyridoxamine, IFABP-PX60 was subsequently studied further [48]. Under single turnover conditions, it converted α-keto glutarate (Figure 9, 9-1) into glutamic acid (7-4) 62-fold faster than free pyridoxamine (7-1b); this was determined by evaluating the extent of conversion at much shorter reaction times. This more rapid reaction rate under single turnover conditions suggested that this construct might be capable of catalytic transamination. Using tyrosine or phenylalanine (9-2) as the amine source to recycle the cofactor from the pyridoxal form (7-3a) back to the pyridoxamine form (7-1), L-glutamate (9-3) was formed with an enantiomeric purity of 93% ee. As many as 50 turnovers were observed with long reaction times (14 days). Both kcat and K M for the reaction were determined by kinetic analysis (Table 1). Comparison with similar parameters obtained from reactions with free pyridoxamine indicated that IFABP-PX60 catalyzed transamination some 200 times more efficiently. Analysis of the specific kinetic constants kcat and K M indicated that the observed rate acceleration was due mostly to an increase in substrate binding Table 1 Kinetic constants and catalytic efficiencies for semi-synthetic transaminases based

on fatty acid binding proteins

PX MPX IFABP-PX60 IFABP-MPX60 hsIFABP-PX60 IFABP-PxK38 IFABP-MPxK38 IFABP-PxK51 IFABP-MPxK51 IFABP-Px126 IFABP-MPx126 IFABP-Px126/14 IFABP-MPx126/14

KM (mM)

kcat (h−1 )

kcat /KM (h−1 mM−1 )

73 38.7 1.8 6.8 10.2 0.81 13.7 0.24 8.9 5.5 40 6.0 48

0.032 0.031 0.29 0.23 0.22 0.44 1.12 0.44 0.52 0.18 1.10 0.11 0.78

4.4 × 10−4 8.0 × 10−4 0.16 0.034 0.022 0.54 0.08 1.83 0.06 0.03 0.03 0.02 0.02


(50-fold), with a smaller effect on the maximal rate (4-fold). While this is an impressive result, the absolute magnitude of kcat /K M (0.02 s−1 M−1 ) makes it clear that this catalyst is still quite primitive compared to natural enzyme systems that occasionally operate with catalytic efficiencies near the diffusion limit. 3.5 Modulation of Cofactor Reactivity with Metal Ions

Model studies with pyridoxamine and related analogs have shown that transamination reactions promoted by these catalysts can be accelerated by the addition of metal ions, including Zn(II), Cu(II), Ni(II) and Al(III). Hypothetically, these cations accomplish this by stabilizing the formation of Schiff base intermediates and by increasing the acidity of the protons that must be removed in the reaction. Interestingly, the addition of metal ions to reactions catalyzed by ALBP- and IFABP-pyridoxamine conjugates resulted in both positive and negative reaction rate perturbations [49]. IFABP-PX104 reacted 4.7-fold faster in the presence of Cu(II) whereas IFABP-PX60 reacted 4.4-fold slower – both rate effects were accompanied by a decrease in reaction enan-tioselectivity. Little change was observed for the reaction catalyzed by IFABP-PX72. UV/Vis spectroscopy experiments that monitored the formation of metal-aldimine intermediates suggested that IFABP-PX60 and IFABP-PX104 but not IFABP-PX72 formed a complex with Cu(II). Thus, metal ions may be used to increase semisynthetic enzyme efficiency, although this did not occur in all cases. These studies also noted that the reactions rates were sensitive to changes in buffer; the use of imidazole resulted in a lower rate than with similar reactions performed in HEPES buffer. These observations raise the possibility that buffer molecules may actually enter the protein cavity and directly participate in the reaction. This makes sense because several proton transfers must occur in a complete transamination reaction and yet there are no functional groups within the cavity that could do this. 3.6 Chemogenetic Approach

In the catalytic mechanism of pyridoxamine-based reactions, the pyridine nitrogen atom must be protonated; this protonation gives the pyridine ring a net positive charge that increases the acidity of the nearby benzylic protons and serves to stabilize a developing negative charge. One way protonation can be effectively enforced is by introducing a permanent positive charge via N-methylation. In some model systems, an N-methylated species accelerates the transamination rate up to 20-fold. In contrast, after reconstitution with N-methylpyridoxamine, alanine aminotransferase lost its activity almost completely (>99.8%). Therefore, N-quaternization may have both positive and negative influences on the reaction rate. To allow a similar modification to be made in these fatty acid binding protein systems, a new reagent containing N-methylpyridoxamine and an activated disulfide (TP-MPX, 6-3) was prepared. A conjugate, IFABP-MPX60 (6-4) was prepared [50] and, under


catalytic conditions, significant reaction (2 turnovers, 41% ee) was observed using α-keto glutarate and phenylalanine as substrates. Kinetic analysis of the reaction showed that the MPX-containing conjugate gave higher K M (3.8-fold higher than the earlier PX construct) but similar kcat . This indicated that N-methylation had no positive effects on the reaction catalyzed by conjugates based on IFABP-V60 C. However, the MPX cofactor proved to be considerably more useful in subsequent experiments with IFABP variants containing lysine mutations (see below for further discussion of those proteins). In those studies, efforts were made to increase their catalytic efficiency by employing a genetic method to enforce N-protonation. Taking a cue from the crystal structure of AATase (aspartate amino transferase), an enzyme that catalyzes transamination using a pyridoxamine cofactor, carboxylatecontaining amino acid residues were introduced close to the pyridine nitrogen locus in two IFABP mutant proteins. It was thought, based on functional studies of AATase, that the positioning of anionic side chains near the pyridine nitrogen center would enforce protonation via an ion pairing mechanism; the presence of the proximal negative charge would favor the protonated pyridine. Several carboxylatecontaining IFABP mutants were prepared for this purpose. Unfortunately, none of these mutants were sufficiently stable for conjugate preparation – they precipitated during purification, and efforts to refold the denatured protein were unsuccessful. However, in separate experiments, the TP-MPX reagent (6-3) was used to prepare MPX-containing constructs instead (see 6-4 for a generic structure). Interestingly, these conjugates showed enhanced catalytic activity compared to their PX progenitors [51]. Kinetic studies indicated that the kcat (1.12 h−1 for IFABP-MPxK38 and 0.52 h−1 for IFABP-MPxK51) and turnover numbers (12.2 turnovers by IFABPMPxK38 and 5.7 by IFABP-MPx51 in 24 h) observed with these constructs under standard conditions are the highest achieved in this system (see Table 1 for a summary of kinetic parameters). The success of these constructs, prepared using a combination of chemical modification of the catalyst structure and genetic manipulation of the protein scaffold, highlights the enormous power and flexibility of this chemogenetic approach for catalyst development. 3.7 Adding Functional Groups within the Cavity

A major goal of research involving these protein scaffolds was to determine whether the flexibility of using a protein scaffold could be fully capitalized upon by using rational design in concert with knowledge of chemical mechanism to improve catalytic efficiency. Taking a cue from Nature, based on biochemical experiments with AATase, lysine residues were introduced into the protein cavity to enable Schiff base formation and to serve as general acids and bases in the reaction cycle. Figure 10 shows these functions in an abbreviated transamination mechanism. Based on the crystal structure of IFABP, molecular modeling was employed to identify possible positions where lysine residues could be introduced to perform the functions noted above. Mutants L38 K,V60 C and E51 K,V60 C were prepared and used to create pyridoxamine conjugates [52]. Figure 11 gives a model of IFABP-PXK51.


Fig. 10 Abbreviated mechanism for transamination reactions promoted by pyridoxal-based catalysts.

The resulting assemblies IFABP-PxK38 and IFABP-PxK51 showed improved kcat and K M . The overall catalytic efficiency (kcat /K M ) of IFABP-PxK51 increased 4200-fold compared to unliganded pyridoxamine phosphate and was 12-fold greater than for IFABP-PX60 while maintaining comparable enantioselectivity (83–94% ee). The principal effect on the kinetic constants for the reactions catalyzed by these mutants was on K M . Each conjugate showed a significant decrease in

Fig. 11 Stereo representation of a model of IFABP-PXK51. The overall protein structure is shown with C60 and the pyridoxal linked via a disulfide bond. The pyridoxal aldehyde group is bonded through a Schiff base to K51 . Color scheme: Protein secondary structure (green), carbon (white), oxygen (red), nitrogen (blue), sulfur (orange).


K M (2.2- and 7.5-fold) and a small increase in kcat (1.5-fold). UV/Vis spectroscopy, fluorescence and electron-spray mass spectrometry verified the catalytic function (Schiff base formation) of the introduced lysine residue in the reaction process. Significantly, in a study of rate versus pH, IFABP-PX60 gave a rate that decreased monotonically with increasing pH while the lysine mutants exhibited a bell-shaped profile with a maximum rate near pH 7.5. Taken together, these results provide compelling evidence that the lysines participate directly in the reaction and that features of enzymatic processes, including covalent catalysis, can be mimicked successfully. Computer modeling together with mutagenesis was also used to identify existing residues in the protein cavity that contributed to the substrate specificity of the IFABP-PX60 catalyst. R126 and Y14 were identified as two important residues that interacted with the γ -carboxylate group of α-keto glutarate when bound to the enzyme via a Schiff base with the PX moiety (Figure 12). Mutants IFABPV60 C,R126 M and -V60 C,R126 M,Y14 F were prepared and pyridoxamine was attached to each mutant [53]. Of particular note, IFABP-PxM126 had a K M three-fold higher than IFABP-PX, while the second mutation at position 14 had no significant effect. The kcat s for both conjugates were 2-fold lower than for the original IFABP-PX60. N-methylated pyridoxamine conjugates were also introduced into these two mutants. Compared to the original IFABP-MPX60 conjugate described above, the

Fig. 12 Putative binding site for the sidechain carboxylate of the substrate "-keto glutarate in IFABP-PX60. Present are Y14 (upper left), R126 (upper right) and the Schiff base complex between "-keto glutarate and the pyridoxamine catalyst. (lower

center). Dashed lines indicate putative hydrogen bonds between the (-carboxylate of "-keto glutarate and the side-chain phenol of Y14 and the guanidinium group from R126 . Color scheme: Carbon (white), oxygen (red), nitrogen (blue), sulfur (orange).


mutations in positions 126 and 14 increased K M as much as seven-fold. IFABPMPxM126 gave a particularly significant result, manifesting a 5-fold higher kcat . While neither the experiments with the lysine mutants nor those with the carboxylate binding site mutants produced catalysts that rival the efficiency of natural enzymes, incremental improvements were observed with many of the conjugates. Moreover, these advances came from constructs that combined mutations in the protein scaffold with synthetic alterations in cofactor structure, thus highlighting the power of the chemogenetic approach. 3.8 Scaffold Redesign

While site-directed mutagenesis for the incorporation of new functional groups into the protein cavity is the most obvious type of genetic modification that can be performed with these systems, more global changes can be made in the scaffold using similar methods. For example, as noted above, the IFABP architecture consists of a β-barrel structure caped off with a α-helical lid. To examine whether entry or exit of substrates or products into or out of this closed cavity was limiting catalytic turnover, a helixless (hs) IFABP mutant was prepared by deleting 17 helical residues (15–31) from the N-terminus and replacing them with a dipeptide linker (Ser-Gly) using site-directed mutagenesis [54]. A previous NMR study showed that this mutant preserved the original β-sheet secondary structure without the α-helical lid and was still relatively stable [55]. The structures of IFABP and hsIFABP are compared in Figure 13; except for the absence of the α-helical lid, the overall protein fold is largely intact. Studies that investigated reactions with this conjugate (hsIFABP-PX60) revealed that in 24 h, approximately two turnovers were observed with a selectivity of 93% ee. A kinetics analysis indicated that kcat s were comparable for both IFABP-PX60 (0.20 h−1 ) and hsIFABP-PX60 (0.22 h−1 ), while the former exhibited a 4-fold increase in K M [56]. After introducing the MPX cofactor into the cavity of helixless protein, 4.9 turnovers in 24 h with 35% ee was observed for the production of Glu under the same conditions. Taken together, these results suggested that removal of the α-helical lid did not affect the maximal rate and enantioselectivity of the respective constructs. However, deletion of this structural element did decrease the substrate binding affinity, as evidenced by an increase in K M . Thus, it appears that while the hsIFABP scaffold was not useful in increasing the efficiency of PX-promoted transamination, these experiments did show that stable constructs could be prepared based on the helixless scaffold. This may have greater utility in reactions with larger cofactor catalysts or substrates. 3.9 Hydrolytic Reactions

Fatty acid binding proteins have also been used as scaffolds for constructing hydrolytic catalysts. Phenanthroline was attached to Cys117 of ALBP using


Fig. 13 Comparison of the structures of IFABP and the helixless variant hsIFABP. Top: Stereo view of IFABP. Bottom: Stereo view of hsIFABP.

iodoacetamido 1,10-phenanthroline and metallated to produce a coordinated Cu(II) ion encapsulated within a chiral protein cavity (Figure 14). The resulting conjugate, ALBP-Phen-Cu(II) was able to promote the hydrolysis of several unactivated amino acid esters under mild conditions (pH 6.1, 25 ◦ C) at rates 32- to 280-fold above the background rate in buffered aqueous solution; these reactions also showed modest stereoselectivity [57]. In 24 h incubations, 0.70–7.6 turnovers were obtained with enantiomeric excesses ranging from 31 to 86% ee. ALBPPhen-Cu(II) could also catalyze the hydrolysis of an activated amide (picolinic

Fig. 14 Reaction used to prepare ALBP-Phen.


acid methyl nitroanilide, PMNA), under slightly more vigorous conditions (37 ◦ C) and after longer incubation times. The rate of amide hydrolysis was 1.6 × 104 fold higher than the background rate [57]. However, kcat obtained with ALBPPhen-Cu(II) was still significantly lower than that obtained with a related Cu(II) bipyridine complex [58]. This rate decrease may reflect a non-optimal, perhaps nonplanar, conformation for PMNA binding within the ALBP cavity. Of related interest, an X-ray crystal structure of ALBP-Phen was obtained. Inspection of this structure showed that the protein could not accommodate the phenanthroline and PMNA within a planar conformation without significant distortion of the protein backbone [45]. This may account for the lower than expected rate of PMNA hydrolysis promoted by ALBP-Phen-Cu(II) compared with non-proteinaceous model complexes. In analogy to the above experiments with pyridoxamine complexes, the 1,10phenan-throline ligand was introduced at several alternative sites (positions 60, 72 and 104, see Figure 8) using the protein scaffold IFABP [59]. Using alanine isopropyl ester as a substrate, IFABP-Phen60 catalyzed ester hydrolysis with less selectivity than ALBP-Phen, while Phen72 promoted the same reaction with higher selectivity. In contrast, hydrolysis of tyrosine methyl ester was catalyzed with higher selectivity by Phen60 and more rapidly by Phen104. These results indicated that the rate enhancement and substrate selectivity of hydrolysis reactions catalyzed by phenanthroline conjugates depended largely on the orientation and environment of the metal ligand within the protein cavity.

3.10 A Flavin-containing Conjugate

Reactions catalyzed by a flavin analog incorporated into IFABP have also been studied by alkylating a Cys residue within the cavity of the helixless variant (hsIFABP) [60]. The conjugate hsIFABP-FL catalyzed the oxidation of several dihydronicotinamides. These experiments were performed mainly to compare results obtained with flavin-IFABP conjugates with similar results acquired with flavopapains; these latter proteins were among the first semisynthetic enzymes produced. Interestingly, while hsIFABP-FL and flavopapain gave comparable rate accelerations (kcat /K M ) for dihydronicotinamide oxidation, hsIFABP-FL manifested much higher K M and kcat than did flavopapain. The low K M observed with flavopapain indicates that it primarily accelerates the reaction rate by enhancing substrate binding, whereas the higher kcat obtained with hsIFABP-FL suggests that its major mode of rate acceleration involves enhancing flavin reactivity. Molecular modeling of hsIFABP-FL indicates that Gln27 is close to N(3)H and O(4) of the flavin (Figure 15). Interaction between the carboxamide of Gln27 and the flavin is likely to alter the redox potential of the isoalloxazine and hence augment its reactivity. Taken together, these results with flavopapain and hsIFABP-FL highlight the diverse ways in which protein scaffolds can be used to modulate catalyst activity.


Fig. 15 Hydrogen bonding to the flavin in hsIFABP-FL.

3.11 Some Limitations

While the examples described above highlight the impressive results that can be achieved with protein as scaffolds for catalyst design, this approach is not without problems. Firstly, there is the issue of cavity size. It would be useful to incorporate large metal–ligand systems into this protein system to generate catalysts for a plethora of interesting reactions. Unfortunately, the enclosed cavity of fatty acid binding proteins (and even the somewhat open cavity of hsIFABP) limits the size of what can be introduced into the protein interior. Efforts to incorporate a heme group or a Mn(III)-salen complex into hsIFABP were not successful. While it was possible to attach these moieties to the protein scaffold under denaturing conditions and refold them, the resulting conjugates were unstable and underwent aggregation and precipitation upon storage. In future work, it will be necessary to identify other protein structures that can serve as effective scaffolds but that are not subject to these size limitations. Secondly, work with fatty acid binding proteins is limited to aqueous systems. This creates two types of problems. The first is that catalytic systems that are sensitive to water cannot be employed. The Mn(III)salen complex described above underwent Schiff base hydrolysis and subsequent ligand decomposition upon prolonged storage in aqueous buffer. While the rate of this hydrolytic degradation can be decreased by employing mixtures of water and organic solvents, fatty acid binding proteins have limited solubility in such solvents. In addition, the inability to work in such solvents also limits the substrate concentrations that can be used and also precludes the use of these catalysts with many types of hydrophobic substrates. However, given the extensive interest and the large body of literature concerning the use of enzymes in organic media, it should be possible to identify other types of proteins that can serve as scaffolds for catalyst design in such solvent systems.

4 Myoglobin as a Starting Point for Oxidase Design 4.1 Artificial Metalloproteins and Myoglobin

Metalloproteins represent a major faction of scaffold proteins in artificial enzyme design. The introduction of a wide range of metals and redox cofactors into protein scaffolds can greatly increase both the diversity of enzyme catalysis and the

4 Myoglobin as a Starting Point for Oxidase Design

application of these artificial metalloproteins. To sequester artificial redox cofactors into protein scaffolds, which would otherwise not bind these chemical catalysts, one may either use non-covalent or covalent attachment [61]. Myoglobin is a wellstudied protein that contains a non-covalently bound heme for oxygen transport. The large heme-binding site provides substantial room to accommodate other ligand systems, and for these reasons serves as a useful starting point for catalyst design. 4.2 Non-covalent Attachment of a Redox Center

Watanabe and co-workers pursued an approach involving the non-covalent placement of Mn (III) and Cr (III) salophen complexes into apo-myoglobin [61]. In this artificial metalloprotein, two residues required mutation to improve the binding affinity of the cofactor and to allow increased access of substrates. The sulfoxidation activity of this Cr(III)-salophen myoglobin complex was studied and showed a sixfold rate increase over free Cr(III)-salophen in solution. The enantioselectivity of free salophen was 0%, while that of the myoglobin catalyst was 13%. While the ee is low for this artificial metalloprotein, Wantabe and co-workers were successful in demonstrating that asymmetric reactions can be accomplished using chiral protein cavities. 4.3 Dual Anchoring Strategy

To increase the enantioselectivity of these myoglobin metalloenzymes, Lu and coworkers have successfully utilized a covalent linkage approach [62]. In an earlier attempt a Mn(III)-salen complex was incorporated into apo-myoglobin by mutating residue 103 to cysteine, followed by modification with a methane thiosulfonate derivative of Mn(III)(salen) (Figure 16). This catalyst showed sulfoxidation activity; however, the ee was only 12%. As such a low ee might be a result of the ability of the bound ligand to exist in multiple conformations within the protein cavity, it was hypothesized that the rotational freedom of the salen complex could be limited if it was anchored at two separate locations, unlike the previous single-point attachment conjugate. The Mn(III)(salen) derivative was modeled into myoglobin to identify two positions that would yield the best fit into the heme pocket (Figure 17). In

Fig. 16 A Mn(III)(salen) derivatizing reagent used to prepare proteinMn(III)(salen) conjugates.

19


Fig. 17 Structure of myoglobin complexed with a heme group. The two residues chosen for mutation, Y103 and L72 , are shown. Left: Side view of myoglobin. Right: Top view of myoglobin. Color scheme: Carbon (white), oxygen (red), nitrogen (blue), iron (pink).

addition to an increase in the rate of sulfoxidation of the substrate thioanisole, the enantioselectivity increased to 51% ee. These results indicate that the dual anchoring strategy may be applied to the design of other artificial metalloenzymes to obtain better enantioselective control. It may also be possible in future work to modulate substrate specificity as well. 5 Antibodies as Scaffolds for Catalyst Design 5.1 Antibodies as Specificity Elements

The immune system of higher eukaryotes produces antibody molecules that recognize a broad range of small molecules. By immunizing animals with specific antigens, produced by conjugating organic compounds to carrier proteins, antibodies that specifically recognize the original small molecule can be generated. This allows protein scaffolds that have binding sites tailored for a specific organic molecule to be produced. Figure 18 shows the hapten binding site of MOPC 315; an impressive number of specific contacts between the antibody and the nitrophenyl amide containing ligand can be seen. While the immunoglobulin-derived scaffolds are quite large, the ability to create high-affinity substrate binding sites without numerous iterations of design makes this an attractive approach for catalyst design. 5.2 Incorporation of an Imidazole Functional Group into an Antibody for Catalysis

The capability of antibodies to provide tailor-made binding sites for catalyst design has been exploited for the preparation of hydrolytic catalysts. The antibody MOPC 315 binds substituted 2,4-dinitrophenyl-containing compounds with association

5 Antibodies as Scaffolds for Catalyst Design 21

Fig. 18 Stereo view of the structure of the antibody MOPC 315 complexed with an amide-containing hapten. Only the region near the hapten binding site is shown. The hapten and Y34 are shown as larger sticks with the remaining residues in the bind-

ing site as smaller sticks. Residues shown include Y34 , W93 and W98 (from VL ) and W333 , H335 , Y339 , R350 , K359 , Y399 , Y401 and S405 (VH ). Color scheme: Carbon (white), oxygen (red) and nitrogen (blue).

constants ranging from 5 × 104 to 1 × 106 M−1 [63]. Imidazole was incorporated into this antibody via a thiol group introduced by chemically modifying K52H in the active site via a disulfide linkage (Figure 19). Using a series of coumarin esters as substrates, multiple (>10) hydrolytic turnovers were observed with no loss of activity. The catalytic efficiency, kcat /K M , for this reaction was over 103 times higher than for a model reaction employing 4-methylimidazole.

Fig. 19 Introduction of imidazole into the ligand binding site of MOPC 315 through chemical modification and site-directed mutagenesis.


5.3 Comparison of Imidazole-containing Antibodies Produced by Chemical Modification and Site-directed Mutagenesis

Since an imidazole group can be incorporated into a protein through site-directed mutagenesis, such an approach was used to prepare a MOPC 315 mutant containing a histidine residue at the corresponding site. In brief, a hybrid Fv fragment of MOPC 315 was constructed by reconstituting a recombinant variable light chain (VL ) produced in E. coli with a variable heavy chain (VH ) from the antibody [64]. Imidazole was introduced into the combining site by substituting Y34 of VL with His using site-directed mutagenesis (see Y34 in Figure 18). This His mutant Fv catalyzed the hydrolysis of the 7-hydroxycoumarin esters of 5-(2,4-dinitrophenyl)aminopentanoic acid 90 000-fold faster than the reaction in the presence of 4-methyl imidazole at pH 7.8. Using that substrate, Fv(Y34HL ) turned over at least eleven times and retained 44% of its activity. The loss of activity probably resulted from the accumulation of the inhibitory reaction product. The mutant Fv bound ε-2, 4-DNP-L-lysine only six-fold less tightly than wild-type protein, suggesting that the substituted Tyr residue is not involved in the DNP recognition process. Compared to the imidazole-catalytic antibody generated by tethering an imidazole chemically, a sixteen-fold greater rate increase was observed for the reactions catalyzed by His mutant Fv protein under similar conditions. This rate enhancement is likely due to the fewer degrees of the freedom of the imidazole moiety inside the combining site (cf. 19-1 and 19-2). While these results alone indicate that the construct prepared by mutagenesis functioned more efficiently than the one obtained from chemical modification, this particular example does not exemplify the most powerful feature of chemical modification, i.e., the ability to incorporate non-natural functionality or spacers into complex protein structures. Considerable use will probably be made of this strategy in future catalyst design.

6 Conclusions

The examples described in this chapter reveal the breadth of methods now used to create protein-based catalysts. Structural analysis via X-ray and NMR techniques has proved to be critical for providing atomic structures that serve as the starting point for design. Computer modeling has become a powerful method in both the planning of new designs and in the interpretation of experimental results. Recombinant DNA techniques allow chemists to make mutations or more global changes in protein scaffolds. Together, these tools have allowed researchers to assemble new protein-based catalysts that can be used to probe biological systems or catalyze useful chemical transformations. While these catalysts do not in general rival the efficiency of natural enzymes, impressive rate accelerations have been obtained in some cases. New methods including phage display [65] and antibody production promise to provide greater access to ligand-specific scaffold development.

References

Peptide/protein ligation techniques offer the possibility of increasing the scope of functionality that can be incorporated into protein structures [66]. As the understanding of how enzymes promote chemical reactions with high efficiency increases, so too will the ability to design more efficient protein-based catalysts. Given the enormous power obtained from combining chemical and genetic methods it is likely that the field of protein-based artificial enzyme design will continue to grow in the near future.

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1

Force Generation in Molecular Motors George Oster University of California, Berkeley, USA

Hongyun Wang University of California, Santa Cruz, USA

Originally published in: Molecular Motors. Edited by Manfred Schliwa. Copyright ľ 2003 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-52730594-0

1 Introduction

Imagine living in a world where a Richter 9 earthquake raged continuously. In such an environment, engines would be unnecessary. You would not need to even pedal your bicycle: you would simply attach a ratchet to the wheel preventing it from going backwards and shake yourself forwards! At the scale of proteins, Brownian motion is even more furious, and proteins evolved to take advantage of this enormous supply of energy. Feynman showed that the familiar mechanical ratchet could not work in an isothermal environment, lest it violate the Second Law of Thermodynamics [1], and so motor proteins must employ a different strategy to convert random thermal fluctuations into a directed force: they use chemical energy via intermolecular forces to capture ‘favorable’ configurations. The way in which proteins do this is dictated by three factors: their size, the strength and range of intermolecular bonds at physiological conditions and the magnitude of the Brownian fluctuations that constitute their thermal environment. These determine the energy, length and time scales on which protein motors can operate. Roughly speaking, motor proteins trap thermal fluctuations in two ways: by biasing diffusion of small, angstrom-sized, steps (‘small ratchets’), and by rectifying nanometer-sized or larger, diffusional displacements (‘big ratchets’). For reasons that will become clear, biasing a sequence of small Brownian fluctuations is generally called a power stroke, while rectifying a large thermal displacement is called a Brownian ratchet. Said another way, Brownian ratchets move down free energy landscapes in steps much larger than kB T, while power strokes move in steps


2 Force Generation in Molecular Motors

comparable to or smaller than kB T1) . The distinction is imprecise, but useful, since it delineates two extremes in the general mechanism by which proteins use intermo-lecular attractions to convert chemical energy into mechanical work.2) There are only a few motors that can be regarded as being pure power stroke motors or pure ratchets; most protein motors employ a combination of the two strategies. However, these ‘thoroughbreds’ are good illustrations of the principles. In fact, evolution has designed one protein that joins both ratchet and power stroke motors into one remarkable device: F 0 F 1 ATPase, or ATP synthase, the machine that manufactures the fuel that powers many other protein machines. We will use this protein as our running example. Our discussion will be mostly qualitative and heuristic. However, it is important to realize that the explanatory cartoons we use are supported by extensive calculations. Omitting them is akin to leaving out the ‘Materials and Methods’ section in an experimental paper: assertions without authority are simply opinions. The supporting quantitative arguments are, perforce, contained in the citations.

2 A Brief Description of ATP Synthase Structure

In order to discuss the workings of the F0 and F1 motors we give a brief account of their structure, which is summarized in Fig. 1. ATP synthase comprises two rotary motors acting in opposition, each operating on an entirely different physical mechanism. The F0 motor is contained in the membrane-bound portion and employs as its energy source a transmembrane electromotive force. The F1 motor is contained in the soluble portion and is driven by the hydrolysis of ATP. The F1 motor is constructed of a coiled-coil shaft (the γ -subunit) surrounded by a hexamer composed of alternating α and β subunits. Nucleotide binding sites nestle in the cleft between subunits; however, only three of the sites are catalytically active, the other three bind, but do not hydrolyze, ATP. The catalytic sites are contained mostly in the β-subunit, with a few—but crucial—residues contributed by the α subunit. The catalytic sites hydrolyze sequentially and drive the rotation of the γ shaft. Movies showing the motion of the hexamer and the rotation of the γ -shaft can be downloaded from a web site given in [1, 2]. The F0 motor is composed of two portions. Between 10 and 14 c-subunits (depending on the species and/or conditions) are assembled into a cylinder attached to the γ -shaft and ε-subunit. It interfaces with a second transmembrane assembly consisting of the a and b subunits. By convention, the γ -cn -ε assemblage is called the ‘rotor’, and the remainder of the protein (the α 3 β 3 hexamer and the δ-b2 -a complex) comprises the ‘stator’, although each rotates in the opposite direction. 1) The quantity kB T measures the thermal energy of Brownian motion; its value is 1 kB T ∼ 4.1 pN nm = 4.1 × 10−21 J at 298 K (25 ◦ C).

2) We have included a short Appendix with several simple examples that illustrate the difference between a ‘power stroke’ and ‘Brownian ratchet’.

3 The F1 Motor: A Power Stroke 3

Fig. 1 The structure of ATP synthase. The panel on the left shows a composite based on the pdb coordinates of the known subunits [23]; the right-hand panel is the corresponding structure represented schemat-

ically showing the relative rotation of the F0 and F1 motors and the direction of ion flow through the a-cn subunit interface. Subscripts denote the subunit stoichiometries.

3 The F1 Motor: A Power Stroke

The study of protein ATPases has led to a few generalizations that help us understand the mechanism by which these molecular motors convert the energy residing in the nucleotide γ -phosphate bond into a directed mechanical force. Ĺ At physiological conditions, the free energy of hydrolysis of one ATP is ∼ 20–24 kB T; of this, about 8–9 kB T is enthalpic, the balance being entropic. Ĺ Almost all nucleotide binding sites are nestled in the cleft between protein subunits. The nucleotide is grasped by loops emanating from a parallel β-sheet. Ĺ In many motors, the force-generating step is associated with the binding of nucleotide to the catalytic site. We propose that this is true of all ATPase motors. In particular, for the F1 motor this is the only way to reconcile all of the biochemical and mechanical measurements with its high mechanical efficiency. Ĺ After the force has been generated by ATP binding, an ATP is tightly bound in the catalytic site. The role of hydrolysis is to break the tightly bound ATP into two products and weaken the binding so the products can be released and the force-generating cycle can be repeated.

In the F1 motor, the ATP binding site lies asymmetrically in the cleft between the β and α subunits, the majority of the catalytic sites residing in the β subunit. The power stroke is accomplished by a hinge bending motion that swings the top of the β subunit down towards the bottom portion. The bending motion of the β subunit can be measured by the motion of helices B and C that emanate from the β sheet of


Fig. 2 The F1 power strokes are accomplished by the hinge bending motions of $ subunits, which are driven by ATP binding to the catalytic sites. (a) During the hinge bending motion, the top part of $ rotates about 30◦ toward the bottom part. This rotation closes the angle between helices B

and C. (b) The hinge bending motion of each $ subunit pushes on the off-axis section of the ( shaft. The rotation of the y shaft is driven by the coordinated hinge bending motions of all three $ subunits [1, 2, 5].

the catalytic site, as shown in the ribbon diagram in Fig. 2a. The bending of the (3 subunit by about 30◦ rotates the central γ shaft by pushing on its off-axis section, much like turning a crankshaft (Fig. 2b). The energy for the power stroke derives from the nucleotide hydrolysis cycle, which consists of four major steps: Binding

Hydrolysis

1

2 ADP Release

CS + ATP CS • ATP PI Release

3

CS • ADP + PI

CS • ADP • PI

CS + ADP

4

(1)

The nucleotide binding step 1 is the force-generating step and should more properly be expressed as a sequence of binding steps: CS + ATP CS • ATP . . . CS • ATP

(2)

Binding Transition

Here the progression from weak to tight binding is symbolized by the increasing size of the bonding symbol indicating the progressive annealing of 15–20 hydrogen bonds (and some hydrophobic interactions at the sugar end). The mechanism driving the hinge bending motion of the β is illustrated schematically in Fig. 3. Immediately after it diffuses into an open catalytic site, the ATP is bound only weakly. The catalytic site wraps around the ATP by forming more bonds which tightens the catalytic site and pulls down the top part of β toward the bottom part. The bonds between the ATP and the catalytic site are formed more or less sequentially, the formation of each bond corresponding to a small drop in binding

3 The F1 Motor: A Power Stroke 5

Fig. 3 The ATP binding transition from weak to tight proceeds as the catalytic site grasps the ATP in a grip of hydrogen bonds (the ‘binding zipper’). As binding progresses, the catalytic site closes up and pulls the top part of the $ subunit toward the bottom part. In this way, the binding free energy is converted to a power stroke

with nearly a constant force. During the power stroke some of the binding energy is stored in the elastic deformation of the p-sheet, which acts ike a spring. This energy is released during the unbending motion to aid product release and return the subunit to its open state.

free energy, that drives a small fraction of the hinge bending motion [1, 3, 4]. When all the bonds have been formed, the ATP is tightly bound in the catalytic site. The overall process from weak to tight binding is called the ‘binding transition’. During the binding transition, the ATP binding free energy is utilized efficiently to generate a bending motion with a nearly constant torque about the hinge region near the β-sheet. The binding transition has two important features [1, 4, 5]: Ĺ The binding free energy decreases (the binding becomes stronger) nearly monotonically and smoothly during the binding transition. This drives the hinge bending motion of the β subunit and consequently the γ shaft rotation in the hydrolysis cycle. In the reverse synthesis cycle, the unbending of the β subunit is driven by the γ shaft rotation in the opposite direction, powered by the F0 motor. When the top of the β subunit is forced up and away from the bottom portion, the


binding free energy increases (the binding becomes weaker). This progressively releases the nucleotide from the catalytic site [6]. Ĺ By the end of the binding transition, approximately 6–10 kB T of elastic energy is stored in the β-sheet whose loops grasp the nucleotide. Note that the catalytic site should be flexible but not elastic, lest it dissipate too much of the binding free energy by elastic recoil. To summarize, the power stroke is driven by progressive capturing of relatively small Brownian motions that anneal the nucleotide into the catalytic site. Product release is accomplished by using the free energy of hydrolysis to weaken the product binding so that thermal fluctuations can knock them out of the catalytic site. Thus Brownian motion drives both the power and exhaust strokes.

4 The F0 Motor: A Brownian Ratchet

The F0 motor is driven by the ion-motive force, µc across a bacterial or mitochondrial inner membrane. The ion-motive force consists of an ion concentration gradient and an electrical potential difference. In most organisms, the ion is a proton. However, much information has been gleaned from anaerobic bacteria whose F0 is driven by a sodium ion motive force. In either case, the transmembrane chemical potential difference in millivolts is given by: µc + = 2.3

kB T e

log10 [c + ] p − log10 [c + ]c +ψ

[mV ]

(3)

pH

where e is the electronic charge, [c+ ]p the ion (sodium or proton) concentration of the periplasm expressed in molars, [c+ ]c the ion concentration of the cytoplasm and ψ is the electrical potential difference across the membrane. Equation 3 is a thermodynamic relationship that implies that a concentration difference is equivalent to an electrical potential at equilibrium. Since motors operate out of equilibrium, this turns out not to be true in general. However, Eq. 3 points out the need for a mechanism for transforming both an entropic and electrical potential into mechanical work. The basic principle that accomplishes this can be illustrated by the ‘toy’ model shown in Fig. 4. 4.1 A Pure Brownian Ratchet

First, consider a rod with a linear array of negative charges (e. g. a DNA strand, etc.) that can freely diffuse through a hydrophilic pore embedded in a membrane as shown in Fig. 4a. If a potential, , is imposed across the membrane, then the rod will be propelled to the left and can perform work against a load force, FL . This can


Fig. 4 Principle of the F0 motor. (a) A pure power stroke. A rod of negative electrica charges passes through a polar transmembrane pore. An electrical potential, imposed across the membrane drives the line of charges to the left. (b) A pure Brownian ratchet. The charged rod passes through a hydrophobic membrane pore. A high concentration of counterion charges (Na+ or H+ ) on the right can bind to the negative sites and neutralize them so that they can pass through the pore. A second positive ion on the left (not shown) neutralizes the membrane potential. The low concentration of counterions on the left ensures that

the bound charges dissociate but do not quickly rebind, so that the bare site cannot re-enter the pore. The attractive bonds between the rod charge and the aqueous solvent molecules rectify the rod’s diffusion. We call this a Brownian ratchet. (c) The power stroke and ratchet can be combined using an L-shaped ‘stator’. An aqueous input channel (colored white) connects via a polar channel (to the right) to the output reservoir The body of the stator is hydrophobic so that ions cannot leak across the membrane. With this design, the motor has both power stroke and ratchet components.

7


be considered as nearly a pure power stroke. The motion of the rod can be viewed as being driven by a driving potential, φ, tilted to the left in Fig. 4a. The slope of the driving potential gives the motor force driving the rod to the left: FM = −dφ/dx. The rod will stall when the motor force equals the load force: FM = FL . Because the rod is very small, its motion is stochastic (indicated in Fig. 4 by the Gaussian force labeled kB T). In fact, the net drift of the rod driven by the potential is deeply buried in its random motion: at any time the rod is only a tiny bit more likely to move to the left than to the right, and the mean square of the instantaneous velocity is several orders of magnitude larger than the net drift velocity. So the net movement down the potential gradient can only be detected by looking at correlations over many steps. This is illustrated by example 3 in the Appendix. 4.2 A Pure Power Stroke

Next, consider the situation in Fig. 4b where the rod passes through a hydrophobic pore. Moreover, there is a difference in the concentration of a positive counterion (e. g. sodium or protons) between the two sides of the membrane. The counterion can bind to and neutralize the negative charges (binding sites) on the rod. A difference in the concentration of a second ion (e.g. potassium), which cannot bind to the sites on the rod, neutralizes the membrane potential. A bare binding site is negatively charged and cannot move into the hydrophobic pore, for that would entail shedding its hydration shell at a considerable energetic cost.3) Because of the high concentration on the right side, the binding sites will be largely neutralized and so the rod can diffuse freely to the left through the pore. However, once a neutralized site has emerged from the pore to the left, the bound positive charge will quickly dissociate and leave the binding site unoccupied. Because of the low concentration on the left side, the binding site is likely to remain unoccupied, which prevents it from moving back into the pore. Thus diffusion to the right is rectified by the ion concentration difference, and the rod moves stochastically to the left, driven by a pure Brownian ratchet. The driving potential, φ, in this case looks like a staircase. When the concentration on the left is much lower than that on the right, each step of the staircase potential is much larger than kB T, so that reverse steps are unlikely. This is illustrated by example 4 in the Appendix. The load force, FL , opposing the motion has the effect of ‘tilting’ the driving potential so that the rod must diffuse uphill, against the load force. The two driving forces can be combined as shown in Fig. 4c. Here the membrane is horizontal and so the motor must be augmented by a fixed ‘stator’ assembly. The stator body is hydrophobic with two exceptions. First, there is a large aqueous channel that permits the ions from the high concentration reservoir to access the binding sites. Second, there is a polar channel connecting the aqueous channel 3) The energy cost of moving a negative charge into the pore is approximately G ∼ 45 kB T [22].


Fig. 5 Operation of the F0 motor [7] (a) Simplified geometry of the sodium driven F0 motor showing the path of ions through the stator. This is the same arrangement as in Fig. 4c, but with the charge array wrapped around a cylinder that is free to rotate in the plane of the membrane with respect to the stator. In addition, a ‘blocking charge’ has been added to the stator to prevent the leakage of charge from the high concentration reservoir (periplasm) to the low concentration reservoir (cytoplasm). (b) Free energy diagram of one rotor site as it passes through the rotor–stator interface. Step 1 → 2, the rotor diffuses to the left, bringing the empty (negatively charged) site into the attractive field of the positive stator charge. 2 → 3, once the site is captured,

the membrane potential biases the thermal escape of the site to the left (by tilting the potential and lowering the left edge). 3 → 4, the site quickly picks up an ion from the input channel neutralizing the rotor. 4 → 5, an occupied site being nearly electrically neutral can pass through the dielectric barrier. If the occupied site diffuses to the right, the ion quickly dissociates back into the input channel as it approaches the stator charge. 5 → 6, upon exiting the stator the site quickly loses its ion. The empty (charged) site binds solvent and cannot pass backwards into the low dielectric of the stator. The cycle decreases the free energy of the system by an amount equal to the electromotive force.

to the low concentration reservoir. Since the aqueous channel is isopotential with the high concentration reservoir, this arrangement converts the transmembrane potential drop from vertical to horizontal. In this way the membrane potential and the ion concentration difference act in tandem to move the rod to the left, the former driving a power stroke and the latter driving a Brownian ratchet. The actual F0 motor works somewhat differently from the idealized version in Fig. 4c. Fig. 5a shows two more modifications that are required to make the arrangement resemble the sodium driven F0 motor of the bacterium P. modestum [7, 8]. The linear array of charged binding sites is first wrapped around a cylinder. This cylinder consists of 10–14 c-subunits, depending on the organisms and/or conditions. Second, a ‘blocking charge’ (R232) is present on the stator that prevents leakage of ions between the two reservoirs that would dissipate the ion gradient unproductively. The presence of this blocking charge alters the picture of the rotation of the rotor with respect to the stator substantially. Fig. 5b shows the potential experienced by one binding site of the F0 motor. The presence of the blocking charge creates an electrostatic potential well that attracts the rotor binding site as soon as it diffuses into the polar channel. Once trapped in the well, the rotor

9


depends on Brownian fluctuations to escape. However, the membrane potential biases escape by lowering the left side of the electrostatic well so that the rotor charge is much more likely to escape to the left than to the right. Once it escapes into the aqueous input channel, it is quickly neutralized by the positive ions so that it can move freely across the hydrophobic barrier. Note that the membrane potential only biases the Brownian fluctuations to the left, but does not actually drive a power stroke as in Fig. 4a. In summary, the F0 motor qualifies as a Brownian ratchet since it rectifies large thermal fluctuations (or long diffusions). The energy for rectification derives from the ion concentration gradient via the short range interactions between ions and the binding sites (binding and unbinding). It also uses the membrane potential to bias thermal fluctuations that release the rotor site from the attraction of the stator blocking charge: it takes less energy to hop out to the left than to the right. This might be thought of as a partial power stroke, so we see that the classification into power stroke and ratchet is largely a question of definition. An interesting class of ratchet motors is those that use the principle of trapping Brownian fluctuations during their assembly to perform a ‘1-shot’ motor task. Examples include the acrosome of Limulus and Thyone, the spasmoneme of Vorticella [9], and the polymerizaton of actin that propels la-mellipodial protrusion and certain intracellular parasites, such as Listeria and Shigella [10, 11]. An important corollary of the ratchet principle, and one that dramatically distinguishes molecular motors from other machines, is that using thermal fluctuations to go uphill in free energy amounts cools off the immediate environment. If the ratchet shown in Fig. 4b is coupled to work against a conservative load force, then the process is endothermic: heat is absorbed from surrounding fluid to increase the potential energy of the external agent that exerts the load force. By contrast, if the motor shown in Fig. 4a is coupled to work against a viscous load, then the process is exothermic: energy from the electrical potential goes to produce the drift velocity of the rod, which in turn is converted to heat by viscous friction. These microscopic thermal energy transactions lead to some surprising properties of molecular motors. For example, it is possible for the motor to perform more work on a viscous load than the free energy it derives from a reaction cycle! These counterintuitive properties are discussed in more detail in the references [5, 12, 13].

5 Coupling and Coordination of Motors

Most ATPase motors have more than one catalytic site. During the motor operation, each catalytic site hydrolyzes ATP and contributes to force generation. These catalytic sites generally do not operate independently, but act in concert with other catalytic sites. Two heads of a kinesin dimer are coordinated with each other to generate unidirectional motion and to ensure processivity. ATP synthase has three catalytic sites. AAA motors are hexamers of ATPases, sometimes stacks of two hexamers. The chaperonin Groel is a stacked pair of heptameric rings, each with seven

5 Coupling and Coordination of Motors

catalytic sites and the portal protein may even be a dodecameric ring of 12 ATPases. Generally, the catalytic sites must act in concert, either firing sequentially, as in F1 , or as in Groel, simultaneously in each ring, but alternating between the rings. This coordination is necessary for the proper operation of the motors, but how is it accomplished? In all cases, motor ATPase catalytic sites are too far apart to communicate in any other way than via elastic strain through the intervening protein structure. Although the details of strain coordination differ, a clue can be found in the correlation between the ADP release at one catalytic site and the ATP binding at another. In F1 , the strain-induced release of ADP arises from two possible sources. First, the γ shaft is asymmetric, so that at every rotational position it strains each catalytic site differently. Second, the intrinsic asymmetry of the protein structure allows the catalytic site to radiate strain differently to the two neighbor sites. The catalytic sites are located in the cleft between adjacent α and β subunits, with the majority of the binding residues in the β subunit [14, 15] This asymmetry allows ATP binding at one catalytic site to propagate a conformational change to the β portion of next catalytic site in the direction of motor rotation, but propagates a different conformational change to the β portion of the previous catalytic site. Thus, one catalytic site can affect the reactions on two neighboring catalytic sites differently. The conformational change directed to the β portion of the next catalytic site can lower the free energy barrier for ADP dissociation, readying it for the next hydrolysis cycle. Indeed this may be the primary structural feature determining the direction of rotation. Because of the unequal symmetry between the F0 and F1 motors, elastic coupling plays an additional role. The F1 motor has three catalytic sites and rotates in three major 120◦ steps, each with a brief pause at 90◦ . On the other hand, the F0 motor has a rotational symmetry that varies between 10 and 14, depending on the source. Therefore, there is no unique ‘stoichiometry’ between the two rotational motions. This symmetry mismatch is not a problem for the motors since the γ -shaft that couples them is torsionally flexible. In synthesis, this allows the stochastic F0 motor to deliver a smooth torque to F1 , which minimizes dissipation as the nucleotide is unzipped from the catalytic site [1, 16]. Thus elastic coupling between subunits of a protein motor provides the means for both coordinating the catalytic cycles and buffering the independent Brownian motions of the subunits. In walking motors the determination of directionality depends on asymmetric structural features of the heads that alternate depending on whether the head is leading or trailing. However, the situation is more complicated since each head has two binding partners: nucleotide and track. One proposal is that strain is generated by binding of the forward head to the track, triggers release of product from that head [17]. Also strain is relayed to the rear head via the connecting structure to lower the energy barrier for a particular step in the reaction cycle (for example, hydrolysis, or product release). This particular reaction step, in turn, triggers the release of rear head from the track [18]. Thus, binding to and release from the track are correlated with the relative positions of the heads. The alternating phases of the two hydrolysis cycles generate unidirectional motion [19].

11


6 Measures of Efficiency

We have discussed the molecular principles by which protein motors convert chemical bond energy into mechanical work. However, while the general principles may be the same for all motors, the detailed mechanisms are quite diverse. Therefore, it is frequently useful to determine the efficiency of a motor to provide clues as to its mechanism. The most common mechanical measurement performed on protein motors is to vary the load (the force resisting the motion) and measure the speed. In general, two kinds of load experiments are carried out on protein motors in order to determine their load–velocity behavior. In one type, a load is applied to resist the motor’s progress using a laser trap or the elastic stylus of an atomic force microscope. In this case the motor is working against a conservative force (i.e. derivable from a potential energy function) that depends only on the displacement of the motor, f = −∂φ/∂ξ ). A second, and generally more experimentally convenient method, is to vary the viscous resistance of the fluid environment of the motor. This is a dissipative force that depends on the motor velocity. The information gleaned from the two kinds of measurements yield different information [1, 12, 13]. The thermodynamic efficiency, ηTD is generally defined as the ratio of the work done by the motor to the energy input: ηT D =

f ·δ −G

(4)

where G is the free energy drop in one reaction cycle (e.g. from hydrolyzing one ATP, or passing one proton through the motor), and f ·δ. is the reversible work done by the motor against the conservative load force, f , in one reaction cycle. Here δ is the average distance covered per reaction cycle, sometimes called the ‘step size’. f ·δ is the energy output from the motor because it goes to increasing the potential energy of the external agent that exerts the conservative force. Thus, the thermodynamic efficiency is the ratio of energy output to energy input and it measures the energy conversion efficiency when the motor is operating reversibly i.e. ‘infinitely slowly’. For a motor working against a constant force (e.g. a laser trap force clamp), Eq. (4) can be generalized to the steady state [1, 13] by defining an efficiency: η≡

f ·v −G·r

(5)

Here r is the average reaction rate (e.g. hydrolysis cycles per second or average proton flux) and (v) is the average velocity. Strictly speaking, Eq. (5) is not a thermodynamic quantity since the steady state need not be the equilibrium state. Nevertheless, it is a well-defined quantity that is less than unity.

6 Measures of Efficiency

In general, the average step size, δ depends on the load force, f . When δ is independent of f we say the motor is tightly coupled. In that case, each reaction cycle is, on average, coupled to a fixed displacement (one motor step) regardless of the load force. A tightly coupled motor has two properties: Ĺ When the motor is stalled the chemical reaction is also stopped. Ĺ Increasing the load beyond the stall force drives the motor in the opposite direction and also reverses the chemical reaction.

For a tightly coupled motor, one can show that the stall force is given by f stall = −G/δ. At stall, the thermodynamic efficiency is 100%. When the motor is operating close to stall, the thermodynamic efficiency approaches 100%, regardless of the motor mechanism. Conversely, a high thermodynamic efficiency suggests only that the motor motion and the chemical reaction are tightly coupled [12]. Equation (5) applies only to the situation where the motor is working against a constant load. For macroscopic motors, inertia is important and the effect of Brownian fluctuations is negligible. Therefore, they tend to move at roughly a constant velocity (at least on short time scales), and so the friction force on the motor is approximately constant. In this case, the friction force can be treated effectively as a conservative force and the efficiency given by Eq. (5) is well defined. Thus, for a macroscopic motor, we generally do not have to worry whether it is working against a conservative force or a friction force. That is, for a macroscopic motor moving with roughly a constant velocity, both a viscous drag and a conservative load simply oppose the motor motion and they have approximately the same effect on the motor. For a protein motor, the situation is very different. Because protein motors are very small, the effect of inertia is negligible and the motor is driven mostly by the random Brownian force. The instantaneous velocity changes direction very rapidly and its absolute value is several orders of magnitude larger than the average velocity. Consequently, the drag force on the motor is stochastic and cannot be treated as a conservative force. The effect of a conservative load force on the motor is different from that of a viscous drag force that simply opposes the motor motion in any direction and whose magnitude increases with the velocity.4) Therefore, for protein motors, it is necessary to distinguish the case where the motor is loaded with a conservative force and the case where the motor is loaded with a viscous drag. When a protein motor works against a viscous load, the thermodynamic efficiency defined above does not apply. A commonly employed measure of efficiency in this case is the Stokes efficiency, defined by replacing f in Eq. (5) with the average viscous 4) A conservative load force tends to drive the motor backwards (opposite to the positive motor direction) and its magnitude is independent of the velocity. A viscous drag simply damps the motor velocity, while the Brownian force and/or the chemical reaction excite the veloc-

ity. If the chemical reaction is halted, the viscous drag will relax the motor to thermal equilibrium with the surrounding fluid, while the conservative force will drive the motor in the opposite direction.

13


drag force, f D = ζ v. Here ζ = kB T/D is the drag coefficient and D is the diffusion coefficient, which can be computed or measured independently [12]: η Stokes =

ζ vδ , −G

or

η Stokes =

ζ v2 −G·r

(6)

Although ζ v2 has the dimension of energy per unit time, ζ v2 is not the rate of the work done by the motor motion on the fluid medium5) . Indeed, it is not clear what energy per unit time ζ v2 measures. However, since ζ v2 increases with v, the quantity ζ v2 does measure some aspect of the motor’s mechanical performance. So the Stokes efficiency is the ratio of this ‘mechanical performance’ index to the energy supply. Of course, for Eq. (6) to be a valid measure of efficiency, it has to satisfy ηStokes ≤ 100 %. This is true, but the proof is not trivial. One can show that Eq. (6) measures how close the motor comes to delivering a constant force [1, 12, 13]. Given the free energy supply — G, the maximum average velocity is achieved if – G is utilized to generate a roughly constant driving force (independent of motor position). When the driving force is constant, the Stokes efficiency is 100%. When the Stokes efficiency is close to 100%, the driving force is close to a constant force [13]. A high Stokes efficiency for the F1 motor was one of the factors that implicated the progressive binding of ATP as the force-generating process, for only in this fashion could a constant torque be generated [1]. To summarize, if the thermodynamic efficiency, ηTD , is close to 1, this means that the motor motion is tightly coupled to the chemical reaction. However, when the Stokes efficiency, ηStokes , is close to 1, this means that the motor is delivering a nearly constant force. Since the Stokes and thermodynamic efficiencies measure different properties of the motor, it is worthwhile to measure both efficiencies experimentally.

7 Discussion

The basic principle underlying force generation in molecular motors seems at first glance pretty simple: proteins use short range attractive intermolecular forces to bias small thermal fluctuations or to rectify larger diffusions. The former we call ‘power strokes’, the latter ‘Brownian ratchets’. But the details are all important, and they are devilishly diverse. At some level of abstraction the operation of a molecular motor can be viewed as the motion of a point moving down a multidimensional free energy surface [1, 20, 21]. Indeed, many models start from this viewpoint and try and deduce what the laws of physics can say about the general properties of 5) ζ v2 should not be confused with ζ v2 . By the equipartition theorem of statistical meachanics, ζ v2 equals ζ kB T m−1 at equilibrium,

where m is the motor mass. ζ v2 is several orders of magnitude larger than ζ v2 .

A1 Example Models to Illustrate the Difference between Ratchets and Power Strokes 15

the surface. While useful from the viewpoint of theory, models at this level of abstraction are likely to be unsatisfying to biologists who seek a more mechanistic understanding, akin to how an engineer would understand the design and operation of an automobile motor. This involves dealing with the details of protein structure. The study of molecular motors, more than most other areas in biology, draws on a diversity of fields. Biochemistry elucidates the kinetics of the energy supplying reactions and thermodynamics establishes constraints on the energetic transactions. Mutation studies isolate the key functional amino acids and mechanical measurements provide criteria that circumscribe possible mechanisms. Finally, microscopy and X-ray crystallography provides the sine qua non structures, for it is nearly impossible to deduce how a machine works without knowing what it looks like. However, all of these studies combined cannot produce a mechanistic theory of how motor proteins work; this requires the unifying power of mathematical modeling. For until the operation of a motor can be formulated as equations and solved, our knowledge is, at best, qualitative and uncertain, hardly better than a plausible cartoon that may, or may not, obey the laws of chemistry and physics and in any event cannot be compared quantitatively with experiments.

Appendix

A1 Example Models to Illustrate the Difference between Ratchets and Power Strokes

Here we discuss four model examples to illustrate the role of Brownian fluctuations and the classification of power strokes and Brownian ratchets. For simplicity, we consider the one-dimensional motion of an object. A1.1 Example 1: A power stroke without Brownian fluctuations

Suppose the object is deterministically moved forward a fixed unit distance, x, per unit time, t (e.g. 0.01 nm in 1 µs). The trajectory of the object is shown in Figure A1a. The object moves forward uniformly with a velocity of (x)/(t). This model example is a pure power stroke without Brownian fluctuations. It is mostly relevant for macroscopic motors. Because of the large inertia, macroscopic motors tend to move at a roughly constant velocity and the effect of Brownian fluctuations is negligible. For protein motors, because of the small size, at room temperature the motion is dominated by Brownian fluctuations. One may argue that this is a hypothetical example of a power stroke protein motor at zero temperature.


Fig. A1 Trajectory of the object for model 1. The object moves forward uniformly with an average velocity of v = x/t = x/1. (b) Trajectory of the object for model 2. The object is driven by a constant force and is subject to Brownian fluctuations. The average velocity of the object is v = x/t. (c) Trajectory of the object for model 3. The object is moved by Brownian fluctuations, and the net drift comes from biasing fluctuations. The average velocity of the object is v = 0.92·x/t. The free energy consumption per unit length is the same as

that in Example 2. Examples 2 and 3 have similar statistical behaviors, and so it is experimentally difficult to distinguish between them. (d) Trajectory of the object for model 4. The object is moved by Brownian fluctuations, and the net drift comes from rectifying large fluctuations. The free energy consumption per unit length is the same as that in Examples 2 and 3 but the average velocity of the object is v = 0.1 x/t, significantly lower than that of Examples 2 and 3.

Since protein motors can only function in a certain temperature range and in a certain fluid medium, this model is not really relevant for protein motors, and so we must modify it to incorporate Brownian fluctuations. A1.2 Example 2: A power stroke with Brownian fluctuations

Suppose, in addition to the deterministic unit forward displacement per unit time, in each unit time the object makes n = 10 independent random moves. Each random move is one unit displacement either forward or backward with equal probability, p (e. g. flipping a fair coin: p = 0.5). This random motion is added to simulate

A1 Example Models to Illustrate the Difference between Ratchets and Power Strokes 17

Brownian fluctuations. The trajectory of the object is shown in Fig. A1b. The average velocity of the object is v = x/t, but the motion is stochastic. If we view the displacement in each unit time as the ‘instantaneous velocity’ for that unit time, then the instantaneous velocity is much larger than the average velocity. The inset shows the details of the trajectory; it is evident that the deterministic drift is buried in the random motions and can be detected only by looking at long time correlations. The number of random moves per unit time is proportional to the diffusion coefficient, D, of the object: 2Dt = n(x)2 . This allows us to calculate the diffusion coefficient from which the drag coefficient, ζ , is computed from the Einstein relation: ζ = kB T/D. In this example, the constant force driving the deterministic forward motion is f = ζ v and the free energy consumption per unit length is δvx. For n = 10, the free energy consumption per unit length is 0.2 kB T. A model such as this is appropriate for describing a charged object, such as a DNA strand, driven through a fluid medium by an electrical potential gradient. A1.3 Example 3: A Brownian ratchet that biases fluctuations

In the two examples above, the net drift is caused directly by a driving force (e. g. an electric potential gradient), and the Brownian fluctuations do not affect the net drift. Next, we consider situations where the net drift is actually caused by biasing or rectifying Brownian fluctuations. In the absence of other influences, the forward and backward fluctuations have the same probability. However, if internal barriers are established to block, partially or completely, the backward fluctuations, then the object is more likely to fluctuate forward. Suppose in each unit time, t, the object makes 10 independent random moves (fluctuations). Each random move is 1 unit displacement, x, either forward or backward with equal probability (p = 0.5). Suppose that each time the object passes a multiple of 5 × x, a barrier is established at that location. At a barrier, the object can fluctuate forward in two ways: a forward Brownian fluctuation and a backward Brownian fluctuation that is reflected by the barrier. The object can move back past the barrier only if a backward Brownian movement ‘breaks’ the barrier. The probability of breaking a barrier depends on the strength of the barrier. Let pf be the probability of forward fluctuation at the barrier and pb be that of backward fluctuation. G/kB T = log(pf /pb ) gives the free energy (in units of kB T) required to break the barrier. If we use pf = 0.73 and pb = 0.27, the corresponding free energy drop at the barrier is 1 kB T Since the barriers are separated by 5 × x, the free energy consumption per unit length is 1/5 = 0.2 kB T, the same as that used in Example 2. The trajectory of the object is shown in Fig. 1Ac. The average velocity of the object is v = 0.92 × x/t, similar to Example 2. Because of the small free energy drop (1 kB T) associated with each barrier, it only partially blocks backward fluctuations. The inset shows the details of the trajectory. This model is an example of a Brownian ratchet that biases small fluctuations. The motion of the object is stochastic and indistinguishable


from that of Example 2 where the object is driven by a constant force and subject to Brownian fluctuations. In this example, the object is directly moved by Brownian fluctuations. The net drift results from biasing fluctuations. For this reason, this model can be classified as a Brownian ratchet. However, it has the same phenomenological behavior as the power stroke motor in Example 2, and so it is very difficult — and unnecessary — to experimentally distinguish it from a power stroke motor. A1.4 Example 4: A Brownian ratchet that rectifies fluctuations

Finally, consider the model Example 3, but now suppose that in each unit time, t, the object makes 10 independent random moves (fluctuations). Each random move is one unit displacement, x, either forward or backward with equal probability (p = 0.5). Now suppose that each time the object passes a multiple of 100 × x, a barrier of 20 kB T is established at that location. The barrier is very high: the probability of a forward fluctuation is pf = 1–3.8 × 10−11 ≈1 and the probability of a backward fluctuation that surmounts the barrier is pb = 3.8 × 10−11 ≈ 0. Since the barriers are 100 × x apart, the free energy consumption per unit length is 20/100 = 0.2 kB T, the same as that in Examples 2 and 3. The trajectory of the object is shown in Fig. A1d. The average velocity of the object is v = 0.1x/t, significantly lower than that of Examples 2 and 3. Because of the large free energy drop associated with each barrier, it almost completely blocks backward fluctuations. Thus, this model is a Brownian ratchet that rectifies large fluctuations. The stochastic motion of the object is different from that of Examples 2 and 3. The object advances slowly in large ratchet steps: once it passes a multiple of 100x, it almost never goes back. One can add a ‘load’ to the above examples by decreasing the probability of a forward fluctuation and increasing that of a backward fluctuation at every location. One can also combine the power stroke and ratchet by adding both a deterministic motion and larger barriers, or alternatively by adding both small barriers and larger barriers. But this would complicate the model and obscure the simplicity of the distinctions we are trying to illustrate. However, we should point out that there is a way to use the time series data itself to estimate how much of the motor driving force can be ascribed to ratchet and power stroke contributions. This involves constructing an Effective Driving Potential from the time series data of the motor; this is discussed in Wang and Oster [12].

A2 A Closer Look at Binding Free Energy

The simple description given in the text of ATP binding to the catalytic site of F1 or of ions binding to the rotor of F0 , conceals a great deal of complexity because it neglects the role of solvent effects. Charges in aqueous solution are always hydrated, surrounded by a shifting cohort of hydrogen bonded waters. Before ATP can bind

A2 A Closer Look at Binding Free Energy

Fig. A2 Enthalpic and entropic changes during desolvation and binding can be followed by plotting—H, vs. TS. In these coordinates, the free energy change, G, is plotted as linear contour lines decreasing up and to the right. The formation of one hydrogen bond between the enzyme and nucleotide can be represented schematically as a reversible path showing a single desolvation and binding process. In state 1, the enzyme and nucleotide sites are hydrated (solid circles). Removing a water molecule from one site entails an enthalpic increase, H12 . This is followed by an entropic increase, TS23 , as the water escapes into solution. Finally, the removed water hydrogen bonds with other waters resulting in an enthalpic decrease, H34 . Similar changes accompany the release of a water molecule

from the other site during the transition from state 4 to state 7. Now the two empty sites must be brought close together, entailing an entropic decrease, TS78 , and an enthalpic decrease, H89 as the sites bind. Thus the overall free energy change, G19 , has enthalpic and entropic components (black bars) that depend on many factors, especially whether the water binds more strongly to the sites than to other waters. Note that the sequence 1 → 9 is only meant to show the enthalpic and entropic changes. It does not represent the actual sequence of what occurs during the desolvation and binding process. In particular, 2 → 3 (the removed water diffusing into solution) and 3 → 4 (the removed water bonding with other waters) occur simultaneously and cannot be separated.

to the catalytic site to initiate the hydrolysis cycle (or sodium binding to the rotor charge in the F0 motor cycle), both must shed their water coats. The shedding of the water coats is progressive as the hydrogen bonds form between ATP and the catalytic site. Before each hydrogen bond can form, the hydration water molecules must be shed from the donor and receptor just before they form the bond (otherwise they will be re-hydrated quickly). Consider the overall process of the formation of one hydrogen bond between ATP and the catalytic site. This entails a number of energetic and entropic transactions. We can plot the process schematically as the path shown in Fig. A2. (This path does not represent the actual non-equilibrium process, but rather a ‘reversible work’ path to illustrate the separate entropic and enthalpic transactions). We see that, even in the simplest case, a single association event entails four enthalpic and four entropic changes when water molecules break their hydrogen bonds to charged sites, escape into solution, re-bond with other

19


waters, and finally two sites associate. Clearly, solvent effects can tip the free energy balance, but it is seldom easy to compute how. The binding transition that generates the power stroke as embodied in Eq. (2) and Fig. 3 is also an oversimplified description. The hydrogen bonds are not arrayed in a linear sequence (as suggested by the term ‘binding zipper’), nor are they either ‘on’ (zipped) or ‘off’ (unzipped). The bonding surfaces are complex, hydrogen bonds have a finite range and angular dependence, and thermal motions create a stochastic pattern of graded bonding interactions. Nevertheless, molecular dynamics studies demonstrate that the free energy changes gradually and nearly linearly as the nucleotide unbinds or binds to the catalytic site.

References 1 OSTER, G. and H. WANG. 2000a. Reverse engineering a protein: The mechanochemistry of ATP synthase. Biochim. Biophys. Acta 1458: 482–510. 2 WANG, H. and G. OSTER. 1998. Energy transduction in the F1 motor of ATP synthase. Nature 396: 279–282. 3 BOCKMANN, R. 2002. Nanoseconds molecular dynamics simulation of primary mechanical energy transfer steps in F1 -ATP synthase. Nat. Struct. Biol. 9: 198–202. 4 SUN, S., et al. 2003. Elastic energy storage in beta sheets with application to F1 -ATPase. Eur. Biophys. J. 32: 676–683. 5 OSTER, G. and H. WANG. 2000b. Why is the efficiency of the F1 ATPase so high? J. Bioenerg. Biomembr. 332: 459–469. 6 ANTES, I., et al. 2003. The unbinding of ATP from F1 -atpase. Biophys. J.. 85: 695–706. 7 DIMROTH, P., et al. 1999. Energy transduction in the sodium F-ATPase of Propionigenium modestum. Proc. Natl. Acad. Sci. USA 96: 4924–4929. 8 OSTER, G., et al. 2000. How F0 -ATPase generates rotary torque. Proc. Roy. Soc. 355: 523–528. 9 MAHADEVAN, L. and P. MATSUDAIRA. 2000. Motility powered by supramolecular springs and ratchets. Science 288: 95–99. 10 MOGILNER, A. and G. OSTER. 1996a. Cell motility driven by actin polymerization. Biophys.J. 71: 3030–3045. 11 MOGILNER, A. and G. OSTER. 1996b. The physics of lamellipodial protrusion. Euro. Biophs. J. 25: 47–53.

12 WANG, H. and G. OSTER. 2002a. Ratchets, power strokes, and molecular motors. Appl. Phys. A 75: 315–323. 13 WANG, H. and G. OSTER. 2002b. The Stokes efficiency for molecular motors and its applications. Europhys. Lett. 57: 134–140. 14 MENZ, R., et al. 2001. Structure of bovine mitochondrial F1 -ATPase with nucleotide bound to all three catalytic sites: implications for the mechanism of rotary catalysis. Cell 106: 331–341. 15 STOCK, D., et al. 2000. The rotary mechanism of ATP synthase. Curr. Opin. Struct. Biol. 10: 672–679. 16 JUNGE, W., et al. 2001. Inter-subunit rotation and elastic power transmission in F0 F1 -ATPase. FEBS Lett. 251: 1–9. 17 UEMURA, S., et al. 2002. Kinesin–microtubule binding depends on both nucleotide state and loading direction. Proc. Natl Acad. Sci. USA 99: 5977–5981. 18 HANCOCK, W. and J. HOWARD. 1999. Kinesin’s processivity results from mechanical and chemical coordination between the ATP hydrolysis cycles of the two motor domains. Proc. Natl Acad. Sci. USA 96: 13147–13152. 19 PESKIN, C. S. and G. OSTER. 1995. Coordinated hydrolysis explains the mechanical behavior ofkinesin. Biophys.J. 68: 202s–210s. 20 BUSTAMANTE, C., et al. 2001. The physics of molecular motors. Acc. Chem. Res. 34: 412–420.

References 21 WANG, H., et al. 1998. Force generation in RNA polymerase. Biophys.J. 74: 1186–1202. 22 ISRAELACHVILI, J. and B. NINHAM. 1977. Intermolecular forces the long and short of it. J. Colloid Interface Sci. 58: 14–25.

23 PEDERSEN, P., et al. 2000. ATP Synthases in the year 2000: evolving views about the structures of these remarkable enzyme complexes. J. Bioenerget. Biomembr. 32: 325–332.

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c02 (JWBK148-Bommarius)

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Enzyme Catalysis Andreas Sebastian Bommarius Georgia Institute of Technology, Atlanta, USA

Bettina R. Riebel Emory University School of Medicine, Atlanta, USA

Orginally published in: Biocatalysis. Andreas Sebastian Bommarius and Bettina R. Riebel. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30344-1

1 Introduction

Enzymes are a class of macromolecules with the ability both to bind small molecules and to effect reaction. Stabilizing forces such as hydrophobic effects only slightly dominate destabilizing forces such as Coulombic forces of equal polarity; thus the Gibbs free enthalpy of formation of proteins, Gformation , is only weakly negative. In an enzyme reaction, initially free enzyme E and free substrate S in their respective ground states initially combine reversibly to an enzyme–substrate (ES) complex. The ES complex passes through a transition state, Gtr,cat , on its way to the enzyme–product (EP) complex and then on to the ground state of free enzyme E and free product P. From the formulation of the reaction sequence, a rate law, properly containing only observables in terms of concentrations, can be derived. In enzyme catalysis, the first rate law was written in 1913 by Michaelis and Menten; therefore, the corresponding kinetics is named the Michaelis–Menten mechanism. The rate law according to Michaelis–Menten features saturation kinetics with respect to substrate (zero order at high, first order at low substrate concentration) and is first order with respect to enzyme. Important milestones in the rationalization of enzyme catalysis were the lockand-key concept [1], Pauling’s postulate (1944) and induced fit [2]. Pauling’s postulate claims that enzymes derive their catalytic power from transition-state stabilization; the postulate can be derived from transition state theory and the idea of a thermodynamic cycle. The Kurz equation, kcat /kuncat ≈ KS /KT , is regarded as the mathematical form of Pauling’s postulate and states that transition states in the case of successful catalysis must bind much more tightly to the enzyme than Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754


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2 Enzyme Catalysis

ground states. Consequences of the Kurz equation include the concepts of effective concentration for intramolecular reactions, cooperativity of numerous interactions between enzyme side chains and substrate molecules, and diffusional control as the upper bound for an enzymatic rate. Enzymes demonstrate rate accelerations (kcat /kuncat ratios) of ≤ 1.4 · 1017 and proficiencies [kcat /(kuncat K M ) (= 1/K T )], of ≤ 10 M−1 . Surprisingly, for a range of enzymatic reactions kcat is within two orders of magnitude whereas kuncat varies by more than six orders of magnitude; the highest rate accelerations are observed for reactions with low kuncat . Every (bio)catalyst can be characterized by the three basic dimensions of merit – activity, selectivity and stability – as characterized by turnover frequency (tof) (= 1/kcat ), enantiomeric ratio (E value) or purity (e.e.), and melting point (T m ) or deactivation rate constant (kd ). The dimensions of merit important for determining, evaluating, or optimizing a process are (i) product yield, (ii) (bio)catalyst productivity, (iii) (bio)catalyst stability, and (iv) reactor productivity. The pertinent quantities are turnover number (TON) (= [S]/[E]) for (ii), total turnover number (TTN) (= mole product/mole catalyst) for (iii) and space–time yield [kg (L · d)−1 ] for iv). Threshold values for good biocatalyst performance are kcat > 1 s−1 , E > 100 or e.e. > 99%, TTN > 104 –105 , and s.t.y. > 0.1 kg (L · d)−1 .

2 Characterization of Enzyme Catalysis 2.1 Basis of the Activity of Enzymes: What is Enzyme Catalysis?

Enzymes are a class of multifunctional, multivalent macromolecules with the ability to bind small molecules and, much more importantly, subsequently to effect reaction. Stabilizing forces such as hydrophobic effects [3] only slightly dominate destabilizing forces such as Coulombic forces of equal polarity; thus the Gibbs free enthalpy of formation, Gformation , is weakly negative. Without exception, enzymes are proteins and consist of one or more linear chains of, with rare exceptions, the common 20 proteinogenic amino acids. The fairly frequent disulfide bonds between cysteines are formed through crosslinking of side-chain interactions but do not constitute branching of the backbone chain. Extremely rarely, an additional genetically coded 21st amino acid is found, such as selenocysteine in formate dehydrogenase or glutathione peroxidase [4, 5], or pyrrolysine, the “22nd” amino acid, in methanogens belonging to the domain of archaea [6, 7], or p-fluorophenylalanine in a redesigned E. coli cell [8]. All three examples used a stop codon to code for the unusual amino acid. Non-proteinogenic amino acids are found much more commonly in non-enzyme proteins. One prominent example is collagen, from gelatin, which mainly consists of triplets of hydroxyproline–glycine–proline, in which the hydroxyproline is generated via posttranslational hydroxylation. Other proteins apart from enzymes


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2 Characterization of Enzyme Catalysis 3

exist that can bind substrates and catalyze reactions, such as catalytic antibodies [9, 10] or ribozymes [11, 12]. On the other hand, there are many proteins that do not catalyze any reaction, and thus are not enzymes, but function as oxygen storage and shuttle units (hemoglobin and myoglobin), receptors (signal transduction proteins such as the EGF-receptor) or as structural proteins (myelin). 2.2 Enzyme Reaction in a Reaction Coordinate Diagram

The idea of an enzyme reaction consisting of both a binding step and a subsequent reaction step is embodied in a free enthalpy-reaction coordinate (G-ξ ) diagram depicting the change in Gibbs free enthalpy G over the extent of reaction ξ (zeta) (Figure 1). In an uncatalyzed reaction, one or more substrates or reactants initially are in their respective ground states; during the reaction, the reactants pass the point of maximum free enthalpy Gtr,uncat , termed the “transition state”, and continue to the ground state of the product(s). In an enzyme reaction, initially free enzyme E and free substrate S likewise are in their respective ground states. From the ground state, enzyme and substrate combine reversibly to an enzyme–substrate (ES) complex. If [S] > K M , the ES complex forms an “intermediate”, i.e., a local minimum of free enthalpy. (If [S] < K M , the free enthalpy of the ES complex is higher than the enzyme’s ground state; at [S] = K M , both free enthalpy levels are the same.) The ES complex passes through another transition state, Gtr,cat , on its way to the enzyme–product (EP) complex and then on to the ground state of free enzyme E and free product P. The maximum of the Gibbs free enthalpy between the ground states of substrate and product forms the Gibbs free enthalpy of activation with the energy difference G= , which determines the rate constant of the reaction. Like every catalyst, an enzyme decreases the value of G= and thus accelerates the reaction. (An agent increasing the value of G= is termed an “anti-catalyst”.)

Fig. 1 Free enthalpy reaction coordinate diagram for an enzyme reaction.


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4 Enzyme Catalysis

2.3 Development of Enzyme Kinetics from Binding and Catalysis

From the idea of enzyme kinetics as a binding and a reaction step with the corresponding course of the energy curve in the Gibbs free enthalpy–reaction coordinate (G – ξ ) diagram, the reaction scheme represented by Eq. (1) can be drawn. E+S⇔ES→EP→(⇔)E+P

(1)

Among the several ways of verifying or disproving such a reaction scheme, the derivation of a rate law linking a product formation rate or substrate consumption rate with pertinent concentrations of reactants, products, and auxiliary agents such as catalysts probably has the greatest utility, as conversion to product can be predicted. A proper rate law contains only observables, and no intermediates or other unobservable parameters. In enzyme catalysis, the first rate law was written in [13] by Michaelis and Menten (the corresponding kinetics is therefore aptly named the Michaelis–Menten (MM) mechanism). The kinetic schemes for the description of enzyme reactions are similar to those for homogeneous and heterogeneous catalysis and were developed roughly contemporaneously. In heterogeneous catalysis, a mechanism postulating a reaction of a mobile substrate molecule on a solid surface is modeled by a rate law named after Langmuir and Hinshelwood, and later also after Hougen and Watson [14] (LHHW). LHHW and MM both assume a limiting number of active sites which can be saturated with reacting molecules from a continuous phase. The kinetic scheme according to Michaelis–Menten for a one-substrate reaction [13] assumes three possible elementary reaction steps: (i) formation of an enzyme–substrate complex (ES complex), (ii) dissociation of the ES complex into E and S, and (iii) irreversible reaction to product P. In this scheme, the product formation step from ES to E + P is assumed to be rate-limiting, so the ES complex is modeled to react directly to the free enzyme and the product molecule, which is assumed to dissociate from the enzyme without the formation of an enzyme–product (EP) complex [Eq. (2)]. E+S⇔ES→E+P

(2)

In the derivation according to Michaelis and Menten, association and dissociation between free enzyme E, free substrate S, and the enzyme–substrate complex ES are assumed to be at equilibrium, K S = [ES]/([E] · [S]). [The Briggs–Haldane derivation (1925), based on the assumption of a steady state, is more general.] With this assumption and a mass balance over all enzyme components ([E]total = [E]free + [ES]), the rate law in Eq. (3) can be derived. v=vmax [S]/(K M +[S])=kcat [E]total [S]/(K M +[S])

(3)


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3 Sources and Reasons for the Activity of Enzymes as Catalysts

In contrast to [E]free , [E]total is observable. Eq. (3) is written with K M , the Michaelis constant, instead of the equilibrium binding constant K S ; unless the enzyme reaction is very fast (Section 4.3.); i.e., in almost all cases, K M ≈ K S . In Eq. (3), the reaction rate is traditionally denoted by v [concentration/time] and kcat is the reaction rate constant [time−1 ]. The equation describes a two-parameter kinetics, with a monotonically rising reaction rate with respect to substrate concentration and saturation at high substrate concentration. The maximum reaction rate at saturation is denoted by vmax , with vmax = kcat [E]. The K M value corresponds to the substrate concentration at half saturation (vmax /2) and is a measure of binding affinity of the substrate to the enzyme: a high K M value corresponds to loose binding, and a low value to tight binding between enzyme and substrate. At low substrate concentration, more precisely if [S] K M , Eq. (3) simplifies to v = vmax [S]/K M and consequently is first order with respect to the substrate concentration [S]. In contrast, at high substrate concentration, more precisely if [S] K M , Eq. (3) simplifies to v = vmax and consequently is zeroth order with respect to [S]. In all situations, v is proportional (first-order with respect) to [E]. 3 Sources and Reasons for the Activity of Enzymes as Catalysts 3.1 Chronology of the Most Important Theories of Enzyme Activity

Just one year after Ostwald’s hypothesis about the existence of catalysts in 1893, when nobody yet had a clear idea of the structure and composition of enzymes, Emil Fischer voiced the idea for the first time that a substrate molecule fits into the pocket of an enzyme, the “lock-and-key hypothesis” [1]. Both the lock (enzyme) as well as the key (substrate) were regarded as rigid. This hypothesis was modified later in many ways. According to Haldane [15], catalysis of a reaction occurs only if a catalyst in the active center is complementary to the transition state of the substrate during the reaction. Therefore, the transition state between substrate and products fits best into a pocket close to the enzyme. The substrate molecule is subject to strain upon binding to the active center and changes its conformation to fit into the active center; the key (the substrate) does not fit completely into the lock but is strained and bent. In a further modification of this concept, a reaction is accelerated if a catalyst stabilizes the transition state; in contrast, stabilization of the ground state leads to a deceleration of the reaction. This concept of transition-state stabilization, formulated first by Haldane, was expanded later by Linus Pauling [16, 17]. The notion of lowering of G= attributed to the stabilization of the transition state by the catalyst or to the destabilization of the ground state in comparison to the transition state is generally accepted nowadays. Instead of assuming a “solid” enzyme, in which active center the substrate molecule is “bent” (the concept of substrate strain), the idea was developed [2, 18]

5


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6 Enzyme Catalysis

that enzymes can embrace the substrate molecule flexibly in the active center and effect reaction by the formation of specific interactions, the so-called “induced fit”. This picture is especially appropriate with allosterically activated enzymes or in situations in which part of the enzyme molecule has to turn or move over longer distances to effect catalysis (hinge movement), as for instance with most NAD(P)(H)dependent enzymes [19]. An equation for the acceleration of the reaction rate by a deliberate catalyst and the relative strength of the binding to the transition state and ground state was formulated in 1963 by Kurz [20], who combined the ideas of the thermodynamic cycle and transition-state theory and whose result, unrecognized by Kurz, can be regarded as a quantification of the Pauling postulate [20]. Just a few years later Jencks originated the idea of transition-state analogs as inhibitors and cited references in the literature [21]. Finally, it was observed [22] that the best inhibitors for serine proteases such as subtilisin, chymotrypsin, trypsin or elastase all led to geometrically similar transition states for the hydrolysis reaction and all bound in the complementary oxyanion hole. All four proteases belonged to the small number of enzymes whose three-dimensional structure was known by then. 3.2 Origin of Enzymatic Activity: Derivation of the Kurz Equation

The key equation resulting from the application of two known theories, the Born–Haber cycle and the transition-state theory, was formulated for any catalyst and without reference to enzymes [20]; a good derivation can be found in the article by Kraut [23]. Transition-state theory is based on two assumptions, the existence of both a dynamic bottleneck and a preceding equilibrium between a transition-state complex and reactants. Eq. (4) results, where k denotes the observed reaction rate constant, κ the transmission coefficient, and ν the mean frequency of crossing the barrier. k=κ·ν·K =

(4)

All correction factors such as tunneling, back-crossing of the barrier, and solvent frictional effects are captured by κ, which is between 0.1 and 1 for the reactions in solution in question here. The equilibrium constant K = is expressed by partition functions, where the mode normal to the reaction coordinate ν is approximated by the term kB T/hν. In the resulting Eq. (5), ν cancels out.

k=κ·(k B T/ h)·K =

(5)

Note that K = is not the thermodynamic equilibrium constant and kB T/h is not a universal frequency for the decomposition of the transition-state complexes into products. The thermodynamic cycle relevant for further discussion is shown in Figure 2.


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3 Sources and Reasons for the Activity of Enzymes as Catalysts E +

S

E + S≠

KS

KT ES

E + P

[ES] ≠

Fig. 2 Thermodynamic (Born–Haber)-cycle.

If one compares the first-order rate constant for the enzyme-catalyzed singlesubstrate elementary reaction ke with the one for the uncatalyzed reaction ku , Eq. (6) is obtained with Eq. (4). ke /ku =(κe ·νe ·K e= )/(κu ·νu ·K u= )

(6)

In the case of a simple elementary enzyme reaction, ke is identical to kcat . By utilizing the thermodynamic cycle the quotient of the constants of formation of the transition state K e = /Ku = can be equated with the quotient of the dissociation constants for substrate KS and transition state K T [Eq. (7)]. kcat /ku =(κe ·νe ·K S )/(κu ·νu ·K T )

(7)

Although only a few data are available for the comparison of κ e ·ν e , and κ u ·ν u , it is assumed that the two terms do not differ much in magnitude, so that Eq. (8) holds approximately. kcat /ku ≈K S /K T

(8)

This is the equation derived by Kurz, expressed here for the case of an enzyme reaction. The ratio of enzyme-catalyzed to uncatalyzed reaction kcat /ku or kcat /kuncat , i.e., the rate acceleration, often reaches orders of magnitude of 1010 to 1012 [24], and in the best case so far 1.4 · 1017 (orotidine monophosphate decarboxylase, OMPD), which translates into a proficiency kcat /(kuncat · K M ) of 1023 M−1 [25]. This underscores the quality of enzymes as catalysts. In comparison, the best value for the rate acceleration of catalytic antibodies is 2.3 · 108 [26], and typical values are 103 to 105 [27]. Radzicka [24] found that kcat for the reactions in question is often within one or two orders of magnitude; interestingly, the highest efficiencies are observed with reactions of very low kuncat . Thus, Eq. (8) means that transition states in the case of successful catalysis must bind much more tightly to the enzyme than ground states; in this way, Eq. (8) is the mathematical form of the Pauling postulate in 1944 [17]. It is emphasized again that the action of enzymes through the stabilization of the transition state is not an independent concept but is derived from transition-state theory and the idea of a thermodynamic cycle. 3.3 Consequences of the Kurz Equation

The Kurz equation [Eq. (8)] has simplified and channeled many discussions in enzymology Many other explanations of enzyme action and many phenomena

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8 Enzyme Catalysis

observed with enzyme reactions can be reduced to the validity of Eq. (8), as shown in the following paragraphs. Ĺ The principle of transition state stabilization can explain why many enzymes show higher activity against larger, sterically more demanding substrate molecules than against smaller ones. Paradoxically, the cause of the enhanced activity was an increased kcat , not a decreased K M value, i.e., an enhanced maximum rate and not a tighter substrate binding. This phenomenon led to the postulation of “induced fit”. But if the enzyme template binds the transition state more tightly than the ground state, it can be expected that peripheral parts of the substrate molecule have an important role for the binding in the transition state but not necessarily for the binding in the ground state. Conformational changes of the enzyme template do not need to be postulated. Frequently, however, conformational changes during binding and catalysis are observed which are then also called “induced fit” by several authors. Ĺ The exact loci of binding and catalysis cannot be distinguished exactly. In subtilisin, the amino acids forming the catalytic triad, Ser221–His64– were replaced in all combinations by Ala. Even the triple mutant without any amino acid from the original catalytic center displayed a 1000-fold higher reaction rate than the uncatalyzed reaction, and the remainder of the enzyme molecule bound the substrate better in the transition state than in the ground state [28]. Ĺ The numerous interactions between enzyme side chains and substrate molecules act synergistically, so the fit is quite exact. This cooperativity enhances substrate specificity, for a small change of substrate can effect a large change in the binding of the transition state. Triose phosphate isomerase is only 1/1000 as effective if the residue Glu165 is moved away from the substrate by even 1 Å, as was demonstrated with the exchange of Glu165 for Asp165 [29]. The change of only one amino acid of the triad in subtilisin causes an activity decrease of 10−6 , whereas the exchange of all three causes a decrease of only 7 × 10−7 and not of 10−18 [28]. The results seem to be explained by an all-or-nothing situation rather than additivity Ĺ Intramolecular reactions proceed much more rapidly than intermolecular ones owing to entropic stabilization. Page and Jencks have calculated the entropies of translation, rotation, and vibration [30]. The entropy loss of the reactants already realized in the enzyme–substrate (ES) complex allows for a much faster reaction than in the situation without complexation, binding, or geometric orientation. Introduction of an effective concentration [S]eff alleviates the trouble of comparing results of intermolecular catalysis with rate constants of dimension [concentration · time]−1 (second-order reaction) with those of intramolecular catalysis with constants of dimension [time]−1 . The high values often achieved for [S]eff (often > 10 M) reflect the efficiency of intramolecular catalysis. In this context, enzymes are often referred to as “entropy traps”, because many contributions to entropy are “frozen in” after binding of the substrate to the enzyme molecule. Ĺ Many enzymatic reaction mechanisms are accelerated or even only made possible by the absence of water or other solvents. The enzyme supports desolvation


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3 Sources and Reasons for the Activity of Enzymes as Catalysts Table 1 Wholly or partially diffusion-controlled enzyme reactions adapted from [52]

Substrate

Enzyme

k cat [s−1 ]

K M [M]

kcat /KM [s−1 · M−1 ]

CO2 ⇔ HCO3 − H2 O2 fumarate benzylpenicillin

Carbonic anhydrase Catalase Fumarase β-Lactamase

1 × 106 4 × 107 800 2 × 103

0.012 1.1 5 × 10 −6 2 × 10−5

8.3 × 107 4 × 107 1.6 × 108 1 × 108

by redirecting part of the possible binding energy liberated from the binding between enzyme and substrate. Ĺ Upper boundary of rate constant of an enzyme reaction: diffusion control The upper boundary of the reaction rate is reached when every collision between substrate and enzyme molecules leads to reaction and thus to product. In this case, the Boltzmann factor, exp(−E a /RT), is equal to 1 in the transition-state theory equations and the reaction is diffusion-limited or diffusion-controlled (owing to the difference in mass, the reaction is controlled only by the rate of diffusion of the substrate molecule). The reaction rate under diffusion control is limited by the number of collisions, the frequency Z of which can be calculated according to the Smoluchowski equation [Smoluchowski, 1915; Eq. (9)]. −dc/dt=−8π RD·c 2

(9)

R is the normalized radius of both participating particles (1/r 1 + 1/r 2 ), D the normalized diffusion coefficient (1/D1 + 1/D2 ). The collision frequency Z is then calculated from Eq. (10). Z=2RT/(3000η·[(r 1 +r 2 )2 /r 1 r 2 ])

(10)

As the enzyme molecule is much larger than the substrate or product molecule (r E r S ) and thus diffuses much more slowly (DE DS ), Eqs. (9) and (10) can be simplified to Eqs. (11) and (12). −dc/dt=−8π RDE ·c 2

(11)

Z=2RT/(3000η·rE )

(12)

In the case of a highly viscous solution the influence of viscosity η can dominate [31]. For simple, uncharged particles in water at 25 ◦ C the second-order rate constant is 3.2 × 10 (M · s)−1 [32, 33]. Some cases of wholly or partially diffusion-controlled enzyme reactions are listed in Table 1. Rearrangement of Eq. (7) results in Eq. (13). kcat /K S =(ku ·κe ·ve )/(κu ·vu ·K T )

(13)

9


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10 Enzyme Catalysis

Eq. (13) is the manifestation of the second-order rate constant for a reaction between free enzyme and free substrate to give free enzyme and free product; thus, the rate constant cannot exceed the maximum value for kdiff , i.e., 3.2 × 109 M−1 · s−1 . A perfectly evolved enzyme therefore features a decreased binding constant K s , which means a stronger binding in the transition state, possibly up to the thermodynamic limit. Limitation of a reaction by translational diffusion in solution is a rather rare case. Much more frequently the limitation of the observed overall reaction rate is by external mass transfer (through a laminar film around a solid macroscopic carrier) or internal mass transfer (diffusion of substrate or product through the pores of a solid carrier or a gel network to an enzyme molecule in the interior of the carrier). 3.4 Efficiency of Enzyme Catalysis: Beyond Pauling’s Postulate

Let us return one more time to the Kurz equation [Eq. (8)], which is regarded as the quantification of Pauling’s postulate that transition-state stabilization, i.e., the tighter binding between enzyme and transition state, expressed by K T , as compared to binding between enzyme and ground state, expressed by K S , is the source of catalytic rate enhancement. kcat /kuncat ≈K S /K T

(8)

Identifying K S with K M , K T approximately equals the ratio (kuncat · K M )/kcat , which is the inverse of termed the “proficiency”, which has been determined experimentally for a number of systems (see Section 4.4 below). If the Kurz equation captured the whole essence of enzyme catalysis, K T should be proportional to (kuncat · K M )/kcat . However, Figure 3 reveals that the correlation of K T (≡ “K TS ”) for some enzyme reactions is much better with kuncat (≡ “knon ”) than with kcat [34]. This seeming contradiction to Pauling’s postulate and the notion of the transitionstate energy and conformation as the sole source of enzymatic catalytic power cast the geometry of the substrate ready to interact with the enzyme, i.e., substrate preorganization, into a new and more important role. While the discussion on this subject is far from settled, a novel concept based on molecular mechanics calculations of the stability of different substrate conformations, termed substrate conformers, has emerged which emphasized substrate preorganization. The substrate conformers which allow reactions to occur most easily clearly resemble the transition state and are called “near-attack conformers” (NACs) [34, 35]. The relative rate constants of anhydride formation krel from mono-p-bromophenyl esters were found to correlate with the mole fractions (P) of ground-state conformers that could be classified as NACs according to Eq. (14) [8]. log krel =0.94log P+7.48 (r 2 =0.915) The results are depicted in Figure 4.

(14)


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4 Performance Criteria for Catalysts, Processes, and Process Routes

Fig. 3 Comparison of the relationship of K TS with the enzymatic reaction rate constant, kcat , and with the uncatalyzed solution or reference reaction rate constant knon [34].

4 Performance Criteria for Catalysts, Processes, and Process Routes 4.1 Basic Performance Criteria for a Catalyst: Activity, Selectivity and Stability of Enzymes

Every catalyst, and thus also every biocatalyst, can be characterized by the three basic dimensions of merit, namely activity, selectivity, and stability. Additional, but not frequently employed, performance criteria beyond the basic dimensions are discussed in Section 4.3. 1. Biocatalysts often feature much better selectivity than non-biological catalysts, so they are developed for use because of their selectivity, be it enantioselectivity, chemo- or regioselectivity. 2. As activity is straightforward to measure and necessary to know for even the basic experimental protocol, enzyme activity is often well studied. In contrast to other catalysts, most enzymes are only active and stable in a very limited temperature and pH range (mostly between 15 and 50 ◦ C or pH 5 and 10). 3. Unlike activity, stability of enzymes is often interpreted simplistically as thermal stability, i.e., a temperature beyond which the enzyme loses stability. Although this quantity is important, first every statement of stability at a certain temperature depends on exposure time and thus is often ambiguous; and second, for biocatalytic process applications, a more important quantity is the process or operational stability, which is the long-term stability under specified conditions.

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12 Enzyme Catalysis

Fig. 4 Log of the relative rate constants of anhydride formation krel from mono-p-bromophenyl esters vs. the log of the probability (P) of NAC formation of each monophenyl ester of the various dicarboxylic acids [34].

4.1.1 Activity The value of the (overall) enzyme activity is usually provided in “International Units” or “Units”:

1 Unit≡1 IU≡1 U≡1 µmol min−1 In most cases a value for the overall activity of enzyme is not very interesting. Much more important are both the specific activity, scaled to the mass of catalyst, and the volumetric activity, based on the activity per unit volume: 1 U (mg protein)−1 =1µmol (min mg protein)−1 ≡specific activity and 1 U mL−1 =1µmol (min mL)−1 ≡volumetric activity All activity data are meaningless without a specification of the conditions of measurement. Connonen, the activity of enzymes is provided at nearly physiological


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conditions at 30 ◦ C and pH 7.5, unless specified otherwise. Any enzyme sold commercially comes with information about the detailed assay conditions used to arrive at the specific (solid preparation) or volumetric (liquid preparation) activity. To judge the quality of a biocatalyst, its specific activity can serve as a guideline: the threshold for a useful biocatalyst is 1 U (mg pure protein)−1 ; the threshold for a biocatalyst developed for plant-scale level, however, lies much higher, rather closer to 100 U (mg pure protein)−1 . The volumetric activity of a (bio)catalyst can be enhanced by simple addition of more catalyst to a system. Specific activity, however, must be improved through optimization of the reaction conditions, or through variation of the structure of the carrier or even of the enzyme. A mass-independent quantity of activity, superior to specific activity, is the turnover frequency (tof), defined as: turnover frequency (tof)=

number of catalytic events time×number of active sites

The turnover frequency allows performance comparison between different catalyst systems, biological and/or non-biological. Its threshold is at 1 event per second per active site. According to the definition, a turnover frequency can be determined only if the number of active sites is known. For an enzyme reaction obeying Michaelis–Menten kinetics, Eq. (15) holds. tof=1/kcat

(15)

4.1.2 Selectivity The notion of selectivity needs to be specified further: point selectivity is the incremental selectivity, usually towards product, at a specific degree of conversion, whereas integral selectivity is the overall average selectivity at the same specific degree of conversion. Owing to the importance of enantiomeric purity of target product molecules in life science applications and the pre-eminent position biocatalysts enjoy with respect to the achievement of that goal, enantioselectivity is the most important kind of selectivity in the context of biocatalysis. The enantiomeric ratio or E value [36] serves as a measure of enantio-selectivity at a certain degree of conversion [Eq. (16)].

E =(kcat /K M )A /(kcat /K M )B =ln([A]/[A]0 )/([B]/[B]0 )

(16)

Although the E value is measured at a specific degree of conversion, nevertheless it is an integral selectivity measure. This is even more obvious in the case of the other measure commonly employed, the enantiomeric excess, e.e. [Eq. (17)], which reflects overall selectivity up to the point of isolation of the product. e.e.=([A]−[B])/([A]+[B])×100%

(17)

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14 Enzyme Catalysis

The threshold for an enantioselective (bio)catalyst is either at E = 100 or at 99% e.e., depending on whether one is evaluating either the reaction itself or a reaction product. [These values are linked to the requirement by the Food and Drug Administration of the United States (FDA), the most important body in the world for setting guidelines for both procedures for novel drug approval and drug performance evaluation, that a separate toxicological study be conducted for any impurity, including an enantiomer of the active drug, exceeding a concentration level of 0.5%.] Besides enantioselectivity, regioselectivity and to a lesser extent, chemoselectivity are also important issues of enzymatic reaction selectivity. 4.1.3 Stability Stability of an enzyme is usually understood to mean temperature stability, although inhibitors, oxygen, an unsuitable pH value, or other factors such as mechanical stress or shear can decisively influence stability. The thermal stability of a protein, often employed in protein biochemistry, is characterized by the melting temperature T m , the temperature at which a protein in equilibrium between native (N) and unfolded (U) species, N ⇔ U, is half unfolded. The melting temperature of a protein is influenced on one hand by its amino acid sequence and the number of disulfide bridges and salt pairs, and on the other hand by solvent, added salt type, and added salt concentration. Protein structural stability was found to correlate also with the Hofmeister series [37–39] [Eq. (18)].

Tm =T0 +K [S]

(18)

In Eq. (18), T 0 is the melting temperature in the absence of salt, [S] the salt concentration, and K the corresponding coefficient. Storage stability over time under fixed conditions of temperature, pH value and concentration of additives often can be expressed by a first-order decay law (analogous to radioactive decay) [Eq. (19)]. [E ]t =[E]0 exp(−kd ·t)

(19)

The validity of a first-order decay law over time for the activity of enzymes according to Eq. (19), with [E]t and [E]0 as the active enzyme concentration at time t or 0, respectively, and kd as the deactivation rate constant, is based on the suitability of thinking of the deactivation process of enzymes in terms of Boltzmann statistics. These statistics cause a certain number of active protein molecules to deactivate momentarily with a rate constant proportional to the amount of active protein [for evidence for such a “catastrophic” decomposition, see Craig [40]]. The relevant stability criteria for process applications (the process stability or operating stability) is discussed in the next section.


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4.2 Performance Criteria for the Process

In evaluating a biocatalyst for a given processing task, there are performance criteria to be met not only for the biocatalyst but also for the process. The dimensions of merit important when determining, evaluating, or optimizing a route for a process are (i) product yield, (ii) (bio)catalyst productivity, (iii) (bio)catalyst stability, and (iv) reactor productivity. 4.2.1 Product Yield Chemical yield to product is most important for the economics of the process. Product yield is inversely proportional to the amount of reactants required per unit of product output. As the key substrate and other raw materials in most mature processes make up more than 50% of the variable cost, a high product yield is indispensable for an economic process. The yield of product y is linked to selectivity σ and the degree of conversion x by Eq. (20).

y=x·σ

(20)

so that the product concentration [P] can be calculated from Eq. (21). [P]=[S]0 ·y=[S]0 ·x·σ

(21)

Yields much less than 100% are no longer acceptable either economically or ecologically. In the separation of a racemate, where the yield per run is limited to 50% per pass, this means that internal or external racemization is necessary unless either the substrate is inexpensive enough to lose up to 50% or the co-product can be marketed in similar amounts. Both of the latter situations are highly unlikely. A threshold for a sufficiently high yield is hard to define, as the threshold value seems to correlate inversely with the unit value of the product. While for basic, large-volume chemicals yields of 98 or 99% are absolutely essential, the situation in fine chemicals calls for 90–95% yield, and in the initial stage of production of extreme performance chemicals, such as pharmaceuticals, yields of > 80% are very acceptable; sometimes values down to 50% have to be encountered. Acceptable yields depend on the number of process steps, including product isolation. If all the steps are assumed to fetch 90% yield, the overall yield depends on the number of steps n as in Eq. (22). yoverall =(ystep )n =(0.9)n

(22)

so that for a one-step process yoverall,1 = 90%, for three-step process yoverall,3 = 73%, and for a ten-step process yoverall,10 = 35%!

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16 Enzyme Catalysis

4.2.2 (Bio)catalyst Productivity If a large amount of (bio)catalyst is added to a substrate solution better results are achieved than if just a small amount of a highly productive catalyst is used. To even call an agent a catalyst, one condition is that the agent be added, as a minimum, in substoichiometric amounts. The smaller the amount of catalyst that has to be added for the same result, the better its performance. The relevant criterion is the dimensionless turnover number, TON [Eq. (23)].

TON=

substrate amount of product = =[S]/[C] amount of catalyst catalyst ratio

(23)

The turnover number is not used frequently in biocatalysis, possibly as the molar mass of the biocatalyst has to be known and taken into account to obtain a dimensionless number, but it is the decisive criterion, besides turnover frequency and selectivity, for evaluation of a catalyst in homogeneous (chemical) catalysis and is thus quoted in almost every pertinent article. Another reason for the low popularity of the turnover number in biocatalysis, apart from the challenge of dimensionality, is the focus on reusability of biocatalysts and the corresponding greater emphasis on performance over the catalyst lifetime instead of in one batch reaction as is common in homogeneous catalysis [41]. For biocatalyst lifetime evaluation, see Section 4.2.3. The turnover number of a catalyst does not refer to the timescale of activity of that catalyst in a process. If the timescale of activity is taken into consideration, the productivity of the catalyst is recovered, expressed as a productivity number, PN, defined in Eq. (24). PN=

mass of product unit catalyst×time

(24)

4.2.3 (Bio)catalyst Stability As in all catalytic processes, catalyst stability is a key process criterion. In contrast to mere temperature or storage stability, which refer to the catalyst independently of a process, the operating stability or process stability is the relevant and decisive dimension of merit. It is determined by comparing the amount of product generated with the amount of catalyst spent. The relevant quantity, also sometimes found in homogeneous catalysis, is the total turnover number (TTN) [Eq. (25)].

TTN=

moles of product produced mole of catalyst spent

(25)

Quite common in applied biocatalysis, where the purity of biocatalyst often is not known, is the expression of biocatalyst stability as an the enzyme consumption number (e.c.n.) [Eq. (26)]. e.c.n.=

g of enzyme amount of enzyme preparation spent = amount of product generated kg (or lb) of product

(26)


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The e.c.n. value depends on process parameters such as temperature, pH value, and concentrations of substrate(s) and product(s). With the molar masses of enzyme and product, e.c.n. and TTN can be interconverted [Eq. (27)]. TTN=(1000/e.c.n.)×(MWenzyme /MWproduct )

(27)

It should be emphasized that the TTN is not a completely suitable quantity for the evaluation of operating stability because the number of moles of biocatalyst is not a suitable reference for the complexity and cost of its manufacture. However, the values for both numerator and denominator in Eq. (27) are usually known and can be expressed in monetary terms. For checking the application of such a biocatalyst, the contribution of the biocatalyst to the overall cost can be assessed readily. The relevant parameter for studies of operating stability of enzymes is the product of active enzyme concentration [E]active and residence time τ , [E]active · τ . In a continuously stirred tank reactor (CSTR) the quantities [E]active and τ are linked by Eq. (28), where [S0 ] denotes the initial substrate concentration, x the degree of conversion and r(x) the conversion-dependent reaction rate [42, 43]. ([E]active ·τ )/[S0 ]=x/r (x)

(28)

For a comparison and discussion of the concepts of stability at rest [Eq. (19)] and operating stability [Eq. (28)]. The threshold value for sufficient biocatalyst stability depends on the application, as does the value for product yield. For any application in synthesis the TTN should exceed 10 000, and for large-scale processing a value of > 1 000 000 is preferred. 4.2.4 Reactor Productivity For the assessment of reactor productivity that is independent of catalyst, the spacetime yield (s.t.y) [Eq. (29)]. is considered, as in any chemical process.

s.t.y.=mass of product generated/(reactor volume×time)[kg(L·d)−1 ]

(29)

An increase in s.t.y. in the same reactor is equivalent to an increase in reaction rate. When considering the Michaelis–Menten equation [Eq. (3): v = vmax [S]/(K M + [S]) = kcat [E][S]/(K M + [S])], there are three possible ways to achieve a maximum reaction rate: 1. High substrate concentration (increase of space-yield). Enzyme reactions can be accelerated by increasing the substrate concentration up to the limit of saturation (≈ 10K M ). If [S] K M , the enzyme is saturated and Eq. (3) is reduced to Eq. (30).

v=vmax =kcat ·[E]

(30)

17


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18 Enzyme Catalysis

At substrate saturation, the reaction is zeroth-order with respect to substrate. With a loosely binding substrate, i.e., a high K M value, the enzyme is often saturated only above the solubility limit of the substrate. To improve the reaction rate or the s.t.y. further, according to Eq. (29) the yield per unit time must be increased, which means enhancing either the biocatalyst concentration and/or the time constant kcat . 2. High enzyme concentration. The reaction rate and s.t.y. can be enhanced by increasing the catalyst concentration [E]; in practice, however, in contrast to the formalism of Eq. (3), owing to an either excessive viscosity increase or excess of deactivated protein in the reactor, a maximum limit of enzyme concentration is reached. 3. Increase in the time constant. According to Eq. (3), kcat signifies the time constant of the enzyme reaction [time−1 ], and the corresponding reaction is performed over a time scale 1/kcat [time]. An increase can be effected through the temperature (Arrhenius behavior) but also for instance through changing a protecting group in peptide synthesis [44]. An increase in kcat also increases the acceleration ratio kcat /kuncat [24] of the enzyme catalyst, in contrast to case 2) (see Section 4.3).

A minimum threshold value for reactor productivity can be set at a space-timeyield of about 100 g (L · d)−1 , a value which tends to be compromised more by lack of substrate solubility than by biocatalyst reactivity. Well-developed biocatalytic process often feature space–time yields of > 500 g (L · d)−1 or even > 1 kg (L · d)−1 [44–46]. 4.3 Links between Enzyme Reaction Performance Parameters 4.3.1 Rate Acceleration While users of biocatalysts are often concerned first and foremost about a high maximum rate vmax , a high kcat value, and possibly also a low K M value, a good (bio)catalyst is one that enhances the chemical background rate by as high a factor as possible, at least 105 , possibly 1010 or even more. The dimensionless ratio of the enzyme catalytic rate constant over the uncatalyzed (i.e., the chemical background) rate constant, kcat /kuncat , is called the rate acceleration. An alternative nomenclature used in the literature, the efficiency or proficiency, should be reserved for the term kcat /(kuncat · K M ) quantifying the total energetic cost of enzymatic catalysis [47]. Surprisingly, when analyzing the source of good rate accelerations, i.e., high kcat /kuncat values, or the source of high proficiencies, i.e., high kcat /(kuncat · K M ) values, kcat was found to be within two orders of magnitude for a variety of enzymes whereas kuncat varied by several orders of magnitude, irrespective of the value of kcat (Figures 5 and 6; [24, 48]). The data provides clear guidance that the biggest improvement in enzyme catalysts can be achieved for reactions with very low chemical background rate constants and not by optimizing rate constants or specificities which are already fairly high.


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10 5 rate constant (e–1)

enzyme rate constants (kcat)

catalyzed

1 t 1/2 1 minute

10 –5 t1/2 1 week uncatalyzed

t 1/2 100 years 10

–10

t 1/2 1 million years 10–15 t 1/2 4 billion years

nonenzymatic rate constants (knon)

Fig. 5 Logarithmic scale comparison of kcat and kuncat (= knon ) for some representative reactions at 25 ◦ C. The length of each vertical bar represents the rate enhancement. [48]. ADC: arginine decarboxylase; ODC: orotidine 5 -phosphate decarboxylase; STN:

staphylococcal nuclease; GLU: sweet potato $-amylase; FUM: fumarase; MAN: mandelate racemase; PEP: carboxypeptodase B; CDA: E. coli cytidine deaminase; KSI: ketosteroid isomerase; CMU: chorismate mutase; CAN: carbonic anhydrase.

4.3.2 Ratio between Catalytic Constant kcat and Deactivation Rate Constant kd The rate equation for a deactivating enzyme in a batch reactor [Eq. (31)] reads:

x·[S]0 −K M ln(1−x)=[E]0 ·(kcat /kd )·{1−exp(−kd ·t)}

(31)

The influence of deactivation depends linearly on the dimensionless ratio kcat /kd , which might serve as a ratio to assess quickly the potential of a deactivating enzyme for synthesis. 4.3.3 Relationship between Deactivation Rate Constant kd and Total Turnover Number TTN Specific enzyme consumption [U per kg product], a hands-on parameter, can be obtained from Eq. (32) [49], where avol,0 [U L−1 ] is the initial volumetric activity).

specific enzyme consumption=(avol,0 ·kd )/(s.t.y.)[U(kg product)−1 ]

(32)

A high value for s.t.y. and low values for [E] (a modified enzyme concentration [g enzyme L−1 ]), avol,0 , or kd all lead to a favorable, i.e., low, specific enzyme consumption. To convert the specific enzyme consumption into the total turnover number TTN, the specific enzyme consumption, sp.e.c. [U (kg product)−1 ] [Eq. (32)] first has to be

19


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20 Enzyme Catalysis 1010 k cat / K m

enzyme efficiencies (k cat /K m)

(M –1 s–1)

catalyzed

10 5

1 t 1/2 1 minute

10 –5 t1/2 1 week uncatalyzed

t 1/2 100 years 10 –10 rate constant (s–1)

k non (s–1)

t 1/2 1 million years 10 –15 t 1/2 4 billion years

nonenzymatic reaction constants (k non)

Fig. 6 Logarithmic scale comparison of kcat /K M and kuncat (≡ knon ) for some representative reactions at 25◦ C. The length of each vertical bar represents the transition-state affinity or catalytic proficiency (its reciprocal). For abbreviations, see Figure 5 [48].

converted into the enzyme consumption number e.c.n. [g enzyme (kg product)−1 ] [Eq. (26)].

e.c.n.=(sp.e.c.·[E] )/avol,0 [g enzyme(kg product)−1 ]

(33)

If Eq. (32) is inserted into Eq.(33) and the resulting equation into Eq. (27),the result is Eq.(34), which with [E] /MWenzyme = [E]0 [mol L−1 ] becomes Eq. (35). TTN[mol product(mol catalyst)−1 ] =(MWenzyme /MWproduct )(100×s.t.y.)/([E] ·kd )

(34)

TTN=(1000×s.t.y.)/MWproduct ·[E]0 ·kd

(35)

A favorable, i.e., high, TTN is achieved by driving up the s.t.y. of a pertinent experiment while keeping both [E]0 and kd low. (The conversion factor of 1000 does not appear if s.t.y. is expressed in [g (L · d)−1 ] and not [kg (L · d)−1 ].)


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4.4 Performance Criteria for Process Schemes, Atom Economy, and Environmental Quotient

Roger Sheldon, of TU Delft (Delft, The Netherlands), has compiled a list of trends for chemical and biochemical process technology [50]; they are towards Ĺ Ĺ Ĺ Ĺ Ĺ

catalytic reactions instead of stoichiometric ones, reactions with 100% selectivity at 100% conversion, high substrate concentrations, no detrimental solvents, and no change of solvent during the process, enhanced use of solid or volatile acids and bases as well as pH-stat techniques.

We recognize the importance of high conversion, high selectivity, and thus high product yields; and of high substrate concentration, often leading to high s.t.y. and TTN. However, there are several additional guidelines which help to optimize a complete process. Some of those criteria have been discussed above in the context of the catalyst or the process, such as enantioselectivity or diastereoselectivity instead of simple selectivity to product, and catalyst performance data such as turnover frequency (tof), turnover number (TON), total turnover number (TTN), and space–time-yield (s.ty). Others have to deal with inputs and outputs of the process apart from reactants, catalysts, and products, such as: 1. the atom economy, i.e., the fraction of carbon (or other elements) of the substrate that is utilized or recovered in the product 2. the specific consumption (consumption per kg product) of solvents, salts, and other auxiliaries such as work-up materials (active carbon, filter aids), and 3. the degree of recovery of all materials, ranging from main process components to solvents, salts, and traces as well as contents of trace streams, often termed ancillary losses.

ad 1) Atom economy The degree of utilization of inputs in the final product is termed the atom economy [51]. An atom-economic process uses as many atoms as possible in the reaction stoichiometry Examples of simple, but atom-economic, chemical processes are the manufacture of maleic anhydride by air-oxidation of butane/butene instead of benzene, or of propylene oxide by air-oxidation of propylene (propene) instead of by ring-closing substitution of epichlorohydrin (with loss of HCl). An example of a process with low atom economy is any synthesis with protection/deprotection steps, such as those that occur during peptide synthesis (while possibly still featuring very high product yields). ad 2) Specific consumption of solvents, salts, and auxiliaries This can be captured by evaluating the environmental quotient, EQ [50], consisting of a relative mass of by-product versus target product [kg kg−1 ] (or for our American readers, lb lb−1 )

21


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22 Enzyme Catalysis Table 2 Comparison of conditions for laboratory- and industrial-scale processes

Medium Substrate pH control Time Temperature, pH gradients

Laboratory-scale processes

Industrial-scale processes

dilute solutions often natural with buffer sufficient for reaction none

(highly) concentrated media often unnatural titration with acid/base; solid acids/bases fast reactions desired heat and mass transfer influence reaction

multiplied by a measure of the environmental unfriendliness. Water or even NaCl might carry a factor of 1 or close to 1, whereas mercury, halogenated solvents, or other toxic by-products might carry a factor of 10 or even 100. ad 3) Recovery of all materials Eco-balances have become increasingly popular with regulators to gauge the impact of discharges. Increasingly, such instruments replace simple levying of costs on discharges or setting inflexible upper limits for concentrations of a range of ecologically unfriendly agents. When aiming for scale-up or when actually already running a process on a pilot or a full industrial scale instead of trying to gain knowledge about a problem under controlled laboratory conditions, one should ensure that the different goals are reflected in different measures taken to obtain data or to develop a catalyst or process for an ultimate large scale. Table 2 contrasts typical laboratory practices with conditions prevailing on production scale. It is prudent, however, for processes to be practiced on a large scale to be developed accordingly, even during the test phase on lab scale. Suggested Further Reading Alan FERSHT, Structure and Mechanism in Protein Science, Freeman, New York, 1999.

William P. JENCKS, Mechanisms in Chemistry and Enzymology, Dover, New York, 1975.

References 1 E. FISCHER, Einfluss der Configuration auf die Wirkung der Enzyme, Ber. dtsch. chem. Ges. 1894, 27, 2985–2993. 2 D. E. KOSHLAND Jr., Application of a theory of enzyme specificity to protein synthesis, Proc. Natl. Acad. Sci. U.S.A., 1958, 44, 98–104. 3 C. TANFORD, Physical Chemistry of Macro-molecules, Wiley, New York, 1961. 4 I. CHAMBERS, J. FRAMPTON, P. GOLDFARB, N. AFFARA, W. MCBAIN, and P. R. HARRISON, The structure of the mouse

glutathione peroxidase gene: the selenocysteine in the active site is encoded by the ‘termination’ codon, TGA, EMBO J. 1986, 5, 1221–1227. 5 F. ZINONI, A. BIRKMANN, T. C. STADTMAN, and A. BOECK, Nucleotide sequence and expression of the selenocysteinecontaining polypeptide of formate dehydrogenase (formate-hydrogenlyase-linked) from Escherichia coli, Proc. Natl. Acad. Sci. USA 1986, 83, 4650–4654.


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References 6 B. HAO, W. GONG, T. K. FERGUSON, C. M. JAMES, J. A. KRZYCKI, and M. K. CHAN, A new UAG-encoded residue in the structure of a methanogen methyltrans-ferase, Science 2002, 296, 1462–1466. 7 G. SRINIVASAN, C. M. JAMES, and J. A. KRZYCKI, Pyrrolysine encoded by UAG in Archaea: charging of a UAG-decoding specialized tRNA, Science 2002, 296, 1459–1462. 8 R. A. MEHL, J. C. ANDERSON, S. W. SANTORO, L. WANG, A. B. MARTIN, D. S. KING, D. M. HORN, and P. G. SCHULTZ, Generation of a bacterium with a 21 amino acid genetic code, J. Am. Chem. Soc. 2003, 125, 935–939. 9 S. J. POLLACK, J. W. JACOBS, and P. G. SCHULTZ, Selective chemical catalysis by an antibody, Science 1986, 234, 1570–1573. 10 A. TRAMONTANO, K. D. JANDA, and R. A. LERNER, Chemical reactivity at an antibody binding site elicited by mechanistic design of a synthetic antigen, Proc. Natl. Acad. Sci. USA 1986, 83, 6736–6740. 11 T. R. CECH, RNA as an enzyme, Sci. Am. 1986, 255(5), 64–75. 12 T. R. CECH, RNA. Fishing for fresh catalysts, Nature, 1993, 365, 204–205. 13 L. MICHAELIS and M. L. MENTEN, Die Kinetik der Invertinwirkung, Biochem. Z. 49, 1913, 333–369. 14 O. A. HOUGEN and K. M. WATSON, Chemical Process Principles, Part III, Wiley, New York. 1947. 15 J. B. S. HALDANE, Enzymes, M.I.T. Press, Cambridge/MA, USA, 1965. 16 L. PAULING, Molecular architecture and biological reactions, Chem. Eng. News 1946, 24, 1375–1377. 17 L. PAULING, Chemical achievement and hope for the future, Am. Sci. 1948, 36, 51–58. 18 D. E. KOSHLAND Jr., G. NEME´ THY, and D. FILMER, Comparison of experimental binding data and theoretical models in proteins containing subunits, Biochemistry 1966, 5, 365–385. 19 T. J. STILLMAN, A. M. B. MIGUEIS, X.-G. WANG, P. J. BAKER, L. BRITTON, P. C. ENGEL, and D. W. RICE, Insights into the mechanism of domain closure and

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substrate sof glutamate dehydrogenase from Clostridium symbosium, J. Mol. Biol. 1999, 285, 875–885. J. L. KURZ, Transition state characterization for catalyzed reactions, J. Am. Chem. Soc. 1963, 85, 987–991. W. P. JENCKS, in: Catalysis in Chemistry and Enzymology, McGraw-Hill, New York, 1969, pp. 288–291. J. KRAUT, Serine proteases: structure and mechanism of catalysis, Ann. Rev. Biochem. 1977, 46, 331–358. J. KRAUT, How do enzymes work?, Science 1988, 242, 533–540. A. RADZICKA and R. WOLFENDEN, A proficient enzyme, Science 1995, 267, 90–93. J. K. LEE and K. N. HOUK, A proficient enzyme revisited: the predicted mechanism for orotidine monophosphate decarboxy-lase, Science 1997, 276, 942–945. G. ZHONG, R. A. LERNER, and C. F. BARBAS III, Enhancement of the repertoire of catalytic antibodies with aldolase activity by combination of reactive immunization and transition state theory, Angew. Chem. Int. Ed. 1999, 111, 3957–3960. J. HASSERODT, Organic synthesis supported by antibody catalysis, Synlett 1999, 12, 2007–2022. P. CARTER and J. A. WELLS, Dissecting the catalytic triad of a serine protease, Nature, 1988, 332, 564–568. D. JOSEPH-MCCARTHY, L. E. ROST, E. A. KOMIVES, and G. A. PETSKO, Crystal structure of the mutant yeast triosephos-phate isomerase in which the catalytic base glutamic acid 165 is changed to aspartic acid, Biochemistry 1994, 33, 2824–2829. M. I. PAGE and W. P. JENCKS, Entropic contributions to rate accelerations in enzymic and intramolecular reactions and the chelate effect, Proc. Natl. Acad. Sci. USA 1971, 68, 1678–1683. U. EICHHORN, A. S. BOMMARIUS, K. DRAUZ, and H.-D. JAKUBKE, Synthesis of dipeptides by suspension-to-suspension conversion via thermolysin catalysis – from analytical to preparative scale, J. Peptide Sci. 1997, 3, 245–251.

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24 Enzyme Catalysis

32 H. GERISCHER, J. F. HOLZWARTH, D. SEIFERT, and L. STROHMAIER, Flow method with integrating observation for very fast irreversible reactions, Ber. Bunsenges. Physik. Chem. 1969, 73, 952–955. 33 C. CREUTZ and N. SUTIN, Vestiges of the inverted region for highly exergonic electron-transfer reactions, J. Am.Chem. Soc. 1977, 99, 241–243. 34 T. C. BRUICE and S. J. BENKOVIC, Chemical basis for enzyme catalysis, Biochemistry 2000, 39, 6267–6274. 35 T. C. BRUICE, A View at the Millenium: the efficiency of enzymatic catalysis, Acc. Chem. Res. 2002, 35, 139–148. 36 C. S. CHEN, Y. FUJIMOTO, G. GIRDAUKAS, and C. SIH, Quantitative analyses of biochemical kinetic resolutions of enantiomers, J. Am. Chem. Soc. 1982, 104, 7294–7299. 37 F. HOFMEISTER, Zur Lehre von der Wirkung der Salze. Zweite Mitteilung, Arch. Exp. Pathol. Pharmakol. 1888, 24, 247–260. 38 P. H. VON HIPPEL and K.-Y. WONG, Neutral salts: the generality of their effects on the stability of macromolecular conformation, Science 1964, 145, 577–580. 39 J. K. KAUSHIK and R. BHAT, A mechanistic analysis of the increase in the thermal stability of proteins in aqueous carboxylic acid salt solutions, Protein Sci. 1999, 8, 222–233. 40 D. B. CRAIG, E. A. ARRIAGA, J. C. Y. WONG, H. LU, and N. J. DOVICHI, Studies on single alkaline phosphatase molecules: reaction rate and activation energy of a reaction catalyzed by a single molecule and the effect of thermal denaturation – the death of an enzyme, J. Am.Chem. Soc. 1996, 118, 5245–5253. 41 H. U. BLASER, F. SPINDLER, and M. STUDER, Enantioselective catalysis in fine chemicals production, Appl. Catal. A: General 2001, 221, 119–143. 42 C. WANDREY, Reaction engineering studies on enzyme catalysts for the development of continuous processes, 913: Habilitation Thesis, TH Hannover, Germany, 1977. 43 A. S. BOMMARIUS, K. DRAUZ, U. GROEGER, and C. WANDREY, 1992, Membrane

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bio-reactors for the production of enantiomer ically pure α-Amino acids, in: Chirality in Industry, A. N. COLLINS, G. N. SHELDRAKE, and J. CROSBY (eds.), Wiley & Sons Ltd., London, 930: Chapter 20, pp. 371–397. A. FISCHER, A. S. BOMMARIUS, K. DRAUZ, and C. WANDREY, A novel approach to enzymatic peptide synthesis using highly solubilizing Nα -protecting groups of amino acids, Biocatalysis 1994, 8, 289–307. J. D. ROZZELL, Biocatalysis at commercial scale: myths and realities, Chimica Oggi 1999, 951: (6/7), 42–47. A. S. BOMMARIUS, M. SCHWARM, and K. DRAUZ, Comparison of different chemo-enzymatic process routes to enantiomeri-cally pure amino acids, Chimia 2001, 55, 50–59. E. A. TAYLOR, D. R. J. PALMER, and J. A. GERLT, The lesser ‘burden borne’ by o-succinyl-benzoate synthase: an ‘easy’ reaction involving a carboxylate carbon acid, J. Am. Chem. Soc. 2001, 123, 5824–5825. R. WOLFENDEN and M. J. SNYDER, The depth of chemical time and the power of enzymes as catalysts, Acc. Chem. Res. 2001, 34, 938–945. U. KRAGL, D. VASIC-RACKI, and C. WANDREY, Continuous production of L-tert-leucine in series of two enzyme membrane reactors, Bioproc. Eng. 1996, 14, 291–297. R. A. SHELDON, Consider the environmental quotient, CHEMTECH 1994, 3, 38–47. B. M. TROST, On inventing reactions for atom economy, Acc. Chem. Res. 2002, 35, 695–705. A. FERSHT, Enzyme Structure and Mechanism, 2nd edition, Freeman, New York, 1985. P. T. ANASTAS and T. C. WILLIAMSON, Frontiers in green chemistry, Green Chemistry 1998, 1–26. M. VON SMOLUCHOWSKI, Versuch einer mathematischen Theorie der Koagulations-kinetik kolloidaler Lösungen, Z. physik. Chem. 1917, 92, 129–168.

1

Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes Ariel B. Lindner, Zelig Eshhar, and Dan S. Tawfik Weizmann Institute of Science, Rehovot, Israel


Abbreviations

TS – transition state; TSA – TS analog; OAS – oxyanion stabilization; OAH – oxyanion hole; AChE – acetylcholinesterase; D-Abs – D-Antibodies (esterolytic antibodies D2.3, D2.4, and D2.5).

1 Introduction

During the past 16 years, knowledge gained from studies of natural hydrolases and the mechanism of hydrolytic reactions led to a variety of strategies for generating antibodies that catalyze hydrolytic reactions. In particular, ester hydrolysis became the most intensely studied antibody-catalyzed reaction. Esterolytic antibodies provided the first ‘proof of principle’ of antibody-based catalysis, spearheading the wide spectrum of antibody catalysts detailed in the chapters of this book. Although esterolytic antibodies do not, in general, exhibit enzyme-like rates, knowledge may be extracted from the many mechanistic and structural studies of hydrolytic antibodies concerning the mechanism of action of hydrolytic enzymes. Comparing hydrolytic antibodies to hydrolytic enzymes is the underlying theme of this chapter. Specific-base-(hydroxyl)-catalyzed and general-base-catalyzed is the mechanism by which most natural esterases act (Fig. 1). The reaction proceeds via relatively low activation barriers (the addition TS being higher than the elimination TS [1]), and a well-defined singular tetrahedral, oxyanionic intermediate (I1 , Fig. 1a). Inherent in hydroxyl-catalyzed ester hydrolysis is the linear of the hydrolytic rates. In implementing these basic mechanisms, which are also seen in solution, enzymes apply complex mechanisms, combining a variety of catalytic forces, as exemplified Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes

Fig. 1 Mechanisms of ester hydrolysis. (a) Hydroxide-catalyzed ester hydrolysis. The rate-limiting step consists of an exogenous hydroxide attack on the ester’s carbonyl, forming, via the addition TS1, a tetrahedral oxyanionic intermediate (I1 ). The latter collapses, via the elimination TS2, to form the carboxylate and alcohol products. Top formula: a representative phosphonate TSA, where, for the D-antibodies, R = glutaryl glycine and R = p-nitrobenzyl [24]. The numbers correspond to bond lengths de-

rived from ab initio calculations [1]. (b) The endogenous nucleophile mechanism of Serine hydrolases. The nucleophilic attack is carried out by the active site’s seryl alkoxide, activated by an active-site general-base residue (B:) to form a tetrahedral oxyanionic intermediate (I2 ). This is followed by a rate-limiting deacylation step mediated by a hydroxide ion generated in situ by the basecatalyzed proton elimination of an activesite water molecule, releasing the products via a second intermediate (I3 ) [2].

by three major families of hydrolytic enzymes: seryl/cysteyl hydrolases (Fig. 1b), aspartyl hydrolases (Fig. 1c), and zinc hydrolases (Fig. 1d). In essence, two major catalytic forces are utilized by these enzymes [2]: i) A nucleophilic attack is mediated by a precisely positioned nucleophile, either an endogenous general base (seryl alkoxide (Fig. 1b, step I)) or an explicitly bound specific base (a hydroxyl generated via general-base-catalyzed

1 Introduction

Fig. 1 (c) The general base – general acid (“push-pull”) mechanism of Aspartic hydrolases. An active-site bound water molecule is activated by a general-base carboxylate residue, whereas the oxyanion tetrahedral intermediate (I4 ) is stabilized by a proton donation from a second aspartyl residue. Residues maintaining the carboxylates’ protonation level [135] are not shown for clarity. An alternative “symmetric” model can be found in [136]. (d) The metal-mediated

mechanism of Zinc hydrolases. The nucleophilic attack is carried out by a zinc-bound water molecule, activated by a general-base Glu residue. The formed oxyanionic intermediate (I5 ) is stabilized by its coordination to the metal ion. Though a similar fivecoordinated zinc intermediate has recently been observed [137], a four-coordinated intermediate was previously suggested [2, 138].

deprotonation of an active-site water molecule; Fig. 1a, step II, 1c and 1d), to yield a tetrahedral, oxyanionic intermediate. ii) Oxyanionic stabilization (OAS) of the tetrahedral, oxyanionic intermediate and the TSs leading to its formation and breakdown. The negative charge developed on the carbonyl oxygen is stabilized by hydrogen bonding (Fig. 1b, c) or proton transfer (general-acid catalysis), or by a positive charge (Fig. 1d). iii) Protonation of the alkoxide leaving group by general-acid catalysis.

The relative positioning of various active-site residues involved in the above mechanisms provides the recognition and stabilization of the tetrahedral transition

3


state (TS). It is therefore expected that, although described as separate forces or contributions, their effect on catalysis is coupled. This coupling is indeed encoded in the enzymes’ active-site structure – for example, the seryl residue (Fig. 3), the aspartyl residues (positioned by a shared a hydrogen bond; Fig. 1b) and the zinc ion (Fig. 1c) all provide both the endogenous nucleophile as well as a hydrogen bond to the TS oxyanion. In addition to mechanism above, enzymes take advantage of a combination of other forces, including medium (solvation/desolvation effects), substrate (groundstate) destabilization, remote recognition, as well as co-ordinated conformational changes [2, 3]. Thus, the remarkable catalytic efficiency of natural enzymes, at times limited only by the rate of diffusion, is the outcome of many forces acting in concert. The separation of these factors and the assessment of their independent contributions to catalysis is far from trivial, as exemplified by various mutagenesis studies (e.g., see [4, 5]). Catalytic antibodies, raised to mimic single facets of the complex enzymatic regimes, provide a unique opportunity to examine and quantify enzymatic forces and mechanisms of action individually. 2 Chapter Overview

The first and most widely applicable strategy for generating esterolytic antibodies is by immunizing with analogs of the tetrahedral, oxyanionic intermediates of ester hydrolysis (Fig. 1a), which are generally referred to as transition state analogs (TSAs). Analysis of the structure, mechanism and catalytic efficiency of these antibodies provides the opportunity to compare and assess oxyanion stabilization (OAS) as the primary source of catalysis in both esterolytic antibodies and natural, hydrolytic enzymes (Section 2.1). Surprisingly, several antibodies directed towards OAS (e.g., 43C9 [6]) have incorporated, haphazardly, an endogenous nucleophile as part of their mechanism of action. These antibodies, together with studies attempting to directly obtain antibodies with an endogenous nucleophile (e.g., reactive immunization [7] and chemical modification strategies [8, 9]), form the basis for our discussion on nucleophilic reactivity (Section 2.2). This is accompanied by an analysis of “chemical rescue” experiments, where strong exogenous nucleophiles supplement the initial OAS activity of various antibodies [10–13] (Section 2.3). Several studies, attempting at the generation of esterolytic antibodies with general-acid/base mechanism either by a “bait and switch” strategy (Section 2.4) or metal incorporation (Section 2.5), are then qualitatively assessed. Finally, the roleof conformational changes, suggested in a number of catalytic antibodies [14–20], is described vis-a-vis known contributions of conformational changes to natural enzymes (Section 2). 2.1 Catalysis by Oxyanion Stabilization (OAS)

The most commonly used TSA haptens for the generation of esterolytic antibodies are phosphonates, although other phosphate derivatives such as phosphinates,

2 Chapter Overview

phos-phonoamidates, and phosphothioates have also been used. This indirect approach for the generation of biocatalysts was first formulated by Jencks [21]. Immunization with these analogs generates antibodies that catalyze ester hydrolysis via preferential binding of the reaction’s tetrahedral, oxyanionic TS compared to the planar, uncharged ester substrate (Fig. 1a). This approach relies on optimal representation of the reaction’s TSs by the TSA. It has been shown, for example, that substrate analogs completely fail to generate esterolytic antibodies [22]. Thus, as discussed below, TSA design is a primary issue. 2.1.1 Fidelity of TSA design Phosphate derivatives that are potent inhibitors of hydrolases were shown to exhibit TSA characteristics, albeit with some important discrepancies ([23] and references therein). Ab initio calculations suggest that these TSAs better present the elimination TS2 , vis-a-vis the addition TS1 , the higher and thus the rate-limiting barrier of the two TSs [1] (Fig. 1a). The TSA P-O bonds overestimate the corresponding lengths of the TS, apart from their being significantly shorter (1.48 Å) than the 2.2 Å distance in the addition TS between the incoming hydroxyl nucleophile and the ester’s carbonyl carbon (Fig. 1a). The symmetrical charge partition between the phosphonates’ oxygens overestimates the charge formed in the actual TS and of the incoming hydroxyl nucleophile. Both of the phosphonates’ oxygens are expected to bait antibodies with H-donor residues. This is crucial to stabilizing the developing oxyanionic charge, yet “baiting” a basic residue would be preferable in the position representing the nucleophilic attack. The latter may act as a general base to activate the incoming nucleophile or as an endogenous nucleophile. Nonetheless, despite the above-described infidelities, phosphonate TSAs (and related derivatives) are fairly close mimics of the TSs of ester hydrolysis and hence elicit antibodies in which OAS is the primary, or often the sole, source of catalysis. 2.1.2 Esterolytic Antibodies Based Solely on Oxyanion Stabilization Our comparison of OAS in esterolytic antibodies with natural enzymes is based on a number of antibodies in which OAS was shown to be the only source of the catalytic activity (group I antibodies). This group includes antibodies D2.3, D2.4 and D2.5 [10, 24–26], 48G7 [27, 28], 29G11 [29, 30], and 6D9 [31–33] (Group I; Fig. 2), which show most if not all of the following characteristics: i) The observed rate acceleration correlates well with the differential affinity toward the TSA vs the substrate: K TSA /K S ≥ kcat /kuncat (Table 1) ii) A linear phase is observed in the pH-rate profile iii) Crystal structure indicating active-site residues that comprise the oxyanion hole iv) Mutation of OAS residues eliminates catalysis (Table 2).

Other antibodies (Group II; Fig. 2) for which structural data is available, CNJ206 [14, 34], 17E8 [29, 30, 35], and 7C8 [11] were all raised against similar or identical TSAs to those of Group I, and exhibit K TSA /K S ≤ kcat /kuncat . This suggests that forces additional to or other than OAS take part (Sections 2.2 and 2.4). Nonetheless, their catalytic efficiencies may be taken as an upper limit to the contribution of OAS to

5


Fig. 2 The “oxyanion hole” of esterolytic antibodies. Oxyanion-stabilizing residues are depicted from the TSA complexes of antibodies CNJ206 (1KNO in PDB), 17E8 (1EAP), D2.3 (1YEC), 48G7 (1AJ7), 6D9

(1HYX), and 7C8 (1CT8), viewing from the oxyanion representing the incoming hydroxyl nucleophile. CPK color codes: gray: carbon, red: oxygen, blue: nitrogen, orange: phosphate, and green: chlorine.

their activity. Antibody 43C9, where an endogenous nucleophile was shown to be involved [6], is not considered, as the two mechanisms that affect its catalysis (OAS and nucleophilic catalysis) are difficult to separate (Section 2.2). Detailed comparisons of the hydrolytic antibodies’ crystal structures in the presence of the respective TSAs are available in several reviews [36–37] . A common motif (“canonical binding array”) of interactions with the TSA oxyanionic core was

2 Chapter Overview Table 1 Oxyanion stabilization by hydrolytic antibodies

kcat /kuncat G‡ cat-uncat G◦ TSA-S Catalytic Antibody × 103 Kcal/mol Kcal/mol OAH residuesa)

Secondary OAH residuesb)

Group I D2.3 D2.4 D2.5 48G7 29G11 6D9

110 36 1.9 16 2.2 0.9

7.0 6.3 4.5 5.8 4.6 4.1

7.0 6.3 4.3 6.3 6.0 4.1

Tyr100dH, Asn34L Tyr100eH, Asn34L Tyr100eH, Ser34L-H2 O His35H, (Tyr96H) Lys93H, (Tyr96H) His27d

Trp95H, Tyr96L-H2 O Trp95H, Tyr96L-H2 O Trp95H, Tyr96L-H2 O Arg96L, Tyr33H His35H, Tyr96L –

a) Residues in hydrogen-bond distance from the phosphonate’s oxyanion representing the ester substrate carbonyl (Fig. 1a), assigned by comparison of the antibodies’ phosphonate- and substrate analog-bound structures (D2.3, D2.4 and D2.5) [128], mutagenesis analysis (see Table 2), pH-dependency analysis (D-antibodies [10], CNJ206 [7, 129]C8 [11]), and docking (48G7, 29G11, and 17E8 [39]). b) Residues in hydrogen-bond distance from the phosphonate’s oxyanion representing the incoming hydroxide nucleophile (Fig. 1a) (see text for assignment).

revealed, identifying the residues contributing to the binding of the two oxyanions (Table 1, Fig. 2). Convergent evolution is evident by the sequence similarities as well as structural homology. Antibodies CNJ206 and 17E8 exhibit only 41–52% differences in the primary sequence of the heavy- and light-variable regions, yet share 8 out of 10 contact residues with the hapten. Antibody 48G7 shares the same light chain with CNJ206 and has 7 of 10 of the heavy chain’s CDR residues in common with CNJ206 (for the detailed analysis see [36–38]). Another example of convergence is seen in the structures of D2.3 and D2.4, sharing the same germline genes with 16 positions difference in their variable regions. In particular, the CDR3 H loop of D2.4 differs in 4 positions and includes an amino acid insertion compared to D2.3, as a result of different D-J segments’ junction. Consequently, their CDR3 H loops adopt Table 2 Oxyanion hole mutants of hydrolytic antibodies

Antibody

OAH residue mutant

k cat /kuncat wt = 1.0

D2.3/4b)

AsnL34 → Gly TyrH100d → Phe TyrH100d → Gly TyrH100d → Lys TyrH100d → Ser His35H → Glu His35H → Gln His27d → Ala

1.1 n.d. n.d. n.d. 0.08 0.04 0.6 n.d.

48G7a) 6D9 a) b)

Mutants expressed as recombinant Fab fragments [28]. Mutants expressed as scFv chimera of D2.4 VL /D2.3 VH [10]. n.d. – catalytic activity not detected.

7


distinctly different structures yet place the critical OAH residue Tyr100d (D2.3) and Tyr100e (D2.4) at the exact same position with respect to the TSA’s oxyanion [26]. The TSAs of Group I antibodies share two epitopes: an aryl or benzyl leaving group, and a phosphonate monoester (Fig. 2). While aromatic moieties are well known as immunogenic epitopes (hence their common use in TSA haptens), Tantillo and Houk suggested that the determining epitope may be the bidentate “transition state epitope” oxyanionic moiety. This is supported by the identification of similar motif in crystal structures of arsonates, sulfonates and DNA binding antibodies. Interestingly, the origin of many of the residues in contact with the TSA phosphonates can be tracked down to the germline genes (e.g. TyrH33 of 48G7, HisH35 of CNJ206 and 48G7, His27d of 6D9). Exceptions do occur though, as in the structures of 6D9 and 7C8 raised against a secondary p-nitrophenyl phosphonate. In contrast to other Group I TSAs, in antibodies 6D9 and 7C8, the linker via which the TSA hapten is linked to a carrier protein, is located on the leaving group and not on the phosphonate (Fig. 2). The active site of 6D9 and 7C8 is shallower, the phosphonate is less buried than in other Group I antibodies, and the active-site residues in contact with the TSA are fewer and different [11]. Despite the evident convergence between Group I’s active sites, the immune system provided some variations within the interactions forming the oxyanion hole (Table 1). The distinction between the position of the oxyanion hole and that of the entering hydroxy nucleophile was determined by the respective structure in the presence of substrate analogs (as in the D-antibodies), by pH profiles pointing to key hydrogen-bond donor residues (D-Abs, CNJ206), and by mutagenesis (D-Abs, 48G7, 17E8), and was complemented by docking in silico (Table 1, Fig. 2) [39]. 2.1.3 Antibody Oxyanion Holes The D-antibodies (D2.3 and D2.4), the most efficient antibodies of Group I, are taken here as a model for examining the contribution of active-site residues acting OAH. Two such residues (Tyr100dH and Asn34L) were suggested to form the crystal structures of the D-antibodies in complex with the TSA and a substrate analog (Fig. 2). In the crystal structure of D2.3 with an amide substrate analog (1YEF in PDB), 3.8 Å is the distance between the aspargine’s δ-amide proton donor and the scissile bond carbonyl-oxygen (compared to the 2.6 Å distance from the oxygen of TyrH100d), suggesting that Asn34L interaction may be specific to the transition state. In addition, D2.5, lacking Asn34L, has 20–50 fold lower catalytic efficiency than D2.3 and D2.4 (Table 1). However, mutating Asn34L to Glycine did not affect the rate of catalysis (Table 2), leaving Tyr100d the primary functional OAH residue. This was further supported by several Tyr100d mutants (Table 2) and pH profiles of binding and catalysis [10]. The pH-binding profiles revealed a 1.6–1.9 units difference in pK a between the substrate-antibody and the TSA-antibody complexes. Thus, the negative charge marking the difference between the substrate and the TSA (and hence the actual TSs of the reaction) dramatically strengthens the tyrosyloxyanion H-bond, thus rendering it the key residue in the D-Abs oxyanion hole [10]. The specific free energies, 6.3 and 7 Kcal/mol for the complexation of D2.4 and D2.3

2 Chapter Overview

with the TSA G◦ TSA –G◦ s ) are identical to the reduction in the activation energy barrier derived for the D-Abs from their rate acceleration (GC‡ cat –G‡ uncat ) (Table 1). This tight correlation suggests that the D-Abs active sites convert binding energy into catalysis with close to 100% efficiency, corresponding to the contribution of the tyrosyl OAH. Judging from Tables 1 and 2, tyrosine is the most efficient single residue for OAS. Further, a hierarchy may be derived where Tyr (contributing up to 7.0 Kcal/mol) > His (4.1 Kcal/mol, assumed from 6D9 single His OAH and48G7 His35HGlu mutant) ≥ Lys (∼3 Kcal/mol, 29G11 Lys-main chain amide OAH combination is weaker than 48G7) > backbone amide (2.2 Kcal/mol, judged by comparison of CNJ206 to wt48G7; 48G7 HisH35Glu mutant’s activity, which is derived from Tyr97H backbone amide, is approximately half that of CNJ206 with two backbone amides). Though highly speculative, this order of activities may be justified by the oxyanion pK a s found for serine hydrolase substrates, ranging between 7 and 10 [40, 41]. It is expected that hydrogen bonds between donors and acceptors with matching pK a would form the strongest hydrogen bonds [42]. Tyrosine’s hydroxyl (pK a ≈10) may afford the closest match to ester’s TS pK a . It is expected that this pK a would be considerably lower than the pK a of the TSs, leading, for example, to amide hydrolysis. The former is often stabilized in nature by OAH comprised of amide bonds that have a pK a much higher than the hydroxyl of tyrosine (see below) [5, 43]. Matching pK a values may also explain the significantly stronger interaction of the D-Abs’ tyrosine (∼7.0 Kcal/mol) compared to the average binding energy of a charged group via a single hydrogen bond (3.5–4.5 Kcal/mol) [44]. 2.1.4 Oxyanion holes–Antibodies vs. Enzymes The versatility of OAS solutions adopted by the immune system is comparable to, or perhaps even higher than, that observed in natural hydrolytic enzymes. Excluding metal hydrolases, which are not easily mimicked by antibodies (Section 2.5), the overall majority of enzymes use a combination of two (e.g., chymotrypsin, trypsin) or three (e.g., AChE, Fig. 3) backbone amides to form their OAHs. Their contribution to the catalytic activity cannot be addressed directly by mutation analysis or separated according to the individual contribution of each backbone NH. OAS could only be assessed by attributing the residual activity of the triad mutants to OAS [e.g., trypsin [45], acetylcholinesterase (AChE) [46]; Table 3]. A minority of enzymes were found to utilize a combination of a backbone NH groups as H-donor together with a side-chain NH group [e.g., Asn in subtilisin (Fig. 3), Gln, or Ser (Table 3)]. Interestingly, the recently characterized prolyl oligopeptidase-like family [43, 47, 48] and the structurally similar cocE esterase (Fig. 3) [49] utilize a tyrosine side chain (Table 3) together with the backbone NH of the seryl nucleophile. Indeed, the efficient OAS by catalytic antibodies such as the D-antibodies may indicate that tyrosine could be as efficient as or even more efficient than NH groups. The contribution of the OAH side chain to the overall catalytic rates of these enzymes was assessed by mutation of the putative side chain (Table 3). Such studies are complicated by mutation-induced structural changes of the active site as clearly

9


Fig. 3 The ‘oxyanion hole’ of hydrolytic enzymes. Oxyanion-stabilizing residues are depicted from the following inhibitor-enzyme crystal structures: acetylcholinesterase (AChE) with m(N,N,N-trimethylammonio)-

2,2,2-trifluoroacetophenone (TMTFA) (1AMN in PDF), cocaine esterase I (cocE) with phenyl boronic acid (1JU3), and subtilisin with D-p-chlorophenyl-1-acetamidoboronic acid (1AVT).

seen in the cocE Tyr44Phe mutant’s structure. As a result of a loss of H-bonding to the mutated tyrosine, three Trp residues change their position, and the Phe ring is tilted by 20◦ compared to the wild-type tyrosyl ring [5]. In addition, given the average bacterial translation error rate of 5 × 10−4 per amino acid [50], wild-type revertants may significantly affect these single-mutants’ analyses. Histidine-containing OAH (as in antibody 6D9; Fig. 2) have not yet been found in nature. The lower efficiency of His in OAS is seen in hydrolytic antibodies (see above). The latter may be due to the relatively low pK a of the His side chain or the more demanding desolvation of the charged His residue compared to the more hydrophobic tyrosine, aspargine, or serine. Interestingly, several structural studies (NMR of trypsin-leupeptin complex [51]; X-ray of proteaseA-chymostatin [52]) identified the oxygen carbonyl of the aldehyde inhibitors pointing toward the active-site’s His, suggesting that the latter has comparable affinity to the oxyanion with respect to the original OAH two backbone NH groups. This observation is in agreement with the OAS efficiency scale postulated from the catalytic antibodies’ data. The estimated values for the individual residue’s contribution to OAS are generally lower for enzymes (Table 3) than for the D-antibodies and most other esterolytic antibodies (Tables 1 and 2). It may be that the enzymes’ mutagenic studies underestimate the OAH side chain’s contribution, as the remaining H-bond by the backbone NH at the OAH is strengthened in the absence of its side-chain partner, hence compensating for its absence. Such compensation may explain the mild effect of 48G7 His35HGlu mutation (40-fold reduction in catalytic rate), compared to 6D9 His27dAla mutation. The latter lacks a backbone NH, which is present in 48G7’s OAH (Table 2). The only natural OAH suggested to have a similar contribution to catalysis as that of the antibodies described above is of AChE, which has three hydrogen donors (Fig. 3) (compared to 1–2 in most antibodies). Did enzyme evolution miss out on an efficient OAS strategy that is easily implemented by antibodies? Few plausible answers come to mind. The OAH may not have evolved independently in nature, but paralleled (or followed) the appearance of a catalytic triad that takes care of nucleophilic and proton-transfer catalysis (Fig. 1b). Once the latter was established,

2 Chapter Overview Table 3 Contribution of oxyanion stabilization to enzymes, mutagenesis studies

Enzyme

OAH

Subtilisin Asn155, Thr220, (Ser221)

CarboxypeptidaseA Trypsin

Gln19

(Ser42) Prolyloligopeptidase Oligo-peptidaseB CocE

AChE

Arg127 (Gly193) (Ser195) (Ser195) Gln19Ala (Cys25) Ser42 Ser42Ala (Gln121) Tyr473 (Asn555) Tyr452 (?)e) Tyr44 (Tyr114) (Gly118) (Gly119) (Ala201)

OASa) ‡ G cat−uncat Kcal/mol

Ref.

Asn155Gly Asn155Ala Thr220Ala Asn155Ala Thr220Ala Ser221Alab) Arg127Alac) Ser195Ala

3 4.2 1.8 6.0

[4] [130] [130] [130]

4.8 6.0 4.2

[4] [131] [45]

3

[132]

3.6

[133]

Tyr473Phe

3.7

[43, 47]

Tyr452Phe Tyr44Phe

3.6 > 4.4d)

[48] [5]

-f)

5–7

[46]

Mutation

a) Contribution to oxyanion stabilization derived from OAH-residue mutation’s effect on catalytic rates or: b) Contribution to oxyanion stabilization derived from residual activity after mutation of the enzyme’s endogenous nucleophile. c) Though Thr220 is only within 4.0 Å of the oxyanion, dynamic simulations suggest this residue’s participation in subtilisin’s OAS. d) Limited by detection of mutant’s activity. Significant structural perturbation of mutant’s active site e) As crystal structure is unavailable, the identity of an assumed second OAH residue (analogous to the homologous Asn555 of prolyloligopeptidase (see separate entry) is unknown. f) Estimated by the authors based on AChE-TSA crystal structure and accumulative data.

together with other mechanisms such as substrate destabilization [46] [52], the hydrolytic activity may have been sufficiently high (diffusion-limited at times [46]) to make further improvements in OAS unnecessary. Alternatively, it was recently suggested that the charged imidazole of the subtilisin triad’s histidine has a major role of lowering the pK a of the TS oxyanion, thereby diminishing the OAH binding energy [40]. In such a scenario, the major role of the OAH is substrate alignment rather than TS stabilization as was initially suggested [53]. It remains to be seen whether these results would extend to other natural enzymes.

11


In conclusion, hydrolytic antibodies validate the fundamental role of OAS in enzyme catalysis. They do so with similar or perhaps better efficiency than enzymes. OAS measured for antibodies could be thus used as an upper limit to the contribution of OAS in natural hydrolytic enzymes, providing a reference value for the contribution of their other mechanistic features (see below).

2.1.5 Antibody Affinity and Rate Acceleration Limitation Despite the limitations of TSA design, our analysis suggests that many antibodies do convert TSA binding energies to catalysis with nearly 100% efficiency (Table 1). This may suggest a facile route to increased catalytic efficiency via antibodies with higher affinity. However, it appears that antibodies with affinities higher than 1 nM may not be easily obtained. Data collected on the affinity of a large number of antibodies [54] suggest an antibody of 0.1–1 nM [55]. This is derived from the upper range for the association rate (< 5 × 10 M−1 s−1 ) and a dissociation rate (koff ) that is sufficient to trigger B-cell activation (10−3 –10−4 s−1 ). These studies suggest (see [54, 55] and references therein) that 0.1–1nM affinities are sufficient for mounting an efficient immunological response, arguing that the immune system is unlikely to produce higher affinity antibodies. By inference, with a low-average K M (≈KS ) in the mM range [56–58] a “catalytic ceiling” of K TS /KS = kcat /kuncat = 106 − 107 may be derived for catalytic antibodies raised against TSA haptens even if these happen to perfectly mimic the reaction TSs. Thus, the turnover number (kcat ) of antibody-mediated hydrolysis of esters commonly targeted by catalytic antibodies (kuncat 10−6 –10−5 s−1 ) is expected to be well under 10s−1 and the specificity constant (kcat /K M ) ≤ 104 M−1 s−1 . Under the immunological constraints limiting K TSA , a lower K M (or higher substrate affinity) would result in a lower rate acceleration, without lowering the specificity constant. Indeed, a survey of the published catalytic antibodies conforms with the above theoretical limitation (Fig. 4). Moreover, in some instances, not all of the TSA binding energy is directed towards TS stabilization, but rather to non-contributing epitopes of the TSA hapten. For example, antibody 48G7 was shown to bind its TSA with 30 000-fold higher affinity compared to its germline counterpart, resulting in only ∼80-fold improvement of kcat /K M [16]. Although independent values of kcat and K M were not reported, it is likely that a significant part of the increase in binding energy upon maturation was directed to both the substrate and the TSA, and therefore led to no increase in rate acceleration. Similarly, attempts to improve the rate of esterolytic antibodies by directed evolution aimed toward higher TSA affinities have shown that major increases in affinity may result in minor rate improvements [59, 60]. Improving the rates of esterolytic antibodies is not a purely academic challenge. The limitations in rate hinder at least two of the most promising biomedical applications of hydrolytic antibodies – cocaine detoxification [61] and pro-drug therapy [62]. Catalytic forces other than OAS may help in closing the gap of ∼100-fold reactivity [5, 62] required for their successful implementation. These are discussed below.

2 Chapter Overview

Fig. 4 Rate enhancements exhibited by catalytic antibodies: a compilation of rate accelerations exhibited by all catalytic antibodies [139]. The theoretical threshold is set by the affinity ceiling of antibodies as described

in the text. The only antibody to surpass the theoretical threshold dictated by the immune system is 38C2 [140], achieved by reactive immunization (see text for details).

2.1.6 Nucleophilic Catalysis The vast majority of esterolytic antibodies make use of a solution (exogenous) hydroxide ion to generate the tetrahedral, oxyanionic intermediate that leads to ester hydrolysis. This is not particularly effective, as at or close to neutral pH, hydroxide ion concentrations are very low. The enzymatic solutions to this problem are basically two: (i) nucleophilic catalysis by an active-site residue (an endogenous nucleophile) via an acyl-enzyme intermediate; (ii) a general-base residue and a bound water molecule leading to the in situ generation of an appropriately positioned hydroxide ion. Inspired by natural hydrolytic enzymes that utilize a covalent catalytic mechanism (Fig. 1b), and with the goal of catalyzing the more demanding reaction of amide hydrolysis, several attempts were made to introduce an endogenous nucleophile into antibodies’ combining sites. But, in fact, the more effective cases of endogenous nucleophilic catalysis seem to be in antibodies in which the endogenous nucleophile was incorporated haphazardly. Antibodies in which endogenous nucleophilic catalysis plays a role are discussed in Section 2.2 below. In the same vein, attempts were made to generate antibodies in which the nucleophilic hydroxide is generated in situ by general-base catalysis (discussed in Section 2.4 below). Alternatively, the efficiency of the exogenous nucleophilic attack can be enhanced by applying nucleophiles more potent than hydroxide (“chemical rescue”; Section 2.3). 2.2 Endogenous Nucleophiles in Hydrolytic Antibodies 2.2.1 Reactive immunization (RI) Several attempts were made to introduce endogenous nucleophiles into antibody combining sites, either by chemical modification [9, 63] or by immunization. The

13


Scheme 1

latter was aimed at “reactive immunization”, whereby the labile phosphonate diester 1 (Scheme 1), a known covalent inhibitor of hydrolases, was used as a hapten [7, 64]. During the immunization process, an RI hapten should covalently trap antibodies with a nucleophilic residue, thereby forming a highly stable complex that is selectable by the immune system. There is, however, a major caveat to this methodology. Hydrolysis of the activated p-methylsulfonate ester substrate via nucleophilic catalysis will result in an that is significantly more stable than the original substrate. For effective catalysis to occur, an would be necessary to activate the hydrolysis of the acyl-enzyme intermediate (Fig. 1b). Such a residue is counter-selected in this system, as it would also hydrolyze the covalent phosphonyl hapten-antibody bond, thus eliminating the selectable advantage of the reactive immunization. Moreover, hapten 1 is labile (estimated half-life mice’s serum is 24 h), resulting in a mixture of haptens, 1 and 2, (the bi- and mono-phosphonate esters, respectively). Unexpectedly, the affinity to the “baiting” hapten 1 is rather low (in the micromolar range) compared to the high nanomolar affinity of antibodies raised against hapten 2. The kinetic parameters of anti-1 antibodies, suggest that OAS (represented by hapten 2) does not account for the RI-derived antibodies’ catalytic efficiencies (e.g., for antibody 15G12, K TSA(2) /K M = 2; kcat /kuncat = 6.6 × 105 ). However, there is no other evidence to support nucleophilic catalysis in the hydrolytic antibodies raised against RI hapten 1 (e.g., burst kinetics, trapping of the covalent intermediate, non-competitive inhibition by hapten 1). Some of the RI antibodies were reportedto catalyze the dephosphonylation of their cognate hapten 1, albeit with a low turnover (< 3), because of product inhibition [7] (such single turnover rate enhancement by

2 Chapter Overview

ester-hydrolytic antibodies, raised against the ground state hapten, was reported as early as 1980 [65]). Schowen has recently suggested that TS flexibility, where the classical bipyramidal dephosphonylation TS may adopt a more tetrahedral-like conformation, renders it available for stabilization by the anti-phosphonate antibodies [66]. It should be mentioned that, although RI achieved a limited success with ester hydrolysis, it has proven highly effective for reactions such as aldol condensation, retro-aldol and Michael addition reactions that proceed via a covalent Schiff-base enzyme-substrate intermediate [67, 68]. 2.2.2 Serendipity and 43C9 Antibody It has been suggested that several hydrolytic antibodies raised against phosphonate TSAs include an endogenous nucleophile as part of their mechanism of action. Unfortunately, in most cases, only partial evidence is available. In general, proving a nucleophilic mechanism is not a trivial matter, and several independent pieces of evidence need to provided, including the direct identification of an acyl-enzyme intermediate, before a definite statement can be made [2]. A “ping pong” mechanism by antibody PCP21H3 (kcat /kuncat not determined) in a transesterification reaction of a vinyl ester has been suggested. The antibody exhibits burst kinetics for the hydrolysis of a related p-nitrophenyl ester [17]. Product inhibition, common to many catalytic antibodies, was detected as well, yet this does not outweigh the evidence for the covalent mechanism. A similar burst in product formation interpolated to a 1:1 molar ratio of product to antibody was reported for antibody 6F11 (kcat /kuncat = 3.4 × 103 ) [69]. Chemical modification oftyrosyl residues resulted incomplete activity loss. Antibody 20G9 (kcat /kuncat = 500) exhibits “burst kinetics”, and this led to the suggestion of nucleophilic catalysis [70–72]. However, the kinetic analysis is complicated by strong product inhibition, and no evidence apart from the “burst” is available. Incubation of the antibody with N-cbz-glycine -O-phenyl ester resulted in multiple acylation of the antibody and a significant decrease in its activity, yet acylation of an active-site residue such as lysine that is not directly involved in catalysis may account for this result [73]. In addition, the reported acid-limb pH profile fitting a tyrosyl pK a (9.6), as well as the loss of activity when chemically modifying tyrosyl residues with tetranitromethane (TNM), could well be explained by this residue’s participation in OAS [10, 73, 74]. Antibody 7C8 (kcat /kuncat = 3.4 × 103 ) also exhibits a pH profile with an acidlimb with a pK a ≈ 9.0. The rate is also insensitive to the addition of a strong nucle-ophile (hydroxylamine; see also Section 2.3 below). These led to the suggestion of nucleophilic catalysis by an active-site tyrosyl hydroxyl [11]. However, D2.3 and D2.4 antibodies also manifest the above characteristics [10], yet their active site tyrosine (H100d) was shown to serve as an OAH rather than a nucleophile. Similarly, 7C8’s crystal structure reveals a sole residue that may serve as an OAH, Tyr95H. Another clear demonstration of the difficulties associated with proving a nucleophilic mechanism is antibody 17E8. Its crystal structure and hydroxylamine partitioning experiments suggested the existence of a Ser-His dyad, reminiscent of natural serine hydrolases [13, 35]. However, direct evidence of an acyl-enzyme

15


intermediate was not provided, and the proposed mechanism involved the disruption of a hydrogen bond between His35H and TrpH47 that is conserved in all known antibody structures [36]. Indeed, nucleophilic catalysis was ruled out when a variant of 17E8 in which the Ser, assumed to act as nucleophile, was replaced by Gly exhibited a similar rate enhancement [30]. Most significantly, analysis of the antibodies described above suggests that even if an is indeed part of their mechanism, the resulting rate enhancements are 2–3 orders of magnitude lower than is achieved with antibodies raised against the same haptens and that act by OAS only (see above). Antibody 43C9, raised against the phosphonoamidate 3 (Scheme 1) [75, 76] stands out from all the above examples, not only because of its high reactivity (it catalyzes the hydrolysis of both a p-nitrophenyl ester and a p-nitroanilide and, although its rate enhancement is within the range of other known esterolytic antibodies, its specificity constant is so far unrivalled (kcat /KM = 2.8 × 107 M−1 s−1 )), but also because of the unambiguous assignment of a nucleophilic mechanism. In the case of antibody 43C9, there is ample evidence to prove a nucleophilic mechanism. The pH-rate profile suggests a change in the rate-limiting step: while at pH > 9 product release is limiting, at more acidic pH an acylation step determines the rate [77]. This was demonstrated by pre-steady-state kinetics, where a burst of antibody-equivalent product release kinetics was observed [77]. Finally, the effect of p-substituents of the phenolic leaving group on the rate of catalysis [78] and the identification by electrospray mass spectroscopy of an acyl-enzyme intermediate (with HisL91 as the acylated residue) unambiguously support the proposed mechanism [6]. A detailed catalytic mechanism based on 43C9 crystal structure was put forward where a nucleophilic attack, carried out by HisL91, is accompanied by OAS via the side chains of HisH35 and ArgL96 (Fig. 5) [79]. The serendipitous selection of a His side chain as the nucleophilic catalyst of 43C9 is quite interesting. Not only is the His’s imidazole side chain a better nucleophile toward activated esters than the seryl alcohol moiety, but also its pK a is optimal for catalysis and theresulting acyl-enzyme intermediate is much more labile than an ester intermediate. Thus, His makes an ideal nucleophilic catalyst. Nevertheless, His was never recorded as a nucleophile in natural hydrolytic enzymes. One possible explanation is that with non-activated ester substrates, imidazole prefers acting as a general base rather than a nucleophile, hinting at nature’s preference for other nucleophiles as alcohols and thiols [80]. The OAH of 43C9 is equivalent to other known antibodies (Table 1, Fig. 2), suggesting a comparable contribution to its rate enhancement. Thus, the nucleophilic mechanism does not seem to contribute much to the rate acceleration of ester hydrolysis. This can be explained by the absence of a general base residue (Fig. 1b) that would catalyze the rate-limiting de-acylation step, thus replacing an activated p-nitrophenyl ester with a less active acyl-imidazolium moiety. However, nucleophilic catalysis may contribute significantly to anilide hydrolysis, where formation of the intermediate, rather than its breakdown, is rate-determining. The nucleophilic mechanism may also release 43C9 from having a kcat /K M limited by low affinity to the ground state substrate, as prescribed for efficient catalytic antibodies utilizing solely the energy gained

2 Chapter Overview

Fig. 5 The mechanism of catalysis by antibody 43C9 [79]. (i) A nucleophilic attack by an endogenous nucleophile – HisL91 side chain. The nucleophilic imidazole is maintained in the correct tautomer by a network of H-bonds with TyrL36 and TyrH95 side chains. (ii) The resulting oxyanionic intermediate is stabilized by H-bonding to His35H and ArgL96. Bond rearrangement, coupled with a proton transfer cascade, re-

sults in the acyl intermediate and the release of the aniline leaving group. The aniline release is mediated by protonation by a water molecule (not shown). (iii) An exogenous nucleophilic attack by a hydroxide ion on the acyl-enzyme intermediate. (iv) Bond rearrangement results in the release of the carboxylate product and the free antibody (see hapten 3 structure for R and R ).

from preferential TS vis-a-vis substrate binding, yielding ca. 2 orders of magnitude higher specificity constant, in parity with natural enzymes’ values. Quantifying the individual contribution of the nucleophilic attack is further complicated by the fact that a HisL91Gln mutation of 43C9 results in total loss of hydrolytic activity, yet retains wild-type levels of affinity to its hapten and substrates [81]. Thus, in agreement with natural enzymes, even in a somewhat simplified model, different catalytic mechanisms seem to be coupled. Importantly, the nucleophilic mechanism endows 43C9 with the ability to hydrolyze the more stable p-nitroanilide substrate, unattainable by the antibodies using OAS as their only source of reactivity. Unlike natural amidases, 43C9 does not catalyze the hydrolysis of non-activated amides. The rate-limiting step in their hydrolysis involves the protonation of the amine-leaving group via a general acid residue (as in natural amidases, Fig. 1b), not found in 43C9 active site.

17


Fig. 6 The tetrahedral intermediate formed by various exogenous nucle-ophiles.

2.3 Exogenous Nucleophiles and Chemical Rescue in Hydrolytic Antibodies

The nucleophilic attack of a solution’s (exogenous) hydroxide anion seems to constitute the rate-limiting step of all hydrolytic antibodies. In most cases, the hydroxide reacts directly with the ester substrate and, in rarer cases, with an acyl-antibody intermediate. This step therefore merits careful examination. One of the most useful analytical tools is the replacement of hydroxide by alternative nucleophiles. Negatively charged, tetrahedral intermediates, homologous to the hydroxyl-mediated catalytic mechanism (Fig. 1a) can be formed by the attack on the ester’s carbonyl by other nucleophiles (Fig. 6). Thus, it may be expected that small nucleophiles, stronger than the hydroxyl anion, may be active in the antibodies’ active sites, resulting in higher catalytic rates. A similar strategy was previously used to “chemically rescue” hydrolytic enzymes where the nucleophilic residue was mutated. The mutants’ catalytic activity can often be restored to rates only 100- to 300-fold lower than the intact enzyme [82–84]. One example of effective “chemical rescue” of hydrolytic antibodies is with the peroxide anion, a ∼200-fold stronger nucleophile than hydroxide, that was found to enhance the turnover rates of D2.3 and D2.4 antibodies to a similar extent (Fig. 7a, Table 4) [10]. The concentration-dependent rate acceleration exhibits a rate ceiling at a kcat that is 500- to 2000-fold higher than that of the antibodies’ hydroxidemediated catalysis. The rate ceiling is due to a change in the rate-limiting step observed at high peroxide concentrations – a conformational change becomes rate limiting instead of the nucleophilic attack. The observed rate enhancement of the D-antibodies in the presence of peroxide compares favorably with the estimated contribution of the deacylation rate-limiting step of various natural hydrolases (e.g., ca. 200-fold for cocaine esterase cocE [10, 49]. The D-Abs’ maximal rate of 6 s−1 is also comparable to many natural esterolytic enzymes, such as cocE [5], thioesterases [85], and carboxylesterases [86], and is within the lower range of typical rates of (1–200 s−1 ). This suggests that a potent exogenous nucleophile can match, in the framework of efficient OAS antibodies, the role of the enzyme’s general-base deacylation mechanism (e.g., the His-Asp dyad in Fig. 1b). The secondorder rate for peroxide anion reactivity with the D-antibodies was calculated to be 6 × 105 M−1 s−1 . This is significantly lower than the directly measured second-order rate for water-mediated deacylation of chymotrypsin (< 4M−1 s−1 ) and compares favorably with its deacylation by an exogenous amine nucleophile (tyrosinamide: 1.7 × 104 M−1 s−1 ), a direct result of the enzyme’s specificity for the leaving group

2 Chapter Overview

Fig. 7 The effect of exogenous nucleophiles on the rate of antibody-mediated ester hydrolysis. Rate acceleration factors are the ratios of net initial rates observed in the presence of the nucleophile at a given con-

centration and in its absence. (a) Effect of peroxide anion on the hydrolysis of pnitrophenyl ester by D2.3 and D2.4 [10]. (b) Effect of hydroxylamine on chloramphenicol ester hydroysis (data derived from [11]).

Table 4 Nucleophile effect on D-Abs’ catalysis

Nucleophilea) (× 106 M−1 )b) − OH −O

2H

NH2 NH2 NH2 OH N3 −

pK a) (× 106 M−1 )b)

Buffer (× 106 M−1 )b)

Antibody D2.3

Antibody D2.4

15.4 11.6 8.1 6.0 4.0

13 2800 0.24 0.45 1 × 10−3

12 1270 0 0 n.d.

14 3200 0.008 0.006 0.01 × 10−3

a) A linear correlation links the nucleophilicity of most nucleophiles to their pK a s. However, the reactivity of the α-nucleophiles cited above (i.e. excluding OH− ) is far higher than predicted from their pK a (e.g. 103 -fold for hydroxylamine). b) Results are given in “molar reactivity” – the rate acceleration factor at 1 M nucleophile as depicted in [10].

19


Scheme 2

[87]. Importantly though, the D-Abs’ rate accelerations (kcat /kuncat ) are relatively unchanged when peroxide replaces hydroxide, suggesting that the D-Abs are equally complementary to the respective TSs of both hydroxide and peroxide. The D-Abs exhibit absolute preference toward small, negatively charged nucleophiles (i.e., hydroxylate and peroxylate) for the displacement of the ester’s alcohol moiety, whilst other nucleophiles (e.g., hydroxylamine) do not affect the rate (Table 4). We believe that this specificity is a result of (i) the selection, by the negatively charged phosphonate anion, of antibodies with a cluster of positively charged residues, creating a channel to the antibody active site, and (ii) the snug fit of the antibody to its hapten and therefore to the TS, preventing the entry of bulky nucleophiles such as azide [10]. These results suggest that it may be possible to optimize the electrostatic properties of the channel through which the nucleophile approaches the active site, perhaps by engineering positively charged residues in and around this channel without interfering with the properties of the active-site core [88], thereby increasing the rate of catalysis. The rate of hydrolysis by antibody 6D9 (raised against hapten 4, Scheme 2) is also increased by the presence of the α-nucleophile hydroxylamine. Unlike the D-abs, both the rate (kcat , or kcat /K M ) and the rate acceleration (kcat /kuncat ) increase, albeit to a minor extent (up to ∼2 fold at 100 mM hydroxylamine) (Fig. 7b) [11]. The source of recognition of hydroxylamine by 6D9 was not addressed, yet the Lys32L residue [11] is located in a position suitable for general-base activation of the nucleophile. Alternatively, it may act as an H-bond donor to orient the nucleophile toward attacking the chloramphenicol ester’s carbonyl. It would thus be interesting to study the effect of Lys32L mutants on the hydroxylamine-mediated reaction, as well as attempting the hydrolysis of the more demanding amide hydrolysis in this system. Another reported case of “chemical rescue” concerns antibody 14–10. Using the heterologous immunization approach, where mice were challenged with both haptens 5 and 6 (Scheme 2), Masamune and coworkers attempted the generation of a p-nitroanilide hydrolyzing antibody that makes use of OAS, general-base catalysis

2 Chapter Overview

Scheme 3

(by “baiting” with a positively-charged amine hapten) and an exogenous phenol nucleophile. The best antibody achieved, 14–10, catalyzed both the lactonization of substrate 7 (Scheme 3) via an intramolecular phenol nucleophilic attack (kcat = 1.5 × 10−2 min−1 , K M = 1.27 mM [89]) and the intermolecular attack of added phenol nucleophile to substrate 8 to produce phenyl propionate, 9 (kcat = 1.3 × 10−4 min−1 , K M(2) = 0.37 mM, K M(phenol) = 0.14 mM [12]). In both cases the phenol does not seem to participate in the background buffercatalyzed reactions. Thus, the antibody catalyzes kinetically “disfavored” reactions, resulting in kcat /kuncat = 2 × 104 and 630 for the intramolecular and intermolecular reactions, respectively (the uncatalyzed rates serving as an upper limit of the background rates). Interestingly, 14–10 does not hydrolyze the ester product 9. This could be only partly explained by the lack of a negatively charged TSA to elicit an OAH, as similar, uncharged haptens (e.g., phosphono-amidates [62], and β-amino alcohols [90]) were shown to elicit efficient hydrolytic antibodies. Nonetheless, as the buffer-mediated hydrolysis rate of 9 proceeds twice as fast as the antibodymediated acyl transfer, the net reaction can be formally considered as the activated amide’s hydrolysis. Future studies may reveal whether the phenol nucleophilicity is indeed enhanced by a general-base residue, as implied by 14–10’s pH optimum (8.0 compared with the phenol’s solution pK a of 10). 2.4 General Acid/Base Mechanisms in Hydrolytic Antibodies

The most notable example of hydrolytic enzymes that make use of general acid and base catalysis is aspartic peptidases. This family of hydrolases utilizes the concerted action of two carboxylate residues (Fig. 1c). The first carboxylate serves as a general base to align and activate the water nucleophile, and subsequently as a general acid to protonate the leaving group. The second serves as the OAH and may be regarded as a general acid, donating a proton to the formed carboxylate. In order to mimic such mechanisms with catalytic antibodies, several “bait and switch”

21

22 Catalytic Antibodies as Mechanistic and Structural Models of Hydrolytic Enzymes Table 5 Kinetic parameters of “bait and switch”-raised antibodies

Antibody

Hapten

kcat a) (min−1 ) pH = 7.8

27A6 30C8 2D5

10 11 12

0.004 0.005 3.5

K M (mM)

kcat /KM M−1 s−1 )

kcat /kuncat b)

Ref.

0.25 1.1 0.5

0.7 0.07 120

1.4 × 105 1.2 × 105 1.3 × 105

[134] [90] [69]

a)

Values for antibodies 30C8 and 27A6 were derived from pH-rate profiles. Although the same substrate and reaction are involved, a ∼1000-fold difference exists between the uncatalyzed rates reported of one group [134] and those of the other [69]. b)

haptens were devised with the aim of “baiting” oppositely charged residues in the antibody’s active site that may serve in a general acid/base mechanism. An example of this strategy (haptens 10–12) and the kinetic rates of the resulting antibodies is summarized in Table 5. Haptens 10–12 (Scheme 4) were designed to bait different types of residues in the elicited antibodies’ binding sites. The positively-charged hapten 11 was designed to bait a general-base residue (e.g., a negatively-charged carboxylate), whereas hapten 10 aims at baiting a general-acid residue that would facilitate the rate-determining protonation of the amine-leaving group of substrate 14. Nonetheless, the catalytic activity of the best antibodies raised against haptens 10 and 11 is remarkably similar (Table 5), suggesting that even if “baiting” did occur, it does not effect to a significant extent the antibodies’ catalytic efficiency. Moreover, whilst the antibodies raised against haptens 10–12 all catalyze the hydrolysis of the ester substrate, 13, with similar kinetic parameters, the amide substrate 14 is hydrolyzed by none

Scheme 4

2 Chapter Overview

of these antibodies, not even by those antibodies raised against hapten 12. The effectiveness of the “bait and switch” strategy in raising hydrolytic antibodies is further questioned by the fact that hapten 12 was designed to bait a general-base catalyst while maintaining the phosphonate tetrahedral negatively charged TSA core for the generation of OAH residues. On the other hand, in haptens 10 and 11, a planar, uncharged carbon represents the tetrahedral TS. The catalytic rate of the best anti-12 antibody, 2D5, is also in agreement with rates of the above-described OAS antibodies, suggesting that even if an active-site general base has been generated, its contribution to catalysis is not very significant. By comparing the catalytic rates of antibodies 30C6 and 27A6 to 2D5, one could conclude that the tetrahedral phosphonate is a better mimic of the TSs of ester hydrolysis than the β-alcohol moiety, or, in other words, the phosphonate TSA “baits” a ∼500-fold more efficient OAH. Other attempts to utilise the “bait and switch” strategy yielded similar rate enhancements to the examples described above, with kcat /kuncat values in the range 102 − 104 (see, for example [91–93]). Future studies of antibodies raised by the “bait and switch” strategy may shed more light on their mechanism and may allow an improvement in their rates. Notably, as is the case with reactive immunization, the “bait and switch” approach was applied to non-hydrolytic reactions with greater success. This is evident, for example, in the Glu residue acting as a general base in antibody 4B2’s mediated catalysis of an allylic rearrangement reaction (PDB code 1F3D [94]). In contrast, the crystal structure of antibody 1D4, raised against a charged β-amino phosphate, does not show the expected positioning of a catalyzing general-base residue in its active site [95]. Finally, it is encouraging to note that a structural model of the chemiluminescent anti-phosphonate 7F11 antibody (kcat /kuncat = 1.2 × 103 , kcat /KM = 5.9 × 105 M−1 s−1 [96]) indicates the presence of two aspartic acid residues in positions resembling the natural aspartyl protease. Based on this model, the authors suggested that the 7F11 mechanism is similar to the natural counterpart, but that its much inferior catalytic efficiency may be due to the imperfect positioning of the apartyl residues. Antibody 7F11 also suffers from severe product inhibition, enabling an estimated turnover of ca. 5 cycles [97]. Nevertheless, if its active-site structure does prove similar to its natural counterpart, it may serve as a starting point for the generation of more potent hydrolases. 2.5 Metal-Activated Catalytic Antibodies

Several studies have attempted to raise catalytic antibodies with metal ion-mediated hydrolysis, following the zinc-protease mechanism (Fig. 1d). Many enzyme models based on zinc have been created that exhibit high rate enhancements [98], suggesting that the generation of metallo-antibodies with enzymatic activities is a promising avenue. However, this approach has thus far been explored with limited success. Lerner and coworkers reckoned that the “bait and switch” hapten 11 may elicit a carboxylate residue in the antibodies’ active site that would serve as a metal-binding ligand. Furthermore, the pyridinium’s methyl group will induce a

23


Scheme 5

cavity for the metal ion. Indeed, in the presence of Zn+ , antibody 84A3 catalyzes with high specificity the hydrolysis of a pyridinium ester (kcat /kuncat = 1200 [99]), the structure of which is related to the immunizing hapten. No catalysis was observed in the absence of the metal ion or with other, structurally similar ester substrates. The antibody suffers from severe substrate inhibition, probably due to non-productive binding of the substrate, probably in a mode not involving a zinc ion, as judged by the affinity difference to the substrate in the presence or absence of Zn+ (3.5 µM and 840 µM, respectively). In an attempt to overcome this hurdle, the mercury(II)-phosphorodithioate unsaturated complex 15 (Scheme 5) was used to elicit Hg-dependent hydrolysis of ester 16 [100]. The best anti-15 antibody, 38G2, exhibited Michaelis-Menten kinetics (kcat /kuncat = 300, K M(Hg) = 87 µM and K M(16) = 345 µM), multiple turnovers and absolute dependency on Hg(II) for catalysis. While far from enzyme-like reactivity, these studies widen the spectra of possible mechanisms that could be tailored by antibody mimics. 2.6 Substrate Destabilization

Remote interactions, away from the ester or amide carbonyl reaction core, play an important role in many natural enzymes. Such interactions may contribute to catalysis via substrate destabilization (for a recent review see [101] and references therein). For example, Sussman and coworkers postulated that as much as 5 × 105 (corresponding to TS stabilization of 8 kcal/mol) of AchE’s spectacular rate enhancement (1013 ) is derived from orienting the substrate to a highly unfavorable conformation, perfectly aligning the substrate for the nucleophilic attack to take place. This is achieved by strong binding to the choline ammonium charged moiety [46]. Several examples of the identification and possible role of substrate destabilization by hydrolytic antibodies are given below. The docking of the activated anilide substrate in the crystal structure of 43C9 suggests that the substrate’s amide bond is twisted by 140◦ out of planarity, thus destabilizing the bond’s resonance and facilitating its hydrolysis [79]. Similarly, 6D9 antibody’s structure suggests that the antibody locks its ester substrate in the sin unfavorable conformation, resulting in substrate destabilization. In both the above examples, a quantitative assessment of the contribution of substrate destabilization to the rate acceleration is not available. A 5-fold increase in the catalytic rate accompanied by a smaller effect of higher K M (2-fold) was seen between the catalysis of the glutaryl-glycine nitrophenylester

3 The Role of Conformational Changes in Catalytic Antibodies

substrate and a caproyl-elongated substrate by antibody D2.3, suggesting a remote effect of the substrate’s linker moiety. Crystal structure analysis revealed different interactions with the two linkers that may account for the kinetic differences [102]. When the linker terminal glycine was replaced by either the D- or L-Ala enantiomers, a ∼20-fold difference in D2.3’s specificity constant (kcat /K M ) was observed in favor of the l-Ala enantiomer [103] (for the enantioselectivity potential of the esterolytic active sites of various antibodies see [39]). Another example is set by antibody 17E8 catalyzing the hydrolysis of n-formylnorleucine (nLeu) phenyl ester. The specificity factors (kcat /K M ) observed with a series of substrates where the nLeu side chain was replaced by different linear and branched alkyl side chains extend over 2 orders of magnitude. But rate enhancements higher than those with the original substrate (namely with a Leu side chain) were not obtained. These results suggested that 17E8 interactions with the side chain are used to stabilise both the TS and the ground state. However, the replacement of nLeu side chain with a more hydrophilic ethyl-cysteyl side chain resulted in a 20-fold increase in kcat (43 vs 2.1 s−1 ) accompanied by a similar decrease in ground state binding (K M = 4 vs 0.18 mM with nLeu). This rendered the new substrate as efficient as the original substrate (kcat /K M ≈ 1.2 × 104 M−1 s−1 ) and suggested that substrate destabilization may play a key role in the mechanism of 17E8 with the ethyl-cysteyl substrate [104].

3 The Role of Conformational Changes in Catalytic Antibodies

Structural dynamics are part of the nature of proteins, though their identification on the molecular level and their mechanistic roles are still a subject of much research. In enzymes, the identification of conformational changes is generally elusive, as they are typically much faster than the chemical step and product release [2]. Nonetheless, structural fluctuations were observed in serine proteases when comparing their free and boronic acid inhibitor-complexed structures. Such plasticity may be an integral part of the catalytic cycle, mediating, amongst other steps, substrate binding, TS formation, stabilization, and product release (for a recent example see [105, 106]). Several hydrolytic enzymes were also shown to utilize an induced-fit mechanism to tune their specificity, either to a broad (e.g., alpha lytic protease) or extremely narrow (e.g., factor D) substrate spectrum (see [101] and reference therein). In addition, a recent kinetic study suggests that conformational changes control the aperture of AChE’s active site and are coupled to its catalytic activity [107, 108]. There had been several speculations regarding the role and effect that conformational changes may have on catalysis by antibodies. Padlan, for example, speculated that the of antibodies may hinder their catalytic efficiency [109], whilst Benkovic and Stewart suggested that antibodies elicited against TSA might be limited by a “lock-and-key” rigidity and will lack the conformational isomerism that characterizes enzymatic catalytic cycles [20]. In a more general context, the issue of

25


Fig. 8. Schematic representation of the two mechanisms of conformational changes linked to antibody-ligand complex formation. Ab and Ab’ represent two antibody isomers exhibiting low and high affinity, respectively.

conformational isomerism in antibodies has been a subject of much research and debate. In the past, kinetic (e.g., [110, 111]) and structural (e.g., [112] and references therein) studies provided strong evidence of the dominance of structural changes in antibodies. Conformational changes can be generally described by the inducedfit model (the ligand binding a low-affinity form of the antibody and inducing a subsequent conformational change that leads to a high affinity complex) or by preequilibrium between two different conformational species of the same antibody, only one of which binds the ligand with high affinity. The two models can be combined in one scheme (Fig. 8). While kinetic evidence supports the existence of both mechanisms, structural evidence has been put forward thus far for induced fit only. However, a recent study combining pre-steady-state kinetics in solution and X-ray crystallography provides conclusive evidence for pre-equilibrium in an antibody [113]. The identification of conformational changes in catalytic antibodies and their possible role in catalysis are discussed in this section.

4 Conformational changes in catalytic antibodies

The advent of catalytic antibodies has brought a renewed interest in antibody structure and evolution. In particular, structures of catalytic antibodies underlined the understanding of their mechanism at the molecular level (over 50 published structures of catalytic antibodies in PDB as of end of 2002). As is the case with many ordinary antibodies, conformational changes are observed with catalytic antibodies as well – in several catalytic antibodies, the structure of the TSA-bound antibody differs from the uncomplexed structure. But, in the absence of kinetic studies in solution, the interpretation of the crystallographic data regarding the mechanism of these con-formational changes is ambiguous. Further complications from the comparison of bound vs unbound antibody crystal structures arise from the effect of packing forces (e.g., the antibodies’ arrangement in the crystal in antibody 1D4 [95], CDR1 packed inside the active site of antibody 43C9, blocking the diffusion of the hapten [79]), and the buffer used for crystallization (e.g., antibody D2.3, see below and [15]). A recurrent structurally observed conformational change is a relative movement between the heavy- and light-chains’ variable regions [114], as exemplified by antibody CNJ206 where a 7◦ rotation and a ∼1 Å translation change were found between the TSA unbound and bound structures. In addition, a distinct isomerization was noted in the active site, where a dramatic change of the entire H3 CDR loop – from an open state to a closed state – is observed.

4 Conformational changes in catalytic antibodies

This changes the binding site completely – from a shallow form in uncomplexed antibody to a deep and narrow cavity in the TSA-bound state. This is assumed to be a TSA-induced conformational change, but in fact the TSA-bound conformation may be pre-existing, as neither the oxyanion hole nor the part of the active site that binds the phenolic leaving group is formed in the free form [14]. Induced-fit movement of CDR-H3 residues (H99-H102) was also suggested to enhance surface complementarity to the TSA vs the substrate and product of the catalyzed retroDiels-Alder reaction, thus implying that better rate enhancement is gained by the conformational changes (antibody 10F11, [19]). Crystal structures of antibody 1D4, catalyzing a syn elimination reaction, indicated conformational changes limited to CDR-H2. The conformational change increases the contacts with the TSA (notably residue Arg58), and may contribute to catalysis by constraining the substrate to an elimination-favorable orientation [95]. The free and TSA-complexed structures of several catalytic antibodies were solved, alongside the structures the putative germline antibodies from which these catatylic antibodies diverged [27, 115–117]. A general trend has been suggested, whereby the germline antibodies exhibit a different free and bound structure, whilst the mature antibodies exhibit the same structure. The observed isomerizations between the free and bound antibodies seem to occur mainly at CDR3H residues. In antibody 48G7’s precursor, tyrosines H98 and H99 shift by up to 5 Å upon TSA binding together, with a 4.5◦ 4 change in the V-regions’ orientation, resulting in better packing of the TSA’s p-nitrophenyl ring by TyrH99 [27, 117]. The hapten of antibody AZ-28, catalyzing an oxy-Cope reaction, induces a rotation of CDR-H3 away from the binding site to yield better π–π stacking with the hapten’s phenyl ring as well as enhanced orbital overlap to the reaction’s TS [115]. In the precursor of the oxidating, periodate-dependent antibody 28B4, better packing of the TSA phenyl ring is also achieved by movement of residues H95–H99 [116]. As judged by crystallography, the matured antibodies related to these germline antibodies exhibit the same structure when free and bound to the TSA. Interestingly, in antibodies 48G7 and 28B4, the TSA’s p-nitrophenyl ring adopts a different conformation in the mature vs the germline-complexed structures, locking them in a different binding pocket. This is surprising, as, in the case of the mature 48G7, the germline’s pocket remains unchanged. The above studies [118] promoted the hypothesis that the germline antibodies exhibit high conformational diversity, and that, during affinity maturation, the hapten binding conformation becomes dominant [110]. However, structural differences between the bound and the free state are commonly observed in mature antibodies ([112] and references therein), and at least one study has shown similar structures of the mature Diels-Alderase antibody 39-A11 and its putative germline precursor in both bound and unbound states [119]. The crystallographic studies were not accompanied by kinetics in solution that indicate the existence of a pre-equilibrium between different unbound conformations, in either the mature or the germline antibodies. Indeed, a kinetic study of mature antibodies that showed no structural differences between their free and TSA-bound states clearly indicated a pre-equilibrium between two antibody conformers, one exhibiting low affinity

27


toward the TSA and another high affinity and catalytic activity. The slow, minutescale isomerization between the two conformers resulted in hysteretic, “memorylike” behavior [15]. Moreover, it has also been shown that the crystallization buffer shifts the equilibrium toward the active conformer, resulting in both the free and TSA-bound antibodies yielding the same structure [15]. Also of interest are the mutations that lead to affinity maturation and their effect on TSA binding and rate of catalysis. In antibodies 48G7 and AZ-28 the mutations occur away from the active site. The importance of remote residues on catalytic efficiency was also noted in mutants of antibody 17E8 selected by phage display for better TSA binding. A double mutant was achieved, with ten-fold improvement of kcat /K M [120]. In antibody 28B4, however, three of the somatic mutations (nine in total), are indirect contact with the hapten [116]. In all three antibodies, somatic mutations increased the TSA-binding affinity (up to 40 000-fold in 48G7 [117]). Interestingly, the catalytic activity of the mature 48G7 was increased compared to its germline counterpart (see Section 2.1), whereas that of antibody AZ-28 was diminished, hinting at the role of structural flexibility in this antibody’s catalytic activity (the catalytic activity of 28B4 germline was not reported). Structural flexibility may also explain the increase in the catalytic activity of the ArgH100aGln mutant of antibody 43C9 (a CDR-H3 mutation remote from the active site), which reduces product inhibition [81]. 4.1 The contribution of structural isomerization to catalysis

The structural studies described above do not indicate a direct role for conformational changes in antibody catalysis, nor do they provide a measure of their contribution to TS stabilization and rate of catalysis. In the first quantitative study of conformational changes in catalytic antibodies, two independent kinetic measurements (one related to TSA binding and another directly to catalysis) performed with the D-Abs (D2.3, D2.4, D2.5) were used to reveal and quantify the contribution of conformational changes to catalysis [10, 15]. Pre-steady-state TSA-binding kinetics of D2.3, D2.4 and D2.5 antibodies, based on the change of intrinsic tryptophan fluorescence upon hapten binding, indicated the formation of a low micromolar affinity primary complex followed by a conformational change toward the final nanomolar affinity complex (Fig. 9a,b and Table 6 [15]). The rate of isomerization (6 s−1 ) revealed by fluorescence quenching measurements with both the 1b and 1p TSAs is significantly higher than the rate of hydrolysis of the D-Abs (0.05 – 0.22 s−1 ) and is therefore invisible in the catalytic, productrelease assay. However by increasing the turnover rate by means of a strongly reactive nucleophile (i.e., peroxide anion, see Section 2.3 above), a “rate ceiling” equal to the isomerization rate was observed (Fig. 9c) [10]. The isomerization provides 25 to 140fold higher affinity to the TSA (Table 6) and a concomitant increase to the D-Abs catalytic rates. This is yet another manifestation of the true representation of the actual TS by the TS analog. The “induced-fit” isomerization contributes not only to rate enhancement but also to specificity. A second,

5 Conclusions

Fig. 9 Induced fit exhibited by the Dantibodies. (a) Two-step kinetics reflected by the stopped-flow trace of fluorescence quenching of antibody D2.4 in the presence of TSA. The fast phase (shaded) is enlarged in the inset. Each phase was fitted to a single exponential rate equation. (b) The effect

of TSA concentration on the slow bimolecular binding step and on the fast isomerization step [15]. (c) D2.4 (full circles) and D2.3 (empty circles) catalytic rates dependency on the peroxide anion exogenous nucleophile’s concentration [10].

minute-scale, isomerization step is unique to antibody D2.3 [15]. This very slow conformational change increases the affinity to the TSA 1b antigen ∼7 fold yet decreases its affinity to TSA 1p by ∼2 fold (Table 6). These results may suggest that the immune system may select high-affinity and high-specificity antibodies that still exhibit a considerable degree of conformational change, and that affinity maturation does not necessarily involve a rigid “lock-and-key” binding mechanism [27, 37].

5 Conclusions

The examples given in this review emphasize the ability of catalytic antibodies to mimic multiple aspects of enzyme catalysis. In addition, several other enzyme characteristics are faithfully mimicked by hydrolytic antibodies but are beyond the scope of this review. Hydrolytic antibodies can exhibit both a fine substrate specificity that rivals natural enzymes (e.g., 2H6 and 21H3 [121], the D-Abs [25]

29

0.24 ± 0.02 0.20 ± 0.02 0.15 ± 0.01 0.3 ± 0.03 0.20 ± 0.03 0.20 ± 0.02 0.6 0.7 0.1 0.6 0.4 0.5

5±1 6±1 1 ± 0.1 6±1 7±2 4±1

0.2 ± 0.1 0.10 ± 0.05 0.01 ± 0.003 0.04 ± 0.01 0.2 ± 0.1 0.08 ± 0.03

Kfast+ s−1b) kslow+ s−1 kslow0.04 0.02 0.01 0.007 0.03 0.02

0.013 ± 0.002 0.001 ± 3 × 104 – – – –

Kslow s−1 kslowest+ s−1

0.002 ± 3 × 104 0.0023 ± 3 × 104 – – – –

kslowest-

0.15 2.3 – – – –

3±1 25 ± 3 1 ± 0.4 4±1 10 ± 2 10 ± 2

Kslowest nM KD c)

b)

Because of high error in its determination kfast+ was back-calculated from the overall dissociation constant (K d ) after measuring all the other kinetic parameters. Calculated from K d . c) Measured by fluorescence equilibrium titration.

a)

4 × 105 3 × 105 2 × 106 5 × 105 5 × 105 4 × 105

TSA µM−1 s−1 kfast+ s−1a) kfast- µM

D2.3 1b 1p D2.4 1b 1p D2.5 1b 1p

Ab

Table 6 Kinetic parameters of D-Abs’ binding to p-nitro-phenyl (1p)- and benzyl (1b)-phosphonate TSAs


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1

Chemical Microarrays for Small Molecule Ligand Screening G. Metz, H. Ottleben, and D. Vetter

Graffinity Pharmaceutical Design GmbH, Heidelberg, Germany

Originally published in: Protein-Ligand Interactions. Edited by Hans-Joachim Böhm, Gisbert Schneider. Copyright ľ 2003 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30521-6

1 Introduction

Since their introduction about a decade ago, combinatorial chemistry and highthroughput screening (HTS) have become indispensable tools in the drug-discovery process. The possibility to synthesize ever-increasing numbers of molecules through novel chemistries and automation is stimulating the development of higher screening capabilities through miniaturization and robotics. Robust biochemical assay development is providing the basis for large-scale screening of biological targets. While in the early days much effort and hope were directed towards managing a numbers game, the focus is shifting from quantity towards quality. For instance, the screening of compound mixtures is being replaced by screening of individual substances in a one-well–one-compound fashion. The design of general-purpose screening libraries as well as corresponding follow-up strategies has become a key aspect in small molecule discovery and optimization. Hits from high-throughput screening enter a selection process to become the subject of medicinal chemistry approaches in lead optimization. Strategies are employed to improve potency, selectivity, and physicochemical profile. If possible, several compound series are generated to allow for alternative routes in case of failure of one. Syntheses of analogues for further exploration are guided by a combination of medicinal chemistry knowledge and intuition, as well as quantitative structure-activity relationships, if available. Such studies help to define particular pharmacophoric features within the hit or lead molecule that constitute the underlying molecular recognition motifs between the ligand and its target and provide a hypothesis for its mode of action. The focus in screening for biological activity assays is on detecting hits with activities in the low micromolar range. Compounds exhibiting this level of activity tend to be of molecular weight in the range of 300–600 Da and of substantial functional Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Chemical Microarrays for Small Molecule Ligand Screening

complexity. A conceptually different strategy can be envisioned that – instead of trying to identify and keep relevant features in a rather complex hit compound – aims at a stepwise discovery starting from molecular fragments. Such fragments need to be screened with techniques suitable for the detection of presumably weak interactions. Guided by early stage structure-affinity information, more potent compounds can then be assembled either from a combination of low affinity binders or by chemical modification of the initial fragments. This article describes the underlying philosophy of fragment-based discovery as well as the experimental approaches suitable for this promising discovery concept. While fragment-based ligand discovery was first adapted in computational methods, several experimental techniques have been introduced recently. Biophysical methods such as nuclear magnetic resonance (NMR) and X-ray crystallography have been successfully applied to fragment discovery. A novel screening technique based on chemical microarrays in combination with label-free affinity detection has emerged and will be discussed in detail.

2 Fragment Approaches 2.1 Conceptual Ideas

Primary screening efforts in drug discovery aim at the identification of hit molecules with the necessary characteristics to be developed into a promising lead molecule. The definition of favorable properties of the starting screening compounds has gained much attention. The design of libraries with drug-like characteristics generally follows the so-called “rule of five” which has been established by retrospective analysis of known drugs and allows a quick assessment based on simple properties, namely, preferred ranges for molecular weight and clogP as well as the number of hydrogen bond donors and acceptors [1]. However, it has been proposed that the ideal profile for hit or lead compounds is different from that of the final drug molecule [2], in particular because hit or lead compounds must be amenable for further optimization. Three categories of lead compounds have been defined based on their physicochemical properties and typical affinities. First, it has been pointed out that hits from drug-like libraries rarely show high ( 96%) [49–51]. To simplify quantitative profiling of proteolytic peptide mixtures from 14 N/15 N-encoded proteins in whole cell lysates, Conrads et al. specifically enriched cysteine-containing peptides via a thiol-reactive affinity tag prior to FTICR-MS analysis [49]. However, a general drawback of cysteine tagging methods [such as the isotope-coded affinity tag (ICAT) technology discussed later in this chapter] is that a large protein fraction does not contain cysteine residues at all and often the cysteine content of the remaining fraction is low. The probability of analyzing a suitable set of isotopically encoded peptides to infer accurate quantitative information on precursor proteins is therefore significantly reduced. With increasing numbers of growth cycles, endogenous proteins become enriched in 15 N atoms when cells are cultured in 15 N media; eventually all 14 N atoms are replaced by 15 N. The corresponding mass shift of 14 N/15 N-labeled proteins or peptides can only be predicted if the amino acid sequences of the proteins are known. This certainly complicates quantitative evaluation of the mass spectral data acquired. Therefore, the use of high-resolution mass spectrometry, such as FTICR, has been recommended [49]. However, a suitable algorithm can predict the expected mass shift of peptides or proteins based on the proteome of the organism studied. 15 N- and 14 N-labeled peptides can easily be distinguished by their isotope distributions observed in mass spectra. Owing to the use of 15 N media of > 96% purity and therefore incomplete replacement of 14 N, the 15 N-labeled peptides exhibit additional isotope peaks in mass spectra. 2-D PAGE is often employed for high-resolution separation of 14 N/15 N-labeled cell lysates followed by MS-based identification and quantitation of distinct proteins. Alternatively, the applicability of a shotgun approach to determine concentration ratios of 14 N/15 N-encoded proteins from yeast on a large scale has been demonstrated [38, 39]. To support data analysis in shotgun proteomics, a correlation algorithm for automated protein quantitation has been developed [52]. In another approach for gel-free proteomics, Wang and coworkers reported an inverse labeling strategy to simplify protein quantitation by 15 N combined with MS [53, 55]. In this experimental approach, both perturbed and control samples are independently cultured in 14 Nand 15 N-enriched media, generating a set of four sample pools as opposed to two in a conventional labeling experiment. Differentially labeled control and perturbed samples are combined, processed, and analyzed by mass spectrometry. Peptide mass spectra obtained were then compared by subtractive analysis to reveal only those peptide pairs which show altered concentrations. This strategy provides confidence in detecting changes in protein concentrations. However, the workload and costs are doubled, so its necessity must be determined on a case by case situation.

6 Metabolic Labeling Approaches

In situ 15 N labeling has successfully been applied to studying bacteria, yeast, and some mammalian cells at moderate costs. Metabolic labeling of Caenorhabditis elegans by feeding 15 N-labeled E. coli cells [54] or 15 N labeling of potato plants for structural protein analysis [55] have also been reported. However, in vivo labeling of higher eukaryotic organisms is expensive and may significantly affect growth and protein expression profiles. A detailed study of the effects of 15 N labeling on these systems has not yet been reported. 6.2 Stable Isotope Labeling by Amino Acids in Cell Culture (SILAC)

Another very elegant in situ method for accurate quantitative assessment of protein or peptide abundances between two given cell lines is the stable isotope labeling by amino acids in cell culture (SILAC) [56]. Mann and coworkers grew mammalian cell lines in media lacking the essential amino acid leucine, but supplemented with its nonradioactive, deuterated form (2 H3 -Leu) [56]. Analysis of cell morphology showed that 2 H3 -Leu caused no differences in the growth of the cells. After five cell cycles, proteins were completely labeled with 2 H3 -Leu. Jiang and English reported metabolic labeling of a yeast strain auxotroph for leucine that was grown on synthetic complete media containing natural abundance Leu or 2 H10 -Leu [57]. Before this concept was harnessed for quantitative proteomics, incorporation of 13 C/15 N/2 H-encoded amino acids during cell culture was utilized to increase confidence in mass fingerprint database searches [58]. To prevent chromatographic differences in peptides containing 1 H/2 H isotopes, the SILAC approach was successfully extended to the use of 13 C6 -arginine [59], improving the accuracy of quantitation. When trypsin is used for protein digestion (note that trypsin cleaves proteins on the carboxy-terminal sides of arginine and lysine residues) in a 13 C6 -arginine experiment, the label is localized at the Cterminus of arginine-containing peptides, while lysine peptides are unlabeled and therefore do not provide quantitative information in peptide mass spectra. Fenselau and coworkers successfully used metabolic labeling with 13 C6 -arginine and 13 C6 lysine for comparative proteomics in drug-resistant and parental breast cancer cell lines [60]. This double isotope labeling approach was previously used to quantify changes in protein phosphorylation in resting versus activated human embryonic kidney cells [61]. Since both 13 C6 -lysine and 13 C6 -arginine were employed during culturing, tryptic digestion of proteins resulted in proteolytic peptide mixtures in which all peptides were labeled at the C-terminus, except for the original C-terminus of the proteins. This is certainly a considerable improvement to the SILAC approach, since it is essential to obtain an adequate set of peptide pairs from each protein for accurate relative quantitation, although there may still be variability from protein to protein. It has been demonstrated very recently that SILAC is applicable to multiplex experiments in order to measure dynamics of protein abundances in cells in response to stimuli such as growth factors or drugs [62]. Henrietta Lacks (HeLa) cells were grown in normal medium or in media lacking normal arginine and

15

16 Mass Spectrometry of Peptides and Proteins

supplemented with either 13 C6 14 N4 - or 13 C6 15 N4 -arginine. After harvesting the cells, differentially labeled cell populations were pooled in a 1 : 1 : 1 ratio for combined sample processing followed by triplicate LC/ESI-MS/MS peptide analyses to test for technical reproducibility. Interestingly, standard deviations obtained in quantitative measurements do vary significantly between proteins, even though the same number of peptide pairs was evaluated. Nevertheless, it was shown that metabolic labeling allows for reliable measurements of alterations in protein abundances of 1.5- to 2-fold in a single biological experiment. To account for biological variations in protein expression profiles, at least three independent biological experiments must be performed. The SILAC approach has also been used to investigate metastatic prostate cancer development at the protein level [63]. The fact that proteins showed altered concentration ratios by quantitative MS was confirmed by western blotting. In addition, proteomic approaches for quantitation of protein phosphorylation via SILAC combined with MS analysis have been described [61, 64, 65]. A recent study reports on identification as well as relative quantitation of in vivo methylation sites of proteins in HeLa cells by stable isotope labeling with 13 C2 H3 -methionine [66]. As shown by the examples given above, SILAC is a very flexible, simple, and accurate procedure for relative quantitation that provides deeper insight into biological systems based on cell culture.

7 Chemical Labeling Approaches

Chemical reactions are used to introduce isotope-coded tags into specific sites in proteins and peptides after sample collection, i.e., following tissue biopsy or cell harvest. In vitro labeling of biomolecules has been achieved in a variety of ways, including enzymatically directed labeling and incorporation of isotope tags prior to or following protein digestion (Figure 3). 7.1 Chemical Isotope Labeling at the Protein Level

Gygi et al. described an approach for accurate quantitation and concurrent identification of individual proteins within complex mixtures from yeast using chemical reagents termed isotope-coded affinity tags (ICATs) and tandem MS [67]. The ICAT reagent consists of a iodoacetyl moiety reacting with cysteine residues, an isotopeencoded linker (1 H8 or 2 H8 ) and a biotin tag. In this method, a set of two samples is differentially tagged and combined in a 1 : 1 ratio, the proteins are separated and then are enzymatically digested. The major innovation here was the selective enrichment of cysteine-containing peptides by the affinity tag, reducing the sample complexity by a factor of about 10 prior to MS analysis. Since the ICAT-encoded peptide pairs differ by at least 8 Da, this labeling method is applicable to MS analysis with low resolution instruments such as ion traps.


The ICAT approach promised to be a widely applicable tool for comparative proteomics of cells and tissues. However, it fails for proteins which are cysteine-free and often the cysteine content of proteins is rather low, rendering them unavailable for accurate quantitative MS analysis. Following this first report on the ICAT strategy [67], a number of studies have further conveyed the ICAT idea in quantitative proteomic research [46, 68–70]. However, there are several drawbacks in the ICAT strategy: (1) ICAT reagents are relatively large (about 500 Da) and their addition to proteins may be sterically hindered; (2) prolonged reaction times are often needed to achieve complete incorporation of ICAT tags into proteins which may lead to partial derivatization of lysine, histidine, methionine, tryptophan, and tyrosine residues; and (3) the chemical stability of the thioether linkage towards molecular oxygen is low, which may result in uncontrolled partial cleavage of the label via βelimination. Smolka et al. systematically evaluated the ICAT method and addressed the need for optimization of labeling conditions [71]. However, even if complete labeling of samples can be achieved, the molecular masses of peptides bearing the ICAT label are significantly increased and the analysis of the corresponding MS/MS spectra (especially for small peptides of fewer than ten amino acids) is complicated due to additional fragmentation of the affinity tag. Nevertheless, improvements in ICAT technology have been made [46], such as the development of a photocleavable ICAT reagent bound to a solid support, thus eliminating the large biotin tag [72, 73], and the introduction of a modified version of the ICAT reagent that uses acid-labile isotope-coded extractants (ALICE) [74]. Finally, 13 C-containing ICAT reagents have been introduced to ensure coelution of peptides during LC separation. This new ICAT generation has been applied to the analysis of a cortical neuron proteome sample to identify proteins regulated by the antitumor drug camptothecin [75]. In 2003, a stable isotope labeling similar to ICAT was reported utilizing a membrane-impermeable biotinylating reagent, which contains a cleavable linker and specifically reacts with primary amino groups, i.e., the ε -amino group of lysine residues and the free amino termini of isolated intact proteins [76]. The capability of this reagent, available in the light and heavy forms for labeling and enrichment of lysine-containing peptides followed by relative quantitative assessment, was shown for two standard proteins. Labeling of proteins was performed under nondenaturing conditions which resulted in incomplete incorporation of the label. Certainly, this considerably limits the reproducibility and accuracy of the method and therefore its usefulness in quantitative proteomics. However, the authors proposed that the method designed would allow for targeting of cell surface proteins (without the need for subcellular fractionation) which could be useful in drug research [76]. Recently, Lottspeich and coworkers reported an alternative approach, termed isotope-coded protein label (ICPL), which is also specific to primary amino groups [77]. Since ICPL is based on stable isotope tagging at the protein level, it is applicable to any protein sample, including extracts from tissues or body fluids. The ICPL tag is an isotope-coded N-nicotinoyloxy-succinimide (Nic-NHS) and different versions such as 1 H4 /2 H4 - and 12 C6 /13 C6 -Nic-NHS have been synthesized for their use

17


in quantitative proteomics on a global scale, the latter ensuring coelution of the isotopic peptide pairs and therefore accurate quantitation by LC/MS analysis. So far, the efficiency of the approach has been demonstrated by comparative analysis of E. coli lysates differentially spiked with standard proteins [77]. Advantages of the ICPL method are the generally high abundance of lysine residues in proteins providing multiple labeling sites and an increase in MS sensitivity for labeled peptides which permits accurate quantitation and yields in improved sequence coverage of proteins. However, modification of proteins by Nic induces a strong shift in pH resulting in a drastic change in the migration of labeled proteins during 2-D PAGE with isoelectric focusing in the first dimension [77]. This may be advantageous in the analysis of very basic proteins (e.g., ribosomal proteins) but considerably reduces the resolution of 2-D PAGE for the bulk of proteins. 7.2 Stable Isotope Labeling at the Peptide Level

Various alternatives to the ICAT and ICPL technologies have been reported but these methods are consistently based on chemical tagging at the peptide level. In this chapter, only the most common and promising peptide labeling approaches will be discussed [45, 78]. Regnier and coworkers coined the term global internal standard technology (GIST) which employs N-acetoxy-[2 H3 ]succinimide or 2 H4 -succinic anhydride as acetylation reagents to modify all primary amino groups in peptides [53, 55, 79–83]. The applicability of GIST to the quantitation of phosphoproteins enriched via immobilized metal affinity chromatography (IMAC) has recently been demonstrated [84]. In this work, relative differences in phosphopeptide concentration between samples were derived from isotope ratio measurements of the peptide isoforms observed in mass spectra. One should bear in mind, however, that the modification of lysine residues reduces the basicity; consequently, peptide ionization efficiencies can be low. In addition, N-terminally blocked peptides containing no further lysine residues are not susceptible to quantitative MS analysis via GIST. To address this issue, a combination of GIST and labeling of the C-terminal carboxyl groups by 18 O has been proposed and tested for the comparison of the relative protein expression level in epidermal cells grown in the presence or absence of epidermal growth factor [85]. Proteolytic peptide labeling with 18 O isotopes was first reported by Schnölzer et al. for its use in de novo sequencing experiments. Chemical labeling of the Ctermini of peptides with 16 O/18 O isotopes leads to a doublet signature for the y-ion series in MS/MS spectra [86]. Since the b-ion series do not contain a label, MS/MS data interpretation is quite straightforward. The usefulness of 18 O labeling for de novo peptide sequencing was shown with MALDI-TOF [87], ESI-qTOF [88], ESI-ion trap [89], and ESI-FTICR [90] instruments. In 2000, Roepstorff and coworkers described 18 O labeling for peptide and protein quantitation [91]. However, the uncontrolled incorporation of one or two 18 O


atoms into peptides and issues related to the overlap of two isotopic distributions complicated the calculation of peptide ratios from the MS spectra. Fenselau and coworkers successfully harnessed 18 O labeling for comparative proteomics using shotgun methods [92]. Generally, two protein samples are proteolytically digested in parallel, one in H2 16 O and the other one in H2 18 O. Alternatively, proteolytic digestion can be conducted prior to 18 O labeling, which increases the flexibility of this method [93]. The incorporation of 18 O atoms in the carboxy-termini of peptides is catalyzed by serine proteases (e.g., trypsin) via hydrolysis [92–94]. Double labeling can be obtained under adequate conditions (sufficient digestion times, high enzyme concentration and high enzyme activity) inducing a fixed mass shift of 4 Da into peptides. High as well as low resolution mass spectrometers can be employed for quantitative analysis of 16 O/18 O-labeled peptide pairs [95, 96]. 18 O labeling as a tool for proteomics was also evaluated by Figeys and coworkers [97] and a number of biologically relevant applications have been reported so far [98–104]. Stable isotope labeling methods commonly impart a mass difference as basis for relative quantitation by evaluation of peptide peaks in MS spectra. These methods are primarily designed for the quantitative analysis of a binary sample set. Even though the feasibility of multiplex protein quantitation from peptide mass spectra via SILAC [62] or ICPL [77] has been demonstrated, the further increase in the overall sample complexity has to be considered. Hence, high-resolution separation prior to MS analysis for the diminution of peak overlapping in peptide mass spectra is a prerequisite for accurate protein quantitation. To address this issue, a new generation of stable isotope tagging reagents, termed iTRAQ, has recently been introduced enabling the simultaneous quantitative comparison of a twofold up to a fourfold sample set [105]. The iTRAQ reagent is an N-hydroxysuccinimide ester which reacts with primary amines in peptides. It links an isotope-encoded tag consisting of a mass balance group (carbonyl) and a reporter group (based on N-methyl-pirazine) to peptides via the formation of an amide bond. In the iTRAQ method, the protein samples to be compared are separately digested with a protease, differentially labeled, pooled, and then jointly separated by chromatography followed by tandem MS analysis. Owing to the specific mass design of the 13 C, 15 N, and 18 O-encoded reporter and balance groups, the overall nominal mass of each of the four different iTRAQ reagents is kept constant and differentially labeled peptides therefore appear as single peaks in MS scans. When iTRAQ tagged peptides are subjected to MS/MS analysis, the amide bond of the tag fragments in a way similar to the cleavage of peptide bonds. The mass balancing carbonyl moiety, however, is lost with no retain of charge (neutral fragment) and liberates the reporter group as isotope-encoded fragment entities of 114.1–117.1 Da. The ion fragments (e.g., b- and y-ion series) of modified peptides are isobaric and fragmentation spectra obtained are usually improved with respect to ion signal intensities and sequence coverage. The relative concentration of the peptides can be deduced from the intensity ratios of reporter ions observed in the MS/MS spectra. The applicability of the iTRAQ method to multiplex quantitative protein profiling has been exemplified by

19


simultaneous relative and absolute quantitative analysis of wild-type yeast versus two isogenic mutant strains [105]. An inherent drawback of this methodology is that quantitative information is only obtained from those peptides which are subjected to MS/MS analysis. This fact stresses the necessity for efficient peptide separation via multidimensional chromatography (e.g., SCX/reversed-phase HPLC) prior to tandem MS analysis. Moreover, ion trap instruments cannot be employed in quantitative iTRAQ experiments since their peptide fragmentation spectra do not contain information on the low m/z range (low-mass cut off) in which reporter ions would appear (Doroshenko and Cotter, [106]). On the other hand, the use of fast scanning mass analyzers would be an advantage in order to increase the fraction of peptides from a total HPLC run that are subjected to MS/MS analysis. In a recent study, DeSouza et al. employed both iTRAQ and cleavable ICAT reagents in combination with multidimensional LC and tandem MS analysis to discover potential markers for endometrial cancer (EmCa) tissues [107]. As can be expected, the use of ICAT led to identification of a higher proportion of lower abundance signaling proteins while iTRAQ was clearly biased towards abundant sample constituents (i.e., ribosomal proteins and transcription factors). From this set of experiments, a total of nine potential markers for EmCa were discovered. Pyruvate kinase was found to be overexpressed in EmCa tissues using both iTRAQ and ICAT labeling [107]. In any case, the researcher should prefer stable isotope labeling methods which permit simple, specific, and complete incorporation of isotope tags at the earliest possible stage of sample processing. Furthermore, one should aspire to acquiring resolved peptide profiles and isotope patterns which imply the use of multidimensional separation technologies and high-resolution mass analyzers (e.g., FTICR, TOF/TOF, or qTOF). However, protein quantitation of high accuracy can also be achieved with low resolution mass spectrometers such as ion trap instruments [95]. In general, qTOF and TOF/TOF analyzers provide more accurate information on peptide abundances than ion trapping instruments due to space-charge limitations of the latter. The improvement in ion statistics and signal-to-noise ratios via prolonged scanning times is particularly beneficial for accurate quantitation of lower abundance constituents. While primarily it is the MS system used that determines accuracy and dynamic range of isotope ratio measurements, reproducibility strongly depends on the entire experimental strategy employed. The latter provides good reason for minimizing the number of separate sample processing steps, and thus for performing stable isotope labeling at the protein level (also in case chemical labeling is required for quantitative analysis of clinical and mammalian specimens). In terms of accuracy, it is essential to perform protein quantitation on the basis of multiple peptide pairs. Comparing stable isotope labeling followed by MS with differential 2-D PAGE using fluorescent dyes (DIGE technology), Heck and coworkers demonstrated that relative protein quantities measured with both methods are generally in good agreement [108]. It should be noted that for verification of altered ratios of protein concentrations in a biological system the analysis of independent repetitions (at least three biological experiments) is a must, irrespective of the quantitation method employed.

8 Quantitative MS for Deciphering Protein-Protein Interactions

8 Quantitative MS for Deciphering Protein-Protein Interactions

Stable isotope labeling in combination with mass spectrometry is on its way to revolutionizing the study of protein-protein interactions in cells covering both stable complexes as well as transient interactions. There is a variety of techniques available for the purification of protein complexes. In-depth information on purification strategies as well as protein complex analysis by MS means is provided in the literature [109–116]. In a conventional approach, the components of a purified complex and a control are separated by 1-D PAGE, proteins are subsequently stained and band patterns are compared. Distinct bands are cut out from the gel followed by in-gel protein digestion and peptide LC/ESI-MS/MS analysis for protein identification. The lists of the individual protein compositions are then compared to distinguish between background proteins and specific interacting partners in the purified complex. However, due to limitations in peptide MS/MS analyses of complex samples performed separately (e.g., MS/MS analysis of only a subset of peptides in a complex mixture), the nonidentification of proteins in a control does not provide convincing evidence for the specificity of protein-protein interactions and therefore further proof is required, e.g., by biochemical methods such as western blotting. To address this issue, quantitative MS methods can be utilized to allow for identification of specific interacting partners in protein complexes with high reliability. In a metabolic labeling approach, two distinct cell populations (e.g., one containing the bait protein fused with a tag for affinity purification, the other one representing the control) are differentially labeled with stable isotopes during culturing, and pooled in a 1:1 ratio followed by cell lysis and isolation of the protein complex via affinity purification (Figure 4). In contrast, in chemical labeling approaches using ICAT or peptide-specific reagents, affinity purification of protein complexes derived from cell cultures or tissues of two distinctive differentiation or developmental states is performed separately followed by stable isotope tagging of proteins or proteolytic peptides. In either case, samples are combined after completion of the labeling step and proteolytic peptide mixtures are jointly analyzed by LC/tandem MS (Figure 4). In such experiments, protein identification is obtained on the basis of sequence-specific information in peptide fragmentation spectra. In addition, specific interacting partners can be distinguished from copurifying proteins via ratio measurements of light peptides versus their heavy counterparts. True binding proteins become specifically enriched via affinity purification of complexes, and therefore peptide ratios are significantly greater than should be observed in mass spectra representing these proteins. In contrast, nonspecific binding partners are present in equal amounts, resulting in peptide ratios of about 1 (Figure 4). In the case of protein complex versus control, direct discrimination between bona fide and nonspecific interacting partners can be achieved by comparing ratios of peptide pairs. A major promise of this approach is that protein complex purification protocols can be simplified and mild washing conditions used; thus, the ability to screen for

21


Fig. 4 Schematic representation of the strategies followed for accurate mapping of protein complexes by stable isotope labeling combined with MS. Relative quantitative MS allows for discrimination between specific interacting partners and contaminants which are often copurified with the protein complex of interest. In the metabolic labeling approach, two cell populations are differentially labeled with stably isotopically encoded amino acids (e.g., 13 C -/12 C -arginine) during culturing. Fol6 6 lowing the labeling experiment, the heavy cell population, which contains the bait protein (termed B) fused with a tag for affinity purification, and the light cell population which is used as control are mixed in a 1:1 ratio followed by cell lysis and protein complex isolation. In the chemical labeling approach, cells from cultures or tissues of two distinctive differentiation or developmental states (termed states A and B) are sepa-

rately lysed followed by affinity purification of protein complexes and stable isotope tagging of proteins. Subsequently, the isotopically tagged protein samples can be pooled in a 1:1 ratio and further processed. In both approaches, proteolytic peptide mixtures are generated and then separated by liquid chromatography (LC) followed by qualitative as well as relative quantitative analysis using MS/MS. Specific interacting partners can be differentiated from copurifying proteins based on ion signal intensities of light peptides versus their heavy counterparts in MS spectra. Peptide ratios significantly greater than 1 point to the presence of specific interacting partners; peptides ratios of about 1 are usually distinctive for nonspecific interacting partners. Protein identification is performed on the basis of sequencespecific information observed in MS/MS spectra.

9 Conclusions

transient or weak interaction partners is improved. This has been demonstrated by Aebersold and coworkers who used ICAT technology combined with MS analysis to identify specific interacting partners of a large RNA polymerase II (Pol II) preinitiation complex (PIC) only partially purified from nuclear extracts by a single-step promoter DNA affinity procedure [117]. Comparative analysis of two or more isolated protein complexes looks for differences in protein composition. Using ICAT, dynamic changes in the composition and abundance of STE12 complexes immunopurified from yeast cells in different states [117] as well as in protein complexes associating with the MafK transcription factor upon erythroid differentiation in MEL cells [118] were detected. Pandey and coworkers have recently provided detailed instructions for using the SILAC approach to study protein complexes, protein-protein interactions, and the dynamics of protein abundance and posttranslational modifications [119]. This in situ labeling approach combined with affinity pull-down experiments was shown to be useful to study (1) signaling complexes involved in the epidermal growth factor receptor (EGFR) pathway of EGF-stimulated versus unstimulated HeLa cells [120]; (2) peptide-protein interactions in EGFR signaling [121] and (3) early events in EGFR signaling in a time-course experiment [122]. Hochleitner et al. have recently reported the determination of absolute protein quantities of the human U1 small nuclear ribonuclearprotein (snRNP) complex by using synthetic peptides and stable isotope peptide labeling combined with MS analyses [123]. The examples given above clearly demonstrate that quantitative MS-based proteomics is a powerful tool for shedding light on the stoichiometry, dynamics and assembly processes of functional protein modules in biological systems. Since drugs can equally be used as affinity baits, this methodology could also provide valuable information on cellular target proteins in drug research.

9 Conclusions

Biological mass spectrometry represents the core technology in proteome research. Instrumental designs and fragmentation techniques are still advancing at an expeditious pace. Novel proteomic strategies, with MS, biochemical as well as molecular cell biological techniques acting in concert, promise to dramatically extend our current knowledge of biological systems in the near future. In quantitative proteomics, it is crucial to consider the variability of biological systems in order to deliver meaningful results regarding protein quantities. This biological variability should not be underestimated but rather be addressed by independent repetitions and advanced statistical data analysis. Certainly, evaluation of large quantitative data sets is very laborious and only feasible using bioinformatics. In our laboratory, the current lack of suitable software tools for accurate large-scale determination of protein concentration ratios deduced from MS or MS/MS spectra was addressed by in-house development of an efficient software package supporting quantitative analysis of

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data obtained in any experiment employing stable isotope labeling. Quantitative MS in combination with epitope tagging opens up an additional major area for future proteomic research: the mapping and characterization of functional protein modules on a large scale. This adds to the great promise of proteomics in drug research. Yet, the final impact of proteomics in the discovery of new drug targets cannot easily be foreseen. Acknowledgements

My special thanks go to Dr. Silke Oeljeklaus for helpful discussion and critical reading of the manuscript and to her and Sebastian Wiese for assistance in the design of the figures presented in this chapter. References 1 BARBER, M., BORDOLI, R. S., SEDGEWICK, R. D. and TYLER, A. N. (1981). Fast-atom bombardment of solids (F. A. B.): a new ion source for mass spectrometry. J. Chem. Soc. Chem. Commun. 293, 325–327. 2 KARAS, M., HILLENKAMP, F. (1988). Laser desorption ionization of proteins with molecular masses exceeding 10,000 daltons. Anal. Chem. 60, 2299–2301. 3 MENG, C. K., MANN, M., FENN, J. B. (1988). On protons of proteins – a beams a beam for a that. Z. Phys. D 10, 361–368. 4 KRAUSE, E., WENSCHUH, H., JUNGBLUT, P. R. (1999). The dominance of arginine-containing peptides in MALDI-derived tryptic mass fingerprints of proteins. Anal. Chem. 71, 4160–4165. 5 ERICSON, C., PHUNG, Q. T., HORN, D. M., PETERS, E. C., FITCHETT, J. R., FICARRO, S. B., SALOMON, A. R., BRILL, L. M., BROCK, A. (2003). An automated noncontact deposition interface for liquid chromatography matrix-assisted laser desorption/ionization mass spectrometry. Anal. Chem. 75, 2309–2315. 6 CHAURAND, P., CAPRIOLI, R. M. (2002). Direct profiling and imaging of peptides and proteins from mammalian cells and tissue sections by mass spectrometry. Electrophoresis 23, 3125–3135. 7 HUTCHENS, T. W., YIP, T. T. (1993). New desorption strategies for the mass

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spectrometric analysis of macromole-cules. Rapid Commun. Mass Spectrom. 7, 576–580. CHAURAND, P., SCHWARTZ, S. A., CAPRIOLI, R. M. (2002). Imaging mass spectrometry: a new tool to investigate the spatial organization of peptides and proteins in mammalian tissue sections. Curr. Opin. Chem. Biol. 6, 676–681. STOECKLI, M., CHAURAND, P., HALLAHAN, D. E., CAPRIOLI, R. M. (2001). Imaging mass spectrometry: a new technology for the analysis of protein expression in mammalian tissues. Nat. Med. 7, 493–496. MEDZIHRADSZKY, K. F., CAMPBELL, J. M., BALDWIN, M. A., FALICK, A. M., JUHASZ, P., VESTAL, M. L., BURLINGAME, A. L. (2000). The characteristics of peptide collision-induced dissociation using a high-performance MALDI-TOF/TOF tandem mass spectrometer. Anal. Chem. 72, 552–558. PETRICOIN III, E. F., ORNSTEIN, D. K., PAWELETZ, C. P., ARDEKANI, A., HACKETT, P. S., HITT, B. A., VELASSCO, A., TRUCCO, C., WIEGAND, L., WOOD, K., et al. (2002). Serum proteomic patterns for detection of prostate cancer. J. Natl. Cancer Inst. 94, 1576–1578. PETRICOIN, E. F., ARDEKANI, A. M., HITT, B. A., LEVINE, P. J., FUSARO, V. A., STEINBERG, S. M., MILLS, G. B., SIMONE, C., FISHMAN, D. A., KOHN, E. C., LIOTTA,

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Difference Gel Electrophoresis (DIGE) Barbara Sitek Ruhr-University Bochum, Bochum, Germany

Burghardt Scheibe GE Healthcare Bio-Sciences, Discovery Systems, Freiburg, Germany

Klaus Jung University of Dortmund, Dortmund, Germany

Alexander Schramm Universitätsklinikum Essen, Essen, Germany

Kai Stühler University of Dortmund, Dortmund, Germany

Originally published in: Proteomics in Drug Research. Edited by Michael Hamacher, Katrin Marcus, Kai Stühler, André van Hall, Bettina Warscheid and Helmut E. Meyer. Copyright ľ 2006 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31226-9 Abstract

In the last years the application of two-dimensional electrophoresis (2-DE) has often been declared outdated and a new century of gel-free proteomics was announced. Nevertheless, 2-DE is still the method of choice when analyzing complex protein mixtures. With a separation of 10 000 proteins, 2-DE gives access to high-resolution proteome analysis. Continuous development has consolidated 2-DE application in proteomics, where the introduction of difference gel electrophoresis (DIGE) is the latest improvement. DIGE is based on fluorescently tagging all proteins in each sample with one set of matched fluorescent dyes designed to minimally interfere with protein mobility during 2-DE. For a DIGE analysis, two different fluorescence dyes, CyDyeő minimal dyes and CyDye saturation dyes, are available. CyDye minimal dyes react with an NHS-ester bond of lysine ε-amino residues and enable coelectrophoresis of up to three different samples in one approach. For special applications, e.g., samples from microdissection, the CyDye saturation dyes allow complete 2-D analysis and


2 Difference Gel Electrophoresis (DIGE)

quantification of protein abundance changes in scarce sample amounts. The dyes react via a maleimide group with all available cysteine residues in the protein sample, giving a high labeling concentration. Here we show the application of both CyDye types for the analysis of relevant clinical samples and an approach for evaluating DIGE data statistically. We used CyDye minimal dyes to detect kinetic proteome changes resulting from ligand activation of either tyrosine kinases TrkA or TrkB in neuroblastoma cells. TrkA and TrkB are members of the family of neurotrophin receptors, which mediate survival, differentiation, growth, and apoptosis of neurons in response to stimulation by their ligands, NGF and BDNF, respectively. In addition, we applied CyDye saturation dyes for the identification of new molecular markers of the pancreatic tumor progression. One thousand microdissected cells were analyzed from different pancreatic intraepithelial neoplasias (PanIN) grades, the precursor lesion of pancreatic ductal adenocarcinoma (PDAC). Based of the multiplexing strategy and the application of an internal standard, DIGE enables one to perform quantitative proteomics with high accuracy allowing statistical approaches for high-confidence data analysis.

1 Introduction

In the early 1990s Marc Wilkins introduced the term proteome as a description for all proteins, which are coded by a genome at specific time points and under certain conditions. It is thought that in addition to the analysis of the genome that proteome analysis, with its high dynamic and complexity, can be used to describe life in more detail. A number of different methods have therefore been established in different proteomics techniques to allow analysis of complex protein mixtures. Currently, two-dimensional electrophoresis (2-DE) is the separation method with highest resolution power for protein samples. Up to 10 000 proteins can be separated in one gel and are then accessible for quantitative analysis (Figure 1 [1]). In 1975, 2-DE was independently developed by Klose et al. [2] and O’Farrell et al. [3] representing the combination of two different separation techniques. In the first dimension the proteins are focused according to their isoelectric points (pIs) (isoelectric focussing (IEF)) and subsequently in the second according to their molecular size (SDS electrophoresis) (see review [4] for more details). For the IEF, two different techniques, carrier ampholyte IEF (CA-IEF) [1] and immobiline pH gradient (IPG) [3, 6], respectively, are available. The immobilization of the pH gradient and fixation on a plastic backing simplified the IEF and therefore allowed the widespread application of 2-DE (reviewed in more detail in, [2, 8, 9]). Meanwhile, a number of different IPG stripes, with narrow or broad as well as linear or nonlinear pH gradient, are provided by different manufacturers. In contrast, CA-IEF, a labor-intensive technique, failed to achieve widespread application but is still used in more specialized laboratories.

1 Introduction

Fig. 1 Schematic representation of 2-dimensional electrophoresis (2-DE). In the first dimension the proteins are focused according to their isoelectric point [isoelectric focusing (IEF)] and subsequently in the second dimension according to their molecular size (SDS electrophoresis or SDS-PAGE).

For quantitative analysis the proteins can be visualized using a number of staining techniques with different sensitivities and linear dynamic detection ranges (Table 1). In contrast to array technologies like, e.g., Affymetrix chips or protein chips where the analyte position is assignable with a high local x, y resolution, the protein (spot) position in a 2-DE gel depends on its physicochemical properties. Therefore, generating 2-DE gels for a differential analysis requires standardized opTable 1 The most commonly used protein staining methods for proteome analysis using

2-DE. Besides the low detection limit of the applied staining methods, the linear dynamic range for protein quantification is another important parameter for a global description of a proteome Staining method Silver Zinc imidazole SyproRuby SyproOrange, SyproRed Colloidal Coomassie DIGE minimal DIGE saturation Phosphor-imaging (32 P, 14 C and 35 S) a

Detection limit (ng)

Linear dynamic range

Reference

1 3–5a 10 1 30

2 orders of magnitude No quantification 3 orders of magnitude 3 orders of magnitude

[38] [40] [37] [42] [44]

3 orders of magnitude 3–5 orders of magnitude 3–5 orders of magnitude 5 orders of magnitude

[41] [15] [43] [39]

8–10 0.1–0.2 0.005–0.010 1

Detection limit of MS-compatible silver staining [25].

3


erating procedures to achieve highly reproducible protein patterns [10]. However, small alterations in 2-DE processing, such as, for instance, during the gel casting process, polymerization reaction and/or temperature changes during the IEF run (21–24 h) results in reproducibility variations which have to be compensated for by appropriate image analysis software. Besides detection and normalization features, classical image analysis software like, for example, ImageMaster, PDQuestő , ProteomWeaverő or Delta2Dő provide appropriate algorithms for matching of the spots between different 2-DE gels to compensate for these variations.

2 Difference Gel Electrophoresis: Next Generation of Protein Detection in 2-DE

An important improvement for the application of 2-DE was the introduction of the so-called difference gel electrophoresis (DIGE) by J. S. Minden’s group in 1997 [11]. DIGE circumvents some basic problems of 2-DE, for example gel-to-gel variations and limited accuracy using different fluorophores (Cy2, Cy3 and Cy5) for a multiplexed analysis. In contrast to classical detection methods (Table 1) this method relies on covalent derivatization of proteins in each sample with one of the set of matched CyDye that do not affect the relative mobility of proteins during electrophoresis. Thus, DIGE allows rapid identification of protein changes between two samples on the same 2-DE gel without influences of gel-to-gel variations. Additionally, DIGE covers a dynamic detection range of 3–5 orders of magnitude; while silver staining can only detect 30-fold changes [1, 9, 11, 13, 15]. For a DIGE analysis the two different fluorescence dyes CyDye minimal dyes and CyDye saturation dyes are available. CyDye minimal dyes that react via an NHS-ester bond with ε-amino residues of lysine enable coelectrophoresis of up to three different samples in one approach. Approximately 3% of the available proteins are labeled on a single lysine per protein, whereas the rest remains unlabeled. This makes the technique robust and labeling optimization is usually unnecessary. The three different CyDye tags add approximately 450 Da to the protein mass when coupled to the protein. The resulting image patterns are comparable to poststained gels and sensitivity of the minimal dyes is similar to most sensitive silver staining. A pooled internal standard can be created by mixing aliquots of all samples to be analyzed. The use of such an internal standard on each gel that comprises equal quantities of each of the samples in the experiment allows calculation of ratios for the same protein spot within one gel as well as between gels. This largely removes experimental gel-to-gel variation leading to improved accuracy of protein quantification between samples from different gels. In a setup of two samples plus internal standard per gel, renunciation of gel repetitions and emphasis on biological repetitions brings down the total number of gels per experiment necessary for quantitative statistics (see Section 2.3). The bottlenecks of classical 2-DE-like methodical variation, laborious image analysis and restricted quantification are therefore minimized in the DIGE workflow (Figure 1).


For special applications, e.g., samples from microdissection (see Section 2.3), the CyDye saturation dyes enable complete 2-DE analysis and quantification of protein abundance changes in scarce sample amounts (5 µg protein/image). The dyes react via a maleimide group with all available cysteine residues in the protein sample, giving a high labeling concentration. Owing to their net zero charge, there is no charge alteration of the labeled protein. As with all DIGE experiments an internal standard sample containing an equal amount of each sample is run on each gel. Sensitivity is 20-fold higher than for silver staining. In contrast to minimal labeling with saturation dyes, a labeling optimization is necessary to determine the appropriate amount of dye. With a confocal fluorescent imager the dye images are acquired at their specific wavelength without crosstalk. Dedicated image analysis software (DeCyder) utilizes a proprietary codetection algorithm (up to triple detection) that permits automatic detection, background subtraction, quantification, normalization and intergel matching of fluorescent images. The experimental design using the internal standard effectively eliminates gel-to-gel variation, allowing detection of small differences in protein levels (Figure 2). Systemic variation as well as inherent biological variation arising from patientto-patient, culture-to-culture differences, etc., can be clearly differentiated from induced biological changes. The DIGE system allows discrimination between true biological differences on the one hand and systemic as well as interindividual differences on the other.

2.1 Application of CyDye DIGE Minimal Fluors (Minimal Labeling with CyDye DIGE Minimal Fluors) 2.1.1 General Procedure In the standard labeling protocol, proteins are first solubilized in a DIGE lysis buffer (30 mM Tris, 7 M urea, 2 M thiourea, 4% (w/v) CHAPS, pH 8.5). Samples produced by acidic precipitation should be lysed with DIGE lysis buffer adjusted to an appropriate pH (e.g., 9.5) resulting in a pH between 8 and 9. Lower pH during the labeling reaction affects labeling efficiency and kinetics in a negative way. The protein concentration should then be determined using a standard protein quantification method. Protein concentrations in the range 1–20 mg/mL have been successfully labeled using the standard protocol (personal communication, Amersham Bioscience). CyDye samples are then added to the protein lysate and incubated on ice in the dark for 30 min so that 50 µg of protein are labeled with 400 pmol of CyDye (prepared in a working solution containing 400 pmol CyDye µL-1 anhydrous dimethylformamide). Then 1 µL of 10 mM lysine is added to stop the reaction and left for 10 min on ice in the dark. Samples can now be stored for at least one month at −70 ◦ C. Lysis buffer in classical 2-DE traditionally contains primary amines (e.g., carrier ampholytes, pharmalytes) and excess of thiols (e.g., DTT, DTE) which should be not present during the labeling reaction.

5


Fig. 2 Difference gel electrophoresis (DIGE). Ettanő DIGE workflow: three-color and two-color experiments including the internal standard. For fluorescence proteins tagging, two different CyDyes techniques are available. Minimal fluors allow consideration of three different CyDyes (Cy2, Cy3 and Cy5) in a multiplexing experiment. Applying sat-

uration fluors multiplexing is only done for internal standardization, but allows working with low concentration protein samples. After image acquisition at CyDyes specific wavelength using a confocal fluorescence scanner (e.g., Typhoon series) protein spot pattern can be analyzed by appropriate image analysis software (DeCyder).

After labeling, 2×lysis buffer [8 M urea, 4% (w/v) CHAPS, 2% (v/v) carrier ampholytes, 2% (w/v) DTT] can be added to the samples in a 1+1 dilution for IEF. Combining the three samples to be separated in one IEF strip or tube gel results in a total protein concentration of 150 µg per gel (50 µg Cy3+50 µg Cy5+50 µg Cy2 labeled). From here methods are virtually identical to classical 2-DE. A peculiarity is that glass cassettes should have low intrinsic fluorescence capacity, since the gel will be scanned still assembled between the plates. The CyDyes are matched for size and charge resulting in an overlay of identical proteins when run on the same 2-DE gel. The lysine amino acid in proteins carries a +1 charge at neutral or acidic pH. CyDye minimal dyes also carry an intrinsic +1 charge which, when coupled to the lysine, replaces the lysine’s +1 charge with its own, ensuring that the pI of the protein does not significantly alter (Figure 3). The experimental design is based on evidence that the experimental variation in a 2-DE experiment is mostly due to gel-to-gel variation. Running multiple samples on a single gel reduces the number of gels required to produce the same amount


Fig. 3 Labeling chemistry of CyDye DIGE minimal fluors. Dye Nhydroxysuccinimid-ester (NHS-ester) reacts with a single g-amino residue of lysine per protein and a positively charged dye molecule replaces the positive charged lysine on the protein so there is no net change in pI.

of data. The recommended protocol suggests that a Cy2-labeled internal standard sample should be run on all gels within an experiment. The standard sample consists of an aliquot of all the different samples in an experiment (Figure 2). This ensures representation of all proteins on all the gels analyzed. The standard sample will increase the reliability in matching between gels and will also allow the generation of accurate spot statistics between gels (see Section 2.3). 2.1.2 Example of Use: Identification of Kinetic Proteome Changes upon Ligand Activation of Trk-Receptors DIGE exerts its full strength in settings comprising highly similar but not identical biological conditions. Potential applications include monitoring of cellular responses to all kinds of external stimuli. Those systems are mostly error-prone in classical 2-DE due to the above-mentioned interexperimental variations. We have chosen the DIGE system to detect kinetic proteome changes resulting from ligand activation of either tyrosine kinases TrkA or TrkB in neuroblastoma cells (Figure 4). TrkA and TrkB are members of the family of neurotrophin receptors, which mediate survival, differentiation, growth, and apoptosis of neurons in response to stimulation by their ligands, NGF and BDNF, respectively (reviewed in [16]). So far, little is known about the molecular mechanisms used by TrkA/TrkB to mediate this different biological behavior. Expression levels of TrkA/TrkB are important prognostic factors in a variety of embryonal tumors including neuroblastoma, the most common solid tumor of childhood [17]. Since TrkA/TrkB exhibit a high level of sequence similarity and use overlapping pathways for signal transduction, the existence of specific effector molecules crucial for receptor and cell-type specific response is likely [18]. To identify these effectors by analyzing biological effects of TrkA and TrkB activation in a defined model, we performed a proteome analysis using the human neuroblastoma SY5Y cell line stably transfected with the TrkA or TrkB cDNA [19]. The resulting phenotypes of these cell lines have been extensively analyzed and are in agreement with the observed biological functions [5, 6].

7


Fig. 4 2DE-gels of SY5Y–TrkA cell lysate (A) or SY5Y–TrkB cell lysate (B). Separation of proteins was performed in the first dimension (horizontal) by IEF and then in the second dimension (vertical) by SDS-PAGE. Representative pictures of whole cell lysates

of SY5Y–TrkA (A) and SY5Y–TrkB (B) are shown. Differentially expressed proteins following activation of neurotrophin receptors TrkA or TrkB by their respective ligands are numbered and correspond to the data presented in Table 2.

Proteomic changes were monitored in a time-course of 0, 0.5, 1, 6 and 24 h following receptor activation. These time points were chosen to monitor immediate responses to neurotrophins (NGF/BDNF) as well as late effector proteins. Considering the biological variation, we focused on identification of significantly regulated protein spots in five biologically independent experiments at each time point. At each time point a differential comparison was performed between neurotrophintreated and untreated cells. Technically, 50 µg protein of untreated and ligand treated cells are labeled with Cy5 and Cy3, respectively. A pool of all analyzed samples was labeled with Cy2, generating the internal standard. Proteins extracted from neurotrophin-treated and untreated SY5Y–TrkA/B cells were labeled with Cy3 and Cy5, respectively, mixed with Cy2-labeled internal standards (Figure 5) and run in one gel. After scanning of gels and manual correction of spot detection, protein spots were matched for statistical analysis and determination of the differentially expressed proteins. In whole cell lysates of SY5Y–TrkA/TrkB, 1700–1900 distinct protein spots were detected by the DeCyder software and subsequent manual correction. The analysis of the expression profiles of SY5Y–TrkB cells resulted in 24 significantly regulated spots induced by BDNF (p < 0.05, ratio > 1.5) and 13 regulated spots induced by NGF in SY5Y–TrkA cells (p < 0.05, ratio > 1.3) (Table 2). A total of 20 spots regulated during the time-course following neurotrophin treatment were specific for SY5Y–TrkB cells, and nine regulated spots were specific for SY5Y–TrkA cells. Four spots were regulated in both cell lines. While three spots demonstrated the same regulation pattern, one was inversely regulated between SY5Y–TrkA and SY5Y–TrkB. The respective protein was identi-


Fig. 5 Schematic workflow of sample preparation for 2D-DIGE. SY5Y–TrkA and SY5Y–TrkB cells were stimulated by neurotrophins NGF (100 ng/mL) or BDNF (50 ng/mL), respectively (“treated cells”). Controls received fresh medium without neurotrophins at t = 0 (“untreated cells”). Samples of the same time point following

neurotrophin addition were labeled, mixed and run in one gel with an equal amount of internal standard (B). This standard was pooled from all samples in an experiment. The lower part of the figure exemplarily depicts the procedure for time point t = 24 h. Experiments were repeated five times to adjust for inter-experimental variations.

fied as tropomyosin-3 by MALDI-PMF. The majority of the proteins differentially expressed upon neurotrophin receptor activation were regulated in the late stimulation phase (10 of 13 protein spots in SY5Y–TrkA and 16 of 24 in SY5Y–TrkB) (Figure 6). Most of the proteins identified were assigned to the GeneOntology (GO) class “cytoskeleton organization and biogenesis”, but proteins involved in signal transduction could also be found. Most prominently, SY5Y–TrkB transfected cells up-regulate hnRNP K, which functions as a “RNA silencer” in immature epithelial cells to suppress translation of proteins only needed in differentiated cells. It is intriguing to speculate about a similar function for hnRNP K in preventing expression of proteins found only in differentiated neuronal cells. A recent report indicated that in fact hnRNP K controls the switch from proliferation to neuronal differentiation through regulation of p21 [22]. In this model system, application of the DIGE system has set the basis for integration of the signaling pathways and functional analysis of effector proteins as well as insights into the underlying courses for the opposing phenotypes caused by activation of different neurotrophin receptors. 2.2 Application of Saturation Labeling with CyDye DIGE Saturation Fluors 2.2.1 General Procedure As this type of labeling method aims to label all available cysteines on each protein, many cysteine residues existing as disulfide bonds in proteins must be unfolded and the disulfide bonds broken. This can be achieved under denaturing conditions

9

10 Difference Gel Electrophoresis (DIGE) Table 2 Summary of significantly regulated proteins following neurotrophin receptor activa-

tion in SY5Y–TrkA or SY5Y–TrkB identified by MALDI-MS Spot No.

Protein

Theoretical

Experimental

pI

MW [kDa]

pI

MW [kDa]

Accession no. NCBI

Regulation/ kinetic group TrkA

TrkB

1

Dynein

5.0

71.5

5.4

96.9

gi/24307879

none

↑A

2 3

5.0 5.0

71.5 72.3

5.5 5.3

95.3 86.8

gi/24307879 gi/16507237

↓C ↑C

↓A ↑C

5.0

72.3

5.3

86.8

gi/16507237

none

↑C

6.8

74.1

6.2

86.6

gi/27436946

none

↑C

5.1

51.0

5.4

72.0

gi/14165437

none

↑B

7

Dynein Heat shock 70kDa protein 5 Heat shock 70kDa protein 5 Lamin A/C isoform 1 precursor Heterogeneous nuclear ribonucleoprotein K isoform a not identified

5.6

67.5

↓C

↓B

8

Lamin A/C isoform 2

6.4

65.1

6.0

62.9

gi/27436946

none

↑C

9

Lamin A/C isoform 2

6.4

65.1

6.1

62.9

gi/27436946

none

↑C

10

JC5704 protein disulfide-isomerase (EC 5.3.4.1) ER60 precursor

5.9

56.8

5.7

60.9

gi/7437388

none

↑C

11

JC5704 protein disulfide-isomerase (EC 5.3.4.1) ER60 precursor

5.9

56.8

5.8

60.4

gi/7437388

none

↑C

12

Dihydrolipoamide dehydrogenase precursor

7.9

54.2

6.3

60.4

gi/4557525

none

↓B

13

Adenylyl cyclase-associated protein Nuclear matrix protein NMP200 related to splicing factor PRP19

8.0

51.7

6.3

57.5

gi/5453595

none

↓B

6.1

55.2

6.1

60.5

gi/7657381

none

↓C

TATA binding protein interacting protein 49 kDa ATP synthase, H+ transporting,

6.0

50.2

6.1

56.6

gi/4506753

none

↓C

9.2

59.8

6.3

55.8

gi/15030240

none

↓C

4 5 6

14

15

16 17

ATP synthase, H+ transporting,

9.2

59.8

6.2

55.8

gi/15030240

none

↓C

18

Calumenin

4.4

37.1

5.0

52.0

gi/4502551

none

↑C

19

A Chain A

8.5

36.7

6.5

35.8

gi/13786849

none

↓C

20

A27674 tropomyosin 3, fibroblast

4.7

32.9

5.1

34.4

gi/88928

↓C

↑C

2 Difference Gel Electrophoresis: Next Generation of Protein Detection in 2-DE Table 2 (Continued)

Spot No.

21

Protein

Theoretical

Experimental

pI

MW [kDa]

pI

MW [kDa]

Rho GDP dissociation inhibitor (GDI) alpha

5.0

23.2

5.3

31.2

Accession no. NCBI

Regulation/ kinetic group TrkA

TrkB

gi/4757768

none

↓C

22

AF487339 1 NM23-H1

5.2

19.7

5.8

22.1

gi/29468184

none

↓B

23

1713410A beta galactoside soluble lectin

5.1

14.6

5.3

12.8

gi/227920

none

↑C

24

not identified

5.4

24

none

↓A

25

6.8

74.1

6.2

86.6

gi/27436946

↑C

none

6.8

74.1

6.2

86.6

gi/27436946

↑A

none

5.1

51.0

5.4

70.5

gi/14165437

↓C

none

28

Lamin A/C isoform 1 precursor Lamin A/C isoform 1 precursor Heterogeneous nuclear ribonucleoprotein K isoform a Lamin B2

5.3

67.7

5.6

71.0

gi/27436951

↑A

none

29

Vimentin

7.9

54.2

5.2

54.2

gi/4557525

↓C

none

30

not identified

5.5

49.4

↓C

none

31

not identified

5.5

49.4

↓C

none

32

ACTB protein

5.5

49.4

↓C

none

33

not identified

5.6

26.4

↓C

none

26 27

5.6

40.2

gi/15277503

with a reducing agent such as tris-(carboxyethyl) phosphine hydrochloride (TCEP) (Figure 7). In some proteins cysteine residues are buried, and thus the extent of labeling will depend on the accessibility of cysteine within the protein under the reaction conditions used. Also protein quantification in the amounts and concentrations used is not easy to carry out. A labeling optimization in a 2-D titration experiment is therefore obligatory to determine the optimum amount of TCEP and dye required for the protein extract being used. The molar ratio of TCEP : dye should always be kept at 1 : 2 to ensure efficient labeling. Typically, 5 µg of protein lysate requires 2 nmol TCEP and 4 nmol dye for the labeling reaction (assuming an average cysteine content of 2%). If the amount of TCEP/dye is too low, available thiol groups on some proteins will not be labeled and show molecular weight trains in the 2-D image. If the amount of TCEP/dye is too high, nonspecificlabeling of the amine groups on lysine residues can occur and have been observed as pI charge trains on the gel. In the standard protocol, proteins are solubilized in lysis buffer (30 mM Tris, 7 M urea, 2 M thiourea, 4% (w/v) CHAPS, pH 8.0). Primary amines or thiols should not be present, because they will compete with the protein for

11


Fig. 6 Typical kinetics of regulated proteins following neurotrophin treatment of SY5Y–TrkA or SY5Y–TrkB. The majority of proteins like for instance galectin-1 were regulated in the late stimulation phase. Circles (control) and triangles (neurotrophintreated) represent single standardized abun-

dance values from one gel. Lines connect the average standardized abundance values. Though experiments were repeated five times, the DeCyder software could not detect the here presented protein spot in all gels. Thus, at some time points less than five circles or triangles are represented.

Fig. 7 Labelling chemistry of CyDye DIGE saturation fluors: dye maleimide group reacts with all available cysteine residues in the protein and due to its net zero charge there is no charge change.


the dye. After lysis, the pH should not deviate from pH 8; protein concentrations should be between 0.55 and 10 mg/mL. Then cysteine residues are reduced at 37 ◦ C for 1 h by incubating with TCEP (2 mM solution) with a volume determined in the labeling optimization experiment. CyDye DIGE Fluor saturation dye (2 mM working solution in anhydrous dimethylformamide) is added for reaction at 37 ◦ C in the dark for a further 30 min. Cy3 is used for the pooled standard and Cy5 for the individual samples. Finally, the reaction is stopped by excess of DTT using 2×lysis buffer (s.a.). 2-DE is according to standard procedure; the alcylation step in the equilibration procedure can be omitted. 2.2.2 Example of Use: Analysis of 1000 Microdissected Cells from PanIN Grades for the Identification of a New Molecular Tumor Marker Using CyDye DIGE Saturation Fluors Pancreatic adenocarcinoma is the fourth leading cause of cancer in the United States [23]. The main problem of combating pancreatic adenocarcinoma arises from a lack of specific symptoms and limitations in detection methods. The overwhelming majority of pancreatic carcinoma patients are discovered at a late clinical tumor stage. Only 10% of patients show a potentially curable resectable tumor. In order to identify new molecular markers of the pancreatic tumor progression we established a proteomics approach analyzing 1000 microdissected cells from different pancreatic intraepithelial neoplasias (PanIN) grades [24]. It is believed that PanINs are the precursor lesions of pancreatic ductal adenocarcinoma (PDAC). PanINs are histologically subdivided into four different grades, namely PanIN1A/B, PanIN-2 and PanIN-3. All PanINs progress from flat to papillary lesions with increasing degrees of dysplasia (Figure 8). To analyze the different PanIN grades we applied microdissection, through which it is possible to analyze the biologically

Fig. 8 Histological images of different PanIN grades stained with H&E. It is believed that pancreatic intraepithelial neoplasias (PanINs) are the precursor lesions of pancreatic ductal adenocarcinoma (PDAC). PanINs are histologically subdivided into four different grades, namely

PanIN-1A/B, PanIN-2 and PanIN-3. All PanINs progress from flat to papillary lesions with increasing degrees of dysplasia. The PanIN classification has been established as a common diagnostic criterion at the National Cancer Institute Think Tank (http://pathology.jhu.edu/pancreas panin).

13


Fig. 9 DIGE analysis of pancreatic ductal epithelium and PanIN-2 microdissected cells. The analysis of 1000 microdissected cells from pancreatic ductal epithelium and PanIN-2 lesions revealed a pattern of A 1.875 and B 2.050 protein spots, respec-

tively. Eight significantly regulated proteins are indicated by arrows and respective spot number. In the zoomed sections two differentially regulated proteins (white box) are displayed in 3D view obtained with the DeCyder software.

relevant cell type, often in the minority when analyzing precursor lesions. Different techniques have been developed to facilitate microdissection [7, 25, 26, 28]. To select for PanIN grades, manual microdissection was chosen because we have found it easier and speedier than automatic processing [24]). Using CyDye DIGE saturation fluors we were able to combine the high resolution power of 2-DE with the high sensitivity of fluorescence imaging for the analysis of the scarce sample amount of 1000 microdissected cells (approximately 2 µg). In this preliminary report we have shown that the established protocol can successfully be applied to the proteome analysis of PanIN-2 grade as well as pancreatic ductal epithelium (Figure 9), resulting in the identification of the first molecular markers of PanIN-2 using LC-ESI-MS/MS (Table 3). Annexin A2 and annexin A4 have been found to be differentially expressed in PanIN-2 grades. These proteins are members of the annexin family of calcium-dependent phospholipid-binding proteins showing a 45–59% identity with other members of the annexin family [12]. Although their functions are still not clearly defined, several members of the annexin family have been implicated in membrane-related events along exocytotic and endocytotic pathways [30]. Furthermore, annexin A2 was found to be significantly up-regulated in PanIN-2 grades. It has been shown that the endothelial cell-surface Ca2+ -binding protein, annexin A2, activates the t-PA-dependent formation of plasmin from plasminogen and promotes tumor cell invasion [4]. In the differential expression profiling of Lu et al., annexin 2A has also been found to be up-regulated in pancreatic carcinoma tissue [22].

2 Difference Gel Electrophoresis: Next Generation of Protein Detection in 2-DE Table 3 First candidate molecular markers for PanIN-2 grade pancreas carcinoma analyzing

1000 microdissected cells Spot No.

Spotname

Accession

Protein

Fold change

T-test

Seq. pI

1 2 3 4

1347 2437 1811 2660

gi/15277503 gi/18645167 gi/4502105 gi/5453740

5 6 7 8

820 1844 2547 823

– – gi/48255907 gi/5030431

Actin, gamma 1 4.0 0.017 5.6 Annexin A2 4.8 0.0008 7.4 Annexin A4 −4.5 0.0017 5.8 Myosin regulatory −2.1 0.00031 4.6 light chain MRCL3 Not identified −2.65 0.0068 – Not identified −13.37 0.0031 – Transgelin −4.5 0.0017 8.9 Vimentin −2.5 0.0055 4.8

Seq. MW (kDa) 40.2 38.6 36.1 19.8 – – 22.6 41.6

Furthermore, a group of three actin-associated proteins down-regulated in PanIN-2 grades were found, whereas actin itself was detected as significantly upregulated. For instance transgelin is an actin-binding protein of unknown function cross-linking actin filaments [33]. It has been shown that down-regulation of transgelin may be an important early event in tumor progression and it is considered a diagnostic marker for breast and colon cancer development [34]. The role of the other actin-associated proteins is at the moment speculative and will be analyzed in more detail when the expression profiles of all PanIN grades and carcinoma samples are available. Currently, there is very little information available concerning the initiation and prevention of pancreatic cancer. But the candidate protein identified applying DIGE saturation labeling for the analysis of PanIN grades will help to find new diagnostic markers for early detection of pancreatic cancer and to decipher processes within tumor biology. After analyzing all of the PanIN stages a widespread catalogue of protein expression in pancreatic tumor progression will be obtained, helping us to understand pancreatic tumor biology in more detail.

2.3 Statistical Aspects of Applying DIGE Proteome Analysis

For the identification of new molecular markers and interaction partners proteome analysis using 2-DE (DIGE) is often applied, but the deduced candidate proteins using the subtractive approach of a differential analysis need further validation. For reasons of time and cost, the number of candidate proteins is limited and only significant proteins changes can be considered for candidate validation. A DIGE experiment produces large amounts of data, or more exactly, volume values for thousands of gel spots. In order to make correct inferences from these data, statistical methods are quite important. Many of the statistical methods used in DNA microarray studies can be adapted for analyses of gel data. In this section we

15


focus on the calibration and normalization of protein expression data as well as on the detection of differentially expressed proteins resulting from a DIGE experiment. 2.3.1 Calibration and Normalization of Protein Expression Data From the DeCyder software it is possible to find the background-subtracted spot volumes, i.e., for a single spot the lowest 10th percentile pixel values on the spot boundary have been excluded. Several features of this raw data require calibration and normalization prior to statistical analysis. The volume values are affected by dye-specific system gains and constant additive biases. In Figure 10 the Cy5 and Cy3 spot volumes of one gel are plotted against each other. It can clearly be seen that the difference between Cy5 and Cy3 spot volumes increases when the spot volumes increase. When using m gels for a DIGE study (with internal standard, treatment and control) and regarding each image of a gel as a distinct gel, one receives a data set with j = 1, . . ., 3 m gels. To calibrate the spot volumes Karp et al. [17] proposed the following statistical model for consideration:

xi j =a j xi j +bi j

(1)

where xi j is the real expression value of the ith spot on the jth gel, xij is the respective measured spot volume, aj the scaling factors that adjust for the dye-specific system gain and bj additive offsets which compensate for the additive biases. These parameters can be estimated by maximum-likelihood estimation. The respective estimation

Fig. 10 Scatterplot of the Cy5 versus the Cy3 spot volumes of a given minimal CyDye gel.


Fig. 11 Graphs of the logarithm and the arsinh for data normalization.

algorithm and the calibration procedure have been implemented within the vsnpackage for the open-source software R (available at http://cran.r-project.org) [15]. The resulting calibrated spot volumes are lognormal-distributed. Most statistical methods, however, are based on the assumption that the data are normaldistributed. Hence, the calibrated volume values must be normalized. A common method for normalization is to apply the logarithm to the data, but the logarithm has a singularity at zero and causes a bias in the Cy3/Cy5 ratios for small values. The mentioned software package uses the arsinh instead, which for high values is equivalent to the logarithm but has no singularity at zero and its smoothness for small values does not cause biases. The graphs of the logarithm and the arsinh can be compared in Figure 11. Applying the logarithm or the arsinh to the data also has the benefit that the variance of the data is stabilized. In the raw data, high volume values usually have a different variance than small volume values. In Figure 12 the effect on data normalization using arsinh transformation is shown. 2.3.2 Detection of Differentially Expressed Proteins In DIGE experiments it is often of interest to find proteins with altered expression in two different mixtures, e.g., treatment and control. Having transformed the data by the logarithm or the arsinh one has to look for differential expression changes and not for relative changes, because ratios become differences when being transformed by the logarithm or the arsinh, i.e., because log (a/b) = log(a) − log(b). If the volume values from treatment and control probes come, for example, from the same patient or, as in DIGE experiments, from the same gel, then there is

17


Fig. 12 Lognormal-distributed volume values before normalization (left) and normal-distributed values after the arsinh-transformation (right).

a dependency between the resulting values. Hence, the t-test for paired samples, which has even a higher statistical power than the t-test for independent samples, can be used here. The term power is explained below. For the paired t-test the difference d between the treatment and control value is used. For each single spot the t-test for paired samples tests the null hypothesis H0 , that there is no expression change for this spot, versus the alternative hypothesis, H1 that there is an expression change. Based on the sample, the test decides either to accept H0 or to reject it and accept H1 . Be aware that the decision of a statistical test does not supply 100% certainty. A differentiation between the test decision and reality must always be made. In Table 4 the two kinds of errors that may occur are shown: a Type I error is to reject the null hypothesis when it is true, and a Type II error is to accept the null Table 4 Possible results of a statistical test

Test decision

Reality

Protein not differentially expressed

Protein differentially expressed

Protein not differenttially expressed

Correct decision

Wrong decision (Type I error α)

Protein differentially expressed

Wrong decision (Type II error β)

Correct decision


hypothesis when it is not true. The probability α for the Type I error is called the level of the test. The probability β for the Type II error depends on this α-level, the sample size, the expression change to be detected and the variance of the measured values. The probability of rejecting the null hypothesis is called the power of the test. It should be very small when the null hypothesis is true and very big when the alternative hypothesis is true. The higher the sample size for the experiment the better becomes the power. Unfortunately it is not possible to keep α and β small at the same time. In practice it is common to specify a small α first, also called the testing level, and determine an appropriate number m of replications to keep β as small as possible afterwards. Within the t-test the data is summarized in the test-statistic which is t-distributed under the assumption that the data for the treatment and control groups was obtained from normal distributions. The t-test is available in most statistical software packages. 2.3.3 Sample Size Determination It is not easy to directly determine a number m of replicates to use for the t-test. Instead it is more convenient to look how the power of the test behaves when using a specified number of replicates. The power of the paired t-test depends (1) on the probability a for the Type I error, (2) on the standard deviation of the

Fig. 13 Power-function of the paired t-test using three replicates (dotted line), four replicates (dashed line) and five replicates (solid line) under the assumption that the standard deviation of the spots differences is s = 0.2.

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differences between the treatment and control values, (3) on the expression change to be detected and (4) on the number m of replicates which are used. Given these four parameters the power can be plotted as function of the expression change to be detected. In Figure 3 the power function is given for different numbers of replicates. On the lower abscissa the relative expression change is given, and on the upper, the differential expression change of the normalized values. From the power one can detect the probability β of the Type II error by the relationship β = 1 − power. If in the case of Figure 13 one wants to detect a relative expression change of 1.5 then β ≈ 0.1 when using five replicates. Hence, the strategy of finding the correct sample size is to specify the above-named four parameters, plot the power function and read the probability β for the Type II error from the graph. If β is too big, plot the power function for a higher number of replicates. Within the open source software R, the power function for the paired t-test can be plotted. 2.3.4 Further Applications There are many other statistical models which can be used for the evaluation of DIGE studies. Inclusion of not only a group factor, but also a time factor in the experiment methods of the analysis of variance (ANOVA) can be applied to find expression changes within the temporal course of the protein expression or to find interactions between the group and time factor. Several multivariate statistical methods are of use, too. Spots with similar expression profiles can be grouped by cluster analysis or, on the other hand, new spots can be assigned to existing groups by the methods of discriminant analysis.

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34 SHIELDS, J. M., ROGERS-GRAHAM, K., DER, C. J. (2002). Loss of transgelin in breast and colon tumors and in RIE-1 cells by Ras deregulation of gene expression through Raf-independent pathways. J. Biol. Chem. 277, 9790–9799. 35 KARP, A. N., KREIL, D. P., LILLEY, K. S. (2004). Determining a significant change in protein expression with DeCyder during a pairwise comparison using two-dimensional difference gel electrophoresis. Proteomics 4, 1421–1432. 36 HUBER, W., HEYDEBRECK, A., VON SÜLTMANN, H., POUSTKA, A., VINGRON, M. (2002). Variance stabilization applied to microarray data calibration and to the quantification of differential expression. Bioinformatics 18, S96-S104. 37 FERNANDEZ-PATRON, C., CASTELLANOS-SERRA, L., HARDY, E., GUERRA, M., ESTEVEZ, E., MEHL, E., FRANK, R. W. (1998). Understanding the mechanism of the zinc-ion stains of biomacromolecules in electrophoresis gels: generalization of the reverse-staining technique. Electrophoresis 19, 2398–2406. 38 HEUKESHOVEN, J., DERNICK, R. (1988). Improved silver staining procedure for fast staining in PhastSystem Development Unit. I. Staining of sodium dodecyl sulfate gels. Electrophoresis 9, 28–32. 39 JOHNSTON, R. F., PICKETT, S. C., BARKER, D. L. (1990). Autoradiography using storage phosphor technology. Electrophoresis 11, 355–360. 40 NESTERENKO, M. V., TILLEY, M., UPTON, S. J. (1994). A simple modification of Blum’s silver stain method allows for 30 minute detection of proteins in polyacrylamide gels. J. Biochem. Biophys. Methods 28, 239–242. 41 NEUHOFF, V., STAMM, R., PARDOWITZ, I., AROLD, N., EHRHARDT, W., TAUBE, D. (1990). Essential problems in quantification of proteins following colloidal staining with coomassie brilliant blue dyes in polyacrylamide gels, and their solution. Electrophoresis 11, 101–117. 42 RABILLOUD, T., STRUB, J. M., LUCHE, S., van DORSSELAER, A., LUNARDI, J. (2001). A

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1

Electrophysiological Analysis of Ion Channels Michael Pusch Instituto di Biofisica, Consiglio Nazionale delle Ricerche, Genova, Italy

Originally published in: Expression and Analysis of Recombinant Ions Channels. Edited by Jeffrey J. Clare and Derek J. Trezise. Copyright ľ 2006 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31209-2 This article outlines electrophysiological methods for extracting biophysical parameters that describe two fundamental properties of ion channels: gating and permeation. The Introduction provides a broad overview of the general concepts of ion channel biophysics and the text a review of the kind of information that can be extracted from electrophysiological recordings. The later sections introduce several methods for the analysis of electrophysiological experiments on heterologously expressed ion channels. Many parts are explicit and can be directly applied “at the bench”. Other, more advanced topics (gating current measurements, singlechannel kinetic analysis) are touched upon only superficially since their application requires further background that can be found in the references.

1 Introduction

The function of ion channels is to rapidly pass – in a passive but selective manner – a large number of ions across biological membranes. This electrogenicity is exploited by excitable cells to quickly change the transmembrane voltage allowing, for example, the conduction of the action potential and the postsynaptic electrical response to chemical neurotransmission. Other cells and organelles exploit the large capacity of charge transport for ion homeostasis and transepithelial transport. For the researcher the electrogenicity is interesting and useful because it allows the measurement of ion channel functioning in “real-time”. It is possible to monitor the action of ion channels both in vivo or in appropriate simplified in vitro systems like brain slices. Intracellular electrodes directly measure the membrane voltage of individual cells while extracellular electrodes monitor cellular electrical activity indirectly. However, the recording of the physiological voltage provides little information regarding the biophysical parameters of the underlying channels, for two main reasons. First, all cells possess a complement of different ion channels and other Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Electrophysiological Analysis of Ion Channels

electrogenic transporters that make it difficult to tease apart their respective contribution to the electrical response. Second, the physiological recordings are currentclamp measurements in which the membrane voltage is “freely floating” and the concentration of neurotransmitters or other ligands is uncontrolled. In order to reliably define a physical parameter of an ion channel (ligand affinity, slope of the voltage-dependence etc.) these key variables have to be fixed, or “clamped”. A further important step is to isolate, as far as possible, a single type of ion channel. In physiological preparations this can be partially achieved using appropriate solutions and blocking agents. Heterologous systems are even better in this respect, even if they carry the risk that some physiologically important molecular component is missing. From the perspective described above the application and interpretation of the voltage-clamp measurements by Hodgkin and Huxley [2] revolutionized the approach to ion channel analysis. Today’s description and interpretation of ion channel function still draws heavily on the principles embodied in this work. One of the most important concepts for ion channels is the “gate”. A gate can be either open or shut (closed). There is no intermediate, half-open gate. At the time of Hodgkin and Huxley, there was no direct evidence for this as practically nothing was known at the level of the single channel. The assumption of a simple open or closed gate allowed a convenient description, as a two-state Markov process associated with a linear differential equation at a fixed voltage or ligand concentration. For the classical “m-gate”, the activation gate of the voltage-gated Na+ channel, this equation reads dm (t)=−βm(t)+α(1−m(t)) dt where m(t) is the probability of the gate being open at time t. The opening and closing rate constants, α and β, respectively, are voltage-dependent and represent a model for the underlying molecular rearrangement leading to gate opening. A verification of the open-closed dichotomy of most ion channels became possible with the patch clamp technique [3, 4] that allowed the real-time visualization of single channel opening and closing in almost all types of animal, plant, and bacterial cells (for simplicity here, the existence of “subconductance” states, “flickering”, and other probably ubiquitous complicated single-channel behavior is ignored). Of course, the description of gating as a two-state process is an idealization, opening is not instantaneous. Also, microscopically, a single “open” state does not exist, but rather an almost infinite number of possible molecular configurations that are macroscopically lumped together into “the open” state, because functionally, they are almost indistinguishable from each other. The transitions from closed to open (and vice versa) appear “instantaneously” on single channel recordings, but in reality take several 100 ns. However, despite the availability of several ion channel structures (see for example Refs. [5–7]) and large computing power it is not yet possible to explore channel gating by molecular dynamics. Even ion permeation that occurs on a much faster time scale cannot be fully simulated, even though some theoretical progress has been made, especially for K+ channels (see for example Ref [8]). Quantitative functional measurements are still essential for a detailed insight into the mechanisms of ion channels. It can be expected that

2 Expression Systems and Related Recording Techniques

many more structural data for ion channels will be available in the near future. The structures will guide computational studies and rational mutagenesis in order to understand the mechanisms of function at a molecular model, to obtain high affinity ligands, and possibly to exploit ion channels as molecular devices in applied technological systems. Computational predictions and structure-based hypotheses have to be tested experimentally with functional data. The present chapter aims to provide an aid to designing, analyzing and interpreting such measurements.


Each type of heterologous expression system determines a range of possible recording techniques. The most popular expression systems are Xenopus oocytes and mammalian cell lines, like HEK293 or CHO cells. A more rarely used system is the incorporation of relatively crude vesicles or purified proteins into planar lipid bilayers. 2.1 Expression in Xenopus Oocytes

The expression in Xenopus oocytes represents an extremely versatile system that allows the application of many different electrophysiological and biochemical methods [see Refs. 9–11]. Normally, in vitro transcribed cRNA is microinjected but expression can also be achieved using nuclear injection of eukaryotic expression plasmids. The oocyte system is popular because electrophysiological recordings can be easily performed by nonexperts employing the two-electrode voltage-clamp technique (TEV) [9]. This method allows a relatively high throughput compared to patch-clamp techniques and is thus often used, for example, for drug screening. A commonly underestimated problem of the TEV technique that is relevant also for qualitative measurements concerns the error introduced by the so-called series resistance (see for example Ref. [12]). The series resistance is caused by a finite conductivity of the oocyte cytoplasm, leading to a voltage drop within the cytoplasm and thus to a voltage error (see Fig. 1). Typical values of the series resistance are of the order of 0.5–1 k. Thus a current of 10 µA will cause a voltage error of the order of 5–10 mV, a value that cannot always be neglected. Furthermore, even when the series resistance error is accounted for, the TEV technique has a limited time resolution of the order of almost 1 ms in most realworld applications. The apparent time resolution can be enhanced but the oocyte nevertheless provides a nonideal space-clamp. “Fast” kinetic parameters that are derived from TEV measurements are therefore seldom comparable to the same parameters measured with the patch clamp technique. Another disadvantage of whole oocytes is that their cytoplasmic content cannot be controlled. This may lead to a significant variability of measurements from different oocytes if the channel properties depend on the cytosolic composition. Also one would often like to

3


Fig. 1 The intracellular series resistance in Xenopus oocytes. Current flowing through the interior of the oocyte leads to a voltage drop caused by the finite resistance of the cytoplasm.

change, or at least to fix, the intracellular solution. Furthermore, in the case of large expression the intracellular ion concentrations can be significantly altered by the voltage-clamp measurements. For example it is very difficult to handle a large expression (>10 µA) of the Cl− selective muscle channel CLC-1, because its kinetics depends strongly on the intracellular Cl− concentration. The disadvantages described above are partially overcome by the “cut-open” oocyte technique [13]. With this method only a small part of the surface area of the oocyte is clamped and the intracellular solution can be exchanged. However, the method is low throughput, necessitates considerable skill, and perfusion of the interior solution is very slow. Thus, this method finds a narrow range of special applications. A general problem with the expression of Ca2+ -permeable channels or channels that depend on intracellular Ca2+ is that Xenopus oocytes endogenously express a large Ca2+ dependent Cl− current, ICl− (Ca2+ ) [14, 15]. With maximal stimulation ICl− (Ca2+ ) can reach several tens of µA of current. Thus activation of Ca2+ permeable channels that leads to an influx of Ca2+ will inevitably activate this endogenous current and confound the measurements. It is also practically impossible to manipulate the intracellular Ca2+ concentration in order to study its effect on expressed channels. Nevertheless, the endogenous current can be exploited as a Ca2+ sensor to test for a possible Ca2+ permeability and also to test if the activation of (expressed) receptors and/or G-proteins results in an increase in the intracellular Ca2+ concentration (see for example Ref. [16]). One other advantage of the oocyte system is that several cRNAs coding for different subunits of an ion channel or other interacting proteins can be co-injected at defined proportions. For example dominant heterozygous genotypes of channelopathies can be simulated by a one to one co-expression of WT and mutant subunits, and possible dominant negative effects can be quantified (see for example Ref [17]). Co-expression of different proteins can also be achieved in transfected cells. However, with the oocyte injection it is easier to precisely control the relative expression of each protein.


Finally, electrical data recorded with the TEV can be correlated for the same oocyte with the surface expression of the expressed protein, using for example an introduced extracellular epitope [18, 19]. This is of particular importance for channelopathies because many disease-causing mutations are pathogenic because they effectively reduce or enhance the plasma membrane expression of the channel (see for example Refs. [18, 20]). The patch clamp technique can also be applied to Xenopus oocytes after the vitelline membrane has been removed [10]. Recordings can be performed in the cell-attached, the inside out and the outside out configuration (see Ref. [4] for a description of these methods). The electrical properties of the obtainable seal are exceptional – seal resistances >100 G can be achieved, which allow very high resolution recordings. The size of the patch pipette range from very small to “giant” [21], allowing single-channel or macroscopic recordings. Rapid solution exchanges can be applied to excised patches allowing a precise investigation of, for example, transmitter activated channels (see for example Ref. [22]). The excellent electrical properties of the cell-free patch-clamp configuration represent a significant advantage over whole cell recordings of small cells that can suffer from limited time-resolution due to the access (series) resistance (see below). 2.2 Expression in Mammalian Cells

Another popular expression system is “transfected” mammalian cell lines like HEK, CHO or many others (see Ref. [23]). The expression of one or more proteins is induced by the introduction of the DNA in an appropriate eukaryotic (often mammalian) expression vector by various chemical or physical methods. Cells can be either transiently or stably transfected. Stable transfection generally requires the integration of one or more copies of the expression construct into the genome and is initially more labor intensive than transient transfection. It is, however, convenient for long term studies on a particular channel or for drug screening where large numbers of cells may be required. Many molecular biological methods exist that increase transfection efficiency and the level of expression. Expression can also be induced with several different kinds of viruses [24]. The patch-clamp technique is the method of choice for studying the function of channels expressed in these small cells. All configurations (cell attached, whole cell, inside out, outside out) can be applied but the whole cell configuration is the most straightforward and widely used. Indeed, several different technical approaches have been taken to automate the whole cell patch-clamp for high throughput drug screening (see Ref. [25]). Several factors have to be considered for the analysis of whole cell data. The time resolution in voltage-jump experiments is limited by the time that is necessary to charge the membrane capacitance, Cm , across the access resistance, Ra , given by τ =Cm Ra

5


Typical values of Cm = 20 pF, Ra = 5 M yield a charging time constant of 0.1 ms. This is adequate for most applications but can lead to problems for very fast kinetics observed for example in voltage-gated Na+ or Cl− channels [26, 27]. The access resistance leads also to an error in the voltage-reading similar to the series resistance problem in the two-electrode voltage-clamp. For a membrane current Im the voltage error amounts to V =Im Ra which for typical values Ra = 5 M, Im = 1 nA amounts to 5 mV and becomes worse for larger currents and/or access resistances. In voltage-jump experiments with large currents and fast kinetics the two kinds of errors described combine to create a complex dynamic error that can render certain measurements uninterpretable. Most amplifiers provide an access (series) resistance compensation that compensates both kinds of errors based on an estimate of Cm and Ra . Since the compensation involves positive feedback elements it increases the noise and is prone to oscillations. Care must therefore be taken with its application (see Ref. [28]). Similar to the oocyte system cell-free patches allow a much better voltage-clamp and also faster solution changes. However, because most mammalian cells are quite small, “macroscopic” recordings are more difficult to achieve in excised patches since large pipette diameters are poorly tolerated. 2.3 Leak and Capacitance Subtraction

In heterologous expression systems unwanted currents can arise either from true leak caused by the recording electrodes or from endogenous ion channels and transporters. These have to be carefully avoided using appropriate solutions and protocols. The subtraction of currents remaining after application of a specific blocker, if available, at a saturating concentration, is a very good but often tedious method. For ligand-gated channels the subtraction of currents at zero concentration of ligand is obviously a good method, because the spontaneous open probability is very small in most cases. For voltage-gated channels studied by step-protocols the responses are additionally distorted by the capacitive transients. These can be assumed to be linear, meaning that their size is proportional to the voltage step but independent of the voltage from which the pulse is delivered. Thus smaller steps applied in a voltage range where channels are closed or steps to the reversal potential can be used to subtract capacitive transients after appropriate scaling (see for example Refs. [29, 30]). The most commonly used protocol for the subtraction of linear leak and capacity currents when measuring voltage-gated sodium and potassium channels is called the “P/4 method” [29], that is a standard feature of most data acquisition programs. For this method four small voltage pulses with a quarter of the size of the main voltage pulse are delivered before or after the main pulse. The small pulses are subthreshold and elicit exclusively leak and capacitive

3 Macroscopic Recordings

currents. The response to the four quarter-sized pulses is summed and subtracted from the main response. Linear components are therefore practically completely subtracted.


In the following sections it is assumed that the ion channel of interest is expressed in a heterologous system and represents the major contribution to the total membrane conductance. The methodologies to extract various biophysical parameters that are useful for a characterization of ion channels are explained. Single channel measurements, if recorded at a sufficient bandwidth, contain, in principle, more information than macroscopic ensemble measurements. However, they are significantly more technically demanding and the very small single channel currents for many ion channels renders their analysis virtually impossible. In addition, fast kinetics are difficult to measure at the single channel level because of the lower signal to noise ratio that has to be compensated by heavier filtering. Thus, for many applications macroscopic recordings represent the only practical approach. The most important relation regarding macroscopic currents is given by I=Ni p

(1)

that describes the total current, I, through a homogenous population of N independent channels. A basic assumption is that the channel under investigation possesses a single open state with current level i, and is without subconductance states. Of course this is an oversimplification for many channels. Without this assumption, however, a practical interpretation of macroscopic currents is almost impossible, because it is very hard to tease apart different conductance levels of a single channel from macroscopic recordings. The parameter p represents the open probability of the channel, that is the time- and/or voltage- and/or ligand-dependent probability of the channel being in a conducting state with an associated current, i. The single parameter p in Eq. (1) summarizes the combined action of all gates of the channels. Often it is useful to think of the gates as independent devices. For example the voltage-gated Na+ channel of Hodgkin and Huxley has 3 m-gates and one h-gate, all independent from each other such that the parameter p is equivalent to p = m3 h. While the independence of different gates is seldom realistic, it is very useful conceptually. Eq. (1) incarnates one of the dogmas of ion channel biophysics: permeation through the open channel, characterized by the parameter i, is independent of channel gating, characterized by the parameter p. This is a very useful conceptual distinction, although in real life it breaks down immediately: the occupancy of the pore by permeating ions generally stabilizes the protein structure and thereby influences gating. However, such effects are generally relatively small with some exceptions. For example, in CLC type Cl− -channels, permeation and gating are

7


strongly coupled [31, 32]. On the other hand, gating has practically no influence on permeation through the open pore, because, by definition “open” implies an open gate. Any influence of closed gates on ion occupancy of pore binding sites vanishes rapidly after opening the gate because the two processes occur on vastly different time scales. This assumption becomes questionable only if a rapid “flicker type” gate is present that opens and closes on a time scale more similar to that of ion conduction. For example in KvLQT1 (KCNQ1) K+ channels a rapid flicker type gate seems to be present that leads to drastic effects of Rb+ ions on the macroscopic current amplitude [33]. In this case it is difficult to distinguish between an effect of Rb+ on permeation or gating because the concept of a “gate” becomes questionable. 3.1 Analysis of Pore Properties–Permeation

Two basic parameters are important when considering permeation properties. One is ion selectivity and the other conductance. Ion selectivity is the ability to favor one ion over another and is expressed as the “permeability” ratio of the two ions, for example PK /PNa for potassium and sodium. It is, in practice, determined from the reversal potential measured in mixtures of the two ions. The simplest situation is when the ions are present at equal concentrations, one on one side of the membrane and the other on the other side – so called bi-ionic conditions. We consider the example of a cationic channel measured with 150 mM NaCl intracellular and 150 mM KCl extracellular. Then the reversal potential, E rev , is given by E rev =RT/(zF )ln(PK /PNa )

(2)

where R is the gas constant, T the absolute temperature, F the Faraday constant and z the valence (z = 1 for the example above). At room temperature the factor RT/F amounts to ∼25 mV, a value that is very useful to remember for electrophysiologists. Inverting Eq. (2) yields PK /PNa =e zF E rev /(RT ) Thus, for example, E rev = 25 mV indicates an e-fold higher permeability of K+ versus Na+ (e ∼2.718). Bi-ionic conditions are preferable but are not easily achieved in some experimental systems. For example in TEV recordings from oocytes the intracellular solution cannot be changed. In this case, the difference of the reversal potential that arises by changing from one extracellular solution to an other is measured. To obtain a permeability ratio, the Goldman-Hodgkin-Katz equation is used E rev = RT/F In((PK [K]ext + PNa [Na]ext )/(PK [K]int + PNa [Na]int ))

(3)

where [K]ext is the extracellular K+ concentration and similarly for Na+ (for anions the sign of the reversal potential has to be inverted). We consider the case where


the extracellular concentrations of Na+ and K+ are changed from [Na]0 to [Na]1 and from [K]0 to [K]1 , respectively, and assume that the measured reversal potential changes from E 0 to E 1 . From Eq. (3) it follows that the permeability ratio is given by PK /PNa =([Na]1 −[Na]0 eφ )/([K]0 eφ −[K]1 ) where φ = F(E 1 −E 0 )/(RT) ∼ (E 1 −E 0 )/(25 mV). The assumption is that the intracellular concentrations do not change. When ions of different valence are compared (for example Na+ and Cl− or Na+ and Ca2+ ) the equations change slightly [1] but the basic type of measurement remains the same. One important and often overlooked problem of reversal potential measurements is the presence of liquid junction potentials that invariably arise when a solution is exchanged for another. The liquid junction potential is caused by the different mobility of different ions and is most pronounced when small inorganic mobile ions (for example Na+ , Cl− ) are exchanged by large organic quite immobile ions (for example NMDG+ , gluconate− ; see Ref. [34] for how to determine and correct for liquid junction potentials). When the Cl− concentration is changed care must be taken because most reference electrodes are Ag/AgCl electrodes that must be shielded from the solution exchange, for example by agar bridges. It may be difficult to determine the reversal potential because the current flowing through the channel is small, such that endogenous background conductances or a leak conductance dominate the effective reversal potential. One reason for this might be that the gates are closed at the expected reversal potential. This is especially a problem for voltage-gated channels. In this case so-called tail-current analysis can be applied to determine the reversal potential and the shape of the single channel current-voltage relationship, as illustrated in Fig. 2. The currents shown in Fig. 2c were simulated based on the two-state scheme shown in Fig. 2a using the pulse-protocol illustrated in Fig. 2b. Opening and closing rate constants are exponentially voltage-dependent such that the channel closes at negative voltages and opens maximally at positive voltages. The channel was assumed to have a linear single-channel current-voltage relationship (i – V) with an imposed reversal potential of E rev = −70 mV. However, from the steady state current-voltage relationship, obtained at the end of the variable pulse (see box in Fig. 2c, squares in Fig. 2d), the reversal potential cannot be obtained, because the open probability is too low. In contrast, the initial current, “immediately” after the end of the activating voltage-step (arrow in Fig. 2c), is measurable at these negative voltages. This “instantaneous tail current” and the resulting instantaneous current–voltage relationship (Fig. 2d, circles) reflect the shape of the single channel current–voltage relationship. This can be seen from the equation Itail (V t )=Npend i(V t )

(4)

where Itail is the instantaneous current at the tail-voltage,V t , pend is the openprobability at the end of the activating prepulse, and i (V t ) is the single channel

9


Fig. 2 Tail current analysis. Macroscopic currents were simulated for the two state scheme shown in A. The reversal potential was −70 mV. From the holding potential of −80 mV a 0.2 s prepulse to 60 mV was followed by variable pulses ranging from −140 to 80 mV as illustrated in B. At negative voltages currents deactivate quickly

such that the reversal potential cannot be reliably obtained from the steady-state currents (squares in D). The initial tail currents (circles in D) faithfully reproduce the linear single-channel current–voltage relationship and allow a precise determination of the reversal potential.

current at V t . It is assumed that the open probability immediately after the voltagestep remains at the value it had at the end of the prepulse (pend ). Then, the factor N pend is independent of the tail voltage, and Itail (V t ) is proportional to i (V t ). If the purpose is only to determine the reversal potential it is not very important to get exactly the initial current, and an average over a short stretch of currents shortly after the voltage jump can be calculated. For measuring the exact shape of the single channel i–V, care must be taken to determine the “correct” initial value. This may be complicated if the deactivation is fast and the voltage jump


is associated with a large capacitance transient. In this case the time course of the deactivating current (after the capacitance transient) is fitted by a suitable function (for example an exponential function) that is then back-extrapolated to time “zero”. The determination of the permeability of a blocking ion can also be difficult. For example the muscle Cl− channel CLC-1 is blocked by iodide while iodide has a significant permeability with a permeability ratio of P1 /PCl ∼ 0.2. If extracellular Cl− is completely exchanged by iodide, however, CLC-1 is totally blocked and the reversal potential is dominated by endogenous background conductances, resulting in a wrong estimate of P1 /PCl . The problem can be solved by partially exchanging extracellular Cl− for iodide (for example change from 100 mM Cl− to a solution containing 20 mM Cl− and 80 mM iodide) leaving enough Cl− to allow for a significant conductance. Eq. (3) can then be used again to quantify the permeability ratio. 3.2 Analysis of Fast Voltage-dependent Block – the Woodhull Model

Tail current analysis is useful to analyze voltage-dependent block by fast blockers. A commonly used model to describe voltage-dependent block is the Woodhull model [1, 35]. In this model it is assumed that the charged blocking particle enters the channel pore to a certain distance, and senses therefore part of the transmembrane electric field. Block is quantified by I(c) 1 = I(0) 1+ K Dc(0) exp(zδV F /(RT ))

(5)

Here I(c) is the current in presence of blocker at concentration c, K D (0) is the dissociation constant at zero voltage, z the valence of the blocking ion and δ the “electrical” distance of the binding site from the bulk solution, that stands for the fraction of the electric field from the bulk solution to the binding site. Eq. (5) describes simultaneously the concentration and the voltage-dependence of block with just two parameters, K D (0) and δ (see Fig. 3). Another way to describe the Woodhull model is in terms of an exponentially voltage-dependent dissociation constant (Fig. 3c). The simplicity of the Woodhull model makes it attractive and it is often a good initial model. Several assumptions are, however, seldom truly satisfied. Firstly, “blocking” ions are often permeable to some extent and may “punch through” at large voltages. Also, almost all ion channels have multi-ion pores in which the ions interact. A blocking ion could displace a permeable ion present at the blocking site within the pore. Such effects add to the intrinsic voltage dependence of block (described by δ) and complicate the picture. The incorporation of such features into mechanistic models is beyond the scope of this chapter. See Ref. [1] for a comprehensive description of blocking mechanisms.

11


Fig. 3 Illustration of the Woodhull model of channel block. A linear single-channel i–V and a zero reversal potential was assumed for the unblocked channel (solid line in A). The parameters of the block were K D (0) = 1 mM and z * = 0.4. Application of increasing amounts of blocker (0.5, 1, and 5 mM) produces the increasing voltage-dependent

block dashed lines in A). The concentration dependence of block is illustrated in B for different voltages (−100, −30, 30, and +100 mV). The exponential voltagedependence of the apparent K D , determined by fitting the curves in B with a simple 1:1 binding curve is illustrated in C.

3.3 Information on Gating Properties from Macroscopic Measurements

Macroscopic currents can be used to obtain an estimate about the open-probability. According to Eq. (1) the macroscopic current is proportional to the open probability, p, but further information or assumptions about the number of channels, N, and the single-channel current, i, are necessary to estimate p from the measured current, I. The number of channels can be assumed to be constant in a typical experiment, as long as its duration is relatively short and no particular maneuvers are undertaken to enhance or decrease protein turnover. Some ion channels can be drastically affected by co-expression and acute manipulation of co-expressed or endogenous regulating proteins, like the ubiquitin-protein ligase Nedd4 [36]. The number of channels can also be nonspecifically affected by agents that lead to a general retrieval of plasma membrane. For example treating Xenopus oocytes with phorbol esters, activators of protein kinase C, leads to an unspecific reduction of expressed conductances via a reduction of the plasma membrane surface [37]. Nevertheless, in most cases, N can be regarded as fixed. If measurements are performed at a fixed voltage, as in the case of ligand activated channels or in “isochronous tail-current” measurements for voltage-gated channels (see below) the single channel current, i, is also fixed. Otherwise, the shape of the single-channel current–voltage relationship (i – V)


has to be taken into account. For the purpose of extracting information about the open probability, the i – V is parametrized in a phenomenological manner, without necessarily interpreting the corresponding parameters mechanistically. In the absence of direct information the i – V is often assumed to be linear i(V)=γ (V−Erev ) with a single-channel conductance, y, and a reversal potential, E rev . This assumption is particularly appropriate if the reversal potential is relatively close to 0 mV, because in this case the intrinsic “Goldman”-rectification [38] has little influence on the shape of the i – V. If the reversal potential is far from zero, and if the channel is highly selective for one ion species present in the solutions, the GoldmanHodgkin-Katz equation is more appropriate to describe the i – V because it takes the concentrations of the permeable ion on the two sides of the membrane into account [1]: i(V )=K φ

exp(z(φ−φr ))−1 exp(zφ)−1

(6)

Where K is a constant depending on the ionic concentrations, φ = VF/(RT) and φ r = E rev F/(RT). The nonlinear shape of the Goldman current-voltage relationship is illustrated in Fig. 4 for a monovalent cation with a reversal potential of −60 mV. It is also not uncommon that some voltage-dependent block or strong rectification is present that has to be taken into account for the description of the i – V. Such a block can be phenomenologically described by a factor that is derived from the Woodhull model. For a Goldman-type rectification with additional block the i – V is described by i(V )=K φ

1 exp(z(φ−φr ))−1 exp(zφ)−1 1+exp((V −V1 )/V2 )

(7)

where V 1 and V 2 are empirical parameters describing the block; an example is shown in Fig. 4 (dashed line).

Fig. 4. The Goldman–Hodgkin–Katz equation. The solid line is drawn according to Eq. (6) with Erev = −60 mV. The dashed line is drawn according to Eq. (7) with Erev = −60 mV and V 1 = V 2 = 50 mV.

13


Fig. 5 Determination of the open probability for voltage gated channels. Currents were simulated according to the 2-state scheme shown in Fig. 2A assuming a reversal potential of 0 mV and applying the pulse protocol shown in A. Steady state cur-

rents measured at the end of the variablevoltage pulse (see arrow in B) are plotted in C as squares. The tail currents at the beginning of the constant tail pulse to 60 mV are shown as circles.

3.3.1 Equilibrium Properties–Voltage-gated Channels Two types of pulse protocols are most often used to extract the overall-open probability from macroscopic measurements. They are illustrated in Fig. 5, together with simulated currents based on the 2-state scheme of Fig. 2. In the direct pulse protocol (Fig. 5a and b) pulses are delivered to various potentials and the maximum current during the pulse is plotted versus voltage (Fig. 5c, squares). Usually, the open-probability of voltage-gated channels is parametrized with a Boltzmann distribution, here written in two different versions

popen =

1 1 = 1+exp((V1/2 −V )/k) 1+exp(zg (V1/2 −V )F /(RT ))

(8)

In both forms the voltage of half-maximal activation,V 1/2 , describes the voltage at which popen = 0.5. The steepness of the voltage dependence is described either by the so-called “slope-factor”, k (in mV), or the “apparent gating valence” zg (dimensionless). These two quantities are inversely related by

k=

RT zg F


The Boltzmann equation derives from the statistical Boltzmann equilibrium that relates the ratio of the probability to be in one of two microscopic states, O and C that differ in free energy by a certain amount G: pO =exp(−G/(RT )) pC The Boltzmann distribution of voltage-gated channels (Eq. (8)) stems from a simple model for voltage-gated channels that assumes that the free energy difference between the open and the closed state is additively composed of an electrical term, determined by the electrical charge, denoted by Q C and Q O , respectively, and a purely chemical term, G0 , such that G is given by G=G 0 +V (QO −QC )=G 0 +V zg F where zg is the apparent gating valence. The larger the charge difference between the closed and open state, the more sensitive is the channel to voltage, and the steeper the popen (V) curve. To extract the gating component (p) from the permeation component (i) for currents obtained from the “direct” I – V (Fig. 5c, squares) the I – V is fitted by the product of the i – V term and the Boltzmann term: I(V )=K φ

1 exp(z(φ−φr ))−1 exp(zφ)−1 1+exp(zg (V1/2 −V )F /(RT ))

(9)

Here, in Eq. (9), the Goldman-Hodgkin-Katz equation (Eq. (6)) was used for a description of the i – V. The four parameters, K, φ r , V 1/2 and zg are obtained from a fit to the macroscopic data, while only the two parameters V 1/2 , and zg are relevant for gating. The tail-pulse protocol illustrated in Fig. 5a and b (see arrow) is often called “isochronous tail protocol” because the fixed tail pulse is applied after a fixed amount of time. The initial tail current is a measure of the open probability at the end of the (variable) pre-pulse (see Eq. (4)). As for the instantaneous I–V (see Section 3.1) a correct determination of the initial tail current may be hindered by the capacitive artifact. A careful back-extrapolation of the time course of the tail current to “time 0” may be necessary to obtain a reliable estimate of the initial tail current. The tail voltage should be chosen such that the relaxations are well resolved with the employed voltage-clamp technique. The resulting initial tail currents are then plotted versus the pre-pulse voltage (Fig. 5c, circles) and fitted by I(V )=

Imax 1+exp(zg (V1/2 −V )F /(RT ))

(10)

Here, Imax , is the maximal current obtained at saturating voltages. It can be determined by normalization with the measured currents or it can be left as an

15


independent parameter. The latter possibility is particularly appropriate if the employed voltage range is not sufficient to saturate channel gating. In this case the plateau of the Boltzmann distribution is not reached and currents can not be normalized by the maximally measured value. Sometimes it happens that currents do not tend to zero at voltages where the channel should close according to Eq. (8). This may be an intrinsic property of the gating mechanism of the channel or may represent an uncompensated leak component. If such a “residual” open probability is an intrinsic property, a description with a Boltzmann distribution (Eq. (8)) is, strictly speaking, not adequate. Nevertheless the shape of the popen (V) curve can often be described phenomenologically by a modified Boltzmann distribution I(V )=Imax (pmin +

1− pmin ) 1+exp(zg (V1/2 −V )F /(RT ))

Where pmin describes the minimal open probability reached at saturating voltages where channels are maximally closed. V 1/2 is no longer the voltage at which popen = 0.5 but where popen = pmin + 0.5(1–pmin ). The isochronous tail-current protocol is, in principle, superior to the direct I – V because it is not influenced by the shape of the i – V. However, in certain cases a direct method has to be employed. For example, voltage-gated Na+ channels are governed by two main gating processes of opposite voltage dependence and one wants to determine separately their respective voltage dependence. Na+ channels inactivate with a voltage-dependent time-course after an activating voltage step. The steady state, isochronous tail current i – V would determine only the “window” current, the region where activation and inactivation gates are both open. To separate the two gates of the Na+ channel two different protocols are used to assess the voltage dependence of the activation and the inactivation gate, respectively. The “peak current” of the direct I – V is generally used as a measure of the activation gate (Fig. 6a and b). This is justified because the time constant of activation is considerably faster than that of inactivation. The inactivation is measured with a two-pulse protocol similar to the isochronous tail current protocol (Fig. 6c and d). 3.3.2 Equilibrium Properties–Ligand Gated Channels Conceptually, the determination of equilibrium properties of ligand activated channels is similar to that of voltage activated channels. The energy driving the conformational change is not supplied by the membrane voltage but by the chemical energy of ligand binding. Accordingly, the relevant intensive variable is the ligand concentration, here denoted by [L]. However, for ligand activated channels the allosteric action of the ligand is much more evident and explicit than is the voltage for voltage gated channels [39] (even though most quantitative models of

I(V )=G Na (V −E rev )

1 1+exp(zg (V1/2 −V )F /(RT ))


Fig. 6 Activation and inactivation of voltage-gated sodium channels. Currents were recorded from tsA201 cells transfected with the cardiac sodium channel and measured using the whole cell configuration of the patch clamp technique. In A currents were elicited by 10 ms voltage steps from −80 to 50 mV (see inset). In B the peak currents are plotted versus the test voltage (symbols) superimposed with a fit of the equation as shown by the lines. C shows

currents from a different cell evoked by a two-pulse protocol as shown in the inset. The response to the 100 ms long prepulse to voltages from −120 to −10 mV is not shown. The currents represent the response to the fixed tail pulse to −10 mV that assays channel availability. The peak currents are plotted in D (symbols) together with a fit of Eq. (10). Note that activation (A, B) and inactivation (C, D) have an opposite voltage-dependence.

voltage gated channels often have an allosteric character). Thus instead of assuming a scheme of the form Scheme 1.

where binding of a ligand directly opens the channel, the minimal scheme applied for ligand activated channels assumes that binding of the ligand favors opening but does not directly open the channel. This scheme is given by Scheme 2.

with a closed, unliganded state U, a liganded (bound) but closed state, B, and an open state, O [40]. Ligand binding occurs with second-order association rate kon and dissociation rate koff , while the allosteric transition is described by rate constants α and β.

17


More complex schemes are necessary to include the possibility that unliganded channels are able to open [39]. Allosteric schemes and equations become even more complex if more than one binding site is present, the usual case in real life. Therefore, mostly for reasons of simplicity, the phenomenological description of ligand activated channels is often expressed in terms of the Hill equation popen =

pmax

(11)

1+( K[L]D )n

Where K D is the apparent dissociation constant of the binding site(s) and n the Hill coefficient, an estimate of the number of binding sites [1]. For the simple 2, equilibrium properties can indeed be expressed in the form of Eq. (11) popen =

1 1+ 1r + 1r K[L]D

=

r/(1+r) K* 1+ [L]D

(12)

Where r = α/β, KD = koff /kon , and the apparent affinity K D * = K D /(1 + r). The maximum open probability is pmax = r/(1 + r) = α/(α + β). Thus, even though the concentration dependence of the macroscopic current strictly follows a 1:1 binding isotherm, the measured apparent affinity can be very different from that of the true affinity of the ligand binding site. K D * is always smaller than K D and only if r is very small (α β) is the apparent affinity equal to the true affinity. But in this case the ligand is not very effective (pmax 1). If r is very large, the ligand is very effective (pmax ∼ 1) but the apparent affinity is much higher than the true affinity (K D * K D ). Since the absolute open probability is difficult to determine from macroscopic equilibrium measurements alone, additional information is necessary to determine true affinities and ligand efficacies. These can stem from kinetic macroscopic measurements, noise analysis, or single-channel analysis (see below). A fundamental difference between voltage gated channels and ligand gated channels is that the latter can be more or less efficiently activated by different types of ligands (for example certain glutamate receptors can be activated also by NMDA), while there is only one stimulus (voltage) for voltage gated channels. In terms of the simple model shown by Scheme 2, quantified by Eq. (12), different ligands have generally a different (true) affinity, and a different efficacy (pmax ). For ligands that occupy the “same” binding sites the true number of binding sites is the same. The Hill coefficient in Eq. (11) may nevertheless be different, since Eq. (11) is an approximate phenomenological description of channel activation. Certain ligands might also counteract the effect of a more potent activator. Furthermore, certain receptors possess different kinds of binding sites. For example certain glutamate receptors need glycine as a co-factor for full activation by glutamate [41]. A good overview over equilibrium properties of ligand gated ion channels with further reference is given by [39].


Many ligand gated channels exhibit desensitization: currents decrease despite the continuous presence of ligand. The degree and kinetics of desensitization vary wildly between different channel types (see Ref. [42] and references therein). This phenomenon is conceptually similar to the “inactivation” of voltage gated channels. The presence of desensitization complicates the determination of activation properties. Experimentally, the most difficult problem, particularly if desensitization is an issue, is the fast application of the ligand (see Section 2.2). 3.3.3 Macroscopic Kinetics Channel kinetics can be evoked by a variety of stimuli. Sometimes, biophysical analysis alone is not sufficient to determine the physiological effect in a complex cellular system, as for example a cardiac myocyte, where numerous channel types contribute to the various phases of the action potential. In such cases, a “physiological” stimulation with an action potential waveform or other stimuli might reveal if, for example, a given mutation leads to action potential shortening (see for example Ref. [43]). However, such physiological stimuli are not well suited to uncovering the underlying mechanistic effect. As outlined in the Introduction, for this purpose clamping the relevant intensive physical parameter (voltage, ligand concentration, temperature, pressure, light intensity,) to a fixed value and performing jump experiments are more informative. Like practically all kinds of conformational changes of proteins, current relaxations of a homogenous population of channels induced by a step-wise change of an intensive parameter, can be described by the sum of a constant term (the steady-state current, I∞ ) and one or more exponential functions

I(t)=I∞ +

n

ai exp(−t/τi )

(13)

i=1

With amplitudes, ai , and time constants, τ i (t is the time after the jump). The kinetics is thus determined by 2n+1 parameters. The time constants, τ i , depend only on the actual value of the relevant physical parameter (the voltage or the ligand concentration after the jump), while the coefficients, ai , depend on the state occupancy before the jump. The exponential time dependence is a mathematical consequence from the Markov property of the conformational changes: once the channel undergoes a conformational change it loses “immediately” the memory of from what state it arrived (if there is more than one possibility to arrive in a certain state) and it has no memory of how long it has already been in a given state. One of the underlying assumptions is, of course, that there exists a finite set of definable, stable “states”. Actually, it is more scientific to turn the argument around: the experimental result that relaxation kinetics for most channels can be well described by Eq. (13) (with a reasonably small and reproducible number, n, of components) suggests that the Markov assumption is valid for ion channels. What is a reasonably small number, n? This is indeed a difficult question. In principle, a gating scheme with N states predicts exponential relaxation with N-1 components (for example a two-state system has single-exponential kinetics). In practice it is very difficult to reliably fit more than two or three exponential components. However,

19


gating schemes often require more than four states. Indeed it is extremely difficult to define a “correct” gating scheme based on fitting of current relaxations. Often gating schemes that have a large number of states can be simplified with symmetry arguments and the number of free parameters can be reduced. A beautiful example are the Hodgkin–Huxley equations (see Ref. [1]), but also recent models of Shaker K+ channels have a relatively small number of parameters, despite a large number of states (see Refs. [44, 45]). Furthermore, current relaxations are often approximately single- or double exponential under certain conditions, even though a full kinetic model requires many states, because most components are negligible. For example the deactivation of Na+ channels at very negative voltages after a brief activating pulse can be well described by a single exponential, even though the general gating involves very many states. Often, kinetics of ion channels is fitted and time constants are determined phenomenologically without necessarily wanting to define a molecular mechanism of gating. In many circumstances, this is the only practical choice, because the underlying mechanism is too complex to be determined reliably from the measurements. One of the most difficult problems in curve fitting with exponentials is to separate components with time constants that differ less than, let us say three-fold. Extreme care has to be taken in such cases and reproducibility has to be tested extensively. Often data can be reasonably well described with the sum of two exponentials, even though the “true” mechanism would require at least three. In such cases, the time constants (and relative weights) of the exponential components determined from the double-exponential fit can be almost meaningless. If the true time course is distorted slightly, for example by voltage-clamp errors (see above), the kinetic analysis becomes even more difficult. Under certain conditions it is impossible to directly follow the kinetics of a process measuring the ionic current. For example, the channel may be closed quickly by one kind of gate while another gate is slowly changing its status. Another reason that renders impossible a direct measurement might be that the voltage is close to the reversal potential or that a strong block occurs. Also, a large capacitive artifact may obscure fast relaxations [27]. In these cases so-called “envelope” protocols can often be applied, as illustrated in Fig. 7 that shows the classical protocol to study the recovery from inactivation of the voltage gated Na+ channel or, completely analogously it illustrates the measurement of the recovery from desensitization of a ligand gated channel. It is insightful to explicitly consider the kinetics of the most simple, two state system, because often more complex schemes can also be simplified to it, allowing an easy quantitative description. Scheme 3.

Opening occurs with rate-constant α, closing with rate constant β. These rate constants depend on the intensive physical parameters (voltage, ligand concentration). The equilibrium open probability is α p∞ = α+β


Fig. 7 Envelope protocol to study recovery from inactivation. The pulse protocol is illustrated in A, current traces are shown in B. Currents during the recovery period are not shown. In C the peak current at the fi-

nal test pulse is plotted versus the recovery time together with a single exponential fit. Currents were simulated with a simplified Hodgkin–Huxley model.

While the relaxation time constant is given by τ=

1 α+β

Knowing, both p and τ allows the determination of α and β: α= p∞ /τ ;β=(1− p∞ )/τ Relaxations that start from a given value of open probability, p0 , proceed in time as α= p∞ +( p0 − p∞ )exp(−t/τ )

The Del Castillo-Katz model for ligand gated channels (Scheme 2) can be reduced to an effective two-state system if the ligand binding/unbinding is much faster than the opening isomerization transition (described by α and β). In this case the receptor is always in equilibrium with the ligand (see Ref. [1]): Scheme 4.

21


Where the effective opening rate is given by αeff =

1 1+ K[L]D

α

Combining steady state-measurements (Eq. (12)) and relaxation measurements with step-changes in ligand concentration allows the determination of all three parameters (K D , α, β) of the model of Scheme 4: for jumps into zero concentration of ligand, the relaxation rate is τ −1 = because α eff = 0 for [L] = 0. For jumps into saturating concentration of ligand the relaxation rate is τ −1 = α + β because α eff = α for [L] KD . These kinetic experiments thus provide estimates for α and β. From the equilibrium measurements the apparent affinity, K D * is determined (Eq. (12)) and using the values for α and β the true affinity, K D , can be calculated from K D = K D * (1 + α/β). This simple example illustrates how the combined use of equilibrium and kinetic measurements in addition to simplifying assumptions can be used to obtain quantitative information about a molecular mechanism.

3.4 Channel Block

All ion channels interact with a variety of smaller molecules (peptides, small organic molecules) that directly or indirectly reduce ion permeation. Such substances are called blockers or inhibitors and are the bread and butter of the pharmaceutical industry interested in ion channel targets. Blockers may directly block the pore and physically impede ion flow. Inhibitors may reduce current flow by stabilizing the closed state of a channel gate, for example, in which case the binding site may be far away from the ionic pore. Such inhibitors are often called “gating modifiers”. This distinction between these two kinds of modulators is actually not so strict - many pore blockers additionally alter the gating by binding more tightly to one or another gating state (state-dependent block). Such blockers may be useful tools to study the properties of channels [46, 47]. Many pore blockers exert a voltage-dependent block that can often be described by the Woodhull model (see Section 3.2). For most practical purposes channel block is quantified by the Hill equation I([B]) 1 = I(0) 1+( K[B] )n D

that quantifies the ratio of current in the presence of blocker at concentration [B] to the current in its absence with an apparent affinity K D and the Hill coefficient, n.


3.5 Nonstationary Noise Analysis

The opening and closing of ion channels is a random process that renders current registrations “noisy”, in particular if few channels are present. The statistical properties of the noise can be used to infer some characteristics of the underlying elementary events. A prerequisite is that the channel-induced noise is significantly above the background noise of the recording system. This condition is, for example, generally not fulfilled in TEV recordings from Xenopus oocytes. The patch-clamp technique, on the other hand, is exceptionally well suited for noise analysis. However, background noise may be large if the series resistance in whole cell recordings is highly compensated (see Section 2.2). For stationary noise analysis the system is recorded for a prolonged time at fixed external conditions (voltage, ligand concentration). The power spectrum is then fitted with the sum of Lorentzian functions. While this method can yield important information [48], nonstationary noise analysis is often faster and easier to perform. Under appropriate conditions, each of the elementary parameters of Eq. (1) can be determined assuming a single open conductance level. This should be verified with single channel analysis. For standard nonstationary noise analysis a step-protocol (that may be a voltagestep or stepwise change in ligand concentration) is applied repeatedly, with enough time passing between individual stimulations to ensure identical initial conditions for each step. Each current response, Ii (t), is recorded (i = 1, . . ., n) (Fig. 8a). From these recordings the mean can be calculated by =

n 1 Ii (t) n i=1

(14)

This is now a much smoother curve than the individual traces (Fig. 8b) and because of this it can be written a =Nip(t) where p(t) is the time course of the (“true”) open probability. The variance of the response, σ 2 (t), for each time point, t, is given by σ 2 (t)=

n 1 (Ii (t)−)2 n i=1

(15)

the standard statistical definition (Fig. 8c). However, the most important “trick” in nonstationary noise analysis is to calculate the variance not as suggested by Eq. (15) but as the squared difference of consecutive records: 1 (Ii+1 (t)−Ii (t))2 2(n−1) i=1 n−1

σ 2 (t)=

(16)

23


Fig. 8 Nonstationary noise analysis. Currents were repeatedly evoked by a test pulse and individual responses are shown in A (currents were simulated with a simplified Hodgkin–Huxley model). The mean and the

variance are shown in B and C, respectively. In D the (binned) variance is plotted versus the respective mean together with a fit of Eq. (18). The horizontal line in D marks the level of the background variance.

(Note the scaling by a factor of 1/2 in Eq. (16)). For a perfectly stable system the results obtained from Eq. (15) and Eq. (16) are identical. However, in reality, small drifts of the total current amplitude (“run-down”, “run-up”) or of the reversal potential or other parameters are practically unavoidable. These small drifts are well cancelled out using Eq. (16) while they artificially increase the variance when calculated by Eq. (16) and may render the noise analysis meaningless, particularly if the single channel conductance is small [49]. Meaningful results can be obtained even with run-down up to a factor of two or more. Since the variance is obtained from the differences of records, leak currents and capacity transients also cancel out (if they are associated with negligible inherent noise). That means that the individual records do not have to be, and should not be, leak-corrected for the application of Eq. (16), in contrast to the calculation of the mean (Eq. (14)). A nice property of the variance is that independent noise source adds independently to it. Thus, total variance is given by 2 2 2 =σchannel +σbackground σtot


and the background variance can usually be assumed to be independent of voltage. It can thus be determined at a voltage where no ion current flows through the channels (for example at the reversal potential or in the absence of ligand for ligand activated channels or at a voltage where channels are closed for voltage gated channels). Having determined the variance and the mean it is now possible to proceed with the “variance-mean analysis”. For this we use the equality σ 2 =Ni2 p(1− p)

(17)

This fundamental equation [48, 49] can be understood intuitively: for p = 0 the variance is null because no current flows. Also for p = 1 (the maximum value it can attain) there is no fluctuation in the current because all channels are permanently open. The largest fluctuations are present for p = 0.5 when channels are half open and half closed on average. Note that (Eq. (17)) is independent of the kinetics of the fluctuation. However, the bandwidth of the recording system must be sufficiently large to resolve the fastest transitions. Combining Eq. (17) with Eq. (1) yields σ 2 =i I−I 2 /N

(18)

that relates the two macroscopic quantities, σ 2 , and mean current I, in a parabolic function that depends on two parameters, the single channel current i, and the number of channels, N, that can be determined by a least-squares fit (Fig. 8d). Often the whole traces are not plotted and fitted against each other but they are first binned in an appropriate manner [49] (see Fig. 8d). The two parameters, i and N, are best defined if a large interval of popen is covered by the relaxation. If only a very small interval of popen is sampled, the two parameters cannot be determined independently: a small variance can be caused by a small number of channels and/or by a small or large popen and vice versa. It may be that the current excursion in the relaxation is substantial but that popen remains significantly smaller than 0.5. In this case the second term in Eq. (18) is negligible, the relationship is linear, and only the single channel current can be determined. If both, i and N, can be determined the absolute popen during the relaxation can be calculated from p(t)=

I(t) Ni

Thus, using a simple experimental protocol (Fig. 8) allows a quite precise determination of fundamental channel parameters, without the need to perform single channel analysis. 3.6 Gating Current Measurements in Voltage Gated Channels

Another way of obtaining additional information about molecular gating mechanisms is to measure the so-called “gating currents” associated with the molecular rearrangements of voltage gated cation channels [50]. These are transient currents,

25


similar to capacitive currents, that reflect the movement of the gating charges within the electric field. Of course, any conformational rearrangement of the channel protein that is associated with charge redistribution gives rise to “transient” gating currents, even if they do not reflect the movement of a “voltage sensor”. However, in voltage gated K+ , Na+ and Ca2+ channels the voltage-sensor movements clearly dominate the gating currents. In order to resolve gating currents that are of small magnitude it is necessary to eliminate the normal ionic currents by applying blockers or by eliminating permeant ions. However, it is necessary to ensure that such maneuvers of completely eliminating ion flow through the pore does not alter significantly the gating process itself, an often difficult task. It is beyond the scope of this chapter to describe the design and the analysis of gating current measurements (see Ref. [51] for review). However, the estimation of the total gating charge of a single voltage gated K+ channel provides a nice example of this approach [52]. The authors determined first the number of channels, N, in an inside out patch using nonstationary noise analysis (see Section 3.5). Then they replaced intracellular K+ with TEA+ , a blocker of K+ channels, to eliminate the ionic currents and measured the total gating charge, Q, by integrating the gating currents. The ratio Q/N, the gating charge per channel, was about 12 elementary charges, consistent with more indirect measurements. Another nice result was obtained by Conti and Stühmer [53], who estimated the size of the charge of a single voltage sensor in Na+ channels. Voltage gated cation channels possess four voltage sensors that move more or less independently of each other. The movement of each sensor produces a spike-like tiny current. The ensemble of many sensors is the random superposition of many such spikes, filtered at the recording frequency. Nonstationary noise analysis yielded an elementary charge of individual spikes of about 2.3 elementary charges, a very reasonable value.

4 Single Channel Analysis

The possibility to observe and analyze the opening and closing of single ion channel molecules marked a revolution in ion channel research and remains one of the few techniques that allow a true single molecule measurement in real time [3, 4]. Single channel recordings can provide a wealth of information and numerous numerical methods for single channel analysis have been developed. The reader is referred to “Single channel Recording” by Sakmann and Neher for detailed information [54]. Here a very broad overview of a typical single channel analysis is provided. Recent papers utilizing more advanced techniques can also be found [55, 56]. 4.1 Amplitude Histogram Analysis

The first step of a single channel analysis is usually to construct an amplitude histogram. Already at this point one ever returning aspect of single channel analysis


becomes important: adequate filtering. Of course the data should be filtered with a good filter (for example an 8-pole Bessel filter) with a cut-off of at least at one half the sample frequencies to avoid aliasing [57]. In order to get, at least in principle, as much information as possible, the sampling rate must be sufficiently high. It is however useless to acquire data at a low signal-to-noise ratio. A dilemma is sometimes that one would like to see online the highly filtered data in order to get an immediate impression of its quality, but one also wants to acquire at a higher frequency for quantitative offline analysis. One possibility is to divert the signal after a primary anti-alias filter into two separately sampled channels. One signal is acquired after only the first filter, while in the second the current signal is subjected to further filtering for immediate inspection. This can be done on an oscilloscope, or, if the acquiring software allows the simultaneous sampling of two channels, the highly filtered signal can be acquired as a second input channel and visualized on the computer screen. For the construction of the amplitude histogram all sample points that fall in a given “current-bin” are counted, resulting in the number of events per bin that are then plotted versus the mid-point of the current bin. This is illustrated in Fig. 9. To each current level of the channel (“closed” and “open” in Fig. 9) corresponds a “peak” in the amplitude histogram. The peaks may not be well separated because noise is large (Fig. 9b). In this case the data can be digitally filtered by a Gaussian filter that has various convenient properties [57] (Fig. 9c). If the baseline is not stable the amplitude histogram becomes distorted. Excessive baseline drift can make the single channel analysis very difficult and must be corrected. Several analysis programs are available that allow baseline correction and many other features. Once an acceptable amplitude histogram has been constructed it is fitted with the sum of Gaussian functions, Gi , one for each peak, i H(I)=

G i (I)=

i

ai (I−µi )2 exp(− ) σi 2σi2 i

(19)

Where each Gaussian functions is characterized by a mean µi a width, σ i , and amplitude, ai . The inclusion of the width, σ i , in the prefactor ai /σ i in the Gaussian fit (Eq. (19)) facilitates the calculation of the relative area, Ai , that is occupied by each Gaussian component: ai Ai =100% j

aj

The relative area, Ai , is a measure of the probability to dwell in the conductance state associated with mean µi . Often it happens that the membrane patch contains an unknown number, N, of identical channels, leading to equidistant peaks of the amplitude histogram at levels ni (n = 0, 1, . . .), where i is the amplitude of a single channel. Even though the absolute open probability cannot easily be determined in such a situation a useful parameter to evaluate effects of drug application or

27


Fig. 9 Amplitude histogram analysis. Currents were simulated based on a simple 2-state scheme with an open conductance level of 1 pA and an added Gaussian noise of 0.4 pA SD (panel A). The baseline and the open conductance level are indicated by horizontal lines. The noisy trace in A seems to be almost useless. However, the amplitude histogram shown in B clearly shows

two peaks at the right amplitudes (0 pA and 1 pA), and the fit with the sum of two Gaussian functions (thick line in B) correctly predicts the amplitude and area of each conductance level. The trace shown in C is strongly filtered and the histogram in D shows two well separated peaks of correct amplitude and weight. The bin width used for the histograms was 10f.A.

other maneuvers is the so-called “NpO ”, i.e. the product of the (unknown) number of channels and their open probability. From the histogram fit the “NpO ” can be calculated as ∞

Np O =

j =0 ∞

jAj Aj

j=0

where the “areas” Aj are obtained from the Gaussian fits as described above, and A0 corresponds to the baseline. 4.2 Kinetic Single Channel Analysis

A comprehensive description of the kinetic analysis of single channel data is beyond the limits of this chapter and only general directions can be given. The strategic decisions that can be taken for kinetic analysis are outlined schematically in Fig. 10.


Fig. 10 Flow diagram of strategies for kinetic single channel analysis.

The most direct way of analysis is depicted on the left of the figure and consists of directly fitting a “hidden Markov model” to the “raw” data [58–60]. A Markov model is a kinetic scheme like those described above (for example, Scheme 2) with possibly many states of various conductance levels connected by rate constants. The rate constants and the conductance levels of the various states are the parameters fitted in this approach. The model is “hidden” for two reasons. First, several kinetic states may be associated with the same conductance. Transitions among these states are therefore not directly visible. Second, the noise can hide shortlived dwell times or low conductance states. One advantage of the hidden Markov model approach is that it takes the noise into account explicitly [59, 60]. Algorithmically, for the hidden Markov model the following question is raised: for a given set of parameters (these parameters include the rate constants of the model, the conductance level of each state, and parameters that describe the noise) what is the probability of observing the currents that have been measured? The parameters of the model and the characteristics of the noise are then adapted to maximize this probability.

29


The calculation of this probability is a formidable task but efficient algorithms have been developed that allow the analysis of quite long data sets. One drawback of the method is that a reasonable kinetic model should a priori be known. The method can be applied for example to study the effect of mutations of an ion channel for which a kinetic scheme has been established previously. Another drawback is that the method is a kind of black box with little possibility of visual evaluation to check if the results are “reasonable”. A further serious problem may occur if the general properties of the noise are not adequately treated [60]. Thus, while the hidden Markov modeling can be a powerful tool, its use requires some experience and results should be checked with other methods. The more traditional methods of analysis do not work directly on the raw data trace but this is first “idealized” (Fig. 11). In the idealization process the events of channel opening and closing are detected either completely automatically or interactively using specially designed computer programs. The noisy raw data trace is thus substituted by the idealized smooth trace (Fig. 11) that can be easily represented as a list with two numbers for each entry in the list: the duration of each dwelling together with the corresponding current level. For idealization one must decide if a current fluctuation represents a transition to another conductance level or if it is just noise. As a criterion the 50% level criterion is most often employed [57]. To apply this criterion, the possible conductance levels are estimated first by the fitting of the amplitude histogram (see above). In addition, events are only accepted if they are of a minimal length. For a reliable assignment of the transitions data usually have to be filtered more than for the hidden Markov approach, at least if the signal to noise level is low. The basic problem in the idealization process is that short events are easily missed (the missed events problem) but short events can also be artifactually introduced if noise is excessive. The problem is double: missing a short closure not only leads to the loss of a closed event but it also lengthens the opening time of the event during which the closure occurred. Similarly, the artifactual introduction of a short closure not only alters the closed times but also shortens, and divides into two the underlying opening. To deal with this problem in generality is not easy. First, a reasonable cut-off is defined such that no events shorter than this are rigorously accepted and the resulting error in the final fitting procedure can then be compensated [56, 61]. For simplicity, we ignore here the missed events problem. The idealized trace can then be analyzed in several ways. A first step is to construct and inspect dwell-time histograms. Several kinds of binning procedures and histograms to construct have been proposed (see for example [62]). In Fig. 11 so-called cumulative dwell time histograms for the two conductance levels are displayed. These histograms can then be fitted with the sum exponential components in order to extract kinetic information from the single channel data. The construction and fitting of histograms is always a first step in data analysis if little is known about the underlying channel and if one wants to obtain an impression about its kinetic behavior, like for example an estimate of the number of open and closed states. Histogram fitting may also provide a purely empirical set of parameters whose variation under the influence of ligands or mutation can be

5 Conclusion

Fig. 11 Dwell-time analysis. In A a short stretch of a simulated trace of a two-state scheme is shown. In B the corresponding idealized trace is displayed. The cumulative dwell-time histograms shown in C and D are based on a total of 408 events each and represent the relative frequency of events to be longer than a given duration. By def-

inition the cumulative distribution equals 1 for a duration of 0. A single exponential dwell-time distribution is represented by a straight line in the logarithmic scaling of Fig. 11. Other representations of dwell-time histograms are probably more common [62] but require a much larger number of events for a satisfactory graphical display.

studied. If a good working hypothesis for a kinetic Markov model for the channel is available it may instead be a good idea to fit directly the likelihood of the idealized channel trace. This approach is similar to the hidden Markov approach described above in that the full kinetic information including possible correlations are exploited. The maximum-likelihood fitting overcomes one of the biggest problems of histogram fitting: it is not clear how the time constants and coefficients extracted from closed and open time histograms have to be weighted in fitting a concrete Markov scheme. In the maximum-likelihood approach the rate constants defining the Markov scheme are directly optimized [56, 61]. Recordings obtained under different conditions can also be fitted simultaneously. This approach thus allows on the one hand an objective estimate of physical parameters similar to the hidden Markov fitting and on the other hand the results can be easily judged visually by comparing the predictions for all kinds of dwell-time histograms [56].

5 Conclusion

This chapter provides a broad overview of current concepts and methods of analysis of electrophysiological data. Several of the methods are incorporated into the free analysis program written by the author that is available for down-

31


load at http://www.ge.cnr.it/ICB/conti moran pusch/programs-pusch/softwaremik.-htm. The simulation program used to generate several of the figures can also be found there.

Acknowledgements

I thank Armando Carpaneto for critically reading the manuscript. The financial support by Telethon Italy (grant GGP04018) and the Italian Research Ministry (FIRB RBAU01PJMS) is gratefully acknowledged.

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43 G. BERECKI et al., HERG Channel (Dys)-function Revealed by Dynamic Action Potential Clamp Technique, Biophys. J. 2005, 88, 566–578. 44 W. N. ZAGOTTA, T. HOSHI, R. W. ALDRICH, Shaker potassium channel gating. III: Evaluation of kinetic models for activation, J. Gen. Physiol. 1994, 103, 321–362. 45 J. ZHENG, F. J. SIGWORTH, Selectivity Changes during Activation of Mutant Shaker Potassium Channels, J. Gen. Physiol. 1997, 110, 101–117. 46 E. D. KOCH, B. M. OLIVERA, H. TERLAU, F. CONTI, The Binding of {kappa}-Conotoxin PVIIA and Fast C-Type Inactivation of Shaker K+ Channels are Mutually Exclusive, Biophys. J. 2004, 86, 191–209. 47 M. PUSCH et al., Mechanism of block of single protopores of the Torpedo chloride channel ClC-0 by 2-(p-chlorophenoxy)butyric acid (CPB), J. Gen. Physiol. 2001, 118, 45–62. 48 F. CONTI, Noise analysis and single-channel recordings, Curr. Top. Membranes Transport 1984, 22, 371–405. 49 S. H. HEINEMANN, F. CONTI, Nonstationary noise analysis and application to patch clamp recordings, Methods Enzymol. 1992, 207, 131–148. 50 C. M. ARMSTRONG, F. BEZANILLA, Currents related to movement of the gating particles of the sodium channels, Nature 1973, 242, 459–461. 51 F. BEZANILLA, The voltage sensor in voltage-dependent ion channels, Physiol. Rev. 2000, 80, 555–592. 52 N. E. SCHOPPA, K. MCCORMACK, M. A. TANOUYE, F. J. SIGWORTH, The size of gating charge in wild-type and mutant Shaker potassium channels, Science 1992, 255, 1712–1715. 53 F. CONTI, W. STU¨ HMER, Quantal charge redistributions accompanying the

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1

Ion Channel Assays Using Fluorescent Probes Jesús E. González, and Jennings Worley III and Fredrick Van Goor Vertex Pharmaceuticals, Inc., San Diego, USA

Originally published in: Expression and Analysis of Recombinant Ions Channels. Edited by Jeffrey J. Clare and Derek J. Trezise. Copyright ľ 2006 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31209-2

1 Introduction

Ion channels are critical for physiological signaling and are the targets of several drugs. Most of these drugs were discovered using in vivo pharmacology models or directly from observation in humans and only later was it determined that their mechanism of action involved modulation of ion channel activity. Ion channels are widely recognized as “druggable” targets due to their modulation by a wide diversity of small molecules. Despite their promise, these targets have historically been difficult to pursue because of limited structural information, low expression levels, requirement of a membrane environment for proper folding and pharmacology, and limitations or absence of high-throughput screening methods. In the last 15 years drug discovery approaches have increasingly relied on high throughput methods to profile a larger number of candidate compounds in increasing numbers of in vitro assays aimed at providing insight into which molecules will be more likely to be efficacious, safe, and able to be administered in humans. Assays for primary targets, safety counter-screens, chemical properties, and in vitro metabolism are key components of modern discovery processes and approaches. Ion channel assays play as an important role and are particularly challenging as they generally require cellular expression. Fluorescence readouts are commonly use for cell-based assays because of their high sensitivity, compatibility with readily available instrumentation and microtiter plates, and the availability of a variety of probes and reagents [1]. The application of physiological indicators of intracellular calcium and membrane potential has made possible functional fluorescence based ion channels assays with the requisite throughput, sensitivity, and reliability required for large scale profiling. In this chapter we will review the range of fluorescence probes, approaches and concepts Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Ion Channel Assays Using Fluorescent Probes

for assaying activity for sodium, calcium, potassium, and chloride channels, including those activated and regulated by ligands and voltage. Within each class of channels the utility and challenges will be discussed. Where appropriate, relevant reviews will be cited and illustrative examples will be given. Finally, we will provide examples and describe areas of where novel fluorescence assays are being developed, with an emphasis on drug discovery applications and new approaches for measuring ion channel trafficking.

2 Membrane Potential Probes

Fluorescent probes of cellular membrane potential were initially developed as physiological indicators and are divided into three major categories. The first includes fast electrochromic probes, such as Di-4-ANEPPS, that are capable of detecting microsecond voltage changes yet have low voltage sensitivity (∼1–10% F/F o per 100 mV) [2]. The second class is comprised of environment-sensitive dyes that distribute between cellular sites and extracellular solution according to the membrane potential. These probes have good sensitivity (1–100% F/F o per 100 mV) yet relatively slow time resolution, requiring minutes to attain maximal fluorescence response [2]. In the last five years new redistribution dyes have been introduced by Molecular Devices that respond substantially faster, in tens of seconds instead of minutes [3, 4]. The third category involves probes based on fluorescence resonance energy transfer (FRET) readout of rapid transmembrane translocation of fluorescent hydrophobic ions [5, 6]. These probes have good sensitivity (20–150% F/F o per 100 mV) and respond much more rapidly than redistribution dyes since the temporal response results from facile transmembrane redistribution of a mobile voltage-sensitive ionic dye and not slow diffusion across multiple membrane/water interfaces. Ion channels in cells throughout the body open and close in microto milliseconds to produce rapid, < 5 s, changes in membrane potential which are important for cell signaling and are not, in many cases, readily detected with slow redistribution dyes. For this reason most applications requiring the highest temporal resolution, such as neuronal signaling, require electrochromic or FRET dyes. Redistribution and FRET probes are generally used for most ion channel drug discovery applications because of their relatively high sensitivity and ease of use. 2.1 Redistribution Probes

Fluorescent redistribution probes are environment-sensitive charged dyes that equilibrate between intracellular hydrophobic sites and extracellular solution according to membrane potential. Fig. 1A is a schematic illustrating the redistribution mechanism and Fig. 1B shows a fluorescence micrograph of cells stained with the oxonol DiSBAC4 (3). Generally, the fluorescence quantum yield is high when the probe binds to hydrophobic sites such as proteins and membranes and very low when


Fig. 1 Oxonol probes that operate via redistribution. (A) Schematic of voltage-sensitive redistribution mechanism. Negativelycharged oxonol molecules (small circles) distribute into a cell (large circles) according to the membrane potential V m . The dye fluorescence is greatly enhanced, represented by the small filled circles, when bound to hydrophobic intracellular protein and membrane sites. At less negative po-

tentials, represented on the right panel, more dye accumulates in the cell and the total fluorescence increases. (B) A fluoresence micrograph of CHO-K1 cells stained with DiSBAC4 (3) showing extensive intracellular staining. The extracellular solution is relatively dark because the quantum efficiency in aqueous solution is neglible. (C) Structures of bis-barbiturate trimethine oxonols.

in aqueous solution. Consequently positively charged probes such as cyanines and rhodamines result in bright fluorescence at negative membrane potentials and are relatively dim at less negative or positive potentials. The opposite is true for negatively charged probes such as oxonol dyes, which make it more challenging to detect significant oxonol fluorescence from cells that hyperpolarize to very negative potentials. Anions translocate across the membranes and into cells more easily than cations because of the dipole that originates from the lipid carbonyl groups. The rate difference due to charge can be orders of magnitude, as has been shown for the isostructural borates and phosphonium ions [7]. This is an important reason why negatively charged probes, such as oxonols, are routinely used and positively charge probes are not. The structures of commonly used bisbarbiturate oxonol dyes are shown in Fig. 1C. One of the first fluorescent assays specifically designed for identifying ion channel modulators used the redistribution oxonol probe DiBAC4 (3) to develop a higher throughput assay for identifying openers of KATP channels [8]. This work played a major role in the development of the fluorometric imaging plate reader (FLIPR) [9]. DiBAC4 (3)’s high sensitivity to temperature and excitation and emission properties were key considerations in the original FLIPR design specifications. Key issues with DiBAC4 (3) are temperature sensitivity, slow temporal response, interference from fluorescent compounds and relatively high false positive rate. Even at 37 ◦ C, the time response often requires 15 min or more to reach the maximal fluorescence

3

4 Ion Channel Assays Using Fluorescent Probes

change. Despite these limitations, DiBAC4 (3) has been broadly applied and has also been used with a microfluidic device, in combination with a cationic dye, to measure ion channel activity in human T lymphocytes [10]. Recently Molecular Devices have introduced a new FLIPR membrane potential (FMP) kit that includes a redistribution dye that offers advantages in temporal response and reduced fluorescence interference compared to DiBAC4 (3). The probe has fluorescence properties similar to known thiobarbiturate oxonols and has redistribution kinetics much faster than DiBAC4 (3) with maximal response achieved in approximately 10–15 s. The kit also includes a second quenching dye that reduces extracellular fluorescence and improves signal to background and potentially reduces the effect of some fluorescent compounds. Another FMP kit advantage for HTS applications is that the dyes can be added directly to cells and do not require additional washing steps, streamlining the process and resulting in greater throughput. 2.2 FRET Probes

Membrane potential sensors based on FRET are useful for high-throughput screening (HTS) of ion channel targets [11–13]. They are comprised of two fluorescent components. The first is a highly fluorescent, hydrophobic ion that binds to the plasma membrane and “senses” the transmembrane electric field. The ion sensor rapidly redistributes between two binding sites on opposite sides of the membrane, establishing a Nernstian equilibrium (10-fold concentration ratio for ∼60 mV). In response to a membrane potential change, the hydrophobic ions electrodiffuse across the membrane and establish a new equilibrium corresponding to the new membrane potential. The voltage-dependent redistribution is converted into a ratiometric fluorescent readout with a second fluorescent molecule that binds specifically to one face of the plasma membrane and functions as a FRET partner to the mobile voltage-sensing ion. A schematic of the mechanism is shown in Fig. 2A. A variety of fluorescent membrane-bound molecules have been designed to function as voltage-sensitive FRET partners with different voltage sensitivities, temporal responses, and wavelengths. In the most commonly used configuration, the two fluorescent dye components are a ChloroCoumarin-labeled phospholipid (e.g. CC1-DMPE and CC2-DMPE) and a bis-(1,3-dialkylthiobarbituric acid) trimethine oxonol (DiSBACn (3)), where n corresponds to the number of carbon atoms in the n-alkyl group, shown in Fig. 2B. Both are very bright fluorophores, the properties are shown in Table 1. CC1/2DMPE selectively partitions into the outer leaflet of the plasma membrane and acts as a fixed FRET donor to the mobile voltage-sensitive and negatively charged oxonol acceptor. CC1/2-DMPE does not cross the bilayer because of two negative charges from the coumarin and the phosphate groups. Cells have negative (inside) resting membrane potentials and under these conditions the majority of the negatively charge oxonols populate the relatively positive extracellular leaflet, resulting in efficient FRET (i.e. quenching of the coumarin


Fig. 2 Voltage-sensitive FRET probes. (A) Schematic of voltage-sensitive FRETmechanism. Fluorescent donor molecules bind selectively to the extracellular leaflet of the plasma membrane, represented by a circle with hatching. Negatively charged acceptor, bold circles, distribute across the plasma membrane according to the membrane potential in a Nernstian manner. At negative resting potentials the acceptors

are predominantly at the extracellular surface and FRET is efficient, as shown on the left panel. Upon depolarization, the transmembrane acceptor equilibrium changes so that more oxonols are at the intracellular side. This causes a decrease in FRET and results in an increase and decrease in donor and acceptor fluorescence, respectively. (B) Structures of FRET donor CC1-DMPE and thiobarbiturate oxonols.

donor and increase in the oxonol acceptor emission). As illustrated in Fig. 2A, depolarization causes translocation of the oxonol to the inner surface of the plasma membrane and an increase in the mean donor and acceptor distance. Because FRET is very sensitive to donor–acceptor distance, this charge movement causes a simultaneous increase and decrease in the CC1/2-DMPE and oxonol fluorescence, respectively. The donor and acceptor fluorescence emission changes are reversed upon repolarization. The oxonol moves reversibly in the membrane with sub-second kinetics, allowing voltage-sensitive FRET to report fast voltage changes. Simultaneous patch-clamp and rapid optical recording have been used to demonstrate the speed, sensitivity and ratiometric nature of voltage-sensitive FRET in cells [5, 6]. The coumarin donor to oxonol acceptor fluorescence emission ratio is independent of the excitation intensity, the number of cells being detected, and the optical path length, providing fewer experimental artifacts compared to single

5

6 Ion Channel Assays Using Fluorescent Probes Table 1 Fluorescence and sensitivity properties of FRET membrane potential probes

Donor CC1/2-DMPE Acceptors DiSBAC2 (3) DiSBAC4 (3) DiSBAC6 (3) DiSBAC2 (5) DiSBAC4 (5) DiSBAC6 (5)

kex (nm)

kem (nm)

Tc (ms)

e (M−1 cm−1 )

Q.Y.

V m sensitivity %R per mV

405

460

na

40 000

1.0

na

540 540 540 640 640 640

560 560 560 660 660 660

500 20 2 50 2 0.40

200 000 200 000 200 000 225 000 225 000 225 000

0.44 0.44 0.44 0.67 0.67 0.67

1–3 0.6–1 0.4–0.8 0.5–2 pH 7.0, or Tris-maleate), and then put in contact for 2–4 h at 37 ◦ C with the biological sample to be analyzed. a Coloration observed after reaction with a solution of Fast Blue BB (N-(4-amino-2,5-diethoxyphenyl)benzamide) in 25% aqueous Tris-HCl containing 10% weight of lauryl sulfate. b Control with biological sample only.

1 Introduction 7

Fig. 4 Principle of PHENOZYM microorganism profiling. The cocktail contains 16 labeled enzyme substrates (see Table 2). Data processing involves determination of percentage conversion of each substrate in the positive (culture) and negative (medium alone) assay.

activity, and interpreted in terms of the presence/absence of the corresponding enzymes [19]. Newer, more exhaustive variations on this theme have been developed. The current format includes 32 different assays and comes in miniaturized, preformated plates [20]. The APIZYM system demonstrates that enzyme activity data allow a functional classification of microorganisms. A more compact version of the APIZYM can be realized in a single assay using a cocktail containing different enzyme substrates, called PHENOZYM (Figure 4 and Table 2) [21]. This format is applicable to extremophilic microorganisms under their optimal growth conditions. Labeled substrates for 16 different enzyme types are combined in this cocktail. The assay involves a single reaction followed by determination of substrate consumption by high-performance liquid chromatography (HPLC) analysis. This allows a rapid identification of multiple enzyme activities, and is compatible with a diversity of growth media and reaction conditions (pH, temperature), including those for thermophilic microorganisms. The functional profiles can be used for a functional classification of the different microbial strains,

8 Enzyme Activity Fingerprinting Methods for Hydrolases Table 2 The PHENOZYM cocktail for profiling thermophilic microorganisms

No. Enzyme 1 2 3 4 5 6 7 8

Phosphatase Amylase β-Galactosidase α-Galactosidase β-Glucosidase β-Glucuronidase N-Acetyl-β-glucosaminidase α-Glucosidase

9 10

α-Mannosidase α-Fucosidase

11 12 13 14 15 16

Valine aminopeptidase Leucine aminopeptidase Chymotrypsin Trypsin Esterase C4 Lipase C8

Substrate

tR

4-NP-phosphate 4-NP-α-D-hexa-(1,4)-glucopyranoside) 4-NP-β-D-galactopyranoside 4-NP-α-D-galactopyranoside 4-NP-β-D-glucopyranoside 4-NP-β-D-glucuronide 4-NP-N-acetyl-β-D-glucosaminide 4-MU-α-D-glucopyranoside 4-Methylumbelliferone 4-MU-α-D-mannopyranoside 4-NP-α-L-fucopyranoside 4-Nitrophenol 4-Nitroaniline L-Valine-paranitroanilide L-Leucine-paranitroanilide L-Phenylalanine-4-nitroanilide N-α-Benzoyl-α-L-arginine-4-nitroanilide 5-(4-Nitrophenoxy)-2-hydroxy-pentyl butanoate 3-(Umbelliferyl)-2-methyl-2-hydroxypropyl octanoate

3.94 7.49 8.98 9.50 10.41 11.53 12.39 13.13 17.38 15.49 19.83 20.12 20.35 23.44 24.83 26.12 27.33 40.23 43.85

a

Analysis conditions: Vydac 218TP54 RP-C18 column, 0.4×22 cm, elution 1.5 mL min−1 , gradient water-acetonitrile +0.1% TFA, detection by UV at 300 nm. b NP: nitrophenyl; MV: methylumbellifenyl. c This substrate gave mostly nitrophenol. Intermediates corresponding to hydrolysis between the glucosyl units were also detected in small amounts. d The 1,2-diol hydrolysis products were not detected and are apparently further degraded under the assay conditions.

resulting in a “phylo-enzymatic” tree consistent with the known phylogenetic classification of the strains. 2 Hydrolase Fingerprinting

The APIZYM and PHENOZYM analyses identify a microorganism on the basis of the enzyme panel it expresses. The principle of enzyme fingerprinting also extends to identifying single enzymes on the basis of their activity profiles against a series of substrates. Substrates for fingerprinting should be sufficiently reactive to return an activity with most enzymes within an enzyme class, yet show differential reactivity to distinguish between different enzymes. The activity detection should also be simple and reproducible to facilitate analysis. 2.1 Fingerprinting with Fluorogenic and Chromogenic Substrates

Fluorogenic and chromogenic substrates provide a very direct, selective, and flexible tool to design enzyme-specific assays. Optimally such substrates react only in

2 Hydrolase Fingerprinting 9

Fig. 5 Fluorogenic and chromogenic substrates using secondary $elimination. The bonds cleaved by the primary enzyme being assayed are marked as wobble lines.

contact with the targeted enzyme or enzyme class, while being unreactive otherwise. The indirect release of a fluorescent or colored phenolate (X− ) by β-elimination from a ketone or aldehyde intermediate provides a general strategy for fluorogenic and chromogenic substrates for a broad variety of enzymes (Figure 5). This principle is suitable for the assay of alcohol dehydrogenases (1) [22], and to measure retro-aldolization in catalytic antibodies (e.g. 2) [23] and transaldolases (e.g. 3) [24]. Similar catalytic antibody assays have been reported, releasing either resorufin from 4 [25], or bromonaphthol from 5 [26]. In the latter case reaction of the released bromonaphthol with brilliant blue forms an insoluble blue precipitate suitable for direct detection in agar plates. Assays using a similar β-elimination strategy to release a fluorescent or colored phenolate have been reported for lipases (6) [27], transketolase (7) [28], Baeyer-Villiger monooxygenases (8) [29], β-lactamase (9) [30], and a fluorescent probe for NADPH and NADH (10) [31]. Indirect release of umbelliferone (7hydroxycoumarin) is also possible via an intermediate hemi-acetal (Figure 6), which leads to assays for lipases and esterases (e.g. 11) [32], lactonases (e.g. 12) and BaeyerVilliger monooxygenases (e.g. 13) [33]. The β-elimination can also be coupled with periodate oxidation of an intermediate 1,2-diol or 1,2-aminoalcohol, thereby allowing the assay of enzymes releasing these as primary products (Figure 7) [34]. The strategy is suitable for lipases and esterases

Fig. 6 Indirect release fluorogenic substrates. The bonds cleaved by the primary enzyme being assayed are marked as wobble lines.

Fig. 7 Fluorogenic and chromogenic substrates for periodate-coupled enzyme assays with $-elimination.


Fig. 8 Naphthaldehyde release assays.

(e.g. 14–15) [35], epoxide hydrolases (e.g. 16) [36], phosphatases (e.g. 17) [37], and acylases and proteases (e.g. 18) [38]. In a similar manner release of the fluorescent 6-methoxy-naphthaldehyde (20) from alcohol precursors can be used in a blue fluorescent assay for alcohol dehydrogenase (19) [39], aldolase antibodies and proline (21) [25], or via the periodate cleavage for esterases or lipases (22), and epoxide hydrolases (23) (Figure 8) [1]. Such substrates do not show any significant background reaction in the absence of specific enzymes, and can be prepared in many variations by simple synthetic steps. A series of mono-acetates, diacetates, cyclic carbonates, and epoxides derived from optically pure versions of fluorogenic and chromogenic 1,2-diols related to 14–16 were prepared and used for fingerprinting hydrolases [1]. The fingerprinting experiment involved incubation of a single enzyme with all the different substrates in parallel in the same microtiter plate, and following the evolution of fluorescence or color simultaneously for all substrates. This fingerprinting procedure for the measurement is repeated at will for each enzyme, and provides reproducible activity profiles across the entire array of substrates. The key advantages of this array are the negligible background reaction without enzyme, the common chromophore/fluorophore release chemistry and the simple signal acquisition using a microtiter plate reader. The array of fluorogenic monoacyl-glycerol analogs shown in Figure 9 was used to analyze the acyl chain length selectivity of lipases and esterases, as discussed below [40]. Esters of nitrophenol and umbelliferone are often used as reference substrate to define activity units of esterases and lipases. These substrates usually show a high level of nonspecific hydrolysis in the absence of enzyme, and therefore cannot be used for fingerprinting. We have recently found that the problem of nonspecific hydrolysis can be circumvented in the case of umbelliferyl esters by using the substrates in a solid-supported format [41]. Umbelliferyl esters (32–47) and oxymethyl ethers (48–51) (Figure 10) can be adsorbed onto silica gel plates

Fig. 9 Lipase fingerprinting using a series of fluorogenic monoacylglycerol equivalents with different acyl chain length. The data from this array are shown in Figure 19.

Fig. 10 Umbelliferyl esters used for fingerprinting on solid support.


Fig. 11 Principle of solid-supported assay.

from dichloromethane solutions to form a homogeneously impregnated layer. The silica gel plates thus prepared are then treated with the enzyme-containing solution in buffer (Figure 11). Under these conditions only active enzymes induce a fluorogenic hydrolysis reaction, while buffer and noncatalytic proteins have no effect. This solid-supported format is advantageous for fingerprinting because only small volumes of enzyme solution are required (1 µL per assay), and simple esters of umbelliferone are prepared in a single synthetic step, or are directly available commercially.

Procedure 1: Esterase/lipase Fingerprinting on Impregnated Silicagel Plates General procedure for the synthesis of esters 32a/b-47a/b: A solution of umbelliferone or 4-methylumbelliferone (1.23 mmol) in anhydrous tetrahydrofuran (THF) (3 mL) was treated with NaH (118 mg, 55% suspension in oil, 2.3 equiv.). After 30 min at 25 ◦ C, the reaction was cooled to 0 ◦ C and the acyl chloride (1.85 mmol, 1.5 equiv.) was added as a solution in dry THF (1 mL). After 2 h at 25 ◦ C, the reaction mixture was poured into aqueous 1 M HCl (50 mL) and extracted with CH2 Cl2 (2x50 mL). The organic phase was dried over Na2 SO4 , evaporated and the residue was purified by flash chromatography to give the pure esters. Fingerprinting: Silica gel thin-layer chromatography (TLC) glass-plates sil G-25 (Macherey-Nagel, Dueren, Germany) (silica gel 40–63 µm, surface 550 m2 g−1 , layer 0.25 mm) were soaked in a solution of substrate (32a/b–51a/b, 2 mM in CH2 Cl2 ) for 2 min and dried. Aliquots (1 µL) of enzyme samples in phosphate-buffered saline (PBS, 10 mM phosphate, 160 mM NaCl, pH 7.4, with 30% v/v glycerol) were spotted at 25 ◦ C. Product formation was recorded after 2–24 h using a fluorescence microtiter plate reader (λex = 360nm, λem = 460nm).

14 Enzyme Activity Fingerprinting Methods for Hydrolases

2.2 Fingerprinting with Indirect Chromogenic Assays

The use of a chromogenic or fluorogenic signal allows multiple quantitative analyses in microtiter plates. However the use of tagged substrates limits the range of structures available to compose the substrate array for fingerprinting. In addition, many chromogenic and fluorogenic substrates are not available commercially, so that the composition of a substrate array for fingerprinting requires a large synthetic effort. A practical alternative consists in using an indirect chromogenic or fluorogenic assay that returns a signal upon reaction progress with an untagged, commercially available substrate. Multisubstrate profiling of lipases and esterases has been realized using a colorimetric assay based on pH indicators, as described by Kazlauskas et al. [42]. Hydrolysis of an ester substrate releases a carboxylic acid, which results in a drop of pH in the assay solution. This pH change can be detected colorimetrically with nitrophenolate as indicator (yellow to colorless upon decreasing pH). In the so-called Quick E variation [43], the reaction rates of separate enantiomers are determined by the pH method in the presence of a competing resorufin ester substrate. Activity profiling with this method across an array of substrates was used to describe the substrate range of four esterases. Indirect fingerprinting of lipases and esterases has also been demonstrated with an assay using the principle of back titration with adrenaline (52) (Figure 12) [44]. In this assay a periodate-resistant substrate, for example a phosphate of carboxylic ester of a 1,2-diol or its epoxide precursor, is converted to a periodate-sensitive reaction product, that is a 1,2-diol, upon enzymatic transformation, that is hydrolysis by an esterase or an epoxide hydrolase. The test solution is treated with a measured amount of sodium periodate, which reacts with the oxidizable functional groups in the products. The unreacted periodate reagent is then revealed by addition

Fig. 12 Indirect chromogenic endpoint assay using back titration of periodate with adrenaline.


of adrenaline (52), which undergoes an instantaneous oxidation with periodate to give adrenochrome (53), a cationic orthoquinone dye with a red absorption maximum in the visible spectrum. This colorimetric assay provides off-the-shelf endpoint assays for lipases using vegetal oils as substrates, phytases using phytic acid as substrate, and epoxide hydrolases using epoxides as substrates. In the latter case, simply plating out a series of epoxides (54a-z) provides an array suitable for fingerprinting of epoxide hydrolase reactivity (Figure 13). The assay can be adapted for fingerprinting hydrolases using an array of acetates (55–88) derived from commercially available polyols such as carbohydrates and glycols (Figure 14) [45]. The resulting activity fingerprints were used to identify unusual reactivities in newly discovered microbial esterases. A related indirect assay based on chelation of copper by a fluorescent chelate or an amino acid is available for screening proteases and peptidases with their natural substrates [46], and could form the basis for similar fingerprinting arrays.

Procedure 2: Lipase/esterase Fingerprinting Using the Adrenaline Test for Enzymes

Substrates were diluted from 10 mM stock solutions in water/acetonitrile mixtures. Enzymes were diluted from 10 mg mL−1 stock solutions in aqueous borate buffer (50 mM, pH 8.0). NaIO4 was added as a freshly prepared 10 mM stock solution in water. Adrenaline (as HCl salt) was added as a 15 mM stock solution in water. Assays (0.1 mL) were conducted in individual wells of 96-well flat-bottom half-area polystyrene microtiter plates (Costar, Cambridge, MA, USA). Conditions: (1) Enzyme in 50 mM aqueous borate pH 8.0, 1 mM NaIO4 , 1 mM substrate, 60 min at 37 ◦ C; (2) 1.5 mM adrenaline, 5 min, 26 ◦ C. The optical density (OD) was recorded using a Spectramax 190 Microplate Spectrophotometer (Molecular Devices, Ismaning/München, Germany λ = 490 nm). Commercial enzyme samples were tested at 1 mg mL−1 , proprietary esterases and lipases samples were used at one-tenth of crude enzyme extracts filtered through size-exclusion chromatography columns.

2.3 Cocktail Fingerprinting

Recording good fingerprinting data requires a simple yet reliable assay procedure. Optimally the reagent should be easy to assemble and use, and it should be possible to check its composition at any time. In addition, fingerprinting should be based on readily available instruments and adaptable to a broad range of reaction conditions, such as to be available to any laboratory working with enzymes. A remarkably straightforward solution to this problem is realized in the form of substrate cocktails, which are mixtures of enzyme substrates chosen such that either the substrates, or the reaction products, or both, are easily separable, selectively detectable, and precisely quantifiable after reaction.


Fig. 13 Array of epoxides used for fingerprinting epoxide hydrolases using the assay described.

The principle of cocktail fingerprinting was first demonstrated with a cocktail of 20 different lipase substrates tagged with a strong UV chromophore, and such that the reaction products are separated by HPLC analysis [47]. The cocktail composition can be checked at any time by HPLC or any spectroscopic analysis method, and the assay involves only a single measurement, which ensures that the relative product distribution is completely conserved for two measurements under identical conditions. The cocktail reagent was used to record the activity fingerprints of 40 different lipases and esterases. All substrates were mono-octanoyl esters of 1,2-diol, which ensured that most substrates showed significant reactivity with the enzymes. Thus, the cocktail ensured activity detection of all enzymes tested, while returning sufficient different reactivity information to uniquely identify each enzyme by a particular reactivity pattern. Cocktail fingerprinting is also suitable for the analysis of protease reactivities [48]. In this case the challenge consists in reducing the enormous diversity of possible peptide sequences to a limited yet representative set of peptides spanning a sufficient range for ensuring broad and differentiated reactivity with various proteases while being separable by HPLC. A series of five hexapeptides including all 25 possible dipeptides between (1) acidic, (2) basic, (3) hydrophobic, (4) aromatic, and (5) small and polar amino acids was realized following a domino-game assembly (Figures 15 and 16). The peptides were fluorescence labeled at the N-terminus, resulting in 31 possible measurable fragments, including the five substrates and


Fig. 14 Array of polyol acetates for lipase/esterase fingerprinting.

Fig. 15 Cocktail fingerprinting of protease with N-terminal labeled hexapeptide substrates. Sequences are selected by domino design.

Fig. 16 Domino design of peptide cocktail. All 25 possible types of dipeptide (upper right) serve as domino pieces to assemble five hexapeptides. Only one possible solution of the domino game is shown and realized as an actual sequence (lower

right). Note that the domino solution selected also realizes 16 of 25 possible 1,3arrangements of amino acid types. The missing 1,3-arrangements are AXA, HXA, HXH, PXP, PXS, NXP, NXN, SXA, SXS.

3 Classification from Fingerprinting Data 19

their N-terminal cleavage products. The resulting cocktail reagent reacts with a broad range of proteases, and returns a specific cleavage pattern in each case (Figure 18).

3 Classification from Fingerprinting Data

The fingerprints provided by the methods above return enzyme-specific or samplespecific activity patterns. The information that can be extracted from such patterns depends on data accuracy, the variability between different samples, and the actual structure of the substrates. In general the fingerprint data consist either of a series of initial reaction rates measured for each substrate in the presence of the enzyme, or of a series of percentages of conversion observed for each substrate after a fixed incubation time. The first step in processing fingerprinting data is to extract this reaction rate information from the primary signal recorded following appropriate calibration curves with reference products. Transformation of the recorded signal to a chemically significant value such as a reaction rate or percentage conversion is not a formal requirement if one is only interested in enzyme similarity by multivariate analysis (see below). 3.1 Fingerprint Representation

A fingerprint data set can be represented in a visual format to compare multiple enzymes and substrates simultaneously. This is done conveniently using grayscale or multicolor array displays associating each square in an array with an enzyme–substrate combination identifiable by its coordinates. The color scale should be adapted for each fingerprint such that the full color scale is used to span reactivity scales from zero to the maximum reaction rate or conversion observed in the fingerprint. Rectangular grids representing the substrate array associated with an enzyme are a convenient option for visual representation. Such grids can be created directly from a data table by conversion to the portable gray map (.pgm) or portable pixel map (.ppm) file formats. Grayscales are suitable for representation of a single data point per square in the array. The representative series of gray-scale fingerprints in Figure 17 was obtained with the carbohydrate acetates array in Figure 14 using the indirect adrenaline test described in Figure 12. The advantage of the array layout is that similarities between enzymes can be quickly assessed by a visual inspection of the data. Such comparisons are not possible if the data are represented as table numbers. One can also represent a series of fingerprints in a combined array where each line corresponds to a different enzyme fingerprint, and each column corresponds to a different substrate in the array. Such combined arrays can be integrated into a table, giving additional information about enzymes and substrates, and the different fingerprints can be ordered according to data clustering as discussed below. The clustered data set of protease cleavage patterns derived from the domino cocktail


Fig. 17 Activity of esterases and lipases towards polyacetates in Figure 14 as determined by the adrenaline test. Each grayscale square corresponds to one ester substrate according to the layout, from white (0%, no activity) to black (100%, maximum signal), after correction from blank values.

is shown as a representative example of such data display (Figure 18). The contrast in these grayscales can be augmented by using multicolor scales as available from many graphic programs, for example the blue–green–yellow–red scale used for coding temperature ranges. One of the most useful multicolor scales combines color shading with color intensity by using gradual desaturation of blue, green, and red channels, which translates into a white–yellow–orange–red–black color scale with increasing data values. Selectivities between pairs of substrates (enantiomers, stereoisomers, or analogs) can be represented using a two-dimensional color code [1]. In this format one square in the array represents data for the substrate pair by associating one color channel to the first substrate, a second color channel to the second substrate, and the third color channel to the average value of the first two channels. Complete selectivity for either substrate appears as a pure color, and the absence of selectivity


Fig. 18 Grayscale-coded protease fingerprints from domino-peptide cocktail. The relative distribution of products is shown by coloring the P1 position of cleavage of each detected N-terminal coumarinlabeled fragment relative to the most abundant fragment detected (shown in black) using the indicated scale. Unreacted substrates are not color coded. Proteases are ordered according to the hier-

archical clustering shown at right (Ward’s clustering using squared Euclidean distances with product percentages as variables. The variables were not normalized). * is the (7-coumaryl)oxyacetyl label and the capital letters are standard amino acid codes. C-termini of the peptide substrates are CONH2 (peptide 1, 3, 5) or COOH (peptide 2, 4).

(equal rates for each of the two substrates) appears as gray. This color scale is very convenient for visual analysis. Taking into account the occurrence of colorblindness in red–green contrast, the best choice of colors is to use the green (G) and blue (B) channel of the RGB code for the substrate values, and the red channel (R) for the average, thus producing a purple-to-green contrast map. An example of color-coded representation of a data set is shown in Figure 19, showing the data set of lipase and esterase fingerprints obtained against the array of enantiomeric fluorogenic monoacyl glycerol analogs in Figure 9. Procedure 3: Data Treatment for Representation of Colored Selectivity Arrays

The file format used to generate the colored selectivity arrays is the portable pixel map (.ppm) format. Each grid position is first assigned three whole numbers corresponding to the RGB color code between 0 (zero intensity, maximum activity) and 255 (maximum intensity, no activity) as follows: the first number is set according to the activity observed with the (R)-enantiomer (or the first of two given stereoisomers), the second number according to the activity observed with the (S)-enantiomer (or second stereoisomer), and the third number is simply the mathematical average of the first two numbers. Thus a grid with X columns and Y lines is coded with 3X columns and Y lines of whole numbers between 0 and 255. The grid of numbers is saved as


Fig. 19 Lipase/esterase fingerprints and hierarchical tree. Each line represents a different enzyme, and each column represents a different substrate. Each colored square represents the reaction rates of two enantiomeric lipase substrates (measured

separately) relative to the maximum rate observed with the corresponding enzyme (line), which is given in pM s−1 (color key at lower right). Hierarchical agglomerative clustering was carried out using Ward’s method on the basis of Euclidean distances.

comma separated value (.csv) file. This file is then opened in a text editor and the following three (or four) lines are inserted at the top of the file: P3 # (optional line with identifier) XY 255 where X is the number of columns in the array and Y the number of lines in the array. The file is then saved from the text editor in the portable pixel map format by simply adding the “.ppm” suffix. This file is opened by imageprocessing software and can be resized and saved in a different format (e.g. bmp). In this format it is possible to permutate the order of the three columns corresponding to the RGB code, and to obtain the color shading orange-to-blue (when G is the average of R and B) and a green-to-purple (when R is the average of G and B, as shown in Figure 19).


3.2 Data Normalization

Data processing should be kept to a minimum, paying attention to staying as close to the actual chemistry taking place in the flask as possible. Raw output data recorded during the experiment are expressed in relative fluorescence units in the case of fluorescence assays, or peak areas when processing HPLC spectra. Such primary data are already suitable for similarity analysis between substrates and between enzymes without any manipulation. Nevertheless one usually converts this primary output into chemically meaningful information such as substrate conversions (%) or reaction rates (pM s−1 ) by means of a calibration curve. With or without such conversion, the unspecific signal recorded in blank assays without enzyme should be subtracted from the primary signal. If the different substrates in the array display very different chemical reactivities, the rate data can be converted to relative rate accelerations over background to report the enzyme-specific contribution only. If there is no observable background reaction, one can calculate the relative rate in relation to a reference nonselective chemical catalyst (Eq. 1). Reporting the enzyme-specific rate acceleration rather than the absolute reaction rate allows a stronger statistical weight to be assigned to unreactive substrates, such as esters of hindered alcohols, where even a small conversion by the enzyme is noteworthy. V(i)rel =V(i)obs /V(i)ref

(1)

where V(i)obs is the observed reaction rate or conversion of substrate i; V(i)ref is the reaction rate or conversion of substrate i with a reference nonselective chemical catalyst; V(i)rel is the relative rate acceleration of substrate i over reference. For the purpose of enzyme similarity analysis, we usually normalize the data such that the sum of all reaction rates or conversions observed with a given enzyme across the different substrates in the array is set to a constant value of 1 or 100% (Eq. 2). In this manner two samples of the same enzyme at two different concentrations or at two different reaction times should appear similar. This normalization enables to conserve the actual ratio between each substrate across the total enzyme reactivity against the array. This allows one to compare biocatalysts independently of their concentration in the test samples. Such a selectivity analysis circumvents the need to define enzyme activity units for each enzyme sample being analyzed. The lipase selectivity data set (Figure 19) was normalized using this method.

V(i)rel =V(i)obs /

n

V(i)

(2)

i=1

n V(i) is the sum of the reaction rates of all the substrates for a given where i=1 enzyme.


Although we have not tested all possible data treatment approaches, we found that the following two data correction methods must be avoided in fingerprint data analysis. First, reporting the data for each substrate relative to the maximum rate or conversion observed in the fingerprint, as is done for the color-coded representation, provides unreliable results in terms of statistical comparisons, probably due to the automatic propagation of any error relating to this maximum peak (Eq. 3). Second, the standard variable normalization methods in multivariate analysis, which consists in transforming each variable such that its average is 0 and its standard deviation is 1, must be avoided because all substrates receive equal statistical weight, in particular substrates with very low conversion where the observed small variations in reaction rate or conversion reflect measurement inaccuracies rather than actual data. V(i)rel =V(i)/Vmax

(3)

where Vmax is the maximum reaction rate observed for an enzyme over the array. Correction of rate data by nonlinear mathematical operations (e.g. logarithmic scales) can be considered when the different reaction rates or conversions comprising the fingerprint span several orders of magnitude yet are all reliably measured. This correction should be considered after normalization of all observed rates of conversion to a constant sum as discussed above, such as to preserve a data set independent of catalyst concentration. 3.3 Hierarchical Clustering of Enzyme Fingerprints

In multivariate analysis, each enzyme is considered as an observation and positioned as a point in a multidimensional space whose dimensions correspond to the reaction rates observed with each substrate in the array, defined as variables (Figure 20). Similarity analysis is based on comparing distances between different enzymes (observations) in this multidimensional space or the different substrates (variables) [49]. This comparison depends on the relative weight assigned to the variables as discussed in the previous section. Hierarchical clustering is an automated procedure for grouping different observations consisting in a set of measured variables according to their similarity (geometrical proximity in the variable space). A number of distance measures and algorithms can be chosen for clustering. For the case of enzyme fingerprints, we have used the agglomerative technique based on Ward’s method which is the most commonly used in statistics [50]. We employed Euclidean and squared Euclidean distances as a measure of similarity. The latter is actually recommended [51], but both resulted in the formation of chemically meaningful clusters. A representative example is the data set in Figure 19. The classification proposed by Ward clustering correctly represents the intuitive reactivity pattern analysis operated visually, and groups esterases operating on short-chain acyl groups at the beginning of the list,


Fig. 20 Multivariate analysis of enzyme rate data. Each substrate defines a dimension, and the enzyme is positioned in the n-dimensional space using the observed reaction rate or conversion with each substrate as coordinate. Hierarchical cluster analysis compares distances in this n-dimensional space.

and lipases active on long-chain acyl groups at the end of this list. The cluster structure is shown in the hierarchical tree. Principal component analysis (PCA) reduces the dimensionality by combining correlated variables linearly into one factor whilst maximizing the variance captured in this factor. For example, PCA identifies if a set of points in a three-dimensional space are grouped in the same plane (two dimensions) or along the same line (one dimension) and determines the corresponding axes, with coefficients for each variable. In most data sets more than 60% of the total variance is explained by the first two principal components, allowing a 2D representation of the intrinsic structure of the clusters. For example, the lipase and esterase data set discussed above can be represented in such a 2D plot, giving a more graphical layout to the data (Figure 21). 3.4 Analysis of Substrate Similarities

The chemical composition of the substrate series composing a fingerprinting array is central to the analysis method, and the structures of these substrates contain the chemical essence of the measurement. As pointed out above, fingerprints based on a limited set of substrates must incorporate substrates displaying a significant reactivity with the targeted enzyme class, such as to avoid “zero” fingerprints devoid of any reactivity information. Most substrates should therefore be variations on a structural type known to be favored by the enzyme class under investigation.


Fig. 21 Principal component analysis (PCA) of lipases and esterases from fingerprint data. The groups A–F identified visually (dashed circles) correspond to those obtained by agglomerative clustering, with the exception of CRL2 and MJL in cluster E, which are assigned to cluster D (see Figure 19).

Any substrate array can be investigated for the functional relevance of its substrates once a data set has been acquired. In particular, the key question is whether the different substrates in the array all return differential reactivity information across the different enzymes tested. The PCA discussed above answers part of the question. Indeed if each substrate in the array behaves differently, the variability of the data set spans as many dimensions as substrates. However, if any two substrates display a similar reactivity pattern across the different enzymes, the diversity observed will be reduced to a lower number of dimensions. In the data set of lipase and esterase fingerprints discussed above, the first two principal components explain 68% of the diversity observed between the different enzymes. A bar diagram representation of the principal component coefficients indicates how each substrate contributes to the principal components of the data set (Figure 22). Substrate similarities can also be directly investigated by hierarchical clustering or PCA of substrates as observations against enzymes as variables. The data set is first normalized against substrates, with the sum of all reaction rates observed for each substrate across all enzymes being set to a constant of 100%. The results of clustering can be visualized as a hierarchical tree, or by direct rendering of the symmetrical distance matrix, which indicates functional similarities between substrates, as illustrated for the lipase data set (Figure 23). Pairs of enantiomers appear very similar in their reactivity against the different enzymes with the exception of the butyryl and hexanoyl esters, suggesting that this particular fingerprinting set

Fig. 22 Coefficients per substrate of the first two principal components (PC), which account for 68% of observed variance: PC1=46% (black bars), PC2=22% (white bars).

Fig. 23 Euclidean distance matrix between substrates 24–31 in the 25-dimensional space of enzyme reaction rate, rendered in grayscale (black = distance 0, white=maximum distance). For each substrate, reactivity for each enzyme was ex-

pressed as per cent of the total reactivity observed with this substrate across all enzymes measured. Substrates were clustered by agglomerative clustering using the group-average method.


could be reduced to a lower number of substrates to return essentially the same analysis results in terms of differentiation between enzymes.

4 Conclusions

Enzyme fingerprints are images of the reactivity profile of an enzyme across the chemical structural space of a given substrate type. The resolution of these images depends on the number and structural diversity of the substrates composing the fingerprinting array. Fingerprinting arrays composed of small sets of substrates ( 50% of primary lung tumors and several immortalized cell lines [32]. Similarly, PPARγ expression was observed in >90% pancreatic adenocarcinomas investigated, in contrast to normal pancreatic duct epithelia which were negative for PPARγ at the protein level [89]. In a study of normal versus malignant brain, breast, and prostate cells, PPARγ expression was consistently higher in the malignant cells but was not regulated by ligand [90]. As well as evidence from expression studies, activation of PPARγ resulted in an inhibition of estrogen production in breast adipose tissue [91] and played a role in tumor regression [92]. Additionally, activation of PPARγ inhibited growth of human hepatocellular [93] and esophageal carcinoma cells though cell cycle arrest and induction of apoptosis [94]. The tumor suppressor functions of PPARγ ligands are reported to be mediated by modulation of expression of the tumor suppressor PTEN [95], a protein known to have an involvement in many cellular functions including proliferation and apoptosis (reviewed in Ref. [96]). 4.3 PPARγ as a Therapeutic Target?

Studies on estrogen production suggest a role for PPARγ ligands in the treatment of breast cancer [91]. In addition, the observation that activation of PPARγ is antiproliferative and correlates with maturational stage in neuroblastoma, vascular smooth muscle cells, and DMBA-induced mammary tumors suggests a potential use for PPARγ in the management of advanced stage vascularised tumors [38, 79, 97]. However, the most compelling evidence for PPARγ as a therapeutic target for cancer is derived from studies of colorectal tumors. Treatment of colorectal cancer cell lines with PPARγ ligands resulted in growth inhibition, promotion of differentiation related markers, and expression of Drg-1 (differentiation-related gene-1), a putative suppressor of metastasis in human colorectal cancer [34, 98]. Activation of PPARγ was also found to inhibit expression of COX-2 in colorectal cancer cell lines, which is in opposition to results observed with PPARα [75, 99]. Furthermore, inactivating mutations in PPARγ were identified in a subset of colorectal tumors, supporting a role for PPARγ as a tumor suppressor of colorectal carcinogenesis [100].

5 PPAR$

Although the majority of studies have demonstrated a role for PPARγ ligands as preventative agents for colorectal cancer, two studies in APCmin/+ mice suggested the converse. These studies showed an increase in tumors or polyps in the colon of these mice after they were fed a diet containing a PPARγ agonist for 8 or 5 weeks, respectively [101, 102]. However, this is in contrast to in vitro and murine xenograft studies using human cell lines in which PPARγ agonists resulted in growth inhibition and induction of differentiation [100, 103]. As already described for PPARα, species differences in response between the human cell lines compared with the APCmin/+ mice offer the most likely explanation for this discrepancy. Additionally, one recent paper suggests that the antitumor effects of TZDs are independent of PPARγ and are mediated instead by inhibition of transcription [104].

5 PPARβ 5.1 Expression and Activation

Expression of PPARβ has been demonstrated in a vast array of tissue and cell types [4, 105, 106]. Unlike PPARα and PPARγ , little is known about the physiological role of PPARβ and several independent studies have linked this receptor to diverse functions. An involvement for PPARβ has been suggested in brain lipid metabolism [107], squamous cell differentiation [105], regulation of other hormone receptors [108], and as a mediator of prostacyclin function in blastocyst implantation in mice [76]. Although several compounds have been demonstrated to activate PPARβ nonspecifically, including a number of PPARα and γ agonists, a selective ligand for PPARβ has yet to be elucidated. 5.2 PPARβ and Cancer

Unlike PPARα and PPARγ , little is known about the role of PPARβ and its involvement, if any, in the development of cancer. As described earlier, activation of this receptor has been associated with processes independent of those identified for either PPARα or PPARγ . The observations that PPARβ can repress human PPARα activity [108] and induce PPARγ expression [109] would suggest that PPARβ can act as a negative regulator of tumorigenesis in certain cell systems. In contrast to this hypothesized antitumorigenic role, PPARβ has been suggested to play a role in the development of colorectal cancer [37, 76, 110]. Expression levels of PPARβ were shown to be higher in colon carcinoma compared to the adjacent surrounding tissue [37, 76]. Furthermore, PPARβ was concentrated in the most differentiated cells at the luminal surface of the mucosal glands in normal mucosa, but was expressed in epithelial cells located throughout the dysplastic glands of the

7

8 PPARs and Cancer

neoplastic tissue [76]. This suggests that PPARβ expression alone is not proneoplastic [76]. Using serial analysis of gene expression (SAGE), PPARβ was identified as a downstream target of the adenomatous polyposis coli (APC) tumor suppressor pathway [111]. Under normal conditions, APC binds to β-catenin and inhibits its ability to form a transcription complex with TCF-4. Conversely, in the majority of colorectal tumors examined, APC is inactivated by truncating mutations giving rise to elevated transcriptional activity of the β-catenin/TCF-4 complex. Similarly, in those tumors with intact APC genes, mutations of β-catenin that render it resistant to the inhibitory effects of APC occur, thereby demonstrating that inhibition of β-catenin/TCF-4-mediated transcription is critical to the tumor suppressive role of APC [37]. Downregulation of PPARβ by APC was demonstrated to be as the result of a direct molecular interaction, since the PPARβ promoter contains TCF-4 binding sites, and PPARβ promoters were repressed by APC as well as stimulated by mutant β-catenin [37]. 5.3 PPARβ as a Therapeutic Target?

Evidence for a potential use of PPARβ ligands in cancer therapy is derived from studies of colorectal tumor development where non-steroidal anti-inflammatory drugs (NSAIDs) can bind and potentially inhibit PPARβ [37], offering an explanation of NSAID-mediated chemoprevention [110]. This theory was supported using a PPARβ-null human colorectal cancer cell line [37]. These cells showed no obvious phenotype in vitro, and no increased sensitivity to NSAIDs. However, they were defective in establishing tumors in vivo [37]. Although the absence of PPARβ affected tumorigenicity of this colorectal cell line, the authors stressed that this effect may be specific to this cell line, and that the role of PPARβ in the chemopreventative effects of NSAIDs may involve more than one mechanism [37]. However, because the APC pathway is mutated with high frequency in colon cancer and overexpression of PPARβ has been seen in many colorectal cancers, it seems likely that increased expression of PPARβ may contribute to the neoplastic process and thus its modulation may present therapeutic opportunities.

6 Conclusions

A review of the last decade of literature reveals an involvement of PPARs in many diverse biological processes from adipocyte differentiation to rat liver cancer. The evidence for a role of PPARs in fat metabolism and energy balance is conclusive whereas the evidence for a role in tumorigenesis is confusing. Nonetheless, it is clear that modulation of PPARγ activity has some effect on the regulation of tumors of the colon, opening many avenues for exploratory research. Perhaps most compelling is the tentative connection between high fat diet, cellular energy balance, and cancer of the colon and maybe of the breast. Although the way forward

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depends on further elucidation of the basic biology of cancer, much is already known about PPAR ligand chemistry and toxicology and the binding specificity of different PPARs [19]. Thus, PPARs provide excellent therapeutic targets since they are receptors with intrinsic thresholds that act directly to modulate gene expression.

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PPARs in Atherosclerosis Jorge Plutzky Harvard Medical School, Boston, MA, USA, 02115

Originally published in: Cellular Proteins and Their Fatty Acids in Health and Disease. Edited by Asim K. Duttaroy and Friedrich Spener. Copyright ľ 2003 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30437-0

1 Atherosclerosis 1.1 Introduction

Atherosclerosis and its protean manifestations – including myocardial infarction, cerebrovascular event, peripheral vascular disease – remains a major cause of morbidity and mortality worldwide [1]. Although a common scourge of Western society, the exportation of atherosclerosis to developing countries reveals obvious and worrisome trends. This immense burden of disease has been countered with extensive efforts to better understand, prevent, and treat atherosclerosis and its complications. Over the years, this has led to an evolution of thinking regarding atherosclerosis [2, 3]. A century ago, atherosclerosis was considered primarily a degenerative disease of the elderly. Over time, it became apparent that atherosclerosis is a clinical entity manifest in a much broader demographic segment of the population. Early observations focused on intra-luminal arterial obstructions as being at the center of an imbalance between arterial supply and tissue demand that precipitated cardiac ischemia and infarction. Although true to some extent for “demand ischemia”, this view of a gradual “hardening of the arteries” gave way to understanding atherosclerotic plaque as a more dynamic lesion [4]. Critical to this was recognition of thrombus formation as an integral element in both the propagation of lesions as well as their complications [5, 6]. Most of acute cardiovascular events derive from plaque rupture and the ensuing occlusive thrombus that results [7]. More recently, the field has focused on two dominant issues regarding the nature of atherosclerotic plaque. The first is atherosclerosis as a chronic inflammatory Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 PPARs in Atherosclerosis

disease [1]. The second is attention to how metabolic pathways and disorders fundamentally and pervasively influence vascular responses and the atherosclerotic process [8]. Both of these trends – inflammation and metabolism as critical components of the atherosclerotic process – underlie the explosive interest in the role of peroxisome proliferator activated receptors (PPARs) in vascular biology and atherosclerosis [9, 10]. We will review the current concepts in atherosclerosis itself before delving into the existing evidence implicating PPARs and their ligands as possible mediators in these responses, looking in turn at PPARα, PPARγ , and PPARδ (also known as PPARβ). 1.2 Atherosclerosis as a Clinical Syndrome

Atherosclerosis can no longer be considered as a single pathologic process accounting for its various clinical manifestations. Two common scenarios highlight this spectrum of disease. The first is the 45-year-old man who presents with sudden death and is found at autopsy to have ruptured a single severe atherosclerotic plaque in his left anterior descending artery, with a subsequent rapid cascade of occlusive thrombus, acute cardiac ischemia, ventricular fibrillation, and death. Contrast this to the octagenarian who has a mild infarct as a complication of hip surgery and is found to have diffuse mild three-vessel disease. Both patients have atherosclerosis, but the manifestations are quite divergent. Similarly, distinctions can be made between atherosclerosis in the coronary arteries, cerebrovasculature, aorta, and femoral vessels. The presence of biomechanical factors, such as shear stress and hemodynamics, which alter cellular responses including gene expression is one of many likely contributors to such differences [11]. Here we will focus largely on coronary atherosclerosis. Atherosclerosis develops through a variety of recognized stages, from its earliest manifestation as a fatty streak through raised lesions to evolution into complicated advanced lesions [6]. An early seminal event in atherosclerosis is the entry of inflammatory cells like monocytes (MO) and their in situ development into macrophages (Mφ) and then foam cells as they take up lipid [1]. This arterial entry of MO/Mφ involves a series of specific cellular interactions heavily dependent on the expression of adhesion molecules on the luminal surface of endothelial cells (EC) [12]. Late-stage complicated lesions are characterized by a large necrotic and highly thrombogenic lipid core, separated from the circulation by the fibrous cap, a reactive response produced primarily by vascular smooth muscle cells (VSMCs) and consisting of collagen and other extracellular matrix materials [10]. Data from pathologic studies as well as thrombolytic trials, in which the culprit lesion leading to myocardial infarction (MI) could be seen by comparing coronary angiograms preand post-thrombolysis, revealed that most MIs occurred not in the most stenotic lesions but rather in those with more modest stenoses, in the 50–70% range [13, 14]. Through such work, the fissuring of the fibrous cap, known as plaque rupture, and the exposure of the circulation to the thrombogenic lipid core result in the occlusive thrombosis inducing most MIs.

1 Atherosclerosis

These observations have focused attention on the nature of the fibrous cap, what maintains it, and the forces and players involved with plaque rupture versus plaque stabilization [7]. One group of proteins highly implicated in plaque destabilization are the matrix metalloproteinases (MMPs) [15]. MMPs represent a large, complex, family of highly regulated matrix-degrading enzymes thought to be especially active in the shoulder regions of ruptured plaques. Another mechanism contributing to lesion formation is the superficial erosion of ECs. 1.3 Cellular Constituents of Atherosclerosis

The complex interplay of factors and forces described above clearly involves unique and critical roles for different cellular constituents of the arterial wall in both normal vessel function and pathologic atherosclerotic responses. One fundamental shift in perspective over the past decade has been in understanding the endothelium as not a simple passive conduit but rather a dynamic organ involved in metacrine, paracrine, and endocrine function [16, 17]. In their critical position lining blood vessels, ECs represent the interface between circulating elements and the ultimate tissue response, serving as a transducer of those reactions. Endothelial changes like adhesion molecule expression are not only among the earliest cellular responses contributing to atherosclerosis, endothelial dysfunction also occurs as an early manifestation of clinical atherosclerosis [18]. Centrally involved in such changes, and a salient example of endothelial roles in vascular responses, is endothelial nitric oxide production, an endogenously produced substance synthesized by endothelial nitric oxide synthetase, that induces arterial relaxation through effects on VSMCs [19, 20]. The studies establishing the dependency on the endothelium for such responses contributed to the Nobel Prize in Medicine being awarded to Furchgott, Murad, and Ignarro [21]. The medial layer of the arterial wall is primarily muscular, comprising VSMCs. These cells help maintain vascular tone and provide an essential structural component. VSMCs are integral to many processes implicated in atherosclerosis [22]. They may contribute to the presence of hypertension or be involved in the reaction to it. The migration of VSMCs from the media to the intima as well as proliferation of VSMCs once present there is a hallmark of the atherosclerotic process. VSMCs also provide the main source of extracellular matrix which forms the fibrous cap. In this role, VSMCs provide an essential function, albeit a reaction to a pathologic state. VSMC dysfunction may contribute to the weakening of the fibrous cap due to less collagen synthesis. Interestingly, such decreases in collagen production are repressed by inflammatory stimuli such as interleukins (IL-1). VSMCs also elaborate matrix metalloproteinases (MMPs), which contribute to the remodeling process [6]. Interestingly, MMP expression appears in part regulated by pro-inflammatory signals such as IL-1 [23, 24]. Two other cell types represent critical players in atherosclerosis, even if they are not cellular constituents of the artery wall under normal conditions. These atheroma-associated cells include monocytes/macrophages and lymphocytes,

3


predominantly T cells [1]. Monocytes are attracted from the circulation to sites of endothelial injury, going through a process of rolling, attachment, and entry into the vessel wall [12]. Extensive data suggest chemokines, or chemoattractant cytokines, as an important signal attracting monocytes and other inflammatory cells to sites of inflammation and injury [25]. Members of this large complex family of proteins are induced by different signals, including inflammatory cytokines acting on various cells, such as EC. Secreted chemokines bind to chemokine receptors present on specific inflammatory cells. Moving down concentration gradients of chemokines, monocytes adhere and enter the vessel wall, and take up lipid, especially low-density lipoproteins (LDL). T lymphocytes are also critical atheromaassociated cells, providing important signals, for example inflammatory cytokines that contribute to monocyte, VSMC, and EC activation [26]. 1.4 Atherosclerosis as an Inflammatory Disorder

As suggested above, an extensive literature has now established atherosclerosis as a chronic inflammatory disease [1]. Although the exact proximal signals remain unclear, with posited contributors ranging from possible infectious etiologies to oxidized LDL, inflammatory cells and inflammatory signals are implicated in essentially every step of atherogenesis and atherosclerosis. Extensive efforts have identified many inflammatory mediators that participate in these responses. Interestingly, these basic science studies now intersect with epidemiologic studies which find that levels of non-specific markers of inflammation like c-reactive protein (cRP) predict the risk of future cardiovascular events [27]. Interestingly, c-RP levels are predictive above and beyond total cholesterol/HDL ratios, and decreased cardiovascular events through lipid-lowering are paralleled by decreases in c-RP levels [28, 29]. Although a multitude of questions persist about the clinical significance of inflammatory markers, their striking correlation with atherosclerosis underscores the prospect of inflammation as a therapeutic target, a notion that has been raised for PPAR agonists. 1.5 Atherosclerosis as a Metabolic Disorder

Coincident with expanding recognition of the dynamic nature of the vasculature and its pathobiology has been the increasing attention to the interaction between various aspects of metabolism and vascular responses [8]. The most obvious connection in this regard has been lipid metabolism and its now well-established role in atherosclerosis. This extends beyond the extensive data establishing LDL as a risk factor, pathogenic mediator, and therapeutic target to other lipoprotein particles [30]. Triglycerides, carried predominantly in VLDL particles (when synthesized by the liver) and chylomicrons (when particles are assembled in the gut) are composed primarily of fatty acids (FAs) [31, 32]. Establishing if triglycerides represent independent risk factors for atheroclerosis has been challenging [33, 34]. Although

2 PPAR in the Vasculature

triglycerides are clearly associated with cardiovascular events, this association tends to weaken or disappear when other parameters are taken into account. This is not surprising given the close interaction between other risk factors like obesity, diabetes, the post-menopausal state and increased triglyceride levels [35, 36]. Triglycerides are also associated, although inversely, with high-density lipoproteins [30]. HDL is thought to participate in reverse cholesterol transport, moving lipoproteins from the periphery back to the liver [37]. The extensive connections between PPARs and lipoprotein metabolism are discussed in detail elsewhere in this book, but remain relevant to the connections between PPARs and atherosclerosis since as PPAR agonists may exert their effects on atherosclerosis through changes in lipids. Recent trials suggest that even in patients with coronary artery disease (CAD) with average to low LDL levels, modestly increased triglycerides, and low HDL, fibrate treatment can decrease cardiovascular events [38]. The possibility that these may occur in part through PPARα activation has been raised [9, 39]. Beyond lipoproteins, the metabolic effects on atherosclerosis are also strikingly evident among diabetic patients [40]. Diabetes mellitus is now considered a cardiovascular risk equivalent based on data that patients with diabetes but no known CAD have the same risk of a future cardiovascular event as non-diabetics with a prior MI [41–43]. Increased cardiovascular risk also extends to patients with the insulin resistance syndrome (syndrome X) and even so-called “pre-diabetes”. A significant percentage of these insulin-resistant subjects will go on to manifest frank diabetes [44, 45]. The fact that so many of these patients will have clinically significant atherosclerosis has suggested the hypothesis that the metabolic aspects of these various syndromes - high triglycerides, low HDL, increased post-prandial hyperglycemia, and lipemia – may exert effects on the vasculature long before hyperglycemia diagnostic of diabetes becomes apparent [46]. Supportive of this hypothesis is the observation that perhaps as many as half of the patients by the time they are diagnosed with diabetes have already had an MI [47, 48]. Considerable attention has been paid to the potential effects, either protective or harmful, of anti-diabetic medications on the vasculature [49, 50].

2 PPAR in the Vasculature 2.1 PPARs in Vascular Biology and Atherosclerosis

From this broad overview, we can now consider the potential role of PPAR pathways in atherosclerosis. PPARs have been considered elsewhere in this book; the focus here is on the potential part these key transcriptional regulators of various aspects of metabolism (Tab. 1) may play in vascular responses, especially those relevant to atherosclerosis. To some extent, this issue has been greatly influenced by the ongoing use of PPAR agonists as therapeutic agents in patients with high risk for atherosclerosis – either those with dyslipidemia receiving fibrates or diabetic

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6 PPARs in Atherosclerosis Table 1 Putative ligands, cellular expression patterns, and pathways in which PPARs are

thought to be involved PPARα

PPARγ

PPARδ

Ligands

Fibrates Fatty acids

Carba/prostacyclins Fatty acids

Expression

Liver Muscle Vascular: ECs, SMCs Atheroma-associated: MPs, MPs, lymphocytes FA oxidation Lipid metabolism/transfer Energy balance

Thiazolidinediones Fatty acids Hydroxyoctadenoic acids Adipose tissue Vascular: ECs, SMCs Atheroma-associated: MPs, lymphocytes Adipogenesis Lipid metabolism/transfer Glucose homeostasis

Inflammation? Lipid metabolism/transfer

Pathways

Ubiquitous

PPAR expression in vascular and atheroma associated cells is now well-established, raising the prospect of synthetic and endogenous PPAR ligands having direct effects on vascular responses. EC, endothelial cell; SMC, smooth muscle cell; FA, fatty acid; MP, macrophages.

patients receiving thiazolidinediones. Early experiments examining these direct effects of PPAR agonists in the vasculature sought to first determine if PPARs were present in the various cellular components of the arterial wall and/or atheromaassociated cells, before considering if PPAR-regulated targets with relevance to vascular biology existed. Work has now evolved into testing if such effects occur in vivo, including humans. Thus, prior and emerging clinical trial data regarding cardiovascular events in patients receiving PPAR agonists can be viewed through this lens. To what extent did changes in cardiovascular events seen in those clinical trials stem from indirect changes in metabolism induced by PPAR agonists versus possible effects of directly activating PPARs in vascular cells. Beyond these pharmacologic issues, a pertinent question also exists regarding the role of endogenous PPAR activation in vascular responses. 2.2 Examining Evidence for PPAR in Vascular Responses

To a certain degree, the data regarding PPARs in the vasculature have been limited by several critical issues. One is the inherent complexity of PPAR action, with multiple levels of control that can vary among different cell types or conditions [51]. These issues include differences in respect to ligands, and the effects of those ligands at different concentrations, differences in PPAR action dependent on cell type, for example due to the presence or absence of certain co-activators or the relative levels of PPAR themselves, and variance in promoter response elements, to name a few. Published results may also differ because of different experimental protocols. Thus, perhaps not surprisingly, various and at times divergent effects

3 PPAR( in Vascular Biology and Atherosclerosis

have been reported in different and sometimes the very same vascular or atheromaassociated cell types. Furthermore, it remains challenging to ever establish that a given effect occurs in response to nuclear receptor activation, and not some other up- or downstream mechanism. These issues must be kept in mind in interpreting many of the results reported in the PPAR literature in general and certainly with regard to PPARs in the vasculature. Thus, elements of confusion, contradiction, and unresolved issues persist in the nascent world of PPARs and their agonists in the vasculature. These issues, perhaps a product of the dizzying pace of progress, will be noted as they arise in the review of the data that follows.

3 PPARγ in Vascular Biology and Atherosclerosis 3.1 In vitro Evidence

PPARγ expression is now recognized in essentially all vascular and atheromaassociated cells seen in the vessel wall [52]. This includes VSMCs, ECs, monocyte/macrophages, and more recently Tcells. PPARγ is also expressed in human atherosclerotic lesions [53]. With the presence of these nuclear receptors established in these cellular settings, the question arises as to the evidence for regulation of relevant targets in the vascular pathways described above. PPARγ expression has been linked to a variety of integral responses in VSMCs. Among these, regulation of matrix metalloproteinase 9 (MMP-9) is perhaps an example of a now canonical PPARγ -regulated target gene, with evidence of transrepression of MMP-9 induction seen across many levels in both VSMCs [54] and monocytes/macrophages [55]. PPARγ agonists repress MMP-9 mRNA induction, protein levels, and gelatinolytic activity on zymograms, a commonly used tool to study MMP responses [54]. Similar PPARγ effects have been reported for the MMP-9 promoter [55]. A net effect of decreased MMP-9 action seems apparent given the absence of PPARγ regulation of tissue inhibitors of MMPs (known as TIMPs) [54]. PPARγ agonists also limit VSMC migration [54] and proliferation [56] with effects on cell cycling [57]. These changes in VSMC responses may contribute to thiazolidinedione effects in cardiovascular settings in which VSMC matrix remodeling may be at work, for example instent restenosis [58, 59]. They may also play a role in the inhibition seen in intimal: medial hyperplasia reported with thiazolidinediones [60]. PPARγ agonists also exert effects on MMP-9 in monocyte/macrophages, where MMPs [61] have been implicated in plaque rupture; it remains to be seen if PPARγ activation contributes to plaque stabilization. In addition to similar effects through PPARγ activation on MMPs in monocytes and macrophages, PPARγ activation also has other effects in these cells as well. Among the earliest reports of PPARγ action in inflammatory cells were studies indicating that PPARγ agonists could limit cytokine induction in

7


monocytes/macrophages [62, 63]. More recent work establishes PPARγ (and PPARα, discussed below) expression in T lymphocytes [64–66], with a variety of effects reported, including apoptosis and inhibition of cytokine production [67]. Provocative results were also reported for PPARγ -mediated induction of CD36, a receptor for oxidized LDL uptake [68, 69]. These effects, which appeared to contribute to foam cell formation, raised concerns for possible pro-atherogenic effects through thiazolidinediones. This would not necessarily be the case, given other possible off-setting effects through sequestering oxidized LDL by CD36 induction in other tissues, and concurrent changes in cholesterol efflux [70]. Furthermore, the nature of the increased lipid in these monocytes was never established and could well have been triglyceride-rich lipoprotein, and not the lipid particles typically associated with foam cells. In fact, subsequent work has established that PPARγ , and apparently PPARα activators also induce the expression of ABC-1, a protein implicated in cholesterol efflux [71–73]. ABC-1 is the gene disrupted in Tangier’s disease, a clinical condition associated with markedly decreased HDL levels [74, 75]. Endothelial cells also express PPARγ as shown by RT-PCR [76], Western blotting and Northern analysis [77]. The anti-inflammatory effects suggested above have also been implicated in ECs on a different limb of inflammation – chemokine expression. We have reported that PPARγ agonists limit IFNγ induction of specific subsets of chemokines implicated in atherosclerosis, at least among those tested. Three different well-established PPARγ agonists limited induction of three different CXC chemokines (IP-10, Mig, and ITAC), but not the CC chemokine MCP-1 [78]. Further analysis found consistent PPARγ effects on the IP-10 promoter and in functional responses to lymphocytes expressing the CXC chemokine receptor. Although no effect was seen on MCP-1, this does not exclude PPARγ effects on the MCP-1 pathway. Two groups have reported responses that either clearly or likely occur by repression of the receptor for MCP-1 – CCR2 [79, 80]. Interestingly, MCP-1 levels were not changed in mouse models of atherosclerosis treated with PPARγ agonists, although MCP-1 levels do appear to fall in patients treated with TZDs [81]. PPARγ agonists may also induce apoptosis in EC [82], although these effects were observed primarily with 15-deoxy12,14 prostaglandin J2 (15-deoxy-12,14 PGJ2 ), which also has PPAR-independent effects [83]. Another relevant PPARγ target in ECs highlights some of the important issues noted earlier regarding challenges of establishing effects in vitro and comparing these to clinical responses. A variety of groups have found that PPARγ agonists could induce plasminogen activator inhibitor-1 (PAI-1), a procoagulant protein implicated in atherosclerosis [77, 84, 85]. Increased PAI-1 expression through PPARγ has been reported in EC in angiogenesis studies [84], in response to putative natural but not synthetic ligands [86] and in adipocytes [85]. In contrast, others find a decrease in PAI-1 in EC. Regardless, in human studies, serum PAI-1 levels appear to fall, possibly due to improved insulin sensitivity, or improved levels of glucose and/or triglycerides [87]. Thus different effects may be seen in vitro while clinical responses may vary due to pleiotropic drug effects, including differences between different agonists as well as PPAR-independent effects.

4 PPAR" in Vascular Biology and Atherosclerosis

3.2 In vivo Evidence

At least four different studies have reported various PPARγ agonists in vivo decrease atherosclerosis in mouse models [81, 88–90]. The first of these, by Glass and colleagues, studied the effects of rosiglitazone and an experimental PPARγ agonist, GW7845, in LDL receptor-deficient animals [81]. The extent of lesions was decreased with both agents, although interestingly, only in male mice. The explanation for this gender difference is unclear, but the female mice were more insulin resistant. This difference between males and females was not observed in other studies employing other models (apoE-deficient and LDLR-deficient mice on high fat or high fructose) and other PPARγ agonists. The ongoing clinical use of PPARγ ligands (pioglitazone or Actos, rosiglitazone or Avandia) as insulin-sensitizing agents in humans affords the possibility of asking if these drugs exert similar effects on atherosclerosis. Such questions are particularly germane given the very high risk for ischemic cardiovascular disease that patients with diabetes face. Despite these opportunities, establishing that the effects of any PPAR agonist derives from activation of its purported nuclear receptor target remains difficult. These agents may influence atherosclerosis indirectly, by improving metabolic status, directly but independent of PPAR activation, or directly through activation of its cognate PPAR. Early studies in humans suggest that these agents can in fact improve both serum and vascular surrogate markers of abnormal vascular responses [60, 87, 91, 92]. In terms of lipids, decreased triglycerides and improved HDL levels have been seen, often with changes that rival or exceed those seen in trials demonstrating decreased cardiac events through HDL-raising therapies [93]. Questions persist if there may also be differential effects on various parameters such as lipids among PPARγ agonists. Beyond this, improvements in inflammatory markers have also been seen, as recently reported. Such effects include changes in markers such as c-reactive protein (C-RP) [94], currently under intensive study as a novel predictor of cardiovascular risk. Arterial reactivity may be improved after TZD treatment, a potentially significant finding given evidence for endothelial dysfunction as a surrogate marker for early changes in atherosclerosis, although much of this data has been in abstract form. Perhaps most impressive have been the improvements in carotid intimal thickness seen in relatively short time frames in studies with limited numbers of patients. These results have contributed to a variety of ongoing studies focusing on cardiovascular endpoints in patients being treated with these agents. 4 PPARα in Vascular Biology and Atherosclerosis 4.1 In vitro Evidence

The central role for PPARα in fatty acid metabolism – controlling enzymes involved with β-oxidation as well as various targets involved in lipid metabolism – places PPARα in a position to be relevant to a host of vascular issues [95]. Such a role is

9


furthered by evidence that certain FAs can act as ligands for PPARα [51, 96]. FAs, present in various lipoproteins to a variable degree and with a varying nature (chain length, degree of saturation, cis/trans conformation), have been implicated in many aspects of EC, VSMC, monocyte/macrophage and even lymphocyte responses [97– 100]. PPARα has been reported in all such cellular settings, thus targets in these locations may come under regulation by synthetic and natural PPAR agonists, exerting direct effects on vascular responses. Such action could occur independently or in synergism with PPARα’s myriad effects on lipid metabolism, which range from regulation of apolipoproteins involved in HDL metabolism to enzymes of lipid metabolism (e.g., lipoprotein lipase), to even possible effects of other lipid drugs on HDL levels (e.g., statin-induction of HDL). Our focus here is on the potential direct vascular effects. Prior observations from our group revealed that certain fatty acids, including the ω-3 FA docosohexanoic acid, could limit adhesion molecule expression, especially VCAM-1, in ECs [101]. We asked if these effects might be occurring through PPARα activation. In fact, PPARα agonists can limit inflammatory cytokine-induced VCAM-1 expression, with effects that do not derive from cellular toxicity or changes in mRNA stability. These effects are demonstrable at the level of the VCAM-1 promoter through likely NFkB effects, as well as in functional assays of adhesion [86]. Although we found this effect to be specific to PPARα, others report that PPARγ agonists can also limit adhesion molecule expression [102, 103]. More recent evidence indicates that oxidation of some ω-3 FA, like eicosopentanoic acid (EPA), can limit leukocyte adhesion through PPARαdependent mechanisms evident in vivo (Fig. 1) [104]. These effects, specific to oxidized EPA and evident in classic PPARα transactivation assays, were absent when such in vivo adhesion studies were repeated in the genetic absence of PPARα (Fig. 1) [104]. Other PPARα-regulated targets in ECs include endothelin 1 as well as various enzymes involved in oxidative processes including superoxide dismutase and phox (p47, p22) [92, 105]. The effects of PPAR agonists in general appear to be an anti-oxidative effect, as also suggested by studies in humans [91, 92]. In contrast to these potentially anti-atherosclerotic effects, at least two reports indicate that certain oxidized lipoproteins may induce adhesion molecule expression through PPARαdependent mechanisms [106], although perhaps only under certain conditions such as in the presence of specific lipolytic enzymes [107]. Integrating these apparently contradictory effects into a unified model remains currently beyond our grasp but certainly worthy of further investigation, and likely a function of the complexity alluded to earlier. PPARα is also expressed in VSMCs. Evidence from Staels and colleagues indicates that PPARα activation can limit IL-6-induced changes in those cells [108]. PPARs clearly play a role in inflammatory cell differentiation and signaling, thus providing another setting in which they are likely players in atherosclerosis. In monocyte/macrophages PPARα has been reported to induce apoptosis, but only after cytokine stimulation, in contrast to PPARγ agonists, which induced apoptosis in the absence of such stimuli [109]. Mentioned earlier was the possible effect of

4 PPAR" in Vascular Biology and Atherosclerosis

Fig. 1 Effect of oxidized EPA on leukocyte adhesion in mesenteric venules in wildtype and PPAR"-deficient mice. Wild-type or PPAR"-deficient mice (PPAR"−/−) were given an intraperitoneal injection of vehicle (Veh) alone, native EPA, or oxidized EPA (oxEPA) one hour prior to injection of LPS. Five hours later mice were anesthetized and mesenteric venules were observed using intra-vital microscopy. (A) Adherent leukocytes were determined (n = 5–7

for each group of mice). *P60 years Infection from bovine prions Infected individuals are generally Met/Met homozygous with respect to the polymorphism at residue 129, which is also Met in bovine PrP Germ-line mutations in the PrP gene Germ-line mutations in the PrP gene

Fatal familial insomnia Iatrogenic CJD

Germ-line mutations in the PrP gene Infection from prion-contaminated human growth hormone, dura mater grafts, etc.

Kuru

Infection through ritual cannibalism (Fore people) Prion diseases in mammals

Bovine spongiform encephalopathy Scrapie (sheep and goats) Transmissible mink encephalopathy Chronic wasting disease (elk, deer) Feline spongiform encephalopathy (cats)

BSE-contaminated food Genetic susceptibility to spontaneous formation of scrapie prions. Transmission among sheep? Infection with prions from cattle or sheep Origin and transmission unknown Infection with prion-contaminated beef

PrPC monomers are by far more stable than PrPSc monomers, such that the small fraction of PrPSc monomers is not observed experimentally. Here, the rate-limiting step in spontaneous PrPSc formation is the oligomerization of the small fraction of PrPSc monomers to an oligomer of critical size, which is sufficiently stabilized by quaternary structure contacts such that it can no longer dissociate. This nucleus is capable of growing further, by pulling additional PrPSc monomers from the equilibrium into the PrPSc oligomer. An important difference between the models is that the nucleation-polymerization model does not necessarily require a direct contact and specific recognition between PrPC and PrPSc . Moreover, it much better accounts for the prion strain phenomenon, as will be discussed below (see Section 5). In addition to these models for the generation of PrPSc , another mechanism of amyloid formation has been postulated for the aggregation of human lysozyme variants in hereditary systemic amyloidosis. Here, single amino acid replacements

4 Biochemistry and Structural Biology of Mammalian Prion Disease

Fig. 1 Schematic representation of the template-assistance model (A) and nucleation-polymerization model (B) of PrPSc propagation.

in lysozyme destabilize the native state relative to partially structured intermediates. This leads to an increase of the fraction of partially folded intermediates that act as amyloid precursors and are pulled from the normal folding equilibrium during amyloid propagation [28]. A further mechanism is observed in the formation of amyloid fibrils of transthyretin (TTR), which are the putative cause of senile systemic amyloidosis (SSA) and familial amyloidotic polyneuropathy (FAP). Here, dissociation of the native TTR homotetramer into monomers is the rate-limiting step in amyloid formation, which is followed by partial monomer denaturation and misassembly into fibrils [29].

2 Properties of PrPC and PrPSc

Mammalian PrPC is a cell surface glycoprotein of about 210 amino acids that is strongly expressed in neurons but is also present in most other tissues. PrPC belongs to the most conserved proteins known, and pairwise alignment of PrPs from different species generally reveals more than 90% sequence identity [30, 31]. In addition, there are three types of invariant posttranslational modifications found in mature PrPC : two N-glycosylation sites at Asn181 and Asn197 [34], a single disulfide bond between Cys179 and Cys214 [33], and a C-terminal glycosyl-phosphatidylinositol (GPI) anchor at residue 231 (amino acid numbering according to human PrP) [16, 17, 32]. Another typical feature of mammalian PrP sequences is a segment of five octapeptide repeats in the N-terminal region (Figure 3A). Although

2 Properties of PrPC and PrPSc

the strong conservation of PrP in mammals suggests an important cellular role of the protein, the biological function of PrPC remains unknown to date. This is mainly due to the fact that PrP knockout mice are essentially healthy [35] and show only subtle phenotypes that are not completely preserved in different knockout strains. Proposed functions for PrP include roles in signal transduction [36], copper storage [37], circadian rhythm regulation [38], and maintenance of synaptic function [39] and Purkinje cell survival [40] (also reviewed in Ref. [8]). PrPC is monomeric, is soluble in non-denaturing detergents, and contains a high fraction of α-helical secondary structure [18]. in vivo, PrPC is enriched in cholesterol and sphingolipid-rich membrane rafts, subject to caveolae-mediated endocytosis [41], and has a half-life of about 3–6 h [42]. There is evidence that raft localization of PrPC and co-localization with PrPSc in the same membrane are required for its conversion into PrPSc [43]. Most PrP molecules synthesized in the cell reach the cell surface and are anchored to the cell membrane via their GPI anchor, but two intracellular transmembrane forms of PrP have been identified that span the ER membrane in different orientations, of which the form with the C-terminus towards the ER lumen (Ctm PrP) may be associated with neurodegeneration in prion-infected individuals [44]. Whether aberrant membrane topology of PrP at the ER has general functional implications still has to be determined. Moreover, retrograde transport of misfolded PrP from the ER to the cytosol normally leads to PrP degradation by the proteasome, but if this degradation is compromised, the accumulation of cytosolic PrP has severe toxic effects on neurons and has been proposed to be the general mechanism underlying the death of neurons during prion pathogenesis [45]. The insoluble, oligomeric scrapie form PrPSc has so far been isolated only from the brain of prion-infected individuals or prion-infected cell cultures. Although the subunits in PrPSc have the same covalent structure and posttranslational modifications as PrPC , it differs from PrPC with respect to all its biochemical properties: it is insoluble in non-denaturing detergents and shows an increased β-sheet content and a decreased α-helix content compared to PrPC [18]. Another important property of PrPSc is its partial resistance to proteinase K (PK). While PrPC is readily degraded by PK, the polypeptide chains in PrPSc exhibit a protease-resistant core ranging from approximately residue 90 to the C-terminal residue 231. In contrast, the N-terminal PrPSc segment from residue 23 to approximately residue 90 is degraded (Figure 2) [46, 47]. As will be discussed in Section 5, this observation has important implications, as it shows that PrPSc is an ordered oligomer in which all subunits appear to have a very similar structural environment that protects residues 90–231 in every subunit from access by PK. This is a strong hint that PrPSc does not represent a nonspecific, inclusion body–like protein aggregate but rather a regular array of subunits with identical tertiary structures in which both the fold of the subunits and their quaternary structure contacts confer protease resistance. This is in good agreement with the nucleation-polymerization model of PrPSc propagation [27], which assumes specific subunit-subunit contacts in the nucleation and further growth of the PrPSc oligomer. The importance of the

5


Fig. 2 Protease resistance of PrPSc . While PrPC is readily degraded by proteinase K into peptide fragments, the subunits of PrPSc contain a protease-resistant core (PrP27–30) from about residue 90 to the C-terminal residue 231 (amino acid numbering according to human PrP). Thus, only

the N-terminal segment 23–90 in PrPSc subunits is accessible to proteinase K degradation. The structure of PrPC , comprised of the flexibly disordered N-terminal tail 23–120, and the structured C-terminal domain PrP(121–231) (cf. Figure 3) are also indicated.

protease-resistant core of PrPSc , termed PrP27–30, is further underlined by the fact that it retains prion infectivity and forms amyloid-like fibers [19]. In addition, transgenic mice exclusively expressing N-terminally truncated forms of PrP that still contain segment 90–231 are susceptible to prions and able to replicate prions [48]. This shows that the octapeptide repeats in the N-terminal segment of PrPC are not required for prion propagation. The identification of PrP27–30 after protei-nase K treatment of brain extracts, protein separation by SDS-PAGE, and immunospecific detection in Western blots represents the basis for both post-mortem and pre-clinical detection of prions, e.g., in BSE-infected cattle, and, as outlined in Section 5, biochemical characterization of prion strains [6, 47]. Interestingly, a high-molecular-weight polysaccharide consisting of 1,4-, 1,6-, and 1,4,6-linked glucose units was found to be associated with infectious preparations of PrP27–30, and it was postulated that this polysaccharide is a scaffold that promotes formation of PrPSc and contributes to its extraordinary physical and chemical stability [49]. Additional structural information on PrPSc has become available from fiber diffraction studies and atomic force microscopy studies indicating that PrPSc contains β-sheets that are arranged perpendicular to the fiber axis [50] and may form parallel β-helices [51].

3 Three-dimensional Structure and Folding of Recombinant PrP

3 Three-dimensional Structure and Folding of Recombinant PrP 3.1 Expression of the Recombinant Prion Protein for Structural and Biophysical Studies

The protein-only hypothesis and all theoretical models of PrPC to PrPSc conversion imply that knowledge of the three-dimensional structures of PrPC and PrPSc is the prerequisite for understanding prion propagation at a molecular level. As PrPSc is insoluble, and because no soluble, low-molecular-weight oligomers with defined stoichiometry have been isolated so far, PrPSc has not been accessible to atomic structure determination by X-ray crystallography or nuclear magnetic resonance (NMR) spectroscopy. Because it is extremely difficult to purify milligram quantities of PrPC from mammalian brain, production of recombinant PrP in Escherichia coli has proved to be necessary to determine the three-dimensional structure of the monomeric protein. Since there is neither N-linked glycosylation nor GPI anchor biogenesis in E. coli, bacterially produced PrP lacks these modifications. Nevertheless, there is no evidence at present that these modifications significantly influence the tertiary structure of the prion protein. To allow formation of the invariant disulfide bond between Cys179 and 214 in the recombinant prion protein, two strategies have been pursued. As disulfide bonds cannot form in the reducing environment of the bacterial cytoplasm, the prion protein and N-terminally truncated fragments were secreted into periplasm with the E. coli OmpA signal sequence [52]. The disulfide bond was formed, but the protein was N-terminally degraded by periplasmic proteases such that only the fragment from residues 121–231 stayed stable in the periplasm. The fragment was termed PrP(121–231) [52]. This was the first evidence that the entire N-terminal segment 23–120 of recombinant PrP does not adopt a defined structure in solution (see below). Because any attempt to produce full-length PrP in the bacterial periplasm has failed so far, the full-length protein was expressed in the cytoplasm where the protein accumulates in the reduced form and nonspecifically aggregates into inclusion bodies. Oxidative refolding from the inclusion body fraction in vitro and further purification yielded large quantities of homogeneous, recombinant PrP for structure analysis and biochemical characterrization [53, 54]. 3.2 Three-dimensional Structures of Recombinant Prion Proteins from Different Species and Their Implications for the Species Barrier of Prion Transmission 3.2.1 Solution Structure of Murine PrP The first three-dimensional structure of a recombinant prion protein was reported in 1996 for the fragment PrP(121–231) of the murine prion protein, which was determined by nuclear magnetic resonance spectroscopy (NMR) [56, 57] (Figure 3B). PrP(121–231) adopts a unique fold that has so far not been found in other proteins. The fold consists of a short, two-stranded, antiparallel β-sheet

7


Fig. 3 Posttranslational modifications of PrPC and three-dimensional structure of recombinant PrP. (A) Mammalian prion proteins contain an N-terminal signal sequence (residues 1–22) that directs the protein to the endoplasmic reticulum, where glycans (CHO) at Asn181 and Asn197 are attached, the C-terminal peptide 232–254 is replaced by a GPI anchor at the carboxylate of residue 231, and the single disulfide bridge between Cys179 and Cys214 is formed (amino acid numbering according to human PrP). Positions of $-strands (S1, S2) and "-helices (H1–H3) in the NMR structure of the recombinant PrP are indicated, as well the length of the protease-

resistant core of PrPSc and the structured C-terminal domain PrP(121–231). (B) Ribbon diagram of the NMR structure of recombinant murine PrP. The protein is composed of a flexibly disordered N-terminal tail (residues 23–120, represented as random coil C" trace) and a structured domain (residues 121–231), which consists of a two-stranded, antiparallel $-sheet and three "-helices. The single disulfide bond (black lines) connects the second and third "-helix and is completely shielded from the solvent. The solvent-accessible residues 90–120, which become protease-resistant in PrPSc , are indicated as red lines.


(residues 128–131 and 161–164) and three α-helices (residues 144–154, 175–194, and 200– 226). The fold is additionally characterized by a tightly packed hydrophobic core of 20 amino acids (residues 134, 137, 139, 141, 158, 161, 175, 176, 179, 180, 184, 198, 203, 205, 206, 209, 210, 213, 214, and 215). The invariant disulfide bond between Cys179 and Cys214 is part of this hydrophobic core and connects the second with the third α-helix. The hydrophobic core is surrounded by a shell of hydrogen-bonded residues that further stabilize the fold [58]. Overall, the structure of PrP(121–231) resembles a flat ellipsoid that is characterized by an uneven distribution of surface charges between the two flat surfaces. The two glycosylation sites at Asn181 and Asn189 are located on the negatively charged side of the molecule, while the opposite surface is positively charged. It has been proposed that the uneven charge distribution may contribute to the orientation of PrPC relative to the cell membrane. One would expect that the protein associates via its positively charged surface with the membrane, such that the two glycosylation sites would be oriented towards the extracellular space [56]. The NMR structure of full-length murine PrP, which was reported in 1997, revealed that residues 23–120 (including the octapeptide repeats), in contrast to the compact domain PrP(121–231), are flexibly disordered in solution (Figure 3B), with all residues in this segment being flexible on a sub-nanosecond timescale [59]. The structure of residues 121–231 in the context of the full-length protein proved to be indistinguishable from that in isolated PrP(121–231). The functional role of the flexible tail is presently unclear. However, a plausible model has been proposed for structure formation within the octapeptide repeat regions upon cooperative binding of Cu2+ ions [60]. As there is increasing evidence that a substantial fraction of cellular proteins are “natively unfolded” in isolation and adopt a defined tertiary structure only in complex with target molecules [61], it appears likely that the flexible tail of PrP is required for the interaction of PrPC with its natural ligands. Overall, the structure of recombinant, non-glycosylated, full-length PrP produced in E. coli is in full agreement with the known properties of PrPC isolated from mammalian cells. As natural PrPC , it predominantly contains α-helical secondary structure and is soluble and monomeric. It is therefore generally accepted that the structure of recombinant PrP corresponds to that of the benign cellular prion protein. As will be discussed in Section 3.2.2, the NMR structure of murine PrP is also very similar to the structures of other mammalian prion proteins. Because the segment 90–231 is protease-resistant in PrPSc , an important conclusion from the structure of full-length PrP(23–231) is that residues 90–120, which are required for prion propagation [36], must adopt a defined three-dimensional structure in PrPSc . This corresponds to the minimal structural change that has to be postulated for a PrP polypeptide when it is converted from PrPC to a subunit of PrPSc (Figure 3B). As one might expect from the flexibly disordered N-terminal tail, attempts to crystallize recombinant prion proteins have failed so far. The only exception is the X-ray structure of human PrP(90–231), which crystallized as a covalently linked homodimer in which α-helices 3 are swapped between the subunits such that αhelix 2 in each subunit is now disulfide-bonded to α-helix 3 from the other subunit (Figure 4B) [62]. This structure may be an experimental artifact that was produced

9


Fig. 4 Comparison of the NMR structure of recombinant murine PrP(121–231) (pdb code: 1AG2) (A) with the X-ray structure of a dimer of recombinant human PrP(90–231) (B) (code1I4M) and the NMR structure of the murine Doppel protein (C) (code 1I17). (B) In the dimer of human PrP(90–231), the C-terminal "-helices are swapped and disul-

fide is cross-linked with the second "-helix of the other subunit. The molecule may be an artifact resulting from oxidative refolding of the recombinant protein from E. coli inclusion bodies under nonnative conditions. (C) Note that the Doppel protein is stabilized by two intramolecular disulfide bonds (dark blue lines).

during oxidative in vitro refolding of the recombinant protein under conditions that favored intermolecular disulfide bond formation. Disulfide-bonded homodimers have not been observed for PrPC , nor have intermolecular disulfide bonds been reported for PrPSc oligomers [33]. As mentioned above, the structure of the C-terminal domain of the mammalian prion protein represents a new fold that has not been observed in other proteins. In this context, it is interesting to discuss the recently solved NMR structure of the mammalian protein with closest homology to the prion protein, termed Doppel (Dpl) [63] (Figure 4C). Dpl lacks the N-terminal octapeptide repeats found in PrP and is most likely not linked with prion diseases, as it is normally not expressed in the central nervous system. In contrast, it was shown to be required for male fertility in mice by controlling male gametogenesis and sperm-egg interaction [64]. Figure 4C shows the NMR structure of murine Dpl [65], which shares 21% amino acid sequence identity with murine PrP and is very similar to the structure of human Dpl [66]. The Dpl fold is similar to that of PrP(121–231) in that the secondary structures fall in the same positions in the two proteins. Differences are observed mainly in the lengths of the α-helices and the positions of the β-strands (in Dpl, the strands are displaced by one to two residues relative to those in the PrP structure). Moreover, Dpl contains two intramolecular disulfide bonds. The disulfide bond


109–143 corresponds to that of PrP(121–231), and the second disul-fide 95–148 connects the segment after β-strand 1 with the C-terminal segment following αhelix 3. The strongest structural difference between the proteins is that the plane of the β-sheet is parallel to the axis of α-helix 2 in Dpl, while it is perpendicular to this axis in the structure of PrP(121–231). Moreover, there is a marked kink in αhelix 2 of Dpl, which is not observed in PrP(121–231). In summary, the structured domains of Dpl and PrP appear to have evolved from the same ancestor by gene duplication but now fulfill different functions. 3.2.2 Comparison of Mammalian Prion Protein Structures and the Species Barrier of Prion Transmission In addition to the structure of murine PrP, the NMR structures of the recombinant prion proteins from hamster [67, 68], human [69], and cattle [70] have been determined. All proteins consist of very similar global architectures, i.e., the flexible N-terminal tail of about 100 residues, followed by the folded C-terminal domain. The folds of C-terminal domains are again very similar but exhibit interesting local differences [57]. For example, α-helix 3 is kinked in the structure of murine PrP but straight in the structures of hamster, bovine, and human PrP. Comparison of the structures of mouse and hamster PrP revealed a better-defined loop segment between β-strand 1 and α-helix 2 and slight local differences between the structures in the segment between α-helix 1 and β-strand 2. In the structure of human PrP, α-helix 3 is also straight and well defined and extends to residue 228. Interestingly, the C-terminal turns of α-helices 2 and 4 appear to be in a local unfolding equilibrium. In addition, short, transient contacts between the flexible tail and the folded C-terminal domain were detected for human PrP, which may slightly contribute to a stabilization of the C-terminal residues of α-helices 2 and 3 against unfolding [57]. The structure of bovine PrP is most closely related to that of human PrP but shows the differences in local backbone conformation from murine and hamster PrP described above (Figure 5). Comparison of the structure of bovine PrP(121–231) with that from humans, mice, and hamsters revealed RMSD values of 0.98 Å, 1.66 Å, and 1.68 Å, respectively [57, 70]. The striking structural similarity between bovine and human PrPC , in conjunction with the fact that BSE prions can be transmitted to humans and cause new variant Creutzfeld-Jakob disease (CJD) (Table 1), suggests that the structural relationship at the level of PrPC inversely correlates with the species barrier of prion transmission. The species-barrier phenomenon has so far been discussed mainly at the level of sequence similarity between PrPs from different species with the view that the transmission barrier decreases with increasing sequence identity [30, 31]. A detailed structural comparison of all available PrP solution structures suggests that the most relevant structural feature with regard to the species-barrier phenomenon, besides the conformational differences described above, are differences in local surface charges, even though the uneven surface charge distribution on both flat surfaces of the C-terminal PrP domain is preserved in all structures [57]. In the context of the nucleation-polymerization mechanism of prion propagation, an inverse correlation between structural similarity of PrPC s from different species

11


Fig. 5 Comparison of the NMR structure of bovine PrP(121–231) (green, pdb code 1DX0) with that of recombinant PrP(121–231) from mice (yellow, code 1AG2), hamsters (pink, code 1B10), and humans (red, code 1E1J). The C" traces are represented as spline functions, and the

thickness of the cylindrical rods represents the mean global displacement per residue after superposition of the backbone atoms in the set of energy-minimized conformers that represents each NMR structure (picture from Dr. R. Riek, ETH Zurich).

and species barrier could mean that those parts of the PrPC structure that are similar between a pair of species but different in other PrPC structures are preserved in the structure of the PrPSc subunits and possibly are involved in subunit-subunit contacts. This could mean that substantial parts of the structure of the PrPC segment 121-231 are still contained in the tertiary structure of PrPSc subunits. In addition, a host-specific factor, called “protein X,” may be responsible for the species-barrier phenomenon. “Protein X” is believed to be contained in every mammalian species and to be required for prion propagation [71]. 3.3 Biophysical Characterization of the Recombinant Prion Protein 3.3.1 Folding and Stability of Recombinant PrP Studies on the folding and thermodynamic stability of murine PrP(121–231) at neutral pH and 25 ◦ C revealed that the C-terminal PrP domain behaves perfectly according to the two-state model as one would expect for a one-domain domain protein. It shows entirely reversible unfolding and refolding transitions in denaturantinduced equilibrium experiments, with a free energy of folding of −30 kJ mol−1 and a cooperativity expected for a 111-residue protein (m = 4.8 kJ mol−1 /M urea) [52]. In the context of full-length PrP(23–231), the stability of the domain is slightly decreased (–26 kJ mol−1 ), which is due to a somewhat lower cooperativity, while the transition midpoint is the same as that of PrP(121-231) (6.3 M urea). The lower folding cooperativity observed for full-length PrP may be due to formation of residual structure in the unfolded state in which residues 23-120 interact with the parts of segment 121–231 [55]. Reversible unfolding transitions at neutral pH were


also confirmed for human PrP(90–231) [72–75]. Hydrogen-exchange experiments on human PrP(90-231) revealed that only a small segment of about 10 residues around the disulfide bond retains residual structure in the unfolded state [74]. Further studies revealed that the stability of recombinant PrP decreases with decreasing pH [87, 96] and shows an unusual salt dependence in that salts destabilize the protein at concentrations below 50 mM increase the stability at high concentrations according to the Hofmeister series [96]. Assuming that folding of recombinant PrP corresponds to that of PrPC , the complete reversibility of PrPC folding nicely explains within the framework of the protein-only hypothesis why treatment of prions with denaturants such as urea or guanidinium chloride (GdmCl) completely abolishes infectivity, even after subsequent removal of the denaturant [76]: high denaturant concentrations dissociate PrPSc into monomers and denature its subunits, so that a solution of dissociated and unfolded PrPSc is identical to a solution of unfolded PrPC . Consequently, unfolded PrPSc subunits will always refold to PrPC . Analysis of the kinetics of folding of murine PrP(121–231) showed that the Cterminal domain belongs to the fastest folding proteins described so far. It folds with a half-time of about 200 µs at neutral pH and 4 ◦ C and unfolds with a halftime of 4.6 min under these conditions [77]. Rapid folding of PrP(121–231) is consistent with the absence of cis-proline peptide bonds in the three-dimensional structure [57, 67–70]. Analysis of the dependence of the rate constants of unfolding and refolding on denaturant concentration showed that the compactness of the transition state of folding is closer to the unfolded state than to the native state. No transient folding intermediates have been detected so far in the kinetics of folding of PrP(121–231). However a kinetic folding intermediate has been identified in the folding of recombinant human PrP(90–231) which nevertheless also folds extremely rapidly [78, 79]. Whether this difference is due to the additional segment 90–120 in human PrP(90–231) or reflects intrinsic differences in the folding of human and murine PrP(121–231) still has to be established. Overall the very rapid folding of PrP seems to exclude that kinetic folding intermediates can serve as a source of PrPSc at neutral pH. However the population of kinetic intermediates is uniformly higher in all recombinant human PrP(90–231) variants that bear mutations linked with inherited human TSEs [79]. PrPSc has been shown to accumulate in lysosomes of prion-infected cells [80–82] where the pH varies between 4 and 6 [83]. This opens the possibility that conversion of PrPC into PrPSc does not occur at the cell surface but rather in the acidic environment of lysosomes. For this reason, folding of recombinant PrP has also been analyzed in detail under acidic conditions, where the properties of the protein indeed change dramatically. At pH 4, the folding mechanism (U↔N) of murine PrP(121–231) changes from a two-state to a three-state equilibrium in which an acid-induced unfolding intermediate is observed by a pronounced plateau phase in the circular dichroism signal at 222 nm [72, 83, 84, 84–86]. At pH 4, the intermediate is maximally populated at 3 M urea and still significantly populated in the absence of urea [85]. In contrast to the α-helical CD spectrum of native PrP(121–231) at pH 4, the intermediate shows far-UV CD spectra of a

13


β-sheet protein with a single minimum at 215 nm that are reminiscent of PrPSc . The transition from two-state to three-state equilibrium transitions has an apparent pKa of 4.5, which indicates that protonation of acidic side chains is required for stabilization of the intermediate [85]. Very similar results were obtained for the folding of the full-length murine PrP(23–231) and human PrP(90–231). Consequently, formation of the acid-induced unfolding intermediate is an intrinsic property of the domain PrP(121–231). Further investigations demonstrated that formation of the intermediate is dependent on protein concentration and that the intermediate is most likely is a homodimer. In addition, ionic strengths above 50 mM are required for the intermediate to be observed. Equilibrium folding of recombinant PrP at neutral and acidic pH can thus by described by the following scheme [87]: pH 4–8, low ionic strength at pH 4–5: U ↔ N pH 4, ionic strength > 50 µM, [PrP] > 20 mM: 2 U ↔ I2 ↔ 2 N An attractive hypothesis resulting from these data is that the intermediate represents an early dimeric intermediate in the formation of oligomeric PrPSc . Indeed, starting with conditions that favor the population of the intermediate, fibrillar aggregates of recombinant PrP(90–231) could be obtained that showed a somewhat increased resistance to proteinase K [88]. Nevertheless, as in all attempts to generate prions de novo from recombinant PrP performed so far, it could not be demonstrated that these aggregates are infectious and relevant for understanding prion propagation. The same holds true for a β-sheet-rich conformation of human PrP(91–231), which is observed after thermal unfolding and cooling of the protein [86]. 3.3.2 Role of the Disulfide Bond in PrP The disulfide bond between the only cysteine residues of PrP, Cys179, and Cys214 is invariant in all mammalian prion protein sequences [31]. It is formed quantitatively in PrPC , and it has been shown unequivocally that free thiol groups are also absent in PrPSc [33]. Thus, there is primarily no indication that reduced PrP is involved in the mechanism of prion propagation. Nevertheless, a series of interesting recent observations argue in favor of a transient reduction of the disulfide bond during PrPSc formation. For example, it could be that the disulfide bond of PrPC , after caveolin-dependent endocytosis into lysosomes, becomes reduced and oligomerizes into reduced PrPSc in which the subunits again become re-oxidized when PrPSc is released into the extracellular space after death of the infected cell. Alternatively, it could be that reduced PrP is required for formation of PrPSc nuclei but that further incorporation of oxidized PrP into these nuclei eventually leads to a PrPSc oligomer in which the vast majority of subunits are oxidized, such that the small fraction of reduced PrP that was initially present is no longer detectable. The characterization of reduced, recombinant prion proteins has indeed revealed that the prion protein has the property of adopting two entirely different tertiary


structures, depending on whether or not the Cys179-Cys214 disulfide is formed. This was first observed for recombinant PrP(91–231) [89, 90]. Reduction of the disulfide bond in human PrP(90–231) at pH 4 and low ionic strength leads to transition of the predominantly α-helical protein to a monomeric polypeptide that shows β-sheet-like CD spectra [90]. Loss of chemical shift dispersion in 1 H–15 N heteronuclear NMR spectra, however, indicates that the tertiary structure content in this conformational state is significantly lower compared to that of the oxidized protein. Increase in ionic strength at acidic pH then leads to formation of amyloidlike fibers of reduced PrP(91–231), which show increased protease resistance [90]. In addition to these in vitro experiments, several studies on the expression of fulllength PrP in the reducing environment of the yeast cytoplasm have revealed that self-replicating amyloid-like fibers of reduced PrP are formed in vivo and show a protease-resistance pattern similar to that of the non-glycosylated PrP from PrPSc [91, 92]. In this context, it is very interesting that an in vitro protocol for transformation of recombinant hamster PrP(90–231) into amyloids could be developed that requires reduction and re-oxidation of the recombinant protein. The subunits in the resulting oligomers were connected by intermolecular disulfide bonds, which can be explained by a sub-domain swapping mechanism according to which each PrP molecule is covalently connected to two other PrPs via its cysteine residues and each oligomer has two non-satisfied ends with a free thiol group [93]. Here, the domain contacts and the structures of the monomers must be different compared to the X-ray structure of homodimeric human PrP(90–231), which lacks free thiols, as each Cys179 is disulfide-bonded with Cys214 of the other subunit of the dimer [62]. In summary, amyloid-like oligomers can be formed with both oxidized and reduced recombinant PrP in vitro. In none of these cases, however, could it be shown that infectious prions were generated. In this context, it should be emphasized that strong evidence has been accumulated during the past few years that nearly any protein, including purely α-helical proteins such as myoglobin that are not related to known amyloid diseases, can be converted into β-sheet-rich amyloid fibrils in vitro [94, 95]. This suggests that protein amyloids are primarily stabilized by β-sheet hydrogen bonds between main chain atoms. Thus, amyloid formation appears to be an intrinsic property of any polypeptide chain, independent of the amino acid sequence [95]. Therefore, it could well be that none of the aggregates of recombinant PrP reported so far are related to infectious PrPSc . 3.3.3 Influence of Point Mutations Linked With Inherited TSEs on the Stability of Recombinant PrP All known inherited prion diseases in humans are associated with mutations in the gene encoding human PrP. There are three different familial human TSEs, all of which are autosomal dominant: the familial Creutzfeld-Jakob diseases (CJD), Gerstmann-Straussler-Scheinker syndrome (GSS), and fatal familial insomnia (FFI) (Table 1) [1]. Figure 6A summarizes the location of amino acid replacements in human PrP linked with the three different disease phenotypes. Interestingly, the

15


mutation D178N causes either CJD or FFI, depending on the polymorphism at position 129 (Val or a Met, respectively). Figure 6B shows that disease-related mutations are mainly located in the folded C-terminal domain of PrP. There is, however, no structural clustering of amino acid replacements related to a specific disease phenotype. As the affected individuals with a point mutation in one of the two PrP alleles spontaneously develop prions without having had contact with external prions, an obvious mechanism underlying this phenomenon could be a destabilization of PrPC due to the mutation and, consequently, a facilitated conversion to PrPSc . Inspection of the NMR structure of recombinant PrP indeed predicts a destabilization in the case of the replacements D178N (loss of a salt bridge to Arg164), T183A (loss of two hydrogen bonds from the hydroxyl group of Thr183 to the HN of Tyr163 and the carbonyl oxygen of Cys179), F198S (generation of a cavity), and Q2017R (loss of a hydrogen bond to the carbonyl atom of Ala133) [57, 58]. However, there are no clear-cut predictions for the other amino acid replacements, and the GSS mutations P102L, P105L, and A117V are located in the flexible tail and should not affect the stability of PrPC at all. Therefore, all disease-related point mutations were introduced into recombinant PrP(121–231), and the thermodynamic stabilities of the variants were analyzed. All variants showed far-UV CD spectra that were indistinguishable from that of the wild-type protein, demonstrating that none of these mutations a priori induces a PrPSc -like, β-sheet-rich conformation [55]. The thermodynamic stabilities of the PrP(121–231) variants with single disease-related amino acid replacements are summarized in Figure 6C and reveal that some but not all mutations destabilize PrPC . A significant destabilization was indeed observed only for the above replacements that were predicted to be destabilizing from analysis of the threedimensional structure. The most destabilizing mutation proved to be T183A, which decreases the free energy of folding relative to the wild-type protein from −30 to −10 kJ mol−1 [55]. Other mutations such as E220K or V210I were completely neutral with respect to PrPC stability. Analogous results were obtained for variants of recombinant PrP(90–231) [62]. Consequently, destabilization of PrPC cannot be the only mechanism underlying the spontaneous formation of prions in inherited human TSEs, and destabilization may trigger disease only in the case of mutations D178N, T183A, F198S, R208H, and Q217R (Figure 6C). Another result from this analysis is that there is no correlation between the thermodynamic stability of a TSE-related PrPC variant and the disease phenotype. For example, both the most destabilizing mutation T183A and the “neutral” mutation E200K are linked with inherited CJD (Figure 6). It follows that there are most likely multiple and independent mechanisms that can lead to spontaneous generation of prions as a consequence of a mutation in PrP. Such mechanisms may, e.g., be an increased stability of PrPSc (“wild-type” PrPSc has a half-life of about two days in vivo) or faster nucleation kinetics of PrPSc . The latter may be caused by the above-mentioned higher fraction of kinetic folding intermediates that has been reported for variants of human PrP(90–231) bearing amino acid replacements associated with inherited TSEs [78, 79].


Fig. 6 Point mutations in human PrP that are associated with inherited prion diseases and their effect on the intrinsic stability of recombinant PrP(121–231). (A) Scheme of the primary structure of mature human PrPC , with the amino acid replacements that are linked with the three inherited human TSEs: CJD (blue), GSS (red), and FFI (pink). The Met/Val polymorphism at residue 129 (gray) determines the phenotype of the disease related to the replacement D178N (FFI or CJD, respectively). (B) Location of residues that are replaced in inherited TSE in the structure of PrP(121–231). The

polymorphism in murine PrP at residue 190 is also shown (same color code as in (A)). (C) Thermodynamic stabilities of recombinant variants of PrP(121–231), with disease-related, single amino acid replacements, relative to PrP(121–231) wild type. Positive Go values indicate that the variant is less stable than wild-type PrP(121231) (same color code as in (A)). Wild-type PrP(121–231) has a free energy of folding of −30 kJ mol−1 . The least stable variant T183A is thus destabilized threefold relative to the wild-type protein.

17


4 Generation of Infectious Prions in vitro: Principal Difficulties in Proving the Protein-Only Hypothesis

Most scientists would consider the in vitro generation of prions from purified PrPC the final proof of the protein-only hypothesis [8]. In principle, there are several ways to perform such an experiment. One approach is the propagation of prions by inoculation of purified PrPC (or recombinant PrP) with catalytic amounts of PrPSc isolated from the brain of prion-infected animals. Another (and possibly even more convincing) experiment is the de novo generation of prions from purified PrPC or recombinant PrP by applying in vitro conditions that favor both spontaneous formation and propagation of PrPSc . The only way to monitor the success of these experiments is to test newly converted PrPSc molecules for infectivity. This is done by injection of prions into the brain of transgenic mice that overexpress PrPC , and are therefore particularly susceptible to prions, and determination of the incubation time until the onset of the disease. Unfortunately, there is only a very rough, inverse linear correlation between the logarithm of the concentration of infectious units and incubation time [48, 97]. In practice, this means that differences in prion concentration between two samples (e.g., before and after an in vitro conversion experiment) can be detected reliably only when the concentrations differ by two to three orders of magnitude. If we assume that a difference in concentration of infectious units by a factor of 100 is minimally required to distinguish two prion preparations, a conversion experiment in which an infectious PrPSc preparation is used as an inoculum has to be designed such that PrPC is used at 100-fold excess over PrPSc and that the conversion would be 100%. It is, however, obvious that the conditions for an in vitro conversion experiment will not be optimal. Thus, if the conversion efficiency was, e.g., only 1%, PrPC would have to be used at an excess of at least 10000-fold over PrPSc . In summary, the low sensitivity of the only available assay for infectious prions requires very high in vitro conversion efficiencies for detection of newly generated prions. The most promising attempts to convert PrPC to the protease-resistant, oligomeric form (which is also termed PrPres ) are experiments in which 35 S-labeled PrPC isolated from mammalian cells is incubated with PrPSc in vitro [47, 98]. Numerous experiments performed according to the following reaction scheme revealed that 35 S-labeled PrPC can indeed be converted in vitro into a protease-resistant form, PrPres , which has the same biochemical properties as proteinase K-treated PrPSc [47]. 35 S-labeled PrPC from mammalian cells + PrPSc from prion-infected individuals Incubation ↓ Proteinase K treatment ↓ Detection of protease-resistant35 S-PrP(Prpres ) ↓

5 Understanding the Strain Phenomenon in the Context of the Protein-Only Hypothesis 19

Further mechanistic studies demonstrated that this in vitro conversion of PrPC to PrPres is characterized by a rapid and specific binding step of PrPC to PrPSc oligomers, followed by a slow conformational transition to PrPres in the PrPSc -bound state [47]. Strikingly, this assay reproduces experimentally determined species barriers of prion diseases, in that PrPSc preparations from one species convert PrPC from another species with similar efficiency, as the disease can be transmitted between the two species [99]. Unfortunately, the seeded in vitro conversion of PrPC to PrPsen generally does not generate more than stoichiometric amounts of PrPres relative to the PrPSc molecules used as seeds [47], which prevents detection of newly generated infectivity. However, the seeded in vitro conversion has been significantly improved recently by a protocol in which growing PrPSc /PrPres oligomers were subjected to repeated rounds of sonification and re-incubation with PrPC , with the idea of breaking growing oligomers into smaller nuclei by the sonification step and thereby increasing the concentration of “growing ends” [100]. Indeed, this PCRlike approach strongly increased the efficiency of the in vitro conversion process, such that a 50-fold molar excess of converted PrPsen over the initial PrPSc seeds was achieved. Despite this significant improvement, it could still not be shown that the concentration of infectivity increased and new prions were generated. In any case, this new method is predicted to improve the sensitivity of prion assays based on the detection of protease-resistant PrP by almost two orders of magnitude. Regarding the above described principle difficulties in proving generation of prions from PrPC when PrPSc is used as an inoculum, the de novo generation of prions from recombinant PrP, which requires only a qualitative proof of existence of infectivity, may be the more promising approach towards the final proof of the protein-only hypothesis. However, the seeded in vitro conversion method has the potential of being further improved through the finding that mammalian RNA preparations stimulate the in vitro amplification of PrPres [101]. This hints at a role for host-encoded stimulatory RNA molecules in prion pathogenesis, which is, e.g., principally possible after retrograde transport of PrP into the cytosol.

5 Understanding the Strain Phenomenon in the Context of the Protein-Only Hypothesis: Are Prions Crystals?

The most frequently raised argument against the protein-only hypothesis is the occurrence of prion strains. This relates to the fact that different prions can be isolated from a single species that differ in incubation time, clinical signs, vacuolar brain pathology, and PrPSc deposition in the brain [8, 23]. These specific phenotypic properties of a given prion strain are reproducibly preserved during consecutive rounds of experimental infection and re-isolation of the strain from the brain of the infected individual. Strain phenomena are typically observed for viruses, which are subject to high mutation rates of the viral genome. The prion strain phenomenon can, however, be explained in the context of the protein-only hypothesis on the basis of the nucleation-polymerization model of prion propagation [47]. This is because


Fig. 7 Explanation of different prion strains within the framework of the protein-only hypothesis: (A) Besides disease symptoms and incubation time, prion strains isolated from mammalians and humans can be distinguished on the basis of the biochemical properties of PrPSc subunits after proteinase K treatment and separation by SDS-PAGE and Western blotting. Specifically, prion strains differ in the fractions of non-glycosylated, mono-glycosylated, and doubly glycosylated PrP contained in PrPSc oligomers, as wel as the N-terminus of the protease-resistant core of the individual subunits. The figure schematically shows the Western blot band patterns of four different human prion strains after proteinase K treatment of PrPSc : (1) sporadic

CJD, Met/Met 129 genotype; (2) sporadic CJD, Val/Val 129 genotype; (3) iatrogenic CJD, Val/Val129 genotype; (4) variant CJD, Met/Met129 genotype. (B) The simplest explanation for the occurrence of different prion strains are crystallike PrPSc oligomers that differ in the fractions of the incorporated PrP glycoforms, the arrangement of subunits in the quaternary structure, and/or slightly different tertiary structures of incorporated subunits. In this model, each prion strain corresponds to a specific crystal form. Only one crystal form of PrPSc would then occur in a prion-infected individual, and only this crystal form would be further propagated when the disease is transmitted to another individual.

there is very convincing evidence that prion strains differ not only with respect to the above phenotypic criteria but also in the biochemical properties of their PrPSc deposits [6, 47, 102]. Figure 7A schematically depicts a Western blot analysis of human PrPSc from the brain of different patients after proteinase K digestion and SDS-PAGE separation [102]. It shows that the proteinase K– resistant core of the PrPSc oligomers that are associated with a given strain differs both in the fractions of the incorporated PrP glycoforms (in mammalian cells, there is always a mixture of non-glycosylated, mono-glycosylated, and di-glycosylated PrPC ) and in the exact length of the proteinase K–resistant polypeptide fragments. Like the phenotypic

6 Conclusions 21

properties described above, the characteristic strain-specific banding patterns are always preserved, even after crossing the species barrier, and thus are diagnostic for each prion strain. For example, a characteristic feature of BSE prions from cattle is the high fraction of di-glycosylated PrP contained in the PrPSc oligomers, which is also observed in PrPSc isolated from patients with variant CJD [102] (Figure 7A, Table 1). The simplest model that explains these similarities is that PrPSc is a crystallike oligomer. Similar to the fact that proteins frequently form different crystal forms, even under identical crystallization conditions, PrPSc appears to be capable of forming multiple macromolecular assemblies that differ in subunit composition and arrangement in their quaternary structures and possibly also in the tertiary structures of their subunits (Figure 7B). Each prion strain thus would correspond to a different PrPSc “crystal form,” and this single crystal form then acts as a seed for formation of new PrPSc oligomers that grow further in the same crystal form. This model can also explain why humans who are heterozygous with respect to the polymorphism at amino acid 129 (Met or Val in humans) are essentially protected from sporadic CJD: although about 51% of the European population is Met/Met or Val/Val homozygous and 49% is Met/Val heterozygous, 95% of all sporadic CJD patients are homozygous [103, 104]. If we assume that residue 129 is solventexposed in PrPSc subunits (as it is in PrPC ; Figure 6B) and involved in subunitsubunit contacts in PrPSc , it is easy to imagine that a PrPSc oligomer composed of identical polypeptide chains could more easily assemble spontaneously into a quaternary structure with a regular array of subunits, compared to a 1:1 mixture of different polypeptides that are incorporated statistically into the oligomer. Another remarkable finding in this context is that residue 129 is exclusively Met in cattle PrP, and variant CJD patients are exclusively Met/Met homozygous [105].

6 Conclusions

Although essentially all available experimental data on TSE prions and their propagation are either in agreement with or can be explained within the framework of the protein-only hypothesis, the most convincing proof of the prion hypothesis, i.e., the experimental replication of TSE prions in vitro, has not been reported to date. Some of the principal difficulties in proving the generation of infectious prions in vitro by seeding experiments have already been addressed in Section 4. In addition, if the “crystal model” of PrPSc that explains the strain-specific fractions of the different PrP glycoforms in PrPSc is correct, it follows that the PrP glycans are critically involved in subunit-subunit contacts that determine the quaternary structure of PrPSc . In the worst case, this means that it will not be possible to produce a PrPSc oligomer that is infectious and identical to PrPSc isolated from prion-infected brains from non-glycosylated, recombinant PrP. Moreover, experiments on the de novo formation of prions from recombinant PrP have so far essentially been performed in solution, while the conversion process in vivo most likely occurs at the surface of membranes. The covalent linkage to the GPI anchor and a membrane


localization of PrP may therefore be additional prerequisites for generation of prions in vitro. A new protocol for the preparation of recombinant PrP covalently attached to liposomes has recently been established [106] that will allow in vitro conversion experiments with membrane-bound, recombinant PrP. In addition, like cellular RNAs [101], the glucose-based polyglycans found to be associated with PrPSc [49] may be another critical cofactor of prion replication. A more general question is which experimental strategies should be pursued to gain mechanistic insights into prion propagation at the molecular level. Because there appears to be an inverse correlation between structural similarity at the level of recombinant PrP and species barrier, the determination of additional structures of recombinant PrPs is certainly a promising approach towards a further understanding of the species barrier. Even more important, however, appears to be the investigation of the oligomeric structure of PrPSc . For example, PrPSc oligomers that are spontaneously formed in inherited human prion diseases can either be homo-oligomers of the mutated prion protein or hetero-oligomers of mutated and non-mutated PrP. Both cases have been reported (see references in [55]), but no comprehensive mechanistic investigation on these phenomena is available. As PrPSc is insoluble and not accessible to structure determination by NMR spectroscopy and X-ray crystallography, the combination of electron microscopy, electron crystallography, and biochemical and genetic studies will be required to obtain further insights into the molecular details of the PrPSc quaternary structure. In contrast to mammalian TSEs, the protein-only hypothesis can be considered proven in the case of the yeast prions Ure2p and Sup35p, which are responsible for the non-genetically transmissible phenotypes [URE3] and [PSI], respectively [107–109]. Specifically, infectious aggregates of Sup35p have been generated from recombinant, bacterial protein in vitro and then successfully transmitted to yeast cells that further propagated the [PSI] phenotype [110]. In addition, the speciesbarrier phenomenon was unequivocally shown to be caused by specific aggregates of Sup35p from different yeasts that replicate independently in the same test tube [111]. Thus, there are no principle reasons that the protein-only hypothesis should not be valid for mammalian TSEs.

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1

Prion Protein

Philippe Derreumaux Institut de Biologie Physico-Chimique–CNRS, Paris, France

Originally published in: Amyloid Proteins. Edited by Jean D. Sipe. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31072-X

1 Introduction

The deposition of amyloid fibrils sharing a common cross-β-sheet structure with the β-strands perpendicular to the fiber axis is a hallmark of several fatal diseases [1–3]. These disorders associated with the failure of proteins to fold correctly can affect the brain, the central nervous system, and various organs such as the liver and heart in humans and animals. The list of fatal diseases includes, among others, primary and secondary systemic amyloidosis, Alzheimer’s, Parkinson’s and Huntington’s diseases, diabetes Type 2, and transmissible spongiform encephalopathies (TSE). The first TSE symptom observed is pronounced astrogliosis [4], followed by spongiform degeneration caused by the formation of vacuoles in neuronal processes and astrocytes [5]. The late step in the TSE disease often involves the formation of amyloid plaques containing PrPSc [6], the abnormal (or scrapie) isoform of the cellular prion protein (PrPC ) encoded by the PrP gene and found predominantly on the outer surface of neurons. The “protein-only” hypothesis states that a single prion (proteinaceous infectious particle) protein, lacking a small nucleic acid, can give rise to multiple isolates or strains with varying infectivity and incubation time, and is able to convert PrPC to PrPSc in an autocatalytic manner [7–9]. This theory is at variance with the virus [10] and virino [11] hypotheses which postulate that the infectious particle consists of a small nucleic acid coated by PrPSc . The original translation product consists of 253 amino acids, but region 1–22 is cleaved as signal peptide during trafficking and region 232–253 is replaced by a glycosylphosphatidylinositol (GPI) anchor at position 231. As seen in Fig. 1, mammalian PrPC is a highly conserved secretory cell surface glycoprotein of approximately 210 amino acids (residues 23–231). PrPC is also characterized by two glycosylation sites at N181 and N197, and a single disulfide bond between the cysteines C179 and C214. PrPC and PrPSc Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Prion Protein

share the same primary structure – no sequence or post-translational differences have been detected [12] – but have distinct biophysical and biochemical properties. PrPC is monomeric, soluble in detergents and sensitive to proteinase K, while PrPSc is oligomeric, insoluble and partially resistant to proteinase K [13, 14]. In addition, the conversion of PrPC to PrPSc involves a large variation in secondary structure as determined by IR spectroscopy: PrPC has 42% α-helix and 3–5% β-sheet, whereas PrPSc has 30% α-helix and 43% β-sheet, and PrPSc 27–30 (i.e. PrPSc with removal of residues 23–90) has 25% α-helix and 48–54% β-sheet [15–18]. Despite decades of study and recent promising discoveries, scientists do not know how to prevent prion diseases or what leads prion proteins to become misfolded and toxic. No purified recombinant PrP has been successfully converted in vitro to infectious PrPSc . Yet, scrapie in sheep and goats was first recognized in the 18th century, and mad cow disease or bovine spongiform encephalopathy (BSE) was discovered in 1986. In humans, different types of TSE were identified during the 20th century. Human TSEs are essentially sporadic (80%) with the modes of natural transmission remaining undetermined; around 15% are inherited [familial forms of Creutzfeldt–Jakob disease (CJD), fatal familial insomnia (FFI) and Gerstmann–Sträussler syndrome (GSS)] by mutations in the human PrP gene within chromosome 20. Sporadic CJD occurs at a rate of 0.5–1 per million population per year and GSS, which is the most common familial TSE, generally occurs in the third or fourth decade of life. New pathogenic mutations in the human prion protein continue to be discovered and the current identified list is given in Fig. 1. Finally, around 5% of human TSEs are infectious through CJD-infected surgical equipment or tissue transplants; kuru, recognized in 1957, results from exposure to contaminated human tissues during endocannibalistic rituals, whereas new variant CJD (vCJD), identified in 1996, is believed to be spread by dietary exposure to the BSE agent [19] and might also be transmitted by blood transmission [20]. In this chapter, we review our current understanding of the underlying factors that may promote the topological change of mammalian prion proteins. We have selected to focus here on the structural, thermodynamic and dynamic aspects of the in vitro and in vivo conversion. Reviews on the link between misfolding, pathogenesis and neurotoxicity on the pathological implications of prion glycosylation, on the differences between mammalian and yeast prions, and, finally, on the possible destinations taken by PrPC can be found elsewhere [21–24].

2 Conformations of PrPC and PrPSc

Knowledge of the three-dimensional structure of PrP in its protease-sensitive and protease-resistant forms is important for our understanding of prion replication. However, both the PrPC and PrPSc structures are still unknown at atomic resolution. Amyloid fibrils are non-crystalline and insoluble, and therefore not amenable to X-ray crystallography and nuclear magnetic resonance (NMR) spectroscopy. Furthermore, there is accumulating evidence that the PrPSc structure is not unique


Fig. 1 Species variations and mutations of the prion protein gene. (A) The octarepeats within the tail are represented by white boxes, the regular secondary structures (helices: H1, H2 and H3; $-strands: S1 and S2) within the NMR PrPC structure [28] are also shown. The disulfide bond (SS) between Cys179 and Cys214 is drawn as a red line; the GPI anchor at position 231 and the two glycosylation sites N181 and N197 are also shown. Polymorphisms and point mutations associated with known human GSS, CJD or FFI are given. The

mutations H187R, T188R, T188A, T188K and P238S are still unclassified [137]. (B) Aligned amino acid sequences of human, bovine, hamster, mouse, sheep, chicken and duck PrP. The multiple alignment was carried using T-Coffee [136]. The reliability of the alignment is given in a color code (e.g. red means very reliable). In the last line below each aligned sequences, the asterisk indicates the strictly conserved residues. Note that the fragment spanning residues 106–126 is fully identical between the seven species.

3

4 Prion Protein

Fig. 2 Mutations of the PrP gene are associated with prion diseases. (A) The NMR recombinant Syrian hamster PrPC NMR structure [28] with the three "-helices H1, H2 and H3, the two $-strands S1 and S2, the disulfide bond between the cysteines C179 and C214, the GPI anchor, and the two glycosylation sites N181 and N197. For

clarity, only residues 120–231 are shown. (B) The positions of all polymorphisms and disease-causing mutations as discussed in Fig. 1. The mutations leading to GSS are given in green; those leading to CJD are in red. Here, all residues spanning PrP90–231 are shown.

and differences in the conformation of the infectious protein determine prion strain variation [25, 26]. Based on far-UV circular dichroism (CD) and limited proteolysis experiments, PrP and recombinant PrPs in membranes (lacking glycosylation sites) or in solution (lacking both GPI anchor and glycosylation sites) seem to adopt similar conformations [27], but no detailed structure is available for exact comparison. Thus far, only the solution structure of recombinant PrP from various mammalian species was characterized by NMR spectroscopy and X-ray diffraction. All NMR solution structures consist of a disordered tail (residues 23–123), and a globular domain with three α-helices and a short antiparallel β-sheet (Fig. 2). The β-strands S1 and S2 comprise residues 128–131 and 161–164, respectively helix H1 spans residues 144–153, helix H2 residues 172–194 and helix H3 residues 200–225 (numbering in Syrian hamster [28]). Although the NMR structures from mouse PrP121–231 [29], bovine PrP23–230 [30], Syrian hamster PrP23–231 [28] and human PrP121–230 at pH 7 and 4.5 [31, 32] superpose well, conformational changes are observed within the loops connecting the helices and even in the length of the helices. The disorder in the loops is an intrinsic property of PrP as deduced by hydrogen-deuterium exchange measurements [33] and is independent of the oligomeric (monomeric or dimeric) state of PrP, as deduced by molecular dynamics (MD) simulations [34]. Helix H3 spans residues 200–222 in mouse PrP121–231 versus 200–228 in human PrP90–231 and helix H2 is 12 residues shorter (173–182) in the human PrP125–228(S170N) variant [35]. Strikingly, the crystal PrP structures do not always superpose on the NMR conformations. The crystal structure of recombinant human PrP119–226 shows a


covalently dimer with an intermolecular disulfide bridge, swapping of helix H3 and formation of an interchain antiparallel β-sheet through residues 190–194, i.e. in helix H2 within the monomeric NMR structure [36]. In contrast to human PrP, the crystal structure of recombinant sheep PrP123–230 points to a monomeric state with an intramolecular disulfide bridge, superposing within 1.7 Å r.m.s. deviation from the NMR human PrP structure [37]. This X-ray structure also shows intermolecular or lattice contacts between the strands S1, allowing for the formation of a four-stranded intermolecular β-sheet and the possibility that the residues 190–194 could easily shift from an α-helix to a β-strand [37]. Whether the dimeric form with swapping of helix H3 and intermolecular disulfide bridge is an artifact of the crystallization protocol or indicates a physiological state remains to be determined. An essential aspect of the recombinant PrP structure is that the entire tail encompassing residues 23 to around 120 is highly flexible and largely disordered for a wide range of pH values. This is not totally surprising since the tail spanning residues 23–90 contains around 40% of glycine residues. Region 90–120 is, however, critical in prion propagation. Residues 108–111 influence prion replication efficiency [38]. PrP with the region 113–120 deleted is not converted to a protease-resistant form when expressed in scrapie-infected neuroblastoma cells [39]. Mice expressing PrP with a deletion of residues 32–106 are not susceptible to infection in vivo [40]. In addition, the mutations P102L and P105L lead to GSS syndrome and the mutation P101L in murine PrP (P102L in human numbering) alters TSE incubation time across three species barriers [41]. The flexible tail contains the octapeptide PHGGGWGQ repeated 4 times from residues 60 to 90 which binds copper within the physiological range [42]. Using PrP-derived peptides, full-length Syrian hamster PrP and electron paramagnetic resonance measurements, copper was found to interact not only with each HGGGW segment of the tail, but also with the PrP92–96 segment GGGTH through the His96 imidazole group and the carboxyl oxygen atom of Gly94 [43]. The 106–126 peptide, cytotoxic in vivo [44] and containing the palindromic sequence VAGAAAAGAV, was shown to form fibrils with parallel β-sheets [45], but the extrapolation of this result to PrP90–231 remains to be determined. Recent NMR studies of recombinant human PrP23–230 at pH 6.2 showed that the octapeptide repeats are structured, with the segments HGGGW and GWGQ adopting a loop conformation and a β-turn, respectively [46]. This result, along with the crystal structure of the copperbinding pentapeptide HGGGW-Cu2+ [47], suggests that the conformations of the HGGGW loop depend on both pH and copper binding. Three PrPSc models were first constructed on the basis of immunological studies and CD spectra. In the first model, the region between residues 90 and 145 was modeled by two consecutive β-hairpins (four β-strands) with β-strand 1 parallel to β-strand 3 [48]. In the second model, the secondary H1, S1 and S2 structural elements were replaced by a Greek key consisting of four adjacent antiparallel β-strands [49], whereas in the last model, PrPSc adopted β-helical conformations [50]. More recently, three other PrPSc models were proposed on the basis of specific criteria and electron microscopy data at 7 Å resolution from in vitro infectious Syrian and mouse PrP two-dimensional protofibril crystals. Fig. 3 shows

5

6 Prion Protein

Fig. 3 Schematic representation of the secondary structures of wild-type recombinant Syrian hamster PrPSc by NMR [28], and the three PrPSc models proposed by Wille et al. [51], Mornon et al. [53] and De Marco

and Daggett [54]. Helices H1, H2 and H3 are presented by grey boxes; S1, S2 and the new $-strands by black boxes; $-helical conformations by cross-hatched boxes.

the fluctuations of the secondary structures within these predicted PrPSc models. Wille et al. proposed a hexamer with each monomer featuring β-helices in the region 90–170 and α-helices H2 and H3 formed using threading techniques onto known β-helical structures. Nevertheless, they did not exclude the possibility that PrPSc could adopt a novel protein fold [51]. Mornon et al., using sequence analysis, identified the TATA box-binding protein fold as a template for PrPSc which allowed for the construction of another hexameric PrPSc model [52, 53]. In their model, the S1H1S2 motif and the N-terminal end of helix H3 are preserved, whereas the helix H2 and its neighboring residues are transformed into three new β-strands. Finally, De Marco and Daggett used MD simulations of the Syrian hamster PrP109–219 (D147N) at low pH to generate β-rich conformations and then docking between the units to build an alternative hexameric PrPSc model [54]. In this model, the three helices are preserved, and the core of the β structure consists of an isolated and new β-strand preceding helix H1, and a new β-strand spanning residues 116–119 packed against the S1 and S2 strands. Since it is possible to generate several PrPSc models within the electron microscopy envelope, other experimental data are needed to discriminate the solution from false positives. These can include differential proteinase K, vibrational spectroscopies and antibodies which recognize selectively PrPC or PrPSc . The model of Wille et al. [51] with β-helices was recently questioned using vibrational Raman optical activity measurements, which pointed to strong similarities between the spectra of PrPSc and flat β-sheet proteins, but not β-helical proteins [55]. The model of Mornon et al. is consistent with a recently discovered antibody recognizing the YYR motif within the strand S2 of PrPSc [56] and with the antibody V5B2, raised against residues 214–226, which recognizes PrPSc , but not PrPSc , indicating a structural rearrangement of the C-terminus [57]. The model of De Marco and Daggett [54] is consistent with peptide-binding studies of PrPSc [58, 59] potently inhibiting the PrP-res induced cell-free conversion of PrP-sen to the protease-resistant state. This model is also consistent with the monoclonal antibody 15B3, which recognizes

3 Stability and Unfolding/Folding of PrPC In Vitro

PrPSc at positions 142–148 (N-terminal half of helix H1 in the normal protein), 162–170 and 214–226, but not PrPC [49]. However, neither Mornon’s or De Marco’s model discuss the conformations of the fragment 90–109, and fail to reproduce, in their present forms, the experimental percentage of β structure. Fourier transform IR studies suggest that 48% of residues (or about 68 residues) [18] are in β-strands within PrPSc 90–231, whereas for instance the De Marco’s model has 37 residues in β-strands within the region 109–219. These differences require study in more detail.

3 Stability and Unfolding/Folding of PrPC In Vitro

An important step towards understanding the conformational conversion of PrPC to PrPSc is to characterize the energy landscape and unfolding/folding pathways of the wild-type and mutant prion proteins in vitro. It is well established that the recombinant prion protein can be folded either to a monomeric α-helical topology or β-rich oligomers and that folding to the PrPC isoform is under kinetic control [60]. Starting from disordered conformations, PrP folds to its PrPC isoform at pH 7, whereas under acidic pH conditions, PrP avoids the kinetic trap and folds to the β-rich isoform. The energy barrier separating the PrPC state and the PrP βrich oligomers was estimated to be 35–45 kcal/mol for wild-type mouse PrP [60]. There is also strong evidence that the full tertiary context of the protein is necessary to stabilize the terminal α-helices and the β-strands within PrPC . Biophysical studies showed that the helices H2 and H3 are largely disordered in the PrP fragment spanning helices H2 and H3 [61]. Monte Carlo simulations suggested that the sequence PrP127–164 can adopt two distinct tertiary folds with different secondary structures [62]. Initial experiments on recombinant mouse PrP121–231 at 298 K, based on stopped-flow fluorescence using the variant F175W which has the same overall structure and stability as wild-type mouse PrP121–231, suggested a two-state folding model at neutral pH, i.e. PrP either unfolded or native. The mouse PrP121–231 fragment was also found to fold very rapidly to PrPC with a half-life of 170 µs [63]. Then, other kinetic studies on human PrP90–231 and mouse PrP121–231 suggested that the change from neutral to acid pH conditions shifted the two-state kinetic to a three-state kinetic by stabilizing a monomeric intermediate with β-sheet structure in equilibrium with the native helical state. An intermediate state was detected at pH 4.0 and 1–1.5 M Gdn-HCl [64], pH 3.6 and 0.75–1.75 M Gdn-HCl [65], pH 5.0 and 2–2.5 M Gdn-HCl [66], and 3.5–4.5 M urea [67]. These β-sheet-rich intermediates, generated under acidic pH conditions (pH below 5), are, however, oligomeric rather than monomeric in character [68–71]. More recently, by following the kinetics of folding and unfolding reactions of human PrP90–231 at 278 K and pH 7, it was again proposed that the prion protein folds by a three-state mechanism involving a monomeric intermediate [72]. Since then, the existence of a folding intermediate, PrP*, has been advocated by a number of studies. These include

7

8 Prion Protein

hydrogen exchanges monitored by NMR [73] and high-pressure NMR experiments on Syrian hamster PrP90–231 [74] – the population of this intermediate was found to be 1% at pH 5.2, 303 K and 1 bar – or mouse PrP121–231 [75]. The point mutations associated with inherited human TSEs are also of interest for characterizing the energy landscape. It was initially believed that mutations in PrP gene could promote the conformational conversion by destabilizing the native structure of PrPC [76]. Fig. 2 shows the positions of polymorphisms and diseasecausing mutations within the NMR human PrP structure. The most frequent polymorphism, modulating disease susceptibility and onset, is located at codon 129. For instance, the M129/N178 allele segregates with FFI, whereas the V129/N178 allele leads to CJD [77]. The codon stop mutation at position 145 which generates an atypical GSS variant is also given [78]. As seen in Fig. 2, most familial mutations are located within the globular domain 124–226: strand S1 (G131V), helix H2 (D178N, V180I, T183A, H187R, T188A or T188K or T188R), loop between H2 and H3 (E196K and F198S), and helix H3 (E200K, D202N, V203I, R208H, V210I, E211Q, Q212P and Q217R). However, three mutations also occur within the disordered N-terminal tail: P102L, P105L and A117V. Although destabilization of the PrPC structure upon mutations has been confirmed for D178N, T183A, F198S, R208H and Q217R mutants by equilibrium unfolding studies in urea [79–81], it is not a general mechanism underlying the formation of PrPSc . The structure of the E200K variant of human PrP superposes well on the wild-type sequence structure [82]. The M129V and P102L variants of human PrP show PrPC -like structural properties from CD analysis [79, 83]. The recently discovered G131V mutation does not lead to any identifiable effects on PrPC secondary structure, as deduced by MD simulations [84]. Clearly, other mechanisms rather than changes in the thermodynamic stability of PrPC are thus important for conversion. One solution is that single-point mutations reduce the energy barrier by stabilizing the transition state. This effect has not been fully explored yet. Another possibility is that the point mutations change the electrostatic energy surface of PrPC and thus alter the interaction (binding or conversion step) with PrPSc . This effect has been proposed for the E200K mutant [82]. Finally, it is possible that the point mutations stabilize a partially folded intermediate. Recent kinetic experiments on a number of human PrP90–231 variants carrying mutations associated with familial forms of prion disease suggest the existence of partially structured intermediates on the refolding pathway of the following PrP variants: F198S, Q217R, V180I, V210I, R208H, D178N/M129 and D178N/V129. In each case, the partially folded state was found to be at least an order of magnitude more populated than the fully unfolded state. The strongest effect was seen for the variant F198S where the ratio of intermediate states relative to the native state is 1/350 versus 1/42,000 for wild-type PrP [81]. In contrast, the P102L mutation did not lead to any increase in the population of the intermediate, leaving unanswered the question of its impact on the energy surface. While it is not a definitive proof for the existence of a monomeric PrP* intermediate, it is interesting that such an intermediate has not been detected for Doppel, a homolog of PrP that does not form infectious prions [73].

4 Mechanisms of Prion Replication In Vivo 9

Little is still known on the structure of the monomeric PrP* intermediate. Highpressure NMR experiments point to an intermediate with the helices H2 and H3 disordered and helix H1 formed at pH 5.2, 303 K and 1 bar [74]. MD simulations of PrP90–231 with either the strand S1 or the strand S2 deleted – these variants do not inhibit prion propagation in a cell-free assay system [85] – suggest partially unfolded intermediates of molten globule type with the helices H2 and H3 disordered, and the helix H1 either fully formed or partially disordered at its C-terminal end [86]. Clearly, the structural features of PrP*, if this intermediate exists in monomeric form, need to be investigated in more detail. Nonetheless, based on MD simulations coupled with φ analysis on globular proteins, it would not be too surprising that PrP* consists of an ensemble of distinct conformations having different secondary structure compositions [87].

4 Mechanisms of Prion Replication In Vivo

Within the protein-only hypothesis, a detailed mechanism for the conformational transition is still unclear. A number of facts are, however, well accepted. Interactions between PrPC and PrPSc provide the main forces to propagate the topological change [60]. The conversion is promoted by partially denaturing conditions and in the presence of low concentrations of urea, i.e. under conditions expected to increase the population of an intermediate. Irrespective of the kinetic model (see below), transition from PrPC to PrPSc is a two-step process, which begins with binding between the two PrP isoforms, followed by conversion of the cellular form to the pathogenic form [88]. By designing specific synthetic peptides which inhibit binding and conversion reaction, it was found that the binding surfaces include residues 119–138 (region preceding helix H1), 165–174 (loop between S2 and H2) and 206–223 (helix H3), while conversion to PrPSc is strongly influenced by residues 139, 155 and 170 [58, 59, 88]. The species barrier, which is the most intriguing feature of prion propagation, limits transmission of prions across species. Mouse scrapie prion can be transmitted to Syrian hamster with an incubation time of 380 days, but transmission in mice is 130 days [89]. At present, two main factors have been identified to contribute to the species barrier from studies on transgenic animals: the conformations of individual strains of PrPSc , and the difference in PrP sequences between the donor and acceptor species [90, 91]. For instance, six positions have been identified in affecting prion transmission between mice and humans, i.e. residues 138, 143, 145, 155 and 166 [29]. Several kinetic models have been proposed to explain why spontaneous formation is an extremely rare event and how infection with PrPSc promotes the conversion of the cellular prion protein (Fig. 4). The template-assisted or heterodimer model [92] postulates that PrPSc is thermodynamically more stable than PrPC , but kinetically inaccessible. PrPC exists in equilibrium with a conformational intermediate PrP* which is able to interact with PrPSc . Binding of PrP* to PrPSc lowers the high activation energy barrier and allows the formation of a PrPSc heterodimer, but other

10 Prion Protein

Fig. 4 Kinetic models for the conformational conversion of PrPC to PrPSc . (A) The template-assisted or heterodimer model [92]. Here, the rate-limiting step during assembly is the conformational conversion between the partially folded PrP intermediate (PrP*) and a newly formed PrPSc . Binding of PrP* to PrPSc under the control of the putative protein X allows the formation

of a PrPSc heterodimer. (B) The polymerization nucleation model [71, 93]. Here the rate-limiting step is the formation of a stable PrPSc nucleus or seed. This is discerned by the observation of a lag phase. The exact size of the nucleus is unknown. PrPSc monomers are stabilized by joining the nucleus, as in seeded crystallization.

cellular factors such as molecular chaperones or prion-disease causing mutations might also contribute by reducing the energy barriers or increasing the stability of the partially folded intermediate species. In this model, the rate-limiting step is the conformational conversion between PrP* and PrPSc (Fig. 4A). The second kinetic model is nucleated polymerization [71, 93]. In this model, PrPC and PrPSc are in equilibrium in solution, but PrPSc is rare and unstable. The rate-limiting step is the formation of a stable PrPSc nucleus or seed. This is discerned by the observation of a lag phase in polymer growth. Once a seed is present, molecular association facilitates the conformational change of PrPC at a rapid rate (Fig. 4B). Within these models, two variants have also emerged recently. The β-nucleation model proposes that unfolding of helix H1 in PrPC , catalyzed by the PrPSc aggregate, is the key event in PrPSc propagation, and that intermolecular salt bridges between the helix H1 D and R residues of adjacent molecules are critical for the conversion [94]. This kinetic model, partially supported by MD simulations of wild-type and its D178N and E200K variants [95], was, however, not validated by cell-free conversion data [96]. The nucleated conformational conversion model, which is a dynamic version of nucleated polymerization, assumes that structurally fluid oligomeric complexes are crucial intermediates and rapid assembly occurs when these complexes conformationally convert upon association with nuclei [97]. At present, computer

4 Mechanisms of Prion Replication In Vivo 11

simulations have not been able to distinguish these kinetic models, independently of the protein models used, because the size of the oligomer is too small [98–102], the energy surface is biased towards the native state [103, 104] or the simulation conditions are not appropriate [105]. Regardless of the kinetic model, template-assisted or nucleation models, several important questions remain to be answered. (1) What is the role played by PrPC plasticity in the conformational change? Based on the experimental percentage of β-sheet within PrPSc [15], both the tail and the protein core undergo a structural change, but does one region initiate the conformational change? To address this issue, several MD simulations focused on the tail [84, 106–109]. In particular, Alonso et al. performed 10-ns MD simulations of Syrian hamster PrP109–219 at 300 K under neutral and low pH conditions [106]. Their results showed that acidic conditions favor the formation of a three-stranded antiparallel β-sheet spanning residues 109–131, whereas neutral conditions maintain the Nterminal region in a disordered state. Santini et al., by using MD simulations on various Syrian hamster PrP90–231 sequences, provided evidence that the ratelimiting factor for the formation of a three-stranded antiparallel β-sheet within the tail is thermodynamic rather than kinetic in character at neutral pH [84]. This βsheet within the tail, which is marginally populated in both PrP90–231 M129V and G131V mutants (30% of β-sheet versus 70% of random conformations) and not populated in wild-type PrP90–231 or its P102L variant, might be stabilized upon binding to the PrPSc template. In parallel, other studies focused on the possible role of the structured core. By using MD studies of PrP in solution [106, 110], molecular mechanics calculations of PrP fragments in monomeric, dimeric and tetrameric forms [94], and NMR structures of sheep PrP142–166 in solution [111], unfolding of helix H1 was suggested to provide the first internal driving force. However, other studies pointed to the high stability of helix H1. These include CD and NMR studies of human and murine prion peptides encompassing helix 1 and flanking sequences under various pH conditions [112–114], high-pressure NMR experiments on Syrian hamster PrP90–231 [74], MD simulations of PrP at low pH and high temperature conditions [115], and MD simulations of two PrP deletion variants at pH 7 and 300 K [86]. Alternatively, other theoretical investigations led to the proposal that helix H2 was the key region in the conformational change [52, 53, 116], because helix H2 is frustrated in the monomeric PrP structure [117, 118]. Frustration in secondary structure elements is defined as the incompatibility between the predicted (e.g. by using neural networks) secondary structure and the experimentally determined structure. Or they concluded using biased MD simulations that there are two populated unfolding routes, the helices H2 and H3 being disordered first with the S1H1S2 motif formed and vice versa [119]. Clearly, this diversity of unfolding and folding routes for PrP is not surprising, but it remains to simulate the dynamics of PrPC in interaction with a PrPSc model to reflect experimental reality. (2) What is the exact role of the β-rich oligomers formed by non reduced recombinant PrP proteins in the conversion? Recent studies have shown that these oligomers, sensitive to digestion by proteinase K, are apparently not on the pathway to amyloid fibril formation [69, 70]. However, it remains possible that these β-rich oligomers may offer a pool of pre-aggregated material for further structural reorganization

12 Prion Protein

through alternative pathways. Furthermore, these oligomers have been observed in other neurodegenerative diseases and there is accumulating evidence that both soluble A β oligomers (early aggregates) and fibrils are toxic in cell cultures [120]. (3) What is the minimal size of the infectious PrPSc particle? Ionizing irradiation experiments have suggested that the minimally infectious PrPSc is a dimer [121], and protease-resistant PrP dimers were observed in hamster brain [122]. On the other hand, recent studies based on size-exclusion chromatography, electrospray mass spectrometry and dynamic light scattering showed that the β-rich oligomers consist of octamers [60, 69, 123], whereas electron crystallography experiments have deduced an hexamer from in vitro infectious Syrian and mouse PrP two-dimensional protofibril crystals [51]. Taken together, these data suggest that infectivity may be associated with several sizes of aggregates under specific external conditions such as pH, temperature, concentration. (4) Finally, what is the role played by copper in the interconversion process and is our inability to generate infectious PrPSc in test tubes due to the absence of cellular factors? There is strong genetic evidence supporting for the importance of the copperbinding domain, since modifications in the number of octa-repeats cause familial prion diseases [124]. The exact role of copper in the interconversion process is far from being elucidated. It is currently believed that the binding of copper ions induces a conformational transition that presumably modulates PrP aggregation [46], thereby enhancing its infectivity [125]. Prior studies on the transmission of human prions to transgenic mice have suggested the role of a cofactor, protein X, in the formation of PrPSc . Protein X appears to bind to the side-chains of residues that form a discontinuous epitope: some amino acids are in the loop composed of residues 165–171 and at the end of helix H2 (Q168 and Q172), whereas others are on the surface of helix H3 (T215 and Q219) [126]. It is intriguing that the protein X epitope coincide with the binding surfaces between PrPSc and PrPC and the disordered regions within the NMR PrPC structures of human, mouse and Syrian hamster. However, all physical attempts to identify the protein X have failed thus far. Recent studies have also indicated that PrPC can associate with various molecular complexes [127]. For instance, Edenhofer et al. found interactions between the molecular chaperone Hsp60 and recombinant glutathione S-transferase-fusion PrPC peptides [128]. In addition, in vitro conversion experiments using a variant of the protein-misfolding cyclic amplification method [129] have shown that specific RNA molecules stimulate PrPSc formation [130]. This raises the possibility that RNA molecules are cellular cofactors and are involved in generating strain diversity. Clearly, advances in mass spectrometry should help clarify the nature of the cellular factors. 5 Conclusions

Since 1997 when Stanley Prusiner was awarded the Nobel Prize for Medicine, important steps have been made in understanding prion replication at the structural

References

level, but many questions remain to be answered. Thus far, many molecules have been identified to be effective for inhibiting prion replication in cell culture essays. For instance, Soto et al. partly reversed in vitro PrPSc to PrPC using β-sheet breaker peptide spanning PrP115–122 [131]. Caughey et al. inhibited the conversion reaction in vitro using diferoylmethane [132]. However, all these inhibitors failed on sick animals. The design of efficient inhibitors blocking prion propagation in mammals requires, at least, a combined effort in three directions: 1. PrP functional role Several studies have pointed to the possible role of PrP in copper transport or metabolism [125], signal transduction [133] and apoptosis [134], to name a few, but the physiological function of PrP remains unknown. Yet, PrP is highly conserved across species, and is expressed in most adult tissues and at high levels in the central nervous system. Thus, PrP is likely to be very useful. 2. Cellular factors Advances in experimental techniques are needed to identify the cellular factors because no purified recombinant PrP has yet been successfully converted in vitro to infectious PrPSc . 3. Structural characterization of the soluble PrP oligomers and insoluble PrPSc fibrils A detailed characterization of the soluble oligomeric intermediates is very difficult because the intermediates are typically short lived and are present in a wide range of conformations and degrees of aggregation; however, these atomic models would greatly facilitate the design of new drugs by computers.

Such a combined effort should not only be useful in prion science but also in Alzheimer’s or Huntington’s diseases because soluble oligomers apparently share a unique common structural feature [135], and, eventually, in understanding the generic rules between amino acid sequences, folded and misfolded structures for all protein coding genes. References 1 SIPE, J. D. Amyloidosis. Annu Rev Biochem 1992, 61, 947–975. 2 KELLY, J. F. The alternative conformations of amyloidogenic proteins and their multistep assembly pathways. Curr Opin Struct Biol 1998, 8, 101–106. 3 DOBSON, C. M. Protein-misfolding diseases: getting out of shape. Nature 2002, 418, 729–730. 4 DIEDRICH, J. F., P. E. BENDHEIM, Y. S. KIM, R. I. CARP and A. T. HAASE. Scrapie-associated prion protein accumulates in astrocytes during scrapie infection. Natl Acad Sci USA 1991, 88, 375–379.

5 JEFFREY, M., J. R. SCOTT, A. WILLIAMS and H. FRASER. Ultrastructural features of spongiform encephalopathy transmitted to mice from three species of biovidae. Acta Neuropathol 1992, 84, 559–569. 6 BENDHEIM, P. E., R. A. BARRY, S. J. DEARMOND, D. P. STITES and S. B. PRUSINER. Antibodies to a scrapie prion protein. Nature 1984, 310, 418–421. 7 ALPER, T., W. A. CRAMP, D. A. HAIG and M. C. CLARKE. Does the agent of scrapie replicate without nucleic acid? Nature 1967, 214, 764–766. 8 GRIFFITH, J. S. Self-replication and scrapie. Nature 1967, 215, 1043–1044.

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1

Protein Aggregates in Neurodegenerative Disorders Giorgio Giaccone

Istituto Nazionale Neurologico Carlo Besta, Milano, Italy

Mario Salmona Istituto di Ricerche Farmacologiche Mario Negri, Milano, Italy

Fabrizio Tagliavini Istituto Nazionale Neurologico Carlo Besta, Milano, Italy

Gianluigi Forloni Istituto di Ricerche Farmacologiche Mario Negri, Milano, Italy


1 Introduction

The presence of protein aggregates has been proposed as a unifying feature of neurodegenerative disorders. The aggregates may consist of fibrils with a β-pleatedsheet conformation, termed amyloid, or be made up of misfolded proteins without the staining and ultrastructural properties of amyloid. This abnormal material can accumulate at the intra- or extracellular levels, but is invariably associated with neuronal degeneration. In some instances, new categories of disorders could be identified on the basis of the misfolded protein. The formation of inclusions may represent an early event or the end stage of a molecular cascade of several steps and the pathogenic role of the aggregates might be variable in different diseases. For several neurodegenerative disorders, genetic variants assist in explaining the pathogenesis of the more common sporadic forms, and in developing mouse and other models. Our understanding of the pathways involved in protein aggregation and in the molecular mechanisms of cellular toxicity is growing rapidly [1, 2]. The presence of extracellular amyloid deposits and of intracellular neurofibrillary tangles is the main neuropathological feature of Alzheimer’s disease. The β amyloid (Aβ) protein, a peptide of 40–42 amino acids, is the major component Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Protein Aggregates in Neurodegenerative Disorders

of amyloid, while neurofibrillary tangles are made up of hyperphosphorylated tau protein. Based on the presence of abnormal hyperphosphorylation and aggregation of tau without other specific neuropathological abnormalities, a heterogeneous group of neurodegenerative disorders clinically characterized by dementia and/or motor syndromes is now unified under the term of “tauopathies”. The group includes Pick’s disease, progressive supranuclear palsy (PSP) and corticobasal degeneration (CBD). The discovery that mutations of protein tau are associated with familial forms of frontotemporal dementia and parkinsonism (FTDP-17) provides strong evidence that the derangement of tau metabolism contributes to neurodegeneration. The accumulation of a misfolded form of the prion protein characterizes prion diseases and plays an essential role also in their transmissibility. Molecular genetic investigations allowed the identification of α-synuclein aggregates as the main component of Lewy bodies, the intracellular inclusions that characterize the degenerating cells in Parkinson’s disease. α-Synuclein is also found in the intraneural inclusions of other neurodegenerative disorders, such as dementia with Lewy bodies (DLB) and multiple system atrophy, which are all now commonly described as “α-synucleinopathies”. The pathogenetic role of protein aggregates found in other groups of neurodegenerative diseases is less defined. Ubiquitinated cytoplasmatic inclusions have been detected in amyotrophic lateral sclerosis, and distinguished morphologically in skein inclusions, hyaline Lewy body-like inclusions and Bunina bodies. The composition of these aggregates is varied: in the familial forms associated with copper–zinc superoxide dismutase (SOD1) mutations, the mutated form of the enzyme was consistently found in the aggregates. Recent studies have proposed a direct involvement of SOD1 misfolding and aggregation in the pathogenesis of this disease. According to this hypothesis, Stathopulos et al. [3] have shown that SOD1 mutants undergo conformational changes that facilitate their polymerization. However, in terms of temporal development of the neurodegenerative process investigated in animal models, the appearance of intracellular inclusions is not an early event. Furthermore, the heterogeneous composition of intracellular inclusions in sporadic cases did not support a seminal role of protein aggregation in the pathogenesis of amyotrophic lateral sclerosis [4–6]. The presence of a trinucleotide (CAG) repeat expansion is characteristic of many neurological disorders including Huntington’s disease, spinobulbar muscular atrophy, dentatorubro-pallidoluysian atrophy and different forms of spinocerebellar ataxia. The expanded polyglutamine [poly(Q)] tract lies in functionally unrelated genes and the accumulation of poly(Q) sequences is a common pathological hallmark of these disorders. In most cases, the accumulation occurs at the level of neuronal nuclei (intranuclear inclusions). The molecular basis of the toxicity of poly(Q) has been explained by two alternative concepts. One implies that the expansion leads to conformational changes creating a misfolded structure with cytotoxic properties [7]; the other that the long poly(Q) sequence increases the capacity of the protein to form amyloid aggregates, according to the hypothesis of nucleation [8]. However, the central role of protein aggregates in poly(Q) disorders has been

2 Neuropathology 3

questioned by researchers who consider them to be a neuronal byproduct that, in some cases, may have protective functions [9, 10].

2 Neuropathology 2.1 Alzheimer’s Disease

The neuropathologic hallmark of Alzheimer’s disease is the co-occurrence of extracellular deposits of amyloid made up of straight fibrils of around 10 nm whose main component is the Aβ polypeptide and the intracellular build-up of abnormal twisted filaments [paired helical filaments (PHFs)], mainly composed of hyperphosphorylated tau protein. Aβ deposition takes place in the neuropil as well as in leptomeningeal and parenchymal vessel walls, whereas PHF formation occurs within perykaria and neurites of selected neuronal populations. Therefore, Alzheimer’s disease is a unique protein misfolding disease, since it is characterized by misfolding of two unrelated proteins, Aβ and tau, that cause two distinct histopathologic changes. This peculiarity, together with the fact that Aβ deposition and neurofibrillary changes can occur independently of each other, is the basis of several questions regarding the pathogenesis, and, in particular, which one of the two lesions is more strictly associated with the development of diffuse neuronal loss and dysfunction, synaptic rarefaction, and the appearance of the clinical signs of dementia. The issue is further complicated by the fact that Aβ deposition and tau pathology, although with differences in severity and topographic distribution, may also be present in non-demented elderly subjects [11]. A main distinction is that, in non-demented subjects, neurofibrillary changes, when present, are less severe and confined to restricted brain areas (the mesial temporal structures), while Aβ deposits may be widespread in the neocortex, in some instances in an amount similar to Alzheimer’s disease. This is in keeping with the concept that neurofibrillary changes evolve in an anatomical stereotypical hierarchical fashion, with the entorhinal cortex being the earliest area affected, followed by the hippocampal formation. With increasing disease progression, neurofibrillary changes occur in the association cortex, while the primary sensory and motor cortices are spared until the very advanced stages of the disease [12, 13] (Fig. 1A and 1B). Aβ is a 40- to 42-residue peptide derived by proteolytic cleavage of a 695- to 770amino-acid Aβ protein precursor (AβPP) [14–16], a transmembrane glycoprotein encoded on chromosome 21, in which Aβ is positioned partly in the extracellular domain and partly in the transmembrane domain. The Aβ deposits in the vessel wall take the form of congophilic (amyloid) angiopathy, while in the neuropil, the deposits determine the formation of the senile plaques (Fig. 2). These are complex lesions whose morphogenesis is influenced by the participation of cellular elements, either neuronal (neurites and synaptic terminals) or glial (reactive astrocytes and activated microglia), as well as by the association of Aβ amyloid with other


Fig. 1 Alzheimer’s disease. Immunohistochemistry with monoclonal antibody AT8 to phosphorylated tau (immunoreactivity corresponds to the brown reaction product) revealed severe involvement of the cerebral cortex by neurofibrillary changes that can spare the primary motor and sensory areas (A) or involve all cortical fields (B). AT8 labeled cell bodies and a network of neu-

ronal processes dispersed in the neuropil of the cerebral cortex (neuropil threads) (C). Clustering of AT8-immunoreactive profiles occurred in direct correspondence to senile plaques (D). Labeled neurons often showed pyramidal morphology, with AT8 immunostaining extending to the apical dendrite, reproducing the picture of flame-shaped NFTs (E).

“chaperone” molecules. These may be associated to all forms of amyloid, such as P component, complement factors, apolipoprotein (Apo) E and J, or more specific for Aβ deposits as α 1 -antichymotrypsin [17]. The introduction of immunohistochemical techniques using antibodies against Aβ revealed a far higher density and wider distribution of Aβ deposits in Alzheimer’s disease than had been appreciated by the use of classic silver impregnation methods or by applying specific staining for amyloid, such as Congo red or Thioflavin S. This was because immunohistochemistry could also recognize Aβ deposits that lacked the staining and ultrastructural properties of amyloid. These amorphous appearing, non-fibrillar plaques were referred to as “diffuse plaques” or “pre-amyloid deposits”, since they might represent an early stage in the morphogenesis of senile plaques, also considering that they are not associated with neuritic or glial changes [18–21] (Fig. 3). This hypothesis is supported by studies on patients with Down’s syndrome (trisomy 21) who invariably develop Alzheimer’s neuropathological changes, starting with pre-amyloid deposits in their teenage years and displaying fully-developed senile plaques with neuritic/glial changes and neurofibrillary tangles (NFTs) about two decades later [22, 23]. Moreover, by immunohistochemistry, it has been demonstrated that, in

2 Neuropathology 5

Fig. 2 Alzheimer’s disease. Double immunohistochemical detection of phosphorylated tau (blue) and A$ protein (brown). Note the intermingling of A$ deposition with tau-immunoreactive neuronal profiles, configuring the senile plaque.

Alzheimer’s disease, Aβ deposition also affects brain regions previously considered to be spared by the pathologic process, such as the cerebellum, striatum and brainstem [24, 25]. Interestingly, non-fibrillar pre-amyloid deposits are the predominant form of Aβ deposition in these regions. The concept of the C-terminal heterogeneity of the Aβ peptides deposited in Alzheimer’s disease brains provided a further distinguishing element between pre-amyloid deposits and senile plaques, since the former are made up almost exclusively by Aβ1–42, while in the latter contain a mixture of Aβ1–40 and Aβ1–42. Aβ1–40 is the main constituent of amyloid in the vessel walls [26, 27]. NFTs are caused by the intracellular accumulation of PHFs, abnormal twisted filaments 80–200 nm thick, whose main component is an abnormally phosphorylated form of tau protein [28–30]. Tau is a microtubule-associated protein, encoded by a gene on chromosome 17 [31, 32], highly expressed in the central nervous system (CNS). In normal conditions, tau is present exclusively in axons, while in Alzheimer’s disease, it accumulates in an insoluble, hyperphosphorylated form ectopically in the perikaria and dendrites. The main biologic function of tau is to bind


Fig. 3 Alzheimer’s disease. Immunohistochemistry with antibodies to A$ (immunoreactivity corresponds to the brown reaction product) revealed the high density of A$ deposits in the cerebral cortex (low magnification, A) and the different morphologies

of A$ deposition: classic senile plaques with a dense core and a more dispersed peripheral halo (B), amorphous amyloid deposit (C), pre-amyloid deposits (D) and amyloid angiopathy close to a senile plaque (E).

microtubules via the C-terminal binding domains, promoting their polymerization and stabilization. Tau therefore plays an important role in maintaining neuronal integrity, axonal transport and polarity [33, 34]. Highly phosphorylated tau has fewer tendencies to bind microtubules [35–38]. NFTs and degenerating neurites of plaques are consistently immunoreactive with antiphosphorylated tau antibodies (Fig. 1D). Moreover, immunohistochemistry with these antibodies is much more sensitive than classic silver impregnation methods in revealing neurofibrillary

2 Neuropathology 7

changes and enabled recognition of an additional form of hyperphosphorylated tau (Fig. 1C–E). These are the “neuropil threads”, a meshwork of randomly oriented, tau-immunoreactive neurites, dispersed in the neuropil, that represent a consistent part of the total burden of neurofibrillary pathology in Alzheimer’s disease [39]. NFT and degenerating neurites of plaques are also consistently immunoreactive for ubiquitin. The molecular mechanism of PHF formation has not been clarified: several kinases and phosphatases have been implicated [40, 41], but it remains to be determined whether hyperphosphorylation is the primary event that makes tau insoluble and resistant to degradation or is a consequence of the aggregation process. In summary, several lines of evidence indicate the cerebral deposition of Aβ amyloid is the seminal event that initiates a cascade of biochemical and cellular changes leading to tau-related alteration of the neuronal cytoskeleton and to neurodegeneration. This scenario – known as the “amyloid cascade hypothesis” [42, 43] – is supported by the observation that quantitative modification of the total levels of Aβ or of the ratio between long forms and short forms of this peptide (i.e. Aβ1–42 versus Aβ1–40) are sufficient to induce the whole neuropathologic phenotype of the disease. The first situation takes place in Down’s syndrome, in which one extra copy of the gene coding for the AβPP is present as a consequence of the chromosome 21 trisomy, while the second corresponds to the familiar cases of Alzheimer’s disease associated with mutations of the genes for AβPP, presenilin1 or presenilin2 (see below) [43]. NFT composed of tau aggregates that are biochemically similar to those of Alzheimer’s disease have been described in many neurodegenerative diseases in which Aβ is absent. These are referred as tauopathies, and include Picks disease, CBD and PSP. A subset of tauopathies is familial and mutations of the tau gene have been demonstrated in a percentage of them [44–47]. 2.2 Tauopathies

Starting from 1994, an autosomal-dominantly inherited form of frontotemporal dementia with parkinsonism was described in several families and linked to the region of chromosome 17 that contains the tau gene, resulting in the denomination “frontotemporal dementia and parkinsonism linked to chromosome 17” (FTDP-17) [44, 48]. The neuropathologic picture of FTDP-17 is characterized by tau-positive inclusions that occur in neurons, but may also be numerous in glial cells. The now widely used term “tauopathy” was introduced to highlight the abundance of tau accumulation, in the absence of Aβ aggregates [45]. In 1998, the first mutations in the tau gene (MAPT) were reported in FTDP-17 families and now the total of the known mutations is more than 30 [46, 47, 49]. Tau protein is encoded by a single gene of 16 exons from which different forms are produced via alternative mRNA splicing. In adult human brain, six tau forms are generated ranging from 352 to 441 amino acids, differing with respect to the presence of three (3R-tau) or four (4R-tau) C-terminal repeat sequences of


31–32 residues. Three different forms of each (3R-tau and 4R-tau) exist, as a result of the presence of none, one or two N-terminal inserts of unknown function [41]. The complex structure of the tau protein is the basis of the variability in the chemicophysical characteristics of the insoluble tau filaments in different tauopathies. The partial elucidation of the relationship between the site of the mutation on the MAPT gene, the tau isoforms engaged in the misfolding process, and the biochemical and structural characteristics of the tau aggregates provided very important insights into the role of tau abnormalities in neurodegenerative diseases, and more generally into the pathogenesis of protein misfolding diseases. The most important implication derived from studies of patients with FDTP-17 is that the mutation of the tau gene is sufficient to cause intracellular tau deposition and to induce neurodegeneration. Moreover, the studies of these rare inherited forms of tauopathies have shed light on the pathogenesis of other more common, sporadic neurodegenerative diseases characterized by tau inclusions (Pick’s disease, CBD and PSP), that can be considered as primary tauopathies, due to the absence of other disease-specific neuropathological abnormalities. In contrast, other brain diseases are known in which tau inclusions are likely secondary to metabolic (Niemann–Pick type C), infective (subacute sclerosing panencephalitis) or traumatic (dementia pugilistica) brain lesions. PSP is characterized clinically by supranuclear gaze palsy and by postural instability [50]. Neuropathologically, the main features are atrophy of the basal ganglia and brainstem, with neuronal loss and gliosis. In these brain areas, there is a high density of tau pathology, including NFTs in neurons and consistent tau aggregates in glial cells, both astrocytes (“tufted astrocytes”) and oligodendrocytes (“coiled bodies”) [51] (Fig. 4C and 4D). In contrast to Alzheimer’s disease, ultrastructurally these aggregates are made up straight fibrils of 15–18 nm [52] and biochemically are mainly composed of 4R-tau isoforms [41]. CBD is a progressive neurodegenerative disease involving the cerebral cortex and deep grey and white matter structures such as the striatum, thalamus, substantia nigra, capsula interna, subcortical frontal white matter and brainstem [53]. The neuropathological features are depigmentation of substantia nigra and asymmetrical frontoparietal atrophy. In affected regions, neuronal loss, gliosis and tau-immunoreactive glial and neuronal inclusions are prominent, and surviving nerve cells are often swollen (“achromatic” or “ballooned” neurons). The neuronal tau inclusions are morphologically pleomorphic. In some neurons they appear as dispersed globose NFTs; in others as small compact inclusion bodies and occasionally tau-immunoreactivity diffusely fills the cytoplasm. A typical finding of CBD is the prominent deposition of abnormal tau in neuronal processes (“threadlike neuritic profiles”) and in glial cells, with consistent involvement of the white matter. In the cerebral cortex, peculiar lesions consisting of annular clusters of tau-immunoreactive processes (“astrocytic plaques”) are considered typical of CBD [54] (Fig. 4E and 4F). The abnormal filaments include both PHF-like and straight filaments [55]. The biochemical profile of insoluble tau is similar to that of PSP, being mainly composed of 4R-tau isoforms [41].

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Fig. 4 Tauopathies. In Pick’s disease the characteristic neuronal, round, welldemarcated inclusions (Pick bodies) are revealed by Bodian silver impregnation (A) and by immunohistochemistry with monoclonal antibody to phosphorylated tau (AT8, immunoreactivity corresponds to the brown reaction product) (B). In PSP the tau pathology is stained by monoclonal antibody AT8 to phosphorylated tau, and includes NFTs in neurons (C) and tau deposi-

tion in glial cells, as the “tufted astrocytes” (D). In CBD the deposition of phosphorylated tau disclosed by AT8 immunohistochemistry is prominent in neuronal perikaria (E), but also in neuritic processes (E). The “astrocytic plaque” (F) is considered typical of this disease. FTDP-17 is characterized by the presence of phosphorylated tau deposition revealed by AT8 immunohistochemistry in neurons (often diffuse in the cytoplasm, G) and in glial cells (H).

Pick’s disease is now a term that should be restricted to forms of frontotemporal degeneration with tau-immunoreactive lesions and, in particular, with the characteristic neuronal, round, well-demarcated argyrophilic inclusions (Pick bodies) [56] (Fig. 4A and 4B). The neuropathologic picture is that of severe frontotemporal


atrophy with marked neuronal loss, gliosis and neuropil vacuolization. Pick bodies are abundant in the dentate gyrus of the hippocampus and focally in layers II, III and VI of the cerebral cortex. They are detected by silver impregnation and by immunohistochemistry with antibodies to hyperphosphorylated tau that reveal additional cytoskeletal changes, including neuritic profiles and astrocytic and oligodendroglial inclusions [57]. Biochemically, the tau aggregates are predominantly made up by the 3R-tau isoforms [41]. It should be recalled that the majority of patients with the clinical features of FTD do not show the neuropathology of Pick’s disease, but a more unspecific picture of atrophy of the frontal and temporal lobes with neuronal loss, gliosis and microvacuolar neuropil degeneration, in the absence of tau-immunoreactive changes. The neuropathology of FTDP-17 is heterogeneous with respect to the morphologic characteristic of the lesions as well as their severity and topographic distribution. It can overlap with the scenario of Pick’s disease, PSP and CBD – the invariable change being the presence of tau deposits in neurons and glial cells in several grey and white matter structures of the CNS (Fig. 4G and 4H). The heterogeneity of the clinical/pathologic phenotypes partially reflects the position of the mutations in the tau gene. They can be separated in two broad categories: (1) coding region missense mutations, and (2) intronic mutations and silent mutations of the coding region. The first group of tau mutations is the ordinary one, in which a nucleotide substitution in the coding region of the MAPTgene changes the corresponding amino acidic residue. Most of the reported missense mutations occur in the highly conserved region within or near the microtubule-binding domains. The effects of these mutations seem to be 2-fold: (1) they reduce the ability of tau protein to bind to microtubules and to promote microtubule assembly in vitro [58, 59], and (2) they increase the tendency of tau to aggregate in insoluble filaments [60, 61]. The structure and biochemical characteristics of the misfolded tau aggregates are dependent on the exon of the mutation. While mutations located outside exon 10 modify all six tau isoforms, those within exon 10 affect only the three tau isoforms bearing four microtubule-binding domains (4R-tau), since the supplementary repeat of this portion of the molecule is coded by exon 10. The second class of mutations is more puzzling. These genetic defects are single base pair substitutions occurring within intron 10 or silent nucleotide changes in the adjacent exon 10. At first it was difficult to explain how mutations that do not affect the sequence of the tau protein could provoke its misfolding. The explanation came from biochemical studies demonstrating that these mutations increase the levels of 4R forms of tau, likely due to a higher proportion of mRNAs in which exon 10 is retained [62, 63], as a consequence of the disruption of a regulatory stem-loop, “splicing enhancer” structure [64]. The mechanisms by which changes in the ratio of 3R-tau to 4R-tau lead to neuronal dysfunction is unclear, but it seems established that a ratio of 3R-tau forms to 4R-tau forms of about unity is necessary for the normal structure and function of microtubules. It has been suggested that some missense mutations in exon 10 close to the downstream intron may also act with this mechanism.

2 Neuropathology 11

2.3 Prion Diseases

Prion diseases are fatal, rapidly progressive, neurodegenerative disorders of humans and animals. They are a unique category of diseases, since may be sporadic or inherited in origin and can be transmitted. They are therefore also called “transmissible spongiform encephalopathies”, a term that underlines their infectious character and the classic main neuropathological hallmark of neuropil vacuolation. The transmissible agent, the prion, is devoid of informational nucleic acid and consists only of protein [65] (see below). Several lines of evidence indicate that prions are composed of an abnormal, pathogenic isoform of the prion protein (PrP). The normal form of PrP (PrPC ) is widely expressed in neurons and glia in the CNS, and little is known about its function(s). The human cellular gene which encodes PrPC has been called PRNP, whose open reading frame is in a single exon. The pathogenic isoform (PrPSc ) results when the normal form undergoes a conformational change, converting α-helical regions to β-sheet motifs, and possesses abnormal physicochemical properties such as detergent insolubility and protease resistance [66]. Therefore, prion diseases are “transmissible protein misfolding diseases”, in which the abnormally folded protein may induce the conversion of normal molecules into the misfolded status. Animal prion diseases include scrapie in sheep and goats and bovine spongiform encephalopathy (BSE) in cattle. In humans, the most common prion disease is Creutzfeldt-Jakob disease (CJD), which can be inherited as autosomal dominant disease associated with mutations of the PRNP gene, acquired iatrogenically through exposure to material contaminated with prions or arise sporadically for no obvious reason. Possible causes of sporadic CJD include spontaneous formation of PrPSc as a rare stochastic event, somatic mutations of PRNP or unrecognized prion exposure. A variant CJD (vCJD) form was reported in 1996 [67], characterized by clustering in the UK and peculiar neuropathological and biochemical features. Geographic and temporal association, as well as transmission studies revealing strong analogies between the vCJD and BSE agents, lead to the conclusion that vCJD is acquired by humans by exposure to BSE contaminated material. The detection of the deposition of PrPSc is now possible not only biochemically by immunoblot analysis, but also in histological specimens following the introduction of effective pre-treatments that make immunohistochemistry with anti-PrP antibodies sensitive and specific [68–70]. These techniques revealed how PrPSc accumulates in the CNS of CJD patients with variable topographic distribution and different patterns of immunostaining – synaptic, perivacuolar, perineuronal and plaque-like (Fig. 5A–D). The accumulation of PrPSc in the CNS is accompanied by neuronal and glial changes. The triad of spongiform changes, neuronal loss and gliosis, in the absence of an inflammatory response, is the classic neuropathological hallmark of CJD. Spongiform change is relatively specific to CJD, but may differ in severity in different patients and from regions to regions of the CNS. It is characterized by small, round or oval vacuoles in the neuropil of the cerebral cortex and other grey structures (Fig. 5E). Glial changes consist of reactive


Fig. 5 Prion diseases. In CJD, most grey structures exhibit PrPSc immunoreactivity (corresponding to the brown reaction product) (A) that may assume different patterns: diffuse synaptic type (B), perineuronal (C) and perivacuolar (D), and is associated with

spongiform changes (hematoxylin & eosin, E) and astrogliosis (anti-glial fibrillary acid immunohistochemistry, F). PrP amyloid deposits are consistent in GSS (Thioflavin S, G) and vCJD (PrP immunohistochemistry, H).

astrogliosis and microglial activation (Fig. 5F). Neuronal loss in the affected cortical and subcortical regions is often severe, sometimes with complete depletion of some neuronal populations [71]. While in classic CJD cases the formation of true amyloid deposits is a rare event and is restricted to specific brain regions, in Gerstmann–Sträussler–Scheinker (GSS) disease this phenomenon is relevant, and PrP amyloid deposits are abundant and widespread in the CNS [72] (Fig. 5G). GSS is determined by specific mutations in the PRNP gene and has a longer disease duration than CJD. Amyloid PrP deposition is also abundant in vCJD, in which the typical lesion is the florid plaques – an amyloid core surrounded by vacuoles of spongiosis (Fig. 5H).

2 Neuropathology 13

The neuropathology of CJD is heterogeneous and it has been shown that this may be related to variations in the tertiary structure of PrPSc , resulting in different conformers of the abnormal protein having distinct physicochemical and pathogenic properties [73]. This is confirmed by the observation of molecular size differences in the protease-resistant core of PrPSc that are due to different sites of proteolytic cleavage reflecting different tertiary structures. In particular, the clinicopathological heterogeneity of sporadic CJD has been linked to two types of PrPSc , termed type 1 and type 2, having a molecular weight of 21 and 19 kDa. Evidence suggests that the PrPSc type in combination with the genotype at codon 129 of the PRNP gene – a common polymorphic site encoding methionine or valine – is a major determinant of deposition pattern (i.e. diffuse or focal) and brain regional distribution of PrPSc . The view that different conformers of PrPSc may possess distinct pathogenic properties is supported by a study on CJD cases with both type 1 and type 2 PrPSc in the same brain. This event is relatively common, involving about 25% of the sporadic patients with a close relationship between PrPSc type, pattern of PrP immunoreactivity and severity of spongiform changes [74]. Further support for the prion hypothesis has been provided by transgenic animal studies; in particular, the finding that mice in which the PrP gene has been ablated (PrP knock-out mice) neither develop neuropathological features of prion diseases nor accumulate PrPSc or propagate the disease after inoculation of prions, demonstrating that the absence of PrPC expression prevents prion spread and neuronal dysfunction [75]. 2.4 Synucleinopathies

Synucleinopathies represent a heterogeneous group of neurodegenerative disorders characterized by the presence of cytoplasmic neuronal and glial inclusions. The synucleinopathies can be divided into Lewy body disorders (Parkinson’s disease and DLB) and disorders with intracytoplasmic inclusions in glial cells (multiple system atrophy). Parkinson’s disease is the second most common neurodegenerative disease after Alzheimer’s disease. Parkinson’s disease is characterized clinically by resting tremor, rigidity and bradykinesia, resulting from the progressive and selective loss of dopaminergic neurons in the pars compacta of the substantia nigra. Histopathologically, it is characterized by the degeneration of specific nerve cell populations that develop filamentous inclusions in the form of Lewy bodies and dilated neurites. The onset of clinical symptoms occurs when the loss of dopaminergic neurons is over 50%. Lewy bodies are found not only in the substantia nigra, but also in other brain structures [76]. The classical Lewy body is a spherical, cytoplasmatic inclusion with a diameter of about 10–30 µm. After hematoxylin & eosin staining, the Lewy body assumes the classical “target shape” with a central core more intensely stained and a more faded external portion (Fig. 6A). At the ultrastructural level, the central portion is formed by tightly packed filamentous and granular material, while in the external part filaments of 7–20 nm are associated with electron-dense material.


Fig. 6 Synucleinopathies. In Parkinson’s disease, Lewy bodies appear as target-like, spherical, cytoplasmic inclusions (hematoxylin & eosin, A) intensely immunoreactive with anti-SYN antibodies (immunore-

activity corresponds to the brown reaction product, B). In MSA, GCIs appear black in sections stained with the Gallyas silver impregnation (C) and are immunolabeled by anti-SYN antibodies (D).

Other round-shaped inclusions (“pale” bodies) are also identified in Parkinson’s disease, and appear as granular and eosinophilic material displacing neuromelanin of pigmented neurons, without the classical “target shape” of Lewy bodies. Pale bodies are considered precursors of Lewy bodies. The identification of familial forms of Parkinson’s disease caused by missense mutations in the α-synuclein gene [77] prompted immunohistochemical studies showing that Lewy bodies are consistently and intensely immunoreactive with anti-SYN antibodies [78] (Fig. 6B). Previously, Lewy bodies were recognized by means of anti-ubiquitin antibodies [79]. Furthermore, parkin, another protein genetically associated to familial Parkinson’s disease, was invariably found in Lewy bodies [80]. SYN immunoreactivity was found not only in perykarial inclusions but also in neurites (Lewy neurites). DLB defines a specific clinical syndrome characterized by progressive dementia with fluctuation of cognitive deficits and hallucinations, mainly visual, associated with the neuropathological finding of Lewy bodies. However, the threshold of Lewy bodies for a DLB diagnosis has not been defined yet. This could lead to the undetermined condition of subjects with a clinical diagnosis of dementia where the typical Alzheimer’s disease pathology (Aβ deposits and NFTs) co-exists with the presence of Lewy bodies. The problem is that the relative contribution to the clinical symptoms of the individual pathological components is difficult to define. Other DLB cases are “pure”, in which Alzheimer-type pathology is absent or minimal. Based on the localization and density of Lewy bodies, three categories have been identified: Lewy bodies at predominance in the brainstem, in the limbic cortex and in the neocortex. Unexpectedly, DLB is not necessary associated with the third category (Lewy bodies at predominance in neocortex). Cortical Lewy bodies after

3 The Neurotoxic Proteins

hematoxylin & eosin show different shapes: round, oval and bean-like, lacking the target shape typical of nigral Lewy bodies. On the other hand, the antigenic characteristics of cortical Lewy bodies are essentially the same of the classical Lewy bodies, as they are immunopositive for SYN, ubiquitin and parkin. Multiple system atrophy (MSA) is clinically associated to disautonomic disturbances, parkinsonism and cerebellar signs. Neuropathologically, it is characterized by intracellular glial inclusions originally recognized by the silver impregnation method of Gallyas [81]. They appear as oval, or triangle-, flame- or sickle-shaped glial cytoplasmic inclusions (GCIs or Papp-Lantos inclusions) localized in oligodendrocytes (Fig. 6C). The size is variable, but the inclusions often completely fill the cytoplasm, confining the nucleus at the periphery. GCIs are immunoreactive with antibodies against SYN [82] (Fig. 6D). At the ultrastructural level, GCIs are composed of tubules or filaments having a diameter of 20–40 nm in association with granular material. In MSA subjects, SYN-immunoreactive inclusions can be found in glial and neuronal cells nuclei and in neurites. GCIs are prominent in the motor and sensory primary cortices, in the putamen, globus pallidus, internal and external capsula, corpus callosum, and several other structures of the brainstem and spinal cord.

3 The Neurotoxic Proteins 3.1 Alzheimer’s Disease

The pathogenic role of Aβ in Alzheimer’s disease has been put forward on the basis of genetic, experimental and biochemical evidence, and an essential contribution to this hypothesis derived from the demonstration that Aβ peptides can exert neurotoxic effects in vitro [42]. The toxicity of Aβ peptides is mediated by apoptosis [83, 84] and was initially associated with the polymerization into fibrils of the peptide [85]. The biological active fragment was found to be the Aβ 25–35 sequence. The fibrillogenic capacity, tested in vitro, increases progressively from peptide Aβ1–40 to Aβ1–42 and Aβ25–35. Aβ peptides containing 40 or 42 residues are those found in the brains of Alzheimer’s disease patients in senile plaques and originate from the precursor of Aβ, AβPP. In the amyloidogenic pathway of catabolism, AβPP is cleaved sequentially by β- and y-secretases at the N- and C-terminus of the Aβ region. β-Secretase cleaves between residues 671 (Met) and 672 (Asp), while the cleavage at the C-terminal by γ -secretases can occur either at residue 711 (Val) or 713 (Iso) to generate Aβ1–40 or Aβ1–42 [86–88]. The amyloid-like fibrils spontaneously formed in vitro by these peptides have morphological, staining and ultrastructural features similar to the amyloid deposits isolated from Alzheimer’s disease brains. The amyloid fibrils are composed of protofilaments that are hydrogen-binding β-sheet structures with the β-strands running perpendicular to the long fibril axis. It is not clear whether the conformational alterations in the oligomers are

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a consequence of the oligomer assembling or whether the misfolded monomers induce the formation of the aggregates. An intermediate possibility is that an initial, unstable, misfolded conformation of peptide monomer can be stabilized by interaction with other proteins to form oligomers that lead the formation of fibrils and aggregates [89]. In this case, the conformational changes are not triggered by oligomerization, but their stable conformation is indissolubly associated with the oligomers. The formation of aggregates in Alzheimer’s disease brains is thought to be seeded by Aβ1–42, which is more prone to form aggregates, whereas Aβ1–40 accumulates subsequently. The essential role of Aβ1–42 is supported by evidence that the common molecular change associated with the mutations of three different genes linked to familial Alzheimer’s disease (AβPP, presenilin1 and presenilin2) is the increase of Aβ1–42 production [90]. The close association between the neurotoxicity and the aggregation of the Aβ peptide was subsequently reconsidered, and the hypothesis that Aβ deposition is a primary factor in the pathogenesis of Alzheimer’s disease has been challenged on the basis of studies showing that the degree of cortical synapse loss and the quantity of neurofibrillary changes predict the severity of dementia more accurately than the density of Aβ deposits [91, 92]. Recently, new data have emerged that lead to reconsideration of the amyloid cascade hypothesis; they support as the central pathogenic element the soluble oligomers of Aβ rather than the fibrillar aggregates of the peptide [93, 94]. Using amidation at the C-terminal of β25–35 to reduce the fibrillogenic capacity of the peptide, we demonstrated that the neurotoxic activity was independent of the aggregation state of the peptide [95]. Accordingly, several other studies suggested that neuronal death better correlates with the presence of oligomer species rather than with aggregates of Aβ peptides [96, 97]. It is possible that senile plaques represent the reservoir of aggregated Aβ that can continuously release diffusible oligomers and protofibrils to induce injury not only in the surrounding cells, but also at a certain distance of the senile plaques. Other theories advance the notion that neuronal death is triggered by intracellular events that occur during Aβ PP processing. In this regard, a role of intraneuronal Aβ in Alzheimer’s disease pathogenesis is suggested by the study of triple transgenic mice (Aβ PP/PS1/Tau). In the hippocampus of these animals, the intraneuronal accumulation of Aβ is an early event that precedes plaque formation and correlates with synaptic dysfunction [98]. A good correlation between total content of Aβ1–42 (biochemically determined in specific cerebral regions) and cognitive decline was recently found in Alzheimer’s disease, in keeping with the closer association of intellectual deterioration with Aβ production rather than with the density of plaques [99]. 3.2 Prion Diseases

As mentioned earlier, the pathogenic mechanism of prion diseases is based on the conformational conversion of the cellular prion protein (PrPC ) into the disease-specific species (PrPSc ). The “protein only” hypothesis on the nature of the


infectious agent (prion) indicates that it is essentially constituted of PrPSc [65] that is able to induce a “conformational transmission” via the host PrPC [100]. It has been postulated that other element(s) (called protein X) facilitate the conversion of PrPC to PrPSc [101]. Although the pathogenetic mechanisms are not fully clarified, the direct role of PrPSc in neuronal degeneration and glial activation is largely accepted. However, based on the evidence obtained in transgenic mice, Chiesa et al. [102] have demonstrated that the disease-associated, infectious form of the prion protein differs from the neurotoxic species. To evaluate the toxicity of PrPSc , in principle, one could directly apply the purified protein to neurons in culture. Although there have been several reports of these experiments [103], they are difficult to interpret because of uncertainties about the physical state of the PrPSc , since detergents that are required to maintain the protein in solution have to be removed prior to application to cell cultures. An alternative strategy has been to analyze the effect on cultured neurons of synthetic peptides derived from the PrP sequence [104]. The concept that misfolding of PrP causes a transmissible neurodegenerative disorder has prompted studies aimed at identifying polypeptide segments that are central to the conversion process. In particular, a synthetic peptide corresponding to the human PrP region 106–126 (PrP106–126) recapitulates several chemicophysical characteristics of PrPSc , including the propensity to form β-sheet-rich, insoluble and protease-resistant fibrils similar to those found in prion diseases [104, 105]. Similar to the studies with synthetic Aβ peptides, 10 years ago PrP106–126 was proposed as a model with which to investigate the biological effects of PrPSc [106]. This approach was successful with respect to the capacity of the synthetic peptide to mimic in vitro several aspects of the disease associated to the presence of PrPSc , including neurodegeneration, glial activation and alteration of membrane fluidity. Although PrP106–126 is not normally found in the brain of individuals with prion diseases, the various N- and C-terminal truncated fragments of PrP produced in the CNS in the course of sporadic and inherited prion diseases invariably contain the 106–126 sequence, suggesting that this region of the protein may possess the ability to trigger a fundamental pathogenic mechanism. The neurotoxic effect of PrP106–126 was abolished or reduced in neurons derived from PrP knock-out mice that are resistant to prion infectivity arguing for a role of the cellular prion protein in the neurotoxic cascade activated by PrP106–126 and underscoring the relevance of this model [107, 108]. Numerous other peptides homologous to PrP fragments were synthesized in wild-type and mutated forms, and their chemicophysical characteristics and biological effects investigated [109]. In some cases, the introduction of missense mutations associated to familial prion diseases (i.e. D178N) increased the fibrillogenic capacity of the PrP peptides [110]. The PrP106–126 peptide is highly fibrillogenic, but its in vitro toxicity was independent of this feature, while the ability to induce astrogliosis seems to be associated with the aggregation state of the peptide [110]. The amidation at the C-terminal combined with the acetylation at the N-terminal strongly reduced the fibrillogenic capacity of PrP106–126, but did not affect the toxicity of the peptide

17


[111]. These results are in agreement with the neuropathological evidence that PrPSc is deposited as amyloid fibrils only in particular conditions, as in GSS and vCJD, while in sporadic CJD PrPSc is assembled in non-fibrillar form, most likely as oligomers, strongly supporting the concept that soluble oligomers rather than mature amyloid fibrils are actually the pathogenetic species that causes neurodegeneration. It is noteworthy, in this regard, that an antibody recently developed and recognizing soluble oligomers of Aβ [112] prevented the toxicity induced by Aβ oligomers. Surprisingly, the antibody also recognized oligomer aggregates of SYN, islet amyloid polypeptide, polyglutamine, insulin and PrP peptides. As for Aβ, the antibody did not react with the monomeric form or the fibrillar version of these proteins, but only with the oligomers, indicating that the antibody recognizes a common structural epitope independent of the amino acid sequence. 3.3 Synucleinopathies

Epidemiological studies suggest that, in the etiology of synucleinopathies, environmental factors are more relevant than genetic ones. Nevertheless, recent data about the existence of rare familiar forms of Parkinson’s disease have been collected, and a pathogenetic role has been demonstrated for several genes, as SYN, parkin and DJ-1, whose mutations have been linked to familiar forms of Parkinson’s disease [113]. Two different missense mutations in the SYN gene (A30P and A53T) cause rare familial forms of Parkinson’s disease [77, 114] and recently a family with a new mutation (E46K) has been described [115]. Based on this evidence, SYN was identified as a major component of Lewy bodies [78]. On the other hand, a recessive form of juvenile Parkinson’s disease was associated with mutations in the gene encoding parkin [116]. Several pieces of evidence, most of which are based on the analysis of transfected cells, support the hypothesis that accumulation of SYN in Parkinson’s disease is a consequence of an alteration of the ubiquitin/proteasome system (UPS) [117, 118]. The self-aggregation capacity of SYN [119, 120] and the ubiquitin ligase activity attributed to parkin [121, 122] suggest that protein aggregation and dysfunction of the UPS might play a causal role in the development of sporadic and familial Parkinson’s disease. SYN is a highly conserved 140-amino-acid protein widespread in the CNS. It interacts with other cerebral proteins (14-3-3, synphilin-1, parkin, tyrosine hydroxylase, dopamine transporter) and it is involved in dopamine vesicle trafficking [123]. In aqueous solution, free SYN shows a natively unfolded conformation that favors protein self-aggregation and toxicity [124]. Experimental injuries reproducing possible Parkinson’s disease triggers (toxins, oxidative stress, proteasome impairment) increase SYN aggregation [125–127]. Like the other amyloidogenic proteins, SYN aggregation follows a multistep process, starting from SYN monomers that form oligomers (protofibrils) whose coalescence generates fibrils that, in turn, can aggregate in inclusion bodies. Recent data suggest that the protofibrils are the toxic intermediate, while the final inclusions may have a protective value [128]. Several in vitro studies have confirmed that SYN mutations


predispose to protein aggregation and the overexpression of the mutated forms of the protein are toxic for the cell [129, 130]. According to this hypothesis, the accumulation of SYN in the Lewy bodies is a toxic event that eventually triggers neuronal death [131]. We found that the fragment corresponding to the residues 61–95 [also known as non-amyloid component of plaques (NAC)] was specifically neurotoxic for the dopaminergic cells and the pre-aggregation of the peptide amplified this effect [132]. Conversely, other studies demonstrated that SYN can exert neuroprotective activity against various cellular stresses like oxidative stress, serum deprivation and apoptosis [133, 134]. In our investigation, we have also noted that PC12 transfected with SYN were more resistant to oxidative stress. This evidence was derived mainly by in vitro overexpression of the wild-type protein whose protective role was reduced by the mutations A30P and A53T. Accordingly, the loss of SYN protective function, rather than a gain of toxic function, may determine the cell death in Parkinson’s disease. Nevertheless, the situation is not completely clear, as other authors have reported a protective role also for the mutated forms of the protein [134]. It is possible that the two apparent opposing actions of SYN are not mutually exclusive. SYN may be at first neuroprotective, but, if the protein self-aggregation becomes relevant, the disease occurs. In this regard, it was recently reported, using an in vitro model that the extracellular administration of wild-type SYN on the nanomolar scale is protective against oxidative stress, while the overexpression of the protein or its extracellular administration on the micromolar scale is toxic [135]. We have confirmed these data using a chimeric SYN associated with the TAT [136] sequence (TAT–SYN) to translocate the protein inside the cells. Mutated (A30P, A53T) and wild-type TAT–SYN sequences equally protected against oxidative stress induced by hydrogen peroxide in PC12 and the effect was mediated by the induction of HSP70 protein [137]. The protective effect by TAT–SYN in SK-NBE neuroblastoma cells was efficacious also against the dopamine-specific neurotoxin 6-hydroxydopamine (6OHDA). A third important point that has been intensively investigated for Parkinson’s disease is the so called “selective vulnerability”, i.e. why only a particular region of the brain (substantia nigra pars compacta) is affected even though SYN is widely expressed in brain and the factors that probably trigger the disease are not region specific. Recent studies have pointed out that a first important factor is dopamine synthesis and secretion that are peculiar features of the neurons of the affected areas [138]. In fact, dopamine is a molecule that is prone to oxidation, which may contribute to generate reactive molecules potentially able to damage cellular components like proteins or lipids. From this point of view several experimental data indicate that SYN is implicated in dopamine metabolism and trafficking [139, 140] suggesting a relationship between protein function, dopamine homeostasis, oxidative stress and neuronal damage. The development of animal models has provided further insight about SYN pathogenetic mechanisms. Transgenic mice overexpressing human SYN (wildtype or mutated) showed intraneuronal inclusions, but the associated phenotype was quite heterogeneous as for the neuronal populations involved and no model recapitulated completely the human pathology. For instance, transgenic mice

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expressing human SYN gene carrying the A53T mutation under Thy-1 promoter control [141] developed a progressive motor impairment starting from 40 days post-birth. Many telencephalic and brainstem neurons and the neurons of the spinal cord show a strong immunoreactivity to SYN antibodies in cellular bodies and dendrites; this picture differs from the axonal and presynaptic distribution of endogenous SYN in control mice. Brainstem neurons and motor neurons seem particularly vulnerable: motor neurons alterations comprise axonal damage and neuromuscular junction denervation, suggesting that an increased SYN expression may interfere with synaptic trophic mechanisms. Otherwise, transgenic mice with an elevated human SYN expression under platelet-derived growth factor promoter control showed a dopaminergic neuronal loss, and SYN- and ubiquitin-positive inclusions formation in different cerebral areas [142]. 4 Conclusions

Although the presence of aggregates made up of misfolded proteins is a common feature of many neurodegenerative disorders, the biological processes responsible for this phenomenon are variable and its significance in the pathogenesis of the diseases is different. As illustrated in this chapter, the neuropathological manifestations may be extremely heterogeneous even within disorders characterized by aggregates made up by the same protein. In some conditions, like Alzheimer’s disease, prion diseases and tauopathies, the evidence supporting a pathogenic role of the aggregates is quite compelling, although the molecular mechanisms triggering the protein accumulation remain to be established. In the synucleinopathies, the responsibility of the pathologic inclusions in determining the degenerative process is more uncertain. In ALS and diseases with CAG repeat expansions the role played by the aggregates is still elusive, although the number of studies focused on their potential pathogenic role is growing. Finally, the consensus around the neurotoxic role of oligomer species rather than fibrils themselves is vast and propose new therapeutic approaches at these diseases. Acknowledgment

Supported by the Italian Ministry of Health, Department of Social Services (PS03–4 and PS-03–10) and by the European Community (LSHM-CT-2004–503039 and Neuroprion FOOD-CT-2004–506579). References 1 TAYLOR, J. P., HARDY, J. and FISCHBECK, J. H. Toxic Proteins in neurodegenerative disease. Science 2002, 296, 1991–1995. 2 SOTO, C. Unfolding the role of protein misfolding in neurodegenerative

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1

Cellular Toxicity of Protein Aggregates Bruce Kagan

University of California, Los Angeles, USA

Originally published in: Amyloid Proteins. Edited by Jean D. Sipe. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3–527–31072-X

1 Introduction

“Amyloid” was first described by Virchow in the 19th century as an amorphous starch-like deposit in tissues that stained with iodine, indicating the presence of carbohydrate. Investigators in the 20th century concluded that amyloid deposits were largely proteinaceous and exhibited a characteristic staining with the dye Congo red. It also became clear that many different proteins could form amyloid deposits and that these deposits were composed of fibrils with a characteristic structure (for review [1]. Fibrils are 80–100 Å in width and often of very extended length. At least 21 known proteins and peptides have been reported to deposit as amyloid (Table 1) and are associated with a wide variety of human illnesses [2]. Although amyloid-forming proteins exhibit no amino acid sequence homology, they all adopt a characteristic β-sheet structure within amyloid fibrils. Amyloid deposits also contain a variety of common components, such as glycosaminoglycans, proteoglycans and the pentraxin serum amyloid P. The structural role of these components in amyloid fibrils and deposits is unclear. Although β-sheet conformation is at the essence of amyloid fibril formation, its role in determining pathology has been less clear. Amyloid deposits were often found to be associated with disease, but were also found in otherwise apparently healthy tissue. Subsequently, studies began to show that amyloid deposits are associated with disease and, perhaps, play an etiologic role [3]. However, the experimental evidence for a correlation of clinical illness with amyloid burden has been irregular and unpersuasive. Recent studies have suggested that smaller, soluble aggregates of proteins, many of which go on to form amyloid fibrils, may be responsible for cellular dysfunction and death. These early aggregates will be referred to as oligomers in this chapter, with the understaning that the term oligomer might refer to species containing as few as two or as many as thousands of molecules. More Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Cellular Toxicity of Protein Aggregates Table 1 Diseases of protein misfolding: amyloidoses

Disease

Protein

Abbreviation

Alzheimer’s disease Down’s Syndrome (trisomy 21) Heredity cerebral angiopathy (Dutch) Kuru Gerstmann-SträusslerScheinker syndrome (GSS) Creutzfeldt-Jacob disease Scrapie (sheep) Bovine spongiform encephalopathy (“mad cow”) Type II diabetes mellitus (adult onset) Dialysis-associated amyloidosis Senile cardiac amyloidosis Familial amyloid polyneuropathy Reactive amyloidosis (familial Mediterranean fever) Familial amyloid polyneuropathy (Finnish) Macroglobulinemia Primary systemic amyloidoses Familial Polyneuropathy – Iowa (Irish) Hereditary cerebral myopathy – Iceland Non-neuropathic hereditary amyloid with renal disease Non-neuropathic hereditary amyloid with renal disease Familial British dementia

amyloid β precursor protein (Aβ1–42)

AβPP (Aβ1–42)

prion protein

PrPC /PrPSc

islet amyloid polypeptide (amylin)

IAPP

β 2 -microglobulin atrial natriuretic protein transthyretin

β2M ANP TTR

serum amyloid A

SAA

gelsolin

AGel

λ1 heavy chain Igλ, Igκ apolipoprotein AI

AH AL ApoAI

cystatin C

ACys

fibrinogen α

AFibA

lysozyme

ALys

FBDP

ABri

precise numbers may come from future studies, but the important distinction is that these oligomers exist in solution (at least for a time), as opposed to fibrils which precipitate out of solution. A number of common, significant, costly, and devastating diseases are amyloidoses. These include Alzheimer’s disease, Type 2 diabetes mellitus and the spongiform encephalopathies [prion diseases, e.g. scrapie, “mad cow” disease, new variant Creutzfeldt-Jakob disease (Table 1)]. The aging of the population in the developed world has caused an epidemic of these age-related diseases to begin, making the search for an understanding of disease pathogenesis more critical than ever. Although a great deal of research has strongly implicated amyloid proteins in the pathogenesis of these illnesses, the molecular mechanisms by which these misfolded proteins cause dysfunction and/or toxicity remains elusive (for review, see [4]). In this article, I will first review the evidence that oligomers (as opposed to fibrils) of amyloid peptides are the pathogenic species. I will go on to

2 Aggregation Table 2 Diseases of protein misfolding: non-amyloidoses

Disease

Protein

Abbreviation

Diffuse Lewy body disease Parkinson’s disease Fronto-temporal dementia Amyotrophic lateral sclerosis Triplet-repeat diseases: Huntington’s spinal and bulbar muscular atrophy spinocerebellar ataxias spinocerebellar ataxia 17

α-synuclein

AS

tau superoxide dismutase-1 polyglutamine tracts in the following proteins: huntingtin androgen receptor

tau SoD-1 PG

ataxins TATA box-binding protein

review the current evidence as to how these oligomers cause cellular dysfunction and/or toxicity. It has been also reported that protein misfolding and aggregation is linked to the pathophysiology of a number of other, non-amyloid diseases (Table 2). Furthermore, it has been reported that non-disease-related proteins can form amyloid oligomers with toxicity. Here, the links amongst these various disease-associated phenomena will be considered.

2 Aggregation

The early findings that amyloid burden in patients did not correlate well with clinical severity of illness was troubling to investigators who were thinking along the lines that amyloid fibrils and deposits play an etiologic role in amyloidoses. Subsequently, the discovery of familial forms of amyloidoses linked to mutations in amyloid proteins gave new impetus to the notion that these proteins “caused” the illness [5, 6]. Further research, initially with the Aβ1–42 peptide of Alzheimer’s disease, showed that amyloid peptides could demonstrate cytotoxicity in isolated cell systems and that these peptides were toxic to the cell types damaged in diseases [7]. Thus, Aβ and PrP106–126 [8] were shown to be toxic to neurons, and IAPP was shown to be toxic to β cells from the islets of Langerhans in the pancreas [9]. Further experiments showed that amyloid fibrils themselves usually lacked toxicity. Monomers, also, seemed to lack cytotoxicity. In early experiments, however, cytotoxicty was poorly reproducible, especially for the Aβ peptides. This was eventually shown to be due to differences in the aggregation status of Aβ under varying experimental conditions [10]. The aggregation of amyloid fibril-forming peptides was found to be a highly complex and variable process [11]. The initial phases of protein aggregation from monomers into small oligomers progressed quite slowly. However, once a critical mass was reached, aggregation proceeded quite rapidly. Thus, growth of fibrils could be “nucleated” by the addition of pre-formed “seeds” to a solution of peptide. The seeds greatly accelerated the aggregation of the amyloid protein. However, it was soon discovered that, while promotion of amyloid peptide aggregation (by

3

4 Cellular Toxicity of Protein Aggregates

acidic pH, high concentration, “aging” or seeding) was necessary to achieve toxicity, extensive aggregation could actually lead to a decline in cytotoxicty [12]. It appeared that an intermediate state of aggregation was associated with the cytotoxicity of amyloid fibril-forming peptides. The notion of amyloid peptide-induced change also broadened. While cytotoxicity was clearly relevant to diseases in which cells died, animal models of Alzheimer’s disease created with mice transgenic for the Aβ precursor protein (AβPP) or Aβ developed amyloid deposits, learning and memory deficits, and synaptic loss, but no frank neuronal loss. Although some criticized the models as inadequate, or attributed the lack of nerve loss to differences between mice and men, others began to think that, perhaps in the early stages of Alzheimer’s disease, there were characteristic dysfunctions that precede neuronal loss. Developmental investigations of transgenic mouse models showed that memory deficits actually preceded amyloid deposits and synaptic loss. Further studies showed that Aβ peptides could inhibit long-term potentiation, a model for memory [13]. Additionally, only oligomeric forms of Aβ could inhibit long-term potentiation, not monomeric or fibrillar Aβ [14]. These observations led to the implication of oligomers in cellular dysfunction relevant to clinical symptoms and to cytotoxicity. In vitro, it was shown that fibroblasts could be killed by Aβ that was “freshly prepared” (and thus less highly aggregated) and that appeared to be “globular” by atomic force microscopy. Toxicity was blocked by antibodies or the channel-blocking zinc ion, but not by antioxidants [15]. A similar story emerged in Huntington’s disease, where early deficits in learning and memory could be linked to the presence of aggregates of the disease related protein huntingtin with a long PG tract. These deficits appear well before any frank neuronal loss in the brains of affected transgenic mice. Electrophysiologic abnormalities accompanied these deficits and could be detected even prior to the appearance of huntingtin aggregates, suggesting that the large, visible aggregates may not be responsible for the early cellular effects of huntingtin with extended PG tracts [14]. It has also been shown that PG aggregates targeted to the nucleus can be directly cytotoxic [17]. Oligomers have also been implicated in the cytotoxicity of AS. Mutations in AS which promote “protofibril” (oligomers) formation also enhance cytotoxicity [18]. Similarly, the “arctic” mutation of Aβ, which enhances oligomer formation, shows increased cytotoxicity in comparison to native Aβ peptide [19]. Toxic oligomers of IAPP were shown by light scattering to contain between 25 and 6000 IAPP molecules [20]. Once again, fibrils and monomers were shown to lack toxicity. An antibody preparation to the “soluble oligomer” conformation was generated, which recognized several different amyloid peptides, but only in this precise conformation [21]. This antibody could block cytotoxicity of a range of amyloid peptides, strongly suggesting that oligomers were the toxic species and that the toxic mechanism (or at least a part of it) was common to all the amyloids. Biological dogma has maintained that the three-dimensional structure and function of proteins is dictated by their primary amino acid sequence. Our knowledge of amyloid diseases has led to the recognition that natively folded proteins may

2 Aggregation

unfold at times and adopt non-native conformations. Proteins may refold to their native conformation (with or without the assistance of chaperones) or they may be targeted for degradation via the ubiquitin–proteasome pathway. Alternately, some partially unfolded or misfolded proteins may aggregate. This aggregation may be driven thermodynamically by new hydrogen bonding possibilities [22] or by the hydrophobic effect, the need to shield hydrophobic parts of proteins from the aqueous environment. Proteins that form amyloid appear to have a propensity to misfold into β-sheet structures. These β-sheets tend to self-aggregate, forming intermolecular bonds that aggregate the proteins or peptides. Proteins that possess hydrogen-bonding defects are more likely to interact with lipid bilayers [22]. It is still not well understood why these complexes are not handled by the cell’s usual defense mechanisms, i.e. refolding to native structures through chaperone assistance or targeting for degradation through ubiquitin–proteasomes. It may be that these processes are at work, but are merely overwhelmed by the amount of misfolded protein they must handle. This might be the place where disease begins. Alternatively, it is possible that the nature of these β-sheet aggregates leads them to attack or insert into cellular membranes. This would render them inaccessible to chaperones and to the ubiquitin–proteasome pathway. Another possibility is that, once seeding or nucleation has occurred, the fibrillation process proceeds so rapidly that the aggregates become too large to be handled by the cell’s defenses. It has been suggested that amyloid fibril formation, with or without vacuolization, is actually a cellular defense mechanism to isolate the misfolded protein and keep it separate from the cell’s vital machinery. Support for this idea comes from experiments showing that amyloid fibrils have little, if any, toxicity to cells, while smaller aggregates appear to possess considerable cytotoxicity. The initial misfolding of a protein may be driven by mutations, metal interactions, environmental conditions such as temperature, acidic pH, oxidation and proteolysis or a simple increase in concentration. All these phenomena can potentially increase the presence of misfolded conformations and increase the probability of aggregation. The presence of membranes may also play a key role in this process, as lipid bilayers have been shown to influence both the conformation of amyloid peptides and the propensity of amyloid peptides to aggregate [23]. Thus, the intracellular component in which an amyloid protein finds itself may significantly affect its ultimate conformational fate. The many amyloid peptides exhibit no primary sequence homology, but they all possess the possibility of converting to relatively high β-sheet content. Sometimes this β-sheet content is unpredictable, as in the prion protein where sequences which would usually be predicted to be α-helical adopt a β-sheet conformation when synthesized [8]. While the earliest stages of amyloid peptide aggregation cannot be easily observed with current imaging techniques, the results of recent electron and atomic force microscopic studies have shown the presence of spherical or globular aggregates early in the process of amyloid fibril formation. The smallest particle appears to aggregate into chains or annular rings referred to as “protofibrils”. These structures appear to be the precursors of fibrils that exist in mature amyloid deposits. The

5


protofibrillar structures possess significant toxicity in comparison to fibrils themselves. It is unclear whether the annular structures or rings are more toxic than their linear brethren, but it is of interest that pathogenic mutations of Aβ amyloid and AS lead to increased formation of these annular structures [18]. 3 Cellular Mechanisms of Oligomeric Toxicity

Recently, it has been reported that proteins and peptides associated with amyloid disease are not unique in their ability to form amyloid. Aggregation and amyloid fibril formation have been achieved for a number of non-disease associated proteins (Table 3) [20]. It has been proposed that under appropriate conditions all proteins might be capable of forming amyloid fibrils. Beyond the structural similarity of these non-disease-associated amyloid proteins to actual amyloid proteins, there are functional similarities as well. For example, HypF is able to aggregate, and the aggregates can kill cells and permeabilize liposomes [95]. Fibrils lack these properties, but aggregation of HypF is required for toxicity and permeabilization, thus implicating smaller oligomers as the cytotoxic, membrane-permeabilizing species. Preceding cytotoxicity, HypF oligomers, but not fibrils, raised intercellular Ca2+ levels and levels of reactive oxygen species [11]. These intracellular effects were reversible, and they were prevented with the omission of Ca2+ from the media or the addition of reducing agents. These physiologic effects are strikingly similar to those observed with disease related amyloid peptides [26, 27]. These results are also consistent with the findings that antibodies raised against soluble pre-fibrillar oligomers of various amyloid peptides recognize that state in other amyloid peptides and can prevent cytotoxicity [21]. This suggests that amyloid peptides share an oligomeric conformation critical to cytotoxicity and independent of amino acid sequence. Thus, a wide variety of structural, biochemical, and physiologic studies suggest that pre-fibrillar amyloid peptide oligomers act to damage and/or kill cells, especially neurons, and that they appear to share a common mechanism of action. As we discuss below, a growing body of biophysical evidence implicates channel formation

Table 3 Non-disease related amyloid-forming proteins/peptides

SH3 domain p 85 Phosphatidylinositol-3-kinase HypF N-terminal domain (E. coli) Apomyoglobin (equine) Endostatin (human) StefinB(human) Fibroblast growth factor (Notophthalmus viridescens) VI domain (murine) Curlin CgsA subunit

fibronectin type III phosphoglycerate kinase acylphosphatase amphoterin (human) apocytochrome c Met aminopeptidase ADA2H apolipoprotein CII B1 domainof IgG-binding protein monellin

7 The Channel Hypothesis 7

by amyloid peptides in cellular membranes as the molecular mechanism of this damage. 4 Loss of Function Hypothesis

This view postulates that cytotoxicity results from a loss of native protein function due to misfolding of native protein and aggregation. This idea seems untenable in light of the fact that so many different, structurally and functionally unrelated proteins are involved in amyloid diseases. Transgenic mouse models in which an amyloid protein-encoding gene is inserted and disease results also argue against this idea. An alternative form of this concept is that aggregated amyloid proteins might bind and inactivate essential cellular proteins, such as transcription factors. However, there is little evidence to suggest that physiologically significant amounts of critical cellular proteins can be found in amyloid deposits or inclusion bodies. Substantial evidence has now accumulated to show that amyloid proteins possess toxic activity toward cells. The molecular mechanism of this toxicity remains unknown. 5 Receptors for Advanced End-products of Glycation (RAGE) Receptors

It has been proposed that RAGE bind various amyloid oligomers, inducing cellular stress and activation of NF-κB [10]. While some experimental evidence supports this idea, it has not been clearly documented that this mechanism accounts for tissue damage in amyloid diseases. Further investigation is warranted. 6 Oxidative Stress

Another hypothesis for the pathogenesis of amyloidosis and amyloid-like diseases is that oxidative stress is induced directly by amyloid peptides, leading to free radical production, mitochondrial dysfunction and death [13]. Although many elements of oxidative stress pathology occur in amyloid induced cytotoxicity, it does not explain how oxidative stress and/or free radicals are generated by amyloid peptides. Initial reports that the peptide itself could generate free radicals have not been confirmed. Oxidative stress clearly occurs in cells affected with amyloid toxicity. It remains to be established where in the chain of toxicity oxidation plays its role. 7 The Channel Hypothesis

A large body of research has focused on the ability of amyloid peptides to interact with membranes. A series of provocative studies has shown that many (if not all)


amyloid peptides can aggregate into β-sheet oligomers capable of spontaneously inserting into lipid membranes. The peptides form relatively permanent ionpermeable channel structures across the cell membrane. These channels are reported to have properties that would damage or kill most cells (Table 4). The channels are reported to be (a) large, (b) non-selective, (c) heterogeneous, (d) voltage independent, (e) irreversible, (f) inhibited by agents that prevented aggregation such as Congo red and (g) blocked by zinc ions. These channels would likely cause a leakage pathway in plasma, mitochondrial, endoplasmic reticulum, lysosomal or other membranes. These leaks could damage cells by (1) disrupting membrane potentials and ion gradients, (2) causing loss of vital intracellular ions such as K+ and Mg+ , (3) allowing influx of toxic ions such as Ca2+ , (4) running down energy stores by forcing ion pumps to work harder, (5) disrupting mitochondrial membrane potential and initiating apoptosis by allowing cytochrome c to leak out of mitochondria, and (6) allowing toxic enzymes and other factors to leak out of lysosomes and peroxisomes. In the sections that follow, the evidence for channel formation by the amyloid peptides which have been most studied by these techniques is reviewed.

8 Aβ

The “channel hypothesis” was first proposed by Arispe et al. [3] after they reported that Aβ1–40 could form cation-selective, calcium permeable channels of various conductances in planar lipid bilayer membranes (BLMs). The channels were formed by Aβ1–42 as well and were large, voltage independent and blocked by tromethamine (Tris+ ) and aluminum [4]. The largest channels observed (4 nS) could potentially change the interior [Na] of a cell by as much as 10 µM/s. They proposed that ionic leaking of Na+ , K+ and Ca2+ could disrupt membrane potential and ionic regulation within a few seconds. While these findings were not immediately confirmed by other laboratories, due to problems with irregular aggregation of Aβ [32], eventually a series of studies found Aβ peptides to be capable of forming channels in BLMs [32, 34], liposomes [32], neurons and oocytes [24], and fibroblasts [36]. The state of Aβ aggregation is critical not only to its cytotoxicity [10, 33], but also to its channel-forming abilities. Indeed, monomers and fibrils of Aβ that are non-toxic fail to form ion channels [32]. Oligomeric species of Aβ cause a variety of channel entities, which can be distinguished by their single-channel conductance, ionic selectivity, kinetics and other channel properties [38]. It was also found that conditions (aging, acidic pH, etc.) that favor the aggregation of monomers into oligomers led to an increase in channel activity [32]. Exposure to organic solvents, which promote monomeric Aβ, led to loss of channel activity. However, channel activity could be recovered by allowing the peptide to “age” in aqueous solution. Although Aβ1–40 and 1–42 are the primary forms of Aβ peptides found in vivo, other Aβ fragments have been of experimental interest. Aβ25–35, a cytotoxic peptide not found in vivo, is a voltage-dependent, non-selective channel former [32]. Studies using variants of Aβ25–35 showed that channel formation was necessary for

Table 4 Channel properties of amyloid peptides

Peptide Aβ25–35 Aβ1–40 Aβ1–40 Aβ1–40 ARC (E22G) Aβ1–42 CT105 (C-terminal fragment of amyloid precursor protein (APP) Islet amyloid polypeptide (Amylin) PrP106–126 PrP106–126 PrP82–146 SAA SAA 2.2 (murine hexamer) C-type natriuretic peptide Atrial natriuretic factor Beta2 -microglobulin Transthyretin Polyglutamine (average molecular weight = 6000) Polyglutamine 40 NAC (AS65–95) AS A30P/A53T CT (human/salmon) Lysozyme 87–114

Single-channel Ion selectivity conductance (pS) (permeability ratio)

Blockade by zinc by Congo red

10–400 10–2000 50–4000

cation (PK /PCl =1.6) + cation (PK /PCl =1.8) + cation (PK /PCl =11.1) +

+

[59, 70] [32] [3, 4], [3, 30] [55]

10–2000 120

cation (PK /PCl =1.8) cation

+ +

+ +

[32] [50]

7.5

cation (PK /PCl = 1.9) +

+

[71]

10–400 140, 900, 1444

+

+

10–1000

cation (PK /PCl =2.5) cation (PK /PCl >10) cation (variable) cation (PK /PCl =2.9)

+

+

21, 63 68, 160, 273 0.5–120 variable 19–220

cation (PK /PCl > 10) + variable non-selective + cation (variable) + non-selective –

+ + + –

[60] [49] [6] [35] [99] [35] [50] [30] [30] [15]

17 10–300

cation variable

+

12/580

non-selective non-selective

15–20/70–100

25/80

20–25/80–120 permeable to β-galactosidase (molecular weight 116 kD) 50/190

Reference

+

[73] [5] [99] [96] [36] 8 A$

Cu/Zn SoD

Ring diameter (inner/outer, in Å, by microscopy)

Chung et al. (2003) 9


cytotoxicity, but not sufficient, i.e. all cytotoxic species formed channels, but there were two channel forming variants of Aβ25–35 that did not kill cells [39]. Channel activity could be enhanced by lipids carrying a net negative surface charge and this effect could be countered by high salt concentrations. Addition of cholesterol, which stiffens membranes, decreased Aβ25–35 channel activity [40]. Aβ25–35 variants could not form channels if they were not at least 10 residues long, indicating a minimum bilayer spanning length of about 30 Å, a result consistent with the β-sheet span lengths of the known channels generated by Staphylococcal α toxin and anthrax toxin [41, 42]. More recently, an extremely short channel-forming Aβ variant (31–35) has been reported [43]. Whether this peptide might form hemi-channels to span the bilayer similar to the peptide Gramicidin is unknown. In vivo, Aβ1–40 or 1–42 can induce currents in rat cortical neurons [25, 44], HNT cells [46] and gonadotrophin-releasing hormone secreting neurons [42]. The channels observed in vivo seem indistinguishable in their properties from those observed in vitro. Aβ1–40 or 1–42 can also kill fibroblasts in a manner inhibited by antibodies, tromethamine or zinc, but not by antioxidants, suggesting that channel formation is the mechanism of cytotoxicity. The freshly prepared Aβ used in these studies appeared “globular:” consistent with an early stage of aggregation [8, 36]. It has also been shown that the cholesterol content of plasma membranes affects a cell’s vulnerability to Aβ1–40 and 1–42 [2], suggesting that the membrane plays a critical role in Aβ cytotoxicity. Also, it has been reported that Aβ can directly induce cytochrome c with release from mitochondria. This could occur through the action of Aβ to decrease mitochondrial membrane potential or even through Aβ channel mediated release of cytochrome c [46].

9 PrP106–126

The prion protein (PrP) has at least two distinct tertiary conformations, PrPC and PrPSc , the latter of which results in a transmissible neurodegenerative disease known as a spongiform encephalopathy. Prion diseases include scrapie in sheep and “mad cow” disease as well as Creutzfeldt-Jakob disease, Gerstmann-SträusslerScheinker syndrome and fatal familial insomnia in humans. These illnesses may be sporadic, infectious or hereditary. The familial versions are associated with mutations in the prion protein [16]. PrPSc deposits in the brains of afflicted organisms in a form that is readily converted in amyloid fibrils in vitro. A critical step in the conformational transition from PrPC to PrPSc is the conversion of α-helical and random coil regions of PrP to β-sheets [52]. One region predicted to be α-helical, PrP106–126, actually forms β-sheets when chemically synthesized and self-aggregates into amyloid fibrils [26]. The β-sheet-rich form of PrP106–126 binds to membranes, unfolds and ultimately disrupts the bilayer [45]. Forloni et al. [23] demonstrated that PrP106–126 was toxic to neurons in culture. Lin et al. [56] reported that PrP106–126 could form ion-permeable channels in planar lipid bilayer membranes at neurotoxic concentrations. PrP106–126 channels were irreversibly associated with the membranes, demonstrated a multiplicity of single

9 PrP106–126 11

channel conductances (10–400 pS in 0.1 M NaCl) and had relatively long lifetimes (seconds to minutes). Ionic selectivity of the channels was meager, with significant permeability being shown to Na+ , K+ , Cl− and Ca+ (PNa /PCl = 2.5). Channel activity could be enhanced dramatically by “aging” of the peptide in aqueous solution, a procedure which promotes aggregation and increases neurotoxicity. Incubation of PrP106–126 at acidic pH also enhanced channel activity by nearly 100 times and shifted the distribution of observed single-channel conductances to higher conductance levels. It has also been reported that acidic pH converts α-helical PrP106–126 to the β-sheet conformation [8]. Kourie and Culverson [49] characterized three distinct channel types formed by PrP106–126. These included: (1) a dithiodipyin sensitive channel of 40 pS with slow kinetic behavior, (2) a giant channel, 900–1500 pS, exhibiting five separate subconductance states and (3) a tetraethyl-ammonium chloride (TEA)-sensitive channel of 140 pS with rapid kinetics. Manunta et al. [58] were unable to observe PrP106–126 neurotoxicity or channel formation, but this may have been a result of the highly variable aggregation state of the PrP106–126 peptide, reminiscent of the variability seen with aggregation of Aβ peptides. Bahadi et al. [6] have reported that PrP82–146, a peptide found in the PrPSc brains of patients with Gerstmann-Sträussler-Scheinker syndrome, can also form ion channels. Scrambling the amino acid sequence of the 106–126 region of this longer peptide abolishes the ability to form ion channels, whereas scrambling the 127–146 region has no effect, thus implicating the 106–126 region as key in channel-forming ability. The electrophysiologic properties of PrP82–145 are very similar to those of PrP106–126. Channel activity could be decreased by the antibiotic rifampicin which had previously been shown to decrease aggregation and toxicity of Aβ peptides. The association of amyloid deposits with prion diseases is variable. Intriguingly, in one prion disease where amyloid fibrils are not found, the mutant prion protein adapts a transmembrane conformation [29]. It is tempting to speculate that this transmembrane protein may be causing ionic leakage across the cell membrane. Lysosomotropic agents have been reported to inhibit PrPSc accumulation in neuroblastoma cells [18]. One of these agents, quinacrine, has been reported to block PrP106–126 channels [6]. Quinacrine is also able to repair the impaired functioning of N-type calcium channels in prion-infected neurons [62]. Thus, it seems likely that channel blockers, such as quinacrine, may be useful as potential therapeutic agents in prion related diseases. Indeed, there is at least one report of quinacrine improving the clinical status of four patients with Creutzfeldt-Jakob disease [63]. Other acridine derivatives and tricyclic compounds may have even better antiprion efficiency [47]. Congo red can inhibit channel formation, block PrP106–126 cytotoxicity and inhibit the development of scrapie [33, 37]. It remains to be seen whether the anti-channel blocking or anti-aggregation activities of quinacrine are key to its anti-prion effects. It has also been reported that a peptide PrP170–175 bearing a prion protein mutation related to schizoaffective disorder [6] increases the permeability of planar lipid bilayers and forms channels with conductance of 8–26 pS in 0.5 M KCl. The native PrP170–175 does not form channels in membranes. This result suggests that yet another segment of PrP may be capable of influencing toxicity via channel formation and that this may be directly relevant to human disease.


10 IAPP

IAPP (amylin) is a 37-residue amyloidogenic hormone which is co-secreted with insulin from β cells in the islets of Langerhans in the pancreas. Amyloid deposits comprising IAPP are found in the islets of patients with Type 2 diabetes, and are positively correlated with β cell loss and clinical insulin requirements [12, 66]. IAPP is cytotoxic to β cells in culture [9]. Although IAPP is α-helical in aqueous solution, exposure to lipid membranes induced a transition to the β-sheet structure [67]. Human IAPP formed ionpermeable channels in planar lipid membranes at cytotoxic concentrations [39]. Rat IAPP, which differs from human IAPP at five amino acid positions, and is non-amyloidogenic and non-toxic, did not form channels. Human IAPP channels could be inserted into membranes irrespective of voltage; however, once inserted, channels rapidly opened at negative voltages and rapidly inactivated at positive voltages (voltages being relative to the IAPP-containing side). Inactivation faded gradually over a time course of several minutes. Open IAPP channels were ohmic and exhibited a single-channel conductance of 7.5 pS in 0.1 M KCl. Channels were permanently associated with the membrane and showed lifetimes of seconds to minutes depending on voltages. Increasing concentrations of net negatively charged lipids in the membrane led to an increase in IAPP channel activity. Increasing salt concentrations in the aqueous solution decreased channel activity. Anguiano et al. [1] showed that liposomes could be permeabilized by IAPP in a graded fashion, allowing Ca2+ to cross the membrane, while not allowing fura-2 (molecular weight = 832) or FITC-dextran (molecular weight = 4400) to escape. IAPP has also been reported to disrupt Ca2+ homeostasis in cells in a manner similar to Aβ and prion-related peptides [27]. Large fibrils of IAPP have been reported to be non-toxic, whereas smaller aggregates are associated with cytotoxicity [20]. The aggregates, but not fibrils, could disrupt planar lipid bilayers. Light scattering showed these oligomers to range in size from 25 to 6000 IAPP molecules. Hirakura et al. [33] showed that Congo red incubation with IAPP, Aβ or PrP106–126, prior to membrane exposure, could inhibit channel formation. They also reported that Zn2+ could reversibly block these channels. The concurrence of channel-forming properties, physiologic effects and cytotoxicity strongly suggests a common mechanism for the channel-forming action for these three amyloid peptides.

11 ANP

A family of hormones, C-type natriuretic peptide (CNP), ANP and brain-derived natriuretic peptide (BNP), helps to regulate fluid and ionic balance. As people age, increasingly large amyloid deposits of ANP are found in their hearts. These fibril-containing deposits are thought to play a deleterious role, perhaps leading

12 SAA 13

to atrial fibrillation and other cardiac pathology [64]. Channels in lipid bilayers have been reported for ANP1–28 [34], CNP-22 and OaC-type natriuretic peptide (18–39) from platypus [48]. ANP channels exhibited a multiplicity of single channel conductances, but all were cation selective. The channels could be divided into three types: (1) Ba2+ -sensitive, fast kinetics with three modes (spike, burst and open), 68 pS, (2) a large conductance channel possessing subconductance states, time dependent inactivation, 273 pS, and (3) transient activation, 160 pS. CNP channels were weakly cation selective, with a high open probability and large single-channel conductance (546 pS). The physiologic properties of these peptides were believed to be concordant with known pathological effects described in animal models and human disease. Although the peptides act through a well-known receptor and second messenger signaling system, it has been proposed that the channel-forming activity of the peptides could play a role in physiologic events. ANP channels would likely hyperpolarize muscle cell membrane and inhibit depolarization driven contractions. Large conductance channels were postulated to degrade membrane potential and ionic balance.

12 SAA

SAA refers to a family of related apolipoproteins. During states of infection or inflammation the acute-phase isoforms of SAA can increase their levels in serum by as much as 3 orders of magnitude. AA fibrils most commonly comprising the N-terminal 76 residues of SAA are found as amyloid deposits in various organs such as spleen, kidney and liver. Patients with chronic infections, such as tuberculosis, or inflammatory diseases, such as rheumatoid arthritis, are particularly at risk. Patients with cancer, arteriosclerosis and Alzheimer’s disease have also been reported to have elevated SAA concentrations [72]. One commercially available, recombinant-generated acute-phase isoform, SAAp, has been reported to form ion-permeable channels in planar lipid bilayer membranes at physiologically relevant concentrations [35]. A wide variety of singlechannel conductances (10–1000 pS) were observed, consistent with a peptide aggregated into multiple oligomeric states. SAA channels were permeable to most physiologic ions including Na+ , K+ , Ca2+ and Cl− , exhibiting only a weak preference for cations over anions. Channel formation could be inhibited by preincubation of SAA with Congo red, but addition of Congo red after channel formation had no effect. Channels could be reversibly blocked by 100 µM Zn2+ . The naturally occurring acute phase isoform SAA1 was reported to lyse bacterial cells when expressed in Escherichia coli, whereas expression of the constitutive isoform SAA4 did not. SAA1 and SAA4 differ in their sequences at approximately 50% of residues. SAA1 has a greater concentration of hydrophobic residues in the N-terminal region. The resemblance of these results to the properties of channel forming toxins such as colicins [74], yeast killer toxins [39], defensins [40] and protegrins [77] led to the proposal that SAA might play a role in host defense


against microbes. Electron microscopic analysis revealed that murine SAA 2.2 can exist as an annular hexamer with a central “pore-like” region [78]. Although membranes were not present, the observed pore diameter of 25 Å was consistent with the physiologic findings of Hirakura et al. [35].

13 AS

Parkinson’s disease is a progressive neurodegenerative disorder characterized by tremor, rigidity and brachykinesia. The hallmark lesion of Parkinson’s disease is the Lewy body, an inclusion body in dopaminergic neurons consisting largely of non-amyloid component (NAC; residues 66–95 of AS). Incorrectly named, NAC is actually a fibril-forming amyloid peptide. Mutations in AS are associated with familial Parkinson’s disease and implicate AS in the pathophysiology of the illness [7]. NAC is also found in Alzheimer’s disease amyloid deposits, which suggests a link between the two amyloid diseases. This notion is supported by the clinical overlap in these illnesses. Alzheimer’s disease patients are commonly found to have motor abnormalities. Parkinson’s disease patients frequently have cognitive problems, such as dementia and depression. Intermediate syndromes such as dementia with Lewy bodies also implicate AS in damage to neurons outside the dopaminergic system [65]. It has been reported that AS can increase the permeability of liposomes in a graded manner to substances of increasing size. This “sieving” action is characteristic of “pore-like” transport systems [81]. Pathogenic mutations in AS such as A30P and A53T have been reported to accelerate the formation of oligomers (protofibrils) capable of permeabilizing activity [81]. Electron microscopy revealed that AS formed annular, pore-like oligomers and that the Parkinson’s diseaserelated mutations enhanced the formation of these structures [18]. The pathogenic “arctic” mutation of Aβ also showed a similar enhancing effect on these annular structures [54]. Electrophysiologic studies of NAC have confirmed the formation of ion-permeable channels in lipid bilayers [5]. These channels have properties strikingly similar to those of other amyloidogenic peptides. Single-channel conductances are heterogeneous. Ionic selectivity is weak. Channels are irreversible and have extended lifetimes. Channel formation is inhibited by Congo red and channels are blocked by Zn2+ .

14 β2M

β 2 M forms amyloid deposits in bones and joints of patients on hemodialysis or peritoneal dialysis, a syndrome referred to as “dialysis-associated amyloidosis”. This 99-residue peptide belongs to the MHC class I complex which is involved in the presentation of foreign antigens to lymphocytes. β 2 M levels can rise 100-fold

16 PG

during states of renal failure [19]. Renal transplantation can result in lower β 2 M levels and clinical symptoms. (β 2 M’s physiologic effects include the induction of Ca2+ , efflux from calvariae, bone resorption and increasing collagenase production [9, 72, 78]. β 2 M is mainly found in amyloid deposits as fulllength native protein. In contrast to other amyloid protein “misfolding” diseases, the misfolding here appears to be solely a function of increased protein concentration in the serum, rather than mutation or proteolysis. Channel formation by β 2 M was reported by Hirakura and Kagan [41]. A multiplicity of single channel conductances were observed, ranging from 0.5 to 120 pS, with 90 pS being the most commonly observed size in 0.1 M KCl. Channel lifetimes were typically extended and ionic selectivity was poor. β 2 M associated irreversibly with the membrane. Incubation of β 2 M with Congo red inhibited formation of channels. Zn2+ could reversibly block inserted channels. Open β 2 M channels exhibited a slight degree of rectification with trans positive currents being somewhat larger than trans negative currents. Channel formation could be accelerated by acidic pH, compatible with the idea that the acidotic/uremic state of renal failure could enhance the generation of pathologic oligomers of β 2 M.

15 AL Amyloidosis

AL (light chain) amyloidosis is characterized by fibrillar deposits of the variable domain of immunoglobulin light chains. Fibril assembly is dependent on environmental conditions, e.g. the process may be different on surfaces versus in solution [36]. AL deposits are frequently found on surfaces such as arterial walls and basement membranes. A recent study of AL aggregation using atomic force microscopy found that AL protein of the variable domain SMA could form annular aggregates similar to those seen with AS (see Section 13). The SMA annular aggregates were significantly larger, however. Acidic pH was critical to the formation of these aggregates. It was suggested that these annular species might form pores in membranes [89].

16 PG

Although triplet-repeat diseases are not classic “amyloidoses”, their pathology seems to involve protein misfolding and the accumulation of toxic protein aggregates. Huntington’s disease is the most common and best known of these hereditary illnesses, which are caused by an expansion of the codon CAG which codes for glutamine. In Huntington’s disease, PG tracts larger than 37 residues cause disease, although this number varies amongst the different triplet-repeat illnesses. Several different proteins are associated with the various triplet-repeat diseases, but all are affected by an extended PG tract with a minimum threshold length for causing

15


clinical symptoms. In Huntington’s disease, the best-studied triplet-repeat disease, the presence of amyloid-like neuronal aggregates of huntingtin, the PG expanded protein, correlates with disease progression in transgenic mice [56]. The toxicity of huntingtin is proportional to the PG tract repeat length. Age of onset of illness is inversely proportional to PG repeat length. However, a PG repeat length = 12 cell line shows vulnerability to apoptosis without visible aggregates of huntingtin (PG repeat length = 150). Thus, visible deposit or aggregates may not be necessary to cause dysfunction or cell death. Indeed, some transgenic mice show electrophysiologic abnormalities in striatum before aggregates are visible [14]. Channel formation by PG was reported almost simultaneously by Hirakura et al. [34] and Monoi et al. [73]. The former group reported channels that were long lived, non-selective and heterogeneous in single channel conductance size, ranging from 19 to 220 pS in 0.1 M KCl. Channel formation was increased by acidic pH. Unlike classic amyloid peptides, Congo red pre-incubation did not inhibit channel activity nor was Zn2+ able to block PG channels. These distinct properties indicate that PG aggregates are clearly different structurally from classic amyloids. Monoi et al. [73] reported that PG40 could form cation-selective, long-lived channels with a single-channel conductance of 17 pS. PG29 could not form channels, consistent with the 37-residue cut-off for clinical illness. The investigators also proposed a structural model, the µ-helix, for PG channels, a model which just spans the bilayer hydrophobic core at a length of 37 residues, again in agreement with the clinical data. Further evidence of possible channel formation by PG in mitochondria was reported by Panov et al. [76], who showed that, in Huntington’s disease, mitochondria exhibited decreased membrane potential and were depolarized at lower Ca2+ levels than control mitochondria. Mitochondria from brain of transgenic mice expressing huntingtin with a pathogenic PG tract exhibited a similar dysfunction. Electron microscopy revealed mutant huntingtin localized to mitochondrial membranes. Most strikingly, the mitochondrial defects could be reproduced by a fusion protein with a long PG repeat. These results suggest that PG tracts are toxic to mitochondria and likely act via a channel-forming mechanism, depolarizing mitochondria and leaving them more vulnerable to apoptosis or metabolic insults. These data are consistent with a report that Aβ peptide can directly induce cytochrome c release in isolated mouse brain mitochondria, by directly inducing a permeability increase in the mitochondrial membrane [46].

17 HypF

The recent studies of non-disease related amyloid proteins has led to a reexamination of the nature of amyloid structure with respect to disease. It has been reported that a wide variety of non-disease proteins can form amyloid fibrils under the “appropriate” conditions [94]. It is a curiosity that the aggregation process of at least one such non-disease-related amyloid protein, HypF, can lead to formation

19 Lysozyme

of oligomeric structures and permeabilization of lipid membranes, without forming amyloid fibrils [83]. These oligomers are cytotoxic as well. These data strongly suggest that it is the oligomer β-sheet conformation itself that leads to membrane insertion, channel formation, cellular dysfunction and, eventually, cytotoxicity. It has been proposed that these are latent properties of all polypeptide chains (See chapter 3). 18 Calcitonin (CT)

Human calcitonin is (hCT) is a 32-residue peptide hormone involved in the regulation of calcium and phosphorous metabolism. It is produced by the C cells of the thyroid gland and is found in fibrillar amyloid deposits in patients with medullary carcinoma of the thyroid. The calcitonin peptide and fragments as small as four residues can form amyloid fibrils when incubated in aqueous solution. Calcitonin has pharmacological use in humans as a treatment for Paget’s disease and osteoporosis. The tendency of hCT to form fibrils limits its utility and has favored the use of salmon calcitonin (sCT), which is less fibrillogenic. A peptide corresponding to residues 16–21 of hCT form antiparallel β-sheets in methanol water and peptides consisting of residues 15–19 are highly amyloidogenic [82]. Fibril formation is accompanied by a conformational charge from α-helix to β-sheet in this central region or a change from random coil to β-sheet in the C-terminal region [27]. CT from several species, including human, salmon, eel and porcine, can form ionpermeable channels in lipid bilayer membranes [96]. These channels were long lived, voltage dependent, non-selective between cations and anions, permeable to Ca2+ , and enhanced by the presence of negatively charged phospholipids. It is possible that the channel-forming activity of hCT could have other physiologic effects on signal transduction, particularly in light of its ability to allow Ca2+ across membranes. Further investigation is needed to investigate the potential physiological role of these channels. 19 Lysozyme

Lysozyme is an enzyme that exhibits antimicrobial activity by cleaving bonds in the outer wall of Gram-positive bacteria. Lysozyme also forms toxic amyloid deposits in human. Mutations or partial protein unfolding leads to aggregation, oligomer formation and fibrilization of lysozyme. Partial unfolding of lysozyme also leads to the development of a non-enzymatic, broad-spectrum antimicrobial activity and a membrane-permeabilizing activity [36]. These activities were localized to a helix–loop–helix peptide at the upper lip of the active site cleft (87–114 of hen lysozyme). Similar peptides from human and chicken lysozyme possessed these activities as well. These results suggest that the unfolding of lysozyme leads to fundamental shifts in protein function and activity. The aggregation of lysozyme

17


monomers into oligomers appears to create a membrane-penetrating, antimicrobial complex which can aggregate into amyloid rings as seen by imaging [62].

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1

Protein Folding and Aggregation in the Expanded Polyglutamine Repeat Diseases Ronald Wetzel University of Tennessee, Knoxville, USA

Originally published in: Protein Folding Handbook. Part II. Edited by Johannes Buchner and Thomas Kiefhaber. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30784-2

1 Introduction

There are at least 15 human diseases associated with the genetic expansion of a chromosomal trinucleotide repeat sequence [1–3]. Some of these occur in noncoding regions, while others are located in open reading frames coding for a variety of proteins. Most of the diseases associated with expressed protein repeats involve CAG repeats coding for polyglutamine (polyGln). The common feature of DNA that underlies the genetic nature of these diseases is the genetic instability of triplet repeats of some of the 64 possible DNA nucleotide triplets [1]. Currently there are nine known diseases involving expansion of a polyGln sequence [1]. While these diseases on average vary one from the other in details of brain pathology and symptoms [1], there are also significant similarities. The polyGln repeat length threshold separating the normal population from at-risk individuals is strikingly similar for eight of the nine diseases [1]. All nine diseases are progressive and uniformly lethal and tend to present in midlife [3]. Of particular significance to this chapter, polyGln aggregates have been demonstrated in each of these diseases [3]. The demonstration of polyGln-containing inclusions in patient material [4] and in animal and cellular models [5–7] also suggested their resemblance to other neurodegenerative diseases [8] involving protein deposition into amyloid or other aggregated deposits. As a subject for studying protein aggregation diseases at the molecular level, however, the polyGln diseases have pluses and minuses. On the one hand, the uniform monotony of glutamine that constitutes the pathogenic core of the disease proteins does not inspire immediate insights into disease


2 Protein Folding and Aggregation in the Expanded Polyglutamine Repeat Diseases

mechanisms or protein-folding mechanisms. Indeed, standard site-directed mutagenesis approaches are technically difficult at the DNA level, due to the lack of unique internal restriction sites [9], and in any case may be of questionable value at the protein level, due to the likely ability of the sequence to shift and slide while aggregating so as to accommodate, and negate the effect of, potential modifying mutations [10]. On the other hand, there is something attractive about the simplicity of a homopolymer as a subject for addressing fundamental problems in protein folding and aggregation; for example, studying a real-world homopolymer should provide data more easily related to theoretical protein-folding studies that are often based on highly simplified model structures. A significant additional attraction to studying these diseases is the prospect of contributing to the development of therapies. As summarized below, protein-folding and aggregation studies of these sequences in vitro are making significant contributions to a better understanding of the molecular basis of these diseases and along the way are providing unique systems for better characterizing the role of protein structure in ordered aggregation processes.

2 Key Features of the Polyglutamine Diseases

The literature on the clinical, genetic, cellular, and biochemical aspects of the expanded polyGln diseases has grown enormously since the discovery of the disease genes over the past decade. There are excellent books reviewing the various aspects of each disease [11, 12], and a steady stream of review articles and essays discuss possible disease mechanisms from various points of view [2, 13–31]. These reviews reflect the lively, ongoing debate on the importance of polyGln oligomers and aggregates in the disease mechanism. The discussion here is very brief and necessarily centered on the toxicity of polyGln peptides, especially the possibility that polyGln aggregates play a toxic role. 2.1 The Variety of Expanded PolyGln Diseases

There are currently nine human diseases linked to a genetic expansion in an in-frame, translated CAG repeat [3], of which the most recently identified is spinocerebellar ataxia 17, involving expansions in the transcription factor TBP, or TATA-binding protein [32]. Table 1 lists the known diseases, the disease protein involved, its molecular weight and subcellular localization, and the CAG repeat lengths for the normal and pathological ranges. The most well known and most common of these diseases is Huntington’s disease (HD). There are a number of genetic ataxias whose disease gene has not yet been identified [33, 34], and at least some of these may eventually be found to also involve CAG repeat expansion.

2 Key Features of the Polyglutamine Diseases Table 1 Overview of expanded CAG repeat diseasesa

Disease

Gene product

Cellular localization

Normal CAG Mutant CAG MW (kDa) repeat range repeat range

Huntington’s DRPLAb SBMAc SCA1d SCA2d SCA3d /MJDe SCA6d SCA7d SCA17d

huntingtin atrophin-1 androgen rec. ataxin-1 ataxin-2 ataxin-3 CACNA1Af ataxin-7 TBPg

Cytoplasm/nucleus Cytoplasm/nucleus Cytoplasm/nucleus Cytoplasm/nucleus Cytoplasm Cytoplasm/nucleus Cytoplasm Cytoplasm/nucleus Nucleus

348 190 104 87 145 46 280 95 38

6–39 3–35 9–33 6–44 13–33 3–40 4–19 4–35 24–44

36–>200 49–88 38–65 39–83 32–>200 54–89 20–33 37–306 46–63

a

Data from Ref. [12]. Dentatorubral-pallidoluysian atrophy c Spinal and bulbar muscular atrophy d Spinocerebellar ataxia e Machado-Joseph Disease f α 1A voltage-dependent calcium channel g TATA box-binding protein b

2.2 Clinical Features

Except for the X-linked spinal and bulbar muscular atrophy (SBMA), involving CAG repeat expansions in the gene for the androgen receptor, the other eight known polyGln diseases exhibit autosomal dominant genetics [3]. Huntington’s disease normally presents as a motor disorder, although a significant minority of patients present with psychiatric symptoms [12]. The expanded CAG repeat ataxias involve a wide range of symptoms and significant variability [3]. Early-onset forms tend to present with distinct, more-aggressive symptoms that set them apart from the later-onset forms. The large variations in the course of disease for patients with the same CAG repeat length suggest important roles for secondary factors, including environment and modifying genes, and these are currently under investigation in those diseases with statistically significant amounts of patient data. 2.2.1 Repeat Expansions and Repeat Length DNA triplet expansions leading to genetic disease can occur in expressed open reading frames or in non-coding regions [1]. Of the possible triplet repeat sequences, only a few are known to be involved in triplet expansion genetic disorders [1]. Although it is presumed that there are unique structural aspects to the DNA of those triplets whose instability during one or more enzymatic process in the cell leads to expansion-based diseases, the structural basis of repeat instability is unclear and continues to be evaluated [35]. The stability of the CAG repeat, in particular, appears to depend on a number of factors, including the length of the repeat before expansion, its sequence context, and whether the donor of that gene

3


is the mother or the father [3, 36]. The normal role of benign repeat lengths of the polyGln sequence in the various proteins in which it is found remains a mystery. In fact, since homologues of the same protein often vary considerably in their polyGln repeat length, while other parts of the protein may show very little variation, it is possible that polyGln in most cases is not functionally important. The hypothesis that normal-length polyGln sequences serve no important function that might be compromised upon sequence expansion is consistent with the widely accepted view that expanded CAG repeat diseases are predominantly gain-of-function disorders [3]. Table 1 shows the clear separation of benign repeat lengths from pathological repeat lengths in the expanded CAG repeat diseases. In all cases but one (SCA 6, which uniquely involves a resident transmembrane channel protein), the pathological cutoff lies in the mid- to upper-30s. In HD, within this critical region, repeat length also influences penetrance, the percentage of individuals with the mutation that eventually develop disease symptoms [37]. Within the pathological repeat length range, age of onset correlates strongly with repeat length, as shown for HD in Figure 1. One of the clinical characteristics of the disease, recognized prior to the discovery of the disease genes, is that of anticipation, in which a parent presents with a disease only after it is already recognized in a child. With the discovery of the dynamic genetic nature of the triplet repeat diseases, anticipation is now understood by the fact that repeat expansions, especially when passed on by the father, can be large, and large repeat expansions tend to dramatically decrease age of onset (Figure 1) [1]. 2.3 The Role of PolyGln and PolyGln Aggregates

Although a few genetic ataxias involve CAG expansions in non-coding regions [33, 34], the bulk of the characterized disorders are associated with expression of the polyGln sequence at the protein level. Loss of function may play a role in some mutations in the gene associated with SCA 6 [3]. Further, a decline in normal levels of active protein, perhaps mediated by aggregation, may also contribute to symptoms in some expanded CAG repeat diseases [38, 39]. On the whole, however, the dominant effect of polyGln expansion appears to be a gain of toxic function. It seems very unlikely that the expanded polyGln disorders, with their often overlapping clinical features and very similar genetics, do not share intimate details of their molecular mechanisms, and it is further unlikely that the molecular mechanisms involving these otherwise unrelated protein sequences are not centered in the behavior of the polyGln sequence. This is further supported by the generation of animal models based on expression of artificial proteins containing in-frame CAG repeats spliced to proteins not known to be associated with disease [5, 40]. Opinion in the field has been divided as to whether expanded polyGln is cytotoxic due to the induction, within the soluble monomer, of a toxic conformation or, alternatively, to the accelerated aggregation of the monomer into a toxic, oligomeric state. Although the ability of a series of antibodies to bind preferentially to expanded

2 Key Features of the Polyglutamine Diseases

Fig. 1 HD CAG repeat length is correlated with the age at onset of neurologic symptoms. The relationship between the expanded HD CAG repeat length (“Number of CAG Repeat Units”) and the age at neurologic onset is given for 1070 HD patients reported in the literature. The mean age at onset for any given HD CAG repeat length is depicted by filled symbol. Power

regression analysis reveals a significant inverse correlation between expanded CAG repeat size and age at onset (r = −0.87, P < 0:0001), although individuals with the same expanded repeat exhibit widely different ages at onset, suggesting the existence of modifiers of the disease process. Figure and analysis courtesy of Marcy MacDonald.

polyGln has been interpreted as evidence for the existence of a novel conformation within the monomer, recent studies suggest that the increased avidity observed for such antibodies can be explained by the multiplicity of independent, oligoGlnbinding sites in the expanded sequence (a linear lattice effect) [41]. Most other data is consistent with no significant change in the conformation of bulk-phase polyGln upon its sequence expansion. In contrast, substantial evidence points to a central role for polyGln aggregation in the disease process [25, 27–29], beginning with the observation of intraneuronal, nuclear inclusions (NII) in cell and animal models of polyGln disease [5–7]. Similar aggregates are found in brain tissue from HD and other diseases listed in Table 1, but not in normal, aged control brain [3, 4]. Genetically identified suppressers of polyGln toxicity, such as PQE1 [42] and dHDJ1 [40], have been shown to be aggregation suppressors in vitro (see Section 8). Further, designed polypeptide-based inhibitors of polyGln aggregation also protect cells from the

5


toxic effects of expanded polyGln sequences [43] and polyGln aggregates [44] (see Section 8.1). As discussed in detail below, the repeat length–dependent aggregation properties of polyGln closely mirror the repeat-length dependence of disease risk shown in Table 1. Also as discussed below, polyGln aggregates made in vitro and delivered to the nuclei of cells in culture are very cytotoxic [45]. Although the appearance of visible aggregates does not always closely correlate with cytotoxicity in cell and animal models [2, 17, 21], there are reasonable explanations for how such observations might still be consistent with cytotoxic aggregates. Some of these rationales will be alluded to in this chapter.

3 PolyGln Peptides in Studies of the Molecular Basis of Expanded Polyglutamine Diseases

If the effect of sequence expansion on the physicochemical properties of the polyGln sequence plays a central role in the expanded CAG repeat diseases, then it is of particular importance to study polyGln behavior with respect to repeat length in well-defined biophysical experiments. Consists of an overview of studies using either chemically synthesized peptides or recombinant proteins to conduct studies on the conformational preferences of soluble polyGln and on the aggregation efficiency of these sequences. The poor solution behavior of the polyGln sequence has provided substantial barriers to conducting such studies, and much has been learned simply in the effort to produce molecules amenable to analysis. 3.1 Conformational Studies

Initial studies on the favored conformation of the polyGln sequence gave conflicting results. Working with chemically synthesized polyGln sequences well below the pathological threshold repeat length, Altschuler et al. found the sequence to exist in random coil [46], while Sharma et al. found peptides of similar length to be dominated by β-sheet [47]. This discrepancy was resolved with the later realization of the importance of applying rigorous methods of peptide disaggregation as part of the solubilization protocol [48]. Using such protocols, Chen et al. showed that polyGln sequences in solution in PBS exhibit the signature CD spectrum of a statistical coil [49]. More importantly, they also showed that the CD spectra of polyGln peptides both below and above the pathological threshold repeat length give essentially identical CD spectra [49], providing the first experimental indication that a pure conformational change within a monomeric polyGln protein is not likely to underlie the repeat-length dependence of clinical features. Similar results were later obtained with various polyGln repeat lengths within the context of E. coli-expressed fusion proteins containing either naked polyGln sequences [50] or polyGln sequences embedded in a fragment of huntingtin [41]. Importantly, these

3 PolyGln Peptides in Studies of the Molecular Basis of Expanded Polyglutamine Diseases

studies also show that results obtained with synthetic peptides, such as those described here, are not compromised by the disaggregation protocol. The above discussion notwithstanding, the CD spectrum generally assigned to the random coil state is not unambiguous. In fact, a recent study has confirmed earlier suggestions that polypeptides exhibiting the random coil CD spectrum might actually exist in the ordered conformation known as polyproline type II helix [51]. Indeed, recent studies suggest that the amino acid glutamine is particularly comfortable in this form of secondary structure [52]. While the exact nature of the conformation of monomeric polyGln may ultimately be of significant importance to understanding disease mechanisms, the main important lesson to date from solution studies is that there appears to be no difference between the solution structures of polyGln sequences below and above the pathological repeat length range. Absent some more subtle conformational difference between short and long polyGln, some other aspect of the polyGln sequence must underlie the repeatlength dependence of disease risk. 3.2 Preliminary in vitro Aggregation Studies

With the revelation that expanded polyGln sequences are responsible for the diseases listed in Table 1, and in light of the importance of protein aggregation in other neurodegenerative diseases such as Alzheimer’s disease, Perutz speculated that polyGln expansion might lead to an increased propensity of proteins containing these sequences to aggregate and thereby cause disease [53]. Working with a Q15 sequence flanked by pairs of charged residues, the Perutz lab reported facile aggregation of the peptide in aqueous solution to yield aggregates exhibiting the cross-β structure characteristic of amyloid [53]. In analogy to the leucine zippers designed by nature to promiscuously self-associate in the creation of dimeric transcription factors, Perutz coined the term “polar zipper” for the tendency of polyGln sequences to aggregate, presenting a model in which hydrogen bonding by side chain amide groups contributes to the stability of the aggregate [53]. Scherzinger et al. were able to overcome many of the limitations encountered in the above preliminary studies by using a recombinantly expressed form of glutathione S-transferase (GST) fused with the exon 1 fragment of huntingtin containing the polyGln sequence [54, 55]. Fusion of GST with htt exon 1 renders the protein reasonably well behaved so that it can be extracted and partially purified while maintaining it in a soluble form. By the ingenious use of an installed protease site, Scherzinger et al. were able to cleave this soluble protein between its GST and htt components in a controlled reaction, releasing the polyGln portion in situ so that its aggregation could be monitored in a synchronized reaction [54]. By doing so, and by following the generation of aggregate using a filter trap assay, they were able to observe a clear increase in the aggressiveness of spontaneous aggregation as polyGln repeat length increased, with a significant increase in aggregation rate as repeat length increased through the pathological cutoff range [54, 55]. This correspondence between the repeat length threshold for disease risk and the threshold

7


for rapid aggregation continues to be one of the most striking features of polyGln behavior in solution, and it has been invoked as supporting evidence for hypotheses involving aggregation in the disease mechanism. 3.3 In Vivo Aggregation Studies

In addition to contributing to our understanding of the disease mechanism, cell and animal models involving recombinant expression of polyGln-containing disease proteins have contributed greatly to our knowledge of the aggregation process itself. The dependence of aggregation efficiency on polyGln repeat length was characterized qualitatively in a number of in vivo experiments comparing a limited variety of repeat length constructs [6, 7, 54, 56]. At the same time, it is also possible for non-pathological repeat length polyGlns to form aggregates under certain conditions in the cell [57], emphasizing that even sequences below the pathological threshold have some potential to aggregate. Transgene experiments have implicated a modulating role for flanking sequences and/or flanking domains in the aggregation of expanded polyGln, consistent with the possibility that proteolytic processing may play an important role in controlling aggregation and toxicity in the cell. For example, mice expressing exon 1–like fragments of htt tend to develop aggregates (and pathology) more rapidly and aggressively than do mice expressing full-length htt [26]. Besides the mammalian systems mentioned here, a number of other organisms, such as yeast [58], D. melanogaster and C. elegans [59], have been used to learn details about how polyglutamine aggregates in the cellular environment. The promiscuity of polyGln aggregation was noted early on in the observation that an aggregate initiated by an expanded polyGln sequence is capable of recruiting the normal-length polyGln protein into the deposit [57]. More recently, recruitment of the normal allele of htt by expanded polyGln htt aggregates has also been observed, suggesting that expanded CAG repeat diseases may include an overlay of loss of function in the disease mechanism [39]. The possible role of generalized polyGln aggregation in the cell was also suggested by the observation that expression of expanded polyGln containing a nuclear localization signal (NLS) could mediate nuclear localization of green fluorescent protein (GFP) linked to a normal-length polyGln [60]. One of the intriguing possible mechanisms of polyGln cytotoxicity is that cellular polyGln aggregates might deplete the soluble cellular pools of other polyGln-containing factors by recruiting them into the aggregate in an ongoing elongation reaction [57, 61, 62]. Such mechanisms are supported by the observation of co-localization of TBP [57, 61] and CBP (CREB-binding protein) [60, 63] to cellular aggregates in tissue from patients and/or from cell and animal models.

4 Analyzing Polyglutamine Behavior With Synthetic Peptides: Practical Aspects

Given the limited success of previous studies of polyGln behavior using synthetic peptides and the comparative success of studies using recombinant proteins in vitro


and in vivo, one might legitimately question the value of putting further efforts into the chemical approach. There are a number of reasons for doing so, however. While polypeptides, even unnatural fusions, expressed in a cellular environment might be considered to be “more native” than chemically synthesized peptides exposed, even transiently, to organic solvents, the situation is complicated by the central role of the prior aggregation state of a protein in biasing its aggregation behavior in vitro. This is illustrated by early experiments in the Alzheimer’s disease field, in which studies of the amyloid peptide Aβ were confused by investigators’ ignorance of the aggregation state of the peptide and the importance it plays in cytotoxicity and aggregation. This led to an odd lore concerning certain manufacturers’ batches of synthetic peptide as being “good” (cytotoxic right out of the vial) or “bad” (nontoxic) batches. Eventually, both cytotoxicity [64] and aggressive aggregation ability [65] were clearly shown to be attributable to traces of aggregates resident in the vialed material. As shown below, methods now exist for rigorously removing preexisting aggregates from synthetic peptides. Such methods are chemically benign, allow complete removal of the organic solvent, and are compatible with peptides whose natural state as a soluble monomer in native buffer is relatively unstructured. In contrast, recombinant proteins are potentially much more complex and, once nonnative assembly occurs, the unnatural oligomeric state may be much more difficult to recognize and reverse. For example, expression of a GST-AT-3 fusion containing a Q27 sequence led to an enriched, soluble protein that, however, migrated as an oligomer on native gel filtration [66]; similar soluble oligomers can be observed in isolation of htt exon-1 fusion proteins (G. Thiagaragan and R. Wetzel, unpublished). Clearly, it is possible, at least with relatively short polyGln repeats, to prepare monomeric, folded proteins by recombinant expression [41], but a great level of care and analytical scrutiny is required to insure that the preparation is aggregate-free, and the challenges increase dramatically with increasing repeat length. Thus, there may be situations in which chemically synthesized peptides, exposed transiently to non-aqueous solvents to effect efficient disaggregation, are “more native” than a recombinant protein prepared and maintained in aqueous buffers. An additional attractive feature of focusing on chemically synthesized polyGln peptides is that optical studies are not complicated by the presence of other segments of polypeptide. For example, this allows relatively clean interpretation of the CD spectra of subject polypeptides without having to be concerned about the contributions of other domains and how those other domains may or may not also be changing during the experiment. Work with synthetic peptides also allows facile use of chemical tags such as biotin [67] and fluorescein [45], as well as non-natural amino acids such as D-amino acids [10]. Synthetic peptides often make it more possible to conduct studies requiring relatively large amounts of material and to accomplish the studies in a shorter time. Finally, while it is clear that the most appropriate studies, if they can be done on pristine protein samples, are those that are chemically and physically most similar to the toxic fragments generated in vivo, it is also true that studies on simple polyGln peptides will allow baselines to be established for their biophysical behavior, so that further studies on the sequence in the natural settings of the disease proteins

9


can be placed into context and the specific effects of sequence context be cleanly appreciated. 4.1 Disaggregation of Synthetic Polyglutamine Peptides

In the review of the aggregation literature in Section 3.2, one troubling inconsistency stands out. Work with recombinant protein fragments yields results on the repeat-length dependence of aggregation that mirror the repeat-length dependence of disease risk in HD and other expanded CAG diseases, with sequences up to about Q35 being relatively resistant to aggregation. In contrast, however, previous work with chemically synthesized peptides established an apparent solubility ceiling of Q15 –Q22 , above which it appeared that peptides could not be studied in solution [46, 47, 53]. This appeared to be so even when charged flanking residues were included in an attempt to improve peptide solubility. However, we now understand that this apparent discrepancy between the transient solubilities of recombinant and chemically synthesized peptides is due to the presence, in the latter, of residual polyGln aggregates, leading to a situation not unlike that described above for Aβ peptides. This is dramatically demonstrated by two examples. Although attempts to directly dissolve a K2 Q44 K2 sequence in aqueous buffer failed to give significant soluble peptide [48], pretreatment of a K2 Q44 K2 peptide (or a K2 Q20 K2 peptide) with a trifluoroacetic acid/hexafluoroisopropanol disaggregation protocol adapted from the Aβ literature [68] yields peptides that display excellent initial solubility in aqueous buffers [48]. Even more dramatic is the effect this protocol has on the peptide K2 Q15 K2 . This peptide readily dissolves directly from the synthetic lyophilized powder in low salt, pH 3 buffer to yield a transparent, low light-scattering solution. When this solution is adjusted to phosphate-buffered saline conditions, however, aggregation occurs very rapidly to form a protofibril-like product [48]. In contrast, when this same peptide is exposed to the TFA/HFIP disaggregation protocol prior to solubilization in water and then is adjusted to PBS conditions, the peptide remains in solution for months at 37 ◦ C, finally reaching an equilibrium position of aggregation after about six months [48]. Although the composition of K2 Q15 K2 directly dissolved in water has not been further investigated, it is most likely that the suspended peptide exists as a mixture of dissolved monomer and microaggregates, the latter of which seed aggregation by the former when the solution is adjusted to PBS conditions. This tediously slow aggregation kinetics of the K2 Q15 K2 peptide after disaggregation now qualitatively matches the behavior of short polyGln sequences in recombinant proteins and is also consistent with the lack of disease risk in humans with short polyGln sequences in htt and other disease proteins. This change in the character of the K2 Q15 K2 peptide is not a consequence of any covalent chemical change in the peptide [48], and no significant amounts of organic solvent remain after the treatment. To ensure that all traces of aggregates are removed, a preparative ultracentrifuge step is included. The sequential treatment of the lyophilized, synthetic peptide with TFA, followed by HFIP, as described


for treatment of Aβ peptides [68], is adequate for disaggregating polyGln peptides up to repeat length 40. Above this, the synthetic peptides do not dissolve well in TFA. However, it was found that peptides of higher repeat length dissolve well in a 1:1 mixture of TFA and HFIP and that, with sufficient incubation time and care, they can be completely disaggregated in this solvent mixture [48]. This treatment also appears to dissolve other aggregation-prone peptides that are resistant to the sequential method. 4.2 Growing and Manipulating Aggregates

Although many studies on disease mechanism in HD and other protein aggregation diseases involve analysis of aggregation behavior, the study of the aggregation products is also important, since it is likely that it is the aggregates themselves, or a continuing aggregation process supported by these aggregates, that are responsible for cytotoxicity. In analogy to the above discussion on working with the peptides, therefore, a few words must also be said about the growth and handling of aggregates. Simple polyGln peptides incubated at 37 ◦ C grow into a variety of morphologically related structures, as reviewed in Section 6.1. In general, these aggregates appear to be variations on the assembly of narrow protofilaments, in analogy to the substructures of amyloid fibrils. The aggregate formation reaction is quite reproducible, so long as the peptides are processed as described above and growth conditions are held constant. The aggregates are quite stable, with critical concentrations well below micromolar for polyGln peptides of repeat length 30 or more [49]. Aggregate suspensions in PBS can be snap-frozen in liquid nitrogen and stored at −80 ◦ C for extended periods without noticeable change in properties. 4.2.1 Polyglutamine Aggregation by Freeze Concentration Simple polyGln peptides dissolved in PBS, when incubated for one day or more in the frozen state at −5 to −20 ◦ C, are highly aggregated upon thawing [66]. This is true even for Q15 peptides, which, when incubated at 37 ◦ C in the completely disaggregated state, require months to achieve significant levels of aggregation. Studies on the mechanism of this effect showed that aggregate formation under these conditions is due to the process of freeze-concentration [69, 70]. Peptide solutions in buffer stored frozen at temperatures above the eutectic points of the buffer components consist of a solid ice phase and fissures of liquid phase containing most of the solutes, including the peptide [69, 70]. Since the liquid phase is only a small volume of the total, the solute concentrations in the liquid phase are enormous, with, for example, NaCl concentrations of several molar resulting from the freezing of a simple PBS ([NaCl] ∼ 0.15 M) solution. These conditions often result in the formation of protein aggregates, which is why it is often advisable to store protein solutions either above 0 ◦ C or below 0 ◦ C with freezing-point depression agents such as glycerol. Interestingly, because ice structure at modest freezing temperatures is rather unstable, solutions of polyGln peptides may be snap-frozen in liquid nitrogen, but if they are then stored at −20 ◦ C, they will still develop

11


Fig. 2 Time course of freeze-concentration aggregation monitored by light scattering. Samples of 10 µM disaggregated K2 Q37 K2 in PBS were incubated at 37 ◦ C (solid black bars), at −10 ◦ C in the liquid state (open

bars), or at −10 ◦ C in the frozen state (gray bars). The atter sample was snap-frozen in liquid nitrogen then transferred to the −10 ◦ C bath. From Ref [66] with permission from the American Chemical Society.

aggregates as the ice structure equilibrates to that normally formed at the −20 ◦ C storage temperature. Figure 2 shows the time course of aggregate formation in the ice state versus liquid state in the −5 to −10 ◦ C range [66]. The freeze-concentration-induced aggregation of polyGln has several implications. First, there is no reason to suspect that any polyGln sequence, even in the context of other protein sequences, will not be susceptible to this effect. We have found that polyGly peptides can be stored for months at −80 ◦ C if they are first disaggregated, adjusted to low concentrations, and snap-frozen in liquid nitrogen before storage. It is recommended that all polyGln-containing proteins be stored with similar care or not frozen at all. Interestingly, we found that the Aβ(1–40) peptide does not form amyloid by freeze-concentration. Thus, the same level of care required for working with polyGln sequences may not be required when working with other amyloid systems. It is important to be aware, however, of the potential for the seemingly benign freezing process to wreck havoc with samples of any protein, especially aggregation-prone proteins. 4.2.2 Preparing Small Aggregates In addition to this potential for undesired aggregation, the freeze-concentrationinduced aggregation of polyGln sequences is important because it generates aggregates with somewhat different properties compared to those grown from the same peptide at 37 ◦ C [66]. As discussed in Section 6, these aggregates appear to be more like the protofibril assembly intermediates of other amyloid growth reactions than like the mature aggregates grown at 37 ◦ C. These aggregates, once formed, are stable and do not assemble further on incubation at 37 ◦ C. On a weight basis, aggregates prepared by freeze-concentration are much better seeds for fibril elongation than are aggregates grown at 37 ◦ C [49]. At the same time, 37 ◦ C aggregation reactions seeded (and hence greatly accelerated) with aggregates prepared

5 In vitro Studies of PolyGln Aggregation

by freeze-concentration produce aggregates indistinguishable from those grown by spontaneous aggregation at 37 ◦ C, indicating that the substructures of these aggregates must be quite similar. We have used freeze-concentration to produce aggregates of small sizes to facilitate their uptake into mammalian cells [45]. After aggregates are grown in frozen buffer, reactions are thawed and the aggregates collected by centrifugation. Aggregates are sonicated with a probe sonicator and then pushed through a series of molecular weight cutoff membrane filters to produce the finest possible aggregate size. This procedure is critical for the ability of mammalian cells to take up naked polyGln aggregates [45] (see Section 7.1.1).


The central importance of the expansion of the polyGln repeat in this family of neurodegenerative diseases logically leads to the study of the biophysical properties of various repeat lengths of polyGln in search of clues to molecular mechanisms. Coincidentally, focusing on the polyGln sequence in isolation also makes accessible certain experimental approaches not possible with more complex proteins. For example, circular dichroism (CD) spectra can be cleanly interpreted in terms of what the polyGln sequence is doing, without being concerned about sequences and other elements of secondary structure. Solution concentrations of non-aggregated peptide can be quantified with great precision and accuracy using reverse-phase HPLC, as described in the experimental Section. Insolubility can be cleanly interpreted in terms of polyGln aggregation, without worrying about the aggregation tendency of other protein sequences in unnatural fusion proteins or truncated native proteins. The data from studies on isolated polyGln sequences establish a baseline for the fundamental behavioral trends of the polyGln sequence, so that further studies with sequences more closely resembling the complex structures of disease proteins can be put into context. 5.1 The Universe of Protein Aggregation Mechanisms

The broad phenomenon of particulate aggregation is systematized in the field of colloidal assembly and coagulation. The aggregation of proteins, especially in response to heat and as a side reaction in protein folding, has been viewed historically as a problem in colloidal assembly [71]. Early generalized models of the kinetics of coagulation involved models of particles undergoing hierarchical assembly into larger and larger clusters. Cluster formation in these models was considered to be controlled either by simple diffusion limits or by an energy barrier that determined the productiveness of each collision, but the assumption in either case was that each productive collision was irreversible [72]. However, since many protein aggregation reactions such as crystallization and amyloid fibril formation reach

13


equilibrium positions in which both aggregates and monomers are populated, the possibility of productive collisions being reversible must be considered, which complicates the modeling [72]. Protein crystallization and amyloid formation may differ from classical colloidal assembly in another respect; rather than a hierarchical assembly mechanism involving the interactions of intermediates of ever-increasing size, protein crystals and amyloids, once formed, might be capable of growing by the addition of monomeric or oligomeric protein units [72]. This could occur, for example, if the protein in the aggregated state projects a docking surface that is capable of recruiting bulk-phase monomer very efficiently, compared to the relatively inefficient process of nucleation. Such a nucleation-dependent polymerization model has been developed [73, 74] and successfully applied to hemoglobin aggregation [75]. The model also takes into account the potential reversibility of steps in the aggregation process and thus treats the nucleation process as an unfavorable pre-equilibrium before a series of increasingly favorable elongation steps; that is, the kinetic nucleus is viewed as the least stable species on the aggregation pathway. The unfavorability of the nucleation step gives rise to a “lag phase,” during which no mature aggregates are formed and negligible amounts of monomer are consumed, followed by a rapid growth phase. Since this is a relatively simple model that has been successfully applied to protein systems, and since there is no evidence from in vitro studies of a more coagulationbased assembly mechanism for polyGln aggregation, this model has been applied to polyGln aggregation data, as described in the following section. 5.2 Basic Studies on Spontaneous Aggregation

Rigorously disaggregated polyglutamine peptides (Section 4.1) incubated in PBS at 37 ◦ C undergo a spontaneous aggregation process that generally exhibits a lag phase typical of nucleated growth polymerization. These lag times become shorter as the repeat length gets longer, with aggregation becoming much more aggressive at repeat lengths about 35 [49]. These results agree with previous observations based on studies of exon I fragments of huntingtin [55]. Following the kinetics by a number of different windows on the aggregation process – light scattering, ThT fluorescence, CD, and solubility – reveals overlapping growth curves (Figure 3), suggesting that there are no major hidden species in the process [76]. The fact that the CD transition, from a spectrum consistent with random coil to one consistent with β-sheet, coincident with the reaction as monitored by aggregate formation and peptide solubility suggests that there is no pre-assembly in the bulk-phase monomer to a β-sheet containing monomeric species before aggregation occurs [76]. This interpretation is reinforced by an experiment in which the aggregation reaction is stopped midway and centrifuged [76]. The resuspended centrifugation pellet gives a typical β-sheet CD spectrum, while the supernatant gives the same random coil signature spectrum as seen in the starting reaction mixture (Figure 4). The implication is that the mechanism by which the bulk-phase polyGln peptides take on β-sheet characteristics occurs either through a concerted mechanism, in


Fig. 3 Spontaneous aggregation of a polyGln peptide monitored by four independent measures. A sample of 66 µM freshly disaggregated K2 Q42 K2 in 10 mM Tris-TFA, pH 7.0, was incubated at 37 ◦ C and aggregation was monitored by circular dichroism

signal at 200 nm (), determining remaining soluble monomer by HPLC (•), light scattering (), and thioflavin T fluorescence (◦). From Ref. [76] with permission from the National Academy of Sciences (USA).

Fig. 4 Circular dichroism spectra of various fractions of a polyGln aggregation reaction midpoint. The reaction described in the legend of Figure 3 was sampled after 90-h incubation and CD spectra were collected after various treatments: no treat-

ment (a); ultracentrifugation supernatant (b); resuspended ultracentrifugation pellet (c); sum of the supernatant and pellet spectra (d). Adapted from Ref. [76] with permission from the National Academy of Sciences (USA).

15


which molecules take on β-structure as they become ensconced onto the end of the growing aggregate, or through a mechanism in which isolated monomers rapidly add to aggregates as soon as they acquire β-structure. 5.3 Nucleation Kinetics of PolyGln

The increasing efficiency with which peptides undergo spontaneous aggregation (see previous section) suggests that it may be this property of the polyGln sequence that is responsible for the critical importance of repeat length in the expanded CAG repeat diseases. The source of the repeat-length dependence of aggregation is not immediately obvious, however. To investigate whether the repeat-length effect is played out at the level of nucleation and to learn more about the nucleation process, we applied a successful model for the nucleation-dependent aggregation of sickle hemoglobin [74] to the polyGln aggregation process [76]. In this model, the formation of an oligomeric kinetic nucleus from a bulk phase of monomeric proteins is treated as a pre-equilibrium, the favorability of which is dependent on the equilibrium constant and the number of monomers that cluster together to form the nucleus. This latter number, which is of particular importance because it becomes an exponential factor in the equation describing nucleation kinetics, is called the “critical nucleus,” n* . Whether any particular nucleus collapses back to n* monomers, or progresses on to committed aggregate growth, depends on the efficiency with which it is stabilized through the elongation of the nucleus into a growing aggregate, as determined by the rate constant describing this elongation step. Since the number of kinetic nuclei formed during a nucleation-dependent polymerization reaction is vanishingly small and cannot be directly observed, the concentration of nuclei present at any one time is inferred by the kinetics with which observable aggregates develop, as described by the rate constant for elongation of aggregates. The overall nucleation kinetics equation (Eq. (1)) emerging from this model describes , the increase in the concentration of monomers that have been converted into aggregates at time t, as it depends on the concentration of monomers in the bulk phase, c, the nucleation (pre-)equilibrium constant K n *, and the rate constants for elongation of the nucleus and aggregate, k+ , which, for simplicity, are assumed to be identical. 2 =1/2k+ K n* c (n*+2) t 2

(1)

Eq. (1) accounts for the time lag observed for nucleation-dependent growth kinetics and for the concentration dependence of the lag phase and the underlying nucleation kinetics. It also predicts two other features of nucleation kinetics: (1) that the kinetics should be dependent on time2 , and (2) that a log-log plot of the term emerging from the t2 plot, with respect to monomer concentration, will yield a line of slope n* + 2, from which n* , the number of monomers in the kinetic nucleus, can be derived. In fact, analysis of polyGln peptides with repeat lengths below and above the pathological threshold region shows linear t2 plots at all peptide


Fig. 5 Critical nucleus analysis for polyGln peptides of various lengths. Nucleation kinetics for each peptide at several concentrations was determined as described in the text and analyzed by t2 plots. The log of the slope of the t2 plot is plotted against the log of peptide concentration. The slope

of the straight line fitting this data is n* + 2, where n* is the critical nucleus. Shown are data for K2 Q28 K2 (), K2 Q36 K2 (•), and K2 Q47 K2 (). Also shown are theoretical lines showing slopes of 3 (◦) and 4 (). From Ref. [76] with permission from the National Academy of Sciences (USA).

concentrations examined [76]. Figure 5 shows that plotting the log of the slopes of the t2 plots at each concentration, against the log of the concentration, gives a straight line for each repeat length. The equivalence of the slopes for peptides above, at, and below the pathological threshold, as shown in Figure 5, shows that, within this range, repeat length does not alter the size of the critical nucleus. Table 2 shows the surprising result that the kinetics analyses for these peptides yield slopes of 3, reducing, therefore, to n* = 1 in each case. Table 2 also shows that the source of the more aggressive nucleation kinetics, as polyGln repeat length increases, is therefore the larger value of the k+ 2 Kn *. term, rather than a change in the value of n*. Since it was previously shown that elongation rates do not change dramatically as polyGln repeat lengths change [49], it can be concluded that the major source of the increased aggressiveness of polyGln aggregation above the

Table 2 Kinetic parameters for polyGln nucleation of aggregationa

Slope of the log-log plot Calculated critical nucleus (n*) from slope k+ 2 K n* (M−2 s−2 ) a

Data from Ref. [76].

Q28

Q36

Q47

2.98 0.98

2.68 0.68

2.59 0.59

0.001128

0.09193

1.8304

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Fig. 6 A model for the nucleationdependent polymerization of polyGln peptides. Reversible formation of the monomeric nucleus, the least stable species on the aggregation pathway, is visualized as an unfavorable folding reaction. Elongation is visualized as a cyclic two-step process

involving addition of bulk-phase polyGln monomer to the nucleus or growing aggregate, followed by consolidation of the structure of the newly added monomer to extend the structure of the aggregate. From Ref. [76] with permission from the National Academy of Sciences (USA).

pathological threshold is an increase in the nucleation (pre-)equilibrium constant Kn* . The result of this analysis, i.e., that the kinetic nucleus for nucleation-dependent polymerization of polyGln sequences contains a single molecule of polyGln, is a surprising, if not shocking, result. The whole theoretical basis for nucleation in nucleation-dependent polymerization and colloidal assembly theories is a model in which the nucleus is itself an oligomer on the assembly pathway. Based on general colloidal assembly models [72] and on water condensation analysis [77], the stability derived from the growth of the nucleus is basically attributed to an improved surface-to-volume quotient for this particulate cluster [78]. The recognition, however, that monomeric polyGln peptides, even long ones, assume a non-β-sheet conformation in solution makes conceptually possible a model in which the nucleation event is an unfavorable folding reaction within the monomer population. Figure 6 illustrates this model schematically, showing bulk-phase monomer as random coil and the nucleus as a monomer collapsed into an antiparallel β-sheet conformation. Interestingly, despite the fact that the model for the nucleation process was developed based on the assumption of an oligomeric critical nucleus, the mathematical analysis growing out of this model is sufficiently robust that it also describes quite well the situation in which n* = 1. The reason for this is that the most fundamental definition of the critical nucleus is more mathematical than physical, stating only that the nucleus is the least stable species on the assembly pathway. The critical nucleus for polyGln assembly into amyloid-like aggregates achieves this distinction by being the product of a highly unfavorable folding reaction. How unfavorable these reactions are in the polyGln series is presently unknown, since the k+ 2 K n* . term cannot be deconvoluted due to (1) a present inability to independently determine the value of k+ and (2) the inability to independently observe the nucleation equilibrium without its leading to aggregation. Direct observation of the nucleation process, determination of K n *, and derivation of molar elongation rate constants remain challenges for future research. The ability of the nucleation kinetics model to accommodate polyGln aggregation made possible the calculated estimation of the expected kinetics curves for


Fig. 7 Calculated nucleation kinetics curves for 100-pM polyGln peptides in PBS at 37 ◦ C, based on kinetic parameters derived from in vitro studies at higher concentrations. From Ref. [76] with permission from the National Academy of Sciences (USA).

nucleation-dependent polymerization of polyGln at relatively low steady-state concentrations, i.e., of magnitudes similar to those expected in vivo. Figure 7 shows the resulting curves for assumed polyGln concentrations of 100 pM, which show predicted ages of onset for the three polyGln repeat lengths studied that differ by orders of magnitude and in a way that is consistent with disease statistics (Figure 1). The actual steady-state concentrations of the aggregation-prone proteolytic fragments of huntingtin or other expanded CAG repeat disease proteins are not known, and it is clear that the molecular complexity of cells and organisms is likely to foil attempts at adequately describing disease through test tube experiments in simple buffers alone. However, the simulation in Figure 7 illustrates how nucleation propensity, as controlled simply by repeat-length variation, is capable of contributing significantly to disease risk and age of onset. Ongoing studies of simple peptides in defined buffers are also providing quantitative descriptions of the roles of molecular crowding, molecular chaperones, downstream amino acid sequences, and other polyGln-containing molecules on impacting aggregation nucleation (A. Thakur, A. Bhattacharyya, G. Thiagaragan, and R. Wetzel, unpublished results). 5.4 Elongation Kinetics

The model for polyGln aggregate growth shown in Figure 6 shows that the nucleus elongates by successive productive encounters with bulk-phase monomer. Although other models have been proposed for fibril elongation that may be followed by some proteins under some solution conditions [79], growth by monomer additions is likely to be the major pathway by which amyloid fibrils grow in vivo, especially in most systems where precursor protein concentrations are quite low. The

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kinetics of this process have been successfully modeled [80] as a simple bimolecular reaction between aggregate and monomer, such that the observed rate depends on monomer concentration, aggregate (molar) concentration, and the second-order rate constant. However, since in a heavily seeded reaction, aggregation mass builds primarily by the lengthening of the seeds, the molar concentration of aggregate (or the molar concentration of growth sites) does not change appreciably over the course of a simple elongation reaction, and the rate expression reduces to a pseudofirst-order kinetics equation involving monomer concentration and the first-order rate constant. (This rate constant is not a universal constant for this peptide, however, since it incorporates the molar concentration of fibrils used as seeds, which is not trivial to determine.) Understanding more about the polyGln aggregate elongation process is important not only because of how this process contributes to the deposition of the disease protein in neurons but also because of the possible importance of the recruitment of other polyGln-containing proteins in the disease mechanism (Section 3.3). If the polyGln aggregate, or the aggregation process itself, is responsible for the neurotoxicity observed in these diseases, it is also important to learn how we might interfere with this process therapeutically. The need to discover and characterize aggregation inhibitors adds to the importance of having available one or more ways of quantifying elongation in vitro. 5.4.1 Microtiter Plate Assay for Elongation Kinetics One convenient way of studying the elongation reaction is by immobilizing preformed aggregate on a surface, incubating the surface in a solution of monomer, and monitoring the growth of the aggregate seed. Aβ elongation has been studied by this approach in two ways, by using either fibrils fixed to microtiter plate wells and following elongation with radio-labeled monomer [81] or fibrils fixed to a dextran surface and following elongation by an increase in fibril mass as detected by surface plasmon resonance [82]. A modification of the microplate approach has been used to develop a polyGln elongation assay, which has been described in detail [67, 83]. In this assay, very small masses (typically 20–100 ng per well) of polyGln aggregates are immobilized onto microtiter plate wells. An aqueous solution of a disaggregated polyGln, with a biotin covalently attached to the N-terminus, is supplied at low concentration (typically 10 nM) and incubated. After incubation, excess biotinylated peptide is washed out and the amount of biotin remaining on the solid phase by virtue of a continuation of the polyGln aggregation process is measured with a streptavidin-linked reagent. High sensitivity is achieved through using streptavidin complexed with europium, which in turn can be measured with great sensitivity using time-resolved fluorescence. Since the microplate must be processed intact, kinetics are accomplished by phasing the start times of multiple reactions so that their common termination will deliver a time course. A typical kinetics progress curve for this polyGln elongation reaction is pictured in Figure 8. The division of the kinetics into fast and slow phases, as seen in Figure 8, has also been observed in the Aβ microplate assay [84]. As predicted by the kinetic model described above, the magnitude of the kinetic phases increases both with


Fig. 8 Kinetics of polyGln aggregate elongation monitored by a microtiter plate– based assay. Twenty nanograms of K2 Q28 K2 aggregates were immobilized by adsorption to each microtiter plate well, and

the wells were incubated for various times with 10 nM biotinyl-K2 Q28 K2 . Signal was developed using europium-tagged streptavidin and time-resolved fluorescence. From Ref. [67] with permission from Elsevier.

the increase of the amount of deposited polyGln aggregate and with an increase in monomer concentration. This assay is proving useful in working out some fundamental aspects of the polyGln elongation reaction [49], in screening for inhibitors of polyGln aggregate elongation (see following section), and in characterizing HD brain material. 5.4.2 Repeat-length and Aggregate-size Dependence of Elongation Rates The repeat-length dependence of spontaneous polyGln aggregation is grounded in the repeat-length dependence of the unfavorable folding of the monomeric kinetic nucleus (Section 5.3) and is the simplest available explanation for how the gross features of the repeat-length effect on disease risk and age of onset are established. The repeat-length effect of the elongation phase of aggregate growth is also important, however. If the ability of polyGln aggregates to cause cell death and/or dysfunction is due to their ability to recruit other polyGln-containing proteins into inactivating aggregates, then the dependence of the efficiency of recruitment on polyGln repeat length will determine the range of polyGln-containing proteins that are at risk of being recruited. Experiments using the microtiter plate elongation assay show that there is, indeed, a length dependence of recruitment kinetics, but it is not nearly as dramatic as that for nucleation-dependent aggregation [49]. PolyGln peptides in the 5–10 repeat length range are not strongly recruited into preexisting aggregates.

21


At repeat lengths in the Q15 to Q20 range, however, elongation rates become more substantial and continue to increase with increasing repeat length up to about Q40 , above which there appears to be little effect on rate of increasing repeat length. This suggests that polyGln proteins with repeat lengths in the 5–10 range are not likely to impact polyGln aggregation processes very much. However, at repeats of Q15 and above, the existence of polyGln proteins, and their relative concentrations and functions in the cell, may well impact expanded CAG repeat diseases. These results are consistent with observations that neuronal polyGln aggregates often contain CREB-binding protein (which has a repeat length of 18 Gln residues), but that the homologous protein P300, which has a polyGln repeat of only 6, is not recruited into cellular aggregates [85]. The rate of aggregate elongation, as determined by polyGln repeat length and concentration, may also significantly contribute to other aspects of polyGln aggregation and pathology (A. M. Bhattacharyya and R. Wetzel, manuscript in preparation). Size also matters in considering the mass-normalized recruitment activity of polyGln aggregates. Microtiter plate experiments show that, on an equal mass basis, the ability of a preformed aggregate to serve as a template for polyGln elongation improves as the average particle size of the aggregate decreases [49]. The data support the idea that not all polyGln aggregates are functionally equivalent and that small aggregates are substantially better than the larger ones at supporting elongation. If polyGln aggregates are, in fact, toxic by virtue of their ability to recruit polyGln-containing proteins and thus remove them from their normal sites of action, it stands to reason that more recruitment-active aggregates would be more toxic. That is, aggregates too small to be easily detected by light/fluorescence microscopy may be far more cytotoxic than the relatively recruitment-inert aggregates that can be so visualized. These conclusions are further supported by staining cellular aggregates for recruitment activity, which will be discussed in Section 6.2.

6 The Structure of PolyGln Aggregates

Like other amyloid-like fibrils, the interior, atomic-level structure of the polyGln aggregate remains opaque, due to its inaccessibility to standard high-resolution methods and the limited resolution of the available methods. The following sections catalog what we do know about polyGln aggregate structure, with an emphasis on the extent to which these aggregates resemble other amyloids. 6.1 Electron Microscopy Analysis

Perutz and coworkers first showed that the polyGln aggregate has a fibrillar construction. The aggregates obtained from a Q15 peptide immediately upon adjusting a pH 3 solution of the synthetic product to pH 7 [53] resemble in the electron microscope the protofibrils observed as assembly intermediates in amyloid


formation by other peptides, such as Aβ [86]. Similar structures are observed from Q15 or Q20 peptides after they are rigorously disaggregated and then incubated in PBS (Figure 9). In addition, aggregates of longer polyGln sequences take on this same protofibril-like appearance when they are grown in PBS at −20 ◦ C by freeze-concentration (Section 4.1) (Figure 9). It is possible that aggregation stops at this stage at −20 ◦ C because the diffusion of the protofibrils is limited by the ice lattice. The generation of protofibril-like structures from simple polyGln peptides in vitro has also been accomplished through the use of nonnative buffer conditions [87]; the relationship of these aggregates to those formed, for example, by freeze-concentration has not been explored. When simple polyGln peptides longer than Q20 are incubated at 37 ◦ C in PBS, they grow into more complex aggregated structures that appear in the EM as ribbons of about 20 nm in diameter and with substructures suggesting assembly from parallel-aligned protofibrils or protofilaments (Figure 9). Attempts to detect protofibril intermediates in these assembly reactions have not been successful, however (unpublished results), suggesting that under native conditions in simple PBS buffers protofibrils either do not exist, or have a very short lifetime. The observation of polyGln-associated protofibril-like structures in cells [88] may be due to the presence of flanking protein sequences and/or other molecules in the milieu altering the assembly pathway. When longer polyGln peptides (Q45 or higher) are incubated in PBS at 37 ◦ C, especially in low salt buffer, they grow into long filamentous aggregates that match the structures of typical amyloid fibrils (Figure 9). Incubation in vitro of fusion proteins containing huntingtin exon 1 fragments with expanded polyGln repeats also generates amyloid fibrils [54], and some EM studies suggest a fibrillar appearance of the substructures of polyGln inclusions in cells [4]. Thus, there appears to be a hierarchy of form in the world of polyGln aggregates, all of which exhibit a similar protofilament substructure that is capable of assembling in higher-ordered structures depending on polyGln repeat length, protein and cellular context, and growth conditions. 6.2 Analysis with Amyloid Dyes Thioflavin T and Congo Red

The early history of amyloids is associated with the discovery of the ability of certain heteroaromatic dyes to bind selectively to fibrils and, through binding, take on altered spectroscopic properties that can be monitored. In 1927, Divry reported that amyloid deposits in tissue, previously detected by a starch-like reaction to iodide staining, could be easily detected by staining the tissue with the dye Congo red (CR) followed by polarized light microscopy [89]. The success of the method, which remains the primary pathological stain for amyloid [90], depends not only on the ability of CR to bind to amyloid fibrils but also on the ability of the fibrils to confer onto the bound CR a macroscopic order that can be detected as birefringence in light microscopy. Since the resolution of light microscopy is limited by the wavelength range of visible light, birefringence is observed only if the ordered

23


Fig. 9 Electron micrographs of various protein aggregates. (A) D2 Q15 K2 aggregates grown in PBS at 37 ◦ C. (B) K2 Q20 K2 aggregates grown in PBS at 37 ◦ C. (C) K2 Q37 K2 aggregates grown in PBS at 37 ◦ C. (D) K2 Q37 K2 aggregates grown in PBS in the frozen state at −20 ◦ C. (E) Fluorescein-

tagged K2 Q42 K2 aggregates grown in PBS at 37 ◦ C. (F) K2 Q42 K2 aggregates grown in Tris-HCl, pH 7.2 at 37 ◦ C. (G) A$(1–40) fibrils grown in PBS at 37 ◦ C. The bar represents 50 nm. From Ref. [66] with permission from the American Chemical Society.

array of CR molecules that produces it extends into that size range. Since amyloid fibril diameters are in the 10 nm range, it is clear that for fibrils to be detected by CR birefringence, the fibrils have to be arranged in a higher level of order, that is, oriented non-randomly in the field. In fact, EM of fibrils often shows them to be loosely bundled together in an approximate parallel arrangement. This type of order is required in tissue for amyloid to be detected by CR birefringence. Perhaps this explains the lack of success in detecting polyGln aggregates in tissue


by amyloid reagents. CR staining of polyGln aggregates grown in vitro produces a red cast, suggesting that the aggregates bind CR, but does not yield significant birefringence [66], suggesting that a lack of macroscopic order, rather than an absence of the fundamental amyloid fold, may be the explanation for the lack of signal. CR binding to amyloid in vitro can be detected by virtue of a spectral shift conferred onto CR upon binding [91], but there appear to be no data on whether polyGln aggregates produce this effect. Another important dye for amyloid analysis is thioflavin T (ThT). ThT has the ability to bind to many different amyloid fibrils [92, 93], and when it does, the binding produces a characteristic change in ThT’s fluorescent properties. The fluorescence yield upon ThT binding is linear with the number of ThT molecules bound, and therefore (under saturating conditions) also with the mass of amyloid, which allows the dye to be used to quantify amyloid in vitro [93]. PolyGln aggregates produce a typical amyloid-like ThT response [49]. PolyGln aggregation kinetics monitored by ThT track exactly, within experimental error, with other measures of aggregate formation (Figure 3). The molecular mechanisms by which CR and ThT bind to polyGln and other amyloids are not known. CR, as a sulfonate, carries a strong negative charge, while ThT carries a positive charge, at neutral pH. The charged nature of these dyes has led to speculation that electrostatics play an important role in their binding to amyloids, most of which are derived from sequences possessing charged amino acids. PolyGln sequences carry only the neutral glutamine side chain, however, casting doubt on any overriding role for electrostatics in the binding of ThT. Although the synthetic polyGln molecules summarized here contain flanking Lys residues for peptide solubility, their aggregates, displaying only positively charged Lys side chains, are still capable of binding the positively charged ThT. 6.3 Circular Dichroism Analysis

As discussed in Section 5.2, the CD spectra of aggregated polyGln peptides are consistent with high levels of β-sheet (Figure 4), and the progress of the CD signal change at 200 nm that marks the progress of the two-state transition from soluble peptide to aggregate overlaps the aggregation kinetic curves determined by other means (Figure 3). The ease of obtaining an interpretable CD spectrum of the polyGln aggregate is somewhat unusual in the amyloid field, where attempts to collect CD spectra of amyloid fibrils in suspension are marred by high levels of light scattering. That polyGln aggregates are better behaved in CD must be due to lower light scattering, presumably due to smaller average particle sizes. In fact, while Rayleigh light scattering in a fluorometer is a very sensitive means of following amyloid fibril formation by Aβ (B. O’Nuallain and R. Wetzel, unpublished observations), equivalent weight concentrations of polyGln aggregates scatter light much less well, although still to a useful extent [48, 76]. When polyGln aggregation reactions are carried beyond the point when the ThT reaction has reached plateau and the solution-phase peptide is essentially depleted, further, relatively small,

25


changes occur in the CD spectra (S. Chen and R. Wetzel, unpublished). Similar changes have been seen in other β-sheet proteins, where they have been interpreted as reflecting increasing degrees of twist in the sheet [94]. 6.4 Presence of a Generic Amyloid Epitope in PolyGln Aggregates

Recently, a class of antibody has been described that has the unusual, unanticipated ability to recognize a conformational epitope that seems to be present to some extent on all amyloid fibrils while not being present in the native state of the monomer [95, 96]. For example, the mouse MAb WO1 was generated by challenging mice with an Aβ(1-40) amyloid fibril immunogen, but the resulting antibody binds not only to Aβ fibrils but also to amyloid fibrils derived from IAPP, β2-microglobulin, transthyretin, immunoglobulin light chain, etc. [96]. This antibody also binds to the amyloid-like aggregates of synthetic polyGln peptides [96], offering another indication of the relatedness of the polyGln aggregate to the generic amyloid motif [66]. Conversely, some MAbs generated from mice immunized with polyGln aggregates are able to bind well to other amyloid fibrils such as Aβ fibrils (B. O’Nuallain, R. Wetzel, S. Ou, J. Ko, P. Patterson, unpublished), further indicating the presence of an amyloid epitope on the polyGln aggregate. Such pan-amyloid antibodies tend to be IgMs, limiting their use as tissue-staining reagents. These antibodies have a number of other uses, however, as diagnostics for the amyloid motif and as a means of quantifying certain features of fibrils [10]. 6.5 Proline Mutagenesis to Dissect the Polyglutamine Fold Within the Aggregate

One of the central questions in amyloid research is how the constituent polypeptide folds upon itself to engage the amyloid structural motif. Amyloid fibrils clearly possess a cross-β structure, in which the peptide chains are perpendicular to the fibril axis, while the H-bonds between strands in the β-sheets are parallel to the axis [97]. The details of how the peptide chains are arranged is space within these simple, minimal restraints have not, however, been established for any amyloid. Mutagenesis, especially with proline residues [98], has been used in other amyloid systems to garner details about how the sequence of the amyloidogenic peptide folds within the fibril. In Aβ, for example, it is clear from a variety of techniques that the N-terminal 10–15 amino acids, as well as the C-terminal 3–4 amino acids, are not involved in tight β-sheet H-bonding [99]. Proline analysis, which is based on the observation that loops, floppy ends, and some turn positions can comfortably accept Pro residues, while β-sheets are destabilized by them, also suggests the location of turns in the Aβ fibril [99]. A major problem confronts the use of mutagenesis to dissect polyGln structure in the aggregate, however. The problem is the prospect that the polyGln sequence will be promiscuous in how it adopts its fold within the aggregate. This means that


any particular sequence position of a Q40 sequence, let’s say position 10, might be found in a number of environments when a series of Q40 molecules folds into the aggregate. Further, should position 10 be favored in a certain kind of structure in the aggregate, a mutation at position 10 that is unfavorable in that type of structure will likely simply lead to adjustments in the polyGln chain as it engages the aggregate structure, to place residue 10 in the least damaging – that is, least destabilizing – place in the aggregate. A modified mutagenesis strategy, taking the above complication into account, has led to some insights into polyGln structure within the aggregate. It is based on the hypothesis that polyGln folds in the aggregate in a series of extended chain segments, of some unknown optimal length, alternating with turn segments [10]. This hypothesis was then tested by analysis of the aggregation kinetics of a series of mutant polyGln peptides composed of alternating segments of oligoGln and Pro-Gly sequences, the latter being especially comfortable in turn elements. It was found that the peptide PGQ9 , consisting of four Q9 segments interspersed with three PG pairs, undergoes spontaneous aggregation essentially as fast as a Q42 peptide of the same length. Since peptides with shorter QN elements aggregate less rapidly, while a PGQ10 peptide aggregates only marginally more rapidly, it was concluded that the best model for how an unbroken polyGln sequence aggregates is PGQ9 . Since each turn may involve one or two Gln residues bordering the PG pair, this corresponds to extended chain segments of 7–8 residues interspersed with turns of 3–4 residues. Two further mutagenesis tests on PGQ9 support such models. To test the interpretation that the PG pairs are located in turns in the aggregate, a PGQ9 peptide was synthesized containing D-prolines in place of L-prolines; this peptide aggregates more rapidly than the L-Pro form, consistent with data showing D-Pro-Gly to be more compatible in β-turns than L-Pro-Gly [100]. To test the interpretation that the oligoGln sequences between the PG pairs are in extended chain in the aggregate, mutant peptides were synthesized in which additional proline residues were inserted in the middle of one or more of these oligoGln sequences. When a single Pro residue is placed in the middle of either the second (PGQ9 P2 ) or third (PGQ9 P3 ) of the four Q9 segments, these peptides are completely incapable of forming aggregates under conditions where PGQ9 aggregates readily. This replicates the strong inhibition of amyloid formation when a single Pro residue is placed in the middle of a β-sheet region in Aβ peptides [98, 99] and strongly supports the model for the aggregate structure of PGQ9 . That the structure of the PGQ9 aggregate is very similar to that of a Q42 aggregate was shown by a number of studies, including EM, WO1 binding (Section 6.4) and the abilities of each aggregate to seed elongation of the other peptide [10]. Interestingly, the aggregation of another Q42 analogue, containing only a single proline residue at the same residue position as the additional proline in PGQ9 P2 , is nearly as rapid as that of an unbroken Q42 peptide. This confirms the concerns about folding promiscuity in polyGln aggregation that guided the design of the PGQ9 series. That is, simple mutagenesis experiments on unbroken polyGln will generally be very difficult to interpret with confidence, because of the ability of the

27


polyGln sequence to accommodate to the mutation by slightly altering its fold in the aggregate. The model of a repeat structure consisting of alternating extended chains and turns is compatible with either an antiparallel β-sheet motif or an irregular parallel β-helix motif of the kind seen in certain globular proteins [101]. Previously, Starikov et al. proposed an antiparallel β-sheet model, noting that the number of residues required to make a four-stranded version of such a model is very close to the pathological threshold for most expanded CAG repeat diseases [102]. Perutz and coworkers proposed a “water-filled nanotube” model for the polyGln aggregate [103], which is related to parallel β-helix models previously proposed for the amyloid structure [101, 104]. It is not clear whether the central core of water would be energetically favorable or whether a more stable variation might be to exclude the water to generate an irregular helix more like those seen in parallel β-helical proteins [101, 105]. However, the “ideal” water-filled nanotube model [103] might be expected to be destabilized by Pro or Pro-Gly insertions, given the uniform extended chain and continuous twisted β-sheet in the model. The ability of polyGln aggregates to tolerate certain periodicities of Pro-Gly insertion is also incompatible with aggregates consisting of very long strands of polyGln extended chain [27]. Interestingly, the mechanism of nucleation of polyGln aggregation does not seem to be altered in these Pro-Gly mutants. Thus, analysis of the concentration dependence of the early phases of spontaneous aggregation for the peptide PGQ9 yields a value of n* = 1 for the critical nucleus [10], the same number obtained for unbroken polyGln sequences.

7 Polyglutamine Aggregates and Cytotoxicity

There is little disagreement that the expansion of the polyGln sequence plays a major role in expanded CAG repeat diseases, but opinions differ considerably as to the mechanisms by which this effect plays out. Many reviews are available that summarize the data and argue for and against an active involvement of polyGln aggregates [2, 13–31]. One of the central barriers to reaching a consensus on the nature of the polyGln effect is the difficulty in characterizing and/or controlling the aggregation state of polyGln in cells. This problem has an impact in various ways. For example, in order to produce htt aggregates in a reasonable time frame in a cell or animal model, it is necessary to express unnaturally high concentrations of monomeric protein in the cell; if toxicity is observed, is it then due to the aggregates or to the high concentration of monomer required to make the aggregate? At the same time, genetic analyses for binding partners of expanded polyGln proteins may in some cases be mediated by an aggregated form of the bait molecule rather than the assumed monomeric form. Semantics and sensitivity of detection may also play roles in the controversy over the toxic species in polyGln diseases. To a protein chemist, an aggregate is any oligomeric protein state held together by noncovalent interactions, even


including native oligomers; thus, a nonnative dimer of huntingtin, for example, would be considered an aggregate. Some workers, however, interpret an absence of large, macroscopic cellular inclusions as definitively ruling out the involvement of aggregates in toxicity. If the neuronal dysfunction and death characteristics of expanded CAG repeat diseases can in fact be attributed to polyGln aggregates, it remains to be determined how aggregates produce their effects at the molecular and cellular level. A number of theories have been put forward to explain the toxicity of protein aggregates in neurodegeneration. Aggregates might saturate and/or otherwise inactivate the cellular machinery, such as molecular chaperones, the ubiquitin-proteasome system [106], and the aggresome system [107], responsible for dealing with protein misfolding. Aggregates might insert into membranes and alter their properties [108]. One mechanism that is uniquely feasible for polyGln aggregation and that has received significant attention and confirmatory data is the recruitment-sequestration model [24, 57, 61, 62, 85]. Since it is well established that amyloid like aggregates of one polyGln sequence are capable of being elongated by the addition of other polyGln sequences, it is possible that polyGln aggregates in the cell might recruit other polyGln-containing proteins. Many such proteins exist in the human genome [109–111], in particular among proteins involved in nucleic acid binding, such as transcription factors. In a pivotal paper, Ross and coworkers showed that expanded polyGln cytotoxicity in a cell model appears to be mediated by the recruitment of the transcription factor CREB-binding protein (CBP), which contains a Q18 repeat, into the polyGln aggregates [85]. Expression of high levels of a functional, polyGln-minus version of CBP protects cells from expanded polyGln toxicity [85]. As discussed below, synthetic polyGln peptides have contributed to the debate about the toxicity of expanded polyGln sequences by addressing some of these technical barriers. 7.1 Direct Cytotoxicity of PolyGln Aggregates

One way to more directly test the toxicity of aggregates would be to deliver polyGln aggregates produced in vitro into cells. This would eliminate the need for producing unnaturally high levels of monomeric polyGln, the toxicity of which then becomes an important question if the experimental result is to be properly interpreted. The ability to produce a variety of well-characterized polyGln aggregates in relative bulk, as described above, makes possible experiments involving delivery of aggregates to cells. In addition, the ability to deliver aggregates essentially composed of only the polyGln sequence addresses the question of the role of other sequence elements in cytotoxicity of expanded CAG repeat proteins. 7.1.1 Delivery of Aggregates into Cells and Cellular Compartments The challenge in analyzing the cytotoxicity of exogenous aggregates is their delivery into the cell. In fact, liposome encapsulation does allow polyGln peptide aggregates to be taken up by cells in a relatively short time [45]. Interestingly, however, a control

29


experiment of polyGln aggregate mixed directly with cells in culture also resulted in very good cellular uptake of the aggregate [45]. Sonication and membrane filtration (Section 4.2.2) to generate smaller average sizes of aggregates improves cellular uptake [45]. Uptake through the cellular membrane was confirmed by confocal microscopy on fluorescently tagged aggregates. The mechanism by which cells take up these aggregates is not clear, but it may be related to an electrostatic attraction between the negatively charged phospholipid membrane and the highly positively charged aggregates, which are composed of peptides consisting of polyGln flanked by pairs of Lys residues. The process does not require polyGln, since a control amyloid fibril is also readily taken up [45]. Since nuclear polyGln inclusions are often observed in cell and animal models, it is also of interest to deliver aggregates into the nucleus. In an attempt to do this, polyGln peptides were prepared that contain an N-terminal nuclear localization signal (NLS) peptide sequence as well as a fluorescein tag. In fact, aggregates of such peptides are not only taken up by the cell but also are transported into the nucleus, as confirmed by confocal microscopy of isolated nuclei [45]. A control amyloid fibril from a peptide containing a NLS is also readily taken up into the nucleus. The ability of the nuclear transporter to manage such aggregates is somewhat surprising, since the aggregates appear to exceed 100 nm in diameter (based on their ability to resist filtration through a 100-nm filter membrane), while the diameter of the nuclear pore is on the order of 26 nm [112]. It is possible that the multiple display of NLS signal peptides along the surface of the aggregate provides a repeat that allows it to “ratchet” through the pore [112]. It is also possible that the large aggregates mixed with and taken up by cells may transiently break apart to individual protofibrillar aggregates, as seen in EMs of these aggregates; these have diameters in the range of 5 nm. It is very unlikely that aggregates get into cells or nuclei by way of full dissociation to monomers followed by reassembly on the other side of the membrane. These polyGln aggregates are very stable in vitro in physiological conditions [45]. Further, such a mechanism could not explain the ability of Q20 aggregates to invade the cell and nucleus, since the very slow aggregation kinetics of the monomer, especially at the concentrations likely to obtain in this scenario, rules out any efficient aggregation after dissociation. 7.1.2 Cell Killing by Nuclear-targeted PolyGln Aggregates The ability to deliver various protein aggregates into cells and their nuclei makes it possible to study the toxicity of these aggregates. PolyGln, as well as control, aggregates delivered to the cytoplasm of either PC12 or Cos-7 cells in culture exhibit little or no toxicity to these cells when examined by a number of standard methods for determining cell death: propidium iodide exclusion, lactate dehydrogenase release, and MTT reduction [45]. In contrast, polyGln aggregates delivered to cell nuclei rapidly kill cells, with very similar results from all three measures of cell killing. Although amyloid fibrils of cold shock protein peptide B-1 (CspB1) are efficiently delivered to cell nuclei, they do not kill cells, indicating the importance


of the polyGln sequence. Interestingly, nuclear-targeted aggregates of Q20 and Q42 peptides are equally effective at killing cells. This strongly suggests that any aggregated polyGln peptide can be toxic when delivered to the nucleus and reinforces the conclusion derived from the aggregation kinetics (Section 5.2) that the distinction between a toxic (Q42 ) and benign (Q20 ) repeat-length sequence is made at the level of aggregation efficiency (Section 5). Although the ability of nuclear-localized polyGln aggregates to rapidly kill cells may thus be responsible for the neuronal loss observed in end-stage expanded CAG repeat brain stem, it remains possible that cytoplasmic polyGln aggregates might also have harmful effects that may be less dramatic and difficult to detect in a one-day cell culture. The cell death induced in cultured cells by nuclear uptake of polyGln aggregates appears to be related to an apoptotic process (W. Yang and R. Wetzel, manuscript in preparation). 7.2 Visualization of Functional, Recruitment-positive Aggregation Foci

One possible explanation for why easily detected polyGln aggregates do not correlate with cytotoxicity in all cases is that the wrong aggregates are being monitored. If the cell killing by expanded polyGln sequences is due to recruitment of cellular polyGln proteins by aggregates of the polyGln disease protein, then the ability of these aggregates to efficiently recruit other polyGln sequences – their “recruitment activity” – is of paramount importance to their cytotoxicity. Recruitment activity can be defined as a measure of the number of monomeric polyGln peptides that can be recruited into a polyGln aggregate of a certain mass in a certain time. Interestingly, even in vitro aggregates of simple polyGln peptides can exhibit different recruitment activities depending on the way they are prepared and on their sizes [49]. Larger aggregates exhibit lower recruitment activity than smaller aggregates, presumably because of the greater surface area of the latter [49] (Section 5.4). Thus, if the recruitment hypothesis is valid, then the best way to gauge the polyGln aggregate burden of a cell or tissue would be to assess the recruitment activity in the tissue, either as a tissue stain or as an activity measurement akin to an enzymatic assay. Both of these approaches are feasible, because of the robustness and specificity of the polyGln elongation reaction, and both have been shown to work when using human brain material. Aggregates in an enriched fraction isolated from HD brain tissue, when fixed to microplate wells, exhibit the same ability to recruit biotinylated polyGln peptides as synthetic aggregates exhibit (V. Berthelier and R. Wetzel, unpublished). HD brain slices incubated with biotinylated polyGln pick up the peptide in small, punctate, intracellular centers called aggregation foci [113]. The ability to characterize cells and tissue for levels of functional polyGln aggregates should provide sharper tools for addressing the question of the role of polyGln aggregates in cell dystrophy and death in expanded CAG repeat diseases.

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8 Inhibitors of PolyGln Aggregation

If polyGln aggregates are toxic to cells, a viable therapeutic approach for expanded CAG repeat diseases would be the use of specific inhibitors of spontaneous polyGln aggregation and/or elongation. Such inhibitors clearly exist in nature. For example, several genetic screens in HD disease models reveal that increased expression of DnaJ class chaperones can protect cells and animals from polyGln toxicity. Investigations of the molecular basis of this effect reveal that recombinant HDJ1, a human DnaJ homologue, is very effective at inhibiting both spontaneous polyGln aggregation and seeded elongation, presumably by binding to nuclei or small aggregates and inhibiting their elongation with additional polyGln monomers (A. Bhattacharyya and R. Wetzel, unpublished). This suggests that inhibitors of polyGln aggregation can have a protective effect. 8.1 Designed Peptide Inhibitors

A similar result has been obtained with a structure-based elongation inhibitor. Not only is the peptide PGQ9 -P2 incapable of spontaneously aggregating (see Section 6.5), it can also inhibit the aggregation of other polyGln peptides. In the microplate assay described earlier, PGQ9 -P2 inhibits Q42 elongation with an IC50 in the low micromolar range (Figure 10) [44]. Furthermore, cells pre-incubated with

Fig. 10 Concentration-dependent inhibition of polyGln aggregate elongation by PGQ9 -based peptide inhibitors. Microtiter plate wells containing adsorbed Q45 polyGln aggregates (100 ng per well) were incubated with 10 nM biotinyl-Q28 and various concentrations of PGQ9 P2 (•) for 45 min at 37 ◦ C. The derived EC50 value is 1.1 µM.

8 Inhibitors of PolyGln Aggregation

PGQ9 -P2 are protected, in a dose-dependent manner, from the cytotoxicity of polyGln aggregates in the cell model described in Section 7.1.2. [44]. This implies not only that aggregates are toxic but also that the basis of their toxicity is their ability to recruit other polyGln peptides in the cell. This in turn suggests that even cells that have already entered the aggregation pathway might be protected from aggregate toxicity by appropriate inhibitors.

8.2 Screening for Inhibitors of PolyGln Elongation

As discussed above, polypeptide-based inhibitors of polyGln aggregation exist. However, small molecules with comparable activities would presumably be better drug candidates. In order to discover and refine such inhibitors, viable assays are needed. A filter blot assay that allows screening of small compound libraries for inhibitors of the aggregation of the polyGln-containing htt exon 1 domain has identified a number of small molecule inhibitors [114]. The microtiter plate elongation assay described in Section 5.4.1 was initially designed for the same purpose [67, 83]. The assay has several important features, including reproducibility, insensitivity to small concentrations of DMSO and small pH changes, good throughput, and the ability to operate at low polyGln concentrations approaching physiological. This assay has been used to screen a number of small libraries of drug-like compounds as well as to determine dose-response curves for hits and analogues (V. Berthelier and R. Wetzel, unpublished). A typical screening result is shown in Figure 11.

Fig. 11 Microtiter plate screen of a small compound library for inhibitors of biotinylpolyGln elongation on adsorbed polyGln aggregates. Amount of elongation for each compound was compared to elongation

without added compound, and the percent inhibition was calculated. In this plate, 1 out of 80 compounds tested gave >50% inhibition. Error bars from triplicate analysis.

33


9 Conclusions

Historically, protein aggregation has been viewed as a nonspecific, amorphous process that is very difficult to study and unlikely to be relevant to events in the cell. Recent studies of amyloid-like phenomena, however, are showing that at least some aggregation processes are highly specific, more closely resembling crystallization phenomena. As described in this article, a logical approach that assumes such crystal-like packing and growth as a starting point for focused analytical experiments can make great progress in understanding the aggregation process both in vitro and in vivo. Far from being alien to the cell environment, it is now known that aggregation is a continuing background problem in the life of the cell. Although cells can normally manage the process pretty well, in some cases, as in the amyloidrelated neurodegenerative diseases, the normal cellular safeguards break down, are overwhelmed, or are isolated by cellular compartmentalization, and protein aggregation proceeds unabated, with devastating consequences. Protein aggregation, long swept under the rug in the protein-folding field, is now very much a viable subject for biophysical studies. While there has long been a perception that protein misfolding and aggregation are shadowy events occurring behind a veil impenetrable to the experimentalist, a variety of analytical methods exist [115], and improved methods are being developed [99, 116–120], that are capable of revealing these processes with analytical clarity.

Acknowledgements

The author would like to thank Geetha Thiagarajan for help with the experimental Sextion of this chapter and Angela Williams, Valerie Berthelier, and Ashwani Thakur for permission to include unpublished data. I am also very grateful to Marcy MacDonald for providing Figure 1 would like to acknowledge past and present members of my laboratory, in particular, Songming Cheh, Valerie Berthelier, J. Bradley Hamilton, Wen Yang, Ashwani Thakur, Anusri Bhattacharya, Brian O’Nuallian, Alex Osmand, Angela Williams, and Tina Richey, for their woek and ideas as developing the body of work summarized here. I gratefully acknowledge the Hereditary Disease Foundation and the NIH (R01 AG19322) for sustained funding and Ethan Signer, Carl Johnson, and Nancy Wexler for their enthusiastic support.

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1

Thermodynamics of Amyloid Formation Ilia V. Baskakov

University of Maryland, Baltimore, USA


1 Introduction

More than 20 systemic and neurodegenerative maladies have been linked to the formation of ordered protein aggregates [1, 2]. When the deposited polymeric form is sufficiently ordered to bind Congo red and Thioflavin T, the term amyloid is used to define these types of aggregation [3]. A common feature of amyloid aggregates is formation of β-sheet-rich polymeric forms organized into highly ordered fibrils or plaques [4]. Recent studies have demonstrated that a broad variety of proteins unrelated to any known conformational disease can adopt β-sheet-rich amyloid forms in vitro and in vivo [5–8]. Medin, a proteolytic fragment of lactadherin, is an example of polypeptides that form amyloid deposits in vivo (amyloid deposits of medin were found in aorta of virtually all individuals studied older than 60 years), but has not been linked to any pathological processes [9]. Amyloidogenic proteins are now found in a variety of organisms including prokaryotes, plants, insect and mammals [10–13]. No consensus sequences that would predetermine the ability to form amyloid have been identified in any of these classes of proteins. Even though the amyloidogenic proteins show no obvious sequence similarity, they share a similar conformational property when converted into the amyloid fibrils, i.e. thermodynamically stable cross-β-pleated sheets [14, 15]. This finding has led to the hypothesis that the ability to fold into amyloid forms is not a unique property of certain proteins associated with degenerative maladies; rather, it may well be a feature of polypeptides in general [16].


2 Thermodynamics of Amyloid Formation

2 Thermodynamic versus Kinetic Control of Protein Folding

In his 1972 Nobel Prize lecture Christian Anfinsen described the “thermodynamic hypothesis” of protein folding [17]. This hypothesis states that “the threedimensional structure of a native protein in its normal physiological milieu is the one in which the Gibbs free energy of the whole system is lowest; that is, that the native conformation is determined by the totality of interatomic interactions and hence by the amino acid sequence, in a given environment”. Although the thermodynamic hypothesis has now been widely established and protein folding is commonly thought to be controlled by thermodynamic preferences, it has been understood by many, including Anfinsen and others, that kinetic issues can alter the folding landscape [18]. Whereas most small globular proteins will refold spontaneously in vitro to a native conformation, in vivo folding often exploits auxiliary molecules and defined subcellular compartments to avoid the deposit of ordered misfolded aggregates. What drives conversion of natively folded proteins into alternative misfolded conformations? Why do some proteins assemble into amyloid fibrils at some point within a protein’s lifetime, while others do not? Direct comparison of the thermodynamic stability of the native state with that of the β-sheet-rich amyloid state is impossible due to the insolubility, heterogeneity and high degree of polymerization of the amyloid fibril aggregates. On the other hand, the high resistance of the amyloid fibril aggregates to denaturation by detergents and to thermal and solvent-induced denaturation serves as a clever illustration of the extremely high thermodynamic stability and remarkable physical properties that the β-pleated sheet conformation imparts to amyloid fibrils. It is noteworthy that nature has learned to exploit these unusual properties of amyloid structures for a variety of physiological functions. For example, the major structural component of the shells of many insects and fish is of amyloid [11, 13]. To protect the developing embryo from temperature variation, mechanical pressure, proteases, bacteria, viruses and dehydration, the ability to construct complex amyloid structures evolved in these organisms through natural evolution and selection [11, 13]. Other examples of naturally occurring amyloid structures include extracellular curly fibrils expressed by Escherichia coli and Salmonella. These fibrils are involved in the colonization of bacteria on surfaces and in biofilm formation [19]. Furthermore, mammalian melanocytes produce a glycoprotein that polymerizes into amyloid-like fibrils, on which melanins are sequestered and concentrated during the multistage process of melanosome biogenesis [20]. These observations illustrate that amyloid formation is an evolutionary preserved biological pathway used to produce natural biomaterials with important physiological functions. While the direct thermodynamic analysis of insoluble amyloid structures is quite complicated, numerous studies have demonstrated that, in addition to insoluble amyloid fibrillar forms, many amyloidogenic proteins also adopt soluble oligomeric β-sheet-rich states [21–25]. Some of these β-sheet-rich oligomeric forms lie on the kinetic pathways to the amyloid fibrils, while others are off the kinetic pathways [24–26]. A direct comparison of the thermodynamic stability of the native and misfolded β-sheet-rich oligomeric isoform illustrates that the native state is not the

2 Thermodynamic versus Kinetic Control of Protein Folding

Fig. 1 Schematic free energy diagram of the conformational transition of the prion protein [27]. The "-helical monomeric isofrom is not the lowest energy state. However, the conformational transition form the "-helical

isoform to a $-sheet-rich oli-gomeric form is controlled by a large energetic barrier and, therefore, is prevented within the protein lifetime.

lowest energy state [27, 28]. It is shown in Fig. 1 that, when the unfolded state is used as a reference in the free energy diagram, the β-sheet-rich state is thermodynamically more stable than the native state. Even though the β-oligomeric states exhibit a thermodynamic stability higher than that of the native states, they might not be the true global energy minimum states, because the β-oligomeric species may undergo an additional time-dependent transition to highly polymeric amyloid forms [24, 25]. Why is the thermodynamically more stable β-sheet-rich isoform not accessible during folding under native conditions? It has been demonstrated that the rate of folding to the β-sheet-rich oligomeric isoform is slower by several orders of magnitude than the rate of folding to the native conformation [27]. To prevent the conformational conversion, the native state has to be separated by a large energetic barrier from the alternative β-sheet-rich state (Fig. 1). Although the free energy diagram does not provide a view of the actual kinetic pathways for the conformational transition, several important observations can be made regarding the origin of the energetic barrier. First, the native form has to unfold substantially on the route to the β-sheet-rich oligomeric isoform. Indeed, several studies have demonstrated that a substantial portion of the energetic barrier is linked to the partial unfolding of the native conformations [27, 29, 30]. The conversion to β-sheet-rich forms can be accelerated by shifting the native-unfolded equilibrium toward the unfolded state [6, 30–32]. Furthermore, the connection between the structural complexity of the pre-transition state and the energetic barrier is demonstrated by numerous observations that conversion of polypeptides with low structural complexity into β-sheet-rich isoforms occurs spontaneously and does not require partially denaturing conditions [21, 33]. A significant contribution to the energetic barrier seems to be associated with the process of assembly. Several studies demonstrated that the accumulation of a β-rich conformation is coupled with oligomerization. Analyses of the kinetic traces indicate that the process of folding to the β-oligomeric

3


isoforms represents a transition with an apparent reaction order of 3 or higher [27, 34]. This high order of reaction suggests that the conformational transition will depend dramatically upon the concentration of the transition state.

3 What Thermodynamic Forces are Responsible for the Exceptional Stability of Amyloid Aggregates?

It has been more than 40 years since Kauzmann described the thermodynamic forces responsible for the folding of proteins to the native conformations [35]. Such forces include hydrogen bonding, electrostatic, van der Waals, conformational entropy and hydrophobic interactions. Upon folding, groups accessible to solvent in the denatured state became newly buried in the native state. On transfer of the denatured state from a denaturing solution environment to physiological conditions, collapse of the denatured state occurs, causing removal of hydrophobic groups from water. This is believed to be an important driving force for folding of proteins into unique native conformations. For hydrophobic forces to play the major role in protein folding, the fraction of the hydrophobic groups destined for burial must be significant in the denatured states of proteins. Fig. 2 presents a analysis of the classes of groups that are buried upon protein folding in terms of their relative proportion as a function of protein molecular mass. Hydrophobic groups account for more than 20% of all buried groups upon protein folding and their relative proportion is consistently maintained, regardless of protein molecular mass. This result strongly supports the importance of the role that hydrophobic effects play in protein folding. Remarkably, it is evident from Fig. 2 that it is the peptide backbone that comprises the largest numerical fraction of groups newly buried on protein folding (numerical

Fig. 2 Number fraction of backbone and side-chain classes that are newly buried upon folding proteins of different molecular masses to their native states. In addition to the peptide backbone class, all side-chains are grouped into three classes: hydrophobic,

polar and charged side-chains. Crystallographic coordinates data were used from the PDB databank in evaluating the number of each group exposed in the native states of the proteins used in the calculations [38].

4 Single Polypeptide Chain-Multiple $-Sheet-rich Abnormal Isoforms

fraction = 55%). Whether the removal of the peptide backbone units from contact with water molecules is thermodynamically favorable or not has been widely debated [36]. The essential feature of the peptide backbone is its ability to form interand intramolecular hydrogen bonds. By forming hydrogen bonds between NH and CO groups in the folded state, polypeptides minimize the unfavorable effects of removing polar groups from water. The requirement that NH and CO groups of the peptide backbone must either form hydrogen bonds with water or with each other provides a strong structural constraint on the pathways toward possible folded states. This holds, especially, for ordered β-sheet-rich aggregates, where the role of hydrophobic interactions seems to be unsubstantial. Analyses of microcrystals of the amyloidogenic peptides demonstrated that amyloid fibril structures are highly hydrogen bonded, nearly anhydrous and densely packed β-sheets [37]. One can speculate that amyloid structures are formed in a way that optimizes formation of hydrogen bonds between strands of newly buried polypeptide backbone. The ability of many proteins to adopt alternative β-sheet-rich polymeric folding in vitro and in vivo argues that the mechanism involved and the thermodynamic forces that stabilize amyloid conformations have to be generic in nature. The fact that the numerical fraction of the newly buried peptide backbone is large and constant with respect to molecular mass illustrates that, under favorable conditions, burial of the peptide backbone could counteract the unfavorable exposure of hydrophobic groups. Considering that the number of newly buried peptide backbone groups predominates over that of newly buried side-chains lends support to the concept that it is hydrogen bonding of the peptide backbone that provides generic force for the stabilization of amyloid fibril forms. As the amyloid fibril aggregates grow and, correspondingly, the ratio of surface accessible to solvent versus the volume occupied by protein fabric decreases, the stabilizing effect increases. In the way that the hydrophobic force is considered one of the major determinants of the unique native conformation, it seems that the chemical nature of the polypeptide backbone, with its capacity for forming inter- and intramolecular hydrogen bonds is the central determinant of amyloid fibril formation [38].

4 Single Polypeptide Chain-Multiple β-Sheet-rich Abnormal Isoforms

Numerous biophysical studies have revealed that amyloidogenic proteins can adopt conformationally distinct non-native β-sheet-rich isoforms in vitro [26, 39]. For example, amyloid fibril formation by the prion protein occurs through a pathway different from the one that leads to formation of the β-oligomeric species [26, 40]. Regardless of which abnormal β-isoforms are biologically relevant, the ability of amyloidogenic proteins to form distinct abnormal conformers reflects the complexity of the energetic landscape of folding, as well as high conformational plasticity. How do different conformers arise from the same amino acid sequence in the absence of cellular cofactors or templates? Recent biophysical studies of model proteins demonstrated that physical properties of the native and denatured states of some proteins may change gradually

5


Fig. 3 Free energy diagram illustrating the thermodynamically variable characters of native and denatured states. Not only do the relative populations of the native (N) and the denatured (D) states change as a function denaturant concentration, but the

physical properties of both states are variable depending of solvent conditions. [N1 , N2 , N3 , N4 ] and [D1 , D2 , D3 , D4 ] illustrate the thermodynamically variable character of native and denatures states, respectively.

with environmental conditions [41]. The gradual change within the native and denatured species is consistent with the thermodynamically variable model, which postulates that the thermodynamic character of the native and/or denatured ensemble changes continuously as a function of the solvent environment [42, 43] (Fig. 3). The gradual change of the thermodynamic character is not always accompanied by conformational rearrangements of secondary structure. Thus, many proteins that display a thermodynamically variable behavior of the native ensembles did not show any changes as monitored by circular dichroism (CD) [42, 44]. However, such changes can be observed by other techniques such as nuclear magnetic resonance (NMR) [45], proton uptake/release [42], size-exclusion chromatography [41, 44] and dynamic light scattering [34]. One may speculate that the diversity of potential misfolding pathways of amyloidogenic proteins is linked to the variable thermodynamic behavior. Thus, a gradual environment-dependent change in the physical properties of the native and/or denatured ensembles may bias the adoption of particular pathway of misfolding under different pathological conditions.

5 Does the Process of Prion Propagation Differ from Formation of Ordered Amyloid Aggregates?

Among many amyloid-related maladies one subclass of the conformational disorders, prion diseases, seems to be distinguished by certain peculiar features: 1. The most unorthodox feature of prion disease is the existence of an infectious isoform of the prion protein, PrPSc [46, 47]. PrPSc propagates its abnormal conformation in an autocatalytic manner using the normal isoform (PrPC ) of the

6 Prion Propagation is an Autocatalytic Process

same protein as a substrate. In addition to the transmission of prion diseases in mammals, the phenomenon of self-propagating conformational transition has been described for prion proteins in yeast and in fungi [48, 49]. In all cases, the abnormal protein conformation acts either as the transmissible agent of disease or as a heritable determinant of phenotype. Reconstitution of mammalian prion infectivity in vitro has been difficult to accomplish for many years. This problem raised growing skepticism over the sufficiency of PrP alone to form an infectious agent. Recent work, however, demonstrated that the amyloid form of recombinant PrP induced a transmissible form of prion diseases in transgenic mice, providing the first compelling evidence for the “protein-only hypothesis” of prion propagation in mammals [50]. 2. Efficient self-propagation of prions requires identity or high homology between the amino acid sequences of PrPC and PrPSc , implying there is a high species specificity associated with their interaction. 3. Another prominent feature of prion propagation is the “strain” phenomenon. When PrPC is converted into pathogenic isoforms, a single unique amino acid sequence is capable of adopting conformationally distinct states, which are known as “strains” of PrPSc .

Interestingly, despite differences in the primary and tertiary structures of yeast and mammalian prions, the yeast prions display all of the characteristic features of mammalian prion proteins [51, 52]. Remarkably, recent studies illustrated that amyloid formation by non-prion proteins can also exhibit some properties that are thought to be peculiar for prion propagation. Thus, the systemic amyloidosis caused by the amyloid deposition of the serum amyloid A protein can be transmitted from animal to animal by a prion-like mechanism [53]. A species barrier, previously thought to be a distinctive feature of prion propagation, was also observed for the non-prion protein α-synuclein [54]. The growth of amyloid fibrils by Aβ peptides displayed striking specificity to cross-seeding by heterologous fibrils [55]. Since the process of amyloid formation is a general property of the polypeptide backbone, it would be of great interest to know how many amyloidogenic proteins are capable of self-propagating conformational transition in a prion-like manner (Fig. 4).

6 Prion Propagation is an Autocatalytic Process

Two competing models have been proposed to explain the autocatalytic conversion of prion folding: (1) the nucleation–polymerization model (NPM) [56] and (2) the template assisted model (TAM) [57, 58] (Fig. 5). Using different terminology, “heterogeneous nucleation” versus “templating”, both models employ an autocatalytic mechanism to explain the process of prion transmission and replication. Phenomenologically, the presence of a catalyst (nucleus or template) accelerates the process of conversion, either by avoiding a rate-limiting step in NPM or by lowering of the energy barrier in TAM. Both models predict low occurrences of sporadic

7


Fig. 4 The multidimensional sequence space defines all possible amino acids sequences. Among all species, only approximately 350,000 polypeptide sequences appear to have evolved through natural evolution and selection (http://protomap.cornell.edu/). Because the ability to adopt an amyloid conforma-

tion seems to be encoded in the physical/chemical nature of the polypeptide backbone, a substantial fraction of naturally occurring proteins would be expected to be amyloidogenic. The number of proteins that are capable of propagating abnormal conformations in a prion-like manner remains to be determined.

form of prion disease, and, correspondingly, a low rate of spontaneous conversion, since spontaneous or non-seeded formation of the first catalytic center is inaccessible either thermodynamically (according to NPM) or kinetically (as follows from TAM). Exogenous administration of catalyst (PrPSc in infectious form of disease) shortens the incubation time to such extent that disease develops within the human

Fig. 5 Schematic diagram showing the nucleation-dependent and the templateassisted mechanisms of the conversion of PrPC to PrPSc . Briefly, NPM postulates that the rate-limiting step is the formation of a nucleus: an oligomeric aggregate of PrP. The nucleus is thermodynamically unstable and this makes the spontaneous process a very rare event. However, as the nucleus is formed, further polymerization is

facile. Exogenous addition of PrPSc bypasses the formation of the nucleus. TAM, on the other hand, argues that PrPC is separated from PrPSc by a substantial energy barrier. The high barrier precludes the formation of PrPSc under normal conditions. However, the process of conversion is facilitated by an exogenous administration of PrPSc , which acts as a catalyst and lowers the energy barrier.

7 Conformational Diversity of Self-propagating Prion Aggregates Table 1 The autocatalytic mechanism predicts three possible outcomes with respect to prion

disease, depending on the multiplication coefficient (r)

Essential kinetic features Kinetics follows formal mechanisms of: Clinical form of disease

r>1

r≈1

r 1 the reaction rate increases exponentially, while the reaction decays when r < 1 (Table 1). Such mechanism postulates a threshold effect when r is very close to 1. In this case slight changes in experimental parameters may switch the reaction from a decay mode to an autoacceleration mode and vice versa. This mechanism predicts that the autocatalytic reaction can be induced even at subthreshold conditions (r < 1) by exogenous addition of catalytic centers. The proposed model is in agreement with the hypothesis that the progression of prion disease and the final outcome are controlled by fine dynamic balance between the rate of self-propagation and the rate of clearance of PrPSc . It would be of great interest to know what mechanisms account for multiplication of catalytic centers in vivo. The ability to control the rate of multiplication of catalytic isoforms of PrP would be a powerful strategy to treat prion diseases.

7 Conformational Diversity of Self-propagating Prion Aggregates

The “protein-only hypothesis” postulates that the given PrP amino acid sequence must be capable of adopting separate states when it is converted into pathological isoforms called “strains” of PrPSc . The existence of different prion “strains” has been recognized for a long time in the history of prion diseases [59, 60]. Each prion “strain” is associated with distinct neuropathologic features, such as the length of incubation time and the distribution of neuronal vacuolation in the brain [61]. These features are stable during serial transmission of PrPSc in a given host species. In the absence of another molecule that assists conversion, the formation

9


of different strains by the same protein can only be accomplished either by covalent modification of PrP or through the adoption of multiple conformations. During the past several years, considerable evidence has been amassed indicating that the properties of prion “strains” are enciphered in the conformation of PrPSc [62–66]. According to the template assisted model the abnormal PrPSc isoform provides a conformational template and guides conversion of PrPC into a conformer identical to PrPSc . Therefore, the conformational properties of each individual “strain” are inherited by nascent PrPSc . Although supported by numerous experiments carried out in vivo, the principle of formation of conformationally distinct self-propagating aggregates by the prion protein within the same primary sequence has yet to be demonstrated in vitro. The questions of fundamental importance are: (1) what factors determine the conformational diversity of self-propagating protein aggregates, (2) what is the role of the primary structure of protein versus template and environmental factors in this process, and (3) how do new conformers (“strains”) arise?

8 High Species Specificity of Prion Propagation

Numerous experiments on transmission of prions between different species demonstrated that prion replication is highly specific with respect to the primary structure of interacting isoforms, PrPC and PrPSc [67]. When the amino acid sequence of the PrPC of a recipient animal is identical to the sequence of PrPSc of a donor animal, the disease has the shortest incubation time. In contrast, if prions are transmitted from one species to another, the incubation time is longer in the first passage and the newly infected animals develop atypical clinical signs and unusual histopathology [68]. Once the initial passage has been accomplished, the incubation period shortens in subsequent passages and becomes fixed, as does histopathology. This phenomenon is called a “species barrier” [69]. From studies in transgenic animals, two factors have been identified that contribute to the species barrier: (1) the difference in PrP sequences between the donor species and the recipient species, and (2) the “strains” of PrPSc . It has been noticed that, when transmitted to a different species, some “strains” overcome the species barrier more easily than others [70]. To explain the species barrier, Caughey et al. proposed that when both homologous and heterologous PrPC are present, the heterologous PrPC (PrPC with a sequence different from PrPSc ) could bind to PrPSc without conversion. By binding to PrPSc , the heterologous PrPC inhibits binding and conversion of homologous PrPC [71]. In this model, the process of conversion, rather than binding, requires amino acid compatibility between PrPC and PrPSc . Although this simple model rationalizes a substantial body of experimental data on the species barrier, it fails to recognize the strain specificity of the species barrier. The questions are: (1) why can some “strains” from donor species be transmitted to recipient species, while other “strains” cannot, and (2) why some, but not all, recipient species propagate certain “strains” from donor species? It is clear now that the strain and the species barrier

9 Conclusions 11

Fig. 6 The new conceptual framework postulates that (1) primary structure of PrP of each individual species (A, B and C) determines conformational diversity of “strains”. (2) However, a template specifies particular conformation within given conformational space. (3) Subsets of conformers (“strains”) formed by PrP molecules from different species may overlap suggesting that some

PrPSc “strains” are more universal and can be shared by two or more species, whereas other “strains” are more species-specific. For instance, “strain” n is shared by species A, B, and C, while “strain” m is shared by species A and B. Only those “strains” that occupy overlapping areas can be propagated by more than one species.

phenomena are closely connected. Therefore, the experimental data related to both phenomena have to be discussed in the context of a common model. Fig. 6 represents a new conceptual framework that explains the strain specificity of the species barrier. This framework may provide general guidance for research conducted on this topic.

9 Conclusions

The observations that many proteins are able to adopt alternative amyloid-like folding require us to revisit many basic issues of protein folding, such as the role of kinetic traps in the folding pathway, the complexity of the energy landscape and the position of the native state in this landscape. An alternative view of protein folding presented in Fig. 7 proposes a complex energy surface with multiple energy minima. Most polypeptides are trapped in a native state that corresponds to a local free energy minimum for the entire protein lifetime, whereas global energy minima are occupied by β-sheet-rich states, which are kinetically inaccessible under physiological conditions and at indefinite protein dilution. However, despite high kinetic barriers, some proteins, including PrP, α-synuclein, parkin, tau and other, find a route to β-sheet-rich multimeric structures with unfortunate consequences. If a β-sheet-rich amyloid structure is an intrinsic preference, particularly at high


Fig. 7 An idealized “volcano energy landscape” of protein folding, to illustrate how a protein could fold fast to the native state, which might not the lowest energy state. Most proteins remain “trapped” in their native states for the entire protein lifetime, because alternative $-sheet-rich

states are separated by a high energetic barrier. Opposite to the native form, the $-sheet-rich state of each individual protein may adopt more than one conformation within given conformational space. N = native state, U = unfolded state; A = amyloid state.

protein concentration, then compartmentalization of proteins during folding and clearance of misfolded proteins should play critical roles in cellular health. In addition, amyloidogenic sidechain patterns, such as alternating polar and nonpolar amino acid residues, will be avoided through natural selection [72]. With many fundamental rules that govern protein folding having been discovered, new questions have to be addressed in the nearest future. How general is the phenomenon of self-propagating conformational transition? Can the prion hypothesis be expanded to non-prion proteins? Are individual non-prion proteins capable of assembly into conformationally distinct self-propagating aggregates (“strains”)? To what extent is sequence identity (or high homology) between native and selfpropagating abnormal isoforms necessary for efficient self-propagation? What is the role of the template in conformational diversity of self-propagating β-sheet-rich states?

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1

Post-translational Chemical Modifications in Amyloid Fibril Formation Melanie R. Nilsson McDaniel College, Westminster, USA


1 Introduction

From a purely na¨ıve point of view, it would seem that an obvious connection between normal protein degradation reactions (i.e. normal protein aging) and diseases associated with aging (e.g. amyloidosis) may exist. This connection, however, has received little attention. The lack of research in this area stems, in part, from the difficulties associated with detecting these subtle modifications and, in part, from the proposition that these modifications are not relevant to disease. The “conformational hypothesis” (the predominant view) suggests that amyloid fibril formation results from destabilization of the native protein as a result of changes in the local solution environment (e.g. pH, ionic strength, accessory proteins) [1–3]. An alternative view, which I will call the “chemical modification hypothesis”, suggests the protein conformational change is facilitated by a change in the primary structure (i.e. chemical modification) of the protein. The local solution environment can dramatically affect the types of chemical modifications that occur, so the solution environment is still a critical factor in fibril formation. Both hypotheses are consistent with Anfinsen’s view (cited in [1–3]) that the primary structure and aqueous environment govern the three-dimensional fold of a protein. In the conformational change hypothesis, the alteration in three-dimensional structure arises from changes in the aqueous environment, whereas in the chemical modification hypothesis, the altered conformation is a result of an altered primary sequence. It is quite possible that both hypotheses may be true, but applicable to different proteins. However, it is important to resolve which mechanism occurs in each disease because unraveling the mechanism of aggregation will have a significant impact on evaluating appropriate therapeutic strategies. The purpose of this article is to present the evidence that supports the view in which post-translational chemical modification of the primary sequence may be an important mechanism of amyloid fibril formation. Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Post-translational Chemical Modifications in Amyloid Fibril Formation Table 1 Modification of amino acids in proteins (reprinted from [4], with permission)

Amino Acid

Type of modification

Alanine Arginine Asparagine Aspartic acid

GPI-anchoring N-methylation, ADP-ribosylation, deimination glycosylation, GPI-anchoring, deamidation phosphorylation, methylation, isomerization, racemization, GPI-anchoring cystine formation, selenocysteine formation, heme linkage, myris-toylation, palmitoylation, S-nitrosylation, oxidation to C-α-formylglycine, deamination, GPI-anchoring γ -carboxyglutamate formation, methylation, polyglycylation deamidation, crosslinking, pyroglutamate formation, ADP-ribosylation, N-methylation GPI-anchoring, myristoylation methylation, diphthamide formation, phosphorylation, deamination N-acetylation, N-methylation oxidation, hydroxylation, crosslinking, palmitoylation, biotinylation, deamination hydroxylation hydroxylation, glycosylation phosphorylation, glycosylation, acetylation, GPI-anchoring, oxidation to C-α-formylglycine, dehydration and formation of a thioether link phosphorylation, glycosylation, dehydration and formation of a thioether link iodination, phosphorylation, sulfation, flavin linkage, nucleotide linkage, o-tyrosine, chloro-, nitrotyrosine and dityrosine formation formylation, acetylation, pyroglutamate formation GPI-anchoring, amidation, polyglycylation

Cysteine

Glutamic acid Glutamine Glycine Histidine Lysine Phenylalanine Proline Serine Threonine Tyrosine N-terminus C-terminus

The primary structure of a protein refers to the covalent arrangement of atoms in a protein. This includes not only the sequence of amino acids, but also posttranslational processing (e.g. formation of disulfides, proteolysis, glycosylation, lipid modification, phosphorylation) in addition to non-enzymatically regulated chemical modifications such as deamidation, racemization and isomerization. Table 1 lists some chemical modifications that are known to occur in proteins [4]. It is important to approach any relationship between primary structure and protein conformation with caution, since these relationships can be extremely difficult to predict. For example, it is not surprising that myoglobin and lysozyme have very different folds (myoglobin is all α-helical and lysozyme is a mixed α/β fold) since the primary structures are not similar. However, there are proteins with significantly different primary structures that all have the same basic fold (e.g. several non-homologous proteins fold into an α/β (TIM) barrel [5]). Another insightful example is the case of mutant sequences of transthyretin (TTR). Some mutations of TTR destabilize the protein conformation and facilitate amyloid deposition. However, there are also mutations that stabilize the native conformation and suppress amyloid formation, and polymorphisms that have no significant impact on

1 Introduction 3

amyloid formation. (For a review on ATTR, see [6].) These examples emphasize that it is not easy to predict the impact a given chemical modification will have on amyloid fibril formation and, furthermore, it cannot be assumed a priori that chemical modifications will play a primary role in all amyloid diseases. Post-translational modifications encompass both enzymatic and non-enzymatic modifications. Enzymatically regulated modifications are a normal part of protein processing and typically are required for protein folding, stability or function. Nonenzymatic chemical modifications, in contrast, are generally not part of normal protein processing and result simply from the favorability of these reactions in the chemical environment of the body. Both types of modifications result in a change in the primary structure of the protein and, therefore, can potentially contribute to amyloid fibril formation. The role of enzymatically regulated modifications in amyloid fibril formation is unclear. There are two possible scenarios by which these modifications could affect fibril formation – one in which the correct processing events do not occur and the other in which there is modification at the wrong place or at the wrong time. The absence of the modifications necessary for normal processing could play a role in amyloid formation by either destabilizing the native fold or by the presence of an unprocessed region that may initiate deposition. For example, incompletely processed islet amyloid polypeptide (IAPP), in which the pro-hormone is not completely proteolytically cleaved, has been identified in amyloid deposits [7–9]. The incorrectly processed peptide contains IAPP with an additional N-terminal peptide. The presence of the N-terminal segment could contribute to amyloid formation by destabilizing IAPP (possibly by preventing the formation of the proper storage form [10]) or the N-terminal peptide may bind to components of the extracellular matrix and act as an initiation site for fibril formation [11]. Similarly, evidence for the precursors of calcitonin [12] and atrial natriuretic peptide [13] have been identified in amyloid deposits. Enzymatically regulated modifications may also contribute to fibril formation by occurring at the wrong time or in the wrong place. For example, tau, the protein component of neurofibrillary tangles in Alzheimer’s disease, is hyperphosphorylated [14]. Phosphorylation is an enzymatically regulated process but, in this case, occurs in excess. The role of non-enzymatic chemical modifications in amyloid fibril formation is also unclear. Proteins spontaneously degrade in vivo and in vitro by a variety of mechanisms. Some of the more common reactions are racemization, oxidation of methionine, and deamidation of asparagine and glutamine. In vitro these modifications are frequently ignored and considered of minor significance. One notable exception, however, is the stability of peptide drugs, which has received considerable attention in an attempt to prolong their shelf life. In vivo the body normally protects itself against these modifications by either preventing the modification, repairing the modified sequences or clearing the damaged proteins. It has been proposed, for example, that the deamidation of asparagine to aspartic/isoaspartic acid may serve as a type of molecular clock to estimate the amount of time a protein has been in the cell [15]. Presumably, after a specified period of time, a signal will be sent to degrade the protein.

4 Post-translational Chemical Modifications in Amyloid Fibril Formation

Alterations due to the labile nature of amino acids can lead to additional functionality, but there is also a danger that these reactions will result in negative consequences. This is reminiscent of the mutability of DNA that allows for evolution but also can lead to disease. Like random mutations in DNA, there is no way to easily predict the outcome of a chemical modification, but the probability of a damaging modification that is not repaired will statistically increase with age (for an overview of chemical damage associated with aging, see [16]). This is a very attractive proposition to explain the familial versus sporadic variability of amyloid disease progression. Familial cases of amyloid diseases occur when a genetic mutation leads to a protein sequence that is more prone to aggregation than the wild-type sequence. There are also, however, late-onset variations in which there are no mutations and the mechanism of destabilization of the native protein is unclear. One possible explanation is that chemical modifications which result from normal age-related protein degradation reactions may destabilize the proteins in a similar fashion as inherited point mutations. 2 Common Modifications that May Play a Significant Role In Vivo 2.1 Cleavage by Proteases or Non-enzymatic Hydrolysis

The cleavage of a peptide or protein can occur as a result of protease activity or nonenzymatic hydrolysis. Four major classes of proteases (aspartic acid, serine, cysteine and metalloproteases) have been identified and named according to the catalytic portion of the active site. Proteases are commonly stored in lysosomes and, therefore, protein cleavage associated with lysosomes has been implicated for many years in amyloidogenesis. For example, Glenner et al. in 1971 proposed a primary role for proteolysis in AL amyloid formation: “The fact that ‘amyloid’ fibrils can be created from some Bence–Jones proteins at a physiologic temperature in the presence of a proteolytic enzyme having an acidic pH optimum suggests that one possible pathogenetic mechanism for amyloid formation may be by means of intralysosomal catheptic digestion of light polypeptide chains of immunoglobulins” [17]. Cleavage of the amide backbone of a peptide or protein, however, does not need to be mediated by proteases. Acidic conditions can be used to facilitate cleavage of a protein into constituent amino acids in preparation for amino acid analysis. Although the conditions employed in this technique are not found in the body, hydrolysis of many amide bonds occurs under physiologically relevant conditions. Amide hydrolysis can be catalyzed by acid or base, but the mechanisms and labile sites are substantially different. In several cases in which the amyloid fibril component is a fragment of a larger precursor, specific proteases have not been identified in the generation of the amyloidogenic fragment, suggesting that some proteins may be cleaved by non-enzymatic reactions. A survey of the amyloid literature results in a multitude of examples in which cleavage of the protein backbone is implicated in amyloid fibril formation. For

2 Common Modifications that May Play a Significant Role In Vivo 5

example, in the most common form of amyloid deposition (which occurs in the aortic smooth muscle), the major protein component is medin, a small fragment of lactadherin [18]. More recently, unusual fragments of kerato-epithelin have been identified in corneal amyloid deposits [19]. Additional examples in which cleavage by proteolysis or hydrolysis may play a significant role in amyloid fibril formation are highlighted below. Non-pathological proteolytically induced amyloid fibril formation has been identified in melanosome biogenesis [20]. Pmel17 is a protein expressed in melanocytes and, as part of normal processing, is cleaved into two unequal pieces (the 80-kDa Mα fragment and the 28-kDa Mβ fragment) by a furin-like convertase. The Mα fragment subsequently forms insoluble fibrils which act as a scaffold for melanin pigments. The full-length protein is unable to form fibrils and assembly only occurs upon cleavage. It is speculated that this tight control mechanism prevents aberrant fibril formation. Other physiologically relevant amyloid processes, such as curli formation in Escherichia coli, are also carefully regulated events [21]. Lithostathine (also known as pancreatic stone protein, S2) is a 144-residue secretory protein that generates a 133-residue amyloidogenic fragment (S1, pancreatic thread protein) upon spontaneous autocleavage or cleavage with trypsin [22]. S1 has been identified in the brain of patients with Alzheimer’s disease [23] and in pancreatic stones [24]. Cleavage of lithostathine to S1 is a pre-fibrillogenic event and is an interesting example of fibril formation that can be induced by proteolysis or by an autocatalytic mechanism. Gelsolin is an actin-binding protein [25]. In familial amyloidosis of the Finnish type (FAF), there is a mutation at residue 187 in which an aspartic acid is converted to an asparagine or tyrosine [26, 27]. The mutations have mild effects on the thermodynamic stability of the protein [28] but lead to a susceptibility to proteolytic cleavage [29]. Cleavage results in the production of an amyloidogenic fragment that encompasses residues 173–223 or 173-225 [26]. In contrast, the wild-type protein is not cleaved and does not form amyloid fibrils. The amyloidogenic fragment has been observed in the cerebrospinal fluid (CSF) [30] and identified as the major circulating form of patients with FAF [31, 32], suggesting that cleavage is the driving force in FAF amyloid deposition. Cystatin C (also known as γ -trace basic protein) is an inhibitor of cysteine proteases. In hereditary cystatin C amyloid angiopathy (HCCAA or HCHWA-I), there is a mutation at position 68 in which a leucine is replaced with a glutamine [33]. Amyloid deposits of this variant protein contain cystatin C lacking the first 10 residues [33]. The soluble truncated form has not been observed in detectable quantities in the CSF [34], but the loss of the 10-residue fragment is consistent with the proposed domain-swapped dimer mechanism of cystatin fibril formation [35]. The dimer interface occurs between strands 2 and 3 (residues 42–59) [35], and the loss of the first 10 residues would eliminate the edge strand (strand 1) and potentially facilitate deposition. In sporadic cerebral amyloid angiopathy with cystatin C deposition (SCCAA), there is no mutation present and full-length cystatin C is found in amyloid deposits. However, in SCCAA it is speculated that cystatin C co-precipitates with the Aβ peptide [36].


N-terminal fragments of different apolipoproteins are associated with a wide variety of amyloid deposits. Fragments of apolipoprotein AIV (ApoAIV) have been co-localized with TTR in cardiac amyloid and the authors have proposed that this fragment may “serve as a local nidus for fibrillogenesis” [37]. An N-terminal fragment of apolipoprotein AI (ApoAI) is the major fibril protein in atherosclerotic plaques and a fragment of variant ApoAI is the major amyloid constituent in FAP Type III (Iowa) [38–40]. Similarly, the N-terminal portion of the apolipoprotein serum amyloid A (ApoSAA) has been isolated from amyloid deposits and this fragment has been generated in vitro upon digestion with cathepsin B, suggesting that proteolysis is a prefibrillogenic event [41, 42]. Hatters and Howlett propose a mechanism by which apolipoproteins may be deposited. They suggest that “oxidative processes may promote protein truncation or modification of apolipoproteins in a manner that perturbs their lipid binding properties and thus by default promote the amyloidogenic folding options” [43]. Familial British dementia (FBD) is associated with amyloid fibrils composed of C-terminal fragments of α- and β-tubulin [44] and fragments of a protein of unknown function named BRI [45, 46]. In FBD, a mutation in the BRI gene results in the production of a protein (BRI-L) that is extended by 11 amino acids at the C-terminus. Cleavage of this protein near the C-terminus results in the release of a 34-amino-acid peptide that readily forms amyloid fibrils [45]. Cleavage of BRI-L is speculated to be mediated by furin [47, 48]. A similar proteolytic event is involved in the formation of a different amyloidogenic peptide derived from BRI that occurs in familial Danish dementia [48]. β 2 -Microglobulin (β 2 M) amyloidosis occurs in patients undergoing long-term dialysis treatment. N-terminally truncated forms of β 2 M are common in ex vivo amyloid. Cleavage occurs between K6 and I7 (around 25%), and to a lesser extent between residues 10–11, 17–18, 19–20, 86–87 and 98–99 [49, 50]. Truncated β 2 M, which corresponds to a loss of the first six residues, has a high tendency to aggregate in vitro [49, 51]. Therefore, it is possible that truncation of β 2 M destabilizes the native fold and leads to amyloid fibril formation. TTR amyloid formation occurs in familial amyloid polyneuropathy (FAP) and senile systemic amyloidosis (SSA). FAP is associated with mutations in TTR [52], while SSA is associated with a fragment of wild-type TTR that results from cleavage between residues 46 and 52 [53]. It has been proposed that the mutations observed in FAP destabilize the native TTR fold and facilitate deposition. The lack of mutations observed in the A and H β-strands of TTR and the demonstration that a synthetic peptide of the A-strand can form amyloid fibrils in vitro [54, 55] led to the hypothesis that the A-strand (and H-strand) may be critical for fibril formation [56]. However, if the A-strand plays a critical role in the fibril structure, removal of this strand would be detrimental to fibril formation. The observation that around 80% of wildtype TTR in ex vivo amyloid fibrils lacks the N-terminal region (residues 1–46, 49, 52 or 53) [53] which contains the A-strand (residues 10–20) is inconsistent with this view. Therefore, wild-type TTR and mutant TTR may form fibrils via different pathways. Mutations of TTR could lead to global destabilization of the native fold that facilitates amyloid fibril formation, resulting in the relatively early age of onset


observed in FAP. In contrast, wild-type TTR may be cleaved as a result of age-related chemical modification or aberrant proteolysis, which would be consistent with the older age of onset of SSA. α-Synuclein (also known as NACP) has been identified in inclusions associated with Parkinson’s disease, dementia with Lewy bodies (DLB), Alzheimer’s disease, multiple system atrophy (MSA) and amyotrophic lateral sclerosis (ALS) (reviewed in [57]). Lewy bodies contain α-Synuclein and in vitro studies reveal α-synuclein fibrils have all the hallmarks of amyloid [58]. Truncated forms of α-Synuclein have been reported in Parkinson’s disease and DLB [59, 60]. Fragments of α-Synuclein (around 2–4 kDa shorter than the full-length protein) have been identified in Lewy bodies, but not in a normal control [60]. Fragments that are around 4 and 10 kDa shorter than full-length α-Synuclein have been observed in soluble fractions of Parkinson’s disease, DLB and normal control extracts [59], suggesting these fragments may be normal breakdown products of α-Synuclein. However, the smaller fragment was also identified in poorly soluble fractions suggesting this fragment may play a role in fibril formation [59]. Immunochemical analysis indicates that the 4- and 10-kDa truncated fragments are derived from the middle region of α-Synuclein [59], which is consistent with the previously identified NAC peptide isolated from the brains of patients with Alzheimer’s disease [61]. Significant Aβ N-terminal and C-terminal heterogeneity has been observed in Alzheimer’s disease amyloid deposits. Aβ is generated by cleavage of a larger precursor, the amyloid β precursor protein (AβPP). In addition to Aβ, cleavage of AβPP can generate two other peptides, C100 and p3 [62]. All three peptides (Aβ, p3 and C100) have been implicated in amyloid formation. Intact C100 is competent to form amyloid fibrils which, upon non-specific proteolysis, results in fibrils composed of Aβ [63]. The peptide p3 is the major component of “preamyloid”, indicating an important role of p3 in amyloidogenesis [64]. Ragged N and C-termini of Aβ have been found in plaques. The C-termini reported include V39, V40, A42 and T43 [65]. Aβ which ends in A42 is highly amyloidogenic and represents the earliest C-terminus observed in deposits, suggesting that an increase in the ratio of A42 to other peptides may facilitate fibril formation [66, 67]. Aβ peptides have been identified with N-termini beginning with –I6, -V3, D1 or isoD1, A2, pE3, F4, S8, G9, pE11 and L17. The earliest N-termini in deposits have been speculated to be L17 (p3) followed by deposition of pE3 and D1 [64, 66]. The full-length human prion protein (PrPc ) consists of residues 23–231 after the signal sequence is removed from the primary translation product and the GPI anchor attached. A protein truncated at approximately residue 80–90 is significantly elevated in Creutzfeldt-Jakob disease [68]. It has been proposed that heterogeneity at the N-terminus could contribute to the strain variation observed in prion diseases [69]. Neuronal inclusions observed in Huntington’s disease are fibrillar [70] and Congo red birefringent [71, 72]. The inclusions contain huntingtin and a 40-kDa N-terminal fragment of huntingtin [70]. In vitro experiments on N-terminal fragments of huntingtin have led to the suggestion that “the proteolytic cleavage event that yields this ‘toxic’ fragment would also provide a rate-limiting step in Huntington’s disease


onset” [73]. Proteolysis has also been proposed as an important factor in other polyglutamine disorders [74]. The neurotoxicity observed in Huntington’s disease and other polyglutamine diseases may be mediated by the binding of polyglutamine proteins (e.g. TATA-binding protein) to the huntingtin deposit which could deplete the necessary supply of these proteins in the brain [72]. Aberrant processing of viral proteins may lead to amyloidogenic fragments that can nucleate amyloid deposition [75]. This has been championed in the case of glycoprotein B (gB) from herpes simplex virus 1 (HSV1). HSV1 DNA has previously been identified in the same regions of the brain that are affected by Alzheimer’s disease [76, 77] and more recently a fragment of gB has been demonstrated to accelerate Aβ fibril formation in vitro [75].

2.2 Deamidation, Isomerization, Racemization and Protein l-Isoaspartyl Methyltransferase (PIMT)

Deamidation of asparagine and glutamine is common under physiological conditions (for reviews, see [78–81]). Both the reaction rate and mechanism are influenced by a host of factors, including solution conditions (e.g. pH, ionic strength, buffers, temperature), primary sequence, secondary structure and tertiary structure. The inherent chemical and thermal lability of these residues is highlighted in the observed degradation products of peptide drugs upon aging (e.g. calcitonin [82], pramlintide [83], insulin [84]) and in the low number of Q/N repeats in thermophiles compared to mesophiles [85]. Deamidation reactions can lead to fragmentation of the amide backbone or result in subtle backbone or side-chain modifications. For example, asparagine deamidation can lead to the formation of L- or D-aspartic acid, L- or D-isoaspartic acid or fragmentation of the protein chain (Fig. 1) [86, 87]. Deamidation of glutamine can yield glutamic acid or, if the residue is at the N-terminus, pyroglutamic acid. The conversion of an asparagine to an aspartic/isoaspartic acid or glutamine to glutamic/pyroglutamic acid results in a protein that has the equivalent of a posttranslational point mutation. From a protein folding point of view, the chemical modification could affect the thermodynamic stability of the native fold, the kinetics of folding or unfolding, or the overall flexibility of the molecule. Deamidation of ribonuclease A (N67isoAsp), for example, significantly decreases the folding rate [88] while deamidation of lysozyme (N103D or N106D) increases the susceptibility to proteolysis [89]. The destabilization of the native state or enhanced flexibility could result in proteins that are more prone to aggregation. Deamidation leads to a change in net charge of the protein. Under physiological conditions (pH ∼7.4), asparagine and glutamine are neutral, but aspartic acid and glutamic acid are negatively charged. Model studies suggest that aggregation is favored when the net charge of the protein is closer to zero [90], so deamidation may facilitate amyloid fibril formation of basic proteins by altering the net charge. Several other post-translational chemical modifications can alter the net charge


Fig. 1 Mechanisms for deamidation, isomerization, and racemization of asparagine and aspartic acid. (Reprinted from [86, 87], with permission)

of a protein, including C-terminal amidation, N-terminal acetylation, advanced glycation (or glycosylation) end-product (AGE) formation and arginylation [91]. Polyglutamine repeat diseases [e.g. Huntington’s disease, spinalbulbar muscular atrophy (Kennedy’s disease), dentatorubral-pallidoluysian atrophy, and spinocerebellar ataxias 1, 2, 3 (Machado-Joseph disease), 6, 7 and 17] are characterized by the aggregation of glutamine-rich peptides/proteins [92, 93]. Huntington’s disease is the most common glutamine-repeat disease, and in vitro analysis of glutaminerich peptides demonstrates that these deposits are fibrillar, β-sheet rich and bind Thioflavin T [94]. Polyglutamine peptides are likely candidates for deamidation, but the effect of deamidation on fibril formation has not yet been investigated. Polyglutamine peptides are, however, susceptible to transglutaminases which can catalyze deamidation of glutamine or crosslinking between glutamine and other amines (e.g. lysine) [95]. These chemical modifications could potentially play a role in fibril formation by promoting nucleation or by stabilizing the resulting fibril. Glutamine- and asparagine-rich domains are common in yeast prions, and it is speculated that these repeats may provide interaction specificity [85]. These proteins, however, will also likely be susceptible to deamidation. Deamidation of PrPC has been shown to occur spontaneously [96] and results in a protein that readily forms protease K-resistant aggregates upon exposure to Cu2 + [97]. Aggregation of other prions may be mediated by a similar pathway.


β 2 M extracted from amyloid deposits has been reported to be deamidated at positions 17 and 42 (Asn to Asp/isoAsp) [98, 99]. In vitro experiments suggest that Asn 17 to Asp is unlikely to accelerate amyloid fibril formation [100], but the other deamidated species have not been investigated. Similarly, deamidation and isomerization of tau has been identified in paired helical filaments, although no specific role in the aggregation process has been identified for these modifications [101]. In familial cataract, amyloid fibrils composed of crystallins are thought to precede cataract formation in vivo [102] and this process is supported by in vitro studies [103]. The crystallins are a group of proteins that reside in the human lens. These proteins do not turn over as the lens ages, so chemical modifications of crystallins are common in both normal and cataractous lenses. Investigations to elucidate differences between normal and cataractous lenses have revealed elevated deamidation of γ S-crystallin in cataractous lenses [104] at asparagine 143 [105]. Significant levels of glutamine deamidation have been reported in ex vivo Aβ amyloid deposits. The glutamine at position 3 of Aβ is significantly deamidated (10–50%) to yield an N-terminal pyroglutamic acid (pE3 Aβ) [65, 106–108]. Pyroglutamic acid at position 11 (pE11 Aβ) has also been reported [65]. The highly variable amounts reported for pE3 Aβ in the brain may, in part, be due to the different localization of this peptide [65, 108]. pE3 Aβ is common in diffuse amyloid plaques (proposed to be an early event in amyloid deposition) and the ratio of Aβ to pE3 Aβ directly correlates with age (younger subjects have higher levels of pE3 Aβ in plaques) [65, 109]. pE3 Aβ has also been shown to aggregate more readily than Aβ in vitro [107, 110]. Taken together, this data suggests that the formation of pE3 Aβ is a pre-fibrillogenic event which can induce amyloid fibril formation of pE3 Aβ followed by the subsequent deposition of Aβ onto these deposits [109]. Isomerization of aspartic acid and asparagine (concurrent with deamidation) is one of the most common non-enzymatic chemical modifications of proteins in vivo. This reaction occurs most readily in regions of high flexibility and with preference to sequences that contain an aspartic acid or asparagine followed by a glycine. Isomerization results in a change in conformation of the peptide/protein backbone [111] which could facilitate fibril formation. Aβ has three aspartic acid residues (D1, D7 and D23) and isomerization of each residue has been reported in ex vivo amyloid deposits [108, 112, 113]. If isomerization occurs as a secondary phenomenon to exposed Asp residues after Aβ fibril formation, a relatively equal distribution of chemically modified Aβ would be anticipated. Interestingly, the distribution of modified forms of Aβ differ between neuritic and vascular amyloid [108, 114]. This variable localization has, therefore, led to the hypothesis that Asp isomerization is a pre-fibrillogenic event [114]. A direct role in nucleation of fibril formation is supported by the observation that peptides with isoAsp at D1 and D7 have a higher β-sheet content [115], and isomerized D23 forms fibrils more readily than unmodified Aβ [113]. Furthermore, the presence of isoAsp (or D-Asp) can prevent protein turnover [112], thereby increasing the local concentration. Alternative explanations for the role of Asp isomerization in Alzheimer’s disease have been proposed in which a central role is attributed to the succinamide intermediate that


is transiently populated during the conversion of Asp to isoAsp. The succinamide intermediate, unlike Asp and isoAsp, is neutral and the loss of charge has been postulated to facilitate aggregation [116]. Furthermore, the succinamide residue adopts a geometry similar to a type II β-turn and may result in the formation of cytotoxic Aβ β-hairpins [117]. Racemization occurs when the stereochemistry at the α carbon is altered from the L configuration to the D configuration. This process is quite common [118] and can be catalyzed by racemases or occur non-enzymatically. Non-enzymatic racemization of amino acids in proteins has been explored as a possible dating technique to estimate the age of shells and other biomineralized materials [119]. Bacteria use racemases to generate cell wall peptides that contain both L- and D-amino acids which are relatively resistant to proteases. Some racemase inhibitors (e.g. cycloserine) produce neurological side-effects which may be a result of interactions with the recently discovered human serine racemase in the brain [120]. The presence of racemized amino acids can affect the protein conformation, have an impact on the folding/unfolding rates, or alter the thermodynamic stability. Although D-amino acids can destabilize β-sheets, peptides with D-amino acids have been shown to readily form amyloid fibrils [121]. Racemization of aspartic acid and serine residues has been observed in Aβ isolated from ex vivo amyloid fibrils [122]. Aspartic acid is very susceptible to spontaneous racemization and D-Asp Aβ has been identified in vitro [123]. D-Asp Aβ aggregates faster than unmodified Aβ, suggesting a possible role for racemization in amyloid fibril nucleation [123, 124]. D-Ser at position 8 likewise facilitates aggregation, but D-Ser at position 26 does not [124]. However, D-Ser 26 Aβ can be proteolyzed into a smaller, highly toxic fragment that may play a role in Alzheimer’s disease [125]. PIMT is an intracellular repair enzyme that facilitates the conversion of D-Asp or L-isoAsp into L-Asp [126]. A lower amount of PIMT activity has been correlated with an earlier age of death [127], suggesting that there may be a relationship between the accumulation of chemically modified proteins and mortality. PIMT knock-out mice exhibit a 9-fold increase in L-isoAsp and a phenotype of fatal epilepsy [128]. Likewise, the examination of human hippocampal tissue reveals dramatically reduced levels (50% lower) of PIMT activity and a 2-fold increase in damaged tubulin in epileptic versus control samples [129]. Cortical samples show similar levels of PIMT activity compared to controls [129], which is consistent with an earlier Alzheimer’s disease study [127]. Thus, local depletion of PIMT may be an interesting area of investigation to determine if excess deamidation or isomerization plays an active role in amyloid fibril formation. 2.3 Oxidative Damage

The reactants and products of oxidation reactions are highly variable, resulting in a staggering array of possible protein modifications that may impact amyloid deposition. Proteins can be oxidized by reacting with metals, lipids and other small


molecules [16]. Oxidative damage has been proposed to play a role in a variety of lateonset diseases including Alzheimer’s disease [130]. For example, lipid peroxides can react with cysteine, histidine, and lysine residues to generate advanced lipoxidation end-products (ALEs) which can affect protein structure, charge, hydrophobicity and crosslinking [16]. Baynes and Thorpe “propose that atherosclerosis is an age-related disease characterized by accelerated lipid peroxidation and lipoxidative aging of proteins in the vascular wall” [16]. Abnormal metabolites (e.g. ketoaldehydes) have also been proposed to similarly react with proteins and affect aggregation [131]. Cysteine side-chains are frequently oxidized to form disulfide bonds as part of normal protein processing, but aberrant disulfide bonds may facilitate amyloid formation [132]. For example, normal lenses contain αA-crystallin in which Cys131 and Cys142 are partially reduced, but cataractous lenses contain only oxidized Cys131 and Cys142 which may impact lens transparency [133]. The release of monomeric TTR from ex vivo amyloid upon reduction suggests that there may be some disulfide bonds formed between TTR monomers in the fibrils [53]. (Note: Cys10 is the only cysteine in wild-type TTR.) The identification of a Cys10Arg TTR mutation that results in FAP [134] and a Cys10Ala mutant that forms amyloid fibrils in vitro [135], however, indicates that an intermolecular disulfide bridge is not necessary for the initiation of amyloid fibril formation. The Cys10 crosslinks may instead play a role in the thermodynamic stabilization of the fibrils. Cysteine residues on proteins can also react with free cysteine, cysteinylglycine, glutathione or sulfite in vivo. Disturbances in cysteine metabolism have been identified in several neurological disorders, including Alzheimer’s disease, Parkinson’s disease, motor neuron disease [136] and Hallervorden-Spatz disease [137]. Although no direct correlation to amyloid fibril formation has been elucidated, it has been proposed that altered thiol metabolism may contribute to the overall oxidative damage observed in these diseases. A more direct relationship between free thiols and amyloid fibril formation has been identified in AL amyloidosis and FAP. Protein extraction from ex vivo amyloid fibrils has revealed cysteinylation of Cys214 in k1 light chain in AL amyloidosis [138], and conjugation of Cys10 of TTR to sulfite, cysteine, cysteinylglycine and glutathione in FAP [139]. Modification of Cys10 with cysteine, cysteinylglycine or glutathione results in destabilization of TTR and enhanced fibril formation [140]. Significantly, these modifications result in TTR molecules that readily form fibrils at pH 4.8, a condition under which wild-type TTR does not aggregate [140]. Methionine oxidation has been reported for several amyloid fibril proteins including lysozyme [141], PrPsc [142], β 2 M [98] and Aβ [143, 144]. While earlier studies suggested that Met35 oxidation of Aβ increased the rate of aggregation [145], more recent work indicates that Met35 oxidation decreases the rate of βfi fibril formation [146–148]. Oxidation of methionine residues in α-synuclein results in a decrease in aggregation, but this inhibition is eliminated in the presence of metals [149–151]. In view of the present evidence, the role of methionine oxidation in amyloidosis remains unclear. It is possible that, in some cases, the oxidation may be protective by slowing fibril formation.


Tyrosine is susceptible to a variety of oxidation reactions that can lead to the formation of chlorotyrosine, nitrotyrosine, o-tyrosine and dityrosine [16]. Tyrosine nitration of α-synuclein in Parkinson’s disease and tau in Alzheimer’s disease has been reported and postulated to contribute to disease progression by affecting fibril formation or stability [152, 153]. Chlorination of tyrosine occurs upon exposure to HOCl, a strong oxidizing agent that is produced by neutrophils and monocytes [154]. Tyrosyl chlorination can promote protein aggregation [155] and has been implicated in formation of atherosclerotic plaques [156]. Dityrosine formation can form covalent crosslinks between peptides and it has been proposed that dityrosine formation of Aβ may stabilize amyloid fibrils in Alzheimer’s disease [157]. 2.4 AGEs

Carbohydrates can react with protein amino groups (e.g. N-terminus, lysine, arginine) to form a Schiff base that can rearrange to an Amadori product and, finally, an AGE. The time required to reach equilibrium is on the order of hours for Schiff base formation, days for Amadori products and weeks to months for AGEs. For an excellent review of AGE chemistry, see Bucala and Cerami [158]. AGEs, therefore, are predominantly identified in proteins with a low turnover rate in vivo. The formation of an AGE may result in the attachment of an adduct to the protein, protein crosslinking (e.g. an argininelysine crosslink or dilysine crosslink) [16] or fragmentation of the backbone [159]. The presence of AGE-modified proteins has been observed in cataracts [158], FAP [160], AA amyloidosis [161], Type 2 diabetes mellitus [162], Alzheimer’s disease [163] and patients undergoing long-term dialysis treatment [164]. AGE modification may promote amyloid fibril formation by decreasing the solubility as a result of the alteration in the net charge of the protein or AGE crosslinks could stabilize the fibrils. AGE modifications of both Aβ [163] and tau [165] have been identified in Alzheimer’s disease. AGE modification likely occurs early in the disease process [166] and may nucleate amyloid deposition as a result of accelerating Aβ aggregation [167]. In contrast, AGE-modified IAPP does not dramatically alter fibril formation kinetics, but does lead to fibrils that are more cytotoxic [168]. 2.5 Phosphorylation

Alzheimer’s disease is characterized by the presence of Aβ amyloid deposits and neurofibrillary tangles (NFTs) in the brain. Similarly, inclusion body myositis (IBM) is characterized by the deposition of Aβ amyloid and tau filaments in muscle. Ultrastructurally, the tau filaments are very similar in Alzheimer’s disease and IBM [169]. In Alzheimer’s disease, these filaments are called paired helical filaments (PHFs) and are the principal structural component of NFTs. In IBM, these filaments accumulate in muscle fibers and are called twisted tubulo filaments (TTFs). PHFs and TTFs are both composed of hyperphosphorylated tau [14, 169]. Phosphorylation


of tau is not required for the formation of PHFs in vitro, but fibrils composed of phosphorylated tau resemble authentic PHFs more closely than do fibrils generated from non-phosphorylated tau [170]. This data suggests that phosphorylation may be an important pre-fibrillogenic event. AGE modification of tau, however, has been proposed to play a greater role in PHF insolubility than phosphorylation [171]. Therefore, it is possible that phosphorylation may nucleate fibril formation and AGE modification may stabilize the resultant fibrils against degradation.

3 Proposed Mechanisms by which Chemical Modifications may Affect Amyloid Deposition

Virtually all amyloid deposits contain some protein that is chemically modified. However, the identification of chemically modified protein in ex vivo amyloid does not a priori indicate that the modifications play a causative role. Therefore, it is necessary to consider the possible ways in which chemical modifications may affect amyloid deposition. There are three fundamentally different ways in which chemical modifications may contribute to amyloidosis. A chemical modification may affect the rate at which fibrils are formed (kinetic effect), the relative stability of the fibrils (thermodynamic effect) or lead to an increase in fibril toxicity (cytotoxic effect). In order to have a kinetic effect on amyloid fibril formation, the chemical modification must occur prior to fibrillogenesis and accelerate the slow step of fibril formation. Most of the mechanistic models of amyloid fibril formation require two basic steps, nucleation and growth (for a review of current models, see [172]). Nucleation involves the conversion of soluble protein (S) into an assembly competent state (A) that forms the nucleus (N) or seed. The formation of the nucleus from the soluble protein is the slow (rate-limiting) step in amyloid fibril formation. Fibril growth then proceeds via the addition of protein to the nucleus. Protein can bind to the nucleus in the soluble form (S) and then rearrange upon binding [173–175], or it can bind in the assembly competent (A) form [176] (Fig. 2). The assembly competent state (A) is generally believed to be a partially folded intermediate state [1–3, 177]. The destabilization of the native state, therefore, can increase the population of this intermediate and drive nucleus formation. Amyloidogenic mutants, for example, have been shown to be less stable than the wild-type proteins and it is proposed that the mutants populate the assembly competent intermediate state more readily [178]. Analogously, it is possible that post-translational chemical modifications can result in a protein (S*) that is less stable than the wild-type, readily populates the assembly competent intermediate state (i.e. S*→A is fast) and facilitates nucleus formation (Fig. 2). The protein generated by post-translational chemical modification could even be the equivalent of the assembly competent protein (i.e. S* = A). Depending on the mechanism of protein addition to the nucleus, the growth of amyloid fibrils by the chemical modification mechanism could require either

3 Proposed Mechanisms by which Chemical Modifications may Affect Amyloid Deposition

Fig. 2 Nucleation and growth mechanisms. Key: S=soluble protein, S*=chemically modified protein, A=assembly competent state, N=nucleus, F=fibril.

small or large amounts of modified protein. If the soluble protein adds directly to the nucleus, then only trace amounts of chemically modified proteins would be required to drive amyloid fibril formation. If, however, the assembly competent state is required for addition, then the majority of the protein in the fibrils would need to be chemically modified. Chemically modified proteins have been observed in ex vivo amyloid deposits in both major and minor quantities, suggesting that both mechanisms may occur in vivo. Nucleus formation is inherently slow as a result of the entropically unfavorable association of the A-state protein. Increasing the concentration of the A-state protein, however, can help to overcome this barrier. The population of the Astate can be increased in two different ways; destabilization of the S-state could shift the equilibrium toward the A-state or the degradation of the A-state could be slowed. Chemical modifications of the primary structure likely affect both pathways. The likelihood of amyloid fibril formation proceeding by the mechanism out lined in Fig. 2b has been strongly supported by examples of proteins in which cross-seeding occurs. Cross-seeding experiments demonstrate that the nucleus does not need to be formed from the protein used in the growth process [179]. A nucleus formed from a fragment of a protein can, in many cases, nucleate fibril formation from the full-length protein (e.g. tau [180], lysozyme [181], β 2 M [182]). This suggests that cleavage of a protein could generate a highly amyloidogenic fragment which nucleates fibril formation of the full-length protein. Nuclei that contain sequences that are very similar (e.g. polyglutamine [183]), differ by only 1 amino acid (e.g. α-synuclein [184], lysozyme [185], protein G BI domain [186], Aβ [187]) or differ significantly (e.g. tau/α-synuclein [188], tubulin/α-Synuclein [189], NAC/Aβ [190], A/β/IAPP [187], Aβ/tau [191], Aβ, TTR fragments, IAPP, silk/serum amyloid A, reviewed in [192]) from the soluble protein have been shown to accelerate fibril formation. Cross-seeding, however, is not necessarily a reciprocal process. A nucleus composed of protein X may be able to seed protein Y, but a nucleus of protein Y may not be able to seed protein X [187, 193]. It is also

15


important to note that there are cases in which a protein may bind to a nucleus but not seed additional fibril formation [194]. The discovery of heterologous seeding has implications for the purity of peptides and proteins used for in vitro experiments designed to understand fibril formation. The batch-to-batch variability of Aβ is well documented and may be a result of different impurities [123]. In some cases, only trace amounts of modified peptide are necessary to seed fibril formation from unmodified protein [195], indicating that experiments need to be performed with exceptionally pure materials to assess mechanistic implications of fibril formation. Furthermore, the protein deposited in the fibrils can also be characterized to determine the extent of modified protein in the amyloid fibrils [196]. The two pieces of evidence necessary to adequately assess the impact of a chemical modification on the kinetics of fibril formation in vivo are the demonstration that the chemical modification is a pre-fibrillogenic event and accelerates nucleus formation. One method to determine if the modification is pre-fibrillogenic in vivo is to determine if the modified protein is present in the circulation. Although this type of experiment is insightful, there are two potential obstacles. First, if modified protein is not observed in the circulation, it is possible that the rate of nucleus formation is so fast that the modified protein is not present in detectable levels. Second, if modified protein is observed in the circulation, it is possible that the protein originated from the dissolution of amyloid fibrils in vivo and the chemical modification was, in fact, a post-fibrillogenic event. Another method to assess if the modification is preor post-fibrillogenic is to examine the distribution of chemically modified protein in the fibrils. If a chemical modification is a post-fibrillogenic event, it is reasonable to assume that the distribution of chemically modified protein in the fibrils will be similar. However, it has been determined that many modifications of Aβ are not equally distributed and have temporal relationship to fibril deposition (see Section 2.2). Finally, the extent of chemically modified protein in the fibrils may provide an insight into whether or not the modification is pre- or post-fibrillogenic. If the chemical modification of a specific residue is a post-fibrillogenic event, over time all of the exposed residues will become modified. Thus, if 50% or more of these residues are modified in aged samples, this may suggest that this residue is surface exposed and may be modified after fibril formation. However, in cases in which only small amounts of modification are present, this chemical modification is either very slow or the residue is in the interior of the fibril. If the residue is in the core of the fibril, chemical modification is likely to be a pre-fibrillogenic event. New methods have recently been explored in which the extent of surface exposed residues of in vitro amyloid fibrils were elucidated by artificial aging of the fibrils [196]. It will be interesting to see if similar methods can be applied to ex vivo fibrils. Another factor that is important to determine the effect of a chemical modification on the kinetics of fibril formation is the extent to which the modification accelerates fibril formation. In vitro studies have been used to determine that many chemical modifications of peptides lead to enhanced rates of fibril formation compared to wild-type as monitored by Thioflavin T fluorescence or Congo red binding. These rates are generally assessed by comparing equal concentrations of modified and

4 Conclusions

wild-type peptides. These results, however, need to be examined in the context of the relative concentrations of modified and wild-type peptides in vivo. For example, Met oxidation of Aβ has been observed in vivo and in vitro and there is evidence that this peptide aggregates more slowly than Aβ in vitro (see Section 2.3). From this information, it is tempting to conclude that Met oxidation of Aβ has a protective effect by inhibiting Aβ fibril formation. However, if the Met-oxidized Aβ peptide is turned over at a significantly slower rate than Aβ, the Met-oxidized peptide could accumulate and nucleate fibril formation. Pre- or post-fibrillogenic chemical modifications can play a role in amyloid fibril formation by altering the thermodynamic stability of the fibrils or the cytotoxicity. The stability can be enhanced by nucleating a different packing arrangement, the formation of covalent crosslinks (e.g. disulfides, AGEs, dityrosine), or decreasing the ability of the immune system to recognize or degrade the fibrils. One method to determine if the chemical modification can nucleate a different packing arrangement is to examine the fibril morphology. Phosphorylated tau, for example, results in fibrils that more closely resemble authentic PHFs than nonphosphorylated tau [170]. Fibrils can be examined for thermostability by comparing the measured solubility of modified and non-modified fibrils upon addition of denaturant. Changes in thermodynamic stability and cytotoxicity could be closely related. Fibrils and small oligomers both appear cytotoxic, but oligomers may be able to diffuse through tissue and, therefore, be toxic to more cells. Therefore, increasing the thermodynamic stability of the fibrils may be both bad and good – bad because it will make the fibrils difficult to clear from the body, good because it may shift the equilibrium toward the fibrils which could decrease the production of cytotoxic oligomers.

4 Conclusions

The majority of amyloid deposits examined to date contain some protein that is post-translationally modified. The role of these chemical modifications in amyloid fibril formation, however, has remained underexplored. A major challenge for the future will be to establish a temporal relationship between chemical modifications and fibril formation, and to assess the impact of the modification on kinetics of fibril formation, thermodynamic stability and cytotoxicity. A clear understanding of the role of chemical modifications in amyloid fibril formation will not only expand our current knowledge of protein misfolding, but may also lead to new avenues for therapeutic intervention. One current strategy to inhibit amyloid fibril formation is to design molecules that can stabilize the native state of the protein. This strategy will also be applicable for cases in which chemical modification plays a role because modifications are less likely in regions of a protein that are not flexible. However, a new strategy to prevent amyloid fibril formation may be to inhibit aberrant protein chemical modifications.

17


5 Acknowledgments

I am grateful to Daniel Moriarty for a critical review of this manuscript, and to Carole Klapper and Randall Morrison for technical assistance in manuscript preparation.

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1

Serum amyloid P Component Steve P. Wood, and Alun R. Coker University of Southampton, Southampton, United Kingdom


1 Introduction

The characteristic amyloid fibril-forming polypeptides and the amyloid P component (AP) are abundant constituents of amyloid deposits, and a growing body of evidence suggests that their interaction is an important feature of amyloid accumulation. When the mouse gene for serum amyloid P component (SAP), the circulating equivalent of AP, is inactivated or knocked-out, then the time course for deposition of amyloid after experimental induction is considerably retarded [1]. This strongly suggests that SAP is centrally involved in some way in fibrillogenesis and/or in stabilizing deposits against clearance. A recently discovered drug, CPHPC, induces rapid clearance of circulating SAP in transgenic mice expressing the human protein, and leads to a reduction in the amount of amyloid associated SAP and to a reduction in the amyloid load in animals where systemic amyloidosis has been induced [2]. These results suggest that SAP contributes to the stability and persistence of fibrils in vivo. Amyloid fibril formation and the calcium-dependent binding of SAP to the fibrils can be reproduced in vitro, and, in these in vitro conditions, it was found that bound SAP protects the fibrils against proteolysis and attack by phagocytic cells, reinforcing the stabilization hypothesis [3]. Physical/chemical stabilization of amyloid fibrils might also result from SAP binding. Amyloid formation in vitro is often driven by harsh protein destabilizing conditions and the process appears to be essentially irreversible. However, regression of deposits in vivo following depletion of circulating fibril precursor pools by immunization [4] or transplantation [5] suggests that they may be more dynamic structures. SAP binding might contribute to “locking in” materials to the deposit. However, the administration of radiolabeled SAP and whole-body scintigraphy, an established and effective procedure for locating amyloid laden organs, depends upon exchange


2 Serum amyloid P Component

of fibril-localized SAP with the circulating tracer pool and is further evidence of a dynamic aspect of amyloid deposits [6]. Although the interaction of SAP with amyloid fibrils appears to be a fundamental feature of amyloid disease pathology, the whole story of amyloid structure is likely to be far more complex. A host of other components are found in association with amyloid deposits including the complement protein C1q, α 1 -antichymotrypsin, apolipoprotein E, ubiquitin and glycosaminoglycans [7]. Their role is far from clear. There is evidence that SAP interacts with some of these constituents in other contexts, such as complement activation and basement membrane structure [8]. Some of the components may well be present in response to the local inflammatory cellular crisis at sites of amyloid deposition. However, the ability to reproduce amyloid recognition by SAP in vitro from pure components suggests that investigation of the interaction at a structural level is both a tractable and valid target. Here, we review what is known of the structure and properties of SAP, and how this is related to current views on the structure of amyloid and its formation, topics covered in greater depth elsewhere in this volume.

2 Amyloid Fibrils and their Formation

For many years, our understanding of fibril structure was limited to the interpretation of X-ray fibre diffraction patterns that reported the presence of cross-β structure. Technical advances in the use of synchrotron radiation for these investigations, the development of cryopreservation techniques and image analysis for electron microscopy and the use of multidimensional nuclear magnetic resonance (NMR) and more recently solid-state NMR are providing a clearer view of amyloid structure and potential motifs for SAP binding [9–13]. Amyloid fibrils are believed to be formed by a concentration dependent, nucleated growth pattern analogous to crystal growth [9]. Structurally perturbed precursors form stable intermolecular hydrogen bonding networks, perhaps involving domain swapping in some cases [14], that, together with any retained β-structure of the precursor, form the extended cross-β structure of the fibril. Strands are believed to run approximately normal to the fibril axis. This regular cross-β structure gives rise to characteristic reflections in the X-ray fibril diffraction pattern [15]. Fibril growth is probably far more complex than sequential monomer addition, as electron microscopy and atomic force microscopy have shown the presence of micelle-like components and beaded protofilaments during growth [16, 17]. Mature fibrils are often assembled as “rope-like” structures of entwined filaments of variable helical pitch and a hollow core [18] (Fig. 1a). Fibrils can be very long, but are usually in the range of 100–150 Å in diameter. The structural destabilization of precursors required to drive fibril growth may not always be extensive. It is clear that a number of mutated proteins associated with aggressive amyloid deposition exhibit reduced thermal stability [19, 20], and experimental conditions that promote fibril growth in vitro lead to partial denaturation and specific strand displacements from native folds.

2 Amyloid Fibrils and their Formation

Fig. 1 (a) A model for ATTR derived from deuterium-exchange protection NMR and electron spin resonance data. Displacement of strands C and D from the native fold creates newly exposed A and B edge strands that together with a slightly modified form of the native intersubunit sheet provides a continuum of $-strands. Disordered loops

and chain termini protrude from the fibril surface. (Reproduced from Oloffson et al. [27], with permission.) (b) Image processed electron microscopy images of insulin amyloid fibrils showing various degrees of intertwining of protofibrils. (Reproduced from Saibil et al. [10], with permission.)

However, models for transthyretin amyloid (ATTR) that fit with much experimental evidence incorporate a good deal of the native β structure [21–24]. Crystal structures of an amyloidogenic TTR mutant and of a κ-light chain variable region show extended β-helix arrangements, generated by the crystal symmetry, that emulate many of the features expected for amyloid fibrils [25, 26]. Recent solution NMR and mass spectrometry studies for TTR, Aβ and β 2 -microglobulin (β 2 M), using deuterium-exchange protection methods, suggest that considerable portions of the polypeptide chain, including chain termini and displaced loops, are probably disordered at the fibril surface providing an interaction surface distinct from that of the native proteins (Fig. 1b) [27–29]. Monoclonal antibodies recognizing cryptic epitopes of TTR that are displayed by amyloid fibrils and by hyper-amyloidogenic mutants in solution, but not the native protein, reaffirm this view [30]. Solidstate NMR studies of Aβ1–40 and fibril proteolysis results suggest that the polar

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N-terminal region and turns are exposed at the surface of the fibril [31, 32] and that the N-terminal region is a primary epitope recognized by antibodies to Aβ fibrils [33]. The hydrogen bonding potential of the β-strands is likely to be fully satisfied by the cross-β structure, except at the fibril ends, and hydrophobic residues are probably buried, if indeed the fibril state represents a low-energy “primordial fold” [34]. Although many proteins, including those with no association with amyloid disease, can be driven to the fibril state by harsh destabilizing conditions, in general, protein structures have probably evolved with variable degrees of success to avoid exposure of sheet edge strands capable of repeating intermolecular interaction in physiological conditions [35]. On the order of 20–30 distinct proteins have been reported to produce amyloid deposits in vivo. Therefore, the universal calcium dependent binding of SAP to these fibrils is unlikely to be highly sequence specific, in view of the variety of the fibrillogenic proteins. Evidence for the existence of a generic amyloid motif, however, is provided by the identification of an antibody that recognizes amyloid derived from different source proteins [36]. In both cases, it is possible that there exists a conformation-dependent discontinuous epitope that obscures identification of important conserved patterns of residues. If the emerging structural models of amyloid fibrils formed by TTR, β 2 M, Aβ1–40 are general in character, then we might expect the surface of the amyloid fibril to display turns between strands, displaced loops and flexible chain termini. These features are likely to be distributed in accord with the twist of the sheets and the inter-twining of the fibrils to produce a fuzz of polypeptide at the fibril surface within which motifs for SAP recognition must reside.

3 The Structure of SAP

The crystal structure of human SAP was first reported in 1994 and was determined by the classical method of isomorphous replacement [37]. Crystallization of the protein had been problematic due to its propensity to aggregate in the presence of calcium ions, a phenomenon that has continued to complicate many aspects of research into the behavior of SAP in solution [38]. SAP is a normal, relatively abundant (around 30 mg/l) plasma glycoprotein that is readily purified by virtue of its calcium-dependent affinity for immobilized phosphoethanolamine. Subsequent experimental manipulations are traditionally carried out in neutral buffers containing EDTA, where the protein exhibits the behavior of a highly soluble decamer of Mr 23,500 subunits. Each subunit comprises 204 amino acids and carries a single Asn32-linked biantennary oligosaccharide chain [39]. The isoelectric point is 5.5, giving the protein a net negative charge at physiological pH; however, this charge is not evenly distributed. Crystallization was carried out in the presence of high concentrations of calcium and acetate ions and at a pH close to the pI.

3 The Structure of SAP

Fig. 2 The SAP pentamer viewed down the 5-fold symmetry axis showing two calcium ions (yellow) bound to each subunit.

The crystallographic structure revealed molecules with cyclic pentameric symmetry packed in a herring-bone fashion and containing only five subunits. These disc-like pentameric molecules are approximately 100 Å in diameter and 35 Å deep, and show a large 20 Å diameter hole through the center, in accord with previous negative stained electron microscopy (Fig. 2) [40]. The polypeptide chain in each subunit is organized as two layers of antiparallel β-strands, sandwiching hydrophobic residues of the core between them (Fig. 3). The first and last four strands sequentially visit the two layers in a “jelly-roll” topology, while the intervening six strands define two β-meanders of three strands at the same end of each layer. This architecture very closely resembles that of the plant lectins, concanavalin A and pea lectin [41, 42]. One of the sheets is flat and has a 10-residue α-helix positioned upon it. The other sheet is rather twisted and loops dipping into the concave surface contribute to the double calcium-binding site of each subunit. The loops connecting strands are generally short and this likely contributes to the remarkable stability of calcium bound SAP in the face of proteolytic digestion [43]. The amino acid sequence of SAP is provided in Fig. 4 in comparison with that of C-reactive protein

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Fig. 3 Ribbon diagrams of orthogonal views of the SAP subunit showing the location of the A-face helix and the concavity of the B-face sheet.

Fig. 4 Amino acid sequences for SAP and CRP with residues involved in calcium binding in red, residues of the hydrophobic pocket bound by MO$DG and CPHPC in blue. The oligosaccharide attachment site (CHO) and site of chymotrypsin cleavage are indicated, as is the distribution of secondary structure elements.

4 The Calcium-binding Site

Fig. 5 Side view of the SAP pentamer showing the 10 bound calcium ions of the B face and five helices of the A face.

(CRP), annotated with secondary structure features and with important residues in color. All of the strands of each subunit approximately populate a plane normal to the 5-fold axis of the pentamer. The subunits interact via van der Waals contacts and hydrogen bonds through the open-core end of the β-sheet sandwich of one subunit and the N- and C-terminal strands of the adjacent subunit, to result in the burying of 20% of the protomer surface on pentamer formation. The pentamer has a polar character, with five α-helices on one face (the A face) and five double calcium sites on the other (B face) (Fig. 5). Only the first ring of the oligosaccharide chain attached to Asn32 was visible in this and several subsequent crystal structure analysis. However, more recently, we have experimentally observed almost the entire length of a single sugar chain for one subunit (unpublished data). In this case the sugar chain was immobilized by crystal packing. If the oligosaccharide chains are fully extended, as suggested by solution X-ray and neutron scattering [44], then their area of influence around the pentamer is considerable, extending the effective molecular diameter from 100 to around 135 Å (Fig. 6)

4 The Calcium-binding Site

Each SAP subunit in the crystal structure contains two calcium ions bound very close together (4.2 Å) in the concavity formed by the buckled B-face sheet. This “tense” electrostatic arrangement is stabilized by a selection of carboxylate and amide protein side-chain ligands to the metals (Fig. 7). Unlike many other calciumbinding proteins, these side-chains come from different parts of the sequence and so it seems likely that the bound metals are important for stabilizing the structure in this region. For instance, Glu136 and Asp138 are positioned on an extended loop that dips into the site, with both side-chains forming bridging interactions with both calcium ions. The two metal ions have different total numbers of stabilizing ligands and are likely to be bound to the protein with quite different affinities. Furthermore,

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Fig. 6 SAP pentamer with fully extended oligosaccharide chains attached to each subunit, extending the effective radius of the molecule by around 40%.

the ligand complement to the metals in this crystal form also included a bound acetate ion in four of the five subunits and an intermolecular interaction involving Glu167 from the prominent A-face helix of an adjacent pentamer. Acetate ions and other small molecules that bind this site inhibit the aggregation of SAP by blocking intermolecular interactions. There are many acidic groups exposed on the surface

Fig. 7 The double calcium ion-binding site of SAP, showing the uneven distribution of amino acid side-chain ligands to the metals and a bound acetate ion.

5 Comparative studies of CRP

of the pentamer that could contribute to intermolecular contacts, but the Glu167 side-chain is of special interest as five copies project from the A face and could simultaneously interact with the five double calcium sites of another pentamer stacked on the same 5-fold symmetry axis. Such stacks would also be expected to be rather stable because the net charges on the contact surfaces are opposite. Limited stacks have been observed on the surface of ex vivo amyloid fibrils by electron microscopy [45], but the most impressive long stacks are reported in the presence of EDTA where, in the absence of calcium, the above stacking mechanism would not be expected to occur [46]. The presence of long strings of SAP pentamers in calcium-free solution seems unlikely as the protein is very soluble, monodisperse and generally well behaved in these conditions. It is possible that stack formation in the absence of calcium is the result of negative stain metal binding during sample preparation. Stack formation is a potentially important property of SAP as it might contribute to SAP accumulation at amyloid sites. Certainly the related troublesome solubility properties of SAP have provided major technical complications for studies of SAP binding to putative target macromolecules. Calcium site I of each SAP subunit is coordinated to the side-chains of Asp58, Asn59, Glu136, Asp138 and the main chain carbonyl of Gln137. The acetate ion provides the seventh ligand to produce a pentagonal bipyramidal arrangement. Calcium site II is more open and has fewer protein ligands deriving from Glu136, Asp138 and Gln148 as well as the acetate ion and two water molecules. The site was found to release calcium easily, by washing crystals in calcium-free buffers, and indeed preferentially bound the much larger cerium ion. It is conceivable that metals larger than calcium, such as copper or zinc, might be preferred in site II in spite of their much lower availability in biological fluids. The integrity of the site probably also depends upon the presence of a bound ligand analogous to the acetate ion seen in the crystal, but no such ligand has been identified in SAP preparations. In the absence of bound metal ions SAP is readily cleaved by chymotrypsin at the 144–145 bond, suggesting that the loop carrying these residues, which is an integral part of the metal binding site, is much more accessible. The degree of protection conferred by calcium is concentration dependent [43]. Protection is complete at 5 mM and absent at 0.1 mM calcium. Protection is not entirely complete at 2 mM calcium and 35 mg/l SAP, conditions close to those expected in vivo.

5 Comparative studies of CRP

CRP is a close sequence and structural homologue of SAP and investigations of its structure and properties provide some intriguing comparisons with SAP, since CRP is not bound by amyloid fibrils. A number of crystal structures of CRP have been determined in the presence and absence of metal ions and ligands [47–50]. These show that the subunit fold of CRP is very similar to SAP. Two closely bound calcium ions reside on each subunit of a pentamer. The distribution of protein ligands to the two calcium ions is much more even in CRP and the metals are held with equal affinity [51]. In structures where no calcium ions are bound, the 140–150

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loop region is no longer well ordered, and, in solution, the 145–146 and 146–147 peptide bonds are readily cleaved by proteolytic enzymes. Although no structure for metal-free SAP has been determined, it seems highly probable that an equivalent disorganization of the metal binding region occurs. The organization of the pentamer of CRP is somewhat different from the SAP pentamer. Each subunit is tilted by 22◦ about an axis through its centre of mass towards the 5-fold axis of the pentamer such that the A-face helices are 5 Å closer to the axis while the five double calcium sites are displaced by an equivalent amount away from the axis. This reorganization may explain why CRP is not as susceptible to calcium-dependent aggregation and to stacking interactions as SAP, because the putative interacting groups are displaced to new pentamer radii differing by 10 Å. Perhaps it also contributes to the inability of CRP to bind motifs with a particular geometry at the surface of amyloid fibrils that are recognized by SAP. When an equivalent subunit rotation is modeled onto the SAP structure, surprisingly few steric clashes occur, suggesting that such rotations could be a dynamic feature of pentraxin structures. The crystal structure of CRP in the complete absence of calcium ions shows two pentamers packed A face to A face on a common 5-fold axis, bringing the helices on each subunit close together. Although there is no evidence for the formation of CRP decamers in solution, this does illustrate a preferred packing arrangement for pentamers of CRP, albeit driven by the supersaturation of the crystallization cocktail.

6 SAP Structure in the Absence of Calcium

In all of the biological contexts in which SAP structure informs function, it is likely that metal ions are present and, thus, metal binding is only likely to be disrupted by proteolytic clipping of the 140–150 loop. In this respect, the decameric structure adopted by SAP in metal-free conditions appears to be of limited interest. However, in many investigations of the possible role of SAP in influencing fibril formation, sugar binding or protein folding (that we will visit later), metal-free conditions have often been employed. The value of observations made in these conditions is that they report on a functionality of the protein that persists when the metal site is severely disrupted. This might be attributed to a completely different part of the protein or to generation of a novel functionality for the disrupted region. The decameric form of SAP that exists in solution in the absence of bound metals very likely involves a face-to-face interaction. The susceptibility of these decamers to proteolysis in the calcium site loop suggests that these loops are exposed and that the pentamers are interacting via their A faces, as they do in crystals of metal-free CRP. Binding of calcium however generates SAP pentamers. It is not clear how the ordering of the metal site would destabilize an A–A-face interaction some 35 Å away unless perhaps subunit rotation equivalent to that of CRP took place. It is conceivable that the SAP decamer involves interactions between pentamer B faces.

7 Binding of Small Molecule Ligands to SAP 11

The loop site of proteolysis is oriented to the outer edge of the pentamer and may not be completely obscured by a B–B-face interaction. Metal binding and ordering of the loop could disrupt a B–B-face decamer. Determination of the structure of metal-free SAP will clearly be informative.

7 Binding of Small Molecule Ligands to SAP

The calcium dependent binding of phosphoethanolamine (PE) and the 4,6-cyclic pyruvate acetal of β-D-galactose (MOβDG) to SAP are well-established observations [8]. Immobilized PE, linked to a matrix through the amino group, and MOβDG as a component of agarose are both effective affinity supports binding SAP from solutions containing calcium ions. The binding sites for the isolated ligands were identified by soaking SAP crystals in the compounds or by co-crystallization and the determination of crystal structures. Subsequently, both studies have been extended to higher resolution [52]. Both compounds were found to bind to the double calcium site of SAP via their acidic moieties. The structure of bound MOβDG was of special interest, as this compound has been shown to inhibit the binding of SAP to amyloid fibrils in vitro [53], and, therefore, has potential both as a lead compound for drug design and as a guide to the nature of the amyloid interaction. The carboxylate of MOβDG interacts with the calcium ions of each SAP subunit and the bridge oxygens of the carboxyethylidine ring hydrogen bond with the side-chain amide nitrogens of Asn59 and Gln148 (Fig. 8). The side-chain carbonyl oxygens of the same residues also interact with the calcium ions. There are also hydrogen bonds between sugar oxygens of C1 and C3 to Gln148 and Lys79, respectively. The methyl group of the bound R isomer of MOβDG locates in a hydrophobic pocket formed by Tyr74, Phe64 and Leu62. In one subunit, crystal packing leaves no room for a molecule of MOβDG to bind and only one calcium ion is bound in site I. This suggests that a bound ligand of some sort is required to stabilize the filling of calcium site II. Isothermal calorimetry measurements indicate a dissociation constant of 50 µM for the interaction of MOβDG with SAP [2], considerably (around 103 -fold) weaker than that with amyloid fibrils. This may be due in large part to the likelihood that multiple attachment points and some associated cooperativity are involved in amyloid recognition. A tight β-turn between polypeptide strands and including aspartic acid at its apex would present a broadly similar array of binding groups, as MOβDG and such turns can be readily built into the binding site, an observation consistent with the supposition that exposed turns at the fibril surface are involved in SAP binding [8]. However, MOβDG would also be expected to interfere with the formation of SAP stacks. Inhibition of stack formation could account for much of the observed inhibition of SAP binding to fibrils in the presence of MOβDG and calcium ions. The importance of cooperative binding effects was demonstrated particularly well for the first time in studies of the calcium-dependent binding of dAMP to SAP


Fig. 8 The structure of the galactose analog (MO$DG) bound into the double calcium site of an SAP subunit, showing direct interaction with the calcium ions and hydrogen bonds to side-chains that also donate ligands to the metals.

[54]. This ligand was identified as an effective inhibitor of SAP aggregation in the presence of calcium ions. The crystal structure of the SAP–dAMP complex revealed decamers of SAP. Each subunit bound a single dAMP molecule via its phosphate group to the double calcium site. The two pentamers were arranged B face to B face, but the dominant interaction stabilizing the complex was the non-covalent stacking of the adenine rings. This complex was stable during gel filtration even though the extent of the interactions at one double calcium site suggests a low-affinity binding, in the same range as MOβDG. High-throughput screening approaches have identified a high-affinity ligand for SAP that clearly reinforces the importance of cooperative binding [2]. The screen selected compounds that inhibited binding of SAP to Aβ1–42 fibrils immobilized on a plastic surface. This approach identified CPHPC, a compound comprising two D-proline residues linked via a five-carbon spacer and that binds SAP with nanomolar affinity. The crystal structure of the SAP–CPHPC complex showed a decameric structure in which the two D-proline head groups of five molecules of the drug bound to the double calcium sites of subunits from two pentamers oriented B face to B face on a common 5-fold axis (Fig. 9). These decamers are stable in solution. The packing of the proline ring into the Leu62, Tyr74, Phe64 hydrophobic pocket and the interaction of the carboxylate with the calcium ions are the major

7 Binding of Small Molecule Ligands to SAP 13

Fig. 9 Two pentamers of SAP ($-strands in green and heices in red) crosslinked by five molecules of the drug CPHPC (left).

stabilizing interactions. Binding of d-proline alone to SAP, where cooperative effects are not involved, is about 103 -fold weaker, although the crystal structure of the N-acetyl D-Pro–SAP complex shows an identical set of interactions to the CPHPC head group complex. As noted earlier, the properties of this compound potentially provide an approach to treating amyloidoses. Although of considerable interest in a clinical context, it is not clear that the discovery of this compound directly aids our understanding of amyloid recognition. Multiple peptide chain C-termini at the fibril surface might bind SAP in a similar fashion to CPHPC and the side-chains of certain hydrophobic amino acids might fill the hydrophobic pocket, but they are unlikely to achieve the particularly snug fit observed with D-proline. L-proline binds considerably less tightly. During studies of the chaperone activity of SAP, we discovered that high concentrations of chloride ions can effectively inhibit the aggregation of SAP in the presence of calcium ions. X-ray structure analysis of crystals grown in these conditions revealed that a single chloride ion can bind close to the calcium ions of each SAP subunit. Although the binding is weak, at sufficiently high concentrations it is capable of blocking the intermolecular interactions that lead to SAP precipitation with calcium ions. This provides a convenient solution condition for comparative investigations of the calcium dependent binding of SAP to other macromolecules (unpublished data). In overview, structural investigations have defined the calcium-binding site of SAP in considerable detail. The most direct interpretation of the calcium dependence of fibril recognition by SAP suggests that this is the site involved although it is conceivable that the structural integrity of the site is required without direct metal


bonding interactions as seen for sugar binding to structurally homologous lectins. The structural features of the binding of small molecule ligands provide clues to the nature of the binding mode of polypeptide ligands likely presented at the fibril surface. It is not clear how the architecture of the amyloid fibril might complement the 5-fold symmetric pattern of binding sites on SAP, although it seems that a multipoint attachment is required to explain the binding affinity observed for such a shallow recognition pocket. It is conceivable that the array of loops, turns and termini at the fibril surface satisfy the multipoint cooperative attachment by virtue of their density rather than their precise organization or sequence. While we have detailed structural information for SAP and its likely amyloid recognition site, and a much clearer view of the structure of fibrils, difficult issues remain in bringing this information together into a coherent hypothesis for the form of decorated fibrils in vivo. The possible motifs for amyloid fibril recognition by SAP discussed above are likely to occur with high frequency due to the tight twisting together of protofilaments in mature fibrils and the relatively large number of repeats of the generally small protein constituents of most amyloid fibrils. A 150Å diameter fibril could bind three SAP pentamers through a face interaction for each 140 Å of its length to fully coat the surface. However, the molar binding ratio of SAP pentamers to synthetic Aβ1–40 fibrils reaches a plateau at around 1:250, corresponding to around 12% of the mass of the fibril – roughly comparable to that found in in vivo amyloid fibrils [3]. Based on calculations of the average mass per unit length of mature synthetic Aβ1–40 fibrils [16], this would only give one SAP molecule every 370 Å along the fibril (Fig. 10). This low binding ratio of SAP to Aβ1–40 suggests that the amyloid fibrils in this model would be rather sparsely coated with SAP. For a fully coated surface providing maximum stabilization, the molar ratios would drop to around 1:20. An explanation for this might reside in the work of MacRaild [55]. Simultaneous attachment of SAP to more than one fibril, crosslinking and generating fibril entanglement might limit the accessibility of potential fibril ligands to further molecules of SAP and provide stabilization and resistance towards proteolytic attack.

Fig. 10 Glycosylated SAP pentamers (around 140 Å diameter) attached through a B-face interaction with a 150-Å diameter fibril of A$ would be around 370 Å apart if SAP made up around 12% of the mass of the complex.

9 SAP, Protein Folding and Amyloid Fibril Formation

8 The Role of Glycosaminoglycans (GAGs)

Heparan sulfate and chondroitin sulfate proteoglycans (HSPG and CSPG) are well-documented constituents of amyloid deposits. In vitro experiments show that SAP interacts in a calcium-dependent fashion with dermatan and heparan sulfate polymers [56]. Kiselevsky et al. [57] have proposed on the basis of in situ electron microscopy that this interaction forms the foundation for the structure of amyloid fibrils in vivo, and that disruptive extraction procedures may explain discrepancies between this and other work. Their model suggests that stacked SAP molecules are coated by a coil of CSPG on top of which lie HSPG chains and surface-exposed 1-nm protofilaments. During water extraction these filaments are freed from the fibril and assemble into the familiar pattern of twisted filaments observed by others and reproduced in in vitro experiments. While all aspects of this model are not in accord with the views of many investigators, it is conceivable that the SAP stacks observed attached to ex vivo fibrils is a relic of a GAG bound structure that has survived fibril extraction from tissues. It is less easy to accept the proposed presence of single strand protofilaments enmeshed in the HSPG layer as a dominant form. TTR fibrils removed from aqueous humor with minimal disturbance clearly show the classical cross-β X-ray fibril diffraction pattern [10] and lend strong support to the concept that in vivo fibril assembly bears strong similarities to that in vitro.

9 SAP, Protein Folding and Amyloid Fibril Formation

SAP recognizes amyloid fibrils formed from a diverse array of source proteins and polypeptides, but there is no evidence to suggest that SAP has any preferential binding to these proteins in their native state. SAP has been reported to bind non-fibrillar β 2 M, serum amyloid A protein (SAA) and Aβ, but the proteins were adsorbed to a plastic surface where some unfolding might be expected [58, 59]. Experiments designed to monitor the effects of SAP on fibril formation and protein folding are complicated by the calcium-induced aggregation of SAP and the technical difficulties arising from the need to employ harsh conditions to initiate fibril growth for some proteins. Synthetic Aβ has often been employed, as fibril growth proceeds spontaneously in dilute aqueous media. Amyloidogenic polypeptides like Aβ and SAA are likely to be metastable in aqueous solution, as their sequences are adapted to quite different environments. SAA preferentially associates with a high-density lipoprotein particle, while Aβ is a fragment of a larger protein and at least a portion of its sequence has evolved to reside within a lipid bilayer. Both materials have poor solubility and are rather difficult to work with. Precise solvent conditioning protocols are recommended by some investigators [29] to ensure reproducible behavior of synthetic Aβ, but these are not followed by all and this, together with the solubility difficulties of SAP, may go some way towards explaining diverging views on the effects of SAP on Aβ fibrillogenesis.

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SAP has been reported to both enhance and to inhibit fibril formation by Aβ in different experimental conditions. Hamazaki [60] reported that SAP enhanced the production of sedimentable Aβ1–40 approximately 3-fold over 16 h in the presence of 1.5 mM calcium ions. Webster and Rodgers [61] reported a doubling of the Thioflavin T fluorescence for Aβ1–42 in the presence of SAP and 2 mM calcium ions over 24 h. In both cases, nanomolar concentrations of Aβ and SAP were employed, approximating to in vivo levels. Janciauskine et al. [62] found that in the absence of calcium ions, SAP completely inhibited the formation of fibrils by Aβ1–42. More recently, MacRaild et al. [55] have investigated the effects of SAP and apolipoprotein E on the state of amyloid fibrils using sedimentation velocity analysis, electron microscopy and rheology. They concluded that both proteins cause amyloid fibrils to associate more densely, becoming highly entangled. The additional stabilization of fibrils provided by calcium-dependent binding and/or crosslinking by SAP would be expected to favor fibril formation. Inhibition of fibril formation, however, suggests a different mode of interaction. There is considerable current interest in identifying small molecules that interfere with extension of the “crossed-β” structure [63–65] and peptide strand terminators offer a plausible guide to the way in which SAP might halt fibril growth. The disordered region made up of residues 140–150 is a unique feature of metal-free SAP and could contribute to this effect, donating a strand that cannot be propagated.

10 Conclusion

The view that some disorganization of amyloidogenic proteins is required for them to restructure into a cross-β fold is well supported by experimental observations and the amyloidoses are often referred to as protein misfolding diseases [66]. The ability of SAP to recognize intermediate and end products of this process shows close parallels with the properties of molecular chaperones. These proteins interact with folding intermediates of other proteins in the cell to prevent aggregation and facilitate efficient production of correctly folded and biologically active materials [67, 68]. SAP has been shown to assist the refolding of chemically denatured lactate dehydrogenase (LDH) and to inhibit the turbulence induced inactivation of the same enzyme [69]. These effects were observed in the absence and presence of calcium ions, with the greatest effect being observed in the presence of the galactose analogue, MOβDG, which blocks amyloid binding. They are not inhibited by the presence of dAMP, which stabilizes a B–B-face interaction, suggesting that some other part of the SAP molecule is involved. The effects are saturable, possibly reflecting partitioning of refolding LDH intermediates between states that bind SAP and others that do not. Only a small proportion of molecules negotiate a direct route through the refolding energy landscape [70] in the absence of SAP, with the majority forming a misfolded and inactive product. SAP may bind misfolding intermediates that would otherwise inactivate correctly folded subunits of the dimeric enzyme and in the case of the turbulence inactivation assay, SAP may inhibit a nucleated

References

aggregation process of inactivation analogous to amyloid formation. One might speculate that these effects of SAP reflect a surveillance role in detecting damaged proteins in vivo, which is overwhelmed in amyloid disease. A selection of other intracellular molecular chaperones have been identified by proteomic analysis of senile plaques, levels of heat-shock protein (Hsp) 90 being particularly elevated [7]. The extracellular chaperone clusterin is also found. This protein has been shown to bind tightly to Aβ and inhibit its aggregation and to inhibit the formation of amyloid fibrils by apolipoprotein CII in vitro [71, 72], effects reminiscent of those exhibited by calcium-free SAP on fibrillogenesis. SAP binding to amyloid may be a pathological accident, perhaps a side-reaction to some other more general chaperone-like function, or SAP may have evolved as a safety mechanism to trap/localize misfolded proteins. A core role of SAP appears to be as an anti-opsonin in chromatin clearance and innate immunity [73] and masking the low level amyloid deposition that appears to be a common feature of ageing [74] may be a further element of this process.

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13 ANTZUTKIN, O. N., J. J. BALBACH and R. TYCKO. Site-specific identification of non-beta-strand conformations in Alzheimer’s beta-amyloid fibrils by solid-state NMR. Biophys J 2003, 84, 3326–3335. 14 STANIFORTH, R. A., et al. Three-dimensional domain swapping in the folded and molten-globule states of cystatins, an amyloid-forming structural superfamily. EMBO J 2001, 20, 4774–4781. 15 SERPELL, L. C. Alzheimer’s amyloid fibrils: structure and assembly. Biochim Biophys Acta 2000, 1502, 16–30. 16 GOLDSBURY, C. S., S. WIRTZ, S. A. MULLER, S. SUNDERJI, P. WICKI, U. AEBI and P. FREY. Studies on the in vitro assembly of A beta 1–40: implications for the search for A beta fibril formation inhibitors. J Struct Biol 2000, 130, 217–231. 17 IONESCU-ZANETTI, C., et al. Monitoring the assembly of Ig light-chain amyloid fibrils by atomic force microscopy. Proc Natl Acad Sci USA 1999, 96, 13175–13179. 18 JIMENEZ, J. L., E. J. NETTLETON, M. BOUCHARD, C. V. ROBINSON, C. M. DOBSON and H. R. SAIBIL. The protofilament structure of insulin amyloid fibrils. Proc Natl Acad Sci USA 2002, 99, 9196–9201. 19 BOOTH, D. R., et al. Instability, unfolding and aggregation of human lysozyme variants underlying amyloid fibrillogenesis. Nature 1997, 385, 787–793. 20 FERRAO-GONZALES, A. D., L. PALMIERI, M. VALORY, J. L. SILVA, H. LASHUEL, J. W. KELLY and D. FOGUEL. Hydration and packing are crucial to amyloidogenesis as revealed by pressure studies on transthyretin variants that either protect or worsen amyloid disease. J Mol Biol 2003, 328, 963–974. 21 INOUYE, H., F. S. DOMINGUES, A. M. DAMAS, M. J. SARAIVA, E. LUNDGREN, O. SANDGREN and D. A. KIRSCHNER. Analysis of X-ray diffraction patterns from amyloid of biopsied vitreous humor and kidney of transthyretin (TTR) Met30

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1

Amyloid Fibril Formation of Natively Unfolded Proteins Vladimir N. Uversky Indiana University School of Medicine, Indianapolis, USA

Anthony L. Fink University of California, Santa Cruz, USA


1 Introduction

The realization that many proteins are intrinsically disordered, or natively unfolded, under physiological conditions (for the purified protein) is a relatively recent one. In fact, current estimates suggest that between 25 and 33% of the human genome encoding proteins may code for intrinsically disordered regions [1–4]! Thus, rather than being rarities, such proteins may be rather common. Several hypotheses have been proposed for the existence of natively unfolded proteins; probably the most widely accepted is that the lack of ordered structure permits greater flexibility in binding partners, a valuable characteristic in a protein that must react with multiple binding partners in its normal function, as is true for many signaling molecules and gene regulators. It is likely that in their normal physiological milieu intrinsically unstructured proteins are not unfolded, due to interactions with other proteins, nucleic acids, membranes or small ligands. This would, at least in part, explain their stability to intracellular proteases. Natively unfolded, or intrinsically disordered, proteins are specifically localized within a unique region of chargehydrophobicity phase space, and are characterized by a combination of low overall hydrophobicity and large net charge [5]. As will be discussed, a variety of conditions can bring about folding of natively unfolded proteins, resulting in either relatively non-compact, partially folded conformations or relatively compact, tightly folded conformations. An informative example is the effect of increasing trimethylamine N-oxide (TMAO) on α-synuclein. TMAO is an osmolyte, and is known to stabilize native states and induce structure in disordered states. When the structure of α-synuclein was examined as a function


2 Amyloid Fibril Formation of Natively Unfolded Proteins

of increasing TMAO, the initially natively unfolded conformation adopted a partially folded intermediate conformation in the vicinity of 1 M TMAO, and became compact and helical in 3 M TMAO. The term conformational disease or protein deposition disease has been given to a number of pathological states in which a specific protein or protein fragment changes from its natural soluble form into stable, insoluble, ordered filamentous protein aggregates, commonly referred to as amyloid fibrils [6–11]. Given the prevalence of natively unfolded proteins it should not be surprising that of the more than two dozen proteins known so far to be involved in protein deposition diseases, more than 50% are either completely unfolded or contain significant unfolded regions under physiological conditions (Tables 1 and 2). For many years it has been generally assumed that the ability to form amyloid fibrils is limited to a relatively small number of proteins, essentially those found in the diseases, and that these proteins possess specific sequence motifs encoding the unique structure of amyloid core. However, recent studies have suggested that the ability of proteins to form amyloid fibrils is probably very common and may be a general feature of the polypeptide chain [7, 12, 13]. In fact, many proteins that are not associated with diseases have been shown to form fibrils in vitro (reviewed in [14]). Overall, there is an increasing belief that the ability to form fibrils is an inherent property of the polypeptide chain, i.e. many proteins, perhaps all, are potentially able to form amyloid fibrils under appropriate conditions [7, 15–17]. Amyloid fibrils are highly organized structures (similar to one-dimensional crystals) in which the essential features of the structure are determined by the physicochemical properties of the polypeptide chain [18]. Furthermore, it has been pointed out that, as with other highly organized compounds (including crystals) whose structures are based on repetitive long-range interactions, the most stable structures are those consisting of a single type of peptide or protein where self-interactions can be optimized [18, 19]. Obviously, this can explain the remarkable specificity of protein aggregation in protein deposition diseases. Although amyloidogenic polypeptides may be rich in β-sheet, α-helix, β-helix or natively unfolded [5, 20–27], the resulting fibrils display many common properties including a core cross-β-sheet structure in which continuous β-sheets are formed with β-strands running perpendicular to the long axis of the fibril [28]. All fibrils have similar morphologies, with mature fibrils usually having a twisted, rope-like structure, which usually involves two to six unbranched protofilaments around 2–3 nm in diameter associated laterally or twisted together to form fibrils of around 10 nm diameter (see, e.g. [29–31]). Since all fibrils, regardless of the original structure of the given amyloidogenic protein, have a common cross-β structure, considerable conformational rearrangements have to occur in order for fibril formation to happen. The molecular basis of amyloid fibril formation from globular proteins is described; briefly, fibril formation can only occur when the rigid native structure is destabilized, favoring partial unfolding and the formation of an amyloidogenic partially unfolded conformation [6–11, 32–35]. This is because the structural rearrangements required for the formation of fibrils cannot take place within the tightly packed native globular

1 Introduction 3 Table 1 Some natively unfolded or significantly disordered amyloidogenic proteins and the

corresponding amyloid-based clinical disorders Amyloidogenic protein

Type of structure

Disease

Prion protein and its fragments

N-terminal fragment (23–121) is natively unfolded; C-terminal domain (121–230) is α-helical (predominantly)

Aβ and its fragments

natively unfolded

ABri Huntingtin

natively unfolded exon 1 is unfolded and forms fibrils ligand-binding and DNAbinding domains are α-helical; N-terminal domain is natively unfolded unknown (natively unfolded)

Creutzfeldt-Jacob disease (CJD) Gerstmann-StrausslerSchneiker syndrome (GSS) fatal familial insomnia (FFI) kuru bovine spongiform encephalopathy (BSE) and scrapie Alzheimer’s disease (AD) Dutch hereditary cerebral hemorrhage with amyloidosis (HCHWA, also known as cerebrovascular amyloidosis) Congophilic angiopathy familial British dementia Huntington disease

Androgen receptor protein

Ataxin-1

DRPLA protein (atrophin-1)

unknown (probably natively unfolded)

IAPP (amylin)

natively unfolded

Calcitonin

natively unfolded

α-Synuclein

natively unfolded

Tau protein

natively unfolded

spinal and bulbar muscular atrophy (SBMA)

spinocerebellar ataxia (SCA) neuronal intranuclear inclusion disease (NIID) hereditary dentatorubral-pal-lidoluysian atrophy (DRPLA) pancreatic islet amyloidosis in late-onset diabetes (type II diabetes mellitus) medullary carcinoma of the thyroid (MCT) Parkinson’s disease (PD) diffuse Lewy bodies disease (DLBD) Lewy bodies variant of Alzheimer’s disease (LBVAD) dementia with Lewy bodies (DLB) multiple system atrophy (MSA) Hallervorden-Spatz disease Alzheimer disease (AD) Pick’s disease progressive supranuclear palsy (PSP)

4 Amyloid Fibril Formation of Natively Unfolded Proteins Table 2 Non-disease-related amyloidogenic proteins and peptides

Protein (peptide)

Type of structure

Prothymosin α [164] Human complement receptor 1, 18–34 fragment [165] GAGA factor [166] Yeast prion Ure2p [167] ApoCII [156, 157, 160] Cold shock protein B, 1–22 fragment [168] Core histones [163] Soluble homopolypeptides [169]: poly(L-lysine) poly(L-glutamic acid) poly(L-threonine)

natively unfolded unfolded native unfolded α-helical/unfolded Natively unfolded Unfolded Natively unfolded

protein, due to the constraints of its tertiary structure. This situation, obviously, is not applicable for natively unfolded proteins, as they are devoid of rigid structure in their starting state. Thus, in contrast to globular proteins, the primary step of fibrillogenesis of natively unfolded proteins is partial folding rather than partial unfolding. The details of the process will be illustrated using results from a detailed analysis of the aggregation of human α-synuclein, a protein whose fibrillogenesis in vitro has been studied extensively. The generality of the results will be emphasized with a variety of other natively unfolded proteins that assemble into amyloid fibrils. 2 Molecular Mechanisms of Amyloid Fibril Formation by a Natively Unfolded Protein: α-Synuclein 2.1 α-Synuclein in Parkinson’s Disease and other Neurodegenerative Disorders

Parkinson’s disease is a progressive disorder resulting from loss of neurons of the substantia nigra, a small area of cells in the mid-brain. Gradual degeneration of these dopaminergic neurons causes a reduction in the release of the neurotransmitter dopamine in the striatum. This, in turn, can produce one or more of the classic signs of Parkinson’s disease: resting tremor on one (or both) side(s) of the body, generalized slowness of movements (bradykinesia), stiffness of limbs (rigidity) and gait or balance problems (postural dysfunction). The precise mechanisms of neuronal death are unknown as yet. Some surviving nigral dopaminergic neurons contain cytosolic filamentous inclusions known as Lewy bodies (LBs) and Lewy neurites (LNs) [36, 37]. LBs are also neurophathological hallmarks of several other diseases collectively termed synucleinopathies. There is a strong link between α-synuclein aggregation and the pathogenesis of Parkinson’s disease, including three different missense mutations in the α-synuclein gene, corresponding to A30P, E46K and A53T substitutions in

2 Molecular Mechanisms of Amyloid Fibril Formation by a Natively Unfolded Protein: "-Synuclein

α-synuclein protein, in a small number of kindreds with autosomal-dominantly inherited, early-onset Parkinson’s disease [38–40]; triplication of the α-synuclein gene locus causes familial autosomal dominant Parkinson’s disease with early onset [41]; overexpression of wild-type (WT) α-synuclein in transgenic mice [42] or of WT, A30P and A53T in transgenic flies [43] leads to motor deficits and neuronal inclusions reminiscent of Parkinson’s disease. 2.2 Key Structural Properties of α-Synuclein: A Natively Unfolded Protein

α-Synucleins from different organisms exhibit a high degree of sequence conservation: human α-synuclein consists of 140 amino acid residues and can be divided into the N-terminal region, residues 1–95, containing six 11-amino-acid imperfect repeats with a highly conserved hexamer motif (KTKEGV), and the C-terminal region, residues 96–140, which is enriched in acidic residues and prolines, suggesting that it adopts a disordered conformation. α-Synuclein is a typical intrinsically unstructured or natively unfolded protein (for recent reviews, see [13, 21, 23, 26, 44, 45]). The sequence of intrinsically disordered proteins is characterized by amino acid compositional bias and the existence of highly predictable flexibility [46]. Further, the majority of the intrinsically disordered proteins, are substantially depleted in I, L, V, W, F, Y, C and N, and enriched in E, K, R, G, Q, S, P and A [21]. These features account for the low hydrophobicity and high net charge of the intrinsically unstructured proteins, a property that allows discrimination of globular (folded) and intrinsically unstructured proteins based solely on their amino acid composition [5]. Several detailed studies show that purified α-synuclein possesses little ordered structure under physiological conditions [45, 47–49] and is characterized by farUV circular dichroism (CD) and Fourier transform IR (FTIR) spectra typical of a substantially unfolded polypeptide pH [49]. The hydrodynamic properties of α-synuclein show that the protein is quite expanded, but is slightly more compact than expected for a random coil and lacks a tightly packed globular structure [45, 49, 50]. The results of pulsed-field gradient nuclear magnetic resonance (NMR; which allows an estimation of the hydrodynamic radius) confirm that α-synuclein is unfolded, but slightly collapsed [51], and a high-resolution NMR analysis revealed that α-synuclein is largely unfolded in solution, but exhibits a region between residues 6 and 37 with a preference for helical conformation [48]. Finally, Raman optical activity spectra suggested that α-synuclein may contain some poly(L-proline) II helical conformation [52]. One of the characteristic features of natively unstructured proteins is a reverse response to changes in their environment compared with that of globular proteins. For example, intrinsically unstructured proteins gain rather than lose ordered structure at extremes of pH or high temperatures. The structure-forming effects of low pH on α-synuclein [45, 49, 50] were attributed to the minimization of the large net negative charge present at neutral pH, thereby decreasing intramolecular charge–charge repulsion and permitting hydrophobic-driven collapse to the

5


partially folded conformation. The effect of elevated temperatures was attributed to increased strength of the hydrophobic interaction at higher temperatures, leading to a stronger hydrophobic driving force for folding [49]. In fact, any changes in the environment of a natively unfolded protein leading to an increase in protein hydrophobicity and/or decrease in its net charge are expected to lead to partial folding. 2.3 Major Structural Characteristics of Partially Folded α-Synuclein

As the pH is decreased (or temperature increased) changes were observed in the shape of the CD spectrum for α-synuclein. Fig. 1(a) shows that the minimum at 196 nm becomes less intense, whereas the negative intensity of the spectrum around 222 nm increases, reflecting pH-induced formation of secondary structure. Similarly, the FTIR spectrum of α-synuclein at pH 7.5 was typical of an unfolded polypeptide (Fig. 1b), whereas a decrease in pH leads to significant spectral changes, indicating an increase in ordered secondary structure. The most evident change is the appearance of a new β-sheet band in the vicinity of 1626 cm–1 . Thus, at acidic pH natively unfolded α-synuclein is transformed into a partially folded conformation with a significant amount of β-structure. Furthermore, Fig. 1(c) shows that a decrease in pH leads to a considerable increase in 1-anilino-8-naphthalene sulfonate (ANS) fluorescence intensity and a large blue shift of the ANS fluorescence maximum (from around 515 to around 475 nm), reflecting the pH-induced transformation of the natively unfolded α-synuclein to a partially folded conformation with solvent-exposed hydrophobic regions. Hydrodynamic methods revealed that this partially folded conformation is accompanied by a substantial decrease in hydrodynamic dimensions (Rs = 27.9 ± 0.4 and Rg = 30 ± 1 Å). Moreover, changes in the profile of the small-angle X-ray scattering Kratky plot at pH 3 were consistent with the development of a tightly packed core (Fig. 1D). This means that protonation of α-synuclein results in the transformation of this natively unfolded protein into a more compact conformation with a significant amount of ordered secondary structure, affinity for ANS and the beginnings of a tightly packed core, all hallmarks of a partially folded intermediate [49]. Comparable structural changes are induced in α-synuclein by high temperatures [49]. The analysis of these data suggests that the partially folded conformation induced in α-synuclein by low pH or high temperature resembles the pre-molten globule state, an intermediate, preceding the molten globule in the refolding of globular proteins. In fact, it has been shown that the pre-molten globule is a denatured state, i.e. a conformation where the protein lacks a rigid three-dimensional structure. It is characterized by some secondary structure, although much less than that of the molten globule or native globular protein. A protein in the pre-molten globule state is considerably less compact than in the molten globule state and does not have a tightly packed globular structure, but it is more compact than the corresponding random coil. Furthermore, the pre-molten globule can interact with the hydrophobic fluorescent probe ANS, although more weakly than a molten globular


Fig. 1 a–c

7


Fig. 1 Major structural characteristics of the natively unfolded and the partially folded pre-molten glubule-like conformation induced in human "-synuclein by low pH. (a) Far-UV CD spectra measured under different conditions. (b) FTIR spectra in the amide I region measured for natively

unfolded, partially folded and fibrillar forms of human alpha-synuclein. (c) ANS spectra measured under different experimental conditions. (d) Kratky plot representation of the results of small angle X-ray scattering analysis of "-synuclein at different experimental conditions.

protein. This means that at least some hydrophobic clusters are already formed in the pre-molten globule state, although there is no globular structure [53–55]. 2.4 Fibril Formation by α-Synuclein and the Partially Folded Amyloidogenic Conformation

The fibrillogenesis of α-synuclein in vitro has been studied extensively (for recent reviews, see [45, 56]). Although α-synuclein is an intrinsically unstructured protein, it forms fibrils of highly organized structure. Fig. 2 shows that both decreasing the pH and increasing the temperature result in accelerated fibrillation of α-synuclein [49]; thus, there is an excellent correlation between the intramolecular conformational change described above and the increased kinetics of fibril formation, suggesting that the partially structured conformation is a key intermediate on the fibril-forming pathway [49]. In contrast to an unfolded polypeptide chain, a partially folded conformation is anticipated to have contiguous hydrophobic patches on its surface, which might foster self-association and fibrillation. Thus, factors that shift the equilibrium in favor of this monomeric partially folded conformation will favor fibril formation. An increase in protein concentration obviously increases the absolute concentration of the intermediate and, thus, is predicted to accelerate fibrillation, as is observed (Fig. 3). There is an inverse linear correlation between the logarithm of α-synuclein concentration and the duration of the lag time (Fig. 3B). This reflects an important aspect of the kinetics of nucleus formation, i.e. the formation of the


Fig. 2 Effect of pH (A) and temperature (B) on fibril formation of human "-synuclein, monitored by Thioflavin T fluorescence. Decreasing pH and increasing temperature accelerate fibrillation, due to decreased net charge and increased hydrophobicity, respectively.

aggregation-prone partially folded intermediate represents the rate-limiting step in α-synuclein nucleation and this process is probably responsible for the observed first-order kinetics. An alternative way to view this is that association to form fibrils involves the association of a rare conformer, the amyloidogenic partially folded intermediate. The data suggest that the key partially folded intermediate, once formed, oligomerizes rapidly to form fibrils. Fig. 3(C) shows that there is also a linear dependence of the first-order rate constant for fibril growth (elongation) on protein concentration. The kinetics of elongation have been shown to follow

9


Fig. 3 The effect of protein concentration on the kinetics of fibril formation of human recombinant "-synuclein. (a) Kinetics of fibrillation at different "-synuclein concentrations. Protein concentrations were 21 (solid circles), 70 (open circles), 105 (solid tri-

angles) and 190 :M (open triangles). (b) Inverse linear dependence of the logarithm of "-synuclein concentration as a function of lag time. (c) Linear dependence of the rate constant kapp for fibril growth on the "-synuclein concentration.

first-order kinetics for some other proteins, including Aβ and insulin [57–60]. The simplest explanation for this fact is that increasing protein concentration leads to increasing numbers of fibrils, due to increasing concentration of nuclei and to increasing concentration of monomeric protein (either in its native-like form or as a partially folded intermediate, see below).

3 Fibrillogenesis of Natively Unfolded Proteins Requires Partial Folding 11

It has been assumed that the aggregation of α-synuclein, leading to the development of Parkinson’s disease and other synucleinopathies, arises from various factors that would significantly increase the concentration of the critical partially folded intermediate [49]. For example, a number of factors have been shown to induce partial folding of α-synuclein and accelerated fibrillation in vitro: these include some common pesticides and herbicides [61–63], metal ions [63, 64], moderate concentrations of osmolytes such as TMAO [65], and some alcohols [66]. Under all these conditions α-synuclein was shown to form fibrils rapidly, thus confirming the correlation between the partially folded conformation and fibril formation (see Figs 2B and 4A). In contrast, Fig. 4(B) shows that the process of fibril formation can be slowed or completely inhibited under conditions favoring formation of more folded conformations [65, 66] or by stabilization of off-pathway oligomers, e.g. via nitration of tyrosines [67] or methionine oxidation [68], or interaction with β- or γ -synucleins [50]. All conditions that populated the aggregation-prone partially folded conformation accelerated both the nucleation and elongation stages of fibril assembly. This means that the partially folded intermediate is likely involved in both the formation of the nucleus and in the subsequent propagation of fibrils. It is worth noting that the formation of amyloid-like fibrils is not the only pathological outcome of protein deposition diseases and in several disorders (as well as in numerous in vitro experiments) protein deposits are composed of amorphous aggregates, without local order, e.g. light chain deposition disease, which involves deposition of immunoglobulin light chains. Similarly, soluble oligomers represent another alternative final product of the aggregation process. The physiological significance of oligomers is that they may be the toxic species. Fig. 5 represents a simplified model of α-synuclein aggregation, demonstrating the fact that aggregation is a very complex process, which can be divided into three major steps. The model shows that the first stage of the aggregation process is the structural transformation of a soluble natively unfolded protein, UN , into the “sticky” aggregation-prone precursor or intermediate (I), which plays a central role in the entire aggregation process. This species can partition kinetically between a minimum of three distinct pathways, leading to fibrils, amorphous aggregates and soluble oligomers. The formation of a nucleus leading to both fibrils and amorphous aggregates is a kinetically disfavored event and is responsible for the lag period preceding significant formation of aggregates. Once a critical nucleus has been generated, the conditions change in favor of a rapid increase in aggregate size.

3 Fibrillogenesis of Natively Unfolded Proteins Requires Partial Folding 3.1 Fibril Formation by Proteins Involved in Conformational Disorders

Data presented in Tables 1 and 2 show that many of the known amyloidogenic proteins are natively unfolded. It is reasonable to assume that such proteins are well


Fig. 4 Effect of different environmental factors on fibrillation of human "-synuclein. (a) Factors accelerating protein fibrillation: control (circles), 1 :M TMAO (inverse triangles), 10% methanol (diamonds), 500 M

paraquat (squares) and 500 :M HgCl2 (triangles). (b) Factors inhibiting fibrillation of "-synuclein: control (circles), nitrated "-synuclein (squares) and 1:1 mixture of "-/$-synucleins (triangles).

suited for amyloidogenesis, as they lack significant secondary and tertiary structure. In the absence of such conformational constraints they would be expected to be substantially more conformationally mobile, and thus more able to polymerize in the form of β-sheets than tightly packed globular proteins. However, this is not always the case and many natively unfolded proteins form fibrils in vitro with similar rates to those of native globular proteins (i.e. within a time frame of a few


Fig. 5 Schematic overview of the aggregation of "-synuclein, showing the central position of the partially folded intermediate and the different final aggregated products.

hours to several days). The delay in the fibril formation of these proteins has been attributed to the requirement for considerable structural rearrangement within the unfolded polypeptide chain, giving rise to a partially folded conformation. The following section considers illustrative examples of fibril formation of natively unfolded proteins. 3.2 Amyloid β protein (Aβ)

Alzheimer’s disease is the most prevalent age-dependent dementia, and is characterized by the accumulation of extracellular amyloid deposits, senile plaques, in the cerebral cortex and of intracellular neurofibrillary tangles. Senile plaques contain Aβ, which is a 40- to 42-residue peptide produced by endoproteolytic cleavage of the Aβ protein precursor (AβPP), whereas neurofibrillary tangles are assembled from the protein tau (see below). Many lines of evidence support the crucial role of Aβ in Alzheimer’s disease, in which the aggregation of Aβ is associated with the development of the cascade of neuropathogenetic events, ending with the appearance of cognitive and behavioral features typical of Alzheimer’s disease. NMR studies have shown that monomers of Aβ1–40 or Aβ1–42 are unfolded under physiological conditions [69]. However, partial folding has been detected at the earliest stages of Aβ fibril formation [69] and Aβ aggregation fits the “standard” pattern of fibril formation, i.e. native(ly unfolded) to partially folded to oligomers to fibrillar species. Many recent studies have shown that Aβ forms small soluble oligomers, in which the polypeptide is partially folded and that appear to be the toxic species [70–78]. 3.3 Tau protein

Tau, a microtubule assembly protein [79–85], represents a family of isoforms with one, two, three or four repeats in the C-terminal region [86]. Post-translational phosphorylation of tau is an additional source of microheterogeneity [87]. Tau


is a significantly disordered protein. Interest in tau dramatically increased with the discovery of its aggregation in neuronal cells in the progress of Alzheimer’s disease and various other neurodegenerative disorders, especially fronto-temporal dementia [88, 89]. In these cases specific tau-containing neurofibrillary tangles paired helical filaments (PHFs)] are formed [90]. Hyperphosphorylation was shown to be a common characteristic of pathological tau [91], and hyperphosphorylated tau isolated from patients with Alzheimer’s disease is unable to bind to microtubules and promote microtubule assembly. However, both of these activities are restored after enzymatic dephosphorylation of tau [92–96]. During brain development, tau is phosphorylated at many residues and by several kinases [97, 98]. Most of the in vitro phosphorylation sites of tau are located within the microtubule interacting region (repeat domain) and sequences flanking the repeat domain. Many of these sites are also phosphorylated in PHF-tau [99, 100]. In fact, 10 major phosphorylation sites have been identified in tau isolated from PHFs from patients with Alzheimer’s disease [99]. All of these sites are located in regions flanking tau’s repeat domain and constitute recognition sites for several Alzheimer’s disease diagnostic antibodies, which may point to an important role for these phosphorylation sites for Alzheimer’s disease pathogenesis. Hyperphosphorylation is accompanied by transformation from the unfolded state of tau into a partially folded conformation [101, 102], dramatically accelerating the self-assembly into paired helical filaments in vitro [93]. To analyze the potential role of tau hyperphosphorylation in tauopathies, mutated tau proteins have been produced, in which all 10 serine/threonine residues known to be highly phosphorylated in PHF-tau were substituted for negatively charged residues, thus producing a model for a defined and permanent hyperphosphorylation-like state [103]. Analogous to hyperphosphorylation, glutamate substitutions induce compact structure elements and sodium dodecylsulfate (SDS)-resistant conformational domains in tau, as well as dramatic acceleration of its fibrillation [103].

3.4 Islet Amyloid Polypeptide (IAPP) or Amylin

In addition to insulin, pancreatic islet cells β produce a peptide called amylin or IAPP [104]. Dysfunction of amylin due to mutation and/or amyloid fibril formation is associated with the development of non-insulin-dependent diabetes mellitus (Type 2 diabetes) [105–107]. Amylin is an unstructured peptide hormone of 37 amino acid residues, which forms fibrils under physiological conditions in vitro. The process of polymerization is relatively fast (lag times of 100 and 50 min for fulllength amylin and its 8–37 fragment, respectively) and results in the appearance of typical amyloid fibrils [108]. Interestingly, both peptides showed formation of a partially folded intermediate early in the fibrillation process. It takes around 90 min for full-length amylin to form such an intermediate, whereas this period was almost half as long for the truncated peptide, showing excellent correlation with the fibrillation lag-times [108].


3.5 Prion Protein

Prion diseases, collectively referred to as the transmissible spongiform encephalopathies (TSEs), are characterized by unique infectious prion particles. TSEs include Creutzfeldt-Jakob disease, scrapie, bovine spongiform encephalopathy, and chronic wasting disease of mule deer and elk [109]. The characteristic pathological features of TSEs are spongiform degeneration of the brain and accumulation of the abnormal, protease-resistant prion protein isoform in the central nervous system, which sometimes forms amyloid-like plaques. The N-terminal region of about 100 amino acids in PrPC is unstructured in the isolated molecule in solution in the absence of copper. The C-terminal domain is folded into a largely α-helical conformation (three α-helices and a short antiparallel β-sheet) and stabilized by a single disulphide bond linking helices 2 and 3 [110]. The central event in the pathogenesis of prion diseases is a major conformational change of the C-terminal region of the prion protein (PrP) from an α-helical (PrPc ) to a β-sheet-rich isoform (PrPSc ) and PrPSc propagates itself by causing the conversion of PrPC to PrPSc . Although unstructured in the isolated molecule, the N-terminal region contains tight binding sites for Cu2+ ions and acquires structure following copper binding [111–115]. Investigations of the steps required for prion propagation and neurodegeneration in transgenic mice expressing chimeric mouse-hamster-mouse or mouse-human-mouse PrP transgenics indicated that the last 50 residues in the disordered N-terminal region play a particularly important role in the interaction of PrPC with PrPSc leading to the conversion of the former to the latter [116, 117]. Those residues are largely unordered or weakly helical in the full-length PrPC [118, 119], but are predicted to be β-structure in PrPSc [120]. These observations emphasize a crucial role of the disordered N-terminal region in the modulation of prion protein aggregation. Several kinetics studies have revealed the existence of partially folded intermediates for the prion protein [121–123] and is reasonable to assume that fibrillation requires partial unfolding of the C-terminal domain prior to self-association. 3.6 Polyglutamine Repeat Diseases

Currently there are eight known hereditary diseases, including Huntington’s disease, in which the expansion of a CAG repeat in the gene leads to neurodegeneration [124, 125]. The neurotoxicity in these diseases is due to the expansion of the CAGn -encoded polyglutamine [poly(Gln)] repeat. The mechanistic hypothesis linking CAG repeat expansion to toxicity involves the tendency of longer poly(Gln) sequences, regardless of protein context, to form insoluble aggregates [126–134]. To help evaluate various possible mechanisms, the biophysical properties of a series of simple poly(Gln) peptides have been analyzed. The CD spectra of poly(Gln) peptides with repeat lengths of 5, 15, 28 and 44 residues were shown to be nearly identical, and were consistent with a high degree of random coil structure,


suggesting that the length dependence of disease is not related to a conformational change in the monomeric states of expanded poly(Gln) sequences [132]. In contrast, there was a dramatic acceleration in the spontaneous formation of ordered, amyloid-like aggregates for poly(Gln) peptides with repeat lengths of greater than 37 residues. Several studies have revealed the existence of partially folded intermediates of poly(Gln)-repeat proteins as key species in fibrillation [133, 135, 136] and these are assumed to be precursors of oligomeric aggregates [132].

4 Fibrillation of Proteins Unrelated to Conformational Disease 4.1 Yeast Prions

There are at least two natural genetic elements, the [PSI+ ] and [URE3] factors, in the budding yeast Saccharomyces cerevisiae that exhibit non-Mendelian inheritance, but can be “cured” by treatment of the cells with low concentrations of the protein denaturant guanidine hydrochloride [137]. The [PSI+ ] factor is a self-perpetuating, conformationally altered form of a cellular protein [138–140]. The inheritance of [PSI+ ] from mother to daughter cells is based on the transmission of conformational information from ordered non-functional Sup35p aggregates (amyloid-like fibrils) to the soluble, functional form of Sup35p, a subunit of the polypeptide chain release complex that is essential for translation termination [141, 142]. In vitro, the prion-determining region of Sup35p, NM, is initially disordered, but slowly converts to a partially folded structure and then fibrils [143–145]. In particular, factors such as elevated temperatures, chemical chaperones and certain mutations that increased the amount of structure increased the amount of aggregation [145]. The acceleration of Sup35p fibril formation is determined by the acceleration of slow conformational changes rather than by providing stable nuclei. Strikingly, inhibitory mutations map exclusively within a short glutamine/asparagine-rich region of Sup35p and all but one occur at polar residues. Even after replacement of this region with poly(Gln), Sup35p retains its ability to form fibrils [140]. This suggests similarities between the prion-like propagation of [PSI+ ] and poly(Gln)-mediated pathogenesis of several neurodegenerative diseases (see above). The [URE3] element is another factor of S. cerevisiae that propagates by a prionlike mechanism and corresponds to the loss of function of the cellular protein Ure2 [146, 147]. The N-terminal region of the protein is flexible and unstructured, while its C-terminal region is compact and folded [148–150]. The overexpression of full-length Ure2p in wild-type S. cerevisiae strains induced a 20- to 200-fold increase in the frequency with which [URE3] arose. On the other hand, expression of just the N-terminal 65 residues of Ure2p increased the frequency of [URE3] induction 6000-fold [148]. These observations emphasize a key role of the disordered N-terminal domain in the fibrillation of Ure2 protein. A dimeric intermediate is populated transiently during refolding and populated under conditions correlating

4 Fibrillation of Proteins Unrelated to Conformational Disease

with fibril formation. Interestingly, the native dimer and dimeric intermediate are significantly more stable than either of their monomeric counterparts. Stabilization of the native state disfavors amyloid nucleation [151]. 4.2 Prothymosin α

This is a very acidic protein, containing around 50% aspartic and glutamic acid, no aromatic or cysteine residues, and very few large hydrophobic aliphatic amino acids [152]. Because of these features, prothymosin α adopts a random coillike conformation with no regular secondary structure at neutral pH ([152, 153], and Tables 1 and 2). However, at acidic pH prothymosin α folds into a partially folded conformation [153]. Interestingly, it has been recently shown that at low pH (below pH 3, i.e. under conditions favoring the formation of the partially folded conformation [153]), prothymosin α is capable of relatively fast formation (lag time around 100 min) of regular elongated fibrils with a flat ribbonlike structure 4–5 nm in height and 12–13 nm in width [154]. 4.3 Apolipoprotein CII (ApoCII)

Human ApoCII is a plasma protein consisting of 79 amino acid residues, whose function is to activate lipoprotein lipase. The structure of ApoCII in the presence of the lipid mimetic, SDS, reveals three regions of well-defined amphi-pathic α-helix with loosely defined intervening regions that may reflect flexible hinge regions [155]. In the absence of lipid or detergent, human ApoCII lacks ordered structure and forms amyloid ribbons after incubation for several days [156–162]. Fibril formation was dramatically accelerated by the addition of phospholipids in submicellar concentrations, which were shown to induce partial folding of the protein into a pre-molten globule-like intermediate [156]. In contrast, the fibrillation of ApoCII was completely inhibited in the presence of micellar phospholipids, i.e. under conditions favoring α-helical conformation [156]. 4.4 Histones

The bovine core histones are natively unfolded proteins in solutions of low ionic strength, due to their high net positive charge at pH 7.5 [163]. Analysis of the structural properties of core histones as a function of pH, ionic strength and organic solvent concentration demonstrated a correlation between conformational change and propensity to aggregate. Overall, the histones are able to adopt at least five different relatively stable conformations: the intrinsically unstructured form (pH 2.0, low protein concentration, low ionic strength), a fully-unfolded conformation (6 M urea), partially folded α-helical dimers (low ionic strength, pH 2.0, high protein concentrations), helical monomers (low ionic strength, pH 2.0, 10% trifluoroethanol)

17


and molten globule-like α-helical oligomers (pH 2.0, high salt) [163]. Under most of the conditions studied the histones formed amyloid fibrils with typical morphology as seen by electron microscopy. In contrast to most aggregation/amyloidogenic systems, the kinetics of fibrillation showed an inverse dependence on histone concentration, which has been attributed to partitioning to a faster pathway leading to non-fibrillar self-associated aggregates at higher protein concentrations. In keeping with the hypothesis that partial folding is required of intrinsically disordered proteins for aggregation, the rate of fibril formation correlated with conditions favoring the stabilization of the partially folded conformation [163].

5 Conclusions

Intrinsically disordered or natively unfolded proteins represent a high fraction of naturally occurring amyloidogenic proteins. This is most likely due to the ease with which they can form the β-sheet topology required in amyloid fibrils, in contrast to folded globular proteins, for which the native conformation places major constraints for the required topological rearrangement. In contrast to tightly folded globular proteins, the first, and critical, step in the aggregation of intrinsically disordered proteins or polypeptides is partial folding. This results in a partially folded conformation/intermediate with hydrophobic surface patches that favors self-association. As with such intermediates formed from the partial folding of globular proteins, such aggregate-prone intermediates can polymerize to form fibrillar or amorphous aggregates, or soluble oligomers.

Acknowledgments

This work was supported by grants INTAS 2001-2347 (V. N.U.), and NIH NS39985, DK55675 and NS43778 (A. L. F.).

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139 PAUSHKIN, S. V., V. V. KUSHNIROV, V. N. SMIRNOV and M. D. TER Avanesyan. Propagation of the yeast prion-like [psi+ ] determinant is mediated by oligomerization of the SUP35-encoded polypeptide chain release factor. EMBO J 1996, 15, 3127–3134. 140 DEPACE, A. H., A. SANTOSO, P. HILLNER and J. S. WEISSMAN. A critical role for aminoterminal glutamine/asparagine repeats in the formation and propagation of a yeast prion. Cell 1998, 93, 1241–1252. 141 SERIO, T. R. and S. L. LINDQUIST. [PSI+ ]: an epigenetic modulator of translation termination efficiency. Annu Rev Cell Dev Biol 1999, 15, 661–703. 142 SERIO, T. R., A. G. CASHIKAR, J. J. MOSLEHI, A. S. KOWAL and S. L. LINDQUIST. Yeast prion [psi+ ] and its determinant, Sup35p. Methods Enzymol 1999, 309, 649–673. 143 GLOVER, J. R., A. S. KOWAL, E. C. SCHIRMER, M. M. PATINO, J. J. LIU and S. LINDQUIST. Self-seeded fibers formed by Sup35, the protein determinant of [PSI+], a heritable prion-like factor of S. cerevisiae. Cell 1997, 89, 811–819. 144 SERIO, T. R., A. G. CASHIKAR, A. S. KOWAL, G. J. SAWICKI, J. J. MOSLEHI, L. SERPELL, M. F. ARNSDORF and S. L. LINDQUIST. Nucleated conformational conversion and the replication of conformational information by a prion determinant. Science 2000, 289, 1317–1321. 145 SCHEIBEL, T. and S. L. LINDQUIST. The role of conformational flexibility in prion propagation and maintenance for Sup35p. Nat Struct Biol 2001, 8, 958–962. 146 WICKNER, R. B. [URE3] as an altered URE2 protein: evidence for a prion analog in Saccharomyces cerevisiae. Science 1994, 264, 566–569. 147 THUAL, C., L. BOUSSET, A. A. KOMAR, S. WALTER, J. BUCHNER, C. CULLIN and R. MELKI. Stability, folding, dimerization and assembly properties of the yeast prion Ure2p. Biochemistry 2001, 40, 1764–1773. 148 MASISON, D. C. and R. B. WICKNER. Prion-inducing domain of yeast Ure2p and protease resistance of Ure2p in

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27

1

Kinesin Superfamily Proteins Nobutaka Hirokawa, and Reiko Takemura University of Tokyo, Tokyo, Japan


1 Introduction

A cell is not simply a bag filled with cytoplasmic fluid surrounded by the plasma membrane in which membranous organelles such as the Golgi apparatus, endoplasmic reticulum and secretory vesicles float and through which newly synthesized proteins diffuse to reach their destination. Instead, cells transport and sort proteins and lipids after their synthesis to their proper destinations at appropriate velocities in membranous organelles and protein complexes using various kinds of motor proteins. The Neuron is composed of a cell body, dendrites and a long axon along the direction of impulse propagation. Because of the lack of protein synthesis machinery in the axon, most of the proteins necessary for the functioning of the axon and synaptic terminal must be transported down the axon after their synthesis into the cell body [1]. Most proteins are conveyed in membranous organelles or protein complexes. In this sense intracellular transport in the axon is fundamental to Neuronal morphogenesis and functioning. Epithelial cells also develop a polarized structure, that is, apical and basolateral regions, to which certain proteins are specifically transported and sorted [2, 3]. Vectorial intracellular transport occurs not only in polarized cells such as Neurons and epithelial cells but rather in all types of cells [4]. Microtubules serve as rails for this transport and have a polarity with a fastgrowing or plus end and a slow-growing or minus end. They are well organized in cells. In nerve axons, microtubules are arranged longitudinally with the plus end oriented away from the cell body. In proximal dendrites, the polarity of microtubules is mixed, while in the distal end, the polarity is the same as in the axon. In epithelial cells, microtubules are organized with the plus end oriented toward the basement Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Kinesin Superfamily Proteins

Fig. 1 Quick-freeze deep-etch electron micrographs showing short cross-bridges between membranous organelles and microtubules that are structural candidates for KIFs. Bar 50 nm. Reproduced from [4], Trends Cell Biol. 6: 135–141, with permission.

membrane. In most other cells such as fibroblasts, microtubules radiate from the cell center with the plus end oriented toward the periphery. Electron microscopy (EM) revealed short cross-bridge structures between the organelles and microtubules [5], which are candidates for microtubule-associated motor proteins conveying the membranous organelles along microtubules (Fig. 1). Moreover, video-enhanced differential interference contrast microscopy revealed bidirectional transport of various types of membranous organelles [6, 7]. Based on these findings, kinesin superfamily proteins have been identified as motor proteins transporting cargoes along microtubule rails. ‘Conventional kinesin’, the first member of the kinesin superfamily that was identified, consists of two 120-kDa heavy chains (KHCs) and two 64-kDa light chains (KLCs), at least in animal species [7–14]. It has a rod-like structure composed of two globular heads (10 nm in diameter), a stalk, and a fan-like end, with a total length of 80 nm. The globular heads are composed of KHCs that bind to microtubules [15, 16]; the KLCs constitute the fan-like end [15, 17]. Complementary DNA (cDNA) encoding KHCs yields a protein of ∼975 amino acids of which ∼350 NH2 -terminal amino acids form the motor domain (which binds to microtubules and has ATPase activity), an α-helical coiled coil-rich stalk domain involved in dimer formation and a tail domain [18].

2 The Kinesin Superfamily Proteins

Fig. 2 Structure of cDNAs from murine kinesin superfamily proteins (KIFs). Purple, motor domain; thin red line, ATP-binding consensus sequence; thick red line, microtubule-binding consensus sequence.

A systematic molecular biological search of kinesin superfamily genes coding for proteins containing ATP-binding and microtubule-binding consensus sequences or screening using anti-pan kinesin antibodies led to the discovery of many kinesin superfamily proteins, KIFs [19–24] (Figs 2 and 3). Combining molecular biological approaches with a BLAST search of proteins in public and private genome databases, a total of 45 KIFs were identified in mouse and human genomes [25]; Figs 4, 5 and Tab. 1. In this chapter, we provide an overview of KIFs.

2 The Kinesin Superfamily Proteins

KIFs have been classified into three major types based on the position of the motor domain: NH2 -terminal motor domain type, middle motor domain type, and COOHterminal motor domain type (referred to as N-kinesin, M-kinesin, and C-kinesin, respectively).

3


Fig. 3 Left: Panels showing main KIFs functioning in intracellular transport, as observed by low-angle rotary shadowing EM. Scale bar, 100 nm. Right: Schematic illustration of the same KIFs based on EM studies or predicted from analysis of primary structures. Reproduced from [26], Science 279: 519–526, with permission.

All KIFs known or predicted to be transcribed in the human and mouse genomes are presented in a phylogenic tree along with the KIFs from S. cerevisiae, Drosophila melanogaster, and Caenorhabditis elegans (Fig. 5). The entire family is classified into 14 classes. N-kinesin consists of 11 classes, comprising 16 families. Most classes are composed of one family except N-3, N-4, N-6, and N-8. The N-3 class consists of Unc104/KIF1, KIF13, and KIF16 families. Members of the Unc104/KIF1 family are mostly monomeric, while members of the KIF13 family have different characteristics. The N-4 class is composed of the KIF3 and Osm3/KIF17 families. KIF3 is heterotrimeric, and Osm3/KIF17 forms homodimers, indicating that these two families are distinct within this class. The N-8 class consists of the KIF18 and Kid/KIF22 families. M-kinesin is composed ofone class, the KIF2 family. Ckinesin is composed of C-1 and C-2 classes, each having one family. Most of the KIFs of other species including plant species can be categorized into these 14 classes.

3 N-Kinesins 3.1 N-1 Kinesins

The first reported kinesin, the kinesin heavy chain (KHC), is included in this group. There are three highly related family members in this group, KIF5A, KIF5B, and

3 N-Kinesins

Fig. 4 All mouse and human KIFs and phylogenetic analysis of mouse and human orthologs. Sequences were analyzed by the neighbor-joining method. Reproduced from [25], PNAS 98: 7004–7011, with permission.

KIF5C, all of which form tetramers consisting of two heavy chains and two light chains [18, 19, 26–30]. KIF5B is ubiquitously expressed in many tissues, while KIF5A and KIF5C are specifically expressed in the nervous system [25, 28, 30]. The members of this family transport various organelles and macromolecular complexes within the cell [25, 26, 31], including anterograde transported vesicles [32] probably important for action potential propagation [33], mitochondria [34], lysosomes [35, 36], endocytic vesicles [37], tubulin oligomers [38], intermediate filament proteins such as vimentin [39, 40] and mRNA complexes [41, 42] (Fig. 6a,b). It may also be involved in cytoplasmic viral transport [43].

5


Fig. 5 Phylogenetic analysis of all KIFs expressed in mouse/human, D. melanogaster, C. elegans and S. cerevisiae. Amino acid sequences were aligned using maximum parisomy. Reproduced from [25], PNAS 98: 7004–7011, with permission.

The exact molecular interaction involved in the binding of KIF5 to these cargoes has not been elucidated, but recently a direct interaction between KIF5 and some of the cargoes has been shown. KIF5 has been shown to bind directly to a group of scaffolding proteins of the JNK signaling pathway namely, the c-jun NH2 -terminal kinase (JNK)-interacting proteins (JIPs), JIP-1, JIP-2, and JIP-3 (also called JSAP1

KIF1A KIF1B KIF1C KIF2A KIF2B KIF2C KIF3A KIF3B KIF3C KIF4A KIF4B KIF5A KIF5B KIF5C KIF6 KIF7 KIF8 KIF11 KIF9 KIF10 KIF12 KIF13A (−) (−) BimC (−) (−) (−) Klp4

(−) (−) Klp61F (−) Cmet Cana CG15844 Kin73

CENP-E

EG5/KNSL1

Unc116

KHC

Y43F4B.6

nKHC uKHC xKHC

KRP85 KRP95

(−) K11D9.1

Unc104

C. elegans

KIF4

CG1453 CG3219

CG8566

D. melanogaster

Klp64D Klp68D CG17461 Klp3A

CAKin/KNSL6

KIF2

ATSV

H. sapiens

(−) (−) Kip1 Cin8 (−) Kip2 (−) (−)

(−)

(−)

(−)

(−)

(−)

S. cerevisiae

AnBo,C XlEg5 CrKlp1

RnnKHC

GgChrkin

CgMCAK XlKCM1 SpKRP85 SpKRP95

RnKIF1D XlXKIF2

Others

Table 1 All KIFs from mouse, human, D. melanogaster, C. elegans, S. cerevisiae, and selected other organisms

(Continued)

[22] [22] [22, 58, 61, 127, 181] [22, 171, 172] [22, 128, 135], FB, [127] [22], FB [22, 77, 182, 183]

[19, 65, 72, 175] [64, 66], FB [22, 63, 176] [19], FB, WB, [153] [25], FB [25, 149, 155, 157] [19, 24, 81, 177] [22], FB [19, 109], WB, [113] [25, 178] [19, 22, 24, 28, 30, 179, 180]

Reference

3 N-Kinesins 7

HSET/KNSL2

KNSL3

Kid/KNSL4 MKLP1/KNSL5

Rab6Kin KlpMPP1

GAKIN HUMORFW Hklp2

T01G1.1 (−) Zen4A Zen4B (−) (−) Vab8 C41G7.2 M01-E11.6 W02-B12.7 Klp3

(−) Klp67A CG9913 CG12298 CG5300 Nod Pav CG17459 (−) (−) Ncd (−)

Osm3 (−) (−) (−)

Klp98A

C. elegans

Klp6 C06G3.2 C33H5.4 (−)

Klp38B (−)

D. melanogaster

FB Flybase http://flybase.bio.indiana.edu:82/; WB, Wormbase, www.wormbase.orgxs

KIFC2 KIFC3

KIF16A KIF16B KIF17 KIF18A KIF18B KIF19A KIF19B KIF20A KIF20B KIF21A KIF21B KIF22 KIF23 KIF24 KIF25 KIF26A KIF26B KIFC1

KIF13B KIF14 KIF15

H. sapiens

(−)

(−) (−) (−) (−) (−) (−) Kar3

(−)

(−) Kip3

(−)

(−) (−)

S. cerevisiae

XlXCTK1

CgCHO2

XlXkid CgCHO1

MmKlp174

XlXklp4

Others

Table 1 All KIFs from mouse, human, D. melanogaster, C. elegans, S. cerevisiae, and selected other organisms (Continued)

[22, 155, 167, 168, 193] [22, 169, 194]

[22, 183] [22, 185] [22, 106, 107] [25], FB, [186] [25], FB [25, 123], FB [25, 187] [114], FB, WB [25, 136–138] [25, 115, 116, 118, 119] [25], FB [25, 188] [25, 143] [22, 189–191], WB, [157, 192]

[22, 78] [22, 184], WB [22, 142], WB

Reference


3 N-Kinesins

Fig. 6 Scheme of KIFs and their cargo organelles in neurons (a) and in cells in general (b). In (b), neuron-specific KIFs and ubiquitous KIFs are illustrated in the same cell. CGN, cis-Golgi network; TGN, trans-Golgi network; ECV, endosomal carrier vesicle. Black arrows indicate the direction of transport.

9


Fig. 7 Schematic drawing of diverse functions of KIF3 in various cell types.

and Sunday Driver, SYD) [44–48]. JIP-1 and JIP-2 interact with tetra-tricopeptide repeat (TPR) motifs of kinesin light chains through their C-termini. JIP-1 and JIP-2 are soluble proteins by themselves. Therefore, they do not directly connect kinesin to membrane organelles. Rather, they may serve as a link between other cargo membrane proteins and KIF5. One possible mechanism of interaction is that JIP-1 and JIP-2 mediate the interaction of KIF5 with vesicles containing ApoER2, the receptor for the Reelin ligand that controls Neuronal migration (Fig. 8). The role of JIP-3 (Sunday Driver, JSAP1) is less clear. JIP-3 interacts with TPR motifs of kinesin light chains through internal sequences, instead of the C-terminus, and is proposed to be a membrane protein by itself [44], but an unequivocal transmembrane domain has not yet been found. Amyloid precursor protein (APP), whose aberrant transport is thought to contribute to the development of Alzheimer’s disease, is another protein that was proposed to interact directly with KIF5 [49]. The interaction between APP and KIF5

3 N-Kinesins

Fig. 8 Scheme of how KIFs recognize and bind cargoes.

is again through the TPR motifs of kinesin light chains (KLCs). It is proposed that APP serves as the membrane cargo receptor for KIF5 which links KIF5 to a particular subset of axon-transported vesicles. Thus far, reports of binding of KIF5 to other molecules seem to be focused on the TPR motifs of kinesin light chains. However, it was not clear whether KLC is the only site for the interaction between kinesin and cargoes. In some species such as fungi [50, 51], kinesin light chains are absent, implying that KHCs alone are sufficient by themselves for binding to some cargoes [52]. An biochemical interaction between KHC and membranous organelles has been suggested [53]. Very recently, GRIP1 (Glutamate Receptor Interacting Protein 1) has been shown to bind to the KHC tail domain and transport AMPA-type glutamate receptor (GluR2)-containing vesicles to dendrites [54] (Fig. 8). Furthermore, this study demonstrated clearly that GRIP1 steers kinesin to dendrites while JSAP1 (JIP-3, Sunday Driver) steers kinesin to axons through its interaction with KLC [54]. KIF5B was also shown to interact with the actin-based myosin motor MyoVA, suggesting a coordination between these two motors [55].

11


3.2 N-2 Kinesins

This class includes KIF11, BimC, Eg5, and plays a role in mitosis [25, 56–62]. KIF11 (Eg5) was shown to form a homotetramer for bipolar spindle formation. The motors localize at the midzone of interpolar microtubules, where microtubules have an anti-parallel orientation. Because the tetramers have motor domains on both ends of the molecule, they are thought to crosslink antiparallel microtubules and to allow them to slide against each other. 3.3 N-3 Kinesins

This class consists of three families, the Unc104/KIF1, KIF13, and KIF16 families [25]. The Unc104/KIF1 family is unique, because most of the proteins in this family form monomeric motors. 3.3.1 The Unc104/KIF1 family There are four members in this family, namely KIF1A, KIF1Bα, KIF1Bβ, and KIF1C [26, 63–66]. KIF1A and KIF1B are monomeric, a unique characteristic of these proteins compared to other KIFs. KIF1A was shown to be able to move processively as a monomer by a biased Brownian movement toward the microtubule plus end [67–70]. C. elegans Unc104 is a homolog of mouse KIF1A [71, 72]. KIF1Bα and KIF1Bβ are splice variants which have similar N-terminal motor domains but are distinct in their cargo-binding domains. Interestingly, the cargo-binding domains of KIF1A and KIF1Bβ have high homology, and both KIFs transport precursors of synaptic vesicles [65, 66] (Fig. 6a). The knock-out mice of KIF1A and KIF1B show an aberrant Neuronal function due to the defect in the transport of precursors of synaptic vesicles [66, 73]. This gene-targeting experiment demonstrated for the first time that defects in axonal transport due to a mutated motor protein can cause human peripheral neuropathy [66]. The KIF1B heterozygotes show progressive muscle weakness and motor disturbance similar to human neuropathies, and it was shown that patients with Charcot–Marie–Tooth disease type 2A carry a loss-offunction mutation in the motor domain of the KIF1B gene [66]. KIF1Bα transports mitochondria, but this role is shared with KIF5B [34] (Fig. 6a). KIF1C is dimeric, has a high homology to the mitochondrial motor KIF1Ba, and is believed to be involved in the transport of vesicles from the Golgi apparatus to the endoplasmic reticulum [74]. KIF1C was shown to be associated with 14-3-3 β, γ , and ζ proteins [75]. However, unexpectedly, cells from KIF1C (−/−) mice showed no significant difference from control cells in the transport of vesicles from the Golgi apparatus to the endoplasmic reticulum, suggesting that KIF1C is dispensable for retrograde transport [63]. Other KIFs may share this function. Recently, KIF1C was reported to mediate mouse macrophage resistance to the anthrax lethal factor [76]. KIF1C does not affect the cellular entry process of anthrax lethal toxin; therefore, events occurring later in the intoxication pathway are probably involved.

3 N-Kinesins

3.3.2 The KIF13 family KIF13A transports vesicles containing the mannose-6-phosphate receptor from the trans-Golgi network to the plasma membrane [77] (Fig. 6b). The intracellular trafficking of the mannose-6-phosphate receptor has been well studied. One of the roles of the mannose-6-phosphate receptor is to function as a recognition signal for the transport of newly synthesized lysosomal enzymes from the trans-Golgi network; it is also involved in the uptake of external ligands on the cell surface. The interaction of KIF13A with the vesicle is mediated via direct interaction between the KIF13A tail and β-1-adaptin, a subunit of the AP-1 adaptor complex [77], and the direct interaction of AP-1 with the mannose-6-phosphate receptor has been demonstrated (Fig. 8). KIF13B (GAKIN) was reported to interact with the human lympohcyte homolog of the Drosophila disc large tumor suppressor protein (hDlg), and may participate in the translocation of hDlg from the cytoplasm to the ‘cap’ structure upon activation [78]. Among the MAGUK superfamily proteins, GAKIN binds to the guanylate kinase-like domain of PSD-95, but not to other MAGUKs. 3.3.3 The KIF16 family KIF14, KIF16A, and KIF16B are members of this family. The function of this group of KIFs has not been clarified yet [25]. Because the tail domains as well as the expression patterns of KIF14, KIF16A, and KIF16B are different, these three KIFs may have separate functions. 3.4 N-4 Kinesins

This class is composed of the KIF3 family, which form heterotrimeric motors, and the Osm3/KIF17 family [25]. 3.4.1 The KIF3 family KIF3A, KIF3B, and KIF3C constitute this family. Members of the KIF3 family form heterotrimeric complexes. Thus the KIF3A–KIF3B heterodimer interacts at its tail domain with a non-motor protein, kinesin superfamily-associated protein 3 (KAP3) [26, 79–84] (Fig. 3). The KIF3A/B-KAP3 complex (also called kinesin II) transports vesicles and macromolecules [85]. In axons of mammalian Neurons, the complex transports vesicles which carry fodrin [86], and choline acetyltransferase, a soluble protein in Drosophila [87]. In melanophores, it is involved in the dispersion of pigment organelles called melanosomes [88–90] (Fig. 7). The KIF3 complex plays a vital role in the formation and maintenance of cilia and flagella in various organisms [91–96]. It transports macromolecular protein complexes from the bottom of the cilium to its distal tip along microtubules by a process called intraflagellar transport (Fig. 7). The components of these macromolecular complexes, called rafts, have been analyzed to some extent; rafts may contain as many as 15 polypeptides, but the details have not been completely clarified [91]. An interesting aspect of the involvement of KIF3 in cilium formation is that it also plays an important role in the determination of the left–right axis in early

13


mouse development [97–99]. During mouse development, there is a monocilium at the nodal cell that rotates to produce unidirectional leftward flow of extraembryonic fluid [98, 99]. This nodal flow could generate concentration gradients of putative secreted morphogens in the extraembryonic fluid at the nodal region, which could switch on a gene cascade strictly expressed on the left side of the body and lead to left–right asymmetry. KIF3 is required for the formation of this monocilium. In kif3A (−/−) or kif3B (−/−) mouse, nodal cilia cannot be generated, nodal flow is absent, and the left–right axis formation is fundamentally impaired [98, 99]. In retinal photoreceptor cells, the KIF3 complex is localized in the connecting cilium, which is located between the outer and inner segment [100]. It was shown that the KIF3 complex participates in the transport of opsin from the inner to the outer segment at the connecting cilium, where microtubule plus-ends are pointed toward the outer segment [101]. At the connecting cilium, opsin is localized at the plasma membrane, and the KIF3 motors probably drag the transmembrane protein along the underlying microtubules. The soluble protein, arrestin, also seems to be transported by the KIF3 complex. In migrating epithelial cells, KAP3 is reported to bind to the tumor suppressor gene, adenomatous polyposis coli (APC), and it is proposed that β-catenin is transported to the edges of migrating epithelial cells through this interaction [102]. There is another form of the KIF3 complex, KIF3A/C–KAP3. KIF3A/C–KAP3 is expressed in the nervous system only [103, 104], while KIF3A/B–KAP3 is ubiquitously expressed in many tissues. However, a gene-targeting experiment showed that the KIF3A/C-KAP3 complex is dispensable; therefore, the function of this KIF3 complex may be redundant [105]. 3.4.2 The Osm3/KIF17 family KIF17 is localized to the dendrites and transports vesicles containing N-methyl-Daspartate (NMDA)-type glutamate receptor 2B towards the microtubule plus-end to the postsynaptic site, where the receptor plays an important role in synaptic plasticity [106] (Fig. 2). In the dendrite, microtubules are of mixed polarity. The large protein complex containing mLin-10 (Mint1), mLin-2 (CASK), mLin-7 (MALS/Velis), and NR2B mediates the attachment of KIF17 to the vesicle. The tail domain of KIF17 interacts directly with the PDZ domain of mLin-10, which then sequentially interacts with mLin-2 (CASK), mLin-7, and the NR2B subunit in this order [106] (Figs 6a and 8). Osm-3 is closely related to KIF3, but forms a dimer [107]. In C. elegans, many Neurons have ciliated dendritic ends, and some of them are chemosensory receptors. Osm-3 transports ciliary components to these ciliated dendritic ends [23, 108]. 3.5 N-5 Kinesins

This class includes KIF4A, KIF4B, KIF21A, and KIF21B, and plays roles in both vesicle transport and mitosis [25, 109]. KIF4A mRNA is abundantly expressed in juvenile Neurons, including differentiated immature Neurons, and the motor

3 N-Kinesins

transports vesicles to growth cones [110]. These vesicles may contain L1, a cell adhesion molecule implicated in axonal elongation in immature Neurons [111]. KIF4 was shown to bind murine leukemia virus Gag proteins and may play a role in the transport of viruses in an infected cell [112]. Chromokinesin, a chicken isolog of KIF4, is associated with chromosome arms and functions as a mitotic motor with DNA as its cargo [113]. What KIF21A and KIF21B transport is not known, but they have different distribution patterns within a Neuron; KIF21A is localized throughout Neurons, but KIF21B is highly expressed in dendrites [114]. 3.6 N-6 Kinesins

CHO1/KIF23 and KIF20/Rab kinesin families constitute this class. 3.6.1 The CHO1/KIF23 family The KIF23 family, called by various names such as CHO1, MKLP, ZEN-4 or Pavarotti, depending on the species, appears to be involved in cytokinesis [115–120]. Members of this family can bind to microtubules at their cargo-binding domain, and are, therefore, capable of bundling anti-parallel microtubules. During mitosis, they exist in the midbody region of the spindle in late anaphase and telophase, and probably establish the structure of the telophase spindle to provide a framework for the assembly of the contractile ring. They are also expressed in cultured Neurons [121, 122]. 3.6.2 The KIF20/Rab6 kinesin family The Rab family of GTP-binding proteins regulates vesicular transport and membrane traffic. KIF20A (Rab6-KIF) was first reported as a motor protein that associates with Rab6 and participates in Golgi-derived vesicle transport [123]. Moreover, it was revealed that KIF20 may also participate in cytokinesis [124, 125]. 3.7 N-7 Kinesins

KIF10 (CENP-E) plays an important role in mitosis [126–129]. It associates with kinetochores and tethers the centromere to spindle microtubules. It is presumed to function in ‘congression’, in which chromosomes attached to the spindle microtubules oscillate and migrate to the spindle equator. Therefore, it plays an essential role in chromosome alignment at the spindle equator by tethering kinetochores to the dynamic microtubule plus end [130–134]. CENP-meta has a similar function in Drosophila [135]. 3.8 N-8 Kinesins

This class includes the Kid/KIF22 and KIF18 families.

15


3.8.1 The Kid/KIF22 family KIF22 (Kid) is involved in mitosis [136]. It binds to the arms of chromosomes, not to the kinetochore, and plays a role in chromosome alignment at the equatorial plate during metaphase [137–139]. It is proposed that Kid is responsible for the polar ejection force acting on chromosome arms and its degeneration is required for chromosome movement during anaphase. A highly homologous KIF, Nod, is required during female meiosis for chromosome alignment [140, 141]. KIF19 is included in this family, but its function is not yet well characterized [25]. 3.8.2 The KIF18 family This most recently discovered family of KIFs has not yet been characterized. 3.9 N-9 Kinesins

KIF12, a member of this family, is highly expressed in the kidney and may have a significant role in kidney cells. However, its function is not yet well characterized [25]. 3.10 N-10 Kinesins

The KIF15 family forms this class that is predominantly expressed in the spleen and testis [25]. Human KIF15 (Hklp2) may interact with the fork-head-associated domain of pKi-67, a widely used cell proliferation marker protein [142]. 3.11 N-11 Kinesins

This class includes the KIF26 family that is related to Cos2 and Vab-8. Vab-8 has been implicated in axon outgrowth and cell migration [143]. Costal2 (Cos2) has been shown to be part of the hedgehog signaling cascade; therefore, it is involved in the determination of cell fate and the patterning of animal development [144– 148]. It forms a large multiprotein complex with Ser/Thr protein kinases, Fused (Fu) and the transcription factor Cubitus interruptus (Ci). Cos2 prevents Ci nuclear translocation by tethering it in the cytoplasm via binding to microtubules, whereas hedgehog stimulates Ci nuclear import.

4 M-Kinesins

M-kinesins have a motor domain at the center of the molecule. The KIF2 family belongs to this class; its members, KIF2A, 2B, and 2C, have functions in vesicle transport, mitosis, and regulation of microtubule dynamics [25, 149, 150];

5 C-Kinesins

Figs 2 and 3). KIF2A plays a significant role in neurite outgrowth and transports vesicles containing a variant of the (β-subunit of the IGF-1 receptor, which is found on growth cones and is implicated in nerve cell development [151]; Fig. 6a). It may also participate in localization of lysosomes [152]. The KIF2C homolog in Xenopus, XKCM1, has a microtubule-destabilizing activity. It influences microtubule dynamics within the cell and is thought to promote chromosome segregation during mitosis by depolymerizing microtubules at the kinetochore [153–155]. XKCM1 acts on a single protofilament, possibly via an interaction of the basic residues in XKCM1 with the acidic tail of tubulin [156]. Mitotic centromere-associated kinesin (MCAK) identified in hamster is also a homolog of KIF2C [157]. It is also a microtubule depolymerizing factor, consistent with its role in promoting chromosome segregation during mitosis [158].

5 C-Kinesins 5.1 C-1 Kinesins

Drosophila Ncd, S. cerevisiae Kar3, and KIFC1 are included in this family [159, 160]. They are microtubule minus-end-directed motors and participate in meiosis, mitosis and karyogamy [25]. They have a microtubule-binding site on the cargo-binding domain. During mitosis, Ncd may function by crosslinking and sliding anti-parallel spindle microtubules relative to one another, pulling them apart, and thereby opposing the effect of bipolar plus-end-directed microtubule motors [161]. In meiosis, Ncd may play a role in establishing bipolar acentrosomal meiotic spindles [162]. Kar3 has an essential role in nuclear fusion during mating or karyogamy in S. cerevisiae. In addition, it has been implicated in several microtubule functions during the vegetative cell cycle including spindle assembly, mitotic chromosome segregation, microtubule depolymerization, kinetochore-motor activity and spindle positioning. These multiple functions may be performed by interaction of Kar3 with different associating proteins such as Cik1p or Viklp [163–166]. 5.2 C-2 Kinesins

The KIFC2/C3 families constitute C-2 kinesins [25, 167, 168]. KIFC2 is localized in the cell body and dendrites of Neurons [168]. It participates in the transport in dendrites of multivesicular body-like membranous organelles, which are morphologically similar to multivesicular bodies but appear to lack markers of endocytic compartments (Fig. 6a). The classic multivesicular body exists in both axons and dendrites; therefore, the organelle transported by KIFC2 may constitute a novel class. It has not been rigorously demonstrated, but it is presumed that KIFC2 acts as a minus-end-directed motor.

17


KIFC3 is a minus-end-directed microtubule motor protein that exists in kidney epithelial cells and other cell types. In epithelial cells, microtubule organization is different from that in other cell types; microtubule minus-ends are located at the apical surface of the cell. KIFC3 accumulates on the apical surface of epithelial cells and transports vesicles associated with annexin XIIIb, a previously characterized membrane protein for apically transported vesicles [169]. A recent gene targeting study showed that KIFC3 plays a complementary role to cytoplasmic dynein in Golgi positioning and integration [170].

6 Orphans

These KIFs have no counterpart in Drosophila or C. elegans. KIF6 and KIF9 are located near the BimC family [25]. KIF9 is reported to interact with the Ras-like GTPase Gem, which suggests that it may play a role in cell shape remodeling [171, 172]. The distance between KIF9 and KIF6 is large. KIF7 has no evident homolog in Drosophila, C. elegans, or S. cerevisiae. Its function is as yet unknown, but it is predominantly expressed in the testis [25].

7 Cargoes of KIFs; Specificity and Redundancy

As mentioned above, KIFs convey various types of cargo. In some cases, a single KIF transports several distinct cargoes. For example, conventional kinesin (KIF5s) transports several cargoes such as mitochondria, lysosomes, vesicles containing Amyloid Precursor Protein (APP) and GAP43 [49, 173], ApoE2, the receptor for the Reelin ligand [48], or the AMPA-type glutamate receptor GluR2 [54]. Kinesin can also convey mRNA complexes [41], tubulin oligomers [38] and vimentin intermediate filament protein complexes [39, 40]. Another example is KIF3. KIF3 also conveys several different cargoes in distinct cell types. KIF3 transports vesicles associated with fodrin, important for neurite elongation in Neurons [86], and it conveys protein complexes for the formation of cilia and flagella along microtubules in various ciliated cell types [92, 93]. KIF3 is also a motor for the transport of pigment granules in pigmented cells in Xenopus [88–90] and is proposed to function in the trafficking of vesicles from the Golgi apparatus to the ER [174]. Thus, KIF3 is another example of a motor that carries out different tasks depending on the type of cell (Fig. 7). How the same KIF can transport different cargoes is an interesting and important question that needs to be addressed. On the other hand, different KIFs sometimes redundantly convey similar cargoes. Mitochondria are transported by KIF1Bα [64] as well as by conventional kinesin (KIF5) [28, 34]. KIF1A’s cargo is synaptic vesicle precursors [65], which are also conveyed by KIF1Bβ [66]. Another example is lysosomes which have at least two

8 Recognition and Binding to Cargoes

microtubule plus-end motors, kinesin and KIF2Aβ [35, 36, 152]. This redundancy could be one of the reasons for subtle phenotypes in knock-out mice in which some KIF genes have been knocked out.

8 Recognition and Binding to Cargoes

In many cases, a KIF has a specific cargo, while sometimes the same KIF, such as conventional kinesin and KIF3, conveys different cargoes. Thus, the fundamental question has been, how do KIFs recognize and bind to their cargoes? Studies of KIF17 and KIF13A clearly indicated for the first time how KIFs recognize and bind cargo molecules [77, 106]. KIF17 transports NMDA-type glutamate receptorcontaining vesicles through an interaction of its tail with mLin10-mLin2-mLin7NR2B. KIF13A, on the other hand, recognizes and binds to mannose-6-phosphate receptor-containing vesicles via an interaction between the KIF13A tail-β1 adaptinAP-1 adaptor complex and mannose-6-phosphate receptor (Fig. 8). In the case of conventional kinesin, it was demonstrated recently that it recognizes and binds cargoes such as APP- and GAP43-containing vesicles through KLC and APP interaction [173], and ApoE2-containing vesicles through interaction between KLC, and JIP1 and JIP2 [48]. Kinesin also recognizes cargoes such as AMPA receptor subunit GluR2-containing vesicles through its interaction with KHC-GRIP1-GluR2 [54] (Fig. 8). Thus, kinesin uses KHC and KLC to discriminate between distinct cargoes. A similar strategy may be used by KIF3. Thus in terms of how KIFs recognize and bind cargoes, two methods can be distinguished. Ĺ KIF tail–scaffolding protein(s) or adaptor protein(s)–membrane protein This is applicable to KIF17-mLin10-mLin2-mLin7-NR2B, KIF13A-13A-β1adaptin-AP1adaptor complex-M6PR, KLC-JIP1/JIP2-ApoE2, the reelin receptor and KIF5 (KHC)-GRIP1-GluR2 (Fig. 8). Ĺ KIF tail–membrane protein Kinesin was reported to bind directly to membrane proteins through its interaction with KLC. Sunday Driver (also called JSAP1 and JIP3) [44] and APP [173] were proposed to be membrane proteins that directly bind to kinesin through KLC, but it needs to be demonstrated whether Sunday Driver (JSAP1 ad JIP3) is indeed a membrane protein.

We are just beginning to understand how KIFs recognize and bind their specific cargoes. So far we only know that rather complex mechanisms are involved in these events. This may be related to how the association and dissociation between KIFs and cargoes are regulated. Another question is how KIFs recognize and bind to protein complexes such as tubulin and intermediate filament proteins, chromosomes and mRNAs. This is obviously a significant question that needs to be answered in the near future.

19


9 How to Determine the Direction of Transport

Another intriguing question is how a cell regulates the direction of transport, for example the transport to axons versus dendrites. Kinesin (KIF5) transports APPcontaining vesicles to the axon while the same motor conveys AMPA-receptorcontaining vesicles to dendrites. In this case it was shown that GRIP1, which binds KHC and JSAP1, which binds KLC, steers kinesin to dendrites and axons, respectively [54]. Thus, GRIP1 and JSAP1 are functioning as drivers for kinesin (KIF5). There are motors that transport cargoes mainly to dendrites. KIF17 conveys NMDA receptors mainly to dendrites [106] and KIFC2 transports a new type of multivesicular body-like organelle to dendrites [168]. KIF21B was also proposed to be a motor for dendritic transport [114]. How differential transport is performed is clearly a fundamental question that needs to be solved in the near future. The mechanism is probably not simple and could involve several distinct events. Thus, interestingly, cells use many KIFs and very precisely control the direction and velocity of transport of various distinct cargoes that are fundamental in basic cellular functions. This field is obviously very important and rapidly advancing. Further studies need to be carried out to fully understand the mechanism of intracellular transport.

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165 SHANKS, R. M., R. J. KAMIENIECKI, and D. S. DAWSON. 2001. The Kar3-interacting protein Cik1p plays a critical role in passage through meiosis I in Saccharomyces cerevisiae. Genetics 159: 939–951. 166 TROXELL, C. L., M. A. SWEEZY, R. R. WEST, K. D. REED, B. D. CARSON, A. L. PIDOUX, W. Z. CANDE, and J. R. MCINTOSH. 2001. pkl1(+) and klp2(+): Two kinesins of the Kar3 subfamily in fission yeast perform different functions in both mitosis and meiosis. Mol. Biol. Cell 12: 3476–3488. 167 HANLON, D. W., Z. YANG, and L. S. GOLDSTEIN. 1997. Characterization of KIFC2, a neuronal kinesin superfamily member in mouse. Neuron 18: 439–451. 168 SAITO, N., Y. OKADA, Y. NODA, Y. KINOSHITA, S. KONDO, and N. HIROKAWA. 1997. KIFC2 is a novel neuron-specific C-terminal type kinesin superfamily motor for dendritic transport of multivesicular body-like organelles. Neuron 18: 425–438. 169 NODA, Y., Y. OKADA, N. SAITO, M. SETOU, Y. XU, Z. ZHANG, and N. HIROKAWA. 2001. KIFC3, a microtubule minus end-directed motor for the apical transport of annexin XIIIb-associated Triton-insoluble membranes. J. Cell Biol. 155: 77–88. 170 XU, Y., S. TAKEDA, T. TANAKA, Y. NODA, Y. TANAKA, and N. HIROKAWA, 2002. Role of KIFC3 motor protein in Golgi positioning and integration. J. Cell Biol. 158: 293–303. 171 BERNSTEIN, M., P. L. BEECH, S. G. KATZ, and J. L. ROSENBAUM. 1994. A new kinesin-like protein (Klp1) localized to a single microtubule of the Chlamydomonas flagellum. J. Cell Biol. 125: 1313–1326. 172 PIDDINI, E., J. A. SCHMID, R. de MARTIN, and D. G. DOTTI. 2001. The Ras-like GTPase Gem is involved in cell shape remodelling and interacts with the novel kinesin-like protein KIF9. EMBO J. 20: 4076–4087. 173 KAMAL, A., A. ALMENAR-QUERALT, J. F. LEBLANC, E. A. ROBERTS, and L. S. GOLDSTEIN 2001. Kinesin-mediated axonal transport of a membrane compartment containing beta-secretase

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1

Myosin Superfamily Proteins Michele C. Kieke and Margaret A. Titus

University of Minnesota, Minneapolis, USA


1 Introduction

Actin filaments and the myosin motors associated with them play important roles in many dynamic biological processes. The classic example of actin filaments and myosin at work is during skeletal muscle contraction. But the functions of actin and myosin extend to many other cellular events, such as motility, adhesion, endocytosis, cytoplasmic streaming, neuron growth, structural maintenance and polarization. Like molecular cars on an actin track, myosins transport organelles and other cellular components, such as mRNA. Myosins can also aid in the formation or maintenance of an organized actin-based structures (such as the stereocilia of hair cells), and play roles in intracellular signal transduction pathways ([1–4], for reviews summarizing myosin functions). Myosins are molecular motor proteins that use the energy from adenosine triphosphate (ATP) hydrolysis to generate force for directed movement along actin filaments. Myosins are composed of one to two heavy chains, and one or more light chains. The heavy chain consists of several major domains and can include various other subdomains or protein motifs (Fig. ??). The relatively conserved N-terminal motor or ‘head’ domain has binding sites for both ATP and F-actin. A short region joining the head and neck (termed the ‘converter domain’) is believed to be responsible for producing the force required for movement. The neck domain contains one to six light chain binding regions termed IQ motifs, repeats of approximately 23 to 30 residues containing the sequence IQXXXRGXXXRK [5]. The divergent C-terminal globular ‘tail’ has been implicated in binding cargo and targeting the myosin to its proper location in the cell [6]. Some myosins also feature a coiled-coil domain that promotes heavy chain dimerization.


2 Myosin Superfamily Proteins

1 Introduction 3

The founding member of the myosin family, filament-forming class II muscle myosin, was discovered nearly a century ago, and its role in muscle contraction has been studied extensively [7, 8]. A combination of biochemical and molecular approaches has led to the identification of over 20 different myosin classes [9]. Because of the extensive amount of knowledge acquired regarding the properties of myosin II it is referred to as ‘conventional’ myosin; all other types of myosin are referred to as ‘unconventional’. The first unconventional myosin, myosin I, was described in 1973 by Pollard and Korn [10, 11]. They isolated a protein with enzymatic properties similar to myosin II (i.e. it exhibited actin-activated Mg+2 -ATPase and ATP-sensitive binding to actin) from the common freshwater amoeba Acanthamoeba castellanii that had a lower molecular weight than muscle myosin II (125 versus 200 kD) and did not form filaments. In addition, it was determined that myosin I had one head rather than two. Careful analysis of this unusual molecule revealed that it was indeed a bona fide myosin [12]. This work provided the first insights into the potential diversity and functions of the myosin superfamily. Myosin superfamily members are grouped into different classes based on phylogenetic analysis of motor domains (Fig. 2) [9, 13, 14]. Each class is designated by a Roman numeral, largely in the order of their discovery (note that we will refer to myosins using Arabic numbers for simplicity). A total of 18 classes have been officially designated, but there are at least six novel myosins that have yet to be classified. Myosins have been found in a variety of organisms but no one class is universally expressed in all phyla. The yeasts Saccharomyces cerevisiae and Schizosaccharomyces pombe have a total of five myosin genes from three classes (M1, M2, M5). These myosins are shared by higher organisms, ranging from Caenorhabditis elegans (C. elegans) to mammals. The human genome includes about 40 myosin genes from 12 classes [9]. M8, M11, and M13 are only found in plants [15]), and M14 myosins are found in parasites such as Toxoplasma gondii and Plasmodium falciparum [9]. A unique class of myosins (as yet undesignated) has been found in the ciliated protozoan Tetrahymena [16, 17], suggesting that these organisms have a distinctive set of myosins. Cells typically express multiple myosins – the expression of at least a dozen myosins in a single cell type has been described [18]. This includes several different classes of myosin, as well as two or more isoforms of several classes. Myosins from the same class can have isoform-specific roles, such as M5 isoforms in mammalian cells and yeast [19], or they can have functionally overlapping roles, such as the M1s in Dictyostelium discoideum and Saccharomyces [20–23].

← Fig. 1 Domain structure schematic for characterized myosin genes. Schematic illustrating the variety of known structural motifs found in myosin genes. A general box diagram is given for each class, although individual members of the same class may vary depending on the organism and/or particular isoform.


Fig. 2 Unrooted myosin superfamily phylogenetic tree. Phylogenetic tree from [9], constructed using myosin motor domain sequences. Species names are listed in the Abbreviations table, and some gene names have been shortened to save space. Se-

quences predicted in full or in part from genomic clones are indicated by an asterisk. Figure and legend text reprinted from Molecular Biology of the Cell (2001, 12: 780–794), with permission from the American Society for Cell Biology.

2 Functional Properties of Myosins

The existence of not only diverse myosin classes but also isoforms within those classes is a clear indication of the range of roles for this motor protein superfamily. Myosins have been implicated in many fundamental biological processes, and their functional significance is accented by the discovery that mutations in unconventional myosin genes in worms, flies, mice, and humans cause severe phenotypes such as deafness, blindness, and sterility. Despite the important and diverse roles for myosins, much remains to be learned about them at the molecular level. Only in the last decade have the intricacies of myosins been realized due to the characterization of properties such as structure, kinetics, localization, directionality, and putative functional roles. In the past five years, the rate of discovery in the myosin field has increased significantly – we are learning much about what myosins do and how they work. This chapter will summarize what is currently known about the myosin superfamily members, including their cellular functions and their roles in various diseases.


The motor domains of myosins are relatively well-conserved, although kinetic properties can vary from class to class. Some of the myosins are short-duty motors that most likely function as part of an assembly of motors (e.g. M2). Others are processive and can move for long distances without releasing from the actin filament (e.g. M5). It should be noted, however, that not all members of the same class have similar properties. For example, mammalian M5 is highly processive while yeast M5 is not [24, 25]. Myosins can move toward one end of the actin filament or the other, a property referred to as ‘directionality’. In contrast to the conservation between motor regions, the tail regions are quite divergent between the different myosin classes. A variety of protein motifs are found in the myosin tail region and a few are also located in the motor domain. These kinetic and structural properties can offer insight into myosin functions. 2.1 Directionality and Processivity

The directionality and processivity of myosins contribute to their diverse cellular functions [26]. Myosins move unidirectionally on actin filaments, either toward the plus-end or the minus-end of actin. The arrangement of actin filaments in the cell periphery is generally with the plus-ends (fast-growing) toward the plasma membrane. Myosins moving toward the plus-ends (i. e. M1, M5) would be expected to carry cargo and/or to localize at the cell periphery. On the other hand, myosins that move in the opposite direction, such as M6 and M9 [27, 28], would be predicted to have complementary roles. Perhaps minus-end directed myosins provide a means to transport cargos into the cell rather than toward the periphery, and thus provide an opposing activity for plus-end directed motors.

5


In addition to the directionality of the motor, the time spent bound to actin filaments is an important consideration when thinking about myosin functions. ‘Duty ratio’ refers to the filament-bound state of myosin, or more specifically to the fraction of the ATPase cycle that myosin spends strongly bound to actin. Myosins that spend a significant proportion of the cycle tightly bound to actin have a ‘highduty ratio’, while those that spend a small fraction of the cycle tightly bound are considered to have a ‘low-duty ratio’. Conventional muscle M2 is a low-duty motor [7], a property reflected in the fact that large assemblies of these motors work in coordination with each other during muscle contraction. A low-duty motor would not be suitable for persistent transport of cargo, as it would detach after a single step on actin. In contrast, motors with a high-duty ratio are good candidates to serve as long-range transport motors. Such single myosin molecules can translocate along an individual actin filament for long distances. It is also worth noting that high-duty ratio is a trait that would be advantageous for a motor suspected to provide force and maintain tension. High-duty ratio motors can be identified by detailed kinetic analyses or by in vitro motility assays of single myosin molecules. Myosins from at least three classes are processive – M5 [24], M6 [29–31], and M9b [27]. M5 and M6 are two-headed myosins, while M9b is a single-headed motor. This structural difference suggests distinct mechanisms for processive movement. 2.2 Protein Motifs Found in Myosins

The myosins possess a variety of known protein motifs at either their N- or Cterminus (see Fig. ??) that offer insight into possible myosin function and localization. These motifs can be generally categorized as membrane localization or targeting, signaling, structural, or myosin-specific. A major effort is currently underway to identify the contribution of each of these regions to myosin function. Several classes of myosin have intriguing N-terminal domains. The motor domain of M3 has a protein kinase domain that is required for its function in the termination of the phototransduction signal cascade in the Drosophila eye [32]. M16 has a series of ankyrin repeats at its N-terminus [33], and M18 has an Nterminal PDZ domain [34, 35]. The role of these two different domains in mediating protein–protein interactions suggests that the head may play a role in myosin localization apart from simply binding to actin. M15 has a unique extension at its N-terminus [36], the function of which remains to be determined but could play a role in its motor function and/or localization. The known membrane localizing motifs in the myosin tail are PH (pleckstrin homology), polybasic regions, and FERM (band 4.1/ezrin/radixin/moesin) domains. PH domains bind to the charged head groups of phospholipids in the plasma membrane and are often involved in signal transduction pathways. Thus, myosins with PH domains (M10 and the novel myoM from Dictyostelium [37–39] might act as adapters or relay proteins in signal transduction cascades, functioning to localize signaling proteins to the plasma membrane. The M1s all have a polybasic


domain that binds to anionic phospholipids with high affinity [2, 40], suggesting that this region directs the localization of this myosin class to a particular subset of intracellular membranes. It should be noted, however, that this domain also binds to actin and might be able to contribute to the association of M1s to actin filaments [41, 42]. FERM domains were first identified as the N-terminal region of the ERM (ezrin, radixin, moesin) family of actin-binding proteins that shares homology with the Nterminus of band 4.1 [43]. The band 4.1 protein family members participate in cell signaling events that regulate processes such as cell growth, cytoskeleton dynamics, and cell-substrate adhesion [44–46]. The FERM domains of ERM proteins bind to phospholipids as well as the cytoplasmic region of transmembrane receptors such as the CD44 adhesion molecule, linking them to the actin cytoskeleton [44, 46]. Talin, another FERM-domain containing protein, binds both the cytoplasmic tail of integrins via its FERM domain and the actin cytoskeleton via its C-terminus [47], forming mechanical links between the two components (similar to the ERMs). The M7, M10, M12, and M15s each have one or two FERM domains in their C-terminus [48]. Several different classes of myosins contain motifs in their tail regions that are commonly found in signaling proteins. These include rho-GAP (rho-GTPaseactivating protein) domains and SH3 (src homology 3) domains. M9s possess a rho-GAP domain [49, 52]. The rho-GTPases have been reported to regulate actin cytoskeleton organization [51], suggesting that M9 may act as both a motor and a signaling molecule. M4 and one subtype of M1 (the amoeboid type) each have a single SH3 domain [48]. The SH3 domain of the amoeboid M1 binds to proteins that recruit the Arp2/3 complex, implicating these motors in regulating or contributing to the directed polymerization of actin in yeast and Dictyostelium [52–55]. There are additional tail motifs that contribute either to the overall structure or function of given myosins. A number of unconventional myosins (M5, M6, M7, M8, M10, M11, M12, and M18) have stretches of predicted coiled-coil, indicating that these myosins function as dimers. The amoeboid M1s have a domain rich in the amino acids glycine, proline, and alanine (or serine or glutamate) that binds to actin with high affinity [56]. The M9s have one or more zinc-binding regions in the tail region [49, 50]. Zinc-binding motifs are also found in several signaling proteins such as protein kinase C, raf kinase, and Vav [57]. Mammalian M5 and M10 both have PEST regions in their tails [37, 58], sequences enriched in the amino acids proline, glutamate, serine, and threonine. It has been suggested that the PEST regions are susceptible to cleavage by calpain, a calcium-dependent protease [59]. If this cleavage occurs in vivo it would result in separation of the motor domain from the tail. One of the most ubiquitous and least characterized domains found in myosins is the MyTH4 (myosin tail homology) domain. MyTH4 domains are found not only in myosins (M4, M7, M10, M12, M15) but also in a kinesin from plants [48] as well as in a protein that participates in signaling processes that occur during axon guidance [60]. The precise role for the MyTH4 motif is not known, but preliminary evidence suggests that MyTH4 domains might bind to microtubules [48, 61]. In many cases

7


the MyTH4 domain immediately precedes a FERM domain, suggesting that these two domains together form a functional unit. Evidence in support of a critical role for this domain comes from the finding that mutations in the MyTH4 domains of M7 and M15 cause disease [36, 62]. Finally, both M5 and M6 have unique globular tail domains at their extreme C-termini that direct the intracellular localization of each of these myosins [19, 63]. In the case of M5a, there are several unique exons found only in M5a expressed in the skin [64]. The skin-specific exon has been shown to play an essential role in targeting M5a to the melanosome [65]. 2.3 Myosin Regulation

Motor function must be precisely regulated. The activity of myosins is regulated by either calcium or phosphorylation [3]. Most myosin light chains (LCs) are calmodulin or members of the calmodulin superfamily that bind to IQ motifs in the neck region. Myosins have from one to six IQ motifs per heavy chain (HC) and the LCs bind in the absence of calcium. Calcium binding to the associated calmodulin can result in a weakening of affinity and may result in LC loss. Binding also influences the actin-activated ATPase activity, actin binding, and can result in either reduced or increased translocation velocity of actin filaments in in vitro motility assays [66, 67]. The relationship between calcium binding to calmodulin LCs, actin binding, actin-activated Mg2+ -ATPase activity (AAA) and in vitro motility is different for each myosin. In the case of M5, micromolar calcium concentration results in higher AAA and an increased affinity for actin, but decreased motility [66, 68]. In contrast, higher calcium levels result in both decreased AAA and motility. The reader is referred to several recent reviews for a more detailed discussion of the effects of calcium-calmodulin regulation of various myosins [3, 67, 69, 70]. Regulation by HC phosphorylation occurs in the motor domain of amoeboid M1 and M6 at a residue called the ‘TEDS rule’ site [71] that resides 16 residues Nterminal from the conserved DALAK sequence located near the actin-binding site sequence. The TEDS site residue is usually either phosphorylatable (amino acids Tor S) or acidic (E or D), thus the ‘TEDS’ designation. The importance of phosphorylation in regulating unconventional myosin activity is highlighted by studies of amoeboid M1s in fungi or amoebae. These motors require phosphorylation by a PAK family kinase that is regulated by small G-proteins for actin-activated ATPase activity [72]. It should be noted that phosphorylation regulates the activity of only the lower eukaryotic forms of M1 because only these M1s have an S or T at the TEDS rule site. Mutation of the TEDS rule site to A in either yeast or Dictyostelium amoeboid M1 abolishes in vivo function [73–75]. Phosphorylation is not essential for the activity of all amoeboid M1s. Mutation of the TEDS rule site of Aspergillus MYOA to A results in a loss of virtually all AAA and motor activity, yet does not impair the in vivo function of this myosin [76]. This suggests that some M1s may play a non-motor role in certain contexts. The higher eukaryotic M1s have an E or D at

3 Diverse Functions for Myosins 9

that site and their regulation is likely to occur via Ca2+ -calmodulin rather than HC phosphorylation. Mammalian M6 is also phosphorylated by a PAK kinase at the TEDS rule site and this has been shown to moderately activate M6 motility in vitro as well the rate of Pi release [29, 77, 78]. The exact relationship between changes in the rates of in vitro motility and alterations in the kinetic properties of phosphorylated M6 are not yet clear, but ongoing investigations should resolve this issue. Recent experiments have revealed that phosphorylation in the myosin tail region can regulate the activity of myosins not by affecting enzymatic activity, but by changing the affinity of the myosin for its cargo. Calcium/calmodulin dependent kinase II (CaMKII) phosphorylates a serine in the C-terminal organelle-binding domain of Xenopus M5 in a cell-cycle dependent manner [79, 80]. The onset of mitosis results in an increase in M5 tail phosphorylation and a concomitant decrease in the level of M5 associated with melanosomes in melanophores. The increased phosphorylation is predicted to cause cargo (melanosome) release from M5 and the treatment of mitotic Xenopus extracts with CaMKII inhibitors strongly inhibits M5-melanosome dissociation. HC phosphorylation in the myosin tail region may be one mechanism for regulating myosins involved in organelle transport.

3 Diverse Functions for Myosins

Myosins participate in diverse biological functions (Tab. 1). While a number of different roles for myosins have been defined over the past few years, it has become increasingly clear with the identification of additional novel myosins that the full spectrum of functions for this family of motors has yet to be appreciated. Thus far, myosins have been implicated in a variety of processes that include cell motility and adhesion, intracellular transport, maintenance of actin-rich structures, membrane trafficking and signal transduction [1, 2]. 3.1 Non-muscle Contractile Structures

M2 forms filaments not only in muscle but in non-muscle cells as well. The nonmuscle M2 filaments are dynamic structures – they can be assembled and disassembled in particular intracellular locations as needed [81]. The critical role for M2 in cell motility will be discussed below; it is also involved in forming actinrich contractile structures during processes such as cytokinesis, morphogenesis and wound healing [82–84]. M2 is found interspersed among actin filaments that form contractile bundles between adjacent epithelial cells and in the cortical actin network that supports cell structure. At the cell periphery, a contracting acto-M2 network is thought to prevent pseudopod formation, assist in the retraction of the rear of the cell and provide resting cortical tension [85–87]. It is also found in actin filament stress fiber bundles that are thought to play a role in anchoring a cell to the substrate [88].

10 Myosin Superfamily Proteins Table 1 Summary of putative myosin functions. A brief listing of the known functions of sev-

eral different myosin classes. The question marks (?) indicate functions that have been inferred based on localization, initial phenotypic analyses and analysis of phenotypes resulting from ectopic expression of wild-type or mutant forms of a given myosin Myosin Putative role M1

M2

M3 M5

M6

Relevant organism or cell type/gene

Cell motility (leading edge) Acanthamoeba, Dictyostelium, mammalian cells Microvilli structure Intestinal epithelial cells/MYO1A Vesicle transport, Intestinal epithelial cells/MYO1A endocytosis Endocytosis Dictyostelium, yeast Endosome recycling Polarized epithelial cells/MYO1D Early endosome recycling Dictyostelium/myoB Signal transduction Stereocilia in mouse/MYO1C Cytokinesis Dictyostelium, mammal. cells, echinoderm Fission yeast/MYO2 and MYP2; budding yeast/MYO1 Morphogenesis Drosophila and Dictyostelium Cortical bundles, stress Mammalian cells fibers Cell motility (rear of cell) Acanthamoeba, Dictyostelium, mammalian cells Cell integrity and structure Drosophila photoceptor cells/ninaC Signal transduction Drosophila eye/ninaC Signal transduction (?) Limulus eye Secretory vesicle transport Budding yeast/MYO2 Vacuole Budding yeast/MYO2 transport/inheritance Golgi and peroxisome Budding yeast/MYO2 inheritance ASH1 mRNA transport Budding yeast/MYO2 Mitotic spindle orientation Budding yeast/MYO2 Particle transport Fisson yeast/MYO4 (MYO52) SER and vesicle transport Neurons Pigment granule transport Frog, fish, mouse/M5a Particle transport, filopod DRG neuron growth cones extentsion Intracellular trafficking Particle transport Transport and membrane organization (?)

Clathrin-mediated endocytosis

Mammal. cells/M5b, M5c Drosophila embryo/95F Drosophila oocyte/95F Drosophila developing sperm/95F C. elegans developing sperm/spe-15 Mouse hair cells, stereocilia/Myo6, sv Epithelial cells Polarized epithelial cells

Reference(s) [101, 102, 106] [13, 182, 184] [185, 206, 207] [20–23] [208] [209] [197, 221, 222, 224] See [89, 90] and references in [83] [92, 93] [84, 96, 97] See reference in [88] [85, 98–100, 106] [187, 188] [186, 217, 218] [219] [120, 124] [131] [133, 134] [137, 138, 141] [135, 136] [129, 257] [148–150, 192] [153, 154, 157–159] [106] [211, 212] [178] [179] [180] [181] [195, 196, 198] [191, 198–200] [63, 213, 214]

3 Diverse Functions for Myosins 11 Table 1 (Continued).

Myosin Putative role

Relevant organism or cell type/gene

Reference(s)

M7

Dictyostelium/myoi

[111]

Sensory hair cells/Myo7a, shaker-1

[197, 202, 204, 205]

Retinal pigment epithelia (RPE) Rat, human (GAP domain)

[175, 176] [49, 226, 227]

Mammalian cells

[193]

Toxoplasma/TgMyoA and Plasmodium

[113–116]

M9 M10 M14

Cell adhesion, filopod extension Lateral links, stereocilia organization Melanosome transport Signaling to actin cytoskeleton (?) Transport (?), filopod extension Gliding motility

Early studies showed the presence of M2 in the contractile ring at the equator of dividing cells and that the injection of function-blocking antibodies resulted in an inhibition of cytokinesis in echinoderm embryos [89, 90]. Subsequent molecular genetic studies in the Dictyostelium and yeast provided direct proof of a role for non-muscle M2 in cytokinesis [83, 91]. While budding yeast and Dictyostelium only have a single M2, the fission yeast Schizosaccharomyces pombe has two M2s (Myo2p and Myp2p) that seem to play non-overlapping roles in cytokinesis [92, 93]. Interestingly, in the case of Dictyostelium, M2 is only essential for cytokinesis when cells are in suspension where mutant cells become large and multinucleate. The cells are able to undergo a M2 independent cytokinesis when plated onto a substrate. Thus, Dictyostelium has more than one distinct mode of cytokinesis. One mode is dependent on M2 (‘cytokinesis A’), while another is M2-independent and instead requires adherence to a substrate (‘cytokinesis B’; [94, 95]). Finally, non-muscle M2 is required for morphogenesis in both Dictyostelium and Drosophila melanogaster [84, 96]. For example, mutations in Drosophila M2 result in embryos with defects in dorsal closure, head involution and axon patterning [84]. Further analysis of the mutant embryos suggests M2 is involved in the maintenance of cell shape and cell sheet movement. Embryos mutant for myosin phosphatase, a negative regulator of non-muscle M2, also exhibit dorsal closure defects [97].

3.2 Cell Motility and Adhesion

Myosins play a central role in cell motility. Immunofluorescence experiments have shown that M2 is concentrated in the lamellae, posterior and cortical region of polarized, locomoting cells [98–100]. Conversely, M1s are located at the leading edges of lamellipodial projections of migrating Dictyostelium and Acanthamoeba [101, 102] and in filopodia, lamellipodia, and growth cones of mammalian cells [103–105]. These findings suggest that M1 is involved in forward extension of the cell and that M2 functions in retraction at the rear of the cell body.


Dictyostelium cells mutant for either M1 or M2 have defects in locomotion [96], further supporting the essential roles for myosins in cell motility. Both M1 and M2 are required for dictating where a pseudopod is formed. M2 filaments at the cortex inhibit protrusion and the loss of this myosin results in the extension of pseudopodia around the periphery of the cell and a 50% decrease in velocity [87]. Dictyostelium lacking the M1s myoA or myoB are not deficient in extending pseudopodia, rather they make excess pseudopodia that are not extended in the typically ordered manner (i. e. one at a time) [106, 107]. Similarly, chromophore-assisted laser inactivation (CALI) of M1 in neuronal growth cones causes expansion of lamellipodia [108]. These observations suggest that M1s have a general role in regulating pseudopod formation, an actin-dependent process. The amoeboid M1s have recently been shown to be linked to the Arp2/3 actin-polymerization machinery and may aid in focusing the site of its action to direct the site of the growing pseudopod [52–54, 109]. Characterization of a Dictyostelium M7 null mutant revealed a role for this class of myosin in cell adhesion [110]. Mutant cells displayed reduced contact between the cell surface and substrate during migration. Loss of adhesion to surfaces results in a defect in phagocytosis [110, 111] due to reduced binding to particles. Preliminary analysis suggests that the loss of adhesion is not due to a loss of cell surface receptors. Since the early stages of phagocytosis and cell motility are similar, it is not completely surprising that M7 is important for both processes. It has been proposed that this myosin participates in adhesion receptor complex assembly and disassembly at the plasma membrane [110]. Apicomplexan parasites such as Toxoplasma gondii and Plasmodium falciparium move by an unusual means termed gliding motility [112]. These cells tightly anchor themselves to their host cell and then propel themselves into the cell by translocating the sites of attachment towards the rear of the cell. No change in cell shape is involved in their movement, in contrast to amoeboid cells. Treatment of cells with actin depolymerizing drugs inhibits gliding motility [113]. These parasites uniquely express M14, a class of myosin that has a motor domain, virtually no neck, and a short tail [114, 115] and one of these, TgMyoA, is localized to the plasma membrane [79, 80]. Recent biochemical analysis of TgMyoA [116] reveals that it is associated with a novel, relatively large LC (31 kD versus 20 kD) that is calmodulin-like and binds to the C-terminal 53 amino acids of the TgMyoA HC. The biochemical properties of this unusual myosin are remarkably similar to those of M2. In spite of a low degree of sequence similarity to other myosins in the converter and LC binding domains, TgMyoA propels actin filaments at a rate of ∼5 µm s−1 and has a step size of 5 nm. These data suggest that this myosin is capable of generating the forces required for propelling gliding motility. The development of an inducible expression system for Toxoplasma [117] should facilitate analysis of the role of this fascinating myosin in gliding motility. Myosins contribute to various aspects of cell migration – directed protrusion of the lamellipodium, adhesion to the substrate and retraction of the rear of the cell. The importance of an actomyosin-based mechanism for the propulsion of a wide array of cells is evidenced by the finding that an unusual myosin, M14, powers


a unique form of motility. Given the complex series of events that occur during migration, it is likely that other myosins might also be found to participate in cell migration as new members of the myosin superfamily are studied. 3.3 Organelle/Cellular Component Transport

The proper localization of intracellular components at the necessary time(s) is important for cell function. Actin- and microtubule-dependent motors each play essential roles in ensuring that cellular components (cargos) reach the right place at the right time. Current evidence suggests that microtubule motors serve to translocate organelles for long distances while the actin-based motors move things locally [118, 119]. Myosins appear to transport cellular cargos that range from vesicles to mRNA, and the best-studied of the transport myosins is M5. Evidence for cargo transport by M5s comes from the study of yeast as well as pigment cells from mouse, fish and frog [118, 119]. The yeast Saccharomyces cerevisiae has two M5 genes (MYO2 and MYO4). The product of the MYO2 gene (Myo2p) is essential for viability Yeast with the temperature sensitive (ts) mutation myo2-66 (a point mutation in the N-terminal actin-binding domain) exhibit an accumulation of secretory vesicles and arrest in a large, unbudded state at the restrictive temperature [120, 121]. It has been shown that Myo2p binds vesicles via its tail region and transports them along actin filaments from the mother cell to the bud [122–124]. Genetic data suggest that the Myo2p tail also binds to a kinesin homolog (Smy1p) and a vesicle-associated Rab protein (Sec4p) [14, 122, 124, 127, 128]. The association between Myo2p and Smy1p is supported by co-localization and yeast two-hybrid experiments [121, 125]. These data contribute to the growing body of evidence that actin filament and microtubule systems cooperate. An M5 from fission yeast, Myo4p (also referred to as Myo52), has also been implicated in organelle transport. Myo4p/Myo52p is localized to small punctae that accumulate at sites of polarized growth at the ends of the cells and is also found at the septum. These particles appear to move rapidly within the cell. Deletion of myo4/myo52 results in rounder cells that exhibited an accumulation of internal vesicles [93, 129]. In addition to its role in transporting materials to sites of cell growth and division, Myo4p/Myo52p has also been found to play a role in vacuole fusion in response to osmotic stress [130]. Further work has revealed a function for M5 in vacuole inheritance in budding yeast [131]. The myo2-66 ts mutant was found to exhibit aberrant vacuole partitioning and inheritance by the daughter cell prior to division. A screen for additional myo2 alleles resulted in the identification of the myo2-2 mutant yeast which was also defective for vacuole inheritance but did not display the cell growth defects observed with myo2-66 [122]. The mutation was found to reside in a region of the Myo2p tail specifically required for vacuole binding [132]. It appears that Myo2p is required for two separate processes in yeast: the transport of secretory vesicles important for polarized cell growth and the segregation of vacuoles (an event not


essential for polarized cell growth). This result has exciting implications for the identification of multiple cargos, and thus multiple cargo binding receptors, for the same molecular motor protein. Like vacuoles, peroxisomes and Golgi elements are also segregated into the yeast bud during cell growth. Movement of these organelles from mother to daughter requires the actin cytoskeleton, and it was demonstrated that Myo2p provides the means for their transport. In cells containing the myo2-66 ts allele, peroxisome movement stops abruptly at the non-permissive temperature [133] and the inheritance of late Golgi elements is inhibited [134]. In addition to its role as a transport motor, yeast Myo2p has been shown to participate in mitotic spindle orientation [135, 136]. Proper spindle orientation is important for accurate chromosome segregation. The tail region of Myo2p binds Kar9p (a microtubule-associated protein) in yeast two-hybrid and co-immunoprecipitation experiments, and Myo2p is required for Kar9p transport into the bud. It appears that Myo2p and Kar9p both play essential roles in spindle orientation. Current models propose that Kar9p binds to both microtubules and Myo2p and that Myo2p transports both into the growing bud [135, 136]. This analysis provides another link between the actin filament and microtubule cytoskeleton systems. The other M5 in budding yeast, Myo4p, has been shown to participate in the asymmetric localization of the transcriptional repressor, Ash1p [137, 138]. Ash1p represses HO endonuclease, the protein that induces mating type switching in yeast [139]. Thus, localization of Ash1p to the budding daughter cells suppresses mating type switching [137, 138]. ASH1 mRNA is asymmetrically segregated to the bud tip of the growing daughter cell [140, 141], and its movement into the bud tip can be monitored by tagging it with green fluorescent protein (GFP) [142, 143]. Myo4p and two other She proteins (She2p and She3p) co-localize with ASH1 mRNA [140], and all four components co-immunoprecipitate from cell extracts [22, 131, 146, 147]. Myo4p co-immunoprecipitates with the She3p adapter in the absence of RNA, while She2p (a second adapter) recruits the Myo4p–She3p complex to ASH1 mRNA [144– 147]. The association between She3p and Myo4p appears to be mediated by their coiled-coil regions [144]. M5 may also have a vesicle transport role in neurons, where it was found to move extruded axon smooth endoplasmic reticulum (SER) and brain vesicles in vitro [148, 149]. Antibodies against the M5 head or tail were able to inhibit this movement [148, 149] and immunogold electron microscopy revealed the co-localization of M5 and a kinesin on the SER vesicles [149], consistent with a model of actin filament and microtubule system cooperation. M5 has also been found to be associated with organelles in live neurons [150]. Analysis of neurons from dilute mice lacking M5a reveals that organelles still undergo bidirectional movement along microtubules. However, loss of this motor causes an accumulation of organelles in regions of the cell enriched for dynamic microtubule ends, suggesting that M5a is responsible for dispersal of these organelles to actin-rich areas of the cell [65, 150]. The role of M5 in pigment granule transport has been well characterized in frog (Xenopus laevis), fish and mouse melanophores – cell types that have traditionally served as model systems to study intracellular transport mechanisms


[151, 152]. Pigment granules in Xenopus melanophores undergo rapid and tightly regulated aggregation and dispersal, creating changes in color in response to external stimuli [151]. Transport of pigment granules seems to involve both the actin filament and microtubule cytoskeletal systems [153, 154]. Purified granules from frog melanosomes contain bound M5, kinesin II, and dynein, and have been shown to move along actin filaments (as well as microtubules) in vitro [80, 155] (Fig. 3a). Recent in vivo analysis of melanosome movement suggests that the motor action of kinesin and dynein are in a ‘tug-of-war’ with M5 and that M5 activity is downregulated during the aggregation phase [156]. Melanosome transport to the cell periphery by M5 has also been studied in mouse melanocytes [152]. Mouse melanosomes containing pigment are transported from the center of the cell to the periphery, where they are taken up by keratinocytes. The pigment is eventually deposited into the hair shaft, causing changes in coat color. Loss of M5a (dilute) in the mouse results in a lightened or ‘diluted’ coat color because the pigment granules are not transported to the dendritic tips of the melanocytes in both cultured cells [157, 158] and in situ [159]. M5 was shown to localize to the melanosomes [160–162]. Expression of the globular tail domain in cultured melanocytes disrupts the binding between endogenous M5 and melanosomes, causing a clumping of these organelles in the central region of the cell that leads to the dilute phenotype [163]. Mutations at two other murine loci (ashen and leaden) cause the same phenotype as dilute – a lightened coat color due to pigment granule transport defects [164]. The ashen gene was found to encode the Rab27a GTPase, and leaden encodes a Rab binding protein named melanophilin [165, 166]. The molecular relationship between the gene products of these loci has only recently been elucidated. The receptor for M5a (dilute) has been identified as a complex of two proteins, Rab27a (ashen) and melanophilin (leaden, also referred to as Slac2-a) [163, 167– 172]. Rab27a was shown to first bind the melanosome, then recruit melanophilin, which subsequently recruits M5a into the complex (Fig. 3b). The C-terminal region of melanophilin binds to the C-terminal domain of M5a [172, 173]. Binding of melanophilin to Rab27a occurs in a GTP-dependent manner, providing a means of controlling M5a association to the melanosome [163, 174]. It is of interest to note that M7a is associated with melanosomes in retinal pigment epithelium (RPE) and that these melanosomes are not correctly distributed in the shaker-1 M7a mutant [175]. This observation, coupled with the recent identification of a rabphilin [176] as an M7a tail binding protein, suggests that M7a may also have a role in melanosome transport in the RPE. M6 has also been implicated in organelle transport, consistent with it being a high-duty ratio motor [177]. The fly M6, designated 95F, is present on moving particles of unknown identity in the syncytial blastoderm. These particles appear to move on linear tracks. An inhibitory polyclonal antibody against 95F was able to block the linear (non-random diffusion) motion of the particles. The effect was similar to what was observed following treatment of the embryos with either actin-depolymerizing agent cytochalasin D or ATP inhibitor dinitrophenol (DNP), suggesting that 95F M6 powers the movement of these particles. These particles appear to be


Fig. 3 Models for melanosome transport by M5a. (a) Schematic of melanosome transport in a Xenopus melanophore, highlighting the involvement of both microtubule-based motors (cytoplasmic dynein and kinesin II), and the actin-based motor M5a. In the dual filament model of transport proposed by Langford (discussed in [151]), melanosomes are transported over a long distance from the center of the melanosome to the cel periphery on microtubules by dynein and kinesis. Once they reach the periphery,

they are transferred to the actin-filament cytoskeleton where local transport occurs via M5a. (b) A more detailed, molecular view of melanosome transport by M5a is diagrammed. A protein complex composed of Rab27a and melanophilin was shown to act as a receptor for M5a, targeting the motor to the melanosome [55, 93, 170, 172]. Rab27a first binds the melanosome then recruits melanophilin, which subsequently recruits M5a into the complex.


associated with the invaginating plasma membrane and may be required for supplying membrane or other material to the growing invagination during the rapid cycles of cell division that occur in the early fly embryo [178]. Fluorescence time-lapse microscopy experiments with anti-95F antibodies revealed a role for fly M6 in the transport of cytoplasmic particles from the nurse cell to the oocyte [179]. Microinjection of mitochondria-specific dyes into the developing oocyte suggested that some of these particles were mitochondria. Partial loss of function mutations in the 95F gene revealed a role for M6 in sperm development [180]. The jaguar mutation disrupts the first exon of the 95F myosin, causing male sterility due to a defect in sperm individualization (the process of enclosing each spermatid with its own membrane from what was once a syncytium). Consistent with this phenotype, 95F is localized to the leading edge of the individualization complex (IC), an actin-rich structure that surrounds the nuclei as the process of individualization proceeds [180]. This region is enriched in vesicles and it is possible that 95F could participate directly in the transport of membranes to the IC. Alternatively, it may contribute to the organization or stabilization of a membrane-rich leading edge of the IC that is required for efficient delivery of plasma membrane to the sperm undergoing individualization. Interestingly, M6 is also critical for sperm development in C. elegans [181]. Animals with a null mutation in the spe-15 gene are sterile and exhibit defects in the segregation of various cellular components during spermatogenesis. M6 is thus essential for transport and/or membrane reorganization during spermatogenesis in both flies and worms, and might play a conserved role despite the significant differences in sperm development between these two organisms. Myosins have also been implicated in nuclear movements. The unique myosin from the ciliated protozoan Tetrahymena thermophila, MYO1, has been found to be required for macronucleus elongation, a process essential for correct segregation of genetic material to daughter cells [17]. This result raises the intriguing possibility that myosins play a role in the transport or motility of nuclei in some contexts. 3.4 Maintenance of Actin-rich Extensions

M1, M5, M6, M7, M10 and M15 are found in actin-rich cell extensions such as microvilli, filopodia and stereocilia. Their exact roles in the generation or maintenance of these structures remain unclear at this time. Several myosins have been proposed to provide materials, force and tension required to maintain these structures. One of the best characterized M1s is the brush border subtype MYO1A (better known as BBM1) that forms links between actin bundles in the microvillar core and overlying plasma membrane in brush-border intestinal microvilli [182, 183]. It has been hypothesized to provide structural support to the microvilli, but recent analysis of GFP-tagged MYO1A in an epithelial cell line suggests that it is rapidly exchanged with a cytosolic pool [184], raising questions about how this myosin might play a structural role. It has also been localized to Golgi-derived vesicles, suggesting that its role is largely to deliver membrane to the growing microvillus


[185] and that it might also contribute to the dynamic growth or changes in flexibility of this structure [184]. The Drosophila M3, ninaC (neither inactivation nor after potential C), was identified by the characterization of a vision mutant defective in photoreception. NINAC is expressed specifically in fly photoreceptor cells as two alternatively spliced proteins [186]. One form localizes to the microvillar extensions of photoreceptors (rhabdomeres), while the other form is found in cell bodies [187, 188]. Cells null for NINAC undergo retinal degeneration [188]. M3 was also recently identified in humans [189], but whether or not it functions in human vision is not yet apparent. M5 is found at the tips of microvilli and is also present in filopodia [190, 191]. CALI of M5 in growth cones of dorsal root ganglion (DRG) neurons results in rapid filopod retraction [108], suggesting a role in delivery of materials required for filopod growth or for growth cone function. Fractionation studies and in vitro motility assays are consistent with this model of M5a action [148, 192]. However, analysis of cultured dilute neurons did not reveal any defects in growth cone dynamics [190]. The discrepancy between the CALI study and the analysis of cultured dilute neurons remains to be resolved, but it is possible that another M5 can compensate partially for the loss of M5a in the mutant mouse neurons. Dictyostelium M7 and mammalian M10 are both localized to the tips of filopodia [37, 110]. Deletion of the Dictyostelium M7 results in a loss of filopodia extension [110]. It is not yet known if this is due to the loss of delivery of materials necessary for making filopodia (as appears to be the case for M5), or if this motor is required for stabilizing this type of actin-rich extension. Overexpression of M10 in COS-7 cells, in contrast, results in the production of longer filopodia in increased numbers [193]. The association of M10 with particles that move up and down along the filopodial shaft suggests that this motor might be required for the targeted delivery of materials to the growing filopod. The inner ear sensory hair cells in mice have actin-rich extensions (modified microvilli) called stereocilia at their apical surface (Fig. 4). These structures are organized into units or bundles that move together when stimulated. The bundles are deflected in response to sound vibrations and gravity, resulting in the opening of ion channels and eventually transduction of electrical signals to the brain [194]. The Snell’s waltzer (sv) mouse mutant carries an intragenic deletion in the gene encoding M6 (Myo6) [195]. These animals are deaf, and also display hyperactivity and circling behavior [196]. M6 is localized to the base of the stereocilia and is enriched in the cuticular plate (a region of densely-packed actin filaments that the stereocilia are embedded into), as well as being diffusely distributed throughout the cytosol [195, 197]. Additionally, it is also found in the pericuticular necklace (a region at the periphery of the apical surface of the cell that is rich in vesicles) and is associated with punctae throughout the cytoplasm of the hair cells [197]. Molecular analysis of the inner ear hair cells in sv mice supports a role for M6 in maintaining stereocilia structure. Stereocilia bundles appear normal at birth, but by 3 days after birth the bundles become fused together and disorganized [198]. One model for this fusion is that M6 provides the force required to anchor the


Fig. 4 Schematic of myosins in the hair cell of the ear. Several myosins play critical roles in the mammalian ear–mutations in M2, M3, M6, M7a and M15 have all been linked to human deafness. The apical tip of the hair cel features actin-rich extensions called stereocilia. M1 localizes near the tip of the stereocilia in close proximity to tip links, which are likely attached to gated ion channels that open upon deflection of the hair cell bundle following stimulation [223]. M6 localizes to the base of the stereocilia, and is also enriched in the cuticular plate and perinuclear necklace [195, 197]. It is

believed to provide the force necessary to anchor the stereociliar membrane at the base of each extension. M7a localizes along the length of each stereocilium and is concentrated at site of lateral links, which are thought to be important for maintaining bundle integrity [197]. M7a may anchor cadherins at lateral link sites via an association with vezatin [204]. Finally, M15 also localizes to stereocilia and is concentrated in the cuticular plate [36]. Its role in the hair cell has not yet been elucidated. Note that the localization of neither M2 nor M3 in the sensory hair cells of the ear is as yet known.

membrane at the base of each individual stereocilium. The organization of actin filaments in the stereocilia and the minus-end directionality of M6 are such that this motor may help pull down the membrane around each stereocilium. In the absence of M6, discrete stereocilia form, but these structures are not preserved as the mice age resulting in stereocilia fusion.


It should be noted that although M6 is expressed ubiquitously in mammals [195, 199], the sv mice have no gross phenotypes other than deafness [195, 196]. However, their intestinal microvilli are shorter than those found in matched controls [198], consistent with M6 playing an important role in the maintenance of these specialized actin-filled structures. Despite the aberrant microvillar length, no gross defects in digestion have been reported. It may be necessary to subject the mice to some type of stress in order to uncover the consequences of shorter microvilli. The hypothesized role for M6 in membrane reorganization during stereocilia bundle formation/maintenance, along with membrane reorganization during fly embryogenesis and sperm development, follows a recurring theme for M6 involvement in membrane remodeling and maintenance. As observed in stereocilia, M6 localizes at the base of microvilli (another type of actin-rich structure) in brush border epithelial cells [191, 199, 200] where it may provide force for anchoring the membrane around each microvillus. A role for M6 in endocytosis could also relate to various biological processes involving membrane restructuring. As a high-duty ratio motor [29], M6 is a motor ideally adapted for maintaining membrane tension and providing force. M7a may also be important for aspects of stereocilia integrity. As with the Snell’s waltzer mouse, mice mutant for M7a (shaker-1) are deaf and display balance dysfunction [201]. Their hair cells extend stereocilia, but these are splayed apart [202]. Immunofluoresence studies have shown that M7a protein localizes along the length of each stereocilium and is concentrated at sites of lateral links that are believed to be important for maintaining the integrity of the stereocilia bundles [203]. Perhaps M7a is involved in organizing the stereocilia into bundles to provide rigidity during bundle deflections. M7a may be required to anchor cadherins at the lateral link sites via an association with vezatin, a ubiquitous, novel transmembrane protein [204]. 3.5 Membrane Trafficking

The M1s have been suggested to play a role in vesicle transport and endocytosis based on localization studies. As mentioned above, MYO1A in intestinal epithelial cells is associated with Golgi-derived vesicles that potentially provide membrane to the base of microvilli [185]. MYO1A is localized to both endosomes and lysosomes and expression of a truncated myosin dominantly inhibits the delivery of endosomal contents to lysosomes [205, 206]. It has also been implicated in the trafficking of basolateral vesicles to the apical plasma membrane in polarized epithelial cells [207]. The structurally similar MYO1D has been shown to play a role in trafficking of recycling endosomes [208]. Interestingly, an amoeboid M1 from Dictyostelium (myoB) has also been implicated in recycling from early endosomes, suggesting that there is some basic conservation of M1 function [209]. Kinetic analysis of MYO1A indicates that this mammalian M1 is a short-duty motor [210]; therefore, if it acts as an organelle motor, it should be present in a complex of motors. Such


a complex has yet to be identified, and indeed a bona fide MYO1A binding protein remains to be found. The different trafficking roles that various M1 isoforms play suggest that there are specific targeting molecules for each one, but these have not yet been identified. Characterization of such proteins should provide interesting insights into how the targeting of individual myosins may contribute to exquisite specification of their function. Two M5 isoforms, M5b and M5c, have also been implicated in intracellular trafficking. M5b co-localizes with transferrin and Rab11a to intracellular perinuclear vesicles [211]. Expression of the M5b tail alone disturbs the recycling of transferrin to the plasma membrane in both polarized and non-polarized cells. Two-hybrid analysis revealed that M5b interacts with Rab11a, but attempts to verify this interaction in vivo have not yet been successful [211]. Similarly, M5c co-localizes with Rab8 on intracellular vesicles and expression of the tail domain alone perturbs transferrin trafficking [212]. The available data suggest that M5c is associated with a compartment distinct from that of M5b, another indication that even within a class of myosins there can be precise functional specificity of individual isoforms. M6 has been suggested to participate in clathrin-mediated endocytosis in cultured polarized epithelial cells [63]. Endocytosis is important for nutrient uptake in the cell, along with immune system defense against microorganisms and cell-surface receptor recycling. The actin cytoskeleton plays a role in endocytosis, perhaps by providing a structural framework for trafficking and/or supplying the track for myosin motors transporting vesicles. M6 tagged with GFP was shown to colocalize with clathrin-coated vesicles via its C-terminal tail in polarized Caco-2 cells [63]. It was found in a protein complex with adaptor protein-2 (AP-2) by pull-down assays and co-immunoprecipitation. The AP-2 protein is a component of clathrin-coated vesicles associated with the plasma membrane. Overexpression of the M6 tail domain reduced the uptake of transferrin in transfected cells by over 50 % [63] In contrast, M6 is found in association with the Golgi complex in non-polarized cells as well as in the actin-rich ruffles of growth factor-stimulated cells [77]. The difference in observed localization when compared to polarized cells has been attributed to the lack of an epithelial cell-specific insert in the globular tail region. These findings reiterate how specific residues in the myosin tail play a significant role in specifying subcellular localization and function. M6 has also been shown by two-hybrid screens and co-immunoprecipitation to associate with Disabled-2 (Dab2), a protein involved in endocytosis and signal transduction [213, 214]. The central region of Dab2 is important for AP-2 binding, while the C-terminal tail mediates binding to M6 [214]. Dab2 transiently co-localizes with members of the low-density lipoprotein receptor (LDLR) family in clathrin-coated pits and is thought to regulate receptor protein trafficking [215]. It has been proposed that the M6–Dab2 interaction provides a link between the actin cytoskeleton and receptors undergoing endocytosis [214]. It remains to be determined if M6 acts as a motor for endocytic vesicles or if it participates in the organization of an actin-based sub-membranous domain where initial sorting of endosomal contents occurs.


3.6 Signal Transduction

The role of myosins in signaling was first eludidated in the fly eye. Illumination of the fly eye with pulses of light results in a distinctive negative potential followed by a rapid return to baseline each time the eye is stimulated. The ninaC (M3) mutant has a larger negative potential upon stimulation followed by a slower return to baseline after cessation of the stimulus [186, 216]. The N-terminal kinase activity, as well as calmodulin binding to sites in the C-terminal region, are required for normal phototransduction [217, 218]. The role of NINAC in normal phototransduction is separate from its role in maintaining the overall integrity of the rhabdomere microvilli [172]. M3 is also found in Limulus polyphemus (horseshoe crab) eyes where it was identified as a phosphoprotein that undergoes clock-regulated phosphorylation (i. e. horseshoe crabs have circadian neural input that changes phototransduction in the eye, among other things) possibly by a cAMP-dependent protein kinase [219]. Interestingly, Limulus NINAC has also been shown to be a phosphoprotein that can be phosphorylated in vitro by protein kinase C (PKC) [219]. Mutation of PKC sites in the tail region results in defects in deactivation after the light stimulus has been shut off. Recent evidence suggests that NINAC is a component of the ‘signalplex’, a cluster of molecules required for phototransduction and that it binds to INAD (a PDZ protein) through its C-terminus [220]. NINAC does not require INAD binding for localization to the rhabdomeres, but their interaction is essential for termination of the electrophysiological response to light [220]. MYO1C (also known as myr 2, myosin 1β) plays an unusual role in signal transduction. This M1 localizes near the tips of the sensory hair cell stereocilia where a fine fiber known as the tip link extends from the side of one stereocilium to the top of the adjacent, shorter stereocilium [197, 221, 222]; Fig. 4. Elegant biophysical studies have shown that the tip link is likely attached to a calcium channel that opens upon deflection of the hair bundle following an auditory stimulation [223]. The cell becomes depolarized, but if the stimulus persists adaptation occurs. Adaptation has been proposed to occur when a myosin molecule climbs up the actin filaments in the core of the stereocilium, acting to reset the tension on the tip link and close the channel. An exciting new chemical genetics approach has been used to test this model of MYO1C function [224]. Introduction of a single mutation in the ATP binding pocket of MYO1C (Y61G) allows the mutant molecule to bind a modified ADP (NMB-ADP) that locks it into the rigor state. Wild-type MYO1C does not bind NMBADP and is not inhibited by it, whereas the Y61G-MYO1C functions normally in the presence of ATP but not with NMB-ADP [225]. Application of NMB-ADP to mouse utricular hair cells expressing small amounts of Y61G-MYO1C has a dramatic effect – it abolishes adaptation [224]. These data provide strong evidence in support of MYO1C being the adaptation motor and suggest a novel role for myosins in the ion channel gating.

4 Myosins in Disease

Human and rat M9s are predicted to contain a GAP domain in their tail regions with approximately 30% sequence identity to GAP proteins of the Rho small Gprotein subfamily [226, 227]. Rho belongs to a family of signaling proteins in the Ras superfamily of monomeric GTPases that act as molecular ‘switches’ whose activity depends on their nucleotide conformation state (GTP-bound versus GDPbound). Activation of Rho leads to the formation of actin filament bundles and focal adhesions, among other processes [51]. GAP proteins negatively regulate these processes by stimulating GTP hydrolysis of Rho, converting it from an active state (GTP-bound) to an inactive (GDP-bound) state. M9s have been shown to stimulate the GTPase activity of Rho A, B and C [50, 57, 226], and as predicted the overexpression of M9 in cultured mammalian cells leads to a loss of stress fibers and focal adhesion contacts [49]. The exact role for M9 in signal transduction is not known, but it is interesting to note that this motor has been shown to be processive and to move toward the minus-end of actin filaments [27]. An unusual myosin from Dictyostelium may also play a role in intracellular signaling. MyoM (to date an undesignated myosin) has a functional GEF domain at its extreme C-terminus [38, 39]. While deletion of this myosin does not result in any overt phenotype, expression of the tail region alone resulted in a decrease in growth rates and a hypersensitivity to osmotic stress [38, 39]. Cells exposed to water extended broad, actin-filled protrusions with the myoM tail at their tip, suggesting that the GEF domain is capable of signaling to the actin cytoskeleton under conditions of stress [38]. It remains to be determined whether this phenotype reflects the true function of myoM, but the fact that the behavior of the cells is only affected under specific conditions suggests that the observed phenotype is not due to non-specific misregulation of GEF activity.


The critical roles for unconventional myosins are highlighted by their connection to a variety of human disorders, primarily those related to sensory dysfunctions. A common theme of structural roles in the inner ear and retina emerge for three classes of myosins. Mutations in genes encoding M3, M6, M7 and M15 have been implicated in human hearing disorders and deafness in mice. Myosin functions in hearing and other cellular processes related to disease phenotypes are just beginning to be uncovered. Perhaps the best-known relationship between myosin mutation and disease is in familial hypertropic cardiomyopathy (FHC; see the Chapter 19 by Konhilas and Leinwand), an autosomal dominant condition characterized by shortness of breath, angina, and heart palpitations that can lead to heart failure, stroke and sudden death [228]. FHC can be caused by mutations in β cardiac myosin HC. Myosin LC mutations have also been associated with human heart myopathies similar to FHC [229].

23


4.1 Griscelli Syndrome

Griscelli syndrome is characterized by severe primary immunodeficiency along with pigment dilution of hair, eyebrows and eyelashes [230, 231]. A role for M5 in this disorder was hypothesized to be due to the phenotypic similarities between Griscelli syndrome patients and the dilute mouse mutant (described above; [232]). For example, pigment accumulation has been observed in the center of Griscelli melanocytes (similar to the accumulation seen in dilute melanocytes; [157, 158, 230]. M5a was subsequently shown to co-localize with melanosomes in cultured human melanocytes [160]. Genetic evidence supports a role for M5 in this disorder. The MYO5A gene maps to the disease locus 15q21 and at least two M5 mutations have been identified in Griscelli patients (type 1 Griscelli syndrome; Pastural et al., [231, 233]). Some Griscelli patients were reported to have immunodeficiency disorders [230, 234], but this phenotype was inconsistent with the mouse dilute phenotype. However, the RAB27A gene is quite close to MYO5A – they are 1.6 cM apart- and RAB27A has also been linked with this syndrome (type 2 Griscelli syndrome; [235]. Rab27a is localized with M5s on melanosomes [168, 172] and these two proteins co-immunoprecipitate from melanocyte extracts [168]. Rab27a plays a role in granule exocytosis while M5a does not [236]. Thus, MYO5A mutations are associated with the pigmentation and neurological aspects of Griscelli disease, while RAB27A mutations are associated with the immunological disease form of Griscelli [235]. Mutations in the non-muscle M2 encoded by MYH9 have recently been implicated in three giant-platelet disorders – May–Hegglin anomaly (MHA), Sebastian syndrome (SBS) and Fechter syndrome [237, 238]. All three disorders are characterized by thrombocytopenia, large platelets and leukocyte inclusions [237, 238]. Based on similarities between the three disorders and mapping experiments, it was proposed that MHA, SBS and FTNS are allelic. One unique feature of FTNS is sensorineural deafness, consistent with a connection between Fechter syndrome and DFNA17 deafness disorder. 4.2 Roles for Myosins in Hearing

Approximately one in 1000 people experience severe or profound deafness at birth or during early childhood, and another one in 1000 children become deaf before they reach adulthood [239]. Non-syndromic deafness patients are afflicted only with hearing loss, while syndromic cases exhibit hearing loss in combination with other defects. There are many genes involved in deafness disorders, including several encoding unconventional myosins (see Fig. 4). M3, M6, M7 and M15 have fundamentally important functions in the auditory system as mutant alleles at these loci have been shown to co-segregate with human deafness. M7 plays a role in hearing and balance in humans, mice and zebrafish [201, 240, 241]. M7a (MYO7A) has a highly restricted tissue distribution in


mammals – it is found almost exclusively in cells that have specialized actin-based structures such as the hair cells of the ear, retina, testis, lung and kidney [203]. Mutations in MYO7A result in two forms of human non-syndromic deafness, mutant loci DFNB2 (autosomal recessive; [242]) and DFNA11 (autosomal dominant; [243]) and a syndromic form of deafness, Usher’s syndrome type IB [241]. Usher’s syndrome type IB is characterized by a gradual degeneration of the rods and cones in the retina, causing loss of vision in addition to hearing loss [244] and [245] for reviews). As mentioned above, the shaker-1 M7a mutant mice are deaf and display balance dysfunction [201]. Mutations in the Danio rerio (zebrafish) M7a gene, mariner, were found to exhibit circling behavior along with hair cell defects similar to those observed in shaker-1 mice [240]. Hair cells did extend stereocilia but these are splayed apart, suggesting that the links between adjacent stereocilia have been lost [240]. These data demonstrate the remarkable functional conservation of M7a through evolution. A recent electrophysiological study has uncovered a role for M7a in gating of transducer channels that cannot be attributed to the disorganization of the hair bundles [246], suggesting that this motor may play more than one role in stereocilia function. While analysis of the shaker-1 mutant mouse has provided insight into the role of M7a in hearing [201, 202, 246], it has not provided a clear explanation for the late onset retinitis pigmentosa exhibited by Usher’s IB patients. A careful analysis of shaker-1 photoreceptors has revealed that opsin transport is decreased [247] and melanosome distribution is aberrant [175]. It has been suggested that the relatively short life-span of the mouse is not long enough for expression of the vision defect to become apparent [247]. Mutations have been documented all along the length of the M7a HC in humans, as well as mice and zebrafish. In addition to confirming the importance of the motor domain, the identification of mutations in both the C-terminal MyTH4 and FERM domains indicate these domains are essential for function [62, 240]. Mutations in M15 also result in deafness both in humans and mice. MYO15 is mutant in human DFNB3, a non-syndromic, congenital deafness disorder [248]. As is the case for M7a, mouse M15 has a restricted tissue distribution. In addition to the cochlea, it is expressed in the pituitary and other neuroendocrine tissue [36, 249]. The M15 homolog in mouse is also associated with a hearing loss phenotype; the shaker-2 mouse was found to have stereocilia that are approximately one-tenth the normal length [250]. M15 is localized to the stereocilia of cochlear hair cells and is concentrated in the cuticular plate [36]. Consistent with its localization, an abnormal organization of actin is also observed in the cochlear hair cells of shaker-2 mice, suggesting that this myosin plays a role in actin organization. M6 was first associated with hearing loss when a null mutation in murine Myo6 was discovered to cause the Snell’s waltzer phenotype [195], as discussed above (Section 3.4). A missense mutation in the motor domain of M6a results in an autosomal dominant non-syndromic form of deafness, DFNA22, indicating that this myosin has a conserved role in hearing [251]. The effect of this single mutation on the motor properties of M6a is not yet known.

25


Less is known regarding the role of other myosins involved in hearing, as mouse models for several human deafness mutations are not yet available for study. The role of M3 in fly vision has already been discussed, but M3 also plays an important sensory role in humans. Mutation of M3a causes progressive hearing loss DFNB30 [252]. MYO3A is expressed in the sensory cells of the mouse ear, but more work remains to be done to determine how this motor protein contributes to the maintenance of hearing. Given the conservation of M3 from flies to Limulus to humans, it is tempting to speculate that it plays a role in regulating some aspect of signaling in hair cells. A mutation in MYH9, a non-muscle M2, has been reported to be responsible for an autosomal dominant form of deafness, DFNA17 [253]. This missense mutation results in a single amino acid change (R705H) in a residue located in the critical SH1 helix, which is known to influence the ATPase activity of myosin. The underlying cause of deafness and the localization of this myosin to sensory cells are not currently known, but it will undoubtedly be of interest to determine how a myosin that plays a role in generating contractile forces contributes to hair cell function.

5 New Myosins and Myosin Functions on the Horizon

A considerable number of myosins have been identified but their functions have not yet been elucidated. The presence of interesting domains in their N- or C-termini and divergence in their motor domains suggests that they have the potential to play unique and interesting roles. As work in the field progresses, these roles will undoubtedly be uncovered. A comprehensive analysis of genes predicted by the human and fly genome projects [9] has revealed the existence of a novel human myosin predicted from contig AC023133. This myosin appears to have a unique N-terminal extension and a short tail region. Two novel myosins were identified in the fly genome, one designated 29CD that has an N-terminal extension and a short, proline-rich tail and another designated 95E that has a large insert in the motor domain loop 1 and a tail region comprised largely of basic residues. The diverse roles played by the cytoskeleton in fly development is likely to catalyze interest in studying the in vivo function of these novel myosins through a search for null mutants or use of double-stranded RNA interference (dsRNAi) to knock out their mRNAs. A novel human myosin, M18 or PDZ-myosin, was identified in a differential screen for genes expressed highly in bone marrow stromal cells that support hematopoietic cells [34]. The tail region of this myosin is comprised largely of predicted coiled-coil sequences (suggesting that it is a dimer) and there is a PDZ domain at the N-terminus. The tissue distribution of this myosin is not yet known. It is localized to cytoskeletal elements in stromal cell lines as well as NIH3T3 cells, where it appears to be associated with both actin and microtubules as well as being distributed throughout the cell.

6 Conclusions 27

The novel M16, also referred to as myr8, was identified in a study of neuronal myosins in the rat [33]. Two forms of this myosin were identified. M16a has a shorter tail and is expressed predominantly in the nervous system. The longer M16b appears to be broadly expressed. Analysis of the tail sequence reveals that it is unique and does not contain any regions of predicted coiled-coil, suggesting M16b is a monomer. The N-terminus contains eight ankryin repeats that bind to protein phosphatases. Immunoprecipitation experiments revealed that M16b indeed associates with protein phosphatase 1 catalytic subunits 1α and 1γ . This myosin has a punctate distribution in both neurons and astrocytes. M4, M12 and a novel unconventional myosin from scallop mantle (tissue comprised of both muscle and non-muscle cells) named ScunM have been identified either at the protein or gene level but no information regarding their tissue distribution (in the case of M12 and ScunM) or subcellular localization is available [1, 254, 255]. These myosins appear to be unique to the organisms in which they were originally identified. The M4 gene was identified in Acanthamoeba and sequence analysis revealed the presence of a C-terminal SH3 domain as well as a single MyTH4 domain [255]. It was shown to co-precipitate with actin under rigor conditions and to be released from actin by ATP, indicating that it has the biochemical properties expected of a myosin. M12 was identified at the gene level through a comprehensive search of the C. elegans genome for myosin genes and is named HUM-4 (heavy chain of unconventional myosin) [256]. HUM-4 has a 150-amino acid extension on the N-terminus. In addition, the P-loop and actin-binding sequences in the motor domain are more divergent than those found in other myosins. There are two MyTH4 domains in the tail but no clear FERM domains have been identified. No known mutations map near the hum-4 gene and efforts to inactivate gene function by dsRNAi have not resulted in any phenotype (J. P. Baker and M. A. Titus, unpublished data; WormBase – www.wormbase.org). ScunM was identified in a search for unconventional myosin genes in scallop mantle tissue [254]. It is one of the smallest myosins, with the HC having a predicted molecular weight of 90 kD. Similar to the M14s, ScunM has a short tail consisting of only 46 amino acids and it lacks any LC binding domains. Phylogenetic analysis, however, reveals that this myosin is the founding member of a novel class and not an M14.

6 Conclusions

It is likely that efforts to sequence the genomes of various organisms will result in the discovery of additional novel myosins. The ability to express myosins in baculovirus and inactivate gene function by dsRNAi in organisms and cultured cells should permit rapid characterization of their biochemical properties and in vivo functions. In addition to identifying myosins based on conserved motor sequences,


it is possible that large-scale efforts directed at crystallizing proteins will identify proteins that do not share the conserved myosin primary structure, but whose structure is highly similar to that of the myosin motor domain. Thus, a convergence of proteomic and genomic approaches is likely to expand our appreciation of the diversity of the myosin superfamily A number of lessons can be learned from examining the myosin superfamily. The most striking one is the diversity of the family – both in terms of the variety of different classes and the number of different isoforms within a class. This diversity is reflected in the wide range of functions observed for myosins of even a single class. There are classes of myosins that appear to be specific for plants and parasites, while only three myosin classes are conserved from fungi to humans. As evolution proceeded, it appears that higher eukaryotes acquired a more sophisticated complement of myosins. Some of which, such as M7, were preserved and others, such as M4 and M12, appear to have been lost. The importance of myosins is highlighted by their essential roles in fungi and their association with a number of human diseases, ranging from cardiomyopathies to deafness. An unexpected surprise that emerged from studies of myosins in mammals is the key role that they play in hearing, perhaps reflecting their ability to participate in an array of cellular functions. Future studies of this large motor family are likely to reveal additional roles and operating principles as well as, undoubtedly, exotic new family members.

Acknowledgements

The authors would like to thank Shawn Galdeen for helpful comments on the manuscript and Dr. Richard Cheney for providing Figure 2. M.C.K. is supported by an NIH postdoctoral fellowship (NRSA) and work in the Titus laboratory is supported by the National Institutes of Health. M.A.T. is an Established Investigator of the American Heart Association.

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257 MOTEGI, F., R. ARAI, and I. MABUCHI. 2001. Identification of two type V myosins in fission yeast, one of which functions in polarized cell growth and moves rapidly in the cell. Mol. Biol. Cell 12: 1367–1380.

258 CHEN, Z. Y., T. HASSON, D. S. ZHANG, B. J. SCHWENDER, B. H. DERFLER, M. S. MOOSEKER, and D. P. COREY. 2001. Myosin VIIb, a novel unconventional myosin, is a constituent of microvilli in transporting epithelia. Genomics 72: 285–296.

1

Plasma-Membrane H+ -ATPase From Yeast Silvia Lecchi and Carolyn W. Slayman Yale University School of Medicine, New Haven, USA

Originally published in: Handbook of ATPase. Edited by Masamitsu Futai, Yoh Wada and Jack H. Kaplan. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30689-3

1 Introduction

The single most abundant protein in the fungal plasma membrane is a protontransporting ATPase, encoded by the PMA1 gene [1] and hydrolyzing as much as one-quarter of cellular ATP [2]. By forming a large electrochemical gradient across the surface membrane, the H+ -ATPase provides energy to an array of protoncoupled cotransporters for sugars, amino acids, and other nutrients; it also contributes to the regulation of intracellular pH (Figure 1). Direct evidence for the ATPase first came several decades ago from microelectrode studies on the filamentous fungus Neurospora crassa, where a large (> 200 mV), metabolically sensitive membrane potential was shown to be generated by ATP-dependent proton pumping [3, 4]. By the early 1980s, the H+ -ATPase had been purified from Schizosaccharomyces pombe [5], Saccharomyces cerevisiae [6], and Neurospora crassa [7, 8] and found to consist of a 100 kDa polypeptide, firmly embedded in the membrane bilayer and requiring detergents for solubilization. Reconstitution into proteoliposomes confirmed that the ATPase did indeed mediate electrogenic proton transport [9–11], with a stoichiometry of 1 H+ /ATP, as predicted from electrophysiological measurements [12]. In parallel, biochemical studies demonstrated that it split ATP by way of a covalent β-aspartyl intermediate [13–16], the hallmark of a widespread group later named the P-type ATPases. Like other members of that group, the H+ -ATPase alternated during its reaction cycle between two major conformational states (E1 and E2 ), which could be distinguished by their different patterns of proteolytic fragments following digestion with low concentrations of trypsin [17, 18]. By 1986, cloning of PMA1 genes from Saccharomyces cerevisiae [1] and Neurospora crassa [19, 20] had revealed clearcut sequence homology between the fungal Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Plasma-Membrane H+ -ATPase From Yeast

Fig. 1 Pma1 H+ -ATPase plays an essential role in yeast cell physiology. Pumping protons (H+ ) across the membrane at the expense of ATP hydrolysis, Pma1 H+ -ATPase establishes the membrane potential (R ) and the pH gradient required for nutrient import and contributes to the regulation of intracellular pH. PM: plasma membrane.

H+ -ATPase, the Kdp K+ -ATPase from Escherichia coli [21], and the plasmamembrane Na+ ,K+ - and sarcoplasmic reticulum Ca2+ -ATPases of animal cells [22–24]. More than 200 P-ATPase genes have since been cloned from archaebacteria, eubacteria, fungi, algae, plants, and animals and analyzed to establish their relationship with one another. In 1995, Lutsenko and Kaplan [25] put forward a useful classification scheme that divides P-type ATPases into three groups based on the cation transported, the number and location of hydrophobic transmembrane segments, and the position of three conserved sequences: DKTGT (where the β-aspartyl phosphointermediate forms), TGES (located upstream of the phosphorylation site in a smaller cytoplasmic loop), and GDGXNDXP (close to the ATP-binding site). According to this scheme, heavy metal pumps are classified as P1 -ATPases; H+ , Na+ /K+ , Mg2+ , and Ca2+ pumps, as P2 -ATPases; and the K+ -ATPase of E. coli, as a lone P3 -ATPase (Table 1). Several years later, Axelsen and Palmgren [26] developed a more complex classification based on the analysis of eight stretches of amino acid sequence shared by all P-type ATPases (Table 1). In this scheme, the heavy metal pumps remain as Type I ATPases, with the K+ -ATPase of E. coli distantly related to them. The Na+ ,K+ -, H+ ,K+ -, and Ca2+ -ATPases and H+ -ATPases still fall in the same general sector of the phylogenetic tree, but are separated into Type II and Type III, respectively. Finally, a place is found for two additional groups of P-ATPases that had become known from genome-sequencing projects: Type IV, involved in the transport of aminophospholipids and other hydrophobic compounds, and Type V, whose function is unknown.

1 Introduction Table 1 Classification of P-Type ATPases

Lutsenko and Kaplan [25]

Cation specificity

Axelsen and Palmgren [26]

Cation specificity

P1

Heavy metal

Type I

P2

Ca2+ Na+ /K+ H+ /K+ H+ Mg2+ K+

Type II

Heavy metal K+ Ca2+ Na+ /K+ H+ /K+ H+ Mg2+

P3

Type III

Type IV Type V

Aminophospholipid Unknown

Within the P-type group, the P2 - or Type II/III ATPases have been most intensively studied. These ATPases have a characteristic topology, with four membranespanning segments (M1-4) towards the N-terminal end of the polypeptide and six such segments (M5-10) towards the C-terminal end (Figure 2). The large cytoplasmic loop between M4 and M5 and the smaller cytoplasmic loop between M2 and M3 assemble to form the catalytically active part of the molecule, as discussed below. The N- and C-termini are also exposed at the cytoplasmic surface of the membrane. This chapter will review how the yeast and Neurospora Pma1 H+ -ATPases have come to serve as simple eukaryotic prototypes of the P2 -ATPases, based on the powerful genetic and cell biological tools that are available to study their reaction mechanism, regulation, and biogenesis.

Fig. 2 Transmembrane topology of Pma1 H+ -ATPase. The ATPase has 10 transmembrane "-helices with both N- and C-termini located in the cytoplasm. Highly conserved amino acid sequences characteristic of Type III ATPases are shown. PM: plasma membrane.

3


2 Structure 2.1 Ca2+ -ATPase as a Model

A landmark event came in 2000 with the publication by Toyoshima and his colleagues of the first high-resolution structure of a P-type ATPase. They successfully crystallized sarcoplasmic reticulum Ca2+ -ATPase in the E1 conformation (with two Ca2+ ions bound) and solved the structure at 2.6 Å [27]. As expected from an earlier cryoelectron microscopic map at 8 Å resolution [28], they found the ATPase to be organized into a cytoplasmic headpiece and a transmembrane or M domain. Within the latter, four of the ten transmembrane α-helices (M4, M5, M6, and M8) assemble to create a pair of Ca2+ binding sites, consistent with the known stoichiometry of the ATPase. Site I is formed by side-chain oxygen atoms in M5 (N768 and E771), M6 (T799 and D800), and M8 (E908), while Site II is formed by main-chain carbonyl oxygens in M4 (V304, A305, I307) and side-chain oxygens in M4 (E309) and M6 (N796 and D800). Local unwinding of transmembrane helices, helped by two Pro residues in M4 and one in M6, plays an essential role in the architecture of both sites. The headpiece of Ca2+ -ATPase is organized into three domains: P (phosphorylation), N (nucleotide-binding), and A (actuator or anchor), quite distinct from one another in the E1 Ca conformation. The P domain is formed by the two ends of the large M4–M5 cytoplasmic loop and contains the aspartyl residue (D351) that undergoes alternate phosphorylation and dephosphorylation during the reaction cycle. It is a typical Rossman fold, with a seven-stranded parallel β-sheet and eight short α-helices. The large N domain consists of the middle portion of the M4–M5 loop, arranged in a seven-stranded antiparallel β-sheet lying between two helix bundles. In crystals soaked with a non-hydrolyzable ATP analog (TMP-AMP), this compound appears within the N domain, surrounded by residues previously implicated in nucleotide binding. Finally, the N-terminal part of the ATPase and the small M2–M3 cytoplasmic loop come together to form the A domain, which is connected to the transmembrane part of the ATPase by long loops. The function of the A domain was unclear at the time of the first Toyoshima paper, although there was evidence from proteolytic cleavage studies that its conformation changes substantially during the reaction cycle. In 2002 the same group published a second high-resolution (3.1 Å) structure of Ca2+ -ATPase, this time stabilized in the E2 conformation by the inhibitor thapsigargin [29]. It is strikingly different from the E1 Ca structure. On the one hand, six of the ten transmembrane helices (M1–M6) move significantly between E1 and E2 , “opening” the M domain, destroying the two Ca2+ binding sites, and, presumably, releasing Ca2+ into the lumen of the sarcoplasmic reticulum. Simultaneously, the A domain rotates ca. 100◦ horizontally and the N domain inclines ca. 90◦ relative to the membrane, shifting the cytoplasmic headpiece into a “closed” conformation. The authors suggest that the return from E2 to E1 requires Ca2+ to bind, fixing the

2 Structure

membrane helices in a way that prohibits the closed conformation of the cytoplasmic headpiece. The headpiece opens; the γ -phosphate of bound ATP can now reach and phosphorylate D351; and the cycle continues. Thus, although much remains to be done to understand the reaction mechanism fully, the two high-resolution structures have made it possible to develop a preliminary picture of how the Ca2+ pump may work. 2.2 Applicability of the Ca2+ -ATPase Structure to Other P2 -ATPase, Including the Pma1 H+ -ATPase

Not surprisingly, there has since been a flurry of activity to determine whether the first (and now the second) Toyoshima structure can serve as a legitimate template for other P2 -ATPases. So far, there is reason for optimism. Sweadner and Donnet [30] found that the sequence of Na+ ,K+ -ATPase could readily be projected onto the 2.6 Å Ca2+ -ATPase structure in a way that superimposes short stretches of homology throughout the cytoplasmic and membrane domains; when this was done, most insertions and deletions appeared (as they presumably should) on the protein surface. The authors concluded that the core regions of both ATPases are likely to fold in the same basic manner. Further support is that conformationally sensitive Fe2+ cleavage sites, identified by Karlish and co-workers for the Na+ ,K+ ATPase, can be mapped logically onto the E1 and E2 structures of Ca2+ -ATPase (reviewed in Ref. [31]). Kühlbrandt and co-workers [32] have recently provided strong evidence that the similarity also extends to the Neurospora and yeast plasma-membrane H+ ATPases. In this case, the authors made a point-by-point comparison between their earlier 8 Å cryoelectron microscopic map of the Neurospora enzyme [33] and Toyoshima’s 2.6 Å crystallographic structure of Ca2+ -ATPase [27]. There was an excellent fit between the two in the M and P domains; the A and N domains could also be accommodated by a rigid-body displacement of 10 Å in the first case and a more substantial rotation of 73◦ in the second case. Within the headpiece, key structural elements involved in ATP binding and formation of the phosphoenzyme reaction intermediate were readily recognizable, and within the membrane, two of the ten cation-ligating residues were conserved in the H+ -ATPase (D730 and E803, corresponding to D800 and E908 of Ca2+ -ATPase). Indeed, only the highly divergent N- and C-termini were impossible to overlay on the Ca2+ -ATPase template. Thus, it seems reasonable to use the Kühlbrandt H+ -ATPase model as a framework in developing experiments and interpreting results on the Pma1 ATPases of yeast, Neurospora, and other fungi. 2.3 H+ -ATPase Oligomers

By contrast with the steady progress made towards a detailed structure for the 100 kDa catalytic subunit, there has been considerable debate about the oligomeric state

5


of the Pma1 ATPase. For the Neurospora enzyme, Scarborough and co-workers have demonstrated that 100 kDa monomers are fully competent to carry out ATP hydrolysis and ATP-dependent proton transport after reconstitution into proteoliposomes [34]. Under other conditions, however, the same enzyme clearly forms hexamers [35], which allow the ready production of two-dimensional crystals for cryoelectron microscopy [33]. Even larger oligomers of yeast Pma1 H+ -ATPase (up to dodecamers) are observed when detergent-solubilized cell extracts are analyzed by gradient centrifugation [36], blue native polyacrylamide gel electrophoresis [37], or co-immunoprecipitation of differently tagged 100 kDa polypeptides [38]. The biological significance of the oligomers is not yet clear. Recent studies by the Chang, Schekman, and Simons laboratories [36–38] suggest that they may play a role in biogenesis, since they are formed soon after the 100 kDa ATPase polypeptide is synthesized and appear to be associated with packaging the ATPase into “lipid rafts” for transport to the plasma membrane (see below). Conversely, Kühlbrandt et al. [32] found crystalline patches of rosette-shaped particles in freeze–fracture replicas of starving yeast and Neurospora cells, and consider them to be a downregulated “storage form” of the enzyme. Further work is required to sort out these possible interpretations, which are not mutually exclusive. It will also be of interest to map the regions of the Pma1 molecule that take part in oligomer formation; based on their structural model of the Neurospora ATPase, Kühlbrandt et al. [32] propose that the C-terminal domain of one monomer (known to be exposed in the cytoplasm [39]) interacts with Q624 and R625 in the P domain of the neighboring monomer. 2.4 Associated Proteolipids

Studies of highly purified Pma1 H+ -ATPase preparations have given no evidence for a tightly bound β subunit of the kind seen in mammalian Na+ ,K+ - and H+ ,K+ ATPases. Conversely, two small proteolipids do associate closely with the Pma1 enzyme and appear to play a physiologically significant role. Information about them began to emerge in 1992, when Navarre and co-workers first observed diffuse 4 and 7.5 kDa bands in Coomassie-stained gels of highly purified yeast H+ -ATPase [40]. Chloroform–methanol extracts of both bands yielded the same 38-amino acid proteolipid, encoded by a gene that the authors named PMP1 (for plasma-membrane proteolipid). A closely related PMP2 gene was later cloned by hybridization with a PMP1 probe [41]. Although the two open reading frames predict slightly different N-termini, post-translational processing yields mature Pmp1 and Pmp2 proteolipids that share all but a single amino acid residue (A21 vs. S21). Their hydropathy profiles are virtually identical, with a hydrophobic stretch of 24 residues followed by a highly basic, 14-amino acid tail. In their overall topology, Pmp1 and Pmp2 call to mind the amphipathic FYXD proteins that co-purify with mammalian Na+ ,K+ -ATPases (reviewed in Ref. [42]) as well as two regulatory proteins, phospholamban and sarcolipin, that are associated with sarcoplasmic reticulum Ca2+ -ATPases [43, 44].

3 Reaction Mechanism

Single deletions of PMP1 or PMP2 have only minor effects on the activity of yeast H+ -ATPase, but deletion of both genes leads to a 50% reduction in activity, suggesting some kind of regulatory role for the two proteolipids [41]. Recent NMR studies by Neumann and co-workers have explored the in vitro interactions between synthetic fragments of Pmp1 and lipid micelles [45, 46]. Based on their results, the authors propose that Pmp1 extends across the plasma membrane bilayer, with its bulky, hydrophobic N-terminal end located alongside the membrane-embedded domain of the ATPase in the outer, sterol- and sphingolipid-rich leaflet, while its C-terminal part sequesters a subset of negatively charged phosphatidylserine lipids in the inner leaflet to create a proper environment for the ATPase at the cytoplasmic surface of the membrane. The relationship between these structural data and stepwise models for ATPase biogenesis is discussed below.

3 Reaction Mechanism 3.1 Overview of the Reaction Cycle

This section covers three interrelated aspects of the reaction mechanism of Pma1 H+ -ATPase: (i) ATP binding and formation of the β-aspartyl phosphoenzyme intermediate; (ii) the E1 –E2 conformational change; and (iii) H+ transport. The Post-Albers scheme, based on more than three decades of investigation into the partial reactions of P-ATPases, describes the generally accepted relationship among these steps. In the top limb of the cycle, as diagrammed for the H+ -ATPase (Figure 3), ATP is bound with high affinity to the E1 form of the enzyme. Intracellular H+ (or a hydronium ion; see below) then binds and stimulates the transfer of γ -phosphate from ATP to create the high-energy β-aspartyl phosphate intermediate; E1 P shifts to E2 P, accompanied by release of the transported H+ to the extracellular medium; and ATP binds with low affinity to the E2 form, accelerating the return to E1 .

Fig. 3 Reaction cycle of Pma1 H+ -ATPase. Based on the classical Post-Albers scheme for P-type ATPases.

7


3.2 ATP Binding and Phosphorylation

Although some of the amino acid residues that contribute to the ATP-binding site of P-type ATPases were first identified by affinity labeling and site-directed mutagenesis (reviewed in Ref. [47]), a much more complete picture has emerged with the crystal structure of Ca2+ -ATPase, where TNP-ATP was used to visualize the nucleotide site directly [27]. As expected, key residues have remained virtually unchanged during evolution. They include F487 in Ca2+ -ATPase (F451 in yeast H+ ATPase), K492 (K456), and K515 (K474), which lie deep in the pocket surrounding the adenosine moiety, and R560 (R509) near the mouth of the pocket, where it probably interacts with the β or γ phosphate of ATP. Consistent with this picture, mutational analysis has demonstrated the functional importance of K474 in the yeast Pma1 ATPase [48], and Pardo and co-workers have shown that K474 of the Neurospora enzyme is labeled by fluorescein 5-isothiocyanate in an ATP- and ADPprotectable way [49]. Overall, however, the N domain has been less well conserved than the rest of the cytoplasmic headpiece, with Ca2+ - and H+ -ATPases sharing only ca. 20 % identity in this region. More work is needed to correlate the structural differences with known differences in catalytic properties. In particular, the yeast H+ -ATPase has a K m for MgATP of ca.1.5 mM and K D s that are too high to measure accurately; its relatively weak affinity may relate to the smaller size of the N domain (ca. 150 residues) than in Ca2+ -ATPase (ca. 240 residues). By contrast, the P domain is similar in size (ca. 165 residues) and displays a very high degree of sequence identity (up to 40%) among P2 -type ATPases. This is not surprising in view of its central role in the reaction cycle. Functionally important P-domain motifs include 347-DKTGTLT in sarcoplasmic reticulum Ca2+ -ATPase (378-DKTGTLT in yeast H+ -ATPase) and 601-DPPR (534-DPPR), both located in or near the “hinge” region between the P and N domains; and 701-TGDGVNDAP (632TGDGVNDAP), located at the interface of the P and A domains. The first of these motifs surrounds the Asp residue that forms the characteristic high-energy E1 P intermediate, while the second and third contain residues involved in phosphoryl group transfer [50–52]. Based on site-directed mutagenesis studies of yeast Pma1 H+ -ATPase, amino acid substitutions within these highly conserved parts of the P domain often lead to a pronounced defect in protein folding, which can be recognized by an elevated sensitivity of the ATPase to trypsin [53]. In severe cases, newly synthesized ATPase fails to leave the endoplasmic reticulum and is eventually degraded by the proteasome (see below). In milder cases, the ATPase manages to reach the plasma membrane, but is then recognized by a yet-to-be-identified quality control mechanism and sent to the vacuole for degradation [54]. Similar folding problems have been reported for Na+ ,K+ -ATPase carrying mutations in the conserved TGDGVNDSP motif [55]. Thus, as might be expected from the crystal structures, the interfaces between the P, N, and A domains play a critical role in protein folding and stability. Valiakhmetov and Perlin [56] have recently reported encouraging progress towards mapping the P domain of yeast Pma1 H+ -ATPase. In this study, oxidative


cleavage by Fe2+ was used to identify parts of the 100 kDa polypeptide that lie close to bound Mg2+ and ATP during phosphoryl group transfer. Of the cleavage sites described, three are near conserved residues (Thr-558 and Lys-615) known to take part in the phosphorylation mechanism; the fourth cleavage site (tentatively located within 335-PVGLPA) provides intriguing evidence that the cytoplasmic end of membrane segment 4 may play a hitherto-unsuspected role in this part of the reaction cycle. 3.3 E1–E2 Conformational Change

Profound conformational changes accompany ATP hydrolysis by P-type ATPases [29], and are intricately involved in the transport mechanism. For simplicity, the reaction cycle is usually drawn as an alternation between two major states, E1 and E2 (Figure 3), even though the shift from one to the other presumably involves a carefully orchestrated sequence of smaller movements. Early evidence for structural differences between E1 and E2 came from proteolytic studies of Na+ ,K+ - and Ca2+ ATPases, in which distinctive cleavage patterns could be seen depending upon the step of the reaction cycle (reviewed in Ref. [57]). Differences were also reported in intrinsic tryptophan fluorescence and in the accessibility of individual amino acid residues to group-specific reagents [57]. Several years ago, Karlish and co-workers introduced a versatile way to probe conformational changes in Na+ ,K+ -ATPase by mapping and comparing the sites of Fe2+ - and Cu2+ -catalyzed oxidative cleavage in the presence of ligands that pull the ATPase into the E1 or E2 state. To date, the results fit well with the E1 and E2 structures of Ca2+ -ATPase, and indeed serve to extend those structures by providing information on the conformation-dependent binding of ATP and Mg2+ (reviewed in Ref. [31]). As expected, Pma1 H+ -ATPase also displays major conformational changes during its reaction cycle. Early work by Scarborough [17] and by our laboratory [18] showed that the tryptic fragmentation pattern of the Neurospora enzyme could be altered by ligands that pull the ATPase into the E1 state (ADP) or E2 state (vanadate). Further insight into the nature of the E1 –E2 conformational change has come from a novel group of yeast H+ -ATPase mutants. Haber and co-workers selected the first example of this group (S368F) based on its resistance to hygromycin B [58], an antibiotic later shown to require a negative membrane potential for uptake into the cell [59]. Not only was H+ -ATPase activity reduced in S368F, but the mutant enzyme also had three kinetic abnormalities: a large increase in IC50 for inhibition by vanadate, a several-fold lowering of the K 1/2 for MgATP, and an alkaline shift in pH optimum [60]. Soon afterwards, three similar mutants were identified by our laboratory as part of a scanning mutagenesis study of membrane segment 4 (I332A, V336A, V341A; Ref. [71]). In this case, the amino acid substitutions were located in the middle of M4, making it difficult to imagine that vanadate binding was affected directly. Much more likely was a shift in the conformational equilibrium of the ATPase from E2 (where vanadate binds tightly as a Pi analogue) towards E1 (which is expected to have low affinity for vanadate but high affinity for ATP

9


and H+ ; see reaction cycle in Figure 3). Mutants with similar kinetic changes have since been described for Na+ ,K+ -ATPase [61, 62], gastric H+ ,K+ -ATPase [63], and sarcoplasmic reticulum Ca2+ -ATPase [64, 65] and interpreted in the same way. The discovery of the vanadate-resistant phenotype has provided a convenient way to map regions of the yeast H+ -ATPase polypeptide that work together to affect the E1 -E2 equilibrium. Not surprisingly, given the global nature of the conformational change [29], “E1 –E2 ” mutations are scattered throughout the 100 kDa polypeptide. Against this general background, however, a remarkable cluster has recently been discovered in a scanning mutagenesis study of the stalk region (S4) that links the cytoplasmic end of membrane-spanning segment 4 to the phosphorylation site, D378. As reported by Ambesi and co-workers [66], mutants at thirteen successive positions from I359 through G371 (including S368F) have undergone a 10- to 200fold increase in the IC50 for vanadate; most of them also show a parallel decrease in the apparent affinity for MgATP and an alkaline shift in pH optimum. In the Ca2+ -ATPase-based homology model of the H+ -ATPase [32], the I359–G371 stretch forms a short α-helix (P1) that is oriented transversely between the membrane and the phosphorylation site (D378). Based on the kinetic behavior of the E1 E2 mutants, it seems likely that this helix plays a central role in the conformational shift that lies at the heart of the transport mechanism.

3.4 H+ Pumping

Sequence identity drops to less than 20% in the M domain of P2 -type ATPases, making it difficult to recognize functional features based on sequence analysis alone. Even before the Toyoshima E1 Ca structure appeared, however, there was considerable evidence from mutagenesis studies that membrane-spanning segments 4, 5, 6, and 8 (M4, M5, M6, and M8) define the actual transport pathway of P2 -ATPases [67–70]. The way in which these four segments cooperate to form the two Ca2+ -binding sites of Ca2+ -ATPase is now clear from the 2.6 Å crystal structure of that enzyme [27]. As summarized above, side-chain oxygen atoms (D, E, N, T) and main-chain carbonyl oxygen atoms (A, V, I) are involved, and the actual coordination geometry is made possible by local unwinding of M4 and M6. Detailed structures are not yet available for other P2 -ATPases, but site-directed mutagenesis has begun to shed light on the residues involved in cation binding and transport. For the yeast Pma1 H+ -ATPase, scanning mutagenesis has been performed along the entire length of M4 [71], M5 [72], M6 and M8 (Petrov et al. and Guerra et al., manuscripts in preparation) and throughout much of M1 and M2 [73]. Among the mutants that have been constructed and analyzed, four types of defects have been noted: i) Protein misfolding, leading to ER retention and eventual degradation. Mutations at 16 intramembrane positions fall into this category, presumably because they disrupt the precise helix packing needed to stabilize the M domain of the H+ ATPase. Not surprisingly, changes in charged residues are poorly tolerated in M5


(R695, H701) and M6 (D730, D739) of the H+ -ATPase (Petrov et al., manuscript in preparation). Kaplan and co-workers have shown that the M5–M6 hairpin of Na+ ,K+ -ATPase is only tenuously embedded in the lipid bilayer, requiring occluded K+ ions to keep it from being lost into the medium after cytoplasmic portions of the polypeptide are removed by proteolysis [74, 75]. M8 appears to play an even more important role in proper maturation of the H+ -ATPase, since alanine substitutions at seven positions along this segment (I794, F796, L797, Q798, I799, L801, I807) lead to misfolding and retention in the ER (Guerra et al., manuscript in preparation). ii) Decrease in ATPase activity. Alanine substitutions in M4 (at G333, L338, V341), M5 (Y691, S699), M6 (L721, V723, F724, I727, F728, L734, Y738) cause H+ ATPase activity to fall below 20 % of the value seen in the wild-type control [71, 72]; Petrov et al., manuscript in preparation. The preponderance of such mutants in M6 again calls attention to the special nature of this membrane segment and points to the importance of bulky hydrophobic residues (Phe, Tyr, Leu, Val, Ile) for its proper functioning. iii) Shift in conformational equilibrium from E1 to E2 . The “E1 –E2 ” phenotype, in which the H+ -ATPase becomes strongly resistant to vanadate, has already been discussed (see above). Six such mutants have been discovered in the membrane domain, and the fact that three of them are located in M4 (I332A, V341A, M346A) points to a possible link between S4 and M4 in mediating the E1 –E2 conformational change (Petrov et al., manuscript in preparation). Similar mutants are also found in M1 (M128C; Ref. [73]), M5 (V692A; Ref. [72]), and M6 (V723A; Petrov et al., manuscript in preparation). iv) Mutational change in coupling between ATP hydrolysis and H+ transport. H+ ATPases pose a special challenge in studies of transport mechanisms since there are no radioisotopic methods to assay unidirectional proton fluxes or to detect protons occluded within the membrane. Instead, experimenters generally rely upon a fluorescent probe such as acridine orange or 9-amino-6-chloro-2methoxyacridine (ACMA) to measure the initial rate of proton uptake by ATPase that has been expressed in inside-out secretory vesicles [76] or purified and reconstituted into proteoliposomes [9–11]. Such measurements gain in reliability if the MgATP concentration is varied over a broad range (say, from 0 to 3.0 mM) and the initial rate of fluorescence quenching plotted as a function of the initial rate of ATP hydrolysis. The resulting plots are typically linear, giving evidence for constant coupling across a wide range of pump velocities [71].

To date, more than 80 mutant H+ -ATPases have been analyzed in this way. For most amino acid substitutions, little or no change is seen in the slope of the quenching vs. hydrolysis plot. However, membrane-spanning segments 5, 6, and 8 contain twelve positions at which mutations have caused the slope (or “coupling ratio”) to fall below 50 % of that seen in the wild-type control [72], Petrov et al., and Guerra et al., manuscripts in preparation. Based on these results, both hydrophobic residues (I712, L700, L708, L713, M791) and polar residues (T733, N792, E703, E803) contribute directly or indirectly to the transport pathway, as do two alanines (A726 and A732) and a glycine (G793).

11


An unexpected effect on coupling has come from mutations at two positions in M8 of the yeast Pma1 ATPase. In the first case (E803, homologous to E908 of Ca2+ ATPase), replacement by Asn or Ala caused pumping to fall to barely detectable levels, but replacement by Gln led to a remarkable two-fold increase in the slope of the quenching vs. hydrolysis plot [77]. Approximately one helical turn away, substitution of S800 by Ala also produced a ca. 2-fold increase in slope (Guerra, L., et al., manuscript in preparation). Control measurements, in which the rate of decay of pH was tracked after the addition of vanadate to turn off the pump, showed no significant change in the passive permeability of the membrane [77]. Thus, the enhanced transport seen in S800A and E803Q appears to reflect an actual improvement in the effective stoichiometry of the pump. Further work is required to understand the relationship between this finding and earlier reports of variable stoichiometries in the Neurospora [78] and yeast [79] Pma1 ATPases. Efforts are already under way to model the transport pathway of Pma1 ATPase, with the longer-range goal of synthesizing the results of site-directed mutagenesis into a clear mechanistic picture of how proton transport actually works. On the one hand, it is tempting to draw analogies with better-understood proton pumps such as bacteriorhodopsin [80] and Fo F1 ATPase [81], where the carboxyl group of an essential Asp or Glu residue is alternately protonated and deprotonated as H+ ions move through the membrane. On the other hand, the clearcut resemblance between Pma1 H+ -ATPase and P2 -type Na+ ,K+ , H+ ,K+ , and Ca2+ -ATPases suggests that hydronium ions (H3 O+ ) may be the actual transported species, crossing the membrane via one or more binding sites analogous to those seen in the crystal structure of sarcoplasmic reticulum Ca2+ -ATPase. Both possibilities are currently being pursued. Based on homology modeling, Bukrinsky and co-workers have proposed that D730 (M6) and I331, I332, and V334 (M4) of yeast Pma1 ATPase may form a binding site for H3 O+ , analogous to site II of sarcoplasmic reticulum Ca2+ -ATPase [82]. In a similar model of Neurospora Pma1 ATPase, however, Radresa and his colleagues have pointed to the relative shortage of negatively charged groups in the region corresponding to Ca2+ sites I and II [83]. Rather, these authors emphasize a bacteriorhodopsin-like string of polar cavities, starting at D378 (the catalytic phosphorylation site), that may conduct protons from the cytoplasmic surface into the interior of Pma1 ATPase. It is unclear whether site-directed mutagenesis will be able to distinguish unambiguously between the two types of mechanistic models or whether other kinds of data, including a longhoped-for high-resolution Pma1 structure, will be required.

4 Biogenesis

Alongside studies of reaction mechanism, there has been great interest in the biogenesis of P2 -type ATPases which, as large polypeptides anchored in the lipid bilayer by multiple hydrophobic helices, pose a special challenge to the membrane insertion and trafficking machinery. The yeast H+ -ATPase has proved to be an

4 Biogenesis

excellent model system. Unlike Na+ ,K+ -ATPase, which has been equally well studied (reviewed in [84, 85]), the yeast ATPase lacks the complication of an auxiliary β-subunit. Work in yeast also benefits from powerful genetic tools that are not yet available in mammalian systems. 4.1 Pma1 Mutants with Defects in Folding and Biogenesis

As expected, yeast Pma1 ATPase is synthesized and inserted into the membrane in the rough endoplasmic reticulum [86]. Studies by Addison on the closely related Neurospora H+ -ATPase have defined the way in which alternating signal anchor (SA) and stop-transfer (ST) sequences serve to thread the 100 kDa polypeptide into the bilayer, helped by hairpin formation between neighboring membrane segments [87, 88]. At the earliest time points in a pulse-chase experiment, the ATPase can already be protected against trypsinolysis by low concentrations of MgADP, MgATP, and vanadate, indicating that its cytoplasmic domain has folded to form recognizable binding sites for nucleotides and inhibitors [89]. It then travels along the well-known secretory pathway to the plasma membrane. The availability of nearly 300 point mutants of the yeast H+ -ATPase has provided a wealth of material to explore the structural requirements for ATPase folding and trafficking [90]. At one end of the spectrum are mutants like D378N, D378S, and D378A, which are severely misfolded [91], become trapped in the ER [91, 92], and are eventually ubiquitinated and degraded in the proteasome [93]. Such mutants act in a dominant lethal fashion to prevent growth when co-expressed with wild-type Pma1-ATPase [91, 92]; this behavior has recently been traced to the formation of mixed oligomers between mutant and wild-type polypeptides [38]. At the other end of the spectrum are mutants that display a modest folding defect and a relatively mild genetic phenotype. By means of confocal microscopy, it has been possible to visualize a transient arrest of one such ATPase (G381A) in prominent punctate structures, which resemble the vesicular-tubular complexes (VTCs) that serve as a normal ER-Golgi intermediate in mammalian cells [94]. Thus, the yeast structures may correspond to specific ER exit ports that become amplified as the mutant ATPase is synthesized. The G381A protein eventually escapes the VTCs and travels to the plasma membrane, still in a misfolded, trypsin-sensitive state. There, its behavior points to an additional, yet-to-be-characterized quality control step that recognizes the ATPase as defective and sends it to the vacuole for degradation [94]. A separate series of experiments has shown that G381A can impose its phenotype on co-expressed wild-type ATPase, transiently retarding the wild-type protein in the ER and later stimulating its degradation in the vacuole [94]. Both effects serve to lower the steady-state amount of wild-type ATPase in the plasma membrane and can thus explain the co-dominant genetic behavior of the G381A mutation. Between the two extremes are numerous genetically intermediate mutants, which display a recessive lethal phenotype but have not yet been studied biochemically. All three kinds of mutations cluster in regions of the H+ -ATPase that are likely to be

13


important for protein folding: in the 231 TGES and 632 TGDGVNDAP motifs, located at the interface between the A and P domains; in the 535 DPPR and 378 DKTGTLT motifs, situated within or immediately adjacent to the “hinge” regions between the N and P domains; and in the membrane-spanning segments, known to play a critical role in protein folding and stability (reviewed in Ref. [90]). 4.2 Use of Pma1 Mutants to Screen for Other Genes that Play a Role in Biogenesis and Quality Control

Because Pma1 ATPase is delivered to the cell surface by way of the secretory pathway, its biogenesis can be probed with the classical collection of temperaturesensitive “sec” mutants that define the basic architecture of that pathway. For example, sec18ts , sec7ts , and sec6ts strains have made it possible to detect stepwise Ser/Thr phosphorylation of the ATPase between the endoplasmic reticulum and the plasma membrane [95]. The sec6ts strain has also permitted the expression of mutant Pma1 ATPases in secretory vesicles, where ATP hydrolysis and ATPdependent proton transport can be examined quantitatively, free of contamination by wild-type ATPase [76]. More recently, genetic screens have been invented to uncover genes that play a specialized role in ATPase biogenesis (Table 2). In one such approach, Haber and his colleagues [96] began with the pma1-114 mutant, which is moderately resistant to hygromycin B due to a low membrane potential; they then used higher concentrations of the same drug to select “mop” (modifier of pma1) mutations leading to increased resistance. Among the isolates were several allelic strains (mop2) with a reduced amount of Pma1 ATPase at the cell surface. The new mutations did not affect transcription of the PMA1 gene, nor did they cause Pma1 ATPase to be arrested intracellularly. Rather, the 108 kDa Mop2 protein (also known as End4 or Sla2) appears to play a role in setting the amount and/or stability of the ATPase at the plasma membrane. In parallel, Chang and co-workers have used misfolded Pma1 mutants to pinpoint other genes involved in ATPase biogenesis and quality control. One novel gene identified in this way is EPS1, which, when disrupted, suppresses the dominant lethal phenotype of pma1-D378N and re-routes the mutant ATPase to the plasma membrane [93]. Interestingly, EPS1 encodes a protein disulfide isomerase that may act as a membrane-bound chaperone. Other genes, when overexpressed [AST1; [97]] or disrupted [SOP; [98]], allow a temperature-sensitive mutant ATPase (pma1–114) to escape quality control and move to the plasma membrane. The nature of these gene products and the specificity of their role in biogenesis are currently under active study. Kaiser and his co-workers [99] have focused genetically on the formation and activity of COPII vesicles, which carry newly synthesized proteins from the ER to the Golgi. Among ten LST (lethal with sec thirteen) genes identified by a method known as “synthetic lethality”, the product of one (Lst1p) helps to package Pma1 ATPase into the vesicles [99, 100]; significantly, it is homologous to a known COPII

4 Biogenesis Table 2 Proteins governing the biogenesis of Pma1 H+ -ATPase

Protein

Subscellular location

Function

effect on Pma1 ATPase

Ref.

Eps1

ER (integral)

Protein disulfide isomerase

[93]

Lst1

ER (peripheral)

Component of COPII vesicle coat

Ast1

Multiple membranes (peripheral)

Targeting to the plasma membrane

Mop2 (End4, Sla2)

Cytoskeleton

Actin filament organization Cell wall organization and biogenesis Endocytosis, exocytosis Polar budding

Deletion of EPS1 prevents the proteasomal degradation of Pma1-D378N ATPase and reroutes it to the plasma membrane Deletion of LST1 inhibits the delivery of Pma1 ATPase to the cell surface Overexpression of AST1 reroutes a temperature-sensitive Pma1 ATPase mutant from the vacuole to the plasma membrane Mutation of MOP2 reduces the amount of Pma1 ATPase in the plasma membrane

Sop1-16

Various

Not known

Deletion of each SOP gene perturbs the endosomal trafficking and re-routes a temperature-sensitive Pma1 ATPase mutant from the vacuole to the plasma membrane

[98]

[99, 100] [97]

[96]

coat protein, Sec24p. More work will be required to understand the exact function of Lst1p, but its discovery hints at differentiation of the secretory machinery to handle large, physiologically important cargo molecules such as the ATPase. 4.3 Role of Lipid Rafts

In addition to the protein components of the secretory pathway, membrane lipids also play a vital role in the biogenesis of Pma1 ATPase. Recent studies by Simons and his colleagues have shown that newly synthesized ATPase and other proteins destined for the plasma membrane are sorted preferentially into lipid rafts, which form an ordered, sphingolipid- and ergosterol-rich phase within the bilayer [101]. If a mutation such as pma1-7 impairs the ability of the 100 kDa ATPase polypeptide to associate with the rafts, the ATPase is mistargeted to the vacuole rather than the plasma membrane [36]. Thus, the rafts are an essential part of the biogenesis

15


process; at the plasma membrane, their high sphingolipid and sterol content may also protect against the stresses of the external environment [36, 101].

5 Regulation

Although P2 -ATPases are structurally and functionally similar in most respects, very different mechanisms have evolved to regulate their activity; this is perhaps not surprising in view of the wide range of physiological roles played by members of the family. In some cases, regulation is mediated by an interacting protein: for example, phospholamban for sarcoplasmic reticulum Ca2+ -ATPase [102] or 14-3-3 protein for plant plasma-membrane H+ -ATPase [103–105]. Small proteolipids are also associated with many P2 -ATPases and appear to modulate activity (see above). In other cases, regulation is accomplished intramolecularly by posttranslational modification – usually phosphorylation of one or more Ser/Thr residues (reviewed in Ref. [106]). The latter mechanism is clearly involved in the well-documented posttranslational regulation of yeast H+ -ATPase activity by glucose. It has been known since 1983 that, when yeast cells are placed in carbon-free medium, there is a rapid 5- to 10-fold decrease in H+ -ATPase activity, and when glucose is added back, activity rebounds completely in less than 5 minutes [107]. Although the mechanism is not yet fully understood, there is growing evidence to implicate the C-terminus of the ATPase, acting as an autoinhibitory domain [108]. Mutations at potential phosphorylation sites near the C-terminus affect the ability of glucose to stimulate ATPase activity [109], and in thermolysin digests of the ATPase, two (as yet unidentified) phosphopeptides decrease in amount during carbon starvation and increase again upon glucose addition [95]. Thus, it has been proposed that the C-terminus becomes dephosphorylated during carbon starvation, allowing it to interact in an inhibitory way with one or more catalytically important parts of the ATPase; upon addition of glucose, the C-terminus is rephosphorylated and the inhibition is released. Recent evidence points to a role for stalk segment 5 (the cytoplasmic extension of M5) in this regulatory process. Cys substitutions at seven positions along one face of S5 lead to strong constitutive activation of the ATPase [110]. Furthermore, labeling studies with a membrane-impermeant fluorescent maleimide have pinpointed three S5 cysteines that fail to react in ATPase from glucose-starved cells but become reactive in ATPase from glucose-metabolizing cells, pointing to a significant conformational change between the two states of the protein [111]. Work is now needed to identify the Ser/Thr residues that undergo glucose-dependent phosphorylation, and, by means of biochemical and biophysical approaches, to probe the interaction of the C-terminus with other parts of the ATPase including S5. Progress is also being made towards defining the elements of the signal transduction pathway. In particular, Portillo and co-workers have recently identified two kinases (Ptk2 and Hrk1) that may mediate Ser phosphorylation of the ATPase in response to glucose [112].

6 Emerging Knowledge of Other Yeast P-type ATPase

6 Emerging Knowledge of Other Yeast P-type ATPase

The sequence of the yeast genome, published in 1996 [113], allowed the first identification of the entire set of P-type ATPases that support the growth and physiological functioning of a simple eukaryotic cell. In all, there are 16 P-ATPases in Saccharomyces cerevisiae [114]; Table 3. The plasma membrane contains Pma and Ena ATPases that mediate the efflux of H+ and Na+ (or K+ ) from the cell. Of them, only Pma1 ATPase (the subject of this chapter) is well expressed and highly active under normal laboratory conditions. Ena1 ATPase is induced during growth at elevated Na+ or Li+ concentrations and serves to protect the cell against excessive accumulation of Na+ , Li+ , or K+ [115–117]. Ca2+ homeostasis in yeast relies upon a complex system of transporters and signaling pathways including two endomembrane P-type ATPases: Pmr1 and Pmc1, which are homologues of mammalian SPCA and PMCA ATPases [118–125]. Both work together to maintain a low Ca2+ concentration in the cytoplasm. In addition, by pumping Ca2+ into the Golgi, Pmr1 creates a lumenal Ca2+ concentration (10 µM) that permits the proper functioning of the secretory pathway [121]. Interestingly, Pmr1 ATPase can also transport Mn2+ ions, and it plays an essential role in the resistance of yeast to millimolar Mn2+ [122]. By site-directed mutagenesis of amino acid residues with oxygen-containing side chains throughout M4, M5, M6, M7, and M8, followed by screening for altered sensitivity to Mn2+ and ion-chelating agents, Rao and co-workers have identified two residues in M6 (Asn-774 and Asp-778) that are likely to be involved in divalent cation binding [122]. Even more intriguing is a third mutant in M6 (Q783A) that transports Ca2+ normally but displays a 60-fold

Table 3 P-type ATPases in Saccharomyces cerevisiae

Specificity

Protein

Type Subcellular location

Size

Ref.

H+ H+ Na+ K+ Ca2+ , Mn2+ Ca2+ Cu2+

Pma1 Pma2 Ena1 (Hor6, Pmr2) to Ena5a Pmr1 (Ldb1) Pmc1 Ccc2 Pca1 (Cad2, Pay2) Drs2 (Fun38, Swa3) Neo1 Dnf1 Dnf2 Dnf3 Spf1 (Cod1, Pio1, Per9) YOR291w

IIIA IIIA IID IIA IIB IB IB IV IV IV IV IV V V

918 aa 947 aa 1091 aa 950 aa 1173 aa 1004 aa 1216 aa 1355 aa 1151 aa 1571 aa 1612 aa 1656 aa 1215 aa 1472 aa

This chapter [135, 136] [115–117] [118–123] [124, 125] [126, 127] [128] [129–131] [131] [131] [131] [131] [132, 133] [134]

PL

Unknown

a

Plasma membrane Plasma membrane Plasma membrane Golgi Vacuolar membrane Golgi Unknown Golgi Unknown Plasma membrane Plasma membrane Golgi ER Unknown

Various strains of Saccharomyces cerevisiae contain a tandem cluster of up to 5 nearly identical ENA genes. PL: Aminophospholipid.

17


reduction in affinity for Mn2+ [122]; results of this kind promise to be useful in modeling the molecular basis for cation selectivity. Studies have barely begun on the remaining yeast P-type ATPases. Two belong to the Type IB subfamily and are related to the Cu2+ -ATPases known to underlie Menkes and Wilson’s diseases; they are Ccc2, located in a late-Golgi or post-Golgi compartment [126, 127], and Pca1, whose location and function have not yet been established [128]. Five (Drs2/Fun38/Swa3, Neo1, and Dnf1-3) are Type IV ATPases, postulated to “flip” aminophosholipids between leaflets of the membrane bilayer [129–131]. One Type V ATPase (Spf1/Cod1/Pio1/Per9), located in the endoplasmic reticulum, has been implicated in Ca2+ homeostasis [132, 133], while another (YOR291W) is known only as an open reading frame [134]. Thus, it seems certain that the P-type ATPases of yeast will provide rich material for studies of subcellular targeting, molecular mechanism, and physiological regulation for many years to come.

Acknowledgments

The authors are grateful to members of the Slayman laboratory for helpful comments on the manuscript. Work in the laboratory has been supported by research grant GM15761 from the National Institute of General Medical Sciences.

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1

Gastric H+ ,K+ -ATPase Jaim Moo Shin, Olga Vagin, Keith Munson, and George Sachs Membrane Biology Laboratory, West Los Angeles VA Medical Center, Los Angeles, USA


1 Gastric H+ ,K+ -ATPase

The gastric H+ ,K+ -ATPase exchanges cytoplasmic protons for extracytoplasmic potassium ions in an electroneutral manner and, by being coupled to a KCl pathway in the apical membrane of the parietal cell, is responsible for the elaboration of HCl by the parietal cell of the gastric mucosa. The transport of H+ across the apical membrane is by means of conformational changes in the protein driven by cyclic phosphorylation and dephosphorylation of the catalytic subunit of the ATPase. This mechanism places the ATPase in the P2 ATPase family. The gastric H+ ,K+ -ATPase is composed of two subunits, the α-subunit and the β-subunit. The α subunit with molecular mass of about 100 kDa has the catalytic site and the β subunit with peptide mass of 35 kDa is strongly but non-covalently associated with the α subunit. The β subunit has six or seven N-glycosylated sites exposed to the extra-cytoplasmic surface. There is about 60% sequence homology between the Na+ ,K+ -ATPase and the H+ ,K+ -ATPase a subunit, while the CaATPase of sarcoplasmic reticulum shows only about 15% overall homology with the H+ ,K+ -ATPase. The hydropathy profile is, however, very similar, as is the 3D structure based on site-directed mutagenesis. The β subunit has 35% homology to the β 2 subunit of the Na+ ,K+ -ATPase. The gastric H+ ,K+ -ATPase has been a therapeutic target in treatment of acidrelated disease for the last 15 years. Substituted benzimidazoles that inhibit the H+ ,K+ -ATPase are the newest and most effective class of anti-ulcer drugs used to treat various diseases of the upper GI tract.


2 Gastric H+ ,K + -ATPase

1.1 α Subunit of Gastric H+ ,K+ -ATPase

The primary sequences of the α subunits deduced from cDNA have been defined for pig [1], rat [2] and rabbit [3]. The hog gastric H+ ,K+ -ATPase α subunit sequence deduced from its cDNA consists of 1034 amino acids and has an Mr of 114,285 [1]. The sequence based on the known N-terminal amino acid sequence is one less than the cDNA derived sequence [4]. Rat gastric H+ ,K+ -ATPase consists of 1033 amino acids and has a Mr of 114,012 [2], and rabbit gastric H+ ,K+ -ATPase consists of 1035 amino acids, with Mr = 114,201 [3]. Notably, the initial publication of the rat sequence had an N-terminal fragment of retinol dehydrogenase that is used in some microarrays as a marker for the ATPase (Agilent). Homology of the α subunits among the animal species is extremely high (over 97% identities). The gene sequence for human and the 5 part of the rat H+ ,K+ -ATPase α subunits have been determined [5–7]. The human gastric H+ ,K+ -ATPase gene has 22 exons and encodes α protein of 1035 residues, including the initiator methionine residue (Mr = 114,047). The N terminal amino acid is actually glycine. These H+ ,K+ ATPase α subunits show high homology (∼60 % identity) with the Na+ ,K+ -ATPase catalytic α subunits [5]. The gastric α subunit has conserved sequences along with the other P type ATPases, the sarcoplasmic reticulum Ca-ATPase and the Na+ ,K+ -ATPase, for the ATP binding site, the phosphorylation site, the pyridoxal 5 -phosphate binding site and the fluorescein isothiocyanate binding site. These sites are thought to be within the ATP binding domain in the large cytoplasmic loop between membranespanning segments 4 and 5 (N domain). Using the hog gastric vesicles containing the H+ ,K+ -ATPase, it was shown that pyridoxal 5 -phosphate is bound at lys497 of the α subunit in the absence but not the presence of ATP [8], suggesting that lys497 is present in the ATP binding site or in its vicinity [9]. The phosphorylation site was observed to be at asp386 [10], which is well conserved in other P type ATPases. Fluorescein isothiocyanate (FITC) covalently labels the gastric H+ ,K+ -ATPase in the absence of ATP [11]. The binding site of FITC was at lys518 [12]. Later, several additional lysines, such as those at 497 and 783, were shown to react with FITC during the inactivation of the Na+ ,K+ ATPase and to be protected from reaction with FITC when ATP was present in the incubation. Based on these data, similar lysines of the H+ ,K+ -ATPase could be near or in the ATP binding site. The membrane topology of the α subunit has been extensively studied. The hydropathy plots is most commonly used to predict the location of membranespanning α helices [13]. These are based on determining α moving average of hydrophobic, neutral and hydrophilic amino acids using α variety of scales. Interpretation of the hydrophobicity plots of the α subunit of the H+ ,K+ -ATPase based on the primary amino acid sequence suggested an 8 or 10 membrane-spanning segment model for the secondary structure. The first 4 membrane-spanning segments in the N terminal one-third of the α subunit are clearly defined; however, in


the C terminal one-third of the protein α prediction from the hydropathy analysis is more difficult. The C-terminal amino acids of the α subunit are tyr-tyr, which therefore can be iodinated with peroxidase–H2 O2 –125 I on the cytoplasmic side of intact hog gastric vesicles. Digestion with carboxypeptidase Y then released about 28% of the counts incorporated into the α subunit, as would be predicted from α cytoplasmic location of the C terminal tyrosines [14]. These data show that there is an even number of transmembrane segments in the α subunit. In either an 8- or 10-transmembrane segment model, each transmembrane pair connecting the luminal loop has at least one cysteine, which allows fluorescent labeling of the cysteines left after complete cleavage of the cytoplasmic domain by trypsin. The N-terminus of each fluorescent peptide fragment is then defined by N-terminal sequencing. The size of the fragment is determined from Mr measurement in the tricine gradient gels used for the separation. This then allows the C-termination of the peptide to be identified using the lys or arg cleavage sites at this end. Four transmembrane pairs connected by their luminal loop were detected in the hog gastric H+ ,K+ -ATPase digest [15, 16]. A tryptic peptide fragment beginning at gln104 represents the H1/loop/H2 sector. The H3/loop/H4 sector was found at a single peptide beginning at thr291, and the H5/loop/H6 sector at a peptide beginning at leu776. The H7/loop/H8 region was found in a single peptide fragment of 11 kDa, beginning at leu853. These represent the first 4 transmembrane segment pairs of the proposed 10 transmembrane segment model [3]. From these studies, 4 membrane segment/loop/membrane segment sectors were identified, corresponding to H1/loop/H2 through H7/loop/H8. Although the hydropathy plot predicts two additional membrane-spanning segments (H9, H10) at the C-terminal region of the enzyme, no evidence was obtained for this pair biochemically, even though H9 is predicted to have 4 cysteine residues that should be able to bind fluorescein 5-maleimide (F-MI). The absence of F-MI labeling of these hydrophobic segments may suggest that the cysteines in H9 are post-translationally modified. Reduction, ester cleavage by hydroxylamine and chelating agents have not been successful in identifying this putative transmembrane pair after tryptic cleavage. The nature of the post-translational modification, if this is the reason for the lack of cysteine reactivity, is obscure. Many monoclonal antibodies have been generated reacting with the H+ ,K+ ATPase. With the sequence of the α subunit deduced from cDNA, it is now possible to define the epitopes, and to determine the sidedness of these epitopes by staining intact or permeable cells with either fluorescent or immunogold techniques. Antibody 95 inhibits ATP hydrolysis in the intact vesicles, and appears to be K+ competitive [17]. Its epitope was identified by Western blotting of an E. coli expression library of fragments of the cDNA encoding the α subunit, as well as by Western analysis of tryptic fragments. The sequence recognized by this antibody was between the amino acid positions 529 and 561. Since it inhibits intact vesicles this epitope must be cytoplasmic. Its epitope is close to the region known to bind the cytoplasmic reagent FITC, namely in the loop between H4 and H5. The ability

3


of this antibody to immunoprecipitate intact gastric vesicles showed that it was on the cytoplasmic surface of the pump. Antibody 1218 was shown by Western analysis of tryptic fragments and by recombinant methodology to have its major epitope between amino acid positions 665 and 689 [18], and the epitope was then more cleanly defined as seven consecutive amino acids, Asp682-Met-Asp-Pro-Ser-GluLeu688 [19]. This epitope is on the cytoplasmic surface of the enzyme also in the loop between H4 and H5. This antibody does not inhibit ATPase activity. A second epitope for mAb 1218 was also identified to locate between amino acid positions 853 and 946 according to Western blot analysis of tryptic fragments. A synthetic peptide (888–907) containing this second epitope displaced mAb 1218 from vesicles adsorbed to the surface of ELISA wells [17]. Monoclonal antibody 146 was generated against intact parietal cells and subsequently purified and shown to react with rat H+ ,K+ -ATPase. In cells it reacts on the outside surface of the canaliculus, as shown by immunogold electron microscopy [20, 21]. Western analysis of rat, rabbit and hog enzyme showed, surprisingly, that it was present on the β subunit of rat and rabbit and absent from the β subunit of hog. Disulfide reduction eliminated reactivity of this antibody. Comparing sequence of the different β subunits, there is an arg, pro substitution in the hog for leu, val in the rat between the disulfide at position 161 and 178. This suggests that the β subunit epitope of mAb 146 is contained within this region of the subunit. Conversely, there was also an epitope on the α subunit of all three species recognized by mAb 146 (also using Western analysis). This epitope was defined, both by tryptic mapping and octamer walking, to be between positions 873 and 877 of the hog α subunit. This is on, or close to, the extra-cytoplasmic face of H7. The finding using Western blotting that there was an epitope both on the α and β subunits was confirmed by expressing the rabbit subunits individually in SF9 cells using baculovirus transfection. The β subunit reacted in the SF9 cells and on Western blots with mAb 146. The α subunit did not react in the cells but did on Western blots as if the epitope in the α subunit was difficult to access in the absence of α denaturing detergent such as SDS [18]. These data may indicate tight binding between the α and β subunits in this region of the enzyme. A molecular biological method was developed to analyze not only for the presence of the membrane segments defined as above, but also to explore the nature of membrane insertion [22]. A cDNA encoding α fusion protein of the 102 Nterminal amino acids of the rabbit H+ ,K+ -ATPase α subunit linked by α variable segment to the 177 most C-terminal amino acids of the rabbit H+ ,K+ -ATPase β subunit was transcribed and translated in a rabbit reticulocyte lysate system using labeled methionine in the absence or presence of microsomes. The cDNA for the variable region containing one or more putative membrane-spanning segments is synthesized using selected primers in a PCR reaction and ligated into the cDNA construct. Since the β subunit region has 5 consensus N-glycosylation sites, translocation of the C-terminal β subunit part of the fusion protein into the interior of the microsomes can be determined by assessment of glycosylation. The presence of glycosylation is evidence for an odd number of transmembrane segments in the variable region preceding the β subunit part. The absence of glycosylation shows


either the presence of an even number of membrane-spanning segments or absence of membrane insertion. From this approach, the first 4 membrane segments are present and are co-inserted with translation. The last 2 hydrophobic sequences, H9 and H10, also appear to be co-inserted. The information within the H5, H6 and H7 sequences is apparently insufficient for these to act on their own as either signal anchor or stop transfer sequences. The information within the H8 sequence is sufficient for this to act as α stop transfer sequence. However, a stop transfer sequence in the absence of a preceding signal anchor sequence may have no implications for membrane insertion of the stop transfer sequence [23]. That H5 through H7 are indeed membrane inserted in the mature enzyme is clear from the biochemical analyses. A reasonable hypothesis that may explain these translation data is that insertion of H5 through H7 is post-translational, depending on insertion of H9/10, and that H8 then acts as a stop transfer sequence. Alternatively, the presence of the β sequence may also be involved in tethering H8 and thence H5 to H7. The presence of the additional pair of membrane segments predicted by hydropathy at the C-terminus is strongly suggested by these translation data. The combination of techniques described above, namely, sided trypsin proteolysis, epitope mapping, iodination, and in vitro translation, provides evidence for a 10 membrane segment model (Figure 1), with α large cytoplasmic loop between H4 and H5 and a large extra-cytoplasmic loop between H7 and H8. The 10 transmembrane segments determined and defined from the above evidence are named as TM1–10. These data are consistent with the 3D structure derived from the crystals of the SR Ca-ATPase. It is also worth noting that the reaction of the thiophilic luminal proton pump inhibitors provided conclusive evidence for the presence of TM5 and TM6, as discussed below.

1.2 β Subunit of Gastric H+ ,K+ -ATPase

The β subunit was first identified in work analyzing the carbohydrates of gastric membranes [24]. Post-embedding electron-microscopic staining techniques showed that wheat germ agglutinin staining occurred on the inside face of the inside-out gastric vesicles. The same β subunit was found by determining the nature of the antigen in autoimmune gastritis [25]. Using α lectin-affinity chromatography, the H+ ,K+ -ATPase α subunit was co-purified with the β subunit, showing that the α subunit interacts with the β subunit [26–28]. By cross-linking with low concentrations of glutaraldehyde, the α subunit was shown to be closely associated with the β subunit [29, 30]. The primary sequences of the β subunits have been reported for rabbit [31], hog [25], rat [32–35], mouse [36, 37], and human [38]. The hydropathy profile of the β subunit appears less ambiguous than the α subunit. There is one transmembranespanning region predicted by the hydropathy analysis, which is located between positions 38 and 63 near the N-terminus. Tryptic digestion of the intact gastric H+ ,K+ -ATPase produces no visible cleavage of the β subunit on SDS gels. Wheat

5


Fig. 1 Proposed model of the gastric H+ ,K+ -ATPase " subunit. There are three lobes in the cytoplasmic domain, N (ATP binding), P (phosphorylation), and A (activation) regions, and ten transmembrane segments. This model is based on the

sr Ca-ATPase structures published by Toyoshima et al. [94]. The $ subunit is shown with " single membrane-spanning domain and regions of interaction are numbered in both subunits.

germ agglutinin (WGA) binding of the β subunit is retained. These data indicate that most of the β subunit is extra-cytoplasmic and glycosylated. When lyophilized hog vesicles are cleaved by trypsin followed by reduction, a small, non-glycosylated peptide fragment is seen on SDS gels with the N terminal sequence AQPHYS, which represents C-terminal region beginning at position 236. This small fragment is not found either after trypsinolysis of intact vesicles or in the absence of reducing agents. A disulfide bridge must therefore connect this cleaved fragment to the β subunit containing the carbohydrates. The C-terminal end of the disulfide is at position 262. This leaves little room for an additional membrane-spanning α helix. Hence it is likely that the β subunit has only one transmembrane-spanning segment. Reduction of the disulfides of the β subunit inhibits the activity of the H+ ,K+ -ATPase [39]. With Na+ ,K+ -ATPase, the effect of reducing agents on the ability of the enzyme to hydrolyze ATP and bind ouabain was quantitatively correlated with the reduction of disulfide bonds in the β subunit [40]. The β subunit of the Na+ ,K+ -ATPase is necessary for targeting the complex from the endoplasmic reticulum to the plasma membrane [41, 42]. It also stabilizes a


functional form of both the Na+ ,K+ -ATPases [43, 44]. The H+ ,K+ -ATPase β-subunit is also necessary to target the membrane after proper assembly and sorting [45–48]. When the α- and β-subunits of Na+ ,K+ -ATPase and H+ ,K+ -ATPase were expressed in Sf9 cells in different combinations, the hybrid ATPase with the Na+ ,K+ -ATPase α-subunit and the H+ ,K+ -ATPase β-subunit showed an ATPase activity, which was 12% of the Na+ ,K+ -ATPase activity, with decreased apparent K+ affinity and about half the turnover number. Another hybrid ATPase with the H+ ,K+ -ATPase α-subunit and the Na+ ,K+ -ATPase β-subunit showed 9% of the H+ ,K+ -ATPase activity but increased the apparent K+ affinity [49]. Another report has shown that both lipid and the β subunit affect the K+ affinity of the gastric H+ ,K+ -ATPase [50]. Since both enzymes are unique in being K+ counter-transporters, perhaps the β is important to force counter-transport. Seven putative N-glycosylated sites (AsnXaaSer and AsnXaaThr) are found in rabbit H+ ,K+ -ATPase β-subunit [31], conserved in rat [32–35] and human [38], but only six putative N-glycosylation sites are present in the hog gastric β-subunit [25]. The structure of N-linked oligosaccharides of the β subunit of rabbit gastric H+ ,K+ ATPase was identified [51, 52]. All seven N-linked AsnX(Ser/Thr) sites at positions 99, 103, 130, 146, 161, 193, and 222 were fully glycosylated. Asn99 was modified exclusively with oligomannosidic-type structures, Man6 GlcNAc2 -Man8 GlcNAc2 , and Asn193 has Man5 GlcNAc2 -Man8 GlcNAc2 and lactosamine-type structures. Asn 103, 146, 161, and 222 contain lactosamine-type structures. All the branches of the lactosamine-type structure were terminated with Galα-Galβ-GlcNAc extensions. Similarly, the structure of complex glycans of the β subunit of the Na+ ,K+ -ATPase contain polylactosaminoglycans [53]. The role of the carbohydrate chains in membrane trafficking has been studied in HEK-293 cells and in polarized cells such as LLCPK and MDCK cells [54]. In HEK293 cells, enzyme activity was not affected by removal of any single carbohydrate site of the β subunit but removal of all the glycosylation sites resulted in the complete loss of activity. The role of glycosylation in sorting and trafficking is discussed below. The H+ ,K+ -ATPase β subunit has the sequence Phe-Arg-His-Tyr in its cytoplasmic domain. This tyrosine is important in initiating the removal of the H+ ,K+ ATPase from the apical membrane of the parietal cells and in ensuring its return to the TVE compartment in order to terminate the process of acid secretion. The participation of a tyrosine-based signal in the retrieval of the H+ ,K+ -ATPase suggests that this process involves interactions with adaptins and is mediated by clathrincoated pit formation [55]. In mice deficient in the H+ ,K+ -ATPase β-subunit, cells that express the H+ ,K+ -ATPase α-subunit had abnormal canaliculi and were devoid of typical tubulovesicular membranes [56]. Chimeric β-subunits between the gastric H+ ,K+ -ATPase and the Na+ ,K+ ATPase were constructed and co-transfected with the H+ ,K+ -ATPase α-subunit cDNA in HEK-293 cells [57]. Whole cytoplasmic and transmembrane domains of H+ ,K+ -ATPase β-subunit can be replaced by those of Na+ ,K+ -ATPase β-subunit without losing the enzyme activity. Also, the extracellular segment between Cys152 and Cys178, which contains the second disulfide bond, was exchangeable between

7


H+ ,K+ -ATPase and Na+ , K+ -ATPase, preserving the ATPase activity intact. However, when four amino acids (76 QLKS79 ) in the ectodomain of H+ ,K+ -ATPase βsubunit were replaced by the corresponding amino acids (72 RVAP75 ) of the Na+ ,K+ ATPase β-subunit, the ATPase activity was abolished. This region appears essential for effective interaction between the two subunits. 1.3 Regions of Association Between the α and β Subunits

For Na+ ,K+ -ATPase, the last 161 amino acids of the α subunit are essential for effective association with the β subunit [58]. Furthermore, the last 4 or 5 C terminal hydrophobic amino acids of the Na+ pump β subunit are also essential for interaction with the α subunit whereas the last few hydrophilic amino acids are not [59]. Expression of sodium pump α subunit along with the β subunit of either sodium or proton pump in Xenopus oocytes has shown that the β subunit of the gastric proton pump can act as a surrogate for the β subunit of the sodium pump as far as membrane targeting and 86 Rb+ uptake, suggesting some homology in the associative domains of the β subunits of the two pumps [60] To biochemically specify the region of the α subunit associated with the β subunit, the tryptic digest was solubilized using non-ionic detergents such as NP-40 or C12 E8 . These detergents allow the holoenzyme to retain ATPase activity. The soluble enzyme was then adsorbed onto a WGA affinity column. Following elution of the peptides not associated with the β subunit binding to the WGA column, elution of the β subunit by 0.1 N acetic acid also eluted almost quantitatively the TM7/loop/TM8 sector of the α subunit. These data show that this region of the α subunit is tightly associated with the β subunit such that non-ionic detergents are unable to dissociate it from the β subunit [61]. The antibody mAb 146-14 also recognizes the region of the α subunit at the extra-cytoplasmic face of the TM7 segment, and also recognizes the β subunit, a finding consistent with the association found by column chromatography. Using a yeast two-hybrid analysis, a fragment Leu855 to Arg922 of the α subunit was identified as interacting with the β subunit [62]. Also, two different domains in the β-subunit, Gln64 to Asn130 and Ala156 to Arg188, were identified as association domains with the α-subunit by this method. 1.4 Regions of Association of the α Subunits

There has been much suggestive evidence that the α-β heterodimeric H+ ,K+ ATPase exists as an oligomer. Such evidence includes target size [63, 64] and unit cell size of the enzyme in two-dimensional crystals [65]. It was shown that the enzyme did indeed exist as an (αβ)2 heterodimeric dimmer, using blue native gel separation, and cross-linking with Cu2+ -phenanthroline. Membrane-bound H+ ,K+ ATPase reacted with Cu-phenanthroline to provide an α-α dimer. No evidence was obtained for β-β dimerization. ATP prevents this Cu2+ -phenanthroline-induced

2 Kinetics of the H+ ,K + -ATPase

α-α dimerization. It was possible, by blocking free SH groups and subsequent reduction and labeling with fluorescein maleimide, to demonstrate that the site of Cu2+ -oxidative cross-linking was either at cys565 or cys616 [66]. Hence this region of the α subunit in the N domain is in close contact with its neighboring α subunit. Expression studies of the Na+ ,K+ -ATPase in insect SF9 cells and immunoprecipitation reached the conclusion that a similar region of the α subunit of this enzyme was also in close contact [67].

2 Kinetics of the H+ ,K+ -ATPase

H+ ,K+ -ATPase exchanges intracellular hydrogen ions for extracellular potassium ions by consuming ATP. The H+ for K+ stoichiometry of the H+ ,K+ -ATPase was reported to be one [68–70] or two [71, 72] per ATP hydrolyzed. The H+ /ATP ratio was independent of external KCl and ATP concentration [71]. When care is taken to use only tight vesicles at pH 6.1 the stoichiometry is 2H+ per ATP. Clearly, at a luminal pH of 0.8, the in vitro pH reached in the canaliculus, the stoichiometry must be 1H+ per ATP. This could be explained by retained protonation at one of the ion-binding site carboxylic acids at a pH well below the pK a of the carboxylic amino acid. The kinetics of the H+ ,K+ -ATPase have defined many reaction steps, similar to those of the Na+ ,K+ -ATPase [73]. The rate of formation of the phosphoenzyme (EP) and the K+ -dependent rate of phosphoenzyme breakdown are sufficiently fast to allow the phosphoenzyme to be an intermediate in the overall ATPase reaction. The initial step is the reversible binding of ATP to the enzyme in the absence of added K+ ion, followed by a Mg2+ (and proton) dependent transfer of the terminal (γ ) phosphate of ATP to asp386 of the catalytic subunit (E1 -P· H+ ). The Mg2+ remains occluded in the P domain near asp730 [74] until dephosphorylation [75, 76]. The addition of K+ to the enzyme-bound acyl phosphate results in a biphasic two-step dephosphorylation. The faster initial step depends on [K+ ], whereas the slower step is not affected by K+ concentration. The second phase of EP breakdown is accelerated in the presence of K+ , but at [K+ ] i 500 µM the rate becomes independent of K+ concentration. This shows that two forms of EP exist. The first form, presumably E1 P, is K+ insensitive and converts spontaneously in the rate-limiting step into E2 P, the K+ -sensitive form. ATP binding to the H+ ,K+ -ATPase occurs in both the E1 and the E2 state, but with a lower affinity in E2 state (2,000 times lower than for E1 ) [77] (Figure 2). H+ or K+ interacts competitively on the cytoplasmic surface of intact vesicles. The effects of H+ and K+ on formation and breakdown of phosphoenzyme were determined using transient kinetics [78]. Increasing hydrogen ion concentrations on the ATP-binding face of the vesicles accelerate phosphorylation, whereas increasing [K+ ] inhibit phosphorylation. Increasing [H+ ] reduces this K+ inhibition of the phosphorylation rate. Decreasing [H+ ] accelerates dephosphorylation in the

9


Fig. 2 Pump cycle of the gastric H+ ,K+ ATPase. The initial step is proton binding to the enzyme (E1 ·H+ ), followed the reversible binding of ATP to the enzyme in the absence of added K+ ion (E1 ·AT P ·H+ ). The phosphorylation step occurs by a Mg2+ (and proton) dependent transfer of the ter-

minal (() phosphate of ATP to the catalytic subunit (E1 -P ·H+ ). After releasing the proton, the enzyme (E2 -P) binds K+ ion, forming E2 -P ·K+ . Dephosphorylation (E2 -P ·K+ → E2 ·K+ ) followed by release of K+ (E2 ·K+ → E1 ) are the last steps for exchange of H+ with K+ .

absence of K+ , and K+ on the luminal surface accelerates dephosphorylation. Increasing [K+ ] at constant ATP decreases the rate of phosphorylation and increasing ATP concentrations at constant [K+ ] accelerates ATPase activity and increases the steady-state phosphoenzyme level [79]. Therefore, inhibition by cations is due to cation stabilization of a dephospho form at a cytosolically accessible cation-binding site. To determine the role of divalent cations in the reaction mechanism of the H+ ,K+ -ATPase, calcium was substituted for magnesium, which is necessary for phosphorylation. Calcium ion inhibits K+ stimulation of the H+ ,K+ -ATPase by binding at a cytoplasmic divalent cation site. Ca·EP dephosphorylates 10–20 times more slowly than Mg·EP in the presence of 10 mM KCl with either 8 mM CDTA or 1 mM ATP. The inability of the Ca·EP to dephosphorylate in the presence of K+ , compared with Mg·EP, demonstrates that the type of divalent cation that occupies the catalytic divalent cation site required for phosphorylation is important for the conformational transition to a K+ sensitive phosphoenzyme. Calcium is tightly bound to the divalent cation site of the phosphoenzyme and the occupation of this site by calcium causes slower phosphoenzyme kinetics. Since the presence of CDTA or EGTA does not change the dephosphorylation kinetics of the EP·Ca form of the enzyme, it was concluded that the divalent cation remains occluded in the enzyme until dephosphorylation occurs [80].

3 Conformations of the H+ ,K + -ATPase

3 Conformations of the H+ ,K+ -ATPase

H+ ,K+ -ATPase generates HCl in the stomach by the electroneutral exchange of H+ for K+ , dependent on conformation changes in the protein. The alteration of enzyme conformation changes the affinity and sidedness of the ion-binding sites during the cycle of phosphorylation and dephosphorylation. The ions transported from the cytoplasmic side are H+ at high pH. Since Na+ is transported as a surrogate for H+ , it is possible that the hydronium ion, rather than the proton per se, is the species transported [81]. The ions transported inwards from the outside face of the pump are Tl+ , K+ , Rb+ or NH4 + [73, 79, 82]. Presumably the change in conformation changes a relatively small ion-binding domain in the outward direction into a larger ion-binding domain in the inward direction. The E1 conformation of the H+ ,K+ -ATPase binds the hydronium ion from the cytoplasmic side at high affinity. Following phosphorylation, the conformation changes from E1 P·H3 O+ to E2 P· H3 O+ form, which has high affinity for K+ and low affinity for H3 O+ allowing release of H3 O+ and binding of K+ from the extracytoplasmic surface of the enzyme. Breakdown of the E2 P form requires K+ or its congeners on the outside face of the enzyme. With dephosphorylation, the E1 K+ conformation is produced with a low affinity for K+ , releasing K+ to the cytoplasmic side, allowing rebinding of H3 O+ [73]. The steps of phosphorylation of the H+ ,K+ ATPase were studied by measuring the inorganic phosphate, P18 O4 and P16 O4 distribution as a function of time at different H+ , K+ , and Pi concentrations [83]. The formation of E·Pi complex that exchanges 18 O with HOH was slower at pH 5.5 than at pH 8 and is not diffusion controlled, suggesting a unimolecular chemical transformation involving an additional intermediate in the phosphorylation mechanism such as, perhaps, a protein conformational change. From competitive binding of ATP and 2 ,3 -O-(2,4,6-trinitrophenylcyclohexadienylidine) adenosine 5 -phosphate (TNP-ATP), two classes of nucleotide binding sites were suggested [84]. TNP-ATP is not a substrate for the H+ ,K+ -ATPase. However, TNP-ATP prevents phosphorylation by ATP and inhibits the K+ -stimulated pNPPase and ATPase activities. The number of TNP-ATP binding sites was twice the stoichiometry of phosphoenzyme formation. Recently, two moles of phosphate were claimed to be liberated from one mole of phosphoenzyme of the gastric H+ ,K+ -ATPase [85]. It was hypothesized that one mole of Pi is from a high-affinity ATP binding site and the other Pi is from enzyme-bound ATP at a low-affinity site during crosstalk between catalytic subunits. All-sites phosphorylation has been proposed by studies using Pi or acetyl phosphate [86, 87], which showed that the stoichiometry of the maximum amount of phosphoenzyme formed from ATP, that from acetyl phosphate, that from inorganic phosphate (Pi), and the maximum amount of ATP binding to the enzyme was close to 1:2:2:2. The phosphoenzyme formed was shown to be turning over. The addition of K+ reduced the amount of phosphoenzyme from ATP to one-tenth but reduced those from acetyl phosphate or Pi to only a half.

11


Fluorescein isothiocyanate (FITC) binds to the H+ ,K+ -ATPase at pH 9.0, inhibiting ATPase activity but not pNPPase activity [11]. Fluorescence of the FITC-labeled enzyme, representing the E1 conformation, was quenched by K+ , Rb+ , and Tl+ [11, 88]. The quenching of the fluorescence by KCl reflects the formation of E2 K+ . FITC binds at lys516 in the hog enzyme sequence [12]. This FITC binding site apparently becomes less hydrophobic when KCl binds to form the E2 ·K conformation. The FITC labeled Na+ ,K+ -ATPase has quite similar properties [89, 90]. Two K+ ions are required to cause the conformational change from E1 to E2. The binding site of FITC was at lys501. However, several additional lysines at positions 480 and 766 were shown to react with FITC during inactivation of the Na+ ,K+ -ATPase. These lysines were also protected from labeling in the presence of ATP. The fluorescent 1-(2-methylphenyl)-4-methylamino-6-methyl-2,3dihydropyrroloquinoline (MDPQ) was shown to inhibit the H+ ,K+ -ATPase and the K+ phosphatase competitively with K+ [75]. MDPQ fluorescence is quenched by the imidazo-pyridine SCH 28080. The imidazopyridine Me-DAZIP binds to the TM1/loop/TM2 sector of the β subunit [91]. MDPQ binding to the extra-cytoplasmic surface of the pump enhances its fluorescence, suggesting that inhibitor binding occurs to a relatively hydrophobic region of the protein. The fluorescence was quenched by K+ , independently of Mg2+ . The binding of Mg-ATP increased the fluorescence due to the formation of an E2 P·[I] complex [75], suggesting that the binding pocket on the outside surface between TM5 and TM6, as discussed later for these compounds, changes conformation or position between the two major conformers of the enzyme. The fluorescent changes with FITC and MDPQ may reflect relative motion of the cytoplasmic domain and outside surface of the enzyme. In the E1 form, the FITC region is relatively closer to the membrane and the extra-cytoplasmic loop relatively hydrophilic. With the formation of the E2 ·K+ form the FITC region is more distant from the membrane, whereas with formation of the E2 P form the MDPQ binding region moves towards the membrane. These postulated conformational changes are therefore reciprocal in the two major conformers of the enzyme. The effect of trypsin on the gastric H+ ,K+ -ATPase provides evidence for conformational changes as a function of ligand binding. Only K+ of the ionic ligands provided significant protection against extensive tryptic hydrolysis [61]. Neither ATP nor ADP affected the tryptic pattern at high trypsin:protein ratios. (J. M. Shin, unpublished observations). Two large fragments, 67 and 33 kDa, were found in the presence of ATP, Mg2+ and SCH 28080 as a K+ surrogate whereas several fragments were produced in the absence of ligands. These data suggest that the E2 K+ , or more particularly the E2 P·[SCH] conformation of the H+ ,K+ -ATPase, severely limits accessibility of trypsin to most of the lysines and arginines in the β subunit [16, 91]. Extensive tryptic digestion of the gastric H+ ,K+ -ATPase in the presence of KCl provided a C-terminal peptide fragment of 20 kDa beginning at the TM7 transmembrane segment, a peptide of 9.4 kDa comprising the TM1/loop/TM2 sector beginning at asp84, and another peptide of 9.4 kDa containing the TM5/loop/M6

4 Functional Residues of the H+ ,K + -ATPase

sector beginning at asn753 [61]. The C-terminal 20 kDa peptide fragment was suggested to be capable of Rb+ occlusion [76]. When these digests in the absence and presence of KCl are compared, some regions near the membrane can be seen to be K+ protected. The region between gly93 and glu104 near the TM1 segment, the region between asn753 and leu776 near the cytoplasmic side of the TM5 segment, and the region after the TM8 segment, especially the region between ile945 and ile963 containing 5 arginines and 1 lysine, are protected from the trypsin digestion in the E2 ·K conformation [61]. Further, there must be protection prior to TM3 and for some distance subsequent to M4, since no fragment containing these segments was found at an Mr of less than 20 kDa. When the gastric H+ ,K+ -ATPase was cleaved by Fe2+ -catalyzed oxidation in the presence of various ligands, the cleavage patterns were different between the conformational changes. There are two Fe2+ cleavage sites. In Fe2+ site-1, the parallel appearance of the fragments at 230 ESE, near 624 MVTGD, and at 728 VNDS upon transition from E1 to E2 (Rb) conformations were observed. Meanwhile, in Fe2+ site-2, the fragment near 299 HFIH was cleaved independently of conformational changes [74]. These cleavage patterns were identical to those of the Na+ ,K+ -ATPase [92, 93]. The cleavage data showed that structural organization and changes in the cytoplasmic domains, association with E1 /E2 transitions, are essentially the same for the H+ ,K+ -ATPase, the Na+ ,K+ -ATPase, and sr Ca-ATPase. Using the well-defined sr Ca-ATPase structure, the N-domain where ATP binds then inclines nearly 90◦ with respect to the membrane, and the A domain rotates by about 110◦ horizontally following E1 to E2 conformational changes [94].

4 Functional Residues of the H+ ,K+ -ATPase

When the gastric H+ ,K+ -ATPase was digested by trypsin in the presence of a high concentration of KCl, the tryptic membrane digest showed capability for Rb+ occlusion [76], like the Na+ ,K+ -ATPase [95]. As described above, some regions near the membrane were K+ protected, such as the region between gly93 and glu104 near the TM1 segment, the region between asn753 and leu776 near the cytoplasmic side of the TM5 segment, and the region after the TM8 segment, especially the region between ile945 and ile963 containing 5 arginines and 1 lysine. Furthermore, when K+ is removed from this membrane digest, the TM5–TM6 hairpin was released from the membrane, showing that this membrane hairpin is stabilized by K+ ions and that this region is more flexible than the other membrane-spanning regions [96]. Proton pump inhibitors such as omeprazole, pantoprazole, lansoprazole, and rabeprazole bind to Cys813 of TM5–TM6, giving inhibition of activity [15, 97]. This biochemical study shows that TM5 and TM6 must be part of the ion pathway. Using site-directed mutagenesis of the gastric H+ ,K+ -ATPase transfected in HEK293 cells, the TM5–TM6 luminal loop was studied in terms of K+ access to the ion binding domain [98]. Mutations of TM5, TM5–TM6, and M6 regions such

13


as P798C, Y802L, P810A, C813A or S, F818C, T823V, and mutations of TM7–TM8 and TM8 such as E914Q, F917Y, G918E, T929L, F932L, reduced the affinity for SCH28080 up to 10-fold without affecting the affinity for the activating cation, NH4 + . The L809F substitution in the loop between TM5 and M6 resulted in about a 100-fold decrease in inhibitor affinity. C813T mutant showed a 9-fold loss of SCH28080 affinity. All these data suggest that the binding domain for SCH28080 contains the surface between L809 and C813 , in the TM5–TM6 loop and the luminal end of TM6. Mutations of C813 and I816 in TM6 and M334 in TM4 also showed that the inhibitor binds close to the luminal surface of the enzyme [99–102]. When negatively charged amino acid residues in the α-subunit were mutated the results showed that carboxyl groups in the membrane-spanning domain are important for cation binding [103]. Mutation of E820Q showed less sensitivity to K+ and the dephosphorylation was not stimulated by either K+ or ADP, indicating that E820 might be involved in K+ binding and transition to the E2 form of the H+ ,K+ -ATPase [104]. Mutation of E795 at the cytoplasmic side of TM5 showed a decrease of the phosphorylation rate and the apparent ATP affinity, indicating that E795 is involved in both K+ and H+ binding [105]. Mutation of E795 and E820 in TM5 and TM6 resulted in a K+ -independent, SCH28080-sensitive ATPase activity, due to a high spontaneous dephosphorylation rate [106, 107]. Similarly, the positively charged lysine in TM5/TM6 region was mutated. The mutants K800A and K800E of the Buffo bladder H+ ,K+ -ATPase showed K+ stimulated and ouabain-sensitive electrogenic transport. When the positive charge was conserved (K800R), no K+ -induced outward current could be measured, but rubidium transport was present. This shows that a single positive charged residue in TM5 can determine the electrogenicity [108]. The sixth transmembrane (TM6) segment of the catalytic subunit plays an important role in the ion recognition and transport in the P2 -type ATPase families. When all amino acid residues in the TM6 segment of gastric H+ ,K+ -ATPase α-subunit were singly mutated with alanine, four mutants, L819A, D826A, I827A, and L833A, completely lost the K+ -ATPase activity. Mutant L819A was phosphorylated but hardly dephosphorylated in the presence of K+ , whereas mutants D826A, I827A, and L833A were not phosphorylated from ATP. Amino acids involved in the phosphorylation are located exclusively in the cytoplasmic half of the TM6 segment and those involved in the K+ -dependent dephosphorylation are in the luminal half. Several mutants, such as I821A, L823A, T825A, and P829A, partly retained the K+ -ATPase activity accompanying the decrease in the rate of phosphorylation [109]. The cation selectivity of the Na+ ,K+ - and H+ ,K+ -ATPase may be generated through a cooperative effort between residues of the transmembrane segments and the flanking loops that connect these transmembrane domains. Substituting three residues in the Na+ ,K+ -ATPase sequence with their H+ ,K+ -ATPase counterparts (L319F, N326Y, T340S) and replacing the TM3–TM4 ectodomain sequence with that of the H+ ,K+ -ATPase result in a pump that gives 50 % of ATPase activity in the absence of Na+ at pH 6. This effect was not seen when the ectodomain alone is replaced [110]. Many of these mutational results can be rationalized based on the crystal structure of the sr Ca-ATPase.

5 Structural Model of the H+ ,K + -ATPase

5 Structural Model of the H+ ,K+ -ATPase 5.1 Crystal Structure of the H+ ,K+ -ATPase

As discussed above, experimental evidence showed that the H+ ,K+ -ATPase has 10 membrane spanning segments in the catalytic α-subunit and 1 membranespanning segment in the β-subunit. The two-dimensional structure crystals of the H+ ,K+ -ATPase formed in an imidazole buffer containing VO3− and Mg+ ions was resolved at about 25 Å [63, 65]. The average cell edge of the H+ ,K+ -ATPase was 115 Å, containing four asymmetric protein units of 50 x 30 Å. The structure of Na+ ,K+ -ATPase was determined by electron crystallography at 9.5 Å from multiple small 2-D crystals induced in purified membranes [111]. The density map shows a protomer stabilized in the E2 conformation that extends approximately 65 x 75 x 150 Å in the asymmetric unit of the P2 type unit cell. The unit cell dimension of the Co(NH3 )4 ATP-induced crystals of Na+ ,K+ -ATPase was 141 Å [112]. Two-dimensional crystals of the Na+ ,K+ -ATPase were reported to be best formed at pH 4.8 in sodium citrate buffer and to represent a unique lattice (a = 108.7, b = 66.2, r = 104.2) by electron cryomicroscopy There are two high contrast parts in one unit cell [113]. The crystal structure of the sarcoplasmic reticulum Ca-ATPase was resolved at 2.6 Å resolution with two calcium ions bound in the transmembrane domain, which consists often α-helices [114]. Also, the calcium-free E2 state of sr Ca-ATPase was compared with the calcium-bound E1Ca state [94]. In the Ca-ATPase, the Ndomain where ATP binds inclines nearly 90◦ with respect to the membrane and the A domain rotates by about 110◦ horizontally with a change from E1 to E2 conformation. Several attempts have been made to define the tertiary structure of this P2 type-ATPase based on the structure of sr Ca-ATPase combined with sitedirected mutagenesis, cleavage patterns of different conformations and molecular modeling [115–117]. 5.2 Molecular Modeling of the Gastric H+ ,K+ -ATPase

Gastric H+ ,K+ -ATPase is a member of the P2 type family of ATPases that transport ions against their concentration gradients across lipid bilayers at the expense of ATP hydrolysis. These enzymes, which differ in their ion and inhibitor specificities, nevertheless contain numerous stretches of identical amino acid sequence that not only place them in the family but imply a conserved three-dimensional structure. The implicit homology of mechanism has been substantiated during the past 30 years by an array of biochemical and spectroscopic results showing common conformational intermediates in all family members. These intermediates occur along a general reaction pathway which includes two fundamental conformations, E1 in which the binding sites for the outwardly transported ion are accessible from

15


the cytoplasm and E2 where the binding sites for the inwardly transported ion are accessible from the side opposite the cytoplasm (the “outside”). Binding of the outwardly transported ion (H+ or H3 O+ for H+ ,K+ -ATPase) favors ATP phosphorylation of a conserved aspartic acid side-chain giving E1 P. This phosphorylation shifts the conformational equilibrium in favor of E2 and a separate conformer, E2 P, predominates. The fundamental reactive difference between these forms is that E1 P can reform ATP from ADP whereas E2 P cannot. Binding of the inwardly transported ion stimulates dephosphorylation and gives an occluded form E2 [K+ ] in the H+ ,K+ -ATPase where the ion is inaccessible from either side of the membrane, similar to that observed in the Na+ ,K+ -ATPase. Relaxation to the E1 state results in release of the K+ ion to the cytoplasm. The ultimate goal of structural chemists in this area of research has been to define the mechanism of active ion transport in terms of the molecular structures of the reaction intermediates and, secondly, to understand the molecular basis of ion and inhibitor specificity. The first of these aims requires high-resolution crystal structures for each of the reaction intermediates of at least one member of the P2 -type ATPase family. A tremendous advance towards this requirement has been met recently with rabbit sr Ca-ATPase where high-resolution crystal structures have been reported for both E1 ·2Ca2+ [114] and E2 [thapsigargin] [94] conformations. These structures have provided a molecular understanding of the general mechanism of ion transport [116]. The general shape of the sr Ca-ATPase is that of a “Y”, where the lower half of the base of the Y (M domain) passes through the membrane and contains the ion transport pathway while the upper half extends out of the membrane to form the “stalk” and contains the site of phosphorylation (P domain). One arm of the “Y” makes up the N (nucleotide) domain which binds ATP and the other assists in the transfer of essential conformational effects from the active site to the M domain and is designated the A (for “activator”) domain. In the E1 ·2Ca2+ conformation the N and A domains are splayed open while the calcium ions are encaged side by side near the middle of the membrane in roughly octahedral complexes formed by side-chains’ ligands from transmembrane (TM) helices 8, 5, and 6 (site-1) or 5, 6 and 4 (site-2). This splaying of the Y allows Ca2+ access to the ion-binding domain. The N domain in this form is too distant from P to allow for phosphorylation by bound ATP and it was surmised that N must be capable of rotation about a hinge to allow the γ -phosphate of ATP to reach the phosphorylated aspartate [118]. Subsequently, in the E2 conformation which binds ATP with low affinity, the N domain was shown to be tilted over the P domain, which itself has tilted to a more vertical orientation with respect to the plane of the membrane, making the M, P and N domains nearly collinear. Further, ATP with a fully extended triphosphate can span the N and P domains in the E2 state [119], proving the ability of N to achieve an orientation with P which is at least close to that in E1 . ATP and, presumably, similar to E1 P (ADP still being bound and capable of reforming ATP). The position of the A domain in E2 is raised and rotated upon conversion from E1 . Thus the A, P and N domains are substantially gathered together in E2 . These effects are transmitted to the membrane domain where the geometry of the ion ligands is altered, with

5 Structural Model of the H+ ,K + -ATPase

TM4 rotating to move its carboxylate ligand away from ion site-2 and TM6 rotating with respect to TM4 and TM5. This rearrangement optimizes the transport site for release of outward ion and for entry of the counterion from the outside, while formation of the ion complex triggers dephosphorylation, occlusion, and return to E1 . Currently, the molecular structures of E1 P and E2 P, which are essential for fully understanding the conversion of energy into vectorial ion transport, remain unknown. In contrast to the sr Ca-ATPase it has not been possible to crystallize either the Na+ ,K+ - or H+ ,K+ -ATPase in a form necessary for high-resolution structural determination. There have been two recent structures reported, however, for the Na+ ,K+ – ATPase at 9.5 [111] and 11 Å [120] resolution determined by cryoelectron microscopy of the E2 . vanadate conformations. In each case the density envelope conformed fairly well to the high-resolution form of the sr Ca-ATPase E1 ·2Ca2+ structure with the differences clearly explained by closer approximation to the E2 [thapsigargin] structure that has also been published [94]. The identity of the A, P, M, and N domains was clearly evident, demonstrating the expected homology in structure. The density contributed by the β subunit was tentatively identified in the transmembrane region of the higher-resolution structure [111]. Models for the N domain of the Na+ ,K+ -ATPase have also been determined by NMR of this fragment expressed in E. coli. Changes in structure were shown in the presence of high concentrations of ATP and an ATP bound conformation was constructed [121]. This work is of particular benefit for homology modeling since the N domain sequences are considerably different in sr Ca-ATPase and Na+ ,K+ -ATPase with several long deletions and insertions [115]. In contrast, the Na+ ,K+ – and H+ ,K+ ATPases show high homology in the N domain, with no insertions or deletions in the sequence alignments. For the closely related Na+ ,K+ - and H+ ,K+ -ATPases the more common strategy has been to use biochemical results to validate homology modeling based on the known sr Ca-ATPase structures [117]. In homology modeling, the peptide backbone of a known high-resolution structure is used to substitute the sequence of a homologous protein based on an amino acid sequence alignment. Steric clashes are removed and energy minimization then gives the final model. This method has been applied to the yeast H+ -ATPase [122], colonic H+ K+ -ATPase [123], and the Na+ ,K+ - [124] and H+ ,K+ -ATPases [125]. The predictions of a model can then tested experimentally. For instance, mutational analyses have shown that most of the positions in the M domain which provide ion ligands are conserved, albeit with some substitutions to other oxygen-containing side-chains. It is assumed that these changes are at least partially responsible for the separate ion specificities, but mutating them to affect specificity changes is ineffective, implying the precise geometry is critically dependent on subtle differences in global conformation. Similarly, site-specific mutagenesis has shown that many of the residues affecting specific inhibitor binding of ouabain to the Na+ ,K+ -ATPase, and K+ competitive imidazopyridine, SCH28080, to the H+ ,K+ -ATPase ([102, 125, 126], are conserved in the two pumps. The binding sites are therefore formed from some of the same

17


Fig. 3 Gastric H+ ,K+ -ATPase a subunit (backbone in ribbon) in the E2 conformation viewed from the membrane (cytoplasmic side up). The model was constructed by substitution of the rabbit amino acid sequence on to the backbone of the known sr Ca-ATPase structure (pdb.1iwo, the E2 [thapsigargin] conformation) whose N domain backbone had been replaced with that of the Na+ ,K+ -ATPase determined by NMR. Energy minimization was then applied to give the structure shown. The site of phosphorylation is adjacent to the ac-

tive site magnesium (violet sphere). Ion transport sites are near the middle of the membrane. Proton pump inhibitors (imidazopyridines) block K+ access and inactivate acid secretion by covalent modification of cys813 in a cavity formed by ala335 and phe332 from TM4 (green), tyr925 from TM8 (orange), pro808 and leu809 from loop TM5/TM6 (yellow), and TM1/TM2 (blue). The structure and location of the $ subunit (not shown) is currently unknown although a major site of interaction is the M7/M8 luminal loop.

residues in the loops between TM3–TM4, TM5–TM6 and TM7–TM8 (Figure 3). Both of these inhibitors bind to E2 P conformations and prevent K+ access to the ion binding site. Specificity presumably arises from a combination of a few side-chain changes near the site and global differences generated allosterically by distant substitutions whose importance for binding is nevertheless demonstrable by mutation. The presence of an inhibitor cavity or vestibule accessible from the outside near the TM5-M6 loop is further shown by covalent labeling of the H+ ,K+ ATPase at Cys813 at the beginning of TM6 by the covalent proton pump inhibitors (e.g. omeprazole). The homology model of the E2 form of the H+ ,K+ -ATPase with SCH28080 bound accounts accurately for the known active conformer of

6 Acid Secretion and the H+ ,K + -ATPase

the inhibitor as well as for the effects given by specific amino acid mutations around the inhibitor site [125]. In addition, the model predicts disulfide bond linkages in the immediate vicinity of the bound inhibitor, which can be engineered into the protein via cysteine substitution. These results validate the accuracy of homology modeling in the membrane based on the sr Ca-ATPase structure. The fit of the model to the density envelopes reported for the homologous Na+ ,K+ -ATPase mentioned above has led to predictions for the site of β subunit interaction and kinase phosphorylation, but these have yet to be tested experimentally.

6 Acid Secretion and the H+ ,K+ -ATPase

The H+ ,K+ -ATPase is present mainly in the gastric parietal cell. In the resting parietal cell it is present in smooth surfaced cytoplasmic membrane tubules. Upon stimulation of acid secretion, the pump is found on the microvilli of the secretory canaliculus of the parietal cell. This morphological change results in a severalfold expansion of the canaliculus [127, 128]. There are actin filaments within the microvilli and the subapical cytoplasm. In the cytoskeleton system, there is an abundance of microtubules among the tubulovesicles. Some microtubules appeared to be associating with tubulovesicles. Numerous electron-dense coated pits and vesicles were observed around the apical membrane vacuoles in cimetidine-treated resting parietal cells, consistent with an active membrane uptake in the resting state. The cultured parietal cells undergo morphological transformation under histamine stimulation, resulting in a great expansion of apical membrane vacuoles. Immunogold labeling of H+ ,K+ -ATPase was present not only on the microvilli of expanded apical plasma membrane vacuoles but also in the electron-dense coated pits [129]. There is activation of a K+ and Cl− conductance in the pump membrane which allows K+ to access the extra-cytoplasmic face of the pump [130]. This allows H+ for K+ exchange to be catalyzed by the ATPase [131]. This conductance is probably due to the presence of individual proteins in the stimulated membrane. Covalent inhibitors of the H+ ,K+ -ATPase that have been developed to treat ulcer disease and esophagitis depend on the presence of acid secreted by the pump. They are also acid-activated prodrugs that accumulate in the acid space of the parietal cell. Hence their initial site of binding is only in the secretory canaliculus of the functioning parietal cell [132]. These data show also that the pump present in the cytoplasmic tubules does not generate HCl. The upstream DNA sequence of the a subunit contains both Ca and cAMP responsive elements in rat H+ ,K+ -ATPase [133]. There are gastric nuclear proteins present that bind selectively to a nucleotide sequence, GATACC, in this region of the gene [34]. These proteins have not been detected in other tissues. Another regulatory site has also been postulated [134]. Stimulation of acid secretion by histamine increases the level of mRNA for the α subunit of the pump [135]. Elevation of serum gastrin, which secondarily

19


stimulates histamine release from the enterochromaffin-like cell in the vicinity of the parietal cell, also stimulates transiently the mRNA levels in the parietal cell [136]. H2 receptor antagonists block the effect of serum gastrin elevation on mRNA levels [137]. It seems, therefore, that activity of the H2 receptor on the parietal cell determines in part gene expression of the ATPase. Chronic stimulation of this receptor might be expected therefore to upregulate pump levels whereas inhibition of the receptor would down-regulate levels of the ATPase. However, chronic administration of these H2 receptor antagonists, such as famotidine, results in an increase in pump protein, whereas chronic administration of omeprazole (which must stimulate histamine release) reduces the level of pump protein in the rabbit [138, 139]. Regulation of pump protein turnover downstream of gene expression must account for these observations. Probably, the pump remains in the tubular state, preventing retrieval and degradation of part of the retrieved protein with receptor antagonist inhibition. This is consistent with the finding that the half-life of the protein is increased with H2 receptor inhibition [155].

7 Inhibitors of the H+ ,K+ -ATPase

Two types of proton pump inhibitors have been developed to react exclusively with the extra-cytoplasmic surface of the gastric H+ ,K+ -ATPase. One class of inhibitors now commercially available is substituted benzimidazoles such as omeprazole, lansoprazole, rabeprazole, and pantoprazole. The other class is a series of K+ competitive reagents, including imidazopyridines that are also known to react on the outside surface of the pump. 7.1 Substituted Benzimidazoles

The H+ ,K+ -ATPase in the parietal cell secretes acid into the secretory canaliculus, generating a pH of < 1.0 in lumen of this structure. The acidity of this space is more than 1000-fold greater than anywhere else in the body, and allows accumulation of weak bases. Weak bases of a pK a less than 4.0 would be accumulated only in this acidic space and no other acidic space in the body. The first compound of this class with inhibitory activity on the enzyme and on acid secretion was the 2-(pyridylmethyl)sulfinylbenzimidazole, timoprazole [140]. Since a substituted benzimidazole was first reported to inhibit the H+ ,K+ -ATPase, many inhibitors of the H+ ,K+ -ATPase have been synthesized. The first pump inhibitor used clinically was 2-[(3,5-dimethyl-4-methoxypyridin-2-yl)methylsulfinyl]5-methoxy-1H-benzimidazole, omeprazole [141]. The covalent inhibitors all belong to the substituted benzimidazole family [142–146] (Figure 4). These reagents are weak base, acid-activated compounds, which form cationic sulfenamides in acidic environments. The sulfenamides formed react with the SH group of cysteines in

7 Inhibitors of the H+ ,K + -ATPase

Fig. 4 Proton pump inhibitors. These are substituted pyridylmethylsulfinyl benzimidazole. For omeprazole R and R are methyl groups and R and R are methoxy groups. For lansoprazole R and R are H, R is a methyl group and R is a trifluo-

roethoxy group. Pantoprazole has R and R as methoxy groups, R is H and R is a difluoromethoxy group. For rabeprazole R and R are H, R is a methyl group and R is a methoxypropoxy group.

proteins to form relatively stable disulfides. Since the pump generates acid on its extra-cytoplasmic surface, only those cysteines available from that surface are accessible to these sulfenamides if labeling is carried out under acid transporting conditions. Omeprazole is an acid-activated prodrug [147]. Omeprazole can be accumulated in the acidic space and easily converted into a reactive cationic sulfenamide species, which binds to SH group of cysteines in the H+ ,K+ -ATPase [147–151]. Omeprazole has a stoichiometry of 2 mol inhibitor bound per mol phosphoenzyme under acid transporting conditions and is bound only to the a subunit even in vivo [150, 151]. Substituted benzimidazole inhibitors show slightly different effects depending on the inhibitor structure. Two irreversible inhibitors that form cysteine-reactive sulfenamides in the acid space generated by the pump, omeprazole and E3810, appear to inhibit the enzyme in different conformations [152]. Both omeprazole and E3810, 2-{[4-(3-methoxypropoxy)-3-methylpyridin-2-yl]methylsulfinyl}1H-benzimidazole, are acid activated in luminal surface to form active sulfenamide derivatives, which can bind cysteines within the H+ ,K+ -ATPase. The omeprazole-bound enzyme has a lower FITC fluorescence, perhaps due to an E2 -like conformation. Both ATPase activity and steady-state phosphorylation were inhibited. The E3810 bound enzyme showed a high FITC fluorescence, more

21


Fig. 5 Proton pump inhibitor reaction pathway. Proton pump inhibitors have a pyridine moiety of pK a ∼ 4.0, which enables them to be selectively accumulated in the acidic space of the active parietal cell. A second protonation on the benzimidazole ring with a pK a ∼ 1.0 results in electron deficiency of C-2 of benzimidazole, where the unpro-

tonated pyridine N can attack intramolecularly. After this rearrangement, PPIs generate either a sulfenic acid or a cyclic sulfenamide, both of which are very reactive with thiol groups (cysteine in protein), giving a product that is a permanent cation, thereby restricting re-entry into the cytoplasm of the parietal cell.

like a E1 conformation. Fluorescence of the E3810 bound enzyme was quenched by K+ in contrast to the omeprazole-derivatized FITC labeled enzyme. It is not known whether these effects are due to differences in structure of the inhibitors or to differences in location of binding site or both. These proton pump inhibitors have different binding sites. Omeprazole (Figure 5) binds to cysteines in the extra-cytoplasmic regions of TM5/TM6 (cys813) and TM7/TM8 (cys892) [16]. Pantoprazole binds only to the cys813 and cys822 in M5/M6 [15] and lansoprazole binds to cysteines in TM3/TM4(cys321), cys813 inTM5/TM6, and cys892 in TM7/TM8 [153]. These data suggest that, of the 28 cysteines in the α subunit, only the cysteines present in the TM5/TM6 region are important for inhibition of acid secretion by the substituted benzimidazoles. The proton pump inhibitors provided different acid recovery rates in vivo. In human, the half-life of the inhibitory effect on acid secretion is ∼13 h for lansoprazole, ∼28 h for omeprazole and ∼46 h for pantoprazole [154]. The half-time of recovery of acid secretion and ATPase in rats following omeprazole treatment is ∼ 15 h whereas the pump protein half-life is 54 h [155, 156]. Covalent inhibition of the ATPase results in inhibition of acid secretion extending for longer than the

7 Inhibitors of the H+ ,K + -ATPase

plasma half-life of the PPIs. Recovery from inhibition of acid secretion could occur in principle by either de novo synthesis of pump protein and/or reduction of the disulfide by an endogenous cellular reducing agent such as glutathione. The latter mechanism of reversal depends on whether glutathione or other reducing agents can gain access to a particular cysteine disulfide bond. Recovery of acid secretion following inhibition by all PPIs, other than pantoprazole, may depend on both protein turnover and reversal of the inhibitory disulfide bond. In contrast, recovery of acid secretion after pantoprazole may depend entirely on new protein synthesis [157].

7.2 Substituted Imidazo[1,2α]pyridines and other K+ -competitive Antagonists

Reversible inhibitors contain protonatable nitrogens and have various structures. One type is represented by the imidazo-pyridine derivatives [158], others are piperidinopyridines [159], substituted 4-phenylaminoquinolines [160], pyrrolo[3,2-c]quinolines [161]g, guanidino-thiazoles [162], substituted pyrimidines[163] and scopadulcic acid [164, 165]g. Some natural products such as cassigarol A [166] and naphthoquinone [167] also showed inhibitory activity. Many of these reversible inhibitors show K+ competitive characteristics, in contrast to the benzimidazole type. Imidazo[1,2α]pyridine derivatives were shown to inhibit gastric secretion more rapidly than the benzimidazole type since the latter require acid secretion, accumulation and acid activation [168, 169]. SCH 28080, 3-cyanomethyl-2-methyl8-(phenylmethoxy)imidazo[1,2α]pyridine, inhibited the H+ ,K+ -ATPase competitively with K+ [170]. SCH 28080 binds to free enzyme extra-cytoplasmically in the absence of substrate to form E2(SCH 28080) complexes. SCH 28080 inhibits ATPase activity with high affinity in the absence of K+ (Figure 6). SCH 28080 has no effect on spontaneous dephosphorylation but inhibits K+ -stimulated dephosphorylation, presumably by forming a E2 -P·[I] complex. Hence SCH 28080 inhibits K+ -stimulated ATPase activity by competing with K+ for binding to E2 P [171]. Steady-state phosphorylation is also reduced by SCH 28080, showing that this compound also binds to the free enzyme. Using a photoaffinity reagent, 8-[(4azidophenyl)-methoxy]-1-tritiomethyl-2,3-dimethylimidazo[1,2α]pyridinium iodide (Me-DAZIP+ ), part of the binding site of this class of K+ competitive inhibitor was identified to be in or close to the loop between the TM1 and TM2 segments [91]. Site-directed mutagenesis shows that the binding region is also in the vestibule bounded by the loop between TM5 and TM6 [102, 125]. This binding region, as discussed earlier, was identified by site-directed mutagenesis. Another type of fluorescent K+ competitive arylquinoline, 1-(2-methylphenyl)4-methylamino-6-methyl-2,3-dihydropyrrolo[3,2-c]quinoline (MDPQ), shows enhanced hydrophobicity of its environment with formation of the E2 -P·[I] conformer of the enzyme, as if the H1/loop/H2 segment moves further into the membrane in the E2 conformation [75].

23


Fig. 6 SCH 28080 and Me-DAZIP+ . SCH 28080, 3-cyanomethyl-2-methyl-8-(phenylmethoxy)imidazo[1,2"]pyridine, inhibited H+ ,K+ -ATPase competitively with K+ . SCH 28080 binds to free enzyme extra-cytoplasmically in the absence of substrate to form E2 (SCH 28080) com-

plexes. SCH 28080 inhibits ATPase activity by forming an E2 -P·[I] complex. Another imidazo-pyridine derivative, 8-[(4azidophenyl)methoxy]-1-tritiomethyl-2,3dimethylimidazo-[1,2"]pyridinium iodide (Me-DAZIP+ ), inhibits enzyme activity as SCH28080 does.

8 Trafficking of the H+ ,K+ -ATPase

Early electron microscopy suggested stimulation of acid secretion was accompanied by a morphological change in the parietal cell whereby membrane vesicles changed locale from the cytoplasm to microvilli of the secretory canaliculus [128, 172]. Disappearance of membrane vesicles from the cytoplasm that is observed by immunoelectron microscopy in association with acid secretion [173] was also consistent with the pump moving into microvilli. Fluorescence microscopy then demonstrated acid secretion takes place into the microvilluslined secretory canaliculus [174]. The association of a cytoskeletal component, ezrin, with stimulated but not with resting membranes [175] and the presence of other SNARE proteins such as Rab 11 [176] in the parietal cell was taken as further support of the vesicle fusion hypothesis. Freeze–fracture images of rapidly frozen fixed tissue indicated the membrane vesicles were in fact tubules [177, 178], or even a network of tubules [179]. These data imply the microvilli result from tubule followed by tubule fusion-eversion, rather than vesicle fusion to form the microvillus. Stimulation of secretion with a tubular network structure presumably results in eversion of this network to form the microvillar network of the stimulated secretory canaliculus without the need even for individual tubule fusion events [180, 181].

8 Trafficking of the H+ ,K + -ATPase

Thus, regulation of acid secretion in gastric parietal cells reflects the appearance of the pump in the microvilli of the secretory canaliculus as compared with a cytoplasmic location. [180, 181]. As mentioned above, a tyrosine-based motif in a cytoplasmic tail of the β-subunit appears to be responsible for the H+ ,K+ -ATPase relocation to the cytoplasmic compartment of the cell [55]. When this tyrosine was mutated to alanine and mutant β-subunit was expressed in mice, transgenic animals continuously secreted acid in a stimulus-independent manner. It was suggested that the same motif might be recognized as a basolateral sorting signal when the H+ ,K+ -ATPase β-subunit was expressed in MDCK cells [182] but the experimental data did not support this suggestion [183]. Parietal cells, similar to other polarized cells, sort proteins into two delivery routes, either to apical or basolateral domains of the plasma membrane. Newly synthesized proteins are sorted to the trans-Golgi network and recycling proteins are sorted in endosomes. Sorting is based on the presence of intrinsic sorting signals that are recognized by specific sorting machinery [184–186]. Information on the sorting signals within the H+ ,K+ -ATPase subunits has been inferred from expression studies. When expressed in LLC-PK1 cells, which do not contain tubulovesicular elements, the gastric H+ ,K+ -ATPase is located exclusively on the apical membrane [187]. Studies in which the H+ ,K+ -ATPase β-subunit or chimeric complexes of the H+ ,K+ -ATPase α- or β-subunit with the appropriate subunit of the Na+ ,K+ -ATPase were expressed in LLC-PK1 cells have shown that both α- and β-subunits of the H+ ,K+ -ATPase contain apical trafficking signals [182, 187]. The apical sorting signal in the α-subunit appears to reside in the fourth transmembrane domain and to act through long-range interactions with its flanking cytoplasmic loop domains [188]. However, the nature of the apical sorting signal(s) within the β-subunit remains unknown. N-Glycosylation sites might be considered as potential candidates for the apical signals since the gastric H+ ,K+ -ATPase βsubunit is heavily glycosylated and N-glycosylation sites act as apical signals in several secreted and membrane proteins [189–191]. If expressed in non-polarized HEK-293 cells, the H+ ,K+ -ATPase is localized on the plasma membrane [101, 192]. Mutational studies in HEK-293 cells have shown that six of the seven glycosylation sites in the gastric H+ ,K+ -ATPase β-subunit are essential for trans-Golgi network to plasma membrane trafficking of the H+ ,K+ -ATPase β-subunit in HEK-293 cells [193]. The second glycosylation site (Asn103), which is not conserved among the β-subunits from different species, is not critical for the plasma delivery of the protein. The trafficking step that is affected by the removal of N-glycosylation sites is the route from the Golgi to the plasma membrane and not from ER to Golgi. It is possible that, similar to polarized cells, HEK-293 cells have machinery which recognizes sorting signal(s) and places the β-subunit into specific cargo vesicles in trans-Golgi network that deliver the protein to the plasma membrane. There is increasing evidence that apical and basolateral sorting and trafficking pathways exist not only in polarized but also in non-polarized cells [194]. When apical and basolateral proteins are expressed in non-polarized cells, in trans-Golgi network they are sorted into different containers that travel separately until they

25


Fig. 7 Hypothetical models of stimulation of acid secretion. The vesicle fusion hypothesis suggests that upon stimulation the H+ ,K+ -ATPase containing vesicles relocate from the cytoplasm to the microvili and fuse with the apical membrane, with

a contribution of cytoskeletal components and SNARE proteins. The tubule-eversion model suggests that stimulation of acid secretion results in fusion and eversion of the H+ ,K+ -ATPase containing tubules rather than individual vesicle fusion events.

fuse with plasma membrane [195–198] (Figure 7). This implies that trans-Golgi network to plasma membrane delivery in HEK-293 cells depends on the presence or absence of apical sorting information encoded by particular glycosylation sites within the extracellular domain of the β-subunit [199]. The decrease in apical surface expression of the H+ ,K+ -ATPase β-subunit in polarized LLC-PK1 due to the mutation of N-glycosylation sites support this conclusion [200].

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1

Proton Translocating ATPases Masamitsu Futai, Ge-Hong Sun-Wada, and Yoh Wada Osaka University, Osaka, Japan


Abbreviations DCCD, dicyclohexylcarbodiimide; Pi, inorganic phosphate.

1 Introduction

The mechanism of ATP synthesis has been a focus of biochemists for more than four decades. ATP synthase was first identified in the mitochondrial inner membrane, and then found successively in chloroplasts and bacterial membranes (for reviews, see Refs. 1–8). This enzyme synthesizes ATP from ADP and phosphate (Pi) coupled with an electrochemical proton gradient, and is also called protontranslocating ATPase or F-ATPase (F-Type ATPase) because of the reversible proton pumping upon ATP hydrolysis. The name F-ATPase originated from the coupling factor of oxidative phosphorylation sensitive to oligomycin [1]. Escherichia coli F-ATPase is composed of a membrane extrinsic F1 sector and a transmembrane Fo , formed from five (α 3 β 3 γ δ ε) and three (a b2 c10–14 ) subunit assemblies with different stoichiometries, respectively (Figure 1). It can also be divided into a catalytic α 3 β 3 hexamer, stalks (γ εab2 ), and membrane (a b2 c10–14 ) domains. Two stalks have been observed by electron microscopy [2]. The mitochondrial enzyme has additional subunits, possibly with regulatory functions. The higher-ordered structure of the bovine α 3 β 3 γ complex has been solved by X-ray crystallography [3]. Following this breakthrough, the structures of crystals of bovine F1 inhibited by efrapeptin, aurovertin, NBD-Cl (7-chloro-4-nitrobenzo2-oxa-1,3-diazole), and DCCD (dicyclohexylcarbodiimide) were solved by the same group [4–7]. Bovine F1 containing 1 mol MgADP-trifluoroaluminate [8] and two moles MgADP trifluoroaluminate [9] has also been crystallized, and the structures Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Proton Translocating ATPases

Fig. 1 Schematic model of F-ATPase, showing the subunit structures of the catalytic hexamer ("3 $3 ), stalk and membrane domain. The membrane extrinsic F1 and transmembrane Fo sector are shown together with energy coupling between the ATP synthesis–hydrolysis and subunit rotation.

determined. These studies, together with biochemical analysis, have contributed greatly to an understanding of the catalytic site and mechanism. The structures of F1 sectors of other origins have been reported [10–13]. The catalytic site is mainly located in the β subunit of the α 3 β 3 hexamer, and the three sites (one in each β) show strong cooperativity. The amino and carboxyl terminal helices of the γ subunit are located in the center of the α 3 β 3 hexamer, and form a central stalk with the ε subunit. The second or peripheral stalk is formed from the a and b subunits, and the δ subunit is located near the top of the α subunit [14]. The proton pathway is located at the interface between the a and c subunits, and the hairpin structure of the purified c subunit has been solved by NMR [15]. A ring structure formed from multiple c subunits was suggested by early studies involving electron [16] and atomic force microscopy [17, 18], and was extensively analyzed recently through NMR structure and genetic approaches [19, 20]. The X-ray structure of yeast F1 with a c subunit ring has also been reported [21]. The mechanism of coupling of proton transport and ATP synthesis or hydrolysis has been a major question in research on this complicated enzyme [14, 22, 23]. The binding change mechanism proposes rotation of the γ subunit relative to the α 3 β 3 hexamer coupled with the chemistry at the catalytic sites [23]. Rotation of the γ εc10–14 complex relative to α 3 β 3 δab2 upon ATP addition was shown recently, indicating that continuous rotation of the assembly of F1 and Fo subunits is involved in the coupling between ATP hydrolysis and proton transport [24–27]. Vacuolar-type ATPase (V-ATPase) with significant similarity to F-ATPase was introduced later to a family of proton-pumping ATPases (for reviews, see Refs. [28– 31]). V-ATPase is apparently different from F-ATPase in its physiological roles, and forms an acidic luminal pH in endomembrane organelles including lysosomes and

2 Catalytic Mechanism of F-ATPase

endosomes, and in extracellular compartments such as resorption lacuna formed between osteoclasts and the bone surface. It has extrinsic membrane V1 and Vo transmembrane domains formed from the A, B, C, D, E, F, G and H subunits, and a, c, c , c and d, respectively. F- and V-ATPase share significant homology, especially in their catalytic subunits and proton pathways, but also some unique differences, including their membrane sector and stalk region subunit compositions. Thus, comparative studies of the two proton pumps are pertinent for understanding the molecular mechanisms of both. In this chapter we discuss recent progress in the understanding of F-ATPase, focusing mainly on the energy coupling between the chemistry and proton transport through subunit rotation, and briefly on comparison with V-ATPase. It should be interesting for readers to follow a series of biochemical studies to show rotational catalysis of the F-ATPase holoenzyme leading to that of V-ATPase.

2 Catalytic Mechanism of F-ATPase

As expected from its complicated structure, F-ATPase is not a simple Michaelis–Menten type enzyme. Furthermore, the overall mechanism includes catalysis (chemistry), subunit rotation and proton translocation. X-Ray structure and kinetic studies, especially substrate binding analysis with an intrinsic tryptophan probe, revealed that the three catalytic sites are asymmetric. Senior and coworkers have recently extensively summarized and discussed the molecular catalytic mechanism of F-ATPase [32]. Thus, we discuss the catalytic mechanism and active site only briefly here. ATP synthesis and hydrolysis could be carried out kinetically at a single site (uni-site catalysis), two sites operating together (bi-site catalysis), or all three sites working together (tri-site catalysis). Uni-site catalysis has only been demonstrated for ATP hydrolysis, and can be measured experimentally with an ATP:F1 ratio of less than 1:3 [33]. This rate is 105 –106 -fold lower than that of steady-state (multisite) catalysis. The enzyme cross-linked chemically, and thus could not rotate, but could still carry out uni-site catalysis [34]. This catalysis is not a part of the steady state, which includes subunit rotation. Catalytic residues have been identified by analyzing uni-site catalysis of the purified E. coli mutant F1 sector (Figure 2). They were discussed in detail previously for mechanistic implication [32, 35–38]. Briefly, βLys155 of the β subunit is required for binding of the γ phosphate moiety, as shown by studies involving affinity labeling [39] and mutant enzymes such as βLys155Ala (βLys155 → Ala) or βLys155Ser [40–43]. The βArg182 residue is also involved in the binding. Enzymes with substitutions of βThr156 showed similar properties to those of βLys155 [40]. The hydroxyl moiety of βThr156 is essential, possibly for Mg2+ binding, since it can only be replaced by a serine residue [40, 44]. βGlu181 is a critical catalytic residue [41]. Its side-chain forms a hydrogen bond with a water molecule located near the γ phosphate of ATP [3]. However, it was shown later that this residue was not

3


Fig. 2 Catalytic sites of F-ATPase and V-ATPase. The catalytic residues of E. coli F-ATPase are shown together with bound ATP. Their positions are cited according to the bovine crystal structure [3]. Corresponding residues of yeast V-ATPase are also shown [109, 110].

involved in nucleotide binding or Mg2+ coordination [42, 45]. βTyr331, which is stacked close to the adenine ring [3], is required for the binding of ADP or ATP [37]. Results of analysis of the tryptophan fluorescence of the βTyr331Trp mutant F1 are consistent with the role of the βTyr331 residue [22, 23]. The bovine residues corresponding to those discussed above are located close to the phosphate moiety or the adenine ring of bound-ATP or ADP in the X-ray structure [3] (Figure 2). Recent results [9, 11, 37, 46–49], including the F1 structure of all three sites filled with nucleotides [9, 11], support tri-site catalysis, in which the three sites are working together during the steady state, and the notion that bi-site catalysis does not occur [32]. We leave convincing discussion of these points to the article of Senior and coworkers [32]. It can be assumed that mutant enzymes defective in steady state catalysis should show impaired uni-site catalysis. In this regard, αArg376 mutant enzymes are of interest [50]. The αArg376 residue of the α subunit is located close to the β- or γ -phosphate of ATP or ADP and Mg at the catalytic site [3] (Figure 2). However, αAg376 does not directly participate in the chemistry of ATP hydrolysis or synthesis, as shown by the uni-site catalysis of mutant F1 sectors [50]. The αArg376Lys r αArg376Ala mutant enzyme showed 2 × 103 -fold lower steady-state ATP hydrolysis than the wild type. However, the mutant enzymes showed essentially the same kinetics for uni-site catalysis as the wild type, suggesting that they can pass through the transition state. These results indicate that αArg376 is essential for promotion of catalysis to the steady-state turnover. This notion is different from the previously suggested roles of αArg376 deduced from the structural model [3] and

3 Roles of the ( subunit: energy coupling by mechanical rotation 5

fluoroaluminate binding, an indicator of the formation of the pentacovalent transition state [51]. The βGlu185 residue, located close to the γ -phosphate and Mg at the catalytic site, may have a similar role to αArg376 because the mutant enzymes maintain uni-site catalysis but are defective in multisite catalysis [52]. However, detailed analysis could not be carried out because the mutant F1 was unstable after solubilization from membranes. The binding change mechanism of Boyer [23] proposes that the three sites are involved sequentially in ATP synthesis or hydrolysis: at a specific time point during the steady state, the chemical reaction occurs reversibly at one site, and ATP release and/or binding of ATP + Pi at two other sites with the expenditure of energy. Evidence supporting catalytic cooperativity and the binding change mechanism has accumulated, as reviewed previously [23]. It includes early kinetic evidence of uni-site and steady-state catalysis showing three K m values and three interacting nucleotide binding sites, 18 O isotope exchange reactions, inhibitor studies, etc. These studies suggested that the chemistry, “ADP + Pi ←→ ATP + H2 O”, at the catalytic site is reversible, and provides essentially no energy change. The mechanism is strongly supported by the X-ray structure showing asymmetric catalytic sites and continuous γ subunit rotation [3]. However, it can not be concluded that the binding change mechanism was proven at the molecular level [32], although the mechanism is conceptually accepted and has been extremely useful for understanding the enzyme.

3 Roles of the γ subunit: energy coupling by mechanical rotation 3.1 Roles of the γ Subunit in Energy Coupling

The essential role of the γ subunit in catalysis and assembly was shown by early reconstitution experiments: a catalytic core complex exhibiting ATPase activity could be reconstituted from the purified E. coli α, β and γ subunits, but not without the γ subunit [53, 54]. Consistently, assembly of the F1 sector is strongly affected by γ subunit mutations, especially those of residues interacting with the β subunit [55]. The isolated α 3 β 3 γ complex could functionally bind to the Fo sector only after its assembly with the δ and ε subunits [54]. These results suggest that the two minor subunits are required for the functional binding of α 3 β 3 γ . Early genetic approaches focused on the γ subunit carboxyl terminal region [56–58]. The amino acid sequences of the γ subunits are weakly conserved among different species [57–59], although their X-ray structures are similar [3, 11–13]. When the known γ sequences were aligned, only 28 of the 286 residues of the E. coli subunit are conserved, and mostly in the carboxyl and amino terminal helices located at the center of the α 3 β 3 hexamer [58]. Seventeen residues are conserved between residues 242 and 286 of the γ subunit. An enzyme with a nonsense mutation lacking 10 residues at the carboxyl terminus is still capable of in vivo ATP


synthesis [57], indicating that the three conserved residues in this region are not required. Structural flexibility of the carboxyl terminus was also demonstrated by a frameshift mutation: the enzyme was still active with the γ subunit having seven additional residues at its carboxyl terminus together with nine altered residues downstream of γ Thr277 [60]. The enzyme with the nonsense mutation (γ Gln269End) was inactive, and substitution of a conserved residue (γ Gln269, γ Thr273, or γ Glu275) between γ Gln269 and γ Leu276 gave enzymes with reduced ATPase activity and energy coupling [57]. We noticed that mutations resulted in different ratios of ATPase catalysis and proton transport; three mutants (βThr277End, βGln269Leu, and βGlu275Lys) exhibited about 15 % of the wild-type ATPase activity, but showed various degrees of ATP-dependent H+ transport and in vivo ATP synthesis. These results suggest active role(s) of the γ subunit in ATPase activity and energy coupling. In addition to the carboxyl terminus, the only other conserved region is near the amino terminus [58]. The importance of this region was first suggested by the mutant lacking residues between γ Lys21 and γ Ala27, which resulted in failure of assembly of the F1 complex [61]. We introduced amino acid substitutions systematically into the amino terminal region of the γ subunit [62]. Most of the changes between γ Ile19 and γ Lys33, γ Asp83 and γ Cys87, or at γ Asp65 had no effect, even with a drastic replacement such as hydrophobic to hydrophilic or acidic to basic. Interesting exceptions were the γ Met23Arg and γ Met23Lys substitutions. These mutants grew only slowly on succinate through oxidative phosphorylation, indicating that they are impaired in ATP synthesis. However, the membranes prepared from the γ Arg23 and γ Lys23 strains showed 100 and 65% of the wild-type ATPase activity, but formed only 32 and 17 % of the electrochemical proton gradient, respectively, indicating that these mutants are defective in energy coupling between ATP hydrolysis and proton transport. In the X-ray structure, the γ Met23 residue is located close to the DELSEED (βAsp380–βAsp386) loop of the β subunit [3]. Thermodynamic and kinetic analyses of the purified γ Met23Lys enzyme suggested that the introduced γ Lys23 residue forms an ionized hydrogen bond with βGlu381 in the loop [63]. Consistent with this interpretation, the phenotype of γ Met23Lys was restored by the second mutation, βGlu381Gln [64]. The βGlu381Lys mutation also caused deficient energy coupling. These results suggest that the interaction between the regions around γ Met23 and βGlu381, and thus the γ subunit residue and β subunit DELSEED loop, are involved in energy coupling. The γ Met23Lys mutation was suppressed by substitution of carboxyl terminal residues, including γ Arg242, γ Glu269, γ Ala270, γ Ile272, γ Thr273 γ Glu278, γ Ile279k, and γ Val280 [65]. From these results, we initially suggested that γ Met23, γ Arg242, and the region between γ Glu269 and γ Val280 are three interacting domains that are required for efficient energy coupling. The X-ray structure shows that γ Met23 located in the amino terminal helix is near γ Arg242, but γ Gly269 and γ Val280 in the carboxyl terminal helix are near the top of the α 3 β 3 hexamer [3]. We further isolated second site mutations that suppressed the primary mutations, γ Gln269Glu and γ Thr273Val [66]. These mutations were mapped to the amino (residues 18, 34, and 35) and carboxyl (residues 236, 238, 242, and 262) termini.


The higher-ordered structure clearly shows that γ Glu269 or γ Thr273 does not interact directly with the residues of the second mutations. The occurrence of suppression at a distance may suggest that the two a helices of the γ subunit located at the center of the α 3 β 3 complex undergo long-range conformational changes during catalysis [67]. As expected, the relative orientations of the γ subunit to the three β subunits (β E , β DP , and β TP ) are different in the crystal structure, strongly supporting γ subunit rotation during ATP hydrolysis or synthesis [3]. 3.2 γ Subunit Rotation

The higher-ordered structure of α 3 β 3 γ indicated that a and β are arranged alternately around the amino- and carboxyl-terminal a helices of the γ subunit [3, 14]. The binding change mechanism predicts that the catalytic sites in the three β subunits participate sequentially in ATP synthesis or hydrolysis via conformation transmission through the γ subunit rotation [23]. This rotation was suggested by experiments on chemical cross-linking between γ Cys87 and βCys380 (originally βAsp380) in the DELSEED loop [68, 69], and analysis of polarized absorption recovery after photobleaching of a probe linked to the carboxyl terminus of the chloroplast γ subunit [70, 71]. The continuous unidirectional γ subunit rotation in F1 was recorded directly by Yoshida and Noji and their colleagues [72] (Figure 3). They immobilized the Bacillus α 3 β 3 γ complex on a glass surface through a histidine-tag introduced into the β subunit. The fluorescent actin filament connected to the γ subunit rotated continuously in an anticlockwise direction during ATP hydrolysis. The rotation became slower with an increase in the filament length, and generated a frictional

Fig. 3 Rotation of the ( subunit of F-ATPase. F1 was immobilized through a histidine tag introduced to the " or $ subunit, and an actin filament was connected to the carboxyl terminus of the ( subunit. ATP-dependent rotation of the wild-type (open circles) and ( Met23Lys mutant (filled circles) F1 sector are shown. Taken from Omote et al. [77].


torque of ∼40 pN nm. The ε subunit rotation was also shown using the same approach [73], consistent with the tight association of γ and ε [21, 74]. A 120◦ step rotation was shown in the presence of a dilute ATP concentration, indicating that the γ subunit rotates, interacting with the three β subunits successively [75]. Furthermore, a refined measurement system involving gold beads revealed that the 120◦ step could be divided into 90◦ and 30◦ steps [76]. These two steps were proposed to correspond to ATP binding and ADP release, respectively, although this model is difficult to prove unambiguously. Interestingly, the rotation rates observed were much higher than those expected from the V max of steady-state catalysis. We could observe the rotation of an actin filament connected to the γ subunit of E. coli F1 [77] using a similar system to that described for the Bacillus α 3 β 3 γ complex [72]. The rotation was anticlockwise, inhibited by azide (F-ATPase inhibitor), and generated a frictional torque of ∼40 pN·nm, as reported for the Bacillus complex. The F1 sector, proven to be a chemically driven motor, should be connected functionally to the Fo sector to complete ATP-driven proton transport. Conversely, proton transport through Fo should be coupled to the γ subunit rotation and chemistry at the catalytic sites. Questions on the energy coupling between chemistry, rotation and proton transport could be answered with the E. coli enzyme by taking advantage of the accumulated genetic and biochemical information. 3.3 Mutational Analysis of the γ Subunit Rotation

We were interested in characterizing the γ rotation of a series of mutant enzymes defective in energy coupling and catalytic cooperativity. As described above, the γ Met23Lys mutant is defective in energy coupling between catalysis and proton transport [62]. We thought that the γ rotation in the mutant F1 may be defective, since the γ residue was replaced. Similar to the original mutant, the γ Me23Lys enzyme engineered for rotation observation exhibited essentially the same ATPase activity as the engineered wild type. However, the enzyme could not form an electrochemical proton gradient in membrane vesicles, or carry out in vivo oxidative phosphorylation [77]. Unexpectedly, an actin filament connected to the γ subunit of γ Met23Lys rotated, and generated essentially the same torque as that of the wild type (Figure 3). These results suggest that the γ Met23Lys mutant F-ATPase could couple between chemistry and rotation, but was defective in transforming mechanical work into proton translocation or vice versa. In this regard, Al-Shawi et al. showed that the γ Met23Lys mutant is defective in communication between F1 and Fo [63]. These results also suggested that analysis of F1 sector rotation is not enough to understand F-ATPase. It became imperative to determine which subunit complex is rotating in the F-ATPase holo enzyme purified or embedded in the membrane. Mutant F1 sectors with substitution of the βSer174 residue and their suppressors have been useful for understanding the rotation mechanism (Figure 4). Replacing βSer-174 with other residues altered the ATPase activity to between 150 and 10 % of the wild-type level, and the larger the side-chain of the residue introduced the lower


Fig. 4 Models of the E. coli $ subunit domain including $Gly149 and $Ser174. (a) Domain structure for the ATP-bound ($TP ) and empty ($E ) $ subunit. The bovine structure [3] was used to model E. coli domains. Positions of ATP and residues discussed

in the text are indicated. Nomenclatures for the a helix and $ sheet are those of previous workers [3]. (b) Models of the $Ser174Phe mutant domain structure. The positions of the residues discussed in the text are shown. Taken from Iko et al. [79].

the ATPase activity observed [78]. Both the βSer174Leu and βSer174Phe enzymes retained about 10 % of the wild-type ATPase activity. However, the two mutants showed a difference in energy coupling: the βSer174Leu mutant could still grow on succinate by oxidative phosphorylation and transport protons into the isolated membrane vesicles, whereas the βSer174Phe mutant could not grow and showed no proton transport. Consistent with these observations, their F1 sectors differed in γ subunit rotation [79]. The F1 sector with βSer174Phe showed apparently slower rotation than the wild type, and generated significantly lower frictional torque (∼17 pN nm), whereas the F1 with βSer174Leu was similar to the wild type. These results suggest that the rotation or torque generation is closely related to the energy coupling with proton transport. Biochemical defects of the βSer174Phe mutation were suppressed by a secondsite mutation, βGly149 to Ser, Cys, or Ala [80]. Double mutants such as βSer174Phe/βGly149Ser showed essentially the same ATPase activity and proton transport as the wild type. As expected from these results, the double mutant F1 generated the wild-type torque (∼40 pN nm) [79]. The high-resolution structure of the bovine F1 predicts that βSer174 is located on the β subunit surface within the loop between an α helix (helix B) and a β sheet (β sheet 4) (Figure 4). βGly149, the first residue of the phosphate binding P loop connected to helix B, is close to the catalytic site. The structure of the βGly149–bSer174 region is significantly


different between the nucleotide-bound and empty β subunits. Thus, it can be assumed that the conformational change of the catalytic site, followed by that of the βGly149–βSer174 domain, leads to the γ subunit rotation for the energy coupling to proton transport. The βSer174Phe mutation was also suppressed by αArg296Cys of the α subunit [78]. It was of interest to analyze the rotation of the double mutant and related strains. However, the F1 sector of βSer174Phe/αArg296Cys was difficult to purify. Energy minimization with a simple potential function of the modeled βSer174Phe mutant F1 predicts that the side-chain of βPhe174 interacts with that of βIle163 or βIle166 of the β subunit with no nucleotide bound [79]. We substituted βIle163 or βIle166 with a less bulky Ala residue in the βSer174Phe mutant. As expected, the F1 with the βIle163Ala/βSer174Phe or βIle166Ala/βSer174Phe double mutant could rotate and generate almost similar torque to the wild type. These results suggest that the βGly149–βSer174 domain plays important roles in the rotation and torque generation essential for energy coupling. The role of the DELSEED loop has been focused on because the conformations of the three β subunits (β E and β T or β D ) are significantly different [3], which is consistent with the roles of γ Met23 [62] and βGlu381 (the second residue of the DELSEED loop) [63] in energy coupling. However, mutagenesis studies involving replacement of residues in the DELSEED loop of thermophilic Bacillus α 3 β 3 γ did not reveal significant effects on the torque generation [81]. The negative results may suggest that the loop is not related to the conformation change driving the β subunit rotation. However, this is difficult to conclude from the directed mutations without structural analysis. Furthermore, the experimental system with actin filaments gives rotation rates and torque values with high deviations. Extensive substitution of related residues and their second mutations should be analyzed for the final conclusion, as discussed above for a region around βSer174. Furthermore, analysis of rotation in the F-ATPase holoenzyme may be necessary, as pointed out for the γ Met23Lys mutation [77].

4 Rotational Catalysis of the F-ATPase Holoenzyme 4.1 Structure of Fo Sector and Proton Transport Pathway

As discussed above, the Fo membrane sector is composed of a, b and c subunits, whose stoichiometry (1:2:10 ± 1, for a:b:c) was first determined from the stained bands on polyacrylamide-gel electrophoresis [82]. Based on structural prediction, Cox et al. proposed a model of Fo in which the a and b subunit helices are surrounded by a ring of c subunits [83]. However, this model was not consistent with the electron [16] and atomic force [17, 18] microscopic images. In the model derived from the images, the a and b subunits are attached to one side of the symmetric ring structure formed by the c subunits. An atomic force microscope image indicated

4 Rotational Catalysis of the F-ATPase Holoenzyme

< 12 c subunits in the ring. The ab2 complex was purified recently after solubilization of Fo from membranes utilizing a histidine tag introduced at the a subunit amino terminus [84]. The proton pathway (Fo ) was reconstituted from the ab2 complex and c subunit. These results confirmed that ab2 and the c ring are two functional subcomplexes of the Fo sector. Subunit a spans the membrane with five helices, and its fourth helix could be cross-linked to the carboxyl terminal helix of a c subunit when Cys residues were introduced at appropriate positions [85, 86]. aArg210 in the fourth helix is involved in proton transport, and all the substitutions, even aArg210 to Lys, gave a Fo sector with no ability to transport protons [19, 20, 87]. The structure of the amino terminus (residues 1–34) of the b subunit, including the trans-membrane helix, was solved by NMR in a chloroform–methanol–water mixture [88], and extramembrane helices interact with the δ and α subunits at the top of the F1 sector [89, 90], which is consistent with the model proposing that the b subunit closely interacts with an α 3 β 3 hexamer [20]. The b subunit dimer interaction with the F1 sector is necessary for a second stalk [88]. The carboxyl terminal helix of a c subunit can be cross-linked to the membrane helix of the b subunit [91]. Fillingame and co-workers solved the structure of the E. coli c subunit by NMR [15]: monomeric c (in a mixture of chloroform, methanol and water, pH 5) give a hairpin-like structure formed from the two helices connected by a polar region that interacts with the γ and ε subunits. Structural models of Fo and the c ring were reviewed recently by Fillingame and coworkers [19, 20]. Thus, we discuss here only the pertinent points of the rotational catalysis. The front face of one c subunit packs with the back face of a second c subunit to form a dimer, consistent with the results of mutant studies [19]. Fillingame and coworkers proposed that dimers form a ring of 12 monomers, with the amino and carboxyl α helices of each c subunit in the interior and at the periphery, respectively, and the ring was modeled from the results of molecular dynamic calculations [92]. The model was supported by cross-linking analysis [93]. However, a ring containing more than 10 monomers was not found in purified Fo F1 [94], and recent recalculation gave essentially the same but a slightly smaller ring formed from 10 copies of the c subunit with the two helices in similar orientations [20, 95]. In an alternative model, the carboxyl and amino terminal α helices form inner and outer rings, respectively [96]. However, this model is not supported by cross-linking experiments [19]. The X ray structure indicated a yeast F1 tightly bound with a c ring formed from ten monomers [21]. These results established that the c subunits form a ring of ten monomers. However, the number of monomers forming the ring is still controversial [21, 24]: atomic force microscopy of chloroplast [97] and bacterial [98] Fo demonstrated rings of 14 and 11 c subunits, respectively. The difference in the copy number may be due to the species difference, loss of a part of the monomers, or reorganization of the ring during purification. Structural studies on F-ATPase or the Fo sector will eventually explain the discrepancy. cAsp61, in the middle of the second transmembrane a helix, is responsible for proton transport, and close to cAla24 and cIle28, of which substitutions by other

11


residues reduced the DCCD reactivity of cAsp61 [18]. The stoichiometry of the a and c subunits (1:10) indicates that one aArg210 and multiple cAsp61 are required for proton translocation in F-ATPase. As the pKa of the cAsp61 carboxyl moiety is 7.1, the NMR c subunit structure at pH 5 is at the fully protonated stage. The interaction between cAsp61 and aArg210 may lower the pKa to facilitate proton release into the proton pathway [19, 20]. Restogi and Girvin solved the c subunit structure at pH 8, with cAsp61 being in an almost completely deprotonated form [99]. The difference between the c structures at pH 5 and 8 is that the carboxyl terminal a helix was rotated by 140◦ with respect to the amino terminal helix. From the two structures and the results of cross-linking experiments, Fillingame et al. suggested that the carboxyl terminal a helix rotates during proton transport, interacts with aArg210, and deprotonates cAsp61 [19]. This c subunit structural change possibly drives stepwise rotation of the c ring.

4.2 Rotational Catalysis of the F-ATPase Holoenzyme

To complete ATP hydrolysis-dependent proton transport, the γ rotation should be transmitted to the Fo membrane sector. Conversely, for ATP synthesis, proton transport through Fo should generate torque to drive the γ subunit rotation coupled to chemistry. The γ rotation should be coupled to protonation–deprotonation of cAsp61 in either direction. Depending on the modeling of Fo (a and b subunit inside the c ring), rotation of a, b, γ , δ and ε relative to the complex of α, β and the c ring once has been suggested [83]. This speculation led to experimental tests, although an alternative model in which the c subunit ring attached by the a and b subunit helices is supported experimentally, as discussed above. Different mechanisms could be proposed for the coupling between γ rotation within the α 3 β 3 hexamer and proton transport through the Fo sector: (a) the γ subunit rotates on the surface of the c ring; and (b) the γ subunit and the c ring rotate as a single complex. Models should be consistent with the proton transport continuously utilizing one aArg210 and 10–14 cAsp61 residues of the a subunit and the c ring, respectively. Consistent with mechanism (b), energy transduction of F-ATPase with a counter-rotating rotor (γ ε c ring) and stator has been proposed [71, 100]. Convincing evidence supporting this mechanism includes the results of chemical cross-linking. Cross-linking between the γ and c subunits did not affect F-ATPase activity, indicating that sequential interaction of the rotating γ subunits with multiple c subunits is not necessary during ATP synthesis-hydrolysis [101]. Similarly, cross-linking between the γ and ε subunits did not affect ATP hydrolysis [102], supporting the rotation of an actin filament connected to either subunit. However, α–γ , α–ε, β–γ and β–ε cross-linking resulted in loss of the activity [20, 103], confirming that ε γ rotates against α 3 β 3 . These results suggest that a complex formed from γ , ε and c subunits is a mechanical unit for relative rotation to the α 3 β 3 hexamer.


Fig. 5 Rotational catalysis of F-ATPase. Fo F1 was immobilized through a histidinetag attached to the a subunit (a) or c ring (b, c), and a fluorescent actin filament was connected as a probe to the c (a), " (b), or

a (c) subunit to observe rotational catalysis. Continuous rotation of the probe connected to the purified Fo F1 (a or b) or membraneembedded Fo F1 (c) was observed upon addition of ATP.

We obtained direct evidence of continuous rotation of the E. coli εγ c10–14 complex during ATP hydrolysis [24] (Figure 5). F-ATPase engineered for rotation was solubilized from membranes, purified, and immobilized through histidine tags introduced into the a subunit. Upon the addition of ATP, the filament connected to the c ring rotated anticlockwise continuously, and generated similar torque to that observed for the γ rotation in the F1 sector. The rotation was inhibited by venturicidin, a specific inhibitor for F-ATPase but not for ATPase activity of the F1 sector. The rotation and ATPase activity became less sensitive to the antibiotic when the cIle38Thr mutation was introduced into the c subunit [26], indicating that antibiotic binding to the c ring inhibited rotation [24]. Pänke et al. also observed c subunit rotation during ATP hydrolysis [25]. They immobilized Fo F1 through a histidine tag introduced into the β subunit, and an actin filament was connected to the c subunit through a streptag and streptavidin. The characteristics of the rotation observed were the same as those with the above systems. These results are consistent with the recent experiments showing that complete cross-linking of the γ , ε and c subunit had no effect on ATP hydrolysis, proton translocation, or ATP synthesis [103]. The important conclusion drawn from these experiments is that the c subunit ring rotates when F-ATPase is immobilized through the α or β subunit. As discussed by Pänke et al. [25] and Wada et al. [104], it is not easy to prove that all the subunits were integrated into F-ATPase rotating under the microscope. However, indirect evidence supports the intactness of the enzyme used for rotation: reconstitution studies indicated that the a, b and c subunit are required to form Fo capable of F1 binding [105], and all F1 subunits are required for binding to the Fo sector [54]. Early genetic and biochemical studies led to the conclusion that all three Fo subunits are required for F1 binding (see Ref. [56] for a review). The intactness of F-ATPase in these observations was also supported by the results for the membrane-bound enzyme [27], which rotated in essentially a similar manner to the purified F-ATPase. As discussed previously [26, 104], criticism [106] regarding

13


the initial observation [24] was based mainly on the negative observation under different experimental conditions. We further addressed the basic question of whether the rotor and stator are interchangeable in F-ATPase [26] (Figure 5b). Thus, F-ATPase was immobilized on a glass surface through a histidine tag introduced into the c subunit, and an actin filament was connected to the β subunit through the biotin-binding domain of transcarboxylase (biotin-tag). We could observe ATP-dependent filament rotation generating the same frictional torque as observed above, similar to the case of F-ATPase immobilized through the α or β subunit and with an actin filament connected to the c subunit [24, 25]. Thus, either of the two subcomplexes (εγ c10–14 or α 3 β 3 δab2 ) could be a rotor or a stator. Their actual roles in vivo will depend on the viscous drag due to the cytosol and membranes. 4.3 Rotational Catalysis of F-ATPase in Membranes

Although rotation of the γ εc10–14 complex has been shown using purified F-ATPase [24–26], a more favorable experimental system is the isolated membrane because the original integrity of the enzyme is maintained. The membrane could be used to answer one of the obvious questions of whether the c ring rotates relative to the a subunit. Such rotation would support the model of proton transport through the interface of the two subunits (Figure 5c). It may be difficult to test the rotation of a probe connected to the c ring embedded in membranes by immobilizing F-ATPase through the α or β subunit. As described above, purified F-ATPase can be immobilized through a histidine-tag connected to the c ring, and an actin filament can be connected to the α or β subunit through a biotin-tag [38]. This experimental system was modified to test the rotation of F-ATPase in the membrane. As histidine and biotin-tags face the periplasm and cytoplasm of the intact E. coli cell, respectively, membrane preparation for rotation should be carefully considered. Right-side out or everted membrane vesicles can not be used for testing rotation, because only one of the two tags in these vesicles is accessible from the medium: the histidine-tag faces the medium for right-side out vesicles, but the biotin-tag is inside the same vesicles. Similarly, the histidine-tag is inside everted vesicles, and thus cannot be used for immobilizing F-ATPase in these vesicles. Therefore, F-ATPase in planar membranes should be used to test for the rotation. We prepared membrane fragments by passing E. coli cells through a French press by a slight modification of the procedure used to prepare everted vesicles [27]. A labeling experiment with streptavidine-conjugated gold particles indicated that the preparation contained a significant number of planar membranes. Using this preparation, we observed ATP-dependent rotation of an actin filament connected to the β subunit. As expected from the previous studies, the rotation was counterclockwise and sensitive to DCCD. To complete a model of proton translocation through the rotation in F-ATPase, it is essential to show rotation of the c ring relative to the a subunit (Figure 5c).


Fig. 6 Relative rotation of the a subunit in membrane-embedded F-ATPase. Time courses of rotating filaments connected to the a subunit of membrane F-ATPase immobilized through the c subunit are shown,

along with time courses of rotating filaments of varying length (a) and typical sequential video images (video interval, 100 ms) (b). Taken from Nishio et al. [27].

The c subunit (cMet65Cys) cross-linked with the a subunit (αAsn214Cys) was highly protected from labeling with 14 C-DCCD [107]. However, when F-ATPase was labeled with 14 C-DCCD and subjected to the ATPase reaction before crosslinking there was significantly increased labeling of the a–c dimer. This experiment supports the rotational catalysis in Fo , although no information on the mechanism could be obtained. We could show rotation of the actin filament connected to the a subunit relative to the c ring using membrane fragments [27] (Figure 6). These results and those obtained with purified Fo F1 or the F1 sector indicate that γ εc10–14 and α 3 β 3 δab2 are mechanical units, i.e., an interchangeable rotor and stator, respectively. Notably, the results obtained with membrane-embedded F-ATPase and those with the purified one are similar. Furthermore, the experimental system will be extremely valuable for studying the mechanism of coupling of rotation and proton transport, although further modification(s) may be necessary. These studies established rotational catalysis by F-ATPase (Figure 7). For ATP hydrolysis, the F-ATPase is a chemically driven motor rotating γ εc10–14 to drive proton transport. Studies on the mechanism of the rotation in the Fo sector were started only recently. The rotation is inhibited by venturicidin or DCCD, suggesting that the tight or covalent binding of these bulky chemicals to the c subunit inhibits mechanical rotation [13]. Junge and coworkers showed that the c subunit ring with the cAsp61Asn mutation can still rotate during ATP hydrolysis, indicating that the proton transport is not obligatory for the chemically driven rotation (ATP hydrolysis-dependent c ring rotation) [108]. For ATP synthesis, the same enzyme is a potential-driven motor rotating γ εc10–14 through an electrochemical proton gradient. This membrane system will be useful for studying the rotation in this direction.

15


Fig. 7 Schematic mechanism of rotational catalysis by F-ATPase. For ATP synthesis, electrochemical proton transport changes the c subunit conformation that drives the rotational movement of the c ring together with the ( and g subunits. Conversely, ATP

hydrolysis drives the ( and g subunit rotation together with the c ring to transport protons. Modified from Oster [100], Fillingame [19, 20], Junge [71] and their coworkers.

5 Rotational catalysis of V-ATPase 5.1 Catalytic Site and Proton Pathway

V-ATPase (vacuolar-type ATPase) acidifies the lumens of endomembrane organelles such as lysosomes, endosomes and synaptic vesicles (for reviews, see Refs. 32–35). The same enzyme in the plasma membranes of specialized cells, including osteoclasts and renal intercalated cells, pumps protons into extra cellular compartments such as resorption lacunae and collecting ducts, respectively. Despite significant physiological differences, V-ATPase exhibits similarities with F-ATPase. The catalytic A subunit of V-ATPase is homologous to F-ATPase β. The homology is striking in the phosphate binding P-loop and other sequences, including the F-ATPase catalytic residues discussed above [29]. E. coli F-ATPase residues at the catalytic site, βLys155, βThr156, βGlu181, βArg182 and βGlu185, correspond to ALys263, AThr264, AGlu266, AArg267 and AGlu290 of the yeast V-ATPase A subunit, respectively (Figure 8). Mutational studies of yeast V-ATPase

5 Rotational catalysis of V-ATPase

Fig. 8 Similarities of the V-ATPase G subunit with the F-ATPase b or * subunit. The subunits are aligned to obtain maximal homology. Identical amino acid residues are boxed.

supported the notion that they are catalytic residues [109, 110]. The kinetics indicate that V-ATPase has three catalytic sites exhibiting cooperativity [111]. The cysteine residue in the V-ATPase P-loop may be involved in regulation [112]. F-ATPase with Cys introduced at the corresponding position became sensitive to sulfhydryl reagents, similar to V-ATPase [113]. The membrane Vo sector of V-ATPase has a more complicated subunit composition: yeast Vo is formed from c, c , c , a and d subunits [28]. Of the three proteolipid subunits, the c and c having four transmembrane helices are a duplicated form of the F-ATPase c subunit, and show 56% identity in amino acid residues with each other. The carboxyl moieties (cGlu137 and c Glu145) for proton transport are located on the fourth helix of the c and c subunits, respectively [114, 115]. The c subunit, exhibiting some homology with c and c , is also required in proton transport: c γ Lu188 in the middle of the third a helix is also implicated for proton transport [114]. The c subunit is conserved in mammalians [116, 117], whereas the c subunit is only found in yeast. The stoichiometry of the c:c :c subunits in yeast is n:1:1 [118]. The three proteolipids may form a ring structure similar to the Fo c ring. Assuming that 4, 1, and 1 molecules of the c, c and c subunits [114], respectively, form a ring, Vo has ∼6 proton translocating residues altogether. This is in contrast with 10–14 residues for Fo . Mutation of V-ATPase aArg735 abolished proton translocation similar to that of F-ATPase aArg210 [119], although the subunits a of Vo and Fo exhibit little homology. Thus, Vo has multiple carboxyl moieties and one arginine essential for proton translocation. The similarities between the two ATPases raised the interesting question of whether V-ATPase can synthesize ATP. Unlike F-ATPase localized with the respiratory chain in the mitochondrial membrane, mammalian or yeast membranes

17


containing V-ATPase do not have a system for generating an electrochemical proton gradient. Does V-ATPase synthesize ATP when a membrane potential or protongradient is generated? To answer this question, we expressed a plant proton translocating pyrophosphatase in yeast vacuoles [120]. Upon hydrolysis of pyrophosphate, the vacuolar membrane vesicles could form an electrochemical proton gradient, which was lowered by the addition of ADP + Pi. As expected from this result, we observed ATP synthesis sensitive to the V-ATPase specific inhibitor bafilomycin. The results further support the similarities between the two ATPases. Plant vacuoles having both V-ATPase and pyrophosphatese may synthesize ATP similar to the hybrid system studied in yeast.

5.2 Subunit Rotation of V-ATPase During Catalysis

Despite the similarities of V- and F-ATPase discussed above, they are also significantly different. Typically, F-ATPase of E. coli has eight subunits, whereas yeast V-ATPase has 13. Most of the subunits unique to V-ATPase are located in the stalk region. Furthermore, the presence of isoforms of subunits B, E, G, d, and a has been found for V-ATPase, whereas the information on isoforms is limited for F-ATPase. As pointed out above, Vo , possibly its ring structure, may be different from Fo . These differences and similarities led us to examine the rotational catalysis in V-ATPase. As described above, rotation of an actin filament connected to the a, α, or β subunit was observed when F-ATPase was immobilized through the c subunit ring. The correspondence of minor subunits in the stalk region between the two ATPases was difficult to determine (Figure 8). The G subunit was shown to exist in the stalk domain of a recent model, and to be accessible from the cytosol [30]. The G subunit and F-ATPase b exhibit ≈24% homology, although the G subunit lacks a transmembrane region. However, recent cross-linking studies suggested that the G subunit is located near the top of V1 [121, 122], similar to the F-ATPase δ subunit [89, 90]. Thus, the G subunit may correspond to the b or δ subunit of F1 , and could be a candidate for connecting a probe to observe rotation, although the homology is quite limited. Based on these considerations, we engineered the yeast chromosome (VMA3 and VMA10 for the c and G subunit, respectively), solubilized V-ATPase from vacuolar membranes, and tested rotational catalysis. An actin filament connected to the G subunit rotated upon ATP hydrolysis [123] (Figure 9). The rotation was inhibited by nitrate and concanamycin, similar to their effects on ATPase activity. Concanamycin inhibition was striking: the rotation terminated within a few seconds after the addition Figure 9, which is consistent with its tight affinity [111]. This antibiotic is similar to bafilomycin, which binds to the Vo sector, possibly to its c subunit [124], indicating that concanamycin, possibly bound to the rotor–stator interface, terminated the rotation. Torque generated by the rotation was similar to that with F-ATPase. We observed rotation in a sonicated vacuolar membrane

6 Conclusion

Fig. 9 Rotation of an actin filament connected to the g subunit of V-ATPase. Solubilized V-ATPase was immobilized on a glass surface through the c subunit, and an actin filament was connected to the G subunit. Rotation was observed upon the addition of ATP, and inhibited by nitrate and concanamycin. Taken from Hirata et al. [123].

preparation; however, we could not obtain enough data for analysis (unpublished observation). These results suggest that the A3 B3 hexamer rotates together with the G subunit, when the c subunit ring is immobilized. An interesting question is “Which part of the V-ATPase rotates in vivo?” In this regard, Holiday and coworkers reported that the B subunit interacts with the cytoskeleton, possibly actin [125]. Thus, the c subunit ring together with the subunit corresponding to the γ (possibly D) subunit may be rotating in vivo, if the V-ATPase holo enzyme is immobilized with the cytoskeleton. However, it is also possible that the rotor–stator is determined only by the slight difference in viscous drag applied to each sub-complex. Rotational catalysis of the peripheral membrane sector of T. thermophilus ATPase was shown recently [126]. This Archaean ATPase is more similar to F-ATPase than V-ATPase, as pointed out by the authors. In this regard, the ATPase of Archae (Archaean bacterial) plasma membranes has been classified as an A-type ATPase [127]. Thus, three distantly related proton translocating ATPases carry out common rotational catalysis.

6 Conclusion

F-ATPase has been a fascinating membrane enzyme for generations of biochemists. As Paul Boyer called it “A splendid molecular machine” [23], it is more than an ordinary enzyme. F-ATPase synthesizes or hydrolyzes ATP coupling between

19


proton translocation and chemistry, and has become a real molecular machine in this sense, since mechanical rotation of its subunit complex was included recently in its mechanism. For the engineering aspect of the mechanism, the F1 sector immobilized on a metal surface can rotate a metal plate [128]. As described above, experimental systems have been established for further studies. Details of the rotation mechanism will be revealed by studies involving E. coli F-ATPase together with the progress regarding physical methods and the higher-ordered X-ray structure determinations of the Fo sector. It was interesting to learn that rotational catalysis has been expanded to V-ATPase and also distantly related Archaean ATPase. The basic mechanism of V-ATPase rotation may be similar to that of F-ATPase, as expected from the similarities between the two enzymes. Thus, the rotational mechanism of E. coli F-ATPase should be studied extensively, using its genetic and biochemical advantages. VATPase may have a more fascinating regulatory mechanism that is supported by a unique Vo structure and a series of isoforms in the stalk region. The regulatory role of stalk subunit(s) including subunit E in energy coupling has already been discussed [129]. Finally, owing to limited space, the present article is not a comprehensive survey, and so the reader is directed to the reviews listed above for references not cited here.

Acknowledgments

We wish to thank our coworkers, whose names appear in the references, from our laboratory. We are grateful to Ms S. Shimamura and M. Nakashima for preparation of the manuscript. The research in our own laboratories was supported by Grantsin-Aid from the Ministry of Education, Science, and Culture of Japan, the Takeda Foundation, and the Japan Science and Technology Corporation.

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126 H. IMAMURA, M. NAKANO, H. NOJI, E. MUREYUKI, S. OHKUMA, M. YOSHIDA, and K. YOKOYAMA, Proc. Natl. Acad. Sci. U. S. A., 2003, 100, 2312–2315. 127 K. IHARA, T. ABE, K. I. SUGIMURA, and Y. MUKOHATA, J. Exp. Biol., 1992, 172, 475–485. 128 R. K. SOONG, G. D. BACHAND, H. P. NEVES, A. G. OLKHOVETS, H. G. CRAIGHEAD, and C. D. MONTEMAGNO, Science, 2000, 290, 1555–1558. 129 G.-H. SUN-WADA, Y. IMAI-SENGA, A. YAMAMOTO, Y. MURATA, T. HIRATA, Y. WADA, and M. FUTAI, J. Biol. Chem., 2002, 277, 18098–18105. 130 R. H. FILLINGAME, Curr. Opin. Struct. Biol., 1996, 6, 491–498.

1

Vacuolar H+ -ATPases: Structure, Mechanism and Regulation Elim Shao and Michael Forgac Tufts University School of Medicine, Boston, USA

Originally published in: Handbook of ATPases. Edited by Masamitsu Futai, Yoh Wada and Jack H. Kaplan. Copyright ľ 2004 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30689-3

1 Introduction

Vacuolar (H+ )-ATPases (or V-ATPases) are ATP-dependent proton pumps that acidify intracellular compartments and, in certain cases, transport protons across the plasma membrane of eukaryotic cells. Acidification of intracellular compartments functions in such processes as receptor-mediated endocytosis, intracellular targeting of lysosomal enzymes, protein processing and degradation, and coupled transport. Plasma membrane V-ATPases are involved in acid–base balance, bone resorption, potassium secretion and tumor metastasis. This chapter will focus on work from our laboratory on the structure, mechanism and regulation of the V-ATPases from clathrin-coated vesicles and yeast. V-ATPases are composed of a peripheral V1 domain responsible for ATP hydrolysis and an integral V0 domain responsible for proton transport. V1 has a molecular mass of 640 kDa and is composed of eight different subunits (subunits A–H) of molecular mass 70–13 kDa, whereas V0 has a molecular mass of 260 kDa and is composed of five different subunits (subunits a, d, c, c and c ) of molecular mass 100–17 kDa. Conventional and cysteine-mediated cross-linking as well as electron microscopy have allowed us to define the overall shape of the V-ATPase and the location of subunits within the complex. Site-directed and random mutagenesis together with chemical modification have been used to identify residues involved in nucleotide binding and hydrolysis, proton translocation and the coupling of these two processes. We have also obtained information about the mechanism of intracellular targeting of V-ATPases and regulation of V-ATPase activity by reversible dissociation.


2 Vacuolar H+ -ATPases: Structure, Mechanism and Regulation

2 Function of V-ATPases

V-ATPases are ATP-dependent proton pumps responsible for acidification of intracellular compartments and, in specialized cell types, proton secretion across the plasma membrane (for reviews see Refs. [1–4]). V-ATPases are present in such intracellular compartments as clathrin-coated vesicles, endosomes, lysosomes, Golgi-derived vesicles, secretory vesicles and the central vacuoles of fungi and plants. A major role of V-ATPases within intracellular compartments is in various membrane traffic processes [1]. V-ATPases within early endosomes create the low pH necessary for ligand–receptor dissociation and recycling of receptors to the cell surface and for the formation of endosomal carrier vesicles that move dissociated ligands along the endocytic pathway. Several viruses (such as influenza virus) and toxins (such as diphtheria toxin) gain access to the cytoplasm of target cells via acidic endosomal compartments. Acidification of late endosomes is required for targeting of newly synthesized lysosomal enzymes from the Golgi to lysosomes. In addition to their role in membrane traffic, V-ATPases also provide the driving force for various coupled transport processes. For example, the proton gradient and membrane potential generated by the V-ATPases drives uptake of neurotransmitters into synaptic vesicles. Finally, V-ATPases in lysosomes and the central vacuoles of fungi and plants provide the acidic environment required for macromolecular degradation as well as the driving force for coupled transport. Plasma membrane V-ATPases have been identified in various cell types, including renal intercalated cells, osteoclasts, macrophages, insect goblet cells and tumor cells, where they function in such processes as renal acidification, bone resorption, pH homeostasis, coupled potassium transport and tumor invasion [1, 4].

3 Overall Structure of V-ATPases and Relationship to F-ATPases

Our current model of the structure of the V-ATPases is shown in Figure 1, and the subunit composition, including the molecular mass of the subunits, the genes encoding these subunits in yeast and information about subunit function, is summarized in Table 1. V-ATPases are composed of a peripheral V1 domain responsible for ATP hydrolysis and an integral V0 domain responsible for proton translocation [1–4]. V1 is a 640 kDa complex that contains eight different subunits (A–H) of molecular mass 70–13 kDa that are present in a stoichiometry of A3 B3 C1 D1 E1 F1 G2 H1–2 [5, 6]. The nucleotide-binding sites are located on the 70 kDa A subunit and the 60 kDa B subunit, with the catalytic sites located on the three A subunits and socalled “non-catalytic” sites located on the three B subunits [7–9]. V 0 is a 260 kDa complex [10] that contains five different subunits (a, d, c, c and c ) of molecular mass 100–17 kDa in a stoichiometry a1 d1 c 4–5 c 1 c 1 [5, 11]. Buried charged residues essential for proton transport are present in both the proteolipid subunits (c, c and c ) and the 100 kDa a subunit [12, 13].

3 Overall Structure of V-ATPases and Relationship to F-ATPases

Fig. 1 Structural model of the V-ATPase complex. The V1 domain (shown in white) is responsible for ATP hydrolysis whereas the V0 domain (shaded) is responsible for proton translocation. ATP hydrolysis at the catalytic nucleotide-binding sites (located on the A subunits) is proposed to drive rotation of a central stalk (composed of the D and F subunits) which in turn drives rotation of the ring of proteolipid subunits (c, c , c ) relative to subunit a. Subunit a

is held fixed relative to the A3 B3 head by a peripheral stalk composed of subunits C, E, G, H and the soluble domain of subunit a. Movement of the ring of proteolipid subunits relative to subunit a is postulated to drive unidirectional proton transport across the membrane. Reprinted with permission

from Ref. [44], copyright 2002, the American Society for Biochemistry and Molecular Biology.

V-ATPases are structurally similar to the F-ATPases that functon in the synthesis of ATP in mitochondria, chloroplasts and bacteria [14–16]. In both cases, the peripheral domain is responsible for nucleotide binding and hydrolysis (or, in the case of the F-ATPases, synthesis), whereas the integral domain is responsible for transport of protons across the membrane. Sequence homology has been identified between the nucleotide-binding subunits of the V and F-ATPases as well as between the proteolipid subunits of these two classes [17–19], suggesting that they have evolved from a common evolutionary ancestor. X-Ray structures have been obtained for F1 from several sources and they reveal a hexameric arrangement of alternating nucleotide-binding subunits (α and β) surrounding a central cavity occupied by a highly α-helical γ subunit, which extends towards the membrane [20, 21]. More recently, an X-ray structure of F1 attached to a ring of proteolipid subunits has been obtained from yeast mitochondria that reveals the γ and ε subunits form a central stalk connecting F1 to a ring of 10 c subunits [22]. F1 and F0 have also been shown to be connected by a peripheral stalk containing the δ subunit and the soluble domains of subunit b [23, 24].

3

4 Vacuolar H+ -ATPases: Structure, Mechanism and Regulation Table 1 Subunit composition of the yeast V-ATPase

V1

V0

Subunit Yeast Gene

Mr (kDa)

Function/location

A B C

VMA1 VMA2 VMA5

69 58 44

D E F G H a

VMA8 VMA4 VMA7 VMA10 VMA13 VPH1

29 26 14 13 54 100

STV1

100

d c

VMA6 VMA3

40 17

c

VMA11

17

c

VMA16

23

Catalytic site, homolog of β of F1 F0 -ATPase Non-catalytic site, homolog of α of F1 F0 -ATPase Peripheral stalk, released from V1 complex during dissociation Central stalk, homolog of γ of F1 F0 -ATPase Peripheral stalk Central stalk Peripheral stalk Peripheral stalk Proton translocation, targeting to vacuole, homolog of a of F1 F0 -ATPase Proton translocation, targeting to late Golgi, homolog of a of F1 F0 -ATPase Cytoplasmic side, non-transmembrane protein Proton translocation, DCCD- and concanamycin-binding sites, homolog of c of F1 F0 -ATPase Proton translocation, homolog of c of F1 F0 -ATPase Proton translocation, homolog of c of F1 F0 -ATPase

The presence of multiple stalks connecting the peripheral and integral domains has been demonstrated for both the V and F-ATPases by electron microscopy [25–27]. The V-ATPase structure, however, appears more complex than the F-ATPase, with many projections emerging from the V1 domain and a knob present on the lumenal side of the membrane [26]. EM images of theV0 domain suggest a ring (presumably of proteolipid subunits) adjacent to a membrane embedded mass that probably corresponds to subunit a [28]. This chapter focuses on our work on the structure and function of individual subunits, their arrangement in the V-ATPase complex, the mechanism of ATP-driven proton transport, the mechanism of targeting of V-ATPases to different intracellular compartments and the mechanisms employed in regulating V-ATPase activity in vivo.

4 Structure and Function of the Nucleotide-binding Subunits

Results from several studies suggest that the catalytic nucleotide-binding sites are located on the 70 kDa A subunits. Modification of the bovine A subunit by sulfhydryl reagents (such as NEM and NBD-Cl) leads to ATP-protectable inhibition of V-ATPase activity [29]. The cysteine residue whose modification is responsible for this inhibition was identified as Cys254 in the bovine A subunit [7]. This cysteine residue is located in a consensus sequence termed the Walker A sequence or

4 Structure and Function of the Nucleotide-binding Subunits

P-loop (corresponding to GXGKTV). The X-ray structure of F1 reveals that this sequence is located at the catalytic nucleotide-binding site on the β subunit, where it surrounds the terminal phosphate groups of ATP [20, 21]. Interestingly, formation of a disulfide bond between Cys254 and Cys532 (located in the C-terminal domain of the A subunit) causes reversible inhibition of V-ATPase activity [30]. It has been proposed that disulfide bond formation between Cys254 and Cys532 leads to inhibition of ATPase activity by preventing this subunit from adopting an open conformation that it must go through during the catalytic cycle [31]. Reversible disulfide bond formation between these cysteine residues in the A subunit has also been proposed to represent an important mechanism of regulating V-ATPase activity in vivo [32]. Studies using the photoactivated nucleotide analog 2-azido-ATP also support the idea that the catalytic nucleotide-binding sites are located on the A subunits [8]. This reagent inhibits V-ATPase activity in an ATP-protectable manner, with complete inhibition of ATPase activity occurring after modification of only one of the three A subunit sites per complex. Further support for the location of the catalytic sites on the A subunit comes from mutagenesis studies. Using molecular modeling based on the F-ATPase structure and the homology between the V and F-ATPase nucleotide-binding subunits, A subunit residues were identified that were predicted to play a role in either ATP binding or hydrolysis at the catalytic sites [33]. Mutagenesis of K263 in the Walker A sequence of the A subunit completely inhibited ATP hydrolysis, as did mutagenesis of E286 [34]. K263 is proposed to stabilize the negatively charged phosphate groups of bound ATP whereas E286 is suggested to serve as a proton acceptor from water during ATP hydrolysis. In addition, mutagenesis studies coupled with photochemical modification by 2-azidoATP have identified four aromatic residues that are thought to form the adenine binding pocket of the catalytic site on the A subunit [33, 35]. More recently, evidence has been obtained for an important role of a novel domain of the V-ATPase A subunit, termed the “non-homologous” region, in coupling of ATP hydrolysis and proton transport and reversible dissociation of the V-ATPase complex [36]. The non-homologous region is a 90 amino acid stretch which is present and conserved in all V-ATPase A subunit sequences but which is absent from the corresponding F-ATPase β subunit [17, 37]. This region is located approximately one-third of the way from the amino terminus of the A subunit and, because the A3 B3 structure (by analogy with F1 ) is predicted to be tightly packed, is likely to be located on the outer surface of the complex. A peripheral location of this domain suggests the possibility that it may contribute to the peripheral stalk connecting the V1 and V0 domains [26]. Consistent with this idea, mutations have been identified in this region that lead to significant changes in coupling of proton transport and ATP hydrolysis [36]. Interestingly, one mutation (P217V) dramatically increased the coupling efficiency, suggesting that the wild-type enzyme is not normally optimally coupled. Several mutations were also identified in the non-homologous region that block reversible dissociation of the V-ATPase complex in response to glucose depletion [36], suggesting that this domain may also play a role in regulating V-ATPase activity in vivo (see below).

5


Several lines of evidence suggest that, in addition to subunit A, the 60 kDa B subunit also participates in nucleotide binding. First, subunit B is selectively modified by the photoactivated nucleotide analog BzATP [9]. Second, the B subunit is homologous to the α and β subunits of F1 and to subunit A of the V-ATPases (approximately 20–25 % amino acid identity), although subunit B lacks the consensus Walker A and B motifs that are often present in nucleotide-binding proteins [18, 38]. Despite the absence of these motifs, the sequences that replace them in the B subunit are highly conserved among species. Modeling studies of the B subunit based upon sequence homology with the α subunit, the X-ray coordinates of F1 and energy minimization have been used to predict a structure for this subunit. This model accurately predicts the presence of a number of residues at the nucleotide-binding site on B, based upon nucleotide-protectable chemical modification of mutant proteins [39]. Unique cysteine residues were introduced into a cys-less form of subunit B and their modification by the sulfhydryl reagent biotin maleimide was tested in the intact V-ATPase in the presence and absence of BzATP. All six cysteine residues which reacted with biotin maleimide displayed protection from labeling by BzATP, which is consistent with their presence at the nucleotide-binding site [39]. Although the function of the “non-catalytic” nucleotide-binding sites on the B subunit is not certain, at least one mutation at this site causes a time-dependent change in ATPase activity, suggesting that they may play a role in regulating activity [33]. Other mutations at these sites appear to have less dramatic effects on activity [40]. As with the F-ATPases [20], the nucleotide-binding sites (both catalytic and non-catalytic) appear to be located at the interface of the alternating A and B (or β and α) subunits [33, 40], although, as indicated above, most residues at the catalytic sites are contributed by the A subunits whereas most residues at the non-catalytic sites are contributed by the B subunits.

5 Structure and Function of other V1 Subunits and Arrangement of Subunits in the V-ATPase Complex

Studies from a number of laboratories have begun to shed light on the function of other subunits in the V1 domain. Mutants have been identified in subunit D (product of the VMA8 gene) that suggest it plays an important role in coupling of proton transport and ATP hydrolysis [41]. The most dramatic effects on coupling are observed for double mutants in which changes near the amino and carboxyl-terminus appear to act synergistically to cause uncoupling. One possible explanation for these results is that the N and C-terminus of the protein interact. Together with the predicted high α-helical content of subunit D, this has led us to postulate that subunit D acts as the γ subunit homolog in the V-ATPases [41]. Mutations causing uncoupling of the F-ATPase have also been identified in the γ subunit of F1 [42], which exists as a coiled-coil structure within the central stalk connecting F1 and F0 [20, 21].

5 Structure and Function of other V1 Subunits and Arrangement of Subunits in the V-ATPase Complex 7

Support for the hypothesis that subunit D is the γ subunit homolog in the VATPases comes from cysteine mutagenesis and covalent cross-linking studies using the bifunctional, photoactivated maleimide reagent maleimido-benzophenone. As indicated above, a molecular model of the A3 B3 hexamer has been derived using the X-ray coordinates of F1 , the sequence homology between the nucleotide-binding subunits of the V and F-ATPases and energy minimization [35, 39]. This model was used to predict the location of unique cysteine residues introduced into a cys-less form of subunit B by site-directed mutagenesis. These unique cysteine residues were first modified using maleimido-benzophenone followed by photoactivated cross-linking. Cross-linked products were then identified by Western blot analysis using subunit-specific antibodies. Results from these studies indicate that subunits B and D are cross-linked only at sites on the B subunit predicted to be oriented towards the interior of the A3 B3 hexamer [43]. By contrast, subunits E and G are cross-linked to subunit B at sites predicted to be facing the exterior of the complex [43, 44]. These results suggest that subunit D is located in the central stalk connecting V1 and V0 whereas subunits E and G form part of a peripheral connection between these domains. Conventional cross-linking studies performed on the coated vesicle V-ATPase using a number of bifunctional cross-linking reagents have demonstrated that subunit D makes contact with subunit F whereas subunit E is in contact with a significant number of subunits, including subunits C, G, H and the soluble domain of subunit a [6]. Disruption of V-ATPase assembly in yeast strains lacking certain V1 subunits gives rise to several subcomplexes, including the heterodimeric species DF and EG [45], providing further support for the structural model shown in Figure 1. Dissociation of the bovine coated vesicle V-ATPase also gives rise to a heterodimeric EC subcomplex, as detected by immunoprecipitation [46]. These results suggest that subunits D and F form a central stalk connecting V1 and V0 whereas subunits C, E, G, H and the soluble domain of subunit a form the peripheral stalk connecting these domains. Considerable information about the function of subunits C, G and H has been obtained from studies performed in other laboratories involving mutagenesis and overexpression of these subunits in yeast. Thus, subunit C appears to activate ATP hydrolysis by V1 whereas subunit H inhibits this activity [47, 48]. Subunit G, which shows some homology to subunit b of the F-ATPases along one helical face [49], has been shown to tolerate short deletions in its sequence without loss of function [50]. This result is similar to those obtained for the F-ATPase b subunit [51], and suggests that subunit G may be the b subunit homolog of the V-ATPases. An X-ray structure of subunit H has recently been obtained [52] and has revealed interesting structural similarities to the importins, proteins involved in nuclear import. Yeast 2-hybrid analysis and co-immunoprecipitation studies have revealed that subunit H makes contact with both subunit A in V1 and the soluble, cytoplasmic domain of subunit a in V0 [53], suggesting that it serves an important bridging function. Moreover, subunit H has been shown to serve an important role as the docking site for the NEF protein, which in turn functions in internalization of the HIV receptor CD4 [54].


6 Structure and Function of V0 Subunits

The V0 domain of the V-ATPases contains five subunits. Three of these correspond to highly hydrophobic proteins termed proteolipids because of their ability to be extracted with organic solvents: subunit c (Vma3p), subunit c (Vma11p) and subunit c (Vma16p). All three of these proteins are required for proton transport by the V-ATPases [12]. Subunit c and c each contain four transmembrane helices with an essential buried glutamate residue present in TM4. Although subunit c was originally predicted to contain five transmembrane helices, recent studies have demonstrated that it instead contains only four, with the critical glutamic acid residue present in TM2 [55]. Interestingly, the region originally predicted to correspond to TM1 has been shown not to be required for function. Topological studies employing expression of epitope-tagged proteins as well as differential modification of unique cysteine residues indicate that the C-terminus of subunit c is present on the lumenal side of the membrane whereas the C-terminus of subunit c is exposed on the cytoplasmic side of the membrane [55, 56]. The buried glutamic acid residues present in the proteolipid subunits are essential for proton transport by both chemical modification by DCCD [31] and site-directed mutagenesis [12]. These residues have therefore been proposed to participate directly in proton translocation. Interestingly, subunit c forms part of the binding site for the specific V-ATPase inhibitor bafilomycin [57]. The three V-ATPase proteolipid subunits are homologous to the F-ATPase subunit c, which is composed of just two transmembrane helices that adopt a helical hairpin structure [58]. The V-ATPase proteolipids appear to have been derived during evolution from a gene duplication and fusion event from an ancestral gene that more closely resembles the F-ATPase c subunit [19]. NMR analysis of the F-ATPase proteolipid suggests that the orientation of the helix containing the essential acidic residue changes upon a change in protonation state [59], leading to the suggestion that this helical swiveling is an important part of the mechanism of proton translocation (see below). The F0 domain from E. coli and yeast mitochondria contain 10 copies of subunit c, which form a ring [22, 60]. Stoichiometry measurements indicate that the V0 domain contains 5–6 copies of subunits c plus c and a single copy of subunit c [5], and epitope-tagging experiments indicate the presence of single copies of both subunits c and c [11]. Thus, these results suggest a subunit stoichiometry for the V0 domain of c4–5 c 1 c 1 . In addition to the proteolipid subunits, the V0 domain contains two additional subunits. Subunit d (Vma6p) is a 40 kDa hydrophilic polypeptide which appears to lack transmembrane segments [61], but which remains tightly bound to V0 upon dissociation of V1 [62]. The fifth V0 subunit is a 100 kDa integral membrane protein called subunit a. In yeast, subunit a is encoded by two genes (VPH1 and STV1), with Vph1p associated with V-ATPase complexes targeted to the vacuole and Stv1p associated with V-ATPases targeted to the late Golgi [63–65]. In mammals, subunit a is encoded by four genes (a1–a4), which are expressed in a tissue specific manner [66–70]. The a3 isoform appears to be responsible for plasma membrane targeting

7 Mechanism of ATP-dependent Proton Transport

of the V-ATPase in osteoclasts [68], and a defect in this gene causes the genetic defect autosomal recessive osteopetrosis [69]. The a4 isoform is responsible for targeting of the V-ATPase to the plasma membrane of renal intercalated cells [67], and disruption of this gene causes the disease renal tubule acidosis [70]. The topology of subunit a has been studied by introduction of unique cysteine residues into a cys-less form of Vph1p followed by evaluation of the exposure of these sites in intact vacuolar membranes using differential reactivity towards membrane permeant and impermeant sulfhydryl reagents [71]. These studies indicate that subunit a contains an amino terminal hydrophilic domain of about 50 kDa oriented towards the cytoplasmic side of the membrane and a carboxyl terminal hydrophobic domain containing nine transmembrane segments, with the C-terminus located on the lumenal side of the membrane. Site-directed mutagenesis of Vph1p has identified a buried, charged residue present in TM7 (Arg735) that is absolutely essential for proton translocation [13]. In addition, there are a series of other buried charged residues present in TM7 and TM9, including E789, H743 and R799, whose mutation causes a reduction in proton transport, suggesting that these residues may also play some role in this process [13, 72, 73]. Although no sequence homology exists between the a subunits of V0 and F0 , we have suggested that the V0 subunit a plays a similar role in proton transport. For subunit a of F0 , Arg210 located in TM4 is critical for proton transport [74]. It is postulated to directly interact with the buried carboxyl groups of the proteolipid c subunits during proton trans-location [75, 76]. The F-ATPase a subunit also contains other buried charged residues that are postulated to form aqueous access channels that allow protons to reach and leave these buried sites [74, 75]. V-ATPase complexes containing different isoforms of subunit a differ in other properties besides intracellular localization [77]. Thus Vph1p-containing complexes show a ca. 10-fold greater assembly of V1 and V0 than Stv1p-containing complexes. In addition, Vph1p-containing complexes are more tightly coupled than those containing Stv1p. These differences may reflect the need to maintain a lower lumenal pH in the vacuole (where Vph1p-containing complexes reside) than in the Golgi (where Stv1p-containing complexes are found). Using chimeric constructs, we have shown that whereas intracellular targeting is controlled by the amino terminal domain, the degree of assembly of V1 and V0 and the tightness of coupling of proton transport and ATP hydrolysis are controlled by the carboxyl-terminal [65].

7 Mechanism of ATP-dependent Proton Transport

Because of their structural similarity, the V and F-ATPases are believed to operate by a similar rotary mechanism [78, 79]. For the F-ATPases, ATP hydrolysis by the β subunits of F1 causes rotation of a central stalk, composed of the γ and ε subunits, which in turn drives rotation of the ring of proteolipid c subunits relative to subunit a. Rotation of both the γ subunit in F1 [80–82] and the ring of c subunits in F1 F0 [83] have been demonstrated. The a subunit is postulated to provide access channels

9


for the protons to reach and leave the buried carboxyl groups on the c subunit ring and to activate release of protons into the egress channel through interaction with a critical arginine residue [16, 74, 75]. Rotation of the c subunit ring relative to subunit a thus causes unidirectional proton transport across the membrane. For the V-ATPases, we suggest that ATP hydrolysis in V1 drives rotation of the D and F subunits together with the proteolipid ring relative to subunit a. Rotation of the D and F subunits of the V1 domain of the V-ATPase from Thermos thermophilus was recently demonstrated [84], which is consistent with the placement of these subunits in the central stalk [43]. Interaction of the proteolipid carboxyl groups with Arg735 on subunit a [13] would then lead to release of protons into the egress channel leading to the lumenal side of the membrane. Further rotation of the c subunit ring would bring these deprotonated carboxyl groups into contact with the cytoplasmic aqueous channel. They would need to pick up protons from this channel before re-entering the hydrophobic phase of the bilayer.

8 Regulation of V-ATPase Activity In Vivo

Control of V-ATPase activity in cells and tissues is critical for the diversity of functions served by these enzymes. Several mechanisms have been proposed to be involved in regulation of V-ATPase activity in vivo. As described above, reversible disulfide bond formation between conserved cysteine residues at the catalytic sites on the A subunit causes reversible inhibition of V-ATPase activity [30, 32], and several studies suggest that this represents an important in vivo mechanism for controlling activity [32, 85]. Differences in coupling efficiency have been demonstrated for V-ATPases containing different a subunit isoforms [77], and mutations in a number of V-ATPase subunits, including subunit D [41], subunit a [13] and the non-homologous domain of subunit A [36], cause changes in coupling efficiency. Moreover, coupling of proton transport and ATPase activity is readily perturbed [86, 87], suggesting that the enzyme may be poised to change the tightness of coupling in vivo. In addition to the mechanisms described above, reversible dissociation of the V-ATPase into its constituent V1 and V0 domains represents an important in vivo regulatory mechanism [88, 89]. In yeast, reversible dissociation occurs in response to glucose depletion [88], although many of the signal transduction pathways activated by glucose withdrawal do not appear to be involved in this process [90]. Glucose-dependent dissociation requires a catalytically active enzyme [35, 90] and an intact microtubular network [91]. Interestingly, however, reassembly of the V-ATPase is not dependent upon microtubules. Several mutations have been identified in the non-homologous region which block dissociation without inhibiting activity [36], suggesting that this domain may play a role in controlling dissociation independent of any effects on activity. Similar mutations have been identified in subunit G, where they appear to cause increased stability of the V-ATPase complex [50].

References

The in vivo dissociation behavior of V-ATPase complexes containing different isoforms of the a subunit has proven to be complex [77]. Thus, Vph1p-containing complexes localized to the vacuole undergo dissociation in response to glucose withdrawal whereas Stv1p-containing complexes localized to the Golgi do not. Re-targeting of Stv1p-containing complexes to the vacuole by overexpression of Stv1p results in a re-appearance of the ability to dissociate in response to glucose depletion. Conversely, prevention of Vph1p-containing complexes from reaching the vacuole by disruption of genes involved in intracellular targeting results in VATPase complexes that show glucose-dependent dissociation, but less completely than when these complexes have a vacuolar localization. Thus, in vivo dissociation appears to be controlled by both the a subunit isoform present in the complex and by the cellular environment in which the complex resides [77]. Recent studies have identified a novel complex (termed RAVE) which includes part of the ubiquitin ligase complex (Skp1p) and which appears to function in both normal and regulated assembly of the V-ATPase [92, 93]. Additional studies will be required to further elucidate the mechanism of regulating in vivo dissociation of the V-ATPase complex.

9 Conclusions

V-ATPases are multisubunit complexes responsible for ATP-driven proton transport in both intracellular compartments and the plasma membrane. They are composed of a peripheral V1 domain responsible for ATP hydrolysis and an integral V0 domain that transports protons. A variety of approaches have begun to elucidate the arrangement and function of subunits in the V-ATPase complex. Regulation of V-ATPase activity in vivo is likely to be complex given the great diversity of functions performed by this family of proton pumps.

Acknowledgments

The authors thank Drs Takao Inoue, Tsuyoshi Nishi and Shoko Kawasaki-Nishi for many helpful discussions. This work was supported by NIH Grant GM34478 (to M. F.).

References 1 T. NISHI and M. FORGAC, Nat. Rev. Mol. Cell. Biol., 2002, 3, 94–103. 2 H. WIECZOREK, D. BROWN, S. GRINSTEIN, J. EHRENFELD, and W. R. HARVEY, Bioeswsays, 1999, 21, 637–648. 3 L. A. GRAHAM, B. POWELL, and T. H. STEVENS, J. Exp. Biol., 2000, 203, 61–70.

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1

Anchoring Proteins of the Erythrocyte Membrane Yoshihito Yawata

Kawasaki College of Allied Health Professions, Kurashiki City, Japan

Originally published in: Cell Membrane. Yoshihito Yawata. Copyright ľ 2003 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30463-9

1 Ankyrin 1.1 Introduction

Ankyrin is one of the major proteins of human red cells, making up approximately 5% of the total membrane proteins [1–3]. Ankyrin is a large, 206 kDa sulfhydryl-rich protein with a molecular size of 8.3 × 10 nm. It is present at a level of 120 × 103 copies per cell. Functionally, ankyrin is connected with β-spectrin through a high affinity linkage (K d : ∼10−7 M), and with the cytoplasmic domain of band 3, which also has a high affinity linkage (K d : ∼10−7 to 10−8 M) [4, 5]. Ankyrin is a polar protein, and is involved in the local segregation of integral membrane proteins. The polarization of membrane proteins appears to be produced by the relative affinities of the various isoforms of ankyrins for target membrane proteins. These ankyrin isoforms appear to be expressed through tissue-specific, developmentally-regulated control [6–10], as discussed below. 1.2 Structure of Red Cell Ankyrin

Ankyrin is composed of three domains [6, 7, 11–13]: (1) a membrane (band 3)binding domain (89 kDa) at its NH2 -terminal (amino acids 2 to 827), (2) a spectrin binding domain (62 kDa) at the central part of the molecule (amino acids 828 to 1382), and (3) a regulatory domain (55 kDa) at its COOH-terminal (amino acids 1383 to 1881) (Fig. 1). The NH2 -terminal band 3-binding domain is basic, the


2 Anchoring Proteins of the Erythrocyte Membrane

Fig. 1 Structure of red cell ankyrin. Protein domain structure of erythroid ankyrin and its binding sites are shown schematically. Genetic basis of ankyrin 2.1 and 2.2 is also shown at the lower-right.

central spectrin-binding domain is neutral, but heavily phosphorylated, and the COOH-terminal regulatory domain is highly acidic. 1.2.1 Membrane (Band 3)-Binding Domain of Ankyrin This 89 kDa domain, which is at the NH2 -terminal portion of the ankyrin molecule, is almost entirely composed of 24 consecutive 33 amino acid tandem repeats (so-called “cdc 10/ankyrin repeats”, or “ankyrin repeats”), which are subdivided into six repeat folding units [12, 13]. These repeats are fairly similar to each other. Binding sites for band 3 and at least six other types of integral membrane proteins are present in this domain. Repeats 7 through 12 (folding unit 2) and especially repeat 13 through 24 (folding units 3 and 4) form two distinct but cooperating binding sites for band 3. These two binding sites of ankyrin for band 3 are able to interact with four band 3 molecules, because normally band 3 is present in a dimeric form on the membrane. This binding of ankyrin to band 3 appears to be critical for maintaining the normal integrity of membrane functions, especially membrane stability, because selective disruption of the ankyrin/band 3 interaction in intact red cells, which are placed at a slightly alkaline pH, decreases membrane stability markedly. The 15 out of 33 amino acids in the ankyrin repeats are highly conserved. Their conserved sequence is “-G-TPLH-AA–GH–V(or A)–LL–GA–N(or D)–”. L-shaped structures of the ankyrin repeats are composed of a pair of α-helices that form an antiparallel coiled-configuration, which is followed by an extended loop perpendicular to the helices and a β-hairpin [8, 14–19]. Very similar repeats have commonly been found in various proteins, in all phyla. Therefore, the ankyrin repeats appear to have been propagated as one of the versatile modules for specific ligands during evolution.

1 Ankyrin 3

1.2.2 Spectrin-Binding Domain of Ankyrin This spectrin-binding domain is also known as the central domain, and was previously known as the 62 kDa domain. The spectrin-binding sites are located in the beginning and middle regions of this domain. The site in the middle portion is highly conserved and appears to be the principal area of binding [20]. These regions are found to be those near the end of the β-spectrin molecule (repeats 15 and 16) which are involved in dimer–tetramer self-association [21]. Although two binding sites for spectrin are available in the ankyrin molecule, each spectrin tetramer appears to bind only one ankyrin molecule, probably because ankyrin is able to bind to the spectrin tetramer approximately ten-fold more strongly than to the spectrin dimer. 1.2.3 Regulatory Domain of Ankyrin This domain (55 kDa) is at the COOH-terminal region of the ankyrin molecule, and is known to contain regulatory sequences that enhance or reduce the extent of interaction of ankyrin with spectrin and band 3. Ankyrin binds to the COOHterminal region of the β-spectrin with its 55 kDa domain and to the cytoplasmic tail of band 3 via repeated sequences in the NH2 -terminal 89 kDa domain. The regulatory domain consists of multiple isoforms of different sizes and functions, which are produced by alternative splicing [8, 9]. One of these distinct isoforms is ankyrin 2.2, which lacks the acidic 162 amino acid sequence from exon 38, which is found in full-sized ankyrin (protein 2.1) [6]. This protein 2.2 (the smaller isoform: ankyrin 2.2) is an activated ankyrin, and enhances ankyrin binding to band 3 and spectrin. Use of alternative promoters has recently been shown to produce a muscle-specific, truncated isoform of ANK 1 [9]. Many alternatively spliced isoforms of ankyrin at the three COOH-terminal exons are known, these are isoforms lacking: (1) exons 38 and 39, (2) exons 36 through 39, and (3) exons 36 through 41. The COOH-terminal exons are highly conserved [8]. 1.3 Functions of Ankyrin

The major function of ankyrin lies in its binding to β-spectrin and band 3 [22]. These bindings create a tight association between spectrin and band 3, but the strength of binding can be modified by the extent of phosphorylation [23]. The presence of seven phosphorylation sites have been proven in vitro for casein kinase I and cyclic AMP-independent protein kinase. Unphosphorylated ankyrin binds preferentially to spectrin tetramers and oligomers rather than to spectrin dimers, but phosphorylation removes this preference. Phosphorylation also reduces the capacity of ankyrin to bind band 3. In addition, ankyrin is phosphorylated by protein kinase A. Stoichiometrically, one ankyrin molecule links each spectrin tetramer to the membrane. Since these binding interactions are cooperative, attachment of ankyrin to band 3 enhances greatly the ability of the molecule to organize the spectrin, to


which it is attached, into tetramers. It is also true that spectrin binding enhances the affinity for band 3. Therefore, ankyrin plays a crucial role in the organization of the network. 1.4 Erythroid and Nonerythroid Ankyrins

Ankyrins are widely expressed, and several ankyrin gene families are known [2]. Red cell ankyrin (ankyrinR , or ANK 1) is expressed not only in red cells, but also in myocytes, endothelial cells, and brain (especially in Purkinje cells of the cerebellum) [6, 7]. The gene is located on 8p11. 2. AnkyrinB (ANK 2) is a neural form, and is present in neuronal cell bodies and dendrites [24, 25]. The gene is located on 4q25–q27. AnkyrinG (ANK 3) is the most widely distributed, mostly in epithelia and axons, but also in megakaryocytes, macrophages, myocytes, melanocytes, hepatocytes, kidney cells, and testicular Leydig’s cells [26, 27]. Nonerythroid ankyrins interact with a variety of integral membrane proteins other than band 3 (AE1), such as Na+ -, K+ -ATPase [28–30], the voltage-dependent axonal Na+ channel [31], the amiloride-sensitive epithelial channel [32], the cardiac Na+ /Ca2+ exchanger [33], H+ -, K+ -ATPase [34], the IP3 receptor [35], CD44 [36], and a group of neuro-fascinrelated brain adhesion molecules [37, 38]. Mice with targeted disruption of ANK 1 or ANK 3 exhibit neurological abnormalities, including Purkinje cell degeneration and ataxia, and those with targeted disruption of ANK 2 also exhibit brain defects, but with more clear-cut abnormalities in brain development [39].

2 Protein 4.2

Red cell protein 4.2 (P4.2) is one of the major components of the red cell membrane skeletal network, which binds to the cytoplasmic domain of anion exchanger band 3 and interacts with ankyrin in red cells [40–43]. Patients with P4.2 deficiency in their red cell membranes suffer from congenital hemolytic anemia with microspherocytosis or other such disorders. This fact suggests that P4.2 plays an important role in maintaining the stability and flexibility of red cells. P4.2 has been reviewed previously by two authors in 1993 [40], 1994 [41, 42] and recently in 2000 [43]. 2.1 Protein Chemistry of Protein 4.2

Protein 4.2 is a membrane protein (Fig. 2) accounting for approximately 5% of the total membrane protein content and for 250 × 103 copies per red cell. It has a molecular weight of 72 kDa on SDS–PAGE [44, 45]. Extraction of protein 4.2 from red cell membranes is more difficult than for any of the other peripheral proteins, even under high and low ionic strength conditions. Therefore, strong basic conditions (pH 11 or above) have been used in combination

2 Protein 4.2 5

Fig. 2 Molecular structure of protein 4.2 and the genetic isoforms. A wild type of protein 4.2 is shown at the second line from the top. Black solid boxes indicate constitutive coding exons, and shaded boxes show alternative coding exons.

with gel filtration with 1 M KI-Sephacryl S-200 as the standard method for the extraction of protein 4.2. This procedure yields 1–2 mg of “type I” protein 4.2 from 500 mL of whole blood with a purity of approximately 85%. This protein is water-soluble, and is difficult to separate from residual ankyrin and protein 4.1. The use of other extraction methods with 10 to 20 mM lithium diiodosalicylate, 6 mM 2,3-dimethylmaleic anhydride, 5 mM p-chloromercuribenzoic acid (pCMB), or 1 mM p-chloromercuribenzoic sulfate (pCMBS) have been reported. The nonionic detergent Triton X-100 can also be used to extract protein 4.2 from red cell membranes. Under these conditions, band 3 is co-extracted along with the portion of protein 4.2, suggesting an association of protein 4.2 with band 3 in red cell membranes in situ [44, 45]. An alternative method using 2 M Tris-HCl (pH 7.6) has also been used to extract protein 4.2 with a purity of greater than 97%. However, this “type II” variety of protein 4.2 is less water-soluble and behaves like an integral protein. It has been speculated that its characteristic hydrophobicity is due to myristylation [46]. Protein 4.2 purified by the standard pH 11 method with 1 M KI-Sephacryl S-200 appears to be heterogeneous in size and probably primarily consists of a mixture of dimers and trimers. Electron microscopically, purified protein 4.2 appears as globular particles with diameters in the range 80–150 Å, and has been suggested as being tetrameric in situ in the membrane.


Protein 4.2 in human red cell membranes is known to be myristylated [47] at a site near the N-terminus, as assayed by the release of myristoyl glycine from partially hydrolyzed protein 4.2. Glycine at the second position appears to be responsible for this myristylation (N-myristoyl glycine). Further studies of its biological functions should be considered. It has also been reported that protein 4.2 is palmitoylated under physiological conditions [48]. After labeling of intact human red cells with 3 [H] palmitic acid, radioactivity was found to be associated with protein 4.2 by immunoprecipitation of peripheral membrane proteins extracted at pH 11 from ghosts with anti-protein 4.2 antibody. The fatty acid linked to protein 4.2 was identified as palmitic acid. Protein 4.2 could be depalmitoylated with hydroxylamine, suggesting a thioester linkage. Depalmitoylated protein 4.2 showed significantly decreased binding to protein 4.2-depleted membranes, as compared with native protein 4.2. Several red cell membrane proteins including ankyrin, band 3, p55, protein 4.1 and spectrin were palmitoylated. Fatty acid acylation of proteins confers an extra hydrophobic moiety on proteins, which promotes hydrophobic protein–membrane and protein–protein interactions. Whereas control protein 4.2 showed a binding capacity of 280 mg per g of vesicle protein (band 3), depalmitoylated protein 4.2 showed a capacity of 108 mg per g of vesicle protein. Therefore, palmitoylation of protein 4.2 appears to favor its interaction with band 3 in the membrane. To date, protein 4.2 has not been crystallized. Crystallography is thus expected to be performed in the future. 2.2 Functions of Protein 4.2

The major functions of protein 4.2 are, in conjunction with other membrane proteins, topographically adjacent to it in situ in red cell membranes, especially band 3, ankyrin, spectrin, and protein 4.1. 2.2.1 Binding Properties of Protein 4.2 2.2.1.1 Interactions of protein 4.2 with band 3 When Triton X-100 extracts of red cell membranes were fractionated by ion exchange chromatography and nondenaturing gel electrophoresis, protein 4.2 was found with band 3 [44]. In addition, direct binding assays have indicated that an excess of the cytoplasmic domain of band 3 eliminated the normal (2–8 × 10−7 M) binding of purified protein 4.2 to red cell inside-out vesicles (IOV). The binding of protein 4.2 to the purified cytoplasmic domain of band 3 usually takes from 6 to 20 h for complete saturation. Therefore, it has been hypothesized that re-binding of protein 4.2 to the membrane probably requires the formation of other types of associations apart from protein–protein contacts at the membrane–medium interface, perhaps the formation of protein 4.2 or band 3 oligomers. Although the major binding site of protein 4.2 has been considered to be the cytoplasmic domain of band 3, no direct evidence has been found, because the state of self-association of purified protein 4.2 is heterogeneous and the exact oligomeric state of band 3 in situ in membranes under the conditions of the

2 Protein 4.2 7

binding assays is unknown. It has been tentatively estimated that the stoichiometry of the protein 4.2 to band 3 interaction is approximately 1:3.9 on a monomer basis. One synthetic peptide of protein 4.2 (P8: L61 FVRRGQPFTIILYF) was found to bind strongly to the cytoplasmic domain of band 3 [49]. Four other peptides (P22: L271 LNKRRGSVPILRQW, P27: G346 EGQRGRIWIFQTST, P41: L556 WRKKLHLTLSANLE, and P48: I661 HRERSYRFRSVWPE) bind less strongly. These peptides have in common a cluster of two or three basic amino acid residues (arginine or lysine) in a region with virtually no acidic residues. The cytoplasmic domain of band 3 bound in a saturated manner to P8 with a K d of 0.16 µM and a capacity of 0.56 mol of the cytoplasmic domain of band 3 monomer per mol of P8. Replacement of R64 R with R64 G, G64 R or G64 G almost completely abolishes the cytoplasmic domain of band 3 binding, suggesting that R64 R is essential for its binding. P8 competitively inhibits binding of purified human red cell P4.2 to the cytoplasmic domain of band 3. Protein 4.2 can be found with band 3 and ankyrin in an immunoprecipitated complex, probably due to association of protein 4.2 with ankyrin and band 3 in situ [40]. A partial deficiency of band 3 has also been found to be accompanied by partial deficiency of protein 4.2 [43]. Furthermore, a cow with total band 3 deficiency and mice with the targeted band 3-knock-out gene clearly demonstrated a loss of protein 4.2 in their red cell membranes [43]. The rotational and lateral mobility of band 3 in red cell membranes from patients with protein 4.2 deficiency is substantially increased [50–52]. In our study with fluorescence recovery after use of the photobleaching method (FRAP), the immobile fraction of band 3, which constitutes about 60% of the total in normal red cells, was totally absent in complete protein 4.2 deficiency. It is also interesting that, with complete deficiency of protein 4.2, the number of intramembrane particles (IMP) has been found to be reduced with a shift to larger sizes, indicating the possibility of increased oligomerization of band 3 molecules in these red cells [53]. Heat treatment considerably enhanced this effect. The structural and functional characteristics of band 3 in these protein 4.2-deficient red cells appeared normal in terms of the cleavage pattern of band 3 fragments and the binding properties of band 3 to protein 4.2 or ankyrin. 2.2.1.2 Interactions of protein 4.2 with ankyrin Considering the association of protein 4.2 with ankyrin, it has been reported that protein 4.2 can bind to 0.65 mol of ankyrin per mol of protein 4.2 with a K d of from 1 to 3.5 × 10−7 M on the Scatchard plot in vitro [45]. However, this binding requires several hours to approach saturation in solution, and no conclusive evidence has yet been shown for an association of protein 4.2 with ankyrin in the membrane in situ. It has also been shown that ankyrin can bind to the cytoplasmic domain of band 3 in IOV without protein 4.2, and that reassociation of ankyrin with IOV is unaffected even when protein 4.2 is removed from the IOV. In some hemolytic anemias, however, a partial deficiency of protein 4.2 has also been reported in ankyrin deficiencies [54]. Decreased protein 4.2 content has been noted in a mouse strain (nb/nb) with an ankyrin deficiency. In a protein 4.2 deficiency with the 142 Ala←Thr point mutation, it was reported that ankyrin was partially released from the patient’s red cell membranes upon preparation of


IOV, suggesting that protein 4.2 might contribute to the stability of the membrane protein association. It has been shown that the red cell membranes of nb/nb mice, which were almost completely deficient in full-length 210 kDa ankyrin due to a defect in the Ank-1 gene on mouse chromosome 8, were severely (up to 73%) deficient in protein 4.2 content [55]. This deficiency of protein 4.2 in nb/nb homozygous mice was not the result of defective protein 4.2 synthesis. Reconstitution of nb/nb to inside-out vesicles with human red cell ankyrin restored ankyrin levels to up to 80% of the normal levels and increased binding of exogenously added human red cell protein 4.2 by approximately 60%. These results suggest that ankyrin is required for normal associations of protein 4.2 with the red cell membrane. 2.2.1.3 Interactions of protein 4.2 with spectrin Normal protein 4.2 has been shown to bind to spectrin in solution and to promote the binding of spectrin to ankyrinstripped inside-out vesicles [51]. Two independent classes of binding sites of protein 4.2 to spectrin have been identified: 1) a high-affinity (a binding coefficient: K d = 7.4 ± 0.2 × 10−9 M−1 ), low-capacity (0.6 ± 0.8 × 10−9 M L−1 ) class of sites; and 2) a low-affinity (K d = 2.8 ± 2.0 × 10−7 M−1 ), high capacity (5.8 ± 1.0 × 10−9 M L−1 ) class of sites. It has been calculated that, at saturation, there is approximately one spectrin binding site per seven protein 4.2 molecules. Therefore, protein 4.2 provides low-affinity binding sites for both band 3 oligomers and spectrin dimers on the human red cell membrane. These observations suggest that protein 4.2 may stabilize skeleton–membrane interactions by providing a direct link between band 3 and spectrin. A spectrin-binding domain of human erythrocyte membrane protein 4.2 has recently been identified [56]. In a disease state, i. e., in red cells with total protein 4.2 deficiency and also in red cells of 4.2−/− mice, the cytoskeletal proteins (spectrins, ankyrin, and protein 4.1) are not deficient [41, 42, 57]. However, the cytoskeletal network in these protein 4.2-deficient red cells appears to be less extended when studied by electron microscopy using the surface replica method and the quick-freeze deep-etching method [53]. Interestingly, the cytoskeletal network in these protein 4.2-deficient red cells becomes markedly disorganized, with the appearance of larger aggregates when heat-treated up to 48 ◦ C [50]. Under these conditions, a marked decrease in red cell membrane deformability has been observed by ektacytometry It should be noted that the spectrin and ankyrin contents were maintained as normal in these cells. Therefore, these abnormalities appear to be independent of spectrin and ankyrin per se, and due to the lack of protein 4.2. These results raise the possibility that protein 4.2 may play a role in connecting the cytoskeletal network to integral proteins (especially band 3) as a type of anchoring protein. 2.2.1.4 Interaction of protein 4.2 with protein 4.1 As to a possible association of protein 4.2 with protein 4.1, it has been reported that protein 4.1 may interact with protein 4.2 in solution, suggesting the masking of the binding domains of these proteins when they are present together in solution [45]. Protein 4.2, protein 4.1 and ankyrin binding are partially inhibited (about 50%) by the presence of these proteins.

2 Protein 4.2 9

The protein 4.1 and protein 4.2 binding sites are localized at the nearby sites on the cytoplasmic domain of band 3. It is possible that, in the absence of protein 4.2, additional binding sites for protein 4.1 on band 3 may be exposed. The content of protein 4.1, however, remains nearly normal both in human red cells when there is a total deficiency of protein 4.2 and in 4.2−/− mouse red cells [41, 42, 57]. It is also worth noting that the content of protein 4.2 appears to be normal in mice with a complete deficiency of all protein 4.1 R isoforms, which had beeen generated by gene knock-out technology [58]. 2.2.2 Transglutaminase Activity of Protein 4.2 Protein 4.2 in human red cells has no transglutaminase activity, although the homology in the gene structure between protein 4.2 and transglutaminase is high (i.e., an overall identity of 32% in a 446 amino-acid overlap with guinea-pig liver transglutaminase, and of 27% in a 639 amino-acid overlap with human coagulation factor XIII subunit a) [59]. It has been speculated that the lack of transglutaminase activity may be due to the presence of an alanine substituted for the cysteine at the active site of the molecule. 2.2.3 Phosphorylation of Protein 4.2 Although phosphorylation has not been observed on protein 4.2 extracted from mature human red cells, protein 4.2 can be phosphorylated, if it is purified by methods that result in exposing phosphorylation sites through alteration of protein 4.2 sulfhydryl groups [40, 46]. Seventeen potential phosphorylation sites have been activated in red cell ghosts, i. e., eight possible protein kinase C sites, seven casein kinase II sites, one tyrosine kinase site, and one cAMP- (or cGMP-) dependent kinase site [47]. It has been suggested that membrane-associated protein 4.2 in human mature red cells is already fully phosphorylated with little or no turnover under normal conditions. The physiological functions of such potential protein 4.2 phosphorylation is unknown. The activities of the major red cell kinases have recently been determined to assess the phosphorylation status in red cells of 4.2−/− mice [57]. Cytosolic protein kinase C (PKC) was significantly decreased with decreased PKC-α and PKC-β I isoforms and normal PKC-β III in 4.2−/− red cells. Cytosolic protein kinase A (PKA) activity was increased in these red cells. Basal phosphorylation was increased and PMAstimulated phosphorylation was reduced in 4.2−/− red cells. Cytosolic casein kinase I (CK I) activity was normal, but cytosolic CK II activity was decreased in these red cells. The functional significance of these activities remains to be clarified at some point in the future. 2.3 Protein 4.2 in Red Cell Membrane Ultrastructure

Although the localization of P4.2 in the red cell membrane structure has still not been elucidated in detail, there are several positive pieces of evidence. With a total deficiency of P4.2, it is now recognized that the intramembrane particles (IMPs) are clustered in the inside-out vesicles (IOVs) of the patient’s red cells [41, 50, 53]. In


addition, electron microscopic studies with the freeze fracture method have shown IMPs on the red cell ghosts to be enlarged, suggesting increased oligomerization of band 3 molecules. This phenomenon has been verified by biophysical analyses [41–43, 53]. Furthermore, analyses with the quick-freeze deep-etching method or the surface replica method have shown the cytoskeletal network to be disrupted in this disorder [41–43, 53]. These results appear to indicate that P4.2 molecules are located near band 3 molecules and membrane proteins making up the cytoskeletal network. Biophysical analyses, especially with ektacytometry, have revealed the increased instability of the cytoskeletal network of the patient’s red cells. It has recently been proven that P4.2 can bind directly to spectrins [51], implying that P4.2 may play an important role as one of the anchoring proteins connecting the cytoskeletal network to the integral proteins (particularly band 3 molecules). The exact location of various membrane proteins in situ in the normal human red cell membrane ultrastructure has been studied by immnuno-electron microscopy with the surface replica method by utilizing antibodies against various membrane proteins: i.e., spectrins, ankyrin, band 3 (the cytoplasmic domain), protein 4.1 and protein 4.2. At first, spectrins were readily detected as major constituents of the cytoskeletal network, and ankyrin and protein 4.1 were also identified on it. Band 3 molecules were found attached to the cytoskeletal network as an immobile form of band 3. Other band 3 molecules were located inside the basic units, as the mobile form of band 3. P4.2, however, could not be detected by this procedure, when the antibody-conjugated immunogold particles were applied to the open red cell ghosts, implying that the epitopes of P4.2 were not exposed. Therefore, normal red cell ghosts were subjected to gentle treatment with Triton X-100 to remove part of the cytoskeletal network. P4.2 was then found to be attached predominantly to the cytoskeletal network. These results suggest that P4.2 is very likely to be present at the outer face of the cytoskeletal network and even under the lipid bilayer, and is attached to spectrins and band 3, as shown by biophysical studies [51]. 2.4 Protein 4.2 Gene 2.4.1 Characteristics of Genomic DNA The human red cell protein 4.2 gene is ∼20 kb in length and contains 13 exons and 12 introns [59–63]. The coding sequence from the genomic DNA is identical to the cDNA sequence. Nucleotide polymorphism has been observed in the normal human protein 4.2 gene. The exons range in size from 104 to 314 base pairs with an average size of 170 base pairs, while the introns vary from 6.4 to 0.3 kb. All the exon–intron boundaries follow the consensus 5 donor–3 acceptor splice junction sequence for eukaryotic genes of gt-ag. The gene is localized at human chromosome 15q15–q21 [62, 63]. The upstream region of the protein 4.2 gene contains several elements that are similar in sequence to the upstream elements of the genes for β-globin and porphobilinogen deaminase, which are also red cell protein genes [59, 62]. The elements are spaced a similar distance from the transcription start site and have

2 Protein 4.2 11

similar relative spacings and orders. These similarities have made it easier to identify five possible regulatory cis-elements in the protein 4.2 gene, starting –20 nucleotides upstream from the transcription start site, i.e.: (1) a possible TATA element, (2) a short G + C-rich domain, which could be an Spl binding site, (3) a possible CAAT box, (4) a CAAC box, and (5) two GF-1 binding domains, one at −23 to −28, and another one at –173 to –178 [59]. These findings suggest the use of common cis-elements in these three erythroid genes, although the identification of these elements as having a regulatory function in protein 4.2 gene expression is highly speculative. It has also been reported [62] that the nucleotides upstream from the cDNA start site (nt 1) are: (1) CAGT (nt –4 to –1), agreeing well with the CA cap signal; (2) nucleotides –26 to –21 upstream from the cDNA start site having a sequence of ATAAAA, which agrees well in sequence and position with the promoter TATA box for eukaryotic genes; (3) a CCAT sequence was noted at nt –89 to –86 within the reported –385 nt upstream sequence, where upstream promoter elements are located; (4) a GC-rich region from nt –85 to –34 (G/C to A/T ratio = 3); (5) within this region, a sequence of CCCACCCC CTCCCCC containing a CACC element (nt –83 to –80) that is a potential binding site for the Spl nuclear factor; and (6) two AGATAA sequences for potential binding of erythroid-specific transcription factor GATA-1 (also known as GF-1, NF-E1, Eryf-1) located at nt –175 to –170 and at nt –28 to –23, respectively. The number of the 5 -CpG-3 di-nucleotide sites appears to be small, unlike the β-spectrin gene that has numerous 5 -CpG-3 sites known as the so-called “CpG islands” [64]. The 5 -CpG-3 sites of the protein 4.2 gene were highly methylated, when genomic DNA was prepared from mononuclear cells in normal human peripheral blood. Alignment of the protein 4.2 amino acid sequence with that of a subunit of human coagulation factor XIII and division of the sequence into exons have revealed a remarkable correspondence, although the gene for the a subunit of human factor XIII, which is on chromosome 6p24–p25, is 160 kb and has 15 exons and 14 introns, while the gene for protein 4.2 is only 20 kb and contains 13 exons and 12 introns [59]. With only one exception, the exons of protein 4.2 are very similar and in many cases identical in size to the exons of the a subunit of factor XIII with which they are paired. In additon, in every case, the corresponding intervening introns are of the same splice junction class. These and other similarities suggest that the gene for protein 4.2 is closely related to and possibly derived from that for the a subunit of factor XIII and that the proteins may share common structural and functional properties. However, it should be noted that, despite this close similarity, purified protein 4.2 has no transglutaminase activity in vitro, and that normal red cell membranes do not contain transglutaminase activity. The lack of protein 4.2 transglutaminase activity is induced by the substitution of the cysteine (GQCWVF) at the highly conserved consensus sequence in the transglutaminase, which is substituted for an alanine (GQAWVL) in protein 4.2. The cysteine appears to be required for transglutaminase activity. It is also possible that the substitution of a leucine for a phenylalanine may also be responsible for a loss of this activity. It has been shown that reticulocytes contain two forms of protein 4.2 mRNA, a small form (P4.2S) encoding a protein of 691 amino acids, and a larger form (P4.2L),


which contains an additional 90 nucleotides following nucleotide +9, encoding a protein of 721 amino acids [59, 62]. Protein 4.2 exon I contains a 5 non-coding sequence, the translation start site, and 99 nucleotides encoding 33 amino acids. These 33 amino acids are identical to the first 33 amino acids of the larger protein 4.2 transcript. The last 90 nucleotides of exon I, coding for 30 amino acids, are removed by splicing in order to generate the smaller transcript, coding for the 691 amino acid protein (a wild type of protein 4.2: 72 kDa on the SDS–PAGE). The genomic organization of the protein 4.2 gene of human red cells contains 13 exons, i.e.: exon I, ut ∼33 residues; II, 34–95; III, 96–173; IV, 174–213; V 214–248; VI, 249–307; VII, 308–354; VIII, 355–388; IX, 389–469; X, 470–569; XI, 570–623; XII, 624–668; and XIII, 669-ut (ut, untranslated sequence) [59, 60] (Fig. 2). The sizes of the introns were 6490 for intron 1, 900 for 2, 580 for 3, 340 for 4, 940 for 5, 320 for 6, 740 for 7, 500 for 8, 390 for 9, 2560 for 10, 2200 for 11, and 3080 for 12. 2.4.2 cDNA of the Protein 4.2 Gene Protein 4.2 complimentary DNA (cDNA) obtained from a human reticulocyte cDNA library has been cloned and sequenced [60–62]. The full-length cDNA was 2.35 kb and contained an open reading frame with a 227–nt untranslated region upstream from the putative ATG start codon. The calculated molecular weight was 76.9 kDa encoding 691 amino acids. The nucleotide sequence CAACCATGC around this initiation site was similar to the consensus sequence for initiation found in higher eukaryocytes, except that the second nt in the P4.2 cDNAs was A rather than C [61]. The presence or absence of the 90 nt insert gave rise to two P4.2 cDNA sequences; that is, a P4.2S from 2073 bp and a P4.2L from 2163 bp [59, 62]. The amino acid sequence derived from the 2.5 kb cDNA contained ∼43% nonpolar, ∼35% polar, ∼10% acidic, and ∼12% basic amino acid residues [61]. The most abundant amino acids were leucine (82 residues) and alanine (60 residues). There were 49 serine and 43 threonine residues, which are potential sites for O-glycosylation and represent 13% of the total residues. There were 16 cysteine residues, six potential N-glycosylation sites (Asn-Xaa-Ser/Thr) at Asn-103, -420, -447, -529, -604, and -705, one potential cAMP-dependent phosphorylation site (basic-basicXaa-Ser) at Ser-278, and nine potential protein kinase C phosphorylation sites (Ser/Thr-Xaa-Arg/Lys) at Ser-7, -57, -58, -154, -224, -449, -455, and -666, and Thr287 [61]. There was one Arg-Gly-Asp sequence at 518–520. Secondary structure analysis predicted that P4.2 should contain ∼3% β-sheet, ∼24% α-helix, and ∼45% reverse turns. Hydropathy analysis of the deduced amino acid sequence revealed a major hydrophobic domain (residues 298–322), which was predicted to be mainly a β-sheet structure with a possible turn. There was a strongly hydrophilic region (residues 438–495). Toward the C terminus of this region, there was a highly charged segment predicted to be an α-helix (residues 470–492) and containing a large number of both positively and negatively charged residues, especially glutamic acid [61]. Elsewhere, it was reported that there were 37% hydrophobic residues and 28% polar residues [60]. Protein 4.2 did not show any obvious repeating primary structure, but a globular protein was suggested [60]. There were no extended stretches of β-sheet or α-helix. Instead, the protein was

2 Protein 4.2 13

characterized by short segments. A hydropathy plot of protein 4.2 showed short alternating regions of hydrophobic and hydrophilic character [60]. The region of protein between amino acids 265 and 475, however, was characterized by two sets of alternating, prominent hydrophobic and hydrophilic domains [60]. 2.4.3 Protein 4.2 Gene in Mouse Red Cells There are substantial discrepancies between the three reports published on the protein 4.2 gene in mouse red cells [65–67]. Korsgren and Cohen (1994) described isolation of a 3.5 kb mouse P4.2 cDNA with the P4.2 transcript of 4.1 kb from mouse reticulocytes [65]. Rybicki et al. (1994), on the other hand, reported isolation of a full-length P4.2 cDNA of 2.2 kb from mouse reticulocytes [66]. Karacay et al. (1995) described an entire P4.2 cDNA sequence consisting of 3465 nt with an open reading frame (ORF) of 691 amino acids, and despite its similarity to human P4.2 cDNA, the mouse cDNA had a longer 3 untranslated region [67]. In addition, they reported that the mouse reticulocyte P4.2 RNA did not exhibit alternative splicing in the region identified in human P4.2 RNA. The P4.2 gene in mice was mapped to murine chromosome 2 [68], in contrast to 15q15–q21 in the human red cell P4.2 gene. 2.4.4 Tissue-Specific Expression of the Mouse Protein 4.2 Gene and the Pallid Mutation Immunoreactive forms of P4.2 with a molecular weight of 72 kDa, in addition to those larger or smaller than 72 kDa, have been detected in nonerythroid cells and tissues [40, 69–71]. Immunologic cross-reactivity between the red cell P4.2 protein and other cellular proteins has also been reported [40, 69–71]. Zhu et al. (1998) recently reported that expression of the mouse P4.2 gene was temporally regulated during embryogenesis and that the P4.2 mRNA expression pattern matched the timing of erythropoietic activity in hematopoietic organs [70]. It should be noted that, contrary to previous reports, P4.2 expression was detected only in the erythroid cell-producing organs and circulating red cells during mouse embryonic development and in adult mice. They first analyzed poly A +RNAs from various adult mice tissues by Northern blot analysis using a 714 bp mouse P4.2 cDNA fragment containing the 3 protein of the P4.2-coding region as the probe. A single 3.5 kb P4.2 transcript was detected at a relatively high level in the spleen, while little or no P4.2 hybridization was seen in other tissues examined (brain, lung, liver, skeletal muscle, kidney, or testis). They extended the use of mice embryos for their P4.2 expression studies. A P4.2 hybridization signal was first detected not at E6.5 days, but instead in primitive erythroid cells in E7.5 embryos. In E10.5 embryos, the P4.2 hybridization signal was detected only in the heart and blood vessels. In E12.5 embryos, there was a switch in the hematopoietic production sites from the yolk sac to the fetal liver. In E 16.5 embryos, the signal was greatly reduced in the liver, and was almost undetectable after birth. Finally, P4.2 gene expression became confined to the red pulp on postnatal day 7. No P4.2 specific labeling was observed in the white pulp, which consisted of germinal centers for lymphocytes, plasma cells, and macrophages. No P4.2 hybridization signal was detected in megakaryocytes.


Therefore, the P4.2 message was specifically expressed in cells of erythroid lineage in postnatal hematopoietic organs [70]. The chromosomal location of the mouse P4.2 gene was near a mouse pallid (pa) mutation [71]. Pallid was found in a mouse with dilution of coat color, increased bleeding time, and abnormal lysosomal enzyme secretion, as a model of the platelet storage pool disease [72, 73]. Therefore, it has been suggested that the P4.2 gene may be related to the pallid mutation gene, and it has even been proposed that the P4.2 gene itself should be nominated as a “pallidin” [40, 65, 71]. Patients with P4.2 deficiency, however, do not have the platelet storage pool deficiency seen in pa/pa mice, and the mutant mice do not exhibit the hemolysis and spherocytosis observed in the P4.2 deficiency [41–43]. It has recently been shown that the P4.2 gene is distinct from the pa gene, and that changes in P4.2 in pallid mice were not responsible for the pallid mutation [68]. There have also been reports of P4.2 transcripts, in addition to spleen, in other tissues (kidney, heart, brain, and liver), and even in other cells (HeLa cells or HT-29 cells). As with the results for the immunoreactive forms of P4.2 previously detected in nonerythroid tissues or cells, neither P4.2 message nor protein 4.2 have been found in nonerythroid tissues and cells [70].

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with homology to tissue-differentiation and cell-cycle control proteins. Nature 344: 36–42. LAMBERT, S. YU, H., PRCHAL, J. T., LAWLER, J., RUFF, P., SPEICHER, D., CHEUNG, M. C., KAN, Y. W., PALEK, J. (1990) cDNA sequence for human erythrocyte ankyrin. Proc. Natl. Acad. Sci. USA 87: 1730–1734. GALLAGHER, P. G., TSE, W. T., SCARPA, A. L., LUX, S. E., FORGET, B. G. (1997) Structure and organization of the human ankyrin-1 gene: Basis for complexity of pre-mRNA processing. J. Biol. Chem. 272: 19220–19228. GALLAGHER, P. G., FORGET, B. G. (1998) An alternate promoter directs expression of a truncated, muscle-specific isoform of the human ankyrin 1 gene. J. Biol. Chem. 273: 1339–1348. GALLAGHER, P. G., SABATINO, D. E., GARETTE, L. J., BODINE, D. M., FORGET, B. G. (1998) Erythroid-specific expression of the human ankyrin 1 (Ank 1) gene in vitro and in vivo is mediated by a promoter that requires GATA-1 and CACCC-binding proteins for its activity. Blood 90: 7a.

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20 PLATT, O. S., LUX, S. E., FALCONE, J. F. (1993) A highly conserved region of human erythrocyte ankyrin contains the capacity to bind spectrin. J. Biol. Chem. 268: 24421–24426. 21 KENNEDY, S. P., WARREN, S. L., FORGET, B. G., MORROW, J. S. (1991) Ankyrin binds to the 15th repetitive unit of erythroid and nonerythroid P-spectrin. J. Cell Biol. 115: 267–277. 22 DAVIS, L. H., BENNETT, V. (1990) Mapping the binding sites of human erythrocyte ankyrin for the anion exchanger and spectrin. J. Biol. Chem. 265: 10589–10596. 23 CIANCI, C. D., GIORGI, M., MORROW, J. S. (1988) Phosphorylation of ankyrin down-regulates its cooperative interaction with spectrin and band 3. J. Cell Biochem. 37: 301–315. 24 OTTO, E., KUNIMOTO, M., MCLAUGHLIN, T., BENNETT, V. (1991) Isolation and characterization of cDNAs encoding human brain ankyrins reveal a family of alternatively spliced genes. J. Cell Biol. 141: 241–253. 25 KUNIMOTO, M., OTTO, E., BENNETT, V. (1991) A new 440-kD isoform is the major ankyrin in neonatal rat brain. J. Cell Biol. 115: 1319–1331. 26 PETERS, L. L., JOHN, K. M., LU, F. M., EICHER, E. M., HIGGINS, A., YIALAMAS, M., TURTZO, L. C., OTSUKA, A. J., LUX, S. E. (1995) Ank 3 (epithelial ankyrin), a widely distributed new member of the ankyrin gene family and the major ankyrin in kidney, is expressed in alternatively spliced forms, including forms that lack the repeat domain. J. Cell Biol. 130: 313–330. 27 KORDELI, E., LAMBERT, S., BENNETT, V. (1995) AnkyrinG. A new ankyrin gene with neural-specific isoforms localized at the axonal initial segment and rode of Ranvier. J. Biol. Chem. 270: 2352–2359. 28 DEVARAJAN, P., SCARAMUZZINO, D. A., MORROW, J. S. (1994) Ankyrin binds to two distinct cytoplasmic domains of Na, K-ATPase α-subunit. Proc. Natl. Acad. Sci. USA 91: 2965–2969. 29 JORDAN, C., PUSCHEL, B., KOOB, R., DRENCKHAHN, D. (1995) Identification of a binding motif for ankyrin on the

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α-subunit of Na+ , K+ -ATPase. J. Biol. Chem. 270: 29971–29975. ZHANG, Z., DEVARAJAN, P., DORFMAN, A. L., MORROW, J. S. (1998) Structure of the ankyrin-binding domain of α-Na, K-ATPase, J. Biol. Chem. 273: 18681–18684. SRINIVASAN, Y., ELMER, L., DAVIS, J., BENNETT, V., ANGELIDES, K. (1988) Ankyrin and spectrin associate with voltage-dependent sodium channels in brain. Nature 333: 177–180. SMITH, P. R., SACCOMANI, G., JOE, E. H., ANGELIDES, K. J., BENOS, D. J. (1991) Amiloride-sensitive sodium channel is linked to the cytoskeleton in renal epithelial cells. Proc. Natl. Acad. Sci. USA 88: 6971–6975. LI, Z. P., BURKE, E. P., FRANK, J. S., BENNETT, V., PHILIPSON, K. D. (1993) The cardiac Na+ -Ca2+ exchanger binds to the cytoskeletal protein ankyrin. J. Biol. Chem. 268: 11489–11491. SMITH, P. R., BRADFORD, A. L., JOE, A. H., ANGELIDES, K. J., BENOS, D. J., SACCOMANI, G. (1993) Gastric parietal cell H+ , K+ -ATPase microsomes are associated with isoforms of ankyrin and spectrin. Am. J. Physiol. 264: C63–70. BOURGUIGNON, L. Y., JIN, H. (1995) Identification of the ankyrin-binding domain of the mouse T-lymphoma cell inositol 1,4,5-triphosphate (IP3) receptor and its role in the regulation of IP3-mediated internal Ca2+ release. J. Biol. Chem. 270: 7257–7260. ZHU, D., BOURGUIGNON, L. Y. (1998) The ankyrin-binding domain of CD44s is involved in regulating hyaluronic acid-mediated functions and prostate tumor cell transformation. Cell Motil. Cytoskeleton 39: 209–222. DUBREUIL, R. R., MACVICAR, G., DISSANAYAKE, S., LIU, C., HOMER, D., HORTSCH, M. (1996) Neuroglian-mediated cell adhesion induces assembly of the membrane skeleton at cell contact sites. J. Cell Biol. 133: 647–655. TUVIA, S., CARVER, T. D., BENNETT, V. (1997) The phosphorylation state of the F1GQY tyrosine of neurofascin determines ankyrin-binding activity and patterns of cell segregation.

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Proc. Natl. Acad. Sci. USA 94: 12957– 12962. DOONER, G. J., BARKER, J. E., GALLAGHER, P. G., DEBATIS, M. E., BROWN, A. H., FORGET, B. G., BECHER, P. S. (2000) Gene transfer to ankyrin-deficient bone marrow corrects spherocytosis in vitro. Exp. Hematol. 28: 765–774. COHEN, C. M., DOTIMAS, E., KORSGREN, C. (1993) Human erythrocyte membrane protein band 4.2 (pallidin). Semin. Hematol. 30: 119–137. YAWATA, Y. (1994) Red cell membrane protein band 4.2: Phenotypic, genetic and electron microscopic aspects. Biochim. Biophys. Acta 1204: 131–148. YAWATA, Y. (1994) Band 4.2 abnormalities in human red cells. Am. J. Med. Sci. 307: 190–243. YAWATA, Y., KANZAKI, A., YAWATA, A. (2000) Genotypic and phenotypic expressions of protein 4.2 in human erythroid cells. Gene Funct. Dis. 2: 61–81. KORSGREN, C., COHEN, C. M., (1986) Purification and properties of human erythrocyte band 4.2. Association with the cytoplasmic domain of band 3. J. Biol. Chem. 261: 5536–5543. KORSGREN, C., COHEN, C. M., (1988) Association of human erythrocyte band 4.2. Binding to ankyrin and to the cytoplasmic domain of band 3. J. Biol. Chem. 263: 10212–10218. DOTIMAS, E., SPEICHER, D. W., GUPTA ROY, B., COHEN, C. M. (1993) Human erythrocyte band 4.2: Structural domain mapping of the protein purified by a novel procedure. Biochim. Biophys. Acta 1148: 19–29. RISINGER, M. A., DOTIMAS, E. M., COHEN, C. M. (1992) Human erythrocyte protein 4.2, a high copy number membrane protein, is N-myristylated. J. Biol. Chem. 267: 5680–5685. DAS, A. K., BHATLACHARYA, R., KUNDU, M., CHAKRABARTI, P., BASU, J. (1994) Human erythrocyte membrane protein 4.2 is palmitoylated. Eur. J. Biochem. 224: 575–580. RYBICKI, A. C., MUSTO, S., SCHWARTZ, R. S. (1995) Identification of a band 3 binding site near the N-terminus of

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erythrocyte membrane protein 4.2. Biochem. J. 309: 677–681. INOUE, T., KANZAKI, A., YAWATA, A., TSUJI, A., ATA, K., OKAMOTO, N., WADA, H., HIGO, I., SUGIHARA, T., YAMADA, O., YAWATA, Y. (1994) Electron microscopic and physicobiochemical studies on disorganization of the cytoskeletal network and integral protein (band 3) in red cells of band 4.2 deficiency with a mutation (codon 142: GCT → ACT). Int. J. Hematol. 59: 157–175. GOLAN, D. E., CORBETT, J. D., KORSGREN, C., THATTE, H. S., HAYETTE, S., YAWATA, Y., COHEN, C. M. (1996) Control of band 3 lateral and rotational mobility by band 4.2 in intact erythrocytes: Release of band 3 oligomers from low-affinity binding sites. Biophys. J. 70: 1534–1542. RYBICKI, A., SCHWARTZ, R. S., HUSTEDT, E. J., COBB, C. E. (1996) Increased rotational mobility and extractability of band 3 from protein 4.2-deficient erythrocyte membranes: Evidence of a role for protein 4.2 in strengthening the band 3-cytoskeleton linkage. Blood 88: 2745–2753. YAWATA, Y., YAWATA, A., KANZAKI, A., INOUE, T., OKAMOTO, N., UEHIRA, K., YASUNAGA, M., NAKAMURA, Y. (1996) Electron microscopic evidence of impaired intramembrane particles and of instability of cytoskeletal network in band 4.2 deficiency in human red cells. Cell Motil. Cytoskeleton 33: 95–105. EBER, S. W., GONZALEZ, J. M., LUX, M. L., SCARPA, A. L., TSE, W. T., DORNWELL, M., ¨ ZCAN, R., HERBERS, J., KUGLER, W., O PEKRUN, A., GALLAGHER, P. G., SCHRO¨ TER, W., FORGET, B.G., LUX, S. E. (1996) Ankyrin-1 mutations are a major cause of dominant and recessive hereditary spherocytosis. Nature Genet. 13: 214–218. RYBICKI, A. C., MUSTO, S., SCHWARTZ, R. S. (1995) Decreased content of protein 4.2 in ankyrin-deficient normoblastosis (nb/nb) mouse red blood cells: Evidence for ankyrin enhancement of protein 4.2 membrane binding. Blood 86: 3583–3589. MANDAL, D., MOITRA, P. K., BASU, J. (2002) Mapping of a spectrin-binding domain of human erythrocyte membrane protein 4.2. Biochem. J. 364: 841–847.

57 PETERS, L. L., JINDL, H. K., GWYNN, B., KORSGREN, C., JOHN, K. M., LUX, S. E., MOHANDAS, N., COHEN, C. M., CHO, M. R., GOLAN, D. E., BRUGNARA, C. (1999) Mild spherocytosis and altered red cell ion transport in protein 4.2-null mice. J. Clin. Invest. 103: 1527–1537. 58 SHI, Z.-T., AFZAL, V., COLLER, B., PATEL, D., CHASIS, J. A., PARRA, M., LEE, G., PASZTY, C., STEVENS, M., WALENSKY, L., PETERS, L. L., MOHANDAS, N., RUBIN, E., CONBOY, J. G. (1999) Protein 4.1 R-deficient mice are viable but have erythroid membrane skeleton abnormalities. J. Clin. Invest. 103: 331–340. 59 KORSGREN, C., COHEN, C. M. (1991) Organization of the gene for human erythrocyte membrane protein 4.2: Structural similarities with the gene for the a subunit of factor XIII. Proc. Natl. Acad. Sci. USA 88: 4840–4844. 60 KORSGREN, C., LAWLER, J., LAMBERT, S., SPEICHER, D., COHEN, C. M. (1990) Complete amino acid sequence and homologies of human erythrocyte membrane protein band 4.2. Proc. Natl. Acad. Sci. USA 87: 613–617. 61 SUNG, L. A., CHIEN, S., CHANG, L.-S., LAMBERT, K., BLISS, S. A., BOUHASSIRA, E. E., NAGEL, R. L., SCHWARTZ, R. S., RYBICKI, A. C. (1990) Molecular cloning of human protein 4.2: A major component of the erythrocyte membrane. Proc. Natl. Acad. Sci. USA 87: 955–959. 62 SUNG, L. A., CHIEN, S., FAN, Y.-S., LIN, C. C., LAMBERT, K., ZHU, L., LAM, J. S., CHANG, L.-S. (1992) Human erythrocyte protein 4.2: Isoform expression, differential splicing, and chromosomal assignment. Blood 79: 2763–2770. 63 NAJFELD, V., BALLARD, S. G., MENNINGER, J., WARD, D. C., BOUHASSIRA, E. E., SCHWARTZ, R. S., NAGEL, R. L., RYBICKI, A. C. (1992) The gene for human erythrocyte protein 4.2 maps to chromosome 15q15. Am J. Hum Genet. 50: 71–75. 64 REMUS, R., ZESCHNIGK, M., ZUTHER, I., KANZAKI, A., WADA, H., YAWATA, A., MUIZNIEKS, I., SCHMITZ, B., SCHELL, G., YAWATA, Y., DOERFLER, W. (2001) The state of DNA methylation in the promoter regions of the human red cell membrane

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69 FRIEDRICHS, B., KOOB, R., KRAEMER, D., DRENCKHAHN, D. (1989) Demonstration of immunoreactive forms of erythrocyte protein 4.2 in nonerythroid cells and tissues. Eur. J. Cell Biol. 48: 121–127. 70 ZHU, L., KAHWASH, S. B., CHANG, L.-S. (1998) Developmental expression of mouse erythrocyte protein 4.2 mRNA: Evidence for specific expression in erythroid cells. Blood 91: 695–705. 71 WHITE, R. A., PETERS, L. L., ADKINSON, L. R., KORSGREN, C., COHEN, C. M., LUX, S. E. (1992) The murine pallid mutation is a platelet storage pool disease associated with the protein 4.2 (pallidin) gene. Nature Genet. 2: 80–83. 72 NOVAK, E. K., HUI, S.-W., SWANK, R. T. (1984) Platelet storage pool deficiency in mouse pigment mutations associated with seven distinct genetic loci. Blood 63: 536–544. 73 REDDINGTON, M., NOVAK, E. K., HURLEY, E., MEDDA, C., MCGARRY, M. P., SWANK, R. T. (1987) Immature dense granules in platelets from mice with platelet storage pool disease. Blood 69: 1300–1306.

1

Skeletal Proteins of the Erythrocyte Membrane Yoshihito Yawata



1 α- and β-Spectrins 1.1 Introduction

Spectrins are the most abundant and largest proteins of red cell membrane skeletal proteins. They constitute approximately 25–30% of the total membrane proteins or 75% of the membrane skeletal proteins, and are present at a concentration of about 200 000 copies per red cell [1–4]. Spectrins are composed of two subunits, the α-chain and the β-chain, which are thus called α-spectrin (2429 amino acids, about 280 kDa) and β-spectrin (2137 amino acids, about 246 kDa) (Fig. 1). The number of copies of both α- and β-spectrins is 240 × 103 per red cell. The α-spectrin gene (SPTA 1) [5–7] is located on chromosome 1 (1q22–q23), and the β-spectrin gene (SPTB) [8–10] is on 14q23–q24.2. They are present as a heterodimer (α 2 β 2 ). The α- and β-spectrins are structurally related, but functionally distinct [11]. The α- and β-chains are aligned side by side in an antiparallel arrangement with respect to their N- and C-terminal ends. Electron microscopically, the spectrin molecule appears to have a slender and twisted rod-like structure. The total length of a spectrin molecule is about 100 nm, when examined in its extended state under an electron microscope with the negative staining method. Spectrins are highly flexible to enable a variety of shape changes. Therefore, spectrins are considered as one of the major determinants of cell shape. Although the theoretical length of spectrin tetramers is about 200 nm, the actual end to end distance is about 76 nm, implying that spectrin tetramers are tightly coiled in the native ultrastructure [12]. These coiled spectrin tetramers are able to extend reversibly to a relaxed state when the membrane is stretched artificially. Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Skeletal Proteins of the Erythrocyte Membrane

Fig. 1 Molecular structure of "- and $-spectrins. Heterotetramers of "- and $-spectrins are arranged in an anti-parallel fashion. The headto-head interaction and the side-by-side interaction (at the nucleation site) are shown in this figure. "-Spectrin is composed of five domains (I to V), and $-spectrin is composed of four domains (I to IV).

1.2 Structure of Red Cell Spectrins

α-Spectrin (280 kDa) can be identified on sodium dodecyl sulfate (SDS)–polyacrylamide gels as a 240 kDa polypeptide [1]. The NH2 -terminal end with an isolated, unpaired helix (helix C) is a self-association site with the COOH-terminal end of the β-spectrin molecule. Nine typical 106 amino acid repeats, which are conformation segments 1 to 9, follow after this self-association site [1–4]. There is a short central segment which lacks homology with the above-mentioned repeats, but is known to be related to SH3 domains as segment 10 [11]. A further 12 more repeats (segment 11 to 22) follow until the COOH-terminal end is reached, where two EF hand structures are present, which are involved in Ca2+ -binding and in regulating Ca2+ action in α-actinin and fodrin. However, the exact role of the SH3 domain and EF hand structures in human red cells in vivo are not known. The α-spectrin chain is divided into five domains, based on the results from limited tryptic digestion of α-spectrin, and designated αI (conformational segments 1 to 6: 80 kDa), αII (7 to 10 including SH3 segment: 46 kDa), αIII (11 to 14: 52 kDa), αIV (15 to 17: 41 kDa), and αV (18 to 22: 41 kDa). This method is useful for isolating and characterizing functional domains of normal and mutated spectrins in disease states, especially hereditary elliptocytosis and hereditary pyropoikilocytosis. β-Spectrin (246 kDa) is isolated as a 220 kDa polypeptide by SDS–polyacrylamide gel electrophoresis (PAGE) [1]. The structure of β-spectrin begins with a nonhomologous NH2 -terminal end that contains an actin-binding domain, followed by 17 homologous 106 amino acid repeat segments [1–4]. Subsequently there is the putative protein 4.1-binding site [13] and a short nonhomologous COOH terminal segment with a consensus sequence for at least four phosphorylation sites for casein kinase 1 [14]. The state of phosphorylation is related to membrane mechanical stability, since it is known that increased phosphorylation decreases membrane

1 "- and $-Spectrins

stability, and also that decreased phosphorylation increases it [15]. Functionally, repeat 15 and part of repeat 16 are arranged in a β sheet structure and form the binding site for ankyrin [16]. β-Spectrin is also divided into four domains, designated as βIV (conformational segments 1 to 7: 74 kDa), βIII (8 to 10: 33 kDa), βII (11 to 16: 65 kDa), and βI (17 down to the COOH end: 23 kDa). In the β-spectrin, no SH3 segment is present. Nucleation sites of α-spectrin and β-spectrin are in αV (conformation segment 19 to 22), and in βIV (1 to 4), respectively. The presence of repeats in α- and β-spectrins strongly suggests that spectrin evolved from the duplication of a single ancestral gene [5–10]. The homologous 106 amino acid repeats in α- and β-spectrin fold into α-helical segments containing three antiparallel helices which are connected by short nonhelical segments. Each repeat has a triple-helical structure that is about 5 nm long and 2 nm wide. The unit is rotated 60◦ (right-handed) relative to the neighboring repeats [17, 18]. Each repeat is composed of three helices, i.e., helix A, helix B, and helix C. The α-helix (helix A) with 28 amino acids in a straight arrangement reverses itself and forms the next α-helix (helix B), which is 34 amino acids long. This is followed by another reverse turn and the third 31 amino acid α-helix (helix C), which bends in the middle. These three helices are in a triangular array and are bound together by both hydrophobic and electrostatic interactions. Hydrophobic amino acids are aligned on one face of each helix, and are spaced every third or fourth residue, because an α-helix makes one turn every 3.6 residues. Additional salt bonds are present between the mostly polar amino acids, particularly between helices A and C, and B and C. The three helices are tilted away from each other by 10◦ to 20◦ , so that the COOH-terminal end of each repeat is wider than the NH2 -terminal end. This molecular arrangement enables the following repeat to be attached without any change to the structure. The repeats connect to each other through the helices A and C, forming one long α-helix. Helix B of the proximal repeat overlaps helix A of the distal repeat, because the connection is tight. Interactions between the two helices appear to restrict the mobility of the repeats at the repeat junction. Several residues appear to be highly conserved in the repeat segments, such as tryptophan at position 45, leucine at position 26, arginine at 22, aspartate at 38, aspartate or glutamate at 41, lysine at 71, histidine at 101, and hydrophobic amino acids at 1, 12, 15, 26, 35, 46, and 68, respectively [11]. Although the presence of homologous 106 amino acid repeats suggests that spectrin evolved from duplication of a single ancestral minigene, the genomic organization of both α- and β-spectrin genes in human beings reveals that the size and position of exons does not necessarily correspond to the structural or conformational unit of spectrin repeats. 1.3 Functions of Red Cell Spectrins

Spectrins are the key protein for the composition of the cytoskeletal network, which regulates cell shape, membrane deformability and stability, and the lateral mobility of band 3 as an integral protein [1–4]. Spectrin has spring-like properties [19].

3


Fig. 2 Demonstration of spectrin dimer (A)and spectrin tetramer (B) by electron microscopy with the shadowing method. By courtesy of the late Professor Jiri Palek.

These important functions of spectrins are mainly mediated by self-association of the spectrins themselves. Spectrin heterodimers (αβ) associate to form spectrin heterotetramers (α 2 β 2 ) and higher oligomers (Fig. 2). On the membrane, tetramer (α 2 β 2 ) is present predominantly. Spectrins stay as the tetramer at a physiological ionic strength and low-temperature (25 ◦ C), whereas they dissociate into dimers at low ionic strength and 37 ◦ C. The equilibrium is kinetically frozen at 0 ◦ C. In normal red cell ghosts, approximately 5% of the extracted spectrin is in the dimeric form and about 50% is in the tetrameric form. The remainder is the spectrin oligomers and the protein complexes of spectrins, actin, protein 4.1, and dematin, which have very high molecular weights. This procedure has been ulitized for elucidating molecular abnormalities of spectrins in the disease states, especially hereditary ellipto-cytosis. For the self-association of spectrins, two mechanisms have been considered: (a) the head-to-head interaction of the spectrins, and (b) their side-by-side interaction (Fig. 3). For the first mechanism, the interconversion of spectrin dimers to tetramers requires a reversible opening of the dimeric bond and formation of two new β attachments. The αβ contact closely resembles the triple-helical structure

Fig. 3 Head-to-head contact of "- and $-spectrins in normal red cells. $-Spectrins are shown as shaded helices, and "-spectrins are denoted as open helices. A polymerization site is composed of the last two helices of $-spectrins and the first one helix of "-spectrin.

1 "- and $-Spectrins

of native spectrin repeats [19, 20]. In the contact site, two of the helices (helices A and B) at the COOH-terminal end of the β-spectrin molecule are bound to one helix (helix C) of the NH2 -terminal end of the α-spectrin molecule [20]. The second mechanism is the side-by-side association of the α- and β-spectrins, which occurs in a zipper-like process. It starts with a defined nucleation site composed of four repeats, that is, α19 to α22 from the α-spectrin, and β1 to β4 from the β-spectrin, respectively [21]. These repeats are located at the tail end of each molecule, to which actin binds. Two of the α repeats and one of the β repeats have eight residue insertions which participate in the interchain interaction [22]. These are located between conformational segments α20/α21 and β2/β3. After the initial tight association of the complementary nucleation sites, a conformational change is initiated that promotes the pairing of the remaining part of the two chains [23]. A common α-spectrin polymorphism, α LELY interferes with normal nucleation and decreases the synthesis of functionally-competent α-spectrin chains [24]. This may influence clinical expression in spectrin mutations [25]. The linkage of spectrins to other red cell membrane proteins is also functionally important. Binding of spectrins to ankyrin requires almost the entire 15th repeat segment and a small portion of the 16th repeat of the β-spectrin molecule [16]. It has been reported that β-spectrin and nonerythroid β-spectrin (β-fodrin) contain an ankyrin-independent site in the NH2 -terminal half of each molecule, which binds to brain membranes. The binding is inhibited by Ca2+ /calmodulin. This site is known as the membrane association domain 1 (MAD1) [26]. β-Fodrin and the muscle/brain isoform of red cell β-spectrin have a second Ca2+ /calmodulinindependent site (MAD2) near to the COOH-terminal end. Another linkage of spectrins to the membrane is mediated by association with the junctional complex that includes spectrin, actin, and protein 4.1, which form a ternary or higher-order complex by linking spectrin tetramers to one another in a tail-to-tail fashion [4]. The actin binding site is located at a region near to the NH2 -terminal end of β-spectrin that contains a 27 amino acid sequence, which is also highly conserved in α-actinin, dystrophin, actin-binding protein (ABP–120), and others. 1.4 Erythroid and Nonerythroid Spectrins

There are a wide variety of spectrin-related proteins in many tissues and animal species [3, 4]. Two closely related, yet distinct mammalian α-subunits of spectrin are known, i.e.: α-spectrin in mature erythroid cells and α-fodrin in all other tissues [27]. In mammals, there are two β-subunits, i.e.: β-spectrin and β-fodrin. A third subunit (β-heavy spectrin) with a molecular weight of 430 kDa is present in Drosophila. β-spectrin exists in both erythroid and muscle/brain isoforms [28–30]. Nonerythroid spectrins are basically composed of two nonidentical, high molecular weight subunits (such as α-spectrin and β-spectrin), which are composed of homologous repeat units of 106 amino acids.

5


Fodrin (synonyms: tissue spectrin, brain spectrin, spectrin II, or spectrin G) is a heterodimer of α- and β-fodrin chains [2]. The biological functions are essentially the same as those in red cell spectrin (spectrin I, or spectrin R). However, there are some differences between them. The α-fodrin has a calmodulin-binding site which is not present in α-spectrin. In addition, β-fodrin contains a pleckstrin homology domain which is a sequence motif in MAD2, but red cell spectrin does not. Erythroid spectrins, nonerythroid spectrins, dystrophin and its homologues, and α-actinin are believed to belong to the so-called spectrin super-family These proteins have the same characteristic features such as the flexible, rod-like shapes of the side-by-side arrangement of the proteins in an antiparallel fashion. They are composed of homologous amino acid repeats of 106, 109, and 120 residues of spectrin, dystrophin, and α-actinin, respectively, which are folded into triple α-helical segments [1, 2]. Amongst these there is homology in the sequence of the triple-helical repeats with conservation of tryptophan at position 45 or 46. The composition of the repeats are homologous mainly between spectrin and dystrophin, whereas that of the actinin repeat is less homologous. The COOH-terminal regions are homologous between α-spectrin, dystrophin, and actinin, which have the potential for Ca2+ -binding, EF hand structures. The NH2 -terminal regions are also homologous in β-spectrin, dystrophin, and α-actinin which demonstrate a potential actin-binding site. Great variability exists with respect to the subcellular localization of erythroid and nonerythroid spectrins [1–4]. The spectrin superfamily proteins are essentially present in the membrane skeleton of the mammalian red cells, but are also expressed in various tissues by developmentally regulated mechanisms, such as neural development, oogenesis, epithelial cell polarity, embryogenesis, viral transformation, the IgG binding to cell surface receptors, and apoptosis. It has been shown that tissue-specific differential modification of the COOH-terminal region of β-spectrin produces various β-spectrin isoforms. The best example is a muscle isoform of β-spectrin [28]. The isoform results from alternative mRNA splicing of the mRNA transcript of the erythroid β-spectrin gene. The splicing alters translation termination near to the COOH-terminus, leading to elongation of the COOH-terminus of the muscle form. At the present time, two α-spectrin isoforms and five β-spectrin isoforms have been reported [29]. A β III isoform is localized on the Golgi apparatus in many cell types [30].

2 Protein 4.1

Protein 4.1 in red cells is a phosphoprotein present in 200 × 103 copies per cell [31, 32]. The cloned protein is globular (5.7 nm diameter) and has a molecular weight of 66 kDa, but migrates as a 78 or 80 kDa protein on SDS–PAGE gels. The protein 4.1 gene (EL1 or EPB41) [33–35] is located on chromosome 1p36.1 [36], which encodes 588 amino acids.

2 Protein 4.1

2.1 Structure of Protein 4.1

Chymotryptic digestion and limited sequencing demonstrate the presence of four domains of red cell protein 4.1 [32]: (1) a 30 kDa domain (residues 1 to ∼300) at the NH2 -terminal end, (2) a 16 kDa domain (residues 300 to 404), (3) a 10 kDa domain (residues 405 to 471), and (4) a 22 to 24 kDa domain (residues 472 to 622) at the COOH end (Fig. 4). In red cells, there are two forms of protein 4.1 with different molecular weights, that is, protein 4.1a (80 kDa) and protein 4.1b (78 kDa). Protein 4.1a, with the higher molecular weight, results from the deamidation of two Asn residues (478 and 502) within the 22 to 24 kDa domain in a non-enzymatic, age-dependent fashion that lowers the mobility of the protein in gels [37]. Therefore, protein 4.1a is hardly apparent in young red cells, and increases as red cells age. Thus, protein 4.1a provides a useful indicator of red cell senescence. Considering the characteristic features of the structure of protein 4.1, clustering of cysteine residues is observed near the NH2 -terminal, and O-linked glycosylation is present in the 10 kDa domain [31]. The glycosylation contributes to its higher apparent molecular weight on polyacrylamide gels than is predicted on the basis of known amino acid sequence. In addition, the NH2 -terminus is definitely basic, whereas the COOH-terminus is clearly acidic. It has been known that red cell protein 4.1 is highly phosphorylated [31]. Phosphorylation sites are located near the COOH-terminal end of the 30 kDa domain and in the middle region of the 10 kDa domain. The latter site appears to be cyclic AMP dependent, but the others may be regulated by protein kinase C. At least one site appears to be a substrate for cdc kinase. 2.2 Binding to Other Membrane Proteins

The most important role of protein 4.1 is in the linkage of the spectrin–actin membrane skeleton to the lipid bilayer by facilitating complex formation between the spectrin–actin fibers [31, 38], the cytoplasmic domain of band 3 [39], p55 [40], and glycophorin C [41]. Protein 4.1 binds tightly (binding coefficient K d ∼ 10−7 M) to β-spectrin very close to the actin-binding site [13]. For this activity, a 21 amino acid within the 10 kDa domain is critical. The interaction of protein 4.1 with spectrin and actin is blocked by protein kinase A phosphorylation at residues in Ser331 in the 16 kDa domain and Ser467 in the 10 kDa domain, and also by tyrosine kinase phosphorylation at Tyr418 in the 10 kDa domain [31]. The ternary complex is also regulated by Ca2+ and calmodulin [42]. Calmodulin binds to a site within the 30 kDa domain of protein 4.1 in a Ca2+ -independent manner (K d ∼ 5 × 10−7 M). The 30 kDa domain of erythroid protein 4.1 is involved in its binding to proteins such as p55, glycophorin C, band 3, and other embedded moieties, e.g., the chloride channel pICln [43]. Protein 4.1 binds directly to glycophorin C to a site near amino

7


Fig. 4 Molecular structure of protein 4.1. A schematic structure of protein 4.1 molecule is shown in (I), and the protein 4.1 mRNA is demonstrated in (II). The details are given in text.

2 Protein 4.1

acids 82 through 98 in the tail of this molecule [41]. It also binds to a more distal portion near residues 112 through 128, by means of the protein p55 [40]. The role of the interaction between protein 4.1 and glycophorin C is well established. Protein 4.1 deficient red cells are also deficient in glycophorin C, but not glycophorin A or band 3. The residual glycophorin C that is only loosely bound to the skeleton becomes tightly bound after protein 4.1 is reconstituted. Both protein 4.1 and glycophorin C bind p55, which enhances or regulates their interaction. The mechanical weakness of glycophorin C-deficient membranes is totally restored by restoration of protein 4.1 or its 10 kDa spectrin–actin binding domain. Under the physiological condition in vivo, approximately 40% of protein 4.1 is bound directly to glycophorin C, 40% is bound indirectly through the protein p55, and 20% is bound to band 3. 2.3 Extensive Alternative Splicings

It is known that protein 4.1 is extremely heterogeneous with respect to molecular weight, abundance, and intracellular localization [31]. Red cell protein 4.1 (P4.1R) isoforms come from a single genomic locus near the Rh locus at chromosome 1p36.1 [36]. Its primary mRNA transcript from the gene (250–300 kb long) is subjected to extensive alternative splicing, producing a diverse protein family through various mRNA (6.5 to 7.0 kb long) [44] (Fig. 5). At the present time, 12 alternatively spliced exons, as well as an important cryptic acceptor site, are known. Protein 4.1 utilizes two different initiation codons, termed “upstream” and “downstream” initiation codons, which translate to isoforms of 135 and 85 kDa, respectively [31]. The 85 kDa isoform is created by the splicing out of a 17 nucleotide motif that contains the upstream translation initiator. In most tissues, this sequence is spliced in, and a downstream 80 base pair motif is spliced out, producing the elongated 135 kDa isoform that contains an additional 209 amino acids attached to the NH2 -terminus of the shorter isoform of protein 4.1. This is because the upstream translation start site, when present, is used almost exclusively [33]. The 85 kDa shorter isoform which is encoded by the downstream initiation codon is found primarily in red cells. In erythroid protein 4.1 (P4.1R), splicings appear to be critical for red cell functions in only two regions, i. e., at the 10 kDa spectrin–actin binding domain, and in the 5 -untranslated region [31]. At the 10 kDa domain, three alternatively spliced exons are located at the 5 -end of this domain, which is the spectrin–actin binding domain. Exon 16, which is one of the three exons and is 63 nucleotides long, encodes for 21 amino acids at the 5 -terminus. This exon 16 is also expressed in muscle and testis [45], which are actually nonerythroid tissues. Two other exons (exons 14 and 15) are adjacent to the one responsible for binding to spectrin–actin. In red cells, the protein 4.1 mRNA contains only exon 16 out of these three exons [33, 46], whereas lymphocytes have

9


Fig. 5 Extensive splicing of the protein 4.1 gene. Complicated splicings are observed around the regions of exons 3 to 5, 14 to 16, and 17A to 20.

2 Protein 4.1

none of the exons, and the brain retains all three exons [45]. Therefore, exon 16 appears to provide erythroid and stage-specific expression.

2.4 Nonerythroid Protein 4.1 Isoforms

Multiple alternative mRNA splicing events generate isoforms with different affinities for membrane and intracellular structures. The high (120–150 kDa) and low molecular weight isoforms (60–90 kDa) coexist in varying ratios in most nonerythroid tissues. Although the functional importance of the high molecular weight isoforms remains unknown, their major part of protein 4.1, which is 80 kDa upstream from the COOH-terminus, is identical to their low molecular weight analogues. Therefore, the additional amino acid residues in the high molecular weight isoforms are equivalent to a headpiece [31, 44]. It is interesting to note that high molecular weight isoforms associate with NuMa (a major organizing protein of the mitotic apparatus) [40, 47, 48], ZO-2 (a component of the tight junctions in epithelial and endothelial cells), and other membrane associated guanylate kinase (MAGUK) family proteins, in addition to p55 [40, 49, 50]. The diverse structure of these spliced isoforms appears to provide for the multiple subcellular locations for their specific functions. Although these isoforms demonstrate their widespread expression and interaction with key components, such as mitotic spindles and tight junctions, there is a surprising fact that erythroid protein 4.1 gene-targeted knockout mice exhibit only anemia and subtle neurological abnormalities [51]. As regards the high molecular weight isoforms, their junctional significance is still unknown [31]. The high molecular weight isoforms are completely absent from mature red cells. mRNA containing the 17 amino acid extension of exon 2, which is required for the synthesis of high molecular weight forms, is essentially absent from early erythroid precursors (younger than proerythroblasts) [33, 45]. Only the downstream translation initiation site is utilized for erythroid protein 4.1 biosynthesis during erythropoiesis. It is now clear that protein 4.1 and its family analogues demonstrate a mosaic of sequence homologues with other proteins [52]. A typical example is the 20–45% homology, which is observed at the NH2 -terminus of protein 4.1 family proteins. The FERM (Four. 1 protein, Ezrin, Radixin, and Moesin) domain, which is present at the NH2 -terminus, defines a family of proteins linking the cytoskeleton including actin to membranes in various cells [53, 54]. The FERM has previously been known as a family of actin binding proteins (moesin, ezrin, radixin, and merlin), which are also homologous with coraclin. This is a Drosophila protein that interacts with a large disk protein (Dlg) homologous with tumor suppressor proteins in mammals [55]. Erythroid protein 4.1 (P4.1R) is the prototype of a much more closely related family of at least four members, such as, P4.1 of a generally distributed type (P4.1G), that of a brain type (P4.1B), or that expressed predominantly in neuron (P4.1N). The extent of the homology among these isoforms is 60–95% [56, 57].

11


3 Actin

Red cell actin (β-actin), which is a 43 kDa protein, is present in abundance (400–500 × 103 copies per red cell). The red cell actin content is 5.5% (w/w) of the total red cell membrane proteins [58–60]. The β-actin gene (ACTB) is located on the chromosome 7 (7p12–p22) encoding 375 amino acids. Red cell actin is similar to other actins in its structure and functions. Although the β-actin is distributed in various nonmuscle cells including red cells, red cell actin is organized as short, double-helical F-actin protofilaments 12 to 13 monomers long and approximately 35 nm in length. Red cell actin interacts with spectrins, adducin, protein 4.1, and tropomyosin for the sake of their stability [38, 58, 61, 62]. The actin also binds to tropomodulin by capping of the slow growing or pointed end of the actin filament [63]. The state of actin polymerization is functionally important to red cell membrane flexibility, which increases when actin polymerization is inhibited. It is also true that increased polymerization of actin makes the red cell membrane more rigid. Spectrin dimers bind to the side of actin filaments at a site near the tail end of the spectrin molecule. On average, six spectrin ends make a complex with each actin oligomer, producing an irregular network, which is approximately hexagonal [60]. Each spectrin–actin junction is stabilized by the formation of a ternary complex with protein 4.1 [38].

4 Other Minor Skeletal Proteins 4.1 The p55 Protein

The p55 protein is a phosphoprotein member of the MAGUK (membraneassociated guanylate kinase) family of proteins [40, 58]. The protein is a 55 kDa skeletal protein, which has 80 × 10 copies per red cell. The p55 gene (MPP1) is located on the X chromosome (Xq28), which encodes 466 amino acids. The protein p55 is the human homolog of dlg, a Drosophila tumor suppressor gene [40]. This protein is composed of three domains: (1) an NH2 -terminal domain of unknown function that is also present in dlg, (2) a central SH3 domain that is embedded in the MAGUK domain, and (3) a COOH-terminal guanylate kinase domain [64, 65]. The protein appears to be present in a dimeric form, extensively palmitoylated, and tightly bound to the membrane [66]. Homologues of the p55 include signal transduction proteins, tumor suppressor genes, and proteins important in cell-to-cell interactions. This protein is expressed throughout erythroid differentiation and is widely expressed in nonerythroid tissues. The p55 binds to the 30 kDa domain of protein 4.1 through a 39 amino acid binding motif in the COOH-terminal MAGUK domain and to the cytoplasmic tail of glycophorin C by means of its single PDZ motif. The p55 binds to the 30 kDa

4 Other Minor Skeletal Proteins 13

domain of protein 4.1 (K d ∼2 × 10−9 M) and to the cytoplasmic domain of glycophorin C (K d ∼7 × 10−9 M). In the disease states, patients who are actually deficient in either protein 4.1 or glycophorin C also lack the p55 [67]. 4.2 Adducin

Adducin, which is a Ca2+ /calmodulin-binding phosphoprotein, is composed of αβ-adducin heterodimers [68, 69]. This protein is located at the spectrin–actin junctional complex [58]. The α and β adducin are structurally similar proteins encoded by separate genes. A theoretical molecular weight of α-adducin, which is calculated from the results of gene sequencing, is 81 kDa, 1% of the total red cell membrane proteins, and with 30 × 103 copies per red cell. However, the actual molecular weight of this protein is 103 kDa on the SDS-PAGE gels, which is quite larger than the theoretical one, because some other components (such as glycosylated chains) are present on this molecule. β-Adducin (80 kDa) is 97 kDa on the SDS-PAGE gels, 1% of the total red cell membranes proteins, and with 30 × 103 copies per red cell. The α-adducin gene (ADDA) is located on chromosome 4 (4p16.3), which encodes 737 amino acids. The β-adducin gene (ADDB) is located on chromosome 2 (2p13–2p14), which encodes 726 amino acids. A third nonerythroid adducin (γ -adducin) is also known. The subunits have three domains: (1) an NH2 -terminal domain (39 kDa) with a globular head, (2) a 9 kDa domain of the connecting neck, and (3) a protease-sensitive domain (33 kDa) with an extended COOH-terminal tail [68, 70]. The last domain contains mainly hydrophilic residues and 22 amino acid segments homologous with the MARKS phosphorylation domain that regulates Ca2+ /calmodulin-regulated capping and bundling of actin filaments [71]. The four head domains cluster in a globular core, and the tail domains extend to interact with spectrins and actin. Adducin increases the binding of spectrin to actin, just like protein 4.1, whereas adducin does not interact directly with spectrin in the absence of actin, unlike protein 4.1. The tails of both α-and β-adducin bind to the actinbinding domain at the N-terminus of β-spectrin, and to the second spectrin repeat [72]. Adducin is expressed at the stage of erythroblasts, but is only incorporated into the red cell membrane structure at a late stage in erythroid development [73]. Adducin contributes to the early assembly of the spectrin–actin complex, which is regulated by phosphorylation of the COOH-terminal domain of adducin by protein kinase C [74, 75]. In this event, adducin does not bind directly to spectrin in the absence of actin. Targeted inactivation of β-adducin in mice produced compensated spherocytic anemia and neurologic abnormalities [76]. For this assembly of a spectrin–actin–adducin ternary complex, the actin-binding domain of β-spectrin and the first two spectrin repeats are required. Adducin is also present in various nonerythroid cells [68], especially at sites of cell-to-cell contact. The protein assembles at these sites in response to extracellular Ca2+ , and dissociates when phosphorylation of the protein is activated by protein kinase C.


4.3 Dematin (Protein 4.9)

Dematin has been known to be associated with actin in erythroid and nonerythroid cells [58]. Human red cell dematin consists of two chains of 48 kDa and 52 kDa, which are present in a ratio of 3 (48 kDa) to 1 (52 kDa). The native protein is a trimer [77]. There are 40 × 103 trimeric copies per cell, and the content of dematin is approximately 1% of the total red cell membrane proteins. The dematin gene (EPB 49) is located on chromosome 8 (8p21.1), which encodes 383 amino acids [77, 78]. Dematin has two binding sites for β-actin, and bundles actin filaments into cables, which is inhibited by phosphorylation with protein kinase A, but not with protein kinase C. The COOH-terminal half of the 48 kDa subunit is similar to villin, which is known as an actin-binding protein that induces growth of microvilli and reorganizes actin filaments in brush borders. The 52 kDa subunit of dematin has an additional 22 amino acid sequence in the C-terminal domain of the 48 kDa subunit [77]. This sequence resembles that in protein 4.2 [79]. Dematin appears to attach to a lipid or integral membrane protein, since it remains associated with the membrane during the extraction of other skeletal proteins. 4.4 Tropomyosin

It is known that tropomyosin is also associated with actin [58]. Red cell tropomyosin is a heterodimer of 27 kDa and 29 kDa subunits on SDS–PAGE gels, on which tropomyosin migrates in the region of band 7 [80, 81]. There are 80 × 103 copies per cell, and its content is approximately 1% of the total red cell membrane proteins. The tropomyosin gene (TPM 3) is located on chromosome 1 (1q31), which encodes 239 amino acids. Stoichiometrically, one copy of tropomyosin binds to every six to eight actin monomers, which is just sufficient to line both grooves of the actin protofilament. The function of tropomyosin appears to be for stabilizing the short erythroid actin filaments and to help spectrin–actin interactions [82]. 4.5 Tropomodulin

Tropomodulin is a 41 kDa protein that binds to tropomyosin in a 2:1 molar ratio with a K d of 5 × 10−7 M [83]. Each protein binds to the NH2 -terminal region of the other. The molecular size of tropomodulin is 43 kDa on SDS–PAGE gels [58]. There are 30 × 103 copies per cell. The tropomodulin gene (TMOD) is located on chromosome 9 (9q22), which encodes 359 amino acids. This protein also binds to actin. Capping is enhanced when the grooves of the actin filament are lined with tropomyosin. Tropomodulin appears to have a tight association with the membrane.

References

4.6 Other Membrane Proteins

Myosin [58, 84, 85] and caldesmon (a 71 kDa calmodulin-binding protein) [86] are known as minor components of the red cell membrane proteins, although their physiological functions have not been elucidated in detail.

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16 KENNEDY, S. P., WARREN, S. L., FORGET, B. G., MORROW, J. S. (1991) Ankyrin binds to the 15th repetitive unit of erythroid and nonerythroid β-spectrin. J. Cell Biol. 115: 267–277. 17 YAN, Y., WINOGRAD, E., VIEL, A., CRONIN, T., HARRISON, S. C., BRANTON, D. (1993) Crystal structure of the repetitive segments of spectrin. Science 262: 2027–2030. 18 WINOGRAD, E., HUME, D., BRANTON, D. (1991) Phasing the conformational unit of spectrin. Proc. Natl. Acad. Sci. USA 88: 10788–10791. 19 ALTMANN, S. M., GRU¨ NBERG, R. G., LENNE, P.-F., YLANNE, J., RAAE, A., HERBERT, K., SARASTE, M., NILGES, M. HORBER, J.K. (2002) Pathways and intermediates in forced unfolding of spectrin repeats. Structure 10: 1085–1096. 20 SPEICHER, D. W., DESILVA, T. M., SPEICHER, K. D., URSITTI, J. A., HEMBACH, P., WEGLARZ, L. (1993) Location of the human red cell spectrin tetramer binding site and detection of a related “closed” hairpin loop dimer using proteolytic footprinting. J. Biol. Chem. 268: 4227–4235. 21 SPEICHER, D. W., WEGLARZ, L., DESILVA, T. M. (1992) Properties of human red cell spectrin heterodimer (side-to-side) assembly and identification of an essential nucleation site. J. Biol. Chem. 267: 14775–14782. 22 URSITTI, J. A., KOTULA, L., DESILVA, T. M., CURTIS, P. J., SPEICHER, D. W. (1996) Mapping the human erythrocyte beta-spectrin dimer initiation site using recombinant peptides and correlation of its phasing with the alpha-actinin dimer site. J. Biol. Chem. 271: 6636–6644. 23 VIEL, A., GEE, M. S., TOMOOKA, L., BRANTON, D. (1998) Motif involved in interchain binding at the tail-end of spectrin. Biochim. Biophys. Acta 1384: 396–404. 24 WILMOTTE, R., HARPER, S. L., URSITTI, J. A., MARECHAL, J., DELAUNAY, J., SPEICHER, D. W. (1997) The exon 4b-encoded sequence is essential for stability of human erythroid alpha-spectrin and heterodimer formation. Blood 90: 4188–4196.

25 ALLOISIO, N., MORLE´ , L., MARE´ CHAL, J., ROUX, A. F., DUCLUZEAU, M. T., GUETARNI, D., POTHIER, B., BAKLOUTI, F., GHANEM, A., KASTALLY, R., DELAUNAY, J. (1991) Spα V/4I : A common spectrin polymorphism at the α IV -α V domain junction. Relevance to the expression level of hereditary elliptocytosis due to α-spectrin variants located in trans. J. Clin. Invest. 87: 2169–2177. 26 LOMBARDO, C. R., WEED, S. A., KENNEDY, S. P., FORGET, B. G., MORROW, J. S. (1994) βII-spectrin (fodrin) and βI 2-spectrin (muscle) contain NH2 - and COOH-terminal membrane association domains (MAD1 and MAD2). J. Biol. Chem. 269: 29212–29219. 27 LETO, T. L., FORTUGNO-ERIKSON, D., BARTON, D., YANG-FENG, T. L., FRANCKE, U., HARRIS, A. S., MORROW, J. S., MARCHESI, V. T., BENZ, E. J., Jr., (1988) Comparison of nonerythroid α-spectrin genes reveals strict homology among diverse species. Mol. Cell Biol. 8: 1–9. 28 WINKELMANN, J. C., COSTA, F. F., LINZIE, B. L., FORGET, B. G. (1990) β-Spectrin in human skeletal muscle: Tissue-specific differential processing of 3 β-spectrin pre-mRNA generates a P-spectrin isoform with a unique carboxyl terminus. J. Biol. Chem. 265: 20449–20454. 29 STABACH, P. R., MORROW, J. S. (2000) Identification and characterization of βV spectrin, a mammalian ortholog of Drosophila βH spectrin. J. Biol. Chem. 275: 21385–21395. 30 STANKEWICH, M. C., TSE, WT., PETERS, L. L., CH’NG, Y., JOHN, K. M., STABACH, P. R., DEVARAJAN, P., MORROW, J. S., LUX, S. E. (1998) A widely expressed beta-III spectrin associated with Golgi and cytoplasmic vesicles. Proc. Natl. Acad. Sci. USA 95: 14158–14163. 31 CONBY, J. G. (1993) Structure, function, and molecular genetics of erythroid membrane skeletal protein 4.1 in normal and abnormal red blood cells. Semin. Hematol. 30: 58–73. 32 LETO, T. L., MARCHESI, V. T. (1984) A structural model of human erythrocyte protein 4.1. J. Biol. Chem. 259: 4603–4608. 33 HUANG, J. P., TANG, C. J., KOU, G. H., MARCHESI, V. T., BENZ, E. J. Jr., TANG, T.

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K. (1993) Genomic structure of the locus encoding protein 4.1. Structural basis for complex combinational patterns of tissue-specific alternative RNA splicing. J. Biol. Chem. 268: 3758–3766. SCHISCHMANOFF P. O., YASWEN, P., PARRA, M. K., LEE, G., CHASIS, J. A., MOHANDAS, N., CONBOY, J. G., (1997) Cell shape-dependent regulation of protein 4.1 alternative pre-mRNA splicing in mammary epithelial cells. J. Biol. Chem. 272: 10254–10259. BAKLOUTI, F., HUANG, S. C., VULLIAMY, TJ., DELAUNAY, J., BENZ, E. J. Jr., (1997) Organization of the human protein 4.1 genomic locus: New insights into the tissue-specific alternative splicing of the pre-mRNA. Genomics 39: 289–302. KIM, A. C., Van HUFFEL, C., LUTCHMAN, M., CHISHTI, A. H. (1998) Radiation hybrid mapping of EPB41L1, a novel protein 4.1 homologue, to human chromosome 20q11.2–q12. Genomics 49: 165–166. INABA, M., GUPTA, K. C., KUWABARA, M., TAKAHASHI, T., BENZ, E. J. Jr., MAEDE, Y. (1992) Deamidation of human erythrocyte protein 4.1: Possible role in aging. Blood 79: 3355–3361. GIMM, J. A., AN, X., NUNOMURA, W., MOHANDAS, N. (2002) Functional characterization of spectrin-actinbinding domains in 4.1 family of proteins. Biochemistry 41: 7275–7282. AN, X. L., TAKAKUWA, Y., NUNOMURA, W., MANNO, S., MOHANDAS, N. (1996) Modulation of band 3-ankyrin interaction by protein 4.1. Functional implications in regulation of erythrocyte membrane mechanical properties. J. Biol. Chem. 271: 33187–33191. CHISHTI, A. H. (1998) Function of p55 and its nonerythroid homologues. Curr. Opin. Hematol. 5: 116–121. HEMMING, N. J., ANSTEE, D. J., STARICOFF, M. A., TANNER, M. J., MOHANDAS, N. (1995) Identification of the membrane attachment sites for protein 4.1 in the human erythrocyte. J. Biol. Chem. 270: 5360–5366. TAKAKUWA, Y., MOHANDAS, N. (1988) Modulation of erythrocyte membrane material properties by Ca2+ and

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G., (1999) Protein 4.1R-deficient mice are viable but have erythroid membrane skeleton abnormalities. J. Clin. Invest. 103: 331–340. TURUNEN, O., SAINIO, M., JAASKELAINEN, J., CARPEN, O., VAHERI, A. (1998) Structure-function relationships in the ezrin family and the effect of tumor-associated point mutations in neurofibromatosis 2 protein. Biochim. Biophys. Acta. 1387: 1–16. TSUKITA, S., YONEMURA, S. (1997) ERM (ezrin/radixin/moesin) family: from cytoskeleton to signal transduction. Curr. Opin. Cell Biol. 9: 70–75. BRETSCHER, A., RECZEK, D., BERRYMAN, M. (1997) Ezrin: a protein requiring conformational activation to link microfilaments to the plasma membrane in the assemble of cell surface structures. J. Cell Sci. 110: 3011–3018. WARD, R. E. 4th, LAMB, R. S., FEHON, R. G. (1998) A conserved functional domain of Drosophila coracle is required for localization at the septate junction and has membrane-organizing activity. J. Cell Biol. 140: 1463–1473. PARRA, M., GASCARD, P., WALENSKY, L. D., SNYDER, S. H., MOHANDAS, N., CONBOY, J. G. (1998) Cloning and characterization of 4.1G (EPB41L2), a new member of the skeletal protein 4.1 (EPB41) gene family. Genomics 49: 298–306. WALENSKY, LD., GASCARD, P., FIELDS, M. E., BLACKSHAW, S., CONBOY, J. G., MOHANDAS, N., SNYDER, S. H. (1998) The 13-kD FK 506 binding protein, FKBP13, interacts with a novel homo-logue of the erythrocyte membrane cytoskeletal protein 4.1. J. Cell Biol. 141: 143–153. GILLIGAN, D. M., BENNETT, V. (1993) The functional complex of the membrane skeleton. Semin. Hematol. 30: 74–83. PINDER, J. C., GRATZER, W. B., (1983) Structural and dynamic states of actin in the eythrocyte. J. Cell Biol. 96: 768–775. BYERS, T. J., BRANTON, D. (1985) Visualization of the protein associations in the erythrocyte membrane skeleton. Proc. Natl. Acad. Sci. USA 82: 6153–6175. KUHLMAN, P. A., HUGHES, C. A., BENNETT, V., FOWLER, V. M. (1996) A new function for adducin.

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Calcium/calmodulin-regulated capping of the barbed ends of actin filaments. J. Biol. Chem. 271: 7986–7991. PICART, C., DISCHER, D. E. (1999) Actin protofilament orientation at the erythrocyte membrane. Biophys. J. 77: 865–878. FOWLER, V. M., SUSSMANN, M. A., MILLER, P. G., FLUCHER, B. E., DANIELS, M. P. (1993) Tropomodulin is associated with the free (pointed) ends of the thin filaments in rat skeletal muscle. J. Cell Biol. 120: 411–420. KIM, A. C., METZENBERG, A. B., SAHR, K. E., MARTATIA, S. M., CHISHTI, A. H. (1996) Complete genomic organization of the human erythroid p55 gene (MPP1), a membrane-associated guanylate kinase homologue. Genomics 31: 223–229. MARFATIA, S. M., MORAIS-CABRAL, J. H., KIM, A. C., BYRON, O., CHISHTI, A. H. (1997) The PDZ domain of human erythrocyte p55 mediates its binding to the cytoplasmic carboxyl terminus of glycophorin C. Analysis of the binding interface by in vitro mutagenesis. J. Biol. Chem. 272: 24191–24197. RUFF, P., SPEICHER, D. W., HUSAIN-CHISHTI, A. (1991) Molecular identification of a major palmitoylated erythrocyte membrane protein containing the src homology 3 motif. Proc. Natl. Acad. Sci. USA 88: 6595–6599. ALLOISIO, N., DALLA VENEZIA, N., RANA, A., ANDRABI, K., TEXIER, P., GILSANZ, F., CARTRON, J. P., DELAUNAY, J., CHISHTI, A. H. (1993) Evidence that red blood cell protein p55 may participate in the skeleton-membrane linkage that involves protein 4.1 and glycophorin C. Blood 82: 1323–1327. JOSHI, R., GILLIGAN, D. M., OTTO, E., MCLAUGHLIN, T., BENNETT, V. (1991) Primary structure and domain organization of human α and β adducin. J. Cell Biol. 115: 665–675. KATAGIRI, T., OZAKI, K., FUJIWARA, T., SHIMIZU, F., KAWAI, A., OKUNO, S., SUZUKI, M., NAKAMURA, Y., TAKAHASHI, E., HIRAI, Y. (1996) Cloning, expression and chromosome mapping of adducinlike 70 (ADDL), a human cDNA highly homologous to human erythrocyte

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19

1

Integral Proteins of the Erythrocyte Membrane Yoshihito Yawata



1 Band 3

Band 3 is the major integral protein of the red cells, and is also known as anion exchanger-1 (AE1) [1–4]. This protein is a transmembrane glycoprotein with a molecular mass of about 100 kDa, and comprises 25 to 30% (w/w) of the total membrane proteins. There are about 1200 × 103 copies per cell. Band 3 migrates as a diffuse band on SDS–PAGE gels due to heterogeneous glycosylation [5, 6]. The human band 3 cDNA is about 4.7 kb in length and encodes a 911 amino acid polypeptide [7, 8]. The gene (EPB3) is located on chromosome 17 (17q21–q22). Structure and functional interpretation of the cytoplasmic domain of erythrocyte membrane band 3 have recently been shown crystallographically [9]. 1.1 Structure of Band 3

Band 3 is composed of the cytoplasmic (NH2 -terminal) domain and helices and β-sheets to form the transmembrane (COOH-terminal) domain [1–5] (Fig. 1). The cytoplasmic (NH2 -terminal) domain of band 3 is composed of the first 403 amino acids. The major part of this domain (amino acids 1 to 359) is released from the membrane by treatment with chymotrypsin or trypsin as a 43 kDa fragment [10]. The two regions can be separated by chymotrypsin cleavage at the inner membrane (position 359). A second chymotryptic site is accessible at the external surface at position 553. This domain is an elongated, water-soluble, 403 amino acid segment with a flexible proline-rich hinge near the center, which is located between amino acids 175 and 190 of this domain. The first 45 amino acids in the cytoplasm are highly acidic. The reminder of the domain is fairly mobile. It extends at a high pH and contracts at a low pH [5]. Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Integral Proteins of the Erythrocyte Membrane

Fig. 1 Schematic model for the molecular structure of human erythroid band 3. Band 3 molecules are composed of their cytoplasmic and transmembrane domains. The cytoplasmic domain demonstrates many binding sites to membrane proteins and

cytoplasmic proteins. The membrane domain contains anion transport sites, such as Lys539, Glu681, and Lys851. Asn642 in the band 3 molecule is the binding site for a sugar chain.

The transmembrane (COOH-terminal) domain (amino acids 404 to 911), which is composed of approximately 52 kDa (17 + 35 kDa), folds into helices and β-sheets to form 12 to 14 membrane-spanning segments containing the anion transport channels. The boundary between the NH2 -terminal (cytoplasmic) tail and the first membrane-spanning segment (amino acids 400 to 404) is highly conserved in red cells of various species [7, 9]. The protein at position 403 is particularly important for creating a β bend or a random coil at the membrane junction, which is known as an inter-domain hinge and gives the tail freedom of movement [11]. 1.2 Functions of Band 3 1.2.1 Membrane Protein Binding by the Cytoplasmic Domain of Band 3 The cytoplasmic (43 kDa) domain of band 3 plays a centrol role in the attachment of the cytoskeleton to the plasma membrane. The inter-domain hinge at this attachment point is important for the flexibility and rigidity of red cells. Three peripheral membrane proteins (ankyrin [12–15], protein 4.1 [16, 17], and protein 4.2 [18]) bind to the cytoplasmic domain of band 3. The ankyrin-binding involves several regions (proximal, middle, and distal) scattered throughout the cytoplasmic domain, implying that the cytoplasmic domain has a complex folded structure. Ankyrin

1 Band 3 3

binds to the flexed conformations of band 3, most probably to band 3 tetramers. It has been suggested that protein 4.1 and protein 4.2 have more than one site of attachment. Protein 4.1 also binds to peptides containing clustered basic residues (LRRRYand IRRRY) that are located at the COOH-terminal end of this domain [17]. Some of the band 3 gene mutations (band 3 Tuscaloosa, band 3 Montefiore, band 3 Fukuoka, and band 3 Coimbra) are known to demonstrate a markedly decreased protein 4.2 content. Band 3 is associated with glycophorin A (GPA) in the membrane [19, 20]. GPA facilitates translocation of band 3 from the endoplasmic reticulum as the site of synthesis of band 3 to the plasma membrane of Xenopus oocytes. Band 3 is critical for GPA synthesis or its stability, since GPA is also deficient in the band 3-deficient red cells [21]. Glycophorins do not appear to be a determinant for the expression of band 3, because when there is a combined deficiency of glycophorin A and B (a glycophorin Mk Mk phenotype) the red cells demonstrate normal amounts of band 3. There is additional evidence for the interaction of band 3 with GPA. The Wright (Wrb ) antigen is caused by interaction of a site on band 3 (Glu658 ) with a site or sites located near the end of the extracellular domain of GPA or in the adjacent transmembrane domain [22]. A monoclonal antibody to the Wrb blood type antigen immunoprecipitates both band 3 and GPA [23]. Furthermore, antibodies to glycophorin decrease the lateral and rotational mobility of band 3, indicating the presence of an interaction of band 3 with glycophorin. Self-association of band 3 is one of its major functions in red cell biology. Band 3, which is extracted from the membrane by a nonionic detergent (octaethylene glycol n-dodecyl monoether), exists as stable dimers (70%), tetramers, and higher-order oligomers (30%) [24]. Tetramers of band 3 are associated with the membrane skeleton. Since isolated membrane domains only form dimers, tetramers appear to be assembled by cross-linking neighboring dimers through the cytoplasmic domain, with ankyrin, or hemichromes. The most efficient physiological functional form in which interactions with cytoplasmic molecules occur appears to be tetrameric. 1.2.2 Binding to Glycolytic Enzymes by the Cytoplasmic Domain of Band 3 The acidic NH2 -terminal sequence has the binding sites for the glycolytic enzymes, that is, glyceraldehyde-3-phosphate dehydrogenase (G-3-PD) [25], phosphoglycerate kinase (PGK) [26], and aldolase [27]. These enzymes are basically cytosolic enzymes, but about 65 % of G-3-PD, 50% of PGK, and 40% of aldolase are bound to band 3 in the intact red cells. The enzymatic activities of these three enzymes are inhibited by membrane attachment, which are regulated by substrates, cofactors, inhibitors, and also phosphorylation of tyrosine 8 [28]. The phosphorus is added by the kinase p72syk and may be removed by a phosphotyrosine phosphatase which is bound to band 3. 1.2.3 Binding to Hemoglobin by the Cytoplasmic Domain of Band 3 The cytoplasmic domain of band 3 also binds hemoglobin. Deoxyhemoglobin binds better than the oxyhemoglobin. As regards this binding, the first five to seven amino acids of band 3 at the NH2 -terminal end are inserted deep into the


2,3-diphosphoglycerate (2,3-DPG)-binding cleft of hemoglobin [29]. Therefore, 2,3DPG inhibits deoxyhemoglobin binding by band 3. Under physiological conditions, approximately half the band 3 molecules have hemoglobin attached. A denatured form of hemoglobin binds better and polymerizes with band 3, forming an aggregate of hemichromes and band 3 [30]. This binding of band 3 with hemichromes appears to stimulate aggregation into patches, which are uniquely recognized by a red cell senescence isoantibody leading to opsonization of the cell and its removal from circulation by the spleen. This mechanism may be one of the important events in red cell ageing. 1.2.4 Anion Exchange Channel by the Transmembrane Domain of Band 3 The transmembrane (52 kDa) domain of band 3 (amino acids 404–911) contains an anion exchange channel [1, 31–34]. The transport capacity is evaluated as being from 1010 to 1011 bicarbonate and chloride anions per second by 1200 × 103 molecules of band 3 per red cell. The bicarbonate is produced by carbonic anhydrase in the red cells. Most of the bicarbonate produced appears to have originated through this channel. In addition, approximately 60 % of CO2 transport from the tissues to the lungs appears to be handled in this way. The hydrogen (H+ ) byproduct by carbonic anhydrase binds to hemoglobin and facilitates oxygen release to the tissues [33]. All of these reactions are reversed in the lungs. The basic structural unit in both the inward- and outward-facing transporters is an oblong band 3 dimer. At the center of the dimer, the two monomers are located close together to form a channel. There is also a flexible subdomain on the far side of each monomer, which might be formed by one or more of the large cytoplasmic loops between transmembrane helices. The anion exchange itself appears to occur through a ping-pong mechanism. An intracellular anion enters the transport channel and is translocated outward and released, with the channel remaining in the outward conformation until an extracellular anion enters and triggers the reverse cycle [1]. It has recently been shown that glutamic acid (Glu681 ) plays the key role in chloride translocation by human band 3, that Glu681 is located near the carboxy terminus of transmembrane segment 8 (TM8) at the region of the chloride channel, and that Lys851 , Ser852 , and Ala858 are also required for this channel [34]. Anion transport requires a high energy and volume of activation. Anion exchange is extremely rapid with a half-life of 50 ms for chloride and bicarbonate, and the specificity of the channel is fairly broad. At much slower rates, large anions (sulfate, phosphate, phosphoenol pyruvate, and superoxide) are also transported through this channel [1, 2]. 1.2.5 Lateral and Rotational Mobility of Band 3 Lateral mobility of band 3 in normal human red cell membranes is constrained by steric hindrance interactions, low affinity binding interactions, and highaffinity binding interactions [35–37]. Steric hindrance interactions between band 3 oligomers and the spectrin-based membrane skeleton put a major constraint on the laterally mobile band 3 fraction, slowing the rate of band 3 lateral diffusion by approximately 50-fold compared with the predicted diffusion rate of free band

1 Band 3 5

3 in membranes devoid of a functional membrane skeleton. The spectrin/band 3 ratio is the major determinant of the lateral diffusion rate of band 3. Fluorescence recovery after use of the photobleaching (FRAP) method (Fig. 2) demonstrates that, in normal red cells, approximately one-third of band 3 is present in the mobile fraction and the remaining two-thirds exist in the immobile fraction, which is fixed to the cytoskeletal network, in particular to ankyrin [35–37]. It has been reported that the mobile fraction was 0.43 ± 0.11 with a lateral diffusion coefficient of (6.86 ± 1.37) × 10−11 cm2 s−1 in normal red cells by the FRAP method. Rotational mobility of band 3 in normal human red cell membranes is constrained by low-affinity and high-affinity binding interactions. The rotationally immobile band 3 fraction apparently represents individual band 3 molecules bound with high affinity to ankyrin. The rapidly rotating band 3 fraction consists of dimers, tetramers, and higher order oligomers of band 3 that are free from rotational constraints other than the viscosity of the lipid bilayer. The slowly rotating band 3 fraction is less well-defined. Rotational constraints applied by low-affinity binding interactions between ankyrin-linked band 3 and other band 3 molecules, and between the cytoplasmic domain of band 3 and membrane skeletal proteins (ankyrin, protein 4.1 and protein 4.2) have been invoked. 1.2.6 Blood Type Antigens and Band 3 On the transmembrane domain of band 3, a complex carbohydrate structure is attached to Asn642 [1, 2, 20]. Within the carbohydrate structure, N-acetylglucosamine, mannose, galactose and other moieties exist. Band 3 carries various polymorphic peptide epitopes including band 3 Memphis (Lys → Glu at position 56), such as the Diego blood group system, Wright (Wra and Wrb ) antigens and several other low frequency antigens [38]. No apparent consequence for band 3 function is observed in these antigenic variants. The Diego (Dia ) allele is observed with varying frequencies in Asian and South American populations, although this allele is exceedingly rare in most Caucasians [20]. Dia and Dib antigens correspond to a proline or a leucine residue at position 854 of the band 3 molecule, respectively. Among the Wright alleles, the Wrb allele is predominantly expressed (higher than 99 %), whereas the Wra allele is extremely rare [20, 22, 23]. Wra and Wrb antigens correspond to a lysine or a glutamine residue, respectively, encoded by codon 658 of band 3. The expression of Wrb is suppressed in glycophorin A deficiency. Band 3 also carries several minor antigens, such as Waldner (Wda ), Redelberger (Rba ), Traversu (Tra ), Wulfsberg (Wu), Moen (Moa ), ELO, and Warrior (WARR). It is known that WARR and Wu correspond to point mutations of Thr 552 → Ile and Gly 565 → Ala of band 3, respectively. ELO, Rb (a+), Tr (a+), and Wa (a+) correspond to Arg 432 → Trp, Pro 548 → Leu, Lys 551 → Asn, and Val 557 → Met substitutions, respectively [20, 38]. On the transmembrane domain of band 3, a complex carbohydrate structure is attached to Asn642 . The Ii antigens are carbohydrate molecules and correspond to portions of the oligosaccharide chains [20]. Molecules with i reactivity correspond to oligosaccharide chains containing at least two repeating N-acetylgalactosamine


Fig. 2 Principle of fluorescence recovery after the photobleaching (FRAP)method to examine lateral mobility of band 3 molecules in red cell membranes in situ. The procedure is shown in (I). After band 3 is labeled by fluorescence (A),it is subjected to a spot laser beam, which abolishes the intensity of fluorescence on band 3 (B). The intensity of fluorescence is recovered by reentry of band 3 molecules with their lateral mobility (C). Relative fluorescence intensity is measured during this course of time.The

extent of the recovery of fluorescence intensity represents a mobile fraction (approximately 30 % of total) of band 3, and the remainder is equivalent to an immobile fraction of band 3. Interpretation of the results is shown schematically in (II). The band 3 molecules attached to the cytoskeletal network with anchoring proteins (ankyrin, and also probably protein 4.2) are immobile (a), and those without any fixation to the network are mobile (b, c, and d).

units, whereas I activity corresponds to a branched oligosaccharide structure formed by an N-acetylgalactosamine unit attached in a β1,6-linkage to a galactose residue within linear lactosamine polymers. Oligosaccharide chains in neonatal red cells are largely unbranched, and those in adult red cells are highly branched. The increase in the I reactivity, with a corresponding decrease in i reactivity, during early infancy corresponds to the elaboration and display of increasing numbers of β1,6-linked lactosamine units. Therefore, red cell expression of these Ii antigens is developmentally regulated. This particular β (1,6) N-acetylglucosaminyltransferase activity and a chain-elongating β (1,3) N-acetylglucosaminyltransferase activity have been identified.

2 Glycophorins

1.3 Band 3 in Nonerythyroid Cells

It is known that, in addition to erythroid band 3 (anion exchanger-1: AE1, or solute carrier family 4A: SLC4A1), two other genes (AE2andAE3) encoding band 3-related anion exchange proteins are present [7, 8, 39–41]. AE2 (or SLC4A2) is the general tissue anion antiporter, and is widely distributed in many tissues and cells. AE3 (or SLC4A3) is expressed in the heart and brain. In AE2 and AE3, approximately 300 amino acids are added to the NH2 -terminus of the AE1 molecule. Therefore, AE2 and AE3 are larger than AE1. Among these three transporters, there is distinct homology, particularly in the membrane domain of their molecules. AE1 is also expressed in tissues other than red cells, such as in the acid-secreting, type Aintercalated cells in the collecting ducts of the kidney, and in cardiac myocytes. The kidney transcript lacks the first 66 amino acids of the cytoplasmic domain of AE1. Thus, it is unable to bind glycolytic enzymes, protein 4.1, or ankyrin.

2 Glycophorins

The glycophorins are known as the most abundant integral membrane glycoproteins in red cells [19, 42, 43]. Glycophorins have a high sialic acid content, and more than 95 % of the periodic acid Schiff (PAS)-staining compounds come from the glycophorins. The five types of glycophorins are known as glycophorins A, B, C, D, and E [19, 42–52]. Among these five glycophorins, glycophorins A, B, and E are encoded by three closely linked genes [49–52], whereas glycophorins C and D arise from a single locus bearing no significant homology to the genes for glycophorins A, and B. Glycophorin D differs from glycophorin C by use of an alternate translation start site created by alternative mRNA splicing [19, 42, 43]. Another gene linked in tandem with those for glycophorins A and B has been isolated, which encodes glycophorin E [49–52]. This appears to have evolved from glycophorin A by homologous recombination at Alu repeats [49, 50]. However, no protein product has been identified regarding the gene for glycophorin E. Characterization of cDNA and genomic clones encoding the glycophorins has revealed that they fall into two distinct subgroups. Therefore, glycophorins are categorized into two groups, that is: (1) glycophorins A, B, and E, and (2) glycophorins C and D [19, 42, 43]. 2.1 Glycophorins A, B, and E 2.1.1 Glycophorin A (GPA) Glycophorin A (GPA) is the major sialoglycoprotein of red cells [19]. It is present at a level of approximately one million copies per cell. Its molecular weight, including carbohydrate, which is estimated from mobilities on SDS gels is 36 × 103 , and the molecular weight which is calculated from its molecular sequence excluding

7


Fig. 3 Molecular structures of glycophorins A, B, C, D, and E. Glycophorins B (GPB) and E (GPE) are basically derived from glycophorin A (GPA). In contrast, glycophorin D (GPD) is derived from glycophorin C (GPC). E denotes exons. Ge: Gerwich blood group antigens.

the carbohydrate chain is 14 × 103 . GPA is approximately 1.6% (w/w) of the total membrane proteins in the red cells. Three GPA complementary DNA transcripts are expressed from the gene (transcripts of 2.8, 1.7, and 1.0 kb) that vary only by the use of alternative polyadenylation sites. The 5 -ends of these transcripts are essentially identical. The gene of GPA (GYPA) is located at chromosome 4q28–q31. GPA is synthesized with a cleavable leader peptide that yields a type I transmembrane protein 131 amino acids long [19, 44, 45]. The protein is heavily glycosylated with a single asparagine (position 26)-linked glycan and 15 serine–threonine-linked oligosaccharide units. Approximately 60 % of the GPA molecule is carbohydrate. GPA gene is composed of seven exons (Fig. 3). Exon 1 yields a leader peptide, whereas exons 2 (amino acids 1–26), 3 (amino acids 27–57), and 4 (amino acids 58–70) encode the extracellular domain. Exon 5 encodes the transmembrane domain (amino acids 71–100). Exons 6 and 7 generate the cytosolic domain (amino acids 101–131) and 3 -untranslated region [19, 44, 45]. The precise biological functions of GPA have not been well defined. Since GPA demonstrates an extensive negative surface charge of the red cells, it is expected that it may modulate interactions between red cells and red cells, or between red cells and endothelial cells. It has been reported that red cells are clumped when sialic acids are removed. The M and N antigens on GPA are determined by amino

2 Glycophorins

acid polymorphism at positions 1 and 5 of the mature polypeptide. The M antigen is defined by a serine at amino acid position 1 and a glycin at position 5, whereas the N antigen is defined by a leucine at position 1 and a glutamine at position 5 [19, 20, 53]. Another example is the Ss phenotype. When methionine is present at amino acid position 29, the S phenotype is expressed [19, 20, 53]. Similarly, the presence of threonine at position 29 produces the s phenotype. GPA is expressed only in erythroid cells, especially after the proerythroblast stage during terminal erythroid maturation. Glycophorin A-deficient red cells, such as an En (a–) type [20, 54], exhibit increased glycosylation of band 3, probably due to the addition of excessive sialic acid, which should have been present on the GPA molecule. Therefore, total surface charge density is not affected, and GPA-deficient red cells maintain the normal red cell shape and deformability. Thus, GPA may not be crucial with respect to the maintenance of mechanical stability, deformability, or shape change. Total deficiency of band 3 in the band 3-knock-out (−/−) mice [55, 56] or in the Japanese cow [57] is associated with complete deficiency of GPA in their red cells. In addition, when red cells bind to immunologically nonspecific ligands, (wheat germ agglutinin) the binding causes aggregation of glycophorin and decreases red cell deformability. Glycophorin-deficient red cells are more resistant to invasion by malaria parasites than are normal red cells [58]. Amino acids 90 to 93 of the membrane domain appear to be critical to the formation of GPA dimers in the membranes. In the transmembrane domain, a GPA dimer showed a small, well-packed interface between the molecules. Van der Waals interactions alone can mediate stable and specific associations between transmembrane helices [59]. 2.1.2 Glycophorin B (GPB) Glycophorin B (GPB) is fairly similar to GPA except for their exoplasmic domains and cytoplasmic tails [46, 48]. The gene for GPB (GYPB) is located on chromosome 4q28–q31. GPB is a structurally similar type I transmembrane protein derived from a gene consisting of five exons. This gene yields a single 0.5 kb transcript. GPB is synthesized with a cleavable single sequence to yield a type I transmembrane protein, which is 72 amino acids in length. There are no asparagine-linked carbohydrate chains, because asparagine in position 26 of exon 3 is spliced out, whereas there are approximately 11 serine–threonine-linked oligosaccharide chains. Approximately 50% of the mass of GPB molecules consists of oligosaccharide. There are only about 150 000 copies per cell. The apparent molecular weight of GPB on the SDS gels is 20 × 103 , and that calculated from protein sequence data is 8 × 103 . GPB is equivalent to 0.2 % (w/w) of the total red cell membrane proteins. The glycophorin B gene is composed of five functional exons (exons 1, 2, 4, 5, and 6) [46, 48] (Fig. 3). Sequences corresponding to exon 3 are designated as pseudoexons, because the sequences are not expressed in GPB transcripts as a consequence of a non-functional splice acceptor sequence at its 3 -border. Exon 1 yields a leading peptide. The extracellular domain is encoded by exons 2 and 4, and exon 5 encodes the transmembrane and short cytosolic segment. Exon 6 gen-

9


Fig. 4 Evolution of glycophorins A, B and E. GPA: glycophorin A, GPB: glycophorin B, and GPE: glycophorin E.

erates the 3 -untranslated region. Amino acid sequence polymorphisms encoded by exon 4 yield an S-specific GPB molecule (methionine at position 29), or an s-specific molecule (threonine at position 29) [20, 53]. The amino acid sequence encoded by exon 2 yields the N antigen, with leucine at position 1 and glutamine at position 5. No biological function has been assigned to GPB other than its association with the Ss blood group [20, 53]. GPB is only present in erythroid cells, as is GPA.

2.1.3 Glycophorin E (GPE) Glycophorin E is a glycophorin that has been identified by molecular cloning [49–52]. The genes for glycophorins A, B, and E are located on the same chromosome 4q28–q31, in the order A, B, and E. The genomic structure and promoters of all three genes are highly conserved, and all three contain cleavable leader peptides. These three genes are oriented in a tandem fashion in a gene cluster (Fig. 4). The glycophorin E gene (GYPE) is predicted to be composed of four functional exons (exons 1, 2, 5, and 6) and two non-utilized pseudoexons (numbers 3, and 4) (Fig. 3). Exon 1 yields a leader peptide, exon 2 (positions 1 to 26) encodes the putative extracellular domain, and exon 5 (positions 26 to 59) encodes the predicted transmembrane segment. Exon 6 produces the 3 -untranslated region. The GYPE locus lacks DNA sequences corresponding to amino acid residues 27 to 39 of GPB, encompassing the position of the Ss amino acid sequence polymorphism in GPB. GYPE also contains a DNA sequence insertion, relative to GYPA and GYPE, at a position corresponding to exon 5 of GYPA. This insertion is predicted to encode eight amino acids not present in GPA or GPB. A polypeptide product corresponding to the GYPE locus has not been identified in vivo, but it could be a 20 000 Da molecule with a length of 59 amino acids, and with residues 1 and

2 Glycophorins

5 occupied by serine and glycine, respectively, corresponding to the M type blood group. 2.2 Gylcophorins C and D 2.2.1 Glycophorin C (GPC) Glycophorin C (GPC) is a glycoprotein with 128 amino acids [42, 43, 47, 48, 60, 61]. The molecular weight on the SDS gels is 32 kDa, but when calculated it is 14 kDa, because extensive posttranslational modification by glycosylation at the single asparagine (residue 8)-linked N-glycosylation site and 12 serine–threoninelinked O-glycosylation sites are present. Approximately 0.1% of the total membrane proteins (w/w) is GPC. There are about 143 × 103 copies of gly-cophorins C (per red cell). The gene for GPC (GYPC) is located on chromosome 2q14–q21. The GPC gene is composed of four exons (Fig. 3). The extracellular domain of GPC (residues 1–57) is encoded by exons 1, 2, and 3. The glycosylation sites are located in this extracellular domain. The transmembrane segment (residues 58–81) is encoded by exons 3 and 4, and the cytosolic domain (residues 82–128) by exon 4. The Gerbich (Ge: 2 and Ge: 3) blood type antigens are located at positions corresponding to exons 2 and 3 of GPC, respectively [20, 42, 62]. It is known that GPC is not erythroid-specific, and is present in multiple nonerythroid tissues in a distinctive fibrillar pattern. During erythroid differentiation, a desialylated form of GPC is present on the surface of erythroid progenitors of the burst forming unit in the erythroid (BFU-E). Normally glycosylated GPC first appears in erythroid progenitors of the colony-forming unit in erythroid (CFU-E). GPC is functionally important, because the cytoplasmic tail of GPC binds to protein 4.1 and p55 resulting in the anchoring of the skeletal network to the membrane. Therefore, GPC plays a crucial role in regulating the stability, deformability and shape of the membrane. 2.2.2 Glycophorin D (GPD) GPD is a truncated form of GPC (Fig. 3), corresponding to residues 22 to 128 of GPC [20, 42, 43, 60]. Therefore, GPD (23 kDa) is approximately 9000 Da smaller than GPC (32 kDa) on the SDS gels. The molecular weight of GPD calculated from the gene (GYPD) sequence is 11 kDa, and roughly 0.02% of the total membrane proteins (w/w) is GPD. There are approximately 82 × 103 copies of GPD per red cell. The same transcript yields both GPC and GPD through a mechanism involving translation initiation at an internal ATG codon. When translation is initiated at the first AUG, GPC is produced. When initiation occurs at the AUG encoding methionine at position 20, GPD as a truncated protein is produced. Residues 22 to 128 are identical in GPC and GPD. GPC does not express a cleavable signal peptide. The genes for GPC and GPD are located on chromosome 2q14–q21. There are six O-linked sugar chains in the GPD molecule, but none of the N-linked sugar chains. GPD expresses only the Ge: 3 determinant of the Gerbich blood group [20, 62].

11


3 Blood Group Antigens

As many as 243 different determinants have been listed as blood group antigens, which belong to one of 19 distinct blood group systems [20, 63–68] (Table 1). Some of these demonstrate the clinical relevance of the antigens especially in red cell transfusion and organ transplant procedures. Therefore, current information on their biochemical and genetic properties and the molecular basis for the inherited polymorphisms in these molecules is critical.

3.1 ABO Blood Group

Human beings are classified into distinct groups by the ABO blood group system (Table 1) depending on the presence or absence of substances in the serum that agglutinate red cells from humans of other classes [69]. The antigens of the ABO system (A, B, and H determinants) are expressed by red cells and by many other tissues. The cells of some human tissues produce water-soluble forms of these molecules as components of the glycans on secreted and soluble glycoproteins, on glycosphingolipids, and on free oligosaccharides. This mechanism is determined by the Secretor (Se) locus. The immunoreactive regions of the ABO blood group determinant are located at the terminal ends of various oligosaccharides, which are components of integral proteins and glycolipids of red cell membranes [69–72]. The A, B, and H blood group molecules are constructed sequentially by distinct glycosyltransferases which are determined by a distinct genetic locus. These glycosyltransferases operate on one of four structurally distinct oligosaccharide precursor types synthesized in human cells. Type 1 oligosaccharide precursors are found at the termini of linear and branched chain oligosaccharides linked to proteins at asparagine residues or at serine or threonine residues. The type 1 oligosaccharide precursors are synthesized only by epithelia of various tissue cells, yielding ABH determinants present in body fluids and secretions. The ABH determinants expressed by red cells are mainly by the type 2 precursor chains, which are also asparagine-linked or serine–threonine-linked oligosaccharides. Type 3 A, B, and H antigens in mucins are not found in human red cells. Type 4 chains are restricted to glycolipids in human red cells. These oligosaccharide precursors are catalyzed by α (1,2) fucosyltransferase in a transglycosylation reaction. These enzymes transfer the nucleotide sugar substrate GDP-fucose to carbon 2 of the galactose molecule at the oligosaccharide precursors [69, 70]. The fucose is attached in an alpha anomeric linkage and forms the blood group H determinant, such as the disaccharide Fuc α (1,2) Gal β-unit. The human genome encodes two different α (1,2) fucosyltransferases in a tissuespecific fashion, which correspond to the products of the H and the Se blood group loci [73].

Table 1 Major human blood group systems

System name/symbol

Gene name

Antigen type

Antigen copy number per red cell

Number of alleles

Chromosome

Chapter No. in text

001

ABO/ABO

ABO

Oligosaccharide

8 × 105 −2 × 106

3 major Several minor (cis-AB, subgroups)

9q34.1–q34.2

3.1

002

MNS/MNS MN (glycophorin A)

∼1 × 106

2 major (M or N) Multiple minor

4q28–q31

2.1

003 004

S (glycophorin B) P/P1 Rh/RH

Glycoproteins GYPA, (GYPE) GYPB P1 Oligosaccharide Protein complex

∼1.5 × 105 ∼103

4q28–q31 22q11.2–qter 1p34.3–p36.1

2.1 3.3 3.2

005 006

D C/c, E/e Lutheran/LU Kell/KEL

RHD RHCE LU Glycoprotein KEL Glycoprotein

∼2 × 104 ∼2 × 104 ∼1600–4100 ∼5 × 103

19q12–q13 7q33

3.4 3.5

007 008 009 010

Lewis/LE Duffy/FY Kidd/JKKidd/JKa , 4> Diego/DI

FUT3 FY JK AE1

Oligosaccharide Glycoprotein Glycoprotein Glycoprotein (band 3)

4500–7300 ∼12 000–17 000 ∼14 000 ∼15 000

19p13.3 1q22–q23 18q11–q12 17q12–q21

3.6 3.7 3.8 3.11

011

Cartwright/YT

Not determined

012

XG/XG

ACHE Glycoprotein (acetylcholinesterase) XG Glycoprotein

2 major (S or s) Multiple minor Complex ∼48 haplotypes Includes minor C/c and E/e alleles 2 major (D+ or D− ) 2 major (C or c, and E or e) 3 major (Lua , Lub , recessive null) 2 major (KEL1, KEL2) Several minor 2 major (Lea , Leb ) 2 major (Fya and Fyb ) 2 major (Jka , Jkb ) Several Dia , Dib Wra , Wrb Other high-frequency antigens 1 major (Yta ) 1 minor (Ytb )

∼9000

Xp22.32

013 014

Scianna/SC Dombrock/DO

SC DO

Not determined Not determined

1 major (Xga ) 1 minor (Xg, a hypothetical null) 3; Scl, Sc2, and Sc3 1 major (Doa ) 1 minor (Dob )

Glycoprotein Glycoprotein (GPI-linked)

7q22

1p36.2–p22.2 unknown

3 Blood Group Antigens 13

ISBT No.

ISBT No.

System name/symbol

Gene name

Antigen type

Antigen copy number per red cell

Number of alleles

Chromosome

015

Colton/CO

AQP1

Not determined

2 major (Coa and Cob )

7p14

016

LandsteinerWeiner/LW

LW

Glycoprotein (aqua-porin-1) Glycoprotein (ICAM-4)

2 major (LWa , LWb ) Rare nulls

19p13.3

017

ChidoRogers/CH/RG

C4A, C4B

∼4400 (on D+ cells) ∼2835–3620 (on D− cells) ∼103

Ch (1 major, several minor) Rg (1 major, several minor)

6p21.3

018

Hh/H

FUT1

See ABO

19q13

019 020

Kx/XK Gerbich/GE

XK Protein GYPC, Glycoproteins GYPD

1 major Several minor (Bombay, para-Bombay) 1 major (Kx ) 1 major (Ge: 1, 2, 3, 4)

021

(glycophorins C and D) Cromer/CROM

DAF

022

Knops/KN

CR1

023 − −

Indian/IN Secretor/Se Ii

CD44 FUT2

ISBT: International Society of Blood Transfusion.

Glycoprotein (4th component of complement; C4A and C4B) Oligosaccharide

Glycoprotein (decay-accelerating factor) Glycoprotein (complement receptor CRI) Glycoprotein Oligosaccharide Oligosaccharide

Not determined ∼1 × 105 (glycophorin C) ∼2 × 104 (glycophorin D) ∼103

Xp21.1 2q14–q21

Chapter No. in text

3.9

2.2

Multiple minor 2 major (Cra , Crb ) Several minor

1q32

Not determined

Several major (Kna , Knb , Mca , Mcb , Yka , Sla )

1q32

Not determined see Lewis see ABO

2 major (Ina , Inb ) 2 major (Secretor, non-Secretor) 1 major (I), at least 1 minor (i)

11p13 19q13 9q21

3.10


Table 1 (Continued).


Glycosyltransferases, which are encoded by the ABO blood group locus, use type 1, 2, 3, or 4 H determinants to form A or B blood group determinants. The A allele at the ABO locus encodes an α (1,3) N-acetylgalactosaminyltransferase that uses H molecules to form the blood group A molecule [69–71]. The B allele encodes an α (1, 3) galactosyltransferase that operates on H-active oligosaccharide precursors to form the blood group B determinant. The O allele is a null allele, which cannot encode a functional glycosyltransferase that will further modify H-active precursors. The ABH determinants in human red cells are mainly associated with membrane glycoproteins, that is, 80% of those (1–2 × 106 molecules per red cell) in band 3, and others in the red cell glucose transporter (band 4.5: 0.5 × 106 molecules per red cell), the Rh-related proteins, and the aquaporin-1 glycoprotein [74]. Red cell membrane glycolipids are also involved with the ABH molecules (0.5 × 106 molecules per red cell). A single asparagine-linked oligosaccharide molecule on band 3 is a branched poly-N-acetylgalactosaminoglycan, whose terminal branches may display several ABH determinants. Ig M antibodies specific for ABO oligosaccharide determinants are not displayed in red cells during infancy. This immune response is a consequence of exposure to microbial oligosaccharide antigens that are structurally similar (or identical) to the A and B blood group molecules. It is also interesting to note that, in most individuals, antibodies directed against H determinants are not formed because a substantial number of the blood group H precursors are not enzymatically converted into A or B determinants. The Ig M isoagglutinins, which occur naturally, efficiently fix complement leading to acute hemolysis of transfused red cells that display the corresponding antigen. There are a substantial number of variants of the ABH blood group antigensss [75, 76]. For example, in A subgroups, the A1 and A2 phenotypes are known, which differ in their molecular structure [70, 71]. The absolute number of immunodominant molecules is greater on A1 cells than it is on A2 cells. The human A transferase is a type II transmembrane protein, and is composed of 353 amino acids with an NH2 -terminal segment (residues 1–15), a hydrophobic segment (residues 16–39), and a COOH-terminal domain (residues 40–353). The COOH-terminal catalytic domain is located within the membranedelimited compartments of the Golgi and the trans-Golgi network, where terminal glycosylation reactions occur. This enzyme has a single potential site for asparaginelinked glycosylation, and functions as a glycosyltransferase [77]. The A transferase is also present as a soluble, catalytically potent polypeptide. In the molecular structure, the NH2 -terminus of the soluble enzyme corresponds to the alanine residue at codon 54, indicating that the soluble form of the A transferase is derived from its transmembrane precursor by proteolysis. Therefore, glycosyltransferase exists in both membrane-associated and soluble, catalytically active forms [70]. As regards the molecular basis for polymorphism at the ABO locus, the blood group O phenotype is produced and is due to a single base pair deletion of one nucleotide in the codon for amino acid 87 of the A transferase. By this frameshift of the reading frame, a termination codon appears at amino acid residue 117 of the A transferase. The truncated protein (116 amino acids) is consequently unable to


modify the H antigen to form A or B structures, resulting in no detectable A or B transferase activity being observed in the sera taken from blood group O individuals (genotype OO). As for the A or B blood group phenotype, seven differences in nucleotide sequence are present within the protein-coding segments of the A transferase cDNA. Three of the seven appear to be functionally neutral polymorphisms. The other four appear to be critical for expressing the A and B transferases. These are at residues 176 (arginine, A; glycine, B), 235 (glycine, A; serine, B), 266 (leucine, A; methionine, B), and 268 (glycine, A; alanine, B). The polymorphisms at positions 266 and 268 are important for enzymatic functions to discriminate between UDPN-acetylgalactosamine and UDP-galactose. Therefore, leucine at 266 and glycine at 268, or methionine at 266 and alanine at 268 generate an A transferase or B transferase, respectively. H blood group oligosaccharide precursors are the most important substrates for the transferases encoded by the ABO locus. H-active precursors display terminal Fuc α (1,2) Gal β linkages, which are an integral part of the A and B antigenic determinants. These linkages are synthesized by α (1,2) fucosyltransferases (GDPfucose: Gal β2-α-L-fucosyltransferase). These transferases can use types 1, 2, 3, and 4 glycoprotein or glycolipid substrates as well as low molecular weight β-D-galactosides [70]. The synthesis of H-active blood group substances is determined by two factors, that is, the H locus and the Secretor (Se) locus [69, 70]. The Se locus determines expression of an α (1,2) fucosyltransferase activity, and H-active blood group substances (membrane-associated, and also soluble). Nearly all of this soluble blood group-active substance is constructed from type 1 precursors and is released mainly from the sublingual and submaxillary glands and the parotid gland. Red cells taken from both secretors and non-secretors maintain an essentially identical complement of H-determinants and also A or B determinants, depending on the ABO locus genotype. In red cell precursors, the synthesis of α (1,2) fucosyltransferase activity, and H determinants is directed by the H locus. The H and Se loci correspond to distinct genes encoding different α (1,2) fucosyltransferases with disparate tissue-specific expression patterns. The H locus is expressed predominantly in erythroid cells. In contrast, the Se locus represents a second α (1,2) fucosyltransferase locus whose expression is restricted to the epithelia of many tissue cells. Individuals with the secretor phenotype maintain at least one functional allele at both the H locus and the Se locus. Individuals with the non-secretor phenotype maintain two null alleles at the Se locus and at least one functional H allele. In human erythroid cells, the Se locus is constructed mainly from type 1 precursors, and the H determinants, which are synthesized by the H-encoded α (1,2) fucosyltransferase, are based on type 2 precursors [69, 73]. The human Se locus and the human H locus are separated only by 35 kb of genomic DNA on the same human chromosome 19. The human H locus encodes a 365 amino acid-long polypeptide as the type II transmembrane glycosyltransferase [78, 79]. The molecular structure is


composed of an NH2 -terminal cytosolic domain (residues 1–8), a hydrophobic domain (residues 9–25) that spans the Golgi membrane, and a COOH-terminal domain (residues 26–365) corresponding to a Golgi-localized catalytic domain, where two potential asparagine-linked glycosylation sites are present. Therefore, the human H locus is an α (1,2) fucosyltransferase. The human Se locus is a 332 or 343 amino acid-long polypeptide, which shares 68% of the amino acid sequence identity with the COOH-terminal 292 residues of the human H blood group α (1,2) fucosyltransferase [79]. The molecular structure is composed of a cytosolic domain with 3 or 14 residues, a 14 residue hydrophobic membrane domain, and a 315 amino acid-long COOH-terminal domain, which is located in the Golgi lumen. There are three potential asparagine-linked glycosylation sites [80, 81].

3.2 Rh Blood Group

The Rh (Rhesus) blood group antigens were initially identified by Landsteiner and Wiener in 1940 in the antisera of immunized guinea pigs and rabbits with red cells taken from Macaca rhesus monkeys [20, 68, 82, 83]. The human alloantibody is called Rh, and the heteroantibody is called LW (Table 1). RH and LW are antigenic systems determined by distinct gene complexes, which are located on chromosomes 1 and 19, respectively. LW and the Rh antigen complex associate in the membrane, and LW expression requires Rh polypeptide expression. A third antigen, Rh50, is also important for normal expression of the Rh, LW, and glycophorin molecules. The incidence of Rh positive is approximately 85 % in whites, and strikingly 99.5 % in the Japanese population. About 15 % of Caucasians and 0.5 % of Japanese are Rh negative. One main determinant of the Rh antigen system is the RhD antigen. Therefore, Rh positive individuals demonstrate the RhD antigen, whereas the Rh negative ones do not express the RhD antigen. In the Rh blood group system, other antigens, that is, the C/c and E/e antigen group, are known [84]. Their expression is determined by a second locus linked extremely tightly to the locus that determines D antigen expression [82, 83]. There are eight common Rh gene complexes considering D, C, c, E, and e alleles. The d antigen does not exist, because there is no product of the hypothetical d allele of the D gene. The C/c and E/e antigens correspond to a single, non-glycosylated, 417 amino acid-long (33 100 Da) polypeptide (RhCE) with 12 membrane spanning domains, encoded by the RHCE gene. The RHCE gene is composed of 10 exons in 70 kb of genomic DNA. The D antigen corresponds to a distinct non-glycosylated, 417 amino acid-long (33, 100 Da) protein (RhD), which is encoded by the RHD gene. This RhD protein shares 90% of the identity of the amino acid sequence with the RhCE protein, and also demonstrates a 12 membrane spanning domain [85]. The RHD and the RHCE genes are closely linked on human chromosome 1p34–p36.


Considering the Rh antigen system, there is a third polypeptide (Rh50), which is a 409 amino acid-long glycoprotein. This peptide is approximately 36 % identical with amino acid sequences of RhD and RhCE, which are also known as the Rh30 polypeptides [86]. This Rh50 peptide has a 12 membrane-spanning domain, and is encoded by the RH50 gene, which is located on human chromosome 6p11–p21.1. The RH50 gene exhibits 10 exons and 32 kb of its size, closely resembling those of the RHD and RHCE genes [87, 88]. Although the Rh50 protein does not itself express Rh antigens, it interacts with the Rh30 polypeptides (RhD and RhCE) in the membrane, and is required to form a heterotetrameric complex, two molecules of Rh50 and one molecule each of RhD and RhCE, which is essential to normal cell surface expression of the Rh30 (RhD and RhCE)-encoded Rh antigens. The RhD and RhCE polypeptides are modified by fatty acylation and are also palmitylated through a thiolester linkage to free sulfhydryl groups on cysteine residues within a consensus tripeptide (Cys-Leu-Pro) in the molecules of the RhD and RhCE. Reactivity of Rh antigens is also regulated by the lipid composition of the membrane in situ. Therefore, alteration of the membrane lipid concentration could induce conformational changes of the Rh peptides. It is also known that the Rh polypeptides interact with the red cell membrane skeleton. It has recently been shown that protein 4.2-deficient red cells lack CD47 implicating an interaction between the Rh complex and the band 3 complex [89]. 3.3 P Blood Group

The P antigens (ISBT No.003: P/PI) are present exclusively on red cell membraneassociated glycosphingolipids [20, 68, 90, 91]. P molecules are produced from lactosyl ceramide by the sequential processing of a series of distinct glycosyltransferases (Table 1). Two distinct pathways have been proposed for the biosynthesis of the P antigen molecule via the step of the P antigen as a precursor, even though so little is known about the corresponding enzymes and genes. The most common phenotype (P1 ) demonstrates full activity, and its frequency is approximately 75 %. The other most common phenotype is P2 with a frequency 25%. Three rare phenotypes have been described: (1) the Pk1 phenotype, which is deficient in P transferase activity, (2) the Pk2 phenotype, which is homozygous for null alleles at the P transferase and the P1 transferase loci, and (3) the p phenotype, which is deficient in all three P antigens (P, P1 , and Pk ). Functions of the P blood group system remain unknown. 3.4 Lutheran Blood Group

In the Lutheran blood group system (ISBT No.005: Lutheran/LU), four major phenotypes are present [20, 68]. Approximately 90 % of normal subjects demonstrates Lu (a–b+), or LU: –1, 2 (ISBT phenotype). Others are Lu (a+b+); LU: 1, 2, Lu (a+b–); LU: 1, –2, and Lu (a−b−); LU: O, which displays no detectable Lutheran antigenic


activity (Table 1). In blood, the Lutheran blood group proteins are restricted in their expression to red cells and to B lymphocytes. The Lutheran antigens are expressed on a pair of membrane glycoproteins with molecular weights of 78 000 and 83 000. The Lutheran blood group antigens are a 597 amino acid-long type 1 transmembrane protein. The extracellular domain contains five potential N-glycosylation sites and five peptide segments that share a primary sequence similarity with members of the immunoglobulin superfamily [92]. Its structural organization is similar to MUC18 (the melanoma-associated, mucinlike protein) and related neural cell adhesion molecules. A single membranespanning segment is followed by a 59 amino acid-long intracellular segment, in which an Src homology 3 domain is present. The Lutheran blood group gene (LU) yields a pair of alternately spliced transcripts, producing two molecular weight isoforms of the protein that differ in their lengths of the cytosolic domains. One isoform of the Lutheran polypeptide is identical to a B-CAM (a basal cell carcinoma/epithelial cancer adhesion molecule). The Lutheran blood group gene corresponds to a 12.5 kb gene with 15 exons [93, 94]. There are several nucleotide polymorphisms in exon 3 at base pair 229 of the coding sequence. In the Lutheran (a+b–) phenotype, the nucleotide A at this position contributes to a histidine codon corresponding to residue 77 of the Lutheran polypeptide. In the (a–b+) phenotype, the nucleotide at this position is G corresponding to an arginine codon at residue 77 of the peptide.

3.5 Kell Blood Group

Although many human blood group alloantigens have been discovered, the KEL1 antigen of the Kell blood group system (ISBT No.006: Kell/KEL) is highly immunogenic, behind the RhD antigen, which is the strongest in its immunogenicity [20, 68, 95, 96]. Anti-KEL1 antibodies account for approximately two-thirds of nonRh immune red cell alloantibodies. There are approximately 5000 Kell determinants per red cell (Table 1). The Kell protein is a 732 amino acid-long 83 000 Da polypeptide. This is a type 1 transmembrane protein with a cytosolic NH2 -terminal segment, a single hydrophobic membrane-spanning segment, and a large COOH-terminal extracellular domain [95, 96]. Six potential asparagine-linked glycosylation sites and 15 cysteine residues are present in the extracellular domain. This protein demonstrates two heptad arrays of leucine residues, with clustered cysteine residues, in a leucine zipper motif that may be involved in protein/protein interactions. There is a primary sequence similarity of the Kell glycoprotein with the neprilysin family of zinc-binding neutral endopeptidases such as bradykinin, neurotensin, enkephalin, oxytocin, and angiotensins I and II. Biochemically, the monospecific anti-Kell antibodies precipitate immunologically a single 93 000 Da red cell membrane protein with two different Kell epitopes. This polypeptide exists as part of a large complex (from 115 × 103 Da to 200 × 103


Da in size) under non-reducing conditions. The Kell glycoprotein appears to exist as a homodimer in the membrane. 3.6 Lewis Blood Group

The Lewis antigens (ISBT No.007: Lewis/LE) expressed in red cells are unique, because these antigens themselves are not synthesized by erythroid precursors. These antigens are Lewis-active glycosphingolipid molecules basically present in plasma, and are adsorbed by the red cell membrane through an apparently passive process [20, 68]. Two forms of the Lewis antigens (Lea and Leb ) are known as complexes with low- and high-density lipoproteins, and also as aqueous dispersions in the plasma (Table 1). There are approximately 4.5–7.3 × 103 Lea molecules per red cell. The molecule associates with red cell membranes through the ceramide moiety. The Lewis antigens are absent in red cells of newborns, and appear approximately 10 days after birth. The full activity of the Lewis antigens (Lea ) in red cells is established at approximately 24 months of age. The Lea and Leb antigens are synthesized by two distinct fucosyltransferases under the control of the Le blood group gene and the Se blood group gene. The Le gene corresponds to an α (1, 3/1, 4) fucosyltransferase gene which is an Fuc-TIII or FUT3 gene [69, 70, 80]. The enzyme encoded by this gene can use oligosaccharide precursors, including unsubstituted type 1 oligosaccharide precursors, to produce the Lea antigen, and type 1 H antigens to generate Leb antigens. Le genedependent expression of Lea and Leb molecules is identified at the epithelia lining the respiratory tract, urinary tract, digestive tract, salivary glands, and bile ducts [69, 70, 73, 80]. These tissues correspond to the tissue types capable of expression of type 1 H molecules, the synthesis of which is determined by the Se locus. α (1,3/1,4) Fucosyltransferase (FUT3) is a 363 amino acid-long type II transmembrane glycoprotein, which is composed of a 15 amino acid-long NH2 -terminal cytosolic segment, a 19 residue transmembrane segment, and a 320 amino acidlong COOH-terminal catalytic segment, which is located in the Golgi apparatus. The enzyme produces several types of α (1,3) – and α (1,4) fucosylated oligosaccharides, which are Lea , Leb , Lex , and Ley molecules, and sialylated forms of the Lea and Lex antigens. The FUT3 gene, as a member of an α (1,3) fucosyltransferase gene family, is located on chromosome 19p13.3. The Lewis α (1,3/1,4) fucosyltransferase can use the oligosaccharide products formed by the Se-determined α (1,2) fucosyltransferase. In addition, the Se and Le fucosyltransferases are expressed in many of the same tissues. Therefore, the genotype at these two genes determines which of the Lewis-active oligosaccharide molecules is constructed. In secretor-positive subjects, type 1 oligosaccharide precursors are first converted into type 1 H molecules, which become substrates for the Lewis locus-encoded α (1,3/1,4) fucosyltransferase [70, 80]. This enzyme converts these into Le -active molecules: that is, an Le (a–b+) phenotype. On the other hand, in non-secretors, type 1 H antigens cannot be produced in secretory epithelia. Thus,


these unsubstituted type 1 molecules are converted into Lea -active oligosaccharides by the Le-encoded α (1,3/1,4) fucosyltransferase: that is an Le (a+b–) phenotype. In homozygotes for null alleles at the Le gene, the phenotype should be Le (a−b−). In this case, two possibilities exist. In Lewis-negative and secretor-positive subjects, type 1 H determinants produced remain unconverted into Le antigens. In Lewisnegative, secretor-negative subjects, the type 1 precursors remain unsubstituted by either blood group fucosyltransferase. 3.7 Duffy Blood Group

There are two major alleles in the Duffy blood group system (ISBT No.008: Duffy/FY), that is, Fya and Fyb with virtually equivalent frequencies (Table 1). Other alleles are Fyx with a weakly reactive form of Fyb , Fy of a null allele, which produces no antigenic activities of Fya and Fyb [20, 68, 97]. The Duffy antigen is a 338 amino acid-long polypeptide with seven membranespanning segments. The Fya and Fyb antigens are localized on a glycoprotein with a molecular weight of from 38 × 103 to 90 × 103 Da. A significant amount of this molecule corresponds to asparagine-linked oligosaccharides, which demonstrate the heterogeneous migration properties in its native condition. The Fya and Fyb alleles differ only by a single amino acid substitution at codon 44: a glycine for the Fya antigen, and aspartic acid for the Fy antigen. Functionally, this protein corresponds to the red cell chemokine receptor, known as DARC (Duffy antigen receptor for chemokines). DARC removes excessive chemokines from the blood and tissues. In the homozygotes for the Fy allele, DARC expression in the bone marrow is defective, in spite of the normal expression of DARC in the extra-marrow tissues. This polymorphism in the tissue-specific expression of DARC is accounted for by a single base pair difference between the Fy allele and the Fya or Fyb alleles [98, 99]. The sequence change is located in the promoter region of the DARC gene, and in the Fy allele, leading to the disruption of a binding site for the erythroid lineage-specific transcription factor GATA-1. Duffy antigens are important for the invasion of human red cells by plasmodium vivax. Successful invasion of the merozoite into the red cells requires the subsequent formation of a junction between the apex of the merozoite and the red cells. Formation of this junction and penetration of the merozoite into the red cells only occur on Duffy-positive red cells [100]. 3.8 Kidd Blood Group

The Kidd antigen of the Kidd blood group system (ISBT No.009: Kidd/Jk) is a 46 000 to 60 000 Da protein [20, 68]. There are approximately 14 000 Kidd molecules in the red cells (Table 1). Two antigens (Jka and Jkb ) are expressed in their virtually equal gene frequencies, that is, 0.514 for the JK a allele and 0.486 for the JK b allele. A single amino acid substitution accounts for the polymorphism in the Kidd blood


group system, that is, Asp at codon 280 of the Kidd gene for Jka , and Asn at the same codon for Jkb , respectively. Approximately half of normal individuals express the Jk (a+b+) phenotype in the red cells, owing to the genotype Jka Jkb . A quarter of individuals (26% or 24%) exhibit the Jk(a+b–) phenotype due to the genotype Jka Jka , or the Jk (a–b+) phenotype due to the genotype Jkb Jkb , respectively The Jk (a−b−) phenotype is rarely observed. The gene mutation of aberrant splicing in the Kidd gene leads to the Kidd null phenotype. It has been shown that the primary sequence of a human red cell urea transporter corresponds to the Kidd antigen [101]. 3.9 LW Blood Group

There are two major allelic antigens with the LW blood group (ISBT No.016: Landsteiner-Weiner/LW), that is, LWa and LWb [20, 68, 102]. The LWa antigen predominates and the LWb antigen is detected in less than 1 % of the total population (Table 1). The two alleles yield the common phenotype LW (a+b–), the much less common phenotypes LW (a–b+) and LW (a+b+), and the rare phenotype LW (a−b−), which is homozygous for null alleles at the LW locus. Expression of the LW polypeptides is dependent on expression of the Rh polypeptides. There are approximately 4400 LW molecules per red cell. The LW antigen corresponds to a 42 kDa red cell glycoprotein with a deglycosylated molecular mass of 25 000 Da. The COOH-terminal region of the LW glycoprotein is present at the surface of the red cells. Two structurally distinct LW proteins are present. The first one is a type I transmembrane protein with a short cytoplasmic tail. Another one is a molecule without the membrane-spanning and cytoplasmic regions of the first longer form. The LW antigens are also one of the ICAM (the intracellular adhesion molecule) family, such as ICAM-4 [103]. This protein shares approximately 30% of the same identity of the protein sequence with other members (ICAMs-1, 2, and 3). The LW gene corresponds to three exons of 2.65 kb on human chromosome 19 [104]. The LW a and LW b alleles are different at a single base pair, in a codon 70 corresponding to one amino acid residue, that is, glutamine for LW a , and arginine for LW b . One LW null allele is known with a 10 base pair deletion, resulting in a truncated protein at a position proximal to its transmembrane and cytosolic regions. 3.10 Ii Blood Group

The commonly expressed antigen of the Ii blood group system is denoted I, and its absence is called i [20, 68, 105, 106] (Table 1). The Ii antigens are carbohydrate molecules. I activity corresponds to branched oligosaccharide structure formed by an N-acetylgalactosamine unit attached in β1,6 linkage to a galactose residue within linear lactosamine polymers, whereas molecules with i reactivity correspond


to oligosaccharide chains containing at least two repeating N-acetylgalactosamine units. It is known that oligosaccharide chains in neonatal red cells are unbranched, and that those in adult red cells are branched. Red cell expression of the Ii blood group system is developmentally regulated. I determinants are deficient in embryonic, and cord red cells, which are highly reactive with anti-i antibodies. During the first 18 months after birth, this relationship is reversed to yield red cells with increased anti-I reactivity and diminished i reactivity, which is usually observed in adult red cells. This phenomenon corresponds to the fact that the increase in the I reactivity and the decrease in the i reactivity during early infancy are associated with the elaboration and display of increasing numbers of β1,6-linked lactosamine units. I reactivity is determined by a locus encoding an N-acetylglucosaminyltransferase that is expressed in a developmentally regulated fashion. The functions of the Ii blood group system are yet to be defined. 3.11 The Diego and Wright Blood Group Antigens on Band 3

Band 3 (anion exchanger 1: AE1) in red cells demonstrates several polymorphic peptide epitopes. The two major ones are the Diego blood group system (ISBT No.010: Diego/DI) and the Wright blood group system [20, 68] (Table 1). The Diego (Dia ) allele occurs relatively frequently in the Japanese population, but is rare in Caucasians. There are two Diego antigens, i. e., Dia and Dib . The Dia antigens correspond to a proline at position 854 of the 911 amino acid-long glycoprotein (band 3), whereas the Dib antigens correspond to a leucine residue at the same position 854 of this molecule [107]. The Wright alleles correspond to the antigens Wra and Wrb . The frequency of the Wr a allele is rare, but the Wr b allele is fairly prevalent. The Wra antigens correspond to a lysine residue at codon 658 of the AE1 gene, and the Wrb antigens to a glutamine residue at the same codon 658, respectively [108]. Wrb expression is suppressed in glycophorin A deficiency. Seven minor antigens are present on band 3, these are: Waldner (Waa ), Redelberger (Rba ), Traversu (Tra ), Wulfsberg (Wu), Moen (Moa ), ELO, and Warrior (WARR). Amino acid substitutions at codon 552 (Thr → Ile), and at codon 565 (Gly → Ala) of the band 3 gene are known as WARR and Wu, respectively. Similarly, amino acid substitutions at codon 432 (Arg → Trp), at codon 548 (Pro → Leu), at codon 551 (Lys → Asn), and at codon 557 (Val → Met) correspond to ELO, Rba , Tra , and Wda antigens, respectively. 3.12 Other Minor Blood Group Antigens

The Chido (Ch) blood group system and the Rodgers (Rg) blood group system are known to be complementary-associated blood group antigens (Table 1).


The decay-accelerating factor (DAF) is one of the complementary regulatory proteins associated with red cell membranes as a glycosylphosphatidylinositol (GPI)-anchor protein. DAF-associated antigens are termed Cromer (Cr)-related antigens. The rare Inab phenotype corresponds to a total deficiency in the Cromerrelated antigens (Table 1). The Knops (Kna ) antigens in red cell membranes are expressed by the complementary receptor 1 protein (CR 1) (Table 1). The Cartwright (Yta and Ytb ) blood group system is associated with acetylcholinesterase in red cells, which is also a GPI-anchor protein. The type III red cells of paroxysmal nocturnal hemoglobinuria is deficient in acetylcholinesterase activity, demonstrating the Cartwright null phenotype (Table 1). The Indian (Ina and Inb ) antigens are expressed on CD44, which is the red cell isoform of the hyaluronan-binding protein (Table 1). The Colton (Coa and Cob ) blood group antigens are carried on an aquaporin-1, which is a red cell glycoprotein (Table 1). The physiological relevance of these antigens remains to be clarified [20, 64, 68].

4 Glycosyl Phoshatidylinositol (GPI) Anchor Proteins

Glycosylphosphatidylinositol (GPI) anchor proteins are proteins with complex glycolipid structures, which are highly conserved in all eukaryotic cells [109–111]. The common core region consists of a molecule of phosphatidylinositol (PI) to which are attached four sugars, that is, one molecule of N-glucosamine and three molecules of mannose (Fig. 5). The last mannose is attached to the carboxyl end of the protein through phosphoethanolamine. A palmitoyl residue is added to the inositol but may later be removed. The N-glucosamine is derived from the N-acetylglucosamine that is added and then deacetylated. The mannose residue is derived from dolichyl phosphoryl mannose. GPI anchor biosynthesis starts on the cytoplasmic side of the rough endoplasmic reticulum [112]. N-Acetylglucosamine is transferred to a phosphatidylinositol acceptor. This product is deacetylated to form glucosamidyl phosphatidylinositol. The first mannose is derived from dolichol phosphoryl mannose, as described above. The phosphoethanolamine is added. Addition of the GPI anchor precursor to the carboxy-terminus of the protein occurs on the luminal side of the endoplasmic reticulum, after amino acid residues 17 to 31 have been cleaved from the protein. When the synthesis of the anchor is completed, it proceeds through the Golgi apparatus and on the membrane surface. There are many membrane proteins which are related to GPI-anchor proteins, such as: (1) enzymes; acetylcholinesterase and leukocyte alkaline phosphatase, (2) complementary defense proteins; decay accelerating factor (DAF and CD55), a membrane inhibitor of reactive lysis (MIRL, CD59, protectin), and C8-binding protein (homologous restriction factor), (3) immunologic proteins; Fcy receptor

References

Fig. 5 Biosynthetic pathway of glycosyl phosphatidylinositol (GPI)anchoring proteins (A), and the molecular structure of the PIG-A gene (B).

IIIa, lymphocyte function-associated antigen-3 (LFA-3, CD58), endotoxin-binding protein receptor (CD14), and CDw 52 (Campath-1), (4) receptors; urokinase (plasminogen activator) receptor, and folate receptor, and (5) granulocyte proteins of unknown function; CD14, CD48, CD66, and Dombrock-Holley/Gregory-bearing protein. It is known that these proteins are missing from the blood cells in paroxysmal nocturnal hemoglobinuria (PNH), in which PNH blood cells are unable to synthesize the mature GPI anchor precursors. In lymphoblastoid cells of PNH, the block always occurs at the step when N-acetylglucosamine is transferred from UDPN-acetylglucosamine to phosphatidylinositol. This is the first step of the pathway and is catalyzed by the α1,6-N-acetylglucosaminyltransferase, which is an enzyne complex formed by four gene products, i. e., PIGA, PIGC, PIGH, and hGP1 [113]. Of these, the PIGA gene has been identified as pathognomonic for PNH [114]. The PIGA (phosphatidylinositol glycan complementation group A) gene consists of six exons and encodes a putative protein of 484 amino acids (approximately 60 kDa). This gene is located on chromosome Xp22.1. Many mutations of this gene have been reported in PNH patients. Recent advances on PNH and related disorders have recently been reviewed in detail [115].

25


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1

Retinoid Receptors RAR and RXR: Structure and Function Alexander Mata de Urquiza, and Thomas Perlmann

Ludwig Institute for Cancer Research, Karolinska Institute, Stockholm, Sweden


1 Retinoids in Development

Vitamin A and its biologically active metabolites, the retinoids, play essential roles in embryonic development, differentiation, and maintenance of homeostasis in the adult organism [1–3]. Adult animals suffering from vitamin A deficiency (VAD) display a number of abnormalities, including impaired vision, fertility, immune response, and epithelial differentiation. Furthermore, altering the levels of retinoic acid (RA) during embryogenesis leads to phenotypical malformations affecting for example cranofacial, CNS, limb, and heart development (reviewed in Refs [4] and [5]), underscoring the importance of correct vitamin A levels during gestation. Some of these abnormalities are thought to arise due to dysregulation of Hox gene expression. Hox genes encode a family of homeobox-containing transcription factors that play crucial roles in informing cells of their position along the anterio-posterior axis (reviewed in Refs [3] and [5]). The overlapping expression domains of these genes along the anterio-posterior axis are thought to specify the positional identity of cells along this axis, thereby enabling them to adopt a correct developmental fate. Interestingly, several Hox genes contain RA response elements (RAREs; see below) in their promoters, indicating that retinoids are involved in regulating their expression. Accordingly, embryos that develop in the absence of RA or in the presence of excess RA, display altered Hox gene expression [5, 6]. The distribution of retinoids in vivo has been analyzed using transgenic mice that express a retinoic acid-inducible reporter gene [7–14]. The results suggest that the brachial and lumbar regions of the developing spinal cord are “hot spots” of RA synthesis. In addition, reporter gene expression has been detected in the developing forebrain, forelimbs, and optic cup, as well as at the midbrain/hindbrain boundary [7, 9–11]. A better understanding of how this highly localized synthesis of Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Retinoid Receptors RAR and RXR: Structure and Function

Fig. 1 (A) Metabolic steps leading from retinol to retinoic acid and one of its oxidation products. The initial step is ratelimiting, and is the only reversible step in this process. 9-cis Retinoic acid (see B) is presumably formed via spontaneous iso-

merization from the all-trans form. (B) Structures of three natural RXR agonists, including 9-cis retinoic acid (left), the fatty acid docosahexaenoic acid (middle), and the chlorophyll metabolite phytanic acid (right) (see text for details).

RA is achieved in tissues has been gained with the cloning of enzymes responsible for RA production and degradation. Dietary all-trans retinol (atROL) is converted to all-trans retinal (atRAL), a reaction catalyzed by one of various ROL dehydrogenases (ROLDH). In the following step, atRAL is converted into atRA by a RAL dehydrogenase (RALDH) (reviewed in Refs [15] and [16]) (see Fig. 1A). 9cRA has been suggested to form by spontaneous isomerization from the all-trans form. Although atRA and 9cRA are the two best-characterized retinoids, other bioactive forms also exist in vivo, for example 3,4-didehydroRA and 4-oxo-RA [17, 18]. 4-oxo-RA is one of several breakdown products of atRA in catabolic reactions catalyzed by a family of cytochrome P450 enzymes, the CYP26 family [19–21], involved in attenuating the RA signal (see Fig. 1A). In early development, most embryonal RA is thought to be synthesized by RALDH2 [22–26]. Intriguingly, a comparison of the expression patterns of RALDH2 and CYP26 suggests that both enzymes are present in complementary and nonoverlapping domains, thereby creating areas differing in their RA levels [19, 24, 25]. CYP26 expression is restricted to the anterior- and posterior-most structures of the embryo, including the presumptive head and tailbud regions, keeping RA levels low. RALDH2 on the other hand is expressed in more central areas of the embryo, from the posterior end of the developing hindbrain to the anterior regions of the developing tail, and is responsible for RA synthesis in this region. Targeted

2 Retinoid Receptors Transduce Retinoic Acid Signals 3

disruption of the CYP26 gene gives rise to embryos where anterior brain structures are transformed to more posterior ones, resembling the morphogenetic defects generated by excess RA administration [5, 27, 28]. Conversely, disruption of the RALDH2 gene produces embryos with phenotypes indicating an expansion of anterior brain structures, reminiscent of the defects generated by vitamin A deficiency [6, 29, 30]. Taken together, these results suggest that disturbances in the graded synthesis of RA leads to posteriorization (excess RA) or anteriorization (shortage of RA) of structures in the developing head, largely due to misexpression of Hox genes. In an elegant study, Koide and co-workers show that active repression of RA target genes needs to be maintained for correct development of anterior regions [31]. This suggests that the mere absence of RA in anterior tissues is not sufficient for normal development, but that active repression of RA-responsive genes per se is essential for correct anterior patterning. Interestingly, retinoid receptors are directly involved in this repression by a mechanism that will be further discussed below (see “co-repressors” below). Within cells, cellular retinol binding proteins I and II (CRBP-I and -II) and cellular retinoic acid binding proteins I and II (CRABP-I and -II), act as cytoplasmic carriers for ROL and RA, respectively. Based on expression studies, it was thought that CRBPs function in protecting retinol from the cellular environment and presenting ROL to RA synthesizing enzymes. In contrast, CRABPs were suggested to be important in sequestering and promoting breakdown of excess RA in embryonic regions sensitive to the teratogenic effects of retinoids (see, for example, Refs [32–34]). However, animals carrying null mutations in both CRABP-I and -II are indistinguishable from wild-type littermates, suggesting that CRABPs are dispensable for normal embryonal development and adult physiology [35]. On the other hand, although not essential for embryonal development, adult mice lacking CRBP-I show decreased liver retinyl ester storage and predisposition to vitamin A deficiency [36], suggesting an important role for CBRP-I during vitamin A-limiting conditions. Interestingly, two CRBP-III genes have recently been identified [37, 38], showing partially overlapping patterns of expression, and it will be interesting to see what roles these proteins might play in vivo.

2 Retinoid Receptors Transduce Retinoic Acid Signals

The cloning and characterization of the nuclear hormone receptors (NRs) that bind and transduce the retinoid signal represent landmarks in our understanding of retinoid physiology (see references in Ref [39]). Two families of retinoid receptors exist, the retinoic acid receptors (RARs) and the retinoid X receptors (RXRs), each present in three isotypes, α, β, and γ (reviewed in Refs [39–41]). In addition, each RAR and RXR isotype exists in several isoforms (for example RARα1 and α2) due to differential promoter usage. Expression studies of RARα, β, and γ show that RARα is ubiquitously expressed, while RARβ and RARγ show a more temporal and spatial restriction, often in a non-overlapping fashion [32–34, 42, 43]. RXRs


are similarly differentially expressed both during development and in adult animals [42–45]. RXRβ is expressed in a general fashion, while RXRα is abundant in tissues associated with lipid metabolism. RXRγ expression is highly restricted to a few distinct areas, including the developing striatum and spinal motor neurons (where it is co-expressed with RARβ). A strong effort has been made during the last few years to understand the specific functions of the different retinoid receptors during development. Despite the unique distributions of the different receptors, genetic ablation studies have revealed a surprising redundancy in function among the different members of each receptor subtype. The results of single and double RAR or RXR mutants, as well as compound RAR/RXR mutants, have been extensively reviewed [41, 46]. Taken together, the results suggest that RARs and RXRs are important for correct regulation of numerous developmental processes. Although many known VAD phenotypes are not apparent in single RAR−/− mice, simultaneous ablation of two RAR genes recapitulates most aspects of VAD. Additionally, compound mutants of RXRα and RARα, β, or γ together reproduce almost the entire VAD phenotype spectrum, supporting the notion that RAR/RXR heterodimers are the active units for retinoid signaling in vivo (summarized in Ref [46]). Interestingly, a number of malformations not described in VAD studies are also observed, either suggesting roles for unliganded receptors or reflecting difficulties in achieving complete VAD by dietary deprivation.

3 Retinoid Receptors Belong to the Nuclear Hormone Receptor Family

NRs comprise a large and evolutionary well conserved family of transcription factors found in organisms as diverse as nematodes, flies, and mammals (reviewed in Refs [47–49]). NRs are thought to function as ligand-activated transcription factors, exerting widely different biological functions by regulating target gene expression positively and/or negatively, and include the receptors for certain small, lipophilic molecules. RARs bind both all-trans and 9-cis retinoic acids, whereas RXRs only bind 9-cRA (see Ref. [50] and references therein). Retinoid receptors activate transcription by recognizing and binding consensus sequences known as RA response elements (RAREs) in the promoters of target genes (see below). RAR binds DNA as a heterodimer with RXR, while RXR also has the ability to bind DNA as a homodimer. Additionally, RXR forms heterodimers with a number of other NRs, including the receptors for thyroid hormone (TR) and vitamin D3 (VDR) [50], thereby coupling retinoid signaling to a multitude of other cellular signaling pathways.

4 Nuclear Receptors Share a Common Structure

As mentioned above, retinoid receptors belong to the NR family of transcription factors. With only a few exceptions, all NRs share a common structure of functionally

4 Nuclear Receptors Share a Common Structure 5

Fig. 2 Nuclear receptor domains and functions. (A) NRs consist of defined domains, with variable degrees of conservation within the NR superfamily. The NTD, which is the most variable, has a ligand-independent trans-activation function (AF-1) shown to be important in basal transcription by some receptors. The more conserved DBD and the LBD are responsible for DNA and ligand binding, respectively. In addition, both domains play important roles in dimerization. A ligand-depen-dent transactivation function (AF-2) is localized to the LBD. NTD,

N-terminal domain; DBD, DNA binding domain; LBD, ligand binding domain; AF-1, activation function 1; AF-2; activation function 2. (B) Structures of the RXR DBD homodimeric complex (left) and of the RXRRAR DBD heterodimeric complex (right) on DR1 DNA response elements. Contacts between receptor partners form over the minor groove of the DNA helix, with additional protein-DNA contacts stabilizing the complex. DR1, direct repeat 1. Modified from Ref. [52].

separable domains, including an N-terminal domain (NTD), a central DNA binding domain (DBD), and a C-terminal ligand binding domain (LBD) (Fig. 2A). The highly conserved DBD is responsible for recognizing and binding to specific DNA sequences in the promoters of target genes, and is also involved in dimerization between receptors. The LBD, besides binding ligand, plays an essential role in the initial interaction between receptor heterodimers, as well as in ligand-dependent transactivation. The LBD also harbors a ligand-dependent activation function (AF-2), mediating ligand-dependent interactions with so-called co-activators (see below). The NTD is less conserved between different NRs, both in amino acid composition and length, and contains a ligand-independent activation function (AF-1), important in the basal transcriptional activity of some receptors. A hallmark of the NR family is the well-conserved DNA binding region. This cysteine-rich zinc binding domain, has been shown to contain structural features important for correct DNA target sequence recognition and binding, as well as dimerization (Fig. 2B) (see Ref. [50] and references therein). Most NRs bind socalled hormone response elements (HREs), arranged as one or two half-sites of the consensus nucleotide sequence 5 -AGGTCA-3 . Half-sites can be arranged as


direct- (DR), inverted- (IR), or everted repeats (ER), with a varying number of nucleotides separating the repeats. Studies of receptor dimers bound to DNA have shown that RXR and its heterodimer partners bind response elements arranged as two direct repeats spaced by one to five nucleotides (DR1 to DR5) ([51]; reviewed in Ref. [52]). Depending on the spacing between the two repeats, different RXR heterodimers will bind and activate transcription. It seems that both heterodimer partners cooperate to ensure correct binding specificity and affinity by making partner-specific protein contacts that stabilize the complex only on a correctly spaced DR motif. Additionally, RXR has the ability to switch its polarity on DR elements, binding either the upstream or the downstream half-site [52]. Importantly, the RXR heterodimer partner not only influences the response element of choice, but also the ability of RXR to become activated by ligand. For example, heterodimers between RXR and the peroxisome proliferator activated receptors (PPARs) are induced by both PPAR and RXR ligands, therefore said to be permissive to RXR activation. RAR-RXR heterodimers, on the other hand, are non-permissive in that they require binding of RAR ligands in order to become responsive to RXR ligands [53–55]. It has been suggested that the initial agonistinduced interaction between RAR and co-activators (see below) is necessary to induce the correct structural rearrangements that allow RXR to become receptive to its ligand [56]. However, several groups have obtained results that are not easily explained by this model. For example, RAR–RXR heterodimers can become responsive to an RXR ligand even after addition of an RAR antagonist, a situation when co-activators are not recruited by RAR [57, 58]. It is therefore still somewhat unclear why some heterodimers are permissive while others are not. An additional dimerization interface important for the initial interaction between NRs is found in the neighboring ligand binding domain (see below). This region has been shown to mediate cooperative binding on all three classes of DNA repeats (direct, inverted, and everted) [59, 60]. In receptor heterodimers, the second dimerization region formed within the DBD restricts receptors to direct repeat elements.

5 The LBD and Ligand-dependent Transactivation

The crystal structure of the LBD of several NRs has been solved, including unliganded (apo) and liganded (holo) RXRα, as well as holo RARγ [61, 62] (reviewed in Ref. [63]). The results reveal a common fold, consisting of 12 α-helices (H1H12) and one β-turn, arranged in a three-layered antiparallel “sandwich” with a hydrophobic core (Fig. 3A). The center of this sandwich contains the ligand binding pocket, lined mostly by hydrophobic and polar residues. As mentioned, the LBD contains the ligand-dependent activation function AF-2, and residues critical for its function have been mapped to helix 12 (the AF-2 core) [64]. Crystallographic studies have provided a model that accounts for the structural transitions involved in ligand activation of NRs (reviewed in Refs [63, 65]). In this so-called “mouse-trap” model, the initial interaction between ligand and LBD

5 The LBD and Ligand-dependent Transactivation

Fig. 3 Schematic drawing of the LBD structure of apo RAR" (A), holo RAR( bound to all-trans retinoic acid (atRA) (B), and antag onist-bound estrogen receptor (ER) " (C). atRA is shown in “stick” form in the center of the LBD in (B), and the ER antagonist,

raloxifene, is depicted as a bent cylinder in (C). Note the different positions of helix 12 (shown in black) in each situation. The "-helices of the LBD are represented as numbered rods. Modified from Ref. [65].

leads to structural changes that trap the ligand within the LBD in an induced fit mechanism. These transitions are subsequently followed by a repositioning of helices 3 and 4, which together with helix 11 now move to form a hydrophobic cleft on the surface of the LBD. The most profound conformational change involves helix 12 (H12) itself (Fig. 3). In the absence of ligand, H12 protrudes from the LBD and is exposed to solvent, whereas in the holo-receptor, it rotates and folds back towards the LBD, thereby compacting its structure. In its final position, H12 seals the pocket, trapping the ligand inside (compare Fig. 3A and B). In addition, in this new conformation, H12 has a major role in positioning so-called co-activator proteins in the hydrophobic cleft formed by residues on helices 3, 4, and 11, a process important in transcriptional activation (discussed below). Once transcription of target genes has been initiated, the hormone response needs to be attenuated in order to control the transcriptional output. Cells can regulate the activity of proteins involved in transcriptional activation by affecting their stability. Several reports show that retinoid receptors are targeted for ubiquitin-mediated degradation upon ligand binding [66–69]. Although somewhat conflicting, the results suggest that only active receptors are degraded. Interestingly, a component of the proteasome that mediates the degradation of ubiquitinated proteins, SUG1, has also been shown to function as a co-activator for certain NRs, including RAR [70], by interacting with the AF-2 helix. This could provide a functional link between ligand-induced transcriptional activation and subsequent degradation of NRs.

7


6 Cross-talk

In addition to acting as ligand-dependent transcription factors, some NRs also become activated or inactivated in the absence of ligand, for example by phosphorylation of the receptor itself or of its co-regulatory proteins. This is especially the case for the steroid hormone receptors (reviewed in Ref [71]), but also the activity of retinoid receptors can be modulated through phosphorylation, both in the presence and absence of ligand (see, for example, Refs [72–74]). Phosphorylation of a serine residue in the RAR DBD has for example been shown to lower the interaction between RAR and RXR, thereby decreasing transcriptional activation, and could be a way of regulating the activity of receptors in vivo [75]. On the other hand, phosphorylation of serine residues in the N-terminal AF-1 of RAR has been shown to have a positive effect on ligand-independent transcription [74, 76]. Perhaps the best established example of cross-talk is the inhibition of AP-1 transcriptional activity by several NRs including retinoid receptors. These modulatory activities are believed to be critical for the common antiproliferative effects of retinoids and other NR ligands (reviewed in Ref. [77]).

7 Co-activators

NRs require accessory factors, so-called co-activators, in order for ligand-dependent activation of target genes to occur. The first co-activators to be described interacted with the estrogen receptor (ER) in the presence of ligand [78]. One such protein, RIP160 (for receptor interacting protein 160), later shown to be identical to SRC-1 (steroid receptor co-activator-1) [79], interacted with several NRs in a hormonedependent manner. Since then, an ever-increasing number of co-activators have been cloned and characterized (reviewed in Ref. [80]). The best-characterized co-activators belong to one of three classes: the p160 family, including SRC-1/NCoA1, GRIP-1/TIF2, and ACTR/pCIP/RAC3/AIB-1; the homologous CBP and p300 co-activators; and the recently isolated TRAP/DRIP complexes (see Ref. [81] for abbreviations and references) (Fig. 4). Members of the p160 family as well as CBP/p300 and p/CAF (CBP associated f actor) show intrinsic histone acetyltransferase (HAT) activity [82–86], suggesting that co-activators may play direct roles in chromatin remodeling at promoters by acetylating histone proteins [87]. The TRAP/DRIP complexes were isolated by their ability to interact with ligand-bound TR and VDR, respectively. They form large multiprotein complexes that share common subunits [88, 89], and are thought to play an important role as bridging molecules between DNA-bound NRs and the basal transcription machinery [90]. Co-activators interact with NRs via one or several leucine-rich β-helices, also known as NR boxes, with the consensus sequence LxxLL (where L corresponds to leucine and x is any amino acid residue) [91, 92]. Structural and functional studies indicate that the co-activator LxxLL helix is accommodated along the hydrophobic

7 Co-activators

Fig. 4 The different steps and proteins involved in NR transcriptional activation. DNA-bound NRs (here exemplified as an RAR-RXR heterodimer) inhibit transcriptional activation by recruiting corepressors with histone deace-tylase activity, keeping promoter DNA packed into histones, i.e. in a silent form. Upon exposure to ligand, co-repressors are released and co-activators of the p160 family and CBP/p300 are recruited, either before or together with complexes like the TRAP/DRIP proteins. The

ATP-dependent activity of the SWI/SNF complex, initially acts to unwind DNA at the promoter. Histone acetylation by coactivators allows stronger interaction between the NRs and DNA, ultimately leading to recruitment of RNA Pol II and other accessory factors. These proteins recognize and bind DNA sequences of the TATA box at the transcriptional initiation site. RNA Pol II is finally released from the promoter and initiates gene transcription. See text for abbreviations and further details.

cleft on the surface of the receptor LBD that forms upon ligand binding [93–96]. Two strictly conserved amino acid residues on the receptor form a “charge clamp” that correctly places the co-activator LxxLL motif on the LBD, leading to transcriptional activation of the receptor. In the crystal structures of ER and RAR bound to antagonists, the AF-2 helix of the receptor is not repositioned correctly as in the ligand-bound receptors. Instead, it is translocated to overlap the co-activator interaction site, thereby preventing co-activator binding [96–99]. This in turn would facilitate the recruitment of another group of regulatory factors, corepres-sors, explaining the molecular mechanism behind antagonistic repression of NRs. The development of novel techniques for studying protein–chromatin interactions at specific promoters has yielded exciting new insights explaining the pro-

9


cess of transcriptional initiation (reviewed in Ref [100]). The results suggest that ligand-bound receptors continuously cycle on and off target promoters, transiently interacting with response elements on DNA, recruiting cofactors and RNA polymerase II to initiate transcription, and subsequently dissociating from DNA again [101]. Furthermore, it seems that cofactors with HAT activity not only acetylate histones at the promoter itself, but also further away on the DNA template to allow better access of proteins to promoter DNA. The exact sequence of events involved in transcriptional initiation is however unclear, and both a stepwise or concerted recruitment of p160 and TRAP/DRIP co-activators to the initiation complex has been suggested [102, 103] (see Fig. 4). 8 Co-repressors

Certain NRs, such as RAR and TR, repress basal transcription in the absence of ligand by binding the promoters of target genes, a process known as silencing [104]. The molecular mechanisms behind this phenomenon involves two related corepressor proteins, N-CoR (nuclear receptor co-repressor) and SMRT (silencing mediator of retinoid and thyroid hormone receptor), which both interact with RAR and TR in a ligand-independent manner [105, 106]. It has recently been shown that corepressors bind a region on the surface of the receptor LBD that overlaps the co-activator-interacting site. N-CoR and SMRT contain α-helical structures similar in sequence to the LxxLL helix of co-activators, with the consensus LxxI/HIxxxI/L (where L is leucine, I is isoleucine, H is histidine, and x is any amino acid residue) [107–109]. Thus, due to sequence similarities, corepressors are able to bind the same region on the NR LBD as co-activators, thereby masking the co-activator site and repressing transcription. Structural transitions in the LBD that occur upon ligand binding move the corepressor and allows AF-2 helix repositioning, which further displaces the corepressor. The extended corepressor motif no longer fits in the cavity due to steric hindrance by the AF-2 helix, and instead the co-activator gains entry to the site. Analogous to co-activators forming large multiprotein complexes, N-CoR and SMRT also interact with other proteins to repress transcription. Histone deacetylases (HDACs 1 and 2) are bridged to unliganded NRs via N-CoR/SMRT and mSin3 co-repressors, thereby mediating silencing (reviewed in Ref. [80]) (Fig. 4). Deacetylation of core histones by HDACs is recognized as a mechanism for keeping chromosome domains transcriptionally silent, and would explain how NRs mediate repression. 9 Nuclear Receptors from an Evolutionary Perspective

Based upon the homologies within the superfamily, NRs have been divided into six subfamilies [110]. The ability to bind ligand and the structure of the ligand in

10 Fatty acids as Endogenous Ligands for RXR 11

several cases seems unrelated to which subfamily the respective receptors belong. For example, RAR and TR, which bind two unrelated ligands, are nonetheless more related in sequence than, for example, RAR and RXR, which both bind retinoic acid isomers, suggesting that ligand-binding ability is independent from the evolutionary origin [111]. The fact that orphan receptors are present in all subfamilies, whereas liganded receptors are not, further suggests that the ancestral receptor did not have a ligand. Interestingly, several recent reports have revealed the unexpected presence of fatty acids/lipids in the ligand-binding pockets of several NR crystals, including RXR, the Drosophila RXR ortholog Ultraspiracle, and retinoic acid-related orphan receptor β (RORβ) [99, 112–114]. Apparently, these LBDs bind lipids derived from the bacterial strains used to express the proteins, and it seems likely that such binding is a prerequisite for a stable LBD conformation. These findings suggest an intriguing model explaining how the ligand-activated receptors of today have evolved. Accordingly, the primordial NR may have used a ubiquitous lipid as a structural element/cofactor to stabilize the LBD in its active state. In this view, the primordial “lipid binding domain” was permissive to those evolutionary adaptations necessary for a regulated domain to evolve, i.e. a structure that is regulated by bona fide ligand interactions. Ligand identity can thus be viewed as the result of an independent, convergent evolutionary process that took place in several different receptors, unrelated to their evolutionary origin within the NR superfamily.

10 Fatty acids as Endogenous Ligands for RXR

The signaling status of RXR in vivo is still a matter of debate, as its proposed natural ligand, 9-cis RA, has proved difficult to identify in mammalian tissue [115]. Nonetheless, several reports show that simultaneous addition of both RXR- and RAR-specific ligands often leads to synergistic biological effects (see, for example, Refs [57, 116, 117]). Therefore, it seems likely that ligand-induced activation of RXR does occur, a conclusion that has been corroborated by experiments in transgenic mice [14, 118]. Recent experiments have identified novel endogenous RXR ligands and expanded the perspectives on RXR functions in vivo. Interestingly, recent data suggests that the chlorophyll metabolite phytanic acid can bind and activate RXR [119, 120] (Fig. 1B). Although phytanic acid is a low-affinity ligand and high serum levels would be required to activate RXR, significant activation might occur in patients with certain metabolic disorders such as Refsum’s disease [119, 120]. Docosahexaenoic acid (DHA or C22:6 cis 4, 7, 10, 13, 16, 19) (Fig. 1B), a long-chain polyunsaturated fatty acid (PUFA), is an RXR ligand, does not activate RAR, TR, or VDR, and promotes the interaction between RXR and the co-activator SRC-1 [121]. Interestingly, other PUFAs including docosatetraenoic (C22:4 cis 7, 10, 13, 16), arachidonic (C20:4 cis 5, 8, 11, 14) and oleic (C18:1 cis 9) acids, also activate


RXR, although DHA is more potent [121]. Unlike 9cRA, which binds and activates RXR with high affinity, DHA is a low-affinity ligand for this receptor, reaching half-maximal activation at a concentration of about 40-50 µM (our unpublished observations). However, while 9cRA has been difficult to identify in vivo, DHA, which accumulates in the mammalian CNS during late gestation and early postnatal development, constitutes between 30 and 50% of total membrane-bound fatty acids in the postnatal brain (reviewed in Refs [122] and [123]). Therefore, it seems likely that sufficiently high intracellular concentrations of free DHA may exist in neurons. Little is known about the mechanisms of DHA release from its phospholipid compartment, although some reports implicate phospholipases A2 and C in this process (see, for example, Ref [124]). This mobilization could potentially supply enough free DHA to compensate for its rather low affinity towards RXR. Deficiencies in DHA have been shown to cause neurological abnormalities, impaired learning abilities and growth retardation (see, for example, Refs [125] and [126]). Memory deficits have recently been shown to improve upon either RA or DHA treatment, suggesting functional overlap between the two pathways [127–129]. Interestingly, the analyses of gene targeted mice lacking one or several genes encoding RXR isotypes have demonstrated overlapping functions with DHA, for example in the process of memory formation [130], vision [131], and reproduction [132]. Moreover, several RXR heterodimerization partners, such as PPARs, liver X receptors and farnesoid X receptor, contribute to energy and nutritional homeostasis in response to their respective ligands (reviewed in Ref. [133]). DHA could thus play an important modulatory role in these processes by binding and influencing the RXR subunit of such heterodimers. Structural analysis of the RXR LBD bound to DHA shows that the receptor LBD adopts the canonical conformation of a ligand bound NR, with the activation function AF-2 helix (H12) packed towards the hydrophobic groove formed on the surface of the protein [134]. The highly flexible fatty acid molecule is optimally accommodated to the ligand cavity of RXR, occupying 80% of the pocket (72% for 9cRA) and making ligand-protein contacts similar to those of 9cRA.

11 Conclusions

A few issues stand out as particularly likely to attract researchers’ attention over the next coming years. For example, although structural studies have provided valuable information as to how nuclear receptors in general, and retinoid receptors in particular, interact with each other, with DNA, and with co-regulatory proteins, most such studies have been performed on isolated domains of the receptors. Notably, crystal structures of receptors bound to DNA have been produced using only the DNA binding domain. Similarly, ligand and co-activator binding studies have focused on the ligand binding domain. Undertaking similar studies using

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1

Human ABC Transporters: Function, Expression, and Regulation Gerd Schmitz and Thomas Langmann

Universität Regensburg, Regensburg, Germany


1 Introduction

Members of the ATP binding cassette (ABC) transporter superfamily, which are found in all three kingdoms of life, namely prokaryotes, archae bacteria and eukaryotes, represent one of the biggest protein families described so far. ABC transporter genes are highly conserved among different genomes and have been sustained throughout the evolutionary tree. They are usually multispan membrane proteins that mediate the active uptake or efflux of specific substrates across various biological membrane systems [1]. The development of these two different directions of transport, import or export, most likely occurred even before the differentiation of eukaryotes from prokaryotes [2]. In bacteria, these bidirectional transport functions either provide a mechanism for nutrition supply, as in the MalK transporter from Escherichia coli [3], or serve as a defense mechanism, allowing bacteria to protect themselves from their own or foreign toxins [4]. During evolution, eukaryotes have developed specialized ABC proteins as a type of early innate immune system, protecting cells from harmful substances. Thus in the human system several ABC proteins (MDRs, MRPs, ABCG2) are responsible for increased drug exclusion in compound-treated tumor cells, providing cellular mechanisms for the development of multidrug resistance [5]. ABC transporters have also received attention because mutations in these molecules are the cause of various inherited human diseases, including familial HDL deficiency (ABCA1) [6–8], some chorioretinal diseases (ABCR or ABCA4) [9], progressive familial intrahepatic cholestasis (PFIC) type II (PFICII: BSEP or ABCB11) and type III (PFICIII: MDR3 or ABCB4) [10–12], Dubin-Johnson syndrome (cMOAT or ABCC2) [13], pseudoxanthoma elasticum (MRP6 or ABCC6) [14], adrenoleukodystrophy (ALDR or ABCD2) [15], and β-sitosterolemia [16]. In Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Human ABC Transporters: Function, Expression, and Regulation

addition, functional polymorphisms have been described in various ABC genes including ABCA1 [17], ABCA4 [18], and ABCG1 [19]. Most interestingly, a significant number of human ABC transporters are critically involved in bile acid, phospholipid, and sterol transport [20–23], whereas the expression of these ABC proteins is itself controlled by lipids. Therefore, it is obvious that ABC transporters are promising target molecules for the treatment of lipid disorders. In this article, we summarize the structural features of human ABC transporters, their role in human disorders and especially highlight the function and regulation of ABC proteins in cellular and total body lipid homeostasis. 2 Structural Features of ATP Binding Cassette (ABC) Transporters

A functional ABC transporter protein usually consists of two transmembrane domains (TMD) and two nucleotide binding domains (NBD) or ATP binding cassettes (ABC). The characteristic ABC domain is composed of two short, conserved peptides, the Walker A and Walker B motifs [24], which are required for ATP binding and which are also found in other ATP binding proteins [25] (Fig. 1). An additional element, the signature motif, is located between both Walker motifs and is characteristic for each ABC subfamily [26]. The TMD and ABC domains are either present in one polypeptide chain (full-size transporter) or in two poly-peptides (half-size transporter) and several arrangements of the TMD and ABC motifs are found in human ABC proteins (Fig. 1).

Fig. 1 Diagram depicting domain arrangements of human ABC transporters. The ATP binding cassette (ABC) consists of Walker A and Walker B motifs, separated by the signature motif characteristic for each ABC transporter subfamily [24, 28]. The membrane spanning domains are depicted as barrels. (A) The TMD0-(TMD-ABC)2 structure of ABCC (MRP) family members is shown. In addition to the regular full-size type containing the (TMD-ABC)2 domain

arrangement, this type displays an additional five trans-membrane domains termed TMD0. (B) Prototype ABC transporter with the (TMD-ABC)2 structure. (C) Two alternative types of half-size molecules, TMDABC and ABC-TMD. Only corresponding half-molecule organizations are able to form heterodimers. (D) The (ABC)2 type of molecules lacking transmem-brane domains is unlikely to function as transporter.

3 Overview of Human ABC Gene Subfamilies

Among the full-size transporters domain structures such as (TMD-ABC)2 as well as TMD0-(TMD-ABC)2 (which contains an additional N-terminal series of five transmembrane spans) occur. (TMD-ABC)2 structures are represented in the ABCA, ABCB, and ABCC families, whereas the TMD0-(TMD-ABC)2 arrangement is solely present in specific members of the ABCC subfamily (Tab. 1). The (ABCTMD)2 is only found in yeast and not present in human ABC molecules. Half-size transporters can either occur in the TMD-ABC organization, as it is the case within the ABCD subfamily, or as ABC-TMD, which is found in the ABCG group of ABC proteins (Tab. 1). In both cases, creation of a functional transporter requires the assembly as a homodimer or heterodimer. Although the final destination of full-size transporters is the plasma membrane, these proteins are also found intracellularly as a result of vesicular trafficking processes. Also, the localization in intracellular membrane-bound vesicles, collectively named multivesicular bodies (MVBs), is conceivable [27]. Most half-size molecules are routed to intracellular membrane systems such as mitochondria, peroxisomes, the endoplasmic reticulum, and the Golgi compartment [28]. However, a member of the ABCG subfamily, ABCG2, has been localized to the plasma membrane [29]. In contrast to these membrane-spanning ABC transporters, proteins from the ABCE and ABCF subfamilies do not harbor TMD at all and contain a (ABC)2 domain structure (Fig. 1). As a consequence, they are not likely to be involved in any membrane transport function. Moreover, ABCE1 binds oligoadenylate, which is produced upon viral infections and seems to be a part of the innate immune system by controlling the RNase L pathway [30]. ABCF1 is associated with ribosomes and interacts with eukaryotic initiation factor 2 (eIF2) and thereby plays a key role in the initiation of mRNA translation [31]. The group of membrane-spanning ABC transporters can be split into two different sections depending on their mode of action. The active transporters or pumps, such as members of the ABCB (MDR/TAP) subfamily, couple the hydrolysis of ATP and the resulting free energy to movement of molecules across membranes against a chemical concentration gradient [32]. In contrast, recent work has identified several ABC proteins that show nucleotide binding and a subsequent conformational change but very low ATP hydrolysis. These molecules mainly function as transport facilitators and include ABCC7 (CFTR) [33], ABCC8 (SUR1), ABCC9 (SUR2) [34], and ABCA1 [35].


A comprehensive description of the currently known human ABC transporters is given in Tab. 1. The list contains fully characterized ABC genes, as well as gene annotations derived from sequence information based on the analysis of the human genome [36, 37] and uses the proposed nomenclature of the Human Gene Nomenclature Committee (HUGO). The synonyms, the chromosomal location, the domain structure, and the tissue specificity and cellular location of each gene are itemized. Furthermore, the lipid-sensitive regulation and the known or putative function of

3

Gene

Alternative name

Location

Domain structure

Tissue expression and cellular location

Lipid regulated

Known or putative transported molecule

ABCA1

ABC1

9q31.1

TMD-ABC) 2

Macrophages, liver

+

ABCA2 ABCA3 ABCA4 ABCA5 ABCA6 ABCA7

ABC2 ABC3 ABCR

9q34 16p13.3 1p22.1–p21 17q24 17q24 19p13.3

(TMD-ABC)2 (TMD-ABC)2 (TMD-ABC)2 (TMD-ABC)2 (TMD-ABC)2 (TMD-ABC)2

Brain Lung Photoreceptors Muscle, heart, testes Liver Spleen, thymus, PBMC

+ + − + + +

Choline-phospholipids and cholesterol Estramustine, steroids Surfactant phospholipids N-Retinylidene-PE

17q24 17q24 17q24 2q34 7p11–q11

(TMD-ABC) 2 (TMD-ABC)2 (TMD-ABC)2 (TMD-ABC)2 (TMD-ABC)2

Ovary Heart Muscle, heart Stomach Low in all tissues

− + − − −

ABCA8 ABCA9 ABCA10 ABCA12 ABCA13

Phospholipids Sphingolipids (e.g. ceramide) and serine-phospholipids (e.g. PS)

ABCB1

MDR1

7p21

(TMD-ABC)2

Excretory organs, apical membrane

+

ABCB2 ABCB3 ABCB4

TAP1 TAP2 MDR 3

6p21 6p21 7q21.1

TMD-ABC TMD-ABC (TMD-ABC)2

− − +

ABCB5 ABCB6 ABCB7

MTABC3 ABC7

7p14 2q36 Xq12–q13

(TMA-ABC)2 TMD-ABC TMD-ABC

Ubiquitous, ER Ubiquitous, ER Liver, apical membrane Ubiquitous Mitochondria Mitochondria

Phospholipids, PAF, aldoster-one, cholesterol, amphiphiles, β-amyloid peptide Peptides Peptides Phosphatidylcholine

− + −

Fe/S clusters Fe/S clusters


Table 1 Overview of human ABC gene subfamilies

− +

Fe/S clusters

− +

Fe/S clusters Monovalent bile salts (e.g. TC)

TMD0-(TMD-ABC) 2

Lung, testes, PBMC

+

10q24

TMD0-(TMD-ABC)2

Liver

+

MRP3

17q21.3

TMD0-(TMD-ABC)2

Lung, intestine, liver, basolateral membrane

−

ABCC4

MRP4

13q32

(TMD-ABC)2

Prostate

+

ABCC5

MRP5

3q27

(TMD-ABC)2

Ubiquitous

+

ABCC6

MRP6

16p13.1

TMD0-(TMD-ABC)2

Kidney, liver

−

ABCC7 ABCC8

CFTR SUR1

7q31.2 11p15.1

(TMD-ABC)2 TMD0-(TMD-ABC)2

Exocrine tissue Pancreas

− −

GSH-, glucuronate-, sulfate-conjugates, GSSG, sphingolipids, LTC4 , PGA1, PGA2 , 17β-glucuronosyl estradiol GSH-, glucuronate-, sulfate-conjugates, bilirubin glucuronide, LTC4 , 17β-glucuronosyl estradiol, taurolithocholate 3-sulfate, anionic drugs glucuronate-, sulfate-conjugates, 17β-glucuronosyl estradiol, taur-olithocholate 3-sulfate Xenobiotics, nucleosides ATP/ADP/AMP/adenosin, GTP/GDP) Xenobiotics, nucleosides (ATP/ADP/AMP/adenosin, GTP/GDP) Anionic cyclopentapeptides (e.g. BQ123) Chlorideions, ATP Sulfonylureas

MAB C1

7q36 12q 24

TMD-ABC TMD-ABC

ABCB10 ABCB11

MTABC2 BSEP

1q42 2q24

TMD-ABC (TMD-ABC)2

ABCC1

MRP1

16p13.1

ABCC2

MRP2

ABCC3


Mitochondria Heart, brain, lysosomes Mitochondria Liver, apical membrane

ABCB8 ABCB9

5

Gene

Alternative name

Location

Domain structure

Tissue expression and cellular location

Lipid regulated

Known or putative transported molecule

ABCC9 ABCC10 ABCC11 ABCC12

SUR2 MRP7 MRP8 MRP9

12p12.1 6p21 16q11–q12 16q11–q12

TMD0-(TMD-ABC)2 TMD0-(TMD-ABC)2 (TMD-ABC)2 (TMD-ABC)2

Heart, muscle Low in all tissues Low in all tissues Low in all tissues

− − − −

Sulfonylureas

ABCD1 ABCD2 ABCD3 ABCD4

ALD ALDR PMP70 PMP69

Xq28 12q11–q12 1p22–p21 134q24.3

TM-ABC TM-ABC TM-ABC TM-ABC

Peroxisomes Peroxisomes Peroxisomes Peroxisomes

+ + − +

Very long-chain fatty acids Very long-chain fatty acids Very long-chain fatty acids Very long-chain fatty acids

ABCE1 ABCF1

OABP ABC50

4q31 6p21.33

(ABC)2 (ABC)2

Ovary, testes, spleen Ubiquitous

− −

7q36 3q25

(ABC)2 (ABC)2

Ubiquitous Ubiquitous

− −

Oligoa denylate Translation elongation initiation factor 2

21q22.3 4q22 11q23 2p21 2p21

ABC-TMD ABC-TMD ABC-TMD ABC-TMD ABC-TMD

Ubiquitous Placenta, intestine Liver Liver, instestine Liver, instestine

+ − + + +

ABCF2 ABCF3 ABCG1 ABCG2 ABCG4 ABCG5 ABCG8

White MXR White2 White3 White 4

Phospholipids, cholesterol Drug resistance Plant and shellfish sterols Plant and shellfish sterols

Notes: The currently known 48 human ABC transporters from six different subfamilies and their typical features are listed. The proposal of the Human Genome Organization (HUGO) for the numbering of human ABC transporter genes has been used and the common names have been included additionally. The domain structure of ABC transporters has been adapted from Klein et al. [28] or by generation of hydrophobicity plots. An excellent regularly updated website established by Michael Müller (http://nutrigene.4t.com/humanabc.htm) provides supplementary information concerning database entries.


Table 1 Continued


human ABC transporters is mentioned. A short outline of each of the six known human ABC gene subfamilies is presented in the following paragraphs. 3.1 The ABCA (ABC1) Subfamily

The ABCA family contains solely full-size transporters (Tab. 1), and with ABCA1, ABCA4 (ABCR), and ABCA2 the largest proteins with 2261, 2273, and 2436 amino acids, respectively. Most of the ABCA proteins are expressed ubiquitously at low levels and also predominantly in specific tissues, such as ABCA1 in macrophages and ABCA4 (ABCR), which seems to be restricted to photoreceptor cells [9]. In contrast to all other ABC subgroups, the ABCA subfamily has no counterpart in yeast and appears for the first time in Caenorhabditis elegans [38]. Based on the genomic locations and phylogenetic analyses [39], two distinct divisions of ABCAs can be formed. The first group contains five genes located in a cluster on chromosome 17q24 (ABCA5, ABCA6, ABCA8, ABCA9, and ABCA10) and the second group consists of seven genes distributed over six different chromosomes (ABCA1, ABCA2, ABCA3, ABCA4, ABCA7, ABCA12, and ABCA13). Interestingly, the transcriptional control of at least seven ABCA members (Tab. 1) is controlled or influenced by lipids [40–45], indicating an important role of the whole ABCA subfamily in cellular lipid transport processes [23, 46]. ABCA1, the founding member of the family, is under extensive investigation and it is now widely accepted that its predominant role is associated to the regulation of cellular phospholipid and cholesterol release via an indirect mechanism, possibly by ATP-sensitive regulation of an as yet uncharacterized molecule [35]. In contrast, ABCA4 is an active retinoid-PE complex transporter which displays strong, lipid-activated ATPase activity [47–49] comparable to active pumps such as ABCB1 (MDR1). In addition to the high expression in neuronal tissues [50], ABCA2 is also present in liver, kidney, and macrophages [45, 51]. ABCA2 co-localizes with endosomal/lysosomal markers and contains a lipocalin signature motif, a feature found in a family of proteins linked to the transport of sterols including retinoids, steroids, lipids, and bilins [52]. Thus it is conceivable that ABCA2 sequesters lipids or lipid-steroid complexes via its lipocalin domain into endoso-mal/lysosomal vesicles, which could serve as a secretory pathway for these molecules [51]. This hypothesis is further supported by the lipid-sensitive induction of ABCA2 in human macrophages [45]. Although ubiquitously expressed, the ABCA7 protein is predominantly found in myelo-lymphatic tissues [43, 44] and a pivotal role in the development of hematopoietic cell lineages has been suggested [53]. Interestingly, there is recent evidence that ABCA7 may be involved in the transport of phosphatidylserine and ceramidespecies and thus be linked to apoptotic processes [54]. The ABCA3 protein is an integral part of the surfactant lamellar body membrane in lung alveolar type II cells [55]. Pulmonary surfactant is a complex mixture consisting of phospholipids, neutral lipids, and specific proteins. It is essential for normal lung function because it reduces surface tension at the air-liquid interface

7


of alveolar spaces. Phospholipids comprise 80% of the mass of surfactant, of which 80–85% are phosphatidylcholines (PC). Among the PC molecular species, dipalmitoyl-PC (PC16:0/16:0) is the principle surface tension-lowering molecule, ranging from 40 to 60 mol% in adult mammals, whereas disaturated palmitoylmyristoyl-PC (PC16:0/14:0), together with the monounsaturated palmitoylpalmitoleoyl-PC (PC16:0/16:1) and palmitoyloleoyl-PC (PC16:0/18:1), comprise up to 38% of total PC [56]. Lung surfactant also contains four unique proteins: surfactant protein A (SP-A), SP-B, SP-C, and SP-D [57]. Lamellar bodies are enriched in SP-B and SP-C and it has been proposed that these hydrophobic proteins are secreted together with the phospholipids. SP-A and SP-D are secreted independently of lamellar bodies. The localization of ABCA3 in lamellar bodies of alveolar type II cells and the finding that raised ATP levels in bronchoalveolar lavage fluid are sufficient to stimulate surfactant secretion [58] implicate ABCA3 in the processing of pulmonary surfactant by transporting phospholipids and/or specific surfactant proteins for secretion. Since lamellar bodies are also important structures in other cells with barrier function such as keratinocytes in the skin and intestinal epithelial cells and because ABCA3 is expressed in other cells as well, a similar function in this cellular system could be envisioned [22, 59].

3.2 The ABCB (MDR/TAP) Subfamily

The ABCB family is the only subgroup of human ABC transporters that contains full-size (ABCB1, ABCB4, ABCB5, and ABCB11) and half-size transporters (ABCB2, ABCB3, ABCB6-ABC10) (Tab. 1). ABCB1 (MDR1) is probably the best studied ABC transporter. It was the first human ABC protein to be cloned [60] and has the ability to mediate multidrug resistance in cancer cells. ABCB1 is localized to the apical membrane of polarized cells and the major sites of expression are found in the liver, the intestine, and the blood-brain barrier. One proposed physiological function of MDR1 is the protection of cells by exporting lipophilic cytotoxic drugs. In addition to ABCB4 (MDR3), which only translocates phosphatidylcholine (PC) across membranes [61], ABCB1 can transport a variety of lipids: PC analogs, phosphatidylethanolamine (PE), sphingomyelin (SM), cholesterol, and glucosylceramide (GlcCer) molecules, which carry a shortened fatty acid at the C2-position of the glycerol or sphingosine backbone [62]. Of particular interest is the finding that ABCB1 (MDR1)-overexpressing cells have elevated levels of cholesterol, GlcCer [63–65] and caveolin 1 [66], all of which are constituents of raft plasma membrane microdomains involved in pathways of lipid efflux from cells. However, these data need further confirmation. Since ABCB1 itself is localized in Triton X-100-insoluble caveolin/cholesterolrich domains [67] and because cholesterol can directly interact with the substrate binding site of ABCB1 [68], it has been suggested that the transport of cytotoxic drugs, which are mostly lipophilic, is coupled to the translocation of cholesterol and sphingolipids [69].


A recent report has indicated that ABCB1 is also involved in the secretion of platelet-activating factor (PAF) [70]. PAF (1-O-alkyl-2-acetyl-sn-glycero-3phosphocholine) is a potent bioactive lipid that is synthesized by a broad range of cells, including circulating infammatory cells, endothelial cells, and epithelial cells. It has a variety of biological effects including activation of inflammatory cells and is involved in many pathological conditions, such as angiogenesis in breast cancers, metastasis, shock, sepsis and multiple organ failure. Since PAF is a naturally occurring short-chain phosphocholine and because MDR1 recognizes short-chain analogs of PC and is expressed in many cell types, including epithelial cells, a model for direct translocation of PAF across the plasma membrane has been proposed. Once present on the cell surface, PAF interacts with the PAF receptor on a neighboring cell and elicits its signaling mechanisms. An unexpected role of ABCB1 in the immune response has been recently identified: mdr1a−/− mice kept under pathogen-free conditions develop spontaneous intestinal inflammation. It is thought that this type of colitis is due to a disturbance of the mucosal layer as a consequence of a defect in the membrane integrity of intestinal epithelial cells [71]. In this context, altered intestinal intraepithelial lymphocyte populations and a disturbed cytokine response has been documented in mdr1a−/− mice [72, 73]. ABCB1 has also been implicated in the efflux of brain β-amyloid protein, since pharmacological blockade of ABCB1 rapidly decreases extracellular levels of βamyloid secretion. Also, in vitro binding studies showed that addition of synthetic human -amyloid peptides to hamster mdr1-bearing vesicles resulted in saturable uptake of these peptides, suggesting that they interact directly with the transporter [74]. These results and the finding that apolipoprotein E (apoE) is also associated with β-amyloid peptides [75] implies that ABCB1 can co-transport apoE and β-amyloid and thereby may contribute to the etiology of Alzheimer’s disease. Two half-size members of the subfamily, ABCB2 (TAP1) and ABCB3 (TAP2), are transporters associated with antigen presentation (TAP) and form a functional heterodimer to transport peptides from the cytoplasm into the endoplasmic reticulum, from where the presentation of peptide antigens via major histocompatibility complex (MHC) I starts [76] (Fig. 2). A transient complex containing a class I heavy chain-β 2 microglobulin (β2m) heterodimer is assembled onto the TAP molecule by numerous interactions with the ER chaperones calnexin, ERp57, calreticulin, and the specialized tetherin molecule, tapasin [77]. Most interestingly, virus-infected and malignant cells have developed strategies to escape immune surveillance by affecting TAP expression or function [78]. The immediate-early gene product ICP47 of herpes simplex virus type I binds to the cytoplasmic face of TAP and thereby blocks peptide entry, whereas the ER-resident human cytomegalovirus protein US6 inhibits TAP function by blocking the ER-luminal part of the transporter (Fig. 2) [79–81]. ABCB9, which is closely related to ABCB2 and ABCB3, has been mainly found in lysosomes [82]. Although ABCB9 has been proposed to be involved in TAP-dependent processes, its exact function is currently unknown. The remaining four ABCB proteins (ABCB6, ABCB7, ABCB8, and ABCB10) are all targeted to the inner mitochondrial membrane and play a role in cellular iron homeostasis by

9


Fig. 2 Proposed role of TAP proteins (ABCB2, ABCB3) in antigen presentation. En-dogenous proteins are degraded in the ubiquitin-proteasome pathway. The peptides are transported into the ER lumen by a fullsize complex composed of TAP1 and TAP2 [76]. The correct folding, assembly and load-

ing of MHC I molecules is mediated by numerous accessory proteins including calnexin, calreticulin, ERp57, tapasin, and TAP [77]. Stable MHC I-peptide complexes leave the ER through the Golgi compartment to the cell surface for recognition by T-cell receptors.

transporting iron-sulfur (Fe/S) cluster precursor proteins [82–85]. In this respect, a mutation in ABCB7, which is located on the X-chromosome, has been linked to X-linked sideroblastic anemia and ataxia (XLSA/A) (see Tab. 2) [86]. 3.3 The ABCC (CFTR/MRP) Subfamily

The ABCC subfamily comprises 12 full-size ABC proteins which perform such diverse functions as drug resistance, ion transport, nucleoside transport, and ion channel regulation (Tab. 2). A special subgroup within the ABCC family can be distinguished by the presence of a TMD0-(TMD-ABC)2 domain arrangement (Fig. 1A). Seven members display this special membrane topology (ABCC1, ABCC2, ABCC3, ABCC6, ABCC8, ABCC9, and ABCC10), whereas the other proteins in this subfamily exhibit the (TMD-ABC)2 structure. Although the TMD0 part is not required for transport activity, a linker region designated L0 is essential for proper

3 Overview of Human ABC Gene Subfamilies Table 2 Human ABC transporter genes and corresponding dieseases or phenotypes

Gene

Alternative name

Disorder or phenotype

Reference

ABCA1 ABCA4

ABC1 ABCR

ABCB1

PGY1, MDR1

6–8 9 9 9 9 9 9 59, 70

ABCB2 ABCB3 ABCB4

TAP1 TAP2 MDR3

ABCB7

ABC7

ABCB11

SPGP, BSEP

Familial HDL deficiency, Tangier disease Stargadt macular dystrophy (STGD) Fundus flavimaculatus (FFM) Retinitis pigmentosa 19 (RP) Cone-rod dystrophy (CRD) Cone dystrophy (CD) Age-related macular degeneration (AMD) Multidrug resistance, inflammatory bowel disease (ulcerative colitis) Immune deficiency Immune deficiency Progressive familial intrahepatic cholestasis type 3 (PFIC-3) Intrahepatic cholestasis of pregnancy (ICP) X-linked sideroblastosis and anemia (XLSA/A) Progressive familial intrahepatic cholestasis type 2 (PFIC-2)

ABCC1 ABCC2 ABCC6 ABCC7 ABCC8

MRP1 MRP2 MRP6 CFTR SUR1

Multidrug resistance Dubin-Johnson Syndrome (DJS) Pseudoxanthoma elasticum (PXE) Cystic fibrosis (CF) Persistent hyperinsulinemic hypoglycemia of infancy (PHHI)

162 13 14 163 184

ABCD1 ABCD3

ALD PXMP1, PMP70

Adrenoleukodystrophy (ALD) Zellweger syndrome 2 (ZWS2)

15 104, 105

ABCG2 ABCG5 ABCG8

ABCP, MXR, BCRP White3 White 4

Multidrug resistance β-Sitosterolemia β-Sitosterolemia

117 16 16

76-78 76-78 10–12 10–12 83 11

ABCC1function [87]. Among the (TMD-ABC)2 molecules, ABCC7 (CFTR) is characterized by an extraordinary domain structure: it contains a regulatory domain, which is controlled by cAMP/PKA-dependent phosphorylation and thereby enables ATP binding and hydrolysis at the nucleotide binding cassettes, which in turn control opening and closing of the chloride channel [88]. Mutations in ABCC7 (CFTR) cause cystic fibrosis by affecting numerous secretion processes. ABCC1, ABCC2, and ABCC3 are all able to transport anticancer drugs, whereby ABCC1 (MRP1) mainly transports glutathione-conjugated (GSH) molecules and therefore has been termed GS-X pump [89]. In addition to cancer drug resistance, the physiologic function of GS-X pumps is closely related with cellular detoxification, oxidative stress, and inflammation [90].

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ABCC2 (MRP2), which is located in the apical membrane of polarized epithelial cells and particularly to the canalicular membrane of hepatocytes, appears to participate in the hepatobiliary secretion of organic anions and has therefore originally called canalicular multispecific organic anion transporter (cMOAT) [91, 92]. ABCC3 (MRP3) is also an organic ion transporter but prefers glucuronate conjugates over GSH conjugates [93]. ABCC4, ABCC5, ABCC11, and ABCC12 are MRP-like proteins which lack the additional N-terminal domain, and ABCC4 and ABCC5 have been shown to function as cellular efflux pumps for nucleosides, including antihuman immunodeficiency virus drugs such as PMEA [94] and nucleotide analogs (e.g. 6-mercaptopurine and thioguanine) [95]. Although the physiological role as well as the potential participation in drug efflux of ABCC6 (MRP6) is still unclear [96], mutations in the gene have been detected in the connective tissue disorder pseudoxanthoma elasticum (PXE, Tab. 2) [97]. Since ABCC6 is highly expressed in liver and kidney cells, sites where PXE is not very pronounced, one hypothesis suggests that ABCC6 may transport or remove toxic metabolites which destroy connective tissue cells [98]. ABCC11 and ABCC12 are most closely related to the ABCC5 gene and are found tandemly duplicated on chromosome 16q12 (Tab. 1) [99]. Since ABCC11 and ABCC12 were mapped to a region harboring gene(s) for paroxysmal kinesi-genic choreoathetosis, a disease characterized by recurrent, brief attacks of involuntary movements induced by sudden movements or startling, ABCC11 and ABCC12 represent positional candidates for this disorder [100, 101]. Interestingly, several paroxysmal neurological manifestations and idiopathic age-dependent seizures are known to be caused by ion channel-related genes [102]. The two remaining members of the ABCC subfamily ABCC8 (SUR1) and ABCC9 (SUR2) bind sulfonylurea with high affinity and interact with potassium inward rectifiers KIR6.1 and KIR6.2, to form a large octameric channel with the stoichiometry (SUR/KIR6.x)4 [103]. These heteromeric channels regulate insulin release in response to glucose metabolism and sulfonylureas are widely used to stimulate insulin secretion in type 2 diabetic patients because they close these ATP-sensitive potassium (KATP ) channels in the pancreatic beta-cell membrane (see Fig. 5) [34].

3.4 The ABCD (ALD) Subfamily

This subfamily is composed of four peroxisomal half-size ABC transporters with a TMD-ABC domain structure. They are involved in very long fatty acid (VFLA) transport. A variable pattern of homo- and heterodimerization for all ABCD members has been proposed [104–106] and mutations in ABCD1 and ABCD3 are associated with adrenoleukodystrophy (ALD) and Zellweger syndrome 2 (ZWS2), respectively (Tab. 2) [107, 108]. An interesting finding is the transcripitonal regulation of ABCD genes by lipids (Tab. 1). In this respect, recent reports have provided evidence that nuclear hormone receptor ligands, especially RXR ligands and PPAR ligands, induce the ABCD2 promoter [109, 110].


3.5 The ABCE (OABP) and ABCF (GCN20) Subfamilies

This subfamily contains four half-size ABC transporters, which are ubiquitously expressed in human tissues and do not possess transmembrane domains. The ABCE1 gene encodes an oligoadenylate binding protein (OABP), which is only found in multicellular eukaryotes and seems to participate in innate immune defense [30]. Oligoadenylates, which are produced from virus-infected cells are activators of RNaseL, which in turn degrades cellular RNAs and thereby blocks protein synthesis in infected cells. ABCE1 binds these oligonucletides and thus inhibits RNAseL, which implies that ABCE1 is involved in the negative control of immune reactions. ABCF1, the human homolog of the yeast GCN20 gene, shares some interesting features with ABCE1. Thus, ABCF1 is involved in the control of protein synthesis and also in the control of the immune system. ABCF1 binds to the translation elongation initiation factor 2 (eIF2) and seems to modulate its phosphorylation state [111]. In addition, ABCF1 has been co-purifed with ribosomal components confirming its role in protein translation [31]. In another interesting study, Richard and colleagues identified ABCF1 as a TNFinduced transcript in synoviocytes. They suggest that this ABC protein could be part of inflammatory processes related to rheumatoid arthritis. Since functionally related genes tend to be clustered on chromosomes and because ABCF1 is located on chromosome 6p21.33 (Tab. 1) in close proximity to class I MHC, the proposition that ABCF1 mediates inflammatory processes is very likely. 3.6 The ABCG (White) Subfamily

The human white or ABCG subfamily consists of five fully cloned genes (ABCG1, ABCG2, ABCG4, ABCG5, and ABCG8) and one gene so far only found in rodents (ABCG3) [22]. The ABCGs are thought to dimerize to form active membrane transporters. Among the half-size molecules ABCG proteins have a peculiar domain organization characterized by a nucleotide binding domain (ATP binding cassette) at the N-terminus followed by six transmembrane-spanning domains (Tab. 1 and Fig. 1). The founding member of this group, ABCG1, was independently described by Chen et al. and Croop et al. as the human homolog of the Dro-sophila white gene [112, 113] and its genomic organization, including the promoter region, has been described recently [114, 115]. Earlier indications linked ABCG1 with the congenital recessive deafness (DFNB10) syndrome, based on its chromosomal localization on chromosome 21q22.3 [116]. However, a recent report [117] has excluded ABCG1 along with five other known genes as candidates for DFNB10. Also, conflicting data exist whether the G2457A polymorphism in the 3UTR of the ABCG1 mRNA is associated with mood and panic disorders and related to suicidal behavior [19, 118]. The most interesting report dealing with ABCG1function came from a study by Klucken et al., which identified ABCG1 as a sterol-induced gene that participates in cholesterol and phospholipid efflux, especially in macrophages and foam cells

13


[41]. The second well-known member of the ABCG subfamily ABCG2 has been identified by different approaches and is known under the names ABCP [119], BCRP [120], and MXR [121]. The protein has been shown to be amplified and overexpressed in human cancer cells and is capable of mediating drug resistance even in the absence of the classical MDR proteins ABCB1 (MDR1) and ABCC1 (MRP1) [121–123]. In contrast to most other half-size ABC transporters, the bulk of the ABCG2 protein has been localized to the plasma membrane, with a minor fractions found within intracellular membranes [29]. It was only a short time ago when two other ABCG transporters ABCG5 and ABCG8 had been identified and linked to the human disease β-sitosterolemia by two independent approaches [124– 126]. The latest paper on an ABCG member has reported the cloning of the complete cDNA of ABCG4 and identifed this transporter as a sterol-sensitive gene [127].

4 Diseases and Phenotypes Caused by ABCM Transporters

Eighteen out of 48 currently known human ABC proteins have been linked to human monogenetic disorders or cause special disease phenotypes (Tab. 2) [98]. Since ABC transporters represent a combination of enzymes and structural proteins, homozygous mutations cause severe human diseases, which are inherited in a recessive manner. As described below, these genetic diseases are found in five of the seven ABC subfamilies (Tab. 2). In addition, heterozygous mutations in ABC genes have been connected with susceptibility to complex, multigenic disorders. It is also worth mentioning that due to the pleiotropic functions of ABC transporters, the disease states affected by mutations in ABC transporter genes are just as complex and diverse as the cellular functions of these proteins. 4.1 Familial HDL-deficiency and ABCA1

The major clue that ABCA1 is involved in cellular cholesterol removal and lipid efflux was the identification of mutations in the human gene as the defect in familial HDL-deficiency syndromes such as classical Tangier disease (Tab. 2) [6–8]. The most striking feature of these patients is the almost complete absence of plasma HDL, low serum cholesterol levels, and a markedly reduced efflux of both cholesterol and phospholipids from cells, strongly supporting the idea that both lipids are co-transported [128, 129]. The lack of ABCA1 function in these patients has a major impact on plasma HDL levels and composition. Thus plasma HDL from TD patients is composed of small pre-β 1 -migrating HDL particles containing solely apoAI and phospholipids but lacking free cholesterol and apoAII [130, 131]. The low HDL levels seen in Tangier disease (TD) are mainly due to an enhanced catabolism of these HDL precursors [131–134]. In addition, the size of the HDL particle strongly correlates with the amount of cholesterol efflux and plasma HDL concentrations [135, 136]. In TD patients, neither cholesterol absorption nor metabolism is


significantly affected, however, the concentration of LDL-cholesterol is only 40% of healthy controls and the particles are often enriched in triglycerides. The reduction in LDL levels is mainly caused by disturbance of the cholesterol ester transfer pathway resulting in changes of LDL composition and size [137]. Interestingly, obligate heterozygotes for TD mutations have approximately 50% of plasma HDL, but normal LDL levels [138]. Studying 13 different mutations in 77 heterozygous individuals, Clee et al. described a more than 3-fold risk of developing coronary artery disease in affected family members and earlier onset compared with unaffected members [139, 140]. However, these results seem to be biased towards the atherosclerotic phenotype, since the prevalence of splenomegaly is much higher in the European group of ABCA1 deficiency patients [46]. These authors also reported an age-dependent modification of the ABCA1 heterozygous phenotype [140]. In addition to the absence of plasma HDL, patients with genetic HDL-deficiency syndromes display accumulation of cholesteryl esters either in the cells of the reticulo-endothelial system (RES), leading to splenomegaly and enlargement of tonsils or lymph nodes, or in the vascular wall, leading to premature atherosclerosis [46]. This indicates differences in macrophage trafficking into tissues in the absence of ABCA1 which may be a reflection of the specific localization of mutations within the ABCA1 gene. In this context, it is of note that the pool size of CD14dim CD16+ monocytes is inversely correlated with plasma HDL-cholesterol levels [141] and the expression of ABCA1 is high in phagocytes [40] but low in antigen-presenting dendritic precursor cells (unpublished observation). These observations may provide clues for a potential interlink between ABCA1function and the control of monocyte differentiation and phagocyte/dendritic cell lineage commitment. Accordingly, we have previously hypothesized that ABCA1 function regulates the differentiation, lineage commitment (phagocytic versus dendritic cells), and targeting of monocytes into the vascular wall of the RES [142]. This concept has been substantiated by recent work from our laboratory demonstrating accumulation of macrophages in liver and spleen in LDL receptor-deficient mouse chimeras that selectively lack ABCA1 in their blood cells [143]. The fact that the absence of ABCA1from leukocytes is sufficient to induce aberrant monocyte recruitment into specific tissues identifies ABCA1 as a critical leukocyte factor in the control of monocyte targeting. In addition to phagocytes, dendritic cells have been shown to be increased in atherosclerotic lesions and have been implicated in T cell activation in atherogenesis [144]. Expression of ABCA1 appears to inhibit monocyte differentiation into macrophages and may thus shift the balance between phagocytic differentiation and dendritic cell differentiation towards the latter [145]. Taking into account that dendritic cells are capable of inducing primary immune responses, ABCA1 may function, through this mechanism, as a modulator of innate immunity in atherogenesis. An interesting clue as to how ABCA1 may be implicated in the control of monocyte/macrophage trafficking at the cellular level comes from the observation that apoAI-mediated lipid efflux in ABCA1-deficient cells is paralleled by the downregulation of the protein Cdc42 and filopodia formation [146]. Cdc42, like

15


rho and rac, is a member of the family of small GTP binding proteins which are sequentially activated by extracellular stimuli in mammalian cells [147]. Cdc42 controls a wide range of cellular functions including cytoskeletal modulation, formation of filopodia and vesicular processing. Rho proteins are known to induce the formation of stress fibers and focal adhesions; rac proteins regulate formation of lamellipodia and membrane ruffles. It is thus tempting to speculate that ABCA1 modulates cellular mobility of monocytes/macrophages through this mechanism and thus may affect recruitment of monocytes into the vessel wall. This regulator function for filopodia formation and cytoskeletal reorganization may even extend to platelet aggregation, vascular smooth muscle cell migration, and endothelial cell integrity, since these cells have been shown to express ABCA1 [148]. 4.2 Retinal Degeneration and ABCA4 (ABCR)

In addition to ABCA1, the ABCA4 (ABCR) gene located on chromosome 1p21 (Tabs 1 and 2) is another example how several mutations in one ABC transporter gene can cause pleiotropic effects. Thus, many different clinical phenotypes, associated with various forms of eye degeneration, and the age of onset as well as disease severity are associated with distinct mutations in ABCA4 [9]. As summarized in Tab. 2, ABCA4 has been found to be a causal gene for a series of retinal diseases. As an effort of several laboratories in 1997 [149–151], mutations in ABCA4 have been identified in Stargadt disease (STGD), a juvenile-onset macular dystrophy characterized by rapid central visual impairment and progressive bilateral atrophy of the retinal pigment epithelium, as well as in the late-onset form termed fundus fla-vimaculatus. Although only 60% of the mutations in the ABCA4 gene of STGD have been determined, all segregated chromosomal regions in these patients have been mapped to a locus between chromosomes 1p13 and 1p22. In addition to the monogenic STGD, ABCA4 mutations have been described in the autosomal recessive diseases conerod dystrophy (CRD) [152, 153] and retinitis pigmentosa (RP) [152, 154–156], which are both genetically and clinically heterogeneous disorders. Conerod dystrophy mainly displays cone degeneration, whereas retinitis pigmentosa affects predominantly rod photoreceptors. Age-related macular degeneration (AMD), the leading cause of severe central visual impairment among the elderly, is the fourth disease state associated with ABCA4 dysfunction. The disease is also characterized by progressive accumulation of large quantities of lipofuscin with retinal pigment epithelial cells and delayed dark adaptation [157]. Athough AMD is strongly influenced by environmental factors such as smoking, heterozygous mutations in ABCA4 have been proposed to increase the susceptibility to develop AMD. Thus, the two most frequent AMD-associated ABCA4 variants D2177N and G1961E, increase the risk of developing AMD by approximately 3-fold and 5-fold, respectively [158, 159]. In addition to the above described results based on the phenotypical analysis of ABCA4 mutations, data from in vitro studies and ABCA4 knockout mice have shed


Fig. 3 Model for the role of ABCA4 (ABCR) in rod outer segments. Left panel: schematic drawing of a rod photoreceptor. Right panel: magnification of rod disc membranes. Rho-dopsin is manufactured from opsin and 11-cis retinal in the Golgi of the rod inner segment and transported to rod outer segment discs. Upon light absorption the 11-cis form of retinal is converted to an all-trans form, which reacts with phosphatidylethanolamine (PE) to form the Schiff-base product N-retinylidene-PE (N-

RPE). ABCA4 is thought to flip N-RPE to the outer leaflet of the disc membrane. There, all-trans retinal is generated by hydrolsysis of N-RPE and subsequently reduced to alltrans retinol by retinol dehydrogenase prior to its delivery to the retinal pigment epithelial cells and re-esterification [62]. Under the effect of short-wave light or in ABCA4 deficiency, all-trans retinal accumulates, causing photooxidative damage and generation of toxic A2E (N-retinyl-N-retinylidene ethanolamine) [63].

light on the transport function of ABCA4 in photoreceptor cells. Recombinant liposome-reconstituted ABCA4 displays all-trans-retinal-stimulated ATPase activity [47, 48, 160] and ABCA4 knockout mice exhibit an acute light-dependent accumulation of all-trans retinal within rod outer segments and a progressive light-dependent culmination of lipofuscin-derived A2E (N-retinyl-N-retinylidene ethanolamine) [161, 162]. Based on these data, a model for the function of ABCA4 in rod disc membranes has been proposed [162]. As summarized in Fig. 3, all-trans retinoids, which are released from rhodopsin by photobleaching, react with the primary amine of PE to form the condensation product N-retinylidene-PE (N-RPE). ABCA4 is thought to flip N-RPE to the outer leaflet of the disc membrane, where all-trans retinal is generated by hydrolysis of N-RPE and subsequently reduced to

17


all-trans retinol by retinol dehydrogenase prior to its delivery to the retinal pigment epithelial cells and re-esterification [163]. 4.3 Cystic Fibrosis (ABCC7/CFTR)

Cystic fibrosis, caused by mutations in ABCC7 (CFTR) (Tab. 2) is one of the most frequent inherited diseases in Caucasian populations with a prevalence of 1:900 to 1:2500, whereas African and Asian individuals are affected to a much lesser extent. Interestingly, a three base pair deletion (AF508) accounts for 70–80% of the mutated alleles in northern European populations. The total number now comprises more than 1000 CFTR mutations (http://www.genet.sickkids.on.ca/cftr/). The spectrum of the disease severity is dependent on the residual function of ABCC7 [164, 165]. Patients with two affected alleles develop a severe disease with a disturbed exocrine function of the pancreas leading to nutritional deficiencies, bacterial lung infections, and a blockade of the vas deferens causing male infertility. In contrast, patients with one partially functional allele retain residuary pancreatic function and have a milder disease phenotype [166]. Before ABCC7/CFTR was identified, it was known in the 1980s that the apical membrane of different epithelia displays a Cl− conductance, which could be activated by cAMP and which was defective in cystic fibrosis. With the identification of the ABCC7/CFTR gene in 1989 [167] and an impressing multitude of publications in the following years, it became more and more evident that ABC transporters are not exclusively ATP-driven pumps, but moreover can exhibit a regulatory and/or channel function. As depicted in Fig. 4, ABCC7/CFTR can act as a cAMP-regulated chloride channel as well as a regulator of outwardly rectifying chloride channels (ORCC) [88, 168]. In both cases targeting to the apical membrane of epithelial cells and activation of the regulatory (R) domain by PKA are a prerequisite for proper ABCC7function. It is now widely recognized that ABCC7 interacts with various proteins in the plasma membrane [169]. Thus, the N-terminus of CFTR binds the coiled-coil protein syntaxin 1a and the C-terminal region of CFTR binds to PDZ domain proteins, a family of proteins containing a 80–90 amino acid motif that binds the C-terminus of a variety of ion channels and receptors [170]. At least three PDZ domain-containing proteins, NHE-RF or EBP50, CAP70, and CAL, bind to the CFTR C-terminus (Fig. 4) [170–174]. Because EBP50 associates with Ezrin, which itself binds the regulatory subunit of PKA and has a binding site for F-actin [175], it is likely that EBP50 anchors CFTR to the actin cytoskeleton at a site where it can be targeted by PKA. CAP70, a subapical protein, is able to bind two ABCC7 molecules simultaneously via PDZ3 and PDZ4 [173] and thus can mediate cross-linking of CFTR dimers and thereby enhance either direct or indirect (ORCC) chloride channel activity. Taken together, the N-terminus of ABCC7 is required for binding of syntaxin 1a and other components of the SNARE-dependent vesicular trafficking machinery, whereas the C-terminus of CFTR is necessary for cytoskeletal fixation via EBP50/Ezrin proteins.


Fig. 4 Schematic diagram summarizing ABCC7 (CFTR) interactions in the plasma membrane. CFTR interacts with various proteins in the plasma membrane including syn-taxin 1a, CAL (CFTR-associated ligand), and EBP50/NHERF [170–175]. Syntaxin 1a contains coiled-coil protein motifs which bind the N-terminal part of CFTR. Syntaxin 1a controls the vesicular trafficking of CFTR through the Golgi to the plasma membrane and thereby inhibits its channel function. CFTR also contains a PSD95/Disc-large/ZO-1 (PDZ)-interacting motif at its C-terminus, which binds the PDZproteins CAL and EBP50/NHERF (ezrin, radixin, and moesin) binding phosphoprotein of 50 kDa/Na+ /H+ exchanger reg-

ulatory factor). CAL has one PDZ domain and two coiled-coil motifs, which organize CFTR into a cluster in the apical membrane. EBP50/NHERF binds to CFTR through its PDZ1 domain and thereby links the protein to the cytoskeleton via ezrin. Ezrin serves as a PKA-anchoring protein and facilitates cAMP-dependent phosphorylation of the CFTR regulatory domain and channel activity. In addition to these direct proteinprotein interactions, CFTR indirectly regulates several ion channels such as ROMK2, ENaC, CaCC, and ORCC. CaCC and ORCC are activated by Ca2+ -dependent purinergic receptors (P2Y2), which are in turn modulated by CFTR-dependent ATP release.

In addition, multimerization of CFTR molecules could be potentially achieved by CAP70-dependent linkage. 4.4 Multidrug Resistance (ABCB1/MDR1, ABCC1/MRP1, ABCG2)

When cells are exposed to toxic compounds, as is the case for tumor cells treated with chemotherapy, resistance against these drugs can occur by a variety of mechanisms. Among these are decreased cellular uptake, increased intracellular detoxification, modification of target proteins, and enhanced extrusion from cells. Although in most cases one compound initially causes these events, cells can become resistant to a variety of drugs with structural similarities to the initial compound. This multidrug resistance (MDR) is mainly caused by three drug efflux pumps, ABCB1 (MDR1), ABCC1 (MRP1), and ABCG2 (MXR, ABCP, BCRP) (Tabs 1 and 2).

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ABCB1, which was described in the mid 1970s by its ability to confer a multidrug resistant phenotype to cancer cells upon chemotherapy [176], is a highly promiscous transporter of hydrophobic drugs, e.g. vinblastine, colchicine, VP16, adriamycin, and of such diverse substrates as lipids, steroids, xenobiotics, and peptides [177]. In small-cell lung carcinoma cells, which did not display high ABCB1 activity, overexpression of ABCC1 (MRP1) was identified [178]. Thereafter a similar substrate pattern for ABCC1 compared to ABCB1 was reported, including doxorubicin, daunorubicin, vincristine, and colchicine. However, in contrast to ABCB1, which has mainly organic cations as substrates, ABCC1 is able to transport organic and anionic compounds, the latter mainly in conjugated forms [20, 95]. Another ABC protein amplified and involved in MDR is ABCG2, which confers resistance to anthracyclin chemotherapeutic drugs such as mitoxantrone. These findings have been supported by in vitro studies: ABCG2-transfected drug-sensitive breast cancer cells are resistant to mitoxantrone, daunorubicin, and doxorubicin, and also display an enhanced rhodamine-123 efflux [120]. Interestingly, an R482G mutation in ABCG2 can significantly alter its substrate specificity and concomitantly change the drug-resistance phenotype [179]. 4.5 Adrenoleukodystrophy (ABCD1/ALD)

The X-linked adrenoleukodystrophy (ALD) is an inherited peroxisomal disorder caused by mutations in ABCD1 (Tab. 2), resulting in progressive neurological dysfunction, occasionally associated with adrenal insufficiency [180]. ALD is characterized by the accumulation of unbranched saturated fatty acids with a chain length of 24–30 carbons, particularly hexacosanoate (C26), in the cholesterol esters of brain white matter and in adrenal cortex and in certain sphingolipids of brain [15, 107]. It took a long period of research for the causative defect to be identified and therapies for ALD to be developed; these include the famous approach developed by the Odone family of dietary treatment with oleic and erucic acids (glyceryl trierucate and trioleate oil), known as Lorenzo’s oil [181, 182]. Adrenoleukodystrophy belongs to a group of defects in peroxisomal β-oxidation [183]. The first step in the oxidation of very-long-chain fatty acids (VLCFA) involves their activation by conversion into CoA esters and the transport into peroxisomes [184]. The ABCD1 protein is thought to mediate this transport process [185] and evidence for this function comes from experiments using overexpression of human cDNAs encoding the ABCD1 protein and its closest relative, the ABCD2 (ALDR) protein. With this approach, Netik et al. could restore the impaired peroxisomal β-oxidation in fibroblasts of ALD patients [186]. The accumulation of very-longchain fatty acids could also be prevented by overexpression of the ABCD2 protein in immortalized ALD cells. Moreover, the peroxisomal β-oxidation defect in the liver of ABCD1-deficient mice could be restored by stimulation of ABCD2 and ABCD4 gene expression through dietary treatment with the PPAR agonist fenofibrate [186]. These results implicate that a therapy of adrenoleukodystrophy might be possible by drug-induced overexpression or ectopic expression of ABCD genes.


4.6 Sulfonylurea Receptor (ABCC8/SUR)

Familial persistent hyperinsulinemic hypoglycemia of infancy (PHHI) is characterized by unregulated insulin secretion from pancreatic beta cells. The defect has been localized to chromosome 11p15.1–p14, a chromosomal region containing the ABCC8 (SUR1) gene and the KCNJ11 (Kir6.2) gene [187]. Subsequently, causal mutations in the ABCC8 gene have been described in PHHI families [188] (Tab. 2); however, no defects in the KCNJ11 gene have found so far. As described earlier in this article (see Section 3.3), ABCC8 and KCNJ11 form together an inwardly rectifying potassium channel (Fig. 5). Under hyperglycemic conditions the

Fig. 5 Schematic model for KATP channelcontrolled insulin secretion from pancreatic $-cells. Entry and metabolism of glucose into pancreatic $-cells leads to increased levels of intracellular ATP and concomitantly decreases ADP levels. The increase in the ATP/ADP ratio causes binding of ATP to the nucleotide binding domains of ABCC8 (SUR1) and to KIR6.2 [34]. Thereby, the KATP channel closes and the plasma membrane is depolarized. The opening of voltage-gated Ca + channels and voltagedependent Na+ channels raises the intracellular Ca2+ concentration by Ca2+ influx and mobilization of intracellular Ca2+ stores, respectively. The increased level of intracellular Ca + stimulates the dephosphorylation

of $2 -syntrophin and the dissociation of 2syntrophin-utrophin-actin complexes from ICA 512 and secretory granules. Following dissociation of $2 -syntrophin, ICA 512 is cleaved by Ca2+ /calmodulin (CaM)-activated calpain, resulting in the mobilization of secretory granules from the cytomatrix and exocytosis of insulin. The pancreatic KATP channels are also regulated by important therapeutic pharmacological agents, such as sulfonylureas and K+ channel openers. Sulfonylureas, widely used in the treatment of NIDDM, stimulate insulin secretion by closing the KATP channels, while K+ channel openers inhibit insulin secretion by opening the KATP channels [103].

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intracellular ATP/ADP ratio increases and thereby causes the ATP-sensitive octameric K+ -channel complex to close, which in turn depolarizes the beta-cell membrane. The subsequent opening of voltage-dependent calcium channels allows calcium influx and initiates insulin release via several successive steps including dephosphorylation of β 2 -syntrophin, dissociation of β 2 -syntrophin-utrophin-actin complexes from the islet cell autoantigen (ICA) 512, ICA 512 cleavage by µ-calpain, and exocytosis of secretory granules. Interestingly, polymorphisms in calpain-10, a related protease also proposed to be involved in the processing of ICA512, affect insulin secretion and have been linked to type 2 diabetes [189, 190]. Hypoglycemia decreases the intracellular ATP/ADP ratio and causes opening of the ABCC8/KCNJ11 complex, which hyperpolarizes the plasma membrane and inhibits Ca + influx and thereby stops insulin secretion [191]. Mutations in ABCC8 lead to defective ATP-sensing of KCNJ11 channels and thereby to abnormal behavior in response to hypoglycemia, resulting in persistent insulin secretion despite low glucose levels. The knowledge of the function of ABCC8 has become very important in clinical practice, since sulfonylureas, drugs widely used as oral hypoglycemics to promote insulin secretion in the treatment of non-insulin-dependent diabetes mellitus (NIDDM), are high-affinity inhibitors of ABCC8 (SUR1) and ABCC9 (SUR2). In addition to the causal mutations in PHHI, polymorphisms in the ABCC8 gene have been associated with hyperinsulinemia and type 2 diabetes in Mexican Americans [192] and French Caucasians [193], respectively.

5 Function and Regulation of ABC Transporters in Lipid Transport

Due to the strong interest in the primary drug-transporting ABC proteins, other aspects of cellular functions, such as lipid homeostasis, of this large transporter superfamily has been for long time remained unknown and unappreciated. The first implication that ABC proteins could participate in lipid binding and/or transport came from mdr2 knockout mice, which displayed a complete absence of phospholipids from bile and as a consequence developed liver disease [194]. Subsequently, the translocation of phospholipids by the human homolog MDR3 (ABCB4) was demonstrated [61, 62, 195, 196]. Only a few years later, the identification of the sterol-responsiveness of ABCA1 [40] and of other ABC family members [41] had paved the way for the identification of the gene defect in HDL deficiency [6–8], which was a major clue in proving the importance of ABC transporters in macrophage cholesterol efflux. In a similar manner, the discovery of the genetic defect in βsitosterolemia has identified ABCG5 and ABCG8 as proteins which extrude dietary sterols from intestinal epithelial cells and from the liver to the bile duct [124, 126]. As is clear from Tab. 1, a significant number of ABC transporters feature a lipidsensitive regulation, which implies that in addition to the currently established ABC proteins, further members of this superfamily could have similar functional


properties. The following section will summarize the current knowledge of ABC transporters in lipid transport with a special emphasis on transport processes in macrophages, liver, and intestine, which reflect the major organ systems in sterol metabolism. 5.1 ABCA1 in Macrophage Lipid Transport

Several factors control the expression and activity of ABCA1. Induced cholesterol influx into macrophage cells has been shown to be a potent inducer of ABCA1 expression [40]. Since the cloning of the complete human and mouse ABCA1 genes, a number of transcriptional control elements acting via alternative promoters have been characterized [197–199] (Fig. 6). The ABCA1 upstream region contains a macrophage-specific promoter preceding exon 1. This sequence binds the repressors ZNF202 and USF1/2, as well as the activating factors Sp1/Sp3 and

Fig. 6 Diagram representing the human ABCA1 and ABCG1 gene promoters Upper panel: The ABCA1 upstream region contains two alternative promoters. Promoter 1 mainly directs macrophage-specific ABCA1 expression and contains binding sites for the transcription factors ZNF202, Sp1, Sp3, E-box binding factors, LXR/RXR, and TATA binding proteins. Promoter 2 is active in liver and steroidogenic tissues and contains putative binding motifs for HNF3, SREBP, LRH/SF1, LXR/RXR, C/EBPs, and a TATA box [198–202]. Lower panel: The ABCG1

gene contains at least three alternative promoters. Binding of ZNF202 to promoter 2 and binding of LXR/RXR to promoter 3 has been determined experimentally. The functionality of promoter 1 has not been demonstrated so far. ZNF202, zinc finger transcription factor 202; Sp1, specificity protein 1; Sp3, specificity protein 3; LXR, liver X receptor; HNF3, hepatic nuclear factor 3; SREBP, sterol regulatory element binding protein; LRH, liver x receptor homolog; SF1, steroidogenic factor 1; N/cFB, nuclear factor kappa B [203, 215].

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the oxysterol-induced RXR/LXR heterodimer [200, 201]. A second promoter located downstream of exon 1 has been recently implicated in the liver/steroidogenic expression of ABCA1 [198] (Fig. 6). The The LXR/RXR-responsive elements in promoter 1 triggers retinoic acid and oxysterol-dependent activation of the ABCA1 promoter and thereby confer the observed induction of ABCA1 during lipid loading of macrophages. The most likely endogenous ligand for LXRa and LXRβ is 27-hydroxycholesterol, since CYP27-deficient cells are not able to upregulate ABCA1 in reponse to sterols and since overexpression of CYP27 activates LXR/RXR [202]. The earlier described LXR ligands 20(S)-hydroxycholesterol, 22(R)-hydroxycholesterol, and 24(S),25-epoxycholesterol are not present in cholesterol-loaded macrophages, rendering them unlikely to be natural ligands of LXR [202]. In contrast to LXR/RXR, the zinc finger transcription factor ZNF202 is a transcriptional repressor of ABCA1 gene expression, which also prevents the induction of the gene by oxysterols by recruiting the universal co-repressor KAP1 (KRAB domain-associated protein 1) [203]. Due to the strong upregulation of ABCA1 expression in response to oxysterols, LXR agonists have been proposed to be promising candidates for therapeutic activation of ABCA1 [199, 204–206] (Fig. 7). It stands to reason that especially under disease conditions such as NIDDM, where the cells have low glucose levels, low ATP levels, and associated low HDL-cholesterol levels, excessive mitochondrial energy production could induce mitochondrial exhaustion. This may ultimately result in cellular ATP shortage, a process that likely enhances the programmed cell death of lesion macrophages. Mitochondrial exhaustion may also inhibit mitochondrial 27-OH sterol synthesis and its export from the mitochondrion, a critical pathway for LXR activation in response to cellular cholesterol stress (Fig. 7) [202]. Since 27-OH sterol is the predominant oxysterol in macro-phage-derived foam cells and atherosclerostic lesions [207], this mechanism may indeed be of pathophysiological significance in atherogenesis. In light of these complexities, treatment with LXR agonists bears the potential risk of inducing mitochondrial failure and pro-apoptotic effects and may thus negatively affect lesion formation. ABCA1 appears to be localized on the plasma membrane and surface expression of ABCA1 is upregulated in macrophages by cholesterol loading [208]. Recent evidence indicates that ABCA1 and Cdc42 are associated with a Lubrol detergentresistant raft subfraction, whereas ABCA1 is not detectable in Triton-resistant rafts [209, 210]. In addition, the fact that ABCA1 is detectable in the cytosol and Golgi compartment of unstimulated fibroblasts also raises the intriguing possibility that it is a mobile molecule that may shuttle between the plasma membrane and the Golgi compartment. Thus, ABCA1 could be a constituent of a vesicular transport route for lipids involving the Cdc42/N-WASP/Arp pathway (Fig. 7). Initial studies on the biologic role of ABCA1 supported the view that this transporter, like MDR1 and MDR3 [62], functions as a translocator of lipids between the inner and outer plasma membrane [211]. This was based on experiments showing an increase in cholesterol and phospholipid export under conditions of forced expression of ABCA1 and ABCA1-null mutant cells from TD individuals that characteristically display the reverse scenario [128, 208]. However, recent work from our laboratories indicated that the ATP turnover of ABCA1 occurs


Fig. 7 Synopsis of ABC lipid transporters, cellular lipid trafficking pathways, and energy-dependent activation of ABCA1. The model view presented highlights the interdependence of ABCA1function and the availability of ATP, thus emphasizing the requirement of mitochondrial integrity for the proper function of ABC transporters. The transcriptional activation of ABCA1 induced by oxidized sterols such as 27-OH cholesterol is shown. ACAT, acyl-CoA:cholesterol acyltransferase; ANT, ade-nine nucleotide translocator; Apaf-1, apoptotic proteaseactivating factor 1; PDH, pyruvate dehydrogenase; PKA, protein kinase A; UC, unesterified cholesterol; VDAC, voltage-dependent anion channel; ABC, ATP binding cassette

transporter; ACS, acyl-CoA synthetase; CE, cholesteryl ester; DAG, diacylglycerol; FA, fatty acid; FABP, fatty acid binding protein; FATP, FA transfer protein; FA-CoA, fatty acid acyl-CoA; GlcCer, glucosylceramide; HE1, Niemann-Pick C2 protein; HSL, hormone-sensitive lipase; L, lysosome; LacCer, lactosylcera-mide; LB, lamellar body; LCAT, lecithin-cholesteryl acyltransferase; Lipo, lipoprotein; MVB, multi-vesicular body; NCEH, neutral cholester-yl ester hydrolase; NPC, Niemann-Pick C protein; PC, phosphatidylcholine; PE, phosphatidylethanolamine; PL, phospholipid; PS, phos-phatidylserine; SPM, sphingomyelin; TG, triglycerides [207–211].

at a very low rate, whereas nucleotide binding induces conformational changes [35]. Based on this information it is likely that ABCA1 acts rather as a facilitator of cholesterol/phospholipid export within the cellular lipid export machinery than exerts bona fide pump function [35]. It will be exciting to elucidate the exact molecular mechanisms by which ABCA1 mediates the export of lipid compounds from the cell. 5.2 ABCG1 and Other ABCG Members in Sterol Homeostasis

Following its cloning in 1996 [112], it was four years before ABCG1 attracted great attention because of its striking similarities with ABCA1 in its expression

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pattern in monocytic cells. Using a differential display approach ABCG1 was identified as a target gene involved in macrophage lipid homeostasis [41]. Like ABCA1 [40], ABCG1 is upregulated during the differentiation process of monocytes into mature macrophages and is strongly induced by foam cell conversion of these macrophages under sterol loading conditions using acLDL. Conversely, cholesterol unloading conditions achieved by further incubation with HDL3 , as the cholesterol acceptor results in the suppression of ABCG1 mRNA and protein expression [41]. In the mean time, these results have been confirmed by other groups as well [212–214]. The observed upregulation of ABCG1 is not restricted to acLDL but is also operative when using other types of modified LDL, such as oxidized LDL or enzymatically modified LDL, but not with free cholesterol or native LDL. Of special interest is the finding that ABCG1 regulation by lipids occurs exclusively in human or murine monomyeloid cells, such as primary human macrophages, THP-1 cells, RAW246.7 cells, peritoneal macrophages, and foam cells of atherosclerotic lesions. The sterolsensitive induction seen in these cells is independent of pro-inflammatory stimuli and the oxidative state of the cell as treatment with TNFα or LPS has no impact on ABCG1 mRNA expression [213]. In addition to lipoprotein-derived lipids, some oxysterols and RXR-specific ligands upregulate ABCG1 expression via the LXR/RXR pathway. Evidence for a significant role of these nuclear receptors in ABCG1 induction comes from two different types of experiments. First, macrophages devoid of LXRα and LXRβ fail to upregulate ABCG1 mRNA upon oxysterol treatment, and secondly, retroviral expression of LXR in RAW246.7 cells facilitates the induction of ABCG1 in response to LXRα and LXR ligands [213]. A first characterization of the ABCG1 promoter (promoter 2 in Fig. 6) demonstrated its functionality and elucidated the minimal region required for liver- and macrophage-specific expression of the gene [114]. Further reports have shown that the ABCG1 gene displays a highly complex transcriptional profile due to the existence of at least three independent promoters (Fig. 6). Whereas the activity of promoter 1 has not been proven so far [115], promoter 3 of ABCG1 has been shown to bind the transcription factors LXR/RXR and thereby mediate the sterol-dependent induction of the gene [215]. In addition to this activating, sterol-regulated pathway, an independent inhibitory mechanism involving the transcriptional repressor ZNF202 and promoter 2 of ABCG1 has been described [203]. ZNF202 regulates a number of genes involved in general lipid metabolism and in particular has been shown to bind to the apoE, ABCA1, and ABCG1 promoters and thereby to modulate cellular lipid efflux. Taken together, transcription from ABCA1 and ABCG1 genes seems to be dominated by sterol-dependent activating mechanisms involving LXR/RXR and by sterol-independent repressory mechanisms mediated by ZNF202. Although the remarkable regulation of ABCG1 gene expression by cellular lipid components revealed its importance in macrophage lipid metabolism, direct evidence for a functional role in lipid trafficking came from an antisense strategy to block ABCG1 expression [41]. Specific antisense oligonucleotides which had no effect on ABCA1 levels caused a 32% and a 25% reduction in macrophage


cholesterol and phospholipid efflux, respectively, thereby directly linking ABCG1 with cellular lipid trafficking (Fig. 7). Since the same ABCG1 antisense oligonucleotides lead also to a significant inhibition of apoE secretion, the pathways involving ABCG1 seem to be at least in part distinct from acceptor mediated lipid efflux. Also, the residual phospholipid and cholesterol efflux present in cells from patients with Tangier disease along with a compensatory upregulation of ABCG1 in these cells further supports a function of ABCG1 in intracellular mobilization of lipid stores [212]. First steps in the elucidation of the localization of ABCG1 showed that the protein is predominantly localized in intracellular compartments mainly associated with the ER and Golgi membranes [41, 206, 216]. The small fraction of ABCG1 surface staining detected in immunocytochemical analysis is presumably due to unspecific binding of polyclonal ABCG1 antibodies to the macrophage receptor, as a ABCG1GFP fusion protein is absent from the plasma membrane [206]. There is still a lack of knowledge regarding the question whether ABCG1 functions as a heterodimer or homodimer. Both forms are conceivable for ABCG1 since both cases have been described within the subfamily, e.g. ABCG2 acts as homodimer, whereas ABCG5 and ABCG8 most likely cooperate as heterodimers. In addition to the above described lipid efflux pathways operative in macrophages and liver cells, two other members of the ABCG subfamily, namely ABCG5 and ABCG8 (Tabs 1 and 2), have been implicated recently in the efflux of dietary sterols from intestinal epithelial cells back into the gut lumen and from the liver to the bile duct (Fig. 9). The sterols in a normal western diet usually consist of cholesterol (250–500 mg) and non-cholesterol sterols (200–400 mg), mainly plant sterols like sitosterol and also sterols from fish. In healthy individuals approximately 50–60% of the cholesterol is absorbed and retained, whereas the retention of non-cholesterol sterols is less than 1% [217, 218]. These subtle mechanisms are disrupted in βsitosterolemia or phytosterolemia or shellfishsterolemia, a rare autosomal recessive disorder first described by Bhattacharyya and Connor in 1974 [219]. The disease is characterized by enhanced trapping of cholesterol and other sterols, including plant and shellfish sterols, within the intestinal cells and the inability to concentrate these sterols in the bile. As a consequence affected individuals have strongly increased plasma levels of plant sterols e.g. β-sitosterol, campesterol, stigmasterol, avenosterol, and 5-saturated stanols, whereas total sterol levels remain normal or are just moderately elevated [220, 221]. Another biochemical feature of β-sitosterolemia is a reduced cholesterol synthesis due to a lack of HMG-CoA reductase. Despite the almost normal total plasma sterol levels, the disease shares several clinical characteristics with homozygous familial hypercholesterinemia (FH). Patients display tendon and tuberous xanthomas at an early age, premature development of atherosclerosis, and coronary artery disease. In some cases hemolytic episodes, hypersplenism, platelet abnormalities, arthralgiasis, and arthritis have been described [222]. In 1998 Patel et al. [223] managed to localize the β-sitosterolemia locus to chromosome 2p21 and a recent fine mapping allowed workers to narrow the gene within a 2 cM region between markers D2S2294 and Afm210ex9 [125]. Using a

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combination of positional cloning and genome database survey, Lee et al. [126] identified ABCG5, which was mutated in nine unrelated β-sitosterolemia patients. Almost at the same time, Berge et al. [124] used a microarray analysis to search for LXR-regulated genes and identified ABCG5. Since ABC transporters are often found in clusters, the group screened nearby regions and found a second new member of the ABCG subfamily, ABCG8, which displayed 61% sequence similarity and was also mutated in sitosterolemia patients. The fact that the translational start sites of both ABC transporter genes are separated by only 374 bp and arranged in a head-to-head orientation led to the assumption that ABCG5 and ABCG8 have a bi-directional promoter and share common regulatory elements [124], although no functional promoter data have been provided so far. The highest expression level of both transporters is found in liver and intestine and high-cholesterol diet feeding in mice induced the expression of both genes. These findings, together with the observed clinical and biochemical features of β-sitosterolemia patients, assume that ABCG5 and ABCG8 play an important role in reducing intestinal absorption and promote biliary excretion of sterols. To date, several mutations and a number of polymorphisms have been identified in ABCG5 and ABCG8 [124, 126, 224, 225]. Interestingly, sequence analysis of both genes showed that the majority of the analyzed patients were homozygous for a single mutation and that the total number of different mutations is very low. This strongly suggests that sitosterolemia has its origin in a limited number of founder individuals. Another striking finding is that mutations in β-sitosterolemia patients occur exclusively either in ABCG5 or ABCG8, but never in both genes together [225, 226]. The coordinate regulation of both genes and the finding that mutations in either gene cause β-sitosterolemia strongly suggest that the ABCG5 and ABCG8 proteins form a functional heterodimer. As depicted in Fig. 8, dietary sterols including cholesterol and plant sterols which enter the intestinal epithelial cells via micellar transport are released along the lysosomal route. β-Sitosterol and other plant sterols are directly transported back to the gut lumen by the heterodimeric ABCG5/ABCG8 complex in a sort of kick-back mechanism, which may also efflux cholesterol, thereby regulating total sterol absorption. The retained sterols are routed along the ACAT pathway in the ER and either stored as cholesteryl esters in lipid droplets or alternatively packed into chylomicrons for further transport back to the liver (Fig. 8). In the liver alternative processes are conceivable. The sterols are either transported to peripheral tissues by VLDL and LDL particles or converted to bile acids. Also, a direct track into the bile duct for excretion exists, possibly mediated by ABCG5 and ABCG8. In addition to ABCG5 and ABCG8, other ABC transporters including ABCG1 and ABCA1 may also participate in intestinal sterol absorption mechanisms. Data from ABCA1−/− mice strongly suggest that ABCA1 is involved in the absorption of cholesterol and in the uptake of lipophilic vitamins [208, 227]. With this respect, it will be of special interest to determine in which membrane compartment, the apical or the basolateral part of intestinal epithelial cells, the ABCA1 molecule is located.


Fig. 8 Proposed role of ABC proteins in intestinal sterol metabolism. ABCG5, ABCG8, and ABCA1 are sterol-induced members of the ABC transporter family. ABCG5 and ABCG8, which are mutated in sitosterolemia, form a heterodimer to mediate the export of absorbed plant sterols and cholesterol into the gut lumen. In contrast, ABCA1 expression and function are required for the uptake of sterols into intestinal ep-

ithelial cells. Implications for the intracellular location and vesicular trafficking of these proteins are presented. Abbreviations not defined in text: CE, cholesteryl ester; DAG, diacylglyceride; DGAT, acyl-CoA: diacylglycerol transferase; HSP70, heat shock protein 70; L, lysosome; MAG, monoacylglyceride; Mic, micelle; MTP, microsomal transfer protein; Sit, sitosterol [22, 124–126].

5.3 ABC Transporters involved in Hepatobiliary Transport

The formation of bile is an elementary physiological function of the liver, which involves numerous transport proteins located in the basolateral (sinusoidal) and apical (canalicular) membranes of hepatocytes (Fig. 9). Bile, which is composed of bile salts, phospholipids, cholesterol, bilirubin and many other small molecules, is necessary for the micellar absorption of lipids from the intestine as well as for the excretion of endogenous and xenobiotic compounds [228]. The first step in hepatobiliary transport, the uptake of compounds into liver cells is mediated by proteins of the solute carrier (SLC) superfamily [229]. Among these, the Na+ /taurocholate co-transporting peptide (NTCP), located in the basolateral membrane is responsible for the uptake of the majority of bile salts in hepatocytes. Small (type I) organic ions (e.g. choline, drugs, and monoamine neurotransmitters) are transported by the organic cation transporter 1 (OCT1), whereas bulky (type II) organic cations,

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Fig. 9 Overview of lipid transport proteins in hepatocytes. Monovalent bile salts, such as taurocholate, are taken up into hepatocytes by the sodium-taurocholate cotransporting poly-peptide (NTCP) [229]. The organic anion transporting polypeptides 1 and 2 (OATP1-2) are responsible for the charge-independent uptake of bulky organic compounds, including bile salts and other organic anions, uncharged cardiac glycosides, steroid hormones, and certain type 2 organic cations [230]. Small, type 1 organic cations are transported by the organic cation transporter OCT1. Several ABC proteins belonging to the ABCB (MDR) subfamily or ABCC (MRP) subfamily are expressed in liver [231]. ABCB1 (MDR1)

is responsible for the excretion of bulky amphi-phatic compounds into bile, whereas ABCB4 (MDR3) is a phosphatidylcholine translocase. Monovalent bile salts are secreted into the bile canaliculi by the bile salt export pump BSEP (ABCB11). ABCC2 (MRP2) functions as a multispecific organic anion transport protein in the canalicular membrane. ABCC1 (MRP1), expressed at very low levels in the basolateral membrane in normal hepatocytes, has a similar substrate specificity to MRP2. ABCC3 (MRP3) preferentially translocates conjugates with glucuronate or sulfate, whereas the physiological substrates for ABCC6 (MRP6) are unknown.

glutathione conjugates, and some amount of bile acids are taken up the organic anion-transporting polypeptide (OATP1) [230]. The subsequent step in hepatobiliary transport, the translocation of compounds from hepatocytes into the bile, involves ABC transporters localized in the hepatocyte apical (canalicular) membrane [231]. These ABC proteins belong to the ABCB (MDR) and ABCC (MRP) subfamilies. Despite the low expression level of ABCB1 (MDR1) in normal human liver [232], data from Mdr1a/1b knockout mice, which are very sensitive to xenobiotics, neurotoxins, and chemotherapeutics, provide evidence that the major function of ABCB1 is the protection of hepatocytes against harmful substances by active translocation into the bile [233, 234]. It is now widely accepted

6 Conclusions 31

that ABCB4 (MDR3), which is exclusively expressed in the liver apical membrane, is a bile canalicular phosphatidylcholine translocase (Fig. 9). This function has been confirmed by a series of experimental data: (1) mice with a target disruption of the Mdr2 gene, the mouse homolog of ABCB4 (MDR3), exhibit a complete absence of PC and strongly decreased levels of cholesterol from bile [194]; (2) transgenic expression of human MDR3 in these mice can fully restore PC secretion into the bile [235]; and (3) mutations in the human ABCB4 (MDR3) gene cause progressive familial intrahepatic cholestasis (PFIC) type 3 [10] (Tab. 2). The third member of the ABCB subfamily involved in hepatobiliary secretion is ABCB11 (SPGP). Gerloff et al. [236] have shown that membrane vesicles isolated from ABCB11-overexpressing Sf9 cells display ATP-dependent taurocholate uptake characteristics similar to those of liver canalicular membrane vesicles, and thus concluded that ABCB11 is the major, if not the only bile salt transporter of mammalian liver, hence the name bile salt export pump (BSEP). Further support for this proposition comes from the findings that the ABCB11 (BSEP) gene is mutated in patients with progressive intrahepatic cholestasis type 3 (PFIC3) [11], a syndrome characterized by very low levels of biliary bile salts and elevated concentrations of serum bile salts. In the ABCC (MRP) subfamily, at least four members have been shown to be expressed in liver cells [95]. In hepatocytes and other polarized epithelial cells, ABCC2 (MRP2) is localized and is highly expressed at the canalicular membrane. In contrast, ABCC1 (MRP1) present at the basolateral membrane domain, is expressed very low in normal liver. As listed in Tab. 1 and displayed in Fig. 9, physiological substrates for ABCC1 and ABCC2 comprise glutathione conjugates (e.g. leukotriene C4), estrogen- and bilirubin-glucuronides, taurolithocholate 3-sulfate, and glutathione disulfide (GSSG). However, due to the differences in the overall expression levels and because of greatly different transport kinetics, ABCC2 seems to be the major transporter of anionic conjugates. Likewise, hereditary defects of ABCC2 in humans cause the Dubin-Johnson syndrome, which is associated with defects in biliary secretion of amphiphilic an-ionic conjugates including bilirubin-glucuronides [237, 238]. Glucuronate- and sulfate-conjugates are also substrates for ABCC3 (MRP3), which has been localized to the basolateral membrane of hepatocytes [239]; however, in contrast to ABCC1 and ABCC2, glutathione conjugates are poor substrates for ABCC3. ABCC6 (MRP6), which has been localized to the lateral hepatocyte membrane [240], is capable of transporting the anionic cyclopentapeptide BQ123, an endothelin receptor antagonist; however, the physiological substrate for ABCC6 has not been elucidated so far.

6 Conclusions

Although our knowledge of lipid-transporting ATP binding cassette transporters has grown substantially over the last few years, the detailed molecular mechanisms by which lipid compounds are transported across cellular membranes still await


clarification. Analysis of the transcriptional and metabolic regulation, the intracellular localization and membrane domain association, the exact substrate specificity, and the functional activity of these proteins will provide helpful hints towards the understanding of working mechanisms of ABC lipid transporters. Based on their complex architecture it can be expected that ABC lipid transporters engage in multifaceted interactions with an array of yet to be identified effector molecules at specialized membrane compartments. The recent finding that ABCA1 is not an active pump but may rather function as a regulator similar to ABCC7 (CFTR) or ABCC8 (SUR1) supports this hypothesis. It will be a fascinating task to characterize the functional partners of ABC lipid transporters and to determine whether these include other ABC lipid transporters. In light of the now documented role of the prototypic cholesterol-responsive ABC molecules ABCA1 and ABCG1, it can be postulated that other ABC transporters which show a cholesterol-dependent regulation in macrophages, especially members of the ABCB and ABCC subfamilies play critical roles in macrophage lipid homeostasis.

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1

The Photoproteins Osamu Shimomura The Photoprotein Laboratory, Falmouth, USA

Originally published in: Photoproteins in Bioanalysis. Edited by Sylvia Daunert and Sapna K. Deo. Copyright ľ 2006 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31016-6

1 Introduction

In 1961, we found an unusual bioluminescent protein in the jellyfish Aequorea and named it “aequorin” after its genus name [1]. The protein had the ability to emit light in aqueous solutions merely by the addition of a trace of Ca2+ . Surprisingly, it luminesced even in the absence of oxygen. After some studies, we discovered that the light is emitted by an intramolecular reaction that takes place inside the protein molecule, and that the total light emitted is proportional to the amount of protein luminesced. At that time, we simply thought that aequorin was an exceptional protein accidentally made in nature. In 1966, however, we found another unusual bioluminescent protein in the parchment tubeworm Chaetopterus [2]. This protein emitted light when a peroxide and a trace of Fe2+ were added in the presence of oxygen, without the participation of any enzyme. The total light emitted was again proportional to the amount of the protein used. These two examples were clearly out of place in the classic concept of the luciferin–luciferase reaction of bioluminescence, wherein luciferin is customarily a relatively heat stable, diffusible organic substrate and luciferase is an enzyme that catalyzes the luminescent oxidation of a luciferin. Considering the possible existence of many similar bioluminescent proteins in luminous organisms, we have introduced the new term “photoprotein” as a convenient, general term to designate unusual bioluminescent proteins such as aequorin and the Chaetopterus bioluminescent protein [2]. Thus, “photoprotein” is a general term for the bioluminescent proteins that occur in the light organs of luminous organisms as the major luminescent component and are capable of emitting light in proportion to the amount of protein [3]. The proportionality of the light emission makes a clear distinction between a photoprotein and a luciferase. Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 The Photoproteins

In a luciferin–luciferase luminescence reaction, the total amount of light emitted is proportional to the amount of luciferin, not to the amount of luciferase. If a luciferin is a protein, the luciferin is a photoprotein, regardless of the existence or nonexistence of a specific luciferase. A photoprotein could be an extraordinarily stable form of enzyme–substrate complex, more stable than its dissociated forms, an enzyme and a substrate. Because of its high stability, a photoprotein, rather than its dissociated forms, occurs as the primary light-emitting component in the light organs. For example, the light organs of the jellyfish Aequorea contain aequorin, which is highly stable in the absence of Ca2+ , but its components coelenterazine and apoaequorin, both unstable, are hardly detectable in any part of the jellyfish. In the cells of luminous bacteria, the bacterial luciferase forms an intermediate by reacting with FMNH2 and O2 , and this intermediate emits light when a fatty aldehyde is added [4, 5]. However, this intermediate is unstable and short-lived; thus, it does not fit the definition of photoprotein. Presently, there are about 30 different types of bioluminescent systems for which substantial biochemical knowledge is available. About half of these types involve a photoprotein (Table 1). These photoproteins include the Ca2+ -sensitive type from various coelenterates (aequorin, obelin, etc.); the superoxide-activation types from a scale worm (polynoidin) and the clam Pholas (pholasin); the H2 O2 -activation type from a brittle star (Ophiopsila); and the ATP-activation type from a Sequoia millipede (Luminodesmus). For analytical applications, the photoproteins of the Ca2+ -sensitive type and the superoxide-sensitive type (pholasin) have been utilized, and the photoprotein aequorin has been in extensive use in various biological studies for the past 35 years. Each of the various photoproteins are briefly described in the next section, followed by a discussion on the extraction and purification of photoproteins and a more detailed account on the photoprotein aequorin.

2 Various Types of Photoproteins Presently Known 2.1 Radiolarian (Protozoa) Photoproteins

A Ca2+ -sensitive photoprotein that resembles coelenterate photoproteins was isolated from the radiolarian Thalassicola sp., but its properties were not investigated in detail [6]. It is of interest as the only known example of a Ca2+ -sensitive photoprotein other than the coelenterate photoproteins. 2.2 Coelenterate Photoproteins

Several kinds of photoprotein, including aequorin and obelin, were isolated from hydrozoan jellyfishes and hydroids. All of them emit blue light when Ca2+ is added,

2 Various Types of Photoproteins Presently Known 3 Table 1 Photoproteins that have been isolated

Source

Protozoa Thalassicola sp.1 Coelenterata Aequorea aequorea2 Halistaura sp.3 Phialidium gregarium Obelia geniculata Obelia geniculata Obelia longissima Ctenophora Mnemiopsis10 sp. Beroe ovata10 Annelida Chaetopterus variopedatus11

Name

Mr

Thalassicolin 21 500

440

Ca2+ Ca2+ Ca2+ Ca2+ Ca2+

465 470 474

Mnemiopsin-1 Mnemiopsin-2 Berovin

24 000 27 500 25 000

Ca2+ Ca2+ Ca2+

485 485 485

120 000 184 000

Fe2+ , hydroperoxide, and O2 Fe2+ , H2 O2 , and O2

455

Peroxidase or Fe2+ , plus O2 Alkaline pH? and O2 Catalase, H2 O2 , and O2

490

60 000

ATP, Mg2+ , and O2

496

45 000

H 2 O2

482

500 000

Mollusca Pholas dactylus13

Pholasin

34 600

Symplectin

60 000

Campbell et al. [7] Shimomura [50] 3 Shimomura et al. [71] 4 Levine and Ward [31] 5 Inouye and Tsuji [76] 6 Stephenson and Sutherland [77] 7 Illarionov et al. [22] 8 Morin and Hastings [36] 9 Markova et al. [34] 10 Ward and Seliger [80] 11 Shimomura and Johnson [57] 12 Nicolas et al. [40] 13 Michelson [35] 14 Fujii et al. [27] 15 Tsuji et al. [79] 16 Shimomura unpublished 17 Shimomura [66, 68] 18 Shimomura [49] 2

Ca2+

23 000 21 600 21 0006 21 000 22 2007

Polynoidin

1

Luminescence maximum (nm)

Aequorin Halistaurin Phialidin 4 , clytin5 Obelin Obelin Obelin

Hamothoe lunulata12

Symplectoteuthis oualaniensis14 Symplectoteuthis luminosa16 Diplopoda Luminodesmus sequoia17 Echinodermata Ophiopsila californica18

Requirements for luminescence

50 000

4758 4859 4959

510

47015

4 The Photoproteins

regardless of the presence or absence of oxygen. The coelenterate photoproteins are suitable for use in the detection and measurement of trace amounts of Ca2+ , and aequorin has been widely used in the studies of Ca2+ in various biological systems, including single cells [2, 4]. The overwhelming popularity of this type of photoprotein compared with the other types sometimes leads to the misconception that the photoproteins are Ca2+ -sensitive bioluminescent proteins. Detailed studies have been made with only three kinds of photoproteins: aequorin obtained from Aequorea, obelin obtained from Obelia, and phialidin (clytin) obtained from Phialidium. Aequorin was isolated from Aequorea aequorea by Shimomura et al. [1], and its purification was described in several papers [5, 9, 10]. The recombinant form of aequorin has been made [12–14]. Obelin was isolated from Obelia geniculata [6] and also from O. australis and O. geniculata [16]. Its recombinant form was prepared by Illarionov et al. [17]. Phialidin was isolated from Phialidium gregarium by Levine and Ward [18] and was cloned by Inouye and Tsuji [19]; the recombinant protein was named clytin. All coelenterate photoproteins have a molecular weight close to 20 000. The concentrated solutions of purified photoproteins are slightly yellowish (weak absorption at about 460 nm) and non-fluorescent except for ordinary protein fluorescence. After Ca2+ -triggered luminescence, the solutions turn colorless and become fluorescent in blue (emission λmax 460 nm). The intensity of the blue fluorescence is dependent on the concentrations of the spent protein and Ca2+ ; however, the fluorescence intensity is not proportional to the concentration of the protein [20]. In the case of aequorin, the emission spectrum of blue fluorescence is almost superimposable on the emission spectrum of Ca2+ -triggered luminescence, suggesting that the blue fluorescent chromophore formed in the luminescence reaction is probably the light emitter [21]. As a Ca2+ indicator, aequorin is useful at a concentration range of Ca2+ between 10−7.5 M and 10−4.5 M [5], whereas obelin is useful at a range between 10−6.5 M and 10−3.5 M under similar conditions [16]. The Ca2+ sensitivity of phialidin is about equal to that of obelin [22]. It should be noted here that the Ca2+ sensitivity and certain other properties of aequorin, and probably of all coelenterate photoproteins, can be modified by replacing the coelenterazine moiety of the photoprotein with its analogues (explained later). The chemistry of the bioluminescence reaction of aequorin has been elucidated in considerable detail and will be described later in this chapter. The reaction mecha nisms of all hydrozoan photoproteins are believed to be essentially identical with that of aequorin. However, the luminescence reaction differs in luminous anthozoans, which are taxonomically closely related to hydrozoan. The luminous species of anthozoans contain a luciferin (coelenterazine) and a species-specific luciferase instead of a photoprotein, although the presence of a small amount of Ca2+ -sensitive photoprotein is suspected in some species, such as the sea pen Ptilosarcus gurneyi and the sea cactus Cavernularia obesa [23]. Spent aequorin that has been luminesced with Ca2+ can be regenerated into the active original form by incubation with coelenterazine in the presence of O2 and a low concentration of 2-mercaptoethanol [24]. The regenerated aequorin is

2 Various Types of Photoproteins Presently Known 5

indistinguishable from the original aequorin in every aspect of its properties. The yield of the regeneration is practically 100% when the protein concentration is over 0.1 mg mL−1 [25]. Thus, a sample of aequorin can be luminesced and recharged repeatedly. The regeneration of spent photoprotein takes place also with obelin [6], as well as with halistaurin and phialidin (unpublished results). 2.3 Ctenophore Photoproteins

The photoproteins mnemiopsin and berovin were isolated from Mnemiopsis sp. and Beroe lovata, respectively [26]. They are Ca2+ -sensitive photo proteins that are similar to aequorin, except that these photoproteins are photosensitive. The absorption maximum of mnemiopsin-2 is 435 nm, which is about 20 nm shorter than that of aequorin. The photosensitivity of ctenophore photoproteins is strikingly different from that of aequorin. Mnemiopsin and berovin are extremely sensitive to light [13], being easily inactivated by a broad spectral range of light (wavelength 230–570 nm) [28]. Aequorin and other hydrozoan photoproteins are not affected by light. Photoinactivated mnemiopsin, as well as spent mnemiopsin after Ca2+ -triggered luminescence, can be regenerated into its active form by incubation with coelenterazine in the presence of oxygen, like aequorin; however, the regeneration takes place only at a narrow pH range around 9.0 [1]. 2.4 Pholasin (Pholas Luciferin)

The boring clam Pholas dactylus is historically important in the field of bioluminescence because it was one of the two luminous species with which Dubois first demonstrated luciferin–luciferase luminescence in 1887. Thus, the luminescence of Pholas was originally considered to be a luciferin–luciferase reaction involving Pholas luciferin and Pholas luciferase. However, Pholas luciferin is a glycoprotein with a molecular weight of 34 600 [19, 21, 32]. Therefore, it is appropriate to call this luciferin a photoprotein. The name “pholasin” was first used by Roberts et al. [33]. The ultraviolet absorption spectrum of pholasin shows a bulge at about 325 nm, in addition to the protein peak at 280 nm. Pholasin emits light (λmax 490 nm) in the presence of various substances such as Pholas luciferase, ferrous ions, H2 O2 , peroxidase, superoxide anions, hypochlorite, and certain other oxidants, all in the presence of molecular oxygen [19–22, 35]. Thus, Pholas luciferase is clearly not an essential component for the luminescence of pholasin. The luminescence reaction of pholasin with Pholas luciferase is optimum at pH 8–9 and at an ionic strength of about 0.5 M, giving a quantum yield of 0.09 for pholasin [32]. According to Reichl et al. [36], the addition of horseradish peroxidase compounds I and II to pholasin induces an intense luminescence. Moreover, the addition of H2 O2 to a mixture of

6 The Photoproteins

myeloperoxidase and pholasin gives an intense burst of light. The chromophore of pholasin is still not chemically identified. The cloning and expression of apopholasin was achieved by Dunstan et al. [9], but attempts to reconstitute the recombinant apopholasin into pholasin by the addition of an acidic methanol extract of Pholas failed, although the mixture gave luminescence by the addition of sodium hypochlorite. Pholasin is commercially available from Knight Scientific, Plymouth, UK. The main application of pholasin is the measurement of oxygen radicals. 2.5 Chaetopterus Photoprotein

The photoprotein of the parchment tubeworm Chaetopterus variopedatus purified by chromatography has a molecular mass of approx. 120-130 kDa [2, 38]. The protein is amorphous when precipitated with ammonium sulfate, but it can be converted into a crystalline form with an increased molecular mass of 184 kDa by slow crystallization with ammonium sulfate. The photoprotein emits light in the presence of Fe2+ , a peroxide, and molecular oxygen. As a peroxide, H2 O2 can be used, but an unidentified hydroperoxide existing in old dioxane or tetrahydrofuran was far more effective. Two kinds of additional activators were found to give brighter luminescence, but they were not identified. The light emission of this photoprotein is strongly affected by the pH of the medium, showing a peak at pH 7.7 with a sharp decrease at both sides (50% decreases at pH 6.5 and pH 8.3); the light intensity is not significantly influenced by the salt concentration up to 1 M when tested with NaCl. The optimum temperature for the luminescence intensity is 22 ◦ C. With this photoprotein, a concentration of Fe2+ as low as 0.1 µM can be detected. The purified photoprotein is practically colorless, and its absorption spectrum shows, in addition to the 280-nm protein absorption peak, a very slight absorption in the region of 330–380 nm, although its significance is unclear. A solution of the photoprotein is moderately blue fluorescent, with a fluorescence emission maximum at 453–455 nm and an excitation maximum at 375 nm, and these peaks do not significantly change after the luminescence reaction. The luminescence spectrum of purified photoprotein (λmax 453–455 nm) closely matched with the fluorescence emission spectrum. 2.6 Polynoidin

A membrane photoprotein isolated from the scales of the scale worm Harmothoe lunulata was named “polynoidin” [39]. The purified photoprotein (Mr 500 000) emits light in the presence of molecular oxygen (λmax 510 nm) by the action of sodium hydrosulfite, the xanthine–xanthine oxidase system, Fenton’s reagent (H2 O2 plus Fe2+ ), or other reagents that produce superoxide radicals. The photoprotein luminescence was 30% brighter in phosphate buffer than in Tris buffer, and the luminescence response was significantly increased by including a complexing agent

2 Various Types of Photoproteins Presently Known 7

such as EGTA. However, the injection of polynoidin solution into the mixture of H2 O2 and Fe2+ failed to produce light; Fe2+ must be added last to initiate light emission. The photoprotein is not fluorescent (except for usual protein fluorescence) after the bioluminescence reaction or before the reaction. The requirements for its luminescence reaction are similar to that of the bioluminescence systems of Pholas and Chaetopterus, suggesting the involvement of a common basic mechanism in these luminescence systems. 2.7 Symplectin

The luminescent substance of the squid Symplectoteuthis oualaniensis was first obtained in the form of insoluble particles by Tsuji et al. [40]. The suspension of the particles emitted light in the presence of monovalent cations such as K+ , Rb+ , Na+ , Cs+ , NH4 + , and Li+ (in decreasing order of effect). Molecular oxygen was needed for the luminescence. Divalent ions such as Ca2+ and Mg2+ did not trigger light emission. The light emission (λmax 470 nm) was optimal in the presence of 0.6 M KCl or NaCl and at a pH of 7.8. The soluble form of the Symplectoteuthis photoprotein was isolated and purified from the granular light organs of the squid and was named “symplectin” [27, 41]. The light organs were first extracted with a pH 6 buffer containing 0.4 M KCl to remove impurities, and then symplectin was extracted from the residue with a pH 6 buffer containing 0.6 M KCl. All solutions used in the experiments contained 0.25 M sucrose, 1 mM dithiothreitol, and 1 mM EDTA. The 0.6 M KCl extract was chromatographed by size-exclusion HPLC on a TSK G3000SW column. Symplectin was eluted as two major components of oligomers, having molecular masses of 200 kDa or more, and a minor component of monomer (60 kDa). All processes of extraction and purification were carried out at 4 ◦ C. Warming up a solution of symplectin, adjusted to pH 8, to room temperature causes the luminescence reaction to begin, and the light emission lasts for hours. A tryptic digestion of the KCl extract increased the content of the 60-kDa species at the expense of the two high-molecular-weight species, accompanied by the formation of 40-kDa and 16-kDa species. SDS-PAGE analysis of the two high-molecular-weight oligomers revealed that they consist mainly of the 60kDa protein. The 60-kDa protein and the 40-kDa protein were fluorescent in the SDS-PAGE analysis. The spent protein of symplectin after luminescence (aposymplectin) could be reconstituted into original symplectin by treatment with dehydrocoelenterazine [27]. 2.8 Luminodesmus Photoprotein

This is presently the only example of a photoprotein of terrestrial origin. The millipede Luminodesmus sequoia [32] emits light from the surface of its whole body

8 The Photoproteins

continuously day and night. The photoprotein extracted and purified from this organism emits light (λmax 496 nm) when ATP and Mg2+ are added in the presence of molecular oxygen [11, 45]. Thus, the luminescence system of Luminodesmus resembles that of the fireflies in that it requires ATP and Mg2+ , but it differs in that it needs only the photoprotein rather than the luciferin and luciferase required in the firefly system. The molecular weight of the photoprotein is 104 000, which is close to the molecular weight of firefly luciferase reported earlier (100 000) but larger than its newer value 62 000 [46]. Although it was suspected that the photoprotein might be a complex of a firefly-type luciferase and firefly luciferin, firefly luciferin itself was not detected in this photoprotein. Recently, the possible presence of a porphyrin chromophore in the photoprotein has been suggested, although the role of this chromophore in the light-emitting reaction is unclear [47]. Using the luminescence system of Luminodesmus, 0.01 µM ATP and 1 µM Mg2+ can be detected. 2.9 Ophiopsila Photoprotein

The brittle star Ophiopsila californica is abundant around Catalina Island, off the coast of Los Angeles [48]. An animal of average size weighs about 3–4 g, and has five arms of about 10 cm long. The purified photoprotein luminesces in the presence of H2 O2 , emitting a greenish-blue light (λmax 482 nm). Molecular oxygen is probably not needed for the luminescence reaction. The molecular weight of Ophiopsila photoprotein is estimated to be about 45 000 by gel filtration. The absorption spectrum of a solution of the photoprotein showed a small peak (λmax 423 nm, with a shoulder at about 450 nm) in addition to the 280-nm protein peak. The 423-nm peak decreased slightly through the H2 O2 -triggered luminescence reaction, accompanied by a slight red shift of the peak. The photoprotein was fluorescent in greenish-blue (emission λmax 482 nm; excitation λmax 437 nm), and the fluorescence emission spectrum exactly coincided with the luminescence spectrum of photoprotein in the presence of H2 O2 , suggesting the possibility that the fluorescent chromophore might be the light emitter. However, the fluorescence emission of the photoprotein did not show any detectable change after the H2 O2 -triggered luminescence reaction; an anticipated increase in the 482-nm fluorescence did not occur.

3 Basic Strategy of Extracting and Purifying Photoproteins

Photoproteins are usually highly reactive, unstable substances, like luciferins. Their luminescence activities are easily lost by spontaneous light emission and various other causes. In isolating active photoproteins, it is extremely important to pay special attention to prevent the loss of the luminescence activity. Compared with the isolation of luciferins, however, techniques available for isolating photoproteins are somewhat limited because of their protein nature.

4 The Photoprotein Aequorin

The basic principle is to extract a photoprotein in an aqueous solution and purify the photoprotein by various means of protein purification, all under conditions that prevent the luminescence and denaturation of the protein molecules. Thus, the luminescence system must be reversibly inhibited during the extraction and purification of a photoprotein. The method of reversible inhibition differs depending on the nature and cofactor requirement of the system to be isolated. For example, the calcium chelator EDTA or EGTA is used to inhibit the luminescence of the Ca2+ -sensitive photoproteins of coelenterates and ctenophores such as aequorin, obelin, and mnemiopsin [1, 6, 13, 26]. Before the discovery of the Ca2+ requirement, however, aequorin was extracted with a pH 4 buffer that reversibly inactivated the photoprotein [1, 49]. In the case of the luminescence systems of Chaetopterus and Pholas, the metal ion inhibitors 8-hydroxyquinoline and diethyldithiocarbamate, respectively, were used [2, 17]. The ionic strength and the pH of buffers are also important, and these conditions should be chosen to optimize the yield of active photoprotein. The use of acidic buffers, pH 5.6–5.8, was effective in suppressing spontaneous luminescence during the extraction of the photoproteins of euphausiids and Luminodesmus [25, 51]. In the case of the membrane photoprotein polynoidin and the squid photoprotein symplectin, easily soluble impurities were all washed out and the substances that cause the luminescence of the photoprotein are completely removed before the solubilization of the photoproteins; thus, inhibitors were not needed [27, 39].

4 The Photoprotein Aequorin 4.1 Extraction and Purification of Aequorin

Aequorin is the best-known photoprotein and has been used widely in various applications. The first step in the extraction of aequorin from the jellyfish Aequorea (average body weight 50 g) is to cut off the circumferential margin of umbrella that contains light organs, making about 2-mm-wide strips commonly called “rings”. This process is important because it eliminates about 99% of unnecessary body parts that do not contain aequorin. The rings can be made efficiently by using specially made cutting devices [5, 29] or, much less efficiently, with scissors. The rings (about 0.5 g each) containing light organs are kept in cold seawater. Then, about 500 rings are shaken vigorously by hand with cold, saturated ammonium sulfate solution containing 50 mM EDTA [28] or with cold seawater [28] to dislodge the particles of light organs from the rings. Then, the rings are removed by filtering through a net of Dacron or Nylon (50–100 mesh), and the light organ particles suspended in the filtrate are collected by filtration on a Büchner funnel with the aid of some Celite. The light organ particles in the filter cake are cytolyzed and the aequorin therein is extracted by shaking with cold 50 mM EDTA (pH 6.5). After filtration, crude aequorin is precipitated by saturation with ammonium sulfate.

9

10 The Photoproteins

Of the two methods of shaking the rings mentioned above, using seawater results in much cleaner crude extracts, with a little less yield, than are obtainable by shaking in saturated ammonium sulfate containing EDTA. On the other hand, saturated ammonium sulfate strongly inhibits the luminescence response of the photogenic particles to mechanical stimulation such as shaking and stirring, and it also salts out and stabilizes aequorin, thus resulting in a better yield of aequorin and less effect on the isoform composition of aequorin extracted, compared with that obtainable by shaking in seawater. With regard to the purification of aequorin, Blinks et al. [5] described a welldesigned method for purifying an aequorin extract that has been obtained by the “seawater shaking method”. The method included gel filtration on Sephadex G-50 and ion-exchange chromatography on DEAE-Sephadex A-50 and QAE-Sephadex A-50. The ion exchangers effectively separated the green fluorescent protein from aequorin. For the purification of the extract obtained by the “saturated ammonium sulfate shaking method”, gel filtration on a column of Sephadex G-75 or G-100, using buffers containing 1 M ammonium sulfate and not containing ammonium sulfate, and ion-exchange chromatography on DEAE cellulose have been used [9, 10, 28]. Aequorin in 1 M ammonium sulfate aggregates to a larger size (Mr > 50 000). Thus, crude aequorin is first chromatographed on Sephadex G-100 with a low-salt buffer not containing ammonium sulfate, then the aequorin fraction obtained is rechromatographed on the same column using a buffer containing 1 M ammonium sulfate to obtain purified aequorin. Using Sephadex G-50 is not recommended in this case, at least for the initial step, because of the presence of a large amount of the aggregated form of impurities. 4.1.1 Hydrophobic Interaction Chromatography Butyl-Sepharose 4 Fast Flow (Pharmacia) is an excellent medium for purifying aequorin, supplementing the methods described above. Aequorin in a buffer solution containing 5–10 mM EDTA and 1.8 M ammonium sulfate is adsorbed on the column, and then aequorin is eluted with a buffer containing decreasing concentrations of ammonium sulfate and 5 mM EDTA. Aequorin elutes at an ammonium sulfate concentration between 1 M and 0.5 M. Because apoaequorin elutes at ammonium sulfate concentrations lower than 0.1 M, aequorin is cleanly separated from apoaequorin. Thus, it is possible to prepare virtually pure samples of aequorin using a single column as follows. The aequorin sample is first luminesced by the addition of a sufficient amount of Ca2+ . The spent solution, after dissolving 1 M ammonium sulfate, is adsorbed on a column of butyl-Sepharose 4. The apoaequorin adsorbed on the column is eluted with decreasing concentration of ammonium sulfate starting from 1 M; apoaequorin elutes at an ammonium sulfate concentration lower than 0.1 M. The apoaequorin eluted is regenerated with coelenterazine in the presence of 5 mM EDTA and 5 mM 2-mercaptoethanol (see the Section 4.4.3). The solution of regenerated aequorin in 1.8 M ammonium sulfate is adsorbed on the same butyl-Sepharose 4 column. Aequorin adsorbed on the column is eluted with a decreasing concentration of ammonium sulfate, yielding highly purified aequorin.


4.2 Properties of Aequorin

Aequorin is a conjugated protein that has a relative molecular mass of approximately 20 000–21 000 [10], and it contains a functional chromophore corresponding to roughly 2% of the total weight. A concentrated solution of aequorin is yellowish because of its absorption peak (λmax about 460 nm), in addition to a protein absorption peak at 280 nm (A1%,1cm 30.0; [50]). Aequorin is non-fluorescent, except for a weak ultraviolet fluorescence that is due to its protein moiety. Natural aequorin is a mixture of isoforms, containing more than a dozen of them, designated aequorins A, B, C, etc. [4, 50]. The isoelectric points of these isoforms lie between 4.2 and 4.9 [3]. The solubility of aequorin in aqueous buffers is generally greater than 30 mg mL−1 [56]. Aequorin can be salted out from aqueous buffers with ammonium sulfate, although the salting out is not complete even after the complete saturation of ammonium sulfate. Usually 1–2% of aequorin remains in the solution. One milligram of aequorin emits 4.3–5.0 × 1015 photons at 25 ◦ C when Ca2+ is added, at a quantum yield of 0.16 [9, 21, 50, 56]. In the presence of an excess of Ca2+ , the luminescence reaction of aequorin has a rate constant of 100–500 s−1 for the rise and 0.6–1.25 s−1 for the decay [15, 33]. 4.2.1 Stability Aequorin is always emitting a low level of luminescence, spontaneously deteriorating by itself. Thus, the information concerning its stability is important when aequorin is used as a calcium probe. The stability of aequorin in aqueous solutions containing EDTA or EGTA varies widely by temperature, pH, concentration of salts, and impurities. To minimize the deterioration of aequorin, it is most important to keep the temperature as low as possible. The half-life of aequorin in 10 mM EDTA, pH 6.5, is about 7 days at 25 ◦ C. At room temperature, aequorin is most stable in solutions containing 2 M ammonium sulfate or when it is precipitated from saturated ammonium sulfate. Freeze-dried aequorin is also stable, but the process of drying always causes a loss of luminescence activity (see below). All forms of aequorin are satisfactorily stable for many years at -50 ◦ C or below, but all deteriorate rapidly at temperatures above 30–35 ◦ C. A solution of aequorin should be stored frozen whenever possible because repeated freeze-thaw cycles do not harm aequorin activity. 4.2.2 Freeze-drying A note on freeze-dried aequorin may be appropriate here, because most commercial preparations of aequorin are sold in a dried form. The process of freeze-drying aequorin always results in some loss of luminescence activity. Therefore, aequorin should not be dried if a fully active aequorin is required. The loss is about 5% at the minimum, typically about 10%. The loss can be slightly lessened by certain additives; the addition of 50–100 mM KCl and some sugar (50–100 mM) in the buffer seems to be beneficial. The buffer composition used for the freeze-drying of aequorin at the author’s laboratory is as follows: 100 mM KCl, 50 mM glucose, 3 mM HEPES, 3 mM Bis-Tris, and 0.05 mM EDTA, pH 7.0.

11


4.2.3 Specificity to Ca2+ Several kinds of cations other than Ca2+ elicit the light emission of aequorin. Some lanthanide ions (such as La3+ and Y3+ ) trigger the luminescence of aequorin as efficiently as Ca2+ . In addition, Sr2+ , Pb2+ , and Cd2+ cause significant levels of luminescence; Cu2+ and Co2+ give some luminescence only in slightly alkaline buffer. However, Be2+ , Ba2+ , Mn2+ , Fe2+ , Fe3+ , and Ni2+ do not elicit any light from aequorin [59]. In testing biological systems, however, aequorin is considered to be highly specific to Ca2+ , because the occurrence of a significant amount of metal ions other than Ca2+ is unlikely. In an in vitro test, all of these metal ions except Ca2+ , Sr2+ , and lanthanoids could be completely masked by including 1 mM sodium diethyldithiocarbamate in the test solution [60]. 4.3 Luminescence of Aequorin by Substances Other Than Divalent Cations

As already mentioned, all forms of aequorin emit photons spontaneously and constantly, regardless of its molecular status or environment conditions, even in the absence of Ca2+ or in the presence of a large excess of EDTA. The light emission results in a gradual deterioration of the luminescence capability of aequorin. A luminescence intensity of this type is quite low at 0 ◦ C, though it can be easily measured with a light meter. The intensity is temperature dependent and steeply increases with rising temperature, reaching a maximum intensity at around 60 ◦ C [56]. Such a temperature-dependent luminescence occurs with aequorin dissolved in aqueous solutions, as well as with freeze-dried aequorin and its suspension in a certain organic solvents, such as toluene, acetone, and diglyme (bis-2-methoxyethyl ether). The quantum yield of the spontaneous luminescence of dried aequorin, when warmed with or without an organic solvent, is generally in the range of 0.003–0.005, whereas that of aequorin in aqueous solutions is considerably less (about 0.001 at 43 ◦ C). Aequorin also emits luminescence in the presence of thiol-modification reagents such as p-benzoquinone, Br2 , I2 , N-bromosuccinimide, N-ethylmaleimide, iodoacetic acid, and p-hydroxymercuribenzoate [20]. The luminescence is probably caused by the conformational change of the protein that results from the modification of cysteine residues (by causing the decomposition of the coelenterazine peroxide moiety). The luminescence is weak but lasts for more than one hour. The quantum yields in this type of luminescence never exceed 0.02 (about 15% of Ca2+ -triggered luminescence) at 23–25 ◦ C. To prevent this type of luminescence, any reagents that might react with an SH group should be avoided. 4.4 Mechanism of Aequorin Luminescence and Regeneration of Aequorin 4.4.1 Structure of Aequorin Aequorin is a globular protein with three “EF-hand” domains to bind Ca2+ , and it accommodates a peroxidized coelenterazine in the central cavity of the protein [16].


The presence of a peroxy group bound to position 2 of the coelenterazine moiety was previously suggested [29] and confirmed by 13 C nuclear magnetic resonance spectroscopy [39]. The protein conformation of aequorin is much more compact and rigid than that of apoaequorin, consistent with the results of the fluorescence polarization studies and the papain digestion of those proteins [30]. The functional group, peroxidized coelenterazine, is shielded from outside solvent. Therefore, no reagent can react with this group without first reacting with the residues of the protein, and any reaction with the protein residues triggers the breakdown of the peroxidized coelenterazine. 4.4.2 Luminescence Reaction In the case of aequorin reacting with Ca2+ , a conformational change of protein takes place when one molecule of aequorin is bound with two Ca2+ ions [52]. The conformational change results in the cyclization of the peroxide of coelenterazine into the corresponding dioxetanone, which instantly decomposes and produces the excited state of coelenteramide and CO2 [20, 29]. When the energy level of the excited state of coelenteramide falls to ground state, light is emitted. A simplified mechanism of the luminescence reaction is illustrated in Fig. 1. The spent solution of the luminescence reaction of aequorin is a mixture of coelenteramide, apoaequorin, and Ca2+ that forms a complex called “blue fluorescent protein” (fluorescence emission maximum about 465–470 nm). The dissociation constant of the complex into coelenteramide plus apoaequorin in the presence of 0.5 mM Ca2+ is 7 × 10−6 M at pH 7.4 and 25 ◦ C by Morise et al. [20]; (based on the molecular weight of aequorin 21 000). Thus, the luminescence reaction product of aequorin is usually blue fluorescent, unless the concentration of aequorin used is too low (much less than 1 µM) to form the fluorescent complex. The blue fluorescence of the complex (λmax 465–470 nm) closely matches the bioluminescence emission of aequorin, giving a basis to the postulation that the fluorescent complex is the light emitter of aequorin bioluminescence [21], although it now seems an oversimplification considering that the conformation of apoaequorin continues to change for several minutes after the light emission. When the light emission of aequorin is measured in low-ionic-strength buffers containing no inhibitor, the log-log plot of the luminescence intensity versus Ca2+ concentration gives a sigmoid curve having a maximum slope of about 2.0 for its middle part [10, 65], indicating that the binding of two Ca2+ ions to one molecule of aequorin is required to trigger the luminescence of aequorin. 4.4.3 Regeneration Apoaequorin can be reconstituted into aequorin by incubation with coelenterazine in the presence of O2 and 2-mercaptoethanol, which the role of the latter substance is to protect the functional sulfhydryl groups of apoaequorin during the regeneration reaction [24]. For the regeneration reaction to occur, there is no need to separate coelenteramide from apoaequorin if the material contains it. Usually, the product of the luminescence reaction is incubated at 0–5 ◦ C in a pH 7.5 buffer solution containing 5 mM EDTA, 3 mM 2-mercaptoethanol, and an excess of coelenterazine

13


Fig. 1 Schematic illustration of a simplified mechanism of the luminescence and regeneration of aequorin. Aequorin (upper left) is a globular protein that contains peroxidized coelenterazine sealed in its central cavity and has three EF-hand Ca2+ -binding sites on the outside. When the protein is bound with two Ca2+ ions, an intramolecular reaction starts, resulting in the formation of coelenteramide and CO2 , accompanied by the emission of blue light

(8max 460 nm) and opening of the protein shell (upper right). The protein part, apoaequorin (bottom), can be regenerated into the original aequorin by incubation with coelenterazine and molecular oxygen in the absence of Ca2+ . In the regeneration reaction, addition of a low concentration of 2-mercaptoethanol increases the yield of regenerated aequorin by protecting the functional cysteine residues of apoprotein.

(at least 2 µg mL−1 more than the calculated amount). The regeneration is usually 50% complete within 30 min and practically 100% complete after 3 h. When the regeneration reaction of apoaequorin is carried out in the presence of an excess of free Ca2+ , rather than in 5 mM EDTA, the result is a continuous, weak light emission from the reaction mixture. This weak luminescence lasts many hours, differing from the short, bright flash of the Ca2+ -triggered luminescence of aequorin. The weak luminescence of the regeneration mixture in the presence of Ca2+ can be intensified several times by including 0.5% diethylmalonate in the reaction medium [45]. During the regeneration in the presence of Ca2+ described above, apoaequorin appears to be acting as an enzyme that catalyzes the luminescent oxidation of coelenterazine. The mechanism involved might be a simple, straightforward one: aequorin is first formed, and then it instantly reacts with Ca2+ to emit light. This simple mechanism, however, has no experimental support at present; the regeneration reaction of aequorin in the presence of EDTA was not activated by diethylmalonate, suggesting either that Ca2+ is needed in the activation by diethylmalonate or that aequorin is not an intermediate in the luminescence reaction in the


presence of Ca2+ [45]. Whatever the mechanism, apoaequorin must be a very sluggish enzyme if it is an enzyme. Apoaequorin has a turnover number of 1–2 per hour [10]. 4.5 Inhibitors of Aequorin Luminescence

All thiol-modification reagents cause weak, spontaneous luminescence of aequorin in the absence of Ca2+ , as already mentioned. They are in effect inhibitors of the Ca2+ -triggered luminescence of aequorin, because the quantum yields of aequorin in the luminescence caused by these reagents (∼0.008) are much lower than that of the Ca2+ -triggered luminescence of aequorin [20]. Bisulfite, dithionite, and p-dimethyaminobenzaldehyde are all strongly inhibitory even at micromolar concentrations [1]. It has been found that the functional group of aequorin, i.e., a peroxide of coelenterazine, decomposes without light emission when the photoprotein is treated with bisulfite or dithionite, resulting in the formation of a corresponding hydroxy-coelenterazine or coelenterazine [63]. A number of inorganic and organic substances at high concentrations (> 50 mM) suppress the luminescence intensity of the Ca2+ -triggered light emission. Thus, KCl (100–150 mM) used in physiological buffers is significantly inhibitory. Magnesium ions are inhibitory at millimolar concentrations, probably by competing with Ca2+ [4]. EDTA and EGTA can inhibit the Ca2+ -triggered luminescence of aequorin in two ways: (1) when free Ca2+ is removed from the reaction medium by chelation, the luminescence reaction is practically stopped; and (2) when the free (unchelated) forms of these chelators directly bind with the molecules of aequorin, inhibition results [30, 44]. The second type of inhibition is strong in solutions of low ionic strength and in the absence of other inhibitor ions such as Mg2+ , but it is relatively weak in the presence of 0.1 M KCl [47], presumably because aequorin is already inhibited by KCl. Therefore, great care must be taken if EDTA or EGTA is to be used in the calibration of the Ca2+ sensitivity of aequorin; this is especially important in the case of low-ionic-strength calcium buffers. It should also be noted that in usual calcium buffers, the lower the Ca2+ concentration, the higher the inhibitory free chelator concentration, resulting in a slope steeper than the true slope in the log–log plot of luminescence intensity versus Ca2+ concentration. 4.6 Recombinant Aequorin

The cloning and expression of apoaequorin cDNA was accomplished by two independent groups in 1985. One of these groups analyzed the cDNA clone AQ440 they obtained and reported that apoaequorin is composed of 189 amino acid residues (Mr 21 400) with an NH2 -terminal valine and a COOH-terminus proline [12, 12], which is consistent with the results of the amino acid sequence analysis of native aequorin reported by Charbonneau et al. [8]. In contrast, the other group reported that the

15


cDNA AEQ1 they obtained contains the entire protein-coding region of 196 amino acid residues, which includes seven additional residues attached to the N-terminus, and the apoaequorin expressed in Escherichia coli showed a molecular weight of 20 600 [13, 14]. The recombinant aequorin of the former group did not exactly match any of the isoforms of natural aequorin in the HPLC mobilities and the properties of Ca2+ -triggered luminescence [70]. No detailed comparison has been made with the recombinant aequorin obtained by the latter group, although a brief test indicated that the recombinant aequorins from both sources are practically identical. 4.7 Semi-synthetic Aequorins

The core cavity of the aequorin molecule can accommodate various synthetic analogues of coelenterazine in place of coelenterazine. The coelenterazine moiety in native aequorin can be replaced by a simple process. First, aequorin is luminesced by the addition of Ca2+ , and then the apoaequorin produced is regenerated with an analogue of coelenterazine in the presence of EDTA, 2-mercaptoethanol (or DTT), and molecular oxygen. The products are called semi-synthetic aequorins and are identified with an italic prefix (see Table 2). Semi-synthetic aequorins can be prepared from both native aequorin and recombinant aequorin, using various synthetic analogues of coelenterazine. A large number of coelenterazine analogues were synthesized, and about 50 kinds of semi-synthetic aequorins have been prepared and tested [70–73]. Some semi-synthetic aequorins are significantly different from the native type of aequorin in various properties, including spectral characteristics [51] and sensitivity to Ca2+ , the rate of luminescence reaction, and the rise time of luminescence (Table 2). The relationship between Ca2+ concentration and the initial light intensity of various semi-synthetic aequorins is shown in Fig. 2. As is apparent from the data of Table 2 and Fig. 2, the tolerance of the central cavity of the aequorin molecule in accommodating the coelenterazine moiety is surprisingly wide. The apparent limitations in the substitution of the coelenterazine moiety, and the changes in the Ca2+ sensitivity caused by the substitution, are as follows. 1. The group R1 must be aromatic. A replacement of the original p-hydroxyphenyl group with a group of larger size tends to decrease Ca2+ sensitivity. 2. The group R2 must be lager than the ethyl group. The replacement of the original phenyl group with a smaller non-aromatic group increases Ca2+ sensitivity. 3. The group R3 must be an OH group, and no substitution is allowed on the phenyl group bearing this OH. 4.7.1 e-Aequorins e-Aequorins, containing a ligand of e-coelenterazine, show properties significantly different from other aequorins [70–73]. In the structure of e-coelenterazines, the 5 position of the imidazopyrazinone structure is bound with the α position of the 6(p-hydroxyphenyl) group through an ethylene linkage, thus restraining the two ring

4 The Photoprotein Aequorin Table 2 Selected semi-synthetic aequorins derived from recombinant aequorin [76]

No. (Prefix)

1 2 (h) 3 (f ) 4 (f2) 6 (cl) 9 (n) 9 (n/J)5 12 (cp) 13 (ch)

Structural modification of coelenterazine1

None R1 : C6 H5 R1 : C6 H4 F(p) R1 : C6 H3 F2 (m,p) R1 : C6 H4 Cl(p) R1 : β-naphthyl R1 : β-naphthyl R2 : cyclopentyl R2 : cyclohexyl R1 : C6 H4 F(p), R2 : n-butyl 17 (fb) R1 : C6 H5 , R2 : cyclopentyl 19 (hcp) R1 : C6 H5 , R2 : cyclohexyl 21 (hch) R1 : C6 H4 F, R2 : cyclohexyl 22 (fch) 23 (m5) R4 : methyl 24 (e) R5 : CH2 CH2 R1 : C6 H4 F(p), R5 : CH2 CH2 26 (ef ) R2 : cyclohexyl, R5 : CH2 CH2 27 (ech) Fluorescein-labeled6

Luminescence max (nm)

Relative luminescence capacity2

Relative intensity at 10−6 or 10−7 M Ca2+3

Half-total time(s)4

466 466 472 470 464 468 467 442 453

1.00 0.75 0.80 0.80 0.92 0.25 0.30 0.63 1.00

1.00 16 20 30 0.6 0.15 0.07 28 15

M M M M 5 5 5 F F

460

0.20

1100

2

445

0.65

500

F

450

0.52

80

F

462 440 405, 472

0.43 0.37 0.50

73 2 6

M M F

405, 470

0.35

40

F

402, 440 528

0.40 1.00

8 2

F M

1 Only the changes from the coelenterazine structure are shown in this column. Those unchanged are shown in parentheses in the above structures. 2 The ratio in luminescence capacity: semi-synthetic aequorin/unmodified aequorin. 3 The ratio in luminescence intensity: semi-synthetic aequorin/unmodified aequorin, in 10−7 M Ca2+ for a value of 1 and larger and in 10−6 M Ca2+ for a value of less than 1. 4 The time required to emit 50% of the total light in 10 mM calcium acetate: F, 0.15–0.3 s; M, 0.4–0.8 s. The half-rise time of luminescence: F, 2–4 ms, all others, 6–20 ms. 5 Prepared from aequorin isoform J. 6 Fluorescein was chemically bound to apoaequorin, followed by regeneration using unmodified coelenterazine.

17


Fig. 2 Relationship between Ca2+ concentration and the initial light intensity of various recombinant semi-synthetic aequorins and n-aequorin J (a semi-synthetic natural aequorin made from an isoform, aequorin J). The curve number corresponds to the photoprotein number used in Table 2. A

photoprotein (3 :g) was added to 3 mL of Ca2+ buffer with various pCa values (pH 7.0) containing 1 mM total EGTA, 100 mM KCl, 1 mM free Mg2+ , and 1 mM MOPS, at 23–24 ◦ C. The data are taken from Shimomura et al. [76].

systems into the same plane. The luminescence reactions of e-aequorins are fast, with a half-rise time of 2–4 ms and a half-total time of 0.15–0.3 s, like ch-aequorins with an 8-cyclohexylmethyl substituent. The luminescence spectra are bimodal, with peaks at 400–405 nm and 440–475 nm. The ratio of the two peaks is variable not only with the type of aequorin but also with the measurement conditions, such as the concentration of Ca2+ and pH. e-Coelenterazines scarcely luminesce in the presence of apoaequorin, Ca2+ , and 2-mercaptoethanol in air [51].

References 1 SHIMOMURA, O., JOHNSON, F. H., SAIGA, Y. Extraction, purification and properties of aequorin, a bioluminescent protein from the luminous hydromedusan, Aequorea. J. Cell. Comp. Physiol. 1962, 59, 223–239. 2 SHIMOMURA, O., JOHNSON, F. H. Partial purification and properties of the

Chaetopterus luminescence system. In Bioluminescence in Progress (JOHNSON, F. H., HANEDA, A., Eds.), pp. 495–521. Princeton University Press: Princeton, NJ, 1966. 3 SHIMOMURA, O. Bioluminescence in the sea: photoprotein systems. Symp. Soc. Exp. Biol. 1985, 39, 351–372.

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37 DUNSTAN, S. L., SALA-NEWBY, G. B., FAJARDO, A. B., TAYLOR, K. M., CAMPBELL, A. K. Cloning and expression of the bioluminescent photoprotein pholasin from the bivalve mollusc Pholas dactylus. J. Biol. Chem. 2000, 275, 9403–9409. 38 SHIMOMURA, O., JOHNSON, F. H. Chaetopterus photoprotein: crystalli-za tion and cofactor requirements for bioluminescence. Science 1968, 159, 1239–1240. 39 NICOLAS, M. T., BASSOT, J. M., SHIMOMURA, O. Polynoidin: a membrane photoprotein isolated from the bioluminescent system of scale-worms. Photochem. Photobiol. 1982, 35, 201–207. 40 TSUJI, F. I., LEISMAN, G. B. K+ /Na+ -triggered bioluminescence in the oceanic squid Symplectoteuthis oualaniensis. Proc. Natl. Acad. Sci. USA 1981, 78, 6719–6723. 41 TAKAHASHI, H., ISOBE, M. Photoprotein of luminous squid, Symplectoteuthis oualaniensis and reconstruction of the luminous system. Chem. Lett. 1994, 5, 843–846. 42 ISOBE, M., FUJII, T., KUSE, M., MIYAMOTO, K., KOGA, K. 19 F-Dehydrocoelenterazine as probe to investigate the active site of symplectin. Tetrahedron 2002, 58, 2117–2126. 43 LOOMIS, H. F., DAVENPORT, D. A lumi nes-cent new xystodesmid milliped from California. J. Wash. Acad. Sci. 1951, 41, 270–272. 44 HASTINGS, J. W., DAVENPORT, D. The luminescence of the millipede, Lumino-desmus sequoiae. Biol. Bull. 1957, 113, 120–128. 45 SHIMOMURA, O. A new type of ATP-activated bioluminescent system in the millipede Luminodesmus sequoiae. FEBS Lett. 1981, 128, 242–244. 46 WOOD, K. V., De WET, J. R., DEWJI, N., DELUCA, M. Synthesis of active firefly luciferase by in vitro translation of RNA obtained from adult lanterns. Biochem. Biophys. Res. Commun. 1984, 124, 592–596. 47 SHIMOMURA, O., SHIMOMURA, A. Effect of calcium chelators on the Ca2+ -dependent luminescence of aequorin. Biochem. J. 1984, 221, 907–910.

References 48 SHIMOMURA, O. Bioluminescence of the brittle star Ophiopsila californica. Photochem. Photobiol. 1986a, 44, 71–674. 49 SHIMOMURA, O. A short story of aequorin. Biol. Bull. 1995c, 189, 1–5. 50 HENRY, J. P., MONNY, C. Proteiprotein interaction in the Pholas dactylus system of bioluminescence. Biochemistry 1977, 16, 2517–2525. 51 SHIMOMURA, O., JOHNSON, F. H. Extraction, purification and properties of the bioluminescence system of the euphausiid shrimp Meganyctiphanes norvegica. Biochemistry 1967, 6, 2293–2306. 52 JOHNSON, F. H., SHIMOMURA, O. Introduction to the bioluminescence of medusae, with special reference to the photoprotein aequorin. Method. Enzymol. 1978, 57, 271–291. 53 JOHNSON, F. H., SHIMOMURA, O. Preparation and use of aequorin for rapid microdetermination of Ca2+ in biological systems. Nature 1972, 237, 287–288. 54 SHIMOMURA, O. Isolation and properties of various molecular forms of aequorin. Biochem. J. 1986b, 234, 271–277. 55 BLINKS, J. R., HARRER, G. C. Multiple forms of the calcium-sensitive bioluminescent protein aequorin. Fed. Proc. 1975, 34, 474. 56 SHIMOMURA, O., JOHNSON, F. H. Chemistry of the calcium-sensitive photoprotein aequorin. In Detection and Measurement of Free Calcium Ions in Cells (ASHLEY, C. C., CAMPBELL, A. K., Eds.), pp. 73–83. Elsevier/North-Holland: Amsterdam, 1979a. 57 LOSCHEN, G., CHANCE, B. Rapid kinetic studies of the light emitting protein aequorin. Nature New Biology 1971, 233, 273–274. 58 HASTINGS, J. W., MITCHEL, G., MATTINGLY, P. H., BLINKS, J. R., van LEEUWEN, M. Response of aequorin bioluminescence to rapid changes in calcium concentration. Nature 1969, 222, 1047–1050. 59 SHIMOMURA, O., JOHNSON, F. H. Further data on the specificity of aequorin luminescence to calcium. Biochem. Biophys. Res. Commun. 1973, 53, 90–494. 60 SHIMOMURA, O., JOHNSON, F. H. Specificity of aequorin bioluminescence

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1

The Outer Mitochondrial Membrane Pore (Voltage-Dependent Anion Channel) Mikhail Vyssokikh Moscow State University, Moscow, Russia

Dieter Brdiczka University of Konstanz, Konstanz, Germany

Originally published in: Bacterial and Eukaryotic Porins. Edited by Roland Benz. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30775-3

1 Introduction

Eukaryotic porins are membrane proteins that form aqueous channels in the cell membrane and the mitochondrial outer membrane. In contrast to bacterial porins that have a similar function, the structure of eukaryotic porins is not known at a useful resolution. The mitochondrial outer membrane pore was first characterized by Marco Colombini [1] as a voltage-dependent anion channel (VDAC). The properties of VDACs have been widely investigated in reconstituted systems by several groups [1, 2] exploring the main differences compared to bacterial porins, i.e. sensitivity to voltage already at 30 mV and to the polarity of the applied voltage. In general, bacterial and mitochondrial outer membranes have segregating functions, and pore proteins in the membranes control limited exchange. However, in bacteria, the outer membrane has more protective functions, whereas in mitochondria, communicative functions are more important. The VDAC plays an important role in the coordination of the communication between mitochondria and cytosol. A substantial aspect of this management is a transient formation of complexes with other proteins. This will be the topic of this article. 2 Structure and Isotypes of VDAC

Although the VDAC amino acid sequence is very different compared to the bacterial porins, it is assumed that the mitochondrial outer membrane pore may form a Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 The Outer Mitochondrial Membrane Pore (Voltage-Dependent Anion Channel)

β-barrel composed of 16 β-strands analogous to the know structure in bacteria [3]. Three different isotypes of VDAC are expressed in cells [4]. Not much is known in mammals about tissue-specific distribution, intracellular localization and function of the different isotypes. All experiments mentioned here relate to the VDAC isotype 1. This isotype might be the main species in the outer mitochondrial membrane. This is deduced from experiments with yeast expressing two VDAC isotypes. When type 1 VDAC was deleted, type II could not replace it, but instead TOM 40, a pore for mitochondrial precursor peptides, was overexpressed [5]. Furthermore, Anflous et al. [6] followed their expression in different tissues by cloning rat heart VDAC isotypes. It was observed that VDAC type 1 was the most abundant.

3 The Influence of Phospholipids on VDAC Structure

Sterols have been found in isolated and purified mitochondrial VDAC such as ergosterol in the case of Neurospora crassa [7] and cholesterol in VDAC from bovine heart [8]. Sterols influence the channel properties of reconstituted VDAC. Popp et al. [9] found them to be essential for proper insertion of soluble precursor VDAC into bilayer membranes and also observed a 10 times increase of conductance after addition of cholesterol to native VDAC. It was postulated by the authors that sterols may increase the hydrophobicity of the surface of the VDAC β-barrel structure exposed to the phospholipid matrix of the membrane. The sterols may thus be required to obtain the high-conducting VDAC structure. Complete saturation of VDAC with sterols may even cause a reduction of voltage-dependent conductance regulation. The outer mitochondrial membrane contains 10% cholesterol, while almost nothing is found in the inner membrane [10]. However, the cholesterol in the outer membrane appears to be clustered. Ardail et al. [11] were able to isolate two contact site fractions differing significantly in cholesterol content (9.4 versus 0.4 %). Both fractions bound hexokinase and, thus, contained the specific binding protein VDAC. Treatment of mitochondria with digitonin that interacts with cholesterol removes only parts of the outer membrane, leaving small vesicles attached at contact sites were hexokinase is bound [12]. This also suggests inhomogeneous distribution of cholesterol in the outer membrane. As with Ardail et al., we were able to characterize different types of contact sites on the basis of their composition by different functionally interacting proteins, as will be described below. We determined the lipid composition of the different contact-forming complexes (Table 1) and found a 5-fold difference of cholesterol content between two contact site-generating complexes. On the basis of this observation we assume the existence of structures like lipid rafts [13] in the outer membrane as a kind of outer membrane-signaling phospholipid domain. The localization of VDAC inside or outside the cholesterol-containing lipid raft-like areas might explain the different properties and structure of VDAC observed in the two complexes, as will be described below.

5 Physiological Significance of the Voltage Dependence Table 1 Lipid composition of hexokinase and creatine kinase complexes from contact sites

of rat kidney mitochondria Fraction Kidney creatine kinase from cristae creatine kinase (periph.) hexokinase complex Liver OMCS IMCS

Cholesterol

Cardiolipin

Cardiolipin/cholesterol

1 10 2

36 26 22

36 2.6 11

9.4 0.4

20.2 21.5

2.15 53.7

In the periphery, mitochondrial creatine kinase interacts with VDAC in the outer membrane and car-diolipin in the peripheral inner membrane. The enzyme is also localized in the cristae, where it interacts with two inner membranes. For comparison, the data from Ardail et al. [11] are shown at the bottom (OMCS = outer membrane contact sites, IMCS = inner membrane contact site). Data are shown as percentage of total lipid mass.

4 VDAC Conductance and Ion Selectivity

Analysis of the isolated VDAC after reconstitution in artificial bilayer membranes resulted in the calculation of the pore diameter of 4 nm at a voltage smaller than 30 mV. In this high conductance state (4 nS at 1 M KCl in the bathing fluid) the pore is anion selective. Above 30 mV, the diameter of the pore is reduced to 2 nm. The conductance decreases to 2 nS and ion selectivity changes to cation selectivity [1, 2]. The voltage-dependent conductance variations are linked to large structural modifications of VDAC. So far, no information about the nature of these changes is available. It has been postulated that a positively charged loop moves out of the channel [14]. However, it is also possible that negative charges move into the mouth of the channel, as has been observed for bacterial porins [15]. Macromolecules such as dextran, that cannot penetrate the pore, have an osmotic effect on the aqueous interior of the channel [16] and thereby increase voltage sensitivity. In the presence of dextran the low-conductance, cation-selective state is adopted already at 10 mV [17]. It has been postulated that the pore acquires this state in the contact sites where the inner membrane potential extends to the outer membrane by capacitive coupling [18]. Recently, a new idea of generation of a potential at the outer membrane was proposed on a basis of metabolic cycling and translocation of ATP, ADP and Pi across the inner membrane, and different resistance in the contact sites [19]. 5 Physiological Significance of the Voltage Dependence

VDAC in the low-conductance, cation-selective state is not permeable for ATP, ADP and other negatively charged small molecules. This has been demonstrated

3


by Colombini et al. investigating VDAC reconstituted in artificial membranes [20]. In isolated mitochondria, enzymes in the inter-membrane space such as adenylate kinase, had no access to external adenine nucleotides in the presence of König’s polyanion [21] that induces the low-conductance state of VDAC [22]. If we convey these results to the physiological situation where more contact sites are formed and macromolecules are present, we have to assume that most VDAC are in the low-conductance, cation-selective state. Indeed, in situ mitochondria of permeable cardiomyocytes or skinned red muscle fibers had a 10 times higher apparent K m for ADP compared to isolated mitochondria from the same tissue. The high K m could be reduced to in vitro values by generating porous outer membranes with digitonin or by the pro-apoptotic protein Bax, suggesting that in situ the outer membrane was not freely permeable for ADP [23]. In addition to pure voltage-dependent regulation, specific proteins in the inter-membrane space or attached to the outer surface of mitochondria have been described that physiologically may as well be involved in regulation of the pore permeability [24, 25].

6 Porins as Specific Binding Sites

In bacteria, porins are receptors for various phages. VDAC at the mitochondrial periphery acts in a similar way as a binding site for enzymes such as hexokinase [26] and glycerol kinase [27]. A shift in trans-membrane topology in the membrane also changes the surface-exposed loops between the β-strands. Considering the voltage dependence of the trans-membrane topology, this means that VDAC in the contact sites might have a role to reflect functional changes of the inner membrane potential at the mitochondrial surface. Alternatively, such a signal at the surface of the outer membrane may be generated by interaction of VDAC with the adenine nucleotide translocator (ANT; see below) if we assume that the ANT would interact exclusively with a certain VDAC state.

7 VDAC senses Inner Membrane Functions in the Contact Site

In support of this idea, it has been observed by immune electron microscopy and binding studies that hexokinase binds with higher capacity to the outer membrane pore in the contact sites compared to pores beyond contacts [28]. Freeze-fracturing analysis of isolated liver mitochondria revealed that contact sites could be induced by dextran or atractyloside and suppressed by glycerol or dinitrophenol [29, 30]. Thus, hexokinase binding to isolated outer membrane or mitochondria with induced or suppressed contact sites was studied. Hexokinase showed sigmoidal-type binding to mitochondria with contact sites, while binding to pure outer membrane or mitochondria with suppressed contact sites led to hyperbolic binding curves [29]. This suggested a cooperative binding of hexokinase to the pore in the contact

9 Isolation and Characterization of VDAC-ANT Complexes

site area. In agreement with this, hexokinase, by binding, was activated and bound hexokinase was found to form tetramers [31]. In a recent investigation, Hashimoto and Wilson were analyzing different epitopes at the surface of bound hexokinase by specific monoclonal antibodies. The authors observed a variation of the bound hexokinase structure that correlated with functional changes of oxidative phosphorylation in the inner membrane [32]. This direct reflection of inner membrane functions at the mitochondrial periphery can be explained by interaction between VDAC in the outer and ANT in the inner mitochondrial membranes [33–35]. Because of the high capacity of contact sites for hexokinase binding, the enzyme was used as a marker for isolation of this membrane fraction from osmotically disrupted mitochondria by density gradient centrifugation [36, 37]. In a recent application, hexokinase was used to indicate structural changes of VDAC correlated to the functional state of ANT. The contact site isolation method in kidney mitochondria revealed that hexokinase activity in the contacts was decreasing when the ANT was shifted to the m-conformation by pre-treatment with bongkrekic acid, whereas it was increasing after induction of the c-conformation by pre-incubation with atractyloside (Figure 1). We do not know how the structure of the VDAC changes and whether the alterations are induced by the membrane potential or by the interaction with the ANT. However, we can summarize the observations of voltage-dependent conductance changes and studies of hexokinase binding as showing that VDAC alters its transmembrane topology correlated to functional mitochondrial states. Because it affects the affinity and capacity of hexokinase binding, we can define a tensed and relaxed state of the pore in analogy to the regulation of allosteric enzymes. The tensed state would be the low-conductance, cationically selective pore in a complex with the ANT, whereas the relaxed state would be characterized by high conductance and anion selectivity without complex formation (Figure 2).

8 Cytochrome c is a Component of the Contact Sites

It was observed that not only hexokinase, but also cytochrome c is a component of the contact sites, and its concentration increased in the presence of atractyloside and decreased upon treatment with bongkrekic acid (Figure 1). Thus, as observed for hexokinase, the structure of the ANT was responsible for the cytochrome c distribution at the surface of the inner membrane.

9 Isolation and Characterization of VDAC-ANT Complexes

Complexes between VDAC and ANT were generated in vitro with isolated porin and ANT proteins [33]. However, they were also isolated from Triton X-100-dissolved brain or kidney membranes by binding the attached hexokinase to an anion

5


Fig. 1 Variation of cytochrome c and hexokinase in the contact site fraction of kidney mitochondria by perturbing ANT structure. After osmotic shock subfractions of rat kidney mitochondria were separated in a sucrose density gradient (54.3 %, left; 30.8 %, right). The distribution of inner membrane

(SDH) and contact site (HK) fractions is shown. Cytochrome c concentration (empty circles) in the different fractions was determined by specific antibodies and varied significantly in 500 :M atractyloside-treated (A), compared to 250 :M bongkrekatetreated (B) mitochondria.

exchanger. It was found that the bound hexokinase complex contained VDAC and ANT isotype I [35]. A second complex composed of porin, creatine kinase and ANT could also be isolated from the same membrane extracts [34, 38]. Based on the separation and purification of the different complexes, two types of VDAC-ANT aggregates could be defined: one complex where VDAC and ANT interacted directly and VDAC achieved higher affinity for hexokinase (Figure 2A), and a second complex where VDAC interacted indirectly with the ANT through a mitochondrial creatine kinase octamer. In this case, VDAC exposed a different structure at the surface that had low affinity for hexokinase (Figure 2D), as hex-okinase activity

10 Reconstitution of VDAC-ANT Complexes

Fig. 2 Different configuration of the VDACANT complex connected to either hexokinase or creatine kinase. Each of the three components of the two kinase complexes can exist in different configurations that support or inhibit complex formation. The ANT (ANT-c, induced by atracty-loside) in its c-conformation faces the cytosol and interacts with the outer membrane pore (VDAC), whereas ANT in the m-conformation (ANT-m induced by bongkrekate) orientates to the matrix and does not interact with VDAC. The ring represents cardiolipin encircling the ANT. Cardiolipin might change from the outer to inner leaflet of the membrane correlated to the ANT structure. The active ANT is a dimer. VDAC-t (tensed) is a dimer when it interacts with the ANT-c in a cholesterol-

free lipid matrix. VDAC-r (relaxed) is in a cholesterol-rich lipid matrix (depicted as bars). It is not bound to the ANTand may also exist as a monomer. This structure of VDAC has low affinity for hexokinase and is present in the complex with the octamer of creatine kinase (CK). (A) Hexokinase (HK) associates to oligomers preferentially in the contact sites formed by the VDACANT-c complex. The oligomerization of hexokinase leads to activation of the enzyme. (B) Monomers of hexokinase bind to VDAC beyond the contacts with less affinity and activity. (C) VDAC-ANT complexes may also exist free without bound kinases. (D) Creatine kinase (CK) binds to VDAC as octamer. The octamer preferentially associates with cardiolipin that is bound to the ANT.

was always absent in the creatine kinase complexes. From cross-linking studies in isolated yeast mitochondria we know that VDAC was a dimer in the intact outer membrane [39]. Determination of the molecular weight of the isolated complexes suggested that tetramers of the active kinases were present in both cases [34]. Hexokinase is active as a monomer of 100 kDa, whereas the functionally active mitochondrial creatine kinase unit is a dimer of 85 kDa. Four dimers of mitochondrial creatine kinase form a cubic structure with identical top and bottom faces that are able to connect two membranes [40]. It is known from in vitro studies that the octamer of mitochondrial creatine kinase interacts directly with the outer membrane pore [41], while the interaction with the ANT may be indirect through the cardiolipin [42] that is known to be tightly bound to the ANT [43] (Figure 2D).

10 Reconstitution of VDAC-ANT Complexes

The hexokinase–porin–ANT complex and the porin-creatine kinase–ANT complex were isolated and functionally reconstituted in vesicles [34]. ATP loaded into the

7


vesicles did not leak out, although VDAC was a component of the complexes. However, the kinases in the complexes had access to the internal ATP through the ANT. This was shown by either glucose-6-phosphate formation from external glucose via hexokinase or creatine phosphate generation by creatine kinase from external creatine. Both reactions were inhibited by blocking the antiporter activity of the ANT by atractyloside. The results suggested that the kinases, after reconstitution, were functionally coupled to the ANT. The ANT was controlling the exchange of adenine nucleotides through VDAC and was working as an antiporter [44]. However, the functional state of the ANT in the reconstituted complexes could be converted into an unspecific uniporter (resembling the permeability transition pore) by addition of Ca2+ , as has been demonstrated also for several related mitochondrial transport systems by Dierks et al. [45].

11 Importance of Metabolic Channeling in Regulation of Energy Metabolism

The tight functional coupling of peripheral kinases to the ANT is important because of two reasons: (i) it regulates the activity of oxidative phosphorylation by ADP and (ii) it increases the free energy (G) in the ATP system. To clarify the first point we may imagine the situation after eating a carbohydrate-rich meal: a high glucose concentration enters the resting muscle cell that must be phosphorylated and converted to UDP-glucose to accomplish incorporation into glycogen. The ATP for this process is more efficiently provided by oxidation of pyruvate in the mitochondria than by lactate formation. Hexokinase bound to the VDAC- ANT complex induces the oxidative pathway by converting intra-mitochondrial ATP into ADP and by that activating oxidative phosphorylation. To describe the second advantage of metabolic channeling, we have to consider that all ATP energy-consuming processes in the cell (including ion pumps) depend on the transfer of the γ -phosphoryl group of ATP according to the reaction: ATP+X = X-P +ADP. In a subsequent step, X-P dissociates when, for example, the ion is pumped. This means the efficiency of ATP to drive these phosphorylation reactions depends on the level of ADP and Pi during the reaction according to the equation of G = Go + RT ln [ADP] [Pi ]/[ATP]. Thus, the power (G) of ATP and the flux through the ATP-dependent reaction would increase if ADP and Pi would be low or be continuously withdrawn from the reaction. In the complex between mitochondrial creatine kinase and the ANT these requirements are perfectly fulfilled. The equilibrium of the reaction will not be reached as the ATP just exported is directly utilized and the ADP produced is immediately taken up into the matrix by the ANT. Pi is excluded from the reaction because of complex formation. Thus, metabolic channeling through functional coupling between ANT and creatine kinase pushes the balance of the reaction between creatine and ATP far to the side of phosphocreatine production. A further advantage of this coupling to the ANT is that the higher free energy of ATP at the surface of the inner membrane is preserved as a higher phosphocreatine/creatine quotient. ATP is electrogenically exported by the ANT at the expense of the membrane

12 The VDAC–ANT Complex as Permeability Transition Pore

potential. It has been calculated that this process increases the free energy of ATP by an additional 12 kJ mol−1 [46]. Thus, the phosphocreatine/creatine ratio could be increased by this quantity through coupling between mitochondrial creatine kinase and the ANT. It has been criticized that the functional interaction between ANT and creatine kinase may be physiologically irrelevant because of the approximately 10-fold difference in ATP turnover of the two partners. However, one has to consider that at least four of the slower ANT dimers may be attached to one of the fast creatine kinase octamers (Figure 2).

12 The VDAC–ANT Complex as Permeability Transition Pore

Mitochondria contain a yet-unidentified structure that forms a large unspecific pore under conditions of high matrix Ca2+ , Pi or oxidative stress [47]. Opening of this so-called permeability transition pore was fully reversible upon withdrawing Ca2+ [48]. Electrophysiological measurements revealed properties of the pore related to VDAC suggesting that it may be a component of the whole structure [49, 50]. The permeability transition pore could be specifically blocked by cyclosporin A, which binds to cyclophilin D, a peptidylprolyl cis–trans isomerase present in the mitochondrial matrix [48, 51]. By employing a cyclophilin D affinity matrix, a complex between VDAC and ANT was isolated by Crompton et al. after extracting heart mitochondrial membranes with the detergent CHAPS [52]. This complex was reconstituted in vesicles, and pore opening by Ca2+ and Pi addition was registered in cyclosporin A-sensitive manner. Comparable results were obtained with the isolated hexokinase–VDAC–ANT complex. Cyclophilin was co-purified through isolation of the complex [38]. The complex was reconstituted in phospholipid vesicles that were loaded with malate. The internal malate was released correlated to increasing Ca2+ concentrations. The process was inhibited by cyclosporin A, suggesting that the reconstituted hexokinase–porin–ANT complex had properties of the permeability transition pore (Figure 3A). Two questions arise: (i) whether the ANT is the only structure that causes permeability transition and (ii) whether the ANT forms the pore only as a complex with VDAC. Concerning the first question, it is possible that all related antiporters such as Pi /OH- or the Glu/Asp exchanger can form a permeability transition pore depending on the actual conditions. As to the second question, it is possible to claim that the purified ANT also forms unspecific pores in the presence of Ca2+ . This was shown by reconstitution of ANT in bilayer membranes [53] or in vesicles [54]. However, these pores were not sensitive to cyclosporin A as cyclophilin D was missing. Cyclophilin D is thus a second important regulatory component of the permeability transition pore [51, 55]. The idea that VDAC may also have regulatory functions of the permeability transition was supported by the analysis of the reconstituted VDAC–creatine kinase–ANT complex. In contrast to the reconstituted hexokinase complex there was no release of enclosed malate from the vesicles by addition of

9


Fig. 3 The VDAC–ANT complex as permeability transition pore. (A, left panel) The scheme depicts the presumable arrangement of the hexokinase porin ANT complex in the contact sites of mitochondria. Hexokinase (HK) binds as a tetramer to the VDAC–ANT complex. Glycerol kinase (GK) and the mitochondrial benzodiazipin receptor (mBr) bind to the same pore configuration. (A, right panel) The complex was isolated from a Triton X-100 extract of brain membranes and was reconstituted in phospholipid vesicles that were loaded with malate or ATP. Cyclophilin was co-purified with the complex and could be extracted from the complex with phosphate buffer, pH 4.5. The enclosed malate was released from the vesicles by Ca2+ between 100 and 600 :M. The malate release was inhibited either by cyclophilin extraction (–Cyp D) or by incubation of the vesicles with 0.5

:M cyclosporin A (CSA) after addition of 200 or 500 :M Ca2+ . According to Brustovetsky and Klingenberg [53], Ca2+ interacts with the cardiolipin (DPG) around the ANT, thus changing the structure of the ANT from a ATP/ADP antiporter to an unspecific uniporter state. The uniporter state of the ANT may act as a permeability transition pore (PTP) in intact mitochondria. (B, left panel) The scheme shows the possible arrangement of a complex between VDAC, the octamer of mitochondrial creatine kinase and the ANT in the contact sites of mitochondria. When the octamer dissociates, VDAC and ANT can directly interact. (B, right panel) The complex was isolated from a Triton X-100 extract of brain membranes and was reconstituted in egg yolk liposomes that were loaded with malate. The malate was not released by increasing Ca2+ concentrations (Octamer). When the

13 The VDAC–ANT Complex as a Target for Bax-dependent Cytochrome c Release

Ca2+ (Figure 3B). As shown schematically in Figure 3B, VDAC is linked to the octamer of creatine kinase that hinders interaction with the ANT. However, direct interaction between VDAC and ANT was possible after dissociation of the creatine kinase octamer. In this case malate was liberated dependent on Ca2+ addition (Figure 3). The release was inhibited by cyclosporin A. The results suggested that VDAC such as cyclophilin D might play a role in regulation of the permeability transition pore. A possible role of VDAC might be to keep the ANT in the c-conformation, exposing the ATP/ADP-binding site to the cytosolic surface of the inner membrane according to the re-orientating carrier model proposed by Klingenberg [56]. In this conformation that is induced by atractyloside, the conversion of the ANT into an unspecific pore by Ca2+ is facilitated and leads to permeability transition. However, if VDAC and the ANT interact with the octamer of mitochondrial creatine kinase, conversion of ANT to the permeability transition pore-like state would be suppressed (Figure 3). In fact this has been observed in a transgenic mouse model expressing mitochondrial creatine kinase in liver mitochondria [57]. The permeability transition pore was opened in isolated mouse liver mitochondria by atractyloside and Ca2+ . In contrast, in the liver mitochondria from transgenic mice containing creatine kinase, the permeability transition was inhibited by substrates that stabilized the octamer of creatine kinase such as creatine and cyclocreatine. It should be mentioned here that mitochondrial creatine kinase, in addition to ts important role of organization in the peripheral mitochondrial compartment, is also localized in the cristae [58] where it has certainly different functions and is not linked to VDAC.

13 The VDAC–ANT Complex as a Target for Bax-dependent Cytochrome c Release

There are several signal- and tissue-specific pathways to induce apoptosis. One of them is a release of cytochrome c from mitochondria [59–61] that activates caspases [60] by binding to a cytoplasmic protein Apaf-1 in the presence of dATP [62]. The mechanism by which external Bax releases cytochrome c is still controversial, and may also depend on the actual localization of cytochrome c in addition to the binding and incorporation of Bax at the mitochondrial surface. VDAC and ANT interact preferentially when adopting a specific molecular conformation. The ANT has to be in the c-conformation as atractyloside induces the complex with the outer membrane pore [35], while bongkrekate shifts the ANT ← Fig. 3 octamer was dissociated into dimer by 20 min incubation of the liposomes with 5 mM MgCl2 , 20 mM creatine, 50 mM KNO3 and 4 mM ADP, VDAC–ANT complexes could form. In these complexes Ca2+ could shift the ANT structure to the unspecific permeability transition pore (PTP) and

the enclosed malate was liberated (Dimer). The malate permeability of the PTP was inhibited by pre-incubation of the vesicles with 100 nM cyclosporin A (dimer + CSA). The octamer-dimer equilibrium of the mitochondrial creatine kinase can be shifted to dimer by reactive oxygen species (ROS) [74].

11


to the m-conformation and by that represses this interaction. The VDAC in the complex with the ANT appears to be in the low-conductance, cationically selective, tensed state. This can be deduced from the observation that heart muscle mitochondria in situ show a reduced response to external ADP [23]. The structural change of the outer membrane pore achieved by interaction with the ANT is recognized at the mitochondrial surface and leads to higher affinity of hexokinase. Recent observations suggest that this tensed pore structure is the preferred target of Bax molecules. Pastorino et al. [63] found that hexokinase and Bax compete for the same binding site, and Capano and Crompton [64] co-precipitated VDAC and ANT when Bax was immune precipitated from extracts of cardiomyocytes.

14 The VDAC–ANT Complexes contain Cytochrome c

As described above it was observed that the contact sites contained cytochrome c. As the VDAC–ANT complexes are derived from the contact sites it was not surprising that the complexes contained a significant amount of cytochrome c [65]. This was found although the complex was eluted from anion exchanger column by 200 mM KCl suggesting that the cytochrome c was not bound by ionic interaction. To investigate whether Bax would be able to interact with the cytochrome c within the VDAC–ANT complexes, the latter were reconstituted in phospholipid vesicles as described above (Figure 4A). After reconstitution, the vesicles were loaded by malate. Bax liberated the endogenous cytochrome c, but did not release the internal malate (Figure 4A). The Bax-dependent liberation of endogenous cytochrome c was abolished when the VDAC–ANT complex was dissociated by bongkrekate (Figure 4B).

15 The Importance of the Kinases in Regulation of Apoptosis

Release of cytochrome c from mitochondria proceeds in two steps. The first involves the release of a small fraction of cytochrome c from the compartment between the outer and peripheral inner membranes and the contact sites. It is induced by Bax at low concentrations [66] in the early phase [64]. The second step includes release of additional fractions of cytochrome c located at the surface of the cristae membranes and results from opening of the permeability transition pore, followed by mitochondrial swelling and membrane disruption [67]. The massive cytochrome c release may also be a consequence of VDAC closure preceding membrane disruption [68]. The observed failure of mitochondria to exchange adenine nucleotides with the cytosol may be a consequence of the transformation of VDAC into the tensed state. The two described kinases bound to VDAC are involved in the regulation of both processes of cytochrome c liberation.

15 The Importance of the Kinases in Regulation of Apoptosis

Fig. 4 Bax-dependent release of cytochrome c from the hexokinase VDAC–ANT complex. (A) Schematic representation of the reconstituted hexokinase (HK)–VDAC (P)–ANT complex in multilayered vesicles. Cytochrome c (circles) that was still attached to the complexes could be released by Bax, as shown in the right panel. However, malate, loaded into the vesicles, was not liberated by Bax, but by 200–500 :M Ca2+

through opening the ANTas a permeability transition pore (Figure 3A). This suggested that Bax was specifically interacting with the cytochrome c in the porin–ANT complex. The sigmoidal curve of cytochrome c release might indicate that Bax is accomplishing this process as an oligomer. (B) Schematic representation of the association/dissociation of the hexokinase–VDAC (P)–ANT complex by atractyloside (ATR)

13


16 Suppression of Bax-dependent Cytochrome c Release and Permeability Transition by Hexokinase

Hexokinase binds to the outer membrane pore in its tensed state, when VDAC interacts with the ANT. In this state VDAC is not freely permeable for adenine nu-cleotides. This has been already observed in mitochondria in the presence of dextran that increases hexokinase binding and contacts between VDAC and ANT [29]. In these mitochondria, external pyruvate kinase had less access to the ADP produced by hexokinase [69]. Although VDAC in the tensed state does not allow ATP transfer, hexokinase utilizes mitochondrial ATP, suggesting a structural modification of VDAC through binding of the enzyme. On the other hand, the activity of mitochondrial-bound hexokinase was found to be important for protein kinase B-linked suppression of cytochrome c release and apoptosis [70]. The experiments suggested that following withdraw of growth hormone, Bax liberated the small fraction of cytochrome c from the contact sites. In complete agreement with this, the Bax-dependent release of cytochrome c bound in the VDAC–ANT complexes described above was inhibited by activity of hexokinase in the presence of glucose and ATP [65]. This was explained by stabilization of hexokinase binding to the VDAC that is also the main target of Bax. In addition to this effect on Bax-dependent cytochrome c release, activation of hexokinase inhibited Ca2+ -dependent opening of the permeability transition pore by ADP production [38].

17 Suppression of Permeability Transition by Mitochondrial Creatine Kinase

Another kinase which binds to VDAC is mitochondrial creatine kinase. This enzyme interacts with VDAC exclusively in the octameric state, while the dimer has only weak affinity to porin [41, 42]. Thus, the association–dissociation equilibrium between octamer and dimer determines the formation of octamer–VDAC complexes. Ligation of the coronary artery in guinea pigs led to decrease of octamer from 85 to 60 % in the infarcted area of the heart [71]. Because of the dissociation of the creatine kinase octamer, the possibility of direct VDAC–ANT interaction was increased. As suggested above, the latter complexes may be the prerequisite of Ca2+ -induced permeability transition, followed by mitochondrial swelling, ← Fig. 4 or bongkrekate (BA) after reconstitution in multilayer vesicles. Treatment of the complexes with bongkrekate changed the ANT into the m-conformation (ANTm). This led to dissociation of the complex and a correlated structural change of VDAC, with a decreased affinity for hexokinase and Bax. Cytochrome c might have redistributed

as well. As shown in the right panel, Bax was unable to release cytochrome c after the structural change of the complex by bongkrekate. All cytochrome c remained in the sediment (BA sed) in contrast to the control, without bongkrekate treatment, and nothing appeared in the supernatant (BA sup).

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18 Conclusion

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51 HALESTRAP A. P., DAVIDSON A. M. Inhibition of Ca + -induced large amplitude swelling of mitochondria by cyclosporin A is probably caused by binding to a matrix peptidylprolyl cis-trans-isomerase and preventing it interacting with the adenine nucleotide trans-locase. Biochem J 1990, 268, 153–160. 52 CROMPTON M., VIRJI S., WARD J. M. Cyclo-philin-D binds strongly to complexes of the voltage-dependent anion channel and the adenine nucleotide trans-locase to form the permeability transition pore. Eur J Biochem 1998, 258, 729–735. 53 BRUSTOVETSKY N., KLINGENBERG M. The mitochondrial ADP/ATP carrier can be reversibly converted into a large channel by Ca2+ . Biochemistry 1996, 35, 8483–8488. 54 RÜCK A., DOLDER M., WALLIMANN T., BRDICZKA D. Reconstituted adenine nucleotide translocase forms a channel for small molecules comparable to the mitochondrial permeability transition pore. FEBS Lett 1998, 426, 97–101. 55 CROMPTON M., ELLINGER H., COSTI A. Inhibition by cyclosporin A of a Ca2+ -dependent pore in heart mitochondria activated by inorganic phosphate and oxidative stress. Biochem J 1888, 255, 257–360. 56 KLINGENBERG M., GREBE K., FALKNER G. Interaction between the binding of 35 S-atractyloside and bongkrekic acid at mitochondrial membranes. FEBS Lett 1971, 16, 301–303. 57 O’GORMAN E., BEUTNER G., DOLDER M., KORETSKY A. P., BRDICZKA D., WALLIMANN T. The role of creatine kinase in inhibition of mitochondrial permeability transition. FEBS Lett 1997, 414, 253–257. 58 KOTTKE M., WALLIMANN Th, BRDICZKA D. Dual localization of mitochondrial creatine kinase in brain mitochondria. Biochem Med Metabol Biol 1994, 51, 105–117. 59 YANG J. L. X., BHALLA K., KIM C. N., IBRADO A. M., CAI J., PENG T. I., JONES D. P., WANG X. Prevention of apoptosis by Bcl-2: release of cytochrome c from

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19

1

The Phytochromes

Shih-Long Tu, and Clark Lagarias University of California at Davis, Davis, USA

Originally published in: Handbook of Photosensory Receptors. Edited by Winslow R. Briggs and John L. Spudich. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31019-7

1 Introduction 1.1 Photomorphogenesis and Phytochromes

Oxygenic photosynthetic organisms have evolved sophisticated mechanisms to adapt to their environment. Dependent upon light as an energy source, these organisms must also cope with too much light, which is especially challenging for highly pigmented species living in an aerobic environment. For this reason, plants have evolved light-receptor systems to recognize and respond to light quality, fluence rate, direction and duration from their environment. Among the many physiological processes under light control are seed germination, seedling growth, synthesis of the photosynthetic apparatus, the timing of flowering, neighbor detection, and senescence. Such light-regulated growth and developmental responses are collectively known as photomorphogenesis [19, 81, 119, 179]. Phytochrome was the first of the photomorphogenetic photoreceptor families to be identified nearly 50 years ago, [18]. In the last decade, research on the physiological, biochemical and functional properties of phytochromes has grown exponentially, [44, 54, 117, 129, 140]. More recent advances in this field can be attributed to the impact of genomics, studies that have revealed phytochrome-related genes in representative organisms from all forms of life on earth except Archaea, [111, 171]. The extended phytochrome family can be categorized into three subfamilies: plant phytochromes (Phy), cyanobacterial phytochrome 1 (Cph1) and cyanobacterial phytochrome 2 (Cph2) families (Figure 1). This review focuses on phytochromes that are found in oxygenic photosynthetic organisms.


2 The Phytochromes

Fig. 1 The phytochrome family. Orthologous photosensory input (GAF, PHY and PAS) and regulatory output (PAS and HKRD) subdomains are depicted in representative members of the phytochrome (Phy), cyanobacterial phytochrome 1/bacte-

riophytochrome (Cph1/BphP) and cyanobacterial phytochrome 2 (Cph2) families (n = 0–2). The linear tetrapyrrole (phytobilin) chromophore is shown associated with the P3 GAF and P4 PHY domains. Adapted from Montgomery and Lagarias (2002).

1.2 The Central Dogma of Phytochrome Action

Synthesized in the red light-absorbing Pr form, all phytochromes are regulated by red light (R) absorption which initiates the photochemical interconversion to the far-red (FR) light-absorbing Pfr form (Figure 2 A). FR promotes the reverse conversion of Pfr to Pr – a process which typically abolishes the R-dependent activation of the photoreceptor. This R/FR photoreversibility is conferred by a linear tetrapyrrole (phytobilin) prosthetic group that is covalently attached to the phytochrome apoprotein. Supporting evidence for the central dogma of phytochrome action, that is that Pfr is the active form, has been accumulating for years. Much of this evidence reflects the strong correlation between the amount of Pfr produced by a given fluence of light and the magnitude of the biological response. While the central dogma appears to hold true for R-FR-reversible low-fluence responses (LFR) and for R-dependent very low fluence responses (VLFR), [147], FR high irradiance responses (HIR) do not conform to this simple view of phytochrome action, [20, 55, 148]. Such data indicate that Pr, Pfr, photocycled-Pr, as well as intermediates produced during Pfr to Pr photoconversion, may all function to transduce the light signal for different phytochromes (Figure 2 B). In addition to photochemical interconversion processes, non-photochemical Pfr-to-Pr dark reversion plays an important role to attenuate the signal output by altering the lifetime of the Pfr form (Figure 2 B). For a given quality and f luence rate of light in the environment, signal output from phytochrome therefore depends on holophytochrome synthesis, the two photochemical interconversion processes, dark reversion as well as protein turnover. It is on these topics that we will mainly focus our discussion.

2 Molecular Properties of Eukaryotic and Prokaryotic Phytochromes 3

Fig. 2 Spectral properties and modes of action of plant phytochromes. Panel A shows absorption spectra of purified oat phytochrome after saturating red (dotted line) and far-red (solid line) irradiation representing a mixture of 87% Pfr/13% Pr and

100% Pr, respectively. The deduced specturm of Pfr (dashed line) was obtained as described (Kelly and Lagarias [80]). Panel B depicts the three modes of phytochrome action in flowering plants that have been described by action spectroscopy.

2 Molecular Properties of Eukaryotic and Prokaryotic Phytochromes 2.1 Molecular Properties of Plant Phytochromes

In flowering plants, phytochromes are encoded by a small nuclear gene family reflecting repeated gene duplication of a eukaryote phytochrome progenitor during its evolution, [102]. In the model plant Arabidopsis thaliana, the phytochrome family consists of 5 genes, denoted phyA–phyE, [27], while monocot species (eg. rice or maize) appear to possess only representatives of the phyA–C families, [103]. The number of phytochrome species is not known for more primitive plants, that is gymnosperms, mosses, ferns, liverworts and algae. However the overall structure of the phytochrome photoreceptor has been preserved in all extant eukaryotic photosynthetic organisms (Figure 1). Although atypical phytochromes have been found in mosses and ferns, these organisms also possess conventional phytochromes, [120, 167]. For this reason, we will limit our discussion to those eukaryotic phytochromes which conform to the consensus structure. Our present understanding of phytochrome structure is mainly based on biochemical analysis of phyAs, but for those studied, other phytochromes appear to possess similar properties. Eukaryotic phytochromes are soluble homodimeric proteins consisting of two ∼120 kDa subunits. Small-angle X-ray scattering analyses indicate an overall protein fold with dimensions similar to mammalian immunoglobin Gs, [118]. Each subunit is composed of two domains that are separated by a protease-sensitive hinge region: the 60–70 kDa N-terminal ‘photosensory’ and 55 kDa C-terminal

4 The Phytochromes

‘regulatory’ domains. The photosensory domain functions to sense the light input and is composed of four sub-domains, that is the serine-rich N-terminal P1 domain, a PAS-related P2 domain, the bilin-binding P3 GAF domain, and the P4 PHY domain, [111]. PAS and GAF domains have been shown to possess structurally-related protein folds, [67], with PAS domains often comprising the binding site for small planar aromatic ligand molecules, [4, 127, 158]. PHY domains, evolutionarily related to PAS and GAF domains, may also adopt a similar three dimensional fold, [111]. The phytobilin prosthetic group of plant phytochromes is covalently bound to a conserved cysteine residue found in the P3 GAF domain via a thioether linkage, [90]. The presence of this conserved cysteine residue is one of the key structural features that distinguishes the phytochrome photoreceptors from members of the phytochrome-related BphP family. Biochemical analyses have shown that the photosensory domain of plant phytochromes adopts a compact globular protein fold with the phytobilin chromophore mostly buried within the protein matrix, [188]. By contrast, the C-terminal regulatory domain of plant phytochromes is elongated and more sensitive to proteolytic degradation. Comprised of four sub-domains, including two PAS domains and two domains related to the transmitter ‘output’ domains found on prokaryotic twocomponent sensor proteins, [145], the regulatory domain of plant phytochrome contains the site(s) of homodimerization, [76]. All four subdomains have been implicated in the high-affinity subunit-subunit interaction suggesting that no single subdomain of the C-terminus is necessary or sufficient, [77, 170]. By analogy to twocomponent sensor proteins, it is reasonable that the R4 H-ATPase-related domain on one subunit interacts with the R3 histidine phosphotransferase (HPT)-related domain on the other subunit, [157]. The potential for heterotypic interactions between R1 and R2 PAS domains on different subunits are also reasonable but has not been directly assessed to date. The critical role of PAS domains to plant phytochrome function is underscored by the many loss-of-function alleles that map to these regions, [130]. The hypothesis that the PAS repeats play a central role in phytochrome signaling is also supported by localization of light-dependent conformational changes to this region, [87] and to the identification of nuclear targeting sequences within this domain, [24, 104]. As depicted in Figure 3, light-induced conformational changes have been shown to occur in both photosensory and regulatory domains of phytochromes with the consensus view being that the P1 domain becomes less exposed while P4, R1, and R2 domains become more exposed upon Pr to Pfr phototransformation, [87]. The surprising observation that the C-terminal domain is dispensable for phytochrome function, if it is replaced with a nuclear-targeted protein capable of homodimerization, indicates that R1–R4 subdomains regulate the subcellular localization, dimerization and overall topology of the N-terminal domain, [104]. Together with structure-function studies using chimeric phytochromes, [175], these results indicate that phytochrome signaling in the nucleus is mediated by protein-protein interactions with its N-terminal photosensory domain. Aside from homodimerization, the presence of transmitter kinase-related R3 and R4 ‘output’ domains on all eukaryotic phytochromes implicates their role in

2 Molecular Properties of Eukaryotic and Prokaryotic Phytochromes 5

Fig. 3 Light induced conformational changes of plant phytochromes. The lightdependent accessibility of various regions of plant phytochrome to modification/interaction by proteases (inverted ar-

rows), protein kinases, and monoclonal antibodies (filled arrowheads) is depicted on the oat phyA subunit. Adapted from McMichael and Lagarias [108].

ATP-dependent phosphotransferase activity. Phytochrome R4 domains possess all of the consensus residues found in H-ATPase domains of the transmitter kinase super-family which is consistent with the observation that plant phytochromes bind ATP, [184]. Plant phytochromes are also autophosphorylating, ATP-dependent serine-threonine protein kinases, [185, 189]. These results, along with the lack of a conserved autophosphorylating histidine residue in the R3 HPT-related domain, indicate that plant phytochromes are atypical transmitter kinases that have gained a new function during evolution, [22]. Resolution of an apparent enzymatic function for the transmitter-related domains of plant phytochromes with their dispensability remains an important, albeit unanswered question. The bonafide catalytic activity of transmitter domains of cyanobacterial phytochromes argue for an ATP-dependent enzymatic role for this region of plant phytochromes in light signaling, [190]. 2.2 Molecular Properties of Cyanobacterial Phytochromes

The Cph1 family encompasses those prokaryotic phytochromes with the greatest similarity to plant phytochromes (Figure 1). All members of this family, which include the bacteriophytochromes, possess three of the four photosensory domains (P2-P4) found on plant phytochromes as well as the two conserved subdomains found in transmitter modules of two-component histidine kinases, [125]. The two regulatory R1 and R2 PAS domains, as well as the serine-threonine rich P1 domain, are missing from all members of the Cph1 family. The first representative of this family to be identified was Cph1, a soluble 85 kDa protein from the cyanobacterium Synechocystis sp. PCC 6803, [73, 190]. Cph1-related genes are present in most cyanobacterial genomes examined; those with conserved cysteine residues in their P3 GAF domains are predicted to encode phytobilin-binding, R/FR photoreversible chromoproteins, [63]. Evidence that the latter is true has been reported for Synechocystis Cph1, [95, 124] and for CphA from Calothrix sp. PCC7601, [78]. Many cyanobacterial species have two representatives of this family in their genome, the other being a bacteriophytochrome, [63].

6 The Phytochromes

Owing to the ability to express and reconstitute large quantities of recombinant Cph1s in bacteria, the biochemical properties of these proteins have been extensively investigated, [78, 92, 95, 124]. Like plant phytochromes, Cph1s bind phytobilins via the conserved cysteine residue in the P3 GAF domain to generate RFR photoreversible chromo-proteins, [71, 78, 92]. Cph1 and CphA are also protein kinases, whose histidine autophosphorylation and aspartate phosphotransferase activities are both light dependent, [72, 190]. By contrast with plant phytochromes, both activities are Pr-dependent; thus light absorption leads to inhibition rather than activation of Cph1’s catalytic function. These studies have shown that Cph1’s enzymatic activities require both HPT and H-ATPase domains which function as phosphohistidine donor/acceptor and ATP-binding sites, respectively, [190]. The overall take-home lessons from biochemical studies on the Cph1 familiy of phytochromes are that 1) Cph1’s P2-P3-P4 photosensory domain structure and phytobilin-binding environments are very similar to those of eukaryotic phytochromes, 2) Cph1’s C-terminal HPT and H-ATPase domains are required for homodimerization, and 3) monomeric and dimeric forms of Cph1 are in dynamic equilibrium – a process that is regulated by both phytobilin binding and light. With regard to the third conclusion, size-exclusion chromatographic analysis has established that phytobilin binding to the Cph1 apoprotein promotes subunit-subunit association, [92, 124]. Taken together, these results support the working hypothesis that phytobilin binding to apoCph1 activates the kinase activity by promoting homodimerization and trans-autophosphorylation. While additional experiments are needed to elucidate the structural basis of light inactivation of Cph1’s kinase activities, this could be accomplished by increasing the subunit-subunit dissociation constant and/or by decreasing the affinity for ATP. These questions remain important topics for future investigation. The second family of prokaryotic phytochromes, that is the Cph2s, also have been found exclusively in cyanobacteria, [111]. With the exception of the founding member, that is Cph2 from Synechocystis sp. PCC6803, all members of the Cph2 family are composed of two to four GAF domains and terminate in a transmitter module containing both HPT and H-ATPase subdomains. The N-terminal GAF domains of Cph2s are most similar to the P3 GAF domains of plant and Cph1 phytochromes in which the conserved cysteine residue is located. Moreover, the GAF domain immediately adjacent to this conserved GAF domain possesses strong sequence similarity with P4 GAFs of both other phytochrome families, [111]. To test the functional significance of this similarity, recombinant Cph2 was shown to encode a phytobilin-binding 1276 amino acid apoprotein that exhibited a characteristic R/FR photoreversible-phytochrome spectral signature, [123]. The phytobilin binding activity and spectroscopic properties of the full length protein were retained within the two N-terminal GAF domains of Cph2, [186]. These results provided compelling support for the hypothesis that the P3 GAF and P4 PHY subdomains of phytochromes delimit the region of phytochromes that is in direct contact with the phytobilin prosthetic group. Through expression of the N-terminal GAF domain of Cph2, it was also shown that the P3 GAF subdomain is sufficient for the phytobilin lyase activity, [186]. Based on these results and the preserved GAF-PHY

3 Photochemical and Nonphotochemical Conversions of Phytochrome 7

molecular architecture for members of all three phytochrome families, we hypothesize that the initial molecular changes that accompany light activation of the phytobilin prosthetic group will be shared by all three classes of phytochromes. The widespread distribution of the Cph1 and Cph2 families of cyanobacterial phytochromes, together with their light-regulated enzymatic activities, suggests that these proteins perform important photosensory roles in these organisms. Interposon mutagenesis of Cph1 and Cph2 genes has so far failed to convincingly identify photoregulatory functions for their protein products although they appear to modulate phototaxis, [180]. Owing to the presence of many phytochrome-related genes in cyanobacterial genomes, the potential for redundant function may account for the lack of a clear phenotype of knockout mutants. Some of the potential functions of these proteins include both photosensory roles (i.e. phototaxis, lightintensity sensing, chromatic adaptation) as well as non-photosensory roles (i.e. regulators of tetrapyrrole and/or iron metabolism or oxygen sensors). There is clearly no dearth of interesting questions remaining to be answered on these two families of prokaryotic phytochromes.

3 Photochemical and Nonphotochemical Conversions of Phytochrome 3.1 The Phytochrome Chromophore

Based upon spectral measurements of phytochrome in 1959, Butler et al. proposed that the phytochrome chromophore was a linear tetrapyrrole similar to those found in phycobiliproteins – the major components of the phycobilisome lightharvesting antennae in cyanobacteria and red algae, [18]. Siegelman and colleagues later reported that the major pigment released from oat phytochrome by refluxing methanol was a linear tetrapyrrole, [149]. An oxidative degradation approach was also used to show that this compound was a 2,3-dihydrobiliverdin ‘phytobilin’ pigment similar to those derived from phycobiliproteins, [83, 137, 138]. The structure of this phytobilin was subsequently confirmed by total chemical synthesis to be 2R, 3E-phytochromobilin (PB) [177]. Together with peptide-mapping studies, [52], these data indicated that the phytobilin chromophore of oat phytochrome was covalently bound to the apoprotein via a thioether linkage – a hypothesis that was later confirmed by 1 H NMR spectroscopy, [90]. The structure and linkage of the Pr chromophore of oat phytochrome based on these studies, and its phytobilin precursor, are shown in Figure 4. The chromophores of no other phytochrome have received direct chemical scrutiny to date. In this regard, it has been generally assumed that the structure and linkages of all flowering-plant phytochrome chromophores are the same. The situation is less clear for phytochromes from more primitive plant species, except for the evidence that the phytochrome from the green alga Mesotaenium caldariorum and Cph1 from the cyanobacterium Synechocystis both possess chromophores

8 The Phytochromes

Fig. 4 The chromophore and phytobilin precursors of plant, green algal and cyanobacterial phytochromes. Panel A illustrates the structure and linkage of the chromophore of oat phytochrome A as deduced by 1 H NMR spectroscopy [90]. The PCB-derived chro-

mophores of green algal and cyanobacterial phytochromes are assumed to have identical linkages and stereochemistry by analogy. Panel B shows the phytobilin precursors of plant, green algal, and cyanobacterial phytochromes.

derived from the more reduced phytobilin prosthetic group, phycocyanobilin (PCB), [71, 187]. Linear tetrapyrroles are flexible molecules, and their spectroscopic properties are strongly influenced by their conformation, protonation state, and chemical environment, [43]. For this reason, the chemical structure of the phytochrome chromophore in situ is still the subject of debate. It is clear however that the phytobilin prosthetic group is protonated and adopts an extended configuration in situ – conclusions that are based on vibrational spectroscopic studies and the large ratio of phytochrome’s red to near UV absorption maxima, [13]. This contrasts with the small ratio observed for bilins in free solution which assume more cyclic, porphyrin-like conformations. Based on vibrational spectroscopic analysis and semi-empirical vibrational energy calculations, the 4Z-anti, 10E-anti, 15Z-syn configuration was proposed for the Pr chromophore, [2]. This contrasts with the proposed 4Z-anti, 10Z-syn, 15Z-anti configuration, also found in chromophores of the phycobiliproteins, that was based on similar arguments, [84]. This issue will likely remain controversial until a phytochrome crystal structure is determined. Regarding the Pfr chromophore structure, 1 H NMR analysis of chromopeptides derived from the Pfr form of oat phytochrome, [139], combined with vibrational spectroscopic studies, strongly implicate the Z to E isomerization of the C15 double bond upon Pr to Pfr photoconversion, [2, 3, 45, 64, 84, 109]. This hypothesis is further supported by the intense fluorescence of covalent adducts between apophytochromes and phycoerythrobilin (PEB), a PB analog that lacks this double bond, [116]. Taken together, these results indicate that the phytobilin chromophore of phytochrome is tightly tethered to the apoprotein in a manner enabling Z to E isomerization of the C15 double bond to be the only efficient mode of radiationless de-excitation.


3.2 Phytochrome Photointerconversions

The photochemical interconversion processes of phytochromes have been extensively studied for many years, [13, 14, 16, 82, 136]. Since Pr and Pfr have overlapping absorption spectra in most regions of the light spectrum, Pr-to-Pfr and Pfr-to-Pr photoconversion processes lead to the formation of a photoequilibium consisting of a mixture of Pr and Pfr forms under saturating illumination. The light fluence needed to produce this photoequilibrium depends on the intensity and wavelength of light used as well as the relative quantum yields for the Pr-to-Pfr and Pfr-to-Pr phototransformation processes, [16]. Photoequilibrium is most rapidly achieved at the absorption maxima of Pr and Pfr – red light (R) producing a mixture of roughly 87% Pfr and 13% Pr for oat and rye phyA, [80, 88]. Owing to the lack of Pr absorption in the far-red (FR) region of the light spectrum, FR irradiation can convert >99% of phytochrome to the Pr form, [101]. This information has been used to calculate the amount of Pfr produced by a given f luence of light providing the basis of support for the central dogma(s) of phytochrome action described previously. From time-resolved absorption and low-temperature trapping spectroscopic techniques, a number of intermediates accompaning the Pr-to-Pfr and Pfr-to-Pr photoin-terconversion processes have been identified. Two nomenclatures have been used to identify these intermediates – one based on the rhodopsin system and the other based upon the wavelength maxima of the intermediates. The former nomenclature will be used herein and in Figure 5. While the following discussion focuses on plant phytochromes (and mostly phyAs), comparative studies on Cph1 suggest that the initial photochemical interconversions are qualitatively similar, [47, 134, 150, 151]. Photoexcitation of Pr yields Lumi-R, the first ‘stable’ photoproduct that absorbs near 700 nm. This conversion occurs very rapidly; the half-life of the Pr excited state(s) lying between 25 and 50 ps, [1, 10, 70]. Based on resonance Raman analyses, it is generally accepted that Z, syn to E, syn isomerization of the C15 double bond occurs during this initial photoconversion, [2, 109]. This mechanistic interpretation is consistent with 1) low-temperature absorption kinetic measurements, [37], 2) the intense fluorescence of PEB-apophytochrome adducts which lack the C15 double bond, [116], and 3) the small deuterium isotope effect on Pr fluorescence yield – results that all but rule out intramolecular proton transfer as the primary mechanism of Pr excited-state decay, [15]. Lumi-R decay occurs in the microsecond timescale to meta-Ra, followed by its conversion into other kinetically distinguishable intermediates (eg. meta-Rc) within the microsecond to millisecond range, ultimately yielding Pfr. The temperatureand solvent-dependence of these non-photochemical interconversions implicate significant changes in phytobilin-apoprotein interactions. At least one of these processes involves the rotation about the C14–C15 single bond to yield the final C15 E,anti conformation of the Pfr chromophore (Figure 5). The chemical structures of these intermediates is subject of debate, with some investigators favoring a deprotonation-reprotonation mechanism while others assert that the chromophore

10 The Phytochromes

Fig. 5 Phytochrome photointerconversion processes. The Pr-to-Pfr photoconversion involves the light-dependent 15Zto-15E isomerization, followed by a lightindependent rotation about the C14–C15 bond and changes in chromophore-protein interactions. The Pfrto-Pr photoconversion

is envisaged to involve a concerted inplane photoisomerization of the C15 double bond, followed by a series of light-independent chromophore-protein relaxation steps. The lifetime and absorption maxima of the various spectroscopically detectible intermediates are shown. Adapted from [3].

remains protonated throughout, [13, 46]. The hypothesis that the phytobilin chromophore is moderately planar in both Pr and lumi-R but becomes more distorted from planarity upon conversion to Pfr is presently the most widely accepted interpretation of the available spectroscopic data, [13]. The overall Pr-to-Pfr photochemical quantum yield (r) has been determined to be in the range of 0.15–0.17 for plant phytochromes, [88]. This quantum yield is in agreement with that determined by femtosecond spectroscopy, [1] and by optoacoustic spectroscopy, [58]. The reverse Pfr-to-Pr photointerconversion is less well characterized for technical reasons but clearly this also proceeds through multiple intermediates, [23]. It is notable that Pfr fluorescence and Pfr-to-Pr photoconversion quantum yields are both lower than those of Pr and Pr-to-Pfr phototransformations, respectively. In this regard, the fluorescence quantum yield for Pfr is vanishingly small, that is less than 10−6 at room temperature, [178], and fr at 0.06–0.08 is less than half of that of r, [88]. Together with the results of resonance Raman analyses and theoretical arguments, [3], these observations have been interpreted to support a mechanism for the Pfr-to-Lumi-F photochemical interconversion that involves a


concerted configurational and conformational isomerization about the C15 double bond (Figure 5). The spectroscopic evidence for strong nonbonded interactions between the C- and D-ring methyl groups in the Pfr chromophore indicate that the excited state of Pfr and its photoproduct are strongly coupled, which may be a major structural feature which favors the concerted C15 E,anti to C15 Z,syn isomerization mechanism, [3]. The small value for fr suggests that one or more of the Pfrto-Pr intermediates can thermally dark-revert back to Pfr-like meta-R intermediates produced in the forward reaction. It also is conceivable that a second reversible photochemical reaction (i.e. proton transfer, amino acid adduct formation) had occurred in parallel with the photoisomerization mechanism to quench the Pfr excited state. Other than their absorption-spectroscopic signatures and thermal stabilities, little is known about the chemical structures of these intermediates and much still remains unknown regarding the reverse photointerconversion process. 3.3 Dark Reversion

From the first spectroscopic measurement of phytochromes, the process of nonphotochemical Pfr-to-Pr dark reversion became evident, [51]. Dark reversion of phytochrome has not only been measured in vitro, but has been observed in vivo, which lead to the hypothesis that this process is of physiological significance, [17]. The bulk of these studies were performed using dark-grown plant seedlings to reduce chlorophyll absorption which strongly overlaps with phytochrome and because these seedlings accumulate high levels of phyA, considerably improving the sensitivity of phytochrome photoassays. Spectrophotometric surveys of etiolated plant tissues revealed considerable variation in the amount of dark reversion for different plant species. While monocot plants showed considerably less dark reversion compared with dicots, the distinct morphology and cell types of these two classes of plants made it difficult to conclude whether this difference reflected the intrinsic properties or cellular environments of phytochromes from the two classes of plants, [51]. The development of methods to isolate and purify phytochromes have facilitated analysis of the molecular basis of dark reversion – a process that has been shown to be modulated by temperature, pH, various denaturants, proteases and reductants, [51]. As improvements have been made in the method of phytochrome purification, the discrepancy between in vivo and in vitro measurements of phytochrome dark reversion has been resolved. Consistent with in vivo measurements, full length monocot phyAs display very little dark reversion in vitro, while proteolytic removal of 50–100 amino acid at phyA’s N-terminus (i.e. the P1 domain) significantly promotes dark reversion, [172]. The stabilizing inf luence of the P1 domain has been seen for most phytochromes examined – its removal invariably leading to a 10–15 nm blue shift of the Pfr absorption maximum, [173]. Indeed, the lack of a P1 domain in Cph1 and Cph2 phytochromes may be responsible for the blueshifted absorption spectra and enhanced dark reversion of these cyanobacterial photoreceptors compared with flowering plant phyAs, [95, 190].

12 The Phytochromes

The low abundance of phyB-E in plants has made it difficult to measure their spectroscopic properties until recently. The development of methods to express and reconstitute recombinant holophytochromes has enabled investigation of the biochemical and spectroscopic properties of phytochromes whose genes have been cloned (see subsequent section). Studies on the Arabidopsis phytochrome family have shown that recombinant phyC and phyE exhibit significantly more dark reversion than recombinant phyA and phyB, [36]. The importance of dark reversion to phytochrome function is further underscored by the observation that the lossof-function phyB-101 mutant displays enhanced dark reversion, [38]. Localization of the phyB-101 mutation to the R2 PAS repeat implicates intra- and inter-molecular interations within the phytochrome dimer to stabilize the ‘active’ Pfr form. The possibility that other loss-of-function alleles of phytochrome may represent enhanced dark reversion has not been carefully assessed in all instances. It is based on these and other biochemical observations that the regulatory interplay between the N- and C-terminal regions of phytochromes is the subject of active ongoing investigation, [154]. The fact that plant phytochromes are all homodimers also raises the possibility that dark reversion of a Pfr chromophore on one subunit may be affected by the chromophore state on the other subunit, [55]. While there is evidence to support this hypothesis, for example that Pfr-Pfr homodimers dark revert more slowly than Pfr-Pr heterodimers, this simple interpretation does not hold true for all phytochromes so far examined, [36]. In summary, dark reversion plays an important role to regulate the phytochrome signal output — a role that future investigations will hopefully better resolve.

4 Phytochrome Biosynthesis and Turnover

Since light perception by all phytochromes depends upon the presence of the linear tetrapyrrole chromophore, the metabolic processes involved in the synthesis of its phytobilin prosthetic group and its assembly with apophytochrome are of fundamental importance to light-mediated plant growth and development. As phytobilin-binding proteins, phytochromes also influence tetrapyrrole metabolism in addition to their photoregulatory function. This section summarizes the current understanding of the pathways involved in the synthesis and assembly of phytobilins to apophytochromes. 4.1 Phytobilin Biosynthesis in Plants and Cyanobacteria

The finding that phytochromes possess linear tetrapyrrole prosthetic groups indicated that its synthesis would share common intermediates with heme and chlorophyll biosynthetic pathways. Early studies exploited the development of specific inhibitors of early steps in the tetrapyrrole pathway which could be

4 Phytochrome Biosynthesis and Turnover 13

overcome by feeding hypothetical intermediates after the blocked step. Oat seedlings treated with gabaculine or 5-aminohexynoic acid, potent inhibitors of glutamate-1-semialdehyde amino transferase (GSAT), were shown to induce both phytochrome and chlorophyll deficiencies, [40, 57]. Supplementation of the medium with 5-aminolevulinic acid (ALA), protoporphyrin or biliverdin IXα (BV) restored phytochrome levels to those of un-inhibited plants, thereby establishing the intermediacy of these compounds in the pathway of phytochrome chromophore biosynthesis, [39, 41, 57, 75, 86]. Based on these studies, it has been well established that the phytobilin biosynthesis shares intermediates with siroheme, protoheme, and chlorophyll biosynthetic pathways that are depicted in Figure 6. All plant tetrapyrroles are derived from glutamate which is converted to ALA via three enzymes: glutamate tRNA synthetase, glutamate tRNA reductase (GluTR) and GSAT, [6, 169]. Found in plants, algae, most eubacteria, and many archaebacteria, the glutamate C5 pathway for ALA synthesis can be contrasted with the C3 Shemin pathway of alpha proteobacteria and non-photosynthetic eukaryotes that utilizes the enzyme ALA synthase (ALAS). In all chlorophyll-based photosynthetic organisms, eight molecules of ALA are metabolized in several steps to yield protoporphyrin IX (Figure 6). A key branch point in chlorophyll and heme biosynthesis, protoporphyrin IX is metalated with magnesium or iron by two distinct chelatase systems to produce Mg-protoporphyrin or iron-protoporphyrin (heme). The intermediacy of heme in phytochrome chromophore biosynthesis was implicated by the observation that Mg-protoporphyrin was unable to restore holo-phytochrome levels in gabaculine-treated plants. Unfortunately, exogenous heme was not incorporated into phytochrome under the conditions used for ALA, protoporphyrin and BV rescue, [166]. Resolution of this impasse was made from two lines of investigation – development of an in organello assay system and molecular cloning of genes involved in phytochrome chromophore biosynthesis. With the knowledge that all plant tetrapyrroles are derived from ALA made in plastids, an isolated plastid system was used to address the hypothesis that linear tetrapyrroles are synthesized in this compartment, [162]. These studies took advantage of the ability of apophytochromes to assemble with phytobilin precursors to yield photochemically active holoproteins [see, [176] and later discussion]. By adding apophytochrome to the assay mixtures, these investigators were able to show that the entire pathway of phytochrome chromophore biosynthesis resides in plastids, [162] and later confirmed the intermediacy of ALA, heme, and biliverdin in its synthesis, [166]. While the low abundance of the enzymes unique to the pathway of phytobilin biosynthesis has proven challenging to their purification and molecular cloning, the key breakthrough on this subject was made with the molecular cloning of HY1 and HY2, [85, 114] – two loci known from previous studies to be involved in the synthesis of the phytochrome chromophore in Arabidopsis, [126]. These studies have confirmed that the phytochrome chromophore is derived from heme via the intermediacy of BV and PB (see Figure 4 for structure). The properties of the plant-specific heme oxygenase and phytobilin synthases are described in the following section.

14 The Phytochromes

Fig. 6 Phytobilin biosynthetic pathways in plants, cryptophytes, and cyanobacteria. Phytobilin biosynthesis shares common intermediates with siroheme, heme, and chlorophyll pathways up to the level of protopor-phyrin IX. Iron insertion leads to heme, which is subsequently converted to BV by heme oxygenase and to phytobilins by a member of the ferredoxin-dependent bilin reductase (FDBR) family. Magnesium insertion leads to the chlorophylls. Heme, BV, phytobilins, and chlorophylls assemble with their respective apoproteins – processes that draw off these intermediates. Alternative pathways for ALA synthesis, heme, and BV metabolism found in animals and some bacteria are shown in the

stipled boxes. Exogenous intermediates that were incorporated into the phytochrome chromophore are underlined. Metabolic steps that are feedback-inhibited by heme are shown with circled Hs. The symbols (H) and (O) indicate that oxidation and reduction steps occur. Abbreviations include: ALA, 5-aminole-vulinic acid; apoBphP, apo-bacteriophytochromes; apoCph, apocyanobacterial phytochromes; apoHP, apohemoproteins; apoLHCP, apo-light harvesting chlorophyll binding proteins; apoPHYs, apo-phytochromes; FDBRs, ferredoxindependent bilin reductases; GSA, glutamate1-semialdehyde; HO, heme oxygenase; tRNAglu, glutamate tRNA

4.1.1 Ferredoxin-dependent Heme Oxygenases Heme oxygenases (HO) from mammals were the first of this family of enzymes to be identified, [159]. Microsomal enzymes, mammalian heme oxygenases catalyze the oxygen-dependent conversion of heme to BV, CO, and Fe2+ products, [121]. The seven electrons required for this reaction are provided by microsomal NADPH-


cytochrome P450 reductases, [141, 160]. Although they catalyze the same reaction, plant HOs are soluble plastid-localized proteins that utilize soluble ferredoxin for reducing power, [115]. Despite their functional differences, the two families of HOs are structurally and evolutionarily related, [50, 163]. Ferredoxin-dependent HOs plays a pivotal role in the biosynthesis of phytobilins in all oxygenic photosynthetic organisms. In plants, BV is reduced by the enzyme phytochromobilin synthase to yield PB, the immediate precursor of the phytochrome chromophore of phytochromes that is critical for its photoactivity, [166]. In cyanobacteria and algae, BV is converted into the phytobilin precursors of the chromophores of the light-harvesting phycobiliproteins for photosynthesis, [5]. These include both PEB and PCB, the latter of which also serves as the immediate precursor of the cyanobacterial phytochromes, Cph1 and Cph2. The first HO identified from a photosynthetic organisms was found in the red alga Cyanidium caldarium by Troxler and colleagues, [168]. These researchers demonstrated that the phycobiliprotein chromophore precursor PCB was derived from heme in a manner similar to that in mammalian systems. More recently, Beale and Cornejo partially purified a Cyanidium HO and reported data establishing its ferredoxin dependence, [7, 31]. Cyanobacterial HO activity was also described by these researchers, [32], who later succeeded to clone and functionally express a ferredoxin-dependent HO from the cyanobacterium Synechocystis sp. PCC 6803, [33]. Other than in organello studies from a variety of plant species, [163], there has been little published on the biochemistry of heme oxygenases isolated from plant sources. The cloning of HY1 has however lead to the identification of three other heme oxygenase-related genes in the Arabidopsis genome, [34, 115]. From genetic and biochemical studies on recombinant proteins, it is clear that HY1 encodes a soluble, ferredoxin-dependent HO that is targeted to plastids, [115]. HO genes have been found in many other plant species, and like HY1, lesions in these genes also lead to phytochrome deficiencies, [161]. Aside from animals, plants, and cyanobacteria, novel HO genes have been identified in a wide variety of bacterial species, [181]. A soluble HO from the bacterial pathogen Neisseria meningitides encoded by the hemO locus was shown to convert heme to ferric-BV in the presence of ascorbate or NADPH-cytochrome P450 reductase, [191]. Biochemical studies of a protein encoded by the HO-related hmuO locus from Corynebacterium diphtheriae suggests that it also encodes an HO enzyme similar to human HO, [26, 144, 182]. Physiological studies on HmuO implicate a vital role in iron acquisition that is essential for survival and pathogenicity of this microorganism, [144]. The HO-related PigA protein from the opportunistic human pathogen Pseudomonas aeruginosa was recently shown to convert heme to the IXβ/γ isomers of biliverdin, [132]. Perhaps the most interesting is the BphO family of HOs, whose genes are invariably found linked to BphPs in a wide variety of bacterial genomes, [8]. Although there is no direct biochemical evidence that shows that BphO is a heme oxygenase, the observation that E. coli cells overexpressing the bphOP operon exhibit green color strongly suggesting that BV is produced by this HO-related protein, [8]. These observations implicate a role for the bacteriophytochrome family in iron homeostasis, [111].

16 The Phytochromes

4.1.2 Ferredoxin-dependent Phytobilin Synthases (Bilin Reductases) In oxygenic photosynthetic organisms exclusively, BV is converted to phytobilins by a family of ferredoxin-dependent bilin reductases (FDBRs) as illustrated in Figure 7. HY2, the founding member of this family, was shown to encode the enzyme PB synthase that mediates the conversion of BV to 3Z-PB, [85]. Unlike the NADPH-dependent biliverdin reductase (BVR) found in mammals, [100] and its cyanobacterial ortholog BvdR, [142], biliverdin reduction by PB synthase appears to proceed via sequential one electron steps involving radical intermediates, [50]. PB synthase isolated from etiolated oat seedlings as well as recombinant HY2 from Arabidopsis both catalyze the ferredoxin-dependent two electron reduction of BV to 3Z-PB, [85, 106]. Formally a phytochromobilin:ferredoxin oxidoreductase (E.C. 1.3.7.4.), PB synthase possesses a submicromolar affinity for its BV substrate and also exhibits a relatively high turnover rate constant of >100 min−1 , [106]. A single-copy gene in Ara-bidopsis, mutations in HY2-related genes found in other plant species all lead to phytochrome deficiencies, [161]. The paucity of HY2 cDNA clones in plant EST databases supports the observation that the HY2 protein is present in vanishingly low amounts in plant tissue, [106]. Together with its very high BV affinity, this observation may account for the high specific activity of PB

Fig. 7 Bilin metabolism in plants, cyanobacteria and mammals. Phytobilins from plants, crytophytes and cyanobacteria are all derived from BV which is metabolized by various members of the ferredoxin-

dependent bilin reductase family, that is HY2, PcyA, PcyB, PebA and PebB [49]). In mammals and cyanobacteria, BV is reduced to bilirubin (BR) by the NADPH-dependent enzymes BVR and BvdR, respectively.


synthase enabling low amounts of enzyme to produce sufficient PB precursor for holophytochrome synthesis. The precursor of the chromophores of green algal and cyanobacterial phytochromes has been shown to be PCB – a phytobilin that is two electrons more reduced than PB, [71, 187]. PCB is produced by the HY2-related enzyme PcyA, phycocyanobilin:ferredoxin oxidoreductase (E.C. 1.3.7.5.), [49]. Found in all phycobiliprotein-producing cyanobacteria as well as the oxygenic photobacterium Prochlorococcus marinus, PcyA genes encode the enzymes that catalyze the ferredoxin-dependent, four electron reduction of BV to a mixture of 3Z- and 3EPCB, [49]. Mechanistic studies on PcyA have established that the reduction of BV by PcyA proceeds via the two-electron reduced intermediate 181 ,182 -dihydrobiliverdin, rather than its isomer PB, [48]. The distinct BV reduction regiospecificity of PcyA was proposed to ensure that cyanobacteria do not produce PB because its misincorporation into phycobiliproteins might alter their spectra and/or stability, [48]. The observation that the green alga Mesotaenium caldariorum possess phytobilin synthase enzymes capable of the reductions of BV to PB and PB to PCB, [187] indicates that an alterative enzyme that we have named PcyB can accomplish one or both of these conversions in this organism, [50]. Although this enzyme has not yet been cloned, the recent demonstration that the PB chromophore of plant phytochromes can be substituted with PCB, and still yield a functional photoreceptor, raise many questions about chromophore selection during the evolution of higher plant phytochromes, [79]. 4.2 Apophytochrome Biosynthesis and Holophytochrome Assembly

Since phytochromes consist of two components, an apoprotein and a phytobilin pigment, phytochrome biosynthesis involves the convergence of two biosynthetic pathways which culminate in the assembly of functional holoprotein. Progress on the synthesis of the phytochrome apoprotein and its assembly with the phytobilin chromophore precursor is summarized below. 4.2.1 Apophytochrome Biosynthesis Although they are encoded by a small gene family of 3–5 members, [27], plant phytochromes can be classified into two groups based upon their light stability. Phytochromes encoded by the phyA gene family are responsible for the light-labile pool, while phyB-E genes encode the light-stable phytochromes, [54]. The pronounced light-lability of phyA proteins is due to two processes – light-dependent transcriptional repression of the phyA gene, [128] and light-dependent protein turnover (see Section 4.3). PhyA protein accumulates to very large levels in dark grown seedlings due to the elevated level of transcription and the pronounced stability of the phytochrome apoprotein. In this regard, the phyA promoter has been shown to be even stronger than the 35S promoter from the caulif lower mosaic virus when used to express heterologous proteins in dark-grown Arabidopsis plants, [152]. It is well established that the light-dependent repression of phyA gene expression is

18 The Phytochromes

mediated by phyA itself, [128]. The pattern of expression of the different phytochrome genes has been extensively studied in Arabidopsis, [59, 153], as has the accumulation of the phytochrome apoproteins, [66, 146]. Taken together, these studies indicate that the five phytochrome apoproteins have an overlapping distribution with phyA and phyB being the dominant species of phytochrome that respectively accumulate in dark-adapted and light-adapted plant tissue. While similar results have been reported for phytochromes in other f lowering plant species, [54, 62], little is known about the regulation of apophytochrome expression in lower plants, [174]. In this regard, it has been reported that phytochrome accumulation in green algae and mosses is light-regulated, although these phenomena may reflect differential protein turnover rather than differential transcription, [42, 112]. Owing to their very low abundance, even less is known about the accumulation of the Cph1 and Cph2 apoproteins in cyanobacteria other than the observation that Cph1 mRNA accumulation is light-regulated, [56]. The regulatory mechanism for this response is distinct from that of phyA in f lowering plants however, and little is presently known how this affects apoCph1 accumulation in cyanobacteria. 4.2.2 Holophytochrome Assembly Phytobilins are covalently bound to both phytochromes and phycobiliproteins via thioether linkages. Consequently, the process of holoprotein assembly has been the subject of extensive analysis, [107]. The assembly of plant phytochromes is autocatalytic and proceeds in two steps – noncovalent phytobilin binding to the apophytochrome followed by thioether linkage formation, [89, 96, 97]. Recent studies with the cyanobacterial phytochrome Cph1 have corroborated this autocatalytic mechanism, [11]. The take-home lesson of these studies is that an A-ring ethylidene in the phytobilin precursor is required for covalent attachment – a substituent that enables Michael addition of the conserved cysteine thiol residue of apophytochromes to this electrophilic double bond. BVs and bilirubins, which lack the A-ring ethylidene moiety, do not form covalent linkages with apophytochrome, [96]. BVs can however non-covalently interact with apophytochrome and act as reversible competitive inhibitors for phytochrome assembly, [97]. These results contrast with the observation that BV covalently binds to bacteriophytochromes, [8, 91, 93, 94]. The autocatalytic mechanism of phytochrome assembly also contrasts with phycobiliprotein assembly which require lyase enzymes to mediate attachment of each phytobilin, [50, 143]. In addition to the ethylidene moiety, both propionic acid moieties have been shown to be necessary for holophytochrome assembly, [9]. The requirement of a C10 methine bridge was based on the observation that rubin analogs of phytobilins fail to form covalent adducts with recombinant apophytochromes, [164]. While most of the published reports on phytochrome assembly have utilized the 3E-phytobilin isomers, the observation that 3Z-phytobilin isomers are also capable of functional assembly with apophytochrome suggests that the 3Z to 3E isomerization is not critical for holophytochrome assembly, [165]. As was discussed in Section 2.1.2, 3Z-PB is the major product of PB synthase, hence its isomerization may not be necessary in vivo. Other studies have shown that bilins with modified substituents in


the D-ring including PCB, PEB and isoPB can assemble with apophytochromes. However, the spectroscopic properties of the resulting phytobilin apophytochrome adducts are significantly altered, [41, 96, 98]. PCB substitution has been shown to produce a functional, albeit blue-shifted plant phytochrome photoreceptor, [79], while PEB adducts of apophytochromes, a.k.a. phytofluors, are intensely f luorescent, [116]. More recent studies using synthetic bilins indicate that, aside from the ethylidene moiety required for thioether linkage formation, [89, 96, 97], modifications in both A- and D-rings can be tolerated, [60, 61]. This work suggests that the phytobilin binding site on apophytochrome can accommodate structural variations, and that it should be possible to alter the spectral properties and photoregulatory function of phytochromes through engineering of new pathways of phytobilin biosynthesis in plants. While the phytobilin specificity for holophytochrome assembly has been actively addressed, less is known regarding catalytic residues on apophytochrome that specify phytobilin attachment. The only absolute requirement for thioether linkage formation appears to be the conserved cysteine itself (i.e. Cys321 in the case of PHYA3), [9, 12]. Deletion analysis of recombinant phytochromes reveals that neither the N-terminal 68 amino acids nor the entire C-terminal regulatory domains are required for assembly, [35, 65]. Comparative domain analysis of phy, cph1 and cph2 families indicate that P1 and P2 domains are dispensable for phytobilin attachment, and in the same study, it was shown the P3 GAF domain delimits the phytobilin lyase domain of phytochromes, [186]. A site-directed phytochrome mutant on the histidine residue adjacent to the conserved cysteine in the P3 GAF domain almost completely abolished phytobilin attachment, demonstrating a potential catalytic role for this residue, [9, 135]. Replacement of this histidine with asparagine did not prevent phytobilin attachment however, indicating that this residue cannot serve as a proton donor/acceptor for catalysis, [155]. Mutagenesis of a conserved glutamate residue (E189) in the P3 GAF domain of Cph1 proved inhibitory to phytobilin attachment implicating its involvement in assembly – possibly as a proton donating residue to the phytobilin, [186]. The detailed knowledge of the residues that participate in both noncovalent and covalent binding of phytobilins to apophytochromes is expected to provide insight into the molecular mechanism of holophytochrome assembly and its regulation, about which little is presently known. 4.3 Phytochrome Turnover

Aside from assembly, photoconversion and dark reversion, phytochrome localization and turnover are two important mechanisms that regulate phytochrome activity. The movement of phytochrome from the cytoplasm to the nucleus is controlled by the photoconversion between Pr and Pfr forms – a process that has been strongly linked to phytochrome signal transduction. The following discussion focusses on the processes that regulate phytochrome stability. Plant phytochromes have been classified into two groups, that is light-labile and light-stable. Accumulating to high levels in plants grown for prolonged periods in

20 The Phytochromes

darkness, the light-labile phyA class of photoreceptors function to regulate gene expression and seed germination under very low light fluences and far-red light enriched canopy environments, [21]. Such conditions are encountered for seeds that germinate and develop underground. When dark-grown seedlings are exposed to light, phyA is rapidly degraded, [117]. The rate of phyA degradation increases 100-fold upon light exposure – from a half-life of over 100 h to less than 1 h after photoconversion to Pfr, [131]. Because of this phenomenon, light-stable phyBEs predominate in light-grown plants, [146]. Light-dependent phyA turnover is preceeded by the formation of sequestered areas of phytochromes or SAPs in the cytoplasm, [99, 156]. Since SAPs are only formed under R and are also detected in phyA preparations in vitro, the hypothesis that SAPs represent self-aggregated Pfr has been proposed, [68, 69]. This proposal has received support from more recent studies showing that phyA must be translocated to the nucleus to initiate signal transduction. In this model, phyA that remains sequestered in the cytosol does not participate in signaling and is ultimately degraded. In the late 1980s, the detection of ubiquitin-phyA conjugates (Ub-P) following light treatment provided evidence that phyA degradation utilizes the ubiquitin/26S pro-teasome pathway [for review see, [29]. Ub-P conjugates appeared rapidly, that is within 5 min, following exposure to red light and disappeared with kinetics similar to those for the loss of Pfr; results that indicated that phyA is degraded in the Pfr form, [25, 74]. This conclusion was more recently corroborated by the pronounced light-stability of phyA in dark-grown phytobilin-deficient plants where phyA accumulates as its apoprotein, [161]. The role of phyA ubiquitination to both signal transduction and turnover has been the subject of great interest. Through deletion analysis of phyA and expression in transgenic plants, several regions important for Pfr degradation have been identified [see review of, [29]. Key regions critical to the light-dependent turnover have been mapped to both N- and C-terminal domains of phytochrome. However, identification of key lysine residues that participate in the light-dependent ubiquitination of phyA remains an elusive challenge, [28]. The recent discovery of cytoplasm-to-nuclear translocation raises the question of in which plant cellular compartment phyA ubiquitination and turnover occurs. This issue will undoubtedly consume countless hours of research in the laboratory. The inability to identify a phyA-specific E3 ubiquitin ligase by both biochemical analysis and genetic screens suggests that a novel mechanism(s) for phyA turnover may be responsible.

5 Molecular Mechanism of Phytochrome Signaling: Future Perspective

Phytochrome signaling requires both translocation to the nucleus and the maintenance of the Pfr form within the nucleus for a sufficient length of time to commit the light signal irreversibly. Based on the protein kinase activities of plant and cyanobacterial phytochromes, it is likely that protein phosphorylation plays a key role in the transmission of the light signal and/or in its regulation. The poten-

5 Molecular Mechanism of Phytochrome Signaling: Future Perspective 21

tial role of phytochromes to regulate tetrapyrrole metabolism is another emerging area of phytochrome signaling studies. Both topics are the subject of the following discussion. 5.1 Regulation of Protein-Protein Interactions by Phosphorylation

Previous studies have shown that the protein kinase activities of both Cph1 and eu-karyotic phytochromes are bilin- and light-regulated, [189, 190]. Phytobilin binding to apoCph1 stimulates its histidine kinase activity, which is consistent with the observation that holoCph1 is mostly dimeric while apoCph1 is a monomer, [92, 124]. Phytobilins are therefore apoCph1 ligands which regulate its trans(auto)phosphorylation – a mechanism that has been well described for other bacterial two-component histidine kinases, [157]. In contrast, the autophosphorylation activity of affinity-purified, recombinant oat phyA is inhibited by bilin attachment, indicating that the ser/thr kinase activity of a eukaryotic phytochrome is also bilin-regulated, [189]. As shown in Figure 8, the polarities of bilin- and light-regulation for plant and Cph1 phytochromes are opposite one another, with Pfr being more active than Pr for plant phytochromes and vice versa for Cph1. These results strongly implicate regulation of the monomer-dimer equilibrium by bilin and light for Cph1, while a different (allosteric) mechanism must be invoked for plant phytochromes which are are obligate dimers. The role of phytochrome phosphorylation activities in light signaling in plants is unknown. However, the observed bilin-stimulated and light-inhibited phosphotransferase activities of Cph1 implicate the regulation of the ATP/ADP-binding affinities, subunit-subunit dissociation constants, and/or ATP-to-histidine and phosphohistidine-to-aspartate equilibrium/rate constants. Little is presently known about the inf luence of bound ATP (and/or ADP) and of the histidine phosphorylation state on the thermody-

Fig. 8 Models for prokaryotic (Cph1) and eukaryotic (phyA-E) phytochrome signal transmission. Bilin and light input signals are perceived by the photosensory domains leading to altered phosphotransfer activities of the regulatory domains. Biochemical studies have shown that Cph1 and eukaryotic phytochromes have opposite responses.

The linear tetrapyrrole (bilin) chromophore is shown associated with the P3 and P4 domains as indicated in Figure 6 Potential downstream targets are indicated in red. The output signal(s) of the phytochrome signal transduction pathways are presently unknown.

22 The Phytochromes

namics and kinetics of the various protein-protein interactions. For higher plant phytochromes, it is reasonable that phytochrome-substrate interactions will be inf luenced by ATP/ADP binding as well as by the phosphorylation status of phytochrome itself and its substrate(s). We envisage that bilin, light, and ATP work together to regulate phytochrome–protein interactions which inf luence its subcellular localization, degradation, dark reversion, gene expression, and lead ultimately to changes in plant growth and development. 5.2 Regulation of Tetrapyrrole Metabolism

The evidence that phytochromes are regulators of tetrapyrrole metabolism in plants has a rich history, [110]. In addition to key enzymes of the chlorophyll pathway that are transcriptionally regulated by phytochrome, such as GluTR, [105] and protochlorophyllide oxidoreductase A (PORA), [113], phytochrome performs a metabolic regulatory role in the tetrapyrrole biosynthetic pathway that likely ref lects a more ancestral role, [111]. Phytochromes coordinate the expression of nuclear genes involved in the biogenesis of the photosynthetic apparatus, most notably the light harvesting chlorophyll a/b binding protein family, with the production of chlorophyll pigments. This is an extremely important function because misregulated chlorophyll synthesis can lead to production of toxic oxygen species via photosensitization, [30]. Indeed, the combination of light, oxygen, and photosensitizing pigments is a deadly recipe that plants must avoid at all costs. In flowering plants, chlorophyll synthesis does not occur in darkness since their protochlorophyllide oxidoreductases are all light-dependent, [183]. This situation contrasts with cryptophytes, that is mosses, ferns and algae, which possess a light-independent protochlorophyllide oxidoreductase system enabling them to synthesize chlorophyll in darkness, [53, 169]. During prolonged dark periods, angiosperm seedlings therefore accumulate protochlorophyllide (Pchlide) which becomes bound to PORA. Upon light exposure, Pchlide bound to the ternary NADPH-PORA-Pchlide complex is photochemically converted to chlorophyllide (Chlide) which is subsequently metabolized to chlorophylls a and b (see Figure 6). The dark accumulation of Pchlide serves to prime chlorophyll biosynthesis and assembly of the photosynthetic apparatus when light becomes available; a process that is of adaptive significance to angiosperm seedlings developing underground, [21]. This program of etiolation can proceed as long as food reserves in the seed last or light becomes available. Highly expressed in dark-grown seedlings, the PORA gene is rapidly down-regulated upon light exposure; a response that has been shown to be mediated by phyA, [113]. The similar pattern of PHYA expression, that is elevated expression in darkness and rapid down regulation in light, suggests that the coordinate expression of PORA and PHYA contribute to this important adaptive process. The metabolic consequence of elevated PHYA expression in dark-grown plants is elevated synthesis of Pchlide which in part is due to removal of heme-derived

5 Molecular Mechanism of Phytochrome Signaling: Future Perspective 23

phytobilins. In this regard, it is well established that heme is a potent feedback inhibitor of ALA synthesis – and in plants, the major target for this inhibition is GluTR [see Figure 6; [30, 122]. For this reason, PORA expression must also be elevated to ensure that all of the Pchlide produced under these conditions is assembled into a PORA-protein complex. The consequence of insufficient PORA expression and the resulting accumulation of free Pchlide, seen in porA mutants and seedlings exposed to prolonged FR in which PORA gene expression is downregulated via the action of phyA, is severe photooxidative damage upon transfer to light, [133]. These results show that PORA and PHYA expression must be balanced during periods of prolonged etiolation, both to prime the system for rapid synthesis/assembly of the photosynthetic apparatus and to avoid photooxidative damage. Beside contributing to the synthesis of light-modulated regulators of gene expression, the phytobilin-binding properties of apophytochromes, as well as the BV-binding properties of apobacteriophytochromes, represent one of many strategies found in nature to modulate heme levels and thereby to modulate tetrapyrrole synthesis. In animals and alpha protoeobacteria, ALA synthesis is also regulated by heme. However in these organisms, ALAS is the target of heme feedback inhibition. The regulation of heme is critical for all aerobic organisms since heme also has pro-oxidant properties. For this reason, nature has invented a wide variety of mechanisms to cope with heme accumulation aside from this feedback regulation. Heme oxygenases play a key role in this regulation. However the mammalian and plant enzymes are inefficient with poor catalytic turnover, unless their BV products are removed. To accomplish heme removal, plants and cyanobacteria utilize the FD-BR family, apophytochromes and apophycobiliproteins, while mammals, fungi, pathogenic and nonpathogenic bacteria have evolved NADPH-dependent BV reductase (BVR-A), bacteriophytochromes, and unique heme oxygenase enzyme systems with distinct heme cleavage specificities [see Figure 6; [50]]. In all of these organisms, removal of heme has the same effect to up-regulate new tetrapyrrole synthesis through metabolite-dependent derepression of ALA synthesis. As for the fate of these newly synthesized tetrapyrroles, this varies from organism to organism – impinging on such processes as photosynthesis, nitrogen fixation, nitrate assimilation, oxygen-detoxification, synthesis/secretion of defense compounds, hormone synthesis/degradation, among many others. In addition to its well-publicized genetic regulatory function, phytochrome’s role to regulate tetrapyrrole metabolism in plants and cyanobacteria is expected to receive increased experimental scrutiny in the years to come. Acknowledgements

We gratefully acknowledge grant support from the National Institutes of Health (RO1 GM068552–01) and the United States Department of Agriculture (AMD0103397) and to members of the Lagarias laboratory for critical reading of this manuscript.

24 The Phytochromes

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143 SCHLUCHTER, W. M. and A.N. GLAZER. 1999. In The Photosynthetic Prokaryotes. (ed. G.A. PESCHEK, W. LOFFELHARDT, and G. SCHMETTERER), pp. 83–95. Kluwer Academic/Plenum Press, New York. 144 SCHMITT, M. P., 1997. J. Bacteriol. 179, 838–845. 145 SCHNEIDER-POETSCH, H. A., B. BRAUN, S. MARX, and A. SCHAUMBURG. 1991. FEBS Lett. 281, 245–249. 146 SHARROCK, R. A. and T. CLACK. 2002. Plant Physiol. 130, 442–456. 147 SHINOMURA, T., A. NAGATANI, H. HANZAWA, M. KUBOTA, M. WATANABE, and M. FURUYA. 1996. Proc. Natl. Acad. Sci. USA 93, 8129–8133. 148 SHINOMURA, T., K. UCHIDA, and M. FURUYA. 2000. Plant Physiol. 122, 147–156. 149 SIEGELMAN, H. W., B. C. TURNER, and S.B. HENDRICKS. 1966. Plant Physiol. 41, 1289–1292. 150 SINESHCHEKOV, V., J. HUGHES, E. HARTMANN, and T. LAMPARTER. 1998. Photochem. Photobiol. 67, 263–267. 151 SINESHCHEKOV, V., L. KOPPEL, B. ESTEBAN, J. HUGHES, and T. LAMPARTER. 2002. J. Photochem. Photobiol. B. 67, 39–50. 152 SOMERS, D. E. and P.H. QUAIL. 1995a. Plant Physiol. 107, 523–534. 153 SOMERS, D. E. and P.H. QUAIL. 1995b. Plant J. 7, 413–427. 154 SONG, P. S., 1999. J. Biochem. Mol. Biol. 32, 215–225. 155 SONG, P. S., M. H. PARK, and M. FURUYA. 1997. Plant Cell Environ. 20, 707–712. 156 SPETH, V., V. OTTO, and E. SCHA¨ FER. 1987. Planta 171, 332–338. 157 STOCK, A. M., V. L. ROBINSON, and P.N. GOUDREAU. 2000. Ann. Rev. Biochem. 69, 183–215. 158 TAYLOR, B. L. and I.B. ZHULIN. 1999. Microbiol. Mol. Biol. Rev. 63, 479–506. 159 TENHUNEN, R., H. S. MARVER, and R. SCHMID. 1968. Proc. Natl. Acad. Sci. USA 61, 748–755. 160 TENHUNEN, R., H. S. MARVER, and R. SCHMID. 1969. J. Biol. Chem. 244, 6388–6393. 161 TERRY, M. J., 1997. Plant Cell Environ. 20, 740–745.

162 TERRY, M. J. and J.C. LAGARIAS. 1991. J. Biol. Chem. 266, 22215–22221. 163 TERRY, M. J., P. J. LINLEY, and T. KOHCHI. 2002. Biochem. Soc. Trans. 30, 604–609. 164 TERRY, M. J., M. D. MAINES, and J.C. LAGARIAS. 1993a. J. Biol. Chem. 268, 26099–26106. 165 TERRY, M. J., M. T. MCDOWELL, and J.C. LAGARIAS. 1995. J. Biol. Chem. 270, 11111–11118. 166 TERRY, M. J., J. A. WAHLEITHNER, and J.C. LAGARIAS. 1993b. Arch. Biochem. Biophys. 306, 1–15. 167 THU¨ MMLER, F., M. DUFNER, P. KREISL, and P. DITTRICH. 1992. Plant Mol. Biol. 20, 1003–1017. 168 TROXLER, R. F., A. S. BROWN, and S.B. BROWN. 1979. J. Biol. Chem. 254, 3411–3418. 169 VAVILIN, D. V. and W.F. VERMAAS. 2002. Physiol. Plant. 115, 9–24. 170 VIERSTRA, R. D., 1993. Plant Physiol. 103, 679–684. 171 VIERSTRA, R. D. and S.J. DAVIS. 2000. Sem. Cell Dev. Biol. 11, 511–521. 172 VIERSTRA, R. D. and P.H. QUAIL. 1982. Planta 156, 158–165. 173 VIERSTRA, R. D. and P.H. QUAIL. 1986. In Photomorphogenesis in Plants (ed. R.E. KENDRICK and G.H.M. KRONENBERG), pp. 35–60. Martinus Nijhoff, Dordrecht. 174 WADA, M., T. KANEGAE, K. NOZUE, and S. FUKUDA. 1997. Plant Cell Environ. 20, 685–690. 175 WAGNER, D., C. D. FAIRCHILD, R. M. KUHN, and P.H. QUAIL. 1996. Proc. Natl. Acad. Sci. USA 93, 4011–4015. 176 WAHLEITHNER, J. A., L. LI, and J.C. LAGARIAS. 1991. Proc. Natl. Acad. Sci. USA 88, 10387–10391. 177 WELLER, J. P. and A. GOSSAUER. 1980. Chem. Ber. 113, 1603–1611. 178 WENDLER, J., A. R. HOLZWARTH, S. E. BRASLAVSKY, and K. SCHAFFNER. 1984. Biochim. Biophys. Acta 786, 213–221. 179 WHITELAM, G. C., S. PATEL, and P.F. DEVLIN. 1998. Phil. Trans. Royal Soc. London Series B, Biological Sciences 353, 1445–1453. 180 WILDE, A., B. FIEDLER, and T. BÖRNER. 2002. Mol. Microbiol. 44, 981–988. 181 WILKS, A., 2002. Antioxid Redox Signal 4, 603–614.

References 182 WILKS, A. and M.P. SCHMITT. 1998. J. Biol. Chem. 273, 837–841. 183 WILLOWS, R. D., 2003. Nat Prod Rep 20, 327–341. 184 WONG, Y. S. and J.C. LAGARIAS. 1989. Proc. Natl. Acad. Sci. USA 86, 3469–3473. 185 WONG, Y. S., R. W. MCMICHAEL, and J.C. LAGARIAS. 1989. Plant Physiol. 91, 709–718. 186 WU, S. H. and J.C. LAGARIAS. 2000. Biochemistry 39, 13487–13495. 187 WU, S. H., M. T. MCDOWELL, and J.C.

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LAGARIAS. 1997. J. Biol. Chem. 272, 25700–25705. YAMAMOTO, K. T., 1990. Botan Mag 103, 469–491. YEH, K. C. and J.C. LAGARIAS. 1998. Proc. Natl. Acad. Sci. USA 95, 13976–13981. YEH, K. C., S. H. WU, J. T. MURPHY, and J.C. LAGARIAS. 1997. Science 277, 1505–1508. ZHU, W. M., A. WILKS, and I. STOJILJKOVIC. 2000. J. Bacteriol. 182, 6783–6790

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The Mammalian Rod Cell Photoreceptor Rhodopsin Najmoutin G. Abdulaev, and Kevin D. Ridge University of Maryland Biotechnology Institute, Rockville, USA


1 Introduction

Visual phototransduction is the process of transforming light energy absorbed by the specialized retinal cells (rods and cones) into an electrical response. The amplitude and duration of the response depends on light intensity and the type of cell. Rods (the dim-light photoreceptors) are more sensitive to light than cones (the bright-light photoreceptors), although the latter demonstrate a faster light response [1]. In rods (Figure 1A), a characteristic two-phase electrical response reflects a multistep, highly amplified series of biochemical reactions expressing activation and recovery phases of signaling. This elegant biochemical machinery is designed to exchange information between the rod outer segment (ROS) plasma membrane (PM) harboring cGMP-regulated ion channels and the light-sensitive pigment, rhodopsin, which is abundantly present in the ROS disk membranes (DM). Rhodopsin is the first link in the chain of biochemical reactions involved in visual phototransduction (Figure 1B). It is also a member of the large family of heptahelical G-protein coupled receptors (GPCRs). In this family, rhodopsin appears to be unique in that it displays very low noise levels and the rapid activation and inactivation kinetics necessary for high sensitivity and fast image processing. These requirements are realized by adopting 11-cis retinal as a chromophore sensitive to visible light. Of the ∼1000 GPCRs identified, bovine rhodopsin is the only receptor for which a three-dimensional structure is available [2–4]. The crystal structure has confirmed a wealth of biochemical and biophysical data on the dark-state of rhodopsin and provided a template for interpreting phenomenological data obtained by affinity labeling and site-directed muta-genesis. In addition, the availability of the rhodopsin structure has provided a useful model for understanding the structure and function of other members of the GPCR family. In this Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754


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2 The Mammalian Rod Cell Photoreceptor Rhodopsin

Fig. 1 Mammalian visual phototransduction. a) Schematic of the highly differentiated rod cell with the outer segment (OS) and inner segment (IS) indicated. The OS is comprised of numerous stacked disks that contain the major phototransduction components. The IS contains the biosynthetic machinery of the cell. b) Signal propagation by light-activated rhodopsin. The major protein components of the disk membrane (DM) that lead to hyperpolarization of the rod cell include rhodopsin (R), the retinal G-protein transducin (Gt ) "$( heterotrimer, light-activated rhodopsin (R* ), the cGMP phosphodiesterase (PDE6) "-, $-, (-, and F-subunits, guanylate cyclase (GC), and gunaylate cyclase activating protein (GCAP). The major protein components of the plasma membrane (PM) that lead to hyperpolarization of the rod cell include the cGMP-gated channel "- and $-subunits

and the Na+ /Ca2+ , K+ exchanger. Also indicated are guanylate kinase (GK) and nucleoside diphosphate kinase (NDPK), which are involved in guanine nucleotide metabolism, and calmodulin, a major Ca2+ binding protein in the ROS. c) Inactivation of light-activated rhodopsin. The major protein components of the DM that lead to the inactivation of R* include the Ca2+ -free and Ca2+ -bound forms of myristoylated recoverin (Rec), rhodopsin kinase (RK), arrestin (Arr), and protein phosphatase 2A (PP2A). In b) and c), R, R* , and opsin (O) are shown as distinctly shaded dimers in order to emphasize their proposed higher order organization. While atomic force microscopy evidence exists for the oligomeric state of murine rhodopsin (R1 R2 ) and opsin (O1 O2 ) in the DM [68, 69], the existence of R* dimers interacting with Gt , Arr, and RK remains an open question.

chapter, various structural and functional aspects of mammalian rod rhodopsin are highlighted. While this by no means represents a comprehensive molecular description of rhodopsin, it is anticipated that the reader will gain a new sense of understanding and appreciation for this fascinating photosensory receptor. 2 Rhodopsin and Mammalian Visual Phototransduction

The process of mammalian visual phototransduction can be divided into three distinct pathways. Only the first two pathways involve rhodopsin: signal propagation


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by light-activated rhodopsin (metarhodopsin II or R* ) and inactivation of R* . The third pathway, inactivation of the catalytic subunit (α-subunit) of the heterotrimeric retinal G-protein, transducin (Gt ), continues to attract much attention and those interested in a perspective on this process are referred to some recent reviews on the subject [5–7].

2.1 Signal Amplification by Light-activated Rhodopsin

Light activation of rhodopsin (R) in the DM to produce R* catalytically promotes the exchange of GDP for GTP in hundreds of Gt molecules to produce the GTPbound form of Gtα (Figure 1B). Gtα -GTP interacts with the γ -subunit of the cGMP phosphodiesterase (PDE6 in rods), removing an inhibitory constraint from the catalytic α or β subunits that results in cGMP hydrolysis. The fast depletion of cGMP results in the closure of cGMP-gated channels in the PM. A drastic reduction in the circulating “dark” current, due to a blockage of Na+ and Ca2+ entry into the rod and Ca2+ depletion of the ROS as a result of continuous function of the PM Na+ , Ca2+ , K+ exchanger, activates the Ca2+ -binding guanylate cyclase activating protein (GCAP). The low intracellular Ca2+ concentration promotes activation of guanylate cyclase (GC) by Ca2+ -free GCAP, catalyzing accelerated synthesis of cGMP from GTP supplied by the guanine nucleotide cycle. The latter is comprised of two distinct nucleotide-binding enzymes, guanylate kinase (GK) and nucleoside diphosphate kinase (NDPK). The release of bound Ca2+ from calmodulin (CaM) leads to its dissociation from the cGMP-gated channel conferring a lower affinity for cGMP. Thus, light-activation of rhodopsin to produce R* leads to changes in the net production of cGMP by perturbing two opposing catalytic activities: degradation of cGMP by PDE6 and synthesis of cGMP by GC. Since cGMP levels govern how many channels are opened, the signal is transformed from a physical stimulus (light) to a biochemical chain of reactions that culminates in the hyperpolarization of the rod.

2.2 Inactivation of Light-activated Rhodopsin

In the dark, when the concentration of intracellular Ca2+ is high, myristoylated recoverin (Rec) is in the Ca2+ -bound state and forms a complex with rhodopsin kinase (RK) at the DM, preventing phosphorylation of R* (Figure 1C). When the Ca2+ concentration in the ROS drops as a result of the light response, Rec releases its bound Ca2+ and dissociates from RK. The interaction between R* and RK leads to rapid phosphorylation of R* . The subsequent binding of arrestin (Arr) blocks R* signaling via Gt . The release of all-trans retinal, accompanied by enzymatic dephosphorylation by protein phosphatase 2A (PP2A), prepares the apoprotein, opsin, for 11-cis retinal binding and restoration of its inactive ground state.

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3 Properties of Rhodopsin 3.1 Isolation of Rhodopsin

Bovine rhodopsin is the best characterized member of the large family of GPCRs. A virtual explosion of information on rhodopsin can be attributed to the high natural abundance of the pigment in the DM and the accessibility of bovine rhodopsin and its mutants in amounts amenable to extensive biochemical, biophysical, and structural studies. Polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate (SDS-PAGE) shows that more than 90% of the protein complement of the DM is accounted for by rhodopsin. In fact, highly purified rhodopsin in the DM can be prepared by sucrose density gradient centrifugation of ROS obtained by simple shaking of bovine retinae in the appropriate buffer and low speed centrifugation [8]. ROS preparations in hypotonic buffers are then subjected to Ficoll f lotation with subsequent high speed centrifugation [9]. Further purification of rhodopsin is typically based on its solubilization in detergents in combination with conventional chromatographic techniques. Although many different detergents effectively solubilize rhodopsin (e.g., digitonin, cetyl trimethylammonium bromide, nonylglucoside), dodecylmaltoside remains the detergent of choice due to very high rhodopsin stability in this media and the retention of many functional properties [10, 11]. Affinity chromatography based on separation with concanavalin A-Sepharose was an early purification method of choice [12]. However, rhodopsin samples prepared by this method are often contaminated with concanavalin A requiring the introduction of additional purification steps. Affinity chromatography based on matrices with immobilized monoclonal antibodies, such as rho 1D4-Sepharose [13], appears to provide an extraordinarily efficient method for the one-step purification of rhodopsin. Immunoaffinity chromatography is essential for purifying mutants of rhodopsin obtained by heterologous eukaryotic cell expression and also appears to be instrumental in separating unfolded or misfolded opsin from properly folded rhodopsin [14]. More recently, methods have been developed for the selective extraction of ROS rhodopsin from the DM using specific divalent cations (Zn2+ or Cd2+ ) in conjunction with alkylglucoside detergents [15]. This method allows for the effective purification of rhodopsin and was instrumental in obtaining diffraction-quality three dimensional crystals [16]. 3.2 Biochemical and Physicochemical Properties of Rhodopsin

Bovine rhodopsin is a single polypeptide of ∼40 kDa consisting of 348 amino acids. Its amino acid sequence is known from analyses of the opsin apoprotein and the corresponding DNA [17–19]. About 65% of the amino acid residues in bovine opsin are hydrophobic residues unevenly distributed along the polypeptide chain (Figure 2). This arrangement appears to be crucial for the membrane disposition of the protein.


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3 Properties of Rhodopsin

Fig. 2 A two-dimensional model for the organization of bovine rhodopsin in the DM. The lengths of the transmembrane (TM) helices (I–VII) are based on the crystal structure of rhodopsin [2, 3]. The cytoplasmic helix, H8, extending from TM VII is also shown. The small hexagons in the N-terminal region represent branched

oligosaccharide chains attached to Asn-2 and Asn-15 and the zigzag lines attached to Cys-322 and Cys-323 in the C-terminal region represent palmitoyl groups. The disulfide bridge between Cys-110 and Cys-187 is also shown. Key amino acid residues mentioned in the text are highlighted in black.

Rhodopsin purity is assessed by comparing the ratios of the absorption maxima at 280 nm (protein) and 500 nm (chromophore) in the dark. This ratio (A280 /A500 ) for purified rhodopsin varies between 1.60 and 1.65. Purified preparations of rhodopsin appear on SDS-PAGE as a single but somewhat broadened and diffuse band. This property is characteristic for many membrane proteins and may represent possible heterogeneity in several post-translational modifications. In bovine rhodopsin, diffusion of the band appears to be primarily due to heterogeneity in asparagine-linked (N-linked) glycosylation (see Section 3.3). Additional factors contributing to the apparent heterogeneity of bovine rhodopsin may include phosphorylated rhodopsin species produced by exposure to light or lack of appropriate dark adaptation before or during retina processing. Finally, defects in fatty acylation (palmitoylation) may also contribute to rhodopsin heterogeneity [20].

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3.3 Post-translational Modifications in Rhodopsin

As indicated in Section 3.2, the opsin polypeptide is subject to several post-translational modifications. First, the amino-terminal sequence, together with the acetylated amino-terminal amino acid, contains two consensus sequences allowing Nglycosylation at Asn-2 and Asn-15 [21]. Carbohydrate analysis of bovine rhodopsin shows that ∼70% of the oligosaccharides are accounted for by Man3 Glc-NAc3 , 10% by Man4 GlcNAc3 , and 20% by Man5 GlcNAc3 [21–23]. What is not clear, however, is the distribution of these carbohydrate components between the two N-linked glycosylation sites. A second type of post-translational modification is palmitoylation of two vicinal cysteine residues at positions 322 and 323 via a thioester linkage [24–26]. Third, two highly conserved cysteine residues, Cys-110 and Cys-187, are disulfide linked [27]. This disulfide bridge is conserved not only in opsins, but also in the entire rhodopsin family of GPCRs and appears to play an important role in protein stability [28–30]. Another type of rhodopsin post-translational modification is the light-dependent phosphorylation of several threonines and serines at the extreme carboxyl terminus of the protein by RK [31, 32]. Despite 30 years of intensive effort, questions as to which of the serine and threonine residues become phosphorylated under physiological conditions remain unanswered. Finally, covalent binding of 11-cis retinal to the ε-amino group of Lys-296 via a protonated aldimine bond, a Schiff base, is a crucial and unique post-translational modification of rhodopsin. 3.4 Membrane Topology of Rhodopsin and Functional Domains

Seven hydrophobic stretches of about 25–30 amino acid residues interrupted by relatively short hydrophilic sequences is the signature feature of rhodopsin (Figure 2), other GPCRs, and numerous retinylidene microbial photoreceptors as well. This odd number of transmembrane (TM) excursions positions the amino- and carboxyl-terminal ends of the protein on the opposing intradiscal (extracellular) and cytoplasmic surfaces, respectively. There is no obvious explanation for these so called “seven pillars of wisdom”. The notion that seven hydrophobic TM helices are required to create a suitable binding pocket for low molecular weight agonists or antagonists (like 11-cis retinal) appears to be consistent with much of the experimental data, but this can not be extended to a vast number of GPCRs which have smaller ligands. Remarkably, a consensus on the heptahelical arrangement of rhodopsin was reached in the late 1970s [33–36] with only limited available information on the primary structure, and subsequently confirmed by the crystal structure of bovine rhodopsin [2–4]. The structure defines three major functional domains within rhodopsin: transmembrane, intradiscal, and cytoplasmic. 3.4.1 Transmembrane Domain of Rhodopsin The TM domain of rhodopsin includes the seven predominantly α-helical segments, termed TM I–TM VII (Figure 2). TMI (residues 34–64) contains a highly conserved


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3 Properties of Rhodopsin

Asn-55 which forms hydrogen bonds with other residues (see Section 5.1). TM II, extending from residue 71 through 100, contains two adjacent glycine residues, Gly-89 and Gly-90. These two residues appear to be particularly susceptible to naturally occurring mutations in both mild and severe visual disorders [37–39]. TM II also contains a conserved Asp-83. TM III encompasses residues 106–139 and contains several important amino acid residues. First, the cytoplasmic end of TM III is capped by the highly conserved and functionally important triad: Glu-134, Arg-135, and Tyr-136 (the E(D)RY motif). The carboxyl group of Glu-113, which is located toward the center of this helix, serves as the dark-state counterion to the protonated retinylidene–Schiff base [40–42]. Next, highly conserved cysteine residue Cys-110 forms a disulfide bond with Cys-187 from the second intradiscal loop [27]. Interestingly, there are also two adjacent glycine residues, Gly-120 and Gly-121, which may contribute to the conformational f lexibility of this helix [43]. TM IV is the shortest of the TMs and extends from residues 150 to 172. Two proline residues, Pro-170 and Pro-171, are located at the cytoplasmic end of this helix. Cys-167, near the center of the helix, is an important component of the chromophore binding pocket [2, 3]. TM V incorporates residues 200–225, and with the exception of His-211 and Phe-212, appears not to be overburdened with functionally important amino acids. TM VI includes residues 244–276. Pro-267 of this helix appears to be essential for providing the necessary flexibility to support the dynamics of receptor activation [44]. Another important residue is Glu-247, which makes contact with Arg-135 of the E(D)RY motif of TM III (see Section 5.1). TM VII extends from residues 286 to 309 and contains several important side chains. First, this helix contains Lys-296, the site of retinal attachment. The ε-amino group of this amino acid reacts with the aldehyde group of 11-cis retinal to form a Schiff base. Second, the cytoplasmic end of this helix contains the conserved NPXXY motif, considered to be a part of a larger functional unit known as the NPXXY(X)5,6 F motif [45]. It has been shown that a portion of this motif becomes accessible to a monoclonal antibody upon light activation of rhodopsin [46]. Finally, Lys-296, Thr-297, Ser-298, and Ala-299 form a 3, 10 helix. All other helices in rhodopsin, both transmembrane and cytoplasmic, are regular α-helices [47].

3.4.2 Intradiscal Domain of Rhodopsin The intradiscal (extracellular) domain of rhodopsin (Figure 2), long suspected in playing an important role in the structural stability of the pigment, incorporates a 33-residue N-glycosylated amino-terminal region and three connecting loops between TM helices II–III (amino acid residues 101–105), IV–V (residues 173–198), and VI–VII (residues 277–285). As indicated in Sections 3.3 and 3.4.1, Cys-187 of the second intradiscal loop forms a disulfide bridge with Cys-110 on the extracellular end of TM helix III [27]. Only three of the seventeen positively charged surface residues in rhodopsin are found in the intradiscal domain, supporting the inside-positive rule characteristic for the arrangement of many polytopic membrane proteins [48]. The intradiscal domain appears to be highly structured and rarely tolerates even minor amino acid substitutions [49].

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3.4.3 Cytoplasmic Domain of Rhodopsin The cytoplasmic domain (Figure 2) includes the carboxyl-terminal tail of about 25 residues and loops connecting TM helices I–II (residues 65–70), III–IV (residues 140–149), and V–VI (residues 226–243). A fourth cytoplasmic loop (residues 311–323), now designated as H8 [2], extends from the cytoplasmic end of TM VII to the palmitoylated Cys-322 and Cys-323. In contrast to the highly structured and rigid intradiscal domain, the cytoplasmic domain appears to be designed to provide transient conformational changes that support activation of signaling and regulatory proteins [2, 50]. Valuable information on the cytoplasmic domain dark-state structure and the dynamics of its light-induced conformational changes became available through the painstaking work of the Khorana and Hubbell laboratories [51]. The use of several experimental approaches, including site-directed spin labeling (SDSL) in combination with electron paramagnetic resonance spectroscopy (EPR) [52], sulfhydryl reactivity [14], and disulfide crosslinking [53], on more than 100 single cysteine mutants and 40 double cysteine mutants of rhodopsin, has allowed important information to be collected on the solution state conformation and the dynamics of this domain, as well as the boundaries of the loops. The resulting borders appear to be in good agreement with the published crystal structure of rhodopsin [2–4], although SDSL/EPR studies suggest that the third cytoplasmic loop might fold up further into TM V and TM VI [54]. Quite interestingly, this latter finding is in agreement with a three-dimensional model generated from an alternative (trigonal) crystal form of rhodopsin [55].

4 Chromophore Binding Pocket and Photolysis of Rhodopsin

The high quantum yield of retinal isomerization in rhodopsin is believed to be supported by the constrained conformation of the chromophore within the protein. Numerous efforts have been directed towards understanding both the groundand excited-states of the rhodopsin chromophore. In the ground-state, the conjugated polyene chain is in the 11-cis configuration with the β-ionone ring primarily oriented to give a 6-s-cis conformer [56–58]. The chromophore binding pocket is predominantly comprised of hydrophobic amino acids with the β-ionone ring positioned between Phe-212 and Phe-261, whereas Trp-265 is closer to the center of the chromophore promoting the 11-cis form of the polyene chain [2, 3]. Although well protected from the aqueous environment, the chromophore binding pocket contains several hydrophilic side chains. Of keen interest is the carboxyl group of Glu-113, which serves as counterion to the protonated the retinylidene–Schiff base [40–42]. This interaction appears to be of key importance in stabilizing ground-state chromophore–protein interactions. Altogether, around 30 amino acid side chains participate in forming the chromophore binding pocket [2]. Light-induced isomerization of 11-cis retinal to its all-trans configuration is the primary step in vision [59]. This ultra-fast (femtosecond) photochemical process is followed by much slower events culminating in increased accessibility of the


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Fig. 3 Photointermediates of rhodopsin studies is shown. The absorbance maxima for photolysis. The photobleaching sequence of the various photointermediates and their aprhodopsin according to kinetic spectroscopy proximate half lives at 20◦ C are indicated.

chromophore binding pocket to the aqueous environment with subsequent hydrolysis of the retinylidene–Schiff base [60]. These two extreme events in the photolysis of rhodopsin are mediated by several thermally stable dark processes resulting in photoproducts with defined spectral, kinetic, and functional properties. Following the light-induced cis → trans isomerization of retinal, the earliest photointermediate observed to date is photorhodopsin [61]. However, this intermediate is highly unstable so little is known about its structure. The first structurally characterized stable photointermediate detected is bathorhodopsin (Figure 3), which thermally decays to the blue-shifted intermediate (BSI), followed by lumirhodopsin, metarhodopsin I, and metarhodopsin II [62, 63]. Of particular interest in this sequence is metarhodopsin II (or R* ), the active form of rhodopsin with bound all-trans retinal and capable of Gt activation. Although the precise mechanism of decay for this signaling photointermediate is not fully understood [64, 65], the two products typically detected are metarhodopsin III and opsin plus free all-trans retinal.

5 Structure of Rhodopsin 5.1 Crystal Structure of Rhodopsin

Having outlined the three topologically distinct functional domains of rhodopsin in two dimensions in Section 3.4, it is now of interest to examine the three-dimensional

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structure of rhodopsin in order to visualize how these domains are arranged to support tertiary contacts that ensure ground-state stability. While several aspects of this issue, including orientation of the TM helices, structural irregularities (e.g., tilts, kinks), chromophore–protein interactions, and the structural consequences of naturally occurring rhodopsin mutations are of particular importance, many of these have already been covered in the original report [2] as well as several recent reviews [43, 50, 81–83]. Here, we focus on some of the key structural features that confer on rhodopsin the unique ability to function as a finely tuned dim-light photoreceptor. The X-ray structure was determined from diffraction-quality crystals of detergent solubilized bovine ROS rhodopsin [2, 16]. A current refined (2.8 Å) model of rhodopsin [3] includes greater than 95% of the amino acid residues as well as post-translational modifications (Figure 4). One of the surprises from the crystal structure is a compact intradiscal arrangement, parts of which fold inwards to enclose the retinal moiety. This “lid” or “plug” over the chromophore binding pocket involves the second intradiscal loop and is comprised of an anti-parallel β-stranded

Fig. 4 Three-dimensional structure of bovine rhodopsin. Ribbon drawing of the rhodopsin structure showing the seven TM helices and the cytoplasmic "-helix, H8, in various colors with the connecting segments in gray. The N-linked oligosaccharide chains attached to Asn-2 and Asn-15 in the N-terminal region and the thioester linked palmitoyl groups attached to Cys322 and Cys-323 in the C-terminal region are shown in green. The 11-cis retinal chro-

mophore is shown in blue. The side chains of key amino acid residues mentioned in the text that are important for maintaining the ground-state structure of rhodopsin or are involved in the mechanism of rhodopsin activation are shown in purple and indicated with arrows. The image was generated using InsightII and PDB entry 1HZX (A chain) and is based on work from Teller et al., [3].


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5 Structure of Rhodopsin

motif. The disulfide forming Cys-187 residue of this loop anchors the latter to Cys-110, near the extracellular surface of TM III. The second part of this loop is proximal to the bound 11-cis retinal, whereas the first part of the loop lies on top of the second strand. As indicated in Section 3.3, the intradiscal domain contains two consensus N-glycosylation sites at Asn-2 and Asn-15. Neither of these sites appears to make structural contacts with the protein (Figure 4), although Asn-15 N-glycosylation has previously been implicated in rhodopsin function [66]. The cytoplasmic domain, in contrast to intradiscal domain, is largely disordered. One exception is H8, which lies almost perpendicular to the carboxyl-terminal end of TM VII (Figure 4). The carboxyl-terminal cytoplasmic tail and portions of the third cytoplasmic loop are poorly resolved in the structure [2–4], while, quite interestingly, the palmitoyl chains attached to Cys-322 and Cys-323 display resolvable density (Figure 4). The TM helices are the sites where a majority of the conserved amino acid residues are found in rhodopsin. For example, Asn-55 of TM I, Asn-78 and Asp-83 of TM II, and Asn-302, Pro-303, and Tyr-306 of TMVII (part of the NPXXY(X)5,6 F motif), are all highly conserved amino acids across the GPCR family that appear to form contacts important for maintaining dark-state stability. While Asn-55 in TM I is hydrogen bonded to Asp-83 in TM II and Asn-78 in TM II is hydrogen bonded to Trp-161 in TM IV (Figure 4), conserved residues from the NPXXY(X)5,6 F motif form a network of interactions that stabilize the cytoplasmic domain [2, 3, 45]. Another important conserved element is the E(D)RY motif at the cytoplasmic end of TM III (Figure 4). Glu-134 and Arg-135 in this motif form ionic interactions with Glu-247 at the cytoplasmic end of TM VI and provide additional dark-state stability to rhodopsin [45]. It should be noted, however, that the electrostatic interaction between Glu-113 and Lys-296, residues which are largely conserved only among opsins, is considered to be the main factor in maintaining the dark-state conformation of rhodopsin [37]. 5.2 Atomic Force Microscopy of Rhodopsin in the Disk Membrane

Oligomerization of GPCRs is now a widely accepted phenomenon. For most GPCRs, evidence for oligomerization is largely indirect and based on results from immuno-precipitation, chemical or disulfide crosslinking, size-exclusion chromatography, and f luorescence or bioluminescence resonance energy transfer studies [67]. In the case of rhodopsin, however, direct evidence for oligomerization has recently been obtained using atomic force microscopy (AFM) [68, 69]. The paracrystalline arrangement of rhodopsin dimers in the murine DM provides compelling evidence for how rhodopsin may be self-organizing, and provide a platform for signal propagation and/or quenching [70]. A current model for the rhodopsin dimer, termed the “IV–V model”, where the dimeric interface is formed between helices IV and V [68, 71], proposes intermolecular contacts (hydrogen bonds) formed by Asn-199 and Ser-202 of both monomers. Asn-199 also extends the hydrogen bond network to Glu-196, and the adjacent Glu-197 residue points its carbonyl oxygen

11


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atom toward Asn-199, thereby contributing to the association of the two rhodopsin monomers. Despite the impressive nature of the AFM and modeling studies, it should be noted that these findings are at variance with the results of earlier biophysical measurements [72, 73], which suggested that rhodopsin rapidly diffuses as a monomeric unit in the DM.

6 Activation Mechanism of Rhodopsin

Light activation of rhodopsin appears to rely on the disruption of dark-state contacts and replacing them with a new set of interactions. This process is directed towards creation of a highly specific and effective binding site for Gt . Since opsin exhibits negligible activity at neutral pH [74], the isomerized retinal chromophore is thought to play a major role in establishing these new interactions. In fact, studies with retinal analogs have shown that removal of the C19 methyl group from 11-cis retinal dramatically shifts the metarhodopsin I-metarhodopsin II equilibrium towards the former [75, 76]. An impairment of metarhodopsin II formation has also been observed when modifications are introduced in or around the β-ionone ring of retinal [77]. Thus, it appears that both the C19 methyl group and β-ionone ring of retinal make important steric contributions to rhodopsin activation. The exact mechanism of rhodopsin activation is not fully understood but the general view is that there is an isomerization-induced separation of the cytoplasmic ends of TM III and TM VI relative to each other [51, 78]. This is thought to be a consequence of charge separation and proton transfer from the retinylidene–Schiff base to the Glu-113 counterion, although a counterion switching mechanism at the metarhodopsin I stage involving Glu-181 has now been proposed [79]. This latter mechanism, however, is at variance with recent NMR results indicating no evidence for any major structural reorganization around the chromophore binding site up to the metarhodopsin I stage [80]. Nonetheless, local conformational changes arising from retinal isomerization appear to be propagated to the cytoplasmic surface causing greater structural changes coupled to the reprotonation and rearrangement around the E(D)RY motif as well as coordinated changes in and around the NPXXY(X)5,6 F motif [45]. A strong distortion in TM VI imposed by Pro-267, one of the most conserved amino acid residues in the family of rhodopsin-like GPCRs [50], is also considered a key element of the activation mechanism.

7 Conclusions

Much has been revealed about the structure and function of mammalian rod rhodopsin over the past several years. As we begin the 21st century, it is likely that further molecular insights into this visual photosensory receptor will emerge. Having now established the basic structural features important for maintaining


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References

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1

Microbial Rhodopsins John L. Spudich and Kwang-Hwan Jung Sogang University, Seoul, Korea


1 Introduction

The first 30 years of research on microbial rhodopsins concerned exclusively four proteins that share the cytoplasmic membrane of the halophilic archaeon Halobacterium salinarum, and a few very close homologs found in related haloarchaea. These four haloarchaeal types were the only microbial retinylidene proteins known prior to 1999: the light-driven ion pumps bacteriorhodopsin BR [1], and halorhodopsin (HR [2, 3]), and the phototaxis receptors sensory rhodopsin I (SRI [4]), and sensory rhodopsin II (SRII [5]). Studies of the haloarchaeal rhodopsins by the most incisive biophysical and biochemical tools available produced a wealth of information making them some of the best understood membrane-embedded proteins in terms of their structure-function relationships. Crystal structures of three [BR [6–8], HR [9], and SRII [10–12]] reveal a common seven-transmembrane α-helical structure with nearly identical helix positions in the membrane, despite their differing functions and identity in only ∼25% of their residues. The positions differ from those of visual pigments, as shown by the crystal structure of bovine rod rhodopsin [13], but their overall topologies are similar, namely the seven helices form an interior binding pocket in the hydrophobic core of the membrane for the retinal chromophore. In both the microbial and visual pigments, the retinal is attached by a protonated Schiff base linkage to a lysine in the middle of the seventh helix and retinal photoisomerization initiates their photochemical reactions. Starting in 1999, genome sequencing of cultivated microorganisms began to reveal the previously unsuspected presence of archaeal rhodopsin homologs in several organisms in the other two domains of life, namely Bacteria and Eukarya [14–16]. Further, in 2001, “environmental genomics” of populations of uncultivated microorganisms in ocean plankton showed the presence of a homolog in marine Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Microbial Rhodopsins

proteobacteria [hence given the name proteorhodopsin [17], which has swiftly expanded so far to ∼800 relatives identified in samples throughout the world’s oceans [18–23]. Microorganisms containing rhodopsin genes inhabit diverse environments including salt flats, soil, fresh water, surface and deep sea water, glacial sea habitats, and human and plant tissues as fungal pathogens. They comprise a broad phylogenetic range of microbial life, including haloarchaea, proteobacteria, cyanobacteria, fungi, dinof lagellates, and green algae. The conservation of residues, especially in the retinal-binding pocket, define a large phylogenetic class, called type 1 rhodopsins to distinguish them from the visual pigments and related retinylidene proteins in higher organisms (type 2 rhodopsins). Analysis of the sequences of the new type 1 rhodopsins, their heterologous expression and study, and in some cases study of the photosensory physiology of the organisms containing them, and spectroscopic analysis of environmental samples, have shown that the newfound pigments fulfill both ion-transport and sensory functions, the latter with a variety of signal-transduction mechanisms. The purpose of this article is to summarize what we have learned regarding the rapidly expanding group of retinylidene pigments comprising the microbial rhodopsin family.

2 Archaeal Rhodopsins

Many laboratories have characterized the four rhodopsins from H. salinarum with a battery of techniques because they provide model systems for the two fundamental functions of membranes: active transport and sensory signaling. Comprehensive reviews on mechanisms of BR [24], HR [25], and the SRs [26, 27] are available. Sixteen variants of BR, HR, SRI and SRII have been documented in related halophilic archaea, such as Natronomonas pharaonis and Haloarcula vallismortis (Table 1). Identification of members of the type-1 family has been based primarily on the conservation of residues in the retinal-binding pocket, which is known from the structures of haloarchaeal members. Atomic resolution structures, which exist for only a small number of membrane proteins, have been obtained from electron microscopy and X-ray crystallography of three of the archaeal rhodopsins: BR and HR from H. salinarum, and SRII from N. pharaonis (“NpSRII”). These proteins share a nearly identical positioning of seven transmembrane helices forming an interior pocket for the chromophore, all-trans retinal. The retinal binding pocket is comprised of residues from each of the seven helices, and it is the conservation of these residues that provides the most definitive identification of archaeal rhodopsin homologs in other organisms. Conservation outside of the pocket is sparse (Figure 1). Even between members of the archaeal branch, conservation outside the pocket is limited. For example, the phototaxis receptor NpSRII is only 27% identical to BR in amino acid sequence, and exhibits typically ∼40% identity with other archaeal sensory rhodopsins; all 4 archaeal rhodopsins exhibit ∼80% identity in the 22 residues that the crystal structures show form the retinal binding pockets in BR, HR, and

2 Archaeal Rhodopsins Table 1 List of microbial rhodopsins with database accession numbers or other sources.

We show all microbial opsin genes found in the NCBI database except proteorhodopsins, for which ∼800 have been identified (see text). We selectively present a subset of proteorhodopsins for which absorption maxima have been published. BR, bacteriorhodopsin; HR, halorhodopsin; SRI & SRII, sensory rhodopsins I and II; PR, proteorhodopsin; NOPI, Neurospora opsin I; CSRA & CSRB, Chlamydomonas sensory rhodopsins A and B Species and Name

Accession Number

Comments

Archaea Haloarcula argentinensis BR Haloarcula japonica BR Haloarcula sp. (Andes) BR Haloarcula vallismortis BR Haloarcula vallismortis HR Haloarcula vallismortis SRI Haloarcula vallismortis SRII Halobacterium marismotui BR

D31880 AB029320 S76743 D31882 D31881 D83748 Z35308 −

Halobacterium marismotui HR

−

Halobacterium marismotui SRI Halobacterium marismotui SRII Halobacterium marismotui BR-2 Halobacterium marismotui SRI-2 Halobacterium salinarum BR Halobacterium salinarum HR Halobacterium salinarum SRI

− − − − V00474 D43765 L05603

Halobacterium salinarum SRII

U62676

Halobacterium salinarum mex BR Halobacterium salinarum mex HR Halobacterium salinarum port BR Halobacterium salinarum port HR Halobacterium salinarum shark BR Halobacterium salinarum shark HR Halobacterium sp. AUS-1 BR Halobacterium sp. AUS-1 SRII Halobacterium sp. AUS-2 BR Halobacterium sp. NRC-1 BR Halobacterium sp. NRC-1 HR Halobacterium sp. NRC-1 SRI

D11056 P33970 D11057 Q48315 D11058 D43765 J05165 AB059748 S56354 NP 280292 NP 279315 AAG19914

Halobacterium sp. NRC-1 SRII

AAG19988

Halobacterium sp. SG1 BR Halobacterium sp. SG1 HR

X70291 X70292

H+ pump H+ pump H+ pump H+ pump Cl− pump phototaxis phototaxis H+ pump by homology with BR, Victor Ng, pers. com. Cl− pump by homology with HR, Victor Ng, pers. com. phototaxis,Victor Ng, pers. com. phototaxis,Victor Ng, pers. com. unknown,Victor Ng, pers. com. unknown,Victor Ng, pers. com. λmax = 568 nm, H+ pump λmax = 576 nm, Cl− pump λmax = 587 nm, phototaxis (attractant/repellent) λmax = 487 nm, phototaxis (repellent) H+ pump Cl− pump H+ pump Cl− pump H+ pump Cl− pump H+ pump phototaxis H+ pump H+ pump Cl− pump λmax = 587 nm, phototaxis (attractant/repellent) λmax = 487 nm, phototaxis (repellent) H+ pump Cl− pump (Continued)

3

4 Microbial Rhodopsins Table 1 (Continued)

Species and Name

Accession Number

Comments

Archaea Halobacterium sp. SG1 SRI Halorubrum sodomense BR Halorubrum sodomense HR Halorubrum sodomense SRI Haloterrigena sp. Arg-4 BR Haloterrigena sp. Arg-4 HR Natronomonas pharaonis HR Natronomonas pharaonis SRII

X70290 D50848 AB009622 AB009623 AB009620 AB009621 J0519 9 Z35086

phototaxis H+ pump Cl− pump phototaxis H+ pump Cl− pump Cl− pump λmax = 497 nm, phototaxis (repellent)

Eubacteria Anabaena sp. PCC7120

AP003592

Gloeobacter violaceus PCC 7421 Magnetospirillum magnetotacticum γ -proteobacterium (BAC31A8) γ -proteobacterium (HOT75m4) γ -proteobacterium (HOT0m1) γ -proteobacterium (PalE6) γ -proteobacterium (eBac64A5) γ -proteobacterium (eBac40E8) γ -proteobacterium (RSr6a5a6) γ -proteobacterium (RS23) γ -proteobacterium (RSr6a5a2)

NP 923144 − AF279106 AF349981 AF349978 AAK30200 AAK30175 AAK30174 AAO21455 AAO21449 −

Also known as Nostoc, λmax 543 nm, photosensory Unicellular cyanobacterium genome.ornl.gov/microbial/mmag λmax = 527 nm, H+ pump; GPR λmax = 490 nm, H+ pump; BPR λmax = 518 nm λmax = 490 nm λmax = 519 nm λmax = 519 nm λmax = 540 nm λmax = 528 nm λmax = 505 nm, RSr6a5a6(V105E), from Oded Béjà

Fungi Botrytis cinerea Botryotinia fuckeliana Cryptococcus neoformans

AL115930 − CF192410

Fusarium sporotrichioides Fusarium graminearum Leptosphaeria maculans Mycosphaerella graminicola Neurospora crassa NR Triticum aestivum Ustilago maydis Algae Chlamydomonas reinhardtii CSRA

BI187800 BU067691 AF290180 A W 180117 AF135863 CA747087 CF642219 AF508965

Chlamydomonas reinhardtii CSRB Pyrocystis lunula Guillardia theta Acetabularia acetabulum

AF508966 AF508258 AW342219 CF259014

cogeme.ex.ac.uk www.genome.ou.edu/cneo.html, Basidiomycetes

two homologs are present λmax = 534 nm Basidiomycetes photomotility for high light intensity photomotility for low light intensity dinof lagellate cryptomonad green alga

3 Clues to Newfound Microbial Rhodopsin Function from Primary Sequence 5

NpSRII. 55–75% identity in these 22 residues is also found in the new rhodopsins (Figure 1). The functions of the four archaeal rhodopsins have been well characterized. BR (λmax = 568 nm) and HR (λmax = 576 nm) are light-driven ion pumps for protons and chloride, respectively, absorbing maximally in the green-orange region of the spectrum [25, 28]. Their electrogenic transport cycles provide energy to the cell under conditions in which respiratory electron transport activity is low. Accordingly, their production in the cells is induced when oxygen is depleted in late exponential/early stationary phase cultures. Both BR and HR hyperpolarize the membrane to generate a positive outside membrane potential, thereby creating inwardly directed proton motive force. HR further contributes to pH homeostasis by hyperpolarizing the membrane by electrogenic chloride uptake rather than proton ejection, thereby providing an electrical potential for net proton uptake especially important in alkaline conditions. SRI and SRII are phototaxis receptors controlling the cell’s swimming behavior in response to changes in light intensity and color [26]. SRI (λmax = 587 nm) is also induced in cells in late exponential/early stationary phase, and attracts the cells to orange light useful to the transport rhodopsins. SRI is unique among known photosensory receptors in that it produces opposite signals (attractant and repellent) depending on the wavelengths of stimulating light. To avoid guiding the cells into light containing harmful near-UV radiation, SRI uses its color-discriminating mechanism to ensure the cells will be attracted to orange light only if that light is not accompanied by near-UV wavelengths [29]. The mechanism is based on photochromic reactions of the protein. If SRI absorbs a single photon (maximal absorption in the orange) it produces a photointermediate species called SRI-M or S373 (λmax = 373 nm) that is interpreted by the cells’ signal transduction machinery as an attractant signal. However, if S373 is photoexcited, it generates a strongly repellent-signaling photointermediate. Therefore single-photon excitation of SRI, such as occurs in orange light, attracts the cells, whereas two-photon excitation, as occurs in white light, repels the cells. SRII absorbs in the mid-visible range (λmax = 487 nm in H. salinarum SRII) and appears to serve only a repellent function [30]. It is the only rhodopsin in H. salinarum produced in cells during vigorous aerobic grow when light is not being used for energy and is therefore best avoided because of possible photooxidative damage.

3 Clues to Newfound Microbial Rhodopsin Function from Primary Sequence Comparison to Archaeal Rhodopsins

A challenge posed by the newfound microbial rhodopsin genes is to identify the photochemical and physiological function of the proteins in the cells containing them, some of which are uncultivated microorganisms. Success has been obtained in several cases: as detailed below, the Monterey Bay surface water-proteorhodopsin functions as a light-driven proton pump for the γ -proteobacterium SAR86 in its



native marine environment [18, 19], and therefore its physiological function is similar to that of BR in H. salinarum. In contrast, the Anabaena (Nostoc) rhodopsin [15] and Chlamydomonas reinhardtii pigments CSRA and CSRB [16] have been demonstrated to serve photosensory rather than transport functions, and therefore are functionally more similar to the archaeal SRI and SRII. These known transport proteins and sensory proteins do not group separately in phylogenetic analyses (Figure 2); therefore the phylogenetic trees do not permit assignment of particular sequences as encoding transport or sensory proteins. Some individual residue differences, however, provide a clue, as do the photochemical reaction cycle kinetics of the proteins. One difference in the primary sequence between BR, HR and the SRs stands out. Asp96 in BR functions as a proton donor, returning a proton to the Schiff base from the cytoplasmic side of the protein during the pumping cycle. This proton transfer improves the pumping efficiency of BR by accelerating the decay of its unprotonated Schiff base photocycle intermediate, M, and is present in all BR homologs in the haloarchaea. In the sensory rhodopsins, the corresponding M intermediates are signaling states of the receptor proteins (demonstrated unequivocally only for HsSRI), and longer M lifetimes increase the signaling efficiencies of the receptors. Accordingly, each of the five known haloarchaeal sensory rhodopsin sequences lacks a carboxylate residue at the position corresponding to Asp96 and contains Tyr or Phe instead. The residue corresponding to Asp85, which is the proton acceptor from the Schiff base, is a carboxylate residue in BR, SRI, and SRII and in each of the newly identified rhodopsins (Figure 1). HR does not produced an unprotonated Schiff base intermediate in its photocycle, and therefore does not contain a carboxylic acid residue in either of the positions corresponding to Asp96 and Asp85. Notably the SAR86 proteorhodopsin demonstrated to be a light-driven proton pump does contain a carboxylate (Glu108) at the position corresponding to Asp96 in BR, and moreover Glu108 has been shown to participate in the reprotonation of the Schiff base in the latter half of the photocycle as does Asp96 in BR [31, 32]. On ← Fig. 1 Primary sequence comparison of 15 microbial rhodopsins. We selected several opsin genes from each domain of life. Archaea- BR: Halobacterium salinarum bacteri-orhodopsin, SRI: H. salinarum sensory rhodopsin I, NpSRII: Natronomonas pharaonis sensory rhodopsin II; BacteriaGPR: (-proteobacterium (BAC31A8) proteorhodopsin, BPR: (-proteobacterium (HOT75m4) proteorhodopsin, Gloeobacter: microbial rhodopsin from Gloeobacter violaceus PCC 7421, Anabaena: sensory rhodopsin from Anabaena (Nostoc) sp. PCC7120; Eukarya- Fungi-rhodopsin from Neurospora crassa, Leptosphaeria maculans,

Botrytis cinerea, and Cryptococcus neoformans, Algae- Pyrocyctis: rhodopsin from Pyrocystis lunula, CSRA & CSRB: Chlamydomonas reinhardtii sensory rhodopsins A and B (N-terminal portions), Guillardia: rhodopsin from Guillardia theta. Conserved residues are marked with black background and the 22 residues in the retinal-binding pocket are marked with asterisks. Bacteriorhodopsin Asp85 and Asp96 in helix C and corresponding residues in the other pigments are marked with blue background (see text). Red-colored KWG residues on helix E are nearly completely conserved in fungal rhodopsins.



the basis of available information, the presence of the carboxylate residue at this position appears to be a necessary, but not a sufficient condition for identification of a new rhodopsin as a proton pump. Neurospora rhodopsin also contains a glutamate at this position, but extensive analysis of the photoactivity of the protein expressed in Pichia pastoris, as well as purified and reconstituted into liposomes, reveals a non-transport photocycle with kinetics indicative of a sensory rhodopsin [33]. A caveat is that one cannot be certain that the protein is folded correctly when expressed heterologously, although the absorption spectrum in the visible range and photochemical reactivity of the expressed protein when reconstituted with all-trans retinal provides some assurance. The demonstrated sensory rhodopsins, namely the archaeal SRI and SRII proteins, the Anabaena rhodopsin, and Chlamydomonas CSRA and CSRB, all lack a carboxylate residue at the homologous position of the BR Schiff base proton donor Asp96, while containing the carboxylate Schiff base proton-acceptor residue corresponding to Asp85 in BR (Figure 1). Hence the absence of a carboxylate in the donor position in Cryptococcus neoformans (alanine in the corresponding position) and in a marine proteorhodopsin sequence recently deposited in GenBank

← Fig. 2 Phylogenetic tree of microbial rhodopsins. A neighbor-joining tree was constructed from CLUSTALX(1.81) alignment of 46 microbial rhodopsin apoproteins. The tree was constructed using the full-length sequences, except in the case of Chlamydomonas sensory opsins, in which only the rhodopsin domains were used (CsoA, 378 N-terminal residues; CsoB, 303 N-terminal residues). Scale represents number of substitutions per site (0.1 indicates 10 nucleotide substitutions per 100 nucleotides). 1000 bootstrap replicates were performed to determine the reliability of the tree topology. The tree was drawn using TreeView1.6.6. Abbreviations: NOPI (Neurospora crassa opsin I), HtHR (Haloterrigena sp. halorhodopsin), HvalHR (Haloarcula vallismortis halorhodopsin), HsHR (Halobacterium salinarum halorhodopsin), HsportHR (Halobacterium salinarum port halorhodopsin), NpHR (Natronomonas pharaonis halorhodopsin), HsodHR (Halorubrum sodomense halorhodopsin), HspSGIHR (Halobacterium sp. SG1 halorhodopsin), AnabaenaSR (Anabaena (Nostoc) sp. PCC7120 sensory rhodopsin), HtBR (Haloterrigena sp. bacteriorhodopsin), HaspBR (Haloarcula sp. bacteriorhodopsin),

HvalBR (Haloarcula vallismortis bacteriorhodopsin), HajaponicaBR (Haloarcula japonica bacteriorhodopsin), HaargentBR (Haloarcula argentinensis bacteriorhodopsin), HsportBR (Halobacterium salinarum port bacteriorhodopsin), HsBR (Halobacterium salinarum bacteriorhodopsin), HsmexBR (Halobacterium salinarum mex bacteriorhodopsin), HspAUS-2BR (Halobacterium sp. AUS-2 bacteriorhodopsin), HsodBR (Halorubrum sodomense bacteriorhodopsin), HspAUS-1BR (Halobacterium sp. AUS1 bacteriorhodopsin), HvalSRII (Haloarcula vallismortis sensory rhodopsin II), NpSRII (Natronomonas pharaonis sensory rhodopsin II), HspAUS-1SRII (Halobacterium sp. AUS-1 sensory rhodopsin II), HsSRII (Halobacterium salinarum sensory rhodopsin II), HsodSRI (Halorubrum sodomense sensory rhodopsin I), HvalSRI (Haloarcula vallismortis sensory rhodopsin I), HsSRI (Halobacterium salinarum sensory rhodopsin I), PR eBAC64A5 ((proteobacterium proteorhodopsin), PR MBP (GPR; (-proteobacterium proteorhodopsin BAC21A8), CSRA and CSRB (Chlamydomonas reinhardtii sensory rhodopsins A and B, respectively).


(gi|42850614|gb|EAA92632.1, threonine in the corresponding position) strongly suggest sensory functions for these proteins. More than 10-fold faster photocycling rates distinguish the archaeal transport from the sensory pigments; the first-found sensory rhodopsin, archaeal SRI, in fact was initially called “slow-cycling rhodopsin” for this reason [4, 34]. The transport rhodopsins are characterized by photocycles typically 300 ms [27]. This large kinetic difference is functionally important since a rapid photocycling rate is advantageous for efficient ion pumping, whereas a slower cycle provides more efficient light detection because signaling states persist for longer times. The photocycle rate difference, which holds firm for the archaeal rhodopsins, may in some cases not be a definitive criterion for assigning a transport versus sensory function. The Anabaena rhodopsin which has been concluded to be a sensory protein based on other criteria has a photocycle half-time of 110 ms [15], intermediate between archaeal transport and sensory proteins. Furthermore, a deep sea proteorhodopsin from a Hawaiian Ocean-Time station plankton sample from 75 m depth exhibits a light-driven proton transport cycle that is ∼10-fold slower (60 ms in cells) than that of the Monterey Bay proteorhodopsin [32]. The slower photocycling rate of the deep sea pigment is explained as an adaptation to the ∼10-fold decreased photon flux rate available to the BPR visible absorption band at 75 m.

4 Bacterial Rhodopsins 4.1 Green-absorbing Proteorhodopsin (“GPR”) from Monterey Bay Surface Plankton

Among the most abundant and widely distributed of the type 1 rhodopsins are the proteorhodopsins, the first of which was identified by genomic analysis of marine proteobacteria in plankton from Pacific coastal surface waters. The proteorhodopsin gene was the first found to encode a eubacterial homolog of the archaeal rhodopsins and was revealed by BAC library construction and sequencing of naturally occurring marine bacterioplankton from Monterey Bay [17]. The gene was functionally expressed in Escherichia coli and bound retinal to form an active, light-driven proton pump. The rRNA sequence on the same DNA fragment identified the organism as an uncultivated γ -proteobacterium (the SAR86 group), and the expressed protein was named proteorhodopsin. Phylogenetic comparison with archaeal rhodopsins placed proteorhodopsin on an independent long branch (Figure 2). The new pigment, designated GPR (λmax = 525 nm), exhibited a photochemical reaction cycle with intermediates and kinetics characteristic of archaeal proton-pumping rhodopsins. Its transport, spectroscopic, and photochemical reactions have now been characterized by a number of laboratories in Escherichia

4 Bacterial Rhodopsins

coli-expressed form, [17, 18, 20, 31, 35–38]. The efficient proton pumping and rapid photocycle (15 ms halftime) of the new pigment strongly suggested that proteorhodopsin functions as a proton pump in its natural environment. Asp97 and Glu108 in GPR function as Schiff base proton acceptor and donor carboxylate residues during the GPR pumping cycle, analogous to Asp85 and Asp96, respectively, at the corresponding positions in BR [31, 32]. The next step was examination of the plankton samples directly for the newfound protein’s activity. Retinylidene pigmentation with photocycle characteristics identical to that of the E. coli-expressed proteorhodopsin gene was demonstrated by flash spec-troscopy in membranes prepared from Monterey Bay picoplankton [18]. Estimated from laser flash-induced absorbance changes, a high density of proteorhodopsin in the SAR86 membrane is indicated, arguing for a significant role of the protein in the physiology of these bacteria. The flash photolysis results provided direct physical evidence for the existence of proteorhodopsin-like pigments and endogenous retinal molecules in the prokaryotic fraction of the Monterey Bay coastal surface waters, and provide compelling evidence that GPR functions as a light-driven proton pump photoenergizing SAR86 cells in their natural environment. Furthermore, the amplitude of the flash-photolysis signals permit a rough estimate of the total rate of solar energy conversion to proton motive force by marine proteorhodopsins; assuming for the calculation that the Monterey Bay sample has the average PR content, the conversion rate is on the order of 1013 –1014 W, a globally significant contribution to the biosphere. Since the initial finding of GPR, a wide variety of similar genes has been identified in picoplankton from very different ocean environments: the Antarctic, Central North Pacific, Mediterranean Sea, Red Sea, and the Atlantic Ocean [18–22]. Genes have been isolated from both surface and deep-water samples, and both coastal and open-sea areas. New members from the PR family were recently reported to be found also in marine α-proteobacteria [19], and based on whole genome “shotgun sequencing” of microbial populations collected en mass on tangential flow and impact filters from sea water samples collected from the Sargasso Sea near Bermuda, a remarkable 782 different partial sequences homologous to proteorhodopsins were identified [23]. Thus, microbial rhodopsin abundance and diversity within marine environments appears to be large. 4.2 Blue-absorbing Proteorhodopsin (“BPR”) from Hawaiian Deep Sea Plankton

One of the variant groups (designated clade II) of proteorhodopsin genes, differing by ∼22% in predicted primary structure from the clade I group defined by the GPR gene and its close relatives, was detected in both the Antarctic and in 75-m deep ocean plankton from Hawaiian water [18]. The Antarctic and Hawaiian PR genes when expressed in E. coli exhibit a blue-shifted absorption spectrum (λmax = 490 nm; hence referred to as “BPR“) with vibrational fine structure, unlike the unstructured spectrum of GPR [18]. The stratification of the surface GPR and 75-m BPR with depth is in accordance with light spectral quality at these depths [18].

11


The different absorption spectra of GPR and BPR have provided an opportunity to examine “spectral tuning” in two rhodopsins with closely similar primary sequence. One of the most notable distinguishing properties of retinal among the various chromophores used in photosensory receptors is the large variation of its absorption spectrum depending on interaction with the apoprotein (“spectral tuning”) [39, 40]. In rhodopsins, retinal is covalently attached to the ε-amino group of a lysine residue forming a protonated retinylidene Schiff base. In methanol a protonated retinylidene Schiff base exhibits a λmax = of 440 nm. The protein microenvironment shifts the λmax [the “opsin shift” [41] to longer wavelengths, e.g. to 527 nm in GPR and to 490 nm in BPR. With structural modelling comparisons and mutagenesis, a single residue difference in the retinal binding pockets at position 105 (Leu in GPR and Gln in BPR) was found to function as a spectral tuning switch and to account for most of the spectral difference between the two pigment families [20]. The mutations at position 105 almost completely interconverted the absorption spectra of BPR and GPR. GPR L105Q shifted to the blue and acquired vibrational fine structure like wild-type BPR, and BPR Q105L shifted to the red and lost the fine structure exhibiting spectra similar to those of GPR. Among both type 1 and type 2 rhodopsins the mechanisms of spectral tuning in general are still not well understood in physical chemical terms and the Q/L switch stands out as a simple spectral tuning model amenable to investigation. Spectral tuning is discussed in more detail in Section 6, below. Another difference between GPR and BPR is their photocycle halftimes, 6.5 ms and 60 ms respectively in E. coli cells [32]. The difference in photocycle rates and their different absorption maxima may be explained as an adaptation to the different light intensities in their respective marine environments, based on measured spectral distributions of intensities of solar illumination at the ocean surface and at various depths [42]. Taking into account the blue shift of BPR, matching the lower photon fluence rate from solar radiation requires a 10-fold slower photocycle in BPR than in GPR. Therefore there is no selective pressure for a photocycle faster than that of BPR at that depth. BPR may function to energize cells by light-driven electrogenic proton pumping, as does GPR. However, the contribution of solar energy capture from BPR is severely limited by the low light intensities in deep waters. This consideration raises the possibility of a regulatory rather than energy harvesting function of BPR, based either on its slow proton pumping or by yet unidentified protein-protein interaction with transducers in its native membrane. 4.3 Anabaena Sensory Rhodopsin

A rhodopsin pigment in a cyanobacterium established that sensory rhodopsins also exist in eubacteria. A gene encoding a homolog of the archaeal rhodopsins was found via a genome-sequencing project of Anabaena (Nostoc) sp. PCC7120 at Kazusa Institute (http://www.kazusa.or.jp/). The opsin gene was expressed in E. coli, and bound all-trans retinal to form a pink pigment (λmax = 543 nm) with a

4 Bacterial Rhodopsins

photochemical reaction cycle containing an M-like photointermediate and 110 ms half-life at pH 6.8 [15]. The opsin gene was found in the genome to be adjacent to another open reading frame separated by 16 base pairs under the same promoter. This operon is predicted to encode a 261-residue protein (the opsin) and a 125-residue (14 kDa) protein. The rate of the photocycle is increased ∼20% when the Anabaena rhodopsin and the soluble protein are co-expressed in E. coli [15], indicating physical interaction between the two proteins. Binding of the 14-kDa protein to Anabaena rhodopsin was confirmed by affinity-enrichment measurements and Biacore interaction analysis. The pigment did not exhibit detectable proton transport activity when expressed in E. coli, and Asp96, the proton donor of BR, is replaced with Ser86 in Anabaena rhodopsin. These observations are compelling that Anabaena opsin functions as a photosensory receptor in its natural environment, and strongly suggest that the 125-residue cytoplasmic soluble protein transduces a signal from the receptor, unlike the archaeal sensory rhodopsins which transmit signals by transmembrane helix–helix interactions with integral membrane transducers (Figure 3). Chimeric constructs have established that the archaeal sensory rhodopsins SRI and SRII transmit signals to their cognate membrane-embedded taxis transducers by interaction with the transducers, transmembrane helices and a short membrane proximal domain [43, 44] and an extensive membrane-embedded transducerbinding region has been observed in the X-ray structure of N. pharaonis SRII [12] co-crystallized with its taxis transducer fragment [10, 45]. The interaction of the soluble 14-kDa protein, likely to be a signal transducer, with Anabaena rhodopsin, therefore would extend the range of signal transduction mechanisms used by microbial sensory rhodopsins. Atomic resolution structures for the Anabaena pigment and its putative transducer have been obtained [46], but the physiological function of Anabaena SR has not been established. Several photophysiological responses of Anabaena with unidentified photosensory receptor(s) have been discussed [15, 47]. One of these is light-modulation of the pigments contained in the light-harvesting complex of Anabaena, a photoresponse called chromatic adaptation. Green light such as absorbed by Anabaena rhodopsin has been found to modulate complementary chromatic adaptation in the related cyanobacteria Calothrix and Fremyella [48]. The adaptation consists of differential biosynthesis of blue-absorbing phycoerythrins and red-absorbing phycocyanins depending on light quality. The genome of Anabaena sp. 7120 contains phycoerythrin subunits α and β(pecA and B), allophycocyanin subunits α and β(apcA and B), and phycocyanin sub-units α and β(cpcA and B). Green light (optimally 540 nm) promotes phycoerythrin synthesis whereas red light (optimally 650 nm) promotes phycocyanin synthesis [49]. A phytochrome would be an attractive candidate for a red light sensor, and 3 phytochrome homologs are present in the Anabaena sp. 7120 genome. The Anabaena rhodopsin (λmax = 543 nm) is a candidate for the green light sensor, and may function alone to discriminate color via its photochronic reactions [46]. The two major biliproteins in Synechocystis sp. 6830 are phycocyanin (λmax = 617 nm) and allophycocyanin (λmax = 650 nm) [50].

13


5 Eukaryotic Microbial Rhodopsins

Interestingly, the phycoerythrin gene which is regulated by green light is missing in the genome of Synechocystis which does not contain an opsin-encoding gene. 4.4 Other Bacterial Rhodopsins 4.4.1 Magnetospirillum Genome sequencing of the α-proteobacterium Magnetospirillum magnetotacticum revealed a microbial opsin gene (Table 1), proceeded by a homolog of brp, which encodes an enzyme for synthesis of retinal from β-carotene in H. salinarum [51]. The two gene-operon contains a single promoter. 4.4.2 Gloeobacter violaceus PCC 7421 Gloeobacter is a unicellular cyanobacterium and the complete genome of this strain has been sequenced [52]. There is only one type 1 rhodopsin gene, which unlike that of Anabaena, encodes a carboxylate (Glu) at the BR D96 position, suggesting a proton-pumping function.

5 Eukaryotic Microbial Rhodopsins 5.1 Fungal Rhodopsins

A genome sequencing project on the filamentous fungus Neurospora crassa revealed the first of the eukaryotic homologs, designated NOP-1 [14], and search of genome databases currently in progress indicates the presence of archaeal rhodopsin homologs in various fungi including plant and human pathogens — Ascomycetes: Botrytis cinerea, Botryotinia fuckeliana (anamorph Botrytis cinerea), Fusarium ← Fig. 3 Functional diversity among microbial rhodopsins. Domains of the two sensory rhodopsins from Chlamydomonas reinhardtii, CSRA and CSRB, based on secondary structure predictions, compared with those for proteorhodopsin, halorhodopsin and sensory rhodopsin I (in a dimeric complex with its cognate dimeric transducer) from haloarchaea and cyanobacterial sensory rhodopsin from Anabaena. The retinal chromophore (shown in red) is covalently linked to a conserved lysine residue in the seventh trans-membrane helix in each protein. Colors shown are approximately the color of the pigments. Residues in helix C of prote-

orhodopsin that are important for proton translocation, Asp-97 and Glu-108, and the amino acid differences at their corresponding positions in the other rhodopsins are highlighted. The corresponding residues are not shown for halorhodopsin (see text) to avoid the impression that they are on the chloride translocation path. Transducer domains or proteins shown in green are presumably involved in the post-receptor signal transduc-tion processes. For the cytoplasmic 14-kDa protein associated with ASR and for CSRA and CSRB, the numbers indicated correspond to the number of amino acid residues in each module.

15


sporotrichioides, Gibberella zeae (anamorph Fusarium graminearum), Leptosphaeria maculans, Mycosphaerella graminicola (two opsin homologs) and Basidiomycetes: Cryptococcus neoformans and Ustilago maydis. Each of these organisms contains genes predicted to encode proteins with the retinal-binding lysine in the seventh helix and high identity in the retinal-binding pocket. The Asp Schiff base counter ion and proton acceptor (Asp85 in BR) is conserved among all fungal opsin homologs and the carboxylate proton donor specific to proton pumps (Asp96 in BR) is also either Asp or Glu except in Cryptococcus which contains an Ala residue. The nop-1 gene was heterologously expressed in the yeast Pichia pastoris, and it encodes a membrane protein that forms with all-trans retinal a green lightabsorbing pigment (λmax = 534 nm) with a spectral shape and bandwidth typical of rhodopsins [33]. Laser-f lash kinetic spectroscopy of the retinal-reconstituted NOP-1 pigment (i.e. Neurospora rhodopsin) in Pichia membranes reveals that it undergoes a seconds-long photocycle with long-lived intermediates spectrally similar to intermediates detected in BR and other members of the type 1 family. The physiological function of Neurospora rhodopsin has not yet been identified. Based on the long lifetime of the intermediates in its photocycle and its apparent lack of ion-transport activities [at least when heterologously expressed [53], it seems likely to serve as a sensory receptor for one or more of the several different light responses exhibited by the organism, such as photocarotenogenesis or light-enhanced conidiation. Neurospora is non-motile, but phototaxis by zoospores of the motile fungus Allomyces reticulatus has been shown to be retinal-dependent [54] and therefore photomotility modulation is a likely photo-sensory function of rhodopsins in this particular fungal species. A blue-green light-induced photocycle in Cryptococcus neoformans native membranes has been detected and confirmed as deriving from the rhodopsin pigment by its absence in an opsin gene-deletion mutant. The photocycle is typical of a microbial rhodopsin exhibiting a blue-shifted intermediate characteristic of a deprotonated Schiff base species and a 100–150 ms half-life (pH 7.0, 25◦ C) (authors, unpublished). 5.2 Algal Rhodopsins

The rhodopsins of the green alga Chlamydomonas reinhardtii are the only ones in eukaryotic microbes to have an identified physiological function, namely photoreception controlling motility behavior [16]. Two type 1 opsin genes were identified in the C. reinhardtii genome. A microbial rhodopsin homolog gene is also present in Guillardia theta, which is a small bif lagellate organism considered both a protozoan and an alga, and opsin genes are also found in the dinof lagellate Pyrocystis lunula and Acetabularia acetabulum which is a unicellular green alga of the order Dasycladales found in warm waters of sheltered lagoons. The CSOA & CSOB (Chlamydomonas sensory opsin A and B) genes encode 712 and 737 amino acid proteins (Figure 1). The N-terminal 300 residues have a significant homology to archaeal rhodopsins with seven transmembrane helices and the conserved

6 Spectral Tuning

retinal binding pocket (Figure 1). The Chlamydomonas rhodopsins provide the first examples of evolution fusing the microbial rhodopsin motif with other domains. Early work established that Chlamydomonas uses retinylidene receptors for photomotility responses. Restoration of photomotility responses by retinal addition to a pigment-deficient mutant of C. reinhardtii first indicated a retinal-containing photoreceptor [55]. Subsequent in vivo reconstitution studies with retinal analogs prevented from isomerizing around specific bonds (“isomer-locked retinals”) in several laboratories further established that the Chlamydomonas rhodopsins governing phototaxis and the photophobic response have the same isomeric configuration (alltrans), photoisomerization across the C13–C14 double bond (all-trans to 13-cis), and 6-s-trans ring-chain conformation, (co-planar) as the archaeal rhodopsins [56–60]. The proteins encoded by csoA and csoB complexed with retinal [16]. RNAi suppression of the genes established that CSRA and CSRB mediate both phototaxis [16] and photophobic reactions [61] to high- and low-intensity light, respectively. The functions of the two rhodopsins were demonstrated by analysis of electrical currents and motility responses in transformants with RNAi directed against each of rhodopsin genes. CSRA has an absorption maximum near 510 nm and mediates a fast photoreceptor current that saturates at high light intensity. In contrast, CSRB absorbs maximally at 470 nm and generates a slow current saturating at low light intensity [16]. The rhodopsin domains of CSRA [62] and CSRB [63] have been shown to exhibit light-induced proton-channel activity in Xenopus oocytes. The relationship of this activity to their control of motility-regulating currents in C. reinhardtii is not clear. To clarify a possibly confusing series of reports in the literature, we mention here a protein that binds radiolabeled retinal, and named on this basis “chlamyrhodopsin,” that had been isolated from Chlamydomonas eyespot preparations [64]. For several years this most abundant protein in the eyespot membranes was assumed and often cited by the authors of that work as the photoreceptor for photomotile responses. However, its gene-predicted primary sequence, as well as that of a similar Volvox protein [65], suggest 2–4 transmembrane helices and no homology to archaeal opsins, nor is a photoactive retinal binding site evident from the sequences. Moreover, recently the “chlamyrhodopsin” has been ruled out as the photoreceptor pigment for either phototaxis or photophobic responses in C. reinhardtii [66].

6 Spectral Tuning

Comparison of the primary sequence, (Figure 1) alone gives hints to distinguishing properties of different microbial rhodopsins, but most properties cannot be deduced from primary structure alone. The Leu/Gln spectral-tuning switch at position 105 in GPR and BPR discussed above was revealed through structural modeling and mutagenesis. However, we were fortunate that a relatively simple single-residue switch is responsible for most of the color difference in that case, and, more

17


generally, detailed knowledge of atomic structure will probably be required to elucidate spectral tuning mechanisms of most microbial rhodopsins. An exemplary case is that of NpSRII [67]. Mutagenic substitution of 10 residues, in or near the retinal-binding pocket with their corresponding BR residues, produced only a 28nm red shift of the NpSRII absorption maximum [68, 69]. Structural differences responsible for the shift are evident in the 2.4-Angstrom resolution structure [12]. One notable change is a displacement of the guanidinium group of Arg72 by 1.1 Å coupled with a rotation away from the Schiff base in NpSRII. This increase in distance reduces the inf luence of Arg72 on the counterion, thus strengthening the Schiff base/counterion interaction, shifting the absorption to shorter wavelengths. In addition the position of the positive charge destabilizes the excited state contributing further blue shift [70]. Arg72 is repositioned as a consequence of several factors, including movement of its helix backbone by 0.9 Å and the cavity created by changes from BR: Phe208 →Ile197 , Glu194 →Pro183 , and Glu204 →Asp192 . Hence the spectral tuning results from precise positioning of retinal binding-pocket residues and the guanidinium of Arg72 , which could not be deduced from primary structure, but required atomic resolution tertiary structure information.

7 A Unified Mechanism for Molecular Function?

The idea that in microbial rhodopsins the sensory signaling mechanisms result from evolution “tweaking” the transport mechanism was suggested by the observation that SRI carries out light-driven proton pumping, but only when it is free of its transducer [71–74]. The HtrI protein was found to close or prevent the opening of a cytoplasmic proton-conducting channel in SRI during its photocycle. This finding led to the notion that a chemotaxis receptor progenitor of HtrI evolved an interaction with a proton-transporter progenitor of SRI, coupling to its pumping mechanism and thereby blocking the pump and converting the transport rhodopsin to a sensory receptor. NpSRII was also observed under some conditions to exhibit light-driven proton transport which was also prevented by its interaction with its transducer, HtrII [75, 76]. That the transducer-inhibition of light-driven transport occurs in both haloarchaeal rhodopsins further supports that the interaction blocking the transport is a critical aspect of the signaling mechanism. A tilting of helices, primarily helix F, contributes to opening a cytoplasmic channel in the latter half of the photocycle in BR [77]. A unifying mechanism is that ion transport and sensory signaling use the same retinal-driven protein structural changes, which is the conformational change that opens the cytoplasmic channel in the proton transport cycle. The key feature of the model is that consequences of retinal photoisomerization, including light-induced disruption of the salt bridge between the protonated Schiff base on helix G and aspartyl counterion on helix C [78], triggers tilting of helix F to which the Htr transmembrane helices are coupled. Supporting this mechanism are (i) the proton-pumping by the sensory rhodopsins, (ii) its inhibition by transducer interaction discussed above, and (iii) light-induced tilting of helix F in N. pharaonis

9 Conclusions

SRII, concluded from site-directed spin-labeling measurements [79]. Furthermore, (iv) genetic evidence supports helix-F involvement in signaling [80]. In addition, (v) substitution of the Schiff base counterion Asp85 with asparagine induces helix-F tilting in the dark in BR and the corresponding substitution in H. salinarium SRII partially activates the receptor in the dark [78]. Finally (vi), helix F interacts with the two transmembrane helices of the HtrII fragment co-crystallized with NpSRII [10]. Another prediction is that, since in the model SR helix tilting is transmitted to the Htr protein by direct helix–helix contacts, alterations in structure must occur in the Htr transmembrane domains between the receptor interaction sites and the cytoplasmic domain of the transducer, where the activity of the bound histidine kinase is controlled. Such structural alterations have been detected as light-induced changes in interactions between spin labels introduced into the NpHtrII transmembrane helices [79] and changes of disulfide bond formation rates between engineered cysteines [81]. In both studies the data indicate that the second transmembrane segment [79], and interaction of the receptor’s E-F loop with the membrane proximal domain of the transducer has been implicated in signal transfer [82, 83]. In summary, the evidence is compelling that the conformational changes in transport and sensory rhodopsins in haloarchaea share essential features despite their differing functions. For study of the newfound microbial rhodopsins, an important question is whether the light-induced conformational change observed in BR and strongly implicated in the haloarchaeal sensory rhodopsin photocycles is a key conserved feature of their functional mechanisms.

8 Opsin-related Proteins without the Retinal-binding Site

Several other genes in the fungi N. crassa [84, 85]. It may be that the conformational switching properties of the archaeal rhodopsins have been preserved in these opsinrelated proteins, while the photoactive site has been lost and its function replaced by another input module such as a protein-protein interaction domain. It is striking that opsin-related proteins lacking the retinal-binding lysine have been observed so far only in fungi and not in any of the many other classes of organisms containing type 1 rhodopsins.

9 Conclusions

Type 1 rhodopsins are present in all three domains of life, and therefore progenitors of these proteins may have existed in early evolution before the divergence of archaea, eubacteria, and eukaryotes. If so, light-driven ion transport as a means of obtaining cellular energy may well have predated the development of photosynthesis, and represent one of the earliest means by which organisms tapped solar radiation as an energy source. As more rhodopsins are identified, their evolution and dissemination into such a wide variety of organisms, whether by

19


divergence from a common progenitor or horizontal gene transfer, should become clearer. There is a much work to be done to understand the physiological roles and molecular mechanisms of the rhodopsins so far identified in the various microbial species. It seems likely that we will see even more members of this family as genomic sequencing becomes ever more rapid. The vast majority of microbial species have never been cultivated in a laboratory. Therefore the use of microbial rhodopsin probes in environmental genomics, which expands the search for homologous genes to uncultivated organisms, is likely to be especially fruitful.

Acknowledgements

We thank Elena Spudich for stimulating discussions. The work by the authors referred to in this review was supported by grants from the National Institutes of Health, National Science Foundation, Human Frontiers Science Program, and the Robert A. Welch Foundation.

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1

Light-activated Proteins Sandra Loudwig and Hagan Bayley University of Oxford, Oxford, United Kingdom

Originally published in: Dynamic Studies in Biology. Edited by Maurice Goeldner and Richard S. Givens. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30783-8

1 Introduction

While there are a considerable number of publications describing small caged molecules for applications in biology (for reviews, see [1, 2]), there are still relatively few papers describing caged proteins (for earlier reviews, see [3–6]). This is perhaps because the preparation of such molecules requires an interdisciplinary approach and often presents considerable technical difficulties. Nevertheless, the potential applications of caged proteins are likely to be highly rewarding and an expansion of work in this area is certainly warranted. We define a caged protein as an inactive protein that can be activated by light (Fig. 1). The definition is best kept loose, and here we make several asides to discuss, for example, reversible activation and light-triggered inactivation. For many applications, there is no alternative to the use of a caged protein. In other cases, the use of a caged small molecule might be considered instead. By comparison with small molecules, the advantages of caged proteins include the fact that the activity in question is precisely pinpointed and that in cellular experiments very low concentrations of a reagent can be used, avoiding, for example, problems with photolysis byproducts. Proteins are usually caged by targeted chemical modification. We aim to point out the issues with which the experimentalist should be familiar and to illustrate them with examples; however, the examples do not in themselves offer a comprehensive review of the literature, which nowadays can be better obtained from databases.


2 Light-activated Proteins

Fig. 1 Typical procedure for caging and uncaging a protein. A key residue (e.g. one in the active site of an enzyme) is derivatized with a caging reagent resulting in inactivation of the protein. The protein can be reactivated by photolysis of the caging group, which is released in an altered form.

2 The Properties of Caged Proteins 2.1 Extents of Caging and Photoactivation

Light-activated proteins are used to obtain control over the timing and location of activation of a biological process, and when necessary the extent of activation. If there are other biomolecules present, they should not be damaged. These requirements raise several important issues, the solutions to which will depend on the experimental system. While it is desirable to efficiently inactivate a protein with a caging reagent, and subsequently recover full activity upon irradiation, this cannot always be done. Therefore, the importance of the extents of caging and uncaging must be considered further, and will in turn affect the choice of caging reagent. For example, in the case of a caged signal transduction protein for injection into target cells, it is highly desirable to begin with a completely inactivated protein and even limited photoregeneration of activity within the cell (e.g. 5%) can be useful. In contrast, in many biophysical experiments, such as those involving time-resolved X-ray diffraction, circular dichroism or IR spectroscopy, a high recovery of activity is usually required (e.g. 95% or above), but complete inactivation is not usually as crucial (e.g. 5% of unmodified protein can often be tolerated). 2.2 The Rate of Uncaging: Continuous Irradiation

The rate of photorelease is often critical in biophysical experiments and should be fast in comparison to the timecourse of the event under observation. The half-time for activation of a caged reagent under continuous irradiation in dilute solution is given by [3]: t1/2 ≈ 0.3/φ p I0 ε

2 The Properties of Caged Proteins

Fig. 2 Illustration of t1/2 and J * for photolysis. During continuous photolysis at relatively low light intensity, the half-life of the caged protein is t1/2 , which is a function of the amount of light absorbed per unit time and the efficiency of conversion of the excited state to the uncaged product. Under

these conditions, the rate at which the excited states (and intermediates when they exist) are converted to product, represented by J *, is not rate determining. After a short intense light flash, the half-time for formation of the uncaged product is given by J *.

where P is the product quantum yield, I0 is the light intensity and ε is the extinction coefficient. t1/2 , ε and I0 are usually measured experimentally to find P . This expression encompasses the rate at which a collection of molecules absorbs photons and the efficiency at which the excited states are converted to uncaged product. t1/2 should be distinguished from the usually more rapid rate of breakdown of the excited states and intermediates (τ *), which will be discussed later (Fig. 2). The relationship given above is valid for a single wavelength, as would be the case for laser irradiation, and it is a reasonable approximation for a narrow band of wavelengths. Where a wide band is used, e.g. as might be the case from an arc lamp, the variation of ε and I0 with wavelength should be taken into account when determining P for the prevailing conditions. This can be done conveniently by summing I0 ε over the wavelength band by using, say, 10-nm intervals. To determine I0 for each interval from the total lamp output (determined by actinometry), both the emission spectrum of the lamp and the filter characteristics must be considered [7]. Clearly, in most circumstances, I0 can be increased far beyond the levels required to achieve photolysis in the desired time. The question is: what happens to other materials in the experimental system at high light intensities? The simple answer is that the consequences are usually deleterious. Most biological molecules are damaged by light, and the damage is exacerbated at short wavelengths and in the presence of oxygen. In a complex system, a wide variety of damage can occur, including crosslinking of nucleic acids and proteins. However, even in a simple biophysical experiment, involving a single protein, inter- and intramolecular crosslinks, polypeptide chain cleavage, and damage to susceptible residues such as tryptophan (Trp) can cause significant problems [8, 9]. At short photolysis times, a measure of the extent of uncaging relative to damage is given by E ud = t1/2d /t1/2u = φu εu /φd εd

3


where the subscripts ‘u’ and ‘d’ indicate uncaging and damage, respectively. At longer photolysis times, as uncaging nears completion, damage will “catch up” with uncaging: a warning to irradiate for no longer than necessary. The larger the value of E ud , the better and the experience of many workers in the field of caged reagents suggests that u ε u should be > 100 M−1 cm−1 , at least in the near-UV (350–400 nm), and even better when > 1000 M−1 cm−1 . In addition, photolysis should be at the longest wavelengths possible (proteins and nucleic acids without prosthetic groups absorb little above 300 nm) and preferably in the absence of oxygen. Values of u and ε u for many caging groups that have or could be used with proteins have been compiled here (Tab. 1). Because these values have not been obtained at identical wavelengths, it can be difficult to compare u ε u values, but we have attempted to quote values at wavelengths that would be used experimentally. The value of u ε u for a class of caging reagents can be manipulated by chemical substitution (Tab. 1). For example, in the case of 2-nitrobenzyl attached at the N atom of carbamoyl choline, the replacement of a methyl group with a carboxylate at the benzylic position increases from 0.25 to 0.8, resulting in an increase in u ε u (312 nm) from ∼300 to ∼1000 [10]. In another case, with a (2-nitrophenyl)ethyl group on the phenolic O atom of phenylephrine, the introduction of methoxy groups in positions 4 and 5 of the aromatic ring increases ε312 from 1300 to 4500 M−1 cm−1 (λmax is increased from 260 to 340 nm), which with a quantum yield = 0.13 gives u ε u (312 nm) ∼600 [11]. By comparison, the un-substituted derivative with = 0.05 gives u ε u (312 nm) ∼65. However, to the chagrin of the investigator, an improvement in ε u often leads to a reduction in u . In one such case, cysteine (Cys)- or thiophosphate-containing peptides were caged on the S atoms with 2nitrobenzyl or 4,5-dimethoxy-2-nitrobenzyl and little was gained from substitution of the ring, which serves to increase ε u [12]. Clearly, we have a great deal to learn about the mechanism(s) of photolysis of 2-nitrobenzyl reagents, which continue to be investigated [2, 13, 14]. A detailed study of substituted coumarins has been made by Furuta et al., who found u ε u (365 nm) values in the range of 100-630, with the highest value assigned to a tribrominated derivative [15]. By comparison, the 7-methoxycoumarins have u ε u (333 nm) values approaching 3000 [16]. Following their successful use of 4-hydroxyphenacyl (HPB) as a caging group [u ε u (300 nm) = 4400] [17], Givens et al. attempted to push the absorbance towards the visible by substitution of the aromatic ring with methoxy groups. The 3-methoxy and 3,5-di-methoxy derivatives have absorption tails above 400 nm, but the values are reduced to 0.03–0.04, which is accompanied by a change in the photolysis mechanism. In contrast to the underivatized 4-hydroxyphenacyl group, the formation of a phenylacetic acid from an initially released spiroketone (Fig. 3) is a minor pathway in the case of the methoxy reagents [18]. Additional examples can be found in Tab. 1. Further, while the determination of ε is a simple matter, the reader is cautioned to note that not all values of that make their way into tables in the literature (including Tab. 1) have been determined with equal rigor: some have been obtained by careful actino-metry, while others


5

Table 1 Caging groups for proteins Caging group

Caging group characteristics

Caged functionality and/or molecule; caging reagent

Comment s

Refs.

25, 33, 2-Nitrobenzyl derivatives, initially described for the protection 34, 141 of carboxylates [141], are the most widely used caging groups. They have been used to cage thiols, thiophosphates, phosphates, carboxylates, amino groups and alcohols. Various modifications at the benzylic position (see NB, NPE, CNB and NPT) or on the aromatic ring (see DMNB, DMNPT, NVOC and MeNPoc) have been made in attempts to improve photochemical properties (see this table). The main drawback of the 2-nitrobenzyl group is the generation of a photolytic byproduct that reacts with thiol groups [25, 33, 34]. Further, when substituted in the benzylic position, the 2-nitrobenzyl group bears an asymmetric center. (312; pH 6) = 0.84; (312; pH 8.5) = 0.14 (312; pH 6) (312; pH8.5)

1090; 180

RCH2S-NB Caged Cys in cAMP-dependent protein kinase (PKA).

NBB 312 = 1300

[3]

(312; pH 5.8) = 0.62; (312; pH 7.2) = 0.14 (312; pH 5.8) (312; pH 7.2)

[12]

810; 180

RCH2S-NB In the heptapeptide C-kemptide, the serine of kemptide is replaced with Cys. This residue is caged with NBB.

A PKA mutant with a single 65 Cys is modified with NBB. Caging with different 2-nitrobenzyl derivatives (BNPA, DMNBB and NBB) and photolysis at two pH values are compared (see this table). Modification with unsubstituted NBB gives the highest extent of inactivation and subsequent photoregeneration of activity. Caging with BNPA and DMNBB 3, 12 (see this table) and photolysis at two pH values are compared. Photolysis of C-kemptide modified with NBB gives the best quantum yield and a slightly higher yield of uncaged product: 70% at pH 5.8 against 62 and 67% for BNPA and DMNBB, respectively.

(Continued)

6 Light-activated Proteins Table 1 (continued) Caging group


312 = 1300


[3]

(312; pH 5.8) = 0.23; (312; pH 7.2) = 0.04 (312; pH 5.8) (312; pH 7.2)

300; 50

[12]

Caged thiophosphorylated serine in the heptapeptide kemptide.

(312; pH 5.8) = 0.37; (312; pH 7.3) = 0.25 (312; pH 5.8) (312; pH 7.3)

480; 325 Caged thiophosphotyrosine in an 11 amino acid peptide.

max = 272

nm

272 = 6200 (300–350; pH 7)

RPhO-NB Caged phenylephrine [11].

= 0.05 312 65 * ND [11]

ROC(O)NH-NB Caged carbamyl(300–350; pH 8) = 0.25 choline. * (nitronate decay) = 1.7 ms (pH 7) 312 325 (pH 8) max = 262

262 = 5200

nm

Comment s

Refs.

Caging with BNPA and DMNBB 3, 12 (see this table) and photolysis at two pH values are compared. Photolysis of thiophosphoserine modified with unsubstituted NBB gives the best quantum yield and the highest yield of uncaged product: 70% at pH 5.8 against 55% for DMNBB. At pH 4, thiophosphate can be selectively modified over Cys with NBB. A thiophosphotyrosine is modi- 31 fied with NBB. Caging with NBB and 4-hydroxyphenacyl bromide (HPB), and photolysis at two pH values are compared. Uncaging of HP gives the best quantum yields and a slightly better yield of uncaged product: 50–70% (at both pH 5.8 and 7.3) against 50– 60% for NB (see below). 11, 142, Photolysis of different 2-nitro143 benzyl derivatives (NB, NPE, DMNB and CNB) are compared (see this table) [11]. Proteins have not been caged directly on hydroxyls, but catalytic serines, with enhanced nucleophilicity, might be selectively modified with NBBlike compounds. Also, caged tyrosines have been incorporated into proteins by unnatural amino acid mutagenesis [142, 143]. Proteins have not been caged as 144 carbamates in this manner, but the work provides a useful study of the pH dependence of breakdown of the nitronate intermediate.


7

Table 1 (continued) Caging group



11 Proteins have not been caged directly in this way, but the work provides a useful comparison of various 2-nitrobenzyl derivatives. and hence values for NPE are about twice those for NB. The breakdown of the nitronate intermediate (Fig. 5.1.3) is slow in this case.

ROC(O)NH-NPE Caged carbamyl(300–350; pH 8) = 0.25 choline. * (nitronate decay) = 67 s (pH 7) 312 325 (pH 8)

Although the and values for 144 carbamylcholine caged with NB and NPE are the same, the decay rate of the nitronate intermediate with NPE is much faster.

RCH2S-CNB In the heptapep270 For Cys in C-kemptide C-kemptide, tide [12] the serine of kemptide is replaced with a Cys. This residue is caged with BNPA. Alternatively, the serine in kemptide is thiophosphorylated and then modified by BNPA [3, 12].

BNPA has been developed as a 3, 12, water-soluble reagent for the 47, 65 modification of nucleophiles in peptides and proteins. While 95% modification of the Cys in C-kemptide is achieved, only 10% modification of a thiophosphorylated serine in kemptide occurs under the same conditions. The value for Cys caged with CNB is much lower than for Cys caged with NB [3, 12]. Further, irradiation of CNB-Cys in PKA leads to 25–30% recovery of activity, while 80–100% recovery is found with PKA caged at Cys with NB [65]. BNPA has also been used for caging the pore-forming protein hemolysin [47].

nm

max = 262

nm

262 = 5200

(312; pH 5.8) = 0.21 312

CNB

Refs.

RPhO-NPE Caged phenyl(300–350; pH 7) = 0.11 ephrine. * (nitronate decay) = 300 ms (pH 7) 312 140 max = 272

272 = 6200

NPE

Comment s

BNPA max = 270

nm

RPhO-CNB Caged phenyl(300–350; pH 7) = 0.28 ephrine. * (nitronate decay) = 0.35 ms 312 365 266 = 6000

The quantum yield for CNB is 11 higher than for NB, NPE and DMNB. The lifetime of the nitronate is short compared to NPE and DMNB in the same context ( * not given for NB).

(Continued)



max = 266

nm

266 = 5200 (300–350; pH 7) = 0.8 * (nitronate decay) = 40 s 312 1040 [144]


Comment s

Refs.

ROC(O)NH-CNB Caged carbamylcholine [144].

is remarkably high and is thus favorable [144]. Also the rate of decay of the nitronate intermediate is faster than for NB and slightly faster than for NPE in the same context [10, 144].

10, 144

RO-NPT and RO-DMNPT Caged alcohols: * = 2.2 and 50 s values are for (biexponential rate for nitronate decay) NPT-choline and DMNPT-arseno312 910 choline. R = OMe: (312; pH 7) = 0.43 312 1935 R = H:

(312; pH 7) = 0.7

R = H, NPT R = OMe, DMNPT

312 = 4500

[3]

(312; pH 5.8) = 0.15 312

675 [12]

RCH2S-DMNB The Cys in C-kemptide is caged with DMNBB [12].

DMNB

DMNBB

(312; pH 5.8) = 0.06 312 = 270

Caged thiophosphorylated serine in kemptide.

145 for NPT is very high. for DMNPT can be as high as 0.62, e.g. for 4-DMNPT -tolylgalactoside. The value for DMNPT is favorable. Proteins have not yet been caged in this manner.

is only slightly lower than for 3, 12, BNPA, but much lower than for 65 NBB. Nevertheless, since the value at 312 nm is higher, is about double that for the NB peptide. The yield of deprotection of 67% at pH 5.8 is similar to that of NB (70%) [12]. However, when DMNBB and NBB are used to cage a Cys in PKA, the photoregeneration of activity with NBPKA is 10–15 times greater than with DMNB-PKA [65]. 3, 12 DMNBB is used to cage thiophosphorylserine in kemptide. for DMNB is lower than that for NB, but (312; pH 5.8) values are about the same (300 for NB). values for NB and DMNB-caged thiophosphoserine are lower than values for caging a Cys residue.


9



max = 330

Comment s


The value for DMNB-phenyl- 11 ephrine is similar to that for NPE-phenylephrine, but is higher. The lifetime of the nitronate lies between the values for CNB and NPE.

nm

RPhO-DMNB Caged phenyl(300–350; pH 7) = 0.13 ephrine. * (nitronate decay) = 2.8 ms 312 585 330 = 5000

max = 350

nm

DMNB is first used with caged 146 phosphate esters. Compared to NB, the absorption maximum is shifted to 350 nm and cyclic nucleotide photorelease is 200 times faster.

O Caged phosphates (cAMP and cGMP).

NH ||

/

likely to be similar to DMNB NVOC (365; pH 7.2) = 0.023 [141]

N H

N H

R

Caged arginine in a peptide inhibitor of protein kinase [141].

NVOC

Refs.

147 In the case of caged Arg, the low value of is compensated by a high , as the NVOC max is around 350 nm [141]. 6-Nitroveratryloxycarbonyl chloride (NVOC-Cl) is used to randomly modify Lys residues in proteins (see Tab. 5.1.2).

NVOC-Cl likely to be similar to DMNB

1-(4,5-Dimethoxy-2-nitrophenyl)- 66 2-nitroethene is used to modify one Cys residue per subunit in -galactosidase. Caged Cys in galactosidase.

(Continued)




Comment s

likely to be similar to DMNB

MeNPoc-OR Photoprotection of 5 hydroxyl groups in deoxyribonucleotides.

148 1-(2-Nitro-4,5-methylenedioxyphenyl)-ethyl-1-oxycarbonyl chloride is used for the protection of deoxyribonucleoside hydroxyl groups during the synthesis of oligonucleotide arrays. Proteins have not yet been caged in this manner, but the oxycarbonyl chloride might be used as an alternative to NVOC-Cl for caging by random modification of Lys residues.

MeNPoc

MeNPoc-Cl

max = 262

nm

259 = 5500

(cyclohexanone ketal) max = 259 nm 259 = 5300 (glycol)

max = 259

Caged aldehydes and ketones.

nm

259 = 7298; 350 = 1289

NNMC

(380; pH 6.7) = 0.63 (nitroso formation) 380 = 510

Model compound.

NNMCC max

260 nm

355 = 400 (365; pH 7.2) = 0.3

[154] 355

NPPOC

120

NPPOC-OR Caged alcohol in thymidine [154].

Refs.

Aldehydes and ketones have 149– been caged as dioxolane rings 153 by using 2-nitrobenzyl derivatives. Ketones have been introduced into proteins by unnatural amino acid mutagenesis and it might be possible to cage them with diol reagents. Alternatively, caged ketones themselves could be introduced by unnatural amino acid mutagenesis. 3-Nitro-2-naphthalenemethanol 50 is irradiated as a model compound to obtain values. 3-Nitro-2-naphthyloxycarbonyl chloride (NNMCC) is then used for random caging of Lys residues in an immunoglobulin, despite the low water solubility of the compound. The NPPOC group has been used in the synthesis of oligonucleotides microarrays [155]. Proteins have not yet been caged in this manner.

154, 155


11


MNI



Comment s

R = OMe: 246 nm max 246 = 17 500 (347; pH 7.2) = 0.085 [156] Two photon uncaging (730 nm), u = 0.06 GM for a caged carboxylate [158]

RCO-MNI Caged carboxylates.

156– Photo-accelerated solvolysis of 1-acyl-7-nitroindoline derivatives 158 is reported [156, 157]. is usually low, but is high, and photolysis occurs in high yield. Two-photon uncaging has been demonstrated [158]. Proteins have not yet been caged in this manner.

MNPC-OR Caged alcohols.

N-Methyl-N-(2-nitrophenyl) carbamoyl chloride is used to cage various alcohols [159] and the catalytic serine of butyrylcholinesterase (see Tab. 5.1.2) [53].

max = 262

nm

(262, pH 7.4) = 5000

is low

MNPCC

Benzoin

R1 = R2 = OMe: max, MeOH = 246 nm 246 = 14250; 282 = 3050 (366, benzene) = 0.64 (benzofuran formation) * 0.1 ns (quenching of triplet state) [161]

Refs.

53, 159

The rate of product formation 160– from the excited state is ex166 tremely fast and the quantum yield of benzofuran release is high [161]. Benzoins have been Photorelease of acetate used to cage phosphates [162], [161]; alcohols [163] and amines [164, benzoin-OC(O)OR, 165]. The chemically inert benzophotorelease of furan byproduct has a huge alcohols [163]; absorbance above 300 nm (for benzoin-OC(O)NHR, R = R = OMe: max, Ephotorelease of amines 1 2 tOH = 301 nm; [164, 165]. 301 = 27480) [161]. Proteins have not yet been caged as benzoins. Potential caging reagents might have solubility problems. The reagent presents a chiral center.

(Continued)




Comment s

Refs.

17, 167, 4-Methoxyphenacyl derivatives were first reported by Sheehan 168 in an attempt to simplify the benzoin protecting group [167]. Givens developed the 4-hydroxy derivative to improve aqueous solubility [17]. Like benzoin, the 4-hydroxy-phenacyl group offers a very short * value. It does not contain or generate a chiral center and a predominant photolysis byproduct is usually released: 4-hydroxyphenylacetic acid. The byproduct, besides being chemically inert, does not absorb at the wavelengths of irradiation (> 300 nm) [168].

HP

(313; pH 7.2) = 0.085 310

670 [169]

RS-HP Caged thiol in 3 -thio2 -deoxythymidine phosphate [169].

HPB

(312; pH 5.8) = 0.65; (312; pH 7.3) = 0.56 310 5100 (pH 5.8); 310 4400 (pH 7.3)

Caged thiophosphotyrosine in a peptide.

4-Hydroxyphenacyl bromide is 31 used to cage a thiophosphorylated tyrosine in an unprotected peptide. Compared with the 2-nitrobenzyl (NB) group, HP exhibits better quantum yields. values are similar at pH 5.8 and 7.3. 4-Hydroxyphenacyl bromide is employed to cage a thiophosphorylated threonine in PKA (see Tab. 5.1.2).

(312; pH 7.3) = 0.21 310

The value is lower than for 54, 169 other 4-hydroxyphenacyl-caged functional groups and several photolysis byproducts are formed [169]. Nevertheless, 4-hydroxyphenacyl bromide (HPB) has proved successful for caging the catalytic Cys of a protein tyrosine phosphatase and about 70% activity is regained on uncaging [54].

1650

Caged thiophosphothreonine in PKA.

7


13



max = 286

nm

286 = 14600

(H2O/CH3CN) [168] (300; pH 7.3) = 0.37 300 4400 * = 1.2 ns (from quenching of triplet state) [17] values for caged phosphate max = 370

nm

304 = 11 730 (300–350; pH 7.2) =

HDMP


0.03–0.04 * = 69 ns 304 410

O

Comment s

Refs.

The very short * value is notable.

17, 167, 168

Caged phosphate (ATP) [17]. RCOO-HP Caged carboxylates [167].

RCOO-HDMP Caged carboxylate in GABA and glutamate.

The introduction of methoxy 18 groups in positions 3 and 5 of the ring leads to a red-shift of the absorbance, but also to a drastic decrease of the quantum yield. The photolysis pathway appears to be altered and the release of the caging group as the 4-hydroxyphenylacetic acid is a minor pathway. The 3-monomethoxy reagent was also examined. Proteins have not yet been caged with methoxy substituted HP. Coumarin derivatives usually do 15 not display high quantum yields, but they do absorb strongly above 300 nm. They can also show an increase in fluorescence upon irradiation. A very interesting feature of several coumarins is susceptibility to two-photon irradiation with IR light, which enables spatial control of photolysis at the micometer level. A drawback is their low aqueous solubility.

(Continued)



max = 325

nm

(325, pH 7.2) = 11600; (365, pH 7.2) = 4100 (365, pH 7.2) = 0.025

HCM

365 = 103 Two-photon uncaging u = 1.07 GM (740 nm); u = 0.13 GM (800 nm) for caged acetate [15] max = 397

nm

(397, pH 7.2) = 15 900; (365, pH 7.2) = 9 700


AcCOO-TBHCM Caged carboxylate (acetate).

365 = 631 u = 0.96

GM (740 nm); u = 3.1 GM (800 nm) values for caged acetate

max = 327

nm

327 = 13 300 (333; pH 7.2) = 0.21

MCM

333 = 2793 for caged cGMP [16]

max = 394

Caged phosphate (cAMP or cGMP).

The introduction of bromine 15 atoms on the coumarin ring of HCM increases both the quantum yield and the absorbance of caged carboxylates. The best results are achieved with three additional bromines, as shown. The two-photon cross section is high. Proteins have not yet been caged with the TBHCM group. Furuta et al. obtain 340 = 670 for caged cAMP in Ringer’s solution [171]. Schade et al. obtain (333, pH 7.2) = 1716 for caged cAMP and 333 = 2793 for caged cGMP in 20% methanol/80% HEPES buffer [16]. Proteins have not yet been caged with the MCM group.

16, 171

DMACM was first described for 172, caging cAMP and cGMP [172]. 173 max is at a favorable wavelength.

nm

394 = 17 200 (333; pH 7.2) = 0.28 for caged AMP [173]

DMACM

Refs.

For caged acetate, the value is 15, 162, low, but the strong absorbance 170 compensates for this [15]. Proteins have not yet been caged Caged phosphate [162] with the HCM group. Cys residues might be modified directly cAMP [170] with 4-halomethyl-7-hydroxyRCOO-HCM [15] RCNHCOO-HCM [15]. coumarins. Two-photon uncaging is demonstrated. The cross-sections are favorable.

(365, pH 7.2) = 0.065

TBHCM

Comment s

Caged phosphates (cAMP, cGMP, ATP, ADP or AMP).


15



max = 370


nm

365 = 12 600–19 500 (365; pH 7.2) = 0.03–0.06 365 = 403–1026 u = 0.51–1.23 GM (740 nm)

BHC-diol

Caged aldehydes and ketones. max = 369

X

nm

369 = 2600

AQMOC-OR Caged alcohol (galactose).

174 Anthraquinon-2-ylmethoxycarbonyl is used to cage galactose. The value is low. However, AQMOC proves to be superior to various arylmethyl carbonates (7-methoxycoumarinyl-4-methyloxycarbonyl, pyren-1-ylmethoxycarbonyl and phenanthren-9ylmethoxycarbonyl) in terms of and .

DANP-OCOR Caged carboxylate.

For R = Me2N-, the quantum 175, 176 yield is very low, although the absorbance is high [175]. The ester is quite unstable in aqueous media (hydrolysis: t1/2 = 99 min at pH 7), but the stability is improved compared to the compound with R = OMe (hydrolysis: t1/2 = 6.1 min at pH 7.1) [176]. The caging group is released as the corresponding phenol.

750 GM (740 nm); u = 0.087 GM (780 nm) for BHQ-OAc u = 0.59

max = 327

nm

350 = 1500

AQMOC

350 = 0.1 (disappearance of starting material) in 50% aqueous THF 350 = 150

R = Me2N400 nm max 400 = 9077 (308–360; pH 7.4) =

DANP

0.03; (450; pH 7.4) = 0.002 * = 5 s (transient absorption change) [175]

41 BHC-diol is used to cage aldehydes and ketones. The quantum yields of uncaging are low, but the strong absorbance leads to high values. BHC-protected compounds can be uncaged by two-photon irradiation with a favorable cross section ( u).

BHQ-protected compounds can 40 be photolysed by two-photon irradiation. The cross-section at 740 nm is quite good. Proteins have not been caged with members of this class of reagents.

(365; pH 7.2) = 0.29

369 = 4100;

Refs.

RCOO-BHQ Caged carboxylates.

365

BHQ

Comment s

(Continued)




Comment s

CINN-OR to modify various serine proteases

177 Cinnamate ester derivatives, CINN-OR have been used to modify the catalytic Ser of various proteases. The nature of the ester used depends on the substrate specificity of the protease. The photochemical data from the different proteases are compared below, together with a model compound. The absorbance and, especially, the quantum yield vary in the microenvironment of the protein.

CINN-OX

(trans to cis) = 0.13 max = 358

nm (366, pH 7.4) = 19 200 366 2500 [177]

CINN-OEt Model ester.

Refs.

177

CINN-O-chymotrypsin Caged catalytic serine. nm (366, pH 7.4) = 25 700 366 4370 [177] * = 199 s (cyclization after photoisomerization) [106]

106, 177

CINN-O-factor Xa Caged catalytic serine. nm (366, pH 7.4) = 26 000 366 5980 [177] * = 151 s (cyclization after photoisomerization) [106]

106, 177

(trans to cis) = 0.17

max = 372


max = 358


CINN-O-thrombin Caged catalytic serine. nm (366, pH 7.4) = 18 600 366 740 [177] * = 287 s (cyclization after photoisomerization) [106] max = 380

106, 177


17




Comment s

Photoprotection of primary and secondary alcohols (X = O) or thiol (X = S).

2-Benzoylbenzoate derivatives 178, have been used to photoprotect 179 alcohols ROH (X=O) and 2-benzoylbenzenethioate derivatives to protect thiols RSH (X=S) [178]. Substituted benzyl 2-benzoylbenzoate derivatives have been used for the modification of the catalytic serine of various proteases, the -XR group varying according to the specificity of the enzyme (see Tab. 5.1.2) [179]. For caged thiols, the photodeprotection yield is 60%, together with 20% formation of disulphide. For ROH, the yield was variable, but generally good.

X = O: 2-benzoylbenzoate derivatives X = S: 2-benzoylbenzenethioate derivatives

Groups that have actually been used to cage proteins are emphasized, although promising potential alternatives are also given. Reagents that are available for directly caging proteins are shown with emphasis on those that can be used to derivatize nucleophiles, especially the thiolate side-chains of Cys residues. Key: λmax , maximum wavelength of absorption; ε xxx , extinction coefficient at XXX nm in M−1 cm−1 ; (xxx; pH Y) , quantum yield at XXX nm and pH Y. Unless otherwise stated, the product quantum yield (P ) is given, which is not always interchangeable with the quantum yield for uncaging (u ). In the text, we have often simply used for the purpose of general discussion; τ *, lifetime of photolytic intermediate, as defined in the text. δ u is the uncaging action cross-section, which is the product of the two-photon absorbance cross-section δ a and the uncaging quantum yield u2 . Ideally, δ u should exceed 0.1 Goeppert-Mayer (GM), where 1 GM is defined as 10−50 cm4 ·s photon−1 [15].

Fig. 3 Intermediates in the photolysis of two common caging groups: 2-nitrobenzyl (nitronate or aci-nitro intermediate) and 4-hydroxyphenacyl (spiroketone intermediate).

Refs.


have been obtained by comparison with other reagents for which the values of may in turn be of dubious origin. 2.3 The Rate of Uncaging: Breakdown of Intermediates

The argument about the half-time for photolysis (t1/2 ), given above, applies to cases in which the breakdown of the photogenerated intermediate is not rate limiting. In some applications, an intermediate might be long lived compared to the process under investigation. Such applications are unlikely to occur in cell biology, where the interest is usually in processes taking seconds to hours, but they can occur in biophysics, where for example there is renewed interest in examining protein folding on the submillisecond timescale using folding intermediates generated by flash photolysis [19–22]. Where they are available, values of the half-lives of caged reagents after the absorption of a photon, τ *, have been tabulated (Tab. 1). The reader must be cautioned to take a careful look at how the measurements were made. Very often the appearance and decay of an intermediate in a flash photolysis experiment is followed by absorption spectroscopy and this does not necessarily indicate the rate of product release (e.g. recent work on 2-nitrobenzyl chemistry [13, 14]). Indeed, the identity of the intermediate is not always clear. Rapid-scan FTIR has a higher information content and might prove to be a method of choice for determining photochemical reaction sequences in the future [13, 14, 23–26]. In some cases, secondary intermediates are expected. For example, unstable carbamic acids are formed upon photolysis of molecules caged with the nitrobenzyloxycarbonyl group. By using a single-molecule approach, we recently examined the lifetimes of both the (presumed) nitronate (aci-nitro compound), generated initially, and the derived carbamic acid at the surface of a protein [27]. The nitronate had a half-life of ∼2 s that was largely independent of pH, while the carbamic acid had a half-life of ∼3 ms at pH 5.5 and ∼5 s at pH 10. Clearly, these values are incompatible with, for example, rapid protein folding experiments. The rates of breakdown of photolytic intermediates can vary hugely even within the same class of reagents. For example, in the case of 2-nitrobenzyl reagents (Fig. 3), the nitronate of the 1-(3,4-dimethoxy-6-nitrophenyl)ethyl ester of glycine breaks down with a unimolecular rate constant of 1 s−1 [28], while the value for the nitronate of the corresponding α-carboxy-2-nitrobenzyl ester is 2 × 105 s−1 [29]. At present, the caging groups known to dissociate most rapidly after the absorption of a photon are the 4-hydroxyphenacyl group, τ * ∼1.2 ns [17, 30] and the benzoins [20], which cleave in < 1 ns. Photoinduced isomerizations can occur even more rapidly (see below). 2.4 The Effects of pH on Photolysis

Because there are often ionizable groups in caging reagents themselves or in photogenerated intermediates, both t1/2 and τ * can be affected by the prevailing


pH. The two possibilities are not always clearly distinguished in the literature. Effects on t1/2 arise because of effects of pH on and ε. The t1/2 values are not meaningful in themselves because I0 (the intensity of the photon source) can vary enormously. The substituted 7-hydroxycoumarins of Furuta et al. provide an example of an effect of pH on t1/2 . Ionization of the coumarins led to a decrease in t1/2 (365 nm), which was largely due to an increase in εu rather than u [15]. In the case of 2-nitrobenzyl derivatives, pH can affect u [3, 31]. In an example of an effect on τ *, the rate of breakdown of the nitronate intermediate in the photolysis of 2-nitrobenzyl derivatives has been reported to be pH dependent over up to four orders of magnitude in certain cases, with faster breakdown at low pH [32]. In other cases, the pH dependence is weak [27, 29]. All these factors, u , ε u , τ * and the effects of pH, will depend on the leaving group, which for proteins will often be the thiolate of a cysteinyl (Cys) residue (see below). Some measurements have been made on proteins derivatized on Cys or thiophosphate (Tab. 1), but more work is needed in this area. At present, the effects of pH should be considered to be unpredictable and the necessary exploratory experiments must be carried out for each new reagent caged protein.

2.5 Problems with the Released Caging Group

The released caging group, which is usually liberated in an altered form, must not interfere with the experiment that is underway. Therefore, the released group should be chemically inert and should not take part in further photochemistry, either by simply absorbing light and screening the remaining caged protein or by forming reactive intermediates. These issues are expected to be less of a problem in the case of cell biology experiments, where caged proteins are most often used at low concentrations (micromolar or less). In these cases, the photoproducts will be dilute and therefore do minimal damage, even if they are chemically reactive, and they will absorb little light, even if they have high ε values. The most widely used caging reagent for proteins has been the 2-nitrobenzyl group (Tab. 1). Its main drawback is that it is released as a reactive nitrosoal-dehyde or nitrosoketone, which has been shown to be deleterious to proteins by reaction with Cys residues in particular [25, 33, 34]. The nitroso compounds can be scavenged by the addition of a substantial concentration (> 10 mM) of a thiol such as dithiothreitol (DTT) or 2-mercaptoethanol, prior to photolysis [34]. Where the caged protein is activated inside a living cell, the millimolar concentrations of intracellular thiols (glutathione, Cys, etc.) will serve to scavenge the reactive products. Semicarbazide at 10 µM has also been used to scavenge 2-nitrosobenzaldehyde, but its efficacy was not documented in detail [35]. Other photoproducts may be more benign. For example, 4-hydroxyphenacyl groups are released as a spiroketone, which rearranges to form 4hydroxyphenylacetic acid, which is water-soluble, chemically inert and blue-shifted in absorbance compared to the caging reagent (Fig. 3) [17, 30]. A detailed

19


comparison of this and other aspects of the nitrophenylethyl and hydroxyphenacyl caging groups has been made [25]. Several chromophores generate reactive oxygen species upon irradiation in the presence of oxygen. Therefore, where possible, uncaging should be carried out in an inert atmosphere, e.g. under nitrogen or argon. Alternatively, scavengers of the reactive species should be used. Thiols are good all round scavengers, but singlet oxygen is more effectively quenched with azide or carotenoids [36–38]. Many of the deleterious effects of short-wavelength light [8, 9] might be eliminated by twophoton uncaging at long wavelengths and this approach is under investigation [15, 39–41]. 2.6 Photochemistry Peculiar to Proteins

While they have been treated here in the context of caged proteins, many of the phenomena described above also occur with small caged reagents. Certain phenomena that might affect the efficiency of photolysis are, however, peculiar to proteins (Fig. 4), e.g. the excited state of the caging group might be quenched by neighboring tryptophan. Conversely, it might be possible to uncage proteins by energy transfer to the caging group from tryptophan residues. This process has been used previously to activate photoaffinity reagents [42]. Other aspects of the microenvironment associated with caged residues in proteins include the local dielectric constant and pH. The dielectric constant can alter both the ground state absorption properties of a caging group, the photo-physics after a photon is absorbed and the subsequent dark chemistry. The importance of pH has been noted above with reference to reactions in bulk solution. However, the “apparent pK a ” of a group in, say, the active site of an enzyme can be quite different from the bulk value, if the local environment differs. In other words, the protonation states of ground states and intermediates may differ significantly when a caged group is buried within a protein. Protonation and deprotonation rates may even be slowed to the extent that their kinetics dominate the deprotection process. Steric hindrance to uncaging might also be of importance. For example, after a photon is absorbed, the caging group might not be able to rotate freely to attain

Fig. 4 The microenvironment at the site of caging in a protein can affect the photochemistry. For example, as depicted, the excited caging group might be quenched by energy transfer. Uncaging might be inhibited by restricted rotation. Local conditions such as pH and the dielectric constant might differ from bulk values.


Fig. 5 Active site photochemistry of 2hydroxycinnamoyl proteases. The enzyme (E-OH) is inhibited by acylation of the active site nucleophile, a serine side-chain. Upon photolysis, trans-cis isomerization oc-

curs such that the phenolic hydroxyl of the caging group can attack the carbonyl of the acyl enzyme. The caging group then cyclizes and is released.

the conformation that is required for bond rearrangement and cleavage (Fig. 4). From a practical viewpoint, these considerations indicate that it is not always possible to assess the properties of a caged protein from related solution photochemistry. Nevertheless, the circumstances suggest that, where possible, a small mobile group should be used to cage a protein. Further, aromatic amino acids, which are potential quenchers, especially tryptophan, should be removed by mutagenesis from the vicinity of the site of modification. Access to solvent will also be important if the photochemistry that occurs in bulk solution is to be approximated. In some cases, the uncaging mechanism relies on active-site chemistry, as in the hydroxycinnamoyl-proteases of Porter et al. (for recent work, see [43–45]) (Fig. 5). For caged enzymes, the photoproducts should be released quickly from the active site and fail to rebind. This can be a tricky issue where a light-removable covalent inhibitor has been directed towards the active site. For example, Cys199 of the catalytic subunit of protein kinase A (PKA), situated at the entrance of the active site, was modified in preference to Cys343, with a substrate-like peptide linked to an α-bromo-2-nitrobenzyl group [46]. While the millimolar affinity of the peptide was sufficient to discriminate kinetically between the two Cys residues in the alkylation reaction (caging), it was, as desired, too weak for the peptide to remain in the active site after photolysis (uncaging). Proteins are chiral and several caging reagents (Tab. 1) are attached through their own chiral centers forming diastereomers. Although the phenomenon has not been observed in practice, it is quite possible that the photochemistries of the two forms of caged protein differ. 2.7 Polypeptide Chain Cleavage

Occasionally, polypeptide chain cleavage has been noted during uncaging. For example, when the pore-forming protein α-hemolysin caged at Cys3 with 2-bromo2-(nitrophenyl)acetic acid (BNPA) was irradiated in the near-UV, an apparently shortened fragment was observed upon sodium dodecylsulfate-polyacrylamide gel electrophoresis (SDS-PAGE) [47]. While other explanations are possible for the shift

21


in electrophoretic mobility, such as intramolecular crosslinking, chain cleavage arising from scission at a site distant in the primary sequence is the most likely possibility. Less cleavage occurred when BNPA was attached at position 104 and in both cases the ratio of cleavage to deprotection was pH dependent [47]. In some cases, chain cleavage is intentional and desirable (see Section 3.10 and [48]). 2.8 Dominant Negative Effect

In the case of multisubunit proteins, “dominant negative” effects are a further consideration. Here, a small fraction of remaining caged protein or protein damaged during uncaging might remain in association with the properly uncaged protein and thereby reduce its activity. For example, in the case of caged α-hemolysin, which forms a heptameric pore, caging was about 60% complete as judged by SDS-PAGE, but only 10–15% of the pore-forming activity based on complete conversion was recovered [47].

3 Sites of Modification in Caged Proteins 3.1 Random Modification

Surprisingly, random modification has been used to cage proteins successfully [49]. For example, antibodies were treated with the oxycarbonylchloride of 1-(2nitrophenyl)ethanol (NPE), which primarily reacts with the e-amino groups of lysine (Lys) residues. When an Fc-specific anti-human IgG was “coated” with 30 NPE groups and the remaining Fc-binding fraction removed by affinity chromatography, a 15% recovery of IgG was obtained in which the specific Fc-binding activity was reduced to 1.5% of the original value. Upon irradiation with near-UV light, the specific binding activity increased to 29% of the original value. 3-Nitro-2naphthalenemethanol has been used in much the same way [50]. For the unconjugated alcohol, u = 0.63 and ε u = 810 M−1 cm−1 (u ε u = 510) in aqueous solution at 380 nm. In this case, the Protein A-binding activity of caged IgG increased from 18% to about 70% upon irradiation and then decreased upon further exposure. While simple in practice, the potential problems of random modification are clear. First, when the target protein has multiple functional domains, all of them will be affected by the caging reaction. Second, because multiple sites are likely to be modified by the caging reagent, uncaging will be a complicated process involving multiple photons, a lag phase and a complex mixture of intermediates with partly restored activity (Fig. 6). If the overall efficiency of removal of a caging group is E (the remaining positions being damaged in some manner), the final extent of uncaging will be ∼E n , where n is the number of groups incorporated initially. However, the effect of multiple sites of caging on the final activity is actually less

3 Sites of Modification in Caged Proteins

Fig. 6 Caging a protein by random modification. A two-domain protein is shown here. The protein is caged by “coating” with many protecting groups. Photolysis occurs through numerous partly deprotected inter-

mediates that may or may not be active. In the case of a two-domain protein, intermediates may be formed in which one domain is active and the other is not.

clear as each position at which caging takes place may have a different value of E and make a different contribution to the activity of the protein. On the positive side, a requirement for activation by multiple photons can aid in the spatial localization of uncaging (see Section 5.3). 3.2 Active-site Directed Modification

In an approach pioneered by Porter et al. [51], enzymes have been caged by making use of active-site chemistry. Various proteases have been inactivated as trans-2hydroxycinnamoyl acyl enzymes by using ester substrates (for recent work, see [43–45]). Upon photolysis, the caging group undergoes trans–cis isomerization and the product then lactonizes in a dark reaction liberating the active site (Fig. 5). In a recent manifestation, the approach has been used to separate a specific enzyme from a mixture of proteases. In this case, the leaving group in the ester substrate confers enzyme specificity (it is an “inverse” substrate) and the acylating group contains an arm carrying biotin. The modified enzyme is extracted from the mixture with an avidin column and then released from the column by irradiation. Tagged acylating agents have been used for proteomic screening [52] and it is possible that photorelease might be useful in this context. Additional active-site-directed reagents have been developed, e.g. the substratelike peptide that reacts with a Cys residue at the entrance to the catalytic site of PKA [46]. However, there may not always be a need to tailor a reagent for an enzyme active site, as the latter often contain hyper-reactive residues associated with the catalytic activity. For example, the serine at the active site of a butyrylcholinesterase can be selectively reacted with the carbamylating reagent N-methyl-

23


N-(2-nitrophenyl)carbamoyl chloride [53] and the catalytic Cys of a protein tyrosine phosphatase can be modified with various α-haloacetophenones, despite the presence of a total of four Cys residues in the protein [54]. It would seem likely that a modification strategy developed for one protein would be applicable to other members of its family, but this is not always so. For example, N-methyl-N-(2-nitrophenyl)carbamoyl chloride efficiently inhibits both acetylcholinesterase and butyrylcholinesterase. However, only butyrylcholinesterase recovers activity efficiently upon irradiation [53]. 3.3 The Use of Crosslinkers

A photocleavable, 2-nitrobenzyl, crosslinker was used to connect a ribosome-inactivating protein (PAP-S) to a monoclonal antibody or to the B-chain (lectin subunit) of ricin for delivery to cells [55]. Attachment to PAP-S was through a chloroformate and non-specific. A sulfhydryl group at the other end of the crosslinker was then deprotected for reaction with the monoclonal antibody, which had been premodified to carry maleimide groups. Alternatively, ricin B-chain was attached through another crosslinker carrying a maleimide. The conjugates were active towards cells in culture, but the activity was increased more than 20-fold in a plating efficiency assay after irradiation in situ at 350 nm, presumably because the liberated PAP-S has greater access to ribosomes (Fig. 7). This promising approach does not appear to have been pursued further.

Fig. 7 Application of a photocleavable crosslinker. (Left) A small toxic protein (PAP-S) is coupled in two steps to a cellspecific antibody through a photo-cleavable crosslinker. (Right) The toxin–antibody con-

jugate recognizes target cells and is internalized, e.g. by endocytosis. Photocleavage of the crosslinker increases the toxicity of the internalized conjugate [55].


There are many other potential applications of photocleavable crosslinkers for proteins. For example, they might be used to for the patterning of surfaces, including protein chips. One important possibility, for reagents carrying biotin on one arm [56], is in the purification of caged proteins. It has already been noted that it is often impossible to completely derivatize a protein with a caging reagent. With a biotinylated caging reagent, it would be possible to extract the derivatized fraction of the protein and this has been explored in the particular case of proteases by using active-site directed reagents [44]. The approach might be generalized by using Cys-directed biotinylated reagents to cage mutated proteins (see below). 3.4 Caging at Cys Residues

The deprotonated thiolate side-chains of Cys residues are by far the most nucleophilic groups in natural polypeptides. Therefore, under normal circumstances, a Cys residue will be selectively modified by an electrophilic caging reagent, even in the presence of hundreds of other reactive side-chains. “Normal circumstances” assumes that the Cys is not buried or that there is not a hyper-reactive active-site nucleophile present in the target protein. The unperturbed pK a value of a Cys residue in a polypeptide is 9.0–9.5. The next issue then is to find or place a suitable Cys residue in the target protein (Fig. 8). In general, Cys residues are rarer than Lys residues and simply “coating” the protein with a Cys-reactive caging reagent is unlikely to work. Generally, a Cys residue in or close to a key functional site in the protein is required and can be introduced by mutagenesis. Occasionally, such a Cys residue will occur naturally; Cys residues are crucial active-site nucleophiles in several classes of enzyme including various proteases and phosphatases. For example, a protein tyrosine phosphatase has been caged with 4-hydroxyphenacyl bromide at the

Fig. 8 Flow chart for caging proteins at Cys residues.

25


active-site Cys [54]. In other cases, a Cys residue in proximity to the active site may suffice and, where a structure is available, the residue is most readily identified by molecular graphics. For example, the catalytic subunit of PKA has been caged at Cys199, which lies near the entrance to the active site and in the so-called activation loop (PKA is activated by phosphorylation at Thr197). Care should be taken when caging enzymes at the periphery of the active site, because not all substrates may be affected equally by the covalent modification. Where structural information is unavailable or unhelpful, cysteine-scanning mutagenesis can be a useful approach. Although it is tedious, the various spinoffs, e.g. mechanistic information and sites for the attachment of fluorescent probes, often make it worthwhile. Cys-scanning mutagenesis was performed on the pore-forming protein α-hemolysin (αHL) before the crystal structure had been determined and when little was known about the assembly mechanism. Wild-type αHL contains no Cys residues. Eighty-three mostly charged residues in the 293-residue polypeptide were changed individually to Cys, and the poreforming activity of the protein was determined before and after modification at Cys with the bulky charged reagent 4-acetamido-4 -((iodoacetyl)amino)stilbene-2,2 -disulfonate, IASD [57]. The information gained in this way proved useful when it came to producing a caged aHL polypeptide [47]. Twenty-eight of 83 modified Cys mutants were substantially inactivated. Through various other negative criteria, including reduced activity before modification, incomplete inactivation by IASD, or a failure to be inactivated by a smaller reagent, the candidate positions were narrowed to four. One of these, Arg104, was investigated in detail and performed well when derivatized with the water-soluble caging reagent BNPA [47]. Subsequent structural information [58, 59] revealed that Arg-104 is in a region that undergoes a critical conformation change, including a 180◦ rotation at the ψ (Cα –C) angle of this residue. In principle, proteins might be inactivated by modification at locations other than an active site. For example, it might be possible to use a caging group to prevent a protein-protein interaction, which could be reestablished upon photolysis. In practice, however, this may be more difficult than it appears. For example, a nitrobenzyl group located at the dimer interface in HIV protease did not prevent the interaction between the subunits [60]. In most cases, it is likely that a bulky reagent will be required for this purpose. For example, polymeric Cys-directed caging reagents might be developed [61] or biotinylated reagents [44, 56] might be used to block the interface after coupling to streptavidin. An additional means of generating Cys residues at specific sites is by chemical ligation of polypeptide fragments, a methodology that is gaining ground for the total synthesis of proteins [62]. In this approach, two or more synthetic or recombinant fragments undergo chemical coupling. In general, the N-terminal fragment terminates in a reactive thioester at the C terminus and the C-terminal fragment contains an N-terminal Cys. For example, active channel proteins have been made in this way [63, 64]. After spontaneous ligation, the new polypeptide contains a Cys residue at the ligation site, which is available for chemical modification with a caging reagent. In this section, the question of how to handle more than one Cys residue in the target protein has been neglected. As noted above, if several caged residues


affect activity, the regeneration of activity upon photolysis will be a non-linear process. Because Cys residues are rare, this will not occur as often as it might with say non-specific modification at Lys. Again, because Cys is rare, it will often be possible to eliminate potentially offending residues by mutagenesis [65] (Fig. 8). In cases of extracellular proteins, Cys residues will often be present as disulfide bonds. The introduction of an additional Cys residue might then present a problem with disulfide scrambling, but often this can be circumvented by refolding in a redox buffer to encourage the formation of the natural disulfides and leave the new residue as a free sulfhydryl. Despite these issues, it is remarkable what can be achieved under potentially difficult circumstances. For instance, each 135 000 Da subunit of the β-galactosidase tetramer contains 16 Cys and 23 Met residues and yet it appears to be inactivated by the attachment of one 1-(4,5-dimethoxy-2nitrophenyl)-2-nitroethene reagent per subunit [66]. Because the alkylation reaction is not reversed by 2-mercaptoethanol, the modification is unlikely to be at the activesite Met502. Upon irradiation, 89% of the enzyme activity was recovered.

3.5 Sites other than Cys for Site-specific Modification

Reactive side-chains other than Cys residues can be introduced into proteins for sitespecific modification. For example, thiophosphate groups in peptides and proteins can be derivatized with various caging reagents. The thiophosphate group is an especially useful target because the activities of many cell-signaling proteins are controlled by phosphorylation. Thiophosphorylation can be carried out at serine and threonine by using the appropriate protein kinase and ATPγ S [adenosine 5 O-(3-thiotriphosphate)] [7]. In the case of tyrosine, the enzymatic phosphorylation conditions must be manipulated by the substitution of alternative divalent metal ions for magnesium [31]. Interestingly, the thiophosphate group is often a poor substrate for phosphatases, so proteins activated by thiophosphorylation rather than phosphorylation are expected to have longer lifetimes in vivo. A second useful feature of the thiophosphate group is that it can be alkylated at low pH in the presence of sulfhydryls [12]. The second pK a value of a thiophosphate monoester is ∼5.5 [67] and therefore at pH 4–5 the reactive monoanion will predominate, while Cys residues (pK a ∼9.0) will be protonated. With groups of similar nucleophilicity, selective modification can be obtained where pK a ∼3 or more. In principle, the wide difference in reactivity between a thiophosphoryl and sulfhydryl group at low pH might allow two different groups to be placed on a protein, e.g. a caging group and a fluorescent probe. Therefore, it is worth considering additional residues that might react orthogonally. A prime candidate is selenocysteine (Sec), which when compared with Cys has a greatly reduced pK a value (estimated to be 6.0–6.5 in a polypeptide chain [68]) and enhanced nucleophilicity [69, 70]. Sec can be introduced into proteins in vivo at TGA codons (although, in bacteria, the required cis signal disrupts the coding region) [71], by chemical ligation [70, 72] or, potentially, by unnatural amino acid mutagenesis. Interestingly, the relative rate of reaction of a selenophosphate over a thiophosphate monoester

27


towards an alkyl iodide has been examined at pH 7.0 and found to be only ∼ 3.5-fold higher [73]. More complex motifs can be derivatized even more selectively, as demonstrated by the work of Tsien et al. on tetracysteine (TC) sequences that can be reacted with bisarsenicals [74–76]. Reaction at these sites is sufficiently selective that proteins bearing them can be derivatized in vivo, although the generality of the approach for all types of cells has been questioned [77]. It is possible, for example, that caged TC motifs might be used to prevent protein–protein interactions in living cells. Other functionalities that might be useful for site-specific modification can be introduced by unnatural amino acid mutagenesis, and include azides [78, 79] and ketones [80]. 3.6 Reagents for Caging by Protein Modification

It will be clear from the discussion above that the overwhelming majority of useful caging reagents for proteins are electrophilic molecules that modify nucleophilic amino acid side-chains (Tab. 1). Preferably, a reagent should be soluble in aqueous media. However, poorly soluble reagents can often be added from a concentrated solution in a water-miscible organic solvent, provided that the final concentration of organic solvent remains low ( 1000-fold reduction in VDX responsiveness and is no longer regulated by physiological concentrations of the hormone. There have been two reports of the use of the VDR structure to design selective ligands that bind the mutant receptor in preference to the wild type VDR [22, 23].

3 The Vitamin D Receptor-Ligand Complexes

In one study, it was proposed that the R274L mutation removed a polar interaction between the protein and the natural ligand VDX and created a hydrophobic hole in its place. SS-III, a derivative of VDX, was designed to fill that hole. It was prepared and found to be 286 times more potent than VDX against the mutant protein [22] (Fig. 18). In another study, computER-αided molecular design was used to generate a focused library of nonsteroidal analogues of the VDR agonist LG190155 (Fig. 19) that were uniquely designed to complement the R274L protein associated with Vitamin D3-resistant rickets. Half of the designed analogues exhibit substantial activity in the R274L mutant. The seven most active designed analogues (such as A-11; Fig. 19) were more than 16 to 526 times more potent than VDX in the

Fig. 18 (a) The R274L VDR mutation fails to bind the natural ligand VDX. (b) The ligand SS-III was designed and found to bind to this mutant protein.

23

24 Molecular Recognition of Nuclear Hormone Receptor-Ligand Complexes

Fig. 19 Structures of ligands for VDR R274L mutant.

mutant receptor. Significantly, the analogues are selective for the nuclear VDR and did not stimulate cellular calcium influx which is associated with activation of the membrane-associated vitamin D receptor (Fig. 19) [23].

4 The Retinoic Acid Receptors RAR and RXR 4.1 Introduction

Retinoid acids (RAs), the active retinoid derivatives of vitamin A regulate complex gene networks The pleiotropic effects of active retinoids are transduced by their cognate nuclear receptors retinoid X receptors (RXRs) and retinoic acid receptors (RARs) which act as transcriptional regulators activated by two stereoisomers of retinoic acid (RA): 9-cis RA (9cRA) and all-trans RA (atRA). Among the nuclear receptors, RXR occupies a central position and plays a crucial role in many intracellular signaling pathways as a ubiquitous heterodimerization partner with numerous other members of this superfamily Whereas RARs bind both retinoic acid isomers RXRs exclusively bind 9cRA The various RAR (RAR-α, -β and -γ ) and RXR (RXR-α -β and -γ ) isotypes are encoded by different genes while their multiple isoforms which differ in their N-terminal region are the results of differential promoter usage and alternative splicing It has been shown that RXRs play a unique role among NHRs since they are able to heterodimerize with a number of members of the NHR superfamily [24] This section will give an overview of the structural biology of retinoid receptor-ligand complexes focusing on the role of the ligand.

4 The Retinoic Acid Receptors RAR and RXR 25

4.2 RAR-γ and RXR-α Retinoid Complexes

The structure of atRA bound to RAR-γ was one of the first reported structures of the ligand-binding domain of an NHR bound to a ligand (PDB entry: 2LBD) [25, 26]. Like the steroid receptors the hydrophobic atRA ligand is buried, but fits loosely, within the ligand binding hydrophobic core of the protein. The protein adopts a typical agonist-bound conformation with helix-12 folding over, and enclosing, the ligand. The carboxylate group of atRA makes hydrogen bonds with both the backbone NH and side chain OH of Ser-289. Ser-289 makes two hydrogen bonds to the guanidine side chain of Arg-278. The carboxylate group of atRA also forms a hydrogen bond with a structural water molecule which is also hydrogen bonded to the backbone carbonyl of Leu-233 Arg-278 is located at the end of helix-5 and Ser-289 is located on a turn between helix-5 and helix-6 (Fig. 20). Comparing the agonist-bound structures of RAR-γ and ER-α shows some significant differences There is a significant change in the positions of helices-1, -3, -6, -7 and -9, and a small change in the positions of helices-11 and -12 relative to helices-4 and -5 In addition the protein conformation between the end of helix-5 and the beginning of helix-7 is very different between ER-α and RAR-γ As with ER-α there is an arginine at the end of helix-5 (Arg-278 in RAR-γ Arg-394 in ER-α) Glu-353 on helix-3 of ER-α is replaced by Cys-237 in RAR-γ Unlike ER and the other steroid receptors this residue in RAR-γ (Cys-237) does not appear to play a key role in ligand recognition As was seen for VDR changes in the gross structure of the protein result in changes in the relative positions of the helices. These changes alter the size and shape of the binding pockets to better accommodate the appropriate ligands. Clearly different NHRs have evolved structures that are complementary to their respective ligands. RAR-γ is a permissive receptor and will also bind 9cRA as an agonist; the structure of the protein in this complex (PDB entry 3LBD) [27] is nearly identical to that

Fig. 20 Schematic of interactions between atRA and RAR-(.


in the atRA complex. The polar carboxylate group of 9cRA interacts with RAR-γ in almost the same manner that atRA does and the rings of 9cRA and atRA fill the same hydrophobic binding site. However, the less than perfect fit of these two ligands to RAR-γ allows the tetraene chain to adopt different conformations (20-methyl group on opposite sides of the pocket) in the two complexes. In each complex the 19-methyl group occupies approximately the same space. The difference in chain conformations, coupled with a loose steric fit, allows for either a cis or trans olefin to be accommodated at the 9–10 position (Fig. 21). In contrast to RAR, RXR can select 9cRA in preference to atRA. The structure of the RXR-α/9cRA complex has been reported (PDB entry 1FBY) [24]. This structure exhibits the typical agonist protein conformation with helix-12 folded over the ligand. The carboxylate of 9cRA forms hydrogen bonds to the backbone NH of Ala-237 and the guanidine group of Arg-316. In addition, the carboxylate makes a hydrogen bond to a water molecule; that water forms hydrogen bonds to the backbone carbonyl of Leu-309 and to another water that is hydrogen bonded to the side chain of Gln-275. Arg-316 is located at the end of helix-5 and is in the equivalent position to Arg-278 in RAR-α and Ala-237 is in the equivalent position

Fig. 21 (a) Schematic of the interactions between 9cRA and RAR-(. (b) Superposition of atRA (black) and 9cRA (gray) bound to RAR-(.


Fig. 22 Schematic of the polar interactions between 9cRA and RXR-". The shape of the RXR-" protein is complementary to 9cRA and explains its selectivity.

to Ser-289 of RAR-α (Fig. 22). Of interest, Gln-275 is located on the same position of helix-3 as Gln-725 in PR. As discussed previously this glutamine is conserved in several other steroid receptors that bind 3-keto steroids and is equivalent to Glu-353 in ER-α. Recall that in the steroid receptors this residue plays a key role in A-ring recognition. However, in RXR-α this glutamine residue is solvent exposed and makes an indirect polar interaction with the ligand. The hydrophobic ligandbinding pocket has a distinct L-shape to it. The cis-olefm of 9cRA allows the ligand to bend and accommodate this binding site. While the fit of 9cRA to RXR-α is not precise, the shape of the binding site precludes the binding of incorrect ligands such as atRA. This readily explains why RXR-α is selective for 9cRA as opposed to at RA. 4.3 Selectivity of RAR Ligands and RAR Isotypes

A very elegant experiment has been reported where enantiomers of an unnatural RAR-γ -selective ligand have been determined [28]. Of special significance is the observation that one of the enantiomers is effectively inactive in the biochemical assays. While acquiring co-crystal structures of many active ligands with the same protein is now commonplace, obtaining structures of inactive (i.e., weakly active) compounds is more challenging. This work provides a high-resolution structure (PDB entry 1EXX) of the complex between an inactive compound (BMS270395, Kd ≈ 500 µM) and RAR-γ . The structure (PDB entry: 1EXA) of the more potent en-antiomer (BMS270394, Kd = 500 nM) and RAR-γ was also determined for direct comparison (Fig. 23). The conformational profiles and energies of the two enantiomers are, by definition, mirror images of each other. Thus, the ligand conformational arguments made by Klaholz et al. [28] are not correct. If one superimposes the mirror image of


Fig. 23 Structures of enantiomeric ligands BMS270394 is a 500 nM RAR-( ligand while its enantiomer BMS270395 is a very poor RAR-( ligand.

the inactive enantiomer on the active isomer it is apparent that the bound conformations of the two ligands are similar. The A-rings (except for the fluorine), amide groups and alcohol moiety superimpose almost exactly. The major difference in the conformations is the torsion angle (C O)-CH-C(Ar)-C(Ar); it is 53◦ and −102◦ for the active and inactive enantiomer, respectively. A quick MM2 calculation predicts a 119◦ torsion angle is a low energy local conformational minimum [29]. These calculations suggest that it is unlikely that the conformation energy difference between the active and inactive enantiomers is close to 4 Kcal mol−1 . It would appear that ligand conformational strain does not play a dominant role in the large difference in affinity between the two enantiomers. Thus, the major component of the large difference in affinity between these two isomers must come from protein-ligand interactions. The structure of the more active enantiomer (BMS270394) shows that the ligand binds to RAR-γ in a typical agonist conformation (Fig. 24). The carboxylate group of the ligand forms hydrogen bonds to Ser-289 and a water molecule in the same

Fig. 24 Key interactions between the active enantiomer BMS270394 and RAR-(. A key interaction involves a hydrogen bond between the ligand hydroxyl group and the sulfur of Met-272. Notice that the linking amide group makes complementary interactions with the enzyme.


fashion that atRA does. The hydrophobic CD ring system makes VDW contacts in the hydrophobic ligand-binding core. The hydroxyl group of the ligand makes a hydrogen bond (3.2 Å) with the thioether of Met-272. The amide group also interacts well with the protein, the NH hydrogen bonds to the backbone carbonyl of Leu-271 and the amide carbonyl interacts with three aromatic H atoms; two of these interactions are much shorter than the third. The shortest of these interactions (3.17 Å) is with Phe-230. The interactions of carbonyl oxygens with aromatic hydrogens are well precedented, favorable interactions in protein-ligand complexes. These are weakly polar interactions between the positive dipole of an aromatic C H bond and the electrons of a carbonyl oxygen. The fluorine atom exhibits short VDW contacts with the Cα and Cβ atoms of Ala-234. For the more active isomer, all of the ligand’s polar groups make complementary polar interactions with the protein, the fluorine atom comes into close VDW contact with the protein, and the hydrophobic groups are buried in hydrophobic parts of the protein. The structure of the weakly active enantiomer (BMS270395) shows that the ligand also binds to RAR-γ in a typical agonist conformation (Fig. 25). The carboxylate group of the ligand makes hydrogen bonds to Ser-289 and a water molecule in the same fashion that BMS270394 does. The hydrophobic CD ring system creates VDW contacts in the hydrophobic ligand-binding core in much the same manner as BMS270394. In addition, the hydroxyl group of the inactive ligand also forms a strong hydrogen bond (3.2 A) with the thioether of Met-272. If the "inactive" and the "active" enantiomer adopted the same bound conformation, the hydroxyl group would still have had room to fit, indicating that the interaction between the hydroxyl group and the methionine is quite significant in determining the structure of the complex. Thus, the "inactive" enantiomer makes many of the same interactions with the protein that the “active” enantiomer does. The major differences between

Fig. 25 Key interactions between the "inactive" enantiomer BMS270395 and RAR-(. Like the active enantiomer, a key interaction involves a hydrogen bond between the ligand hydroxyl group and the sulfur of Met-

272. In this isomer the amide group does not make favorable interactions with the protein and the A-ring shows some disorder in the positions of the fluorine atoms.


the two are the position of the linking amide group and the position and occupancy of the fluorine atom and the conformation of the ligand. In the "inactive" enantiomer complex, the amide group does not make complementary interactions with the protein, and unlike the "active" enantiomer, the polar amide group is buried in the protein without making favorable compensatory interactions. Also, unlike the "active" enantiomer, BMS270395 shows two different orientations for the fluorine atom, in both orientations there are sub-optimal interactions between the fluorine and the protein. While the aromatic A-rings are in nearly the same position in both ligands, subtle differences in the position of the A-ring appear to be responsible for the disorder in the fluorine atoms. Furthermore, while the bound conformations of the two ligands are different, as discussed above, the conformational differences are not likely to be too severe. It appears that the burial of the polar amide and the less than optimal fit of the fluorine atom are the key determinant of enantiomer selectivity. Both of these structures show a hydrogen bond to the thioether of Met-272. Clearly, this is an important recognition element for binding to RAR-γ . In both RAR-α and RAR-β, the residue corresponding to Met-272 is an isoleucine. BMS270394 is much less potent against both RAR-α and RAR-β. While the structure of BMS270394 with either RAR-α or RAR-β has not been reported, one can speculate that the burial of the hydroxyl group near the isoleucine is responsible for the weaker affinity of these ligands for these isotypes. A paper has been published that discusses the structural basis for isotype selectivity of the RARs [30]. This paper reported structures of three RAR ligands bound to RAR-γ , BMS184394 (PDB entry: 1FCX), an RAR-γ -selective agonist, CD564 (PDB entry 1FCY), an RAR-β/γ co-agonist and BMS181156 (PDB entry 1FCZ), a panagonist (Fig. 26). Residues that directly contact the ligand are mostly conserved

Fig. 26 Structures of RAR ligands. BMS184394 is an RAR-(-selective ligand, CD564 is an RAR–$/( co-agonist, and BMS181156 is an RAR panagonist.


in all three isotypes. The only differences are that Met-272 in RAR-γ is replaced by an isoleucine in both RAR-α and -β, and that Ala-234 in RAR-γ is replaced by a serine in RAR-α. BMS181156 has approximately the same activity against all three RAR isotypes and it is the most potent of these three ligands against RAR-γ . The structare of its complex bound to RAR-γ shows the bicyclic ring system bound in a hydrophobic site similar to BMS270394 and the carboxylate interacting with Ser-289 in the same manner as other RAR-γ ligands. This ligand binds to RAR-γ in a manner similar to 9cRA with the carbonyl group of BMS181156 occupying the same space as the C19 methyl group of 9cRA. In the BMS181156 complex, the carbonyl group makes close contacts with the Cε atom of Phe-304 and the Cε atom of Met-272. These close contacts have been described as C H O C hydrogen bonds [30]. The conformation of the Met-272 side chain is different in this complex to that in the BMS270394 and BMS270395 complexes, in the BMS181156 complex the sulfur atom does not interact directly with the ligand. CD564 shows comparable affinity towards RAR-γ and RAR-β but is 40-fold less potent against RAR-α. It is 5-fold less potent against RAR-γ than BMS181156. The structare of its complex bound to RAR-γ shows the bicyclic ring system bound in a hydrophobic site similar to BMS181156 and the carboxylate interacting with Ser-289 in the same manner as other RAR-γ ligands. The carbonyl group of CD564 makes a close contact with Cζ of Phe-304, which is in a different conformation than it is in the BMS181156 complex. The loss in potency against RAR-α is attributed to a steric clash between the naphthalene ring of the ligand and a serine side chain. In RAR-γ (and RAR-fl), this residue is Ala-234 and, in the complex structare, this residue is in close contact with the naphthalene ring. For BMS181156 the acyclic linker avoids this steric clash allowing potent binding to RAR-α. BMS184394, a mixture of enantiomers, is 100-fold less potent than BMS181156 against RAR-γ ; however, it shows 10-fold selectivity over RAR-β and 100-fold selectivity over RAR-α. This ligand binds to RAR-γ , as a single enantiomer, in a manner nearly identical to CD564. The major difference in the ligand is the difference between a ketone and an alcohol. As with BMS270394 and BMS270395, the alcohol group of BMS184394 forms a hydrogen bond to the thioether of Met-272. The side chain of Met-272 is in the same conformation as it is in the BMS270394 and BMS270395 structures. Thus, this side chain adopts different conformations depending upon the presence or absence of a hydrogen bond between the ligand and the thioether. The data suggests that the interaction of a ligand alcohol group with the thioether of Met-272 plays a key role in RAR-γ selectivity. The significant loss of affinity upon going from a ketone (CD564) to an alcohol (BMS184394) suggests that the observed C H O C hydrogen bonds in the ketones play an important role in stabilizing the complex and the alcohols cannot take advantage of this type of interaction. For RAR-γ , the alcohol-containing ligands can gain sufficient affinity by forming a hydrogen bond with Met-272 and thus becomes a selective ligand. SR11254, the oxime of CD564, is a potent (4 nM) RAR-γ -selective agonist [31]. Its structure in complex with RAR-γ (PDB entry 1FD0) shows that it binds in a


Fig. 27 Key interactions of the RAR-(-selective agonist with RAR-(.

manner nearly identical to CD564 (Fig. 27) [32]. The ligand binds as a ∼1:1 mixture of E and Z oximes. In both isomers, the hydroxyl group forms a hydrogen bond to the Met-272 thioether. The side chain of Met-272 resembles the conformation found in other complexes in which the ligand forms a hydrogen bond to the Met272 side chain. In this structure, close contacts between the oxime oxygens and protein methyl groups are observed. The Z-isomer interacts with the CE methyl group of Met-272 and the E-isomer interacts with the Cγ 2 methyl group of Ile-275. These have been characterized as C H O hydrogen bonds and are proposed to help stabilize the complex. The Klaholz and Moras paper is also noteworthy for its discussion of C H O hydrogen bonds [32]. The above studies provide a structural explanation for how very similar NHRs (i.e., the isotypes of RAR) can selectively bind different ligands. In each case a key polar interaction (ligand carboxylate) and the burial of a substantial amount of hydrophobic surface allows binding to occur. Tight binding occurs when there are no bad steric or electrostatic interactions. The case of enantiomer selectivity (BMS270394 vs. BMS270395) occurred when only one of the enantiomers could bury an amide group and make complementary interactions. RAR-γ and RAR-β differ by only one amino acid group in the ligand-binding pocket (RAR-γ Met272 is an lie in RAR-β). Ligands that can form a hydrogen bond to the thioether of Met-272 tend to be selective for RAR-γ . RAR-β and RAR-α differ by only one amino acid group in the ligand-binding pocket (RAR-γ Ala-234 is an Ala in RAR-β and a Ser in RAR-α). Selectivity for RAR-β over RAR-α can be achieved with

5 PPAR: Isotype-Selective Ligands

Fig. 28 Structures of RXR ligands.

ligands that make close VDW contacts with the side chain of Ala-234 in RAR-γ (or RAR-β). These ligands presumably bind poorly to RAR-α due to a bad steric contact. Thus, it is possible to achieve selectivity against very similar isotypes of the same NHR and it is possible to rationalize these selectivities based on structure. It is likely that in the future subtle selectivity will be rationally designed into ligands. 4.3.1 RXR Complexes with Unnatural Ligands The structures of two structurally related RXR-selective ligands have been reported. The complex between BMS-649, RXR-α and a co-activator peptide (PDB entry 1MVC) [33] and between LG-268 and RXR-β (PDB entry 1H9U) [34] are similar and will be discussed briefly below. The carboxylate group of both ligands interacts with the backbone NH of Ala-327 (RXR-α, Ala-398 RXR-β) and Arg-316 (RXR-α, Arg-387 RXR-β in the same manner that 9cRA does. The D-ring binds in the same place that the ring of 9cRA does (Fig. 28). The shape of the ligand is complementary to the L-shaped RXR ligand-binding pocket. This shape complimentarity explains why these ligands are selective for RXR over RAR. One significant difference between these two structures is the position of helix12; in the complex between RXR-α, BMS-649 and a co-activator peptide, helix-12 is in the standard agonist position. In the complex between LG-268 and RXR-β helix-12 is in a novel position not seen in other NHR structures. In this position LG-268 does not fold over and cap the ligand. Additional data indicate that LG-268 is unable to release co-repressors from RXR unless co-activators are also present; this suggests that certain RXR ligands alone may be inefficient at repositioning helix-12 [34].


Peroxisome proliferator-activated receptors (PPARs), are members of the steroid/retinoid nuclear receptor family of ligand-activated transcription factors. There are three isotypes of the PPAR family, PPAR-α, PPAR-δ and PPAR-γ . Various natural fatty acids (FAs) and eicosanoids serve as ligands for the PPARs.

33


Interestingly, several unsaturated FAs that activate the PPARs in vitro have pharmacological effects similar to those reported for the synthetic PPAR ligands. Based upon these observations and the promiscuous ligand-binding properties of the PPARs, it has been suggested that these receptors serve as physiological sensors of lipid levels, linking FA concentrations to glucose and lipid homeostasis [35]. The biological profiles of each have led to a number of ligated structures having been reported. This section will focus upon how ligands can select between these three isotypes. Unlike the RARs, there are several amino acid differences in the subtypes that directly contact the ligand. Thus, there are more options for obtaining isotype selective ligands and it may be more challenging to obtain ligands that show high affinity for all isotypes. The structure of the FA eicosapentaenoic acid (EPA) bound to PPAR-δ has been determined (PDB entry 3GWX) [35]. The ligand-binding pocket of PPAR-δ assumes roughly a "Y" shape with each of the three arms approximately 12 Å in length. One arm of the Y is composed of a mix of hydrophobic residues and polar residues and is capped by two residues from helix-12. This is the only arm of the Y that is substantially polar in character. Another arm of the Y is composed of hydrophobic residues and is sealed by the last residue of helix-1, and the loop between helices-1 and -2. The third arm of the Y-shaped pocket is composed mainly of hydrophobic residues with a few polar residues. The interior of the ligand-binding pocket is accessible via a channel that is exposed to solvent [35] The EPA complex crystal has two molecules in the asymmetric unit; each independent molecule has two distinct conformations for the EPA ligand (Fig. 29). For EPA in each complex molecule, the acid group and the first eight carbons adopt very similar conformations. The hydrophobic tails of EPA diverge in the different bound conformations with each occupying a different arm of the Y. The carboxylate head group of EPA interacts with several polar residues in one arm of the Y. Surprisingly, the details of these interactions are different in each of the four copies of the EPA in the crystal. There are three polar residues that interact with the carboxylates, His-323 from helix-5, His-449 from helix-11 and Tyr-473 from helix-12. His-449 forms a buried hydrogen bond to the side chain of Lys-367. PPARA and the PPARs in general, use a helix-12 residue to make a specific hydrogen bond to agonist ligands, unlike many of the other NHRs. The structure of the fibrate ligand GW2433, a PPAR-α/δ co-agonist, has also been reported bound to PPAPwJ (PDB entry: 1GWX; Fig. 30) [35]. In the structure, the carboxylate forms hydrogen bonds to two histidines (His-323 and His-449) and to Tyr-473 from helix-12. The rest of the molecule makes substantial VDW contacts with all three branches of the Y-shaped ligand-binding pocket. These are mostly hydrophobic interactions between ligand and protein; however, a hydrogen bond between the ligand urea NH and the side chain of Cys-285 is observed. While GW2433 is a PPAR-α/δ co-agonist, the structure of its complex with PPAR-α has not been disclosed. As will be discussed below, the residue corresponding to His-323 in PPAR-α is a tyrosine (Tyr-314). AZ-242 is a PPAR-α/γ co-agonist and its structure has been determined in complex with both PPAR-α (PDB entry 1I7G) and PPAR-γ (PDB entry 1171) [36]. Like


Fig. 29 Interactions of EPA with PPA* (a) The carboxylate head group of the ligand interacts with three protein residues His-323 His-449 and Tyr-473 (b) Two distinct confermations of the hydrophobic tail of the ligand were observed The ligand-binding site of PPARU5 is Y-shaped and each conformation of EPA fills a different branch of the Y.

PPAR-α, both PPAR-α and PPAR-γ have Y-shaped ligand binding sites; however, AZ-242 only fills two of the three sites. PPAR-α has four polar residues that interact with the carboxylates, Tyr-314 from helix-5, His-440 from helix-11, Tyr-464 from helix-12 and Ser-280 from helix-3. His-449 forms a buried hydrogen bond to the side chain of Lys-358. Tyr-314, His-440 and Tyr-464 are in the equivalent positions as His-323, His-449 and Tyr-473 in PPAR-δ. Thus, a major difference between PPAR-α and PPAR-δ is the change of a histidine in PPAR-δ to a tyrosine in PPARα. The dihydrocinnamate group of AZ-242 has a different local conformation than that of the phenoxy acid group of GW2433, as a result the carboxy groups of both ligands interact with the protein in distinct manners. This may explain why AZ-242 is a PPAR-α agonist and not a PPAR-α5 agonist (Fig. 31). The structure of AZ-242 bound to PPAR-γ has also been reported. The conformation of the dihydrocinnamate group resembles the conformation found in the

35


Fig. 30 Interactions of the PPAR-"/* co-agonist with PPAR-,5 The Yshape of the ligand is complementary to the Y-shape of the ligand binding pocket One of the three arms of the Y makes polar interactions with the carboxy-late head group of the ligand the interactions in the other two arms are mostly hydrophobic.

PPAR-α complex. In PPAR-γ , Tyr-314 in PPAR-α is replaced by His-323, thus the carboxylate interacts with two histidines (His-323 and His-449) and one tyrosine (Tyr-473). This arrangement of polar groups is the same as found in PPAR-δ; however, there are subtle differences in the position of each residue relative to each other in the two isotypes. In addition, one of the carboxylate oxygens also forms a

Fig. 31 Key polar interactions of PPAR-"/( co-agonist AZ-242 with PPAR-".


Fig. 32 Key polar interactions of PPAR-"/( co-agonist AZ-242 with PPAR-(. Notice that the equivalent residue of PPAR-" Tyr-314 is His323 in PPAR-(.

hydrogen bond with Ser-289. One difference between the two AZ-242 complexes is the conformation of the phenoxy "tail" of the ligand; this results in slight differences in the final position of the phenyl sulfonate group. One difference between PPAR-γ and PPAR-α is the presence of Cys-275 in PPAR-α, which is Gly-284 in PPAR-γ . The different tail conformations of the phenoxy linker allow the sulfonate group of AZ-242 to avoid the Cys-275 side chain in PPAR-α (Fig. 32). The thiazolidinediones (TZDs) are a class of antidiabetic agents that were subsequently shown to be potent PPAR-γ agonists. TZDs are usually selective for PPAR-γ although exceptions are known [37, 38]. The structure of the TZD rosiglitazone in complex with PPAR-γ has been reported both in the presence (PDB entry 1FM6) [3] and absence (PDB entry 2PRG) [39] of RXR-α ligated with 9-cis-retinoic acid. Both complexes are quite similar and the discussion will center around the heterodimeric complex. Like AZ-242 rosiglitazone occupies two arms of the Y-shaped liand-binding site. The polar TZD head forms hydrogen bonds with His-323 and Tyr-473 and also makes longer polar interactions with His-449 and Ser-289 (Fig 33). Thus, the TZD group mimics a carboxylate group. The nonpolar part of rosiglitazone makes VDW contacts within the ligand-binding site. The larger size of a TZD group compared with a carboxylate group usually allows it to be selective for PPARγ over the other isotypes. For PPAR-δ, the carboxylate binding site is narrower than in the other isotypes; thus the larger TZD head group cannot interact with the polar groups on the PPAR-S protein. The presence of the Tyr-314 precludes the TZD head group from making the proper polar contacts with the PPAR-α protein. Consistent with this proposal, the PPAR-γ H323Y mutant binds rosiglitazone 50-fold weaker than wild type PPAR-γ . In addition, the PPAR-α Y314H mutant binds rosiglitazone

37


Fig. 33 Key polar interactions of PPAR-(-selective agonist rosiglitazone with PPAR-( The TZD group acts as a carboxylate mimetic.

weakly while the wild type PPAR-α does not bind it to any measurable extent [40]. Clearly, the steric demands of the carboxylate-binding pocket play a major role in determining isotype selectivity. It has been speculated that the shift in the TZD head group caused by Tyr-314 in PPAR-α results in an unfavorable conformation in the remainder of the TZD ligand. The TZD KRP-297 has been reported to show dual PPAR-α/γ activity [41]. Unlike rosiglitazone, KRP-297 has a meta-substituted side chain across the central phenyl rings. The meta-substituted side chain may allow an improved fit in the PPAR-α protein [40]. Farglitazar (GI262570) is a PPAR-γ /α co-agonist that shows 1000-fold selectivity for PPAR-γ over the α-isotype. The structure of its complex with PPAR-γ has been reported in the heterodimeric complex with ligated RXR-α (PDB entry: 1FM9) [3]. Farglitazar binds to PPAR-γ in a similar manner that AZ242 does; the dihydrocinnamate moieties bind nearly identically and, like AZ242, the carboxylate interacts with His-449, Tyr-473, His-323 and Ser-289 (Fig. 34). The tails of the two ligands bind by VDW contacts in the same pocket although the details of the binding are different. Thus, only two of the three branches of the Y-shaped pocket are occupied. The most dramatic difference between these ligands is the size of the substituent adjacent to the carboxylate. For AZ242 it is an O-ethyl group, and for farglitazar it is the much larger amino benzophenone group. The side chain of Phe-363 moves in the farglitazar structure relative to the AZ242 structure to open a pocket that accommodates the second ring of the benzophenone group. A close analogue of farglitazar, GW409544, is also a co-agonist that binds only ten times poorer towards PPAR-α (Fig. 35). Neither farglitazar nor GW409544 bind to, or activate, PPAR-δ to any measurable extent. The structures of GW409544 bound to both PPAR-γ (PDB entry: 1K74) and to PPAR-α (PDB entry: 1K7L) have


Fig. 34 Key polar interactions of PPAR-(-selective agonist farglitazar with PPAR-(. The larger benzophenone group fits into a pocket created by the movement of Phe-363 relative to its position in the rosiglitazone complex.

Fig. 35 Structure of the PPAR-"/( co-agonist CW409544 and the PPAR-*-selective agonist CW501516.

39


been reported [40]. GW409544 binds to PPAR-γ in a manner very similar to, but not identical to, the way farglitazar binds to PPAR-γ . The α-amino side chains bind differentiy in the two structures. GW409544 binds to PPAR-α and PPAR-γ in nearly identical manners despite slight changes in key protein residues contacting the ligand. The equivalent residue to PPAR-γ Phe-363 is Ile-354 in PPAR-α, and as previously discussed PPAR-α His-323 is substituted by the larger Tyr-314 in PPAR-α. A modeling exercise proposed that for farglitazar to interact with the larger Tyr-314 in PPAR-α a shift in the ligand position would result in a steric clash between a benzophenone ring and Phe-273. The structure of the complex between GW409544 and PPAR-α suggests that adding three atoms to the vinylogous amide of GW409544 to generate farglitazar would result in a steric clash with Phe-273. Consistent with this proposal, the PPAR-α Y314H mutant potently binds farglitazar [40]. A very interesting discussion about the key factors that result in PPAR subtype specificity has been published by the Glaxo group [40]. This discussion is now summarized below. The PPAR-α and PPAR-δ ligand-binding pockets are significantly larger than the PPAR-δ pocket because of the narrowing of the pocket adjacent to helix-12. It is notable that only a handful of potent PPAR-δ ligands have been described. Ligands such as TZDs and L-tyrosine-based agonists like farglitazar show little or no binding to PPAR-α. In both cases, their acidic head groups seem to be too large to fit within the narrow PPAR-δ pocket. In contrast, the potent PPAR-δ agonist GW501516 contains an unsubstituted phenoxy-acetic acid head group that complements the narrow PPAR-δ ligand-binding pocket. Fibrate ligands, which generally bind to PPAR-δ only at high micromolar concentrations, contain small alkyl substituents adjacent to the carboxylate group. A PPAR-δ mutant M417V, which allows fibrate ligands to bind to PPAR-δ, has been reported [42]. This mutation is likely to increase the size of the PPAR-δ pocket to facilitate the binding of the small alkyl substituents adjacent to the carboxylate. Thus, the reduced size of the PPAR-δ pocket is a major determinant of ligand binding to this subtype. In comparison with PPAR-α, the PPAR-α and PPAR-γ ligand-binding pockets are closer in size and shape to each other. The Glaxo group has found that a major determinant of selectivity between these two subtypes is the substitution of Tyr-314 in PPAR-α for His-323 in PPAR-γ . These amino acids form part of the network of hydrogen-bonding residues that are involved in the activation of the receptor by its acidic ligands. Overlay of the PPAR-α and PPAR-γ crystal structures reveals that the larger volume of the Tyr-314 side chain in PPAR-α forces a 1.5 A shift in the position of the high-affinity ligand GW409544. The structurally related ligand farglitazar is unable to accommodate this shift because of a steric interaction with PPAR-α Phe-273. As a result, farglitazar shows 1000-fold selectivity for PPAR-γ over PPAR-α. The point mutations of Y314H in PPAR-α and H323Y in PPAR-γ demonstrate that these single amino acids are, in large part, responsible for determining the subtype selectivity of farglitazar. In each case, a 10–100-fold shift in the potency of the ligand was observed. Compared with farglitazar, GW409544 has three atoms removed to allow it to shift within the PPAR-α pocket without clashing

6 Conclusions

with Phe-273. The potent dual PPAR-α/γ agonist activity of GW409544 results from a complementary match of the re-engineered ligand with both the PPAR-α and PPAR-γ ligand-binding pockets. TZD ligands also respond to the point mutation of Y314H in PPAR-α and H323Y in PPAR-γ with a corresponding increase in PPAR-α and decrease in PPAR-γ activity, respectively. These data suggest that the TZDs, which do not contain the large N-substituents present in the L-tyrosinebased ligands, also have difficulty accommodating the 1.5 Å shift required to bind to PPAR-α. It was speculated that the shift in the TZD head group results in an unfavorable conformation in the remainder of the molecule. It is interesting to note that a single amino acid difference in PPARs has such a dramatic impact on ligand selectivity, given that the PPAR pocket is composed of more than 25 amino acids [40].

6 Conclusions

NHRs function as either activators or repressors of gene expression. Small molecule ligands (hormones) are the regulators of these proteins. Structural studies have characterized at least two distinct forms of ligated NHRs, agonist-bound NHR with a peptide derived from a co-activator and antagonist-bound NHR with a peptide derived from a co-repressor. These define two states for the NHR, the first one is the "on" state where genes are expressed and the second is the "off-state where gene expression is repressed. There are other structures that may, or may not, represent intermediate states between "on" and "off". The discussions above, while not comprehensive, have focused on how ligands can bind to and alter the conformational state of the NHR. More importantly, the discussion has focused on the important issue of ligand selectivity for a specific NHR. A system that is regulated by ligands is only effective if the proteins that are being regulated respond only to a specific ligand, or to just a few key ligands. Thus, the ability of estradiol to regulate ER-α and ER-β and not to regulate other related proteins such as GR and AR is the key to this regulatory system. The key trends about how ligands selectively bind to and agonize their NHR target protein will now be summarized. A striking observation for many of these agonist NHR complexes is the lack of high steric complementarity between the mostly hydrophobic ligand and the ligandbinding core of the protein. The complex between estradiol and ER-α shows that there are large unoccupied cavities between the ligand and the protein. In spite of this, estradiol is a potent and selective ligand for ER-α and the closely related ER-β. Another good example is found in the complexes between PPAR-γ and rosiglitazone (Fig. 33) and the larger ligand farglitazar (Fig. 34). Both are potent selective ligands; however, the larger ligand is 90 times more potent. While the polar thiazolidedione moiety of rosiglitazone plays a key role for its isotype selectivity for PPAR-γ , the larger benzophenone moiety of farglitazar is required for PPAR-γ

41


selectivity. Thus, for farglitazar a precise relative fit of the polar carboxylate functionality and one of the apolar groups work together to achieve PPAR-γ selectivity. As discussed previously, the Glaxo group has shown that changes to the benzophenone moiety of farglitazar produce ligands that agonize both PPAR-γ and PPAR-α. They have also determined that a single polar amino acid change between PPARγ (His-323) and PPAR-α (Tyr-314, equivalent to PPAR-γ His-323) is the primary determinant of PPAR-γ selectivity for both rosiglitazone and farglitazar. The conclusion reached is that the PPAR isotype selectivity is determined by the polar ligand group making a precise interaction with the protein, with the remainder of the ligand being well tolerated by the protein. Many steroid hormones are mostly hydrophobic molecules that have polar groups at opposite ends of the molecule (A- and D-rings). For these steroid receptors a key polar residue on a common position of helix-3 helps the estrogen receptors (Glu353 in ER-α) select for A-ring phenols, while PR, AR, GR and MR, which have a glutamine at this position, are selective for 3-keto A-rings (Figs. 4 and 12). Further ligand discrimination occurs primarily by additional polar interactions near the Dring (Figs. 12, 13, 14, and 15). For the steroid receptors, agonist selectivity appears to be determined by matching of polar groups on the ends of the ligand with complementary protein polar groups. The hydrophobic-α of the ligands and the apolar nature of the ligand binding site provides a mechanism for obtaining high affinity, provided there are no polar mismatches, in spite of less than optimal steric complementarity. Another good example of this is the case of two enantiomers (BMS270394 and BMS270394, Fig. 23) of the same ligand binding to RAR-γ . Both ligands orient the carboxylate, hydroxyl group, the A-, C- and D-rings in approximately the same manner. The linking amide group of the more potent isomer makes complementary interactions with the protein (Fig. 24) while the less potent isomer does not (Fig. 25). Thus we see a vivid example of where effective agonist binding occurs when a hydrophobic ligand makes a less than optimal steric fit in the core of the NHR and has the matching of polar ligand groups with complementary protein polar groups. The "inactive" isomer must bury the polar amide group in the protein interior and thus bind very weakly. Comparison of RAR and RXR shows distinctly different shapes of their ligandbinding pockets. Thus, RAR can effectively bind either all-trans-retinoic acid or 9-cis-retinoic acid as agonists, while RXR can only accommodate the bent ligand 9-cis-retinoic acid. These data make it tempting to speculate that selectivity is determined not by a precise match of the "right" ligand with the "right" NHR, but rather by the exclusion of "wrong" ligands by a mismatch of polar groups or an impossible steric fit. To conclude this chapter, the molecular recognition of ligands by NHRs is determined by a number of subtle factors. Agonism or antagonism is due to the ability of a given ligand to bind to the appropriate form of the protein. It is likely that nonpolar interactions provide binding affinity even in the absence of good shape complementarity. Selectivity appears to be guided by the avoidance of interactions, either polar or apolar, that are unfavorable.

References

Acknowledgements

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1

The Determination of Equilibrium Dissociation Constants of Protein-Ligand Complexes by NMR Gordon C.K. Roberts

University of Leicester, Leicester, United Kingdom

Originally published in: BioNMR in Drug Research. Edited by Oliver Zerbe. Copyright ľ 2002 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30465-7

1 Introduction

In general terms, the essential requirement for measuring the dissociation constant of a protein-ligand complex spectroscopically is the identification of a signal (from either the protein or the ligand) which has a different value in the complex from that in the free protein or ligand. In this respect, NMR has real advantages, since of the many resonances from the protein and the ligand several are sure to change – in shift, relaxation and/or scalar coupling – on complex formation. Most NMR studies of complex formation are carried out in order to determine the magnitude and direction of these changes in NMR parameters and to use them to draw conclusions about the nature of the complex. Here we will consider only the more limited use of NMR to determine the equilibrium dissociation constant for complex formation. As we shall see, NMR is particularly useful for the measurement of relatively weak binding, typically dissociation constants (K d ) 10−5 M. Accurate measurement of K d requires the use of protein concentrations ≤K d , and the low sensitivity of NMR in terms of the concentrations required obviously sets a practical limit to the range of dissociation constants which can be measured; this is discussed further below. A key factor in the use of NMR for measuring dissociation constants is its sensitivity to the rate of “chemical exchange”. Complex formation necessarily involves the exchange of the nuclei or molecules being observed between (at least) two states – the free ligand or protein and the complex. The fact that the appearance of the NMR spectrum is sensitive not only to the position of this equilibrium but also to the rates involved has a major influence on the design of NMR experiments for measuring dissociation constants and on the analysis of such data.


2 The Determination of Equilibrium Dissociation Constants of Protein-Ligand Complexes by NMR

2 Chemical Exchange and NMR

In the context of NMR, “chemical exchange” refers to any process in which a nucleus exchanges between two or more environments in which its NMR parameters (chemical shift, scalar or dipolar coupling, relaxation) differ. The effect of this exchange process on the appearance of the NMR spectrum depends on the rate of exchange relative to the magnitude of the difference in NMR parameters between the two states. The effects which exchange can have on the appearance of the spectra are illustrated in Fig. 1 for the simple case of exchange between two equally populated states. When the exchange is very slow on the “NMR timescale” – here relative to the magnitude of the chemical shift difference between the two states – two separate resonances are seen at the positions corresponding to the chemical shifts characteristic of the two states. At the other extreme, when the exchange is very fast relative to the chemical shift difference, a single resonance is seen, whose position is the average of the chemical shifts of the two states, weighted by their relative populations (see below). Between these extremes there are complex changes in line shape, which are very sensitive to the precise value of the rate of exchange. Detailed descriptions of these exchange effects can be found in a number of textbooks and reviews [1–7].

Fig. 1 Effects of the exchange rate on the appearance of the NMR spectrum from a system in which a nucleus is exchanging between two equally populated states in which it has a different chemical shift.

2 Chemical Exchange and NMR 3

When attempting to use NMR to measure a dissociation constant, the basic experiment will be to vary the ligand concentration in the presence of a fixed concentration of protein. (The converse experiment, varying the protein concentration, may sometimes be carried out, but is generally less satisfactory because of problems with protein solubility and aggregation.) What one sees in this experiment will depend critically on the rate of exchange of the molecules between the free state and the complex. If the exchange is slow on the “NMR timescale”, the two resonances, corresponding to the bound and free states, will remain in the same positions, but their relative intensities will change. If the exchange is very fast, the position of the single resonance will change progressively as the relative amounts of free protein or ligand change. It is important to recognise that the “NMR timescale” is a relative one. For a given equilibrium, different resonances will show slow, intermediate or fast exchange behavior, depending on how much their chemical shifts differ between the free and bound states. An example of this is shown in Fig. 2. This shows a small region of a superposition of a series of 1 H-15 N HSQC spectra of the calcium-binding regulatory protein S110B, alone and in the presence of increasing concentrations of a peptide corresponding to its binding site on the actin-capping protein CapZ [8]. Three residues whose 1 H and 15 N chemical shifts change on peptide binding are indicated. For Glu72, the chemical shift changes are small, and a progressive movement of the cross peak is seen, indicative of very fast exchange. This is also the case for Ser62, where the shift changes are larger, but here the progressive shift of the cross peak is accompanied by a marked broadening at intermediate concentrations, indicating that for this peak the exchange regime is between fast

Fig. 2 Expanded region of the overlaid 1 H15 N HSQC spectra from a titration of 15 Nlabeled S100B with CapZ peptide. The protein concentration was 1 mM, and the spectra shown are for peptide concentrations of

0, 0.25, 0.5, 0.75, 1.0, 1.5, 2.5 and 3.0 mM. Chemical shift changes for the cross peaks of Ser62, Glu72 and Ala78 are indicated. From Ref. [8] with permission.


and intermediate exchange. Finally, for Ala78, which shows the largest changes in 1 H and 15 N shifts, the broadening at intermediate peptide concentrations is such that the peak disappears completely, characteristic of intermediate exchange rates (cf. Fig. 1). These line-broadening effects can be very valuable as a means of determining the dissociation rate constant of the complex; detailed procedures for doing this are described elsewhere [9]. As described below, they can, however, present complications in the accurate determination of dissociation constants. Bearing in mind the relativity of the “NMR timescale”, it is useful to estimate the range of rates which will produce line-broadening effects. For chemical shift differences1) in the range 10–1000 Hz, intermediate exchange behavior will be observed for dissociation rate constants in the range 102 –104 s−1 ; assuming that the association rate constant is diffusion-limited, ∼107 –108 M−1 s−1 , this will correspond to dissociation constants in the range 10−3 –10−6 M.

3 The Basic Equations

The following descriptions will be couched primarily in terms of chemical shift changes, which are the most widely used for the measurement of dissociation constants; the general principles apply equally to measurements based on other NMR parameters, and these will be touched on later. k+1

E + L EL k−1

(1)

kd = kk −+ 11 = [E][L] [EL]

In discussing the effects of exchange on NMR spectra, an important parameter is the lifetime of a particular state: For a nucleus in the ligand molecule: Lifetime in state EL,

τEL = 1/k − 1

Lifetime in state L, τL = 1/k + 1 [E]

For a nucleus in the protein molecule: Lifetime in state EL,

τEL = 1/k − 1

Lifetime in state E, τE = 1/k + 1 [L]

Using Eq. (1), the lifetimes in the free state of the ligand and protein can be expressed more conveniently as: Lifetime in state L,

τL =

p k − 1 PEL

1) Note that in the context of exchange effects on NMR spectra chemical shift differences

Lifetime in state E, τE =

pE k − 1 PEL

must be measured in Hertz, not parts per million.

3 The Basic Equations

Where pX is the fractional population of species X, so that, for a ligand resonance, PEL =

[EL] LT

(2)

and pL =

[L] = 1 − pEL, LT

(3)

while for a protein resonance pEL =

[EL] ET

(4)

and pL =

[E] = 1 − pEL , ET

(5)

where LT and ET represent the total concentrations of ligand and protein, respectively. From Eq. (1), for a ligand resonance, 2 [EL] (ET + LT + Kd ) − (ET + LT + Kd ) − 4ET LT = pEL = LT 2LT

(6)

with an analogous equation for an enzyme resonance. In discussions of exchange effects in NMR, a single lifetime is often used to characterize the exchange process 1/τ = 1/τEL + 1/τL = k − 1 (1 + pEL / pL ) = k − 1 / pL

(7)

The spectrum in the presence of exchange in the absence of scalar coupling can be simulated by using McConnell’s modification of the Bloch equations [10–12]. (When there is strong scalar coupling, or when the coupling changes because of the exchange process, the density matrix approach must be used [5]; a number of line-shape fitting programs based on this approach are available [5].) For the simple case, the line shape (amplitude as a function of frequency, v) is given by the imaginary part of G(v), G(v) = iC[2PL PEL τ −τ 2 (PL αEL + PEL αL )] PL PEL −τ 2 αL αEL

(8)

where C is a scaling factor and αL = 2πi(vL − v) + 1/T2,L + pEL /τ ,

αEL = 2πi(vEL − v) + 1/T2,EL + pL /τ

5


In general terms, the three exchange regimes are defined by: Slow exchange 1/τ 2π|vL − vEL | Intermediate exchange 1/τ ∼ 2π|vL − vEL | Fast exchange 1τ 2π|vL − vEL | Simpler expressions for the line shape are available for both the fast and slow exchange regimes (see Ref [9]). The possibility of measuring the dissociation equilibrium constant in each of these three regimes will now be considered.

4 Slow Exchange

In this regime, separate peaks are observed for the free and bound states (cf Fig. 1); when the relative amounts of these two states are altered by varying the ligand concentration, the positions of these two signals remain unaltered; only their relative intensity changes. In principle, this change in relative intensity can be used to estimate the dissociation constant; in practice this is only of any value under very restrictive conditions. First, the signals must be sufficiently well resolved to allow accurate intensity measurements; this can be achieved by appropriate isotope labeling – for example, using per-deuterated protein to study ligand resonances. Second, the signal-to-noise ratio must be sufficient to allow accurate intensity measurements at sub-stoichiometric ligand concentrations ([EL] ∼ 0.1ET ). This latter problem is compounded by the fact that accurate measurement of K d requires that ET < K d . Slow exchange implies a slow rate of dissociation, which in turn implies a relatively low value of K d and hence the need for a low protein concentration. For example, for a chemical shift difference of 100 Hz, observation of slow exchange would imply that k–1 ≤ 3×102 s−1 , so that (assuming a diffusion-limited association rate) K d ≤ 3×10−5 M, and one would need to make accurate intensity measurements at protein concentrations of ∼5 µM. Given the insensitivity of NMR, it is therefore very rarely practical to make reliable estimates of K d from spectra in the slow exchange regime.

5 Intermediate Exchange

The intermediate exchange regime covers a relatively narrow range of exchange rates, no more than a factor of two either side of the “coalescence point” (Eq. (9)). Within this regime the full line-shape equation must be used in order to extract the relative populations of the bound and free states, their chemical shifts and the rate of exchange between them. In principle good estimates of these parameters can be obtained from fitting a series of spectra at different ligand concentrations, but in practice limitations of resolution and signal-to-noise ratio – the signals can

6 Fast Exchange

be very broad at the coalescence point – mean that this is rarely achievable. In most circumstances a change in spectrometer frequency will be sufficient to shift the system into either slow exchange (at higher spectrometer frequency) or fast exchange (at lower spectrometer frequency), where the analysis will be easier.

6 Fast Exchange

The fast exchange regime is by far the most useful for the measurement of dissociation constants from changes in chemical shift, relaxation or other NMR parameters – a wide range of K d values, from 10 µM (or perhaps a little less) to ≥10 mM, can be accessible. 6.1 Very Fast Exchange

When the exchange rate is very fast, the chemical shifts and relaxation rates of the nuclei of the ligand and the protein are completely averaged between the bound and free states, and a weighted average spectrum is observed. For ligand resonances δobs,L = pL δL + pEL δEL

R2,obs,L = pL R2,L + pEL R2,EL,

(9)

(10)

where δ obs,L , δ L and δ EL are, respectively, the observed ligand chemical shift and the chemical shifts of the free ligand and of the ligand in the complex, with analogous definitions for R2 (=1/T 2 ), the transverse relaxation rate. Similarly for protein resonances δobs,E = pE δE + pEL δEL

(11)

R2,obs,E = pE R2,E + pEL R2,EL

(12)

From Eq. (9), for example, for a ligand resonance pEL =

δobs − δL δEL − δL

(13)

and this can be combined with Eq. (6) to give 2 δobs − δL (ET + LT + Kd ) − (ET + Lt + Kd ) − 4Et LT = δEL − δL 2LT

(14)

7


and fitting Eq. (14) (or the analogous equations derived from Eqs. (10)–(12)) to the data (δ obs as a function of LT ) will yield estimates for K d and δ EL . This is the commonest approach used to determine K d by NMR; a detailed protocol for experiments of this kind is given elsewhere [9]. 6.2 The General Case of Fast Exchange

It is important to note that Eqs. (9)–(12) are valid only for very fast exchange. If the exchange is somewhat slower (“moderately fast” exchange), there will be an exchange contribution to the line width, and Eq. (10) for the ligand resonance2) will be replaced by R2,obs = PL R2,L + PEL R2,EL +

PEL PL2 4π 2 (δEL − δL )2 k−1

(15)

This equation predicts a different dependence of the line width on the ligand concentration from that predicted by Eq. (10). For large k–1 , Eq. (15) clearly reduces to Eq. (10), and, if changes in relaxation are being used to estimate K d , it is generally sensible to fit the data using Eq. (15) rather than Eq. (10) (the only exception being paramagnetic relaxation, see below). Most important, if the exchange contribution to the line width (the third term on the right hand side of Eq. (15)) is significant, Eqs. (9) and (14) will not be an accurate description of the change in chemical shift with ligand concentration. It has been shown by simulation experiments that the erroneous use of Eqs. (9) and (14) when exchange is not very fast can lead to errors of at least an order of magnitude in K d [13]. For example, Fig. 3 A shows a series of simulated spectra of a ligand resonance at increasing values of LT ; the chemical shift difference between the bound and free states is 100 Hz and k–1 is 500 s−1 . There is clearly a progressive change in chemical shift as the ligand concentration is increased, as would be expected for fast exchange, and the signals appear to have Lorentzian shapes. However, broadening of the resonance line in the middle of the titration is also clearly evident; this arises from the exchange contribution in Eq. (15), which has a maximum when pEL ≈ 1/3. While the chemical shifts obtained from these simulated spectra seem to be described reasonably satisfactorily by Eq. (14), as shown in Fig. 3 B, the K d value estimated from the least-squares fitting of Eq. (14) to the data is in error by almost a factor of two, even when, as here, data is available for a wide range of ligand concentrations (from 0.1ET to 30ET ) [13]. When, more realistically, only ligand concentrations greater than the protein concentration are used in the analysis, the error becomes a factor often for the parameters used in Fig. 3 and can be as much as 100 for somewhat lower values of k–1 /(δ EL – δ L ) [13].

2) Eq. (15) and the following discussion are couched in terms of ligand resonances, but ex-

actly the same considerations apply to protein resonances.

6 Fast Exchange

Fig. 3 Simulation of spectra of a ligand resonance in a system involving proteinligand complex formation. The parameters used for the simulation were: (*EL –*L ) 100 Hz, k−1 500 s−1 , K d 10−5 M, ET 10−3 M, LT (0.1–30) 10−3 M, free line width 5 Hz, bound line width 20 Hz. A (left): Simulated line shapes for LT 0.5 mM (◦), 1.5 mM (), 2.0 mM (+), 2.5 mM (×), 10 mM ().

B (right): Observed chemical shift (*obs –*L ) as a function of total ligand concentration, LT . The points are values calculated from the exact simulation, and the curve is the best fit to Eq. (14), assuming no exchange contribution; the value of K d estimated from the fit is 6.2×10−6 M. From Ref [13] with permission.

This is a potentially serious source of error for combinations of δ EL – δ L ), k–1 and K d which are very likely to be encountered in practice. It is thus essential to establish when it is safe to use Eq. (14) to analyze chemical shift changes. The simplest way to do this is to measure the line width of the resonance of interest as well as its position, and to analyze its dependence on ligand concentration (as described in Refs. [9, 13]) to estimate the magnitude of the exchange contribution (Eq. (15)); only when the exchange contribution is negligible – the very fast exchange condition is satisfied – is it safe to use Eq. (14). In the more general case of “moderately fast” exchange, where the simple Eq. (14) cannot be used, the alternative is either a full line-shape analysis or, more practically, a comparison of measured chemical shifts and line widths as a function of ligand concentration with those obtained from simulated spectra; detailed protocols for this approach are available [9]. Both in order to identify the exchange regime and to obtain accurate and precise values of K d , it is important to make measurements at as low a ligand concentration as possible. 6.3 Paramagnetic Relaxation

For those proteins which contain a paramagnetic center (e.g., heme iron, copper), usually at the active site, the bound ligand will experience a marked increase in relaxation rate due to dipolar interaction with the unpaired electron(s) of the paramagnetic center. While the paramagnetic relaxation effect is commonly used

9


to obtain structural information (the distances between the paramagnetic center and ligand nuclei), it can also be used for the determination of K d (see, for example, Refs. [14, 15]). It is mentioned separately here because, since the magnetic moment of the electron is 1800 times that of the proton, it is, when present, a dominant effect. It magnitude is such that it can only be measured under fast exchange conditions, with a large excess of ligand over protein. This can simplify the analysis (for LT ET , pL ≈1), and it also allows one to use low enzyme concentrations, so that somewhat lower K d values can be measured. On the other hand, these same experimental conditions mean that a long extrapolation to R2,EL or R1,EL is required, and this limits the precision of the measurement.

7 Conclusions

As discussed above, NMR can be a useful tool for the determination of dissociation constants provided that its strengths and limitations are recognized. In practice, it is only useful under fast exchange conditions, and thus for relatively weak binding: K d 10 µM. The upper limit to the dissociation constants which can be measured is usually set by the solubility of the ligand molecule, since concentrations 10 K d are needed to obtain reliable estimates. Modest amounts of cosolvent (e.g., DMSO, dimethyl sulfoxide) are often used to ameliorate this problem, but most proteins will only tolerate small amounts < 10% of such cosolvents, and for accurate data it is important to keep the cosolvent concentration constant during the titration. For dissociation constants in the 10 µM–10 mM range, NMR is certainly competitive with other available methods; it is not dependent on the presence of a chromophore and does not require the separation of bound and free ligand. Provided that simple precautions, described above, are taken to ensure that the appropriate analysis is used, K d measurement by NMR is rapid, simple and accurate. The emphasis in the discussion has been on the analysis of changes in chemical shift and relaxation rate because these are the parameters most commonly used for K d measurements. Any protein-ligand binding process is bound to be accompanied by some change in chemical shift of a resonance or resonances from both the protein and the ligand, although of course the magnitude of such a shift change cannot in general be predicted. A number of resonances and indeed different spectrometer frequencies may need to be examined to find one with a shift large enough to be useful but small enough to be in fast exchange. Changes in relaxation rate are likely to be almost universal for ligand resonances, since the rotational correlation time of the ligand will increase substantially on binding to a macromolecule, though the magnitude of the effect can be complicated by internal motion in the complex. For the protein, on the other hand, changes in relaxation may occur but will be wholly dependent on local changes in mobility. The use of changes in relaxation to study ligand binding to proteins was one of the earliest biological applications of NMR [3, 16] and remains a valuable approach. In fact, of course, any NMR parameter which changes on complex formation and which shows fast exchange

References

behavior (being described by an equation analogous to Eqs. (9)–(12)) can be used to measure K d . Scalar coupling constants are likely to change if the conformation of the ligand or of protein side-chains changes on complex formation, and this can provide useful structural information; however, the magnitude of such changes is rarely large enough to be useful for K d determination. On the other hand, NMR experiments which are used to screen for binding, such as pulsed-fieldgradient measurements of translational diffusion or the measurement of the sign of intramolecular NOEs in the ligand (see, e.g., Ref [17]), can also readily be adapted for quantitative determination of K d – for example, see Ref. [18] for the use of translational diffusion measurements in this way.

References 1 KAPLAN, J. I. and FRAENKEL, G., NMR of Chemically Exchanging Systems. Academic Press, New York, 1980. 2 SANDSTROM, J., Dynamic NMR Spectroscopy. Academic Press, London, 1982. 3 JARDETZKY, O., and ROBERTS, G. C. K., NMR in Molecular Biology. Academic Press, New York, 1981. 4 SANDERS, J. K. M. and HUNTER, B. K., Modern NMR Spectroscopy. Oxford University Press, Oxford, 1987. 5 NAGESWARA RAO, B. D., Methods Enzymol. 1989, 176, 279–311. 6 LED, J. J., GESMAR, H., and ABILDGAARD, F., Methods Enzymol. 1989, 176, 311–329. 7 BERKOWITZ, B., and BALABAN, RS., Methods Enzymol. 1989, 176, 330–341. 8 KILBY, P. M., Van ELDIK, L. J., and ROBERTS, G. C. K., Protein Sci. 1997, 6, 2494–2503. 9 LIAN, L.-Y., and ROBERTS, G. C. K., in NMR of Macromolecules (ed. ROBERTS, G. C. K.), Oxford University Press, Oxford,

pp. 153–182, 1993. 10 MCCONNELL, H. M., J. Chem. Phys. 1958, 28, 430–435. 11 LEIGH, J. S., Jr., J. Magn. Reson. 1971, 4, 308–318. 12 MCLAUGHLIN, A. C. and LEIGH, J. S., Jr., J. Magn. Reson. 1973, 9, 296–305. 13 FEENEY, J., BATCHELOR, J. G., ALBRAND, J. P. and ROBERTS, G. C. K., J. Magn. Reson. 1979, 33, 519–529. 14 MODI, S., PRIMROSE, W. U., BOYLE, J. M. B., GIBSON, C. F., LIAN, L.-Y., and ROBERTS, G. C. K. Biochemistry 1995, 34, 8982–8988. 15 MODI, S., PAINE, M. J. I., SUTCLIFFE, M. J., LIAN, L.-Y., PRIMROSE, W. U., WOLF, C. R.. and ROBERTS, G. C. K., Biochemistry 1996, 35, 4540–4550. 16 FISCHER, J. J. and JARDETZKY, O., J. Am. Chem. Soc. 1965, 87, 3237–3281. 17 ROBERTS, G. C. K., Drug Discov. Today 2000, 5, 230–240. 18 TILLETT, M. L., HORSFIELD, M. A., LIAN L.-Y. and NORWOOD, T. J., J. Magn. Reson. 1998, 133, 379–384.

11

1

Protein-Lipid Interactions during Virus Entry by Membrane Fusion Alex L. Lai, Yinling Li, and Lukas K. Tamm University of Virginia, Charlottesville, USA

Originally published in: Protein-Lipid Interactions. Edited by Lukas K. Tamm. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-31151-4

1 Introduction

A large number of animal viruses are enveloped by lipid bilayer membranes. Since enveloped viruses bud during biogenesis from specialized areas of the host cell surface, viral membrane envelopes are also highly specialized in terms of their lipid and protein compositions. Like the cell membranes from which they derive, they are enriched in sphingomyelin, phosphatidylcholine (PC) and cholesterol in the outer leaflet, and phosphatidylethanolamine (PE) and various negatively charged lipids in the inner leaflet. Viral membrane envelopes contain only a very small number, typically one to three, different integral membrane proteins. Most viral membrane envelope proteins are glycosylated and appear as elongated projections or spikes in electron micrographs of these viruses. The different viral spike glycoproteins have a number of specific tasks in viral cell entry. One function is to attach viruses to receptors on the surface of target cells. A second major function is to promote the fusion of viral and target cell membranes, either directly at the cell surface or after endocytosis with the endosomal membrane in a reaction that proceeds only at the lower pH that prevails in the endosome. In some viruses, both functions are packaged into a single protein, whereas in other viruses they are distributed between two different proteins. The atomic structures of the ectodomains of several viral spike glycoproteins have been solved by X-ray crystallography. According to these structures, viral membrane fusion proteins are generally grouped into two classes. Class I viral fusion proteins are elongated proteins characterized by trimeric bundles of helical hairpins with coiled-coil α-helical cores. In contrast, class II viral fusion proteins consist of three β-sheet domains that pair into dimers and form relatively flat lattices on the viral membrane surfaces (Fig. 1).


2 Protein-Lipid Interactions during Virus Entry by Membrane Fusion

Fig. 1 Diagrams of representative membrane-enveloped viruses with type I and type II fusion proteins. (A) Influenza virus. The type I fusion spike glycoprotein is HA, which is a trimer consisting of three identical subunits. Each subunit is composed of HA1 and HA2 chains. The coiled-coil structures (green) in the stem are formed by the HA2 chains and the globular domains (yellow) at the tip of the molecule are formed by the HA1 chains. The virus also contains the matrix M1 protein (pink) and eight segments of single (−) strand RNA (blue) that together with the nucleocapsid protein NP (grey) form

ribonucleo-protein complexes. The envelope also contains neuraminidase NA (blue) and the proton channel M2 protein (not shown) in relatively small copy numbers. (B) Semliki Forest virus. The type II fusion glycoprotein is the E1 protein (red) that forms together with the receptor binding protein E2 (yellow) a T=4 icosahedral lattice on the membrane surface. The nucleocapsid consists of 240 copies of the capsid protein C (pink). A single (+) strand RNA (blue) contains the entire genome of this alpha-virus. The envelope also contains the small soluble E3 protein in stoichiometric amounts (not shown).

2 Fusion of Pure Lipid Bilayers

2 Fusion of Pure Lipid Bilayers

Pure lipid bilayers do not spontaneously fuse. The headgroups of phospholipids are highly hydrated and hydration repulsion prevents the spontaneous fusion of uncharged lipid bilayers. If bilayers are negatively charged, charge repulsion adds to the repulsive energy between lipid bilayers. At equilibrium, fluid-phase PC binds about 34 molecules of water and PE binds about eight waters per lipid [1]. Therefore, the equilibrium distance measured from headgroup phosphate to headgroup phosphate between two fluid-phase lipid bilayers is about 30 Å for PC bilayers and about 10 Å for PE bilayers. The hydration of negatively charged phosphatidylserine (PS) is similar or slightly larger than that of PC [2]. This hydration barrier must be overcome en route to membrane fusion. There are a few different views of how this could happen. These will be briefly discussed in the following. It has been postulated as early as in the late 1970s and early 1980s that membrane fusion may proceed through “point defects” [3, 4]. The nature of these point defects, however, was not so clear. Markin et al. proposed in a theoretical study that membrane fusion could proceed through an hourglass-shaped “lipid stalk” intermediate [5]. Since lipid stalks are thought to be transient structures that may exist only briefly, these structures have never been observed experimentally in membrane fusion. However, arrays of lipid stalks have recently been observed in X-ray diffraction experiments of aligned stacks of membranes at submaximal hydration [6, 7]. Under specialized experimental conditions, pure lipid bilayers can also be observed to form “hemifusion” diaphragms when they are closely apposed to each other. In hemifusion, the two distal leaflets of two approaching bilayers join to form a single bilayer in the central hemifused region. Depending on the experimental set-up, hemifusion diaphragms may be quite extended. Hemifusion has been observed electrophysiologically in planar bilayer experiments [8, 9] and in the surface forces apparatus when two mica-supported lipid bilayers are mechanically pushed together [10, 11]. The combination of electrophysiological and theoretical studies on pure lipid bilayer fusion led to the “stalk–pore” model of membrane fusion (reviewed in [12]). In this model, lipid stalks radially expand until the two distal monolayers contact each other to form a hemifusion diaphragm. The hemifusion diaphragm is thought to break at some point to form an initial fusion pore. The stalk and initial pore intermediates have different curvatures, whose energies have been calculated in numerous theoretical studies. The effect of lipid additives that alter the spontaneous curvature of lipid bilayers are consistent with the stalk–pore model. For example, lyso-PC (induces positive curvature) added to the proximal monolayers and oleic acid (induces negative curvature) added to the distal monolayers promote fusion, whereas lyso-PC added to the distal monolayers and oleic acid added to the proximal monolayers inhibit fusion [12]. These and a multitude of similar studies on many different reconstituted and biological fusion systems are often taken as direct evidence for the stalk–pore model of membrane fusion. However, the reader should be cautioned: neither theoretical calculations of energies of stalks nor the correlation between the effect of curvature agents on the calculated energy of stalks and membrane fusion prove that neatly organized stalks actually exist as

3


intermediates in fusion. They just prove that calculated energies with sometimes somewhat arbitrarily chosen boundary conditions are compatible with certain nonbilayer lipid structures and that certain lipophilic agents produce membrane situations (curvature, hydration, defects, etc.) that are conducive/inhibitory to membrane fusion. It is important to make this distinction because the stalk–pore model has become so popular recently that many take it for a fact rather than a model. One should also be aware that it is virtually impossible to distinguish experimentally between different stalk and hemifusion intermediates in fusion of biological membranes because, by definition, lipids can exchange freely between the two membranes through all hemi-fusion intermediates. If well-organized stalks exist, they are probably short-lived transient intermediates that according to the newer models progress directly to fusion pre-pores without the need for a transmonolayer contact hemifusion intermediate [13]. Extended hemifusion diaphragms may occur rather as off-pathway side-products under suboptimal conditions in biological membrane fusion [14–18]. Lentz et al. developed sophisticated methods to monitor many important details of the kinetics of fusion between pure lipid bilayers [19]. In this work, small unilamellar vesicles were induced to fuse by the addition of 5–20% polyethylene glycol of molecular weight 8000. Polyethylene glycol aggregates the vesicles by reducing the activity of water. It has been found that bilayers composed of PC, PE, sphingomyelin and cholesterol at a molar ratio of 35:30:15:20 fuse quite efficiently and in a non-leaky fashion [20]. The kinetics of outer leaflet lipid mixing (fast), transbilayer lipid movement (intermediate), inner leaflet mixing (intermediate) and contents mixing (slow) have been resolved [21]. Clearly, outer leaflet lipid mixing (around 0.5 min) and pore formation (4–6 min) are two distinct kinetic processes in these welldefined pure lipid bilayer model systems. A “problem” of the original stalk–pore model for membrane fusion is that hydrophobic “voids”, i.e. volumes of hydrophobic mismatch are created in the stalk, which may be energetically very costly [22, 23]. Different authors have tried to get around this problem by different means. Newer calculations have simply eliminated the voids by assuming intermediate structures with tilted lipids [13, 24] or reduced their effects by relaxing the geometry of the stalk [25]. In another effort to rescue the original stalk model, it has been postulated that the “voids” have to be filled by hydrophobic substances that reduce the high energy of the stalk and make it a viable intermediate in lipid bilayer fusion. Indeed, when small amounts of very long-chain lipids or alkanes were added to the lipid mixtures, the rates of fusion (contents mixing), but not hemifusion (lipid mixing) of pure lipid bilayers were increased up to about 2-fold [26]. This rate increase was additional to that achieved by negative curvature inducing lipids. Even if the rates are increased by these lipid additions, these experiments do not explain why bilayers in their absence are still able to fuse at fairly rapid rates. Finally self-consistent field theory of flexible amphiphilic chains has been used to model the formation of traditional lipid stalks [27]. The energetic barrier to forming a stalk derived from this theory was significantly smaller than that derived from the phenomenological continuum theories, but a large energetic barrier, which depended on the spontaneous intrinsic curvature of the amphiphile, was associated with the radial expansion of the stalk into a hemifusion diaphragm.

3 Viral Protein Sequences that Mediate Lipid Bilayer Fusion

Coarse-grained molecular simulations have been used recently to provide more insight into the microscopic details of transitions at lipid bilayer fusion junctions. In one study, lipids were modeled as amphiphilic three-segment rods and studied by Brownian dynamics [28]. In another set of studies, the amphiphiles were modeled as flexible co-polymer chains in a hydrophilic polymer solvent and their transformation from bilayers into fusion intermediates was studied by Monte-Carlo lattice simulations [29, 30]. Interestingly, the outcome of both approaches was similar and suggested a new fusion mechanism, which was different from the classical stalk–pore mechanism. After formation of an initial quite disordered lipid stalk between two closely apposed bilayers, a hole appeared in either one or both parent membranes next to the stalk and the stalk then grew asymmetrically around these holes to form the initial fusion pore. These initial small holes appear in the bilayer because the line tension is high near disorganized lipid stalks. Coarse-grained molecular dynamics simulations revealed a few additional new aspects of microscopic details in the evolution of lipid stalks and fusion pores [31, 32]. In one study [31], the stalk was initiated by the displacement of a few lipid molecules from their normal position in one of the two bilayers. Fusion could then proceed through the classical stalk–pore or the new stalk-hole mechanism. Which mechanism prevailed depended on the headgroup composition of the bilayers with negative curvatureinducing lipids favoring the stalk–pore and bilayer-forming lipids favoring the stalk-hole mechanism. In the other study [32], the lipid molecules became tilted and eventually splayed their aliphatic tails such that each chain was resident in different opposing leaflets. The nucleus of the initial stalk formed by lipid tail splaying was again seen to expand asymmetrically in a circle to gradually enclose an initial fusion pore. Whether specific details of each of these mechanisms will hold up for bilayers made of “real” phospholipids (i.e. not coarse-grained models) remains to be seen. However, the fact that so many different oversimplified models and methods of simulation independently led to very similar general results is very promising. Perhaps the recently observed spurious leakage in influenza hemagglutinin (HA)-mediated membrane fusion is a reflection of hole formation in the new stalk-hole model of membrane fusion and thus may provide experimental support for this mechanism [33]. New details of how fusion between pure lipid bilayers proceeds at the microscopic level will almost certainly emerge as computational simulation methods continue to be further refined and as computer power increases year-by-year.

3 Viral Protein Sequences that Mediate Lipid Bilayer Fusion 3.1 Fusion Peptides

Photoaffinity labeling studies have shown that the major regions of viral spike glycoproteins that interact with lipid bilayers are domains called “fusion peptides”

5


as well as the transmembrane anchors of the viral spike glycoproteins. While the transmembrane anchor domains are permanently inserted into the viral lipid bilayer, the fusion peptides interact with the lipid bilayer of target (and/or viral) membranes only upon activation of the fusion process. Although fusion peptides are quite hydrophobic, these polypeptide sequences are protected by other parts of the viral spike glycoproteins and therefore do not interact with membranes in their resting state. Many fusion peptides like those of the influenza and human immunodeficiency viruses are located at the extreme N-terminus of the fusion subunit of the viral spike glycoproteins. Others such as those of Ebola or Dengue virus are internal sequences of the respective fusion proteins. A list of a few selected fusion peptides is shown in Fig. 2 (see also [34, 35]). The sequences of fusion peptides are extremely well conserved within each family of viruses, but not between different families. There are, however, some general features that are common between fusion peptides of the different virus families: the propensities of glycines and alanines are high, large bulky aromatic residues are frequently found, and hydrophilic residues are found interspersed towards the Cterminal end of N-terminal fusion peptides and towards both ends of internal fusion

Fig. 2 Sequences of class I and II Nterminal and internal fusion peptides. Glycines and alanines, which are frequent in these sequences are highlighted. Bulky aromatic residues are underlined and hy-

drophilic residues are marked with a dot. HIV, human immunodeficiency virus; ASLV, avian sarcoma leukemia virus; TBE, tickborne encephalitis virus; SFV, Semliki Forest virus.

3 Viral Protein Sequences that Mediate Lipid Bilayer Fusion

peptides. Although it is not always totally clear where a fusion peptide sequence begins and ends, site-directed mutagenesis has shown that quite dramatic fusion phenotype changes are found with some only relatively mild single amino acid changes in the fusion peptide region. Most evidence for this comes from work with influenza virus [36–40], whose fusion protein HA contains a receptor binding subunit (HA1) and a fusion subunit (HA2). Glycines 1 and 4 of HA2 are particularly susceptible to fusion defects. For example, the G1S mutant causes hemifusion, whereas a G1V replacement is completely defective in fusion [39]. Large bulky hydrophobic residues can be replaced with other large bulky hydrophobic residues, but not with glycines [40]. It appears that a proper balance and spacing of glycines and large bulky hydrophobic residues in the fusion peptide is important to confer fusion activity to influenza HA2. Even a deletion of the first glycine is not tolerated in this fusion protein [41]. 3.2 Transmembrane Domains

A first indication that the transmembrane domain is also very important for fusion and not just for anchoring the fusion protein in the viral membrane came from a study in which the transmembrane domain of influenza HA2 was replaced with a glycosylphosphatidylinositol anchor [14]. This construct was able to induce hemifusion between HA expressing and red blood cells, but it was unable to complete the reaction to develop a full fusion pore. Subsequent studies in several laboratories established that there was little requirement on the actual sequence of the transmembrane domain, but that its length mattered. For example, HA constructs with only the N- or C-terminal half of the transmembrane domain present mediated hemifusion, but not full fusion, and full fusion was gradually recovered as the domain length was increased to its full length [42]. 3.3 Other Regions of the Fusion Protein

Some fusion proteins with N-terminal fusion peptides contain other segments of the ectodomain that may interact with lipids in the process of membrane fusion. For example, the kink region of the ectodomain of influenza HA2 (residues 106–112) that connects the inner and outer layer α-helices in the pH5 structure has been proposed to contribute to membrane fusion [43] either by pH-dependent lipid interactions [44] or protein–protein interactions [45]. Similarly, it has been suggested that paramyxoviruses contain an internal membrane-interactive segment in addition to the N-terminal fusion peptide [46]. This putative internal secondary fusion peptide, which comprises residues 208–229 of the Sendai virus F protein is located in between the N- and C-terminal heptad repeats that form coiled coils in the post-fusion structure [47]. This sequence also maps to the C-terminal end of the N-terminal heptad repeat and a short helix (N1) that is part of the neck in the pre-fusion structure of the F protein of the related Newcastle disease virus [48].

7


It has been reported that a tryptophan-rich region just N-terminal to the transmembrane domain of HIVgp41 contributes to membrane fusion [49, 50]. This segment has been predicted to lie in the interface of the external leaflet of the viral membrane. A similar interfacial juxta-membrane domain has been predicted to exist in Ebola virus [51]. It is interesting to note that juxta-membrane domains have also been implicated to contribute to fusion in the SNARE proteins syntaxin 1a and synaptobrevin 2, and that the aromatic residues of this segment may serve an important structural and functional role in this process [52]. Finally, HIV and other lenti- and retroviruses contain three consecutive helices on the cytoplasmic or inner side of the viral membrane that exhibit large hydrophobic moments that promote strong interactions with the inner surfaces of the viral membranes. However, rather than contributing to fusion, the predominant roles of these amphipathic helices appear to direct the intracellular targeting of the Env glycoproteins in virus assembly and their targeting to the perinuclear envelope after host cell entry [53]. In the case of influenza HA2, the cytoplasmic domain comprises only 13 residues, three of which are palmitoylated cysteines. Palmitoylation appears to be particularly important in intracellular vesicle traffic for correctly targeting HAs to the apical membrane and for subsequent virus assembly [54, 55]. However, a minor modulatory role in fusion pore opening has also been ascribed to the palmitoylated cytoplasmic tail of influenza HA2 [56].

4 Interactions of Fusion Peptides with Lipid Bilayers

N-terminal fusion peptides likely form independent folding units in membranes and, therefore, are sometimes also called fusion domains. The reason for this contention is that the linkers between fusion peptides and the structured ectodomains (1) contain several glycines, (2) are not ordered in crystal structures even if the residues are present, and (3) are susceptible to protease digestion. Moreover, the lipid bilayer and its interface constitutes a very different folding environment for membrane proteins or inserted peptides than the aqueous environment, in which ectodomains fold [57]. However, to obtain meaningful folding units of fusion peptide models in lipid bilayers, it is important to choose peptides that comprise the full length of the membrane-interactive fusion domain. Peptides can be quite polymorphic in membrane environments and different conformations and molecular properties may be expressed if shorter than full-length peptide models are chosen. This has been a particular problem in studies of the HIV envelope glycoprotein gp41 fusion peptide in lipid bilayers, where different authors have found quite diverging results. As seen in Fig. 2, this fusion peptide is quite long, but several studies have used relatively short model peptides to study the interactions with lipid bilayers. The hydrophobicity of the full-length HIV gp41 fusion peptide also makes it quite prone to aggregation in solution and in membranes, which adds further difficulties in its handling and potential reasons for differences in results that have been reported for this peptide. Therefore, choosing a certain length of a fusion


peptide model often represents a trade-off between a short sequence that is more easily handled and a long sequence that may better represent the membrane-bound structure of the full-length protein, but may form non-physiological aggregates before it is incorporated into the membrane. The results that are summarized below should be judged in the light of this background. 4.1 HIV Fusion Peptide-Bilayer Interactions

Some of the earliest studies on the HIV gp41 fusion peptide reported that the 16 most N-terminal residues inserted as an oblique α-helix into lipid bilayers [58, 59]. The oblique insertion correlated with the fusion activity because mutations with reduced activity inserted more parallel to the membrane surface as determined by polarized Fourier transform infrared (FTIR) spectroscopy. This finding was in agreement with a prediction from computer models of several viral fusion peptides that a tilted insertion into membranes might be a common feature of many viral fusion peptides [60]. The N-terminal segment of the simian immunodeficiency virus (SIV) fusion peptide analog was also found by neutron diffraction to insert as an oblique α-helix into lipid bilayers [61]. The first 23 residues of the HIV fusion peptide were also predominantly α-helical at low concentrations in membranes, but adopted an extended β-structure at higher concentrations [62, 63]. A still longer HIV fusion peptide analog (first 33 residues) was found by circular dichroism (CD) and FTIR spectroscopy to consist of about 30% α-helical and 50% β-structures [64]. Solution-state nuclear magnetic resonance (NMR) studies of the first 23-residue peptide revealed that these segments adopted predominantly α-helical structures in sodium dodecylsulfate (SDS) [65] and dodecylphosphocholine (DPC) micelles [66], but solid-state NMR found the first 23 residues of the HIV fusion peptide in parallel and antiparallel laterally associated β-sheet conformations when bound to lipid bilayer membranes [67–69]. Combining site-directed 13 C-FTIR spectroscopy and molecular modeling, Gordon et al. [70] suggested that the first 16 residues of the HIV fusion peptide are α-helical and the next seven residues extended under dilute conditions in lipid bilayers. In solution and at higher loading on membranes, the same peptide assembles into antiparallel β-sheets [71]. 4.2 Influenza Fusion Peptide Structure

CD and FTIR spectroscopy showed that the fusion peptide of influenza virus HA2 adopts about 50% helix in lipid bilayers [72–74]. This has been confirmed with 23residue models of the HA fusion peptides [75]. Several critical single site mutations in positions 1, 5 and 7 induce larger amounts of β-structure, when these peptides are incorporated into lipid bilayers [75, 76]. The relative proportion of β-structure is roughly anti-correlated with fusion activity when the same mutation is introduced into the full-length HA and fusion is measured between HA-expressing cells. The fusion activity is not so easily measured with the peptide models themselves because

9


generally only lipid mixing, but not contents mixing, are observed in liposome fusion assays with peptides. In contrast to their full-length analogs, fusion peptides cause contents leakage, i.e. pore formation, when added to lipid bilayers [77, 78]. Short of testing the effect of mutations in full-length fusion proteins, the best correlation between fusion-active and inactive peptide models is their ability to cause hemolysis in red blood cells [74, 76]. Why hemolysis induced by mutant fusion peptides is a better indicator for fusion activity of the full-length protein with the same mutation than lipid mixing and leakage assays of peptides with liposomes is not entirely clear. The influenza HA2 fusion peptide models discussed so far were all only poorly soluble in water and had to be added to liposomes from dimethylsulfoxide or combined with lipids in organic solvents for structural and functional measurements. To eliminate potential artifacts that might arise from solvent exposure of the peptides, we designed a new generation of fusion peptide models which had a very polar carrier peptide appended to the C-terminus of the fusion peptide via a flexible, i.e. glycine-rich, linker. The polar carrier peptide in a sense replaces the ectodomain of the fusion protein in these models. This strategy generates fusion peptides that are highly soluble in water or buffer, while retaining their ability to bind and insert into lipid bilayers with high affinity [79]. The solubilized 20-residue fusion peptide of influenza HA2 is largely random coil in solution, but adopts about 50% α-helical structure when bound to lipid bilayers as do the first-generation fusion peptides [79]. At very high surface concentrations on lipid bilayers, a fraction of the peptide is converted to extended self-associated β-structures at the membrane surface [80]. The new peptide design has permitted their atomic structures to be solved by solution NMR in DPC detergent micelle solutions at pH 7 and 5 [81]. The pH 5 structure is the physiologically relevant structure when influenza HA interacts with target membranes in the endosome, but the pH 7 structure is also informative because the differences between the two pH structures explain why influenza virus requires a low pH not only for releasing the fusion peptide by the conformational change of its ectodomain, but also for membrane fusion. Both structures are characterized by a well-defined N-terminal α-helix that extends to glutamate 11 (Fig. 3). Residues 11–13 form a turn and redirect the polypeptide chain so that it forms a “V” or “boomerang” with an opening angle of about 105◦ . The C-terminal arm does not form a regular secondary structure at pH 7, but residues 14–18 form a 310 -helix at pH 5. The inner volume of the boomerang is filled with bulky hydrophobic residues, a ridge of conserved glycines lines the N-terminal outer face and polar residues characterize the C-terminal outer face of the boomerang at pH 5, but not at pH 7. Therefore, folding of the C-terminal arm at pH 5 likely drives the N-terminal arm deeper into the membrane. These structures determined in detergent micelles by NMR have been confirmed by site-directed spin-labeling in lipid bilayers [81]. The spin-label electron paramagnetic resonance (EPR) measurements also determine the angle between the N-terminal α-helix and the membrane plane (23◦ at pH 7 and 38◦ at pH 5) and the depth of membrane penetration of the fusion peptide. These studies confirmed and substantially refined earlier spin-label studies on a HA2 construct with the fusion peptide and a differently designed fusion peptide

4 Interactions of Fusion Peptides with Lipid Bilayers 11

Fig. 3 Structures of the influenza HA fusion peptide in a lipid bilayer membrane at pH 5. The atomic structures of the wildtype, the glycine1-to-valine mutant (which causes a complete fusion defect), and the glycine1-to-serine mutant (which causes

hemifusion) fusion peptides are shown. All structures were determined by NMR in DPC micelles and their dispositions in lipid bilayers were measured by site-directed spinlabeling (adapted from [81, 86]).

[82, 83]. Fig. 3 shows that the Cα of Asn 12, which forms the apex of the boomerang, is located in the phosphate plane of a fluid lipid bilayer. The N-terminal arm penetrates to about the mid-plane of the lipid bilayer at pH 5, but to a shallower depth at pH 7. Two NMR structures of a different influenza HA2 fusion peptide analog are also available [84, 85]. Since two of the conserved glycines were replaced with glutamates (to make the peptides more soluble) and since in one study the peptide was bound to highly negatively charged SDS micelles, it is not surprising that many details of the structures are quite different, although some general aspects of the wild-type structure of Fig. 3 are preserved. 4.3 Influenza Fusion Peptide Mutants

The structures of the hemifusion-inducing fusion peptide mutant G1S and the fusion-blocking mutant G1V have also been determined recently by NMR in DPC micelles using the polar C-terminal carrier peptide approach [86]. The NMR structure of G1S is very similar to that of the wild-type (Fig. 3). It also forms an amphipathic boomerang with all bulky hydrophobic residues sequestered into the hydrophobic pocket of the structure. Only the glycine ridge on the N-terminal outer edge is disrupted. In contrast, G1V forms an irregular approximately linear


amphipathic helix (Fig. 3). Site-directed spin-labeling shows that this helix is oriented more parallel to the membrane surface. Apparently, an angled and deeply membrane inserted boomerang structure is necessary to promote hemifusion, but a preserved glycine ridge is further required to drive the hemifusion intermediate into a full fusion product. 4.4 Binding of Fusion Peptides to Lipid Bilayers

The binding of the solubilized fusion peptides to lipid bilayers has been measured by fluorescence spectroscopy and isothermal titration calorimetry [79, 80, 87]. The free energy of partitioning of the wild-type influenza HA2 fusion peptide into fluid lipid bilayers is −7.6 kcal mol−1 . In small unilamellar vesicles, this interaction is driven by enthalpy (−18.0 kcal mol−1 ) and opposed by entropy (10.4 kcal mol−1 ). Apparently, there is a real energetic affinity of the fusion peptide for highly curved membrane interfaces, rather than a classical hydrophobic effect-driven peptide association with the lipid bilayer. The entropy loss (−34.8 kcal mol−1 ) could be due to a combination of folding of the peptide and its partial immobilization (reduction of dimensionality of degrees of motional freedom) upon membrane binding. Interestingly, the enthalpy of binding of G1V is much reduced (−9.2 kcal mol−1 ), but that of G1S is similar to that of the wild-type fusion peptide (−15.9 kcal mol−1 ). Recent results from our laboratory show that measurements of binding enthalpy are more sensitive than measurements of free energy to discriminate between active and inactive fusion peptides (A. L. Lai and L. K. Tamm, unpublished results). 4.5 Sendai, Measles and Ebola Fusion Peptide–Bilayer Interactions

The secondary structures and lipid interactions of the N-terminal fusion peptides of the F proteins of two paramyxoviruses, Sendai and measles virus, have also been studied. The fusion peptide of Sendai virus is about 50% α-helical in lipid bilayers [88], but that of measles virus exhibited only weak membrane interactions, which is surprising given its close sequence similarity with the Sendai fusion peptide [89]. A study of the internal fusion peptide of the Ebola virus glycoprotein revealed only a limited amount of secondary structure, but attributed an important structural role to the central conserved proline [90]. A peptide corresponding to the putative secondary internal fusion peptide of Sendai virus is helical in membrane mimetic environments and promotes lipid mixing between liposomes [46]. A similar internal secondary fusion peptide has been postulated to exist in the F protein of measles virus and its interactions with lipid bilayers have been found to be stronger than those of the N-terminal fusion peptide [89]. This anomalous result may be explained if the more hydrophobic N-terminal fusion peptide was not adequately solubilized prior to membrane binding in these experiments. Whether the internal secondary “fusion peptides” of the F proteins of paramyxoviruses really contribute to fusion


by lipid interactions in the context of the full-length proteins is not yet known. Even for the primary internal fusion peptides, such as those shown in Fig. 2, it is presently not clear how well peptide models represent their true structures in the context of the membrane-bound forms of the entire fusion proteins. It may be possible to design looped peptide models with appropriately constrained distances between the ends of the loops in the cases of class II fusion proteins, for which the atomic structures of the post-fusion conformations of the entire ectodomains are known from X-ray crystallography such as, for example, for Semliki Forest and Dengue viruses (see Figs. 1 and 4) [91, 92].

4.6 Perturbation of Bilayer Structure by Fusion Peptides

How do fusion peptides alter the structure of lipid bilayers? How do they induce non-bilayer structures in lipid bilayers, which must occur in some form at intermediate stages of membrane fusion? These are questions of much current debate with few definite answers mostly because it is difficult to trap lipid–protein fusion intermediates for structural studies. A frequently held notion is that fusion peptides alter membrane curvature and make it more negative as required in the stalk–pore model of membrane fusion. For example, 20-residue models of the influenza HA2 fusion peptide decrease the transition temperature from bilayer to curved hexagonal phases and thus stabilize the negatively curved lipid phase [93]. The same result has been observed for the interaction of the 12-residue SIV gp32 fusion peptide with lipid bilayers that are prone to hexagonal phase formation [94]. Given the structure and position of the fusion peptide shown in Fig. 3, it is difficult to envisage how this structure could promote negative curvature in lipid bilayers. If anything, it is expected to promote positive curvature. The experimentally observed depression of the bilayer-to-hexagonal phase transition temperature may be explained if the hydrophobic peptides escaped into the interstitial spaces in the hexagonal phases, which they could not do if they were attached to polar ectodomains that must tie the fusion peptides to the membrane surface. Influenza HA2 fusion peptides have also been found to alter the hydration properties of bilayers at the level of the lipid ester carbonyl groups as measured by FTIR spectroscopy [75, 76]. It is likely that the observed lipid signals reflect a partial dehydration of the membrane surface by the peptides because they also induce more order in the lipid acyl chains [75, 76], which is a common consequence of headgroup dehydration in lipid bilayers. As discussed in Section 2, lipid dehydration could lead to rather disorganized lipid stalks connecting two bilayers. Such dynamically disorganized stalks may exhibit quite different structures than the lipid stalks with neatly curved surfaces that have been proposed in the standard stalk–pore mechanism of membrane fusion. In addition to changing lipid hydration, fusion peptides also decrease the rupture tension of lipid bilayers as has been demonstrated with the influenza HA2 fusion peptide [95]. A trimeric version of the HA2 fusion peptide is more effective in rupturing membranes than its monomeric version [96]. Similarly, a trimeric HIV gp41 fusion

13


Fig. 4 Structures of influenza virus HA (left) and Semliki Forest virus E1 protein (right) in their post-fusion state modeled into a possible fusion intermediate (A) and into a fusion pore (B). The membraneinserted fusion peptides are shown in black and the transmembrane domains in red. Note the large conformational changes of both proteins when compared to their

prefusion structures depicted on the virus surfaces in Fig. 1. The pink dots in the influenza HA fusion peptide structure denote glycines that may mediate helix interactions and the blue squares denote glutamates that may be responsible for the pHdependence of the fusion peptide penetration into lipid bilayers.

peptide construct induced more rapid lipid mixing than a fusion peptide dimer, which in turn was more active than the monomer [97]. It may be expected that further microscopic detail about fusion peptide–lipid interactions will be learned from molecular dynamics simulations, particularly those that are run for long enough times and large enough systems to allow for

5 Interactions of Transmembrane Domains with Lipid Bilayers

extended membrane transformations. Encouraging first steps in this direction have been taken by several groups. Kamath and Wong [98] have simulated the 16-residue HIV gp41 fusion peptide as an α-helix in a POPE bilayer. The peptide maintained an α-helical structure and became tilted in the bilayer during the 1.5 ns of the simulation. The fusion peptide increased the thickness of the proximal leaflet of the bilayer, but left the distal monolayer unperturbed. Huang et al. [99] performed an 18-ns simulation of the 20-residue influenza HA2 fusion peptide in a DMPC bilayer. Although they started from a helical rod structure, the peptide adopted a kinked and tilted helical structure in the bilayer similar to that observed by NMR and EPR spectroscopy [81]. In this study, the lipids of the proximal layer and closest to the N-terminus of the peptide were compressed in both leaflets of the bilayer relative to lipids in unperturbed bilayers or farther away from the N-terminus. Vaccaro et al. [100] started from the NMR structure of the influenza HA2 fusion peptide and simulated it for 5 ns in a POPC bilayer. The kinked tilted α-helical structure was maintained throughout the simulation and the order parameters of the lipid acyl chains were decreased leading to bilayer thinning. No differences between proximal and distal lipids have been reported in this study.

5 Interactions of Transmembrane Domains with Lipid Bilayers

Numerous mutagenesis studies indicate that the transmembrane domains of viral spike glycoproteins are more than just simple anchoring devices to attach these proteins to the viral membrane surfaces. A first indication for this comes from studies on expressed influenza HA, in which the transmembrane domain, i.e. a single α-helix, has been replaced with a glycosylphosphatidylinositol lipid anchor [14]. Cells expressing this mutant are arrested at the hemifusion stage in fusion assays with red blood cells. A similar result has been found for the parainfluenza type 2 fusion protein [101]. Since a deletion of the 12-residue cytoplasmic tail of HA2 still promotes membrane fusion [102], it is clear that the transmembrane domain contributes to the fusion reaction in a major way. This is true even if the cytoplasmic tail influences details of fusion pore opening, which are not yet completely understood [56]. The transmembrane domain must span both leaflets of the lipid bilayer because versions that span only half of the lipid bilayer cause hemifusion just as do the lipid-anchored HAs [42]. Despite the requirement for a near full-length transmembrane domain, there may be no specific sequence requirement of the influenza HA2 transmembrane domain to support membrane fusion [42]. However, some sequence requirements in this domain and appropriate acylations of three cysteines in the short cytoplasmic domain must be met to correctly target influenza HA to cholesterol/sphingomyelin-rich domains in the apical membrane during the biogenesis of influenza virus particles [54, 55]. In retroviruses, a conserved arginine or lysine in the middle of the transmembrane domain of the envelope glycoprotein appears to be important for folding and assembly of the protein in the membrane [103, 104] and/or its ability to support membrane fusion [105, 106]. Similarly, the

15


murine leukemia virus envelope glycoprotein requires a proline [107] and the vesicular stomatitis virus (VSV) G glycoprotein requires two glycines in the middle of the transmembrane domain in order to support the transition from the hemifusion intermediate to the full fusion product [108]. Perhaps, flexibility or kinks are necessary in the transmembrane domains of some fusion proteins for the completion of full membrane fusion. The transmembrane domain of influenza virus HA2 by itself is α-helical and oriented at an angle of 15◦ or less from the membrane normal [109]. It forms oligomers (dimers, trimers and tetramers) in SDS micelles and readily exchanges amide hydrogens when embedded in lipid bilayers. Therefore, it may form a small partially water-accessible helical bundle in membranes. Helical contacts and perhaps water accessibility may be mediated by three conserved serines (or two serines and a cysteine in some strains) that occur in heptad repeats in the central part of the transmembrane domain. Interestingly, the synthetic transmembrane domains of influenza HA2 also increase the lipid chain order parameters [109], which could be a consequence of a partial dehydration at the membrane surface as discussed above in the context of fusion peptide-membrane interactions. Lipid interactions of the tryptophan-rich juxta-membrane domains of HIV gp41 have also been examined. Peptides corresponding to this domain insert into membranes as amphipathic helices parallel to the membrane surface and competitively inhibit fusion promoted by full-length gp41 [110, 111].

6 Structure–Function (Fusion) Relationships of Membrane-interactive Viral Fusion Protein Domains 6.1 Fusion Peptide Mutants

Studies establishing correlations between the structures of membrane-interactive segments of fusion proteins in membranes and their ability to support membrane fusion are only in their beginning stages. The first residue, a glycine, of the influenza HA2 fusion peptide appears to be particularly important for supporting fusion. If glycine 1 is missing, fusion and viral infectivity is completely aborted [40, 41]. Molecular dynamics simulations indicate that the fusion peptide lacking the first glycine is more linear in structure and lies more parallel to the membrane surface compared to the wild-type fusion peptide structure [100]. This mutant also has a higher tendency to self-associate into β-structures on membrane surfaces than the wild-type fusion peptide [75]. If glycine 1 is substituted with a valine, fusion is also completely blocked [39]. This functional defect correlates with a more linear and more surface-located structure of the G1V fusion peptide in lipid bilayers ([86, 87], see also Fig. 3). The defective G1V fusion peptide also has an increased propensity for self-association on membrane surfaces. Substitution of glycine 1 with a serine causes a milder fusion phenotype: fusion can proceed to hemifusion, but not to

6 Structure–Function (Fusion) Relationships of Membrane-interactive Viral Fusion Protein Domains

full fusion [39]. Structural analysis reveals that the G1S fusion peptide still forms a kinked boomerang structure in lipid bilayers similar to that of the wild-type fusion peptide, but that the glycine ridge on the upper face of the N-terminal arm of the V-shaped molecule is disrupted ([86, 87], see also Fig. 3). Very recent data show that mutation of tryptophan 14 to an alanine also completely blocks fusion and that peptides with the W14A substitution are very flexible and lack a fixed angle between the two arms of the boomerang structure of the wild-type (A. Lai et al., unpublished results). Therefore, it appears that a deep insertion into the bilayer at a fixed oblique angle as accomplished by the boomerang structure is required to promote the initial joining of two bilayers (hemifusion), but that an intact glycine ridge may be additionally required to proceed to the formation and expansion of the fusion pore. Further experiments are needed to examine whether or not an intact glycine ridge on the N-terminal arm of the boomerang is an absolute structural requirement for influenza HA-mediated full membrane fusion. Glycine residues in the fusion peptide of HIV gp41 also appear to be critically important to support the fusion activity of the expressed fusion protein and infectivity of the virus [112]. The highly conserved glycines 10 and 13 are particularly sensitive to mutation, whereas the less conserved glycines 3 and 5 are more permissive to substitutions. Substitution of the bulky aliphatic side-chains of valine 2 and leucine 9 with the charged residues glutamate and arginine, respectively, also reduces the fusion activity of gp41 and the infectivity of HIV particles bearing these mutations [113]. The V2E substitution is particularly severe, whereas the L9R and A15E substitutions are milder and reduce syncytium formation to about 50%. The relatively conservative mutation of phenylalanine 11 to a tyrosine blocks syncytium formation of HIV gp41 expressing cells almost completely [114]. The V2E, L9R and F11Y mutations have been introduced into peptide models [115]. All three mutant peptides induced decreased levels of hemolysis of red blood cells compared to hemolysis induced by the wild-type gp41 fusion peptide. The V2E, L9R and F11Y mutant fusion peptides also exhibit significantly decreased contents of α-helix and increased contents of β-structure in lipid bilayers when compared to the wild-type fusion peptide [115]. Molecular dynamics simulations indicate that the V2E and L9R peptides may lie parallel on the membrane surface whereas the wild-type fusion peptide inserts as an oblique helix into lipid bilayers [98]. Therefore, it appears that some general structure-function relationships described for the influenza HA fusion peptide are recapitulated in the HIV gp41 fusion peptide. 6.2 Transmembrane Domain Mutants

As discussed above, the structural requirements on the transmembrane domain are likely less stringent than those on the fusion peptides. It appears that in some cases a full-length transmembrane domain of an almost generic sequence is sufficient to support fusion. However, there may be requirements for some flexibility in the middle of the transmembrane domain and perhaps the presence of some polar/apolar residues in heptad repeats to allow for appropriate homotypic and

17


heterotypic helix–helix interactions. An alignment of the sequences of the transmembrane domains of fourteen different subtype strains of influenza virus HAs shows that the N-terminal half of the domain is more conserved than the C-terminal half [109]. A heptad repeat with the conserved motif ILsIYSsbssSL or ILWISFsbssFL (s = small semipolar, i.e. G, A, S, T or C; b = branched aliphatic, i.e. I or V; underlines denote heptad repeats of apolar residues) is found in the N-terminal half, i.e. the half that resides in the outer leaflet of the viral lipid bilayer, in all fourteen strains. A conserved glycine or alanine is found eight or seven residues, respectively, downstream from these motifs in all strains. The C-terminal half of the transmembrane domain that resides in the inner leaflet of the viral lipid bilayer is otherwise highly variable between the strains and does not show features that distinguishes it from any generic transmembrane domain. Despite their good conservation, a few point mutations (e.g. W3 → A or s9 → L) in the above motifs did not affect fusion, nor did deletion of the first five residues of this motif [42]. As noted above, tryptophan 3 is not totally conserved and is a “s” in the other motif. Significant lipid–protein mismatch may occur in the deletion mutant and thereby destabilize the viral membrane and induce fusion in an unusual manner. The insensitivity of the G9L mutation in an H3 subtype HA is significant and interesting [42]. In contrast, changing glycine 10 into a leucine in an H2 subtype HA leads to a hemifusion phenotype [116, 117]. Apparently, a single glycine-to-leucine substitution in this region with several s residues is sufficient in some, but not in all, cases to reduce full fusion to hemifusion phenotypes. The high conservation of unusual motifs in the N-terminal half of the transmembrane domain of influenza HA remains intriguing and more drastic motif changes may lead to interesting new fusion mutants. Transmembrane domain glycines also seem to be important for fusion mediated by the VSV G protein. Two critical glycine substitutions in the transmembrane domain of this fusion protein block the transition from hemifusion to full fusion [108]. In the case of HIV gp41 and some other retroviruses, a conserved arginine or lysine residue in the center of the transmembrane domain appears to be important for fusion and viral infection [103–106].

7 Possible Mechanisms for Initiating the Formation of Viral Fusion Pores

Fusion is initiated by quite dramatic conformational changes in the ectodomains of the viral fusion proteins. These conformational changes are triggered by low pH in the endosome after receptor-mediated endocytosis for some viruses or by a receptor-mediated activation mechanism in other cases. Two examples for possible intermediate and final fusion structures of a class I and a class II fusion protein are shown in Fig. 4. These structures and their lateral organization in between two fusing membranes should be compared to the dramatically different resting structures that are present on the viral membrane surfaces and that are depicted in Fig. 1. In class I proteins, exemplified by influenza HA2, the coiled coils refold by executing large jack-knife motions (see, e.g. Fig. 2 in [118]). The pH 7 structure

7 Possible Mechanisms for Initiating the Formation of Viral Fusion Pores

of influenza HA2 is metastable. Energy is gained when HA2 assumes the pH 5 structure, which is why this conformational change has been characterized as “spring loaded” [119]. An important feature of the conformational change of class I fusion proteins is that the hydrophobic fusion peptide, which is protected in a hydrophobic pocket in the resting structure, becomes exposed and available for interaction with the target membrane upon fusion activation. In influenza HA2, for which most structural information is available, the conformational change is thought to occur in two steps: the core coiled-coils of HA2 extend in the Nterminal direction and thereby translocate the fusion peptides towards the top of the molecule where they become available for interaction with the target membrane [120]. Next, the outer layer helices refold to form helical hairpins with the core helices and thereby redirect the C-terminal ends with the attached transmembrane domains toward the N-terminal ends with the attached fusion peptides. These are the states shown in Fig. 4 on the left. Interestingly, the N- and C-terminal ends of the ectodomains form a tight cap structure [121] and disruption of this cap by mutagenesis also disrupts fusion and, therefore, is functionally important [122]. This is clear evidence that, at least in the case of influenza HA2, the fusion peptide and transmembrane domain, which each are only about nine residues away from the cap, end up inserted into the fused membranes in very close proximity to each other (Fig. 4, bottom left). Although not quite as much information is yet available on class II proteins, the final situation upon completion of fusion is likely very similar in class I and II fusion proteins [91, 92]. Class II proteins start out as lattices of dimers that lie almost flat on the viral membrane surface (Fig. 1). The dimer contacts and/or the receptor protein (E2 in the case of Semliki Forest virus) protect the fusion peptide loops from hydrophobic exposure. Each subunit in the dimer consists of four domains, i.e. a N-terminal domain, a middle domain that contains the fusion peptide at its tip, a C-terminal domain and the most C-terminal transmembrane domain. Upon activation, the N-terminal and middle domains re-associate into a trimer with the fusion peptide loops exposed close to each other on the tip of the pear-shaped molecule (Fig. 4, top right). The C-terminal domains become redirected and pack into the grooves between neighboring subunits at the base of the trimer. The C-terminal ends of the ectodomain, to which the transmembrane domains are attached, are not visible in the crystal structures, but likely pack into the upper parts of the grooves and thereby position the transmembrane domains very close to the fusion peptides (Fig. 4, bottom right). Therefore, a major purpose of the refolding of the ectodomains of class I and II viral membrane fusion proteins appears to be a mechanical device to (1) insert the fusion peptides into the target membrane, and (2) bring the viral membrane-inserted transmembrane domains into close proximity of the target membrane-inserted fusion peptides and thereby induce a merger of the two membranes. In the following, we briefly discuss protein–lipid and protein–protein interactions that may lead to hemifused and fully fused states similar to those depicted in Fig. 4. It is clear that the ectodomains with attached highly specific fusion peptides are sufficient to proceed to hemifusion, at least in the case of influenza HA-mediated

19


membrane fusion. A specific transmembrane domain is not required and a simple lipid anchor suffices to bring the two membranes into hemifusion contact, presumably by inducing “nipples” in one or both membranes and tilting the ectodomains. Nipple membrane protrusions have been observed by electron microscopy [123], and a tilting of the influenza HA ectodomains relative to the viral and target membranes has been measured by polarized IR spectroscopy under fusion conditions [124–126]. We do not believe that the boomerang-shaped structure of the fusion peptide is compatible with the membrane curvature required to induce a classical hourglass-shaped lipid stalk structure. We rather believe that a high concentration of fusion peptides in the contact zone between two membranes perturbs the bilayer structure by removing water molecules from this region. Therefore, the lipid stalk may be much more dynamic than previously thought and may resemble a “lipid mixer” rather than a highly organized fluid ordered structure. A cartoon of such a dynamic lipid (“mixer”) stalk is depicted in Fig. 5 for the case of influenza HA [127]. Perhaps the fusion loops of class II viral fusion proteins create similar perturbations of the lipids in between the two membranes that are about to fuse. We have postulated that the fusion peptides of class I viral fusion proteins may adopt a full transmembrane orientation and interact directly (presumably as continuous α-helices) with the transmembrane domains [118, 120]. This postulate is based on circumstantial evidence and direct evidence for a direct interaction of the fusion peptides and transmembrane domains in membranes is still lacking as is a detailed structure of the fusion pore. Proteinaceous pores of fusion proteins have been previously suggested for fusion mediated by influenza HA [128] and SNARE proteins [129]. The fundamental difference between the model suggested here and these earlier “gap junction-type” models is that in our model the different transmembrane segments are postulated to interact laterally in the same membrane and not in trans across two different membranes. Obviously, lateral helix association models cannot explain full fusion of type II viral fusion proteins. It appears that

Fig. 5 Dynamic lipid mixer stalk that may be responsible for the hemifusion intermediate in influenza HA-mediated membrane fusion (adapted from [127]). The glycines in the fusion peptide structures are indi-

cated with small dots. The larger spheres in the center are the headgroups of perturbed lipids whose apolar sidechains may rapidly flip between the upper and lower membranes.

References

in this case, insertion of the fusion loops into the membrane interface must suffice to attract the transmembrane domains to this destabilized membrane region. However, heteromeric lateral helix associations may still occur in class II fusion proteins because many of these proteins have other transmembrane domains in their vicinity in stoichiometric amounts, i.e. those of the receptor subunits – E2 in the case of Semliki Forest virus. The discussion about the structures that lead to the opening of fusion pores and the dilation of these pores to complete the membrane fusion reaction is reminiscent of a similar vivid discussion on the action of lytic peptides. Some antimicrobial peptides induce purely proteinaceous pores (barrel stave model), whereas others induce pores with more loosely arranged peptides with many interspersed lipids (carpet model). Clearly, more work needs to be done to define how exactly class I and II fusion proteins work on membranes and creative new combinations of structural and biophysical experimentation will be needed to establish what exact structures lead to membrane fusion in both classes of viral fusion proteins.

8 Acknowledgments

We thank the many members of the Tamm laboratory, present and past, who have contributed to our current understanding of the mechanisms of membrane fusion. This work was funded in part by grants from the National Institutes of Health.

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1

The Calponin Homology (CH) Domain Mario Gimona Consorzio Mario Negri Sud, Santa Maria Imbaro, Italy

Steven J. Winder University of Sheffield, Sheffield, United Kingdom

Originally published in: Modular Protein Domains. Edited by Giovanni Cesareni, Mario Gimona, Marius Sudol and Michael Yaffe. Copyright ľ 2005 Wiley-VCH Verlag GmbH & Co. KGaA Weinheim. Print ISBN: 3-527-30813-2

1 Introduction and Brief History

“Does Vav bind to F-actin through a CH domain?” This title of a 1995 paper by Castresana and Saraste [1] marks the birth of the calponin homology (CH) domain. With the help of structure-based sequence alignments the authors described a 100 residue long protein module that they found in a number of signaling and cytoskeletal proteins. The title of the manuscript prejudiced in a peculiar way the developments in the years to come. Not only did very few people know that the guanine nucleotide exchange factor Vav existed or even interacted with the actin cytoskeleton, but also the calponin community was surprised to find that the functionally almost dispensable N-terminal region of the calponin molecule would serve as the ‘mother of actin-binding modules’. At any rate, once the CH domain was born it stirred up the fields of cytoskeleton research and signaling and made a significant contribution towards a greater mutual understanding of the importance of upstream and downstream targets, respectively. Prior to this historical landmark, partial sequence similarities between the actinbinding domains of classical actin cross-linkers like α-actinin, filamin, spectrin, or fimbrin were noted [2–4] and the name-giving protein, calponin, likewise showed sequence similarities in parts of its N-terminal domain (see Figure 1 for sequence alignments and type classification). With the initial establishment of the CH domain, researchers began to look at actin-binding sites from a new perspective, and soon novel CH-domain family members were identified.


Fig. 1

2 The Calponin Homology (CH) Domain

2 Structure of the Domain – The CH Domain Fold 3

However, the early years of the CH domain were troublesome. A persistent misapprehension of CH domain function, based on an oversimplified interpretation of actin-binding data, delayed a more intellectual discourse with this fascinating protein module [5]. The CH domain was stuck in mediating actin association for all kinds of proteins, irrespective of their subcellular localization or molecular context. More recent and more careful annotations of additional functional sites present on all types of CH domains not only sharpened researchers’ minds but also led to the revival of long-forgotten questions with respect to the regulation of actin binding. Recent years have seen a greater appreciation for and a more liberal view of the functional polymorphism displayed by CH-domain–containing molecules. The realization that actin-binding CH domains can bind the filaments in multiple ways [6–9] considerably shaped the current view on actin-binding modules of the CH-domain family (reviewed in [10]). The final hits aiding in establishing the CH domain as a multifaceted tool were added by the unequivocal determination of the microtubule-anchoring function of the EB1 CH domain [11] and the identification of CH domains in proteins of the nuclear mitotic apparatus (NuMA) [12].

2 Structure of the Domain – The CH Domain Fold

The CH domain is a compact globular fold comprising four major (A, C, E, G) and two or three minor helices (B, D, F) interconnected by loops of variable lengths (Figure 2a). The four major helices of between 10 and 20 residues form the core of the domain, with the smaller helices making minor structural contributions. Helices C, E, and G run roughly parallel to each other, with the N-terminal helix A being roughly perpendicular to them. The A and G helices are the ‘bread and butter’ sandwiching the ‘jam’ represented by the hydrophobic helices C and E (Figure 2a). Among the 15 different CH domain structures currently known (Table 1), which range from those present in a single copy to those of the double tandem domain in fimbrin and represent actin-binding, signaling, and microtubule binding proteins, the structural unit is highly conserved with an rmsd of ∼1 Å between Cα atoms of the four helices in all CH domains [13]. Despite strong structural conservation, CH domains can nonetheless be divided into distinct families, based on structural [7, 14] or sequence alignments [14]. In either situation, families arise due to differences in the lengths and positions of the minor helices and sequence ← Fig. 1 Sequence alignment of CH domains representing the individual types. Invariant key residues are highlighted, and the grey bars at the bottom indicate the positions of helices. The key residues tryptophan (W) at position 11 and the aspartate at the beginning of helix G are the most-conserved residues. Helix C usually follows the con-

sensus DGXXLXXL. The proline (P) residue terminating helix C is invariant in type2 and type3 CH domains and also in the fimtype and EBtype CH domains, but is missing in type1, type4, and type5 CH domains. The asparagine (N) at or within helix E is conserved, but the position varies significantly among the different CH domain types.


Fig. 2 Representative structures of single and tandem CH domains. (a) The single CH domain of calponin [13] with major helices colored individually, from blue at the N terminus (n) to red at the C terminus (c), and labeled from A to G. A short 310 helix unique to calponin is found between helices E and F, and there is no helix D. (b) The compact tandem CH-domain pair of plectin [20], colored and labeled as above.

CH1 is on the left and CH2 on the right; as in calponin there is no helix D in CH1. In comparison to other tandem CH-domain structures, helix A in CH1 is very long. This is probably not a feature unique to plectin, since other studies have suggested a long N-terminal helix for utrophin [19], but the extended amino terminal part of the helix is not part of the true CH domain fold.

variations in interconnecting loops. In all instances the integrity of the domain is maintained by hydrophobic interactions between the core helices, with only a tryptophan residue in helix A being absolutely conserved in all CH domains. This Trp residue generally forms nonpolar interactions with other aromatic or aliphatic residues in helices E and G, thus stabilizing helix A with respect to the triple-helical bundle formed by helices C, E, and G. The four or five residues that are conserved in character in all CH domains are mostly involved in helical packing of the core structure. Table 1 CH domain structures

Protein source Type of analysis

CH domain subtype

Calponin EB1 Rng2 β-Spectrin Utrophin α-Actinin Plectin Dystrophin Utrophin Fimbrin

type3 EBtype type3 type2 type2 type1/type2 tandem type1/type2 tandem type1/type2 tandem type1/type2 tandem type1/type2 tandem

NMR crystal crystal crystal crystal crystal crystal crystal crystal crystal

Reference 1H67 1PA7, 1UEG 1P2X, IP5S 1AA2, 1BKR 1BHD unpublished 1MB8 1DXX 1QAG 1AOA

[13] [17] [18, 76] [14, 16] [7] [22] [20] [23] [6] [21]

2 Structure of the Domain – The CH Domain Fold 5

2.1 Structures of Single CH Domains

The first structures of single CH domains to be solved were of CH2 in spectrin and utrophin [7, 16]; however, these were single CH domains belonging to actin-binding domains containing tandem pairs of CH domains and are discussed in more detail below. The first structure of a true single CH domain, and to date the only solution structure of a CH domain, was that of the archetypal CH domain from chicken gizzard calponin [13] (Figure 2a); more recently this was followed by the crystal structures of EB1, a microtubule binding protein [17], and Rng2, an IQGAP protein from yeast [18]. In all instances the CH domains of these proteins are not thought to be necessary for actin binding, despite the association of Rng2 and calponin with actin or actin-containing structures. Calponin does not require its CH domain for actin-binding and, as discussed below, the single CH domain is not an actinbinding domain per se. In EB1 however, this protein binds to microtubules and not to F-actin, but it is postulated that this interaction is mediated by hydrophobic residues in a similar manner to CH-domain interactions with F-actin [17]. It has been suggested that the interhelical loops, which in structural and sequence terms are the elements that show the most variation among CH domains, might confer the different properties on CH domains [19] and that flexibility in these regions might also confer unique properties, particularly in fimbrin. The solution structure of the calponin CH domain displayed very little if any flexibility in interhelical loops [13], and it is now accepted that it is probably conserved residues in the core helices that allow single CH domains to ‘locate’ on an actin filament. 2.2 Structures of Tandem CH Domains

The basic structure of individual CH domains within the tandem CH domaincontaining proteins recapitulates the sequence-derived phylogeny, in that the CH1 domains are structurally more similar to CH1 domains in other proteins than to the CH2 domain in the same protein. The main differences stem from the relative lengths of the core helices and the number and position of the secondary helices, including short 310 helices flanking helix C in CH2 domains and the presence of an additional helix D in CH2. These differences notwithstanding, the CH domains are remarkably similar in core structure [13]. The most striking and perhaps controversial feature of the tandem CH-domain structures so far elucidated is the positions of the first and second CH domains relative to each other. Fimbrin, plectin, and α-actinin all crystallized as compact monomers with a single molecule in the asymmetric unit [20–22], whereas dystrophin and utrophin crystallized in a more extended conformation and as antiparallel dimers [6, 23]. Biochemically, all the isolated actin-binding domains of these proteins are monomeric; furthermore, the interface between the antiparallel utrophin and dystrophin molecules was such that CH1 of one chain was juxtaposed with CH2 of the other chain in an orientation identical to the orientation of CH domains in fimbrin, α-actinin, and plectin. This


sort of conservation of interactions that exists between domains in proteins that adopt two different states, here, crystallographic monomer versus crystallographic dimer, is known as 3D domain swapping [24]. As such, this is not unusual – the controversy arises, however, as to whether these tandem CH-containing actinbinding domains can adopt different conformations in solution and whether there is significant reorganization of the CH domains upon binding to actin [25]. To add fuel to the controversy, several cryoelectron microscopy reconstructions of tandem CH domains with F-actin have yielded different configurations for the CH-domain orientation on actin. Initially this difference was thought to be a consequence of the length of the inter-CH domain linker [6], because in the first two tandem CH-domain structures to be published, those of fimbrin and utrophin [6, 21], the long linker of fimbrin was believed to allow the two CH domains to fold back on each other and the shorter linker in utrophin resulted in a more open structure. α-Actinin however, which has one of the shortest interdomain linkers of all, crystallized as a compact monomer [22], effectively ruling out the linker hypothesis. An alternative view might be that the conformation of the actin-binding domain reflects the function of the whole protein, i.e., actin-bundling proteins like fimbrin and α-actinin require a compact formation because they bind at right angles to the actin filament [26, 27], and dystrophin and utrophin, which are more akin to side-binding proteins [28, 29], require an open conformation. This argument would seem to support available cryoelectron microscopic data (see [10]); however, the recent crystal structure of plectin [20], which is likely to also be an F-actin side-binding protein to some extent, was found to be a compact monomer (Figure 2b). Although Pereda and colleagues demonstrated rather elegantly [20], as had been suggested previously for utrophin [8, 30], that the two CH domains can undergo movement and can rearrange upon binding to actin [31], further direct experimentation is required to resolve this fascinating problem.

3 Molecular and Signaling Function

CH domains are found in a wide range of molecules, but the unifying theme among them is their involvement in cytoskeletal structure, dynamics, and signaling. The enormous functional plasticity displayed by the otherwise structurally highly conserved domain argues that the CH domain represents a platform for a plethora of functional sites. 3.1 Actin-binding Domains

Sequence- and structure-based profiling has led to the established classification of CH domains into at least five distinct subfamilies [15, 32]. By far the largest number of CH domains belongs to the type1 and type2 classes. With a single exception (smoothelin), these CH domains occur in tandem, and this dual module


forms a high-affinity actin-binding domain (ABD) in a large variety of actin-binding and cross-linking proteins. The amino acid sequences of fimbrin-type CH domains are sufficiently different to place them in a separate subfamily, yet they follow the general consensus and arrangement of ABDs formed by the type1/type2 CH domains. Type1 and type2 CH domains differ not only in sequence, but also in their affinities for actin, as has been shown for the actin cross-linking protein α-actinin. Notably, the ‘type1 fim’ CH domains of fimbrin are able to associate additionally with the intermediate filament component vimentin [33] at a site encompassing residues 143–188 (corresponding to the type1 CH domain in the first ABD). ABDs bind one actin monomer in the filament, with affinities typically in the low micromolar range. When analyzed in isolation, CH domains from ABDs have divergent actin-binding characteristics. All type1 CH domains bind actin with significant affinity, but for type2 CH domains actin binding is almost undetectable. Nevertheless, both N- and C-terminal CH domains are required for the generation of a fully functional ABD, in which the type2 CH domain appears to contribute to the overall stability of the module [34]. Similarly, the ABD in plectin requires only the CH1 for actin binding and dimerization [35]. Here, the crucial importance of the ABS2 residing in the most C-terminal helix of the CH1 was shown for the first time. Interestingly, the CH2 in plectin appears to have a negative influence on actin binding of the ABD, because deletion of the second half increases actin binding of the plectin ABD; however, this may be a reflection of alternatively spliced exons in the first CH domain because, depending on the spliced exon, the affinity for actin can vary considerably [36]. Structural studies of CH domain proteins and their interactions with actin filaments have suggested that a conserved hydrophobic surface is implicated in binding to actin filaments. It is therefore believed that the general mode of binding and the molecular interface employed for contacting the actin filament is conserved among actin-binding domains formed by a CH domain tandem. When the isolated ABD from the actin cross-linking protein α-actinin was used as a molecular targeting vehicle, a variety of otherwise cytoplasmic components (e.g., GFP, Vav) could be targeted to the thin filaments of transfected fibroblasts. However, the similarly arranged ABD from the actin-network-stabilizing molecule filamin failed to serve as a strong targeting vehicle, although the domain is undoubtedly required for filamin binding to actin [37]. This example illustrates that there are functional differences among ABDs from different subfamilies of CH-domain proteins, likely reflecting the differences displayed in the amino acid sequence of this module [38]. One may thus hypothesize that functional diversity occurs among type1/type2 CH domain ABDs and that these differences may account for the subtle differences in actin affinity, mode of cross-linking, and site of attachment along the actin filament. 3.2 Single EB-type CH Domains Function as Microtubule Anchors

In total contrast to the CaP-family and Vav-family CH domains, the CH domains of EB-family proteins (namely EB1, EB2, EB3, RB1, and the yeast EB1 homolog

7


Bim1) have been shown to contact growing microtubule ends. End-binding (EB) proteins are evolutionarily conserved proteins that modulate microtubule dynamics by regulating dynamic instability. In particular, EB1 targets growing microtubule ends, where it is favorably positioned to regulate microtubule polymerization [39] and to confer molecular recognition of the microtubule end [40, 41]. Immunodepletion of EB1 from Xenopus egg extracts has been shown to reduce microtubule length, and this effect was reversed by readdition of recombinant EB1 [42]. EB1 also decreased microtubule catastrophe, resulting in increased polymerization and stable microtubules in interphase cells. The effect of EB1 on microtubule dynamics is highly conserved, suggesting that this protein family belongs to a core set of regulatory factors conserved in higher organisms. In Drosophila, EB1 has been shown to play a crucial role in mitosis by its ability to promote the growth and interactions of microtubules within the central spindle and at the cell cortex [43]. Finally, in Dictyostelium, the largest known EB1 homolog (57 kDa), termed DdEB1, also localizes along microtubules and at microtubule tips, centrosomes, and protruding pseudopods and was found at the spindle, spindle poles, and kineto-chores during mitosis. In addition, EB1 is involved in regulating the interaction of the tumor suppressor adenomatous polyposis coli (APC) with the microtubule apparatus [44, 45]. EB1 binds to the C terminus of the APC protein and may regulate accumulation in cortical clusters of extending membranes, but endogenous EB1 does not accumulate in the APC clusters [46]. 3.3 Kinases, Phospholipids, and Other Cytoskeletal Components

Tandem CH domains are also present in the actopaxin/parvin family [47, 48], but here, both CH domains branch into separate subfamilies (type4, type5) and expose (overlapping) binding sites for F-actin and the focal adhesion proteins paxillin and integrin-linked kinase (ILK) [49, 50]. Thus, the CH domains in this family display functions different from those of type1, type2, or type3 CH domains. Unfortunately, more detailed molecular information about the residues involved in these interactions is lacking. CH domains display different functions when present in single as compared to tandem motifs, and different proteins contain CH domains of different ‘types’. All ABDs strictly follow the consensus type1/type2 arrangement. However, the relative contribution of each CH domain to actin binding is not known. Detailed correlative sequence analysis has revealed that CH domains in ABDs can harbor additional conserved binding motifs for phosphatidylinositol (type2 CH domains; see below) and also novel autonomous actin-binding sequences, like the recently discovered DFRxxL motif from myosin light chain kinase [51], reviewed in [15]. These findings strongly suggest that even CH domains in ABDs may expose additional, as yet unidentified, features that contribute to the selectivity and specificity of binding to actin and possibly to other thin-filament-associated components. A perfect example of this are the actin-binding domains from plectin and dystonin, which bind β4 integrin [52]. Mutations in the integrin binding pocket, formed


by a sequence stretch unique to plectin and dystonin and corresponding to the region preceding and partially overlapping the ABS2 (residing in the C-terminal helix in the CH1 of plectin), significantly decrease β4 integrin binding but do not influence the F-actin binding ability of plectin’s ABD. Interestingly, actin binding and integrin β4 association are mutually exclusive in plectin, suggesting that CH domain function can be switched ‘on site’ — the mechanism, however, remains obscure. In contrast to the situation in type1 and type2 CH domains, the actin binding affinities of type3 CH domains are in the millimolar range, and researchers thus earlier questioned whether their physiological function is indeed related to actin binding. Although the contribution of the CH domain in calponin-family proteins is still an open question, we have seen an accumulation of data in recent years that support the original skepticism. It is clear now that the CH domain is neither sufficient nor necessary for actin binding activity in this protein family [53–56]. This development has helped to make manifest the view that type3 CH domains may serve a primarily regulatory function, and indeed several laboratories have identified a plethora of binding partners for type3 CH domains. In the namegiving protein calponin (CaP), for example, the type3 CH domain interacts with extracellular regulated kinases ERK1 and ERK2 in vitro [57] and is hypothesized to be modulated by an association with LIM kinase in vivo [58]. Work from the Gusev laboratory revealed that Hsp90 binds directly to the CH domain of smooth muscle h1CaP and affects actin binding. The authors hypothesized that, in the presence of Hsp90, CaP is trapped in a complex, which makes the molecule unavailable for interaction with G-actin. In this way, Hsp90 could decrease the CaP-induced polymerization of actin [59, 60]. Calponin interacts in vitro with the dimeric S100family members calcyclin (S100A6), which exposes two functional Ca2+ -binding EFhands per monomer, and S100A2. EF-hands have a similar amino acid arrangement as zinc fingers and require the presence of conserved Cys or His residues in a particular spatial arrangement. EF-hands of S100A2 can also bind Zn2+ , suggesting that Zn2+ binding is involved in the regulation of CaP via S100 molecules. In agreement with this scenario, the type3 CH domain in the Rho family nucleotide exchange factor Vav binds intramolecularly to a zinc-finger–like domain [61] and has been shown to interact with the guanine nucleotide dissociation inhibitor (RhoGDI) in a two-hybrid interaction screen [62]. Vav proteins are activated by an N-terminal deletion that removes all (in Vav-2) or part (in Vav-1) of the CH domain, and this deletion occurs naturally in the onco-vav gene [63]. The CH domain is required for the modulation or down-regulation of Vav’s GEF activity [64, 65] by mediating a conformational switch in the molecule, which involves interaction with the C-terminal zinc-finger domain and results in a steric block of the catalytic DH-PH domains [66]. Notably, the short loop connecting helices A and B in the Vav CH domain is responsible for this Vav-specific function, and replacement of the loop with the homologous region from CaP makes the molecule constitutively active [67]. Binding of the phosphoinositide PtdIns(4,5)-P2 negatively regulates actin binding and bundling activity of the antiparallel actin filament cross linker α-actinin, likely

9


by mediating changes in the molecular structure [68]. The conserved PiP2 binding site resides in the CH2 of α-actinin, in good agreement with the supporting function of CH2 domains in ABDs. Site-directed mutagenesis in this region has revealed three critical basic residues. Mutant proteins carrying sequence alterations in these amino acids, leading to defective PiP2 binding, display increased actin binding and bundling activity in vitro. 3.4 CH Domain-containing Proteins and Human Diseases 3.4.1 The Dystrophin ABD and Muscular Dystrophy Dystrophin is a large cytoskeletal linker protein that connects the subsarcolemmal actin cytoskeleton of skeletal muscle to the transmembrane adhesion receptor dystroglycan [69]. Mutations in dystrophin give rise to the crippling and fatal Xlinked disease Duchenne muscular dystrophy (DMD). The majority of mutations in the DMD gene give rise to premature stop codons, resulting in transcript instability and complete loss of protein. The milder allelic form of DMD, Becker muscular dystrophy, which is caused by in-frame deletion or missense point mutations, does allow the synthesis of mutated protein. Most point mutations in the CH domaincontaining actin-binding region of dystrophin lead to a relatively severe phenotype [70–72], emphasizing the importance of the actin-binding domain for dystrophin function. However, it is doubtful that in the majority of cases the actin-binding domain is functional at all, because the four missense mutations; L54R, A171P, A168D, and Y321N and in-frame deletions of exon 3 (residues 32–62) and exon 5 (residues 89–119) are all expected to disrupt either the hydrophobic core of the protein or its overall structure [23]. Consequently, these mutations have not been particularly useful in determining function. 3.4.2 The Filamin ABD and Otopalatodigital Syndromes Otopalatodigital (OPD) syndromes and related disorders are a diverse group of X-linked diseases affecting craniofacial, skeletal, brain, visceral, and urogenital structures. The affected gene encodes the cytoskeletal protein filamin A, with approximately half of the 17 mutations so far described being missense mutations residing in the CH2 domain of the filamin ABD [73]. No structure of a filamin CH domain is yet available, but mapping the mutations to a model of filamin based on the structures of spectrin or dystrophin CH2 suggested that some mutations were likely to not simply result in loss of actin-binding function [73], which points to additional and as-yet-unidentified roles for CH domains. 3.4.3 The α-Actinin ABD and Glomerulosclerosis Mutations in α-actinin 4 have been described in familial focal segmental glomerulosclerosis (FSGS). FSGS is a common nonspecific renal lesion characterized by decreasing kidney function and often leading to end-stage renal failure. α-Actinin 4 has been implicated in some cases of autosomal dominant FSGS, with point mutations occurring in helix G of CH2 [74]. All three of the mutations characterized –

4 Emerging Research Directions and Recent Developments

K228E, T232I, and S235P – are on the solvent-accessible surface of helix G and are not expected to affect core structure, but are also not in a region implicated in direct interactions with actin, and so presumably are involved in some other as-yet-unidentified role of α-actinin in the kidney. 3.4.4 The β-Spectrin ABD and Spherocytosis Hereditary spherocytosis (HS) includes a group of heterogeneous hemolytic anemias ranging in severity from asymptomatic to severe. In all cases the red blood cell has a distinct morphology, with varying degrees of surface area reduction leading to a spherocytic phenotype and osmotic fragility. Of the four characterized subsets of HS patients, two are characterized by a deficiency in β-spectrin. Several mutations in spectrin have been described and shown to be the molecular defect in HS with spectrin deficiency [75]. Of these mutations, two were found in the second CH domain, W182G and I220V, with both residues being important for maintaining the hydrophobic core of the CH domain. Changing Trp182 to Gly in the short helix B in particular would be expected to have a severe effect on the stability of the CH domain, with consequent effects on the function of the whole ABD. Many other proteins that contain CH domains are implicated in diseases, for example EB1 is a tumor-suppressor protein, and plectin is involved in epidermolysis bullosa with muscular dystrophy, but to date, disease-causing mutations in these proteins have not been identified within the CH domains.


The presence of a CH domain in any given protein is still taken as a strong indication that the molecule associates with the actin cytoskeleton, despite controversial interpretations of binding data and subcellular localization studies which call this simplified view into question. It is more than evident, from the diverse list of binding partners that have been identified for the various CH domains, that the module is a platform for a number of interaction sites with cytoskeleton and signaling components. The most interesting study of the past decade may be the identification of the EB1 CH domain. EB CH domains strictly follow the consensus of conserved residues and, even though the protein has not been ascribed any direct association with the actin cytoskeleton, are placed in the CH-domain family tree, representing a separate branch. The work of Hayashi and Ikura has demonstrated that the module folds almost identically to the CH domains described thus far for spectrin, fimbrin, and calponin – but the CH domain in EB1 is a microtubule anchor instead. It is difficult to envisage any similarity in the surface profiles between an actin filament and a microtubule. Hence, morphofunctional plasticity, established during a coevolutionary process of the diverse eukaryotic cytoskeleton filament systems, and the factors regulating their assembly and dynamics, may be the key to understanding this apparent paradox. Other important work, such as with the plectin ABD [52], should serve as future guidelines on how meaningful studies can

11


be tailored to increase our knowledge of CH domain function and interactions. Future research activities might put greater emphasis on identifying the in vivo functions of the diverse CH domains and aim at determining in more detail the (sequence) parameters that drive functional plasticity in this family.

5 Conclusions

Not even ten years have gone by since the CH domain was delineated in detail. Like all other domains and modules analyzed in this book, the CH domain obeys the rule of protein module definition. It is a stable fold, and the function can be transported to other molecules by simple fusion of the coding sequence. We must, however, be aware of the fact that protein linguistics and positional semantics may lead us into novel territory in which the strict functional definitions may not hold. There is rapidly increasing evidence for the existence of a novel class of modules that fulfill the criteria of autonomous function but not of folding. These intrinsically unfolded protein (IUP) modules appear to fold exclusively at their ligands – and a good portion of these associate with the cytoskeleton! One should therefore keep in mind that the definition of domain borders is perhaps the most relevant parameter for assessing the proper in vivo function and that structurally dispensable flanking regions may contribute significantly to the fine-tuning of domain function.

Acknowledgements

Work cited in this chapter was funded by the BBSRC, MRC, and Wellcome Trust (SJW). We are grateful to Mike Broderick for critical reading of the manuscript and to Jose Pereda for providing Figure 1b. MG is recipient of the Marie Curie Excellence Grant MEXT-CT-2003-002573 of the European Union.

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36 FUCHS, P., ZORER, M., REZNICZEK, G. A., SPAZIERER, D., OEHLER, S., CASTANON, M. J., HAUPTMANN, R., WICHE, G., Unusual 5 transcript complexity of plectin isoforms: novel tissue-specific exons modulate actin-binding activity. Hum. Mol. Genet. 1999, 8, 2461–2472. 37 GIMONA, M., unpublished. 38 STRADAL, T., KRANEWITTER, W., WINDER, S. J., GIMONA, M., CH domains revisited. FEBS Lett. 1998, 432, 134–137. 39 NAKAMURA, M., ZHOU, X. Z., LU, K. P., Critical role for the EB1 and APC interaction in the regulation of microtubule polymerization. Curr. Biol. 2001, 11, 1062–1067. 40 MORRISON, E. E., MONCUR, P. M., ASKHAM, J. M., EB1 identifies sites of microtubule polymerisation during neurite development. Brain Res. Mol. Brain Res. 2002, 98, 145–152. 41 BIENZ, M., The subcellular destinations of APC proteins. Nat. Rev. Mol. Cell Biol. 2002 3, 328–338. 42 TIRNAUER, J. S., GREGO, S., SALMON, E. D., MITCHISON, T. J., EB1–microtubule interactions in Xenopus egg extracts: role of EB1 in microtubule stabilization and mechanisms of targeting to microtubules. Mol. Biol. Cell 2002, 13, 3614–3626. 43 ROGERS, S. L., ROGERS, G. C., SHARP, D. J., VALE, R. D., Drosophila EB1 is important for proper assembly, dynamics, and positioning of the mitotic spindle. J. Cell Biol. 2002, 158, 873–884. 44 MIMORI-KIYOSUE, Y., SHIINA, N., TSUKITA, S., The dynamic behavior of the APC-binding protein EB1 on the distal ends of microtubules. Curr. Biol. 2000, 10, 865–868. 45 ASKHAM, J. M., MONCUR, P., MARKHAM, A. F., MORRISON, E. E., Regulation and function of the interaction between the APC tumour suppressor protein and EB1. Oncogene 2000, 19, 1950–1958. 46 BIENZ, M., Spindle cotton on to junctions, APC and EB1. Nat. Cell Biol. 2001, 3, E67–E69. 47 NIKOLOPOULOS, S. N., TURNER, C. E., Actopaxin, a new focal adhesion protein that binds paxillin LD motifs and actin and regulates cell adhesion. J. Cell Biol. 2000, 151, 1435–1448.

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activity. J. Biol. Chem. 2003, 278, 24039–24045. WINDER, S. J., The membrane–cytoskeleton interface: the role of dystrophin and utrophin. J. Muscle Res. Cell Motil. 1997, 18, 617–629. KAPLAN, J. M., et al., Mutations in ACTN4, encoding α-actinin-4, cause familial focal segmental glomerulo-sclerosis. Nat. Genet. 2000, 24, 251–256. BEGGS, A. H., et al., Exploring the 73 molecular basis for variability among patients with Becker muscular dystrophy: dystrophin gene and protein studies. Am. J. Hum. Genet. 1991, 49, 54–67. PRIOR, T. W., BARTOLO, C., PEARL, D. K., 75 PAPP, A. C., SNYDER, P. J., SEDRA, M. S., BURGHES, A. H. M., MENDELL, J. R., Spectrum of small mutations in the dystrophin coding region. Am. J. Hum. Genet. 1995, 57, 22–33. ROBERTS, R. G., GARDNER, R. J., BOBROW, M., Searching for the 1 in 2,400,000: a

review of dystrophin gene point mutations. Hum. Mutat. 1994, 4, 1–11. 74 ROBERTSON, S. P., et al., Localized mutations in the gene encoding the cytoskeletal protein filamin A cause diverse malformations in humans. Nat. Genet. 2003, 33, 487–491. 75 HASSOUN, H., et al., Characterization of the underlying molecular defect in hereditary spherocytosis associated with spectrin deficiency. Blood 1997, 90, 398–406. 76 DOKLAND, T., personal communication.

Websites Directly Related to the Domain http://www.proteinmodules.org General platform site for protein domains and modules and official web site of the Protein Modules Consortium.

1

PDZ Domains: Intracellular Mediators of Carboxy-Terminal Protein Recognition and Scaffolding Laurence A. Lasky, Nicholas J. Skelton, and Sachdev S. Sidhu


1 Introdution

Protein–protein binding interfaces are involved with virtually every aspect of intraand extracellular physiology. These interfaces range in size from the very large binding sites formed between antibody combining regions and their cognate antigens to the very small interfaces recognized by a variety of more compact motifs including, for example, the SH2 and SH3 domains. The vast majority of these protein modules bind to unmodified or post-translationally modified regions within proteins, as opposed to sites at the very amino- or carboxy-terminal ends of proteins. The potential benefits of recognition via the beginnings or ends of proteins are two-fold. First, recognition of these regions would allow for enhanced specificity by taking advantage of binding interactions involving the free ends of proteins. This might allow for a relatively small binding site, since specificity can be induced both by mainchain and sidechain interactions as well as by amino- or carboxy-type interactions. Second, recognition of terminal regions of proteins allows for the rest of the protein to be unencumbered by an internally bound protein-recognition motif. This would enable efficient scaffolding with other functionally related proteins together with simultaneous activity of the terminally-bound protein. It thus appeared likely to many investigators that scaffolding proteins that recognized the ends of other proteins were likely to exist. The discovery of a large family of proteins containing a compact domain, termed the PDZ domain, has fulfilled this expectation, at least with respect to C-terminal interactions. Determination of the sequences of several large proteins in the early 1990s led to the discovery of a 90–100 amino acid long domain that was repeated within the proteins and was conserved in many other molecules derived from a variety of organisms and cell types. The initial discovery of this motif was in three polypeptides: the mammalian protein post-synaptic density-95 (PSD-95), the Drosophila Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 PDZ Domains: Intracellular Mediators of Carboxy-Terminal Protein Recognition and Scaffolding

melanogaster epithelial tumor suppressor protein, Discs Large (DLG), and the mammalian epithelial tight junction protein, zonula occludens-1 (ZO-1) [1–7]. Initial sequence comparisons revealed a reasonable degree of conservation between these diverse domains, including a highly conserved amino acid sequence, GLGF, and the motifs were initially referred to as GLGF domains. Subsequently, because this short sequence is not identical in all of these domains, as well as to honor the initial sites of discovery, the motifs were renamed PDZ domains (PSD-95, DLG, ZO-1). Large-scale genomic analyses, together with in silico bioinformatics work, has allowed for the determination of the number of potential PDZ domains in the organisms for which we have reasonably complete genomic DNA sequence information. Interestingly, and in contrast to both total gene numbers and genome sizes, the numbers of PDZ domains in various organisms seem to increase dramatically with increasing complexity, although the number of genomes so far analyzed is far too small to be certain of the validity of this trend. However, there are ∼90 domains in the nematode Caenorhabditis elegans, ∼130 in the fly D. melanogaster, and over 400 in the human genome. Some of this increased complexity can be explained by increasingly larger gene families. For example, although the nematode appears to have only two members of the LAP (leucine-rich repeat and PDZ domain) family, higher organisms appear to have additional homologs encoded within their genomes. This appears to be so with many other PDZ domain-containing proteins, including DLG, ZO-type, the MAGI proteins, and LIN-7, among others. The potential reasons for this increased complexity, including tissue specificity, diversity of function, etc., remain to be fully elucidated. PDZ domains are virtually always embedded in proteins that are assembled from multiple protein motifs (a complete list of PDZ-containing proteins together with other domains, functions, and annotations can be found at http://smart.emblheidelberg.de/). These protein motifs can include other PDZ domains, and there are examples of proteins containing only PDZ domains, such as MUPP1, in which the protein consists of 13 PDZ motifs [8]. Other multi-PDZ domain-containing proteins include INAD, par-3, and NHERF. It seems likely that this type of multi-PDZ protein would be involved in the assembly of functionally related proteins via C-terminal recognition, and this has been proven both genetically and biochemically for the INAD protein of Drosophila (see below), as well as several others. A second large family of PDZ domain-containing proteins is the MAGUK (membrane-associated guanylate kinase) family. This large subgroup of PDZ-containing proteins contains the founding members of the PDZ family, and it is unified by the inclusion of a protein motif with distant homology to guanylate kinases (the GUK domain), although there is no evidence that this domain has enzymatic activity. Many of these proteins, including DLG, ZO-1, and the MAGI (membrane-associated guanylate kinases with inverted orientation) proteins, are associated with the tight junctions of epithelial cells and are presumably involved in assembly and maintenance of these important structures. This large family contains proteins with other potential protein interaction domains, including, for example, SH3, WW, calmodulin kinase (CAMK), and L27 motifs. Finally, a third large group of PDZ proteins contains a variety of other sequence motifs (but not a

2 Structural Analysis of PDZ Domains 3

guanylate kinase-like domain) juxtaposed with one or more PDZ domains. Included in this family are proteins containing leucine-rich repeats (the LAP proteins), LIM, or crib motifs. A unifying feature of all of these PDZ-containing proteins is that the molecules contain a variety of protein interaction motifs with no clear evidence of enzymatic activity in any of the family members. These data, together with their localization to diverse intracellular sites, such as epithelial junctions and the neuronal post-synaptic density, suggest that the PDZ-containing proteins are predominately site-specific scaffolding proteins that assemble functional complexes, many of which appear to be important for the assembly, maintenance, and function of these subcellular anatomies.

2 Structural Analysis of PDZ Domains

Structural studies of many PDZ domains have identified a common 3D fold that undoubtedly provides the physical underpinning for the sequence motifs that can be used to identify the domains [2, 9]. Today, 34 independent structures of 20 different PDZ domains have been reported, and all consist of a six-stranded β barrel (strands βA through βF) capped by one short (α1) and one long (α2) helix (Figure 1) [10, 11]. Although the β-barrel core is common to all PDZ domain structures, there is sequence variation within the elements of regular secondary structure and also in the length and composition of the loops connecting them. In addition, a number of the domains also contain additional N- or C-terminal appendages that pack against the conserved core [9, 12, 13].

Fig. 1 Schematic view of the PDZ domain fold. In this structure of the Erbin PDZ domain, $ strands are shown as arrows and " helices as coils. The phage-derived peptide ligand (acTGWETWVCOOH ) is shown in stick form with the C-terminal carboxylate at the top of the view. Coordinates are taken from PDB accession number 1N7T.


Over half of the PDZ entries present in the structural database have a peptide ligand associated with the domain. In all instances, the GLGF motif of the domain wraps around the C-terminal carboxylate of the ligand [14]. The remainder of the ligand lies in the cleft between strand β2 and helix α2, with the peptide forming an antiparallel β-sheet interaction with β2. As a consequence of this fixed backbone hydrogen-bonding pattern, the ligand sidechains maintain a fixed register with respect to the PDZ domain and are able to contact only a limited subset of the domain sidechains (Figure 1). To standardize the categorization of all PDZ interactions, the ligand residues are numbered from 0 at the C terminus counting by increasingly negative integers toward the N terminus [15]. The importance of interactions at the 0 and −2 sites was appreciated from the earliest studies [16– 19] and led to the classification of PDZ domains into two classes based on their ligand preference at the −2 site. Subsequent studies have shown that the −1 site can also make a crucial contribution to binding [20–23]. Moreover, PDZ domains having a preference for ligands with a particular −3 sidechain are common [16, 20– 22, 24, 25], and preference for residues N-terminal to −3 have been observed, especially if the β2–β3 loop is long [13, 22, 26, 27]. Since the sidechains of ligand residues 0, −1, −2, and −3 all have relatively fixed interactions with their PDZ domain host, establishing the ‘rules’ for predicting ligand affinity and selectivity should be possible, given a suitable training set of PDZ domain–ligand interactions (e.g., see [28]). In addition to the ‘canonical’ ligand binding mode, the co-complex between αsyntrophin and neuronal nitric oxide synthetase (nNOS) PDZ domains indicates that non-C-terminal interactions are also possible. In this particular example, an ancillary β hairpin at the N terminus of nNOS lies in the peptide binding groove of α-syntrophin with the reverse turn of the hairpin positioned approximately where a C-terminal carboxylate would ordinarily reside [29]. More recently, structural studies of the seventh PDZ domain from GRIP demonstrate that its peptide-binding groove is not well-formed [30]. NMR chemical shift mapping suggests that GRASP1, the cognate ligand, binds to a hydrophobic surface formed by the strand β4–β5 hairpin and the face of helix α2 away from the peptide binding groove; precise details of the interaction were not reported. Finally, there have been reports that not all PDZ domains behave as independently folding units. Structures determined for tandem PDZ domains from PSD-95 and syntenin suggest that the domains can have a relatively fixed orientation with respect to each other, which may have implications for their function as scaffold proteins [31–33]. The quaternary structure observed for PDZ6 from GRIP-1 suggest alternative modes of such supramolecular assembly [34]. Finally, the PDZ4 and PDZ5 domains of GRIP-1 could not be expressed efficiently alone and had NMR spectra indicative of partial folding. However, the tandem domain expressed well and appeared highly folded by the same criteria [35]. Subsequent studies have shown that the two domains are intimately packed (with a different geometry than that observed in PSD-95 and syntenin tandem domains); the occluded peptide binding pocket of PDZ4 suggests that its primary function is not ligand binding, but rather, stabilization of the PDZ5 structure [36].

3 Analysis of PDZ Domain–Ligand Interactions with Mutagenesis and Synthetic Peptides 5

3 Analysis of PDZ Domain–Ligand Interactions with Mutagenesis and Synthetic Peptides

In vitro studies of PDZ domain–ligand interactions have proven invaluable for complementing and extending the knowledge acquired from structural analyses, and also for refining and even revising our views on physiologically relevant in vivo interactions. Studies with combinatorial peptide libraries, in particular, have been instrumental in defining the fine specificity of individual domains and revealing binding contributions from up to six C-terminal ligand residues. Early studies made use of synthetic peptide libraries [16] or a ‘peptides-on-plasmids’ library generated in Escherichia coli [24, 27–39], but more recently, C-terminal peptide libraries displayed on M13 [20] or lambda phage [40] have also been developed. Peptides fused to the C terminus of the D-capsid protein of lambda phage resulted in high valency display that enabled the selection of even very low-affinity ligands for the seven PDZ domains of the human INAD-like (INADL) protein [40]. Different degrees of consensus were observed, with different PDZ domains selecting ligands with homology at two, three, or four C-terminal residues. Unfortunately, the resulting low homology data, which may have been hampered by the high display levels of the peptides, were not as enlightening as the genetic/biochemical data derived from the homologous Drosophila protein INAD (see below). In contrast, low valency display achieved through C-terminal fusion to the M13 major coat protein yielded high-affinity peptide ligands that bound to MAGI-3 PDZ2 with affinities in the submicromolar range and showed a clear consensus in all four C-terminal positions [20]. C-terminal M13 phage display was also used to study the binding specificity of the Erbin PDZ domain, which had originally been isolated as a putative binding partner for ErbB-2 in a yeast two-hybrid screen (see below) [41]. Interestingly, the Erbin PDZ-binding consensus defined by phage display ([D/E][T/S]WVCOOH ) differed significantly from the C terminus of ErbB-2 (DVPVCOOH ) and instead matched the conserved C termini of three p120-related catenins (DSWVCOOH ). Subsequent in vitro affinity measurements with synthetic peptides showed that the Erbin PDZ domain binds to the catenin C-terminal sequence at least 100 times tighter than to the ErbB-2 C terminus. Although the yeast two-hybrid method is a powerful technology for discovering natural protein–protein interactions, experience with the Erbin PDZ domain shows that the method can be highly sensitive to even extremely low-affinity PDZ domain–ligand interactions and, thus, it is worthwhile corroborating yeast two-hybrid results with the results of other methods. In a subsequent study, NMR spectroscopy, phage display, and in vitro affinity measurements were combined to provide a detailed analysis of the structure and function of the Erbin PDZ domain [22]. The NMR co-complex with a phageoptimized peptide revealed five distinct binding sites on the protein, which accommodated the five C-terminal ligand residues (Figure 2). Binding assays with a panel of synthetic peptides showed that each of the five ligand sidechains contributes to binding, with the last two sidechains and the C-terminal carboxylate providing


Fig. 2 Summary of alanine-scanning mutagenesis data for Erbin PDZ domain binding to the phage-optimized peptide ligand. The solvent-accessible surface of Erbin residues that were mutated to alanine in a combinatorial phage library are colored according to their effect on ligand binding, with red, yellow, and blue indicating a large, moder-

ate, or no effect on binding, respectively. Likewise, the ligand sidechains are colored according to the effect that peptide alanine substitutions had on binding to wild-type Erbin PDZ domain, with red and yellow indicating a greater than 100-fold or greater than 10-fold effect on IC50 , respectively.

the majority of the binding energy. A combinatorial mutagenesis strategy was also used to assess the effects of alanine substitutions for 44 PDZ domain sidechains in and around the peptide-binding site. When the results of the peptide affinity and protein mutagenesis studies were mapped onto the NMR structure, they provided an extremely comprehensive view of the molecular elements involved in PDZ domain–ligand recognition (Figure 2). More traditional point-mutation studies have been used to understand the binding of α-syntrophin to both C-terminal peptides and nNOS, and also to investigate the impact of charge substitutions on ligand binding [42, 43]. In an interesting extension of such an approach, Ranganathan and colleagues looked at sites of pairwise covariation among several hundred PDZ domains [44]. This analysis identified energetically coupled pathways emanating from the peptide-binding groove to sites on the opposite side of the domain. Experimental validation of these pathways was afforded by ligand-binding measurements in double-mutant cycles; their presence raises the possibility of functionally relevant allosteric processes associated with ligand binding [44]. Finally, progress has also been made on the development of computational algorithms for predicting and engineering PDZ domain specificities [28]. Given the importance of PDZ domains in mediating numerous protein assemblies, attempts have been made to antagonize their interactions so as to elicit a biological response. Optimal peptide ligands identified from phage display

4 Molecular and Signaling Functions of PDZ Domains 7

libraries have been used to such ends, to block the interaction between Erbin and δ-catenin [21]; such reagents were able to induce distinct phenotypes in neuron outgrowth assays (K. Kosik, personal communication). Small peptides have also been used to disrupt the in vivo interaction between an NMDA receptor and a PDZ domain of PSD-95 [45]. Recently, organic mimics of C-terminal peptide ligands have been proposed that can bind in a selective manner to MAGI-3 PDZ2 and inhibit formation of complexes with PTEN peptides [46]. In a further recent development, halothane anesthetics have been found to bind to PSD-95 and PSD-93 and to inhibit interaction with NMDA receptor or nNOS. The binding was localized to the peptide-binding groove, explaining the mechanism of antagonism of PDZmediated protein assembly, and possibly also explaining the mechanism by which these anesthetics exert their physiological response [47].

4 Molecular and Signaling Functions of PDZ Domains

Although an ever-increasing diversity of PDZ-mediated interactions has emerged in the literature (for example, see [7]), we focus here on a few examples of interactions that have been validated by biochemical as well as more physiological methods, particularly genetics. As outlined in the structure section above, many of the interactions determined by biochemical techniques, such as the yeast two-hybrid and coprecipitation methods, are encumbered with numerous potential artifacts, such as very low, probably nonphysiological, binding affinities. Thus, although it is clear that many of the interactions predicted by these approaches are undoubtedly correct, there are likely to be many incorrectly described interactions as well. Luckily, there are a variety of physiologically well characterized PDZ-mediated interactions that allow for a number of interesting conclusions regarding the diverse functions of these motifs. 4.1 INAD as a Molecular Scaffold

Anyone who has attempted to swat a fly cannot help but marvel at the insect’s ability to escape certain death. This capability is in large part due to the rapid and efficient transduction of visual information. As in other organisms, vision in the fly D. melanogaster is initiated by the absorption of photons by the G protein-coupled receptor, rhodopsin. This results in a conformational change in the receptor, which induces the association with a Gq type of GTPase signaling protein. This interaction induces a modulation in downstream signaling molecules, including protein kinase C (PKC) and phospholipase C (PLC), which results in opening the transient receptor potential (TRP) and transient receptor potential-like (TRPL) calcium ion channels. The flow of ions through these channels is subsequently transmitted to the central nervous system, where the appropriate response to the initial visual input is engendered. Once the visual input signal is abolished, the system


undergoes a negative feedback response to intracellular calcium concentrations by down-regulating the activities of rhodopsin and phospholipase C [48]. It is not surprising that the efficiency of visual signal transduction as regulated by this large number of signaling proteins would be enhanced by placing the proteins very close to one another. Genetic studies of strains of Drosophila carrying mutations at the InaD locus suggested that this gene was critical for proper function of the light-gathering ommatidial cells in the eye [49–52]. Morphological analysis of TRP channel localization demonstrated that this critical visual component appears to be mis-localized in the eyes of flies with various InaD mutations [53]. Isolation of the gene and analysis of the protein encoded by the InaD locus demonstrated that this gene encodes a molecule containing five PDZ domains [49–52]. This result immediately suggested the possibility that the INAD protein was involved in assembly of components of the visual signal transduction system into a macromolecular structure that enhanced signaling efficiency by closely associating these components. Biochemical analysis has demonstrated that at least seven proteins bind to the INAD protein, with many of the proteins binding via an interaction between their C termini and the INAD PDZ domains [38, 54–59]. These results suggest that INAD functions as a scaffold to assemble the components of the fly visual signal transduction system into a ‘signalosome’ (Figure 3). Functional analyses suggest, as expected, that the INAD signalosome functions to enhance signaling efficiency, since mutations in the scaffolding protein result in significantly increased latency between visual stimulus and cellular response. Furthermore, this macromolecular complex appears to be also involved in termination of the visual response [62, 63]. In summary, INAD meets the expectations that multi-PDZ proteins can function as scaffolds to assemble macromolecular complexes that enhance signaling.

Fig. 3 The Drosophila visual signal transduction complex is assembled by the multiPDZ protein INAD (adapted from [48]). This figure illustrates that various cell-surface and intracellular proteins involved in signal transduction, including G proteins (Gq"), phospholipases (PLC), ion channels (TRP), are physically brought together by interactions between their C termini and the multi-

PDZ protein INAD. The complex is maintained in an appropriate subcellular location by interactions between the NINAC protein and the cytoskeleton (F-actin). In addition, PDZ–PDZ interactions appear to hold separate complexes together into a supercomplex, although this type of interaction is not mediated by C-terminal binding.


4.2 LIN-7-Receptor Tyrosine Kinase Interactions and Subcellular Localization

Vulval development in C. elegans has provided an elegant system for analysis of the genetics and biochemistry of complex organ formation [62, 63]. Important early work in this system demonstrated that a homolog of the epidermal growth factor (EGF) receptor family, called LET-23, was critical for the formation of this intricate structure [64]. The activation of this receptor by LIN-3, an EGF-like molecule expressed by basolaterally localized gonadal anchor cells, suggested that the receptor was found on the basolateral surface of the vulval precursor cells, and in fact, this was found to be true [65]. The asymmetric basolateral orientation of this receptor was critical for completion of the vulval developmental program, since it allowed the receptor access to the adjacently produced activating factor. It appeared likely that the asymmetric localization of LET-23 was accomplished by a combination of both specific intracellular trafficking and retention mechanisms. An elegant analysis by the Kim laboratory soon provided strong evidence that the basolateral localization of LET-23 was accomplished by an interaction with a PDZ domain-containing protein, LIN-7 [66, 67]. This protein was found to be associated in a complex with two other PDZ domain-containing proteins, LIN-2 and LIN-10 [66, 68]. The assembly of this complex did not involve PDZ-binding interactions, a finding that suggested that the PDZ domains within the complex were free to interact with the C termini of other proteins. Importantly, it was clearly demonstrated that the C terminus of LET-23 was a type 1 PDZ-binding motif (TCLCOOH ), and it was established that this motif bound to the PDZ domain of LIN-7 [66–68]. Further genetic studies suggested that this interaction was critical for basolateral targeting of LET-23 and vulval development, providing the first molecular and genetic evidence for the role of PDZ-containing proteins in the subcellular targeting of signaling receptors in an epithelial cell [66–68] (Figure 4). Additional work suggested that the role of this interaction was to inhibit internalization of the bound receptor, consistent with the suggestion that the LIN-7-PDZ interaction was required for retention of the receptor at the basolateral surface of the polarized epithelial cell [66–68]. Because C. elegans LET-23 is a prototype of the EGF receptor family of higher organisms, it was important to establish the mechanisms by which these receptors, which include the oncogenically associated HER2-neu/ErbB-2 receptor, are asymmetrically maintained in polarized epithelial cells. An interesting initial study in this field was that of Borg and colleagues, who showed that Erbin, a member of the LAP family of PDZ-containing proteins, was involved in delivery and/or retention of the ErbB-2 receptor at the basolateral surface of epithelial cells [41]. However, as described above, further work demonstrated that the affinity of the ErbB-2-Erbin interaction was likely to be far too low to efficiently accomplish intracellular binding [21, 22]. In addition, several laboratories reported a high-affinity set of protein ligands for the Erbin PDZ domain, which included a variety of p120-type catenins [21, 69–71]. It thus appeared likely that Erbin was not involved in subcellular targeting of ErbB-2 [72], although initial data of Borg et al. did suggest that the ErbB-2 C


Fig. 4 The EGF-like receptor LET-23 is localized in the basolateral region of the epithelial cell by the LIN-2, -7, -10 complex of PDZ proteins. This figure illustrates the role of the multi-PDZ protein complex containing LIN-2, -7, and -10 in mediating the subcellular localization of the Caenorhabditis elegans EGF-like receptor LET-23. LET-23 is kept in the basolateral part of the vulval epithelial progenitor cell, where it binds to an EGF-like ligand and induces differentiation. LIN-7 binds to both the C terminus of LET-23 and potentially, by analogy with the mammalian system, to a site in the kinase domain and determines trafficking by the golgi complex to the basolateral region.

The PDZ complex is also involved in the inhibition of receptor-ligand endocytosis. In the absence of LIN-7, the default trafficking pathway for LET-23 is to the apical region of the cell. The figure also illustrates that Erbin, a PDZ protein previously thought to be involved in the subcellular localization of a LET-23-related receptor, HER2-neu (erb2), is now thought to interact with catenin-like proteins at the adherens junction (A. J.) of the epithelial cell (see Figure 5). Finally, the figure also suggests that a similar trafficking system is involved in the appropriate subcellular localization of other EGF-like receptors, including those in the EGFR family (i.e., erb2, EGFR, etc.).

terminus was critical for basolateral localization [41]. This conundrum was recently resolved in a report that clearly demonstrated that the mammalian homolog of the LIN-7 protein was involved in the basolateral targeting and retention of the ErbB2 protein [73]. Importantly, these investigators demonstrated that this targeting was accomplished by a bipartite signal in LIN-7, with an N-terminal domain involved in basolateral sorting and the PDZ domain involved in basolateral retention (Figure 4). Thus, the subcellular targeting of the EGF receptor tyrosine kinases, as well as, potentially, other receptor kinases, appears to involve PDZ domain interactions that are conserved from worms to humans [74]. Finally, it is likely that


appropriate subcellular localization of other cell surface signaling molecules, such as for example, TGFα and the cystic fibrosis transmembrane regulator, is accomplished through interactions with PDZ-containing proteins such as GRASP and NHERF or CAP-70, respectively [75, 76]. 4.3 PDZ Domain Proteins and Epithelial Polarity Induction and Maintenance

The contrast between the aesthetically beautiful structure of the polarized epithelium and the grossly malformed appearance of the oncogenically transformed tissue could not be greater. In general, oncogenic transformation of epithelial cells is accompanied by a loss of normal polarization accompanied by increased proliferation, loss of the normal flat, single-cell epithelial morphology, and invasion of adjacent normal tissues by tumor cells [77]. Epithelial cells separate their polarized apical and basolateral regions by using complex junctional structures composed of adherens junctions and tight (septate) junctions [78]. The molecular mechanisms involved in formation of the polarized epithelium are complex and involve a multitude of intracellular and extracellular proteins, including a variety of signaling, scaffolding, and adhesion molecules. Notably, PDZ domain-containing molecules play major roles in the appropriate assembly and maintenance of the epithelium of all metazoan organisms [79–82]. It is not surprising that the powerful genetics of lower metazoan organisms, such as the fly and the nematode, have played a critical part in elucidating the roles of various PDZ domain-containing molecules in the assembly and maintenance of the epithelium [78]. Interest in PDZ proteins was stimulated early on, when it was found that mutations in two different molecules, discs large (DLG), a MAGUK-type PDZ protein, and Scribble, a LAP-type PDZ protein, resulted in the loss of epithelial integrity and a tumorigenic phenotype in Drosophila [83–87]. Furthermore, of the ∼50 known tumor suppressors in Drosophila, these two loci are the only places in which mutation leads to neoplasia in both imaginal discs and brain. These exciting results suggested that PDZ-containing molecules were likely to be involved in the assembly and/or maintenance of epithelial structures and that their loss correlated with a lack of morphological integrity as well as cellular hyper-proliferation and invasion, all hallmarks of oncogenically transformed cells. Importantly, further analysis revealed that not every cell with a PDZ protein mutation gave rise to a tumor, suggesting the need for additional oncogenic insults. Scribble and DLG both appear to be localized in the basolateral region of the epithelial cell. Another group of PDZ proteins, including the MAGUK protein Stardust (PALS-1), and the multi-PDZ protein Bazooka (Par-3) appear to be involved in formation of apical regions of the epithelial cell [88–90]. Elegant genetic and biochemical work from the Perrimon group has gone the farthest toward elucidating the roles of these various PDZ proteins in epithelial formation and maintenance [91]. These studies demonstrated that Bazooka is involved in the ‘apicalization’ of the cell, while the Scribble complex served to antagonize the Bazooka-induced formation of apical regions by repressing apicalization along the basolateral side of


the cell. These data were consistent with the subcellular localization of each of these proteins. Earlier work had demonstrated that the overexpression of a cell-surface protein called Crumbs increased apical formation, and a PDZ-mediated complex between Crumbs and the Stardust MAGUK counteracted the basolateral induction by Scribble [92–95]. Together, these data suggest the importance of PDZ domain proteins in the determination of this critical subcellular morphology, although the molecular mechanisms by which this control is mediated are still poorly understood (Figure 5). Although some of the PDZ interactions involved in this important signaling pathway have been elucidated, it will be fascinating to use newer methods, such as C-terminal phage display [20], to further dissect the molecular interactions

Fig. 5 The apicalbasolateral polarity of epithelial cells is in large part determined by a variety of PDZ proteins. Elegant genetic analyses in Drosophila and Caenorhabditis have demonstrated that a variety of PDZ-containing proteins, including Scribble (scrib), Discs Large (DLG), Lethal Giant Larvae (LGL), Erbin, and Bazooka are involved in the induction and maintenance of the apical-basal polarity of the epithelial cell. Scrib, LGL, and DLG are localized to the adherens junction, and their mutation leads to loss of apical–basal polarity and hyperproliferation that is tumor-like. Importantly, the oncogenic HPV viruses encode an E6 protein that is involved in mediating the degradation of these adherens junction components. Crumbs, an apical cell surface

protein that appears to induce apicalization, interacts with Bazooka via a PDZ-mediated binding event. Scrib induces a basolateral polarity, and the balance between scrib and the Bazooka–Crumbs complex appears to control apical-basal polarization. Erbin interacts via a PDZ-mediated interaction with p120-like catenins (i.e., *-catenin), and this interaction has been shown to regulate epithelial formation in C. elegans, possibly via an interaction with Ras-type GTPases. Finally, tight junction assembly and maintenance appears to be under the control of the ubiquitously expressed (in epithelia) ZO-1 and MAGI family of MAGUK-type PDZ proteins, which are also sensitive to E6-mediated degradation.


induced by these proteins. In addition, a variety of other proteins, including the ZO-type MAGUKs as well as the MAGI family of proteins (for example, see [96]), are localized to the tight junctions of virtually all epithelial cells and are therefore likely to play an important role in epithelialization. Finally, although genetic evidence for a role for these PDZ proteins in human tumor formation is still lacking, recent data suggest that oncogenic human papilloma viruses (HPV) target a variety of PDZ proteins, including Scribble, DLG, and the MAGI proteins, for ubiquitinmediated proteasome degradation [97–101]. This targeting occurs via interactions between the C terminus of viral E6 protein, a ubiquitin ligase complex-associated protein, and various PDZ domains, suggesting that viruses can hijack cellular PDZ domains for their own nefarious purposes. A hallmark of HPV infection is loss of epithelial structure, hyperproliferation, and invasion again consistent with a role of these PDZ proteins in epithelial control. Although Scribble, a LAP family member, has clearly been shown to be critical for epithelial formation, other members of this family, including Erbin, are also likely to be important for epithelialization. The strongest genetic data for the function of Erbin comes from C. elegans, where the LET-413 protein, a likely homolog of mammalian Erbin, was clearly shown to be involved in epithelial formation [102, 103]. As described above, the probable binding partners for the Erbin PDZ domain are a group of p120-like catenins, including p0071, δ-catenin, and ARVCF, all of which contain a conserved C terminus (DSWVCOOH ), which was found to be a high-affinity Erbin PDZ-binding peptide [21, 22, 69–71]. Other important genetic data from C. elegans have recently been reported regarding JAC-1, the nematode homolog of the mammalian p120-related catenins. Mutation of this protein gives an epithelial phenotype that is reminiscent of the LET-413 mutations [104]. This result becomes even more interesting when viewed in the context of the work of Laura and colleagues, who predicted that the PDZ domain of LET-413 should bind with high affinity to the C terminus of JAC-1 (DSWVCOOH ) [21]. Together, these data suggest that a complex between LET-413 and JAC-1 is likely to be involved in epithelial regulation. Although the mechanism of this regulation remains to be fully elucidated, recent work by Huang and colleagues has demonstrated that the leucine-rich repeats of Erbin appear to be involved in the regulation of Ras GTPase activity [105, 106]. Epithelial integrity and intracellular communication are, in part, mediated by the cadherin family of adhesion molecules. Because Erbin is linked to this adhesion system via its PDZ-mediated catenin binding, the data of Huang et al. suggest a mechanism whereby epithelial adhesion might regulate Ras subcellular localization and activity [107]. 4.4 A Few Miscellaneous Examples: The Synapse, Disheveled, CARD MAGUKs, and Beta Adrenergic Receptors

In addition to the polarized epithelium, a second highly polarized cell type is the axon. In this cell type, the synapse corresponds to the highly polarized apical region of the epithelial cell, and it has been demonstrated that the synaptic and


post-synaptic regions of neurons contain a variety of signaling molecules, including neural transmitters and the many channels that they bind to. Importantly, the subcellular localization, signaling efficiency, and signalosome assembly of the various synaptic and post-synaptic signaling proteins appear to be accomplished by a variety of PDZ proteins [11, 108, 109]. Many potential PDZ-mediated interactions in neurons have been identified, but we discuss only two examples for which there is some genetic analysis of their function. By biochemical studies, a variety of excitatory receptors, including NMDA and glutamate receptors, have been found to be associated with PSD-95, one of the prototypical MAGUK-type PDZ proteins, and it has been assumed that their clustering and/or localization to the post-synaptic density requires an interaction with this MAGUK. Murine gene knockout studies of PSD-95, however, revealed that these receptors were correctly localized, but their signaling pathways with respect to synaptic plasticity seemed affected, although subtly [110]. Other ion channels, such as the potassium channel, also appear to interact via a C-terminal–PDZ domain interaction [11, 108, 109]. This interaction appeared, in cell culture assays, to be critical for the clustering and activity of these channels. However, recent murine knockout experiments have shown that the localization of these channels to the juxtaparanodal regions of myelinated axons does not require PSD-95 [111]. Thus, although PSD-95 is colocalized to these sites with the potassium channels, its presence is clearly not required for the localization of these channels. Together, these two examples highlight the importance of in vivo analysis in assessing the true function of PDZ-containing proteins. The signaling pathway regulated by the Wnt family of proteins has emerged as one of the most critical for numerous aspects of development, including the maintenance and expansion of stem cells. A critical component of this pathway is disheveled (Dsh/Dvl), a scaffolding protein containing a PDZ domain as well as a variety of other protein interaction motifs [112]. This protein is a critical component of the numerous pathways controlled by Wnt signaling, and mutations of the Dsh/Dvl gene result in various phenotypic changes in these pathways that mimic the loss of either Wnt protein or Wnt receptors [113–115]. Genetic analyses demonstrated that Dsh/Dvl was downstream from the Wnt/Wnt receptor but upstream from the β-catenin destruction complex, suggesting that Dsh/Dvl plays an important role in regulating this complex and its transcriptional activator, β-catenin. The role that the single Dsh/Dvl PDZ domain plays in regulating the Wnt pathway is controversial and appears to depend upon assay conditions, although much of this heterogeneity may be due to differences in the types of PDZ mutants that have been examined. Oddly, a bewildering array of proteins have been found to bind to the Dsh/Dvl PDZ domain [112]. Interestingly, these proteins appear to all have completely different C termini, suggesting that either this PDZ domain is unusually promiscuous or that most (or all) of these binding partners are artifactual. Recent data from our laboratory using the C-terminal phage-display method suggest that peptides that bind with high affinity to the Dsh/Dvl PDZ domain are quite different from the C-terminal sequences of these putative protein ligands (Zhang and Sidhu, unpublished observations). Thus, as with many other PDZ domain interactions reported in the literature, careful analysis of binding affinities is required


to determine the validity of potential PDZ binding partners. Finally, a recently published study suggests that Dsh/Dvl is overexpressed in mesothelioma tumor cells, and this overexpression enhances β-catenin levels and cellular transformation [116]. As described above, the MAGUK-type PDZ domain-containing proteins are involved in the assembly of complexes that appear to be involved in polarization in both epithelial and neuronal cells. The CARD MAGUKs are a recently described family of proteins that contain MAGUK-type domains (including PDZ, SH3, and guanylate kinase domains) as well as a CARD (caspase recruitment domain) domain and a coiled-coil motif [117–121]. These proteins are encoded by multiple genes in mammals, and early data demonstrated that one CARMA-1 is expressed in cells of the immune system. Interesting early work demonstrated the function of CARMA-1 in the stabilization and activation of NFκB transcription factors, suggesting that this MAGUK may be involved in immune system function. This hypothesis was spectacularly supported by data from two groups who inactivated the CARMA-1 gene by using traditional knockout as well as a novel ENU-mediated point-mutation technique [122, 123]. Although some of the resultant phenotypes were different (likely due to the fact that one group introduced a point mutation in the coiled-coil domain and the other completely abolished expression), both types of mutant mice showed profound defects in immune system function, including loss of antigen receptor signaling in both B and T cells. Further analysis of the defects demonstrated that these mutant animals had defects in NFκB activation that appeared to be due to stabilization of the IκB inhibitor. Together, these data suggest that CARD MAGUKs are involved in antigen receptor signaling by controlling the stability of the inhibitor of NFκB. One possibility is that antigen receptor activation induces CARMA-1 to bring a protein degradation complex close to the IκB inhibitor. If the PDZ domain is critical to this event, inhibition of its binding activity by small-molecule antagonists may result in a decreased immune response, a potentially important result for diseases ranging from autoimmune syndromes to transplantation reactions. Heart failure is a major unmet medical problem with little or no effective treatment. The three beta-adrenergic receptors are G-coupled proteins that are regulated by the sympathetic nervous system, and it is likely that their modulation plays a role in the pathogenesis of heart failure [124]. Both beta 1 and 2 adrenergic receptors appear to be localized to specific subcellular localizations and are associated with downstream intracellular signaling molecules including, for example, G proteins, beta arrestin, and various kinases. This is highly reminiscent of other proteins that utilize PDZ domain-containing molecules to accomplish subcellular localization and signalosome assembly, and it has been shown that the C-terminal PDZ binding motifs of these two G-coupled proteins associate with PSD-95/MAGI-2 (beta 1) and the sodium–hydrogen exchange regulatory factor (NHERF) (beta 2) [125, 126]. Two major functional aspects of these interactions appear to be the control of subcellular localization and endocytosis as well as coupling to various G proteins [127, 128]. Interestingly, mutation of the PDZ binding site in the beta 2 receptor or blocking the interaction with a C-terminally derived peptide both result


in a change in intracellular coupling that is accompanied by increased contractility in response to ligand binding [129]. Increased contractility might be a useful response for heart failure patients, although other data suggest that modulation of intracellular coupling in this manner might also be detrimental to myocyte survival. In summary, these results provide for yet another interesting mechanism by which PDZ-containing scaffold or localization proteins play a critical role in cellular physiology and which might provide for a clinically useful application.

5 Conclusions

PDZ domains and the proteins they are found embedded in have emerged as major components of mechanisms for subcellular localization, signaling, and polarity induction. Although this chapter has not exhaustively discussed the huge number of reported PDZ interactions, we have attempted to discuss representative examples that have strong biological and/or genetic support. In addition, a number of examples exist, with more being presented all the time, which suggest that PDZ domains may be involved in a variety of pathogenic situations. For example, disruption of the interaction between the C terminus of an NMDA receptor and a PDZ domain of PSD-95 by a small peptide results in an inhibition of glutamate-mediated damage in an ischemic stroke model [45], suggesting that the detrimental effects of excitotoxicity may depend on PDZ-mediated receptor localization and/or signaling. The structural data discussed here suggest that the interface between the PDZ domain and its ligand(s) is likely to be among the smallest of protein–protein binding sites. Thus, although the dream of disrupting protein–protein interactions appears quite daunting for most targets, the possibility of identifying and enhancing the binding activity of small-molecule antagonists of PDZ-mediated interactions seems quite likely. It will be interesting in the future to identify such small-molecule antagonists in both model and disease-related systems and to examine their effects on PDZ-mediated biology. If these studies are successful, these inhibitors might be among the first drugs to target pathogenic protein interfaces. Acknowledgment

We thank David Wood for help with the figures.

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1

PTB Domains Ben Margolis University of Michigan Medical School, Ann Arbor, USA

Linton M. Traub University of Pittsburgh School of Medicine, Pittsburgh, USA


1 Introduction

The phosphotyrosine binding (PTB) domain also known as the phosphotyrosine interaction domain (PID) was the second domain (after the SH2 domain) found to bind phosphotyrosine-containing peptides. Over time, scientists have come to realize that this domain does more than mediate phosphotyrosine-related signaling, because its binding is not always dependent on the presence of phosphotyrosine. The domain was first identified in the amino terminus of the Shc protein [1–3], where it was found to bind to an Asp-Pro-any amino acid-pTyr (NPxpoY) motif found in many activated growth factor receptors. Simultaneously, a binding domain on insulin receptor substrate 1 (IRS1) and insulin receptor substrate 2 (IRS2) was identified that also bound the NPxpoY motif [2, 4]. Although the NPxpoY binding regions in the IRS and Shc proteins do not have significant primary sequence homology, they have similar binding specificity and 3D structure. PTB domains with sequence homology to those found in IRS proteins are referred to as PTBI for PTB-domain-IRS-like to differentiate them from PTB domains that have sequence homology with the Shc PTB domain (Simple Modular Architecture Research Tool; http://smart.embl-heidelberg.de). A relatively small number of proteins have been identified that contain PTBI domains, and these primarily function in tyrosine kinase signal transduction. Many more PTB domains with sequence similarity to the Shc PTB domain have been identified [5], and they have more diverse cellular functions. Evolutionarily, PTB domains are not found in yeast or plants but appear in Drosophila and Caenorhabditis elegans genomes. This chapter discusses the


2 PTB Domains

function of proteins containing PTB and PTBI domains and examines the structural basis of their interactions with peptide and phospholipid ligands.

2 Function of PTB Domain Proteins

Proteins with PTB and PTBI domains are involved in multiple cellular functions. The following sections discuss their role in tyrosine kinase-dependent and -independent signaling, protein trafficking, and cell adhesion. A schematic representation of several PTB and PTBI domain proteins is displayed in Figure 1. 2.1 Role of PTB Domain Proteins in Tyrosine Kinase Signaling 2.1.1 Shc As described above, PTB domains are known to play an important role in tyrosine kinase signal transduction. This of course relates to the initial identification of the PTB domain in the tyrosine kinase substrate, ShcA, and has been reviewed previously [6]. The classical role of the Shc PTB domain is to allow Shc to bind to growth factor receptors, facilitating Shc phosphorylation [7, 8]. Once Shc is phosphorylated, it can bind to other downstream signal-transduction molecules. For example, the nerve growth factor receptor, TrkA, cannot bind efficiently to Grb2 and its binding partner, the Ras guanyl nucleotide exchange factor, Son of Sevenless (Sos). Grb2-Sos is the primary protein complex used by growth factor receptors to activate the Ras G protein. However, TrkA does have an NPxY motif that, once phosphorylated, can bind the PTB domain of Shc [8]. Once TrkA binds Shc, it mediates Shc tyrosine phosphorylation, allowing it to bind to the Grb2-Sos complex and activate Ras. Many growth factor receptors are known to utilize Shc and its paralogs as a crucial intermediary in signal transduction [9]. One of the better examples for the role of the PTB domain is found in Drosophila Shc [10]. In a genetic screen, Nüsslein-Volhard and coworkers identified Shc as a maternal gene important for embryonic development. They found that Shc signaled downstream of the Drosophila EGF and TOR tyrosine kinase receptors. Interestingly, one Shc mutant isolated had a mutation in the PTB domain and functioned similar to a null allele, confirming the crucial role of the PTB domain in Shc function. A second aspect to the function of the Shc PTB domain and other PTB domains is the ability to bind to phospholipids [11]. The PTB domain is highly related in structure to the pleckstrin homology (PH) domain, a domain known to bind to phospholipids [12]; indeed, the protein binding capabilities of the PTB domain may have evolved from its initial ability to bind phospholipids as a PH domain [13]. The lipid binding region of the PTB domain is distinct from the peptide binding region, indicating that the PTB domain can bind both lipids and proteins simultaneously. This can assist the targeting of Shc to the membrane while binding to tyrosinephosphorylated receptors. It has been shown that specific mutations within the


Fig. 1 Domain architecture of several representative PTB domain-containing proteins and other modular domains within the PTB and PTBI protein families. Many of the other modular domains are discussed in other chapters in this volume. More details on those not covered in this book can be found at one of the numerous domain databases such as SMART (http://smart.embl-heidelberg.de). PTB = phosphotyrosine binding domain;

PTBI = phosphotyrosine binding domain IRS-like; SH2 = Src homology 2 domain; WW = domain with two conserved tryptophans; PDZ = postsynaptic density 95/discs large/zona occludens-1 domain; SH3 = Src homology 3 domain; RGS = regulator of G protein signaling domain, RBD = Raf-like Ras-binding domain; Go Loco = G"I/O -Loco motif; SAM : sterile alpha motif; JBD = JNK binding domain; PH = pleckstrin homology domain.

PTB domain that disrupt phospholipid but not phosphotyrosine binding impair Shc tyrosine phosphorylation by growth factor receptors [11]. A unique aspect to Shc is that it also contains an SH2 domain in addition to a PTB domain (Figure 1). This organization is conserved throughout evolution, suggesting some important role for the SH2 domain. Yet there are very few examples of the SH2 domain

3

4 PTB Domains

rather than the PTB domain coupling Shc to tyrosine phosphorylated proteins. This suggests that the Shc SH2 domain may have an alternative role in Shc function, but the exact nature of this is unclear [9]. 2.1.2 Proteins with PTBI Domains Proteins with PTBI domains play a prominent role in signaling by receptor tyrosine kinases. As discussed previously, PTBI domains have a 3D structure similar to that of PTB domains, but there is limited primary sequence similarity between these domains. The classic family of proteins having this domain is the insulin receptor substrates (IRS). It had been known for several years that the insulin receptor had an NPxY motif centered on Tyr960 that was phosphorylated. It was also known that this motif was necessary for insulin-mediated signal transduction as well as phosphorylation of a 185-kDa substrate protein [14]. This substrate protein was later identified as IRS-1 [15] and shown to have a domain that can interact with the insulin receptor [16]. A similar domain was also identified in the related IRS-2 protein [4], and this domain is now referred to as the PTBI domain. The IRS family of proteins functions as adapters similar to Shc. However, although both Shc PTB and the IRS PTBI domains bind NPxpoY motifs, there are differences in the sequences surrounding the NPxpoY motif that are required for high-affinity binding [17]. IRS proteins contain multiple tyrosine-phosphorylation sites and can mediate multiple downstream signaling pathways. A complete discussion of IRS signaling is beyond the scope of this review, but it has been described in other publications [18]. However, a few points specifically related to the PTBI domains of these proteins can be noted. One of the surprises is that the role of PTBI in signaling by IRS proteins has been difficult to demonstrate. The amino terminus of IRS proteins has one PTBI domain that sits just carboxy-terminal to the PH domain (Figure 1). As mentioned previously, a primary function of PH domains is to bind phospholipid headgroups. In IRS-1, the PH domain is necessary for IRS tyrosine phosphorylation after insulin stimulation [19]. The PTB domain seems to be essential for high-affinity interactions between IRS-1 and insulin receptor, but the actual necessity for this interaction in insulin signaling has been difficult to demonstrate [19]. The picture becomes more confusing when one considers that the PTB domain of IRS proteins can bind phospholipids [20] and that the PH domain of IRS-1 also has protein binding partners [21]. The most reasonable explanation for the function of these domains comes from structural studies, which suggest that cooperation between PTB and PH domains mediates a high-affinity interaction between insulin receptor and IRS-1 [22]. Many of the studies using mutagenesis of these domains relied on overexpression and thus may miss the cooperative nature of domain interactions when the proteins are expressed at physiologic levels. Two other families of proteins with PTBI domains are also important in growth factor receptor signal transduction. One family is referred to as the fibroblast growth factor receptor substrate (FRS) family [23]. These proteins are also called Sucassociated neurotrophic factor-induced tyrosine-phosphorylated target (SNT). This protein family was first identified by its binding to p13suc1 agarose, an affinity reagent that binds to certain cyclin-dependent kinases [24]. This suggested that


this tyrosine-phosphorylated substrate of fibroblast growth factor (FGF) and Trk receptors might directly control the cell cycle, but later purification indicated it was a large scaffold protein that bound to growth factor receptors at the cell surface [23]. Proteins in this family have a single PTBI domain at their amino terminus that mediates interactions with FGF and Trk receptors, as well as other growth factor receptors [23, 25, 26]. Like Shc and IRS proteins, the FRS proteins become tyrosine-phosphorylated after association with growth factor receptors and then recruit signaling molecules onto the tyrosine phosphorylation sites [27, 28]. The binding specificity of the FRS PTBI domain is unique. On TrkA it binds to phosphorylated Y490, a classic NPxpoY site, and can compete with Shc for binding to this site [29]. In contrast, the binding to FGF receptor appears to be constitutive and not dependent on receptor phosphorylation [25, 27]. The peptide from the juxtamembrane domain of the FGF receptor binds to the FRS PTBI domain in a fashion completely different from that seen with the NPxpoY motif from the TrkA receptor [30] and represents a novel peptide-PTB domain interaction. One trend seen in previously discussed PTB domain proteins is also seen with the FRS family of proteins: both Shc and IRS proteins have the ability to bind membranes and protein peptide motifs either with their PTB domain alone or with the PTB domain in combination with another domain. The FRS proteins address this problem, not with an additional domain, but by adding a myristate group at the amino terminus that links them to membranes [23]. The other family of PTBI domain proteins is the DOK (downstream of kinase) proteins (Figure 1). The first DOK family member was identified as a 62-kDa tyrosine-phosphorylated protein that binds to the p120 RasGAP protein and contains an amino-terminal PH and PTB domain [31, 32]. Since then, an additional four members of the DOK family have been identified [33–40]. The PTB domain of DOK proteins can bind to the canonical NPxpoY motifs and to highly related motifs present in EGF receptor [41] and other tyrosine-phosphorylated proteins [38, 42]. However, there also appears to be unique phosphotyrosine-containing motifs that can bind these PTB domains, such as those found in Tie1 receptor [43]. As in IRS-1, there appears to be a need for cooperation between the PH and PTB domains to mediate the growth factor-induced tyrosine phosphorylation of DOK proteins [41]. However, in contrast to IRS and FRS family members, the primary role of DOK proteins focuses on the inhibition of growth-promoting signals. This is particularly evident for DOK signaling in cells of the hematopoietic lineage, where DOK family members inhibit signaling from tyrosine kinases to the MAP kinase-Erk pathway [36, 44]. However, there have also been reports of positive roles for DOK proteins in signal transduction pathways [38] and cell migration [45]. 2.1.3 Additional PTB Domain Proteins Involved in Tyrosine Kinase Signaling Other proteins with PTB domains appear to play a role in tyrosine kinase signaling, although the role of the PTB domain in these processes is not as clear. The EPS8 protein was first identified as a protein that became tyrosine-phosphorylated after EGF stimulation [46]. EPS8 appears to bind to the EGF receptor, but surprisingly, the PTB domain does not appear to be involved in this process [47]. EPS8 protein

5

6 PTB Domains

also contains an SH3 domain (Figure 1). An important insight came when it was determined that EPS8 can localize to sites of actin polymerization [48]. When EPS8 is knocked out in mice, the mice appear to develop normally, but there are defects in actin polymerization in response to growth factor receptors in knockout fibroblasts [49]. Based on several studies, it was concluded that EPS8 is in a complex with several proteins, including Sos, and mediates Rac activation after growth factor receptor stimulation in a PI-3 kinase-dependent fashion [50, 51]. EPS8 may also play a role in the endocytosis of growth factor receptors [52]. Another PTB domain known to be phosphorylated after EGF stimulation is Odin [53]. Odin also has SAM domains as well as multiple ankyrin repeats (Figure 1). It appears in the protein database in multiple isoforms with and without ankyrin repeats. The function of the PTB domain in this protein is unclear, because its removal does not affect Odin tyrosine phosphorylation, and it is not certain that this protein even associates with growth factor receptors [53]. A highly related gene called EB-1 was identified as a gene that is induced in hematopoietic cells transformed by the E2A-PBX oncoprotein, but the function of EB-1, as well as of Odin, is unknown [54]. Regulator of G protein signaling 12 (RGS12) is another signaling molecule that contains a PTB domain and is involved in tyrosine kinase signaling (Figure 1). RGS proteins function as GTPase activating proteins for the α subunits of heterotrimeric G proteins [55]. In RGS12, it has been reported that the PTB domain of this protein can bind to tyrosine-phosphorylated calcium channels [56]. The binding of the PTB domain to the calcium channel may modulate calcium channel activity as well as regulate the heterotrimeric G proteins that control the channel. 2.2 PTB Domain Proteins That Function Independent of Phosphotyrosine

The above signaling pathways involve the binding of PTB and PTBI domains to tyrosine-phosphorylated proteins. Indeed, these were the early signaling pathways that defined the function of PTB and PTBI domains. However, soon after the identification of this domain, it was found that PTB domains can also bind NPxY motifs independent of tyrosine phosphorylation [57]. This led to an expanded appreciation of the role of PTB domains in signaling, cell adhesion, and protein processing and trafficking. 2.2.1 PTB Domain Proteins That Bind APP The realization that PTB domains can bind nonphosphorylated NPxY motifs came with studies of amyloid precursor protein (APP). APP is of great interest because it is processed via proteases to amyloid beta, a protein deposited in the brains of Alzheimer disease patients. Evidence obtained over the past several years has strongly pointed to the deposition of amyloid beta as an important causative factor in Alzheimer’s disease [58]. Despite the known role of APP processing in Alzheimer’s disease, the physiological role of APP and highly related proteins is unclear.


However, the binding of PTB domains to APP has provided unique insights into the normal function of this protein. Initial studies pointed to the binding of two PTB domain proteins, FE65 and X11/Mint, to the NPxY motif of APP [57, 59, 60]. Later, several other PTB domain proteins, including Disabled (Dab), JIP-1, and Numb, were identified as proteins that can bind APP [61–66]. There are also reports of APP becoming tyrosinephosphorylated and binding Shc via an NPxpoY motif, but the physiologic relevance of this interaction is uncertain [67, 68]. A large number of studies have examined the effect of PTB domain proteins on APP processing to amyloid beta [64, 69– 72]. A weakness of all these studies is that they employ overexpression of the PTB domain proteins, and thus it is not clear that these proteins physiologically regulate APP processing normally in the brain. Nonetheless, a large amount of data has been obtained on the effects of X11/Mint proteins on the processing of APP. X11/Mint proteins have one PTB domain and two PDZ domains (Figure 1; see also Chapter 3). X11/Mint protein overexpression slows APP processing and reduces amyloid beta production, both in tissue culture and in vivo [69, 70, 73]. The mechanism of X11/Mint action in this regard is unclear; however, studies on the C. elegans homolog of X11/Mint proteins, known as Lin-10, are instructive [74, 75]. Lin-10 plays a role in proper trafficking of proteins in worm epithelia and brain, suggesting that X11/Mint might have a similar role in mammalian cells. The localization of X11/Mint in the Golgi fits with such a role for this protein [76, 77]. In addition, X11/Mint proteins also interact with Munc-18, a protein that modulates SNARE interactions and thus possibly vesicle fusion [78]. Munc-18 interaction also modulates the effects of X11/Mint on APP processing, adding to the hypothesis that altered trafficking of APP is at the core of X11 actions on APP processing [79, 80]. The processing of APP is known to occur in discrete intracellular compartments, and rerouting of APP in the cell by X11/Mint proteins could alter amyloid beta production [81]. A more exciting story has emerged on the interaction of the FE65 family of proteins with APP. FE65 family members have two PTB domains and an aminoterminal WW domain (Figure 1). The effects of FE65 on APP processing have also been studied, and it appears that FE65 usually increases amyloid beta production, but the effects may be variable depending on the situation [71, 82]. FE65 also interacts with another transmembrane protein called lipoprotein receptor-related protein (LRP), a member of the lipoprotein receptor family [83]. The amino-terminal PTB domain of FE65 interacts with LRP while the carboxy-terminal PTB binds to APP, allowing FE65 to act as a bridge linking LRP to APP [83, 84]. This is of significance, because a large body of literature suggests that LRP can control APP processing, and FE65 is an important link between these two proteins [85, 86]. Further studies revealed that FE65 is a nuclear protein and that its localization to the nucleus is restricted by binding to APP [87]. A breakthrough study by Cao and Sudhof [88] then demonstrated that processing of APP leads to translocation of FE65 to the nucleus bound to the cleaved intracellular domain of APP. It was demonstrated that this complex of FE65 bound to the intracellular domain of APP can affect gene transcription by binding to the Tip60 protein, a histone

7

8 PTB Domains

acetyltransferase [88]. This pathway can be modified by the interaction of APP with X11/Mint [89] and by the interaction of FE65 with LRP [90]. The concept of FE65 and APP as a unit that can control transcriptional activity has been confirmed by several other groups [91, 92]. These findings suggest a signal-transduction pathway in which the processing of APP and translocation of its intracellular domain to the nucleus leads to control of gene expression. Further work will be necessary to define what mechanisms are utilized to control the processing of APP and the nature of the genes induced. Other roles have also been invoked for FE65, including the control of actin bundling and cell migration [93], but these findings still await confirmation. APP has also been reported to interact with the PTB-domain proteins JNK interacting protein-1 (JIP-1) and JNK interacting protein-2 (JIP-2). JIPs were first identified as scaffolding proteins that can regulate the MAP kinase pathway that leads to JNK activation [94, 95]. There are several theories as to how JIP might control JNK activity, and it may have both inhibitory and activating effects on JNK activation [94–96]. JIP-1 and JIP-2 contain a PTB domain that can interact with APP [63, 64] and a carboxy terminus that can interact with the light chain of conventional kinesin, kinesin-1 [97]. In this fashion, APP, a transmembrane protein, can connect transport vesicles to a kinesin motor and assist vesicle movement along microtubules. Thus, another physiological role for APP, in addition to control of transcription, may involve vesicle trafficking along microtubules in neurons. Indeed, several reports have linked APP to kinesins, both directly and indirectly [98, 99]. The exact role of JIPs, and more specifically, the associated MAP pathway kinases in control of kinesin transport is an area of active investigation. It is interesting to note that another APP binding partner, X11/MINT, has also been implicated as binding to kinesins [100], indicating that APP may have a general role in coupling transport vesicles to kinesin motors. Thus, in conclusion, several PTB-containing proteins interact with the NPxY motifs of APP and give important clues to the physiological role of APP and its paralogs in transcription control and vesicle trafficking. The studies may also provide important additional clues to the pathogenesis of Alzheimer’s disease. 2.2.2 PTB Domain Proteins That Bind Integrins Another family of transmembrane proteins known to contain NPxY motifs are beta integrins, which serve to link the intracellular actin cytoskeleton with the extracellular matrix [101]. NPxY motifs are found in multiple copies in several of the beta integrins’ cytoplasmic tails [102, 103]. Recent studies have shown that several PTB-domain proteins can interact with integrins [103]. However, the best data are found for the interaction of integrin NPxY motifs with two proteins, integrins’ cytoplasmic domain associated protein-1 (ICAP1) [104, 105] and talin [106]. ICAP1 was first identified by yeast two-hybrid screening using the intracellular domain of beta integrins as bait [104, 105]. ICAP1α is a small phosphoprotein of 200 amino acids (Figure 1), with a C-terminal PTB domain and an N terminus that is the site of threonine phosphorylation [104, 107]. Several roles have been ascribed to ICAP1. One set of studies found that ICAP1 can function as a guanyl dissociation


inhibitor, sequestering the small G proteins, Cdc42 and Rac, in the cytoplasm and leading to altered actin dynamics [108]. In this regard it has been demonstrated that modulation of small G protein function can regulate ICAP phosphorylation [104]. ICAP may also alter cell adhesion and cell spreading by modulating the interactions of integrins with other members of the actin cytoskeleton [109]. One of the members of the actin cytoskeleton whose binding to integrins may be affected by ICAP1 is talin. Talin is a large protein (> 2500 amino acids) that interacts with integrins via its FERM domain [110]. FERM domains were originally identified in members of the Band 4.1 family of actin-binding proteins [111]. The FERM domain of talin interacts with the NPxY motifs in beta integrins [106]. It is of great interest that a crystal structure of this interaction reveals that this FERM domain interaction is very similar to PTB domain interactions and that a region of the FERM domain has a structure very similar to that of PTB and PH domains [112, 113]. Thus, some FERM-domain proteins may have interactions that mimic PTB domains and may greatly broaden the proteins that can interact with NPxY motifs. 2.2.3 PTB Domain Proteins That Control Endocytosis Nearly a decade before the discovery of the PTB domain, the Goldstein and Brown laboratory [114] showed that a single point mutation (the JD mutation) in the gene encoding the low density lipoprotein (LDL) receptor can prevent receptor internalization. The LDL receptor governs the internalization of plasma LDL particles, primarily in the liver, and targets internalized LDL for lysosomal degradation. The JD mutation substitutes a tyrosine for a cysteine residue within the cytosolic domain of the receptor [114], causing stagnation of the receptor on the cell surface and leading to hypercholesterolemia and coronary artery disease. More complete mapping revealed that an FxNPxY internalization sequence is required to drive the efficient endocytic uptake of the LDL receptor and that this sequence is both autonomous and transplantable [102]. Most other members of the LDL receptor superfamily contain at least one FxNPxY motif within the cytosolic portion but, because the PTB domain was originally presumed to be pTyr-specific, a link between PTB domain proteins and endocytosis from the cell surface was not immediately appreciated. It is now known that several PTB domain proteins, including Disabled-1 (Dab1), Disabled-2 (Dab2), FE65, and JIP-1/2, can bind to these nonphosphorylated FxNPxY sequences directly [61, 83, 115–117]. The PTB domain–endocytosis connection became apparex nt when Dab2 was found to colocalize with clathrin at internalization sites on the cell surface [117]. The C-terminal region of Dab2, following the PTB domain, is essentially unstructured and contains multiple interaction motifs that bind physically to clathrin, the clathrin-associated AP-2 adaptor complex, and several other endocytic components [117–119]. Clathrin and AP-2 are the two principal components of a sorting coat that assembles on a class of endocytic transport vesicles termed clathrin-coated vesicles [120]. Like the signaling PTB-domain scaffolding proteins, Dab2 appears to mesh LDL receptor cargo with assembling coated vesicles by simultaneously binding to an NPxY internalization signal, PtdIns(4,5)P2 , and the clathrin coat machinery. In so doing, Dab2 expands the cargo selective capacity of the major

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sorting adaptor AP-2. Indeed, overexpression of a tandem Dab2 PTB domain prevents uptake of LDL but not of transferrin, because the transferrin receptor uses a distinct internalization sequence that binds directly to the AP-2 adaptor [118]. In mice, targeted disruption of the Dab2 gene is lethal, but conditional disruption only in the embryo leads to viable animals with a proteinuria similar to, albeit milder than, that seen in megalin nullizygous mice [121, 122]. A member of the LDL receptor superfamily, megalin is a scavenger receptor that plays a vital role in the recovery of protein and vitamins from the renal glomerular filtrate [122]. Although there is evidence for Dab2 participating in signal transduction [121, 123–125], the trafficking activity appears phylogenetically conserved. Ce-Dab-1, the single Disabled ortholog in C. elegans, acts together with two LDL receptor-related proteins (Ce-LRP-1 and Ce-LRP-2) in the proper biosynthetic delivery of Egl17, a fibroblast growth factor [126]. The recent identification of a gene responsible for a rare form of hypercholesterolemia characterized by a recessive pattern of inheritance has fortified the connection between PTB domains and endocytosis. Autosomal recessive hypercholesterolemia patients have a clinical phenotype remarkably similar to that of familial hypercholesterolemia patients, but have no mutations in the LDL receptor. Instead, the defective gene encodes a 308-amino-acid adaptor protein, termed autosomal recessive hypercholesterolemia (ARH), with an N-terminal PTB domain related to the Dab1/2 and Numb PTB domains (Figure 1) [127, 128]. ARH patient lymphoblasts exhibit defective LDL uptake despite normal levels of LDL receptor, but retroviralmediated expression of exogenous ARH rescues LDL receptor activity in these cells [128]. The hypercholesterolemic phenotype of ARH−/− mice fed a high-fat diet and the prominent accumulation of the LDL receptor on the sinusoidal surface of hepatocytes in these animals also support a role for ARH in regulating the endocytic activity of the LDL receptor [129]. The ARH PTB domain interacts with FxNPxY sequences while the C-terminal region physically binds to AP-2 and clathrin [130–132]. In Xenopus oocytes, production of mutant ARH that cannot engage the endocytic machinery severely blunts the uptake of vitellogenin [132], a lipoprotein endocytosed by the vitellogenin receptor, a member of the LDL receptor superfamily with an FxNPxY internalization signal. Because ARH patients do not exhibit defects in LDL uptake in all tissues [133, 134] and the Dab2−/− phenotype is rather mild [121], it is possible that Dab2 and ARH are functionally redundant to some extent. The function of Numb is perhaps best understood in terms of the development of external sensory organs in Drosophila. Sensory bristles on the fly head are composed of four different cell types that all originate from a single sensory organ precursor (SOP) cell by asymmetric cell division. This results in the formation of two nonequivalent daughter cells with different cell fates. Numb is a modular PTB domain protein (Figure 1) that antagonizes signaling by the transmembrane receptor Notch [135]. The bristle phenotype of notch mutants is similar to that seen on Numb overexpression [136], and there is some evidence for Numb binding directly to Notch [137, 138]. During SOP cell mitosis, Numb partitions asymmetrically, repressing Notch activity in the cell preferentially enriched in

3 PTB Domain Structure

Numb, which is instrumental in establishing the different identities of the daughter cells. The PTB domain of Numb is necessary and sufficient for membrane association and asymmetric localization [139]. As in Dab2, the C-terminal region of Numb is likely disordered, and there are binary associations between Numb and the AP-2 adaptor [136, 140]. In fact, certain AP-2 (α-adaptin) mutant alleles phenocopy numb mutations, and Numb is necessary to drive asymmetric partitioning of AP-2 in the mitotic SOP cell [136]. Epistasis analysis reveals that AP-2 acts between Numb and Notch, leading to a model in which Numb-dependent endocytosis and degradation of Notch underlies the different levels of Notch signaling that cause distinct cell fates. The asymmetric distribution of Numb in mitotic cells may be controlled by NIP (Numb-interacting protein), a transmembrane protein that binds the PTB domain via two redundant NPxF-type sequences [141]. That the PTB domain is important in asymmetric cell division is also demonstrated by the gene dosage-dependent effect of overexpression of Numb-associated kinase (Nak), which engages the PTB domain via an FSNMSF sequence [142]. A significant complication of the Notch down-regulation model is that it has not been convincingly shown that the levels of Notch differ significantly between the two daughter cells. In a modified model, Numb may regulate Notch activity by binding to the multispanning transmembrane protein Sanpodo and promoting its internalization [143]. Endocytosis of Sanpodo prevents the protein from localizing to the plasma membrane where it can bind and assist Notch in signal transduction [143]. It is not yet known whether endocytic uptake of Sanpodo requires the Numb PTB domain, but the YTNPAF sequence at the N terminus of Sanpodo is highly suggestive. Irrespective, it seems clear that Numb, ARH, and Dab2 are globally similar adaptors that use the PTB domain to select designated cargo while synchronously interacting with the endocytic machinery.


The high-resolution atomic structures of the PTB domain from eight proteins (Shc [13], IRS-1 [144, 145], Numb [146], X11 [147], SNT-1 [30], Disabled-1 (Dab1) [148, 149], Disabled-2 (Dab2) [149], and Dok1 [150]) currently available reveal that, despite low primary sequence identity and conservation, all adopt a common basic fold. The canonical PTB domain core is composed of seven central β strands, folded into two antiparallel β sheets oriented nearly orthogonally to one another and capped by an α helix at one end of the β sandwich [151]. Typically, one sheet is composed of strands β1–β4 and the other of β5–β7, although β1 can be long and arched, allowing this strand to contribute to the β5–β7 sheet as well (Figures 2 and 3). As mentioned previously, the overall topological arrangement is homologous to that of a phosphoinositide interaction module, the PH domain [12], making the PTB domain a member of the PH superfold. Indeed, the core Cα atoms of the IRS-1 PTB domain and the dynamin, phospholipase C-δ1, and spectrin PH domains superimpose with an rms deviation of ∼1.0 Å [144].

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Fig. 2 Schematic representation of the structure of the Shc PTB domain bound to a TrkA NPxpoY phosphopeptide. Ribbon diagram with " helices colored green, $ strands cyan, and connecting loops gray. The TrkA phosphopeptide (HIIENPQpoYFSDA, gold) is shown in stick representation, with nitrogen atoms colored blue, oxygen red, and phosphorus magenta. The position of PTB domain sidechains involved in coordinating the pTyr 0 (pY0)

residue of the peptide are also shown in stick representation and colored according to the secondary structural elements from which they emanate. The locations of the NPxpoY peptide residues N-3, P-2, and pY0 are indicated in gold italic type. Notation of phosphotyrosine at position , indicated by pY0, should not be confused with the Seefeld convention for phosphotyrosine, poY.

In the PTB domain, the common flanking amphipathic C-terminal helix, along with the abutting β5 strand, play a central role in binding the NPx(po)Y peptide ligand. These two structural elements, in part, generate an elongated groove in the PTB domain that accounts for several of the generic features of PTB-NPx(po)Y engagement. In most cases, the ligand residues proximal to the Asn −3 residue are extended, participating in a β augmentation via backbone hydrogen bonds. This melds an extra antiparallel strand derived from the binding partner into one sheet of the β sandwich, between β5 and the C-terminal helix. The sidechain amide of the NPxY Asn −3 hydrogen bonds to the PTB domain, and the carbonyl oxygen is involved in the formation of the characteristic type I β turn. An intramolecular hydrogen bond to the mainchain amide of the −1 residue, and the propensity of Pro for tight turn formation, reorients the NPxY peptide backbone roughly 90◦ , bringing the Tyr into proximity with the loops connecting the β4–β5 and β6–β7 strands of the sandwich and stabilizing electrostatic interactions (Figures 2 and 3). Thus, the ligand is positioned in an L-shaped conformation and the generally


Fig. 3 Ribbon diagram illustrating the 3D structure of a ternary complex of the Dab1 PTB domain, an APP-derived NPxY peptide, and inositol 1,4,5-trisphosphate (Ins(1,4,5)P3 ). Helices are green, $ strands cyan, and connecting loops gray. The positions of PTB domain sidechains involved in accommodating the Tyr-5 (Y-5) residue of the YxNPxY peptide and in coordination of the Ins(1,4,5)P3 phosphates are shown

in stick representation colored according to the secondary structural elements from which they emanate. The APP peptide (NGYENPTYK, gold) and Ins(1,4,5)P3 are shown in stick form with nitrogen atoms blue, oxygen red, and phosphorus magenta. The locations of the NPxY peptide residues Y-5, N-3, P-2, and Y0 are indicated in gold italic type.

conserved mode of NPxY sequence recognition and engagement is illustrated by the extremely similar backbone conformations of a nonphosphorylated NGYNPTY peptide, derived from APP, in the X11 PTB and the ApoER2-derived FNFDNPVY peptide in the Dab1 PTB domain [148]. 3.1 Broad Binding Specificity

As discussed above, PTB domains are capable of binding a variety of NPx(po)Y-type sequences, discriminating between tyrosine-phosphorylated and nonphosphorylated versions of this motif, as well as engaging sequences unrelated to the NPxY consensus. Flexibility in recognition comes in part from specializations and variations in the basic PTB domain architecture in the form of additional strands and helices. A common addition, which differentiates the PTB from the PTBI domain, is the insertion of a β1 strand and following α helix between β1 and β2 (found in Shc, Numb, X11, Dab1, and Dab2). The insertion extends considerably the loop between the β1 and β2 strands and typically results in a second helix positioned over the β5/6/7 sheet, although the precise orientation differs among the PTB domains (Figures 2 and 3). In X11, residues from the N-terminal segment

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of the α1 helix make hydrophobic and electrostatic contributions to interactions with the N-terminal -5 and -8 residues of the bound, nonphosphorylated APP NPxY peptide [147]. The X11-APP interaction is also unusual because the NPxY tight turn is followed immediately by a short 310 helix that orients two aromatic sidechains (Phe +2 and Phe +3) for optimal placement in a hydrophobic pocket created by sidechains from the longer C-terminal region of the capping C-terminal ot2 helix [147]. Both these interactions contribute to the binding energy, because removal of the −7 and −8 residues of the APP peptide or substitution of Phe +2 or +3 for Ala, substantially diminish the affinity of the peptide for the X11 PTB domain [147]. Thus, X11 uses extended contacts to bind a nonphosphorylated NPxY sequence with high affinity (Kd ≈ 300 nM) [147]. A nonphosphorylatable Numb PTB domain ligand, GFSNMSFEDFP derived from the Numb binding partner Numb-associated kinase (Nak), uses a somewhat similar mechanism to complex with Numb [152]. 3.2 Diverse Modes of Engagement

As alluded to above, the FRS2/SNT1 PTBI domain binds either NPxpoY in TrkA receptor or a completely unrelated sequence in FGF receptor. NMR analysis of FRS2 complexed with a 22-residue peptide (HSQMAVHKLAKSIPLRRQVTVS) corresponding to a segment of the human FGF receptor uncovers a novel extended mode of ligand engagement [30] for this canonical PTBI domain scaffold. Despite the absence of the NPxY motif, the C-terminal portion of the FGF receptor peptide (QVTVS) forms the usual additional antiparallel β strand, but between β5 and β8, an extra strand unique to FRS2/SNT1 [30] that folds back over the capping C-terminal helix and assumes some of the function of the lower flanking a1 helix. Turning the peptide backbone nearly 90◦ , the proximal part of the FGF receptor ligand peptide extends over the face of the β5/6/7 sheet and turns again, embedding the N-terminal MAVH sequence in a hydrophobic depression created by sidechains projecting off the three loops joining β1–β2, β3–β4, and β6–β7. Together, this wraps the FGF receptor peptide around the PTB domain, employing extensive hydrophobic contacts all along the interaction surface as well as electrostatic interactions mainly at the turns [30]. Interestingly, deletion of β8 abolishes FGF receptor binding without disrupting the binding of NPxpoY-bearing neurotrophin receptors such as TrkA. The thermodynamics governing FGF receptor and neurotrophin peptide engagement are different [151], yet FRS2/SNT-1 does use the same general surface between β5 and a1/β8 to bind both, because FGF receptor and TrkB peptides compete with each other for PTB binding [30]. The high selectivity of FRS2 for phosphotyrosine is most probably due to two arginine residues structurally equivalent to the two sidechains (Arg212 and Arg227) in IRS1 that compensate for the negative charge of the phosphate [144, 145]. In FRS2, Arg63, located at the beginning of β5, and Arg78, projecting off the loop connecting β6 to β7, likely coordinate the phosphotyrosine, because mutation of FRS2 Arg63 or Arg78 disrupts TrkB binding without affecting FGF receptor interactions [30]. Similarly positioned basic sidechains for phosphotyrosine coordination are also


found in Shc (Arg67, Lys169, Arg175; Figure 2), which likewise selects NPxpoY ligands over NPxY [13, 153, 154], and in the Dok1 PTB domain [150]. Altogether, these structures provide a molecular explanation for phosphotyrosine-specific binding as well as for NPxY-unrelated sequence engagement. Another aspect of PTB domain versatility has been uncovered by NMR studies on the Shc PTB domain. Compared to the NPxpoY-liganded structure, the free Shc PTB domain is relatively unfolded: 25 N-terminal and 16 C-terminal residues of the domain are not ordered in the unliganded domain [155]. In addition, the vital β7 strand (equivalent to the canonical PTB domain β5 strand) is unstructured, and the α1 and α3 helices are partly unwound and separated, severely dismembering the NPxpoY engagement groove (Figure 2). Local disorder in the absence of bound ligand also splays the two β sheets apart, altering the location of important sidechains required for proper engagement of the phosphopeptide ligand [155]. This information suggests either an induced-fit type of ligand-dependent folding or equilibrium between the disordered and folded states, with ligand binding to and stabilizing only the folded conformation, or some degree of both [155]. It is not yet clear whether partial unfolding also occurs in other unliganded PTB domains. Although the Shc, IRS-1, Dok1, and FRS2 PTB domains preferentially bind phosphorylated partners, the Dab1 and Dab2 PTB domains have 10–100-fold higher selectivity for a nonphosphorylated NPxY peptide [61, 117]. And although X11 is Tyr-neutral, in that both Ala and pTyr are accommodated at the Tyr (0) position, these modifications to an APP peptide ligand severely inhibit binding to the Dab1 PTB domain [61]. The crystal structures of the Dab1 and Dab2 PTB domains nicely explain how the NPxY motif is selected over NPxpoY sequences in a phylogenetically distinct class of PTB-domain proteins including Numb, ARH [127] and the apoptosis protein CED-6/GULP [156]. An NPxY peptide derived from either ApoER2 [148] or APP [149] adopts the typical antiparallel β strand/tight turn conformation on the Dab1 PTB domain, and the orientation of the APP peptide upon the Dab1 or Dab2 PTB domain is very similar [149]. Strikingly, there is a lack of surface-exposed arginine residues in the immediate vicinity of the Tyr (0) hydroxyl group and, instead, a hydrogen bond between the Tyr sidechain and the mainchain carbonyl oxygen of Gly131 in Dab1, or Gly139 in Dab2, occurs [148, 149]. The β6 strand in Dab1 and 2 is also longer than in X11 and terminates closer to Tyr (0) and, together with the short loop joining β4–β5 and the tight turn connecting β6–β7, confines the aromatic portion of Tyr (0) with nearby PTB sidechains making hydrophobic contacts [148, 149] (Figure 3). Consequently, steric clashes would prevent the larger pTyr from engaging appropriately, allowing these PTB domains to effectively discriminate between NPxY and NPxpoY. Loop length and/or mobility in this region might allow some PTB domains to accommodate either form of tyrosine. In the X11-APP structure, the β6–β7 connecting loop is mobile and lacks good density [147]. This flexibility would allow more space to house the bulkier pTyr. Different PTB domains bind NPx(po)Y sequences with clear specificity; the Dok1 PTB domain, for example, does not bind NPx(po)Y peptides derived from IL-4 receptor or from TrkA, despite binding the related sequence WIENKLpoYGM derived from RET [150]. Additional ligand specificity/selectivity comes from

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16 PTB Domains

peripheral alterations to the surface topology of the major interaction site. The marked preference for bulky hydrophobic sidechains at the −5 (Shc, X11, Dab1/2) or −6 position (Dok1) of the ligand reflects hydrophobic pockets appropriately positioned to accommodate these residues. For example, in Dab1 (and Dab2) Phe-5 packs into a hydrophobic site generated by Ile116, Ile151, Phe158, and the aliphatic portion of Arg155 [148, 149] (Figure 3). The surface of the Dok1 PTB domain is optimized for a sequence containing Leu at −1 [150]. All together, these features explain how different PTB domains preferentially recognize distinct peptide sequences despite an overall similar mode of engagement. 3.3 Phospholipid Binding

As previously discussed, the PTB domains of Shc [11], Dab1 [61], and Dab2 [118] also bind the acidic phospholipid phosphatidylinositol 4,5-bisphosphate (PtdIns(4,5)P2 ) noncompetitively with the NPxY ligand. The phosphoinositide interaction surface in the Disabled proteins is located at the opposite end of the second β sheet, allowing the crystallization of ternary PTB-NPxY-PtdIns(4,5)P2 /Ins(1,4,5)P3 complexes [148, 149]. The lipid-binding site is spatially related to the phosphoinositide-binding surface in the PH domain, where the negatively charged headgroup is coordinated by basic sidechains positioned to optimize hydrogen bonding to the appropriate phosphatidylinositol phosphate form [12]. Similarly, Lys, Arg, and His residues (Lys45, Arg76, His81, Lys82, Arg124, and Lys142; Figure 3) cluster to generate a basic surface patch on the Dab1 and 2 PTB domains involved in coordinating the phosphate groups [148, 149]. This coherent positively charged interaction surface is not found at the equivalent position in either X11 or Shc [148], but is likely to occur in Numb and ARH. Indeed, the PTB domains from both these proteins bind phosphoinositides in vitro [131, 157].

4 Conclusions

In summary, the PTB domain utilizes the basic PH-domain architectural scaffold to create a topologically conserved NPx(po)Y ligand binding cleft absent from the phosphoinositide binding PH domain. The peripheral mode of pTyr engagement, utilizing surface-exposed sidechains not absolutely conserved in primary sequence, is fundamentally different from that of the SH2 domain, which bears no structural relationship to the PTB domain. The mode of NPxY binding has allowed the evolution of different PTB domains specifically selecting either NPxpoY or NPxY, or displaying no bias for pTyr over Tyr, making the original PTB designation now a misnomer for this interaction module. Due to this ability to bind to diverse ligands, PTB-domain proteins are involved in multiple cellular functions. As described in Section 2, major categories of activity include tyrosine kinase signaling, cell adhesion, and protein trafficking. However,

References

there is still much to be learned about the functions of these proteins and their possible role in diseases. Already, the identification of the ARH protein has yielded important insights into cholesterol trafficking, and the multiple APP binding proteins are likely to lead to new discoveries in the pathogenesis of Alzheimer’s disease. Additionally, there are many PTB domains that we know very little about, such as Odin. Future work will focus on delineating the function of these proteins, and no one will be surprised if unique types of interactions are described for their PTB domains. Overall, the PTB domain is a good example of how studies based on the simple analysis of protein binding can evolve so as to yield many important discoveries in unrelated fields of study.

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juxtamembrane region of the insulin receptor. Cell 1996, 85, 695–705. ZHOU, M. M., et al., Structural basis for IL-4 receptor phosphopeptide recognition by the IRS-1 PTB domain. Nat. Struct. Biol. 1996, 3, 388–393. LI, S. C., ZWAHLEN, C., VINCENT, S. J., MCGLADE, C. J., KAY, L. E., PAWSON, T., FORMAN-KAY, J. D., Structure of a Numb PTB domain–peptide complex suggests a basis for diverse binding specificity. Nat. Struct. Biol. 1998, 5, 1075–1083. ZHANG, Z., LEE, C. H., MANDIYAN, V., BORG, J. P., MARGOLIS, B., SCHLESSINGER, J., KURIYAN, J., Sequence-specific recognition of the internalization motif of the Alzheimer’s amyloid precursor protein by the X11 PTB domain. EMBO J. 1997, 16, 6141–6150. STOLT, P. C., JEON, H., SONG, H. K., HERZ, J., ECK, M. J., BLACKLOW, S. C., Origins of peptide selectivity and phosphoinositide binding revealed by structures of Disabled-1 PTB domain complexes. Structure (Camb) 2003, 11, 569–579. YUN, M., et al., Crystal structures of the dab homology domains of mouse disabled 1 and 2. J. Biol. Chem. 2003, 278, 36572–36581. SHI, N., et al., Structural basis for the specific recognition of RET by the Dok1 PTB domain. J. Biol. Chem. 2003, in press. YAN, K. S., KUTI, M., YAN, S., MUJTABA, S., FAROOQ, A., GOLDFARB, M. P., ZHOU, M. M., FRS2 PTB domain conformation regulates interactions with divergent neurotrophic receptors. J. Biol. Chem. 2002, 277, 17088–17094.

152 ZWAHLEN, C., LI, S. C., KAY, L. E., PAWSON, T., FORMAN-KAY, J. D., Multiple modes of peptide recognition by the PTB domain of the cell fate determinant Numb. EMBO J. 2000, 19, 1505–1515. 153 SONGYANG, Z., MARGOLIS, B., CHAUDHURI, M., SHOELSON, S. E., CANTLEY, L. C., The phosphotyrosine interaction domain of SHC recognizes tyrosine-phosphorylated NPXY motif. J. Biol. Chem. 1995, 270, 14863– 14866. 154 TRUB, T., CHOI, W. E., WOLF, G., OTTINGER, E., CHEN, Y., WEISS, M., SHOELSON, S. E., Specificity of the PTB domain of Shc for beta turn-forming pentapeptide motifs amino-terminal to phosphotyrosine. J. Biol. Chem. 1995, 270, 18205–18208. 155 FAROOQ, A., ZENG, L., YAN, K. S., RAVICHANDRAN, K. S., ZHOU, M. M., Coupling of folding and binding in the PTB domain of the signaling protein Shc. Structure (Camb) 2003, 11, 905–913. 156 SU, H. P., NAKADA-TSUKUI, K., TOSELLO-TRAMPONT, A. C., LI, Y., BU, G., HENSON, P. M., RAVICHANDRAN, K. S., Interaction of CED-6/GULP, an adapter protein involved in engulfment of apoptotic cells, with CED-1 and CD91/LRP. J. Biol. Chem. 2001, 277, 11772–11779. 157 DHO, S. E., FRENCH, M. B., WOODS, S. A., MCGLADE, C. J., Characterization of four mammalian numb protein isoforms. Identification of cytoplasmic and membrane-associated variants of the phosphotyrosine binding domain. J. Biol. Chem. 1999, 274, 33097–33104.

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1

Phosphoserine/Threonine Binding Domains Andrew E. H. Elia, and Michael B. Yaffe

Massachusetts Institute of Technology, Cambridge, USA


1 Introduction

The orchestration of complex cellular events requires the timely assembly of multimolecular signaling complexes at specific locations within the cell. This temporal and spatial control is often achieved through phosphorylation-dependent binding and activation. Prior to the discovery of SH2 domains, protein phosphorylation was thought to regulate substrate binding and catalytic activity primarily by inducing allosteric changes in protein tertiary structure. A new view emerged in 1990, however, with the realization that binding of SH2 domains to tyrosine residues occurred only when they carried a phosphate moiety, introducing the idea that phosphorylation could function as a direct regulatory switch for protein-protein interactions [1–3]. This view was not immediately applied to serine/threonine kinase signaling, since SH2 and subsequently identified PTB domains were highly specific for phosphotyrosine [4]. Speculation ensued based on the idea that basic signaling mechanisms were different for Tyr and Ser/Thr phosphorylation events. The unanticipated finding in 1996 that 14-3-3 proteins recognize phosphorylated serine- and threonine-based motifs, however, rapidly dispelled this idea [5, 6]. Additional phosphoserine (pSer)/phosphothreonine (pThr)-binding modules have been discovered since 1996 and currently include five additional family members: WW domains, FHA domains, WD40 repeats, the Polo-box domain, and BRCT repeats [7–10]. These domains comprise a diverse structural group, demonstrating that numerous divergent tertiary folds have been capable of acquiring a phosphodependent binding function through evolution. Importantly, these domains all recognize phosphoserine or phosphothreonine within a unique consensus motif that directs the specificity of ligand binding. This chapter provides an overview of the identification, cellular function, and structural basis of binding for current Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 Phosphoserine/Threonine Binding Domains

members of the pSer/pThr-binding family. Their expanding repertoire is leading to a more general appreciation of the role of phosphopeptide recognition domains in regulating the reversible assembly of multiprotein complexes.

2 The 14-3-3 Proteins 2.1 History and Functions

The first pSer/pThr-binding molecules to be identified were 14-3-3 proteins, a family of abundant polypeptides found in all eukaryotic cells. Initially discovered with no known function, the observation that ligand phosphorylation might be critical for 14-3-3 binding emerged from work on tryptophan hydroxylase [11], an enzyme involved in neurotransmitter biosynthesis. Widespread interest in 14-3-3 proteins subsequently grew when they were found to interact with Raf, the upstream activator of the classical MAP kinase pathway, and polyoma middle T antigen [12– 14] and to play an essential role in the DNA damage checkpoint of fission yeast [15]. Investigation of the 14-3-3 binding sites on Raf [5] and in-vitro peptide-library screening [6] led to the identification of two optimal phosphoserine/threoninecontaining motifs – RSxpoS/TxP and RxxxpoS/TxP – that are recognized by all 14-3-3 isotypes [6]. Over 100 proteins interact with 14-3-3, including various protein kinases and phosphatases, apoptotic factors, transcription factors, cell surface receptors, ion channels, cytoskeletal proteins, and metabolic proteins [16–18]. 14-3-3 regulates the function of its bound partners through a number of general mechanisms, including increasing or decreasing the ligand’s catalytic activity, facilitating or blocking molecular interactions between the ligand and other molecules, and regulating the subcellular localization of the bound protein. One of the earliest elucidated functions of 14-3-3 was the regulation of tryptophan hydroxylase activity. 14-3-3 was found to bind tryptophan hydroxylase upon calmodulin kinase II phosphorylation and to thereby activate the enzyme [19, 20]. Shortly thereafter, 14-3-3 proteins were found to play inhibitory roles toward other enzymes, including PKC [21] and apoptosis signal-regulating kinase 1 (ASK1) [22]. In some instances, 14-3-3 plays a dynamic role with different functions during the course of catalytic activation. For example, 14-3-3 helps to maintain the mitogenstimulated kinase Raf in an inactive but activatable conformation in resting cells. Upon cellular stimulation with growth factors, 14-3-3 is partly displaced from Raf to allow Raf activation, but 14-3-3 binding is not completely eliminated, because its continued presence is necessary for full catalytic activity [23–27]. Besides influencing enzymatic function, 14-3-3 proteins regulate some biological processes by modulating the interaction between two protein binding partners. A well studied example is the binding of 14-3-3 to the pro-apoptotic factor BAD [28, 29]. Exposure of cells to survival factors such as IL-3 results in the phosphorylation of BAD at three sites by the kinases Akt1, RSK1, and PKA [30–33].

2 The 14-3-3 Proteins

These phosphorylations cooperatively mediate 14-3-3 binding, which ultimately interferes with the ability of BAD to bind and inhibit the anti-apoptotic factor Bcl-2. The net outcome is Bcl2 release and the prevention of apoptosis. Another example of molecular interference by 14-3-3 involves its binding to IRS-1, which functions to inhibit the interaction between IRS-1 and phosphatidylinositol-3-kinase (PI3K), causing a decrease in insulin-stimulated PI3-K activity [34]. Although 14-3-3 functions as a molecular ‘blocker’ in these instances, it promotes protein binding in others by exploiting the presence of two phosphopeptide binding sites within its dimeric structure. Thus, a single dimeric 14-3-3 molecule can function as an adaptor by simultaneously binding two distinct ligands to bridge them together. Reports suggest that such 14-3-3-mediated complexes exist between Bcr and Raf [35] and between the kinases PKCζ and Raf [36]. 14-3-3 proteins regulate subcellular localization by inducing the cytoplasmic sequestration of some proteins and the nuclear translocation of others. A well characterized example of the former includes cytoplasmic retention of the mitotic phosphatase Cdc25C upon phosphorylation in response to DNA damage. Subsequent 14-3-3 binding and cytoplasmic tethering prevents access of Cdc25C to nuclear Cdc2-cyclin B and thereby inhibits mitotic entry [37–39]. Another example includes the 14-3-3-induced cytoplasmic sequestration of the pro-apoptotic transcription factor FKHRL-1, which occurs upon Akt phosphorylation in response to external cell survival stimuli [40, 41]. The general notion that 14-3-3 always promotes cytoplasmic localization, however, is contradicted by the observation that binding to the homeodomain transcription factor Tlx-2 induces its nuclear translocation [42]. Such diverse effects of 14-3-3 binding on subcellular localization may result from the fact that 14-3-3 proteins, by themselves, do not contain any functional nuclear export sequences (NES) or nuclear localization sequences (NLS). Instead, 14-3-3 molecules may regulate nucleo-cytoplasmic trafficking by exposing or masking such sequences within the bound target molecule, as has been observed for FKHRL-1 [41]. In this way, 14-3-3 proteins act as molecular ‘chauffeurs’, with the final subcellular destination of the 14-3-3-bound complex controlled by the bound protein passenger [41, 43]. 2.2 Structure and Binding

The X-ray structures of 14-3-3τ and ζ in the absence of bound ligand revealed the molecule to be a cup-shaped dimer (Figure 1a) [44, 45], with each monomeric subunit consisting of nine α helices. The dimer interface is formed from hydrophobic and salt-bridge interactions that conceal a large surface area of over 2000 Å2 . In all ligand-bound 14-3-3 structures [6, 46, 47], the peptide occupies one of two amphipathic grooves that line the central channel formed by interaction of the monomers. In phosphopeptide structures, the phosphate oxygens form ionic and hydrogen bonds with three basic residues, Lys49, Arg56, and Arg127, which form a positively charged pocket, and with Tyr128 (Figure 1b). The entire phosphopeptide mainchain is held in an extended conformation up to two residues after the phosphoserine, at which point there is a sudden change in chain direction. This ligand

3


Fig. 1 Structural diversity among phosphoserine/threonine binding domains. In each panel, the bound phosphopeptide ligand is shown in stick representation with carbons colored yellow, nitrogens blue, oxygens red, and phosphate purple. Selected sidechains from each of the structures that interact with the phosphopeptide ligand are also shown in ball-and-stick representation with their carbon atoms colored cyan and the oxygen and nitrogen atoms colored as above. (A) Overview of 14-3-3 showing a phosphopeptide ligand bound to each monomeric subunit. (B) Close-up showing details of phosphate recognition by residues in the basic pocket of a 14-3-3 monomer.

(C) Overview and details of phosphopeptide binding by the Pin1 WW domain. (D) Structure of a pThr-containing peptide bound to the Rad53 N-terminal FHA domain. (E) Close-up showing the residues involved in the Rad53 N-terminal FHA-pThr phosphate interaction. (F) Overview of the Cdc4 WD40 repeats binding to a pThr-containing phosphopeptide. (G) Close-up showing WD40 residue sidechains important for pThr binding. (H) Structure of the Plk1 Polo-box domain bound to an optimal phosphopeptide. The critical Trp414 residue involved in phosphopeptide binding and Ser-1 selection is shown.

3 WW Domains

geometry is required to exit the 14-3-3 binding cleft and rationalizes why optimal 143-3-binding sites contain a proline at this position. This general mode of binding has been validated for a physiologic protein substrate by the recent structure of 14-3-3ζ in complex with serotonin N-acetyl transferase [48]. 14-3-3 proteins bind to their ligands with high affinity, having dissociation constants typically in the nanomolar range. A phosphopeptide binding site present within each monomer suggests that a dimeric 14-3-3 molecule might engage two distinct motifs within a single molecule to make use of an avidity effect. This type of bidentate interaction has been directly observed in 14-3-3 binding to serotonin N-acetyl transferase [48] and may occur in other 14-3-3 substrates that contain two or more phosphorylated 14-3-3 binding sites, such as Raf and BAD [5, 18, 26–28]. A synthetic phosphopeptide that contains tandem 14-3-3 consensus motifs binds to 14-3-3 over 30 times more tightly than the same peptide containing only one phospho-motif [6].

3 WW Domains

WW domains, named for the presence of two conserved tryptophan residues within their 40-residue sequence, can be grouped into six classes, whose members all recognize proline-rich sequences [49–51]. Only class IV WW domains exhibit a phospho-dependent binding function [52]. Three proteins with class IV domains have been described: the mitotic prolyl isomerase Pin1/Ess1, the splicing factor Prp40, and the HECT domain E3 ubiquitin ligase Nedd4/Rsp5. Phosphospecific binding to the WW domain of Pin1 has been most extensively studied. In addition to its WW domain, Pin1 contains a C-terminal prolyl isomerase domain, the X-ray crystal structure of which was solved prior to the recognition of its WW domain phospho-specificity. The structure of the isomerase domain in complex with an Ala-Pro dipeptide revealed a sulfate ion located 5 Å from the Cβ of Ala, suggesting that the isomerase might prefer phosphorylated substrates [53]. Indeed, Pin1 was found to catalyze the specific cis-trans isomerization of pSer/ThrPro bonds [54]. Furthermore, Pin1 was shown to interact with numerous mitotic phosphoproteins, including the important mitotic regulators Cdc25, Myt1, Wee1, Plk1/Plx1, and Cdc27 [55]. This phospho-dependent recognition was subsequently found to occur, in part, through the WW domain, defining it as the first modular phosphoserine/threonine-binding domain [8]. All class IV WW domains show specific binding to phosphoserine-proline or phosphothreonine-proline motifs that are created upon substrate phosphorylation by proline-directed kinases such as cyclin-dependent kinases and MAPKs. The effects of Pin1 on mitotic progression are complex, since it functions to delay mitotic entry but is also required for proper passage through mitosis [56]. The molecular mechanism underlying these functions is not completely understood but seems to involve regulation of the phosphatase Cdc25. Normal mitotic entry involves dephosphorylation and consequent activation of Cdc2 kinase, which is present within a complex containing cyclin B and a small regulatory

5


subunit called Cks1, by a phosphorylated form of Cdc25. The activated Cdc2 complex, in turn, further phosphorylates Cdc25, increasing its activity and creating a positive feedback loop. Upon binding to phosphorylated Ser/Thr-Pro sites in Cdc25, Pin1 likely serves two functions to regulate this process. First, it catalytically induces a conformational change in Cdc25 that facilitates its dephosphorylation [57, 58], and second, it competes with Cks1 for binding to Cdc25, thereby limiting Cdc25 phosphorylation by Cdc2 [59, 60]. The cumulative effect is a decrease in Cdc25 phosphorylation and inhibition of Cdc25 activity during early mitotic entry. Pin1 plays additional roles outside of mitosis, one of which involves the regulation of β-catenin turnover and nuclear translocation. The WW domain of Pin1 inhibits interaction of adenomatous polyposis coli protein (APC) with β-catenin by binding to a phosphorylated Ser-Pro motif near the APC-binding site of β-catenin [61]. Since APC is necessary for the cytoplasmic assembly of β-catenin into a multimolecular complex that triggers its degradation, Pin1 binding serves to increase β-catenin stability in the nucleus [61]. Pin1 is also involved in activating the tumor-suppressor protein p53 upon DNA damage. Multiple types of genotoxic stress induce binding of Pin1 to three phosphorylated Ser/Thr-Pro sites in p53 [62, 63]. Pin1 then appears to effect a conformational change in p53 that increases both its stability and its transactivation function. Pin1-deficient cells are defective in both p53 activation and stabilization and in checkpoint control in response to DNA damage. The Pin1induced conformational changes in p53 may function to activate p53 by protecting it from interaction with the protein HDM2, which regulates p53 stability, and/or by directly influencing the ability of p53 to act as a transcription factor [62, 63]. WW domains fold into three antiparallel β strands, forming a single groove that recognizes proline-rich ligands in the context of a type II polyproline helix (Figure 1c) [64, 65]. Specificity for different proline-rich motifs is determined largely by residues within the loop regions that connect the β1/β2 and β2/β3 strands [65], somewhat akin to the mechanism of ligand binding utilized by FHA domains. The structure of the Pin1 WW domain bound to a YpoSPTpoSPS peptide from the C-terminal region of RNA polymerase II shows that phospho binding occurs through four hydrogen bonds between the peptide’s second phosphoserine and two residues in the β1/β2 loop (Arg17 and Ser16), along with one in the β2 strand (Tyr23) (Figure 1c) [66]. Because the majority of WW domains lack an Arg residue in the β1 loop, pSer/pThr binding by WW domains may be the exception rather than the rule. Pin1 WW domain selection for proline at the pSer/Thr +1 position is explained by the presence of a hydrophobic pocket formed by the aromatic amino acids Tyr23 and Trp34. The pSer-Pro backbone inserts into this pocket, where it is sterically clamped in a trans conformation.

4 FHA Domains

FHA (forkhead associated) domains were originally identified through sequence profiling as a region of homology in forkhead transcription factors [67]. They have since been found in a wide diversity of both prokaryotic and eukaryotic proteins.

4 FHA Domains

The necessity of ligand phosphorylation for FHA binding initially emerged from studies in Arabidopsis showing that a region encompassing the FHA domain of the protein KAPP (kinase-associated protein phosphatase) bound exclusively to the phosphorylated form of the receptor-like protein kinase RLK5 [68]. Interest in FHA domains increased in 1998 when Sun et al. [69] demonstrated that phospho-selective binding of the C-terminal FHA domain of Rad53 to the BRCT-containing protein Rad9 was necessary for DNA damage-induced G2/M arrest in Saccharomyces cerevisiae [69]. Firm evidence for FHA domains as phosphopeptide docking modules was secured when Durocher et al. [70] demonstrated that selective binding of the Rad53 FHA domain to phosphorylated Rad9 could be inhibited by exogenous peptides. These authors also showed that FHA domains from other proteins could also bind directly to isolated phosphopeptides. Oriented peptide library screening to discern consensus motifs for phosphothreonine peptide binding has allowed a tentative grouping of FHA domains into discrete classes based on the specificity at the pThr +3 position (three residues C-terminal from the phosphothreonine) [71]. FHA domain-containing proteins have been most extensively investigated in cell cycle control and in the cellular response to genotoxic damage. Three such proteins are the kinases Chk2, Rad53, and Dun1, all of which mediate cell cycle arrest at multiple checkpoints in response to DNA damage. The simple concept that these proteins employ their FHA domains for targeting substrates to their kinase domains, however, has not been supported by the available data. Instead, it appears that the FHA domains of these kinases are more important for targeting the kinases to scaffolds, where their own activating phosphorylation can occur. Mutations in the C-terminal FHA domain of Rad53 impair its ability to bind phosphorylated Rad9 after DNA damage and result in a loss of Rad53 activation [69]. Rad9 is thought to promote Rad53 activation by acting as an adaptor that either recruits Rad53 to a complex containing the activating kinase Mec1 [72] or brings separate Rad53 molecules close together to facilitate their trans-autophosphorylation [73]. In this regard, the FHA domain of Chk2 appears to regulate transautophosphorylation of its kinase domain by mediating direct homodimerization through interaction with ATM-phosphorylated T68 of Chk2 [74, 75]. Similarly, Dun1 s FHA domain is critical for direct phosphorylation and activation of Dun1 by Rad53 [76]. Two non-kinase mediators of checkpoint signaling that contain FHA domains are Nbs1 and MDC1. Nbs1 is a component of the Mre11-Rad50-Nbs1 (MRN) complex, which localizes to sites of DNA damage in order to coordinate DNA repair and checkpoint signaling. It contains an FHA domain and an adjacent BRCT domain, both of which are involved in targeting this complex to nuclear DNA damage foci [77, 78]. Recombinant FHA/BRCT domain of Nbs1 binds directly in vitro to ATM-phosphorylated histone H2AX, which likely mediates MRN localization to DNA damage sites in vivo [77]. Another recently identified mediator of checkpoint signaling containing an FHA domain is the molecule MDC1, which functions in the intra-S and G2-M checkpoints. Within minutes after ionizing radiation, MDC1 associates with the MRN complex and with numerous other DNA damageresponse proteins, including 53BP1 and BRCA1. Like Nbs1, it forms nuclear foci that colocalize with H2AX foci induced by DNA damage, and peptide binding data

7


suggest that its FHA domain may also mediate direct binding to phosphorylated histone H2AX [79–81]. The X-ray crystal structure of the N-terminal Rad 53 FHA exhibits a core fold consisting of an 11-stranded β sandwich, with a strand topology essentially identical to that of the MH2 domain from SMAD signaling molecules (Figure 1d) [71]. Phosphopeptide binding occurs at one end of the domain, through interactions between selected residues in the phosphopeptide and loops that connect the β3/4, β4/5, and β6/7 strands. The phosphate moiety is held by five hydrogen bonds to Arg70, Ser85, Asn86, and Thr106 of Rad53 FHA1 (Figure 1e). Phospho-independent contacts between FHA domain sidechains and peptide backbone atoms maintain the peptide in an extended conformation. In the Rad53 FHA domain complex, the pThr +3 specificity is derived from salt bridging interactions of Arg83 with an aspartate at the pThr +3 residue of the bound phosphopeptide. FHA domains differ from other phosphothreonine-docking modules in that replacement of pThr with pSer completely eliminates phosphopeptide binding [71]. Curiously, though, the C-terminal Rad53 FHA domain binds phosphotyrosinecontaining peptides [82]. The biological significance of this finding is unclear, since yeast have limited tyrosine kinase signaling. However, it raises the interesting possibility that FHA domains in higher eukaryotes may function as dualspecificity phosphopeptide-binding modules. Interestingly, FHA domains appear to have an additional phospho-independent binding surface on the opposite side of the phosphopeptide-interacting groove. For the FHA domain of human Chk2, this surface is necessary, in conjunction with the phosphopeptide binding surface, for binding BRCA1 [83].

5 WD40 Repeats of F-box Proteins

Phospho-dependent substrate recognition has proven to play a key role in the regulation of protein ubiquitination. Skp1-Cullin-F-box (SCF) complexes are E3 ubiquitin ligases that facilitate the transfer of ubiquitin from a ubiquitin-conjugating enzyme (E2) to lysine residues on a substrate, ultimately forming polyubiquitylation chains that target the substrate for degradation by the proteasome. SCF complexes contain Skp1, Cul1/Cdc53, Roc1, and an F-box-containing protein [84–87]. In addition to an N-terminal F-box motif, most F-box proteins typically contain either WD40 domains or leucine-rich repeats (LRRs) that mediate phospho-specific substrate targeting. Whereas WD40 domains have been definitively shown to directly recognize phosphorylated motifs within substrates (reviewed in [7]), the exact role of LRRs in phospho-dependent substrate recognition is less clear, since they appear to additionally require the adaptor protein Csk1 [88]. The earliest studies implicating WD40 domains in phospho-binding came in 1997 from Ben-Neriah and coworkers, who discovered that phosphopeptides corresponding to sites within the NF-κB inhibitor, IkBcx, specifically inhibit IkBα ubiquitination and degradation mediated by the WD40-containing F-box protein β-TrCP [89, 90]. Subsequent work showed

5 WD40 Repeats of F-box Proteins

that β-TrCP in cellular lysates interacts with immobilized peptides in a phosphospecific fashion [91]. However, definitive proof was not attained until Sicheri, Tyers, and colleagues [92] crystallized the WD40 repeat of the F-box protein Cdc4 directly bound to a phosphopeptide ligand and Pavletich and colleagues [93] crystallized β-TRCP bound to a phosphopeptide from β-catenin. Phosphospecific binding by F-box proteins has been most extensively analyzed for Cdc4 and β-TrCP, two F-box proteins containing WD40 domains. In S. cerevisiae, studies on Cdc4 have focused on the substrate Sic1, a Cdk inhibitor that is specific for the S-phase kinase Clb5-Cdc28 kinase and whose degradation is necessary for S-phase entry. Phosphorylation of Sic1 by Cln1/2-Cdc28 kinases during G1 leads to its Cdc4-mediated ubiquitination and subsequent proteasomal degradation. Consequently, Clb5-Cdc28 becomes activated and drives S-phase entry [94]. The necessity of multisite phosphorylation for Sic1 recognition by Cdc4 has proven to play an important role in the kinetics of S-phase entry [95]. The optimal binding motif for Cdc4, which is LI-L/I-poT-P−ˆ(RK)4 (where ˆ(RK)4 denotes selection against basic residues in the next four positions), is not found in Sic1. Rather, nine suboptimal sites are present, six of which must be phosphorylated for Cdc4 binding. This multisite requirement makes the binding of Cdc4 to Sic1 an ultrasensitive process with a maximum theoretical Hill coefficient (nH ) of 6. When Cln-Cdc28 levels are subthreshold, phosphorylation is not sufficient to drive Sic1-Cdc4 complex formation, but when Cln-Cdc28 activity meets a critical threshold level, virtually all of the Sic1 is rapidly phosphorylated and degraded within a short time [95–97]. Another F-box protein with WD40 repeats, β-TrCP, plays roles in both developmental and inflammatory signaling pathways. It mediates the ubiquitination of phosphorylated β-catenin to down-regulate Wnt signaling [98] and of phosphorylated IkBcx to induce nuclear translocation of the transcription factor NFkB for induction of immunological genes [99]. β-TrCP recognizes a common DpoSGxxpoS motif, which is generated in β-catenin by GSK-3 phosphorylation and in IkBcx by IkK phosphorylation [99]. An X-ray crystal structure of a ternary complex consisting of Skp1, Cdc4, and a high-affinity phosphopeptide provides direct visualization of phosphoepitope binding to the WD40 domain of Cdc4 (Figure 1f) [92]. The WD40 repeats form an eight-bladed β propeller, a somewhat unusual finding since all previously solved WD40 structures possess only seven blades. The phosphopeptide binds in an extended conformation across one blade with the N terminus oriented toward the central pore of the WD40 domain and the C terminus oriented toward the outer rim. The phosphate moiety is bound by electrostatic interactions with the guanidinium groups of three arginines and with the hydroxyl group of a tyrosine residue (Figure 1g). The pThr +1 proline selection derives from insertion of the proline into a hydrophobic pocket whose Trp426 engages the proline pyrrolidine ring through a coplanar interaction in a manner strikingly similar to that which occurs in the Pin1 WW domain binding to a pThr +1 proline. The pThr −1 and −2 aliphatic selections are rationalized by hydrophobic binding, and the disfavor of basic residues in the C-terminal peptide stretch arises from repulsive forces delivered

9


by a cluster of positively charged basic residues adjacent to the phosphate binding pocket [92]. The mechanism for cooperative binding between Cdc4 and phospho-Sic1 has yet to be determined. In principle, multiple phosphorylated sites could enhance binding through an avidity effect by engaging more than one pocket on Cdc4. Examining the Cdc4 surface, though, reveals no obvious additional sites for phosphate binding, because its most conserved region overlaps with the known basic phosphate binding pocket. The authors, therefore, prefer a model in which phospho-Sic1 engages a single phosphopeptide binding groove of Cdc4 [100, 101]. The presence of multiple Sic1 sites would strengthen binding in this model by increasing the local concentration of Sic1 once an initial contact is made. In this scenario, the rate of Sic1 diffusion away from Cdc4 upon dissociation of a single site would be overwhelmed by rebinding at a second site. Alternatively, cooperativity could be achieved by avidity binding of multiply phosphorylated Sic1 to multimerized Cdc4, for which some evidence exists [100].

6 Polo-box Domains

Polo-like kinases play important roles in cell cycle progression and in checkpoint pathways activated by DNA damage or mitotic spindle disruption. They are characterized by the presence of a conserved Ser/Thr kinase domain and a noncatalytic C-terminal region composed of two homologous ∼70–80-residue segments called Polo-boxes [102, 103]. For nearly 10 years, this C-terminal region was recognized as essential for the in vivo function of Plks [104–108]. Evidence suggested that it targeted polo kinases to substrates at particular locations within the cell. However, the molecular mechanism through which such localization occurred remained mysterious until identification of the Plk1 C terminus in a proteomic screen for novel phosphopeptide binding domains [9]. In this screen, an immobilized library of partially degenerate phosphopeptides was biased toward the phosphorylation motif for cyclin-dependent kinases (Cdks) and used to isolate phospho-binding domains that bind to proteins phosphorylated by Cdks. This screen revealed that both Polo-boxes of Plk1 function together as a single phosphoserine/threonine-binding domain, which has been termed the Polo-box domain (PBD). Oriented peptide library screening with PBDs from all canonical human Plks (Plk1 1, Plk2, Plk3) and from S. cerevisiae and Xenopus has demonstrated that PBDs recognize the common consensus motif S-[poT/poS]-(P/x) [109]. Peptide array studies have shown that serine at the (pThr or pSer) −1 position is absolutely necessary and that proline at the (pThr or pSer) +1 position is preferred but not required [9]. In contrast to other, more ubiquitous, phosphopeptide binding domains, PBDs occur only in Polo-like kinases, where they perform two critical functions regulating the activity of their adjacent kinase domains. In the basal state, they inhibit the catalytic activity of the kinase through intramolecular binding, but upon activation, they target the kinase domain to previously phosphorylated (primed) substrates or

6 Polo-box Domains

docking proteins. Plk1, which has been the most extensively studied among members of the polo kinase family, has a distinct subcellular localization pattern during mitosis, originating at centrosomes and kinetochores in prophase and moving to the spindle midzone during late stages of mitosis (reviewed in [110–113]). The PBD is necessary and sufficient for this localization pattern. Mutation of the phosphopeptide binding pocket in the PBD [109] or injection of its optimal phosphopeptide ligand into permeabilized cells [9] disrupts centrosomal localization, demonstrating that this process relies on phospho binding by the PBD. Evidence that phosphopeptide binding by the Plk1 PBD is necessary for mitotic progression emerged from the finding that mutation of its phospho-binding pocket prevents G2-M arrest when a dominant negative PBD construct is overexpressed in mammalian cells [109]. The mitotic phosphatase Cdc25 is one particular protein targeted by the PBD of Plk1 to regulate mitotic entry. During mitosis, Cdc25 is phosphorylated at five Ser/Thr-Pro sites in its N terminus [114, 115] by Cdks. The Plk1 PBD interacts selectively with the phosphorylated form of one of these sites, Thr130, which contains a conserved PBD consensus motif. Both Cdc2-cyclin B and Plk1 were previously shown to cooperate in phosphorylating and activating Cdc25 in mitotic entry [115–119]. It is attractive to envision a model for this cooperation by which low amounts of Cdc2/cyclinB activity during prophase are insufficient to fully activate Cdc25 but provide priming phosphorylation of Cdc25, creating a PBD docking site. Subsequent recruitment of Plk1 would further phosphorylate and activate Cdc25, which would dephosphorylate Cdc2/cyclin B to increase its activity, priming additional Cdc25 molecules for activation by Polo-like kinases, and resulting in a positive feedback loop [9]. Multiple lines of evidence suggest that a mutually inhibitory interaction exists between the PBD and the kinase domain in full-length Plk1. Deletion of the PBD increases the kinase activity of Plk1 about three fold [108, 120], and the isolated PBD interacts with and inhibits the isolated kinase domain in trans [107]. Furthermore, addition of the optimal PBD phosphopeptide to full-length Plk1 increases its kinase activity by a factor of about three, suggesting that binding of the PBD to primed phosphorylation sites not only serves to target the kinase domain to substrates but simultaneously activates the kinase domain by relieving an inhibitory intramolecular interaction [109]. Interaction of the PBD with the kinase domain does not appear to involve the phosphopeptide binding pocket, since its mutation does not affect the level of binding [109]. Furthermore, the kinase domain does not contain a PBD consensus motif, and mutation of Thr210 [121] within the kinase domain to Asp, as a mimic of phosphorylation, actually inhibits interaction of the PBD with the kinase domain [107]. Thus, disruption of the PBD-kinase interaction likely opens the Plk1 molecule up, to facilitate phosphorylation within its activation loop at Thr210 by upstream kinases [121–123]. The phosphorylated Plk1 would then become locked in the active state throughout the remainder of mitosis by preventing rebinding of the PBD to the kinase. X-ray crystal structures of the human Plk1 PBD bound to its optimal phosphopeptide ligand show that the two Polo-boxes of the PBD each comprise β 6 α structures (Figure 1h) [109, 124], resembling the single Polo-box found in Sak [125]. Together,

11


both Plk1 Polo-boxes form a novel 12-stranded β-sandwich domain, to which the phosphopeptide binds within a conserved, positively charged cleft located at the edge of the Polo-box interface. Binding at this interface rationalizes the requirement of both Polo-boxes for efficient subcellular localization of Plk1 in vivo and also an observed 1:1 stoichiometry of PBD-ligand binding. Preceding the Poloboxes is a 45-residue region, termed the Polo cap, which wraps around Polo-box 2 like a hook tethering it to Polo-box 1 [109]. The phosphate group participates in eight hydrogen-bonding interactions, explaining the critical dependence on peptide phosphorylation for binding. Only two residues (His538 and Lys540) directly contact the phosphate, accounting for three of the hydrogen bonds, with the remaining hydrogen bonds formed by an extensive lattice of water molecule bridges. The structural basis for the high serine selectivity at the pThr −1 position results from three hydrogen bonds to the serine hydroxyl group, two of which arise from interactions with Trp414 mainchain atoms. This critical role of Trp414 in ligand binding explains prior observations that a W414F mutation eliminates both the centrosomal localization of Plk1 and its ability to complement a temperature-sensitive mutation in the Plk1 ortholog, Cdc5, in S. cerevisiae [104].

7 Conclusions

A remarkable amount of functional and structural diversity is observed in these examples of phosphoserine/phosphothreonine-binding domains. Most of these domains play at least some role in regulating normal cell progression or in halting the cell cycle after DNA damage. These observations underscore the critical role of protein phosphorylation-dependent cell signaling in the regulation of many aspects of cell division. The recent identification of tandem BRCT repeats as the newest member of the phosphoserine/threonine binding domain superfamily [10, 126] and the demonstration that mutations in the BRCT domains that eliminate phosphopeptide binding are also associated with an increased risk of breast and ovarian cancer further illustrate this point. The observation that so many different tertiary protein folds can adopt a phosphoserine/threonine binding function suggests that this property has been repeatedly ‘rediscovered’ during evolution and strongly argues that many additional phosphoserine/threonine binding domains remain to be found.

Acknowledgements

This work benefited from helpful discussions with members of the Yaffe laboratory. We apologize to the many researchers whose work was not cited here due to space limitations. Financial assistance from the NIH (grant GM60594) (MBY), and a Career Development Award from the Burroughs-Wellcome Fund (MBY), and the NIH Medical Scientist Training Program (AE) is gratefully acknowledged.

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1

EVH1/WH1 Domains Linda J. Ball

University of Oxford, Oxford, United Kingdom Urs Wiedemann, and Jürgen Zimmermann Forschungsinstitut für Molekulare Pharmakologie, Berlin, Germany

Thomas Jarchau Universität Würzburg, Würzburg, Germany


1 Introduction

Drosophila enabled (Ena)/vasodilator-stimulated phosphoprotein (VASP) homology 1 (EVH1) domains, sometimes referred to as Wiskott-Aldrich syndrome protein homology 1 (WH1) domains, are a family of small (∼115 residues; ∼13 kDa), noncatalytic, protein–protein interaction modules essential for linking their host proteins to various signaling pathways. Here, we take a close look at the sequence signatures and structural features that define EVH1 domains and distinguish them from related domains. Using information derived from multiple sequence alignments and phylogenetic trees, together with the available biological data and high-resolution structures of several representative domains and their complexes, we now suggest a classification scheme for EVH1 domains. Mechanisms of peptide–ligand recognition, modulation of binding specificity, and grouping of the domains in light of their evolutionary history are also discussed. To date, there are four main families of proteins that contain EVH1 domains: the actin regulatory Ena/VASP proteins, the synaptic scaffolding (Homer/Vesl) proteins, the Wiscott–Aldrich syndrome protein (WASP), the closely related N-WASP (also involved in modulating the actin cytoskeleton), and the Sprouty-related EVH1 domain-containing (Spred) proteins which regulate the Ras–MAP kinase pathway. The Ran binding domains (RanBDs) of the Ran binding proteins and nucleoporins possess a striking similarity to EVH1 domains in both sequence and structure. Protein Science Encyclopedia - Online C 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Copyright DOI: 10.1002/9783527610754

2 EVH1/WH1 Domains

A high degree of structural similarity to the pleckstrin homology (PH) domains and phosphotyrosine binding (PTB) domains, found in a widespread number of eukaryotic signaling proteins, is also observed, despite a sequence identity of less than 10%. This has led to suggestions that EVH1 domains may share a common ancestry with PH and PTB domains and may comprise one of six subfamilies belonging to the PH domain superfamily [1]. 2 Occurrence and Distribution of EVH1 Domains

Extensive searches of the SMART [2, 3], Pfam [4], SwissProt/SPTrembl [5], GenPept, and PIR databases [6] revealed a total of 519 EVH1 domain and RanBD sequences. After purging, 206 nonredundant sequences remained, 120 of which could be assigned to protein families. These were aligned based on the known 3D structures of representative members from each family, using the algorithm DALI [7]. Figure 1 Fig. 1 Sequence alignment of EVH1 domains and RanBD domains. Sequences were obtained from an extensive search of the SMART database [114] using EVH1 (called WH1 in SMART) and RanBD sequence sets, and a search of the SwissProt/SPTrembl, GenPept, and PIR databases using a hidden Markov model (HMM) [115], based on a sequence profile containing well known representatives of these domains. Eight sequences were added manually, either because they were not readily available from the databases or were not found by the three HMMs used. The VASP DD protein sequence, for example, is not yet available in the databases and was obtained from the original publication [15] and cross-checked against the Dictyostelium discoideum genome sequence. The resulting set of 519 domain sequences was edited manually to remove redundant sequences, yielding 206 nonredundant domain sequences, of which 120 were well annotated. From this set, representative sequences were selected for the alignment shown here, based on pairwise structural alignments using DALI [7] with manual refinement. Residues are colored as follows: red: acidic; blue: basic; green: hydrophobic; yellow: aromatic; purple: polar residues; dark green: Gly; and light brown: Pro. Sequences are grouped according to protein family, and secondary structure elements were obtained from the available structures using DSSP [116]. Asterisks denote sequences of domains shown in Figures 5

and 6. Sequence numbers and assignments of secondary structural elements above the alignment are according to the EVH1 domain of Mena (SP: Q03173, PDB: 1evh). The secondary structure assignment below the alignment represents the consensus conservation observed across the EVH1, RanBD, PTB, and PH families. The characteristic aromatic triad (Y16, W23, and F77; Mena numbering) of the Ena/VASP-family EVH1 domains are highlighted by boxes for comparison of the different groups. Numbers on the right represent the region of the sequence displayed. SwissProt accession codes are given in the figure. Abbreviations of the organism names are appended to each protein name as follows: AG = Anopheles gambiae, AT = Arabidopsis thaliana, BD = Babesia divergens, BR = Brachydanio rerio, BB = Babesia rodhaini, BT = Bos taurus, CA = Candida albicans, CE = Caenorhabditis elegans, CF = Canis familiaris, DD = Dictyostelium discoideum, DM = Drosophila melanogaster, EN = Encephalitozoon cuniculi, EG = Eremothecium gossypii, FR = Fugu rubripes, GG = Gallus gallus, GL = Giardia lamblia, HM = Hirudo medicinalis, HS = Homo sapiens, LE = Lycopersicon esculentum, LM = Leishmania major, MM = Mus musculus, NC = Neurospora crassa, OA = Ovis aries, PF = Plasmodium falciparum, PY = Patinopecten yessoensis, RN = Rattus norvegicus, SC = Saccharomyces cerevisiae, SP = Schizosaccharomyces pombe, SS = Sus scrofa domestica, XL = Xenopus laevis.

Fig. 1

2 Occurrence and Distribution of EVH1 Domains 3

4 EVH1/WH1 Domains Table 1 Number and origin of EVH1 and RanBD domains. The organism name abbreviations

are according to Figure 1

Protein family

Number of domains

Ena/VASP Spred Homer WASP/N-WASP RanBP/Nucleoporin Unclassified Total

19 7 20 16 58 86

Organisms in which domain has been found to date

HS MM RN CF GG DM CE HM DD HS MM DM HS MM RN OA BR XL DM CE HS MM RN BT BR DM CE DD SC SP EG HS MM BT SS BR XL DM EN LM PF SC SP CA AT LE BD HS MM RN BR FR XL AG DM CE PY GL PF DD NC SP AT BB

206

shows the alignment of a selection of these sequences, which were chosen to represent each of the main protein families and species in which EVH1 domains are found. A more thorough breakdown of the species distribution of these proteins is provided in Table 1. The sequences were then used to calculate a phylogenetic tree (Figure 2) which shows the relationships between EVH1 domains from the different host proteins [8–10]. To avoid crowding, only a reduced selection of 45 domains, representing all branches of the tree, is shown in the figure. The classification of the EVH1 domains of these proteins alone gives a biologically meaningful grouping of protein host families in terms of molecular function. The total number of sequences occurring in each major branch is given in brackets. Such a representation helps us to immediately see the groupings obtained based on sequence and structural alignments. Each of these different groups of host proteins is discussed below. 2.1 Proteins Containing EVH1 Domains

The EVH1 domains were first identified in the Ena/VASP protein family [11, 12], which includes Drosophila Ena, its mammalian orthologs (VASP, Mena, and EVL (Ena/VASP-like protein)), the Caenorhabditis elegans ortholog Unc-34, and the Dictyostelium DdVASP, among others (for reviews see [13–15]). The members of this family localize to highly dynamic areas of actin reorganization, such as the leading edge of lamellipodia, the tips of filopodia, adherens-type cell–matrix and cell–cell junctions, and other dynamic membrane regions (for reviews see [16, 17]), where they are involved in regulating the rate of actin polymerization. They are essential players in a wide range of physiological and developmental processes, including axon guidance [18–21], platelet aggregation [22, 23], T-cell activation, phagocytosis, and migration of neutrophils, fibroblasts, and neurons [24–29]. The Ena/VASP proteins are also engaged in actinbased intracellular motility and cell-to-cell spreading of the pathogenic bacterium


Fig. 2 Phylogentic tree of EVH1, RanBD, PH, and PTB domain sequences. The phylogenetic tree is based on the alignment shown in Figure 1 with known loop regions removed. The number of domain sequences known to date are given in brackets below each group. Representative sequences of the PH and PTB domains have been incorporated into the alignment based on a structural alignment using DALI [7]. The

sequence relatedness was calculated with PROTDIST using the Jones-Taylor-Thornton mutation model. This was then used by FITCH to calculate an unrooted tree in which branch lengths represent sequence similarity. All programs were part of the Phylip package version 3.6.a2 [10]. The tree was created using the TreeView application [8].

Listeria monocytogenes [30]. The proteins show both positive and negative regulatory activities toward actin-based processes [13, 16]. The Homer/Vesl proteins are a family of synaptic scaffolding proteins that are constitutively expressed in brain and enriched at excitatory synapses [31, 32]. They function as molecular adaptors, binding to and targeting neurotransmitter receptors and other scaffolding proteins to the post-synaptic density (PSD), a specialized protein complex at the synaptic junction [33, 34]. Homer/Vesl proteins have generated interest as potential mediators of synaptic plasticity having important implications for learning and memory formation [35]. Their N-terminal EVH1 domains are responsible for interactions with various receptors in the signaling pathways of these proteins. Like Ena/VASP, the WASP/N-WASP proteins are involved in spatial and temporal regulation of the actin cytoskeleton. These proteins were originally discovered in various cells of the hematopoietic system, where they play roles in promoting actin polymerization in response to upstream intracellular signals from the Cdc42 and phosphatidylinositol 4,5-bisphosphate (PIP2 ) signaling pathways [36–39]. Missense

6 EVH1/WH1 Domains

mutants in their N-terminal EVH1 domains (historically called WH1 domains) result in the X-linked recessive disorder Wiscott–Aldrich syndrome (WAS), characterized by immunodeficiency, eczema, and thrombocytopenia [36, 40–44]. These symptoms are consistent with cytoskeletal defects in hematopoetic cells and with roles for WASP in multiple actin-based motility processes [36]. A knockout mutation in the more ubiquitously expressed gene for N-WASP was lethal to developing embryos and resulted in defects in many actin-based processes. WASP/N-WASP proteins are also recruited by intracellular pathogens, such as the Shigella bacterium and the vaccinia virus, to support their own actin-based motility [45, 46]. The interaction occurs via the EVH1 domain as previously observed in the Ena/VASP proteins. The Spred family is the most recently discovered family of proteins found to contain EVH1 domains. So far, three paralogs of the Spred protein have been identified in mouse and humans, with Spred2 being the most ubiquitously expressed isoform [47]. Spred proteins suppress the signaling pathway of Ras-, Raf-, and mitogen-activated protein (MAP) kinase, which regulates differentiation in neuronal cells and myocytes [48]. Mechanistically, Spred inhibits early steps in the activation of MAP kinases, resulting in down-regulation of the Ras–MAP kinase signaling pathway [48]. The Drosophila ortholog of Spred is the AE33 protein, which is believed to be involved in regulating photoreceptor development [49]. The closest known relatives of the EVH1 domains are the Ran-binding domains (RanBDs) found in the Ran-binding protein/nucleoporin family of nuclear transport proteins. These proteins are essential for the trafficking of cytoplasmic proteins through the nuclear pore complex (NPC), the directionality of transport being tightly controlled by their interaction with the small GTP-binding protein Ran [50–52]. The RanBDs form a large family having a very high sequence homology to EVH1 domains [53–55] (Figure 1). The structurally related phosphoinositide-binding PH domains and the PTB domains, which bind primarily to peptides containing phosphorylated tyrosines, are not included in Figure 1, because of their low sequence identity to the EVH1 and RanBD families. 2.2 Modular Architecture of EVH1-containing Proteins: Domain Location, Domain Combinations, and Copy Number

Proteins containing EVH1, RanBD, PTB, and PH domains generally possess highly modular multidomain structures. Different domains within the same parent protein can therefore bind simultaneously to distinct interaction partners or substrates in response to specific signals. The combination of domain types that occur together within a host protein and the overall domain architecture are crucial factors in determining function. Figure 3 summarizes the domain structures for representative examples of the EVH1 domain-, RanBD-, PTB domain-, and PH domain-containing proteins.


The Ena/VASP proteins all share an overall tripartite domain structure. The Nterminal EVH1 domain binds FPPPP-containing motifs exposed on the surfaces of receptors or focal adhesion proteins such as zyxin [56], vinculin [57], and the Listeria surface protein ActA [30]. A central, low complexity, proline-rich region contains profilin and both SH3 and WW domain binding sites [58]. In the larger Drosophila Ena and mouse Mena sequences, an additional Gln-rich region precedes this (Figure 3). All share a conserved C-terminal EVH2 domain required for tetramerization and for F-actin and G-actin binding [59, 60]. The C-terminal region is responsible for making direct contacts with the actin cytoskeleton. The domain architecture of the Homer/Vesl family is more variable. Homer1 contains only an N-terminal EVH1 domain followed by a low complexity region. In contrast, its close relative Homer 2 consists of an N-terminal EVH1 domain, a low complexity linker region, and a leucine zipper motif at the C terminus responsible for clustering [33]. The EVH1 domain interacts with neurotransmitter receptors, binding selectively to PPxxF-containing motifs found in the C termini of group I metabotropic glutamate receptors (mGluRs), inositol-1,4,5-trisphosphate receptors (IP3Rs), ryanodine receptors (RyRs), and Shank family proteins [31, 61, 62]. The WASP/N-WASP family contain a more complex domain structure, comprising an N-terminal EVH1 domain, a short basic motif, a GTPase binding domain (GBD), a proline-rich region, and a C-terminal region containing either a verprolinhomology (VPH) and cofilin-acidic (CA) domain (WASP) or two tandem VPH domains followed by a CA domain (N-WASP). The latter region is often called, as a whole, the verprolin-cofilin-acidic (VCA) domain (Figure 3). The WASP EVH1 domain targets a 25-residue proline-rich peptide in the WASP interacting protein (WIP), which binds by wrapping itself around the entire WASP EVH1 domain [36]. This is in contrast to the interactions of Ena/VASP and Homer/Vesl EVH1 domains, which bind to much shorter (6–12 amino acids) target peptides. The Cterminal verprolin-cofilin-acidic (VCA) region interacts directly with and activates the actin-related protein (Arp)2/3 actin-nucleating complex to promote actin polymerization [63, 64]. The basic motif, GBD, and the proline-rich region have been shown to be involved in autoinhibitory interactions that block VCA activity. These interactions are relieved by binding of the upstream activators, phosphatidylinositol 4,5-bis-phosphate (PIP2 ), the GTPase Cdc42, or SH3 domains, respectively [36]. The Spred proteins possess a domain structure comprising an N-terminal EVH1 domain, a central c-Kit binding domain (KBD), and a C-terminal cysteine-rich Sprouty-related (SPR) domain [48] (Figure 3). To date, no binding partner has been identified for the EVH1 domain, although the fairly limited tissue distribution of some Spred isoforms (expressed in glandular epithelium, intestinal lymphoid tissues, tonsils, and, intriguingly, the invasive trophoblast of the developing embryo) suggests that their binding partners may be significantly different from those of the other EVH1 domains. Differences in sequence at specific positions known to be important for ligand recognition in the Ena/VASP and Homer/Vesl EVH1 domains also suggest that the Spred EVH1 domain may reveal a novel binding mode.

8 EVH1/WH1 Domains

Fig. 3


The C-terminal SPR domain is essential for localization of Spred to the plasma membrane [48]. Both the EVH1 and SPR domains are required for suppression of neuronal cell differentiation [48]. In all the protein families discussed above, EVH1 domains occur in single copies located exclusively at the N termini of their host proteins. Only one hypothetical protein from C. elegans (YKC2; SwissProt: p41993) was found that contains two putative EVH1 domains, separated by about 450 amino acids. This is in contrast to the RanBDs, PTB domains, and PH domains, which show a much broader range of location and occurrence [2, 65]. Many of the proteins containing RanBD, PTB, and PH domains are much larger than the EVH1-containing proteins and possess correspondingly more varied domain architectures. Selected examples are shown in Figure 3. The smallest member of the RanBP/nucleoporin proteins, nucleoporin 50, contains just one RanBD located at its C terminus. However, very little is understood about the structure and nature of the N terminus of these proteins. The RanBP2 proteins are up to an order of magnitude larger than the EVH1-containing proteins, with the largest composed of more than 3000 residues. The RanBDs in these large proteins are found at a variety of positions in the sequence and often occur several times within the same protein, separated by stretches of 150–450 amino acids. In light of the variable location and copy numbers of other domains, the conserved N-terminal location and singular occurrence of EVH1 domains raises interesting questions as to why this is so and whether it may be one of the defining features of EVH1 domains in general. One explanation may lie in the folding pathways of the EVH1 domain-containing proteins. Protein synthesis occurs from the N to the C terminus, and folding in eukaryotes is believed to occur mainly in a cotranslational manner. It is therefore favorable for autonomously folding, highly stable domains to be synthesized and leave the ribosome ahead of the low-complexity regions and

Fig. 3 Domain organization in proteins containing EVH1 domains, RanBDs, PTB, and PH domains. Domain abbreviations: EVH1 = Ena/VASP-homology 1; EVH2 = Ena/VASP-homology 2; GBD = GTP-binding domain; VPH = verprolin homology domain; CA = cofilin homology and acidic domain; KBD = kinase binding domain; SPR = sprouty-related domain; RanBD = ran binding domain; ZnF-RBZ = zinc finger domains; PPIase = proline isomerase; TPR (Pfam) = tetratricopeptide repeat; GRIP (Pfam) = golgin-97, RanBP2alpha, Imh1p, and P230/golgin-245; PTB = phosphotyrosine binding; PDZ = postsynaptic density/disc-large/ZO-1; SH2 = Src homology 2; SH3 = Src homology 3; PH = pleckstrin homology; RhoGAP = GTPase-

activator protein for Rho-like GTPases; DYNc = dynamin GTPase c; GED = dynamin GTPase effector domain. SwissProt accession codes for the specific examples shown in the figure are: VASP/Evl = P50552 (human); Ena/Mena = Q03173 (mouse); WASP = P42768 (human); N-WASP = O00401 (human); Homer1 = Q9Z216 (mouse); Homer2 = Q9UNT7 (human); Spred = Q7Z698 (human); nucleoporin 50 = O08587 (rat); RanBP2/Nup358 like = P49782 (human); RanBP2L/BS63 like = Q99666 (human); amyloid $ binding protein = P98084 (mouse); tensin = Q9UPS7 (human); Fe65 = O00213 (human); Rho GTPase-activating protein = Q7ZWQ2 (Xenopus); syntrophin = Q8BNW6 (mouse); dynamin-1 = Q05193 (human).

10 EVH1/WH1 Domains

oligomerization regions, which are also present in the EVH1 domain-containing proteins. Such regions are largely unstructured and, in the absence of their binding partners, are more prone to proteolysis and nonspecific aggregation, once released into the cytoplasm. In contrast, the domains that coexist in the host proteins of RanBDs, PTB, and PH domains are generally highly stable modules capable of independent folding. EVHI domains bind specifically to proline-rich motifs (PRMs) in peptides exposed on the surfaces of their binding partners. The N-terminal location of EVH1 domains is likely to facilitate their access to PRMs on bulky target molecules and may contribute to a segmental polarity of the EVH1 host protein that separates the adaptor module (EVH1) from its various effector domains. Certainly, given this strong preference for an N-terminal location, it is clear that EVH1 domains can occur only once in any given protein. The use of domain repeats, either in a cis configuration on the same polypeptide chain or in a trans configuration on identical chains in a quarternary structure, can achieve synergistic functions for the host protein, such as allowing the clustering of multiple binding partners or increasing the binding affinity for a single ligand. Since the Ena/VASP proteins tetramerize via their C-terminal EVH2 domains [59, 60], this brings together four EVH1 domains in each tetramer, thereby providing a mechanism for clustering of Ena/VASP proteins. Often, the target PRMs of EVH1 domains occur in close tandem repeats in the sequence of the binding partner. This could provide an additional mechanism for increasing binding affinity. Similarly, the Homer2 protein contains a leucine zipper motif at its C terminus, which is also involved in clustering [32, 66]. The EVH1 domains of each polypeptide chain in the oligomerized protein must therefore be located far away from the C-terminal oligomerization regions in the final structure, to allow unhindered access to their target peptides. This is clearly easier to achieve when these domains are well separated in the sequence. In summary, a conserved, unique N-terminal location appears to be a characteristic feature of EVH1 domains, setting them clearly apart from structurally related domains such as RanBDs, PTB, and PH domains. The EVH1 domains may confer a segmental polarity to their hosts that is required for functional or biogenetic reasons, resulting in the topological separation of this exposed terminal adaptor domain from the different types of genetically fused effector domains with which they coexist. 2.3 Classification of EVH1 Domains

To date, there is considerable discrepancy in the nomenclature and classification of EVH1 domains in the various databases. The name ‘EVH1 domain’ is frequently used interchangeably with the name ‘WH1 domain’, although the EVH1 nomenclature is generally the most widely used in the fields of biochemistry and molecular cell biology, owing to the early functional connotations resulting from the


identification of the first ligand [67]. Further confusion arises when different databases (for example, SMART, Pfam, InterPro [68]) refer to only one of these different names and classify them in conflicting schemes. Here, we refer to EVH1/WH1 domains as EVH1 domains and classify them based on their sequence conservation, domain co-occurrence, structural similarities, and ligand binding preferences, where this information is available. From the sequence alignment and phylogenetic analysis (Figures 1 and 2), it is clear that the EVH1 domains cluster into four main groups, which we have named after their primary host proteins: the Ena/VASP class, the Homer/Vesl class, the WASP/N-WASP class, and the Spred class. The Ena/VASP and Homer/Vesl classes have already been referred to as Class 1 and Class 2 EVH1 domains, based on their selectivities for peptides containing either FPPPP or PPxxF motifs, respectively [69]. We suggest keeping this nomenclature and adding the more distantly related WASP/N-WASP EVH1 domains (Figure 2), which bind a LPPPEPY-containing peptide in WIP, as Class 3, and the more recently described family of Spred EVH1 domains, which contain distinct putative ligand binding residues but for which no binding partner is yet known, as Class 4. Several EVH1 domains fall between these main classes, such as those from the Drosophila Still-life type 1 protein [70], the C. elegans hypothetical YKC2 protein, and the Dictyostelium RasGEFS. These could either provide putative evolutionary links between the different classes already known or be singular representatives of new, so-far-undiscovered families. Further detailed information on their structures, functions, and ligandbinding characteristics is needed before solid conclusions can be drawn about these proteins. The RanBDs cluster to form a large class of their own, which includes domains from the Ran-binding proteins and nucleoporins, as well as the HBA1 protein from the yeast Schizosaccharomyces pombe. Both the alignment in Figure 1 and the tree in Figure 2 show clearly that the RanBDs are most closely related to the EVH1 domains of the Homer/Vesl family, as well as to the RasGEFS and Drosophila Stilllife type 1 protein. However, despite this relationship, it would be inaccurate to include RanBDs as a subclass of EVH1 domains based on the criteria used here. It is possible that the RanBDs and EVH1 domains may instead stem from a common ancestor, which at some distant time may have descended from the same origin as the other PH and PH-like domains. The specific binding of the Homer EVH1 domain to C-terminal sequences has previously led to the suggestion that this EVH1 domain may be a divergent PDZ domain [31, 66]. Inspection of Homer EVH1 and PDZ domain sequences, however, reveals almost no similarity apart from a common four-amino-acid ‘typical’ motif constituted by turn-promoting residues, an invariant Phe, and a hydrophobic residue (GLGF in Homer). This motif is involved in C-terminal COOH recognition in PDZ domains [71, 72]. However, the different locations of this motif in the PDZ and Homer EVH1 structures rule out any functional relationship and make it unlikely that there is a meaningful evolutionary link between the two types of domain [53].

12 EVH1/WH1 Domains

3 Structures of EVH1 Domains and Their Complexes 3.1 High-resolution Structures of EVH1 and Related Domains

High resolution 3D structures of seven EVH1 domains representing three of the four classes of EVH1 domain, from the Ena/VASP, WASP/N-WASP, and Homer/Vesl families, have now been solved, several in complex with their target proline-rich ligand [33, 36, 73–77]. Structures of the RanBDs from both the large RanBP2 and the smaller, nucleoporin-like RanBP1 proteins are also known: the former in complex with Ran-GTP [54] and the latter in a ternary complex with Ran-GTP bound to RanGAP [78]. The structures of 5 PTB and 16 PH domains from a wide range of different host proteins are also available (for reviews see [79, 80]). Superposition of the Cα traces of representatives from each of the three classes of EVH1 domain for which structures are now known and the related RanBDs, PTB domains, and PH domains with the Class 1 EVH1 domain from murine Ena (Mena) are shown in Figure 4. The overall folds are essentially the same, forming a compact, parallel β sandwich capped along one side by a long α helix. Mutagenesis studies have highlighted several specific core residues as being important for stabilization of the EVH1 fold [81]. The main differences between the different families occur, not surprisingly, in the loop regions, where sequence variability is also highest (Figure 1). The structures of the different classes of EVH1 domains show very little difference (mean backbone rmsd values of VASP, Homer, and WASP EVH1 domains relative to the EVH1 domain of Mena are 1.39, 2.82, and 2.0 Å, respectively). The similarity to the RanBD fold is quite remarkable (rmsd 1.6 Å). Despite low sequence identity (∼10%), the PTB and PH domains show high structural homology to both EVH1 domains and RanBDs. The agreement is closest in the most highly structured regions (average rmsd values relative to Mena EVH1 are 2.7 and 2.4 Å, respectively) but much less in the loop regions, where long insertions and deletions occur. Additional elements of secondary structure are also present in several members of these more distantly related families (for example, the PTB domain of the human SHC protein shown in the figure). A distinguishing feature of EVH1 domains is the highly conserved triad of surface-exposed aromatic sidechains, Y16, W23, and F77 (outlined by boxes in the alignments in Figure 1; numbering relative to Mena). These come together in the 3D structure to form an aromatic cluster (shown in black in Figure 4), which provides a hydrophobic docking site for the proline-rich peptide ligands targeted by EVH1 domains. From the degree of conservation alone, it is clear that W23 is a highly important residue. It is completely conserved in all families of EVH1 domains and in almost all of the RanBDs. From inspection of the 3D structures, one can see that a large area of this sidechain makes important hydrophobic contacts with the core, thus involving it in fold stabilization. The mutation W23L in human VASP results in an insoluble, aggregated protein [75], in agreement with similar inactivating mutations analyzed in a yeast two-hybrid system [82]. Nevertheless,

3 Structures of EVH1 Domains and Their Complexes 13

Fig. 4 Comparison of 3D structures. Superpositions of the backbone (N, C", and C ) atoms of the EVH1 domain of Mena with representative members of each of the related groups shown in the phylogenetic tree. The Mena EVH1 domain (blue) is overlaid with (a) the Class 1 EVH1 domain from VASP (red); (b) the Class 2 Homer EVH1 domain (green); (c) the Class 3 N-WASP EVH1 domain (orange); (d) the first RanBD (RanBD1) of Nup358 (magenta); (e) the

PTB domain of Numb (yellow); and (f) the PH domain of DAPP1/PHISH (cyan). Structures were aligned by using DALI [7]. The locations of the exposed aromatic clusters are shown in black. PDB (and SwissProt) accession codes are Mena. EVH1 = 1evh (Q03173); VASP EVH1 = 1egx (P50552); Homer EVH1 = 1ddv (O88800); N-WASP EVH1 = 1mke (O08816); RanBD = 1rrp (P49792); PTB = 2nmb (P16554); PH = 1fao (Q9UHF2).

the indole proton of W23 (H1) is oriented toward the surface and is available as a H-bond-donating group in ligand interactions. Thus, in EVH1 domains and RanBDs, W23 fulfils a crucial role in domain stabilization while simultaneously exposing one functional group for ligand recognition. The other two residues of the triad are less strictly conserved. Variation in these allow corresponding variations in the geometry and properties of the binding site, enabling the different families of EVH1 domains to bind specifically to distinct consensus ligands. The conservation in fold and of important functional residues relates the above domains very closely. Furthermore, as this fold has been found only in signaling proteins and only in eukaryotes, this suggests that EVH1, Ran-binding, and PTB domains all comprise subfamilies of the PH superfamily, which may have arisen from a common distant ancestor [65]. Over time, these subfamilies have diverged, but have retained the stable PH fold as a scaffold upon which to build distinct

14 EVH1/WH1 Domains

specialized ligand-recognition sites, leading to the rich degree of functional diversity observed today [65]. 3.2 Structures of EVH1 Complexes and Determinants of Ligand Specificity

The EVH1 domains of Ena/VASP and Homer/Vesl families bind peptides that are 6–12 amino acids long and contain proline-rich motifs (PRMs) 4–6 amino acids long. The binding affinities of the core motifs in isolation are extremely low (K d values in the millimolar range), but are increased to biologically significant levels by the presence of additional core-flanking epitopes, which make additional contacts with the domain surface. The Ena/VASP EVH1 domains bind specifically to FPPPP motifs found in focal adhesion-associated proteins like zyxin [56], vinculin [57], and the Listeria ActA [30], whereas the Homer/Vesl EVH1 domains bind specifically to PPxxF motifs from the group I mGluRs [31], IP3R receptors [61], ryanodine receptors, and Shank proteins [62, 83, 84]. In contrast, the N-WASP EVH1 domain binds a much longer proline-rich peptide, having a minimum length of 25 residues (residues 461–485) and does not bind a 10-residue ligand of the Mena EVH1 domain from ActA, which contains a PRM (DFPPPPT) very similar to that found in the WIP peptide (DLPPPEP) [67]. It should be noted that the peptide-N-WASP complex was expressed from a single construct in which the 25-residue WIP peptide was fused by a 5-amino-acid linker to the N terminus of N-WASP [36]. This clearly favors binding for entropic reasons. It is not known whether binding occurs to an equivalent independent peptide, because isolated N-WASP EVH1 domains were found by these authors to be insoluble. The structures of representative complexes from each of the three families of EVH1 domains for which structures are now available are shown in Figure 5 [36, 73, 76]. No structure is yet available for the Spred EVH1 domains. The overall construction of the ligand-recognition sites is generally similar in all classes of EVH1 domains. The exposed Trp sidechain (W23; Mena numbering) is usually located at the centre of the aromatic triad (Figures 4 to 6) and is oriented in a plane almost perpendicular to the domain surface. On one or both sides, at approximately 90◦ to this plane and almost parallel to the domain surface, lie the flat rings of either Tyr or Phe sidechains. This perpendicular arrangement of aromatic rings results in rectangular hydrophobic pockets on each side of the Trp, well suited to the recognition of peptide ligands that adopt structures close to that of the left-handed PPII helix structure, characterized by backbone angles = −78◦ and = +146◦ [85–87]. The FPPPP-containing peptides bound by the Ena/VASP EVH1 domains and the LPPPEP region of the WIP peptide bound by the N-WASP EVH1 domain are good examples of this. The indole proton of the central Trp forms a hydrogen bond to a backbone carbonyl oxygen in the peptide, which anchors the ligand into place. The sidechains of the peptide residues surrounding this carbonyl (usually prolines) then pack closely into the rectangular hydrophobic pockets on either side of the W23 sidechain (Figure 6). In the Ena/VASP EVH1 domains, Y16 and F77 (Mena numbering) comprise the Trp-flanking aromatic residues, and the peptide


Fig. 5 Structures of complexes of EVH1 and related domains with their respective ligands. (a) Mena EVH1 domain with FPPPP peptide. (b) Homer EVH1 domain with the TPPSPF peptide. (c) N-WASP EVH1 domain with the minimal 25-residue peptide from WIP fused to its N terminus. Only the N-terminal 11 amino acids of this peptide (DLPPPEPYNQT) that bind in the ‘PRM-binding groove’ are shown. (d) The first RanBD (RanBD1) from Nup358 with the Ran protein. Only the C-terminal 10 amino acids (EVAQTTALPD) of Ran relevant to this discussion are shown. (e) The

PTB domain of the SHC protein with the 12-residue peptide HIIENPQpoYFSDA phosphorylated at tyrosine. (f) The PH domain from DAPP1/PHISH with inositol-1,3,4,5tetrakisphosphate. All classes of EVH1 domains and the RanBD share similar exposed clusters of aromatic sidechains in their peptide binding grooves, as described in the text. Numbers with asterisks refer to the equivalent positions in the sequence of Mena. PDB accession codes are as in Figure 4, except for the PTB domain complex (PDB: 1shc; SwissProt: P29353).

P(2) and P(5) of the Class 1 EVH1-binding motif FPPPP binds to either side of W23 (underlined residues are those whose sidechains make the closest contacts with the domain; yellow in Figure 6). The indole proton of W23 makes a hydrogen bond to the carbonyl of P(3). An additional hydrogen bond between the domain’s Q79 sidechain and the ligand P(2) supports the interaction. In each class of EVH1 domain, an H-bond–donating residue is almost always located at this position (Figure 1). The N-terminal F(1) of this sequence makes a further close hydrophobic contact with the domain, which is important for anchoring the peptide and for determining the orientation of the otherwise highly symmetric ligand (Figure 6). Variations in the geometries of the PRM binding sites give rise to the observed differences in ligand preference between the different domain families. The complex of the Homer EVH1 domain bound to the Class 2 EVH1-binding mo-

16 EVH1/WH1 Domains

Fig. 6 Comparison of peptide binding interfaces for the different classes of EVH1 domains (a-c) and RanBDs (d). Hydrogen bonds in the interface are shown by dotted lines where distances are less than 2.6 Å. The conserved W23 (Mena numbering) is colored pink. In all EVH1 domains the conserved W23 forms an important hydrogen bond via its indole proton to a carbonyl oxygen in the peptide backbone. The second conserved H-bond-donating residue Q79 (Mena) is shown in green. The location

of this residue is conserved in Ena/VASP and Homer/Vesl EVH1 domains, but differs in the EVH1 domains of N-WASP. (Asterisks indicate numbering when aligned to Mena). Ligand sidechains that make close hydrophobic contacts with the domain are colored yellow. The peptide sequence is shown above each complex. Hbond-acceptor residues are underlined and colored as follows: red = hydrogen bond to Trp; green = hydrogen bond to Gln (EVH1) or Arg (RanBD).

tif TPPxxF shows a very different binding mode from that of Ena/VASP EVH1 domains bound to their FPPPP motifs (Figure 6). In the Homer/Vesl proteins, I16 replaces Y16 (Mena), thereby altering the shape of the binding pocket that accommodates the sidechain of P(2) in the Ena/VASP EVH1 complexes [69]. Simultaneously, the hydrophobic residue M14 of Ena/VASP domain is replaced by the aromatic F14 in Homer/Vesl (Figures 1 and 5) providing a new binding pocket for the sidechain of P(3) in the Homer peptide. Hence, the loss of an aromatic sidechain at position 16 and the gain of a new aromatic sidechain at position 14 change the location and geometry of the PRM-recognition triad. The two consecutive prolines and terminal Phe of the TPPxxF motif make the closest hy-


drophobic contacts to the Homer EVH1 domain surface, with the exposed indole proton of Homer W24 forming a hydrogen bond to the carbonyl oxygen of the N-terminal peptide Thr (Figure 6). The complex of the N-WASP EVH1 domain with the N-terminal 11 amino acids of the 25-residue WIP minimal binding sequence 1 DLPPPEPYNQT11 is shown in Figures 5 and 6. The remainder of the peptide wraps around the back of the EVH1 domain [36] and is omitted from the figures for clarity. The PRM-binding triad of N-WASP is most closely related to that of Homer (also apparent from the sequence alignment in Figure 1). As in Homer, the conserved Y16 residue of the Ena/VASP EVH1 domains is lost (replaced by A48; equivalent to I16 in Homer) and thus no longer forms part of the peptide binding site. Instead, residues Y46 (14*), W54 (23*), and F104 (77*) in the N-WASP EVH1 domain are equivalent to F14, W24, and F74 in Homer (numbers with asterisks refer to the equivalent positions in Mena), forming almost identical aromatic clusters in the two proteins. However, the WIP peptide does not bind N-WASP in the same manner as observed for the Homer complex. Instead of making close contacts with Y46 (14*), the WIP peptide contacts a groove comprising W54 (23*), F104 (77*), T106 (79*), and Q113 (86*) on the N-WASP surface. The peptide P(3) and P(5) flank the exposed W54 sidechain on either side of the hydrogen bond between this residue and the carbonyl of P(3). An additional hydrogen bond from Q113 (86*) to the peptide E(6) helps to anchor the ligand into position (Figure 6). The most striking difference between the N-WASPWIP complex and the Ena/VASP and Homer/Vesl EVH1-peptide complexes is that the WIP peptide binds in the reverse orientation to that observed in the other complexes. One reason for this may be the Y(8) sidechain of the WIP peptide, which makes several close contacts with the domain surface, just as the F(1) of the FPPPP motif contacts the Class 1 EVH1 domains. Y(8) is also oriented so that a hydrogen bond may be possible from its hydroxyl proton to the backbone carbonyl of N-WASP G109 (82*). It is therefore likely that Y(8) plays a role in orienting the WIP ligand, and it will be interesting to find out whether other WASP ligands exist in which this pattern is conserved. Binding studies to date have shown that the Spred EVH1 domain does not bind the FPPPP motif recognized by the Ena/VASP EVH1 domains (Zimmermann et al., personal communication). The replacement of the conserved Class 1 EVH1 domain Y16 with an Arg in Spred is almost certainly an important factor in determining the ligand preference of this family (Figures 1, 4 and 5). Work in progress will reveal more details on the binding mode of this class of EVH1 domains in the future. All the substitutions seen in the exposed PRM-recognition sites are very conservative and have no noticeable effect on the overall stability of the fold. Nevertheless, they are critically placed and provide sufficient external variation to tailor the different families of EVH1 domains to recognize highly specific consensus target sequences. This allows EVH1 domains to be used as molecular adaptors by diverse families of host proteins to mediate their localization to very different signaling proteins. Interestingly, the PRM-binding interfaces of the EVH1 domains described above and in Figure 6 have many features in common with those of other protein

18 EVH1/WH1 Domains

interaction domains that bind specifically to proline-rich sequences. Examples include the well known families of the SH3, WW, GYF, and UEV domains, as well as the small, actin-binding, profilin protein. This mechanism for proline-rich peptide recognition is therefore not specific to EVH1 domains, but is rather widely used in many different signaling pathways (Ball et al., 2004).

3.3 Comparisons with RanBDs, PTB Domains, and PH Domains

There are four sites of contact between Ran and the RanBD. One of these involves a groove on the RanBD surface, which binds the C-terminal fragment of Ran. The interaction surface is analogous to the peptide interaction surface of the EVH1 domains. Figure 5 shows the first RanBD (RanBD1) of the nuclear pore complex protein Nup358 in complex with the Ran protein [54]. For clarity, only the 11 Cterminal amino acids 1 LEVAQTTALPD11 of Ran, which bind the groove relevant to this discussion, are shown in the figure. The RanBDs are the closest relatives to the EVH1 domains (as shown by the phylogenetic tree in Figure 2) and employ very similar mechanisms of ligand recognition. Both families of domains utilize clusters of exposed aromatic sidechains to recognize their peptide ligands. The completely conserved W23 of the EVH1 domain (Mena numbering) is also conserved in the majority of RanBDs, where it is also necessary for fold stabilization. As in the EVH1 domains, this important residue is flanked in RanBDs by two additional aromatics to form hydrophobic binding pockets that accommodate specific peptide sidechains (yellow in Figure 6). The interaction is supported by one or more additional H-bond-donating residues exposed at the ligand binding interface. The W1211 (W23*) sidechain of the Nup358 RanBD1 is positioned so that it can form a hydrogen bond with the backbone carbonyl of the peptide T(7) in the same way as observed for the EVH1 domains, but this distance in the NMR structure of the N-WASP–WIP complex is longer than usually expected for hydrogen bonds and the geometry of the interaction is not ideal. The interaction of the ligand with the domain is stabilized by an additional hydrogen bond between the domain R1284 (R90*) sidechain and the ligand P(10). The Ran protein binds RanBD in the same orientation as the Ena/VASP and Homer EVH1 domains. However, since the peptide fragment shown here is only a small fragment of a much larger ligand, there are likely to be other factors that also contribute to its binding orientation and affinity, which shall not be discussed here. For comparison, the PTB domain of the human SHC protein with a phosphotyrosine peptide from a TRKA receptor [88] and the PH domain of DAPP1/PHISH with inositol 1,3,4,5-tetrakisphosphate [89] are also shown in Figure 6. Although their folds are visibly very similar, it is clear that the binding sites of these domains have little if anything in common with those of the EVH1 domains or RanBDs. Thus, the aromatic peptide recognition clusters clearly evolved long after the above domains had diverged from the PH family.

4 Biological Function and Signaling Pathways Involving EVH1 Domains 19

4 Biological Function and Signaling Pathways Involving EVH1 Domains

The ability of proteins to target their binding partners in a highly specific manner is the basis for the assembly of multiprotein complexes required for signaling in all living cells. EVH1 domains mediate protein–protein interactions in a diverse range of signaling cascades, depending on their host protein and site of action. 4.1 Ena/VASP Interactions

The Ena/VASP proteins are involved in the generation and maintenance of cell polarization processes by localized actin polymerization and reorganization in a variety of specialized cytoskeletal substructures [14, 16]. During epithelial sheet formation, they bind FPPPP motifs within zyxin and vinculin via their EVH1 domains. This localizes the Ena/VASP proteins to premature epithelial contact sites called adhesion zippers, which are involved in sealing epithelial layers [29, 90]. Ena/VASP proteins are also involved in barrier formation in the endothelium [91, 92] and in regulating integrin-mediated platelet adhesion [22, 23]. Interactions between activated T-cells and antigen-presenting cells lead to the formation of a contact structure called the immunological synapse. Ena/VASP proteins are recruited by FPPPP motifs within the EVH1-binding protein Fyb/SLAP to a signaling complex which, together with WASP, supports localized actin-filament polymerization at these synapses [28]. Additionally, phagocytosis by macrophages involves assembly of phagocytic cups, to which Ena/VASP proteins are again recruited by the Fyb/Slap protein. This leads to the Fc cell-surface-receptor-mediated remodeling of the actin cytoskeleton that accompanies particle internalization [26]. The Ena/VASP proteins also have important roles in migration of neurons, neutrophils, and fibroblasts [14, 16]. During the establishment of synaptic contacts in developing nervous systems, growth cones of axons are navigated by receptormediated recognition of guidance proteins [93]. FPxxP binding sites for Class 1 EVH1 domains are found in the intracellular portion of the Drosophila and C. elegans axon guidance receptor Robo/Sax-3 [20, 94]. In the human and murine transmembrane semaphorin Sema6A-1 guidance protein [95], the corresponding EVH1 binding motifs are VPPKP. Furthermore, the C. elegans Ena/VASP ortholog Unc-34 is required for appropriate response of the axon guidance receptors Unc-5 and Unc-40/DCC to the guidance protein netrin [94, 96]. The locomotion of fibroblasts is a complex process requiring coordination of forward protrusion, attachment, contraction, and rear detachment, which translocates the cell body. Ena/VASP proteins were found to be negative regulators of cell migration [25]. At the subcellular level, they enhance F-actin polymerization at the leading edge of highly dynamic, transiently formed lamellipodia, to which they are localized by EVH1-mediated interactions [97]. Molecular understanding of these processes has greatly benefited from studies of the intracellular pathogenic bacterium Listeria monocytogenes [98, 99]. The EVH1 domains of the Ena/VASP

20 EVH1/WH1 Domains

proteins are recruited to the bacterial cell surface by tandem FPPPP motifs in the Listeria surface protein ActA [67]. There, they are involved in actin polymerization, resulting in the assembly of dynamic ‘comet’ tails, which generate the motile force necessary for bacterial propulsion through the cytoplasm [98, 99]. In summary, Ena/VASP proteins form a key link between signaling pathways and cytoskeletal dynamics by acting as regulators of actin filament assembly. These proteins are components of the actin polymerization machinery and are believed to function mechanistically by delivery of polymerization-competent actin monomers, exclusion of polymerization-terminating proteins, detachment of F-actin filaments, and/or inhibition of ‘Y’ branch formation in actin filament arrays, depending on the cellular context in which they are found [13, 100]. 4.2 Homer/Vesl Interactions

Homer/Vesl proteins are adaptor proteins involved in clustering, anchoring, and modulation of neurotransmitter receptor proteins in excitatory glutamatergic synapses of the central nervous system [84], which are among the most elaborate junctions existing between cells. The apical postsynaptic plasma membrane region of dendritic spines differentiates into an electrondense subcellular compartment called the postsynaptic density (PSD). The PSD contains, together with various multidomain PSD scaffold proteins, the NMDA (N-methyl-D-aspartate) and metabotropic glutamate receptors (mGluRs) [34]. The Class 2 EVH1 domains of Homer/Vesl proteins bind a PPxxF motif in the multidomain scaffold protein Shank/ProSAP [83], which is engaged in further interaction with NMDA receptors [62]. They also bind the Class 2 EVH1-binding motif (TPPSPF) in the C termini of group 1 mGluRs [31]. Thus, the clustering of Homer/Vesl proteins [59, 60] provides a mechanism for indirectly linking two glutamate receptor types. This may be relevant to synaptogenesis or to specific types of receptor crosstalk. The Homer/Vesl EVH1 domains are also known to bind the PPKKF motif within inositol trisphosphate receptors (IPRs) and the C termini of group 1 mGluRs [61]. Here, they are involved in the glutamate-induced release of Ca2+ from intracellular stores. The monomeric Homer1 isoform, which lacks a self-association domain [32, 66], is upregulated by neural activity [35]. This protein is expected to competitively disrupt the signaling complexes assembled by the oligomeric Homer/Vesl isoforms [61]. The Homer/Vesl proteins thus regulate the composition and stability of multiprotein complexes comprising different synaptic receptor proteins. This implies a role in the modulation of synaptic plasticity, which is clearly important with respect to learning and memory formation. 4.3 WASP/N-WASP Interactions

The founding member of the WASP/N-WASP proteins, the Wiskott-Aldrich syndrome (WAS) protein, was originally discovered during a search for the genetic

4 Biological Function and Signaling Pathways Involving EVH1 Domains 21

defect responsible for WAS, a rare, X-linked, recessive immunodeficiency disease with altered functions of many types of hematopoetic cells [101]. Among other roles, WASP is expected to be involved in biogenesis of the immunological synapse and in the early stages of T-cell cytoskeletal polarization (see above) [101]. The protein–protein interactions involved in these processes are not yet well understood. Understanding of the molecular function of WASP has been gained from analysis of cellular model systems [101], which identified WASP/N-WASP proteins as highly regulated effectors of Rho GTPases. Following release from an autoinhibitory state by Rho GTPases, WASP activates the actin-nucleating Arp2/3 complex via its C-terminal VCA effector domain. The EVH1 domain of N-WASP binds a consensus LPPPEPY motif in the C-terminal region of WIP and its homologs, CR16 and verprolin [36], to regulate N-WASP-mediated actin polymerization and filopodium formation. Various viral and bacterial pathogens have evolved strategies to hijack the actin-nucleating activity of Arp2/3, using pathogen-encoded virulence factors that are direct or indirect upstream activators of N-WASP [98]. Whereas the IcsA protein of Shigella flexneri directly mimics the host cell Rho GTPase Cdc42, to activate N-WASP [102], the viral membrane protein A36R of vaccinia virus [103] and the bacterial protein Tir of extracellular enteropathic Escherichia coli (EPEC) [104, 105] both recruit the host-encoded adaptor protein Nck, together with WIP and N-WASP, to initiate localized actin polymerization, which then supports either intracellular motility (Shigella, vaccinia) or contact formation to the host cell (EPEC). Pathogen-triggered binding of the EVH1 domain of N-WASP to PRMs of host proteins therefore functionally resembles the interaction of Ena/VASP EVH1 domains with the PRMs of the Listeria monocytogenes virulence factor ActA, discussed above. However, unlike the other virulence factors, ActA is able to activate the Arp2/3 complex directly.

4.4 Spred Interactions

Spred proteins were originally isolated in a screen for new tyrosine-kinase-binding proteins. The proteins are involved in regulation of differentiation in neuronal cells and myocytes [48]. The proteins were recently described as negative regulators of the signaling pathways of Ras, Raf, and mitogen-activated protein (MAP) kinase [47, 48]. Mechanistically, Spred inhibits early steps in the activation of MAP kinases by association with Ras, thereby suppressing phosphorylation and activation of Raf. This results in down-regulation of the Ras–MAP kinase signaling pathway [48]. The specific binding partner of the Spred EVH1 domain is currently unknown, although Spreds’ inhibitory activity is lost when their EVH1 domain is substituted for a Class 3 EVH1 domain of WASP, indicating a highly specific interaction. To date, only one other Spred family member, the Drosophila AE33 [49], has been identified. It is known to be involved in photoreceptor cell specification during Drosophila eye development. However, its role in these developmental pathways has not yet been elucidated.

22 EVH1/WH1 Domains


One of the most important reasons for studying sequence similarities and obtaining detailed high-resolution structures of proteins and their complexes is that comparisons may then be made with proteins for which little functional data are available. This information also facilitates the annotation and classification of hypothetical proteins revealed by genome sequencing projects and provides a rational basis for the prediction of protein structures and the identification of candidate ligands for novel proteins. 5.1 Use of Sequence and Structural Data in Prediction of Binding Partners

Comparisons of sequence and fold reveal important relationships between different domains and provide insights into their evolutionary origins. Since the overall fold of a domain is determined by a small number of key residues in the hydrophobic core, structural alignments performed with algorithms such as DALI [106] and FSSP [107, 108] can provide important clues about function and ancestry even where sequence identity is very low. However, a fold similarity on its own is insufficient for the prediction of binding partners, as demonstrated in detail for the EVH1 domains discussed in this chapter. For this, close inspection of the conserved residues exposed on the surface of the domain can be very revealing. Often, groups of surface-exposed residues, although separated by many amino acids in the primary sequence, come together in the 3D structure to form a ligand binding site. These residues are usually conserved within a given domain family, and sometimes across many families that share common mechanisms of ligand recognition. Good examples are the PRM-binding domains, which include the SH3, WW, EVH1, GYF, and UEV domains and the single-domain actin-binding protein profilin. All six families of PRM-binding domains contain exposed groups of aromatic residues termed ‘aromatic clusters’ or ‘aromatic cradles’, which are responsible for binding proline-rich motifs [53, 109]. The presence of this cluster is now considered a signature for PRM recognition. Thus, if the structure of a new domain is solved, detailed comparisons with similar structures for which the function is better understood can narrow the search for binding partners significantly. A thorough understanding of the common features that define different types of domain–ligand interfaces is ultimately needed to enable us to derive rules for the prediction of binding partners from sequence information alone. 5.2 Use of Structural Data from Complexes to Guide the Rational Design of New Ligands

Knowledge of the molecular determinants of binding affinity and specificity is necessary for rational structure-based design of inhibitors to hinder interactions between specific proteins and to modulate cellular processes. To understand the

6 Conclusions 23

subtle factors that modify specificity and affinity, it is necessary to characterize, structurally and biochemically, the most important features of the interactions we wish to inhibit. Only then can this knowledge be used to guide the design of novel target peptides or nonpeptide molecules having new activities. SPOT analyses [110] have helped enormously in understanding which residues of a known binding sequence are critical for binding and which may be replaced with little or no consequence. This allows the derivation of consensus ligand sequences, as demonstrated for the FPPPP motifs recognized by Class 1 EVH1 domains. Here, the consensus peptide sequence was found to be FPxP (where is a hydrophobic residue and x is any residue), meaning that the central two residues can be replaced in searches for higher-affinity partners [69, 75]. Knowledge of variable and conserved residues is a crucial first step toward the prediction, modification, and design of peptide binding partners, as successfully demonstrated experimentally for EVH1 domains [111]. The design of inhibitors for EVH1 domains is currently under way. Such inhibitors should be useful in several different ways: (1) to study the effects of modulating EVH1-mediated signaling cascades; (2) as molecular tags to monitor the formation and dissociation of EVH1-mediated interactions within the cell; and (3) as lead molecules for the development of future generations of novel therapeutics. The dose-dependent modulation of EVH1 domain binding activity would mean that it should be possible to develop treatments of diseases for which partial inhibition of EVH1-mediated events would be desirable (for example, pathologically altered adhesion and motility in inflammatory diseases and metastatic states, or even the spreading of intracellular pathogens).

6 Conclusions

In this chapter we have taken a close look at the sequence signatures, domain occurrence/co-occurrence, and structural features that define the EVH1 domain. Subtle differences in the patterns and nature of exposed residues have been shown to lead to different ligand specificities. Combining sequence similarity, phylogeny, structural information, and ligand preference, we have classified the known EVH1 domains into four main categories. The stability of any domain fold is clearly an important factor in its evolution, and EVH1 domains are no exception. This is demonstrated by the high conservation of residues that comprise the hydrophobic core. The specific functionality then built upon the stable scaffold depends on subtle variations in the solvent-exposed residues, particularly those located within and around ligand binding sites. These can alter specificities dramatically, depending on their charge, hydrophobicity, size, and H-bond forming abilities. More distant relatives show greater differences in their patterns of surface residues, leading to widely different functions although they utilize a common global fold. Even seemingly minor variations at critical sequence locations can alter ligand binding specificities considerably. This provides an economical evolutionary mechanism for

24 EVH1/WH1 Domains

gradual functional diversification starting from a stable fold, as has been observed repeatedly during studies of the protein universe [112, 113]. Detailed studies of these variations provide clues about the domain’s ancestry and phylogeny. Furthermore, knowing which residues can be varied without destabilizing the domain fold, and which contextual combinations of domain folds are tolerated in different host protein repertoires, is a crucial first step in the understanding of protein evolution and protein–protein interactions in general. Such knowledge is essential for the rational design of inhibitors, interaction interfaces, and novel proteins in the future.

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