90 Structure and Bonding
Metal Sites in Protein and Models Redox Centres
Volume Editors: H. A. O. Hill, P. J. Sadler, A. J. Thomson
Springer
Berlin Heidelberg New York
The series Structure and Bonding publishes critical reviews on topics of research concerned with chemical structure and bonding. The scope of the series spans the entire Periodic Table. It focuses attention on new and developing areas of modern structural and theoretical chemistry such as nanostructures, molecular electronics, designed molecular solids, surfaces, metal clusters and supramolecular structures. Physical and spectroscopic techniques used to determine, examine and model structures fall within the purview of Structure and Bonding to the extent that the focus is on the scientific results obtained and not on specialist information concerning the techniques themselves. Issues associated with the development of bonding models and generalizations that illuminate the reactivity pathways and rates of chemical processes are also relevant. As a rule, contributions are specially commissioned. The editors and publishers will, however, always be pleased to receive suggestions and supplementary information. Papers are accepted for Structure and Bonding in English. In references Structure and Bonding is abbreviated Struct Bond and is cited as a journal.
Springer WWW home page: http://www.springeronline.com Visit the SB content at http://www.springerlink.com
ISSN 0081-5993 (Print) ISSN 1616-8550 (Online) ISBN-13 978-3-540-62888-0 DOI 10.1007/3-540-62888-6 Springer-Verlag Berlin Heidelberg 1998 Printed in Germany
Preface
Controlling the movement of electrons in time and space is vital to biology. How do proteins achieve this? In this volume (the third in a three-part special series on Metal Sites in Proteins and Models, volumes 88, 89 and 90) metals from all three transition metal series are highlighted: V, Mn, Fe, Ni and Cu from the first, Mo from the second and W from the third series. Without manganese and photosynthesis, and the conversion of water into dioxygen, there would not be abundant chemical energy! and Penner-Hahn critically reviews the evidence for Mn clusters and cofactors in the oxygenevolving complex of Photosystem II - does it contain one or two clusters.~ what are the Mn oxidation states? do we need to invoke the presence of additional metals.~ Copper, too, is sometimes found in clusters in proteins, for example the purple dicopper in cytochrome o oxidase, and trinuclear sites of some other copper oxidases; Messerschmidt shows how the structures of copper centres are related to their properties and to an overall classification of copper sites. He also discusses the first structure of a vanadium-containing protein, the enzyme chloroperoxidase, and its unexpected coordination geometry. (The wider chemistry and biochemistry of vanadium is reviewed in detail in Volume 89) An important role for molybdenum involves the catalysis of oxygen atom transfer reactions coupled to electron transfer between substrate and cofactors such as flavins, Fe/S centres and hemes. About 70 molybdenum oxotransferase enzymes are known, which are widely distributed amongst eukaryotes, prokaryotes, and archaea. They belong to the families: xanthine oxidase, sulfite oxidases and assimilatory nitrate reductases, and DMSO reductases. The first representative structure of a member of the xanthine oxidase family (aldehyde oxido-reductase from Desulfovibrio gigas) is described here by Romeo and Huber; it contains a catalytically essential Mo = S group, and a molybdopterin cofactor, both in close proximity to Fe/S centres, but curiously no direct Moprotein bonds. Hydrogenases (which catalyze the two-electron oxidation of H 2) are enzymes of enormous biotechnological interest, providing potential sources of clean energy, and being vital to a wide variety of bacteria. Frey describes the first 3D structure of a Ni-Fe hydrogenase with its unusual Fe site and three bound diatomic ligands, and a Ni site which can bind hydride. Capozzi, Ciurli and Luchinat analyze in detail how protein-induced modulations of metal-metal interactions allow the redox potentials of iron-sulfur centres to span a wide range of redox potentials, and elegantly demonstrate how individual iron
VIII
Preface
atoms in clusters can be distinguished. The remarkable versatility of iron sites is also addressed in Volume 88 of this series. Finally, Hagen and Arendsen argue that tungsten has a widespread catalytic role in nature. There is no doubt that is proving to be the case in certain bacteria, but will it also be true for eukaryotes.~ And if Mo and W have such roles, what about Cr? That topic will have to wait for a future volume! We hope you will enjoy reading these articles and find them as stimulating as we have. H. Allen O. Hill, Peter ]. Sadler and Andrew J. Thomson
Contents
Structural Characterization of the Mn Site in the Photosynthetic Oxygen-Evolving Complex ]. E. Penner-Hahn . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Metal Sites in Small Blue Copper Proteins, Blue Copper Oxidases and Vanadium-Containing Enzymes A. Messerschmidt . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
37
Structure and Function of the Xanthine-Oxidase Family of Molybdenum Enzymes M. ]. Romeo, R. Huber . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
69
Nickel-Iron Hydrogenases: Structural and Functional Properties M. Frey . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
97
Coordination Sphere Versus Protein Environment as Determinants of Electronic and Functional Properties of Iron-Sulfur Proteins F. Capozzi, S. Ciurli, C. Luchinat . . . . . . . . . . . . . . . . . . . . . . .
127
The Bio-Inorganic Chemistry of Tungsten W.R. Hagen, A. E Arendsen . . . . . . . . . . . . . . . . . . . . . . . . . .
161
Author Index Volumes 1 - 90 . . . . . . . . . . . . . . . . . . . . . . . . .
193
Contents of Volume 88 Metal Sites in Proteins and Models Iron Centres Volume Editors: H. A. O. Hill, P. ]. Sadler, A.]. T h o m s o n
Polyiron Oxides, Oxyhydroxides and Hydroxides as Models for Biomineralisation Processes A. K. Powell Heme: The Most Versatile Redox Centre in Biology.~ S. K. Chapman, S. Daff, A.W. Munro Rationalisation of Metal-Binding to Transferrin: Prediction of Metal-Protein Stability Constants H. Sun, M. C. Cox, H. Li, P. ]. Sadler Metal Centres of Bacterioferritins or Non-Heam-Iron-Containing Cytochromes b557 N. E. Le Brun, A. ]. Thomson, G. R. Moore Ribonucleotide Reductases - A Group of Enzymes with Different Metallosites and a Similar Reaction Mechanism B.-M. Sj6berg Protein Engineering of Cytochrome P450cam L.-L. Wong, A. C. G. Westlake, D. P. Nickerson
Contents of Volume 89 Metal Sites in Proteins and Models Phosphatases, LewisAcidsand Vanadium Volume Editors: H. A. O. Hill, P.J. Sadler, A. J. T h o m s o n
Advances in Zinc Enzyme Models by Small, Mononudear Zinc(II) Complexes E. Kimura, T. Koike, M. Shionoya Zinc Catalysis in Metalloproteases D. S. Auld Modeling the Biological Chemistry of Vanadium: Structural and Reactivity Studies Elucidating Biological Function C. Slebodnick, B. J. Hamstra, V. L. Pecoraro Vanadium Bromperoxidase and Functional Mimics A. Butler, A. H. Baldwin Metal Ions in the Mechanism of Enzyme Catalysed Phosphate Monoester Hydrolyses D. Gani, J. Wilkie The Dimetal Center in Purple Acid Phosphatases % Klabunde, B. Krebs
Structural Characterization of the Mn Site in the Photosynthetic Oxygen-Evolving Complex James E. P e n n e r - H a h n Department of Chemistry, 930 N. University Avenue, Ann Arbor, Michigan 48109-1055, USA
E-mail:
[email protected] The photosynthetic conversion of solar to chemical energy is based on light-driven charge separation in a chlorophyll-based pigment. In higher-plants, the electrons required for this process are extracted from H20, ultimately producing 02 as a waste by-product of photosynthesis. The photosynthetic oxidation of water takes place at the oxygen evolving complex (OEC) on the donor (lumenal) side of Photosystem II. The OEC contains four Mn ions, together with calcium and chloride as essential inorganic cofactors. The techniques which have proven most useful in characterizing the nature of the OEC are X-ray absorption spectroscopy and EPR. Recent results from both techniques are reviewed. Key Words: Photosynthesis; oxygen evolution; manganese; X-ray absorption spectroscopy (EXAFS, XANES); electron paramagnetic resonance (EPR)
1
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2
1.1 1.2 1.3 1.4
Importance of Photosynthesis ................... Reactions in P h o t o s y s t e m II . . . . . . . . . . . . . . . . . . . . The Kok Cycle . . . . . . . . . . . . . . . . . . . . . Basic Elements o f P h o t o s y s t e m II . . . . . . . . . . . . . . . . .
2 3 4 5
2
Physical M e t h o d s for C h a r a c t e r i z i n g t h e OEC . . . . . . . . . . .
5
2.1 2.2 2.3
X-Ray A b s o r p t i o n S p e c t r o s c o p y . . . . . . . . . . . . . . . . . . X-Ray A b s o r p t i o n Near Edge S t r u c t u r e . . . . . . . . . . . . . . E x t e n d e d X-Ray A b s o r p t i o n Fine S t r u c t u r e . . . . . . . . . . . .
6 7 8
3
Structural C h a r a c t e r i z a t i o n o f t h e OEC . . . . . . . . . . . . . .
9
3.1 3.1.1 3.1.2 3.1.3 3.1.4 3.1.5 3.2 3.3 3.3.1 3.3.2
Magnetic Properties . . . . . . . . . . . . . . . . . . . . . D i m e r I n t e r p r e t a t i o n s of the Multiline Signal . . . . . . . . . . . T e t r a m e r I n t e r p r e t a t i o n s o f the Multiline Signal . . . . . . . . . I n t e r p r e t a t i o n s o f the g ~ 4.1 Signal . . . . . . . . . . . . . . . . The $1 State EPR Signal . . . . . . . . . . . . . . . . . . . The "$3" State EPR Signals . . . . . . . . . . . . . . . . . . O x i d a t i o n State A s s i g n m e n t s . . . . . . . . . . . . . . . . . . . . Atomic Arrangements . . . . . . . . . . . . . . . . . . . . Mn-Nearest Neighbor Interactions ................ Mn-Mn Interactions . . . . . . . . . . . . . . . . . . . . .
......
. . .
. . . . . . . . . . . .
9 10 10 12 14 15 16 16 17 18
Structure and Bonding,Vo].90 © SpringerVerlag BerlinHeidelberg1998
2
James E. Penner-Hahn
3.3.3 3.4 3.4.1 3.4.2 3.4.3 3.4.4 3.4.5
Outer Shell Interactions . . . . . . . . . . . . . . . . . . . . . . S-State Dependence of the OEC . . . . . . . . . . . . . . . . . . TheS2 State . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The $3 State - XANES . . . . . . . . . . . . . . . . . . . . . . . The $3 State - other measures of the oxidation state The $3 State - EXAFS . . . . . . . . . . . . . . . . . . . . . . . The So State . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4
Chemical Perturbations of the OEC . . . . . . . . . . . . . . . .
27
4.1 4.2 4.3
Inhibitors of Oxygen Evolution . . . . . . . . . . . . . . . . . . Modifications at the Chloride Site . . . . . . . . . . . . . . . . . Reduced Derivatives of the OEC . . . . . . . . . . . . . . . . . .
27 28 28
5
Structural Possibilities . . . . . . . . . . . . . . . . . . . . . . .
30
5.1 5.2 5.3
One vs. Two Clusters . . . . . . . . . . . . . . . . . . . . . . . . Arrangements of Mn Ions . . . . . . . . . . . . . . . . . . . . . Mn oxidation States . . . . . . . . . . . . . . . . . . . . . . . .
31 32 32
Conclusions and Future Prospects References
.......
................
. . . . . . . . . . . . . . . . . . . . . . . . . . . . .
19 21 21 22 25 26 27
33 33
Symbols and Abbreviations OEC XAS EXAFS XANES EPR ESEEM ESE-ENDOR PSII
oxygen evolving complex X-ray absorption spectroscopy extended X-ray absorption fine structure X-ray absorption near edge structure electron paramagnetic resonance electron spin echo envelope modulation electron spin echo detected electron nuclear double resonance Photosystem II
1 Introduction 1.1
Importance of Photosynthesis The ukimate source of most of the chemical energy used in life is solar energy, which is captured and converted to a chemically useful form by photosynthetic organisms. The high-energy compounds that are produced in this process are typically reduced hydrocarbons, with the required reducing equivalents coming, in the case of plants and cyanobacteria, from water (Eq. 1). The waste product of this reaction is oxidized water, i.e. 02. Oxygenic photosynthesis produc-
StructuralCharacterizationof the Mn Site in the PhotosyntheticOxygen-EvolvingComplex
3
ed most of the oxygen in the atmosphere, and in the process has dramatically akered the earth's environment. 2 H20 + light --> 02 + 4 H + + 4 e-
(1)
This chapter describes recent progress in characterizing the photosynthetic oxygen evolving complex (OEC). There have been numerous recent reviews of various aspects of the OEC (see for example [1], [2], [3], and [4]). The purpose of the present review is not to provide a repetition of earlier reviews but rather to provide a critical assessment of the available structural data, particularly recent results, and to use this to assess possible models for the oxygen evolving complex. 1.2 Reactions in Photosystem II
Oxygenic photosynthesis involves reactions at two separate photosystems, with oxygen evolution taking place at Photosystem II (PSII). The net reaction catalyzed by PSII is the transfer of electrons from water to plastoquinone. A schematic illustration of the principal polypeptides involved in Photosystem II and the routes of electron flow through the complex are illustrated in Fig. 1 (for details see [2]). The photochemical reaction at the heart of Fig. 1 is the charge separation that takes place at a chlorophyll pigment known as P680. The acceptor side of PSII is believed to be similar to that in the reaction centers of purple non-sulfur bacteria [5], with the electron flowing through a pheophytin to the plastoquinone QA and then eventually to a second exchangeable plastoquinone Qb. A second charge separation event then reduces Qb to the hydroquinone
(28
"/
CP4;
1
zl''N 4H÷+
Fig. 1. Schematicillustration of the polypeptides of PhotosystemII. Arrows indicate the electron flowfrom (top half) and to (bottom half) P680followingexcitation
4
]ames E. Penner-Hahn
form, which exchanges with the oxidized plastoquinone pool. On the donor side of P680,tyrosine Yz reduces P~80,resetting the system for another charge separation. Yz" is, in turn, reduced with an electron from water, via the Mn in the OEC. From Fig. 1 it is apparent that a basic problem in oxygenic photosynthesis is how to couple the one-electron charge separation reactions of P680to the fourelectron oxidation of water. 1.3 The Kok Cycle
The broad outlines of the solution to this problem were first recognized with the demonstration by ]oliot and co-workers [6] that photosynthetic membranes which are illuminated with short (ca. 5 ps) but saturating flashes of light evolve oxygen maximally every fourth flash, starting with the third flash. This demonstrated that each reaction center acts independently to acquire four oxidizing equivalents in a sequential manner and led Kok and co-workers to propose [7] the model shown in Fig. 2. This model, which has guided nearly thirty years of research into photosynthesis, is based on four kinetically resolvable intermediates known as Sn states, where the subscript refers to the number of stored oxidizing equivalents. The dark stable state of the OEC is the S1 state. Three flashes oxidize this to the transient $4 state, which evolves 02, regenerating So and beginning a new period 4 oscillation. Although $4 is postulated, only So through $3 have been observed spectroscopically. After a number of flashes, the period 4 oscillations are gradually damped. This damping is generally attributed to a combination of double hits (advance of two rather than one S state) and misses (no productive photooxidation). Recently, Shinkarev and Wraight [8, 9] have noted that a complete discussion of charge separation, and in particular the miss fraction, must consider both the donor side (the OEC Sn states) and the acceptor side (the quinones). The involvement of both a non-exchangeable quinone (QA) and an exchangeable quinone (Qb) (see Fig. 1) means that there will be a period two oscillation on the acceptor side, with different probabilities of charge separation depending on whether the acceptor is oxidized or reduced. This two-fold oscillation is superimposed on the four-fold oscillation resulting from the donor side (S-state) cycling. The im-
[S4]
4H+~
t
+j
I
% Sl
;;rP68°'~--'/
} --" 2H20
",,.. s 2/
3 "~,,,.,..,.~..,,,~ S
Fig. 2.
Kok cycle for oxygenic photosynthesis. The S1 state is the dark-stable state. The S 4 state is unstable with respect to production of 02 and the SOstate. Solid lines show changes in the OEC, dashed lines indicate electron flow
StructuralCharacterizationof the Mn Sitein the PhotosyntheticOxygen-EvolvingComplex
5
portant consequence of the acceptor side cycling is that the probability of S state advance (i.e. the miss fraction) shows a pronounced 2-fold periodicity. When all of the kinetic and thermodynamic factors are considered [8], the apparent miss fraction, particularly on the $2-~ $3 and $3 ~ So transitions, can be quite large. 1.4 Basic elements of Photosystem II
Initial work on the OEC used thylakoid membranes. An important advance came with the development of the so-called BBY preparation [10, 11], which removed most of the contaminating polypeptides. The essential elements of PSII are illustrated in Fig. 1. Peptides D1 and D2 are homologous with the L and M subunits of bacterial reactions centers [2]. These bind the primary redox cofactors P680,QA, Qb, and Yz, together with a non-heme iron bound between QA and Qb and a second redox-active tyrosine YD. In addition, D1 and D2 probably provide some of the ligands to the Mn and/or Ca ions. The CP43 and CP47 intrinsic membrane polypeptides are part of the light-harvesting antennae, peptides a and fl bind cytochrome b559, and the 28 kDa peptide plays a role in Qb binding [2]. In addition to these intrinsic transmembrane polypeptides, there are three extrinsic polypeptides, with molecular masses of approximately 17, 24, and 33 kDa. The 33 kDa polypeptide helps to stabilize the Mn, while the two lighter extrinsic polypeptides enhance the binding of Ca 2+ and C1-.A more highly purified preparation of the OEC, known as the reaction center complex (RCC) [12], lacks the 17 and 24 kDa extrinsic polypeptides. This preparation requires added Ca 2+ and C1- for activity, but appears to be spectroscopically similar or identical to BBY preparations. An important advantage of RCC preparations is that they have an approximately 3-fold higher Mn concentration than BBY preparations, and consequently give spectra with a better signal/noise ratio. Although there have been other suggestions [13], it is generally accepted that the OEC contains four Mn ions [2]. Full activity also requires at least one Ca2+ and one C1- ion per reaction center. Among the outstanding questions regarding the OEC are the nuclearity of the Mn cluster; the geometry of the cluster; the ligands to the Mn and Ca ions; the role of the inorganic cofactors; and the oxidation state of the Mn. As discussed below, the answers to many of these questions remain in dispute, although in some cases a consensus is beginning to emerge. 2 Physical Methods for Characterizing the OEC
Characterization of the OEC is inherently difficult since the portions of interest (Mn, C1, Ca) are elements that are generally hard to study. Most of the techniques imaginable have been turned to OEC at one time or the other. Some are indirect (e. g. thermoluminescence), some are difficult to attribute directly to the Mn/Ca/C1 cluster (FTIR) and some are extremely insensitive
6
James E. Penner-Hahn
(UV-vis). When they have been used, many of these techniques have been (and in some cases continue to be) controversial, with very different conclusions drawn from similar data. In many cases, a generally accepted consensus of which results and which interpretations are reliable has not yet been established. This review focuses on the application of magnetic resonance and Xray absorption spectroscopy to characterization of the OEC. The attraction of these methods is that they are highly specific either to paramagnetic centers or to a specific element (e.g. Mn). Most of the recent structural models that have been proposed for the OEC have been based on results from magnetic resonance and X-ray absorption. Before reviewing these results it is worth reviewing the limitations of these techniques. For magnetic resonance, the limitations are straightforward - it is only possible to obtain information about paramagnetic centers, and, perhaps more importantly, not all paramagnetic centers give rise to detectable magnetic resonance signals. With X-ray absorption, the limitations, most of which are related to the fact that X-ray spectroscopy is really a spectroscopically detected scattering method as opposed to a more conventional spectroscopy, are somewhat more subtle and warrant additional discussion.
2.1 X-Ray Absorption Spectroscopy X-ray absorption spectroscopy (XAS) refers to the structured absorption on the high-energy side of an X-ray absorption "edge" (the abrupt increase in absorption resulting from excitation of a core electron) [14-16]. For Mn, the "K" (ls initial state) edge occurs at ca. 6500 eV ~ 2 ~. X-ray absorption spectra are frequently divided into the X-ray absorption near edge structure (XANES) region, for structures within ca. 50 eV of the edge, and the extended X-ray absorption fine structure (EXAFS) region, for structures from -50 eV to 1000 eV above the edge. The structure in X-ray absorption spectra is due to the modulations of X-ray photoabsorption cross-section that occur as a consequence of interference between the X-ray excited and the back-scattered photoelectron waves. The principal attraction of XAS, particularly in the context of the OEC, is that it is able to provide element-specific structural information about non-crystalline systems. Unlike magnetic resonance, XAS spectra are always detectable, regardless of the spin-state of the sample. This is, however, also a potential weakness. All of the metal in a sample (e.g. all of the Mn in the OEC) contributes to the observed XAS spectrum. This makes it difficult, if not impossible, to determine the structure of each of the metal sites in a protein. Instead, only the average structure can be determined. The physical principles responsible for XAS structure are the same regardless of whether one is concerned with the XANES or the EXAFS regions. Although this equivalence has been recognized for some time, it has continued to be useful to maintain a distinction between XANES and EXAFS based on the fact that the lower-energy XANES structure is more sensitive to the geometry and the long-range structure of the absorbing site. This is a conse-
StructuralCharacterizationof the Mn Sitein the PhotosyntheticOxygen-EvolvingComplex
7
quence of the longer mean-free-path of the X-ray-excited photoelectron at low photoelectron kinetic energies. In contrast, the EXAFS region can typically be modeled by considering only the simpler, short-range structure. With modern ab initio theory, EXAFS spectra can be modeled to a fairly high degree of precision [17, 18]; similar treatments of the XANES region are not yet possible. 2.2 X-Ray Absorption Near Edge Structure
It is generally accepted that the energy at which an X-ray absorption edge occurs is correlated with the oxidation state of the absorbing atom, with higher oxidation states giving higher energy edge thresholds [19-21]. The physical basis of this effect is open to debate, but probably involves both electrostatic effects (edge energy increases with increasing nuclear charge as it becomes more difficult to ionize the core electron) [19] and continuum resonance effects (edge energy.~ 1/R2 where R = absorber-scatterer distance) [22-24]. It is extremely difficult to separate these effects, since higher oxidation state complexes typically have shorter bond lengths, and are thus expected to have higher energy edges according to either interpretation. The significance of the fact that bond length can affect edge energy is that this has the potential to confound oxidation state assignments: If a complex has unusually short bond lengths, it might have a higher edge energy than would otherwise be expected. Despite the uncertainty in interpretation, and the potential for misleading results, there is ample empirical data which demonstrate an excellent correlation between XANES energy and oxidation state [19, 20, 25]. One of the difficulties with XANES analyses is that it is hard to define exactly what is meant by"the" edge energy. Most XANES spectra consist of numerous poorly resolved transitions and cannot really be said to have a single unique edge energy. A variety of definitions have been used, including the energy at half height [26, 27] and the energy of the first inflection point, as defined either by the maximum [28] in the first derivative or the zero-crossing [29] in the second derivative of the XANES spectrum. All three of these definitions suffer from the fact that they are extremely sensitive to minor variations in the shape and/or resolution of the XANES spectrum. This is illustrated by the simulation in Fig. 3. The spectra in Fig. 3 are the XANES for a Mn(III) model compound subjected to various amounts of broadening. This could represent either instrumental (monochromator) resolution or broadening due to chemical heterogeneity. It is apparent that relatively small changes in resolution can result in large changes in the energy of the first inflection point. An alternative approach to XANES analysis is to fit a spectrum using linear combinations of spectra drawn from a library of model compounds of known oxidation state and structure [30]. Such analyses are substantially less sensitive to minor variations in shape, but suffer the disadvantage of being dependent on the library-of model compounds that are used to fit the unknown spectrum.
8
James E. Penner-Hahn 4020-
-1200 -
1000
-800~>
c~
-20~ ~'~-40!
600~ -400 ~' •200 •
-60i -801
...........................
6530
6550
6 5 7 0 6590 eV Fig. 3. Effect of spectral resolution on the XANES for an Mn model complex. XANES data are in bold (right hand scale), second derivative of the XANES is in light (right hand scale). The edge energy is the negative-going zero crossing in the second derivative at ca. 6554 eV. As resolution improves, new first inflection points are resolved. Even if the same inflection points are followed, the energy shifts by ca. 0.5 eV with resolution
2.3 Extended X-Ray Absorption Fine Structure EXAFS spectra contain oscillations from all of the neighbors around the absorbing atom, with each neighbor contributing oscillations of a characteristic frequency 2R when spectra are plotted in k space (k = 2~-mme (E-Eo)/t~ 2 is the photoelectron wave vector, Eo is the threshold energy for photoelectron excitation). Spectra are often shown as their Fourier transform, so that each "shell" of scatterers shows a peak at R + a, where a is a phase-shift due to the photoelectron scattering. Here a "shell" refers to a group of atoms of similar atomic type (e.g. N or O) at the same distance from the absorbing atom. It is generally accepted that EXAFS can be used to determine bond lengths with an accuracy of 0.02 ,~, coordination numbers to one atom in four or five, and atomic numbers to within one row of the periodic table [15, 16], at least for analyses of firstcoordination shells. Somewhat lower accuracy is generally obtained for more distant atoms. An important limitation of EXAFS concerns the resolution. Like all scattering methods, the ability of EXAFS to resolve two shells is limited by the range of the data. This limitation is particularly severe for the Mn edge, since there are at least two endogenous Fe atoms in the OEC (see Fig. 1). The presence of the Fe K absorption edge (7100 eV) limits the accessible Mn EXAFS range to kmax < _ ~ 11.9 Jr-.1 This means that, under the best conditions, the closest distance spacing that can be resolved is AR/2Ak ~ 0.14 A. In practice, the true resolution limit is generally somewhat worse than this [31]. A second consequence of the limited range of OEC EXAFS spectra is that only a limited amount of information is available. Because EXAFS spectra are band-limited, the amount of information contained in a spectrum is determined by the range of the spectrum (Ak) and by the range of distances that contribute to the spectrum (AR) [32, 33]. The number of independent data points, Nidp = 2 AkAR/n is typically , where $12 and $34 are the effective dimer spins, and ST is the total spin. The authors were able to obtain good fits to the native and the NH3-modified multiline signals signal using [7/2, 4; 1/2> and [7/2, 3; 1/2> basis sets, respectively. An important conclusion is that, if the simulations are limited to normal Mn hyperfine couplings, the ca. 1900 spectral width can only be explained by invoking large dimer spins (e.g., $12=7/2, $34=3,4). This appears to rule out pairwise coupling schemes such as that in Fig. 5c, where there is interdimer coupling between only one Mn of each dimer [51]. Like the dimer simulations, the distorted tetrahedral simulations appear to be inconsistent with other data about the OEC. They require a Mn(III)3Mn(IV) oxidation state while other data (see below) favor a Mn(III)Mn(IV)3 oxidation state for the $2 state. They require that the sign of the hyperfine anisotropy for Mn(III) be reversed from that observed in oxo-bridged Mn(III) model compounds, implying either trigonal bipyramidal or a very unusual tetragonally compressed Mn(III) environment; as well as ferromagnetic intradimer exchange coupling, to give 812 = 7/2 and $34= 3 or 4. There are no examples of ferromagnetic coupling in di-p-oxo bridged dimers. However, in a tetrahedral coupling scheme (Fig. 5 a) apparent ferromagnetic intradimer coupling can result, if the interdimer couplings are larger than the intradimer coupling: 1/4(J13+ J23+ J14+ J24) >[J34[. This condition also seems unlikely. The 1- 2 and 3- 4 dimers are generally considered to have di-poxo bridges (see below), which, in the model compounds made to date, are strongly antiferromagnetically coupled. The $12= 7/2 ground state would thus appear to require unexpectedly large interdimer coupling. Zheng and Dismukes suggest that a possible structural interpretation of their EPR model might be that the Mn ions in the dimers are bridged by hydroxide rather than oxide ligands. The EXAFS data (below) are probably inconsistent with this structure. The ability to simulate two different multiline signals with only small changes in coupling [51] is an important advance in understanding the OEC. These simulations were not able to account for the EPR spectrum observed in Ca2+-depleted, EGTA-treated samples. This could indicate that the Ca2+-deplet -
12
lames E. Penner-Hahn
ed samples have a unique structure, or could reflect the need to adopt a different coupling scheme. In addition, it is not clear that the coupling models in Fig. 5 can account for the existence [39] of excited spin states lying as close as a few cm -1 above the ground state. Although both the dimer simulation [47] and the dimer or dimers simulations [51] can account for the observed EPR spectra, neither appears consistent with other known properties of the OEC. The availability of several different multiline spectra, 55Mn ENDOR data [48], experimentally calibrated estimates of Mn hyperfine anisotropy [51, 52], and careful characterization of excited spin-state energies [39, 49] provide stringent criteria against which future multiline simulations will be judged. In addition, EXAFS data (see below) limit the range of possible OEC structures. 3.1.3 Interpretations of the g = 4.1 Signal
The g = 4.1 signal was originally attributed to an S = 3/2 mononuclear Mn(IV). However, the discovery [53] that the g ~ 4.1 signal for NH3-inhibited, oriented OEC multilayers shows resolved multiline structure demonstrated that at least this form of the g = 4.1 signal must also come from a multinuclear Mn cluster. Based on an analysis of the signal shape, g = 4.1 has been assigned to the middle Kramers transitions of a rhombic S = 5/2 state [54]. More recently, it has been suggested that an S = 3/2 description is required to account for the multiple-frequency EPR spectra of this signal [39]. Since the g = 4.1 signal arises from a ground state [50] and since an Mn(III)/Mn(IV) dimer (see below for oxidation state assignments) would have either an S = 1/2 or S = 7/2 ground state, the g = 4.1 signal must arise minimally from an Mn trimer. Since the g = 4.1 signal is produced at low temperature and converts to the multiline signal at higher temperatures, an early interpretation of the g--4.1 signal was that it represents a precursor to the multiline signal. This precursor state could be either a redox intermediate (Fig 6.I) or a conformational intermediate (Fig 6.II). Recently, Rutherford and co-workers [55] have been able to separate formation of the multiline from formation of the g = 4.1 signal. They found that near-IR (ca. 820 nm) radiation specifically converts the multiline form of $2 to the g -- 4.1 form. Since most earlier illumination studies used white light, samples were exposed to both visible and IR components. The near-IR result might be consistent with Figs. 6.I and II, if it were possible to drive the reaction backwards using near-IR light. However, Rutherford and co-workers also demonstrated that it is possible to prepare samples in the multiline $2 state using 130 K illumination, if no near-IR radiation is present. This demonstrates that the g = 4.1 form is not an intermediate between $1 and the multiline form of $2, but does not distinguish between a redox conversion (Fig. 6.III) and a conformational conversion (Fig. 6.IV) interpretation of the near-IR transition. The near-IR conversion shows a maximum at 150 K. Above 200 K there is no net conversion due to rapid relaxation of the g = 4.1 form back to the multiline form. Below 100 K, the conversion does not proceed, suggesting either that it is an excited state of the species giving rise to the multiline signal that is converted
Structural Characterization of the Mn Site in the Photosynthetic Oxygen-EvolvingComplex
1 T
A
130 K illumination>
Aox
200 K annealing>
B
c
130 K illumination >
multiline
Cox
200 K annealing>
g=4.1
A
visible light
III B
820 nm, 150 K>
A
ZOO K annealing
Box
< B
g=4.1
multiline
IV
c
Co: multiline
Aox
>
A Box
g=4.1
II
13
visible light
820 nm, 150 K> >
Cox (
multiline
200 K annealing
Cox
g=4.1
Fig. 6. Possible interpretations of the production of multiline and g = 4.1 EPR signals for $2. I and II are the initial proposal based on the observation that formation of a g = 4.1 signal appeared to precede formation of a multiline signal; III and IV are modified models in the light of the recent finding that the multiline signal precedes the g = 4.1 signal [55]. Models I and III assume that the multiline and g = 4.1 signals originate from oxidation of different sites (A and B, respectively). Models II and IV assume that the mukiline and g = 4.1 signals originate from different conformations of a single cluster C (C and C', respectively)
by IR radiation, or that there is an activation barrier which m u s t be overcome in the conversion. If it is an excited state that is responsible for the IR conversion, this state m u s t lie significantly higher than the ca. 35 cm -1 excited spinstate found in relaxation studies [49]. The action s p e c t r u m of the multiline -+ g = 4.1 conversion has a m a x i m u m at about 820 n m [55]. The authors assigned this to an intervalence transition, converting for example an Mn(III)/Mn(IV) dimer to an Mn(IV)/Mn(III) dimer. The valence isomer might either be responsible itself for the g = 4.1 signal or might trigger a further redox reaction. Thus, if an Mn(III)/Mn(IV) dimer represented Aox in Fig. 6.III, the valence isomer Mn(IV)/Mn(III) might be able to oxidize B. Equivalent schemes can be drawn for possible valence isomerization within a single cluster (Fig. 6.IV), e.g. M n ( I I I ) / M n ( I V ) - M n ( I V ) / M n ( I V ) --> M n ( I V ) / M n ( I I I ) - M n ( I V ) / M n ( I V ) , possibly with further conversion to Mn(IV)/ Mn(IV)-Mn(III)/Mn(IV). Interestingly, low-temperature near-IR irradiation was able to convert the native multiline, the Sr2+-substituted multiline and the Ca2+-depleted, chelator-
14
James E. Penner-Hahn
treated multiline forms to g 4.1 forms, but was not able to convert the N H 3m o d i f i e d multiline to a g = 4.1 form [55]. This suggests that the species giving rise to the NH3-modified mukiline is in some way unique and different, perhaps in structure or ligation, from the other multiline forms. This is in contrast to the conclusion, based on multiline simulations [51], that it was the Ca2+-depleted form that appeared to have a unique structure. =
3.1.4
The ST State EPR Signal Although the $1 state is EPR-silent under normal conditions, it has been reported to have an EPR signal centered at g = 4.8 with a peak-to-peak width of ca. 600 G [56]. This signal, which is consistent with an integer spin, S = 1 state [57, 58], is seen only in parallel polarization EPR. The $1 EPR signal converts to the multiline EPR signal on illumination at 200 K. However, the most striking observation about the $1 EPR signal is that low-temperature illumination gave the g=4.1 S2 EPR signal (ca. 30% of maximum intensity, as judged by the extent of conversion to multiline on annealing at 200 K) without significant loss of the S1 EPR signal. If this observation is correct, it requires that the g = 4.1 $2 signal and the multiline $2 signal come from different clusters, not different conformations of a single cluster. The S1 EPR resuks were originally attributed to intermediacy of the g = 4.1 species between the oxidant (YZ.) and the multiline state [56]. With the discovery (above) that low-temperature production of the g = 4.1 signal is driven by a near-IR transition [55], this model requires modification. The scheme shown in Fig. 7 would be consistent both with the near-IR results and the observations made on the S~ EPR signal. In this scheme, A is the center that gives rise to the S~ EPR signal when reduced and the multiline signal when oxi-
30% >
o
>
A OX
St EPR )-
AOX
multiline
)
multiline
B
A S 1 EPR
B
g~.~. l
200 K annealing
visible
near-IR
light, 130 K
light, 130 K
200 K illumination
B
A 70%
S 1 EPR
B
A Sj EPR
B
Aox multiline
B
Fig. 7. A possible model for OEC redox conversions, incorporating results of both near-IR
conversion [55] and $1 EPR measurements [56] (see text for details)
StructuralCharacterizationof the Mn Site in the PhotosyntheticOxygen-EvolvingComplex
15
dized, while B is the center that gives the g = 4.1 signal when oxidized. The production of only 30% oxidized centers following 130 K illumination is due to competition from other, non-manganese electron donors at low temperature [591. More recently, it has been reported, based on measurements of spin-lattice relaxation rates for the YD" tyrosine radical, that the $1 state of the OEC is diamagnetic [60]. This would appear to be inconsistent with the existence of an EPR signal for $1. However, the relaxation measurements were made on longterm dark adapted samples, thus it is possible [4, 60] that a paramagnetic form of $1 (which gives rise to the $I EPR signal) converts, on long-term dark adaptation, to a diamagnetic resting form. 3.1 .S The "$3" State EPR Signals
A variety of treatments, most involving modification of the Ca :+ site, the C1site, or both, inhibit water oxidation by the OEC. When inhibited samples are illuminated they show a novel broad (100-230 G) radical EPR signal centered at g = 2.0. This effect was originally discovered for Ca2÷-depleted samples [42], but has subsequently been found [61-63] to result from a variety of inhibitory treatments, including C1- depletion, and fluoride, acetate, or ammonia treatment (which may operate by C1- displacement). A variety of lines of evidence, including flash yield and the ability to regenerate $2 on readdition of the disrupted cofactor (cf. [2, 64] for reviews) suggest that the new signal arises from a state that is one electron more oxidized than $2, thus making this formally an S3 EPR signal. The new signal has very limited stability at room temperature (hence the need to freeze under illumination in some of the preparations) and has limited stability even at low temperature [64]. The new signal lacks the breadth and hyperfine structure that would be expected for an Mn-centered paramagnetic center, but is much broader and much harder to saturate than would be expected for an isolated organic radical. It is generally agreed that the new signal can be explained as an organic radical that is dipole-coupled to a fast-relaxing paramagnetic center, the latter presumably being the OEC Mn cluster. This is supported by the observation that formation of S3 leads to loss of the S2 multiline signal as detected by cw EPR, but not by field-swept spin-echo EPR [65]. The new 100-230 G signal can thus be described as being due either to $3 or to S/X +, where X ÷ is the organic radical. Although the distinction is to some extent only semantic, the S3 designation implies that the state giving rise to the EPR signal is on the normal path to water oxidation, albeit altered in some way that makes it EPR-detectable. In contrast, the $2X÷ designation implies that the oxidation of X is an artefact of the inhibitory treatment, and is not necessarily relevant to the normal functioning of the OEC. It has been suggested, based on UV-visible difference spectra, that oxidized histidine is the radical responsible for the new EPR signal [66]. This could be consistent with either the $3 or the S2X+ designations, although a specific role for His- in water oxidation has been strongly favored by some authors. There have, however, been several suggestions that His. is not responsible for the "$3"
16
James E. Penner-Hahn
signal [67]. Recently, Britt and co-workers have reported ESEEM spectra for the "$3" signal that is produced in acetate-inhibited OEC samples [68]. By using a cyanobacterial (Synechocystis) source for the OEC, the authors were able to specifically deuterate the tyrosine residues, and to provide conclusive evidence that a tyrosyl radical is responsible for the "$3" EPR signal. Since Yz is the tyrosine that is close to the Mn cluster, at an estimated distance of 4.5 A [69], this is presumably the tyrosyl radical that is dipole-coupled to the Mn cluster. A more appropriate designation for the state that gives rise to the radical signal is thus S2Yz', a state on the normal path to water oxidation. The structural change that gives rise to the new signal appears to involve some modification of the Mn site so that it can no longer be oxidized byYz.. On removal of the inhibition (e.g., addition of Ca2+), the Yz" can oxidize the OEC to $3, thus continuing the Kok cycle. 3.2 Oxidation State Assignments It is only possible to have an S = 1/2 EPR signal for $2 if the cluster that gives rise to this signal contains an odd number of half-integer spin ions. For Mn, the only commonly available oxidation states with an odd number of electrons are Mn(II) and Mn(IV). The structural evidence (see below) that the OEC contains Mn(p-O)2Mn units, and the observation that di-p-oxo bridged structures are only found for Mn(III) and Mn(IV), has led to the suggestion that the $2 state must have an oxidation state of either Mn(III)3Mn(IV) or Mn(III)Mn(IV) 3. If S~ is one electron more reduced than $2 (see Sect. 3.4), this implies that $1 is either Mn(III) 4 or Mn(III)2Mn(IV)2. Several groups have used XANES to determine the oxidation state of S1, and the consensus is that the XANES data for S1 are most consistent with the latter oxidation state assignment [4, 26, 29, 70]. In model studies, it is relatively difficuk to distinguish between Mn(III) and Mn(IV) XANES spectra, such that the uncertainty in absolute (although not necessarily relative) oxidation state assignments is ca. +25% [70]. Although relatively poor, this precision is sufficiently good to distinguish unambiguously between average oxidation states of 3.0 and 3.5. It is perhaps worth recalling, however, that the ability to reduce the oxidation state question to one of distinguishing between average oxidation states of 3.0 and 3.5 is based on the assumption that only Mn spins contribute to the overall spin of the OEC. If another paramagnetic center was coupled to the cluster (as proposed e. g. in [39]), then it would be possible, from a spin-counting perspective, to have S~ as either Mn(III)3Mn(IV) or Mn(III)Mn(IV) 3. The best fit to the OEC XANES gives an average oxidation state of 3.5 + 0.25 for $1 [70], which could also be consistent with either of these possibilities. 3.3 Atomic Arrangements The Fourier transform of a recent EXAFS spectrum for the S1 state of the OEC is shown in Fig. 8. This spectrum is dominated by three peaks, correspond!ng to three principal shells of scatterers. The first two of these, at R + a = 1.6 A and
17
Structural Characterization of the Mn Site in the Photosynthetic Oxygen-Evolving Complex o
12
'
I
'
'
'
I
'
'
'
I
I
Mn-O nearestneighbors Mn-Mn at = 2.7 A
:18 ~ ~
4
•
2
~
0
I
0
2
Mn-Mn at = 3.3 A
|
I|
New interaction.*
4 6 Radius + a (/~_)
8
10
Fig. 8. Fourier transform of the EXAFS spectrum for the $1 state. Redrawn using data from [74]
2.3 A, are due to Mn-nearest neighbor interactions at an average distance of 1.9 A and Mn-Mn interactions at ca. 2.7 ~. These peaks were seen in the first EXAFS measurements of the OEC [71], and have dominated the structural models that have been proposed. The 2.7 A Mn-Mn distance is typical of the Mn-Mn distances found in di-p-oxo bridged Mn dimers, suggesting that this unit is a basic building block of the OEC Mn cluster. The distance for the Mnnearest neighbor peak is typical of Mn-O and Mn-N ligation (which cannot be distinguished by EXAFS), and is much shorter than Mn-S or Mn-C1 distances, thus suggesting that the Mn in the OEC must be ligated primarily by O- or Ncontaining ligands. Since only a single histidine nitrogen is seen in ESEEM measurements [72, 73], it is generally assumed that the Mn ions have predominantly oxygen ligation (oxo, hydroxo, aqua, carboxylato and possibly phenoxo or alkoxo ligands). 3.3.1
Mn-Neorest Neighbor Interactions
If the nearest neighbor interactions are modeled by a single Mn-O shell, the best fit is with ca. 3 oxygen atoms at an average distance of ca. 1.9 A [74]. This is clearly not a complete description of the nearest neighbor environment for at least two reasons. Firstly, the apparent coordination number is much too small. Secondly, if there are Mn(p-O)2Mn dimers in the OEC, there must be Mn-oxo bonds, which are exl~ected to be about 1.75/~ long, and there must be longer Mn-O bonds at ca. 2 A. This recognition has led many groups to analyze the first shell EXAFS in terms of two shells of scatterers [75- 79]. These fits typically give a shell of about two Mn-O at 1.75-1.8 • together with a second shell of variable coordination number at 1.90-2.25 A. The difficuky with these fits is that the number of variable parameters comes very close to the number of independent degrees of freedom in the data (see Sect. 2.3). If an attempt is made to correct
18
James E. Penner-Hahn
this when fitting the data [33], the significance of the second shell is marginal at best [79], and the two shell fits are probably not justified [74, 80]. We have argued [74] that the large (0.35 .~) range of distances reported for the second Mn-O shell reflects this fact. Much has been written regarding the question of whether there are one or two shells of nearest neighbors in the OEC. To some extent, this is an argument that is of interest only to EXAFS aficionados and is of little relevance to the OEC. After all, there is general agreement that the EXAFS data point to ligation by a disordered shell of primarily oxygen ligands. There are, however, two reasons why this issue is important. Firstly, if it is possible to resolve the Mn-oxo shell, then it is possible to use the apparent number of Mn-oxo interactions to define the structure of the OEC. Thus, it has been argued [4] that structural models with more than two Mn(p-O)2Mn units are unlikely, since these would give a higher Mn-oxo coordination number than is observed. If the two shell fits are not justified, than this argument cannot be used. Secondly, there is important information contained in the observation that two-shell fits are not significantly better than one-shell fits, and this information can be overlooked when two shells are forced to fit the data. Many Mn models and the Mn(p-O)zMn unit in Mn catalase do show two well-resolved shells of scatterers [31, 74, 81]. The models that do not show two well-resolved shells are those that have intermediate-range Mn-O interactions (ca. 1.9 A) in addition to both short Mn-O distances (N 1.8 A) and long Mn-(O/N) distances (N2.1 A) [82]. Thus, for example, the EXAFS data for the [Mn(IV)(salpn)(p-O)]2 dimer are best fitted by a single shell of 3 - 4 Mn-O scatterers at ~ 1.90 A, despite the presence of both short Mn-Ooxo distances and longer Mn-O/N distances [83, 84]. A similar situation may exist for the OEC [74], with Mn-O distances of 1.9 A coming from hydroxo, alkoxo, or phenoxo ligation. Unfortunately, the resolution of the OEC EXAFS data (ARN0.16 _~) is not sufficient to justify the use of three shells of nearest neighbors. 3.3.2
Mn-Mn Interactions
The apparent coordination number for the 2.7 ~ Mn-Mn peak is approximately 1.2 or 1.3 [4, 74, 75, 79]. This represents the number of Mn neighbors seen by each Mn, and thus corresponds to 2 or 3 Mn-Mn interactions for the Mn 4 cluster. This has generally been interpreted as evidence for two Mn(l.t-O)zMn dimers within the OEC [4, 77], a structure which would give an average coordination number of 1.0. EXAFS coordination numbers are not well defined. Nevertheless, it is intriguing that the apparent coordination number is consistently somewhat larger than expected for data measured by different groups, on different preparations, and analyzed using different parameters. Although disorder can easily lead to underestimation of EXAFS coordination numbers, it is rare to overestimate the coordination number. Thus, the apparent Mn-Mn number of 1.2-1.3 may indicate that there is an additional contribution to the 2.7 ]k peak. The most likely possibilities are either Mn-C from bidentate carboxylate residues or a third Mn-Mn interaction.
StructuralCharacterizationof the Mn Sitein the PhotosyntheticOxygen-EvolvingComplex
19
An important advance in characterizing the OEC structure was the development of polarized EXAFS measurements on oriented OEC multilayers. The initial measurements demonstrated that the 2.7 A peak is relatively isotropic, while the 3.3 A peak (see Sect. 3.3.3) has more pronounced orientation dependence [85]. A subsequent repetition of this work gave an average angle of 60 ° between the Mn-Mn vectors and the membrane normal [86]. Interestingly, the latter work found that the average Mn-Mn distance varied from 2.71 ,~ when the X-ray polarization was at 15° to the membrane normal to 2.74 A when it was at 75° [86]. Although this difference is small, the precision of EXAFS distance determinations is usually very good [74, 87], thus this difference is likely to be significant. The difference in Mn-Mn distance, which disappeared on oxidation to the S2 state (see Sect. 3.4), was interpreted [86] as evidence for two slightly different Mn-Mn distances in $1, which might be attributed to a Mn(III) 2 dimer and a Mn(IV)2 dimer. Subsequent EXAFS studies of inhibited samples (see Sect. 4) have confirmed the presence of two different Mn-Mn distances and have given refined Mn-Mn angles of 55° and 67° for the two different 2.7 ~ Mn-Mn vectors relative to the membrane normal [88]. The observed distance for the Mn-Mn shell is simply the weighted average of the different Mn-Mn distances that contribute to the shell. If the angles of 55° and 67° are attributed to dimers "A" and "B" respectively, R A (the Mn-Mn distance for dimer A) will contribute more when the X-ray polarization is parallel to the membrane normal, while RB will contribute more when the X-ray polarization is perpendicular to the membrane normal, thus accounting for the anisotropy in distance. However, angles of 55° and 67° give very nearly isotropic EXAFS and thus should not produce significant anisotropy. The anisotropy predicted by the measured angles [88] is so small that the true Mn-Mn distances would have to differ by > 0.2 .~ in order for two Mn-Mn vectors to give 0.03 A anisotropy in distance. An 0.2 A spread in distances is inconsistent with the isotropic EXAFS. It is not clear what this observation means for the OEC structure. It is possible that either the reported distance anisotropy or the reported angles are wrong. An alternative, however, is that there is a third scatterer (probably MnMn) which also contributes to the 2.7 A peak. If this interaction were oriented approximately parallel to the membrane normal (i.e., if it were very anisotropic) with a slightly shorter Mn-Mn distance, it would be possible to account for the angular anisotropy in distance [86] but still explain the lack of two resolvable 2.7 A Mn-Mn distances in the isotropic EXAFS. In this model, the apparent coordination number of 1.2 would be attributed to a slight underestimation of the true coordination number, due to slight disorder in the three Mn-Mn distances. 3.3.3 Outer Shell Interactions
From Fig. 8, it is clear that there is a third peak that is well above the noise level and there may even be a fourth peak. The latter is unconfirmed and will not be discussed further. The third shell peak reflects the long range structure in the
20
lames E. Penner-Hahn
OEC cluster. Interpretation of this peak has been controversial. The first suggestion that there might be scatterers at > 3 A was made in 1987 [89], but this suggestion was subsequently withdrawn and the peak was re-attributed to noisy data [28]. Later work using lower temperature measurements, showed reproducible features at R > 3 A, first in oriented [90] and subsequently in isotropic samples [30]. The observation that the third shell feature is seen only at low temperatures suggests that this feature arises from a fairly weak interaction. The third shell interaction shows greater dichroism than the 2.7 A Mn-Mn vectors, with an average angle of 43 ° relative to the membrane normal [86]. The relatively large intensity of the third shell peak, together with the absence of a significant number of histidine ligands (which give outer shell peaks) suggests that the third shell peak in the OEC is due to a Mn-metal interaction. The likely candidates for the metal are Mn and Ca. Although it has been reported that Mn-Ca interactions give better fits to this peak than do Mn-Mn interactions [78], it is now generally agreed that it is probably not possible to distinguish Mn-Mn from Mn-Ca scattering, at least not in these samples [79, 91]. Although the peak can be fitted with two shells (e. g. Mn-Mn+Mn-Ca), these fits were difficult to justify compared to single shell fits [79, 88]. This does not, of course, exclude the possibility that Mn, Ca (and potentially C) all contribute to the third shell [86, 92]. In addition to uncertainty over the identity of the distant scatterers, there is even uncertainty over the Mn-metal distance. We [30, 82, 87], together with George et al. [90] and the Klein/Sauer group [4, 77, 79, 86, 92] have consistently found a Mn-metal distance of approximately 3.3 A. In contrast, Nugent and coworkers have reported that the distance is 3.7 A [78, 91]. The origin of this discrepancy is not known. One possibility is that one of the reported distances is a result of the existence of multiple minima in EXAFS fits. Thus, for example, we have found that our data can be fitted using either distance, although we find the 3.3 A fits to be better [82]. In an effort to determine unambiguously whether Ca scattering contributes to the 3.3 A feature, several groups have removed Ca 2+ or replaced it with other metals and determined the effect of these perturbations on the EXAFS. One such study [93] found that replacement of Ca 2÷ by Sr 2÷ gave an increase in the amplitude of the 3.3 A interaction and that fits to the substituted data were slightly better with Sr + Mn scattering than with Ca+Mn scattering. Similarly, a decrease in amplitude was observed when Ca 2÷ was replaced by Na ÷ [91]. Both results suggest that at least some of the outer shell scattering is due to a Mn-Ca interaction. In contrast, we found no change in the third shell peak when Ca 2÷ was replaced by Sr2+, La3÷, or Dy3÷ [87]. In some cases, the fits were slightly better using Sr rather than Ca as the scatterer. However, this was true both for control samples that had never been exposed to Sr and for Sr-substituted samples, thus illustrating the difficulty in using fit quality to distinguish between fits for outer shell scattering [31]. One possible explanation for these apparently different results is that the samples were biochemically different. Our experiments were done in the absence of the 17 and 24 kDa extrinsic polypeptides, while these peptides were present in the other studies. If one or both peptides contributes ligands that
StructuralCharacterizationof the Mn Site in the PhotosyntheticOxygen-EvolvingComplex
21
/0 Mn~'!/NMn~=3. 3/~ ? '
H~
/M O
M=Ca2+,Sr2+,Dy3+,La3+
H
9. One possible explanation for the changein 2.7 the/~ Mn-Mn distancewhen the metal in the Ca site is changed Fig.
bridge the Mn-Ca interaction, the interaction would be expected to be stronger (and thus more readily detectable) in the presence of the extrinsic polypeptides. Akhough we did not find any changes in the 3.3 .~ feature, we did observe small but reproducible changes in the 2.7/~ peak when Ca was replaced [87]. The average Mn-Mn distance decreased slightly (0.013 _+0.004 .~) when Ca was replaced with Sr and increased slightly (0.015 + 0.004 .~) when Ca was replaced with a lanthanide. Similar results seem to have been observed in other studies [93], albeit with somewhat lower precision due to variability in E0 [87]. This is direct evidence of a role for Ca in controlling the M n core structure. The average Mn-Mn distance appears to increase with increasing Lewis acidity of the cation that is bound in the Ca site. Based on model studies, the observed changes in RMn.Mnare too small to be due to protonation [83, 84]. One possible explanation is that the Mn-Mn distance is affected by hydrogen bonding, with the Ca site modulating the acidity of a proton bound near an oxo bridge of one of the di-p-oxo bridged dimers (Fig. 9). A model such as Fig. 9 would explain the observation that Caa÷ is bound most tightly in $1 [3, 94, 95], since the basicity of the oxo-bridge, and thus the strength of the hydrogen bond, is expected to decrease when the Mn is oxidized [84]. The change in the multiline EPR spectrum following Sr substitution would be consistent with the fact that protonation alters intradimer exchange couplings [84]. Finally, the inability to form $2 when lanthanides are bound at the Ca site [96] and the stability of $2 when Ca is depleted [42] might be related, at least in part, to alterations in the redox potential of the Mn(p-O)/Mn dimers as a function of variations in hydrogen bonding. 3.4 S State Dependence of the OEC 3.4.1 The S2 State
As described in Sect. 3.1, the $2 state is EPR-active while S1 is EPR-silent under normal conditions. This suggested that Mn was oxidized in the $1~ $2 transition, and this was confirmed by early XANES measurements showing an i n -
22
James E. Penner-Hahn
crease in edge energy on formation of S 2 [97]. Very similar changes in energy are observed regardless of whether the multiline or the g ~ 4.1 forms of $2 are prepared [98, 99], although the details of the edge shape and the EXAFS are slightly different for the two forms of $2 [92]. This suggests that Mn is always oxidized in the S~-4 $2 transition, but that slightly different structures are formed, as expected given the different magnetic properties of the multiline and g-~4.1 forms. Several of the most recent reports of S state edge energies are summarized in Table 1. Although slightly different edge energies are obtained depending on the details of the data analysis (and possibly on sample preparation), similar results have been obtained in all of the studies of the $1-4 $2 transition, strongly supporting the conclusion that Mn is oxidized in this S-state transition. The 2.7 A Mn-Mn peak in the EXAFS for the g ~ 4.1 form of $2 is about 30% smaller than that seen for the multiline form of $2 [92]. This decrease in amplitude was modeled using two different Mn-Mn distances, at 2.72 and 2.85 A. This is probably not a unique fit since the spread in the apparent Mn-Mn distance is smaller than the resolution of the data. Nevertheless, it suggests a structural origin for the different spin-states of the two $2 forms. It is intriguing that the ratio of the coordination numbers for the short (2.72 ~k) and long (2.85 ~,) Mn-Mn shells is approximately 2:1. Similar behavior has been observed for inhibited $2 samples (see below). 3.4.2 The S3 state - XANES
In contrast to the general consensus on the S1--~ S2 transition, characterization of the $2-4 $3 transition has been controversial. There are three approaches to preparation of $3: The S1 state can be illuminated under conditions that allow two, and only two, turnovers; samples can be inhibited so that they cannot advance beyond $3; or illuminated with two short, saturating flashes and then frozen rapidly before they can be $3 can decay. The double-turnover approach was the first to be applied to the OEC [28]. Using chemical treatments to introduce a high-potential acceptor (oxidized non-heme Fe), it was possible to prepare samples with only 35 % of the normal $2 multiline EPR signal. The decrease in multiline intensity was attributed to formation of S3 in 65 % of the centers. The observed edge energy (see Table 1) was unchanged from that in $2, suggesting that there was no oxidation of Mn in the $2-4 $3 transition. Different results were obtained using inhibited samples. The advantage of inhibited samples is that they may permit preparation of more homogeneous samples. The disadvantage is that these samples really represent S*, i.e. a potentially modified $3. The observed edge energies increase by approximately 1 eV in both the S1 -~ $2 and the $2 -~ $3 transitions (see bottom of Table 1), suggesting that for the inhibited samples, Mn oxidation occurs in both transitions [100]. As described in Sect. 3.1.5, the inhibited $3 state is characterized by a broad radical signal in the EPR, now shown to be Yz'. This is consistent with the oxidation of Mn if the inhibited S* states contain a mixture of states, many of which have oxidized Mn and account for the shift in edge energy, and some
Structural Characterization of the Mn Site in the Photosynthetic Oxygen-Evolving Complex Table 1.
23
Edge energies for the OECa
State
Edge
S1 $2 $3 S~ $2 S~ $2 $3 $1 $2 $3 S~ S2 $3
6551.4 6552.3 6552.1 6551.6 6552.6 6551.7 6553.5 6553.8 6551.7 6552.5 6553.7 6551.8 6552.7 6553.2
Inhibited samples Acetate S* S* S* NaC1 S* S* S* Ammonia S* S* S*
6552.6 6553.2 6553.8 6551.0 6551.9 6553.2 6550.6 6551.5 6552.5
AE (eV)b
Ref. [28, 76]
0.9 0.0 [100] 1.0 [29] 1.8 0.3 [26] 0.8 1.2 [27] 0.9 0.5 [100] 0.6 0.6 [100] 0.9 1.3 [100] 0.9 1.0
a $3 state in [28] was trapped by chemical oxidation; other uninhibited S3 data were for flashed samples. b Change in energy between $2 and $1 or $3 and $2. of which have Yz" instead of oxidized Mn, and account for the broad radical in the EPR [100]. Recently, two groups have used flash illumination to prepare $3 and So [26, 29], reaching opposite conclusions. These experiments are inherently difficult because of the inevitable dephasing of S state composition as successive flashes are given. Dephasing can result from misses (centers that do not advance), double hits (centers that advance two S states on a single flash) or from centers that don't cycle (e. g. are inactive). In order to obtain "true" S state spectra, both groups were forced to estimate the S state composition after each flash and use these populations to deconvolute their measured spectra. The assumptions used in both of the deconvolutions are potentially problematic and may be responsible for the different conclusions. Neither group included a fraction of centers that did not turnover. This could potentially lead to an underestimation of the edge shifts. However, Ono et al., who have the smallest range of edge energies and thus would be most likely to have suffered from non-active centers, are the group that finds an energy
24 Table 2.
James E. Penner-Hahn Comparison of observed and fitted S state edge energies for flash experiments
Sample
Edge
Native
Observed 6551.7 Fit 6551.7 Observed 6551.7 Fit 6551.7 Observed 6551.8 Observed 6552.1
6552.4 6552.5 6552.4 6552.5 6552.7 6552.8
Observed 6551.7 Fit 6551.7
6553.5 6553.7 6553.5 6553.8
Native Native C1-depleted Native
0 flash/S, 1 flash/S2 2 flash/S3 3 flash/S0 4 flash 5 flash 6553.3 6553.7 6553.1 6553.6 6553.2 6552.9
6551.5 6550.7 6551.6 6550.9 6551.8 6552.7
6551.7
Ref. [26]
6551.7 6552.0 [101] 6551.6 6552.5
6550.5 6551.0 6552.9 6550.1
[27] [29]
See text for discussion of procedures used to calculate true S state spectra. change on $2--~ $3 (see Table 2 for a summary of reported edge energies). It thus seems unlikely that the different conclusions can be related to different fractions of inactive centers. Ono et al. illuminated their samples using Nd-YAG laser flashes (7 ns) and thus did not consider double hits [26, 101]. The rate of reduction of S states by YD was estimated from literature values and the percentage of misses (9 %) was refined to give the best fit to the observed edge energies. This approach has been criticized for lacking an independent measure of the S state composition [29]. However, the clear period-four oscillation in the measured edge energies (Table 2) suggests that there is not significant scrambling of the S state composition. In particular, the observed edge energy after 3 flashes is slightly lower than the starting energy ($1). This is the step at which virtually all of the models (see below) predict Mn should be reduced relative to $1. If the composition was too far off, the authors should not have seen reduction in edge energy at this step. Recently, Ono and co-workers have repeated these measurements on samples that were inhibited by either C1- [27] or Ca2÷ depletion [102]. The latter showed a resting (0-flash) edge energy that was decreased by ca. I eV relative to the control. This suggests the presence of Mn(II), and thus these data may be difficuk to interpret. For the C1--depleted data, however, the 0- and 1-flash energies are very similar to those in the control (see Table 2). In the C1--depleted samples, which do not advance beyond $2, the edge energy increases by only 0.1 eV on the second flash, while for the control, the second flash gives an increase of 0.5-0.9 eV. This difference is consistent with a model in which Mn oxidation is blocked in C1--depleted samples, and thus supports the proposal that Mn is oxidized in the $2---~$3 transition. However, these data are relatively noisy and must certainly be confirmed. Klein, Sauer, and co-workers [29] measured the relative concentration of $2 directly following each flash by using the intensity of the S2 multiline signal. With these data available for five flashes, they were able to treat both the miss fraction (12 %) and the double hit fraction (5 %) as adjustable parameters (the larger double hit fraction resuked from use of a flash lamp with a 14 l~s
Structural Characterization of the Mn Site in the Photosynthetic Oxygen-Evolving Complex
25
FWHM). This is clearly a more precise solution to the problem of S state deconvolution. It is not clear, however, that five flashes provide enough information to permit an accurate determination of the miss and double hit fractions. With data for only five flashes it is not possible to account for different miss fractions on different S state transitions (see Sect. 1.3). Perhaps more importanfly, the use of an average double hit parameter may underestimate the S state scrambling if, as has been found in some cases, the double hit fraction is large only on the first flash [8, 9]. A large double hit fraction would be one explanation for the 1.8 eV shift observed on the first flash, which is large for oxidation of only 25 % of the Mn. In model compounds, the change in edge energy is typically about 3 eV between Mn(III) and Mn(IV) [82]. If the $ 1 ~ Sa transition corresponds to oxidation of one out of four Mn atoms, it should show a shift of approximately 0.75 eV. If the $2 energy is overestimated, this would, in turn, lead to underestimation of the $2--~ $3 transition energy change. In addition to deconvolution differences, a second difference in these data concerns the method used to define the edge energy. Ono et al. used half-height, while Klein, Sauer, and their co-workers used the first inflection point. There is a rather pronounced change in the shape of the XANES spectrum between S~ and Sa (see Fig. 4 in [29]). The appearance of a new feature on the rising edge of the $2 spectrum may be, in part, responsible for the unusually large change in energy between S1 and S2. The change in edge shape (i. e. the new feature in the second derivative) causes the first inflection point, defined as the zero crossing in the second derivative, to shift to higher energy. In principle, this could lead to overestimation of the $2 energy. In summary, the XANES data for S3 are contradictory. Some of the data clearly suggest Mn oxidation in both $1---~$2 and $2---~$3. Other data show a very large shift in energy in $1---~S2 and a very small shift in Sz---~$3. This has been interpreted as evidence that Mn is not oxidized on the $2---~S3 transition [29], but might alternatively be interpreted as evidence that Mn is oxidized in both steps, but that the apparent S2 edge energy is too high. At the very least, it seems safe to conclude that the present XANES data do not provide compelling evidence for concluding that Mn is not oxidized in the $2---~$3 transition. 3.4.3 The $3 State - Other Measures of the Oxidation State
In addition to XANES, NMR, EPR and UV-visible spectroscopy have all been used to investigate the Mn oxidation state changes that accompany the S state transitions. The solvent 1H relaxation rate, which can be measured by NMR, should increase if Mn(II) or Mn(IV) ions are formed. The solvent relaxation rate increases in the $1 -~ $2 transition but not in the S2 --~ $3 transition [103]. This was interpreted as evidence for a mononuclear Mn(IV) in $2 and $3, but not in $1. However, strong relaxers other than mononuclear Mn(IV) might also be consistent with these data. Data for So are less well defined. It appears, however, that there is a further increase in relaxation rate in So. This was at-
26
James E. Penner-Hahn
tributed to formation of mononuclear Mn(II). The NMR data thus suggest that Mn is oxidized on So ~ $1 and $I ~ $2. The lack of change in relaxation rate on the $ 2 ~ $3 transition suggests either that Mn is not oxidized in this transition, or, if an Mn atom is oxidized that it does not contribute to the solvent water relaxation rate. The relaxation rate of the inactive tyrosyl radical, YD', has been measured by EPR relaxation [104, 105]. As with the NMR, the fastest relaxation rates were found for So, with $2 ~ $3 lower, and $1 lowest. If a single Mn dimer dominates the relaxation rate for YD', these data would suggest oxidation states of Mn(II) Mn(III) for S0, Mn(III) 2 for So, and Mn(III)Mn(IV) for Sa and $3. As with the NMR data, these data indicate that if an Mn atom is oxidized in the $2--~ $3 transition, it must not affect the spin-lattice relaxation rate for YD'. Both the NMR and EPR measurements are only indirectly sensitive to Mn oxidation state changes and may not necessarily be responsive to all of the Mn in the OEC. In contrast, UV-visible changes should be more directly sensitive to the Mn. These data have been controversial (see [106] for a review). There is now general consensus on spectral differences for S1--~ Sa and Sa---~$3. Both are characterized by a broad peak centered around 300 nm with AE-~ 4000-6000 M-~cm-~. Based on the similarity in peak wavelength, bandwidth, and amplitude, Dekker has interpreted both as M n ( I I I ) ~ Mn(IV) transitions [106]. The S0--~ S1 transition is the hardest to study, since the SOstate is the hardest to make pure and thus most susceptible to differences in data deconvolution. The S0---~$1 difference can also be interpreted as Mn(III)--~ Mn(IV), however this is by no means conclusive [106].
3.4.4 The Ss State - EXAFS
The first studies of S 3 found no change in the average Mn-Mn distance (2.72 A) on formation of $3 [28], although there was some increase in disorder of this shell. Fitting results suggested that if the disorder was due to the presence of 2 Mn-Mn distances, the spread in distance was approximately 0.15 .~. More recent measurements, using two saturating flashes to produce $3, suggest that there is a significant change in the Mn-Mn interactions when $3 is formed [4, 107], with both an increase in average distance and a decrease in the size of the Mn-Mn peak. The data can be fitted using either a single shell of two Mn-Mn interactions at 2.8 A, or with two Mn-Mn shells at 2.8 A and 3.0/~, giving an average distance of 2.9 ~. It is not clear why such different results have been obtained from $3. Although both experiments required deconvolution to obtain the true $3 spectrum, the%S3 that was formed in the two cases was very similar (65 % vs 70 %). Bond length differences of this magnitude are well outside the uncertainty of EXAFS, thus suggesting a difference of some sort between the samples. If the observation of a significant increase in both Mn-Mn distances is correct, this implies a significant structural change in the $2-~ $3 transition.
StructuralCharacterizationof the Mn Sitein the PhotosyntheticOxygen-EvolvingComplex
27
3.4.5
The So State The So state is the hardest to prepare by flash oxidation due to the dephasing which occurs. The XANES data for So all agree that this state is more reduced than S1 (see Table 2). Based on analysis of the shape of the XANES second derivative, it has been argued that Mn(II) is present in So [4, 29]. 4 Chemical Perturbations of the OEC
In addition to illumination, a variety of chemical perturbations have been used to probe the structure of the OEC. Inhibitors include potential mimics for water binding (NH3) or for H20 a (NH2OH), together with cofactor substitutions (e. g. F- for El-). 4.1 Inhibitors of Oxygen Evolution
Ammonia is a potential analog of substrate water. There are two independent NH 3 sites [108]. One, accessible to many amines, is competitive with CI-, while the other, accessible only to ammonia, is not competitive with C1-. In NH3-inhibited samples, NH 3 appears to be a ligand to Mn based on EPR [40] and ESEEM [109] measurements, and has been proposed to bind as a bridging amide (NHe) based on the observed asymmetry in the 14N quadrupolar interaction. Binding at the C1--independent site (which may be a water site) does not occur in SOor $1, or at low temperature, and gives rise to a modified multiline signal. The EXAFS data for such samples show a significant decrease in the amplitude of the Mn-Mn peak [88]. This decrease can be modeled either by doubling the Debye-Waller disorder term while keeping the distance the same (2.72 ~), or by including two different Mn-Mn shells at 2.71 and 2.86 A. This spread in distances is too small to allow reliable refinement given the limited resolution of the data. Consistent with this, the fit quality, when corrected for the number of variable parameters, is worse for two Mn-Mn shells than for a single shell. This does not mean that there are not two different distances but rather that they cannot be refined using isotropic data. However, the presence of two different Mn-Mn distances was confirmed by polarized measurements on oriented samples. The apparent distances were now 2.72 and 2.87 ~,, at angles of 55 ° and 67 ° respectively, relative to the membrane normal. The apparent coordination number was consistently larger (ca. 0.8) for the 2.72 A shell than it was for the 2.87 A shell. The larger contribution from the shorter Mn-Mn shell may account for the surprising result that the single shell fits gave a 2.71 ~ distance, not an average (ca. 2.8 A) distance. One interpretation of these data is that NH 3 has replaced an oxo bridge in one of the Mn(p-O)zMn dimers. This would explain both the ESEEM resuks (above) and the increase in the Mn-Mn distance. In addition, the ability to now
28
]ames E. Penner-Hahn
distinguish two different Mn-Mn distances in the polarized measurements supports the conclusion (above) that there are two different Mn dimers. 4.2 Modifications at the Chloride Site
Chloride is essential for full OEC activity [3]. Recent data suggest a single C1- is bound at the OEC, in one of two conformations [110]. In intact centers, C1- is bound in a tight-binding, slowly-exchanging site. In C1--free centers (i. e. once C1- has been removed), rebinding initially occurs at a low-affinity, fast-exchanging site which subsequently converts to a tight-binding site. Based on the correlation of 02 evolution rate with light intensity, it appears that the OEC is active in the absence of CI-, but at only 35 % of the rate seen for control samples [110]. In principle, it might be possible to use EXAFS to prove (or disprove) ligation of C1 to Mn. In reality, however, this is not possible using Mn EXAFS with the presently achievable data quality [30], although it may be possible to answer this question using CI (or Br) EXAFS. The best evidence that C1- is bound close to, if not necessarily on, an Mn atom, is the recent observation that replacement of C1- with F- causes a change in the Mn-Mn shell for the S2 state [111]. The change induced by F- is similar to, but much smaller than, the damping induced by NH3 (see Sect. 4.1). This can again be modeled using either a single shell of Mn with an increased Debye-Waller factor, or two shells of Mn separated by ca. 0.12 A. As with the NH3-inhibited samples, the two shell fits were generally worse than one shell fits (using a goodness-of-fit parameter corrected for the number of variable parameters), although marginally improved two shell fits could be obtained by constraining the Mn-Mn coordination numbers. A final similarity to the NH3-inhibited samples is that, in unconstrained fits, the apparent Mn-Mn coordination number for the longer shell is approximately half as large as that for the shorter shell. Polarized measurements to confirm the presence of two shells have not been reported for these samples. 4.3 Reduced Derivatives of the OEC
The OEC shows a two-flash delay in production of oxygen after treatment with potential substrate analogs such as NH2OH, N2H4, or H202 [112-114]. The delay is consistent either with a two-electron reduction of the Mn cluster to form an "S_I" state (see [ 115, 116] and references therein) or with formation of a state such as $1" NH2OH, where the bound NHzOH serves to reduce a higher S state (e.g., $2) following photooxidation [117, 118]. Initial XANES measurements suggested that NH2OH-treated samples were not reduced in the dark [76]. More recently, we have reported XANES data consistent with reduction of the Mn in the dark, and reoxidation to $I on exposure to light [70, 74]. The data in the two reports are, in fact, very similar. The different conclusions depend on whether the observed edge shifts are attributed to reduction of Mn to S_1 or to production of inactive Mn(II). In the latter study, the NHzOH effect was photoreversi-
Structural Characterization of the Mn Site in the PhotosyntheticOxygen-EvolvingComplex
29
ble, demonstrating that there was no significant decomposition, and suggesting that the observed edge shifts represent reduction of Mn by NH2OH in the dark. Regardless of the kinetic scheme (above) that is used to explain the two-flash NH2OH delay, the state produced by single turnover illumination should be So (or, more properly, S*, since this is not necessarily identical to the normal So). We found that NH2OH reduced the OEC to Mn(III) in S_~,as judged both by XANES [70] and by the lack of change in the 2.7 ~ Mn-Mn interactions on reduction [74]. Reduction to Mn(II) would be expected to cause loss of one of the MnMn interactions. An Mn(III) 4 oxidation state for S_1 implies an Mn(III)3Mn(IV) oxidation state for S0. In contrast, Guiles et al. found [76] evidence for Mn(II) when their NHaOH-treated samples were illuminated. This has been taken as evidence for Mn(II) in So, a conclusion that would be consistent with some [29] but not all [26] of the flash measurements of So. In addition to NH2OH, larger reductants, such as hydroquinone (H2Q), can also reduce the OEC. The latter gives a form of the OEC that contains ca. 50 % Mn(II), as judged by a six-line EPR signal, but that is nevertheless completely photoreversible, thus demonstrating that no Mn has been lost [119]. The Mn(II) produced by H/Q is not EDTA-extractable and does not cause enhancement of the solvent 1H-NMR relaxation, leading to the conclusion that this Mn(II) is sequestered near its active site [119]. Hydroquinone reduction gives a significant
Site A Low Potential Sl
Mrl~ !
MnIII O~'"~2QO~~ J"Mnll .... Mnll 1
n'¢ n'V
Site B
High Potential NH2OH(fast)/ N2H4 (siow)~ "So: ~0~, ~." •" " I Mnlll MnTM
S-1
I.o. ....
"~Of
~1/ Fast Mnll O~. S_ 1* MnIv MnlV ~,O d ~ NHpOH (fast)
Mnll
I ~04 I --O~
Mn~,
~ n Ill
O
HzOH
M nlP-----Mnll
"Mnl!
I
s_z I .o. Mn III
Mnlll
"~ 0 f'
NH20H
..........
Mnl1
) Mn release
~) ,Mnl~......~_~MnIU
S-s (unstable) Fig.10. Schemethat could account for the effectsof different reductants on the OEC structure within the "dimer of dimers" model. Alternate schemes based on different nuclearity would also be consistent. The connection between the two dimers is drawn for consistency with EPR data, but is not required by X-ray absorption measurements. Redrawn with permission from [74]
30
lames E. Penner-Hahn
decrease in XANES edge energy that is best modeled [70] by ca. 50% Mn(II), consistent with the EPR. As expected, the EXAFS spectra for H2Q-reduced samples show major structural changes [74], consistent with loss of the Mn-Mn interaction as a result of reduction of one Mn(p-O)zMn unit. The NHzOH and H2Q treatments show strong synergism in their ability to attack the OEC [119]. This, together with the EXAFS results, suggested [74] the two-site model shown in Fig. 10. Although the scheme is drawn using a dimer of dimers model, other arrangements of Mn would be consistent with observations. In this scheme, large reductants (e.g. H2Q) are only able to attack one site (A in Fig. 10). NH2OH preferentially attacks at a different site (B in Fig. 10) but will also slowly attack site A. When both A and B are reduced, the OEC is unstable and loses Mn to solution, thus accounting for the inactivation that accompanies long-term NH2OH incubation or simultaneous NH2OH + H2Q incubation. In this model, the structural change that accompanies H2Q reduction blocks the internal electron transfer (dashed line) that would normally prevent the production of different products when different reducing agents are used [74]. The significance of the two-site reduction model is that it suggests that, at least in the $1 and lower states, the OEC Mn is organized into two functionally distinct units. This would be consistent with the report (see Sect. 3.1.4) of two magnetically distinct Mn sites. If this model is correct and if it applies also to the higher oxidation states, it would suggest a two-step mechanism for water oxidation.
5 Structural Possibilities A wide variety of models for the OEC have been proposed at various times (see [2, 4] for discussion of possible structural models). Initial efforts centered on high-symmetry models, many of which have subsequently been shown [4, 79] to be inconsistent with the EXAFS data. A model that has dominated much of the recent discussion of the OEC has been the "dimer of dimers" model [77]. This contains two Mn (la-O)zMn units with 2.7 A Mn-Mn distances separated by a single 3.3 ]l oxo-bridged Mn-Mn, to give a C-shaped molecule. This is consistent with the EXAFS for S~, with the observation of two different Mn-Mn distances in inhibited samples and with suggestions (from reductant studies) that there are two separate sites of action. As discussed in Section 3.1.2, a dimer of dimers magnetic model can give an excellent fit to several different multiline EPR spectra [51]. The difficulty with the dimer of dimers model is that it is only possible to reproduce the EPR data by using magnetic couplings and Mn oxidation states that appear to be inconsistent with the EXAFS and XANES data. This incompatibility suggests that the dimer of dimers model requires modification in order to be consistent with both the EXAFS and the EPR. The goal of this section is to identify the key experimental observations that define the structure and to suggest ways in which existing models might be altered to be consistent with most, if not all, of the available data. At the outset, it is important to recall that EXAFS gives only average structural information. The consensus in the OEC field is that 4 Mn and 1 Ca are the
StructuralCharacterizationof the Mn Site in the PhotosyntheticOxygen-EvolvingComplex
31
metals present in the OEC [2] and this assumption is used in all of the interpretations of the EXAFS. It has been suggested [13], although not generally accepted (see [2] for a summary of the arguments) that there are more than 4 Mn. If there were more Mn, then the interpretation of the EXAFS coordination numbers would be different: An apparent coordination number of 1.2 - 1.3 with 6 Mn would represent 4 Mn-Mn interactions at 2.7/~. Similarly, contributions from metals other than Mn or Ca would not be distinguishable by EXAFS, but would alter significantly the interpretations of the EPR. One way to reconcile the EXAFS and EPR results would be to incorporate additional metals. Certainly, the history of bioinorganic chemistry contains many examples of enzymes whose true metal stoichiometry is larger than initially believed. However, it is premature to conclude that additional metals must be invoked in order to explain the observed data. The questions to be considered are: Does the OEC contain one or two Mn clusters? Is there an arrangement of Mn ions that would be consistent with both the EXAFS and the EPR? What are the Mn oxidation states? 5.1 One vs Two Clusters
At various times, the possibility of two different Mn clusters has been suggested. The presence of two dusters would be consistent with (although not proof of) a mechanism in which one cluster oxidizes water to H202, or a similar species, while the second cluster oxidizes H202 to Oz. It is thus important to establish whether two clusters are present. The key piece of evidence suggesting that there could be two independent clusters in the OEC is the observation that the $1 EPR signal converts to the multiline signal, but is uncorrelated to production of the g = 4.1 signal [56]. This observation requires that the g = 4.1 and multiline signals come from magnetically distinct species. The presence of two clusters would provide a ready explanation for the observed synergism between hydroquinone and NHaOH [119] and for the production of different reduced species [74]. However, these observations can also be explained within a single-cluster model. Similarly, the finding that it is possible to prepare both ground state and excited state g = 4.1 forms of Sa has been interpreted as evidence for two distinct clusters [39], but could also be consistent with multiple conformations of a single cluster. The principal weakness in the two-cluster model is that no additional reports of an $1 EPR signal have appeared. While this does not prove that this observation is incorrect, it raises the question of its reproducibility. The case for a single Mn cluster is based on two EPR observations: 1) The finding that, if the 55Mn ENDOR data are used to constrain the Mn parameters, the breadth of the multiline EPR signal can only be explained using at least 3 and probably 4 Mn ions [51]; and 2) The observation of multiline structure in the g = 4.1 signal, demonstrating that this signal must come from a cluster containing at least 3 Mn [53]. Thus, the g = 4.1 and multiline signals could only come from two different clusters if the OEC contained a minimum of 6 Mn ions. The discovery of a near-IR conversion from the multiline to the g = 4.1 forms of
32
James E. Penner-Hahn
Sa [55] removes any need to invoke two clusters (with the g ~ 4.1 signal coming from oxidation of an intermediate cluster). This does not, however, exclude the possibility of two clusters (see Fig. 6). The principal weakness in the singlecluster model is that mukiline character has only been observed for oriented, NH3-inhibited samples. This leaves open the possibility that the g ~ 4.1 signal in NH3-inhibited samples arises from a different source than the other g -- 4.1 signals (e. g., with NH3 bridging between the two putative clusters). Certainly, the structural changes observed on NH 3 inhibition [88] are larger than those seen for other inhibited samples. If there were two clusters, they would presumably be arranged as a monomer (giving the non-mukiline g ~ 4.1 signal) and a trimer (giving the multiline signals), since independent dimers appear inconsistent with the ENDOR data for the multiline form [48]. In summary, the data can probably be stretched to accommodate a twocluster model. The only reason to do so, however, is the S~ EPR signal. It is important to confirm or correct this report.
Note added in proof: A very recent report [120] confirms the presence of an $1 EPR signal that converts to the multiline $2 signal but not to the g ~ 4.1 $2 signal. The S~ signal arises from an excite state that is 2.5 K above the S = 0 ground state. 5.2 Arrangements of Mn Ions
It appears that the dimer of dimers scheme cannot account for both the EXAFS and EPR using a single structural model (i.e., oxo bridges for the EXAFS, hydroxo bridges for the EPR, as described above). One obvious way to modify this model would be to add a third Mn-Mn interaction at 2.7 A. This would give, for example, a strongly coupled triangle of Mn-Mn, perhaps with both p2-O and p3-O bridges. The spin-frustration that would be expected for such an arrangement might account for the small energy differences between the different OEC conformations that are seen in the EPR. The presence of three rather than two Mn-Mn interactions would explain the EXAFS coordination numbers. The presence of p3-0xo ligands, which typically have longer Mn-O bond lengths than pz-OXO, would explain both the disorder in the first shell EXAFS and the relatively long Mn-Mn distance (2.72 A for the OEC, vs 2.68 It for Mn catalase [81]). Finally, the presence of three Mn-Mn interactions, one of which is elongated in inhibited samples, would explain the apparent coordination numbers for inhibited samples and the anisotropy in Mn-Mn distances that was seen for S~. 5.3 Mn Oxidation States
If Mn is oxidized in each S state transition, then the mechanism of water oxidation can rely on conventional Mn redox chemistry. In contrast, if a non-manganese radical is produced in the higher S-state transitions, then this will have important mechanistic implications. Several models have been proposed in
Structural Characterization of the Mn Site in the Photosynthetic Oxygen-Evolving Complex
33
which a non-Mn radical plays a crucial role in the mechanism [4]. It is therefore important to evaluate the evidence for non-Mn radicals. The original suggestion that Mn is not oxidized in the $2~$3 transition came from XANES data [28]. This conclusion is inconsistent with many of the subsequent XANES measurements. Moreover, it is not clear that even the original data were necessarily proof that Mn was not oxidized, since, based on model studies, it seems possible that th e $1-~ Sa transition is anomalously large, thus making the apparent $2--~S3 energy difference appear small. The second line of evidence that Mn is not oxidized on $2--~$3 was the observation of a radical signal in the EPR. If an organic radical was produced on $2-4 S3, then Mn could not be oxidized in this transition. With the finding that the "$3 radical" is actually Yz" [68], this evidence against Mn oxidation disappears. It thus appears most likely that Mn is oxidized in all four S state transitions. A second redox question concerns the oxidation state of S0. It has been suggested [4] that one Mn dimer is redox-inert, with all of the water oxidation chemistry taking place on a second Mn dimer. The presence of Mn(II) in SO is consistent with this proposal [29]. However, if it is correct that the S_~ produced by NH2OH contains all Mn(III) [74], this would imply that at least 3, and probably all 4 of the Mn are redox-active (given $2 = Mn(III)Mn(IV)3; $3 = Mn(IV)4). As discussed above, these different conclusions may be related either to the difficulty of deconvoluting So in flash experiments or to the question of whether S_~truly represents a physiologically relevant state. Experiments of this sort should eventually clarify the role of Mn oxidation in water oxidation.
6 Conclusions and Future Prospects Despite a great deal of effort over many years, the detailed structure of the OEC remains elusive. Numerous models have been suggested and, as experimental methods have improved, most of these models have been shown to require modification. As the set of incorrect structures expands, the possibilities for the true structure become increasingly limited, and thus better defined. Recent EXAFS and EPR data appear to be inconsistent with the currently popular dimer-of-dimers model. Modifications based around triangular clusters may be one way to improve agreement with the available data. Acknowledgments. Supported in part by the NIH (GM-45205). I thank Profs. Charles Yocum, Vince Pecoraro, and Gerry Babcock for useful discussions.
References 1. Ghanotakis DF, Yocum CF (1990) Annu Rev Plant Physiol Plant Mol Biol 2. Debus RJ (1992) BBA 1102:269 3. Yocum CF (1992) In: V.L. Pecoraro (eds) Manganese Redox Enzymes VCH Publishers, New York, p 4. Yachandra VK, Sauer K, Klein MP (1996) Chem Rev (Washington, D C 96:2927 5. Deisenhofer J, Michel H (1991) Annu Rev Biophys Biophys Chem 20:247
34 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46. 47. 48. 49.
James E. Penner-Hahn Joliot P, Barbieri G, Chabaud R (1969) Photochem Photobiol 10:309 Kok B, Forbush B, McGloin M (1970) Photochem Photobiol 11:457 Shinkarev VP, Wraight CA (1993) Photosynth Res 38: 315 Shinkarev VP, Wraight CA (•993) Proc Nail Acad Sci U S A 90:1834 Berthold DA, Babcock GT, Yocum CF (1981) FEBS Letters 134:231 Kuwabara T, Murata N (1982) Plant Cell Physio123: 533 Ghanotakis DF, Demetriou DM, Yocum CF (1987) Biochimica et Biophysica Acta 891:15 Pauly S, Witt HT (1992) BBA 1099:211 Lee PA, Citrin PH, Eisenberger P, Kincaid BM (1981) Rev Mod Plays 53:769 Scott RA (1985) Methods Enzymol 117:414 Teo BK (1986) EXAFS: Basic Principles and Data Analysis, Springer-Verlag, New York Rehr JJ, Mustre dLJ, Zabinsky SI, Albers RC (1991) J Am Chem Soc 113:5135 Binsted N, Hasnain SS (1996) J Synchr Rad 3:185 Kirby JA, Goodin JA, Wydrynski T, Robertson AS, Klein MP (1981) J Am Chem Soc 103: 5537 Bart JCJ (1986) Adv Catal 34:203 Shulman RG, Yafet Y, Eisenberger P, Blumberg WE (1976) Proc Nail Acad Sci USA 73:1384 Wong J, Lyfle FW, Messmer RP, Maylotte DH (1984) Plays Rev B30: 5596 Rehr JJ, Albers RC, Zabinsky SI (1992) Phys Rev Lett 69:3397 Natoli CR (1983) In: A Bianconi, L Incoccia, and S Stipcich (eds) Springer Series in Chemical Physics Springer-Verlag, Berlin, p 43. Cramer SP, Eccles TK, Kutzler FW, Hodgson KO, Mortenson LE (1976) J Am Chem Soc 98:1287 Ono T-a, Noguchi T, Inoue Y, Kusunoki M, Matsushita T, Oyanagi H (1992) Science 258:1335 Ono T-a, Noguchi T, Inoue Y, Kusunoki M, Yamaguchi H, Oyanagi H (1995) J Am Chem Soc 117:6386 Guiles RD, Zimmermann JL, McDermott AE, Yachandra VK, Cole JL, Dexheimer SL, Britt RD, Wieghardt K, Bossek U, Sauer K, Klein MP (1990) Biochemistry 29:471 Roelofs TA, Liang W, Latimer MJ, Cinco RM, Rompel A, Andrews JC, Sauer K, Yachandra VK, Klein M (1996) Proc Nail Acad Sci USA 93:3335 Penner-Hahn JE, Fronko RM, Pecoraro VL, Yocum CF, Betts SD, Bowlby NR (1990) J Am Chem Soc 112: 2549 Riggs-Gelasco PJ, Stemmler TL, Penner-Hahn JE (1995) Coord Chem Rev 144:245 Lyfle FW, Sayers DE, Stern EA (1989) Physica B 158:701 Bunker G, Hasnain S, Sayers D (1991) In: S. Samar Hasnain (eds) X-ray Absorption Fine Structure Ellis Horwood, New York, p 751. Dismukes GC, Siderer Y (1980) FEBS Lett 121:78 Rutherford AW, Bouss ac A, Zimmermann JL ( 1991) New J Chem 15: 491 Miller A-F, Brudvig GW (1991) Biochim Biophys Acta 1056:1 Zimmermann JL, Rutherford AW (1984) Biochim Biophys Acta 767:160 Casey JL, Sauer K (1984) Biochim Biophys Acta 767:21 Smith PJ, Pace RJ (1996) Biochim Biophys Acta 1275:213 Beck WF, De Paula JC, Brudvig GW (1986) J Am Chem Soc 108:4018 Boussac A, Rutherford AW (1988) Biochemistry 27:3476 Boussac A, Zimmermann JL, Rutherford AW (1989) Biochemistry 28: 8984 Sivaraja M, Tso J, Dismukes GC (1989) Biochemistry 28:9459 Pace RJ, Smith P, Bramley R, Stehlik D (1991) Biochim Biophys Acta 1058:161 Zheng M, Dismukes GC (1992) Res Photosynth, Proc Int Congr Photosynth, 9th 2: Bonvoisin J, Bloudin G, Girerd JJ, Zimmermann JL (1992) Biophys J 61 : 1076 Ahrling KA, Pace RJ (1995) Biophys J 68:2081 Randall DW, Sturgeon BE, Ball JA, Lorigan GA, Chan MK, Klein MP, Armstrong WH, Britt RD (1995) J Am Chem Soc 117:11780 Lorigan GA, Britt RD (1994) Biochemistry 33:12072
Structural Characterization of the Mn Site in the Photosynthetic Oxygen-Evolving Complex
50. 51. 52. 53. 54. 55. 56. 57. 58. 59. 60. 61. 62. 63. 64. 65. 66. 67. 68. 69. 70. 71. 72. 73. 74. 75. 76. 77. 78. 79. 80. 81. 82. 83. 84. 85. 86. 87. 88. 89.
35
Hansson (),Aasa R,V~inng~trd T (1987) Biophys J 1987:825 Zheng M, Dismukes GC (1996) Inorg Chem 35:3307 Zheng M, Khangulov SV, Dismukes GC, Barynin VV (1994) Inorg Chem 33: Kim DH, Britt RD, Klein MP, Sauer K (1990) ] Am Chem Soc 112:9389 Haddy A, Dunham WR, Sands RH, Aasa R (1992) Biochim Biophys Acta 1099:25 Boussac A, Girerd J-J, Rutherford AW (1996) Biochemistry 35: 6984 Dexheimer SL, Klein MP (1992) ] Am Chem Soc 114:2821 Chan MK, Armstrong WH (1991) J Am Chem Soc 113:5055 Armstrong WH (1992) In: V.L. Fecoraro (eds) Manganese Redox Enzymes VCH Publishers, New York, p 261 de Paula ]C, Li PM, Miller A-F, Wu BW, Brudvig GW (1986) Biochemistry 25:6487 Koulougliotis D, Hirsh DJ, Brudvig GW (1992) J Am Chem Soc 114:8322 Andreasson LE, Lindberg K (1992) Biochim Biophys Acta 1100:177 MacLachlan DJ, Nugent JHA (1993) Biochemistry 32:9772 Baumgarten M, Philo IS, Dismukes GC (1990) Biochemistry 29:10814 SzalaiVA, Brudvig GW (1996) Biochemistry 35:1946 Zimmermann JL, Boussac A, Rutherford AW (1993) Biochemistry 32:4831 Boussac A, Zimmermann J-L, Rutherford AW, Lavergne J (1990) Nature 347:303 Hallahan BJ, Nugent JHA, Warden JT, Evans MCW (1992) Biochemistry 31:4562 Tang XS, Randall DW, Force DA, Diner BA, Britt RD (1996) ] Am Chem Soc 118:7638 Gilchrist ML, Ball JA, Randall DW, Britt RD (1995) Proc Natl Acad Sci, USA 92:9545 Riggs P, J., Yocum CF, Penner-Hahn JE, Mei R (1992) J Am Chem Soc 114:10650 Kirby ]A, Robertson JP, Smith AC, Cooper SR, Klein MP (1981) J Am Chem Soc 103: 5529 Britt RD, Tang XS, Gilchrist ML, Lorigan GA, Larsen BS, Diner BA (1994) Biochem Soc Trans 22: 343 Tang XS, Diner BA, Larsen BS, Gilchrist MLJ, Lorigan GA, Britt RD (1994) Proc Natl Acad Sci USA 91:704 Riggs-Gelasco PJ, Mei R, Yocum CF, Penner-Hahn ]E (1996) J Am Chem Soc 118:2387 Yachandra VK, Guiles RD, McDermott A, Britt RD, Dexheimer SL, Sauer K, Klein MP (1986) Biochim Biophys Acta 850:324 Guiles RD, Yachandra VK, McDermott AE, Cole JL, Dexheimer SL, Britt RD, Sauer K, Klein MP (1990) Biochemistry 29: 486 Yachandra VK, DeRose VJ, Latimer MJ, Mukerji I, Sauer K, Klein MP (1993) Science 260: 675 MacLachlan DJ, Hallahan BJ, Ruffle SV, Nugent JHA, Evans MCW, Strange RW, Hasnain SS (1992) Biochem J 285:569 DeRose VJ, Mukerji I, Latimer M], Yachandra VK, Sauer K, Klein MP (1994) J Am Chem Soc 116:5239 Penner-Hahn JE, Fronko RM, Waldo GS, Yocum CF, Bowlby NR, Betts SD (1990) Curt Res Photosynth, Proc Int Conf. Photosynth, 8th 1: Waldo GS, Yu S, Penner HJE (1992) ] Am Chem Soc 114:5869 Riggs-Gelasco PJ (1995) PhD thesis, University of Michigan Larson EJ, Riggs PJ, Penner-Hahn JE, Pecoraro VL (1992) J Chem Soc, Chem Commun 102 Baldwin MJ, Stemmler TL, Riggs GPJ, Kirk ML, Penner-Hahn JE, Pecoraro VL (1994) ] Am Chem Soc 116:11349 George GN, Prince RC, Frey TG, Cramer SP (1989) Physica B 158:81 Mukerji I, Andrews JC, DeRose VJ, Latimer MJ, Yachandra VK, Sauer K, Klein MP (1994) Biochemistry 33: 9712 Riggs-Gelasco P,Mei R, Ghanotakis D, Yocum CF, Penner-Hahn JE (1996) J Am Chem Soc 118: 2400 Dau H, Andrews JC, Roelofs TA, Latimer MJ, Liang W, Yachandra VK, Sauer K, Klein MP (1995) Biochemistry 34: 5274 Guiles RD, Yachandra VK, McDermott AE, Britt RD, Dexheimer SL, Sauer K, Klein MP (1987) Prog Photosynth Res, Proc Int Congr Photosynth, 7th 1:
36
James E. Penner-Hahn
90. George GN, Prince RC, Cramer SP (1989) Science 243:789 91. MacLachlan D], Nugent JHA, Bratt P], Evans MCW (1994) Biochim Biophys Acta 1186:186 92. Liang W, Latimer MJ, Dau H, Roelofs TA, Yachandra VK, Sauer K, Klein MP (1994) Biochemistry 33:4923 93. Latimer MJ, DeRose VJ, Mukerji I,Yachandra VK, Sauer K, Klein MP (1995) Biochemistry 34:10898 94. )~delroth P, Lindberg K, Andreasson LE (1995) Biochemistry 34:9021 95. Seidler A, Rutherford AW (1996) Biochemistry 35:12104 96. Bakou A, Ghanotakis DF (1993) Biochim Biophys Acta 1141:303 97. Goodin DB, Yachandra VK, Britt RD, Sauer K, Klein M (1984) Biochimica Biophysica Acta 767:209 98. Yachandra VK, Guiles RD, McDermott AE, Cole JL, Britt RD, Dexheimer SL, Sauer K, Klein MP (1987) Biochemistry 26: 5974 99. Cole ],Yachandra VK, Guiles RD, McDermott AE, Britt RD, Dexheimer SL, Sauer K, Klein MP (1987) Biochim Biophys Acta 890:395 100. MacLachlan DJ, Nugent JHA, Evans MCW (1994) Biochim Biophys Acta 1185:103 101. Kusunoki M, Ono T, Noguchi T, Inoue Y, Oyanagi H (1993) Photosynth Res 38:331 102. Ono Ta, Noguchi T, Inoue Y, Kusunoki M, Yamaguchi H, Oyanagi H (1993) FEBS Lett 330: 28 103. Sharp RR (1992) In: VL Pecoraro (eds) Manganese Redox Enzymes. VCH Publishers, New York, p 177 104. Bvelo RG, Styring S, Rutherford AW, HoffA] (1989) Biochim Biophys Acta 973:428 105. Styring SA, Rutherford AW (1988) Biochemistry 27:4915 106. Dekker JP (1992) In: V.L. Pecoraro (eds) Manganese Redox Enzymes VCH Publishers, New York, p 85 107. Liang W, Roelofs TA, Olsen GT, Latimer MJ, Cinco RM, Rompel A, Sauer K, Yachandra VK, Klein MP (1995) In: P. Mathis (eds) Photosynthesis: From Light to Biosphere Kluwer, Netherlands, p 413. 108. Sandusky PO, Yocum CF (1986) Biochim Biophys Acta 849:85 109. Britt RD, Zimmerman J-L, Sauer K, Klein MP (1989) J Am Chem Soc 111:3522 110. Lindberg K, Andreasson LE (1996) Biochemistry 35:14259 111. DeRose VJ, Latimer MJ, Zimmermann JL, Mukerji I, Yachandra VK, Sauer K, Klein MP (1995) Chem Phys 194:443 112. Bouges B (1971) Biochim Biophys Acta 234:103 113. Renger G, Bader KP, Schmid GH (1990) Biochimica et Biophysica Acta 1015:288 114. Mano J, Takahashi M-a, Asada K (1987) Biochemistry 26: 2495 115. Messinger J, Renger, G. (1993) Biochemistry 32:9379 116. Kretschmann H, Pauly S, Witt HT (1991) Biochim Biophys Acta 1059:208 117. Radmer R (1981) Biochimica et Biophysica Acta 637:80 118. Radmer R, Ollinger O (1982) FEBS Letters 144:162 119. Mei R, Yocum CF (1992) Biochemistry 31:8449 120. Yamauchi T, Mino H, Matsukawa T, Kawamori A, Ono T-a (1997) Biochemistry 35:7520
Metal Sites in Small Blue Copper Proteins, Blue Copper Oxidases and Vanadium-ContainingEnzymes Albrecht Messerschmidt Max-Planck-Institut ffir Biochemie, Am Klopferspitz 18A, D-82152 Martinsried
E-mail:
[email protected] The coordination geometries of metal sites in cupredoxins, mutants and metal derivatives of cupredoxins, multi-copper oxidases and a vanadium-containing chloroperoxidase as derived from X-ray crystallography are described. Correlations with their spectroscopic, electrochemical, electron transfer and catalytic properties are discussed. X-ray crystallography,EPR and Resonance Raman spectroscopy of copper sites in cupredoxins and mutants have led to a classification ranging from type 1 trigonal, type 1 distorted tetrahedral, type 1.5 to type 2. The mutation of copper ligands in azurin or amino acids close to the copper site changes the redox potential in a range of_+140 mV, only. The high redox potential of rusticyanin of 680 mV (azurin, 380 mV) should be mainly due to the special protein environment of the copper site (high proportion of hydrophobic residues). The type 1 and trinuclear copper centres of the multi-copper oxidases ascorbate oxidase, laccase and ceruloplasmin are presented. The metal sites of type 2 depleted, fully-reduced, peroxide and azide forms of ascorbate oxidase, as determined by X-ray crystallography, are discussed in terms of the mechanistic properties of these enzymes. The first X-ray structure of a vanadium-containing protein, namely of a chloroperoxidase from the fungus Curvularia inaequalis, is briefly discussed. The protein fold is mainly a-helical with two four-helix bundles. In the X-ray structure, which is an azide:enzyme complex, the vanadium exhibits a simple unexpected coordination geometry, namely, a trigonal bipyramidal coordination with three non-protein oxygen ligands (VO3 group), one nitrogen ligand from a histidine and one nitrogen from the exogenous azide ligand.
Keywords:protein crystallography, small blue copper proteins, multi-copper oxidases, vanadium-containing enzyme, electron transfer
List of Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . 1
Metal Sites in Small Blue Copper Proteins
38 ..............
39
1.3 1.4
Introduction ................................ Spatial S t r u c t u r e s of C u p r e d o x i n s a n d Mutants . . . . . . . . . . . The C u p r e d o x i n Fold . . . . . . . . . . . . . . . . . . . . . . . . . . . G e o m e t r i e s o f C u p r e d o x i n C o p p e r Sites . . . . . . . . . . . . . . . . S u b s t i t u t i o n o f C o p p e r b y Hg, Zn, C d a n d Ni . . . . . . . . . . . . . The CuA D i n u c l e a r C o p p e r Site . . . . . . . . . . . . . . . . . . . . . Spectroscopy of Cupredoxins ...................... Redox Potentials a n d Electron Transfer P r o p e r t i e s . . . . . . . . .
39 39 39 40 45 47 48 50
2
Metal Sites in Multi-Copper Oxidases . . . . . . . . . . . . . . . . .
52
2.1 2.1.1
The Trinuclear C o p p e r Active Site . . . . . . . . . . . . . . . . . . . Native O x i d i z e d E n z y m e . . . . . . . . . . . . . . . . . . . . . . . . .
52 52
1.1 1.2 1.2.1 1.2.2 1.2.3
1.2.4
Structure and Bonding, Vol. 90 © Springer Verlag Berlin Heidelberg 1998
38 2.1.2 2.1.3 2.1.4 2.1.5 2.2
Albrecht Messerschmidt Type-2 Depleted (T2D) Form of Ascorbate Oxidase . . . . . . . . . Fully-Reduced Form of Ascorbate Oxidase . . . . . . . . . . . . . . Peroxide Form of Ascorbate Oxidase . . . . . . . . . . . . . . . . . . Azide Form of Ascorbate Oxidase . . . . . . . . . . . . . . . . . . . Aspects of Electron Transfer and Mechanistic Properties . . . . . . . .
55 56 57 60 61
First X-ray Structure of a Vanadium-Containing Enzyme: Chloroperoxidase from the Fungus C u r v u l a r i a inaequalis
62
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
....
65
List of Abbreviations AzPwt AzADwt AzAXwt AzPH35Q AzPH35L AzPH35F AzADM121Q AzADM121H AzPwt: Zn AzADwt red AzPF 114A, AzPW48M: Ni AzPN47D: Zn AzPI7S AzPFll0S PcP PcE PcP: Hg ~p-AzAF AmPD AmTV CBP Scc AzAD: Cd NiR
AO TIHg
wild-type azurin from Pseudomonas aeruginosa (PA) [20, 21] wild-type azurin from Alcaligenes denitrificans (AD) [22] wild-type azurin from Alcaligenes xylosoxidans [23] azurin from PA, H35Q mutant [24] azurin from PA, H35L mutant [24] azurin from PA, H35F mutant [21] azurin from AD, M121Q mutant [25] azurin from AD, M121H mutant [26] wild-type azurin from PA containing Zn [27] wild-type azurin from AD, reduced form [28] azurin from PA, F114A mutant [29] azurin from PA, W48 M mutant containing Ni [30] azurin from PA, N47D mutant containing Zn [31] azurin from PA, I7S mutant [32] azurin from PA, F110S mutant [32] plastocyanin from Populus nigra [33, 34] plastocyanin from Enteromorpha prolifera [35] plastocyanin from Populus nigra, Hg substituted [36] pseudoazurin from Alcaligenesfaecalis [37, 38] amicyanin from Paracoccus denitrificans [39] amicyanin from Thiobacillus versutus [40] cucumber basic protein [41] cucumber stellacyanin [6] azurin from AD containing cadmium [42] copper-containing nitrite reductase from Achromobacter cycloclastes [15] ascorbate oxidase from Cucurbita pepo medullosa [43] mercury substituted type 1 site
Metal Sitesin Small BlueCopperProteins,BlueCopperOxidasesand Vanadium-ContainingEnzymes
39
1 Metal Sites in Small Blue Copper Proteins 1.1
Introduction
The unique properties of the small blue copper proteins have fascinated chemists for a long time. These proteins contain a mononuclear copper site with three characteristic properties: (1) an intense blue colour, ~ -600 nm, with absorption coefficients of 2000-6000 M-1 cm -1, due to an S(Cys)--~Cu(II) charge transfer; (2) an unusually narrow hyperfine coupling (All values of 0.0035-0.0063 cm -1) in the EPR spectrum of the Cu(II) protein; and (3) high reduction potentials (range 184-680 mV) as compared with the aqua Cu(II/I) couple (115 mV). This copper site has been classified as a type 1 copper site. The small blue copper proteins function as electron transfer proteins in such fundamental processes as photosynthesis and respiration. They have been termed cupredoxins to emphasise their common origin and their role as electron mediators in analogy to the iron-containing ferredoxins [1]. The research on cupredoxins has been summarised in several reviews in the past (see [2-4], for example). During the last few years cupredoxin research has benefited from the application of genetic engineering techniques in combination with modern spectroscopic techniques, X-ray crystallography, electrochemistry, kinetics and quantum theory. The main genetic engineering work has been done on azurins from Pseudomonas aeruginosa (PA) and Alcaligenes denitrificans (AD) in Leiden by Canters and co-workers (see [4], for example) and in G6teborg by V~nngard, Malmstr6m, Lundberg and colleagues (see [5], for example). Mutations were introduced in the direct copper ligand sphere and in the second coordination sphere to test their influence on the properties of the metal site. Mutations in different parts of the molecule have been introduced to probe their influence on other molecular properties. The first part of this article will focus only on new cupredoxin spatial structures, mutants of cupredoxins which influence the properties of the metal site, as well as the relation of the structure of the metal sites to their physicochemical properties. 1.2 Spatial Structures of Cupredoxins and Mutants 1.2.1 The CupredoxinFold
Cupredoxins fold into a single domain consisting mainly of a fl-sandwich or flbarrel. This fl-sandwich may comprise 6 to 13 fl-strands. The arrangement of the fl-strands is basically antiparallel showing the Greek-key motif. A typical cupredoxin fold represented by azurin is depicted in Fig. 1. In this case, the flsandwich consists of 8 fl-strands. The fl-sandwich of stellacyanin from cucumber, whose X-ray structure has been determined very recently, has only 6 flstrands [6]. This stellacyanin belongs to a subgroup of cupredoxins called
40
AlbrechtMesserschmidt
phytocyanins which are plant proteins. A characteristic feature of these phytocyanins is a disulphide bridge between a cysteine in the middle part of the amino acid sequence and a cysteine adjacent to the C-terminal histidine copper ligand as well as a 6- to 7-stranded fl-sandwich. The phytocyanin cucumber basic protein, whose X-ray structure is also known, has a 7-stranded fl-sandwich [7]. On the other hand, rusticyanin from Thiobacillusferrooxidans is characterised by a compact 13-stranded fl-sandwich, as determined by multidimensional NMR techniques [8] and X-ray crystallography [8a]. This compact fl-sandwich creates a hydrophobic core particularly rich in aromatic residues. Rusticyanin has a very high redox-potential (680 mV) and operates at extremely low pH (2.0). It is assumed that the unusual acid stability of the protein results from the high degree of secondary structure and the special hydrophobic core. Subunits II in cytochrome c oxidase [9, 10] and quinol oxidase [11] exhibit a cupredoxin fold in their membrane-exposed domain. These domains contain a binuclear copper site denoted as the CUA site. The cupredoxin domains of cytochrome c oxidase have a 10-stranded fl-sandwich. In quinol oxidase it is an 11-stranded fl-sandwich. The helical content in cupredoxins varies from 0 % in amicyanin to 14% in azurin. The blue, multi-copper oxidases ascorbate oxidase, laccase and ceruloplasmin consist of three (ascorbate oxidase, laccase) and six (ceruloplasmin) domains with the cupredoxin fold (see [13], for example). The mononuclear blue copper site is located in the C-terminal domain of ascorbate oxidase and laccase. Ceruloplasmin has three mononuclear coppers bound to domains 2, 4 and 6 [14]. A copper-containing nitrite reductase from Achromobacter cycloclastes consists of two domains with a cupredoxin fold which arrange to the enzymatically active trimer (six domains) [15]. The mononuclear copper site is found in the N-terminal domain. The cupredoxins and their multidomain relatives form a large evolutionary family. This family also comprises proteins that have lost their copper-binding capability during evolution (see [13], for example). 1.2.2
Geometriesof Cupredoxin CopperSites
The type 1 copper centre is located close to the surface of the protein (Fig. 1). In native type 1 proteins the copper has four protein side-chain ligands and in some cases (e.g. azurin) a weak main-chain carbonyl oxygen as the fifth ligand. The four canonical type 1 copper ligands are His, Cys, His, Met arranged in this sequence on the polypeptide chain. The Met ligand may be a glutamine in the subgroup of phytocyanins (e.g. stellacyanin from cucumber). The Cys, His, Met ligands come from a loop between the two C-terminal strands of the fl-sandwich. They are arranged in the sequence (Cys-Xn-His-Xm-Met(Gln)) with n and m varying from one protein to the next. The first histidine sits on a strand adjacent to these, but this strand is more than 30 residues away in the N-terminal direction due to the Greek key topology of the fl-sandwich. The methionine is always sandwiched between two hydrophobic residues, two residues apart, coming from a loop between two fl-strands at the N-terminus. One or more NH...S hydrogen bonds are formed between main-chain amide nitrogens and
Metal Sites in Small Blue Copper Proteins,Blue CopperOxidasesand Vanadium-Containing Enzymes
41
Fig. 1. Ribbon-type representation of an azurin from Pseudomonas aeruginosa, a typical small blue copper protein, type 1 copper binding site and disulphide bridge are included, MOLSCRIPT[12]
the cysteine ligand. The copper centres are usually not buried by more than 8 A. The C-terminal histidine, known as northern histidine, protrudes through a more or less extensive hydrophobic surface patch, which is probably one of the surface regions through which electron transfer takes place. The type 1 copper sites can be subdivided into T1 trigonal and T1 distorted tetrahedral based on the optical, EPR and RR spectroscopy [16-18], as well as X-ray crystallography [3]. The most abundant geometry of type 1 copper sites is between T1 trigonal and T1 distorted tetrahedral and found in plastocyanin, ascorbate oxidase, amicyanin, pseudoazurin and cucumber basic protein. This unusual copper coordination is a compromise between the preferred tetrahedral Cu(I) and tetragonal Cu(II) coordination. It reduces the reorganisation energy between both redox states which helps to speed up electron transfer [19]. As already mentioned, the genetic engineering work has been mainly focused on azurin. A relatively large number of high resolution X-ray structure analyses of different azurins, azurin mutants, azurin metal derivatives and apo-azurins has been carried out including structure determinations at different pH values and/or redox states. Their copper ligand bond distances are shown in Table 1. Most of these values are derived from high-resolution X-ray structures (resolution better
42
Albrecht Messerschmidt
than 2.0 ~) and the accuracy of the metal ligand bond distances has been estimated to be about + 0.1 ]k [22]. The copper coordination in azurin is slightly different to the T1 distorted tetrahedral coordination. It is T1 trigonal, a trigonal-bipyramidal coordination with the ND 1 atoms of the two histidines and the SG atom of the cysteine in the trigonal equatorial plane, the SD atom of the methionine and a main-chain carbonyl oxygen at the apices of the bipyramid (see Fig. 2). From attempts to correlate the spectroscopic properties with the coordination geometries of the type i sites it emerged that the cysteinate copper bond length and the distance of the copper ion from the N(His)-N(His)-S(Cys) plane may play a crucial role. These out-of-plane values are indicated in Table 1 for the cases where they were given in the corresponding references. Their meaning will be discussed in the context of the spectroscopy of the type 1 sites later. The Cu-S(Cys) bond of = 2.12 .~ in both T1 subtypes is extremely short as a consequence of having only three strong ligands. The T1 trigonal sites are further characterised by long Cu-S bonds to the axial methionine (>2.8 A) and out-of-plane values in the range of 0.02 to 0.13 _A.PA azurin, wild type and some mutant structures have Cu-S(Cys) bonds that are = 0.1 A longer than the normal value of 2.12 A in the other type 1 copper proteins. This is especially surprising among the azurin structures from different sources. It may be due to differences in the amino acid sequence
Fig.2. Type-1copper site in wild-type azurin from PA,MOLSCRIPT[12]
Metal Sites in Small Blue Copper Proteins, Blue Copper Oxidasesand Vanadium-Containing Enzymes
43
Table 1. Bond distances of type 1 copper sites and metal derivatives
AzPwt, pH 5.5 AzPwt, pH 5.5, red AzPwt, pH 9.0 AzPwt, pH 9.0, red AzADwt AzADwt, red AzAXwt AzPH35Q AzPH35L AzPH35F AzPI7S AzPFll0S AzPFl14A AzADM121H, HP AzPwt: Zn AzPN47D: Zn AzAD" Cd AzPW48 M: Ni PcP PcP red, pH 7.8 PcE PcP: Hg AO ~o-AzAF ~o-AzAF red, pH 7.8 ~-AzAF red, pH 4.4 NiR AmPD AmTV CPB ScC AzADM121Q AzADM121Qred
CuN(H46)
CuS(Cl12)
CuN(Hll7)
CuS(M121)
Cu-O (G45)
out of NNSplane
2.11 2.14 2.09 2.14 2.08 2.13 2,02 2.03 2.09 1.94 2.10 2.09 2.19 2.06 2.01 2.09 2.25 2.15 1.91 2.12 1.89 2.34 2.11 2.16 2.16 2.19 2.06 1.95 2.04 1.93 1.96 1.94 1.97
2.25 2.29 2.26 2.27 2.14 2.26 2.12 2.05 2.20 2.08 2.24 2.10 2.23 2.15 2.30 2.27 2.39 2.49 2.07 2.11 2.12 2.38 2.08 2.16 2.17 2.16 2.17 2.11 2.13 2.16 2.18 2.12 2.09
2.03 2.10 2.04 2.15 2.00 2.05 2.02 2.05 2.03 2.12 1.94 1.96 2.46 2.06 2.07 2.04 2.20 2.07 2.06 2.25 2,17 2.36 2.08 2.13 2.29 3.09 2.00 2.03 2.13 1.95 2.04 2.05 2.67
3.15 3.25 3.12 3.17 3.14 3.23 3.26 3.04 3.01 3.06 3.10 3.18 3.04 2.22(N 8) 3.40 3.44 3.23 3.34 2.82 2.90 2.92 3,02 2.87 2.76 2.91 2.42 2.55 2.90 2.84 2.61 2.21(O e) 2.26(0 ~) 2.72(0 ~)
2.97 3.02 2.95 3.10 3.13 3.23 2.75 3.09 3,09 3.00 3.00 3.16 2.99 3.92 2.32 2.36 2.76 2.35 3.89 4.81 3.79 3.83 4.01 3.90 3.90 3.85 3.37 3.33
0.10 0.10 0.08 0.02 0.10 0.13 0.11 0.60 0.15 0.36 0.46 0.36 0.43 0.37
0.77 0.54 0.30 0.39 -
The resolution of the X-ray structure analyses is in most cases better than 2.0 ~..The omitting values were not given in the corresponding reference. The proteins are in the oxidized state if not indicated otherwise.
n e a r H i s 3 5 . H i s 3 5 is i n v a n d e r W a a l s c o n t a c t w i t h H i s 4 6 , a c o p p e r l i g a n d . I n PA a z u r i n a n H - b o n d is f o r m e d b e t w e e n t h e m a i n - c h a i n c a r b o n y l o f p r o l i n e 36 t o N D 1 o f H i s 3 5 . I n a z u r i n f r o m t h e Alcaligenes s p e c i e s denitrificans o r xylosoxidans the p r o l i n e is r e p l a c e d b y a v a l i n e w h i c h c a u s e s a c h a n g e i n t h e m a i n - c h a i n c o n f o r m a t i o n a t t h i s r e g i o n a n d t h e H - b o n d t o h i s t i d i n e is n o l o n g e r f o r m e d . The akered charge distribution at His35 could have an influence on the electron i c s t a t e o f t h e c o p p e r site g i v i n g r i s e t o t h e s e s m a l l a l t e r a t i o n s i n t h e c o p p e r site g e o m e t r y .
44
Albrecht Messerschmidt
The crystal structures of wild-type azurin at two pH values in both redox states have been determined [20, 21]. There are little changes in the COl~per site geometry. The bond lengths are slightly increased by about 0.05 to 0.1 A as also observed in reduced AD azurin [28], reduced poplar plastocyanin at pH 7.8 [34] and pseudoazurin from Alcaligenesfaecalis (AF) at pH 7.8 [38]. This is in line with a determination of the electronic structure of the reduced type 1 copper site [44]. In the reduced form of AF pseudoazurin at pH 4.4 the copper ion moves 0.69 A, mainly towards the SD atom of the axial methionine, and the imidazole of the northern histidine rotates by 26 ° around its CB-CG bond. The metal moves in the opposite direction with respect to its movement in the pH 7.8 reduced structure. The movements at the copper site resemble those in plastocyanin but the extent is quite different in these two cases [34]. The low-pH forms of PA azurin exhibit a pH-induced Pro36-Gly37 main-chain peptide bond flip. At the lower pH, the protonated imidazole ND1 of His35 forms a strong hydrogen bond with the carbonyl oxygen from Pro36, while at alkaline pH the deprotonated ND 1 acts as an acceptor of a weak hydrogen bond from the amide nitrogen of Gly37. In the crystal structures of the PA azurin mutants H35Q, H35L, H35F, I7S, Fll0S no remarkable influence on the copper site geometry can be noticed. This is not valid for the PA azurin mutant F114A [29]. The mutation was performed on residue Fl14, which exhibits a n-electron overlap with the copper ligand His117, to investigate its suggested role in the electron self-exchange reaction. Removal of steric constraints from the phenylalanine side chain created a somewhat different geometry around the copper site with an increased mobility of His117 resulting in an enlarged Cu-N bond length, which may be responsible for the slight differences obtained in the spectral properties of the mutant compared with the wild-type protein. Mutation of the axial methionine ligand in AD azurin to histidine generated a protein that has a green colour at higher pH values (6-7) and is blue below pH 3.8 [45]. The copper site geometry of the high-pH form (pH 7.0), as determined by X-ray crystallography [26], is shown in Fig. 3. The mutation generates a distorted tetrahedral copper coordination with an 0.6 ]~ out-of-plane value for the copper ion. The imidazole ND1 of His121 forms a strong bond with Cu, which leads to the distorted tetrahedral copper site. Copper sites that resuked from the replacement of the axial methionine by lysine, glutamic acid and histidine were found to have spectroscopic properties between those of type 1 and type 2 sites [46-48]. These sites were named type 1.5 sites [29, 42], but a clear understanding of their properties has still been lacking. The type 1 copper site of nitrite reductase from Achromobacter cycloclastes displays subtle geometric differences [15]. The bonds to the equatorial ligands of the type 1 site in oxidised nitrite reductase expand relative to those in plastocyanin with 2.17 ]k Cu-S(Cys) and 2.06 and 2.00 A Cu-N(His) bond lengths, while the axial Cu-S(Met) bond contracts to 2.55 •. Furthermore, the copper ion is raised further out of the equatorial NNS plane (0.54 A), and angular changes at the site, particularly with respect to the Cys and Met ligation, are evident. This perturbed geometry confers the protein a green colour.
Metal Sitesin Small BlueCopperProteins,BlueCopperOxidasesand Vanadium-ContainingEnzymes
45
Fig. 3. Copper site in M121Hmutant of azurin from AD, pH 7.0,MOLSCRIPT[12]
The copper sites of the M121Q mutant ofAD azurin both in the oxidised and reduced state, as derived from X-ray crystallography, are displayed in Fig. 4 [25]. The axial methionine ligand was mutated to glutamine because there was evidence from amino acid sequence alignments among the phytocyanins and molecular modelling based on the spatial structure of cucumber basic protein that glutamine is the axial ligand in stellacyanin [49] and the study of the mutant may shed new light on the structure of the copper site in the latter protein. In the oxidised form (Fig. 4a), the histidine and cysteine copper ligand distances and angles in the equatorial plane around the copper are very similar to the wild-type protein. Gln121 is coordinated in a monodentate fashion via its side-chain oxygen atom at a distance of 2.26 ~. The distance between the copper and the carbonyl group of Gly45 is increased to 3.37 A resulting in a distorted NNSO copper coordination. These values are equal, within experimental error, to the copper ligand bond distances in cucumber stellacyanin, whose Xray structure has been determined very recently [6]. In contrast to wild-type azurin, the copper site in M121Q azurin undergoes significant structural changes upon reduction (Fig. 4b). An increase of the Cu-OE Gln121 and ND1 His117 bond lengths to 2.73 and 2.68 A, respectively, with an SG Cys112-Cu(I)ND1 His46 angle of 156° produces an almost linear Cu(I) site with two strong bonds to the thiolate sull~hur atom of Cys112 (2.09/~) and the imidazole nitrogen atom of His46 (1.97 A), and two weak interactions with OE Gln121 and ND His117.
46
Albrecht Messerschmidt
Fig. 4. Copper site in M121Q mutant of azurin from AD. a) oxidised form, b) reduced form MOLSCRIPT [12]
Metal Sites in Small Blue Copper Proteins, Blue Copper Oxidasesand Vanadium-Containing Enzymes
47
1.2.3 Substitution of Copper by Hg, Zn, Cd and Ni The apo-forms of the small blue copper proteins are able to bind other metals like Hg, Zn, Cd and Ni at their metal binding site. The Cu(II) has been substituted by Hg(II) in crystals of poplar plastocyanin and the crystal structure determined [36]. The quasi-tetrahedral coordination geometry found in Cu(II)plastocyanin remains almost unchanged in Hg(II)-plastocyanin, apart from the slight enlargement of the coordination polyhedron required to accommodate the mercury atom. When Hg(II) replaces Cu(II) in plastocyanin, a ring flip at Pro36 is observed. This ring flip is not unique. Pro36 undergoes a similar conformational change in the transition between the high and low-pH forms of reduced plastocyanin [34], and moves even further towards the CG-exo conformation in apoplastocyanin [50]. A zinc-containing azurin has been obtained as a by-product of heterologous expression of the gene encoding PA azurin in Escherichia coli and its X-ray structure determined [27]. The geometry at the metal binding site has been changed to some extent. The largest difference between Zn-azurin and Cu-azurin is at Gly45 O, which forms a weak bond (2.9 ~) with Cu but a strong bond (2.3 ~) with Zn. The movement of this atom and the slight adjustment of backbone atoms connected to it is probably induced by the preference of Zn for a tetrahedral coordination. A crystal structure of the PA azurin mutant N47D, which was prepared to test the influence of this mutation on the redox potential, showed that zinc was bound instead of copper. The zinc site of this mutant structure is almost identical to the wild-type zinc azurin structure [31]. A Cd-substituted AD azurin has been characterised by X-ray crystallography [42]. The Cd ion is not so close to the main-chain carbonyl of Gly45 (2.76 A) as the zinc ion in zinc azurin (2.32 ~). The distance between the Cd ion and the methionine ligand is 3.23 ~ compared to 3.40 .~ for the zinc-methionine bond distance in zinc azurin. This means that Cd behaves more like Cu than Zn in this protein. The structural data of this metal derivative are of interest for investigations on azurin and azurin mutants using i11mCd-perturbed angular correlation spectroscopy [51]. Copper can also be replaced by nickel in type 1 copper proteins. A crystal structure of the nickel-substituted W48M azurin mutant has been determined [30]. The nickel metal site of this mutant structure is in principle similar to the zinc metal site in wild-type zinc azurin, however, the Ni-S(Cys) bond length is increased to 2.49 A. 1.2.4
The CuAOinudear Copper Site The purple CuA copper site is one of four metal sites in cytochrome c oxidases and functions as a primary electron acceptor for cytochrome c (see [19, 52], for example). N20 reductase, another cupredoxin domain-containing enzyme, also contains a CuA site as one of its metal sites and has been characterised both in cytochrome c oxidase and N20 reductase as a mixed valence, Cu(1.5)-Cu(1.5) redox state, due to its seven-line EPR spectrum and other similar spectroscopic
48
Albrecht Messerschmidt
Fig.5. Schematic drawing of the dinuclear Cua site in CyoA(with permission from [11]) properties [53, 54]. Cua sites have been incorporated into amicyanin [55], azurin [56] and purple Cyo A (membrane-exposed domain from a quinol oxidase) [11]. The structure of the CUAsite in purple Cyo A, as derived from the crystal structure [11], is shown in Fig. 5. Two thiolate groups from different cysteines bridge the two coppers. Two histidines act as terminal ligands to each copper. The arrangement of these four ligands is symmetrical with respect to the copper pair. Asymmetry is introduced by the coordination of a methionine and main-chain carbonyl oxygen which each complete the distorted tetrahedral coordination of the individual coppers. All four Cu-S(Cys) bonds are ~- 2.2 A long. The copper-copper distance is 2.5 ~. The CuA sites in cytochrome c oxidases resemble that of Cyo A. The copper-copper distances in the X-ray structures of cytochrome c oxidase from Paracoccus denitrificans and bovine heart are 2.6 and 2.7 ~, respectively. It is remarkable that the distorted tetrahedral coordination of each copper is conserved and that the Cu2S2Im2 cluster allows complete electron delocalization, thereby ensuring that the small reorganisation energy is spread over both copper ions [19]. 1.3 Spectroscopy of Cupredoxins Cu(II) proteins have been classified as type 1, type 2, type 3 or CUA.The type 1, type 2 and CuAclassification is based on the nature and magnitude of their EPR hyperfine coupling and their VISible spectra (Fig. 6), whereas the type 3 copper is EPR-silent due to antiferromagnetic coupling of the two copper ions that constitute this copper species. The trigonal or bipyramidal type 1 site, as found in wild-type azurin, exhibits an axial EPR spectrum. This corresponds to the effective C3v symmetry of
49
Metal Sites in Small Blue Copper Proteins, Blue Copper Oxidasesand Vanadium-Containing Enzymes
Type 1
Type 2
N~,
,,~N
"-cu .... S~f"
"~N
CuA
L,
""- .cu. . . ~mS/,,,
N/
S~"
/
,N
"""
Examples
Plastocyanin Azurin
EngineeredAzurin EngineeredSOD
Cytochrome c Oxidase N20 Reductase
Color Absorption max. in nm
Blue Green 460, 600
Yellow 400
Purple 480, 530, 800
EPR A H in 104 cm "1
< 90 (4 lines)
> 140 (4 lines)
< 30 (7 lines)
"L
Fig. 6. Copper coordination geometries in copper cysteinate proteins. Ligands denoted as S for cysteine thiolate,N for histidine imidazole and L for more weaklycoordinated methionine thioether or backbone carbonyl (with permission from [64])
the electronic distribution of the site and indicates that the detailed geometry and nature of the ligands in the NNS plane only weakly influence the shape of the electronic distribution. Distorted tetrahedral type 1 sites (the geometries of the different cupredoxin sites have been described in the previous paragraph) show a rhombic EPR spectrum consistent with a ligand field symmetry lowered to C2v. Stellacyanin, some type 1 copper site mutants, pseudoazurin and nitrite reductase belong to this subgroup. It is assumed that in these cases the rhombicity is due to a strong axial ligand field component [4]. W-band EPR, ENDOR and ESEEM studies have been carried out on frozen solutions and single crystals of wild-type azurin and the M121Q azurin mutant [57-60]. These studies more precisely define the character and position of the g-tensor and other spectroscopically relevant parameters. The data obtained can be used as a basis for an improved quantum-mechanical characterisation of the relevant copper sites. A strong band around 600 nm and a weaker one at 450 nm are found in the VISible region of the spectrum of type 1 sites. The strong absorption around 600 nm is mainly caused by a sulphur to Cu ligand-to-metal charge transfer (LMCT) transition. New theoretical studies identify the electron from a sulphur pn-orbital as being involved in the 600 nm LCMT transition [61]. The band around 450 nm has been assigned to a histidine to Cu LMCT transition but from recent resonance Raman (RR) studies on wild-type and mutant azurin as well as on superoxide dimutase mutants it is more probable that this band is caused by a second sulphur to Cu LMCT transition [62]. It is plausible that small changes in the copper coordination alter the electronic structure of the site and, as a consequence of this, the LMCT transfer properties. Associated with this is a change in the relative intensities of the 450 nm and 600 nm bands. Naturally occurring type 2 copper proteins do not contain sulphur ligands. Using genetic engineering, type 2 copper sites that contain a cysteinate as one of their ligands have been generated in azurin [63] and superoxide dismutase
50
Albrecht Messerschmidt
©
T1 Trigonal
T1 Distorted Tetrahedral
(430 - 405 cm-1)
(405 - 355 cm-1)
T1.5 Tetrahedral (360 - 340 cm-1)
/
T2 Distorted Tetragonal (365 - 340 cm-1)
T2 Tetragonal (320 - 300 cm-1)
Fig.7. Detailed coordination geometries in mononuclear copper-sulphur proteins (with per-
mission from [64]). Range of RR frequencies for the Cu-S(Cys) stretch in parentheses [16]. They exhibit a yellow colour (maximum absorbance at 400 nm). RR spectroscopy has been used to predict the coordination geometry in copper-sulphur proteins [64]. From these investigations it turns out that the mononuclear copper sites in copper-sulphur proteins can be subdivided in T1 trigonal, T1 distorted tetrahedral, T1.5 tetrahedral, T2 distorted tetragonal and T2 tetragonal (Fig. 7) [64]. This classification according to RR-spectroscopy is mainly based on the characteristic variation of the predominant Cu-S(Cys) stretching frequency and analyses of its correlation with ~460/E600absorption ratios and the geometry of the metal sites. 1.4 Redox Potentials and Electron Transfer Properties
The redox potential of type 1 copper sites in proteins varies from 185 mV for stellacyanin to 680 mV for rusticyanin, whereas the values for most are in the range from 260 mV to 380 mV. This is remarkably high as the redox potential
Metal Sitesin Small BlueCopperProteins,BlueCopperOxidasesand Vanadium-ContainingEnzymes
51
for the aqueous Cu(I)/Cu(II) pair (150 mV). It is thought that the stabilisation of the tetrahedral coordination in type 1 copper proteins is responsible for the higher redox potential because it is known from copper coordination chemistry that Cu(I) prefers a tetrahedral coordination and Cu(II) a tetragonal one. It was assumed that the nature of the axial ligand (usually methionine) would be responsible for tuning the redox potential [65]. This picture had to be extended as mutations of coordinating histidines or outside the first copper coordination sphere also changed the redox characteristics [66, 67]. An unexpected result was observed with the AD M121Q mutant. Met121 had been replaced to mimic a stellacyanin blue type 1 site. But the redox potential changed only by -23 mV with respect to the wild-type protein. The explanation came from the X-ray structure determination of both the oxidised and reduced state of the mutant. As already described in a previous paragraph, the oxidised form exhibits a strong axial coordination of the copper by OE of Gln121 stabilising the Cu(II) state. In the reduced form, a mainly two-fold coordination of the copper is found, which stabilises the Cu(I) state. Compared with the wild-type, both oxidation states are stabilised, resulting in an only moderate change in the redox potential. In PA azurin Met121 has been replaced by all other amino acids [46]. The redox potentials for this group of mutants varies from -105 mV to + 138 mV compared with the wild-type value at pH 7.0 [47]. Replacements of other copper ligands or in the second copper coordination sphere do not have drastic effects on the redox potential (changes are in the range for the Met121 mutations) (see [4], for example). Finally, titrating groups may have a considerable influence on the redox potential. The relatively low variation of the redox potential in the large number of azurin mutants designed to study the influence of the mutation on the redox potential supports the idea that the protein fold of the protein matrix around the redox site is also of importance. This is evident from the spatial structure of rusticyanin with its unusually high redox potential of 680 mV. The copper ligands in rusticyanin have been confirmed to be His, Cys, His, Met as in plastocyanin and many other blue type 1 copper proteins, for example [68]. Although the detailed geometry of the copper coordination site is not known yet it should be the significant abundance of hydrophobic residues in the loops closest to the copper site which causes this high redox potential by drastically changing the charge distribution in the environment of the redox centre. It is not the aim of this article to discuss the electron transfer properties of cupredoxins in detail. These have been described in several review articles (see [2, 4, 69], for example). Briefly, the type 1 copper sites in natural cupredoxins have been designed to make a rapid electron transfer possible. This has been accomplished by a copper site geometry that undergoes few changes upon alteration of the redox site (small reorganisation energy). The molecules have one or two surface regions for electron entrance and/or electron exit. One of these surface areas is a hydrophobic patch around the northern histidine as found in plastocyanin, azurin, and amicyanin. The structure of the electron transfer complex between methylamine dehydrogenase and amicyanin from Paracoccus denitrificans has been determined by X-ray crystallography [70]. The amicyanin docks at the light subunit of methylamine dehydrogenase with its hydro-
52
Albrecht Messerschmidt
phobic patch close to the northern histidine. The counterpart of the methylamine dehydrogenase is a hydrophobic surface area near the tryptophan tryptophylquinone (TTQ) cofactor. The copper atom of amicyanin and the redox factor of methylamine dehydrogenase are about 9.4 A apart, enabling a rapid electron transfer. The northern histidine is located between the two redox centres and may facilitate electron transfer between them. Plastocyanin has the hydrophobic patch and a remote acidic surface area. Cytochrome f, one of the natural redox partners, reacts at this site with plastocyanin [71]. The azurin system has been used to study long-range intramolecular electron transfer reactions. Farver, Pecht and other investigators started this research by following the electron transfer between the copper centre and a remote disulphide bridge [72-74]. Electron transfer rates were calculated using an electron pathway analysis based on the theory of Beratan and Onuchic [75]. The distant coupling problem has been studied in detail by Gray, Onuchic and associates [76]. In PA azurin positions 122, 124 and 126 of the C-terminal/3strand have been mutated to histidines. A [Ru(2,2"-bipyridine)2(imidazole)] 2÷ complex has been attached to the imidazole group of these residues and to His83 in the wild-type protein. The coupling between the copper ion and the Ru-complex coordinated to a histidine on the protein surface has been determined. In the case of the Ru-complex attached to histidines in positions 122, 124 and 126, the intervening medium was a single fl-strand, in the wild-type Rucomplex attached to His83 a section of a/3-sheet. On the basis of the results of these experiments a new theoretical approach for electron transfer beyond the single-pathway analysis has been developed. In the new theory, the protein matrix is reduced to only those relevant parts (tubes) which mediate the tunnelling matrix element. Such a tube is a tightly grouped family of pathways. These may be relevant for the coupling of a single tube only or for multiple tubes interfering with each other.
2 Metal Sites in Multi-Copper Oxidases 2.1 The Trinuclear Copper Active Site 2.1.1
Native Oxidized Enzyme The blue multi-copper oxidases ascorbate oxidase, laccase and ceruloplasmin catalyse the four-electron reduction of dioxygen to water with concomitant one-electron oxidation of the reducing substrate. They consist of domains with a cupredoxin fold. Ascorbate oxidase and laccase (their monomers have about 550 amino acids) are formed by three and ceruloplasmin (monomer about 1,000 amino acids) by six domains. The blue multi-copper oxidases have recently been reviewed [13] and a whole book has been devoted to the subject [77]. Apart from the blue type 1 copper sites, whose nature has been discussed in the previous section, the blue oxidases contain a trinuclear copper site which is located between the N- and C-terminal domains. The globular fold and the
Metal Sitesin Small BlueCopperProteins,BlueCopperOxidasesand Vanadium-ContainingEnzymes
53
Fig. 8. Schematic presentation of the monomer structure of ascorbate oxidase. The mono-
nuclear type 1 copper site is in the C-terminal domain whereas the trinuclear copper species is located between the N- and C-terminal domains
positions of the copper sites in the three-domain ascorbate oxidase are shown in Fig. 8. The atomic structure of the trinuclear copper site for ascorbate oxidase [43] is displayed in Fig. 9. The trinuclear cluster has eight histidine ligands symmetrically provided by the N- and C-terminal domains. It may be subdivided into a pair of copper atoms with histidine ligands whose coordinating N atoms (5 NE2 atoms and one ND1 atom) exhibit a trigonal prismatic arrangement. This pair is the putative type 3 copper. The remaining copper has two ligands and is the putative spectroscopic type 2 copper. Two oxygens are bound to the trinuclear species; as OH- or O2-, bridging the putative type 3 copper pair, and as OH- or H20 to the putative type 2 copper trans to the copper pair. An oxygen ligand in the centre of the three copper ions could not be detected. The bond lengths within the trinuclear copper site are similar to comparable binuclear model compounds [78, 79]. The average copper-copper distance in the trinuclear copper site of ascorbate oxidase is 3.74/~ and the individual distances do not deviate by more than 0.16 A from this mean value. Spectroscopic studies on laccase indicate tetragonal geometries for all three coppers in the cluster [80]. The tetragonal coordination geometries for all three coppers is not consistent with the structure. The coppers of the pair are both tetrahedrally coordinated, whereas the type 2 copper has three ligands. The existence of a central oxygen ligand would give rise to a pentacoordination of both copper pair atoms (but not a tetragonal-pyramidal coordination) and a square-planar coordination for the spectroscopic type 2-copper.
54
Albrecht Messerschmidt
t~
O
O
,.Q
8 0A
o~
Metal Sitesin Small BlueCopperProteins,BlueCopperOxidasesand Vanadium-ContainingEnzymes
55
There is experimental evidence in earlier studies that the type 2 copper is close to the type 3 copper and forms a trinuclear active copper site [81-86]. Solomon and associates described this metal binding site as a trinuclear active site, based on spectroscopic studies of azide binding to tree laccase [85, 86]. In ascorbate oxidase, the putative binding site for the reducing substrate is the type 1 copper [43]. Two channels which provide access from the solvent to the trinuclear copper site, the putative binding site of the dioxygen, could be identified. The structure of the trinuclear copper site in human ceruloplasmin, as determined by X-ray crystallography [14], is similar to that in ascorbate oxidase. However, in the current model of the 3.2 ~ resolution structure all histidine ligands are coordinated to the coppers by their RE2 atoms. 2.1.2
Type-2 Depleted (T2D) Form of Ascorbote Oxidose It is possible to selectively remove copper from the trinuclear species in the blue oxidases. This is documented by a loss of the type 2 EPR signal and of about 25% of the bound copper. The 2.5 .~ resolution X-ray structure of T2D ascorbate oxidase shows that about 1.3 copper ions per ascorbate monomer are removed [87]. The copper is lost from all three copper sites of the trinuclear copper species whereby the EPR-active type 2 copper is depleted somewhat preferentially (see Fig 10). Type-1 copper is not affected. The EPR spectra from polycrystalline samples of the native and T2D ascorbate oxidase were recorded. The native spectrum exhibits type 1 and type 2 EPR signals in a ratio of about 1:1 as expected from the crystal structure. The T2D spectrum reveals the characteristic resonances of the type 1 copper centre, as was also observed for T2D ascorbate oxidase in frozen solution, as well as the complete disappearance of the spectroscopic type 2 copper. The X-ray crystallography and the EPR spectroscopy seem to present a dilemma [88]. The X-ray structure shows all three sites depleted,
i2
°
!
Fig.lO. AveragedFOT2D-FCT2D-differenceelectrondensitymap plus atomicmodelaroundthe trinuclear copper site. Gontourlevels: - 18.0 solid line, 18.0 dashed line, magnitudesof holes less than -35.0
56
Albrecht Messerschmidt
whereas the EPR spectrum indicated selective removal of the type 2 copper. However, the EPR spectroscopy of the T2D form seems to be more complicated than usually assumed. A removal of copper from the trinuclear species means that we have a statistic mixture of copper pairs with the hole sitting on each copper, respectively. The relevant copper pairs are antiferromagnetically coupled and therefore EPR-silent. As a result of this, the T2D depleted duster does not produce an EPR signal. It is also conceivable that the remaining copper pairs are in the reduced state. In line with this are studies on T2D laccase which had been specificallylabelled with 63Cu and 65Cu.They show that there is either migration of Cu between the sites or a flip of ligands between the coppers [89].
2.1.3 Fully-ReducedForm of Ascorbate Oxidase The 2.2 ~ resolution X-ray structure analysis of fully-reduced ascorbate oxidase gave the following results [90]: The geometry at the type 1 copper remains much the same compared to the oxidised form as mentioned already in the previous section. The mean copper-ligand bond lengths of both subunits are increased by 0.04 A on average which is insignificant but may indicate a trend. A schematic drawing of the reduced form of ascorbate oxidase is shown in Fig 11.
~
,,P....~ ~, ,rto506
,
!
t
> HlO4
) Fig.11. Schematic drawing of the reduced form of ascorbate oxidase around the trinuclear copper site
Metal Sitesin Small BlueCopperProteins,BlueCopperOxidasesand Vanadium-ContainingEnzymes
57
The structural changes are considerable at the trinuclear copper site. Thus on reduction the bridging oxygen ligand OH1 is released and the two coppers CU2 and CU3 move towards their respective histidines and become tricoordinate, a preferred stereochemistry for Cu(I). CU2 and CU3 are each trigonally planar coordinated by their respective histidine ligands with equal bond lengths and bond angles within the accuracy of this X-ray structure determination. The copper-coopper distances increase from an average of 3.7 A to 5.1 A for CU2CU3, 4.4 A for CU2-CU4 and 4.1 A for CU3-CU4. The mean values of the copper-ligand distances of the trinuclear copper site are comparable to native oxidised ascorbate oxidase. CU4 remains virtually unchanged between reduced and oxidised forms. Coordinatively unsaturated copper(I) complexes are known from the literature. Linear bicoordinated [91] and T-shaped tricoordinated [92] copper(I) compounds have been reported. The copper nitrogen distances for both linearly arranged nitrogens are about 1.9 A, about 0.1 A shorter than copper nitrogen bond lengths in copper(II) complexes. In Tshaped copper(I) complexes, the bond length of the third ligand is increased. The copper ion CU4 has a T-shaped threefold coordination not unusual for copper(I) compounds. The structure of the fully reduced trinuclear copper site is quite different therefore from that of the fully oxidised resting form of the enzyme. 2.1.4
Peroxide Form of Ascorbote Oxidose
The 2.6 A resolution X-ray structure analysis of the peroxide form of ascorbate oxidase is illustrated in Fig. 12 [90]. The geometry at the type i copper site is not changed compared to the oxidised form. The copper-ligand average bond distances for both subunits show no significant deviations from those of the oxidised form. As in the reduced form, the structural changes are remarkable at the trinuclear copper site. The bridging oxygen ligand OH1 is absent, the peroxide binds terminally to the copper atom CU2 as hydroperoxide and the copper-copper distances increase from an average of 3.7 A to 4.8 A for CU2-CU3 and 4.5 ]l for CU2-CU4. The distance CU3-CU4 remains 3.7 A. The mean values of the copper-ligand distances of the trinuclear copper site are again comparable to native oxidised ascorbate oxidase and corresponding copper model compounds. The copper ion CU3 is tricoordinated, as in the reduced form, but the coordination by the ligating N atoms of the corresponding histidines is not exactly trigonal-planar and the CU3 atom is at the apex of a flat trigonal pyramid. The coordination sphere around CU4 is not affected and similar in all three forms. The copper atom CU2 is tetracoordinated to the NE2 atoms of the three histidines, as in the oxidised form, and by one oxygen atom of the terminally bound peroxide molecule in a distorted tetrahedral geometry. Its distance to CU3 increases from 4.8 A in the oxidised peroxide derivative to 5.1 A in the fully reduced enzyme. The bound peroxide molecule is directly accessible to solvent through a channel leading from the surface of the protein to the CU2-CU3 copper pair. This channel has already been described in [43] and its possible role as
58
Mbrecht Messerschmidt CU!
C507
J
f
H45o
H~II
~
/
\
x~104 Fig.
12. Schematicdrawingof the peroxide form of ascorbate oxidase around the trinuclear copper site
a dioxygen transfer channel has been discussed. An interesting feature is the close proximity of the imidazole ring of histidine 506 to the peroxide molecule. Histidine 506 is part of one possible electron transfer pathway from the type 1 copper to the trinuclear copper site and could indicate a direct electron pathway from CU1 to dioxygen. It may also help to stabilise important intermediate states in the reduction of dioxygen. The strong positive peaks at CU2 in both FONATrFOwoxand FOREDu-FOwox electron density maps could not be explained by a shift of CU2 alone. Occupancies of the copper atoms as well as of the oxygen atoms OH3 and the peroxide molecule were refined. Type-1 copper CU1 is almost unaffected. Copper atoms CU3 and CU4 are only partly removed, but copper atom CU2 is about 50% depleted. The oxygen ligands exhibit full occupancy. The treatment of crystals of ascorbate oxidase with hydrogen peroxide not only generates a well-defined peroxide binding but also a preferential depletion of the copper atom position CU2. In the copper-depleted molecules the coordinating histidine 106 adopts an alternative side-chain conformation as detected in the 2 FOFC-map calculated with the final peroxide derivative model coordinates.
Metal Sites in Small Blue Copper Proteins, Blue Copper Oxidasesand Vanadium-Containing Enzymes
59
This map shows that Hisl06 moves away when the copper atom CU2 is removed and opens the trinuclear site even more. From the T2D crystal structure of ascorbate oxidase it is apparent that copper from all three metal binding sites of the trinuclear copper species are removed to different extents. The movement of the His 106 side-chain could explain how this process is accomplished. Copper depletion may also cause instability of the protein towards hydrogen peroxide. Reaction of hydrogen peroxide with ascorbate oxidase in solution in excess leads to a rapid degradation of the enzyme [93].This can be monitored in the UV/VIS PEOX-NATI difference spectrum by a negative band at 610 nm and a positive band at 305 nm. Adding four equivalents of hydrogen peroxide per monomer ascorbate oxidase does not lead to enzyme degradation and gives a positive peak at 305 nm indicative of peroxide binding. Unfortunately, it was not possible to monitor the UWVIS spectrum of dissolved crystals after X-ray data collection because of the dissociation of the bound peroxide in solution. The reaction of dioxygen with laccase or ascorbate oxidase has been investigated by several groups and is reviewed in [43] where the possible binding modes of dioxygen to binuclear and trinudear copper centres are also discussed. A novel mode of dioxygen binding to a binuclear copper complex was found in a compound synthesised by Kitajimia et al. [94] which should be mentioned in detail. The complex contains the peroxide in the p-e2:e2 mode i. e. side-on between the two copper(II) ions. Such a binding mode for dioxygen has been detected in the crystal structure of the oxidised form of Limulus Polyphemus subunit II hemocyanin [95]. However, the binding mode of dioxygen to the trinuclear copper site in the blue oxidases appears to be different, as can be seen from the Xray structure of the peroxide derivative of ascorbate oxidase. During its reaction with fully reduced laccase dioxygen binds to the trinuclear copper species and three electrons are very rapidly transferred to it resuking in the formation of an"oxygen intermediate" with a characteristic optical absorption near 360 nm [96, 97] and a broad low temperature EPR signal near g = 1.7 [98, 99]. The type 1 copper is concomitantly reoxidized when the low-temperature EPR signal is formed. The oxygen intermediate decays very slowly (tm ~- i to 15 s) correlated with the appearance of the type 2 EPR signal [100]. Solomon and co-workers [101-103] have identified and spectroscopically characterised an oxygen intermediate during the reaction of either fully reduced native tree laccase or T1Hg laccase with dioxygen. They concluded from their spectroscopic data that the intermediate binds as 1,1-/2 hydroperoxide between either CU2 and CU4 or CU3 and CU4. As it is unlikely that the dioxygen migrates or rearranges coordination during reduction, Messerschmidt et al. [90] proposed that the binding site and mode determined in the peroxide derivative of ascorbate oxidase is representative for all reaction intermediates of dioxygen and, using homology arguments, valid in all blue oxidases. Recently, a dicopper complex was synthesised that performs the reversible cleavage and formation of the dioxygen O-O bond within the complex [104]. The copper-copper distance is 3.56 A in the Cu2(/.I-~2:~2-O2) core where the O-O bond is not cleaved and 2.79 ~ in the Cu2(p-O)2 moiety with the cleaved O-O bond. This mechanism will be valid for oxygen-activating dinuclear copper enzymes, such as tyrosinase, but not for the trinuclear active
60
Albrecht Messerschmidt
copper site of the multi-copper oxidases. The copper-cop~per distance for the copper pair in fully reduced ascorbate oxidase is at 5.1 A m u c h too large to enable this binding mode. Furthermore, the end-on binding of the peroxide in ascorbate oxidase is indicative of an asymmetric action of the dioxygen or at least of its reaction intermediates.
2.1.5 Azide Form of Ascorbote Oxidose
The results of the 2.3 i resolution X-ray structure analysis are shown in Fig. 13 [90]. The geometry at the type 1 copper site is not changed compared to the native form. The copper-ligand bond distances averaged over both subunits show no significant deviations from those of the native form. Again, the structural changes are large at the trinuclear copper site. The bridging oxygen ligand OH 1 and water molecule 145 have been removed, CU2 moves towards the coordinating histidines and two azide molecules bind terminally to it. The copper-copper
//,=,,..~H 506
,,oo H448~_... ""
OH3---cu
~",.~z
"..
!i~/ 4
i l
• ~.~
/
I
\
104
Fig. 13. Schematicdrawing of the azideform of ascorbate oxidasearound the trinuclear copper site of ascorbate oxidase
Metal Sitesin Small BlueCopperProteins,BlueCopperOxidasesand Vanadium-ContainingEnzymes
61
distances increase from an average of 3.7 _~ to 5.1 ~ for CU2-CU3 and 4.6 ,~ for CU2-CU4. The distance CU3-CU4 is decreased to 3.6 A. The mean values for the copper-ligand distances of the trinuclear copper site are again comparable to native ascorbate oxidase and corresponding copper model compounds. The coordination of CU3 resembles that of the peroxide form. The threefold coordination by histidines is a very flat trigonal pyramid. The coordination sphere around CU4 is not affected. CU2 is pentacoordinated to the NE2 atoms of the three histidines, as in the reduced form, and to the two azide molecules. The two azide molecules are terminally bound at the apices of a trigonal bipyramid. Both azide molecules bind to the copper atom CU2, which is easily accessible from the broad channel leading from the surface of the protein to the CU2CU3 copper pair. It is not unexpected that the second azide molecule (az2 in Fig. 13) binds in a similar manner as the peroxide molecule, becausse azide is regarded as a dioxygen analogue. There is no azide molecule bridging either CU2 with CU4 or CU3 with CU4. The binding of azide in laccase as well as to ascorbate oxidase has been studied extensively by Solomon and co-workers [101,105-107] and by Marchesini and associates [108, 109] by spectroscopic techniques. The derived spectroscopic models involve the binding of two azide molecules for laccase and three azide molecules for ascorbate oxidase with different affinities. As the binding of the high-affinity azide molecules seemed to generate spectral features related to the type 2 and type 3 coppers, the spectroscopic data were interpreted as the binding of at least one azide molecule as a 1,3-/2 bridge between the type 3 copper ions and the type 2 copper ion. There are many structural studies of copper coordination compounds with azide ligands from mainly mononuclear and binuclear copper complexes but few with trinuclear copper complexes. A comprehensive review of copper coordination chemistry has been written by Hathaway (see [110], for example). Azide binds only terminally to mononuclear systems. Pentacoordination of nitrogen ligands, including azide, to Cu(II) is frequently found arranged as a trigonal bipyramid. In binuclear systems azide may bind terminally as 1,1-/2 or bridging as 1,3-/2. Similarly two azides may bind di-l,1-/2 or di-1,3-/2. The interaction with all three copper ions of a trinuclear complex may be either terminally as 1,1,1-/2 or bridging as 1,1,3-/2. In the X-ray crystal structure of ascorbate oxidase two azide molecules bind terminally to the type 3 CU2. Azide binding in ascorbate oxidase resembles therefore the binding of azide to an isolated copper ion. In fact there is little interaction of CU2 with CU3 and CU4 which are 5.1 ~ and 4.6/~ away, respectively. The coordination of the copper ion CU4 in the native oxidised structure is of some interest. It has only three ligating atoms at close distances forming a Tshaped coordination which is known for Cu(I) complexes (see the discussion of reduced the form). However, the ligand field is completed if we take into account the n-electron systems of the imidazole rings of histidines 62 and 450 (see Fig. 9). A ligand field with tetragonal-pyramidal symmetry around CU4 is then formed. The shortest distances of CU4 are 3.4/~ to CD2 450 and 3.6 .~ to CG 62. These distances are too long for strong copper n-electron interactions but the histidines will contribute to the CU4 ligand field.
62
Albrecht Messerschmidt
2.2 Aspectsof ElectronTransfer and MechanisticProperties The arrangement of the type 1 and the trinuclear copper redox centres in the muki-copper oxidases is crucial for their mechanistic properties. This arrangement as found in ascorbate oxidase [43] is illustrated in Fig. 14. The distances from the type 1 copper CU1 to the individual coppers of the trinuclear copper cluster, CU2, CU3 and CU4, are 12.2 A, 12.7 A and 14.9 A, respectively. The His506-Cys507-HisS08 amino acid sequence segment (ascorbate oxidase numbering) links the type 1 copper centre and the type 3 coppers as a bridging ligand. The same situation is found in the X-ray structure of ceruloplasmin [14]. The three type I copper sites of ceruloplasmin are arranged in a triangle whose corners are about 18 A aopart. The two type 1 copper sites located in domains 2 and 4 are more than 18 A away from the trinuclear cluster reducing their capability to directly interact with the trinuclear centre. Electron transfer properties and a proposal for the catalytic mechanism for ascorbate oxidase, which should be valid in principle for all blue oxidases, have been described in a recent review [111]. Briefly, the type 1 site in the C-terminal domain is where the electrons enter from the reducing substrate. In ceruloplasmin, the reducing substrate may react with the two other type 1 sites as well but the main entrance site will be the type 1 copper centre in the C-terminal domain. After reduction of the type 1 centre an intramolecular electron transfer to the trinuclear copper site takes place. The electron tunnelling may be completely through-space or follow individual electron transfer pathways. These may be through-bond, through-space or a combination of both. A through-bond pathway is available for both branches each with 11 bonds (Fig. 14). Two combined through-bond pathways can be chosen. The first one from CU1 to CU2 of the trinuclear centre involves a transfer from the SG atom of Cys507 to the main-chain carbonyl of Cys507 and through the hydrogen bond of this carbonyl to the ND1 atom of HisS06 (blue arrows in Fig. 14). The second one branches at the main-chain carbonyl of Cys507 and follows through bonds to CU3 (green arrows in Fig. 14). The catalytic mechanism contains the following stages: oxidised resting form; fully reduced form, OH-bridging ligand released, dioxygen binds probably to copper ion CU2; hydroperoxide intermediate, dioxygen has been reduced by two reduction equivalents, dioxygen double bond has been broken; oxygen radical intermediate O- coordinated to CU2, O-O bond cleaved, first water molecule released, CU1, CU3 and CU4 reoxidized; continuation of reduction of CU1 by a fifth reduction equivalent and electron transfer to CU3, release of second water molecule; continuation of reduction and attainment of the fully reduced state of the next catalytic cycle.
3 First X-ray Structure of a Vanadium-Containing Enzyme: Chloroperoxidase from the Fungus Curvularia inaequalis The first X-ray structure of a vanadium-containing protein, a chloroperoxidase from the fungus Curvularia inaequalis has been solved very recently [112]. The
Metal Sitesin Small BlueCopperProteins,BlueCopperOxidasesand Vanadium-ContainingEnzymes
63
Fig. 14. Region of the atomic model of ascorbate oxidase containing the type 1 and the trinuclear copper centres
chloroperoxidase from this fungus belongs to a class of vanadium enzymes that oxidise halides in the presence of hydrogen peroxide to the corresponding hypohalous acids. The mature enzyme consists of 609 amino acid residues with a calculated molecular mass of 67,488 Da. The 2.1 A crystal structure (R-value = 20%) of an azide chloroperoxidase complex reveals the geometry of the catalytic vanadium centre (Fig. 15). In the structure with a 2 mM azide mother liquor concentration, the vanadium has a trigonal bipyramidal coordination with three non-protein oxygen ligands (bond distances of about 1.65 A), one nitrogen ligand (NE2 atom) from His496 (bond distance of 2.25 ~) and an exogenous azide ligand (bond distance to coordinating nitrogen of 1.98 A). The negative charge of the VO3 group with vanadium in the oxidation state (V) is compensated by hydrogen bonds to several positively charged or hydrophilic protein side-chains and the main-chain amide nitrogen of Gly403. Oxygen OWl of the VO3 group forms hydrogen bonds to nitrogens NH1 of Arg360 (2.94 ~) and NH2 of Arg490 (2.93 A), oxygen OW2 to nitrogen NZ of Lys353 (2.72 A) and nitrogen N of Gly403 (2.99 A), and oxygen OW3 to oxygen OG of Ser402 (2.71 A)
64
Albrecht Messerschmidt
Fig. 15. Plot of the vanadium active site of chloroperoxidase from Curvularia inaequalis, MOLSCRIPT[12]
and nitrogen NE of Arg490 (3.04 ~). A water molecule from the solvent is hydrogen-bonded to the nitrogen atom N 1 of the bound azide molecule. The binding of vanadium as hydrogen vanadate (V) has now been confirmed by a difference Fourier map of the azide-free form. This difference map contains the VO4 group as its highest peak and one water molecule hydrogen-bonded to the apical oxygen atom. The difference electron density has the shape of a trigonal pyramid with the vanadium in the centre of the trigonal base plane, three oxygens at the vertices of this plane, and a fourth oxygen at the apex of the pyramid. The NE2 atom from His496 completes the trigonal bipyramidal coordination of the vanadium. The protein fold is mainly a-helical with two four-helix bundles as main structural motifs (Fig. 16) and an overall structure different from other structures. The vanadium centre is located on top of the second four-helix bundle. The molecule has an overall cylindrical shape with a length of 80 A and a diameter of 55 A. The secondary structure is mainly helical (about 44% of the atomic model), consisting of 20 helices, a small proportion of fl-structures and the rest extended strand and loop regions. Three residues near the azide or putative chloride binding site deserve special interest, the hydrophobic side chains of Trp350 and Phe397 as well as the imidazole ring of His404 (see Fig. 15). Both hydrophobic side-chains provide a hydrophobic environment, which seems to be necessary to stabilise chloride binding. His404 is on the other side of the putative chloride binding site and may play a crucial role as an acid-base group in catalysis. Steady-state kinetic data for vanadium chloroperoxidase show that the binding of peroxide is inhibited when a group with a pKa larger than 5 is protonated [113]. It may be that when His404 is protonated, binding of peroxide to the metal site is no longer possible as is also seen in heme iron peroxidases. This suggests some analogy to
Metal Sitesin SmallBlueCopperProteins,BlueCopperOxidasesand Vanadium-ContainingEnzymes
65
Fig. 16. Ribbon-type representation of the chloroperoxidase molecule including the vanadium binding site, MOLSCRIPT [12]
the reaction of heme-containing cytochrome c peroxidase with peroxide and the vanadium enzyme. A detailed catalytic reaction scheme has not been elaborated due to the lack of structural information about catalytic intermediates. The high thermostability of the enzyme is mainly due to the packing of the helices to a compact molecule. An amino acid sequence comparison with vanadium-containing bromoperoxidase from the seaweed Ascophyllum nodosum shows high similarities in the regions of the metal-binding site, with all hydrogen vanadate(V) interacting residues conserved except for Lys353, which is Asn. The crystal structure of the vanadium-containing bromoperoxidase from Ascophyllum nodosum has recently been solved by the Schomburg's group in Braunschweig, Germany, but has not been published yet. Acknowledgements. The author wishes to thank Prof. R. Huber for supporting the work on the reviewed projects and for his valuable suggestions. The Deutsche Forschungsgemeinschaft (Schwerpunktthema: Bioanorganische Chemie) is thanked for financial support.
References 1. Adman ET (1985) In: Harrison P (ed) Metalloproteins. Verlag Chemie, Weinheim, Germany, p 1 2. Sykes AG (1991) Adv Inorg Chem 36:377 3. Adman ET (1991) Adv Protein Chem 42:145 4. Canters GW, Gilardi G (1993) FEBS Lett 325:39
66
Albrecht Messerschmidt
5. Pascher T, Bergstr6m J, Malmstr6m BG, V/inngard T, Lundberg LG (1989) FEBS Lett 258: 266 6. Hart PJ, Nersissian AM, Herrmann RG, Nalbandyan RM,Valentine IS Eisenberg D (1996) Protein Science 5: 2175 7. Guss ]M, Merritt EA, Phizackerly RP, Hedman B, Murata M, Hodgson KO, Freeman HC (1988) Science 241:806 8. Hunt AH, Toy-Palmer A, Assa-Munt N, Cavanagh J, Blake II RC, Dyson HJ (1994) J Mol Biol 244: 370 8a. Walter RL, Ealick SE, Friedman AM, Blake II RC, Proctor C, Shoham M (1996) J Mol Biol 263: 730 9. Iwata S, Ostermeier C, Ludwig B, Michel H (1995). Nature 376:660 10. Tsukihara T, Aoyama H, Yamashita E, Tomizaki T, Yamaguchi H, Shinzawa-Itoh K, Nakashima R, Yaono R, Yoshikawa S (1996) Science 272:1136 11. Wilmans M, Lappalainen P, Kelly M, Sauer-Eriksson E, Saraste M (1995) Proc Natl Acad Sci USA 92:11955 12. Kraulis P (1991) J Appl Cryst 24:946 13. Messerschmidt A (1993) Adv Inorg Chem 40:121 14. Zaitseva I, Zaitsev V, Card G, Moshkov K, Bax B, Ralph A, Lindley P (1996) J Biol Inorg Chem 1: 15 15. Adman ET, Godden JW, Turley SJ (1995) ] Biol Chem 270:27458 16. Han J, Loehr, TM, Lu Y,Valentine JS, Averill BA, Sanders-Loehr (1993) J Amer Chem Soc 115:4256 17. Lu Y, LaCroix LB, Lowery MD, Solomon EI, Bender CJ, Peisach ], Poe JA, Gralla EB, Valentine JS (1993) J Amer Chem Soc 115:5907 18. Andrew CR, Yeom H, Valentine JS, Karlsson BG, Bonander N, van Pouderoyen G, Canters GW, Loehr TM, Sanders-Loehr J (1994) J Amer Chem Soc 116:11489 19. Ramirez BE, Malmstr6m BG, Winkler JR, Gray HB (1995) Proc Natl Acad Sci USA 92:11949 20. Nat H, Messerschmidt A, Huber R, van de Kamp M, Canters GW (1991) J Mol Biol 221 : 765 21. Nar H (1992) PhD thesis, Technical University Mfinchen 22. Baker EN (1988) J Mol Biol 203:1071 23. Dodd FE, Hasnain SS, Abraham ZHL, Eady RR, Smith BE (1995) Acta Cryst D51 : 1052 24. Nat H, Messerschmidt A, Huber R, van de Kamp M, Canters GW (1991) ] Mol Biol 218:427 25. Romero A, Hoitink CWG, Nar H, Huber R, Messerschmidt A, Canters GW (1993) J Mol Bio1229:1007 26. Messerschmidt A (1996) unpublished results 27. Nar H, Huber R, Messerschmidt A, Filippou AC, Barth M, Jaquinod M, van de Kamp M, Canters GW (1992) Eur J Biochem 205:1123 28. Shepard WEB,Anderson BF, Lewandowski DA, Norris GE, Baker EN (1990) J Amer Chem Soc 112:7817 29. Tsai L-C, Sj61inL, Langer V, Pascher T, Nar H (1995) Acta Cryst D51 : 168 30. Tsai L-C, Sj61inL, Langer V, Bonander N, Karlsson BG, V~inngard T, Hammann C, Nar H (1995) Acta Cryst DS1:711 31. Sj61inL, Tsai L-C, Langer V, Pascher T, Karlsson G, Nordling M, Nar H (1993) Acta Cryst D49:449 32. Hammann C, Messerschmidt A, Huber R, Nar H, Gilardi G, Canters GW (1996) J Mol Biol 255: 362 33. Guss JM, Bartunik HD, Freeman HC (1992) Acta Cryst B48:790 34. Guss JM, Harrowell PR, Murata M, Norris VA, Freeman HC (1986) J Mol Biol 192:61 35. Collyer CA, Guss JM, Sugimura Y,Yoshizaki F, Freeman HC (1990) J Mol Biol 211 :617 36. Church WB, Guss ]M, Potter ]J, Freeman HC (1986) J Biol Chem 261 : 34 37. Petratos K, Dauter D, Wilson KS (1988) Acta Cryst B44:628 38. Vakoufari E, Wilson KS, Petratos K (1994) FEBS Lett 347:203
Metal Sites in Small Blue Copper Proteins, Blue Copper Oxidasesand Vanadium-Containing Enzymes
67
39. Cunane LM, Chen Z-W, Durley RCE, Mathews FS (1996) Acta Cryst D52:676 40. Romero A, Nar H, Huber R, Messerschmidt A, Kalverda AP, Canters GW, Durley R, Mathews FS (1994) J Mol Bio1236:1196 41. Guss JM, Merritt EA, Phizackerley RP, Freeman HC (1996) unpublished results 42. Baker EN, Anderson BF, Blackwell KA (1995) Protein Data Bank Reference 1AIZ 43. Messerschmidt A, Ladenstein R, Huber R, Bolognesi M, Avigliano L, Petruzzelli R, Rossi A, Finazzi-Agr6 A (1992) J Mol Biol 224:179 44. Guckert JA, Lowery MD, Solomon EI (1995) J Amer Chem Soc 117:2817 45. Kroes SJ, Hoitink, CWG,Andrew CR, Ai J, Sanders-Loehr, Messerschmidt A, Hagen WR, Canters GW (1996) Eur J Biochem 240:342 46. Karlsson BG, Nordling M, Pascher T, Tsai L-C, Sj61inL, Lundberg LG (1991) Protein Eng 4:343 47. Pascher T, Karlsson BG, Nordling M, Malmstr6m BG,V~inngard T (1993) Eur J Biochem 212:289 48. Kroes SJ, Andrew CR, Sanders-Loehr J, Canters GW (1995) J Inorg Biochem 59:661 49. Fields BA, Cuss JM, Freeman HC (1991) J Mol Biol 222:1053 50. Garrett TPJ, Clingeleffer DJ, Guss JM, Rogers SJ, Freeman HC (1984) J Biol Chem 259: 2822 51. Danielsen E, Bauer R, Hemmingsen L, Andersen M-L, Bjerrum MJ, Butz T, Troeger W, Canters GW, Hoitink CWG,Karlsson G, Hansson O, Messerschmidt A (1995) J Biol Chem 270:573 52. Gennis R, Ferguson-Miller S (1995) Science 269:1063 53. Kroneck PMH, Antholine WE, Riester J, Zumft WG (1988) FEBS Lett 242:70 54. Kroneck PMH, Antholine WE, Kastrau DHW, Buse G, Steffens GCM, Zumft WG (1990) FEBS Lett 268: 274 55. Dennison C, Vijgenboom E, de Vries S, van der Oost J, Canters GW (1995) FEBS Lett 365: 92 56. Hay M, Richards JH, Lu Y (1995) Proc Natl Acad Sci USA 93:461 57. Coremans, JWA, Poluetkov OG, Groenen EJJ, Canters GW, Nar H, Messerschmidt A (1994) J Amer Chem Soc 116:3097 58. Coremans JWA, van Gastel M, Poluetkov OG, Groenen EJJ, den Blaauwen T, van Pouderoyen G, Canters GW, Nar H, Hammann C, Messerschmidt A (1995) Chem Phys Lett 235: 202 59. Coremans JWA,Poluetkov OG, Groenen EJJ, Canters GW,Nar H, Messerschmidt A (1996) J Amer Chem Soc 118:12141 60. Coremans JWA, Poluetkov OG, Groenen EJJ, Warmerdam G, Canters GW, Nar H, Messerschmidt A (1996) J Phys Chem 100:19706 61. Solomon EI, Baldwin MJ, Lowery MD (1992) Chem Rev 92:521 62. Sanders-Loehr J (1993) In: Karlin KD, Tyeklar Z (eds) Bioinorganic Chemistry of Copper. Chapman & Hall, New York, NY, p 51 63. den Blaauwen T, Hoitink CWG, Canters GW, Han J, Loehr TM, Sanders-Loehr J (1993) Biochemistry 32:12455 64. Andrew CR, Sanders-Loehr J (1996) Acc Chem Res 29:365 65. Gray HB Malmstr6m BG (1983) Comments Inorg Chem 2:203 66. Hoitink CWG, Canters GW (1992) J Biol Chem 267:13836 67. Nishayama M, Suzuki J, Ohnuki T, Chang HC, Horinouchi S, Turley S, Adman ET, Beppu T (1992) Prot Eng 5:117. 68. Casimiro DR, Tyo-Palmer A, Blake RC, Dyson HJ (1995) Biochemistry 34:6640 69. Sykes AG (1990) Struct Bond 75:175 70. Chen L, Durley R, Poliks BI, Hamada K, Chen Z, Mathews FS, Davidson VL, Satow Y, Huizinga E, Vellieux FMD, Hol WGJ (1992) Biochemistry 31 : 4959 71. Beoku-Betts D, Chapman SK, Knox CV, Sykes AG (1985) Inorg Chem 24:1677 72. Farver O, Pecht I (1992) J Amer Chem Soc 114:5764 73. Farver O, Skov LK, Nar H, van de Kamp M, Canters GW, Pecht I (1992) Eur J Biochem 210:399
68
Albrecht Messerschmidt
74. Farver O, Pecht Iet al (1993) Biochemistry 32:7317 75. Beratan DN, Onuchic JN, Winkler JP, Gray HB (1992) Science 258:1740 76. Regan J], di Bilio A], Langen R, Skov LK, Winkler JR, Gray HB, Onuchic JN (1995) Chem & Biol 2:489 77. Messerschmidt A (1997) Multi-Copper Oxidases. World Scientific, Singapore 78. Karlin KD, Hayes JC, Gultneh Y, Cruse RW, McKnown JW, Hutchinson JP, Zubieta J (1984) J Amer Chem Soc 106:2121 79. Chaudhuri P, Ventor D, Wieghardt K, Peters E, Peters K, Simon A (1985) Angew Chem 97:55 80. Cole JL, Clark PA, Solomon EI (1990) J Amer Chem Soc 112:9534 81. Br/inden R, Deinum J (1977) FEBS Lett 73:144 82. Martin CT, Morse RH, Kanne RM, Gray HB, Malmstr6m BG, Chan SI (1981) Biochemistry 20: 5147 83. Morpurgo L, Desideri A, Rotilio G (1982) Biochem 1 207:625 84. Winkler ME, Spira DJ, LuBien CD, Thamann TJ, Solomon EI (1982) Biochem Biophys Res Com 107:727 85. Allendorf MD, Spira DJ, Solomon EI (1985) Proc Natl Acad Sci USA 82:3063 86. Spira-Solomon DJ, Allendorf MD, Solomon EI (1986) 108:5318 87. Messerschmidt A, Steigemann W, Huber R, Lang G, Kroneck PMH (1992) Eur J Biochem 209:597 88. Beinert H (1996) J Inorg Biochem 64:79 89. McMillin DR, Eggleston MK (1997) In: Messerschmidt A (ed) Multi-Copper Oxidases. World Scientific, Singapore, p 129 90. Messerschmidt A, Luecke H, Huber R (1993) J Mol Biol 230:997 91. Schilstra MJ, Birker PJMW,Verschoor GC, Reedijk J (1982) Inorg Chem 21:2637 92. Sorell TN, Malachowski MP (1983) Inorg Chem 22:1883 93. Marchesini A, Kroneck PMH (1979) Eur J Biochem 101:65 94. Kitajima N, Fujisawa K, Moro-oka Y (1989) J Amer Chem Soc 111: 8975 95. Magnus K, Ton-That H (1992) J Inorg Biochem 47:20 96. Andreasson L-E, Br~indenR, Malmstr6m BG, V~inngard T (1973) 32:187 97. Andreasson L-E, Br[inden R, Reinhammar B (1976) Biochim Biophys Acta 438:370 98. Aasa R, Br/inden R, Deinum J, Malmstr6m BG, Reinhammar B, V~inngard (1976) FEBS Lett 61:115 99. Aasa R, Br~inden R, Deinum J, Malmstr6m BG, Reinhammar B, V/inngard (1976) Biochem Biophys Res Com 70:1204 100. Br~inden R, Deinum J (1978) Biochim Biophys Acta 524:297 101. Cole JL, Clark PA, Solomon EI (1990) J Amer Chem Soc 112:9534 102. Cole JL, Ballou DP, Solomon EI (1991) J Amer Chem Soc 113:8544 103. Clark PA, Solomon EI (1992) J Amer Chem Soc 114:1108 104. Halfen ]A, Mahapatra S, Wilkinson EC, Kaderli S, Young Jr VG, Que Jr L, Zuberbuehler AD, Tolman WB (1996) Science 271:1397 105. Allendorf MD, Spira DJ, Solomon EI (1985) Proc Natl Acad Sci USA 82:3063 106. Spira-Solomon DJ, Allendorf MD, Solomon EI (1986) J Amer Chem Soc 108:5318 107. Cole ]L, Avigliano L, Morpurgo L Solomon EI (1991) J Amer Chem Soc 113:9080 108. Casella L, Gulotti M, Pallanza G, Pintar A, Marchesini A (1988) Biochem J 251:441 109. Casella L, Gulotti M, Pintar A, Pallanza G, Marchesini A (1989) J Inorg Biochem 37:105 110. Hathaway BJ (1987) Compr Coord Chem 5: 111. Messerschmidt A (1997) In: Messerschmidt A (ed) Multi-Copper Oxidases. World Scientific, Singapore, p 23 112. Messerschmidt A, Wever R (1996) Proc Natl Acad Sci USA 93: 392 113. van Schijndel JWPM, Barnett P, Roelse J, Vollenbroek EGM, Wever R (1994) Eur J Biochem 225:151
Structure and Function of the Xanthine-Oxidase Family of Molybdenum Enzymes M a r i a Joao Rom~to 1. a n d R o b e r t H u b e r 2 1 Instituto de Tecnologia Quimica e Biol6gica, Apt. 127, 2780 Oeiras and Instituto Superior T6cnico, Dep. Qulmica, 1096 Lisboa Codex, Portugal, E-maih
[email protected] 2 Max-Planck-Institut ffir Biochemie, am Klopferspitz 18a, D-82152 Martinsried, Germany
This work gives an account of the recent achievements which have contributed towards the understanding of the structure and function of the xanthine oxidase family of enzymes-the molybdenum hydroxylases. It is based essentially on the crystallographic data of the aldehyde oxido-reductase from Desulfovibrio (D.) gigas, a member of that family,whose structure is described in detail. Comparisons are made, whenever appropriate, with spectroscopic, kinetic and model compound studies. Mechanistic implications of the crystal structure of the D. gigas enzyme are considered and extended to the xanthine oxidase family. Keywords: Hydroxylase, molybdoenzymes, molybdopterin, protein crystallography, xanthine oxidase
List of Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
70
1
Introduction ..................................
70
2
Overview of the Xanthine Oxidase Family of Molybdenum Enzymes
71
3
Crystal Structure o f t h e A l d e h y d e O x i d o - R e d u c t a s e f r o m
Desulfovibrio gigas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
75
3.1 3.2 3.3 3.4 3.5 3.6
S t r u c t u r a l a n d D o m a i n A r r a n g e m e n t of the Protein . . . . . . . . . . The [2Fe-2S] Centers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The M o l y b d o p t e r i n Cofactor . . . . . . . . . . . . . . . . . . . . . . . . S t r u c t u r e a n d E n v i r o n m e n t o f the Metal Centers . . . . . . . . . . . . The M o l y b d e n u m Site a n d Its E n v i r o n m e n t . . . . . . . . . . . . . . . The D i m e r . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
76 79 79 81 87 88
4
C o m p a r i s o n o f t h e A l d e h y d e O x i d o - R e d u c t a s e f r o m Desulfovibrio gigas w i t h the X a n t h i n e O x i d a s e F a m i l y o f E n z y m e s . . . . . . . . . .
89
5
Structure-Based Catalytic Mechanism
90
6
Conclusions
7
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
..................................
..................
92 93
* Corresponding Author. Structure and Bonding, Vol. 90 © Springer Verlag Berlin Heidelberg 1998
70
Maria Jofio Romgo • Robert H u b e r
List of Abbreviations AOR DMSOR Mop Mod MPT MCD MGD MAD MHD XO XDH
aldehyde ferredoxin oxido-reductase from Pyrococcusfuriosus dimethylsulfoxide reductase aldehyde oxido-reductase from Desulfovibrio gigas aldehyde oxido-reductase from Desulfovibrio desulfuricans ATCC27774 mononucleotide form of molybdopterin molybdopterin cytosine dinucleotide molybdopterin guanine dinucleotide molybdopterin adenine dinucleotide molybdopterin hypoxanthine dinucleotide xanthine oxidase xanthine dehydrogenase
1 Introduction Molybdenum-containing enzymes can be grouped into two classes: nitrogenase, which catalyzes the reduction of dinitrogen to ammonia and where molybdenum is part of a heterometal FeMo-cofactor [1]; and hydroxylases or oxotransferases, which promote a variety of two-electron oxidation-reduction reactions, whereby oxygen (oxo) atom transfer occurs. An essential role of molybdenum is the catalysis of the controlled oxo-transfer reaction coupled to an electron transfer between substrate and other cofactors, such as Fe/S centers, hemes or flavins. Coupling of both functions leads to a formal direct transfer of an oxygen atom from the metal center to the substrate (oxotransferase activity [2]). Among the molybdenum hydroxylases which are currently known (see [3] for a review), xanthine oxidases, sulfite oxidases, nitrate reductases, aldehyde oxidases and DMSO reductases have been characterized in greater detail. Molybdenum oxotransferase enzymes catalyze the following general reactions, where water has been shown to be the source of the incorporated oxygen atom [4], such that reducing equivalents are generated rather then consumed [60, 61] RH + H20 ~
ROH + 2e- + 2H +
(1.1)
RH = aldehyde or aromatic heterocycle
RR'E + H20 __k_>RR'E = O + 2e- + 2H +
(1.2)
E = N, S (simple oxo transfer without cleavage of a C-H bond)
Xanthine oxidase from bovine milk, due to its ready availability, is the prototype for molybdenum hydroxylases and has been intensively studied for the past forty years. Many other oxomolybdenum enzymes have been investigated within the last decade. In a recent review on molybdenum oxotransferases [3], an extensive list of 68 enzymes is given, according to the information available at the end of 1995. These enzymes have been classified into three families on the basis of the reactions they catalyze, as well as on the basis of the characteristics of their molybdenum centers: (1) The xanthine oxidase family, (2) the sulfite
Structureand Functionof The Xanthine-OxidaseFamilyof MolybdenumEnzymes
71
oxidase and assimilatory nitrate reductase family and (3) the DMSO reductase family. This classification is supported by amino acid homologies within these protein families, which are found in a wide range of organisms. The large xanthine oxidase family of enzymes may be considered one of the true hydroxylases, with the substructure dithiolene of the molybdopterin cofactor (fac) coordinated to a MoOS (H20) unit. The MoOS coordination was suggested by EXAFS experiments [8, 9, 57], but the presence of an additional water ligand was established by crystallography [6, 56]. Members of this family have been found broadly distributed within eukaryotes, prokaryotes and archaea. They catalyze the oxidative hydroxylation of aldehydes and aromatic heterocycles in reactions involving C-H bond cleavage (reaction 1.1). Sulfite oxidase and assimilatory nitrate reductases possess afacMo02 unit. There is uncertainty about additional coordination positions of the molybdenum. They have been found in eukaryotes and catalyze a simple oxo transfer to the lone pair of sulrite in sulfite oxidase (E=S, reaction 1.2-a) or the reverse reaction in nitrate reductase (E=N, reaction 1.2-b). In the DMSO reductase family, other enzymes such as the biotin-S-oxide reductase and bacterial dissimilatory nitrate reductases have been included, which also follow the overall stoichiometry of reaction 1.2-b. This family of enzymes possesses the Mo-cofactor with a bisdithiolene coordination of the molybdenum, established by recent X-ray structural data [59, 65]. These studies showed that the Mo atom is also coordinated by the side-chain oxygen of a serine (Ser 147). This class of enzymes seems to be structurally more diverse when compared to the other two, and its members have only been found in bacteria and archaea. Enzymes have been included in this class on the basis of significant sequence homology or spectral similarity. It is the purpose of this review to focus on the xanthine oxidase family of molybdenum enzymes with particular emphasis on the crystal structure of the aldehyde oxido-reductase from Desulfovibrio gigas (Mop), as the first representative of this group of enzymes for which a three-dimensional structure is available [6]. In the following, structural data on Mop will be provided and, whenever possible, comparisons will be made with literature data derived from spectroscopic - EPR, EXAFS, resonance Raman, MCD, ENDOR - and kinetic techniques, as well as from electron transfer studies. The mechanistic implications of the Mop structure for a general catalytic mechanism for the xanthine oxidase family of molybdenum enzymes are presented.
2 Overview of the Xanthine Oxidase Family of Molybdenum Enzymes Molybdenum oxotransferases in general possess a common Mo=O group in the metal coordination sphere, which is the reason why they have been called oxomolybdenum enzymes [7]. The molybdenum hydroxylases, which constitute the xanthine oxidase family (containing an MoOS unit), have been assigned on the basis of their irreversible inhibition by cyanide which reacts with Mo=S, releasing thiocyanate. Until X-ray structural data were available for this class of enzymes, the main techniques used to study the molybdenum coordination sphere were EXAFS [8, 9] ([58] for Mop) and EPR from the Mov species [10]
72
Maria Jofio Romio • Robert Huber
([11, 52] for Mop). EXAFS m e a s u r e m e n t s have shown that the metal center in its oxidized form has at least one oxo group with an Mo=O distance of about 1.7 ~, as well as (at least) two thiolate ligands at distances of about 2.4 A (see Fig. 1). The latter feature is c o m m o n to all o x o m o l y b d e n u m enzymes, and is due to the coordination of m o l y b d e n u m to an organic cofactor c o m m o n l y designated as " m o l y b d o p t e r i n ' . In spite of the lability of this cofactor when isolated from the protein matrix, its basic pterin ring structure and dithiolene side-chain were proposed on the basis of chemical and spectroscopic analysis of the cofactor iso2.33/~ (2.35)
2.15 A (2.18)
~
1.67A (1.74)
f
1.66/k (1.68)
___=.
2.00/k (2.14)
2.44 A (2.47) (1.98A )
b) reduced enzyme (Mo TM) [81([57])
a) oxidized enzyme (MoVO [81([57]) 1.67/~. (1.70)
SH-~
1.66 A
(~\ .H
>o
fs,,,
2.38/k (2.36) 2.27/k (2.34) c) reduced enzyme-alloxanthine complex (MOW)
[91([571)
SH-')
2.40 A
~_..-N~_LNH
r
Y--
2.08 A
d) reduced enzyme-violapterin complex (MOW) [91 2.11/~
O
,----s
~
C 1.66/~ kgH~,
1.68 A
! #o -
2.47/~
'~ ~-1.90a 1.68/~
e) oxidized Mop (desulfo) (MovI)
[581
1.
~..S_.~.~M0...~ ? 2.4oA f) reduced Mop (desulfo) (Mo w) [58]
Fig. The molybdenum coordination, as seen in Mop, with metal ligand distances obtained by EXAFS for various derivatives of XO and Mop: a xanthine oxidase, oxidized [8, 57]; b xanthine oxidase, reduced [8, 57]; c complex of xanthine oxidase with alloxanthine, reduced [9, 57]; d complex of xanthine oxidase with violapterin, reduced [9]; e desulfo form of Mop, oxidized [58]; f desulfo form of Mop, reduced [58]
Structureand Functionof The Xanthine-OxidaseFamilyof MolybdenumEnzymes
73
lated from the enzymes in different modified or inactivated forms [12-15]. However, the presence of a pyran ring in the structure was established by crystallography for the tungsten-containing aldehyde oxido-reductase from Pyrococcus furiosus [16], for the aldehyde oxido-reductase from Vesulfovibrio gigas (Mop) [6] and for the DMSO reductases from two Rhodobacter species [59, 65]. The pyran ring closure may occur by attack on the side-chain 9"OH to C7 of a dihydropterin, possibly after incorporation and binding to the enzyme. The molybdopterin cofactor is coordinated to the metal via its dithiolene function and may be present either in dinucleotide forms or in the simpler monophosphate form. In Mop, the cofactor is the dinucleotide of cytosine molybdopterin cytosine dinucleotide (MCD) -, but in other enzymes from prokaryotic sources is found as the guanine (MGD) [17], adenine (MAD) [18] or hypoxanthine (MHD) [18] dinucleotide. The simpler monophosphate form (MPT), also found in some bacterial enzymes [3], is the only form present in all known enzymes from eukaryotic sources, and the diversity of the pterin cofactor within known molybdenum-containing enzymes seems to be related to the species of origin rather than to the enzymatic function, as shown within the xanthine oxidase family, where MPT has been found in eukaryotic enzymes and MCD in bacterial enzymes reported so far [3]. Members of the xanthine oxidase family show about 25 % sequence identity and 60-70 % sequence similarity, with higher conservation in segments involved in the binding of the metal centers and redox-active sites. Enzymes belonging to this class are either: homodimers, a2, or dimers of heterotrimers, a2/J2y2. Mop, aldehyde oxidases and eukaryotic xanthine oxidases and xanthine dehydrogenases are organized as a2 homodimers, with all redox-active cofactors confined within a single polypeptide chain. The common folding pattern for this group of enzymes, starts with the two [2Fe-2S] domains, followed by a flavin domain (which is absent in Mop and replaced by an extended connecting segment, cf. Sect. 3.2) and finishes with two large domains responsible for binding the molybdopterin cofactor [5, 19]. Other groups of hydroxylases have been included in the xanthine oxidase family of enzymes, due to analogies in their molybdenum centers and the reactions catalyzed: the CO dehydrogenases [20-22], the isoquinoline oxidoreductases [23-29] and nicotine dehydrogenases [30-33], all from bacterial and archaeal sources. They are organized as a2fl2y2 structures, where the a subunit harbors the two [2Fe-2S] centers, the fl subunit the flavin and the y subunit the molybdenum cofactor. Xanthine oxidase is the prototype of the molybdenum hydroxylases and has been intensively studied using spectroscopic and kinetic techniques, which have contributed to characterizing it both structurally and functionally. Relations to a number of inorganic model systems could be deduced from such studies. These results have been described and summarized in a number of recent reviews [3, 34, 35, 101]. We will now focus on some general features of xanthine oxidase, highlighting relevant points which can be illustrated with details from the crystal structure of Mop. Xanthine oxidase catalyses the oxidation of xanthine to uric acid, using dioxygen as the physiological electron acceptor (or NAD + in the case of xanthine dehydrogenases).
74
Maria Jo~o Romgo • Robert Huber O
o¢-
O
2 H2°*+"
NH
" O~NH
N
> NH
O*H
+ H202
In addition to xanthine, it can also oxidize other aromatic heterocycles and aldehydes, although with little specificity. Xanthine is oxidized at the molybdenum center, whereby MovI is reduced to MoTM and reducing equivalents are transferred to 02 at the flavin site. Electron transfer between the molybdenum center and the FAD is mediated by the Fe/S centers. Intramolecular electron transfer within the xanthine oxidase family of molybdenum enzymes is an essential aspect of catalysis and has been studied by different techniques: flash photolysis [36, 37], pulse radiolysis [38, 39] and pH-jump perturbation [40-42]. Rate constants for electron transfer between redox-active centers have been determined and show that the equilibration of reducing equivalents within xanthine oxidase is rapid in comparison to the overall catalysis and is not rate-determining [40]. EPR was one of the first tools employed for the study of the molybdenum center of xanthine oxidase and a variety of Mov species were detected, either in the course of equilibrium reductive titrations, or transiently within reaction with substrate [10]. Other characteristic EPR signals have been observed from complexes of the enzyme with inhibitors such as arsenite [43-45], methanol [46], ethylene glycol [47] and alloxanthine [48]. The alloxanthine complex, characterized also by EXAFS (Fig. 1), gives, upon a one-electron reduction, an EPR signal resembling the so-called "very rapid" signal, observed very early and transiently in the course of the reaction of xanthine oxidase with excess alloxanthine. While this "very rapid" signal is quite anisotropic and exhibits no proton hyperfine coupling, the so-called"rapid" Mov EPR signal, which also arises in the course of reaction with substrate, belongs to two different types, "rapid type 1" and "rapid type 2", depending on the nature of the proton hyperfine coupling observed [7, 10]. The Mov EPR signals of ,type 2", exhibited by xanthine oxidase upon reaction with xanthine [49] (gl.2,3= 1.9951, 1.9712, 1.9616), are quantitatively very similar to the corresponding signals reported for Mop upon reaction with salicylaldehyde [11] (gl,a,3= 1.9882(3), 1.9702(3), 1.9643(3)), suggesting similarity in their active sites. Also the two [2Fe, 2S] centers - Fe/S I and Fe/S II - which are clearly distinguished on the basis of their characteristic EPR signals observed at low temperature, display similar features in Mop [52] and xanthine oxidases. The so-called Fe/S type I shows characteristic g-values similar within eukaryotic xanthine oxidases (milk XO [46] gl,2,3=2.022, 1.935; 1.899) and Mop [52] (gl,2,3 = 2.021, 1.938; 1.919) typical of spinach ferredoxin [3] (g1,2,3=2.02(1), 1.93(1), 1.90(1)). The center Fe/S II, on the other hand, exhibits broader lines than Fe/S I, with g-values comparable in Mop [52] (2.057, 1.970, 1.900) and milk xanthine oxidase [53] (2.12, 2.01, 1.91), but displaying larger variations within members of the xanthine oxidase family than the Fe/S I center. M6ssbauer spectroscopy also allows one to distinguish between both Fe/S centers in the reduced enzymes: one of the centers (probably Fe/S I) exhibits a
Structureand Functionof The Xanthine-OxidaseFamilyof MolybdenumEnzymes
75
rather normal quadrupole splitting AEQ of 2.4 mm/s and 2.69(2) mm/s for the ferrous site in xanthine oxidase (at 175 K) [54] and Mop (at 180 K) [55] respectively, while the other center exhibits an unusually large quadrupole splitting of 3.2 mm/s and 3.14(2) mm/s for xanthine oxidase and Mop respectively.
3 Crystal Structure of the Aldehyde Oxido-Reductase from Desulfovibrio gigas (Mop) The crystal structure of the aldehyde oxido-reductase from Desulfovibrio gigas (Mop) represents the first structure of a molybdenum oxotransferase and is a valid model for the interpretation of the weakh of experimental data for xanthine oxidase and related enzymes. It was solved at high resolution in its native desulfo form [6], as well as in its sulfo, oxidized, reduced and alcohol-bound forms [56], allowing a detailed look at the several structural aspects of the molybdenum hydroxylases relevant for catalysis: domain architecture; structure of the cofactors and binding mode within the polypeptide chain; molybdenum center environment; metal coordination and its role in catalysis. In analogy to eukaryotic xanthine oxidases, xanthine dehydrogenases and aldehyde oxidases, Mop is an ot2 homodimer [63] consisting of two 100 kDa subunits (2 × 907 amino acids) [62]. The redox-active cofactors are found in discrete domains within a single polypeptide chain. Mop possesses, per subunit,
o!s) o 4~
9 o
o 'o' 2""
l[
3-
SH
HSx~[
O
NH~/J~NH
(Dihydropterin) Fig.2. The MCD molybdopterin cofactor as established by crystallography in Mop [6]. The open-chain form as suggested by chemical analysis is also shown and an equilibrium between these two forms suggested in solution
76
M a r i a Joao R o m e o • Robert H u b e r
two different kinds of [2Fe-2S] centers [52], as well as a molybdopterin cofactor (molybdopterin cytosine dinucleotide-MCD) (Fig. 2), but lacks the flavin and its domain which is present in most molybdenum hydroxylases. 3.1 Structural and Domain Arrangement of the Protein
From the electron density map interpretation the two [2Fe-2S] centers were recognized in the earlier stages of the analysis, while the molybdopterin cofactor was identified only at a later stage of the structure solution as molybdopterin cytosine dinucleotide (MCD), later confirmed by chemical means [64]. For some of the higher resolution data sets which were analyzed [56], isopropanol molecules as well as magnesium ions, both from the crystallization solution, were identified in density and accordingly refined. The presence of alcohol molecules, particularly the one close to the molybdenum site, was important for modelling the Michaelis complex of an aldehyde substrate molecule (cf. below). The protein molecule is roughly globular with an approximate diameter of 75 .~ (Fig. 3). Its secondary structure shows 28% a-helical and 21%//-sheet conformations, with a total of 33 helical and 27 fl-strand segments longer than 3 amino acid residues. The molecule folds into four distinct domains (Fe/Sa, Fe/Sb, Mol and Mo2) (Fig. 3) of which the first two (Fe/Sa and Fe/Sb) bind the two iron-sulfur clusters, while the larger domains Mol and Mo2 bind the MCD cofactor in extended conformation by a network of hydrogen bonding interactions: Mol contributes with two single molybdopterin binding segments and Mo2 binds the other side of the pterin system and provides all of the dinucleotide binding segments (Fig. 4). These two large domains also surround the molybdenum catalytic site and define, at their interface, a 15 k-deep tunnel, wide open at the surface and constricted in the middle, which leads substrate molecules into the buried molybdenum catalytic site (Fig. 5).
Fig. 3. Stereo plot o f the m o l e c u l a r s t r u c t u r e of Mop w i t h the four i n d e p e n d e n t d o m a i n s rep r e s e n t e d in different colors a n d cofactors s h o w n as colored spheres. Fe/Sb - blue, residues 1 76; Fe/Sa - red, residues 84 156; c o n n e c t i n g peptide - white, residues 158 195; M o l - green, residues 196 581; Mo2 - purple, residues 582 907
Structureand Functionof The Xanthine-OxidaseFamilyof MolybdenumEnzymes
77 c~ 0
2=
8 =
2-0
0
~
,-~
0
0
=
,'~ ~.~ 0
.
0
=8
"~ ~ ~ 0 ~:~0 0 ~
=
~
E~
0
78
Maria Jo~o Romao • Robert Huber
Fig. 5. Stereo Cc' representation of the Mop molecule with cofactors highlighted as colored spheres. Two isopropanol molecules are also represented as well as the three buried waters close to the molybdenum site. The entrance of the tunnel is marked by one of the isopropanol molecules (IPP1) and by Pro258
The first Fe/Sb (residues 1 to 76) domain shows the CXXGXCXXC motif common to the plant-type ferredoxin class of iron-sulfur proteins [ 19]. Also the chain fold is similar to that of plant (Spirulinaplatensis) [66], alga (Aphanothece sacrum) [80] and cyanobacterium Anabaena [81] [2Fe-2S] ferredoxins. It shows the topology of a five-stranded fl-half barrel with an a-helix running almost perpendicular to the strand direction. Apart from the absence of a 19-residue loop toward the C-terminus of this domain and a / J - t u r n , shorter by six-residues, between the first two strands, the overall fold can be superimposed to Spirulina platensis ferredoxin [66] with a deviation of less than 0.5 A in the C° atom positions of the iron-sulfur cluster-binding turns. The second iron-sulfur domain Fe/S a (residues 84 to 156) reveals a new ferredoxin-type fold: a two-fold symmetric four-helical bundle with two longer central helices flanked by two oblique shorter a-helices. The iron-sulfur cluster lies at the N-termini of the two central helices. Domain Fe/S a is connected to the molybdenum-binding domain Mol via a long, extended segment with irregular secondary structure which juts about 50 A across a rather concave region of the surface of the molecule. This concave region extends from one side of the protein to the other and appears to be the most likely site for the insertion of the flavin domain present in xanthine oxidases. This region of the molecule also appears to be the most probable site of interaction with flavodoxin, which has been shown to be able to accept electrons from Mop/aldehydes in vitro [67]. The two larger Moco binding domains, Mol (residues 196 to 581) and Mo2 (residues 582 to 907) are in close contact to each other, accommodating the molybdenum catalytic site at their interface. Domain Mol is rather elongated (ca 75 ~ long and 28 .~ wide) and organized in two subdomains: a larger N-ter-
Structureand Functionof The Xanthine-OxidaseFamilyof MolybdenumEnzymes
79
minal part, which consists of a seven-stranded incomplete fl-barrel with one ahelix filling its central cavity, and two additional helices flanking the barrel and exposed to solvent. The smaller C-terminal subdomain consists of a fivestranded mixed parallel-antiparallel fl-sheet flanked by two helices which run approximately parallel to the strand direction. Domain Mo2 is also organized in two subdomains, each with a similar basic fold (a four-stranded fl-sheet, which bends around a pair of helices), and in part dyad related. Both subdomains resemble two large wings spanning over 80 A and with the cofactor MCD lying at the intersection of the two wings. 3.2 The [2Fe-2S] Centers
Both 2Fe-2S clusters appear planar with the iron atoms and cysteine sulfur atoms defining a plane orthogonal to the plane of the [2Fe-2S] group. All iron atoms are tetrahedrally coordinated by the sulfur atoms of the cysteines, with Fe-S bond lengths of 2.2 A and 2.3 A for the iron-sulfide and for the iron-cysteine SVrespectively. The N-terminal plant-type ferredoxin iron cluster (Fe/Sb) has one of the iron atoms linked to C40 and C45, while the other iron atom is coordinated by C48 and C60. The second iron cluster (Fe/Sa) has one of the iron atoms bound to C100 and C139 and the other to C103 and C137.While the Fe/Sb center is close to the protein surface with its Gys S ~ 0 exposed to the solvent, the FeS a center is deeply buried and in contact with the molybdopterin. The assignment of both Fe/S centers to the two spectroscopically distinguishable [47, 52, 53] iron-sulfur centers, type I and type II, is still unclear and under debate [3]. Data from magnetic coupling with molybdenum [52, 53, 78, 79] and electron transfer rate studies [3, 40], suggest that Fe/S I corresponds to Fe/Sa (helix bundle) and Fe/S II to Fe/Sb (plant-type ferredoxin). This assignment is also in agreement with the distribution of reducing equivalents among the redox centers of xanthine oxidase, which is kinetically fast in comparison to the catalytic turnover and follows their relative redox potentials [70]: for XO at pH 7.7 [42] MoVI/MoV-373mV, MoV/MorV-377mV, Fe/S 1-310mV, Fe/S II-255mV, FAD/FADH'-332mV, FADH"/FADH2-234 mV, for Mop at pH 7.6 [67, 69] MoVr/MoV-450mV, MoV/MoIV-530mV, Fe/S 1-280mV, Fe/S II-285mV. However, center type I is the one with g values quite similar to spinach ferredoxin. A possible explanation for this apparent contradiction is that details of coordination protein environment and solvent accessibility, rather than the general polypeptide fold, determine the EPR features of the iron centers [3]. For these reasons, we maintain the designation Fe/S a and Fe/Sb throughout this work. 3.3 The Molybdopterin Cofactor
'
The structure of the molybdopterin cofactor was established for Mop by crystallography as a cytosine dinucleotide (MCD) (Fig. 2). The bicyclic pterin structure and dithiolene side-chain had been proposed by chemical and spectroscopic analysis in general [12-15], but the fused pyran ring resulting from a
80
Maria Joao Romao • Robert Huber
tricyclic structure had remained undetected. It is now well established by crystallography and has been found in three different enzymes: in the aldehyde ferredoxin oxido-reductase (AOR) from Pyrococcusfuriosus [16], in Mop [6] and, more recently, in the DMSO reductase (DMSOR) from Rhodobacter sphaeroides [59] and capsulatus [65]. In both AOR and DMSOR, the metal atom (W and Mo respectively) coordinates two molybdopterins through their dithiolene groups but in distinct ways. The AOR from Pyrococcusfuriosus has two molybdopterins coordinating the tungsten atom and additionally linked by a magnesium ion bound to their phosphate groups [16]. In DMSOR (Fig. 6) we find two molybdopterin guanine dinucleotides (MGD) at the molybdenum atom resulting in a very elongated structure (N 35 ~) [59, 65]. Neither the tungstoprotein nor the DMSO reductases share homology with Mop or the xanthine oxidase family; they are members of three different protein families, of which the DMSO reductase family of oxomolybdenum enzymes is likely to be structurally more diverse than the xanthine oxidase family [3]. Nevertheless, the basic structure of the tricyclic system of the cofactor is similar in all three enzyme families. When isolated, the cofactor is bicyclic, suggesting that the pyran ring is closed by a reversible intramolecular nucleophilic addition of the hydroxyl 9'-OH to the C7 carbon atom of the double bond of a dihydropterin system [77]. The pyran ring closure may occur in situ subsequent to binding of the open-chain cofactor to the enzyme favored by the numerous interactions between the cofactor and the surrounding polypeptide chain as observed in Mop [6] (Fig. 4), DMSORs [59, 65] and AOR [16]. In all structures the pyran is approximately 40° tilted relative to the two ring pterin system (
[Ala-1831
°
Fig. 8. Idealized octahedral representation of metal coordination in W- and Mo-enzymes as
deduced from X-ray crystallography (cf [11, 13-15]). The encircled P symbolizesthe protein cording to the EPR redox titration [66]. It thus appears premature to draw conclusions about the stereochemical consequences of the tungsten coordination for the mechanism of action [ 11 ]. This stereochemistry has been reviewed recently for Mo enzymes based on the crystal-structural information [13-15] on the assumption that oxidized crystals have Mo(VI) and reduced crystals have Mo(IV) [129]. Unfortunately, the Mo enzymes are not readily comparable in detail to P. furiosus W-AOR, because D. gigas Mo-AOR has a five-coordinated Mo(VI?) with a single pterin, and in R. sphaeroides DMSO reductase one of the ligands to the Mo(VI?) is oxygen from a protein residue (cf. Fig. 8). The W-AOR of P.furiosus contains a single iron-sulfur cluster per subunit coordinated by four Gys residues, in a pattern of three, C-X-X-C-X-X-X-C, with a fourth Cys further downstream. Although this pattern, which is conserved in T. litoralis FOR [128], is not exactly the canonical ferredoxin-type sequence, the cluster appears to be a regular [4Fe-4S]/2+;z+/, single-electron transferring cubane. The reduction potential is Era,8= - 410 mV at 20 °C [87]; the ground state of the oxidized cluster is diamagnetic; the reduced, mixed-valence cluster is a physical mixture of S = 312 and S = 1/2 [66, 87]. The cubane is the proposed electronic link between the enzyme"s natural redox partner(s), presumably ferredoxin, and the tungstopterin active center. The distances to the protein surface and to the tungstopterin (Fig. 7) are consistent with this assignment.
186
W.R.Hagen. A.F.Arendsen
The third, subunit-bridging metal center is enigmatic. In the original paper: "the metal site is tentatively identified as iron on the basis of anomalous scattering of the atom and a metal analysis of the AOR protein although direct confirmation of this assignment by M6ssbauer spectroscopy is needed" [11]. Unfortunately, details of the anomalous scattering experiment are not given. In the present reviewers view anomalous scattering would allow for a more straightforward and specific assignment than M6ssbauer spectroscopy (one out of nine tetrahedral iron sites per protein dimer). Also, details of metal analysis are not given, however, again, to differentiate eight versus nine Fe in a protein of 132 kDa is far from a trivial matter. We have briefly reported on EPR studies of the anaerobically oxidized enzyme that indicated the bridging metal to be highspin (S=5/2) ferric iron in an environment of intermediate rhombicity [130]. However, our subsequent studies indicate that this signal is not always found in AOR preparations and does not appear to be related in a simple way to activity (unpublished resuks). For the metal in reduced enzyme Adams and collaborators have recently claimed on one occasion high-spin ferrous, S=2 [92] and on another occasion low-spin ferrous, S =0 [16]. The two ligands per subunit to the metal are Glu-332 and His-383, and they occur in two separate E-X-X-H patterns per subunit. In their recent review Kletzin and Adams note that in the protein sequence a total of three of these patterns are found, and they recall that a double E-X-X-H pattern is a putative binding motif for a dinuclear Fe-O-Fe cluster. They also call two of the patterns "conserved". However, this is not borne out by their comparison to the sequence of T. litoralis FOR [16, 128]. Finally, we know of no evidence, spectroscopic or otherwise, for the presence of dinuclear iron-oxo clusters in tungsten proteins. At this time we tentatively conclude that ,,the subunit-bridging iron" in P. furiosus AOR appears to be a nonfunctional accident of purification. 5.4 Redox Chemistry at the Active Site
The fact that the reduction-potential values E(V/VI) and E(IV/V) in Mo/W coordination complexes are frequently found to be close, or even crossed over, combined with the relative stability of the Mo/W-oxo group in the hexavalent state, naturally suggests a central role for the metal of Mo/W enzymes in biological OAT or CEPT reactions: group transfer and localization of valence. Thus, it is a long-standing paradigm of molybdenum biochemistry that the metal shuttles between Mo(VI) and Mo(IV) (e.g. [125, 126]) and that the pterin cofactor does not enter in the redox chemistry [131]. It would appear to be only natural to extend these ideas also to tungsten enzymes. However, several independent observations have been recently made that force us to reconsider this central paradigm of Mo/W biochemistry. The AOR of P.furiosus was initially purified in an inactive form on the basis of its reddish color in the presence of dithionite: RTP or red tungsten protein [87]. This protein was studied in EPR-monitored pH- and temperature-dependent redox titrations. No W(V) signals were observed in RTP. The oxidized protein was EPR-silent; reduction gave the S--3/2 and S = 1/2 mixture of the [4Fe-
The Bin-InorganicChemistryof Tungsten
187
4S] 1+ cubane; upon further reduction to very low potentials,
