Membrane structural biology: with biochemical and biophysical foundations

3 NVHBII\I3 11\1

STRUCTURAL BIOLOGY WI H BlOCH M CAL AND BIOPHYSICAL FOUNDATIONS

MARY LUCKEY San Francisco State University

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CAMBRIDGE UNIVERSITY PRE S

CAMBRIDGE UNIVERSITY PRESS

Cambl'idge, Ne\\' York, Melbourne, MilclT-iu, Cape Town, Singapol'e, Sao Palllo, Delhi Cnnlbridgc University Pres~

32 Avenue or the America" New York, NY 10013-2473, USA \\·ww.calnbridge.org Jn~ormalion on

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o Man' LuckeI' 2008 This publication is in copvrighl. Subjeci to statuton exception and to the provisions of relevant collecrive licensing agl'cemenls, no reproduction or anv pan mil\' take place without the wrillen permission of Cambridge Universilv Press. First published 2008 PI'inted in Canada bv Friesen, rl cala/o,~ ,.cco,.d {()/' Ihis puhlicrllioll is nvai!a!JIc {iOiIl Ihe B,./Ii,/, Lihron·.

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Luckev, Mal'." Membl'ane structural biology: \\'itb biochemical anel bioflh~'sical foundations I Marv Luckev. p,; cm. Includes bibliographical rderences and index. ISBN 978-0-521-85655-3 (hardback) I. Membranes (Biology) 2. Membrane lipids. 3. IVlembrane proteins. 1. Title. [DNLM: l. Cell Membrane - phvsiologv. 2. iV1cmbrane Lipids - phvsiologv. 3. Membl'ane Proteins - phvsiologv. QU 350 1..941 m 2008] QH60l.L75 2008 571.6'4-dc22 2007031145 ISBN

978-0-521-85655-3 hardback

Cambl'idge University Press has no responsibilitv for the persistence or accuracy of URLs for external or third-partv Internet Web sile, refern::d to in thi, publication and doe~ not gu(\rantee that al1,v content on .such

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The title page shows high-resolution stl1.JClures of membrane proteins incorporated into a simulated lipid bilavel'. The proteins are, fl'Om left to r'ight: vitamin B '2 transponel's BlLrCD with BtuF, the light harvester LH2 with some chlorophylls, the mechanosensilive channel MscS, lactose permease, BtuB from the outer membrane, rhe pore domain of Kv1.2, aquilporin. ilnd Cal, -ATPase. Substr-,

0.

ell 20

/ ./

0

40

~

:l

'0 if)

~

c;:20, and the resulting heterogeneity can be quite deleterious, especially in crystal formation. For homogeneous alternatives, a series of alkyl polyoxyethylenes of defined chain length (CxE N , where X is the number of C atoms in the alkyl group and N is the number of oxyethylene monomers in the headgroup) is used. Commercially available detergents may have problems of impurities; for example, SDS often contains n-dodecanol and polyoxyethylene-based detergents may contain peroxide and aldehyde, which necessitates additional purification steps or the purchase of "protein-grade" or "especially purified" quality. A few detergents, including sodium cholate and

ANIONIC Sodium dodecy\ sulFale (Sodium lauryl sulFale)

Sodium dodecyl-N-sarcosinale (SodiuIl11auryl-N-sarcosinale) (Sarkosyl L) CH 3

o

0

I

II

II

~C /N 'CH /CO-Na+ 3

~O-S-O-Na+

II

II

o

o CH 3

CATIONIC Celyl lrimelhylammonium bromide (Hexadecvl lrimelhylammonium bromide) (CTAB)

I

~N+-CH3Br­

I CH 3

ZWITTERIONIC Lauryldimelhylamine oxide (LDAO) (Dodecvlamine N-oxide)

CHAP~S 0 HO

CH 3

,

I

~N+-O-

I

HO

CH 3

~H3

~

N~N+~S-O-

I

I

II

H

CH 3

0

OH

SulFobetaines (Zwiuergent bl'and)

0

CH 3

II

I

~N+~S-O-

I

II

0

CH 3 UNCHARGED

~

Digilonln

Polyoxyelhylene alcohols (denoted CxENl (I) Brij series (2) Lubrol (vVX,PX)

HoroL...J'OB Glc-GIc-Gal-Gal-Xyl-O

~(O

: H

I3-D-oclylglucoside

~~

~O

~OHO

OB

CH OH

I3-D-Dodecylmalloside (laury! maltoside)

CH 20H

OH

Fatly acid ester~ of polyoxyelhylene sorbitan (denoled C,-SOI bltan-E n ) .

CH 3 CH 3 )n -OH

CH OH 2

~2 ~ HO

0

HO

OH 0

OH

(0 CH CH ), - OH 2 2 J~

I

II

TlVe~ C -

OH

(0 CH 3 CB 3 )" - 0 - CH 2 - CH

r

(0 CH 2 CH 2)y - OH

(n = w+ x + Y + z) Alkyl-N-melhylglucamides (MEGA"" brand) CH 3 I OH OH OB

~rN;YY o

OH OH OH

(0 CH CH2)z - OH 2 Polyoxyelhylene p leu octylphenols (denoted IeI'I - C80 E,,) (I) Trion X-lOO, n = 9.6 (2) Trion X-114, n = 7.8 (3) Nonidet PAD, n= 9

~ ( O CH 2 CH 2)n - OH

BILE SALTS Sodium cholale

0

&C'ON,'

Sodium deoxycholate

HO

'OH

0

§C'ON"

II

II

HO

3.1. Structures of some detergents used to solubilize membrane components. From Gennis, R. B., Biomembranes, New York: Springer-Verlag, 1989, pp. 90-91.

Detergents

45

c.

B.

~ zCO

Na+

~O r-0 / HN HN

Y

FFFFFF

S~H F

F F

F F

F

OA~H

Ht00H OH

3.2. Three new alternatives to detergents. A. The tripod amphiphile that extracts bacteriorhodopsin. Redrawn with permission from Yu, S. M., et aI., Prot Sci. 2000,9:2518-2527. B. An example of an amphipol called A8-35 with variation in the polymeric backbone giving x = 35%, y = 25%, and z = 40%. Redrawn with permission from Gohon, Y, et al. Anal Biochem. 2004, 334:318-334. C. A hemifluorinated surfactant called HF-TAC [Cz Hs C6 H1ZCZ H 4 -S-poly-Tris-(hydroxymethyl)aminomethane] with a single hemifluorinated hydrocarbon chain. Redrawn with permission from Breyton, c., et al., FEBS Lett. 2004, 564:312-318.

l3-octylglucoside, can be purified by crystallization prior to use. Because no detergent satisfies all criteria desired by researchers, alternatives to the traditional detergents are being designed and tested (Figure 3.2). One of the tripod amphiphiles (Figure 3.2A), which have three hydrophobic tails and a hydrophilic headgroup, has been used to solubilize bacteriorhodopsin. The amphipols (Figure 3.2B) are small amphiphilic polymers of various structures that form water-soluble complexes with membrane proteins, presumably by wrapping around their nonpolar domains. Hemifluorinated surfactants (Figure 3.2C) are unlike detergents because the perfluorinated regions of their chains are hydrophobic but stilllipophobic. The nonfluorinated tails enable them to interact with membrane proteins while presumably allowing the interactions between the proteins and lipids to remain intact in aqueous solutions.

The action of most detergents involves micelle fonnation. Micelles are roughly spherical assemblies of surfactant molecules, in which most of the nonpolar tails are sequestered from the aqueous environment in a disorganized (liquid-like) hydrophobic interior. Thus the chains are not fully extended like the spokes of a wheel, and the radius of the micelle is 10% to 30% smaller than the fully extended monomer (Figure 3.3A). Furthermore, the surface is rough and heterogeneous rather than smoothly covered by polar headgroups: NMR studies of SDS micelles revealed that only onethird of the surface was covered by hydrophilic headgroups (Figure 3.3B). At high concentrations of detergent, micelles change shape to become elliptical or rod-like; this occurs at lower concentrations for surfactants with weakJy polar headgroups. Micelles of small B.

A.

(a)

Mechanism of Detergent Action

(b)

3.3. A. Cross-sectional views of detergent micelles. The old view (a) incorrectly portrays the chains as ordered like spokes, whereas they are actually disordered and fluid (b), resulting in an uneven surface. Redrawn with permission from Menger, F. M., R. Zana, and B. Lindman, J Chem Educ. 1998,75:93 and 115. B. Model of the surface of a micelle, showing the uneven surface at the detergent/water interface. Redrawn from Lindman, B. et al., in J.-J. Delpuech (ed.), Dynamics of Solutions and Fluid Mixtures by NMR, Wiley & Sons, 1995, p. 249. © 1995 by John Wiley & Sons Limited. Reprinted with permission from John Wiley & Sons Limited.

TABLE 3.1. Properties of micelles derived from some commonly used detergents Monomeric MW

Detergent Octyl-(3-D-glucoside Dodecyl-maltoside Lauryldimethylamine oxide (LDAO) Lauramido-N,N-dimethyl-3-n-propylamineoxide (LAPAO) Dodecyl-N-betaine (zwittergent 3-12) Tetradecyl-N-betaine (zwittergent 3-14) Myristoylphosphoglycerocholine Palmitoylphosphoglycerocholine 3-[[3-cholamidopropyl)-dimethylammonio]-1propanesulfonate (CHAPS) Deoxycholic acid Cholic acid Taurodeoxycholic acid Glycocholic acid Sodium dodecylsulfate (SDS) in 50 mM NaCI Dodecylammonium ClGanglioside GM , PEG-dodecanol Polyoxyethylene glycol detergents CsE 6

ClO E6 C'2 E6 C12 ES C'2 & 14 E9.5{Lubrol PX) C'2 E12 C,2E23 (Brij 35) C,6&1SE17 (Lubrol WX) tert-p-Cs0E9.5(Triton X-1 00) tert-p-Cs0R7.S (Triton X-114) C12 sorbitan E20 (Tween 20) C1S:1 sorbitan E20 (Tween 80) Cetyltrimethylammonium bromide (CTAB)

Critical micelle concentration (M)

Aggregation number

x 10- 2 x 10-4

292 528 229 302 336 350 486 500 615

2.5 1.7 2.2 3.3

393 409 500 466

3 x 10- 3 1 x 10- 2 1.3 x 10- 3

27 140 75

x 10- 3 x 10- 3

8 x 10- 2 6 9 1 5

x x x x

87 130

10- 3 10- 5 10- 5 10- 3

22 4 20 6 62

8 x 10- 3 15 x 10- 3

55

10-9

150 130

x 10- 4

394 422 450 538 620 710 1200

1 x 10- 2 9 x 10- 4 8.2 x 10- 5 8.7 x 10- 5

32 73 105 120 100 80 40 90 140

9 x 10- 5 9 x 10- 5

1000 1625 540 1240 1320 364

3 x 10- 4 2 x 10- 4 6 x 10- 5 1.2 x 10- 5 9.2 x 10- 4

60 169

MW, molecular weight. Source: Jain, M. K., and R. C. Wagner, Introduction to Biological Membranes, 2nd ed. New York: Wiley, 1988, p. 71.

TABLE 3.2. Effect of ionic strength on micelle formation by ionic surfactants in aqueous solutions at 25: C Surfactant Anionic Sodium n-octylsulfate Sodium n-decylsulfate Sodium n-dodecylsulfate (SDS) Sodium n-dodecylsulfate Sodium n-dodecylsulfate Sodium n-dodecylsulfate Cationic n-Dodecyltrimethylammonium bromide n-Dodecyltrimethylammonium bromide n-Dodecyltrimethylammonium bromide n-Dodecyltrimethylammonium bromide

Medium

CMC (mM)

N

piN

H2 O H2O H2 O 0.1 M NaCI 0.2 M NaCI 0.4 M NaCI

130 33 8.1 1.4 0.83 0.52

58 91 105 129

018 0.12 014 013

14.8 10.4 7.0 4.65

43 71 76 78

0.17 0.17 0.16 016

H2O 0.0175 M NaBr 0.05 M NaBr 0.10 M NaBr

N, aggregation number; p, micellar charge. Source: Jones, M. N., and D. Chapman, Micelles, Mono/ayers, and Biomembranes. New York: Wiley-Liss, 1995, p. 68

Detergents

47

I I I I I I I I I I I I I /

~

2

(/)

V

..c

c: c:

.:2 0:1

'c:"" Q)

U

Micelles >,

t: V

0. 0 0. '""

c:

Monomers

.g :l

(3

------~-

C/)

/

/

/.

c:

0

u CMC

Total concentration 3.4. The critical micellar concentration. As detergent (or surfactant) is added to an aqueous solvent, the concentration of dissolved monomers increases until the critical micellar concentration (CMC) is reached. At that concentration, micelles form. Further addition of detergent increases the concentration of micelles without appreciably affecting the concentration of monomers. Redrawn with permission from Helenius, A., and K. Simons, Biochim Biophys Acta. 1975, 415:38.

detergents exhibit even more fluctuations in shape as they can deform, split, and fuse over time. Micelle formation is a direct consequence of the degree of amphiphilicity of surfactants. The surfactant molecules that form micelles are more water soluble than most lipids but still contain nonpolar groups with a propensity to form hydrophobic domains. They also tend to have conical shapes with bulky headgroups relative to their nonpolar groups (see Figure 2.17). In addition to detergents, Iysophospholipids (phosphol ipids lacking one acyl chain) form micelles, as do PLs with very short acyl chains (e.g., PC with four to nine carbon chains) under certain conditions. Self-association of detergents into micelles is strongly cooperative and occurs at a defined concentration called the critical micellar concentration, or CMC (Table 3.1). Below the CMC, the amphipath dissolves as monomers; as its concentration increases beyond the CMC, ideally the monomer concentration is unchanged while the concentration of micelles increases (Figure 3.4). The CMC can be detected by measuring surface tension or other aqueous properties, such as conductivity or turbidity (Figure 3.5). Micelle formation is dynamic, allowing constant interchange between constituents of aggregates and soluble monomers. For ionic surfactants, it is strongly affected by ionic strength (see Table 3.2). Micelle formation is also a function of temperature. The critical micellar temperature (CMT) is defined as the temperature above which micelles form (Figure 3.6). The Krafft point, also called the cloud point, is the temperature at which a turbid solution of surfactant becomes clear due to the formation of micelles.

Concentration

3.5. Variation in surface tension (y), specific conductivity (K), and turbidity (T) as a function of detergent concentration. The schematic plots show the dependence on concentration of detergent (surfactant) in solution of properties commonly used to find the CMC. (Note that conductivity only applies to ionic surfactants.) At the CMC, denoted by the dashed line, there is a break in the line for each property. Redrawn from Jones, M. N., and D. Chapman, Micelles, Mono/ayers and Biomembranes, Wiley-Liss, 1995, p. 65. © 1995. Reprinted with permission from John Wiley & Sons, Inc.

Thus the Krafft point falls at the intersection of the lines for the CMT and the CMC, and at the Krafft point the temperature dependence of solubili ty rises steeply as the result of micelle formation. At the Krafft point, insoluble crystalline detergent is in equilibrium ,\lith monomers and micelles, so if the temperature is lowered, the detergent crystallizes out of solution. A familiar illustration is the precipitation of SDS in aqueous solutions below 4°C (its Krafft point). The CMT CMT ~

E

d

Detergent crystals

Detergent miceJles

.g 0:1

l::::

c:Q)

u

c: ou

L-----rCMIC Detergent monomers Temperature,OC

3.6. Detergent phase diagram. At temperatures below the Krafft point, the detergent exists as monomers at very low concentrations and insoluble crystals at higher concentrations. Raising the temperature increases the monomer concentration until the critical micellar temperature (CMT) is reached, when micelles form. At (and above) that temperature, the solution clears at temperatures because the only two phases present are micelles and monomers. The Krafft point falls at the intersection of the lines for the CMT and the CMC, where the temperature dependence of solubility rises steeply due to micelle formation, Redrawn from Helenius, A., and K. Simons, Biochim Biophys Acta. 1975,415:37. © 1975 by Elsevier. Reprinted with permission from Elsevier.

Membrane

strongly dependent on the ionic strength of the aqueous medium (see Table 3.2), as well as the kind of counterions available to shield the charged headgroups. Membrane Solubilization

+Detergent

Membrane "vith bound detergent

+More detergent

Mixed micelles: Detergent-Ii pid-protei n complexes

+More detergent

+

Mixed micelles: Detergent-protein complexes and detergent-lipid complexes

3.7. The stages in membrane solubilization. This schematic illustration follows the addition of increasing amounts of detergent to a membrane. Initially, integral membrane proteins are embedded in the lipid bilayer. At low concentrations of detergent, some detergent molecules penetrate the bilayer but do not disrupt it. As more detergent is added, disruption of the bilayer results in mixed micelles containing detergent, lipid and protein. At even higher detergent concentrations, most of the lipid is removed from the protein, prodUCing detergent-protein complexes, along with detergent-lipid complexes.

for nonionic surfactants and the common bile salts is below G°c. The size of detergent micelles is usually described by the aggregation number (N), the average number of surfactant moJecu les per micelle, although for some situations the molecular weight or hydrodynamic radius is reported (Table 3.J). The aggregation numbers given in the literature are averages, and the size distribution may be quite large. Micelle size can be determined by light scattering, ultracentrifugation, viscometry, and gel filtration. It varies widely, reflecting the size of the nonpolar domain: N increases with increasing tail length for a series of surfactants in which only the hydrocarbon chain length is varied. For ionic surfactants, N is

Detergents are used to extract membrane lipids and proteins into an aqueous suspension. When a low concentration of detergent is added to a membrane. the detergent molecules intercalate into the bilayer. When a higher concentration is added, the detergent disrupts the bilayer and forms mixed micelles containing lipid, protein, and detergent (Figure 3.7). Mixed micelles vary considerably in structure and size. The detergent concentration must be kept above its CMC to maintain the mixed micelles. Sometimes adding still higher concentrations displaces the lipid completely, producing detergent-protein complexes Free of lipid. Thus both the detergent concentration and the detergentto-protein ratio are important variables that influence how a particular membrane protein will be extracted from the membrane. The behavior of the membrane protein in further purification and characterization steps will depend on detergent-protein and detergentdetergent interactions, along with detergent-lipid and lipid-protein interactions if lipid remains. The amount of a particular detergent that solubilizes the membrane is roughly propoltional to its CMC. Bile-type detergents solubilize segments or

PC

Logiludinal

vie\v

Cmss-seclional

vie\."

3.8. Schematic illustration of the structure of mixed micelles of bile salts and phospholipid sandwiches of bile salt detergent with lipids. Redrawn from Jones, M. N, and D. Chapman, Micelles, Mono/ayers and Biomembranes, Wiley-Liss, 1995, p. 97. © 1980 by American Chemical Society. Reprinted with permission from American Chemical Society.

Detergents

49 vQ) V

Q)

N

5

13

(a)

-

1i 4 ::I 1i "0 ::I VJ v 3 "0 VJ N

C

9

0-

"0 'v .£ 0 0- 2 .... p...

~

'"

5

VJ

0

.£

p...

~

a

0.0

0.2

0.4

0.6

0.8

1.0 a 2 4

Triton X-lOa concentration

6 8 10 12 14 16

Detergent concentration

3.9. Ratio of protein to phospholipid solubilized from epithelial cells by four different detergents. A spike at the low detergent concentrations indicates protein leaking out before the lipid is solubilized. 0, Triton X-1 00; _, sodium dodecylsulfate; 6, dodecyltrimethylammonium bromide; .... , sodium cholate. Redrawn from Jones, M. N., and D. Chapman, Micelles, Monofayers and Biomembranes, Wiley-Liss, 1995, p. 148. © 1991 by Elsevier. Reprinted with permission from Elsevier.

the membrane as detergent/bilayer sandwiches (Figure 3.8). The success of an extraction procedure is determined by checking the amount and composition of the desired component (usually protein) in the supernatant following sedimentation of the membrane. The

ratio of phospholipid to protein solubilized can indicate whether proteins leak from the membrane before it is completely disrupted, revealing whether a detergent concentration is suf-ficient to disrupt the membrane or only to solubilize segments of it (Figure 3.9).

3.10. Belts of detergents around purified membrane proteins. The positions of detergent molecules in solutions of detergent-solubilized proteins are revealed in neutron diffraction density maps obtained at different H20/D20 ratios to provide contrast variation. This image obtained with OmpF porin in ClODAO also shows C<x traces for protein obtained from the x-ray structure (protein is pink and detergent is green). From Pebay-Peyroula, E., et al., Structure. 1995,3:1053. © 1995 by Elsevier. Reprinted with permission from Elsevier.

Fixed barrier

Moveable balTier

I

'Y

~_!--~.-l'\'.l.~.l.l ...

.. .l.,L./..

'Yo

I

Aqueous phase

I~~~~

,

Trough

!

3.11. Formation of a monolayer in the Langmuir trough. The moveable barrier allows the area covered by the monolayer to vary. Redrawn from Jones, M. N., and D. Chapman, Micelles, Mono/ayers and Biomembranes, Wiley-Liss, 1995 p. 26. © 1995. Reprinted with permission from John Wiley & Sons, Inc.

For reconstitution experiments, it is often desirable to replace the solubilizing detergent with lipids to better imitate biological conditions. Methods for detergent removal include dialysis, gel filtration, adsorption to polystyrene beads, and pH changes. Detergents that have a low CMC, like Triton X-I 00 (CMC = 0.24 mM), are much more difficult to remove by dialysis than detergents like OG (CMC = 25 mM) because so little of the detergent is present as monomers. Gel filtration is most effective when there is a large difference in the sizes of the detergent micelle and the detergent-protein mixed micelle, and thus works best with detergents with small values of N (and high CMCs). Adsol-ption to polystyrene beads (e.g., SM 2 Bio-Beads) is effective for most detergents, including Triton X-I 00, OG, DDM, cholate, CHAPS, and C I2 E g . Of course it is a problem if the protein of interest also adsorbs to the beads, in which case the beads can be placed outside a dialysis chamber. Some ionic detergents, such as cholate and deoxycholate, precipitate at about one pH unit above their pKas (5.2 and 6.2, respectively), which greatly simplifies their removal provided the mildly acidic conditions do not harm the protein being studied. SDS can be precipitated after exchange of sodium for potassium, as potassium dodecylsulfate is insoluble at room temperature. For some hardy proteins (such as bacterial porins), complete removal of detergent is effected by precipitating the protein with organic solvents. Although detergents have been widely used in pUlifying proteins for crystallization studies, their disorder prevents their resolution in the resulting highresolution structures obtained by x-ray diffraction. To visualize the detergent in protein-detergent complexes, low-resolution images of the structure of detergent domains in crystals can be obtained by neutron diffraction with H 2 0ID 2 0 contrast variation (see Box 8.1 on neutron diffraction). In this procedure, several crystals are prepared that vary in their H 2 0ID 2 0 content, giving relative contrasts to the protein and detergent with respect to the solvent to allow visualization of individual components. Discrete belts of detergent around the nonpolar portion of the protein are clearly detected by neutron diffraction studies of membrane proteins

such as the l3-barrel OmpF porin (Figure 3.10). In the case of OmpF protein, the "hardness" of the detergent torus affected the observed shape: with softer detergents, such as OG, the belts of detergents fuse with those of their nearest neighbors. Clearly the size and shape of the detergent molecules - along with smaller additives, such as heptane - are crucial to the success of the crystallization process (see Chapter 8), which has led to much interest in new detergents. Lipid Removal

Although it is rare for detergent extraction to completely remove bound lipid from membrane proteins, lipid removal may lead to loss of biological activity. There are dozens of examples of proteins (including succinate dehydrogenase and other components of the electron transport chain, several ATPases, numerous transferases, and receptors) that are inactivated when stripped of lipid by detergent or organic solvent and are reactivated by addition of lipid (see Table 4.2 for more examples). The lipid requirement may be quite specific, such as the absolute requirement of l3-hydroxybutyrate dehydrogenase extracted from mitochondrial inner membrane for Pc. Even for the cases of a general lipid requirement, it is clear that a portion of the lipids in a biomembrane associates dynamically with membrane proteins. This boundary lipid, which differs in mobility from the bulk lipid of the bilayer, may be functionally important and can be studied in model membranes.

MODEL MEMBRANES

The functions of the membrane and of many of its components are lost upon its disruption, necessitating reconstitution of the membrane in an in vitro model system for studies to elucidate mechanisms of transport and energy transduction, to measure enzyme kinetics and ion flows, and to explore phase changes and microdomains. The availability of a wide variety of model membrane systems is fortunate as no single system is suitable for all the membrane components being characterized or for all the techniques used to study them. Hundreds of papers describe the applications of each classical system, while the promise of the newest membrane mimetics has yet to be realized. Monolayers

Amphipathic lipid molecules with sufficiently large hydrophobic portions will line up at an air-water interface with their hydrophobic tails in the air. Such monolayers are commonly formed in a Langmuir trough, a container with a movable barrier on one side that allows control of the area and measurement of the pressure of the monolayer (Figure 3.11). The surface pressure (n)

Model Membranes

51

expanded

gaseous

____ solid ZOO

400

A2/molecule

I

E

z

E l=

expanded

20

30

40

50

70

60

80

AZ/molecule 3.12. A surface pressure-versus-area isotherm for a monolayer of a fatty acid at the air-water interface. A diagram of the surface pressure (TI) versus the area per molecule shows the relationship between TI and the area of the molecules forming the monolayer. The isotherm reveals phase changes: the monolayer approaches a solid state at high pressure, changes to liquid states at lower pressures, and changes to gaseous states at very low pressures, as shown in the inset. Redrawn from Jones, M. N., and D. Chapman, Micelles, Mono/ayers and Biomembranes, Wiley-Liss, 1995, p. 27. © 1990. Reprinted with permission from John Wiley & Sons, Inc.

A.

is created by the dilference between the surface tension of the monolayer (y) and that of the air-water interface (Yo): 7t = 1'0 - y. The high surface tension of water means that it takes work to cover the area of the air-water interface. To decrease that work, the monolayer spreads over the surface, putting pressure on the movable barrier. Inward movement of the barrier to decrease the area increases the surface pressure of the monolayer. In forming monolayers, the composition is controlled and the amount of lipid is known. A surface pressure-versus-area isotherm, showing the relationship between 7t and the area of the molecules forming the monolayer, reflects phase changes (Figure 3.12). Highly compressed molecules are so condensed they approach a solid state, whereas at very low pressures, the molecules are so spread out they do not interact and are considered to be in a two-dimensional gaseous state. Between these extremes, the monolayer is in a Iluid phase described as liquid. The effect of chain length on the phase of the monolayer is revealed in the pressurearea isotherms for a series of monolayers composed of PC with varying acyl chains (Figure 3.13A). Two fluid phases, called L E (liquid expanded) and L c (liquid condensed), are evident in the curve for DPPC, but only B.

50

50 r x - - - - - , - - - - - - , . - - - - - , - - - - - ,

x

~

x

40

40

E 30

E 30 E '--'

Z

Z

E

(j)

....

(j)

'-< :::l

:::l if) if)

(j)

20

if) if)

(j)

'-
10,000 nm.

Aqueous compartment 3.25. The structures and dimensions of three types of liposomes. Multilamellar liposomes (MLVs) have many more layers than indicated. Comparison of small unilamellar liposomes (SUVs) and large unilamellar liposomes (LUVs) reveals the difference in curvature that results in more loosely packed acyl chains in SUVs. Redrawn with permission from Jones, M. N., and D. Chapman, Micelles, Mono/ayers and Biomembranes, Wiley-Liss, 1995, p. 119. © 1995. Reprinted with permission from John Wiley & Sons, Inc.

Multilamellar vesicles (MLVs) contain concentric spheres of lamellae and may be made by simply shaking a thoroughly dried lipid film into an aqueous solution. They are usually polydisperse, with diameters from 0.2 to 50 IJ.m, and have as many as 20 concentric layers of membranes. Their internal volume is unknown but quite small. To increase the internal volume, they can be converted to vesicles with one to four lamellae by extrusion through polycarbonate filters. They have been used in studies of lipid phase transitions by DSC and in many studies of enzyme and peptide binding. Their osmotic sensitivity allows quantitative measurements of solute uptake rates from turbidity changes.

Example (Luckey, M., and H Nikaido, Specificity of diffusion channels produced by lambda phage receptor protein o(Escherichia coli. Proc Nat] Acad Sci USA. 1980, 77:167-171) The liposome swelling assay {ollows absorbance changes as the MLVs respond to osmotic pressure; when the liposomes are suspended in hypotonic solutions,

Model Membranes

swelling results li'omwater entry that pushes the concentric bilayers further apart, causing a decrease in light scattering. In isotonic solution, swelling does not occur without solute uptake, so MLVs can be used to measure the rate of uptake, which is more sensitive than methods that measure the extent of uptake after reaching equilibrium. This liposome swelling assay was crucial to discovering the specificity of maltoporin (LamB protein) because it could distinguish among uptake rate::; of disaccharides (Figure 3.26).

61

0.7

Small Unilamellar Vesicles

Example (Beschiaschvili, G., and 1. Seelig, Peptide binding to lipid bilayers. Nonclassical hydrophobic effect and membrane-induced pK shifis. Biochemistry. 1992, 31: 10044-1 0053) This study elnployed SUVs made using PO PC with and witlwllt POPG, which had diameters of around 30 nm, to compare the binding of a small al11phiphilic peptide to that in larger vesicles. Since the acyl chains are less tightly packed in the small vesicles than in the larger vesicles, they have less lateral tension, which is related to membrane elasticity. High-::;ensitivity titrati011 calorimetry was used to measure the heats of reaction {or binding the cyclic peptide, an analog of somatostatin called SMS that is an amphiphilic peptide with a positive charge. l\!lonolayer studies had already indicated that SMS intercalated into lipid with little change in its conformation according to circular dichroism, and 2H NMR studies had shown it could diffuse rapidly on the surface when bound. The binding enthalpy for SMS was -7.3 kca 1111101 for SUVs, independmt of pH or lipid composition, in contrast to the enthalpy of binding to larger vesicles of -1.4 kcallmol. The ditfermce in t::.G for binding to the two classes of vesicles was less than 1 kcal/mol, indicating there is a large enthalpy-entropy compensation, which means el1tropy is not important in SUV binding but is the driving force for binding the larger vesicles. The entropy effect is explained in terms of the increased area of the bilayer that accompanies peptide binding, which lessens

Suc=",

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o Small unilamellar vesicles (SUVs) with diameters of 20 to 50 nm result from extensive sonication of MLVs. They are also made by extrusion through polycarbonate filters of defined pore size and can be further sized by gel filtration or gradient centrifugation. Another method to make SUVs is by injection of lipids in organic solvent into aqueous media, followed by removal of the organic solvent. SUVs are vel)' asymmetric due to their extreme curvature. For example, SUVs of PC are 22 nm in diameter and have 1900 and 1100 molecules in the outer and inner leaflet, respectively. Although the acyl chains are less tightly packed than in larger liposomes, the extreme curvature of SUVs makes it difficult to incorporate proteins.

+ LamB protein

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Time, min 3.26. The liposome swelling assay follows a decrease in optical density (OD) at 500 nm upon mixing MLVs with permeant solutes. The optical density tracings in the control panel show very little change observed in the absence of maltoporin (LamB protein). When MLVs have incorporated purified maltoporin (3 IJ.g/mg PL). the rate of uptake of maltose is significantly higher than that of lactose and sucrose. Redrawn with permission from Luckey, M .. and H. Nikaido. Proc Natl Acad Sci. USA. 1980. 77167-171.

the intemal bilayer tension of the more tightly packed acyl chains in larger vesicles. Large Unilamellar Vesicles

Large unilamellar vesicles (LUVs) with diameters hom 100 nm to 5 ~m can be made by freeze-thaw methods that induce fusion of SUVs. Since the introduction of commercial extruders that force the liposomes under nitrogen pressure through polycarbonate filters of defined pore size, more uniform LUV sizes have been obtained, especially with repeated extrusions. Other procedures make proteoliposomes in this size range by mixing the protein in detergent with an excess of lipid and then gently removing most of the detergent by dialysis or dilution. Dialysis is slow (often requiring days) and is successful for detergents with high CMCs and small aggregation numbers, such as OG, sodium cholate, and CHAPS. Dilution to well below the detel-gent CMC is rapid; micelles break up and proteins (along with detergent monomers) incorporate into the lipid vesicles, which are collected by centrifugation. The rate of detergent removal affects how well the proteins are distributed. Gel fiJtration is used to size the vesicles, as well as to separate proteoliposomes from excess detergent. LUVs have the advantage of large encapsulated volumes, up to 50 Llmol of lipid, but their disadvantages include heterogeneous size distributions and fragility of larger vesicles.

Example (Costello, M. 1., et aI., Morphology of proteoliposomes reconstituted with purified lac carrier protein li'om Escherichia coli. J Bioi Chem. 1984, 259: 1557915586,' Costello, M. 1., et al., Purified lac permease and cytochrome-o oxidase are functional as monomers. J Bioi Chem. 1987, 262:17072-17082)

3.27. A series of pictures showing the effect of Ca 2 + on the elastic compressibility of GUVs manipulated with micropipettes. The suction pressure is sufficient to hold the vesicles firmly on the pipette tips (A). When the suction pressure is reduced as they are brought together, the vesicles in 3 mM Ca 2 + (B) do not adhere to each other, while in 0.12 M sucrose (e), they do. From Akasyi, K., et aI., Biophys J. 1998, 74:2973. © 1998 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

Lactose permease (also called the LacY protein; see Chapter 10) was originally reconstituted by DC dilution followed by fi~eez.e-thaw/sonication to make proteoliposomes that were examined by fi'eeze-fracture EM. At a molar protein-to-lipid ratio o[ 1:2500, the majority of the proteoliposomes had diameters of 30 to 150 n111 and exhibited fairly even distributions of protein particles. Quantitation ofthe siz.e and distribution ofthe lactose permease with variation of protein/lipid ratios led to the conclusion that the protein was incorporated as a monDma To answer the question o[ the quaternary state of lactose permease during active transport, proteoliposomes were reconstituted with both lactose pennease and cytochrome o (a tenninal oxidase of the E. coli respiratoly chain) and energized by providing ubiquinol, generating an elecli'ical gradient ofaboU! -130 mV Alternatively, proteoliposomes containing the lactose pennease were suspended in bu[fers containing high levels ofK+ Q/1Cllater treated with valinomycin, which carries K+ across the 111embranes to dissipate the K+ gradient. Changes in energiz.ed states did not lead to dimerizat ion o[ the lactose permease, proving the reconstituted lactose permease was capable o[ both passive and active lactose transport as a I1101wmer. For many subsequent experimel1ts, LUVs were used to compare the activities o[ lactose pennease variants made when evelY residue of Lac Y protein was altered by mutation. Short-Chain/Long-Chain Unilamellar Vesicles Short-chain/long-chain uniJamellar vesicles (SLUVs) form spontaneously from aqueous suspensions of longchain phospholipid (saturated PC, PE, and sphingomyelin with acyl chain lengths of at least 14 carbon atoms) mixed with small amounts of short-chain lecithin (acyl chain lengths of six to eight carbons). They range in diameter from 10 nm to > 100 nm, depending on the ratio of short-chain to long-chain compo-

nents (increasing short-chain PL produces smaller vesicles). Inclusion of cholesterol can increase the size of the SLUV. While they have not been of general importance, SLUVs have been employed in functional studies of lipolytic enzymes because they are superior as substrates for the water-soluble phospholipases (phospholipase C and phospholipase A 2 ; see Chapter 4). Giant Unilamellar Vesicles Giant unilamellar vesicles (GUVs) are 5 to 300 !J.m in diameter. These giant liposomes are cell-size vesicles that are large enough to insert a microelectrode or to visualize surface sections by optical microscopy. They can be manipulated by micropipettes to test their elastic compressibility by their adherence to other vesicles (Figure 3.27). While they are generally viewed as excellent membrane mimetics, their large internal volume may be a disadvantage. Also, proteoliposomes of this size are very fragile. GUVs can be made by slowly hydrating lipid at low ionic strength and high lipid concentration, followed by sedimentation through sucrose to eliminate MLVs and amorphous material. Alternatively, a homogeneous population of < 100 !J.m diameter can be prepared by electroswelling, applying a voltage to the solution of lipids in 100 mM sucrose at 60 v C. Preparation of GUVs at high ionic strengths (comparable to physiological salt concentrations) requires 10% to 20% of a charged PL and millimolar concentrations of Mg 2+ or Ca 2+. GUVs may also be made from native membrane with addition of lipid: for E. coLi membrane, the optimal lipid concentration is 90 mg/ml. To better incorporate membrane proteins into GUVs, a fusion technique has been devised. First LUVs are prepared with the proteins and then coupled to fusion-inducing peptides. One such fusogenic peptide is a short ex-helix called WAE; since it is negatively charged, a positively charged target is

incorporated in the GUV to facilitate docking of the LUVs. Thousands of LUVs dock onto the surface of a GUV, and after a few minutes they fuse, as demonstrated by free diffusion of the lipids between them.

Example (Korlach, I, et aI., Characterization of lipid bilayer phases by confocal microscopy and fluorescence correlation spectroscopy. Proc atl Acad Sci US A.1999, 96:8461-8466) Two fluorescent probes were incorporated into CUVs of DLPClDPPClcholesterol. The probe Dil-C2o (1,1'dieicosanoyl-3,3,3',3'-tetramethylindolcarbocyanine perchlorate) partitions preferentially (3:1) in the La phase and the probe Bodipy-PC (2-(4,4-difluoro-5,7dimethyl-4-bora-3a, 4a-diaza-s-indacene-3-pentanoyl)-Ihexadeca11.0yl-sl1-glycero-3-phosphocholine) partitions preferentially (4:1) in the Ld phase. Their appearance in complementary regions of the images obtained by confocal microscopy facilitated phase assignments for a set of CUVs of varying compositions and gave conclusive evidence for the coexistence of separate lipid phases (see Chapter 2).

Mixed Micelles and Bicelles

Because micelles have little resemblance to bilayers, they are not generally considered to be good membrane mime tics. Even so, mixed micelles of phospholipid and detergent have been used in a multitude of studies of membrane proteins, especially when either detergent removal led to denaturation or detergent inclusion gave better activity. Micelles have been used to determine quaternary structure of membrane proteins by gel filtration and electrophoresis. Mixed Triton X- LOO micelles are especially useful for kinetic analysis of enzymes with phospholipid substrates, because variations in PL concentration (up to 15 mol %) have little effect on the micelle structure. Finally, due to their small size, micelles form isotropic solu tions that are advantageous for NMR studies of membrane-associated peptides and small proteins. An exciting advance for NMR studies that provided a better imitation of the membrane and avoided the severe curvature of micelles was the development of bilayered micelles, or bicelles. Bicelles are discoidal lipid aggregates composed of long-chain phospholipid and either detergent or short-chain phospholipid. The center of the disc is a lipid bilayer with its edges stabilized by the detergent or short-chain lipid (Figure 3.28). Bicelles made with detergent have a much lower detergent content than mixed micelles. Varying the longchain lipid can alter features of the bilayer, changing both headgroups (typically DMPC doped with DMPS or OMPG) and length of acyl chains (OM PC, OPPC, and OLPC) in ways that allow investigation of these features of membranes. The size of the bicelles is dependent on both the ratio of long-chain to short-chain PL (q) and

A.

4nm

20-40 nm 3.28. Schematic cross sections of bicelles. Bicelles contain a mixture of long-chain phospholipids, such as DMPC, and either shortchain PLs, such as DHPC (Al. or bile salt detergents, such as CHAPSO (B). The planar region is composed mainly of long-chain PLs, while the rim is formed by a monolayer of the short-chain PL (in A) or the bile salt detergent (in B). A polytopic membrane protein is shown incorporated in the bicelles in B. Redrawn from Sanders, C. R., and R. S. Prosser, Structure. 1998,6:1227-1234. © 1998 by Elsevier. Reprinted with permission from Elsevier.

the total concentration of PL (Cl). When q > 3 and CL is 15% to 20%, the bicelle diameter is 500 A and the bicelles orient in strong magnetic fields, allowing them to be used for solid-state NMR. When q < 1 and CL is 5% to 15%, the bicelle diameter is only 80 Aand these form isotropic suspensions suitable for high-resolution MR. Recently, bicelles have been used for crystallization of bacteriorhodopsin.

Example (Czerski, L., and C. R. Sa l1ders, Functionality ofa membrane protein in bicelles. Anal Biochem. 2000, 284:327-333) Kinetic analysis of the integral membrane protein diacylglycerol kinase (DCK, which functions to phosph01ylate the lipid diacylglycerol using Mg-ATP; see "Membrane Enzymes" in Chapter 6) shows its activity is optimal in mixed micelles containing decyl maltoside and cardiolipin and decreases in bicelles. The DCK activity in bicelles is dependent 011 lipid composition, demonstrating a preference for DMPC or DPPC with 3-([3-cholamidopropyl]dimethylammonio)2-hydroxy-l-propanesulfonale (CHAPSO). The kinetic data show a reduced V",ax rather than changes in KI1I , suggesting lillie perturbation at the subst rate-binding site. The enzyme activity exhibited by DCK in bicelles validates the use of this system for NMR studies. Blebs and Blisters

The goal of most reconstitution systems is to reproduce the membrane environment in a model system. A different approach is to use protrusions from the membrane, which are called blebs or blisters. Because they allow experimentalists to examine portions of membranes still attached to living cells, blebs are model membranes with a different set of advantages. Blebs are composed of the physiological mixture of lipids,

3.29. Bleb on Xenopus laevus oocyte surface viewed by confocal microscope. After using hypertonic stress to induce blebbing, the sample was treated with NBD-phallacidin, a fluorescent dye that stains actin. The bleb is readily visualized in the light image (A) and is absent in the fluorescence image (BJ, where the plasma membrane from which it derived is clearly stained. Similar results are obtained with a stain for tubulin, indicating the bleb lacks an associated cytoskeleton. Redravvn with permission from Xhang, Y, et aI., J Physio/. 2000, 523: 117-130. © 2000. Reprinted with permission from Blackwell Publishing.

providing a native environment for other constituents free of detergents. They preserve the asymmetric orientation of membrane proteins and may also preserve the distribution of lipids in inner and outer leaflets. Because they are still connected to the cell, they may be reached by diffusible intracellular compounds that modulate important membrane properties. Finally, they allow comparisons of different cells to test the effect of specific mutations or of up-regulation or down-regulation of membrane components. The formation of such protrusions of the plasma membrane is one of the many changes induced by eukaryotic cell injury. When a bleb bursts, the loss of the permeability barrier triggers the onset of cell death, but the events leading up to rupture are reversible. Cell surface blebbing has been observed '.\lith numerous cell types and may be caused by mechanical or chemical (e.g., depletion of ATP with potassium cyanide, injection of polar organic solvents, addition of iodoacetic acid) treatments, as well as bacterial infection of macro phages. The absence of actin and tubulin from blebs formed on oocytes of Xenopus laevus clea r1y indicates the bleb membrane is detached from the cell cytoskeleton (Figure 3.29). For this reason, the lateral mobility of certain integral membrane proteins measured by FRAP reveals faster diffusion rates in membrane blebs than in intact cells. Mass spectrometry shows a large number of (if not most) cell membrane lipids are found in the lipid composition of vesicles derived from blebs. Blebs have also been studied by confocal fluorescence microscopy, immunocytochemistry, EM, and patch clamp techniques. Example (Baumgart, T., et aI., Large-scale fluid/fluid phase separation of proteins and lipids i/1 giam plasma membrane vesicles. Proc Natl Acad Sci USA. 2007, 104:3/65-3170)

Membrane microdo111ains were observed in blebs induced by treat ing cultured fibroblast cells and leukemia cells with polar organic solvents. Imaging of two fluorescent dyes, napthopyrene that partitions preferentially in La phase and rhodamine-B-DOPE that partitions preferentially into Ld phase, shows that Lo-Ld fluid-fluid phase coexistence occurs in blebs at room tempemture as well as at 4°C (Figure 3.30). The variation in shape of the observed regions provides evidence for phase boundmy line tension. Selective partitioning into L o domains of the blebs by lipid-anchored proteins associated with rafis, obtained using antibodies against specific membrane proteins and green fluorescem protein/membrane protein chimeras, supports the hypothesis that the ordered regions ofthe blebs are rafis, which makes itthetirst physical demonstration of rafis in biological membranes on the micrometer scale.

In pathogenic bacteria, such as Neisseria gonorrhoeae, Pseudomonas aeruginosa, and Borrelia burgdorferi (Lyme disease), membrane blebs occur as part of the growth cycle and lead to the shedding of' membrane vesicles that probably have a role in the spread of infection. These blebs have been visualized by various microscopic techniques bu t have not been extensively used as model membranes. Bleb-like stnlctures called blisters have been made on giant liposomes containing reconstituted E. coli membrane fractions for patch clamp studies. Treatment with 20 mM Mg 2+ induces collapse of the liposomes, followed by the emergence of blisters of 50 to 100 ~lm diameters that are stable for several hours. Example (/yer, R., and A. H. Delcow; Complex inhibition of011lpF and OmpC bacterial porins by polyamines. J Bioi Chem. 1997,272:18595-/860/; Samartzidou, H., and A. H. Delcow; Distinct sensitivities of OmpF and PhoE poril1s to charged modulators. FEBS Lett. /999, 444:65-70)

3.30. Evidence for coexisting Ld and lo domains in blebs derived from mammalian plasma membranes. Direct visualization of large lateral domains in membrane blebs of cultured mammalian cells at 25"C is achieved by labeling with specific dyes, napthopyrene in A and rhodamine-DOPE in B. Previous experiments have shown that naphthopyrene preferentially labels the Lo phase and rhodamine-DOPE preferentially labels the lei (L,,) phase. Blebs are induced with injection of 4% (VN) ethanol. Scale bars 5 >J,m. From Baumgart, T., et ai, Proc Natl Acad Sci. USA. 2007, 104:36153170. Reprinted with permission of PNAS.

Model Membranes

65

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Proteins at the Bilayer Surface

81

A.Cl

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4.16. Four types of membrane-binding domains found in hundreds of peripheral proteins involved in signal transduction. The x-ray structures reveal major features of four membrane-binding domains: A. C1 domain from protein kinase C; B. C2 domain from PLA2; C. FYVE domain from Vps27p (a yeast protein for endosomal maturation); and D. PH domain of phospholipase C. Selective residues that make contact with the surface are labeled, and specifically recognized lipids are modeled. PI3P is phosphatidylinositol 3-phosphate and PI(4,S)P2 is phosphatidylinositol 4,S-bisphosphate. The membrane leaflet is divided into an interfacial zone and the hydrocarbon core. Hydrophobic residues are colored green and basic residues are blue. From Hurley, J. H., and S. Misra, Annu Rev Biophys Biomol Struct 2000,29:49-79. © 2000 by Annual Reviews. Reprinted with permission from the Annual Review of Biophysics and Biomolecular Structure, www.annualreviews.org.

Zn 2+ very tightly. As described for PKC (above), this domain binds DAG and phorbol esters. The binding occurs at the tip of the domain, unzipping the two 13strands to expose the binding site. This groove is located in a hydrophobic end of the domain that is adjacent to a ring of basic residues posi tioned so that membrane penetration by the hydrophobic tip allows the basic ring to contact the membrane surface. Most Cl domains are not targeted to the membrane if they lack specific binding sites for DAG, although there are a few atypical Cl

domains that do not require DAG. Most PKCs and DAG kinases have pairs of Cl domains, and in some, DAG is an aHosteric activator. C2 domains, identified by a conserved sequence motif that binds Ca2+ reversibly, have been found in >400 proteins, including many involved in signal transduction, inflammation, synaptic vesicle trafficking, and membrane fusion. The C2 domain is a f3-sandwich like the immunoglobulin fold, with the Ca 2+ -binding sites formed by three loops at a tip analogous to the

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4.17. Helical wheel of the amphipathic helix of cytidyltransferase. Since the pH at the surface of anionic (but not zwitterionic) membranes is lower than the bulk pH due to the attraction of protons to the negative surface, the probability of protonation of three Glu residues increases, which effectively increases the hydrophobicity of the surface of the peptide. Redrawn from Johnson, J. E. et aI., J BioI Chem. 2003, 278:514-522. © 2003 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

antigen-binding site. It is not easy to generalize about this domain. There are five different C2 domain structures that Fall into two permutations of this fold. Some are hydrophobic enough to penetrate the membrane, while others are not; most require acidic PLs, while the C2 domain of the c isoform of PLA 2 prefers neutral lipids, especially PC. There are even C2 domains that do not bind calcium! The FYVE domain, identified in ~60 proteins, is noted For its specificity in binding phosphatidylinositol 3-phosphate (P1P3), which enables it to target proteins to endosomal membranes that are enriched in PIP 3 . This domain consists of 70 to 80 residues, forming two small double-stranded l3-sheets and an <x-helix, with a conserved RKHHCR motif that binds PIP 3 . It also contains eight Cys or seven Cys with one His that coordinates two Zn 2+ ions, two Leu residues at one end that protrude into the membrane, and a few less-conserved Lys residues that probably contact the membrane surface. The PH domains bind different phosphoinositides with varying degrees of specificity and thus respond to signaling that interconverts phosphoinositides having different phosphorylation patterns. Found in >500 proteins, this domain consists of two curved l3-sheets of three or four strands capped by an <x-helix. It has different subsites that bind phosphate groups to make up the substrate-binding site of varying a ffinity and specifici ty. It has a positively charged face that interacts with acidic lipids in the membrane; mutations that strengthen this

interaction can result in constitutive activation, while mutations that decrease it can lead to loss of function. Some PH domains also participate in proteinprotein interactions and some provide allosteric regulation. Like the other domains, adjacent portions of the PH domain contribute nonspecific interactions to give variable interplay with the membrane. A less common mechanism for binding to the membrane is the insertion of an amphipathic <x-helix parallel to the plane of the bilayer, observed in a miscellaneous group of proteins including DnaA, adenosine diphosphate (ADP)-ribosylation factOl~ vinculin, epsin, several regulators of G protein signaling, and cytidyltransferase (CT). The hydrophobic interactions between the hydrophobic face of the helix and the nonpolar core of the bilayer provide the driving force for insertion, which is opposed by the resulting perturbation of lipid packing. Formation of the <x-helix concomitant with insertion can provide a significant additional driving force (see below). CT is a well-characterized representative of this group of amphitropic proteins. It carries out the transfer of a cytidyl group fTom cytidine triphosphate (CTP) to phosphocholine in the key regulatory step for the synthesis of Pc. Studies employing circular dichroism indicate that its ~60-residue membrane-binding domain changes from a mix of l3-strand, l3-turn, and disordered conformations to <x-helix upon binding the membrane. The amphipathic helix has basic residues on one face and a mixture of acidic and basic residues on another,

Proteins at the Bilayer Surface

83

with a center strip dominated by acidic residues. A nonpolar face contains a total of 18 hydrophobic residues giving a hydrophobic surface area of ~2500 A2 .lnterestingly, three glutamate residues positioned at the interfacial region contribute to the selectivity of CT for anionic lipids because they become protonated in the low pH milieu at the surface of anionic, but not zwitterionic, membranes (Figure 4.17). Studies of CT bindi ng to multilamellar vesicles (MLVs) suggest that the first step of membrane association is electrostatic: CT binds when the MLVs are in gel phase, but can only insert its amphipathic helix upon raising temperature above the transition to liquid crystalline phase. Modulation of Binding Since membrane binding regulates the actIvIty of amphitropic proteins and thus controls many key processes of the cell, modulation of reversible binding to the membrane is crucial. It is accomplished by one or more of several mechanisms that respond to signaling kinases, altered levels of ions or effector molecules, or changes in local compositions of the membrane that are sometimes linked to trafficking, which is the targeted movement of specific molecules to particular regions or organelles. A simple mechanism is the disruption of the electrostatic interactions between basic groups of the protein and anionic lipids by the addition of negative charges when the protein is phosphorylated on serine or tyrosine residues in the membrane-binding region. In the example of the Nterminal region of Src, the phosphorylation site is a serine located between the clusters of lysine and arginine residues. Because the phosphate can be cleaved by phosphatases, the reversible change in charge provides an "electrostatic switch" that affects membrane binding. Another mode of regulation is by the enzymatic addition ctnd removal of an acyl chain. In these cases, the first lipid anchor is permanently added in a posttranslational modification and barely provides enough energy for the protein to bind to the bilayer. Modification with a second acyl chain (or prenyl group) doubles the hydrophobic interactions with the bilayer, converting a weak association into a strong one; thus its addition and removal regulate localization to the membrane. For example, after the ex subunit of the G, protein complex is myristoylated in a posttranslationaJ modification that remains for the lifetime of the protein, specific addition and removal of a palmitate chain by acyltransferase and thioesterase activities control its affinity for the membrane. A third major type of regulation involves binding ligands such as nucleotides or calcium. The binding of guanosine triphosphate (GTP) to the G pl-otein ADPribosylation factor causes a conformational change that exposes its amphipathic helix, which then inserts

4.18. Model for the Ca 2 + -triggered extrusion of a Trp residue in annexin V. The ribbon diagrams show annexin V viewed from the side with the membrane-binding surface face up and the mobile Trp residue at the right. Ca 2 + ions are blue spheres. A. If Ca 2 + is absent from domain 3, the Trp side chain is buried. B. When Ca2+ binds domain 3, the Trp side chain emerges from its buried position to interact with the lipid bilayer. From Seaton, B. A., and M. F. Roberts, in K. Merz and B. Roux, Biological Membranes, Birkhauser, 1996, p. 38S. © 1996 by Springer Verlag. With kind permission of Springer Science and Business Media.

into the bilayer. Calcium binding can trigger structural changes in the protein that affect its membrane-binding surface, as seen in annexin V. The crystal structure of annexin V shows a flattened molecule with opposing convex and concave faces. EM studies suggest the convex side flattens on the surface of the membrane, aJlowing multiple Ca2+ -binding loops to contact the surface. Ca2+ binding triggers the rotation of a single tryptophan side chain, moving it from a buried position to a protrusion that intercalates into the bilayer (Figure 4.18). Binding calcium has a very different effect on the retinal protein recoverin. When recoverin binds Ca 2+, its conformational change induces the extrusion of a bound myristoyl group that becomes a membrane anchor (Figure 4. J 9). This "myristoyl switch" has been observed in other peripheral membrane proteins that are involved in signaltTansduction. Finally, those amphitropic proteins that bind a specific lipid component of the membrane, such as DAG or polyphosphol)dated inositols, are subject to temporal changes in the concentrations of their ligands in the bilayer, and these concentrations are controlled by the phospholipases in response to signaling. Furthermore, the different concentrations of the specific lipid ligands in various membranes ofeukaryotic cells can contribute

Calcium-myristoyl switch

2 Ca 2 + ~

•

T

R

4.19. Diagram of the calcium-myristoyl switch. A conformational change in recoverin enables it to extrude its bound myristoyl group upon binding calcium. The protruding acyl chain interacts with the lipid bilayer and activates the protein to prolong the photoresponse of rod and cone cells. From Seaton, B. A., and M. F Roberts, in K. Merz and B. Roux, Biological Membranes, Birkhauser, 1996, p. 393. © 1996. Reprinted with permission from J. Ames, M. Ikura, and L. Stryer.

to membrane-selective targeting, as seen in the enrichment of endosomal membranes for PIP,.

carries out its translocation. DT has three domains, a catalytic C domain at the N terminus, a receptorbinding R domain at the C terminus, and between them a T domain involved in translocation (Figure 4.20). After binding to a receptor on the cell surface, DT is cleaved into the A fragment consisting of the C domain and the B fragment containing both T and R domains; A and B are linked by two disulfide bridges. Because DT is internalized by endocytosis, it enters the cell in acidic membrane-bound compartments called endosomes. The low pH of the endosomes triggers an acid-induced conformational change that enables the T domain to insert into the endosomal membrane and translocate the A domain into the cytosol where it carries out ADP-ribosylation of elongation factor 2, inhibiting protein synthesis and leading to cell death. The T domain of DT is a bundle of 10 ex-helices, two of which are hydrophobic and insert as a helical hairpin that has been shown to span the membrane. In planar lipid bilayers at low pH (::;6), DT as well as its B fragment and isolated T domain have all been observed to form cation-selective channels, but the structure of the pore and how it translocates the A fragment is not known.

PROTEINS AND PEPTIDES THAT INSERT INTO THE MEMBRANE

While a fe\-\' amphitropic proteins have small segments that insen into the nonpolar domain of the bilayer, there are soluble proteins and peptides that insert more extensive portions to cross the bilayer, often via major conformational changes. Insertion of these TM domains is generally governed by the same forces that drive the insertion o[ TM segments of integral membrane proteins, described in detail below. Besides the hydrophobic and electrostatic forces, insertion of a peptide across the bilayer involves the perturbation of the acyl chains in the membrane, immobilization of the peptide, and possibly its unfolding or refolding. Clearly the state and lateral tension of the lipid are important: the enthalpy of peptide insertion into small unilamellar vesicles was greater for insertion into tightly packed vesicles than loosely packed ones. How soluble proteins and peptides such as toxins, colicins, and ionophores move into the membrane milieu to accomplish their functions is fascinating.

Toxins Some protein toxins that come from outside the cell and affect cytosol ic targets provide their own mechanism of translocation across the membrane. Decades of research on diphtheria toxin (DT) established the AB model, in which part A of the toxin carries out its catalytic attack on an intracellular target and part B

4.20. High-resolution structure of diphtheria toxin. Diphtheria toxin has three domains: the C (catalytic), R (receptor-binding), and T (translocation) domains. The active site cleft of the C domain contains the endogenous dinucleotide ApUp. Two helices of the T domain insert into the membrane as a helical hairpin. From Collier, R. J., Toxicon. 2001,39:1793-1801. © 2001 by The International Society on Toxinology. Reprinted with permission from Elsevier.

Proteins and Peptides that Insert into the Membrane

85

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4.21. Steps in the internalization of anthrax toxin. Anthrax toxin is made up of three proteins, PA (protective antigen), EF (edema factor), and LF (lethal factor). (1) PA binds to a receptor, either ATR or CMG2 (2) Cleavage by a furin protease removes PA20. (3) PA 6 3 self-associates to form the heptameric prepare. (4) Up to three molecules of EF and/or LF bind to the prepore. (5) Endocytosis brings the complex to an acidic intracellular compartment. (6) The low pH triggers conversion of the prepore to a pore, and EF and LF are translocated to the cytosol, where EF catalyzes the formation of cAMP and LF proteolytically inactivates MAPKKs. Redrawn from Collier, J. R., and J. A. T. Young, Annu Rev Cell Dev Bioi. 2003, 19:45-70. © 2003 by Annual Reviews. Reprinted with permission from the Annual Review of Cell and Developmental Biology, www.annualreviews.org.

The family of AB toxins has grown to include cholera toxin, pertussis toxin, tetanus and botulinum neurotoxins, and recently, anthrax toxin, although it is structurally more complicated than the others. Anthrax toxin is made up or three different proteins, PA (protective antigen), EF (edema factor), and LF (lethal factor). PA is the antigen for the anthrax vaccine in current use. PA binds to a receptor on the cell surface and then gets cleaved, releasing a 20-kDa fragment. The change in conformation of the remaining 63-kDa PA fragment allows it to form heptamers that can bind up to three molecules of EF and/or LF (Figure 4.2\). The low pH of the endosome triggers a conformational change in PA, enabling each protomer to insert t,vo f3-strands into the membrane, forming a pore that is a 14-stranded 13barrel. The other two proteins, EF and LF, unfold at least partially to translocate through this pore into the cytosol, where they exert their lethal inhibitory actions. The structure of the pore formed by anthrax toxin PA is very similar to that formed by (X-hemolysin ((XHL), a hemolytic toxin secreted by Sraphy/ococcus aureus in a soluble form. A major difference is that (XHL requires no catalytic domain or other inhibitors; it simply lyses human erythrocytes and other cells by pore formation, causing leakage. Another difference is that each 33.4-kDa polypeptide inserts into the membrane

before forming the heptameric prepore at the surface of the bilayer. Once the heptamer forms, each monomer extends a f3-hairpin, forming a f3-barrel pore (see Chapter 5) that is 52 A long and 26 A in diameter, with an inner diameter that is only ~ 15 Ain the narrowest part (Figure 4.22). The rest of the l3-sheet structure fonns a much wider cap, with hydrophobic residues at the rim that contacts the membrane bilayer. This mushroomlike structure was first observed for aerolysin, another l3-barrel channel-forming toxin, which is a virulence factor of Aeromonas bacteria that cause gastrointestinal disease and wound infections. Calicins

Colicins are bacteriocins, a class of toxins synthesized and released by bacteria to kill competing microorganisms. More than a third of E. coli strains produce colicins. These bacteria harbor plasm ids that encode specific colicins, along with specific immunity proteins, which are membrane proteins that ensure their own protection from the lethal action of the colicins. Colicins enter their target cells utilizing outer membrane receptors and either the Tol or Ton intermembrane translocation systems (see Chapter I J on TolC). Those in the channel-forming subfamily then insert into the

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A:coJicin Ja

'l

B:colicin B

C:colicin N

E:colicin A (C-domain)

F:colicin E1 (C-domain)

4.23. Structures of four colicins and the C domains from two others. Pore-forming domains are shown in blue and brown, connecting helices in red, translocation domains in green, and one immunity protein in purple. From Zakharov, S. D., et ai, Biochim Biophys Acta. 2004, 1666:239-249. © 2004 by Elsevier. Reprinted with permission from Elsevier.

I

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S

Proteins and Peptides that Insert into the Membrane cytoplasmic membrane to form voltage-gated channels that leak ions at a very great rate (> 10 6 ions channel-I seC I), depolarizing the membrane and eventually killing the cell. Colicins are proteins of around 60 kDa organized into three functional domains: the N-terminal T domain, which mediates translocation across the cell envelope; the R domain for receptor binding; and the C-terminal domain called C for cytotoxicity, or more specifically, P for pore formation. The x-ray structures of four colicins reveal a central helix or pair of helices usually connecting two structural domains in a dumbbell fashion and giving a very elongated shape to the molecules (Figure 4.23). The lengths o( the coiled coils of colicins 1a and E3 are 160 'A and 100 'A, respectively! In most of the channel-forming colicins, the C domain is a 10-helix globule, which characteristically contains a pair of hydrophobic helices and is very similar to the T domain of DT (compare E and F in Figure 4.23 to T in Figure 4.20). Although the average length of the C domain exhelices is ~ 13 amino acids - clearly not enough to span the bilayer - isolated C domains from many different channel-forming colicins have been shown to form voltage-gated pores in planar bilayers. Experiments with biotin-labeled single cysteine mu tants of the C domains of both colicin Ia and colicin A indicate that a large portion of the peptide chain (115 and 70 amino acids, respectively) moves to the opposite side of the membrane duling insertion and voltage-gated channel opening. Therefore, the sequence of events in channel formation is postulated to be binding to the membrane, unfolding to an extended flexible helical network in the interfacial layer of the membrane, helix elongation, and then insertion (Figure 4.24). It is possible that lipid curvature is involved in pore forma tion, as suggested for some peptides that insert into membranes.

Peptides Peptides that insert into membranes include many antimicrobial pep tides and peptide toxins - some well characterized, like melittin of bee venom, and others newly identified, such as the peptide toxins of spiders and sea anemones. Numerous antimicrobial peptides are under study not only for their role in the natural immune defense of mammals but also their potential as valuable new therapeutics either by themselves or as transporters, as in the case of the family called 'Trojan pep tides," which can deliver other agents into cells. Other pep tides capable of membrane insertion are the ionophores, named for their affinity for ions, which can be highly specific. While some ionophores simply chelate the ion, surrounding it with a lipid-soluble coat, others insert into the bilayer to make ion channels. Alamethicin and gramicidin are two widely stud-

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4.24. Model of two steps in pore formation of channel-forming colicins. A. After binding the membrane surface, the C domains unfold into a helical network that extends in the ~ 15 A-thick interfacial layer of the lipid headgroups, anchored by insertion of the hydrophobic helical hairpin. B. Insertion is predicted to involve helix elongation as well as toroidal configuration of lipids to make the open pore. In A, the membrane potential is trans-positive or is absent, and in B it is trans-negative. Anionic lipids that interact with basic side chains in the pore lining of the toroidal pore structure are red. From Zakharov, S. D., et aI., Biochim Biophys Acta. 2004, 1666:239-249. © 2004 by Elsevier. Reprinted with permission from Elsevier.

ied examples of channel-forming ionophores that have provided many insights for the understanding of protein ion channels (see Chapter 10). In addition, many synthetic peptides have been designed to insert into membrane bilayers. The wealth of data on peptide insertion into model membranes [Tom studies using conductance, Fourier transform infrared (FT1R) spectroscopy and oriented circular dichroism, solid-state NMR, neutron diffraction, and other techniques emphasizes the importance of the lipid composition, temperature, extent of hydration, and the peptide-to-lipid ratio. In general, the peptides can insert in one of two orientations, parallel to the plane of membrane or perpendicular to it, and then permeabilize the membrane by one of four possible mechanisms. In the Carpet mechanism, pep tides bind as ex-helices to the membrane surface and embed in the headgroup region oriented parallel to the bilayer plane. Although they remain in this orientation, at high concentrations they disrupt the membrane integrity

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the plane of the membrane even at high peptide-to-lipid ratios. Alamethicin is a 20-r-esidue peptide with eight helix-stabilizing amino isobutyric acid residues, only one charged amino acid (GluI8), an internal proline (Pro14l. an acetylated N terminus, and an alcohol (phenylaJaninol) at the C terminus. An amphipathic (Xhelix that can bend at Pro14, it forms voltage-gated ion channels of the barrel-stave configuration, as indicated by much experimental evidence. Its single-channel conductance is characterized by multiple discrete states, suggesting the channel is oligomeric and changes its conductance state when a single alamethicin molecule joins or leaves the aggregate. The pore dimensions determined by neutron scattering give it a thickness that matches the helix diameter. Solid-state NMR results indicate that in the nonconductive state, the helices are tilted by 10° to 20 from the bilayer normal, suggesting a possible "preaggregate" state that leads to oligomerization and an open channel. Gramicidin A, which forms channels that are specific for monovalent cations, is composed of 15 nonpolar amino acids of alternating Land D configurations. Gramicidin A can form ~-helices, which are helical structures made up of 13-sheets twisted into cylinders, with hydrogen bonding of the backbone N-H and car~ bonyl groups roughly parallel to the axis of the helix. The ~-helices have hydrophobic exteriors since with alternating L- and D-amino acids, all the side chains are on the outside. The Nand C termini are blocked, so with the lack of polar side chains, the most polar part of the molecule is the peptide backbone; indeed the ion path in gramicidin Ainvolves the carbonyl oxygens. Different gramicidin structures are observed depending on the solvent (lipid, organic solvent, or detergent) and ions present. Detailed structures obtained by x-ray crystallography and NMR have been classified as either a double ~-helical pore that can span the bi layer or a ~-helical dimel~ both of which can have open and closed states (Figure 4.26). Since conditions under which the double helix forms a conducting pore are velY limited, the helical dimer is probably the major conducting form and exemplifies the fourth mechanism for peptide channel formation. There are actually several species of natural gramicidins with slight differences in amino acid composition, as well as numerous synthetic analogs. Because of its self-associating dimer, gramicidin A has been found to be a suitable nanodevice for membrane biochips. 0

4.25. Two mechanisms for pore formation by inserted peptides: the barrel-stave model (A) and the toroidal model (B). See text for details. Redrawn from Yang, L., et al., Biophys J. 2001,81 :14751485. © 2001 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

(without forming pores). In two of the insertion mechanisms, the peptides bind in parallel orientation but oligomerize when a critical concentration is reached, changing their orientation to approximately perpendicular to the bilayer and resulting in pore formation. There are two mechanisms of oligomeric pore formation (Figure 4.25). In the barrel-stave model, the peptides span the two leaflets of the bilayer to line the pore like the staves of a barrel. In the other mechanism, the peptides form a toroidal pore when the insertion of helices stimulates the lipid monolayer to bend back on itself like the inside of a torus (a mathematical term for a surface containing a hole). In contrast to the barrelstave model, the toroidal pore has a continuous bending of a PL monolayer, stabilized by the peptides. In the fourth mechanism, the peptides insert to span the two leaflets of the membrane and then form an open pore when they align as dimer-s. Melillin is an amphipathic ex-helix of 26 amino acids, with five basic residues along its polar side and a bend of ~I20° at its internal proline. Because of its large pore size (4.2 nm inner diameter and 7.7 nm outer diameter), the hydration and temperature dependence of pore formation, and the lack of discrete conductance steps, melittin is considered to form a toroidal hole. In contrast, a solid-state NMR study of the antimicrobial peptide ovispirin supports the Carpet mechanism for its membrane disruption by showing it remains parallel to

SecA: Protein Acrobatics

Perhaps the best example of a protein that is so versatile it can move readily between the cytosol, the membrane surface, and the membrane interior is SecA, the motor for the translocon that exports proteins across the

Proteins and Peptides that Insert into the Membrane

89

A. Channel (helical dimer)

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Open

B. Pore (double helix)

.... Closed

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4.26. Channel and pore structures of gramicidin A. A. Gramicidin A forms a helical dimer when two f)-helices form a channel by associating end to end, as diagrammed schematically on the left. The hydrogen bonds are intramolecular except at the interaction between the two N-terminal groups. On the right is the high-resolution structure of the open helical dimer in SDS as determined by NMR. The polypeptides are portrayed with ball-and-stick models (one molecule is white and the other yellow), and the interior lumen of the channel is indicated by overlapping green spheres. B. In the pore form, gramicidin A forms a double helix with intermolecular hydrogen bonding along two antiparallel f)-strands, which opens in response to environmental factors (such as cations), as diagrammed on the left. The high-resolution structure of the cesium-containing open-pore double helix shown to the right was determined by x-ray crystallography. From Wallace, B. A., J Struct BioI. 1998, 121: 123-141. © 1998 by Elsevier. Reprinted with permission from Elsevier.

bacterial plasma membrane (described in Chapters 7 and 12), SecA is a remarkably flexible protein composed of an N-terminal ATPase domain and a C-terminal domain that binds membrane lipids as well as the preprotein to be exported and the cytoplasmic chaperone SecB (Figure 4.27). SecA is quite concentrated in the cytoplasm (5 ~M inside E. coli). Its membrane bind-

ing is dependent on the presence of anionic PLs, except at high temperatures, and is enhanced by nonlamellar lipids. It binds the SecYEG translocon with high affinity (20-40 nM KJ) and inserts repeatedly into the translocon in cycles that couple ATP hydrolysis to movements of the exported polypeptide chain (see Chapter 7). The resulting insertion is so extensive that several regions of

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4.27. The domain structure of SecA. The major SecA domains are the ATPase motor, the C-domain that binds lipids and SecB, and the substrate specificity domain (SSD) that binds to the preprotein. Three views are shown of the space-filling model of the x-ray structure of SecA from B. subtilis. While SecA is shown here as a monomer, two different x-ray structures show SecA dimers of quite different organization. From Vrontou, E., and A. Economou, Biochim Biophys Acta. 2004, 1694:67-80. © 2004 by Elsevier. Reprinted with permission from Elsevier.

the SecA protein can be chemically modified from the outside (periplasmic side). SecA can be cross-linked to portions of both SecY and SecE, although not to membrane lipids.

PROTEINS EMBEDDED IN THE MEMBRANE The operational definition of integral (or intrinsic) membrane proteins implies that they are embedded in the membrane, since disruption of the membrane is required to solubilize them. The exceptions are those peripheral proteins that are held in the membrane by two or more lipid anchors, binding them to the membrane with enough strength to require its disruption for their release (see above). Embedded membrane proteins include monotopic proteins, which insert into the membrane but do not span it, and proteins with one or more TM segments. Monotopic Proteins

There are only a few well-characterized examples of monotopic proteins. Some enzymes involved in lipid metabolism access their substrates by integrating into one leanet of the membrane. Structures have been solved for three of these: prostaglandin H 2 synthase (described in Chapter 9), squalene-hopene cyclase, and fatty acid amide hydrolase, which are all important pharmaceutical drug targets. A high-resolution structure is available for another monotopic protein with clinical importance: monoamine oxidase, which binds to the outer membrane of mitochondria. It is important for inactivation of several neurotransmitters, such as serotonin and dopamine, as well as catabolism of monoamines ingested in foods.

Another example of monotopic proteins is the caveolins, which associate with rafts to form caveolae (see Chapter 2). Caveolin is inserted into the plasma membrane from vesicles derived from the Golgi and remains in the inner leatlet, strongly immobilized by associations with the cytoskeleton. It forms dimers that bind cholesterol and form a striated coat as the membrane invaginates for endocytosis (see Figure 2.26). Integral Membrane Proteins

Most integral membrane proteins have one or more TM segments. Bitopic proteins span the bilayer one time and are classified by their topology as type I, \vith their N terminus outside, or type II, with their C terminus outside. Poly topic proteins are called type III integral membrane proteins and have multiple spans connected by loops. When several bitopic integral membrane proteins oligomerize with interacting TM segments, they are called type IV (Figure 4.28). Chapters 9, J 0, and J J present examples of detailed structures of integral membrane proteins. Up-todate lists of the membrane proteins with structures solved at or near atom ic levels can be found at http:// blanco. biomol. uci .ed u/Mem brane_Protei ns_xtal. h tml and http://wwwmpibp-frankfurt.mpg.de/m ichel/pu bl ic/ memprotstruct.html. The possible folds of integral mem brane proteins are dictated by the process of export to and assembly in the bilayer as well as by the stability of the embedded protein. These constraints on their folds may explain why all of the type III integral membrane proteins whose detailed structures have been solved are of two structural types, bundles of ex-helices and ~-barrels, described in detail in the next chapter. Since most of these structures were solved by x-ray crystallography, it is also possible that crystallization

Proteins Embedded in the Membrane

91

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TABLE 4.1. Properties of the cytosolic and membrane

environments that affect proteins

Bitopic

Polytopic TypenI

Oligomeric Type IV

Inside

Outside

4.28. Classification of integral membrane proteins by topology. Both type I and type II are bitopic proteins with only one TM helix. Type I has the N terminus outside while type II has it inside. Type III proteins have multiple TM segments in a single polypeptide. In contrast, type IV proteins are oligomers assembled from several polypeptides, each having one TM helix. From Nelson, D. L., and M. M. Cox, Lehninger Principles of Biochemistry, 4th ed., W. H. Freeman, 2005, p. 374. © 2005 by W. H. Freeman and Company. Used with permission.

methods favor these two structural types. With known atomic structures for fewer than 10% of all predicted integral membrane proteins, a more varied menu of structures, such as the I)-helix, will likely be revealed in [l.lture work. As many of the following chapters will illustrate, the TM ex-helix is currently the focal point for most researchers dedicated to understanding the nature of these intriguing proteins. The structures of membrane-spanning proteins must cope with many chemical and physical differences from soluble proteins, as indicated in a list of important environmental factors (Table 4.1). Notable among these are the lack of homogeneity and isotropy; the presence in most membranes of gradients of pH, electric field, pressure, dielectric constant, and redox potential; the paucity of solvent; and the very low dielectric constant. Some properties vary as a function of the depth in the membrane (see Chapter 8). Together these factors significantly alter the ~GO of functions associated with folding processes. Overall, it is much harder to break a main-chain hydrogen bond, ionize a side chain, or break a salt bridge of a protein domain in the membrane interior than in the cytosol; of course it is easier to expose a hydrophobic group, as well as to bring subunits in close proximity.

Property

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Plasma membrane'

Solvent chemical homogeneity Chemical groups available

Yes

No

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~Yes Isotropy pH gradient No ~O Electric field (V.m- 1 ) Pressure gradient No No Dielectric constant gradient Redox potential gradient No ~17 Volume or surface occupancy [protein/solvent (%)JC Separation between two proteins: Distance (A) ~50 Intervening solvent ~ 15-20 molecules ~1O-11 Exchange time between solvent molecules (s)d Viscosity at 20°C 0.001 (Tl; N.s.m -2) Dimensions 3 Translational diffusion: e ~lO-'O Dla, (m 2 .s -') ~250 Average range explored in 1 ~lS (x; A) Dielectric constant (c) 80 t.Go (kcal.mole- 1 ) for: ~O Breaking a main-chain H-bond Deprotonating a Glu side -4 chain (pH7) Opening a salt bridge 30 60 ~O

~O

5

a For properties that vary as a function of the depth in the membrane, the data correspond to those at the membrane center. b In most but not all membranes. C Estimated from data for the cytosol and plasma membrane of an E. coli cell. Calculations for the cytosol assume a 1:2.5 w/w ratio of RNA to protein; calculations for the plasma membrane assume the average integral protein (ohen an oligomer) to comprise ~ 12 TM helices and to have about half of its volume buried into the membrane. Estimates published in the literature vary from 17% to 50%. d In pure solvent. e For a middle-sized protein (~50 kDa) in either pure water or pure lipids; in the cytosol and in real membrances, diffusion coefficients vary with the distance range considered. Source: Popot, J. L., and D. M. Engelman, Annu Rev Biochem. 2000, 69:881-922. © 2000 by Annual Reviews. Reprinted with permission from the Annual Review a Biochemistry, www.annualreviews.org.

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The membrane milieu strongly favors the formation of secondary structure in TM segments. The low dielectric constant of the nonpolar domain of the membrane and the scarcity of water molecules favor formation of hydrogen-bonded secondary structure. This is readily shown by computing the change in free energy of transfer (~Glr) for peptide bonds with and without H-bonds:

Non-H-bonded-NH-C=O H-bonded-NH-C=O

~Glt- from water to alkane +6.4 kcal/mol +2.1 kcallmol

Therefore the per-residue cost of disrupting Hbonds in the membrane is ~4 kcaJlmol. For a TM segment of 20 amino acids, this is 80 kcaJlmol driving the formation of an (X-helix! Analysis of the solved integral membrane protein structures and mutagenesis of particular residues in them have led to some generalizations about the locations of amino acids in the nonpolar membrane domain: I. Nonpolar amino acids are typically found in membrane-spanning (X-helices with their side chains pointing into the hydrophobic interiOl- of the bilayer. This is expected from thermodynamic arguments and was tested when the small bitopic protein phospholamban from sarcoplasmic reticulum was engineered to replace nonpolar residues in its TM helix with polar ones, resulting in a water-soluble analog. 2. Acidic and basic amino acids either (i) remain uncharged due to the effect of the low dielectric environment on their pKas, (ii) form ion pairs that neutralize their charges, or (iii) playa special role, for example, in transport of protons or electrons or in binding a cofactor such as heme or retinal. Polar residues are not completely excluded from the nonpolar region, since the partitioning of many hydrophobic side chains into the interior is so favorable it can overcome the cost of including a few less-favorable groups. In addition, the charged residues found in TM segments can move their polar groups toward the interface by snorkeling, adopting configurations that orient their polar atoms to partially escape from the hydrophobic membrane core toward the interface. Snorkeling can be quantified by measuring the displacement of the polar atom(s) from the f)-carbon of the amino acid, which shows that the largest snorkeling distances are achieved by lysine residues. However, side chains of Arg, Tyr, Asp, Glu, Asn, and GIn also snorkel, in addition to Trp residues at the in terfaces. 3. Hydrogen-bond formers often use H-bonds to link their side chains to backbone carbonyl groups. These can provide "caps" for the ends of helices, as well as stabilizing the interactions between helices in a heli-

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cal bundle or oligomer. Even the hydrogens on the main-chain (X-carbons can form hydrogen bonds with backbone or side chain oxygen atoms. Several of these bonds, each wi th about half the strength of a conventional hydrogen bond, can stabilize a helix in the nonpolar bilayer interior and are often found in glycine-, alanine-, serine-, and threonine-rich packing interfaces. 4. Of the amino acids considered to be helix breakers, glycine and proline are found frequently in TM helices, often at conserved positions. Glycine ,-esidues are important for allowing the close packing that minimizes interhelical distances in bundles of (X-helices (see below). While proline residues are rare in (X-helices of soluble proteins, they are common in TM (X-helices and are often located near the center of the bilayer. The restricted backbone angles of the peptide chain at proline causes a kink in the helix. Inclusion of proline also leaves one carbonyl of the helix without an intrachain H-bond. The kink is a bend in the chain of ~120' in the direction away from the missing backbone Hbond. Interestingly, mutagenesis of bacteriorhodopsin showed that substitution of alanine for proline does not remove the kink, indicating that the tertiary structure of the integral membrane proteins maintains the helix distortion. The conservation of prolines at kinks in many integral membrane proteins led to the suggestion that the TM segments with kinks that do not have prolines evolved from TM segments with proline, since homologous proteins do have proline in the position of the kinks. 5. Aromatic amino acids, especiallyTrp and Tyr, play a special role at the interface of the hydrophilic and nonpolar domains in both (X-helical and f)-baITel integral membrane proteins. NMR studies with model indole compounds reveal that this is not due to their dipole moment or H-bonding ability but rather to their flat rigid ring and aromaticity that lead to complex electrostatic interactions with the hydrocarbon core. The well-characterized integral membrane proteins that are bundles of (X-helices typically have an even number of helices, with the notable exception of the family of seven-helix bundles that are involved in signal transduction, such as bacteriorhodopsin (see Chapters 5 and 6). Analysis of predicted membrane proteins from 26 genomes (see Chapter 6) shows the number of predicted TM helices is distributed over all integers from two to 13, with the occurrence decreasing as the number increases except for spikes at foul~ seven, and 12. When all the inner membrane proteins of E. coli are similarly analyzed, by far the highest incidence is for bundles of 12 predicted helices, with the next-largest groups havi ng two and six predicted TM helices and significant numbers with four, five, and 10 predicted TM helices.

Proteins Embedded in the Membrane

93

c.

D.

4.29. Distortions of a-helices in TM segments of integral membrane proteins. A. n-Bulge at Ala215 in helix G of bacteriorhodopsin, which causes the peptide plane to tilt away from the helix axis locally. B. Unwinding of helix M4 in the calcium ATPase of the sarcoplasmic reticulum exposes backbone carbonyl groups that participate in coordinating Ca 2+. C. Proline kink in helix C of bacteriorhodopsin, resulting in a lack of hydrogen bond to the carbonyl of Leu87. D. Half-helices in the glycerol facilitator, numbered 3 and 7. From Ubarretxena-Belandia, I., and D. M. Engelman, Curr Opin Struct Bioi. 2001, 11 :370-376. © 2001 by Elsevier. Reprinted with permission from Elsevier.

Distortions from the classical ex-helix are not uncommon in the TM segments seen in the x-ray structures (Figure 4.29). Many of the distortions probably have a functional role; alternatively they may serve to facilitate folding by preventing off-pathway intermediates (see Chapter 7). The proline-induced kinks described above are one class of distortions seen often in TM helices. Anothertype is a 7f bulge, where one backbone carbonyl is not H-bonded, such as a site involved in retinal binding in bacteriorhodopsin (see Chapter 5). Helix unwinding is a third kind of distortion, seen in the Ca 2 + pump from sarcoplasmic reticula (see Chapter 10), vvhere the unwinding frees up carbonyls to coordinate Ca 2+. Hal f-helices, where two short helices that do not individually span the bilayer stack end-toend to span the bilayer, have been observed, for example, in the glycerol facilitator and the aquaporins (see Chapter 10). A number of approaches have been taken to study helix-helix interactions. In model peptides that form TM helices, inserting glutamine in the middle of the TM sequence drives formation of oligomers. Indeed, any amino acid capable of acting simultaneously as both donor and acceptor of hydrogen bonds (Asp, Glu, Asn, and His) promotes oligomerization, while serine, threo-

nine, or tyrosine does nol.ln type III integral membrane proteins, H-bonds play an important role in the tertiary structure. For example, there is at least one hydrogen bond between each pair of helices in bacteriorhodopsin (see Chapter 5). Glycophorin A, the primary sialoglycoprotein of human erythrocyte membranes, has a single TM helix with a critical Gxx.x.G amino acid sequence that is needed for the helix-helix interaction of dimer formation (Figure 4.30). Finding this sequence in numerous other TM peptides has defined a TM-oligomerization motif of GxxxG, along with the less common Gxx.'\.A, which clearly reflects the importance of small amino acids at positions buried between the helices. In glycophorin A the critical sequence is LlxxGVxxGVxxT. Two helices cross at an angle of 40°, making a righthanded coiled coil in which the helices mesh closely by "knob-into-hole" interactions. The knobs are formed by isoleucine and valine residues and the holes by glycine residues. These interactions bring the helices close enough for important van del' Waals interactions along the coiled coil (see Figures 4.30 and 7.5). Even though its [unction is unknown, the detailed analysis of glycophorin A by saturation mutagenesis, NMR, and computational approaches provides a good model for

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coo4.30. The TM helices in a dimer of glycophorin A. Two views of the dimmer show the intermolecular contacts with residues colored as shown in the legend on the right. The two views of the TM helices differ by 90°. From Arkin, I. T., Biochim Biophys Acta. 2002, 1565:347-363. © 2002 by Elsevier. Reprinted with permission from Elsevier. B. The figure on the right shows the position of the TM helix in the entire polypeptide. From Garrett, R. H., and C. M. Grisham, Biochemistry, 2nd ed., Brooks/Cole, 1999, p. 271. © 1999. Reprinted with permission from Brooks/Cole, a division of Thomson Learning: www.thomsonrights.com. Fax 800 730-2215.

type IV proteins as well as for type III integral membrane proteins with multiple TM helices.

PROTEIN-LIPID INTERACTIONS

The properties of membrane proteins are best understood in the context of their lipid surroundings. Indeed, the contacts between integral membrane proteins and lipids must be very tight to maintain the seal of tbe membrane as a permeability barrier. The presence of a protein has essentially no effect on distant lipids but has a large effect on the shell or ring (annulus) of lipids that surround it, forming the interface between it and the rest of the membrane. These lipids are called annular or boundary lipids and can be distinguished experimentally from the bulk lipids of the bilayer. In addition to bulk lipids and annular lipids, there is a third class of lipids comprising those which are tightly bound in crevices or between subunits of the proteins. These lipids are called nonannular lipids (not to be confused with bulk lipids) or lipid cofactors, as they are frequently required for activity. Experiments with purified proteins have shown that many membrane proteins require specific lipids or classes of lipids to stably bind or insert into bilayers, while numerous enzymes require specific lipids for activity (Table 4.2). The activity of Ca2+ -ATPase, for example, increases as lipid is added up

to 30 moles of I ipid per mole of ATPase. Both the nature of interactions with annular lipids and the influences of the physical state of the lipid bilayer on the functions of membrane proteins have been extensively studied using EPR, fluorescence quenching and energy transfel~ and molecular dynamics simulations. Additional information can be gleaned from those lipids detected in highresolution structures obtained by x-ray crystallography described in Chapter 8. EPR is especially suited for studying boundary lipids because it can readily detect two populations of membrane lipids (Figure 4.31; Box 4.2). Since the mobility of the acyl chains is greatest near the center of the membrane (see Figure 2.11), inCOllJOration of a nitroxyl spin label close to the terminal methyl of the chain gives an EPR spectrum with quite narrow line widths in a pure lipid bilayel~ The rotation around the C-C bonds (trans-gauche isomerism, see Chapter 2) is fast (~1O-'O seconds) and averages out, so the spectrum results [Tom the axial rotation of the lipid molecule as a whole (10- 8 -10- 9 seconds). With proteins present, the axial rotation of a spin-labeled lipid molecule in the annular layer is hindered. Because it is relatively immobilized, it produces broader line widths, resulting in a second component most easily seen in the "outer wings" of the EPR spectrum. The selectivity of a protein for annular lipids can be determined From the relative intensity of the peaks in the outer wings, as obse,'ved for

Protein-Lipid Interactions

95

TABLE 4.2. Specific lipid requirements of membrane proteins and enzymes assessed by various techniques A. Lipid specificity for reactivation of delipidated enzymes Enzyme

Source

Cytochrome-c oxidase

Bovine heart mitochondria

j3-Hydroxybutyrate dehydrogenase

Bovine heart mitochondria

Sarcoplasmic reticulum Ca z+ -ATPase Monoamine oxidase

Delipidation by

Reactivation by

Reference

Cardiolipin, not PE, not PC

Sedlak and Robinson, Biochemistry 1999, 3814966

Phospholipase A

Only PCs

Sandermann et al., J Bioi Chem 1986, 261 :6201

Rabbit skeletal muscle

Cholate extraction

Phosphatidylinositol-4phosphate

Starling et aI., J Bioi Chem 1995,270:14467

Rat brain mitochondria

PLAz

PI, negatively charged PLs

J Bioi Chem 1981,

Huang and Faulkner, 256:9211

B. Lipid specificity in reconstitution of membrane proteins Protein

Source

Reconstituted in

Specific requirements

Reference

Acetylcholine receptor

Torpedo californica

DOPC vesicles

cholesterol, PA

Fong and McNamee, Biochemistry 1986, 25:830

Rhodopsin

Bovine retinal rod

Egg PC or DOPC/DOPE supported bilayers

PE (favors activated M2 state)

Alves et aI., Biophys J 2005,88:198

C. Lipid requirement of amphitropic proteins for binding and activation Protein

Source

Binding to

Specific requirements

Reference

Protein kinase C

Rat brain

PClPS LUVs

DAG and PS, anionic lipids

Slater et aI., J Bioi Chem 1994,269:4866

MARCKS

Mouse

PC/PS monolayer

Phosphocholinecytidylyltra nsferase

Rat

PC LUVs or SUVs

a series of spin-labeled lipids reconstituted with myelin proteolipid protein (Figure 4.32). The selectivity for differenl lipids is a reOection of different exchange rates, since there is constant exchange of lipids between the bulk and the annular layer (Figure 4.33). For the exchange equilibrium, LNP + L* ++ L N-! L'P + L, the lipid association constant, K" is ([L'P] [L])/([LP] [L'J). (Typically the concentration of the spin-labeled lipid, [L'"J, is less than 1 mole % of [L].) The reference lipid in these studies is PC, the most abundant PL in most animal cell membranes; thus the K, for PC in this example is 1.0. When there is no selectivity, K, = 1; with fairly high selectivity, K, approaches 10. The restriction of the bilayer to two dimensions produces a high effective concentration of lipids that can further enhance the selectivity. Since K, gives an average of affinities of a particular lipid, which may be due to several sites of quite different binding

Wang et aI., J Bioi Chem 2001,276:5013 Anionic lipids, unsaturated PE, DAG

Arnold and Cornell, Biochemistry 1996, 35:9917; Davies et aI., Biochemistry 2001, 40:10522

affinities, it may mask the presence of a tightly binding site for the lipid. A comparison of the lipid selectivity of different proteins shows that other proteins do not have the large variation in K r seen with myelin proteolipid protein and some, like rhodopsin, do not discriminate at all (Figure 4.34). Thus in general, the composition of lipids in the boundary layer appears to be quite similar to the composition in the bulk lipid bilayer. The exchange into the annulus (on-rate) is diffusion limited (~108 sec!) and thus is the same for different lipids, while the off-rate reflects the specificity of interaction with the protein and can be slowed to 10 7 or even 10 6 sec l . By performing experiments at different lipid-to-protein ratios, both K, and the fraction of spin-labeled lipid associated with protein can be determined. The stoichiometry of annular lipids for a number of different proteins has been found to correlate well

noterns In or at me tlilayer

96

Time scale 10- 9 sec

10- 8 sec

10- 7 sec 10- 6 sec Fluid lipids

•

•

Restricted lipids

-----==-==-- - - -

--:;;;.---------1\ l\

ij /.

_,-h

II

4.31. EPR detection of two populations of lipids: bulk and annular. A. Diagram of components of the membrane with time scales for the rotational mobility of each. The mobility of the annular lipid is about 1OO-fold slower than that of the bulk lipid. B. EPR spectrum of a lipid spin labeled on (14 (see Box 4.2). The solid line is the spectrum arising from contributions of two components, annular and bulk lipids, shown by dashed lines. The spectral ranges of the two components are identified over the spectrum, with the black indicating the more fluid (bulk) lipid and the gray indicating the restricted (annular) lipid. From Marsh, D, and L. I. Horvath, Biochim Biophys Acta. 1998. 1376:267-296. © 1998 by Elsevier. Reprinted with permission from Elsevier.

with the shape of the protein. Given a helix diameter of ~I nm and a lipid diameter of ~0.5 nm, a bitopic protein with one TM helix has 10 lipids in the shell around the helix. For poly topic proteins, the number of annular lipids is proportional to the number of TM helices and dependent on whether the geometric arrangement of the helices is sandwiches or polygons (Figure 4.35). An example is the Ca 2+ -ATPase with an estimated circumference of 14 nm, which was calculated to require a lipid shell of ~30 lipid molecules and was found by EPR

measurements to have 32 annular lipids. Similar calculations may be done for oligomeric proteins by treating the geometry of the arrangement of subunits instead of the TM helices. In integral membrane proteins that are /3-barrels, the TM segments are more extended and thinner, with diameters of ~0.5 nm. The predicted number of annular lipids works out to be the same as the number of /3strands if there is no tilt as they cross the bilayer; if there is a 60 tilt, there are about twice as many annular 0

Protein-Lipid Interactions

97

BOX 4.2. Electron paramagnetic resonance Electron paramagnetic resonance (EPR), also called electron spin resonance (ESRl, detects the orientation of unpaired electrons on paramagnetic molecules placed in a strong magnetic field. EPR can be used for biochemical substances containing paramagnetic transition metals or free radicals. To broaden its applicability, researchers employ free radical probes that are spin-labeled analogs of the molecules of A. interest, such as DPPC with a nitroxyl group. Such a compound, phosphatidylcholine carrying a spin probe on carbon 14 of the acyl chain (14PCSL), is shown in part A of Figure 4.2.1 o The absorption spectrum created by irradiation of the sample in the magnetic 4PCSL field is displayed as a first derivative, characterized by its intensity, line width, g6PCSL value (for positions), and multiplet structure. The nitroxide group gives three lines, 8PCSL whose line widths (related to the spin relaxJOPCSL ation times, T1 and T2) indicate the mobility of the molecule carrying the unpaired 12PCSL electron, which can vary from freely tumbling to strongly immobilized. In a series 14PCSL of PCSL probes, the position of the probe on the acyl chain determines its mobility. Placing the spin probe near the terminal methyl group gives relatively narrow lines due to the angular fluctuations from rotations around C-C bonds all along the chain, as shown in part A. The sharp lines that result are typical of isotropic mobilDescription of Approx rotational B. ity. Of course in a lipid bilayer, the spin spectra tumbling times (ns) probes do not rotate isotropically, but their large fluctuations average the orientational Freely tumbling O.J anisotropy when they are labeled near the end of the acyl chains. Cooling the lipid sample will slow the movement of the spin Weakly probe, broadening the lines, until it even0.6 immobilized tually produces a rigid glass or powder, as shown in part B of Figure 4.2.1. When the spin label is not free to tumble in all directions, it has anisotropic motion. The Moderately 2.5 frequent trans-gauche isomerizations along immobilized the acyl chains give motional averaging, while the axial rotation of the lipid is slower and is characterized by the rotational correlation times TRII and TR.L as shown in 5.0 part A. In addition to its application in studies of lipid-protein interactions described in this chapter, EPR is also used to probe protein Strongly -300 conformations with a procedure called siteimmobilized directed spin labeling (SDSL). The first step in these studies is the use of site-directed Rigid glass mutagenesis to create a single reactive site, -100°C >300 or powder typically by replacing an individual residue in I a protein with cysteine. Then reaction with a sulfhydryl-reactive spin label positions a 4.2.1. A. Dependence of the EPR spectra of nitroxide-Iabeled spin-labeled side chain at that site to proDPPC on the location of the spin probe. B. Effect of temperature on vide information about structure, orientathe mobility of a spin-labeled Pc. Both redrawn from Campbell, I. tion, and conformational changes in memD., and R. A. Dwek, Biofogica/Spectroscopy, Benjamin Cummings, 1984, pp. 197 and 192. © 1984 by lain D. Campbell and Raymond brane proteins. A. Dwek. Reprinted with permission from the authors.

- ~V-O:-;:T

rrOH~InS

98

14-SASL

14-PASL

14-PSSL

14-PGSL

14-PCSL

In or dl Ule Dlldyer

EPR experiments have addressed the detailed nature of the selectivity for annular lipids by comparing phospholipids with different headgroups and varying the ionic strength and pH, as well as by examining the importance of the glycerol backbone and the length of the acyl chains. In general, most proteins are found to prefer negatively charged lipids. Tn some cases, this is simply an electrostatic effect that is overcome by high ionic strengths, and in other cases the selectivity for the headgroup holds even in high ionic strength. (The few examples of headgroups resolved in highresolution structures reveal that extensive electrostatic and hydrogen-bonding interactions stabilize them in binding pockets, described in Chapter 8.) There is little or no difference when sphingomyelin and gangliosides are compared to PC, indicating the glycerol backbone is not a factor in selectivity. On the other hand, the acyl chain length is important: the free energy of association shows a linear dependence on chain length from 13 to 17 carbons. Hydrophobic Mismatch

4.32. EPR spectra of different lipids reconstituted with myelin proteolipid protein in DMPC. The protein/DMPC ratio is 23:1 and the temperature is 30°C. All the lipids contain a spin label on C14. They are stearic acid (14-SASLl, phosphatidic acid (14PASLj, phosphatidylserine (14-PSSU, phosphatidylglycerol (14PGSL), and phosphatidylcholine (14-PCSLl, where SL stands for the spin label in each case. The increasing relative intensity of the outer peaks arising from motion ally restricted lipids indicates increasing selectivity for the protein. Redrawn from Marsh, D., and L. I. Horvath, Biochim Biophys Acta. 1998, 1376:267-296. © 1998 by Elsevier. Reprinted with permission from Elsevier.

lipids as the number of I)-strands. The exchange rate was found to be slo\ver for annular lipids of I)-strands, presumably because lipids aligned along the I)-strands are more extended and less flexible than annular lipids around (X-helices. This result was determined with M13 coat protein, whose TM domains can be either (X-helices or I)-strands, and the exchange mte was four to five times slower for lipids associating with M 13 coat in 1)strand conformation than those associating with M13 coat in (X-helix.

© tt

~

4.33. Lipid exchange between bulk and annular lipids. Two lipids exchange at one "site" on the surface of a membrane protein, as L leaves and L' takes its place. Redrawn from Lee, A. G., Biochim BiophysActa. 2003, 1612:1-40.

The importance of acyl chain length on lipid-protein interactions produced the concept of hydrophobic mismatch, which results when the nonpolar region of the bilayer is thinner or thicker than the hydrophobic thickness l-equired by an integral membrane protein. The thickness of a bilayer is strongly influenced by its lipid composition: for example, a PC bilayer with saturated chains is 2.5 A wider than a bilayer with unsaturated chains of the same number of carbon atoms (see Chapter 2). If the hydrophobic regions of the protein and lipid do not match, either the lipid bilayer must stretch or compress to match the hydrophobic thickness of the protein (Figure 4.36), or' the protein must change by tilting helices or rotating side chains to fit to the bilayer to avoid exposing nonpolar groups to the aqueous environment. Since proteins are more rigid than lipids, the bilayer might be expected to deform to accommodate the dimensions of their TM segments, contributing to the lateral tension of the bilayer. This is observed when the thickness of lipid bilayers changes to accommodate gramicidin channels: insertion of gramicidin causes a DMPC bilayer to become 2.6 A thinner and a DLPC bilayer to thicken by 1.3 A. The perturbation of the membrane due to the mismatch creates a tension that contributes to its free energy: the t.GC' for bilayer deformation has been calculated to be ~ 1.2 kcallmol for a large hydrophobic mismatch of loA. While lipid bilayers adjust to accommodate gramicidin, they do not similarly accommodate single TM helices. Synthetic peptides designed to be TM helices of different lengths have no effect on the thickness of model bilayers. Rather, NMR measurements revealed

Protein-Lipid Interactions

99

PA

/' /' /'

/'

/'

/'

/' /'

/'

/'

/'

/'

/'

/'

/' /'

Kr

4.34. Patterns of lipid selectivity of different proteins. Kr , the relative association constant between each protein and each lipid, varies from 1 (shown in light gray) to > 6 (shown in dark gray). The data show the Kr for each protein {listed along the right edge}, with each of the lipids identified at the top. Lipid selectivity increases from front (with rhodopsin exhibiting almost no lipid selectivity) to the back (highest selectivity with PLP, the myelin proteolipid protein). Redrawn from Marsh, D., and L. I. Horvath, Biochim Biophys Acta. 1998, 1376:267-296. © 1998 by Elsevier. Reprinted with permission from Elsevier.

that TM helical peptides tilt with respect to the bilayer normal to match the hydrophobic thickness of the lipids. The peptides have sufficient flexibility of orientation to accommodate to the bilayer and not deform it. These findings suggest that proteins that cross the membrane with a small number of lX-helices are likely to accommodate the bilayer thickness by helix tilting. However, larger proteins or proteins with less flexibility impact the lipid enough for hydrophobic mismatch to induce changes in bilayer thickness. The latter includes proteins that cross the bilayer as l3-barrels (see Chapter 5), which have structural constraints thai prevent them from adapting to the lipid bilayer and thus are

more likely to select for lipids that provide hydrophobic matching (see Chapter 7). A comparison of relative binding constants for PCs with acyl chains of different lengths indicates that some integral membrane proteins bind more strongly to lipid that requires no change in bilayer thickness than to

A

dp

\,J

B.

Helical sandwich

Polygon

4.35. Geometries considered for determining the lipid-toprotein stoichiometry. Two geometries suffice to describe the stoichiometry of lipid to protein for integral membrane proteins with up to six TM helices. (With seven or more, there may be centrally located helices that do not contact the lipid.) On the left is a helical sandwich and on the right is a regular polygon. From Dc dl), the surface area they occupy decreases; when they compress (d l > d p ), it increases. Redrawn from Lee, A. G., Biochim Biophys Acta. 2004, 1666:62-87. © 2004 by Elsevier. Reprinted with permission from Elsevier.

IIUlt::lIl;:) III UI

lUU

Cll

1I1~

Ulldy'-=l

T ~

4.37. The consequence of hydrophobic mismatch in biological membranes may be a high-energy state as lipids and proteins try to compensate by extension of acyl chains (E), compression of acyl chains (C), and/or tilting of TM helices (T). Redrawn from Mitra, K., et aI., Proc Natl Acad Sci USA. 2004, 101 :4083-4088. © 2004 by National Academy of Sciences, USA. Reprinted by permission of PNAS.

lipid that requires such a change. Preference for chain length has been demonstrated with both rhodopsin and the photosynthetic reaction center. When covalently spin-labeled rhodopsin was reconstituted in PC with different-length saturated chains, it was active in DMPC (CJ4), segregated into protein-rich domains in DLPC (CI2), and aggregated in DSPC (CI8). Similarly, \-vhen the incorpora tion of photosynthetic reaction cen tel' into lipid bilayers was monitored by DSC, the T m for DLPC (CI2) increased by goC whereas that for DPPC (CI6) decreased by 3°e. These differences suggested that the protein partitioned into the gel phase with the shorter acyl chains and into the liquid crystalline phase with the longer acyl chains, since the bilayer is thicker in gel phase than in fluid phase. Such findings su pport the idea that hydrophobic mismatch could drive integral membrane proteins to regions of the bilayel- of appropriate thickness, which could be important in raft formation. Hydrophobic mismatch could also be involved in sorting membrane proteins to different membrane compartments. For example, along the secretory pathway that carries proteins to the plasma membrane in eukaryotic cells, some proteins remain in the Golgi apparatus, where they glycosylate secreted proteins. These Golgi-resident proteins have TM domains that are typically five amino acid residues shorter than the TM domains of proteins of the plasma membrane. This length difference was shown to be critical to sorting when constructs were engineered with shorter and longer TM segments. The bilayer thicknesses of the membranes of the secretory pathway have been determined by x-ray scattering. The ER, Golgi, basolateral, and apical plasma membranes from rat hepatocytes were treated with proteases (and puromycin and ribonuclease [RNase] as appropriate to remove ribosomes) prior to measuring their distances from P atom to P atom to determine bilayer thickness. The thickness of these membranes was expected to increase along the pathway, proportional to their cholesterol contenl. While the thickness does increase from the ER to the Golgi to the apical

plasma membrane, the basolateral plasma membrane is significantly thinner than the others. Since proteins targeted to the apical plasma membrane of the rat hepatocyte pass through the basolateral plasma membrane, hydrophobic mismatch must occur along the pathway. It is possible that the strain of hydrophobic mismatch puts the membrane in a high-energy state useful for vital functions such as fusion or protein insertion (Figure 4.37). In addition to protein sorting, hydrophobic mismatch is involved in membrane protein folding (see Chapter 7). The stress induced by mismatch is likely to affect the environment in which integral membrane proteins fold and assemble, which may at least partially account for the need for specific lipids in folding certain proteins. For example, the E. coLi transporter lactose permease (see Chapter 10) requires PE for correct folding but does not require PE for function. Thus it is proposed that the lipids have the role of chaperone in the folding process. With this understanding of how the special environment of integral membrane proteins constrains their structure and how they interact closely and dynamically with their boundary lipids, Chaptet- 5 focuses on the properties of some very well-characterized proteins. The following chapters describe the kinds of functions membrane proteins carry out, the structural principles used to predict their structures, and their folding and biogenesis.

FOR FURTHER READING

Reviews Peripheral Proteins

Gerke, v., C. E. Creutz, and S. E. Moss, Annexins: linking Ca 2+ signaJJing to membrane dynamics. Nal Rev Mol Cell Bioi. 2005, 6:449-461. Heimburg, T., and D. Marsh, Thermodynamics of the interaction of proleins with lipid membranes, in K. Men and B. Roux (eds.), Biological Membra l1es. Cambridge, Mass.: Birkhauser, 1996, pp. 405-462.

For Further Reading Hurley, J. H., and S. Misra, Signaling and subcellular targeting by membrane-binding domains. Annu Rev Biophys Biomol Struct. 2000, 29:49-70. Johnson, J. E., and R. B. Cornell, Amphoteric proteins: regulation by reversible membrane interactions. Biochim Biophys Acta. 1999,16:217-235. Mayor, S., and H. Riezman, Sorting GPI-anchored proteins. Nat Rev Mol Cell BioI. 2004,5:110-119. McLaughlin, S., and A. Aderem. The myristoyl-electrostatic switch: a modulator of r'eversible protein-membrane interactions. Ii-ends Biochem Sci. 1995,20:272-276. Seaton, B. A., and M. F. Roberts, Peripheral membrane proteins, in K. Merz and B. Roux (eds.), Biological Me1'l'lbrClnes. Cambridge, Mass.: Birkhauser, 1996, pp. 355-403.

101

Zakharov, S. D., et aI., On the role of lipid in colicin pore Formation. Biochim Biophys Acta. 2004,1666:239-249. General Features of Integral Membrane Proteins Curran. A. R., and D. M. Engelman, Sequence motifs. polar interactions and conformational changes in helical membrane proteins. CWT Opin Struct Bioi. 2003.13:412-417. Popot. J. L., and D. M. Engelman, Helical membrane protein Folding, stability and evolution. Annu Rev Biochem. 2000, 69:881-922. White, S. H., and G. von Heijne, Transmembrane helices before, during and after insertion. Curl' Opin Struct Bioi. 2005, 15:378-386. White, S. H., et aI., How membranes shape protein structure . .1 Bioi Chern. 2001,276:32395-32398.

Toxins and Colicins Collier, J. R., and J. A. T. Young, Anthrax toxin. Amw Rev Cell Dev Bioi. 2003, 19:45-70. Falnes, P.O., and K. Sandvig, Penetration of protein toxins into cells. Curr Opin Cell Bioi. 2000, 12:407-413. Gouaux, E., ex-Hemolysin from Staphylococcus aureus: an archetype of l3-barrel, channel-forming toxins. J Struct Bioi. 1998, 121: 110-122. Zakharov, S. D., and W. A. Cramer, Colicin crystal structures: pathways and mechanisms for colicin insertion into membranes. Biochim Biophvs Acta. 2002, 1565:333-346. Zakharov, S. D., and W. A. Cramer, Insertion intermediates of pore-forming colicins in membrane two-dimensional space. Biochimie. 2002,84:465-475.

Protein-Lipid Interactions Lee, A. G., How lipids affect the activities of integral membrane proteins. Biochim Biophys Acta. 2004, 1666:62-87. Lee, A. G., Lipid-protein interactions in biological membranes: a structural perspective. Biochim Biophys Acta. 2003,1612:1-40. Marsh, D., and L. 1. Horvath, Structure, dynamics and composition of the lipid-protein interface. Perspectives from spin-labelling. Biochim Biophys Acta. 1998, 1376:267296. Marsh, D., and T. Pali, The protein-lipid interface: perspectives ITom magnetic resonance and crystal structures. Biochim Biophys Acta. 2004, 1666: 118-141.

5

Bundles and Barrels

B. Structures of helical bundle and j3-barrelmembrilne proteins d Her in many respects, seen ,n the nbbon diagrams of the photosynthetic reaclion center from Rb. sphaeroldes (A) and themallopo.inlrimer frol11 E. coli outer membrane (B) A redrawn fro III Jones, M. R., et aI., Biochirn Biophys Acta. 2002. 1565:206-214 ,i 2002 by ElseVier. Reprinted Witt, permiSSIon from ElseVier. B redrawn 2003 by Elsevier. Reprinted wltl permiSSion from Elsevier from Wrmley, W. C, CurT Opin Struci Bioi. 2003, 13:404-411

The thermodynamic arguments discussed in the previous chapter make it clear that the TM segments of proteins will utilize secondary structure to satisfy the hydrogen bond needs of the peptide backbone. While a variety of combinations of secondary structures might be imagined in type ITl membrane proteins, all known protein structures cross the bi layer wi th ei ther ex-hel ices or l3-strands, producing either helical bundles or 13barrels. This chapter looks at how understanding structure and fu nction for a few proteins has provided the paradigms for these two known classes of integral membrane proteins.

x-ray structu re solved for mem brane proteins, that of the photosynthetic reaction center (RC). The majodty of integral membrane proteins whose high-resolution structures have been solved by x-I-ay crystallography exhibit the helical bundle motif (see examples in Chapters 9, 10, and 11). Helix-helix interactions have been analyzed in many of these, providing details of both tertiary and quaternary interactions. Identification of new integral membrane proteins in the proteome relies heavily on prediction of TM helices, as described in Chapter 6. Bacteriorhodopsin

HELICAL BUNDLES

Transmembrane (TM) ex-helices have dominated the picture of membrane proteins, guided by early stn.1Ctural information on bacteriorhodopsin and by the first

If a single protein dominated the thinking about structure, dynamics, and assembly of membrane proteins in the decades following 1970, that protein was bacteriorhodopsin (BR) from the purple membranes of the salt-loving bacterium Halobacler salinarum. From early

electrochemical proton gradient that supports the synthesis of AT? The ability of reconstituted vesicles containing BR and beef heart mitochondrial AT? synthase to synthesize AT? in response to light provided crucial early support for Mitchell's chemiosmotic hypothesis that the energy of an electrochemical gradient across the membrane could be lIsed to do work (Figure 5.2). Like rhodopsin. the light-absorbing protein in the rod outer segments of the eye's retina (see Chapter 9), BR has seven helices labeled A to G that span the membrane, and a retinal that is bound to a lysine residue in helix G via a protonated Schiff base (Figure 5.3). Whereas BR undergoes a light-induced photocycle involving conversion of the retinal from all-trans to J3-cis accompanied by conformational changes in the Light

+.+.• •

Bacteriorhodopsin

•

Cell~

wall

•

+

·.41~-~-~-[f.;-~ +

FJageJla 5.1. Schematic showing the different membrane domains of a halobacterium cell. The patches of purple membrane containing bacteriorhodopsin (BR) are separate from the regions of membrane containing the respiratory chain and the ATP synthase. Protons are pumped out of the cell in response either to light absorption by BR in the purple membrane or to cytosolic substrates for the electron transport chain. The ATP synthase normally uses the uptake of protons to drive the synthesis of ATP, although it can act as an ATPase and eject protons at the expense of ATP. Redrawn from Stoeckenius, w., Sci Am. 1976,234:38-46. © 1976 by Scientific American. Reprinted with permission from Scientific American.

structural images and spectroscopic characterizations, BR became the paradigm for ion transport proteins and indeed for ex·helical TM proteins in general. From the wealth of studies of its structure and function, a truly detailed understanding of this membrane protein has emerged. BR is the only protein species in the discrete membrane domains called purple membranes, the lightsensitive regions of the plasma membranes of H. salinarum (Figure 5. J). Together with specialized lipids, this protein forms functional trimers that pack as ordered two-dimensional arrays on the bacterial cell. In photophosphorylation BR functions to pump protons out of the cell in response to the absorption of light by its chromophore, retinal, converting light energy into an

Lipid vesicle

AD? +

Y

Mitochondrial FIFo-AT? synthase

~TP H

5.2. Schematic of reconstituted vesicles containing BR and ATP synthase. When the light is turned on, ATP is synthesized from ADP + P;. When the light is turned off, ATP synthesis stops. These vesicles gave important evidence to support the chemiosmotic theory of Peter Mitchell. Redrawn from Garrett, R. H., and C. M. Grisham, Biochemistry, 2nd ed., Brooks/Cole, 1999, p. 897. © 1999. Reprinted with permission from Brooks/Cole, a division of Thomson Learning: www.thomsonrights.com. Fax 800 730221S.

Bundles and Barrels

104

C4

Cl8

Cl9

C20

I

I

I

rCS~ /C7~ . . . . . C9.:::::::. /clr~ . . . . .C13..::::::....C6 C8 CIO Cl2 Cl4

I

C.- Lys216

I

C3 CI 'C2/ \~CI7 Cl6

~ hv

CIS

CI9

I C4

I

C20

I

/C5.:::::::.

C6

/C7~

I

. . . . . C9.:::::::.

C8

CIO

. . . . . Cll~

CI2

. . . . .CJ3~

CI4

I

C3 CI ' 0 / \~CI7 CI6

5.3. Retinal bound to lysine 216 in bacteriorhodopsin. The Schiff base linkage (shaded) between the aldehyde of retinal and the Eamino group of lysine 216 is protonated, as shown, before light stimulation. The all-trans retinal converts to 13-cis retinal upon absorption of a photon. Redrawn from Neutze, R., et aI., Biochim BiophysActa. 2002,1565:144-167. © 2002 by Elsevier. Reprinted with permission from Elsevier.

protein that result in proton transfer across the membrane, visual rhodopsin's activation cascade involves an II-cis to all-trans conversion of the retinal, followed by its dissociation from the protein. In addition to BR, H. salinaru111 contains three related rhodopsins, and similar molecules are found in eubacteria and unicellular eu karyotes. A.

B.

The purple membrane is composed of 75% protein and 25% lipid by weight, with 10 halobacteriallipids per BR monomer. These native lipids are based on archeol (see Figure 2.6), so they differ in headgroups but not in length of acyl chains. Delipidation by treatment with a mild detergent affects the kinetics of the BR reaction; addition of halobacterial lipids but not phospholipids restores activity. When BR is crystallized (see below) the bound lipids that are retained from the membrane fit well into the grooves along the protein surface (see Figure 8.9). BR was the first integral membrane protein whose topological organization in the membrane was elucidated. Electron diffraction of the native twodimensional crystalline alTays of purple membrane provided early images of BR trimers, revealing the monomer structure to be seven TM ex-helices arranged in an arc-like double crescent in the plane of the bilayer (Figure 5.4A and B). Models fitting the primary structure of the protein, with 70% of its 248 residues being hydrophobic, to the observed images were aided by the sensitivity of exposed loop residues to partial proteolysis in situ (carried out on BR in the membrane), although the precise beginning and end of each helix were uncertain for years. Such studies also showed that a few N-terminal amino acids are exposed to the exterior and the last 17 to 24 amino acids of the C terminus

c.

B

A

c

o

E

F

Gr I MOAO I

5.4. EM structure of BR. A. The electron density profile of the 2D-crystalline purple membrane shows arrays of BR trimers. Each trimer is arc-shaped with three well-resolved peaks in the inner layer and four less resolved peaks in the outer layer. By Unwin and Henderson. Redrawn from Garrett, R. H., and C. M. Grisham, Biochemistry, 2nd ed., Brooks/Cole, 1999, p. 273. C9 1999. Reprinted with permission from Brooks/Cole, a division of Thomson Learning: www.thomsonrights.com. Fax 800 730-2215. B. Seven helices are modeled to correspond to the seven peaks of a BR monomer. Based on neutron diffraction data, a retinal has been placed in the center of the protein. From Subramaniam, S., and R. Henderson, Biochim Biophys Acta. 2000, 1450:157-165. © 2000 by Elsevier. Reprinted with permission from Elsevier. C. A topology model of BR shows the predicted sequence composition of the seven helices and their connecting loops. The model was adjusted periodically based on genetic mutations of targeted residues (colored boxes) until the x-ray structure was solved. From Khorana, H. G., j Bioi Chem. 1988,263:7439-7442. © 1988 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

G

Helical Bundles

105

= 5 ms

T

BR ~4 ps A l11ax = 570 nm ~

o

K

Amax = 640 nm An/ax = 590 nm \ T

= 5 ms

7

(

N~Amax

=

T

560 nm

f-

1 "'

L

Ama.x = 550 nm

= 5 ms

T

= 40

J1.s

AlIlax = 410 nm

M z _ Mt T

= 350

J1.S

5.5. The photocycle of bacteriorhodopsin. In response to light,

BR undergoes a series of transitions through intermediates K, L, M1, M2, N, and 0, which have different lifetimes (T) and different absorbance maxima as shown. The photocycle is initiated by isomerization of the retinal from all trans to 13-cis (BR --+ K, L), followed by transfer of a proton (L --+ M), the conformational change that switches the accessibility of the Schiff base from the extracellular side to the cytoplasmic side (M1 --+ M2), another proton transfer (M --+ N), and conversion of the retinal back to all trans (N --+ 0). From Neutze, R., et aI., Biochim Biophys Acta. 2002, 1565:144-167. © 2002 by Elsevier. Reprinted with permission from Elsevier.

are accessible in the cytoplasm. The location of retinal in the center of the protein (Figure 5.4B) was determined by neutron diffraction, and the lysine to which it binds was identified by reduction of the Schiff base with NaBH 4 . Once it was clear that the retinal binds to Lys2 J6, nearby residues were investigated by sitedirected mutagenesis, which identified residues that interact with the retinal, such as Asp212 and Arg82, as well as residues crucial to proton pumping, such as Asp85 and Asp96. These results led to a model for the topology of BR that was further modified as genetic studies defined the positions of many residues (Figure SAC). Extensive mutagenesis of the gene for BR provided a large collection of mutant proteins that could be studied in cell suspensions or reconstituted in lipid vesicles, with changes of pH, temperature, and salt conditions used to further characterize the protein function. In addition, a variety of retinal analogs were incorporated to observe their effects on the absorption spectrum and activity. Researchers used a number of pH-sensitive dyes to investigate the stoichiometry of proton pumping. And over many years, increasingly sophisticated instrumentation for visible and ultraviolet absorbance, fluorescence, circular dichroism. Raman, and infTared spectroscopy have been employed to follow the response of BR to light. The primary event when BR absorbs a photon is the isomerization of retinal. This event triggers subse-

quent structural changes and pKa shifts in the protein that allow deprotonation of the Schiff base, vectorial transfer of the proton to the extracellular side of the membrane, and uptake of a proton from the cytosol. These processes are accompanied by differences in the absorbance spectru mol' BR, allowing detection of intermediates with lifetimes varying from a few picoseconds (ps, 10- 12 sec) to a few milliseconds (msec, 10- 3 sec). The light-induced changes in BR are summarized in a photoreaction cycle, or photocycJe (Figure 5.5). BR in its resting state has a )'ma, of 570 nm (purple); when it absorbs a photon, it rapidly isomerizes to the K intermediate ()'max 590 nm) and then converts to the L intermediate (),.max 550 nm). The transition from L to M ()'m", 410 nm) occurs when the proton fTom the Schiff base is transferTed to Asp85, the primary acceptor. At this point a structural rearrangement occurs to switch the accessibili ty of the Schi ff base, described as M I -> M2, which is essential for vectorial proton transport by preventing reprotonation from the extracellular side that

Cytoplasmic

A

Extracellular 5.6. The first high-resolution structure of BR. This overview of the structure shows the seven TM helices labeled A to G, and the residues involved in proton translocation as well as the retinal. From Pebay-Peyroula, E., et al., Biochim Biophys Acta. 2000, 1460: 119-132. © 2000 by Elsevier. Reprinted with permission from Elsevier.

Bundles and Barrels

106

B.

A.

Cytoplasmic side

) Helix G

Q7

ca

~

82

3.29

W408

0

3 • 11

_

2.36

3.25

.J

Extracellular side -../

5.7. Proton path in BR. A. The ribbon diagram for the seven TM helices, labeled A to G, is marked to show proton transfer steps indicated by arrows numbered in chronological order from 1 to 5. Step 1 is release of a proton from the Schiff base to Asp85. In step 2 a proton is released to the extracellular medium, possibly via Glu204 or Glu194. In step 3 the Schiff base is reprotonated by Asp96. Step 4 is the reprotonation of Asp96 from the cytoplasmic medium. Step 6 is the final proton transfer step from Asp85 to the group involved in proton release at the extracellular side, either Glu204 or Glu194. From Neutze, R., et al., Biochim Biophys Acta. 2002, 1565:144-167. © 2002 by Elsevier. Reprinted with permission from Elsevier. B. Details of the proton path on the extracellular side of the Schiff base reveal a network of hydrogen bonds between Asp85, Asp212, Arg82, Glu194, and Glu204 and discrete water molecules (red, labeled W). Interatomic distances are given in angstroms. From Pebay-Peyroula, E., el aI., Biochim Biophys Acta. 2000, 1460: 119-132. © 2000 by Elsevier. Reprinted with permission from Elsevier.

would result in a zero net effect. The next three steps are s!owel', each taking around 5 msec. Transfer of a proton to the Schiff base from Asp96 creates the N intermediate U.max 560 nm). With the return from 13-cis to alltrans retinal, the 0 intel-mediate o..max 640 nm) is formed and the release of a proton from Asp85 completes the cycle. The first high-resolution structure of BR was achieved by x-ray crystallography of microcrystals prepared in bicontinuous cubic phase lipids, either monoolein or monopalmitolein. (These are not phospholipids but rather racemic mixtures of glycerol esterified to one acyl chain, either oleoyl or palmitoleoyl, on Cl.) In cubic phase-grown crystals, BR trimers are stacked in layers that have the same ori-

entation and lipid content as that observed in purple membrane. Furthermore, spectroscopic studies show that BR in cubo undergoes the main steps of the photocycle. As expected from the electron density images, the seven TM hel ices cross the mem brane nearly perpend icular to the plane of the bilayer and are packed closely together and connected by short loops (Figure 5.6). The structure has now been refined to better than 2 A resolution and provides sufficient detail to trace the proton channel, including several important water molecules (Figure 5.7A). The central cavity that contains retinal in Schiff base linkage to Lys21 6 is quite rigid, with the n-bulge in helix G stabilized by an H-bond From Ala215 to a water molecule. In the resting state, the positive charge on the protonated Schiff base

Helical Bundles (pKa ~13.5) is stabilized by the nearby deprotonated carboxylate groups of Asp85 and Asp212. Polar side chains and 'vvater molecules make a clear proton path in the extracellular half of the molecule from Asp85, the proton acceptor, to the extracellular surface where the proton is released (Figure 5.7B). At the cytoplasmic surface are several acidic groups that may be involved in transferring protons from the cytoplasm, but no clear proton path connects the central cavity to them. The pKo of Asp96, the proton donor during reprotonation of the retinal from the cytoplasmic side, is very high due to its nonpolar environment and to its side chain H-bond to the side chain ofThr46. This part of the protein is more flexible than the extracellular half and must undergo a conformational change to open a proton path between the cytoplasmic surface and Asp96. To relate the elegant structure of BR in its resting state to the dynamic events of its photocycle, structural information has been obtained for different intermediates in the photocycle by crystallization of mutants prevented from completing the photocycle (such as D96N, which stops at the late M state) and by "kinetic crystallography" of wild-type crystals, which uses low temperatures and different wavelengths of light to Irap a significant population of the molecules in the crystals in one state. The structures show that the geometric and electrostatic effects of photoisomerization of retinal produce tensions in the protein molecule, which responds with small motions of residues and movements of discrete water molecules, as well as movements of helices G, F, and B (Figure 5.8). These detailed structures reveal a high-resolution "movie" that complements the spectroscopic data to present an exciting view of the dynamic mechanism of this light-driven proton pump, which is nature's simplest photosynthetic machine.

107

A.

Trp182

0/

.W407

B.

Photosynthetic Reaction Center

Nearly a decade before the first high-resolution x-ray structure for BR was published, the 1988 Nobel Prize in Chemistry was awarded to Hartmut Michel, Johann Deisenhofer, and Robert Huber for the elucidation of the x-ray structure of the photosynthetic reaction center (RC) from Rhodopseudomol1as viridis, Ihe first highresolution structure achieved for integral membrane proteins. When Michel and coworkers first crystallized the RC, the gene sequences encoding its protein constituents were not even available' The beautiful struclure of this multicomponent complex provided specific descriptors of its protein domains including TM helices, along with the locations of cofactors involved in light absorption and electron transfer. In photosynthesis light energy is converted into chemical energy when the absorption of a photon drives an electron transfer thai is otherwise

5.8. Examples of the structural shifts that occur during the photocycle in SR. Small differences are revealed when the high resolution structures of the active sites of the K and L intermediates are overlain on the structure of the ground state. A. The K intermediate, obtained for wild-type SR illuminated with green light at 11Oo K, shows disordering of a water (W402) and slight movements of Asp85 and Lys216. The circle indicates where another water molecule may appear. Positive and negative difference electron densities are shown in blue and yellow, respectively. B. Structural models for two different intermediates, K (blue) and L (red), are overlain with the ground state (green backbone with colored residues). The larger shifts in L include reorientation of the guanidinium group of Arg82, flexing of the backbone of helix C, and movement of the side chain of Trp182 toward the cytoplasm. From Neutze, R., et al., Biochim Biophys Acta. 2002, 1565: 144167. © 2002 by Elsevier Reprinted with permission from Elsevier.

Bundles and Barrels

108

5.9. The structure of the photosynthetic reaction center from Blastochloris viridis, a group II reaction center. The complex contains four protein subunits, L, M, H, and a cytochrome, and 14 cofactors (red). The TM helices are highlighted in yellow. Compare it with the group I reaction center shown in the chapter frontispiece. Redrawn from Nelson, D. L., and M. M. Cox, Lehninger Principles of Biochemistry, 4th ed., w. H. Freeman, 2005, p. 376. © 2005 by W. H. Freeman and Company. Used with permission.

thermodynamically unfavorable. The subsequent passage of electrons through spatially arranged carriers is coupled wi th the expulsion of protons, just as it is in oxidative phosphorylation, thus providing the proton gradient that drives the ATP synthase. Plants have two types of photosystems, called PSI and PSII, which di ffer in the electron acceptors used. The much simpler photosynthetic RCs of purple bacteria are considered the ancestors of PSII. Recent x-ray structures (at somewhat lower resolution) of both PSI and PSII reveal common structural features with the RCs in spite of their enormous size and complexity. RCs were discovered in photosynthetic purple bacteria and characterized by biophysical techniques such as EPR (see Box 4.2) and optical spectroscopy before they proved amenable to crystallization. Soon aher elucidation of the structure from R. viridis (renamed BlaslOchloris viridis), another high-resolution structure was obtained for the RC [yom Rhodopseudomonas sphaeroides (renamed Rhodobac/er sphaeroides). More recently, the x-ray structure for the RC from Ther-

l11ochromariul11 repidul11 was solved at 2.2 A resolution. While the cofactor-protein interactions are nearly the same in all three complexes, they represent two types of RCs. The Rb. sphaeroides RC is a member of group I and contains three protein subunits called L, M, and H, along with 10 cofactors (see Frontispiece). The other two RCs are members of group II, and they contain an additional subunit, a c-type cytochrome with its four heme cofactors (Figure 5.9). The three protein subunits, L (light), M (medium), and H (heavy), were named for their apparent molecular weights determ ined wi th SDS gel electrophoresis.

The Proteins The B. viridis RC has a size of ~ 130 A by ~ 70 A; its TM domain consists of five <x-helices from L, five C(helices from M, and one <x-helix from H. The remainder of the H subunit caps the structure on the cytoplasmic side, and the c-type cytochrome lies on the peripJasmic side (Figure 5.9). The TM helices composed of 21 to 28 amino acids cross nearly perpendicular to the plane of

Helical Bundles

109

L

H

M

cyt

weo

5.10. The peptide backbones of L, M, H, and cytochrome subunits of the photosynthetic reaction center from B. viridis. From Deisenhofer, J, et aI., Nature. 1985, 318:618-624. © 1985. Reprinted by permission of Macmillan Publishers Ltd.

the membrane. Three helices from each Land M subunit are nearly straight, while one is curved and one is bent more than 30 at a proline residue and ends with a 3 10 helix (a slightly narrower helix with three amino acids per turn). The structural similarity of Land M (Figure 5.10) gives a high degree of twofold symmetry in the TM domain in spite of only 26% sequence identity. The cytochrome with its two pairs of hemes also has twofold symmetry, unrelated to that ofL and M.lts com· pact structure consists of five segments: the N-terminal segment (residues Cl to C66), the first heme-binding segment (C67 to CI42), a connecting segment (C143 to C225), the second heme-binding segment (C226 to C315), and the C-terminal segment (C316to C336). As the first available detailed structure of integral membrane proteins, the RC confirmed expectations about distribution of amino acids on its surface, yet it revealed a new concept of interior polarity. The surface of the RC complex is polar in the peripheral subunits, with a net negative charge on the peri plasmic side and a net positive charge on the cytoplasmic side. The mem brane-spanning surface is very hydrophobic. There are no charged amino acids in this 30 A-wide band around the center of the RC, and very few water molecules associate with it. Like other membrane proteins, it has tyrosine and tryptophan residues distributed at the interfacial borders. Surprisingly, the polarity of the interior of the membrane-spanning G

domain is like that of the interior of soluble proteins, intermediate between the polarities of amino acids exposed to water and the hydrophobic interiOl- of the bilayer. The 10 TM helices of Land M together have 74 polar side chains. Furthermore, most of these polar residues do not appear to be involved in forming hydrogen bonds, as there are at most two hydrogen bonds between any pair of TM helices. Their predominant roles seem to be interactions with cofactors and protein subunits. Comparison of related RC sequences indicates that residues buried in the interior of the structure are conserved more than are residues on the surface. The M, L, and H subunits of the Rh. sphaeroides RC have 59%,49%, and 39% homology to subunits in B. viridis, respectively, and are very similar in structure. While some differences in amino acid sequence lead to differences in the interactions of the cofactors with the peptide chains, the complex has the same approximate twofold symmetry. The site-specific mutants first available in Rb. sphaeroides revealed the roles of GluL212 and SerL223 for reduction and protonation of the quinone and TyrM21 0 for efficient electron transfer. Lipids Based on its size and shape, the RC is predicted by EPR to have 30 to 35 annular lipids (see Chapter 4). However, most of the lipids are replaced by detergent during purification. For crystallization, the RC is solubilized with LDAO (N,N-c1imethyl-dodecylamine-N-oxide), and a few detergent molecules are included in the crystal structure. While it is often difficult to ascertain whether an acyl chain detected in the structure is from a lipid or a detergent, the x-ray structures give evidence for specific lipids (see Chapter 8), including a cardiolipin and a PE, which fit closely into hydrophobic grooves at the surface of the protein exposed to the nonpolar membrane domain.

The Cofactors The proteins provide a scaffold for the cofactors, holding them in the same spatial arrangement in all three RCs crystallized. The RC core has 10 cofactors: Four bacteriochlorophyJls (BChl), which resemble heme except for the replacement of iron by Mg2+ ions, a cyclopentenone ring fused to one pyrrole ring, and different substituents off two of the pyrrole lings Two bacteriopheophytins (BPh), which are BChl with two protons in place of the Mg 2+ Two qui nones (one ubiquinone and one menaquinone in B. viridis) A nonheme ferrous ion A carotenoid, which is a largely linear C40 polyene such as l3-carotene

110

There are two types of both BChl and BPh, a and b. The a type is found in the RC from Rh. sphaeroides and has either a phytyl or geranylgeranyl side chain, whereas the b type, found in the RC from B. viridis and T tepidum, has only a phytyl side chain and has one more C=C in the side chain of ring II. The cofactors form two symmetrically related branches within the hydrophobic environment of the closely packed TM helices (Figure 5.11). At the top, two molecules of BChl are positioned so close together that the edges of their tetrapyrrole rings overlap. Called the special pair, they receive the photon of light and release an electron in the primary event of photosynthesis. Each branch has another BChl (called the accessory BChl), a BPh, and a quinone. The nonheme iron is between the two quinones. The two branches follow the same local symmetry displayed by the Land M chains. The symmetry is not perfect, and the two branches have different electron transfer properties - in fact, electron transfer uses only the branch that associates with the L subunit. The cofactors are differentiated by groups of the protein that alter their environments. For example, the two quinones play different roles (see below) and the primary quinone is in a more hydrophobic environment than the secondary quinone. After being reduced and protonated, the secondary quinone (now quinol) diffuses from the RC, its leaving facilitated by nearby carboxylate groups. The similarities in the arrangement of cofactors in all photosystems, including the ironsulfur type PSI complexes, allow definition of a common motif: a dimer of tetrapYITole molecules nanked

5.11. The arrangement of the RC cofactors. The tetrapyrrole rings of BChl-b, BPh-b, and the quinone headgroups follow the same local symmetry displayed by the Land M chains. The figure shows the special pair, PA and PB (coral); the accessory BChl molecules, BA and BB (rose); the two BPh, HA and HB (cyan); the ubiquinones, OA and OB (yellow); carotenoid, Crt (purple); and non-heme iron atom (gray). The electron path utilizes only the A half, indicated by the arrows. From Jones, M. R., et al., Biochim Biophys Acta. 2002, 1565:206-214. (-< f-
ro

~

~

2

o 5.18. The structure of OmpX, a small, closed l3-barrel protein, Because OmpX is monomeric, all of its exposed side chains go into the bilayer. Additionally, four of its eight l3-strands protrude on the outside of the cell. The barrel interior is filled with polar residues that form a hydrogen-bonding network. A Ribbon drawing shows nonpolar side chains (yellow) on the surface of OmpX. From Schulz, G. E., Curr Opin Struct Bioi. 2000, 10:443-447. © 2000 by Elsevier. Reprinted with permission from Elsevier. B, Topology model shows the primary structure and distribution of amino acids in OmpX, with lipidexposed (red), interior (black), and external loops (green). The y-axis gives the transbilayer location with the center of the bilayer as 0, To show the interstrand hydrogen bonds between each pair of strands, the eighth strand is repeated on the left (gray). C. Bar graphs show the distribution of amino acid residues on the external surface (left) and in the interior (right). Band C from Wimley, W. C, Curr Opin Struct Bioi. 2003, 13:404-411. © 2003 by Elsevier. Reprinted with permission from Elsevier.

is thought to be a part of the virulence mechanism of E. coli. There is much variation in quaternary structure among the known (3-barrels. Several of them are monomers, while all the porins are homo-oligomers (usuaJJy trimers) with very tight interactions between the subunits. The enzyme activity of outer membrane

phospholipase A (OMPLA; see Chapter 9) is regulated by its quaternary structure: it is only active as a homodimer. The (3-barrel formed by ct-hemolysin (see Chapter 4) results from a completely different motif, in that each subunit of the heptamer contributes a (3hairpin to make the 14-stranded barrel that inserts into target membranes. The rich diversity of structure and

116

Bundles and Barrels

BOX 5.1. NMR determination of membrane protein structure Structure determination by nuclear magnetic resonance (NMR) spectroscopy is limited to fairly small proteins and is even more difficult for membrane proteins in detergent micelles with their increased particle sizes. Complete structures have been solved for only a few small membrane proteins (~20 kDa or less) in detergent micelles. While there has long been hope for application of solid-state NMR to solve the structures of membrane proteins, the structures obtained to date are from solution NMR. These successes were made possible by the development of TROSY (transverse relaxation optimized spectroscopy) by Kurt Wuthrich, for which he was awarded the Nobel Prize in Chemistry in 2002. NMR

has the potential to measure dynamics within the protein structure, as well as to give information about protein-lipid interactions.• Structures of f)-barrels are easier to solve by NMR than those of helical bundles because (1) the 1 H chemical shift

A detailed explanation of these NMR methods is beyond the scope of this book. See Fernandez, C., and G. Wider, TROSY in NMR studies of the structure and function of large biological macromolecules, Curr Opin Struct BioI. 2003, 13:57Q.-580. For a general introduction, read MagnetiC Resonance in Chemistry and Medicine by Ray Freeman, published by Oxford University Press, 2003.

5,1,1. OmpX structure solved by solution NMR. A. NMR spectra of OmpX in DHPC micelles obtained using 'H lsN COSY (a) and 'H 1sN TROSY (b). B. Structure of OmpX determined by NMR. When the NMR assignments are mapped onto the x-ray structure of OmpX (a), the barrel portion is well-structured (red) and three loops are disordered (yellow). The flexibility of the loops is clear in the NMR structure represented by the superposition of 20 conformers (b). From Fernandez c., et aI., FEBS Lett. 2001, 504: 173-178. © 2001 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

I)-Barrels

117

BOX 5.1. (continued)

G1448 G42,' ~G'126 • IJ .. • G36.4i. 1'48 • E 134

A.

tGJJ2

GI651 T95

110.0

..

,..,

GS4'~ ~ GI710

VI27"

• • TSO

•

",.

"

0

•

#.

."

IS-

Q

AJt~'

GIO

T137"

..."...,

O'~:4°~

115.0

.f

fIIC'

~ A'

KI2

DS9'!":

iii '. N~6 NI.596t. . • FI5 W. ''1''. T'pS'•• t : .. . '. 51030' F40 ·:O;'j.e.

11

E0. 120,0

.4' ,..~ ,..".

I

\'166'\~~~~'~~~, ..·}~·:tt·~" l

-8'

.v.~)

~, ~

lr

3

'\

'1'168" .

Z

"

RI3~~]i aO• L:

V"l

M53" •

125.0

~

•

IV'

, . K82 •

"

.i~'f·"'.

:'l... • : AI7/If ~N69 ~79

RI69'O~'

130.0

,

J.

60 157

....

\'45 044

\,129.

,•

LS3

G41'CI

10.00

"L91

1131NIl4

AI36. • ,

A"

· .

,fl A176

• 00 &"

16 LI62

'

T6

o

.~?GI2S

AI3C\:ltGI73 • 0 ;,.1';...

9.00 8.00 'H w2 (ppm)

7.00

B.

L2

(a) 5.1.2. OmpA structure solved by NMR. A. TROSY-based 1 H_ 15 N spectrum of the TM domain of OmpA in DPC micelles. B. NMR structure of OmpA. The NMR solution structure of OmpA is represented by the superposition of the ten confomers having lowest energy (a). Again the l3-barrel (red) is well-defined. while the loops are disordered. Four flexible loops are resolved in the ribbon diagram based on the NMR results (b). C. Comparison of the NMR (red) and x-ray (blue) structures for OmpA show good agreement on the l3-strands and considerable differences in the loops. From Arora, A., et aI., Nat Struct Bioi. 2001, 8:334-338. © 2001. Reprinted by permission of Macmillan Publishers Ltd.

(continued)

Bundles and Barrels

118

BOX 5.1. (continuec/)

c.

~C(X77

Outer membrane

(Figure 5.1.1 A) shows the enhancement obtained with TROSY. which allowed the eight-stranded fold of the polypeptide backbone of OmpX to be determined from 107 nuclear Overhauser effect (NOE)-derived distance constraints and 140 dihedral angle constraints. The structure was further refined with selective protonation of the methyl groups of Val. Leu, and lie on a perdeuterated background. which enabled assignment of 526 NOE distance constraints. When 20 NMR conformers are superimposed. the result clearly reveals the f)-barrel with its flexible loops (part b of Figure 5.1.1 B). The ribbon diagram in part a of the figure maps the NMR results on the x-ray structure and distinguishes the well-structured regions (red) from the disordered loops (yellow). The NMR result is strikingly similar to the structure from x-ray crystallography and in particular confirmed the extension of f)-strands on the exterior of the protein.

OmpA TM Domain from E. coli

Periplasmic end

C N

5.1.2. (continued)

dispersion is larger for f)-sheets than for lX-helices. especially with alternating hydrophilic/hydrophobic residues. and (2) f)-barrels have higher thermal stability. so they can withstand higher temperatures for the longer times needed to get well-resolved NMR spectra. Furthermore. the two f)-barrel proteins described here could be overexpressed in E. coli and purified in denatured form from inclusion bodies before refolding in detergent micelles (see Chapter 7). resulting in a mixed micelle particle size of 60 to 80 kOa. Uniform labeling of the proteins with 2H. 13C. and 15N allows detection of the protein NMR signals with little or no interference from the signals of the unlabeled detergent molecules. OmpX from E. coli Triply labeled OmpX. a 148-residue protein. was solubilized from inclusion bodies with guanidinium chloride and reconstituted in dihexanoyl-PC (OHPC) micelles. Comparison of the 20 COSY (Correlation Spectroscopy) and TROSY spectra

function among 13-barrels continues to be uncovered, yet the paradigm for this group of membrane proteins is the porins.

Porins The outer membranes of Gram-negative bacteria protect the cells from harmful agents while allowing nutrient uptake via porins and other transport proteins. Porins are pore-forming TM proteins that function as passive diffusion channels and thus allow rapid diffusion of their solutes. even at O°c' Porins are grouped in two categories: general porins. like OmpF and OmpC, and specific pOl-ins, like PhoE (phosphoporin), LamB (maltoporin). ScrY (sucrose porin). and Tsx (the nude-

The f)-barrel domain of OmpA (residues 172-325) was purified after denaturation in urea and refolding into dodecylphosphocholine (OPC) micelles. The protein was labeled with 15N, 13C. and 2H, and TROSY experiments were carried out at 600 and 750 MHz. The protein in a large excess of OPC gave the best NMR spectra, shown in Figure 5.1.2A with several of the assigned resonances labeled. TROSY experiments were carried out with several specific amino acid-labeled samples to aid in assignments. The backbone fold of the OmpA TM domain was initially calculated from 91 NOE distance constraints and 142 torsional angle constraints. It was refined by introducing 116 H-bond constraints between adjacent f)strands that were identified in the initial fold calculations. The structure of the eight-stranded f)-barrel is well defined. Figure 5.1.2B shows 10 superimposed conformers (a) and the corresponding ribbon diagram of the solution structure (b) of OmpA. Like the OmpX structure. the NMR structure of OmpA shows much agreement with its x-ray structure, illustrated in Figure 5.1.2C, where the solution structure determined by NMR (red) is overlain with the x-ray crystal structure (blue). As NMR gives information about protein dynamics. some of the poorly defined loops are thought to have intrinsic high mobility. A study of the dynamics of the backbone from 15N relaxation times indicates the H-bonded core is not completely rigid but moves on the microsecond-millisecond time scale.

oside channel). The channels of general par'ins do not discriminate among solutes thal are hydrophilic. under ~600 Da. and not highly charged. which allows them to take up many nutrients such as mono- and disaccharides. In contrast. the specific porins have channels that are selective for their solutes. although the line of demarcation is blurred as described below for PhoE. Note that aquaporin, which is a TM channel for water molecules in plasma membranes (see Chapter 10). is not a 13-barrel. Porins were first detected by the permeability of the outer membrane of Gram-negative bacteria to hydrophilic antibiotics. Thus the first assay was based on hydrolysis of 13-lactams (penicillin and its derivatives) in intact cells. since their rate of permeation

~-Barrels

through the outer membrane determines their availability for hydrolysis by the enzyme ~-Iactamase in the periplasm. Channel activity of pOl'ins is evident when the purified proteins are reconstituted with lipids: porins make lipid vesicles permeable to small, hydrophilic solutes and make voltage-gated conductance channels in black films. OmpF and Om pC Typically Gram-negative bacteria have ~lOs copies of general porins per cell, with the number of species of porins varying in different strains. The outer membrane of E. coli K12 is dominated by OmpF and OmpC pOl'ins, both homotrimers of around lIS kDa whose levels are controlled by osmolarity of the growth medium. This regulation is carried out by a two-component system made up of EnvZ, the protein sensor of osmolarity, and OmpR, the transcriptional activator for their two genes. OmpC has been called osmoporin, because its expression is induced by high osmolarity as well as high pH and high temperature. On the other hand, higher levels of OmpF are expressed in medium of low osmolarity. OmpF has a larger channel diameter than does OmpC, and although the size difference is only ~IO%, the flux of larger solutes through the OmpF channel is significantly faster than that through OmpC. This difference allows the bacteria to respond to two very different habitats: in mammalian hosts, high osmolal-ity and high temperature induce OmpC, whose narrower channel keeps out some of the body's inhibitory substances, \-vhereas outside the host the wider channel of OmpF speeds the uptake of nutrients h'om very dilute environments, such as ponds and rivers. Purification of porins is facilitated by their strong, noncovalent association with peptidoglycan, as this complex can be isolated by extraction of the cell envelope with SDS at 60°C: while most of the membrane proteins are solubilized, the porin remains in homogeneous two-dimensional crystalline aggregates. These aggregates have porins in hexagonal arrays (with patches of phospholipids between porin trimers) and give typical conductance properties when incorporated into planar bilayers. To remove the peptidoglycan, further purification involves salt treatment before gel filtration or ion exchange chromatography. POI-ins can also be solubilized [Tom cell envelope by extraction with non ionic detergents such as octyl-POE (see Chapter 3). Detergent-solubilized porin binds 0.6 g detergent per gram of protein, which amounts to around 200 detergent monomers per protein trimer. The protein is remarkably stable: it is resistant to proteases, chaotropic agents and most organic solvents, in addition to SDS and other detergents. It is even functional after lyophilization and storage at -20°C.

119

Crystals of OmpF from E. coli were described as early as 1980 but due to their unusually symmetric packing, the phase problem could not be solved. lL was another decade before a crystal structure of a porin \,vas solved, that of the analogous porin from Rb. capsulaIus. The high-resolution structure of OmpF, as well as that of PhoE, was then determined. The highly homologous OmpC resists crystallization but is expected to share many of the structural features of Rb. capsulalus porin and OmpF. Even a porin with weak homology can have very similar architecture, as demonstrated in the x-ray structure of the porin h'om Rhodopseudomonas blaslica. These porins are homotrimers, in which each subunit forms a ~-barrel (Figure 5.19). The barrel consists of 16 antiparallel ~-strands tilted by 4SO, with a salt bridge between the amino and carboxyl termini to complete the cyclic structure. At the extracellular end one loop latches onto the next subunit, and one or more of the loops fold back. in to the channel. Loop 3 in OmpF has the highly conserved sequence motif PEFGG and folds into the channel to constrict the pore at the middle of the barrel to IS x 22 'A. This eyelet of the pore has clusters of acidic residues on one side and basic residues on the other, which create a local transverse electric field that accounts for the slight cation specificity observed in conductance studies (Figure 5.20A). As discussed in Chapter 3, conductance measurements can detect Single-channel openings, as well as channel closures at voltages above 100 mV (see Figure 3.16), although the mechanism of channel closing and its significance under physiological conditions are unknown. Characterization of structure and function of porins with mutations affecting the constriction site of the channel has enabled properties such as single-channel conductance and transport rates to be correlated with precise changes in the crystal structures. None of the mutations altered the barrel framework of the protein. Single amino acid substitutions for each of the charged residues of the eyelet alter the ion selectivity but produce little change in pore size and conductance (Table 5.1). If, however, both acidic residues are mutated (DI13N/E 1170), the conductance drops by 50% and the cation selectivity is removed. Simulations of ion flow by Brownian dynamics illustrate the role of the acidic residues in drawing cations through the constriction site. The change in pore size does not always COlTelate with changes in conductance but is well reflected in uptake rates for disaccharides measured by the liposome-swelling assay, as seen in the effects of deletion of six residues of Loop 3 (6109-114) and substitution of neutral amino acids for five charged residues (R42A1R82A/RI32A1DI13N/EI170). Clearly both pore dimensions and the constellation of charged residues

A.

Loop3

D.

323

,ft,,n, '1[1

,i[1

i ~''.j1 K

S

"

'ij' , I

I I

V -,-,' l' . ~v

,; HY.j.~e

I

Lt,- t'e t,' t,'

r,-

~l,~e,~'e~ ,

Gy.jei~t;

y>vc'1&r.~& t. t,' 9N

340

1

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5.19. Structure of OmpF porin. A. Ribbon diagram of the OmpF monomer showing the long loops facing the cell exterior and the l3 loop that folds inside the barrel, as well as the proximity of the Nand C-terminal residues. B. View of the OmpF trimer from the top (outside) showing the constriction of each pore by the l3 loop. C. Slice through the center of the OmpF trimer using a stick model enhanced with the molecular surfaces, showing the dimensions of the pores. A through C from Schirmer, T., j Struct Bioi. 1998, 121: 101-1 09. © 1998 by Elsevier Reprinted with permission from Elsevier. D. The topology of OmpF as an unrolled barrel with the last two l3-strands repeated to emphaSize the cyclic nature of the structure. Residues in l3-strands are in diamonds, residues in !X-helices are in rectangles and residues in turns and loops are in circles. From Cowan, S. W., et al., Nature. 1992,358:727-733. © 1992. Reprinted by permission of Macmillan Publishers Ltd.

at its narrowest site determine the transport properties of the general pm-ins. Specific Porins

Sharing many characteristics of general pOI-ins, the specific porins add to them the ability to discriminate

among solutes. Since they carry out passive diffusion, they cannot rely on energized conformational changes to release substrates from high-affinity binding sites (see below and Chapter 10). Rather, the specific porins have low affinities for their substrates and have achieved selectivity through features of their channel architecture. Their specificity is apparent only with

~-Barrels

121

B.

A.

5.20. Key amino acid residues in the constriction zones of porins. A. Viewed in cross-section, the channel of the OmpF porin monomer is constricted by Loop 3, forming an eyelet lined by clusters of basic residues (K16, R42, R82, and R132) and acidic residues (D113 and E117). B. The channel constriction in PhoE, also formed by Loop 3, has two additional basic residues, here numbered K18 and K131. Note the residue numbers in PhoE have been adjusted to match those in OmpF. From Karshikoff, A., et aI., j Mol BioI. 1994, 240:372-384. © 1994 by Elsevier. Reprinted with permission from Elsevier.

TABLE 5.1. Channel properties of OmpF, PhoE, and OmpF mutants

Protein OmpF PhoE D111G E117Qe D113N/E117Q(NQ) D113N/E117Q/R42A/R82A/R132A (NQAAA) R42C R82C R132P V18K1G131K(KK)

Mimimal cross-section a (%)

Conductance b (nS)

SelectivityC PNa/PC!

Disaccharide permeation d Relative swelling rates

100 70 129 100 94 177

0.S4 ±0.06 0.63±0.06 0.SO±0.06 0.64±0.02 0.40±0.02 0.64 ±0.04

4.5 ± O.S 0.44 ±0.05 1.4±0.1 2.9 ±0.2 1.0±003 12.3 ±0.9

5 2 2 2 12±1.5 7±3 NT 37 ± 1.5

127 114 109 75

07S±0.07 0.76±0.06 o 77±0.04 0.75±0.02

9.7 ±2.0 2.1 05 nO±1.9 2.1 ±O.OS

NT NT 1O±3 100 single-conductance step measurements. nS, nanosiemen. C PNa/P CI (the relative rates of permeation by Na+ and (1-) was determined from at least four different preparations. d Disaccharide uptake is measured by the liposome-swelling assay in MLVs and given as the mean swelling rates for sucrose, lactose, melibiose. and maltose. NT, not tested. e The liposome-swelling assay utilized E117C instead of E117Q. Source: Phale, P. S., et aI., Biochemistry. 2001, 40:6319-6325. © 2001 by American Chemical Society. Reprinted with permission from American Chemical Society.

Bundles and Barrels

122

Cell exterior

-PeripJasm 5.21. X-ray structure of a monomer of LamB, the maltoporin. Ribbon diagram of the LamB monomer viewed from the membrane-exposed surface, with the external chains given in ball representation. (Oxygen atoms are in black.) The arrows point to the aromatic girdles at the boundaries between the nonpolar and interfacial regions of the bilayer, and the vertical bar (~25 A) denotes the nonpolar region. The charged side chains are mainly in the outer region of the barrel, where they could interact with lipopolysaccharide. From Schirmer, T., J Struct Bioi. 1998, 121: 101-1 09. © 1998 by Elsevier. Reprinted with permission from Elsevier.

larger molecules because small solutes like arsenate and glucose easily permeate the nonspecific channel interiors. Clues to their specific functions come from their regulation: for example, in E. coli, PhoE protein is induced under phosphate starvation and LamB protein is induced by growth on the carbon source maltose. PhoE, the Phosphoporin

In spite of the very high homology between PhoE, OmpF, and OmpC, PhoE has a specific transport function. When their functions are compared in whole cells, mutants constructed to have only PhoE take up phosphate and phosphorylated compounds much more efficiently (,vith a ninefold decrease in K m of tl-ansport) than mutants with only OmpF or OmpC. Conductance studies with purified PhoE protein demonstrate a strong anion selectivity. Furthermore, polyphosphates such as ATP inhibit the flux of small ions through reconstituted PhoE channels. Like OmpF protein, the PhoE protein is a 16stranded j3-barrel; in fact, the barrel structures of the two proteins are superimposable, and the differences in their folds are confined to the loop regions and a single short turn. The constriction zone of the PhoE pore has two additional basic groups (Figure 5.20B),

and the calculated electrostatic potential of the pore is more strongly positive than that for the pore of OmpF. A genetic approach was taken to identify critical residues of PhoE that might form a phosphatebinding site, targeting basic residues that replace neutral or acidic amino acids in OmpC or OmpF. Indeed the single mutation K125E changes the ion selectivity of the PhoE pore. In the crystal structure of PhoE, this lysine residue is in Loop 3 at the constriction site (as KI25 in PhoE corresponds to amino acid 131 of OmpF; it is labeled Lys131 in Figure 5.20B). Another unique lysine is at the mouth of the pore. Howevel~ introduction of lysine residues at these locations in OmpF (OmpF mutant V18KJG131K) did not completely convert it to a PhoE-like pOl-e (Table 5.1). Other residues in PhoE must contribute to its selectivity; for example, Serl15 contributes by changing the position of the backbone in Loop 3 to make room for the side chain of Lys 131. Electrostatic contributions from other residues are probably important, as in the pore in OmpF. Given the lack of a specific binding site, PhoE might be viewed as a general porin. [ts selectivity is achieved with nonsaturable binding sites of low affinity. Howevel~ it clearly facilitates the transport of phosphorylated compounds into cells, and its blockage by polyphosphates is velY similar to the inhibition of the LamB pore by maltodextrins. LamB, the Maltoporin

LamB protein is named for its role as the receptor for phage A.It is required for growth of E. coli in chemos tats with limiting maltose as the sole carbon source, which provided early evidence for its role in maltose transport. The specificity of the LamB channel for maltose and maltodextrins can be detected by comparing rates of sugar uptake in the liposome-swelling assay (see Figure 3.26). The reconstituted LamB pore shows little discrimination among monosacchal-ides, but among disaccharides uptake of maltose is > 10 times faster than that of lactose and 40 times faster than that of sucrose. Other than maltodextrins, sugars larger than d isaccharides do not permeate the channel. The affinity for maltodextrins can be quantitated by their inhibition of glucose uptake as well as by binding to immobilized starch; both show a weak affinity (K,s in the low mM range). Maltodextrins also block conductance through the LamB channel when reconstituted in black films. Overall, the high-resolution structure of LamB protein shows similar architecture to the other porins: it is a homotrimer in which each monomer has 18 strands in the f)-barrel (Frontispiece and Figure 5.21). Three of the loops fold in to constrict the channel to a minimum diameter of 5 A. Six aromatic residues line one side of the channel, forming the "greasy slide," a smooth hydrophobic path through the otherwise aqueous pore

0-Barrels

123

(Figure 5.22). Soaking the maltoporin crystals in maltodextrin and different disaccharide substrates showed the substrates threading lengthwise through the channel with the hydrophobic sides of their pyranose rings along the greasy slide. The specificity for the maltose configuration is determined by numerous hydrogen bonds to charged side chains on the other side of the channel. When sucrose diffuses into the crystals, it gets stuck above the channel constriction, which explains its very low rate of uptake into liposomes. Interestingly, a sucrose-specific porin, called SrcY, is homologous to LamB protein, with a few strategic differences in the residues revealed in its high-resolution structure. Thus, like PhoE, the selectivity of these pores for their sugar substrates is achieved by the specificity built into their channels.

Iron Receptors 5.22.

Side view of the LamB monomer showing the "greasy slide". Maltodextrins pass through the pore down a slide made up of six aromatic residues: Trp74 from the adjacent monomer, Tyr41 , Tyr6, Trp420, Trp358 and Phe227 (in purple, with oxygen atoms red and nitrogen atoms blue). Tyrl18 (green) constricts the channel from the other side. From Koebnik, R., et ai, Mol Microbiol. 2000, 37:239-253 © 2000. Reprinted with permission from Blackwell Publishing.

Receptors involved in iron transport are 0-barrel proteins with an entirely different transport mechanism. Iron is abundant but unavailable in the environment due to its insolubility as ferric hydroxides. To solubilize this essential mineral, microorganisms synthesize and secrete siderophores, iron cheJators of 500 to 1500 Da with extremely high affinities for ferric ions. The

5.23. Structures of FepA and FhuA proteins involved in iron transport. A. Side views with the extracellular surface on top and the periplasmic end on the bottom. B. Views from the external solvent show the blockage of the interior. The FhuA l3-barrel is blue and plug domain is yellow; the FepA l3-barrel is green and plug domain is orange. From Ferguson, A. D., and J. Deisenhofer, Biochim Biophys Acta. 2002, 1565:318-381. © 2002 by Elsevier. Reprinted with permission from Elsevier.

124

enteric bacteria, including E. coli, synthesize and transport an iron chelator called enterobactin and also have transport systems for siderophores secreted by other microorganisms, such as ferrichrome. These transport systems consist of outer membrane receptors, periplasmic binding proteins, and inner membrane transport proteins. Unlike the porins, the outer membrane iron receptors bind their substrates with high affinities (K! ~O.l I-lM) and pump them into the periplasm at the expense of energy. The energy is provided via a complex of three proteins, TonB, ExbB, and ExbD, anchored in the inner membrane. In E. coli the iron receptors, along with another TonB-dependent protein, BtuB, the receptor for vitamin B 12 , interact with TonB at a conserved sequence of five amino acids near the N terminus called the ''TonB box" (TXXV[S/T], where X is a hydrophobic residue; see Chapter 11). While these interactions are well characterized, the mechanism of energy delivery is unknown. ExbB and ExbD are candidates for a proton translocation apparatus that couples chemiosmotic potential with conformational changes in TonB. In this scenal-io, the energized TonB then binds FhuA or FepA to open a high-conductance channel either by moving the plug domain into the peri plasm as postulated for the BtuB protein (see Figure 11.41) or by conformational changes more like those of other transporters, such as the sugar transpOl-ters LacY and GlpT described in Chapter 10. High-resolution structures show many similarities between the E. coli receptors for enterobactin and for ferrichrome, the FepA protein and the FhuA protein, respectively. They each have t\,vo domains, a C-terminal domain that makes a 22-stranded f)-barrel, and a globular N-terminal domain of ~ 150 residues that fills the interior; making a plug (Figure 5.23). The barrel has a diameter of ~40 Aand extends beyond the bilayeron the outside. Some of the 11 loops on the external membrane surface are unusually long: they range from seven to 37 l-esidues in FepA. The plug domain has a four-stranded I)-sheet and interspersed LX-helices and loops, including two loops that extend 20 A beyond the outer membrane interface to frame a pocket with the binding site for the siderophores. The largest difference between the two \-eceptors is the nature of this site, which is tailored for siderophores that differ in composition and charge. Comparison of the x-ray structures of FhuA in the presence and absence of substrates indicates that a conformational change takes place in the N-terminal domain upon ligand binding. Small movements of the loops in the binding pocket trigger unwinding of an LXhelix and a large movement of the polypeptide chain to the opposite side of the barrel wall (Figure 5.24). These changes do not open a transport channel, but

Bundles and Barrels

5.24. Conformational change in FhuA when it binds its siderophore. The (3-barrels of unliganded and liganded FhuA are shown as wire frame models. The plug domain of unliganded FhuA is blue and that of liganded FhuA is red. From Ferguson, A. D., and J Deisenhofer, Biochim BiophysActa. 2002, 1565:318332. © 2002 by Elsevier. Reprinted with permission from Elsevier.

could enable the binding of TonB to do so (see Chapter 11). The structural information from studies of FepA and FhuA sharpens the understanding of iron transport in E. coli and also provides models for its investigation in other microorganisms. Certainly iron receptors in pathogenic bacteria are imponant from a medical standpoint. Acquisition of iron is critical for invading microbes, which extract iron from host proteins such as transferrin and hemoglobin. In addition, the iron receptors are used by an ti biotics as well as col ici ns to enter the cell. Finally, siderophore-drug conjugates show potential for targeted drug delivery. From the porins to the iron receptors, tremendous progress has produced elegant structures of I)-barrel membrane proteins along with enhanced understanding of their transport mechanisms. Even so, it is likely they represent a small h-action of the proteins in this class of membrane proteins. If the genomic analyses are correct, there are many more I)-barrel proteins to be characterized. The next chapter looks at bioinformatics techniques used to predict and analyze membrane proteins and shows how they al-e grouped in families.

For Further Reading FOR FURTHER READING

Bacteriorhodopsin Reviews Haupts, U., J. Tittor, and D. Oesterhelt, Closing in on bacteriorhodopsin. Ann Rev Biophys BioI/wi Strltcl. 1999,28:367399. Lanyi, J. K., and H. Luecke, Bacteriorhodopsin. CUlT Opil1 Strltct Bioi. 2001,11:415-419. Lanyi, J. K., X-ray diffraction of bacteriorhodopsin photocyde intermediates. l\!lol NJembr Bioi. 2004, 21:143-150. Neutze, R., E. Pebay-Peyroula, K. Edman, A. Royant, J. Navarro, and E. M. Landau, Bacteriorhodopsin: a highresolution structural view of vectorial proton transport. Biochim Biophys Acta. 2002,1565:144-167. Seminal Papers Henderson, R., and P. N. T Unwin, Three-dimensional model of purple membrane obtained by electron microscopy. Natllre. 1975,257:28-32. Oesterhelt, D., and W. Stoeckenius, Functions of a new photoreceptor membrane. Proc Natl Acad Sci USA. 1973, 70:2853-2857. Pebay-Peyroula, E., G. Rummel, J. P. Rosenbusch, and E. M. Landau, X-ray structure of bacteriorhodopsin at 2.5 angstroms h'om microcrystals grown in lipidic cubic phases. Science. 1997,277:1676-1681. Winget, G. D., N. Kanner, and E. Racker, Formation of ATP bv the adenosine triphosphatase complex from spinach chloroplasts reconstituted together with bacteriorhodopsin. Biochiln Biophys Acta. 1977,460:490-499. Photosynthetic Reaction Centers Reviews Deisenhofer, J., and H. Michel, Structures of bacterial photosynthetic reaction centers. Al/lnt Rev Cell Bioi. 1991, 7: 1-23. Nogi, T, and K. Mild, Structural basis of bacterial photosynthetic reaction centers. ] Biochem (Tokyo). 200 I, 130:319329. Vermeglio, A., and P. Joliot, The photosynthetic apparatus of Rhodobacter sphaeroides. Trends NJicrobiol. 1999,7:435440. Seminal Papers Deisenhofer, J., O. Epp, 1. Sinning, and H. Michel, Crystallographic refinement at 2.3 A. resolution and refined model of the photosynthetic reaction centre from RhodopseudOlnonas viridis. ] Iv101 Bioi. 1995,246:429-457. Deisenhofer, J., O. Epp, K. Miki, R. Huber, and H. Michel, Structure of the protein subunits in the photosynthetic reaction centre of Rhodopseudol1lcmas viridis at 3A. resolution. Nature. 1985,318:618-624. I3-Barrels Buchanan, S. K., [3-Barrel proteins from bacterial outer membranes: stnJCture, function and refolding. ClilT Opin Struct Bioi. 1999,9:455-461.

125

Schulz, G. E., [3-Barrel membrane proteins. Curl' Opin Struct Bioi. 2000, 10:443-447. Wimley, W. c., The versatile [3-barrel membrane protein. Curr Opin Struct Bioi. 2003, 13:404-411. Porins Achouak, W, T Heulin, and J.-M. Pages, Multiple facets of bacterial porins, FEMS Microbiol Lett. 200 I, 199: 1-7. Delcour, A. H., Solute uptake through general porins. Frontiers Biosci. 2003, 8:1055-1071. Dutzler, R., Y-F. Wang, P. J. Rizkallah, J. P. Rosenbusch, and T Schirmer, Crystal structures of various maltooligosaccharides bound to maltoporin reveal a specific sugar translocation pathway. Structure. 1996, 4: 127-134. Nikaido, H. Porins and specific channels of bacterial outer membranes. Mol Microbiol. 1992,6:435-442. Schirmer, T, General and specific porins h'om bacterial outer membranes. ] Struct Bioi. 1998, 12\: I 0 I-I 09. Iron Receptors Cao, Z., and P. E. Klebba, Mechanisms of colicin binding and transport through outer membrane proteins. Biochimie. 2002, 84:399-412. Clarke, T E., L. W Tari, and H. J. Vogel, Structural biology of bacterial iron uptake systems. CUlT Top Med Chem. 2001, 1:7-30. Ferguson, A. D., and J. Deisenhofer, TonB-dependent receptors - structural perspectives. Biochim Biophys Acta. 2002, 1565:318-332. Locher, K. P., B. Rees, R. Koepnik, A. Mitschler, L. Moulinier, J. Rosenbusch, and D. Moras, Transmembrane signaling across the ligand-gated FhuA receptor: crystal structures of fTee and felTichrome-bound states reveal allosteric changes. Cell. 1998,95:771-776. First Crystal Structures of Proteins Discussed in Chapter 5 Buchanan, S. K., B. S. Smith, L. Venkatramani, D. Xia, L. Esser, M. Palnitkar, R. Chakraborty, D. van del' Helm, and J. Deisenhofer, Crystal structure of the outer membrane active transporter FepA from Escherichia coli. Nat Struct Bioi. 1999,6:56-63. Cowan S. W., T Schirmer, G. Rummel, M. Steiert, R. Ghosh, R. A. Pauptit, J. N. Jansonius, and J. P. Rosenbusch, Crvstal structures explain functional properties of two E. coli pOl'ins. Nature. 1992,358:727-733. Deisenhofer, J., O. Epp, K. Miki, R. Huber, and H. Michel, Structure of the protein subunits in the photosynthetic reaction centre of Rhodopseudomonas viridis at 3A. resolution. Nature. 1985,318:618-624. Ferguson, A. D., E. Hofmann, J. W. Coulton, K. Diederiche, and W. Welte. Siderophore-mediated iron transport: crystal structure of FhuA with bound lipopolysaccharide. Science. 1998,282:2215-2220. Forst, D., W. Welte, T Wacker, and K. Diederichs, Structure of the sucrose-specific porin ScrY from Salmonella tvphinntriu111 and its complex with sucrose. Nat Struct Bioi. 1998, 5:37-45.

126

Kreusch, A., A. Neubuser, E. Schiltz, J. Weckesser, and G. E. Schulz, Structure of the membrane channel porin from Rhodopseudol11on£ls blastica at2.0 Aresolution. Protein Sci. 1994, 3:58-63. Pebay-Peyroula, E., G. Rummel, J. P. Rosenbusch, and E. M. Landau, X-ray structure of bacteriorhodopsin at 2.5 angstroms from microcrystaJs grown in lipidic cubic phases. Science. \997,277:1676-1681.

Bundles and Barrels Schirmer, T., T. A. Kellel-, Y-F. Wang, and J. P. Rosenbusch, Structural basis [01' sugar translocation through maltopOlin channels at 3.\ A resolution. Science. \995,267:512514. Weiss, M. S., A. Kreusch, E. Schiltz, U. Nestel, W. Welte, J. Weckesser, and G. E. Schulz, The stn.1Cture of porin from Rhodobactercapsulalus all.8 Aresolution. FEBS Lell. 199\, 280:379-382.

6 x

Functions and Families

10 3

50

100

150

200

250

50

100

150

200

250

Amino terminus

Q)

-0

.:: ;>.,

-5 ell

00 H -0

a

;>.,

::r:: -3

10

Residue number A hyclrcpil'ny plot pred,-ts . day r-sF r nit g leg'ons u! m "bru _ proteir s lik bactcriorhooopsln, s own I ere w th Its TM helices colored to maId he corr.

0

k,

k2

E+A .... EA

L,

Surface step

k3

EA + B .... EAB .... EA + Q,

L

2

L3

where E is the enzyme, A is the mixed micelle, EA is the enzyme-mixed micelle complex, B is the substrate, EAB is the catalytic complex, and Q is the product. (Note that the equation for the first step applies whether the enzyme binds nonspecifically, in which case A is the sum of the molar concentrations of the lipid and the detergent, or specifically to a phospholipid species, in which case A is the molar concentration of that lipid.) Once bound, the association between EA and B is a function of their surface concentrations, expressed in units of mole fraction or mole percent. For a water-insoluble integral membrane enzyme, the protein is delivered to the assay as a detergent-protein mixed micelle, which is likely to fuse with lipid micelles. In this case, E represents the concentration of the enzyme-detergent complex. The kinetic equation becomes v

=

Vmax[AJ[B]

-

Ks A Km B + Km B [AJ + [AJ[BI

}

where the dissociation constant, KsA = k.., /k" and the interfacial Michaelis constant, KmB = (k_ 2 + k3)/k 2 , are expressed in surface concentration units.

Constant mol percent phosphatidylserine

1.2

:J-

Bulk step

~ 1.0

l:>.

l:>.

.D ~~

eU

c:J

.D

~ 0.8

CIl

~

:.a;Q -

s:;

2.... 'v

0.6

~

o

E::

2 0.4

0. ::r:: ~ .....,

Constant bulk concentration phosphatidylserine

""""'0

'-'

0.2 0

0

0.25

0.5

0.75 1.0 1.25 1.5 Triton X-IOO (WN %)

1.75

2.0

6.1.1. Surface dilution effect on the lipid-dependent enzyme, PKC. Redrawn from Gennis, R. B., Biomembranes: Molecular Structure and Function, Springer-Verlag, 1989, p. 228. © 1989 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

Membrane Enzymes

129

([substrate]/{[substrate] + [total lipid]}) or mole percent (mole fraction x 100). Then the classical MichaelisMenten equation is applicable, with surface concentration units used for the substrate concentration (see Box 6.1). When an enzyme that loses activity upon dilution of the lipid is solubilized and reconstituted into micelles or liposomes, the amount of lipid remaining will affect its activity. For this reason, the extent of separation of lipid and protein components during solubilization of the membrane components (see Chapter 3) is critical in studies of membrane enzymes. Only a few high-resolution structures of integral membrane proteins that are classical enzymes are available (see Chapter 9) in addition to those involved in energy transduction and transport. HO'vvever, many membrane enzymes have been extensively characterized biochemically. In addition, some enzymes that are integral membrane proteins have extensive soluble portions that can be removed by proteolytic cleavage and crystallized. When the soluble portion of the enzyme carries out the catalysis, its structure reveals the binding site and catalytic groups to give a picture of the enzyme Function, even though it is missing the portion that anchors the enzyme in the membrane and perhaps plays a regulatory role. Diacylglycerol kinase (DGK) is an example of a well-characterized mem brane enzyme lacking a complete high-resolution structure. Some of the P450 cytochromes provide examples of membrane enzymes whose soluble portions have been crystallized and their structure solved. Both of these examples are enzymes that occur in mammals in numerous isoforms, different Forms of the enzymes that are encoded by difFerent genes. IsoForms, also called isozymes, are catalytically and structurally similar and are typically located in different tissues oFthe organism, where they respond to different regulators. Diacylglycerol Kinase

Diacylglycerol kinase carries out the reaction Diacylglycerol

+ MgATP

---+

Phosphatidic acid

+ MgADP with Michaelis-Menten kinetics and rates limited by substrate diffusion. Both the substrate and product of the DGK reaction are allosteric effectors and second messengers in signal transduction in mammals, which have 10 isoForms of DGK. Localized to the cytosol or the nucleus, the mammalian DGKs are peripheral proteins that dock on the membrane to access their substrate. Two of the isozymes are activated by both PE and cholesterol and inhibited by sphingomyelin when reconstituted in large unilamellar vesicles. All the mammalian isozymes appear to have specialized roles in signaling based on their diFFerent sites and pat-

terns of expression. Since lower organisms such as the nematode worm Cael10rhabditis elegans and the fruit fly Drosophila melal10gaster have only a few isozymes of DGK, and none has been detected in yeast, the mammalian isoforms appear to be involved in processes of development, neural networking, and immune functions that are essential in higher vertebrates. The E. coli DGK provides an example of a very well-characterized integral membrane enzyme whose stnlcture has not been determined at high resolution. Located in the inner membrane, DGK functions to replenish phosphatidic acid. The phosphatidic acid is needed in a surprising turnover of membrane phospholipid that provides the cell with osmoprotectants called membrane-derived oligosaccharides (MDOs). MDOs are made in the peri plasm under conditions of low osmolarity, when they can account For up to 5% of cell dry weight. Because they are water soluble and too large to diffuse through the porins, MDOs stay in the peri plasm and keep it from shrinking too much. MDOs contain six to 12 glucose units joined by ~-1,2 and ~-I ,6 linkages that are variously substituted with sl1-1-phosphoglycerol, phosphoethanolamine, and 0succinyl ester residues. The phosphoglycerol and phosphoethanolamine are enzymatically added from PG and PE, respectively, leaving diacylglycerol. It is the job of DGK to phosphorylate the diacylglycerol to return it to the phospholipid pool in the membrane. The activity of DGK in E. coli is determined by the rate of TM flip-flop supplying diacylglycerols from the outer to the inner leaflet of the plasma membrane. The smallest known kinase, E. coli DGK is a homotrimer of 13-kDa subunits, with three active sites at the subunit-subunit interfaces. It has been purified and reconstituted in detergent micelles and in phospholipid vesicles; in the latter, the enzyme activity depends on the structure of the surrounding lipids. Spectroscopic studies using FTIR spectroscopy and circular dichroism indicate that DGK is ~90% <x-helical, and topology predictions using fusions with f3-1actamase and ~-galactosidase(see below) indicate it has three TM helices (Figure 6.1). Many mutants of DGK have been made, including some that exhibit a remarkable increase in stability. Patterns of disulfide bond formation between singlecysteine mutants indicate the second TM segment from each subunit mediates trimerization by forming a bundle within the trimer. Indeed, addition of a free peptide corresponding to this TM segment interferes with trimer formation. In contrast, the first TM helix appears to be a passive membrane anchor since its replacement with polyaJanine produced an active enzyme. with the polyalanine (influenced by the flanking residues of the protein) spanning the membrane. Several highly conserved residues that are critical for activity « 10% activity when mutated) cluster near the N terminus in a

Functions and Families

130

A.

Periplasl11

Cytoplasln

B. TM-l: 31

50

KKKK-WINEAAFRQEGVAVLLAWIACWLDV-KKKK TM-2: 50

75

KKK-VDAITRVLLISSVMLVMIVEILNSAI-KKK TM-3: 95

121

KKKK-DMGSAAVLIAIIVAVITWAILLWSHFG-KKKK 6.1. A and B. Model of the predicted structure of DGK. The protein has two amphipathic helices at the membrane surface and three TM helices, whose sequences are given in (8). The encircled numbers on the model show where each given sequence begins and ends. From Partridge, A. W., et aI., J BioI Chem. 2003, 278:22056-22060. © 2003 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

region (residues 9-28) shown by NMR to form a helix at the interface just before TMJ. NMR studies also indicate that TM J and TM2 are connected by a shon loop (residues 48-52) that is exposed to the aqueous environment. In spite of years of effort, no highly diffracting crystals of DGK suitable for x-ray crystallography have been obtained. However, nanocrystals of the enzyme give promising results with solid-state NMR. P450 Cytochromes

P450 cytochromes are a ubiquitous superfamily of heme-containing monooxygenases that are named for their absorption band at 450 nm. They are involved in metabolism of an unusually wide range of endogenous and exogenous substances. They panicipate in the metabolism of steroids, bile acids, fatty acids, eicosanoids, and fat-soluble vitamins, and they convert lipophilic xenobiotics (foreign compounds) to more polar compounds for funher metabolism and excretion. P450 cytochromes catalyze hydmxylation of an organic substrate, RH, to R-OH with the incorporation of one oxygen a tom of O 2 , while reducing the other oxygen atom to H 2 0. Their source of reducing power is NAD(P)H, with electrons either donated directly fmm a flavin-containing reductase (class II) or shuttled to the P450 by smaJI soluble electron carrier proteins (class I; Figul-e 6.2). Some self-sufficient P450s in bacteria have

been found to contain heme, flavin, and iron-sul fur centers in one polypeptide. In eukaryotes, most P450 cytochromes are bound to the mitochondrial inner membrane or to the ER. The P450s in the ER are integral membrane pmteins, each bound by a single N-terminal TM domain. Truncation of the N-terminal domain is not sufficient to express soluble protein for crystallization and must be accompanied by several point mutations to disrupt a pel-ipheral membrane-binding site. Even so, detergent is needed to prevent aggregation. High-resolution structures of such constructs show the large soluble domain is a triangular prism shape with ~-sheets along part of one side and ex-helices forming the rest (see Figure 6.2). Structures of these P450 catalytic domains in the presence and absence of substrates reveal that dramatic conformational flexibility must be needed for binding such a variety of compounds. About half of the 57 P450 cytochmmes in the human genome metabolize endogenous compounds, while many of the rest metabolize drugs and other xenobiotics. Some plants have around 300 or more P450s. The genes are classified into families based on sequence identity: to the root symbol CYP is added a number for the family (one of more than 200 gmups with >40% sequence identity), followed by a letter for subfamilies (having> 55% identity), followed by a number for the gene. For example, sterol 27-hydmxylase

Class I

Class II

Heme

e

Fe-S

Heme

e

FAD/NAD(P)H

FMN/FAD/NADPH

6.2. Examples of the two classes of P450 cytochromes. The classes are distinguished by their redox partners. Class I is represented by the P450 system from Pseudomonas putida with P450cam (with heme), putidaredoxin (with Fe-Sl, and putidaredoxin reductase (with FAD/NAD(P)H). Class II is represented by P450BM3 from Bacillus megaterium (with heme) and cytochrome P450 reductase from rat liver (with FMN/FAD/NADPH). The high-resolution structures for P450cam and P450BM3 have been solved for proteins lacking their TM segments. From Li, H., and 1. L. Poulos, Curr Top Med Chern. 2004, 4: 1789-1802. ~, 2004. Reprinted by permission of Bentham Science Publishers Ltd.

is CYP27A and vitamin D 24-hydroxylase is CYP27B, because their amino acid sequences are >40% identical (placing them both in CYP27) but ,

::r:

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c.

-20

V

-

i

I

I

I

I

I

I

I

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c

Current prediction

~

.. .. ..

"

· I I. I

.

a

.., .

... 5 .

'{j sO

Input layer

~ S1

First or hidden layer

~ s2

Second or output layer

6.4.1. Neural networks compute relationships between residues in layers to make predictions based on aligned sequences. From Rost, B., and C. Sander, Proc Natl Acad Sci USA. 1993,90:7558-7562. © 1993 by National Academy of Sciences, USA. Reprinted by permission of PNAS.

~

Q. ::J ...... o

3 OJ

.-+

n' VI -j

Sequence information [rom protein family

Profile delived [rom multiple alignmenl for a window of HI adjacent residues

local alignmellf

1'1 :ldj;l(elll residues

A 100 100 0 0 66

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res , ..

(., r':

.

C

I

L

G

S

0 0 0 0 0 0 0 0 0100 0 0 o 33 66 0 0 0 o 33 o 66 0 0 0 0 0 0 o 33

0 0 0 0 0 33

o

V Nirs NEl 0 0 0 0 33 0 0 o 33 0 0 0 0 0 0 0 0 0 (Xi 0 0

C1:n3

o

Q...

VI

......

o.,

~

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input, local in sequence AA.!l. .F:..A.

Two levels of neural network system

CD

3 0.,

hidden

OJ

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bycr

1.17 0.42 0.92 0.74 1.17 0.74 0.48

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-0

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~.

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,

illlJllt, glohal ill sequcllce {'.Iobal 'I; 1\ /\ statistics L.cngtll wltok 6N·trl1l protein L\C'llllll

frequcncy of' cach amino acid in prolcin Icnglh of prolein ('5' 60. :;;120. ::;240. >NO) disl:H1CC cenl.rc 10 N-Ierm (S40,S.'O,S20.SIO) dislance ccnlre 10 C-Ierm (54lJ.~-'lJ,52U.SIO) firsllevcl:

sc qu cn rc -I 0- S ( I'll CIII re nclwork

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second Icvel: sl rllclll rc-Io-sl rllclll re nctwork iI/pilI: outrul of lSI kvd: Cons. Lell~lh. 'Ic i\ ,\, l~ N-leml. t. C-term fur 11 aJi:lcCIlI rcsidlil'S

6.4.2. PHD uses two levels of neural networks to score structure preferences generated from multiple sequence alignments. From Rost. B" et aI., Protein Sci. 1995, 4:521-533. © 1995, Reprinted with permission from Protein Science.

n-o ;a

5' c

Cl>

.eo

.... .,.. 00

Functions and Families

150

BOX 6.4 (continued)

Amino acid seq:

MGDVCDTEFGILVA ... SVALRPRKHGRWIV ... FWVDNGTEQ ... PEHMTKLHMM ...

State seq:

o oooooooohhhhh ... hhhhi iii iii hhh ... hhhooooOO ... OOOoooohh h ...

Topology:

Tail Out

Tail-Tail

Hetix

SlWl't loop

Helix

Tail - Loop - Tail Long loop

6.4.3. Structural states defined for HMMTOP, where thick lines represent tails while thin lines are loops. Redrawn from Tusnady, G. E., and I. Simon, J Mol Bioi. 1998,283:489-506. © 1998 by Elsevier. Reprinted with permission from Elsevier.

When PHD was tested with an initial dataset of 69 membrane proteins with experimentally determined locations of TM segments, its accuracy was >95%. Assessments of its accuracy when tested on a dataset not used in setting up the analysis are considerably lower. One weakness of the system is the step it uses to filter out TM helices that are too long (>35) or too short «17). If too long, they are split in the middle into two helices; if too short, they are deleted or elongated.

Hidden Markov Models Hidden Markov models (HMMs) apply statistical profiles to describe a series of states connected by transition probabilities. For proteins, each state corresponds to the columns of a multiple sequence alignment, which are intermediate steps in the algorithm that the user does not see. A matrix describes the possible states and the transitions between the states, and an algorithm is employed for a "random walk" through the states, that is, one that derives each possibility from the previous state. For sequence alignment, the HMM generates profiles compiled of high scores (if sequence is highly conserved), low scores (if sequence is weakly conserved), and negative scores (if sequence is unconserved) and identifies sequences with the highest scores. In HMMs for membrane protein topology, structural states are defined to describe portions of the protein. HMMTOP

uses five structural states: inside loop (I), inside tail (i, the region of a loop that is close to the TM helix), TM helix (h), outside tail (0), and outside loop (0) (Figure 6.4.3). TMHMM further divides the residues in TM helices according to whether they are in the center of a helix (helix core) or on one end of a helix (helix cap). Thus seven states are considered in TMHMM: helix core, helix caps, short loop on inside, short and long loop on outside, and globular regions. Each state has a probability distribution over the 20 amino acids, based on the dataset with known topologies. The overall layout of the HMM is a function of the different structural states (Figure 6.4.4). Arrows show the possible connections between the different states, which are limited by the constraints of the protein structure as depicted in (A), with boxes corresponding to one or more states in the model. The connectivity of the different states varies. For the inside and outside loops and helix caps the connectivity is shown in (8), and for the helix core it is shown in (C). This model allows the helix core to be between five and 25 residues, which makes the entire length of TM helices (including caps) between 15 and 35 residues. The program follows rules of "grammar" that state a helix must be followed by a loop, and inside and outside loops must alternate. Then it calculates probabilities for the sequences of these states. This is depicted in the architecture of HMMTOP (Figure 6.4.5), which shows states within the same transition matrices (gray = helix states, yellow = tail states, red = loop states). The rectangular areas represent fixed-length states,

Bioinformatics Tools for Membrane Protein Families

151

BOX 6.4 (continued)

A.

Cytoplasmic side

Me.mbrane

non-cytoplasmic side

Loop

C.

cap

Helix core 22

2J

24

2S

6.4.4. The layout of the hidden Markov model. Redrawn from Krogh, A, et aI., J Mol Bioi. 2001,305:567-580.

"f MA,'(L, ()

MINL,

tail L, MAXL"

h MINL"

helix L" MAXL,

MINL,

({Ii! L,

6.4.5. The architecture of HMMTOP. Redrawn from Tusnady, G. E., and I. Simon, J Mol BioI. 1998, 283:489-506.

© 1998

by Elsevier.

Reprinted with permission from Elsevier.

(continued)

Functions and Families

152

BOX 6.4 (continued) which include the helices whose lengths need to be sufficient to cross the nonpolar domain of the bilayer and the tails that are defined to be the short segments of loops adjacent to ends of helices. The hexagonal areas represent the nonfixed-length states, which are the loops that are allowed to be any length. By considering fixed-length states and non-fixed-

length states, the methods allow realistic length constraints on TM helices. While the current programs using these methods (listed in Table 6.4) are very powerful, new computational approaches can be expected to make them even more useful in the near future.

highest fraction (around three fourths) were correctly predicted with two algorithms using hidden Markov model, TMHMM, and HMMTOP, while MEMSAT and TOPPRED had less success and the lowest fraction (around one half) were correctly predicted with a neural network predictor, PHD. The best results were achieved by carrying out analyses by all five and searching for consensus: a very high rate of correct predictions occurs when all five or four of the five agree. A new algorithm that compares the results of nine different methods is called CONPRED. The success of the predictions is still consistently higher with prokaryotic genomes than with eu kalyotic genomes. Analysis of 26 genomes for membrane proteins produced a total of 637 families of poly topic TM domains, based on a combination of TM helices predicted using

TMHMM and those annotated in Swiss-Prot. The domains were classified based on homologies with known families or characterized using sequence alignment, hydrophobicity plots with the GES scale, and identification of consensus sequences for TM segments (Figure 6.18). When the families are sorted by the number ofTM helices, the number of families with domains of a given number of TM helices decreases as the number of helices increases (Figure 6.19). Within this trend, the plot shows the highest occurrences of two- and four-TM helices, a slight excess of seven-TM helices, and a major spike at twelve-TM helices due to the prevalence of this topology among transporters and channels. Interestingly, the total number of membrane protein domains in the genome is roughly proportional to the number of open reading fTames in all of the genomes

TM·helix

TM-helix

PF01618

•.... ,~","m.,~=,.~",~",["~.=,,,"~,~,~,,~.,~,~,u~~"u~"""'I ..~",=m.,,,~_ v.;ILn:r':Oll

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n=.r."'Y'GLt.;:;t'rtt:ILlt7T':l:l.o'''''~~'''~'ill~'''''~'''~''=-':J'' 'F~''''~''~~ill~'''~''''''''''''

",L1I1lD}U_I" C'1L1l1lliH·_U

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5

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6.18. Classification of a family of polytopic membrane domains. The example shown is family PF01618. The steps involved are (top to bottom) sequence alignment, hydrophobicity plot based on GES scale, consensus sequence displayed by sequence logo, and consensus sequences of TM helices, where the nonconserved amino acids are represented by "x." From Liu Y, et al., Genome Biology 2002, 3:research0054.1-0054.12. © 2002 by Yang Lui, Donald M. Engelman, and Mark Gerstein. Reprinted with permission from the authors.

Bioinformatics Tools for Membrane Protein Families

153

30,--,----------------------,

25 Vl

::l

20

t:

15

cQ)

::l

v

C5 10 5 O""::""~~,!QJ,,~=L.J...I!:j~:w...l!~~~~~

7 8 9 10 11 12 13 Number of TM helices 6.19. The number of TM helices in families of polytopic membrane domains. The number of families of polytopic membrane domains is plotted on the y-axis as a function of the number of TM helices, plotted on the x-axis. The green bars are the numbers from all studied families from the Pfam-A database. The yellow bars are the numbers from families from the Pfam-A database that are annotated as transporters and channels. Redrawn from Liu, Y, et al., Genome Biology, 2002, 3:research0054.1-0054.12. © 2002 by Yang Lui, Donald M. Engelman, and Mark Gerstein. Reprinted with permission from the authors.

D 1% K

A. All residues

except that of the nematode worm C. elegans, which has a huge number (754) of seven-TM chemoreceptors. In fact, the genome of C. elegan5 conforms to the general picture when its three large families of chemoreceptors are removed from the total. Clearly this organism that lacks sight and hearing has finely developed chemosensation to find its food l Approximately half of the membrane proteins identified by genome analysis have an even number of TM helices with the Nin-C in topology. The other three combinations of N- and C-terminal locations are about equally represented in the other half. The high proportion of proteins with both Nand C termini inside is attribu ted to the mechanism of biogenesis, because this topology results from insertion of helical hairpins (see Chapter 7). Amino acid distributions in the TM segments of the putative membrane proteins in the 26 genomes show the expected high amounts of nonpolar amino acids, along with the polar residues Ser and Thr that participate in hydrogen bonding (see below; Figure 6.20A).

B. Positionally conserved residues K

]%

L

16%

I%C

R 1% E 1%0 C

~--rr'r--L

1%

M R D 1%2%2% E

2% 2% H 2%

2%0

2% N 3%

M 4%

H 3%

T 4%

L 12%

V

4%

G 8%

P 9%

A

8%

6.20. Amino acid distributions in TM helices. Amino acid distributions were determined for the TM segments from the 168 families from the Pfam-A database that have more than 20 members. A. A pie diagram of the amino acid compositions of TM helices shows their high content of nonpolar residues, along with glycine, serine, and threonine. B. Consensus sequences identified as shown in Figure 6.18 allow comparison of the amino acid residues in conserved positions of the TM helices. The diagram of the compositions of these positionally conserved residues shows that the prevalence of three amino acids (Gly, Pro, and Tyr) has increased significantly, indicating they are the ones whose positions are highly conserved. Redrawn from Liu, Y, et al., Genome Biology, 2002, 3:research0054.1-0054.12. © 2002 by Yang Lui, Donald M. Engelman, and Mark Gerstein. Reprinted with permission from the authors.

,. .'

Functions and Families

154

Similarly, the composItion of nearly 50,000 TM segments annotated in the Swiss-Prot database (see Box 6.2) reveals that the six amino acids Leu, Ile, Val, Phe, AJa, and Gly make up two thirds of TM residues. When the sequences are aligned to determine conserved residues (see Figure 6.18), the nonpolar amino acids are not prevalent in conserved positions, presumably because they al-e quite interchangeable. Interestingly, the amino acids that are prevalent at highly conserved positions in the helices are Gly, Pro, and Tyr (Figure 6.20B). The proline residues form kinks in the helices, which are conserved even after mutation of the Pro residues, while tyrosine residues playa special role near the interfaces due to their electronic propel1ies (see Chapter 4). The positions of glycine residues are often highly conserved in soluble proteins because they occur at positions where the proteins do not accommodate larger side chains. This is also the case in TM helices, where Gly is commonly observed where two helices are in close contact. The GxxxG motif, llrst observed in glycophorin dimers, places both Gly l-esidues on the same side of the <X-hel ix (see Figure 4.30). Genomic analysis for pair motifs shows a very high presence of GxxxG and GxxxxxxG pairs, as well as similar motifs with Ala or Ser replacing Gly residues. The resulting location of small side chains is expected to be important fOI' helix-helix interactions in a broad range of membrane proteins. Helix-Helix Interactions

The prevalence of the GxxxG motif in poly topic membrane proteins in the genome reflects both the close packing of TM helices observed in many helix-bundle proteins and the importance of protein-protein interactions in oligomeric membrane proteins and complexes. To analyze helical packing patterns within proteins, a comparison of pairs of helices in membrane proteins and pairs in soluble proteins shows that most helixhelix pairs in membrane proteins have homologs in soluble proteins. The exceptions to this correlation are the irregular helices of some membrane proteins (described in Chapter 4). The high occurrence of GxxxG and GxxxA in helices of membrane proteins allows the helices to approach more closely than those in soluble proteins, forming "knob-into-hole" interactions, as well as to interact over longer distances, that is, over the width of the bilayer. The information on helix pairs within proteins probably also applies to helixhelix interactions between subunits because the TM helices from different subunits often align as much as TM helices within one subunit. This can be seen with oligomeric proteins, where a cross-section through the middle of the membrane shows the extensive interaction of helices from different subunits (Figure 6.21): the

Photosynthetic react ion center

(Rhodopseudomonas virdis) Deisenhofer et al. (1995)

I

~J.

,

• liS Cytochrome-e oxidases in bovine hearlmitochondria Tsukihara et al. (1996)

Cytochrome-be I complexes in bovine heart mitochondria Iwata et al. (1998)

6.21. Helix interactions viewed from the membrane midplane. Positions of five-residue sections at the middle of TM helices are shown for photosynthetic reaction center, cytochrome-c oxidase, and cytochrome-bcl complex, as labeled. The subunits are colored differently. The gray-scale image of cytochrome-c oxidase shows that the subunit composition cannot be inferred from the relationships of the helices. Redrawn from Liu, Y, et aI., Proc Natl Acad Sci USA 2004. 101:3495-3497. © 2004 by National Academy of Sciences, USA. Reprinted by permission of PNAS.

gray-scale image of cytochrome-c oxidase in the figure shows how difficult it is to assign subunits from the relationships of the helices! Additional information about helix-helix intemctions comes from the analysis of interhelical three-body interactions to find "triplets" where three atoms from at least two different helices are closely packed. A program called INTERFACE-3, which computes interhelical atomic triplets based on algorithms for geometric shapes and distances, detects six diffel-ent types of triplets in the glycophOlin dimer: GGV, GTV, GVV, ILL, nT, and ITT (Figure 6.22A). Likewise, bacteriorhodopsin and its homologs halorhodopsin and sensory rhodopsin II have three triplets that are conserved in sequence and in structure (Figure 6.22B). Furthermore, an additional 13 triplets are conserved structul-ally but not fOl-med by identical residues, allowing conservative mutations that suggest that the triplet interaction has been conserved to maintain the orientation of the helices in the three highly homologous proteins. Comparison with a set of soluble <x-helical proteins identified triplets that are unique to membrane proteins, such as AGF, AGG, GLL, and GFF. Such triplets are often found in regions of the closest contact between helices, and are therefore often correia ted with GxxxG-type motifs. Close contact between TM helices is also stabilized by two types of hydrogen bonding. The hydrogen bonds

Bioinformatics Tools for Membrane Protein Families

B.

A.

a

b G79:C V80:Ca

V80:Ca

6.22. Three-body contacts, or triplets, in helix-helix interactions. A. The TM helices (residues 73-91) of a glycophorin dimer with one of the triplets illustrated in space-filling representation. The atoms involved, C from Gly79 and Ccx from Val80 on both chains, are shown in space-filling representation (a), and viewed from the top (b). (Orange, C from G79, chain A; green, Ccx from V80, chain A; blue, Ccx from V80, chain S.) B. A conserved triplet in the archaei rhodopsins is shown by superposition of helices C, E, and F from bacteria rhodopsin, halorhodopsin, and sensory rhodopsin II. The residues in the triplet are two Leu and a Thr, corresponding to L97, L152, and T178 in SR, again in space-filling representation. The retinal is drawn in green From Adamian, L., et aI., J Mol Bioi. 2003, 327:251-272. © 2003 by Elsevier. Reprinted with permission from Elsevier.

observed in glycophorin dimers are relatively weak bonds because they use the CO( proton as the donor. Each of these hydrogen bonds contributes less than 1 kcal/mol to the stability of the dimer. The second type B.

A.

155

of hydrogen bond makes use of polar residues in TM segments; such bonds are detected in high-resolution structures of poly topic membrane proteins such as bacteriorhodopsi n. Interactions between polar residues account for around 4% of all atomic interhelical contacts in membrane proteins, as ,.veil as in soluble proteins. In contrast to soluble proteins, where interacting ionized residues typically form salt bridges, the types of polar interactions are more varied in TM segments of membrane proteins, which also have H-bonds bet,veen ionizable and polar residues (the most common are DY and Y-R) and between polar nonionizable residues (the most common are Q-S and S-S). How prevalent are these interhelical hydrogen bonds? In a dataset based on the high-resolution structures of 13 membrane proteins, 134 unique TM helices form nearly 300 helical pairs, 53% of which are connected by at least one H-bond. In the 299 interhelical H-bonds identified, almost half involve Ser, Tyr, Thr, and His. Hydrogen bonds between two side chains and hydrogen bonds between a side chain and a backbone carbonyl oxygen occur at the same frequency. The majority of these helical pairs have one H-bond between two amino acids from two neighboring helices. However, two other types ofH-bonding patterns emerge. One type is a "seline zipper" motif, named for its similarity to a leucine zipper (Figure 6.23). A search for homologous serine zippers by PSI-BLAST identified more than 100 sequences with highly conseJ\led Ser residues in positions 7, 14, and 21 of one helix and 1,8, and 15 of the other. A three-body analysis also found triplets of two serine residues with a leucine. The other type of H-bonding pattern is called a polar clamp because the

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6.23. The serine zipper motif in helix-helix interactions. A. Schematic representation of hydrogen bonding between two serine residues on two adjacent helices. B. An example of a serine zipper in bovine cytochrome-c oxidase. The two helices shown are helices III and IV, and the hydrogenbonded serine residues are 5101-5156, 5108-5149, and 5115-5142. C. The pairs of serines can be predicted from helical wheels of helices III and IV from subunit 1 of bovine cytochrome-c oxidase. A helical wheel designates residues i and i + 3 as a and d and shows how they fall on the same side of the helix. Nonpolar residues are shaded pink. Three 5er-5er pairs and two Leu-Leu pairs form a zipper at different a and d positions in this example. A and B redrawn from Adamian, L., and J. Liang, Proteins. 2002, 47:209-218. © 2002. Reprinted with permission from John Wiley & Sons, Inc.; C from Adamian, L., et aI., J Mol Bioi. 2003, 327:251-272. © 2003 by Elsevier. Reprinted with permission from Elsevier.

Functions and Families

156

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6.24. Polar clamp hydrogen bonding in helix-helix interactions. A. Three residues in rhodopsin, W161, T160, and N78, form a clamp between helix IV and helix II. The side chain of N78 (on helix II) makes two hydrogen bonds: its 0 atom is hydrogen bonded to the N atom from W161, and one of its amide hydrogen atoms is hydrogen bonded to the oxygen of T160. B. A polar clamp in subunit I of cytochrome-c oxidase from Thermus thermophifus involves residues S155 and S159 from helix 0:4 and 086 from helix 0:2. From Adamian, L., and J. Liang, Proteins. 2002, 47:209-218. © 2002. Reprinted with permission from John Wiley & Sons, Inc.

side chain of an amino acid at a given posItIOn I that is capable of forming at least two hydrogen bonds (i.e., E, K N, Q, R, S, T) is "clamped" by H-bonding to two other residues, one at position i + 1 or i + 4 and one on the othel- helix (Figure 6.24). The polar clamp shown in rhodopsin, involving l-esidues T160 and WI61 (both on helix IV), and N78 (on helix U), is highly conserved in the GPCR family. When the numbers of intermolecular and intramolecular helix-helix interactions are estimated for membrane proteins in whole genomes, the totals are in the millions. Clearly an understanding of them will deepen as the number of high-resolution stnlctures increases and will provide in turn a more complete insight for predicting membrane protein structure. Furthermore, helix-helix interactions are a critical step in the assembly of polytopic membrane proteins, as desclibed in the next chapter. Since the nonpolar residues are quite nonspecific in their interactions, the often conserved hydrogen bonds between polar residues in TM segments must be crucial in helix alignments. Proteomics of Membrane Proteins

With thousands of membrane proteins predicted by genomic analysis, many questions are raised about their

identities and interactions. Using TMHMM predictions combined with PhoA/GFP fusions to localize the C termini, a global topology analysis of E. coli membrane proteins detected 601 inner membrane proteins. When these proteins are sorted by their known or predicted functions, 40% of them are involved in transport, while nearly 20% are involved in metabolism, biogenesis, and signaling (Figure 6.25). Over one third are "orphans" with unknown functions. The functions of some orphan memb.-ane proteins can be ascertained from their association with known proteins in complexes. (The yeast two-hybrid analvsis, an important proteomic tool for identifying proteinprotein interactions in complexes, does not work with membrane proteins because loss of function in the fusion proteins generated can result from loss of correct compartmentalization, topology, or orientation in the membrane.) Complexes of membrane proteins can be detected by two-dimensional gel electrophoresis using native gel electrophoresis for the first dimension, combined with mass spectrometry to identify the proteins. To carry out this procedure with Gram-negative bacteria such as E. coli, the inner and outer membrane fractions were first separated by gradient centrifugation and solubilized in a mild detergent, such as 0.5% 11dodecyl-0-D-lllaltoside. When native gel electrophoresis was carried out in Coomassie blue, complexes involving 44 inner membrane proteins and 12 oute.- membrane proteins were isolated. The results enabled roles to be assigned to six Ol-phan proteins of unknown function, in addition to identifying a number of Imown proteins. A majority of the inner membrane proteins in complexes are members of .-espiratory chains (e.g., succinate dehydrogenase, F I Fo-ATP synthase, and cytochrome b0 3 ubiquinol oxidase). Others are involved in biogenesis (including the SecYEG translocon; see Chapter 7) and

Metabolism - 7% Channels - Go u, in kcal/mol) as a function of the hydrophobic thickness of the PC bilayer. For saturated and monounsaturated chains (filled circles), the L'>Go u increases linearly with chain length (thickness), while for cis double-unsaturated acyl chains (open circles), it decreases linearly with chain length. From Hong, H., and L. K. Tamm, Proc Natl Acad Sci USA. 2004, 101 :4065-4070. © 2004 by National Academy of Sciences, USA. Reprinted by permission of PNAS.

Classical il1 vitro folding studies in lipids or detergents have now been carried out with several other membrane proteins, including the enzyme DGK, the lactose permease, and the potassium channel, KcsA. In addition, some of these studies employed different strategies to gain different types of information about folding membrane proteins. For example. the stability of various mutants of DGK was assessed with thermodynamic analysis of refolding the partially denatured pmtein. When DGK is reversibly denatured in SDS, it retains most of its a-helical content (as is typical of soluble proteins in SDS). The refolding that takes place upon the removal of SDS is sufficient to differentiate the stability of different mutants.

Protein Folding and Biogenesis

170

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7.11. Ribbon diagram showing the lowest energy conformer of Mistic determined by NMR. From Roosild, 1. P., et aI., Science. 2005, 301: 1317-1321. © 2005. Reprinted with permission from

MAS

To study the TM portion of the KcsA potassium channel (see Chapter 9), a novel approach combined pallia] de novo synthesis with folding. The semisynthetic TM protein was constructed by fusion of a recombinant peptide (residues 1-73) with a synthetic peptide (residues 74-125). The refolding of this construct was then used to define the lipid requirement for folding. Folding of membrane proteins can be a very important step of theil- purification after they are harvested in denatured form from insoluble inclusion bodies, which are lipid-bounded storage sites within the cytoplasm of bacterial cells that are overproducing the proteins from high-expression vectors. An example of a membrane protein that was refolded after it was obtained in inclusion bodies is the enzyme OMPLA (described in Chapter 10). The denatured OMPLA was solubilized in 8 M urea and diluted into Triton X-lOa, which produced a mixtlll'e of folded and unfolded OMPLA that was resolved by ion-exchange chromatography to recover the native enzyme. The choice of detergent is critical, as refolding studies under different conditions revealed a strong preference for Triton X-lOa in the case of OMPLA. Often, however, refolding from inclusion bodies is not effective for membrane proteins. An alternative procedure is now available that makes use of Mistic (the acronym for Membrane Integrating Sequence for Translation of Integral Membrane Protein Constructs), a l3-kDa protein from Bacillus subtilis that spontaneously inserts into membranes in vivo. The structure of Mistic \.vas determined using sophisticated NMR methods along with site-directed spin labeling, which revealed a four-helix bundle with an exposed periplasmic C terminus (Figure 7.11). In spite of its hydrophilic character, Mistic associates tightly with the membrane. When it is expressed as a fusion with an integral membrane protein, it has a unique ability to insert the TM domains from the membrane proteins. Over 20 eukaryotic membrane proteins have now been overexpressed

in E. coli as Mistic fusions, and those tested had functionality. The use of Mistic might overcome the usual problems caused by expressing high amounts of eukaryotic membrane proteins in bacteria: (l) their targeting signals may not be recognized by the host, and (2) their overexpression is toxic because it clogs the machinery for inserting the bacterium's own membrane proteins or causes other problems like proton leaks. Finally, the use of in vitro transcription/translation systems to synthesize mem brane proteins in the presence of lipid vesicles or micelles has revealed conditions needed for folding and insertion. This approach was pioneered in studies of the E. coli outer membrane protein PhoE (see Chaptel- 5) and demonstrated that folding and insertion of PhoE require lipopolysaccharide. In similar studies of the E. coli lactose permease (see Chapter 10), protein folding took place only with inside-out vesicles, mimicking the vectorial insertion of the lactose permease into the inner membrane.

BIOGENESIS OF MEMBRANE PROTEINS

Folding and insertion of membrane proteins are just two elements of the very complex process needed for the assembly of proteins into cell membranes. The first steps of that process are shared by membrane and secreted proteins and utilize a complex apparatus called a translocon that has been conserved through eukaryotic, bacterial, and archaei kingdoms. In later steps integral membrane proteins are differentiated from secreted proteins by a process that allows their lateral integration into the lipid bilayer and detel-mines their topology. The process of integration and topogenesis (the genesis of topology) will be examined after describing the process for export and the steps involved in translocation. Export from the Cytoplasm

Since membrane proteins are not synthesized by a special population of ribosomes, they must be targeted for their destinations as they exit the ribosomes. In eukalYotes, they may enter the ER and end up in the plasma membrane or endocytic organelles (the ER pathway) or they may follow other pathways for mitochondrial, chloroplast, and nuclear membranes. The destinations in prokaryotes are the plasma membrane and, in Gramnegative bacteria, the outer membrane. Once targeted for export, the processes of translation and translocation may be coupled (cotranslational translocation) or sequential (posttranslational translocation). Three types of signals initiate the export of membrane proteins: cleavable signal sequences, signal. anchors, and reverse signal-anchors. (Since reverse signal-anchors differ only in the resulting orientation

Biogenesis of Membrane Proteins

171

BOX 7.2. EVidence for cleavable signal sequences Involved in protein translocation The protease protection assay distinguishes between translocated membrane proteins and secreted proteins and provides evidence that both can have cleavable signal sequences. The experiment uses SDS polyacrylamide gel electrophoresis (PAGE) to distinguish products of in vitro protein synthesis in the presence or absence of membrane vesicles, with and without added protease. When translation of a protein with a classical N-terminal cleavable signal sequence is carried out in vitro in the absence of membranes, a precursor containing the signal is produced. The precursor is not protected by membranes, so it is digested by added protease (sample 1 in Figure 7.2.1). When it is carried out in the presence of membrane vesicles, the signal insert into the membrane, along with any TM segments of the protein (samples 2

and 3). Signal peptidase on the membrane cleaves the signal from both secreted and inserted proteins, resulting in mature forms that migrate faster on SDS-PAGE as shown. To determine whether the protein is integrated into the membrane or secreted across it, a nonspecific protease such as proteinase K is added. If the protein is soluble and outside the vesicles, it is fully digested (sample 1); if it is integrated into the membrane, the external portion is digested, resulting in a smaller protein or fragments (sample 2); and if it is fully transported into the vesicle (secreted), it is protected from the protease and retains its size (unaltered mobility on the gel, sample 3). Many applications of this basic protocol have revealed the nature and role of signal sequences on hundreds of proteins.

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9.33. The environment of 11-ci5 retinal in bovine rhodopsin. A. The amino acid residues in the vicinity of the 11-c;5 retinal (pink) include Lys296 on H7, which forms the Schiff base linkage to retinal, and Glu 113 on H3, its counter-ion that is H-bonded to the peptide N of Cys187 in f)4 near the plug to the extracellular surface. From Palczewski, K., Annu Rev Biochem. 2006, 75:743-767. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org. B. The retinal binding pocket is defined by the electron density map in bovine rhodopsin. C. A sketch of the residues that surround the retinal indicates ligand bonds, hydrogen bonds, and polar and hydrophobic contacts. From Li, J., et ai, j Mol BioI.; 2004, 343: 1409-1438. © 2004 by Elsevier. Reprinted with permission from Elsevier.

Membrane Enzymes and Transducers

234

C3 Loop 9.34. The cytoplasmic domain that interacts with transducin. The cytoplasmic region consists of the four segments that project above the lipid phosphate groups: three cytoplasmic loops, C1, C2, and C3; the peripheral helix H8 (purple); and the (truncated) C-terminal tail. Side chains of individual residues that interact with transducin are shown, including those that mutate to give constitutive activation (green). those that have been cross-linked to the transducin ex subunit (Gtex; red). those that inhibit stabilization of Metall by Gtex (pink). those that reduce activation (yellow), and those that regain function upon reverse mutation from Ala (greenish yellow). The footprint of retinal (light pink) from the TM region of rhodopsin illustrates its alignment with key residues at the interface. Interhelical cross-links that inhibit activation are indicated by chains from H6 to nearby helices. From Li, J, et aI., J Mol BioI. 2004, 343: 1409-1438. © 2004 by Elsevier. Reprinted with permission from Elsevier.

retinal fits at the second kink in helix 2, bordered by hydrophobic groups, except for Glu 122. Some of the polar residues in the binding pocket are those that vary in the color pigments, alJowing the pigments to absorb light of different wavelengths. After its isomerization is triggered by absorption of a photon, retinal no longer fits the pocket, and conformational changes occur in the active intermediate that enable it to leave by hydrolysis of the Schiff base. These structuml changes are the subject of much current investigation employing various NMR, EPR, and mutagenic approaches since there is no high-resolution structure of the active intermediate called MetaU. Mutagenesis to introduce Cys residues For disulfide formation and spin-labeling studies has allowed comparisons between distances across the activated and ground-state structures. The results suggest that activation results in an expansion and opening of the cytoplasmic end, which could open a crevice For Gprotein binding. This activation mechanism is thought to be a general Feature of GPCR activation. As the portion of rhodopsin that interacts with transducin, the cytoplasmic domain receives much interest. This region includes the three cytoplasmic loops and adjoining ends of the TM helices, along with the peripheral helix and the C-terminal tail (Figure 9.34). It contains many of the residues that are highly conserved among different GPCRs, including the (D/E)R(YIW) and NPXXY motifs described above, and

is the site of many mutations that affect rhodopsi n activity. It is also the site of the main differences between the two crystal structures of bovine rhodopsin, and its loops have the highest crystallographic B factol~ suggesting it is the most flexible portion of the structure (see Figure 8.28 and Frontispiece.) A Fairly new question about the interaction of rhodopsin with transducin is the oligomerization state of rhodopsin in native membranes. Long considered a monomel~ rhodopsin has been shown by SDS-PAGE, EM, and AFM to be dimeric. The dimer contact is thought to involve helices 4 and 5 (Figure 9.35). The data suggest higher oligomers are likely in disc membranes and these may form and dissociate dynamicaJJy and may have important functional roles. If rhodopsin dimers bind transducin, rhodopsin is an even better prototype F01- a general signal transduction mechanism since many other GPCRs appear to function as dimers. Genomic and proteomic studies of the GPCRs indicate they have strong evolutionary relationships in spite of their diverse functions. Since they respond to different ligands or stimulants, the portion of the molecules that diverges the most is the ligand-binding domain. This domain is usually on the extracellula.- surface, unlike the buried [-etinal of rhodopsin. The binding sites of many GPCRs have been characterized by Cys scanning to probe accessibility of different residues to sulfhydryl reagents in the absence and presence of lig· ands. With many GPCRs, the binding site is the most studied domain, as it is valuable for designing new

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9.35. Model of a rhodopsin dimer. The dimer is viewed from the cytoplasm and includes an inactive rhodopsin subunit (pink) and an activated subunit (yellow, marked with an asterisk). (Acidic residues are red and basic residues are blue.) The model has the constraints from the x-ray structure fitted to the AFM data from native membranes. Cross-linking data were applied to model the activated form. From Palczewski, K., Annu Rev Biochem. 2006, 75:743-767 11;) 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

Transducers

9.36. Model of the structure of another GPCR, the atl angiotensin receptor. based on information from the rhodopsin structure. The functional regions are shown in space-filling models, with ligand binding (red), signal propagation (green), and G-protein binding (blue) from the crystal structure of rhodopsin. The positions of these regions are based on locations of important residues in the sequence of the atl receptor. From Filipek, S, et al., Annu Rev Biophys Biomol Struct. 2003, 32:375-397 © 2003 by Annual Reviews. Reprinted with permission from the Annual Review of Biophysics and Biomolecular Structure. www.annualreviews.org.

drugs that interFere with binding. Since details about the TM domain are lacking, the structure of other GPCRs are modeled based on the crystal structure of rhodopsin, as exemplified in a model [or the angiotensin receptor (Figure 9.36). Thus. as the first high-resolution structure of a GPCR, the structure of rhodopsin is a highly significant prototype.

235

Bacteria maintain a slight outward turgor pressure as they respond to varying osmotic conditions. When osmotic pressure increases, the cells accumulate solutes such as betaine, proline, and potassium ions to balance the pressure and minimize water efflux. When osmotic pressure drops, they avoid rupture by opening MS channels For solute efflux. MS channels were discovered in 1987 in patch clamp experiments on giant bacterial spheroplasts (see Chapter 3). E. coli has three types of nonspecific MS channels, named for their different single-channel properties: MscL (large), MscS (small), and MscM (mini; Figure 9.38). The smallest channels, MscM, open at lo'vv tensions, are not essential, and are poorly characterized. Two species of the small MS channels that open next have been identified, MscK, which opens a K+ channel, and MscS, which is anion specific. At high tension, near that which would rupture the cell, the large nonselective channel MscL opens. Double mutants thal lack both MscL and MscS do nOl survive osmotic downs hocks. Furthermore, introducing MscL from E. coli into marine bacteria gives them increased resistance to drops in osmotic pressure. MscL MscL, the large MS channel of the E. coli inner membrane, Forms a nonselective ion channel that is activated by quite high levels of membrane tension. The MscL pore is large, giving a conductance of 2.5 nS in vitro. AFter crystallization attempts with a dozen MscL homologs from different bacteria, the MscL Crom Mycobacterium tuberculosis was crystal Iized fTom dodecylmaltoside and its structure solved at 3.5 A resolution. This protein, called Tb-MscL, has 37% sequence identity with the E. coli MscL, Eco-MscL, and its structure is consistent with cross-linking and EM data on Eco-MscL. Tb-MscL is a homopentamer consisting of two domains, a TM domain and a cytoplasmic domain

Mechanosensitive Ion Channels

Mechanosensitive (MS) channels respond to mechanical stresses applied to the membrane or to membraneattached elements of the cytoskeleton, enabling organisms to respond to touch, sound, pressure, and gravity. They fall into two broad classes, depending on whether or not cytoskeletal elements are involved (Figure 9.37). MS transduction in vertebrate auditory hair cells provicks examples of fast and sensitive responses involving the cytoskeleton. The prokaryotic MS channels that respond to deCl-eased osmotic pressure exemplify the class that does not involve the cytoskeleton. Crystal structures of two of the prokaryotic MS channels illustrate the best understood models for how membrane tension can drive conformational change and lead to channel opening.

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Membrane Enzymes and Transducers

236

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and van der Waals contacts involving many conserved residues (Figure 10.17). The cavity has several patches of basic residues located at positions ranging from the entrance Lo the bottom. It also has a ladder of tyrosine residues (Tyrl94, Tyr190, and Tyr186) entering the cavity along helix 4. At the third basic patch the cavity is constricted to 8 A. by Tyr186 and three basic residues (Lys22, Arg79, and Arg279). Many of the charged and polar residues are linked by extensive hydrogen-bond

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10.18. A large hydrogen bond network in the AAC at the binding site for the inhibitor CATR. Highly conserved polar, acidic, and basic residues, along with main-chain carbonyls (labeled CO) are hydrogen bonded, often via numerous water molecules. From Nury, H., et aI., Annu Rev Biochem. 2006, 75:713-741. «;; 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

networks (Figure 10.18). These networks link all the TM helices except helix 4, which deviates from the threefold symmetry and has the bulk of the buried aromatic residues. The inhibitor binds at or near the binding site for ADP, as ATR blocks uptake of ADP from the IMS, so the cavity can be analyzed as a nucleotide-binding site. The patches of positive charge \vill attract the anionic nucleotide to the entrance. The funnel-shaped opening

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10.17. Two-dimensional projection of the residues present at the surface of the cavity in the AAC basket. Each circle represents an atom of a residue located within the cavity, with a size proportional to its solvent accessibility and a color indicative of the type of residue (basic, blue; acidic, red; aromatic, gray; hydrophobic, yellow; polar, green). The tyrosine ladder is marked by Y and four positive patches are circled and numbered. From Nury, H., et aI., Annu Rev Biochem. 2006, 75:713-741. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

Transporters and Channels

252 A.

10.19. Conserved residues on external surfaces of Me. Sequence similarities within ADP/ATP carriers are mapped onto the structure of the bovine MC and colored to show extent of homology (0% to 100%, white to red). The highly homologous regions include residues lining the cavity and are well defined on both the end of MC that faces the IMS (A) and that facing the matrix (B). From Nury, H., et al., Annu Rev Biochem. 2006, 75:713741. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

Homology

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of the cavity will orient an incoming nucleotide substrate with stacking interactions with the row of tyrosine residues along helix 4. Selectivity of the transport for adenosyl and not guanylyl nucleotides can be explained by specific interactions with conserved residues. The adenosyl amino groups and sugar hydroxyl groups will form hydrogen bonds with polar residues, while the phosphoryl groups of the nucleotides will interact with the basic residues. Hypothesized Transport Mechanism The exchange process catalyzed by AAC requires that it bind to and release substrates on opposite sides of the membrane. Binding ATP from the matrix is predicted to disrupt the salt bridge between Arg236 and Glu264 (shown in Figure 10.16), which will shi ft the extensive hydrogen-bond networks stabilizing this conformation. Thus binding of negatively charged substrates perturbs the salt bridges at the bottom of the cavity and shifts the network of polar interactions to trigger conformational changes in the carrier molecule. Interestingly, among AACs fyom different species the most conserved residues are inside the cavity and include several basic residues, along with aromatic residues to an unusual degree (Figure 10.19). It is clear from the high-resolution structure of the conformation that opens to the IMS that AAC must undergo large conformational changes during transport. Large movements of TM helices are needed to invert the "basket" when AAC opens to the matrix. Most likely the proline residues that form kinks in the odd-numbered helices are the hinges that permit these conformational changes (Figure 10.20). This may be a general mechanism for mitochondrial carriers, since these proline residues are part of a highly conserved sequence found in each of the three repeats of AAC and other members of the MCF. The MCF motif is PxD/ExxKlRxKlR-(20-30 residues)-D/EGxx.;xxaKlRG, where the letter x denotes any amino acid and the letter a denotes an aromatic residue. In the AAC structure this motif is found in the bottom of the cavity where matrix loops close off the basket. Specifically, in the third repeat

of the AAC. the MCF motif includes the Glu264-Arg236 salt bridge along with Arg271, which clamp together the matrix ends of helix 5 and helix 6 at the bottom of the basket. The role of the MCF motif in shaping the "basket" seen in the structure of AAC suggests a common architecture and mechanism for mitochondrial calTiers. Many mitochondrial carriers are proposed to hmction as dimers. and a variety of biochemical and biophysical data have indicated that AAC forms a dimeI'. Covalent dimers made by fusing two tandem copies of the yeast gene for AAC produced functional carriers. However, the crystal structures strongJy suggest that a monomer of AAC can perform its transport lntelmembrane

space

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Matrix 10.20. Proline-induced kink in helix 1 of the Me. This example shows the kink after Pr027 in helix 1, where the carbonyl of Thr23 hydrogen bonds with Trp70. which interacts with cardiolipin (green). Hydrogen bonds are indicated by gray lines. From Pebay-Peyroula, E., and G. Brandolin, CurrOpin Struct BioI. 2004, 14:420-425. © 2004 by Elsevier. Reprinted with permission from Elsevier.

Channels

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of the pore itself, in addition to special features like vestibules (both external and internal) that attract ions and selectivity filters that exclude other ions from the pores. Further, many channels have gating mechanisms that enable them to open in response to outside signals communicated via ligand binding, electric potentials, and/or pH. Channel gating involves the response mechanism itself and the conformational change it stimulates. All these characteristics are better understood thanks to high-resolution structures of several AQPs and potassium channels. Both families of channels can)' out passive transport: the AQPs transport water in response to an osmotic pressure gradient, while potassium channels are driven by the electrochemical K+ gradient created by the Na+K+-ATPase. The chapter ends with a description of the calcium pump, an ion channel that undergoes large conformational changes of its cytoplasmic domains driven by the hydrolysis of ATP. Aquaporins and Glyceroaquaporins

10.21. Protein-protein interactions mediated by lipid in crystalline arrays of AAC. The crystal packing of AAC monomers allow them to interact via cardiolipin molecules (gray), as seen both from the side (A) and from the plane of the membrane (B). From Nury, H., et al., Annu Rev Biochem. 2006, 75:713-741. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

function and also show that protein-protein interactions are mediated by endogenous lipid (Figure 10.21). The oligomeric state of AAC in its native environment is still unclear. If dimers are functional in vivo, they could face opposite orientations, allowing them to be loaded simultaneously with an internal and an external nucleotide. Trapped in open conformations that resemble a heart, a trapezoid, and a basket, the three transporters described appear to have the common general mechanism of alternating access for a single binding site. Thus their functions necessitate quite large conformational changes to open to the other side of the membrane. While this mechanism distinguishes them from channels, mutations can stabilize them in an open conformation that allows passive diffusion, giving them channel characteristics.

CHANNELS In contrast to the flexibility of transporters, channels do not require large conformational changes to allow their substrates to cross the membrane, since they contain conduction pores. The passage of ions and molecules through the pores is not uncomplicated, however. The pore characteristics depend on the nature of the walls

Although predicted in the 1950s, water channels in biological membranes were not discovered until 1992, when a protein purified from the er)'throcyte membrane was reported to greatly increase the water flux in response to a gradient of osmotic pressure. Since the discover)' of this first AQP, over 350 different AQPs have been identified in all forms of life. Mammals have 11 isoforms, designated AQPO to AQPI0, that faIJ into two classes, AQPs and glyceroaquaporins. The latter group aIJows entry of glycerol and a few other small molecules in addition to water (see below). Four of the human AQPs (AQP3, AQP7, AQP9, and AQP10) are glyceroaquaporins. The physiological roles of the different human AQPs val)' widely because they differ in cellular locations as well as in modes of regulation. For example, in secretory glands, AQP5 is specifically expressed in the apical membrane where water passes into secretions such as tears, saliva, and sweat, while in respiratol)' epithelia of the lungs, different cell types express different AQPs (Figure 10.22). The role of AQPs in the lungs was explored in the interesting case of a few asymptomatic AQP I-null individuals, whose pulmonar)' capillaries swelled normally in response to saline but their surrounding tissue did not accu mulate fluid (Figure 10.23). In addition to the basic need for all cells to keep water in balance, the AQPs are involved in several illnesses, including abnormalities of kidney function, loss of vision, onset of brain edema, and starvation. Water channels are typically bidirectional, allowing influx and efflux of water molecules in response to changing osmotic conditions. They are extremely fast: water Hovvs through a single AQPI molecule at a rate of three billion molecules per second. And they are very selective, allowing the passage of water molecules without protons (or other ions). AQPs transport only

Transporters and Channels

254

structure arose from a gene duplication event in evolution. The highly conserved sequence motif NPA is repeated near the center of each half of the primary structure, and several other conserved residues, such as GJu 14 and Glu 152, are repeated near the beginning of each segment.

A.

AQP3 AQP5 Basement/ membrane

AQP4 H 20

Respiratory airspace

B.

Structure of Aquaporins Early images of AQP I in lipid bilayers obtained by both AFM and EM gave evidence for pores. The structure of AQP 1 from hu man red blood cells was determined by EM at a resolution of 3.8 Ain the same year that the crystal structure of GlpF, the glyceroaquaporin from E. coli, was reported at 2.2 A resolution. These structures A.

130 Goblet cell AQPI-null control

v 120

AQP4

>:::

(\$':'::::

Surface layer

v v '-

CIl

'fl

CIl

-~ V ~ ~ >:::

110

v v

>

~ v

8 JOO

Basal layer

90

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bsln

IL

2L

3L

Normal saline challenge B.

~':

150 AQPI

Capillary

10.22. Localizations of different human aquaporins. A. In secretory glands, AOP3 and AOP4 are found in the basement membrane, while AOP5 resides in the apical membrane where water passes into secretions. B. In epithelial tissues of the lungs, AOP4 is expressed in surface columnar cells, AOP3 in basal cells, and AOP1 in underlying fibroblasts and capillaries, while no AOPs are expressed in goblet cells. From Agre, P., Proc Am Thorac Soc. 2006, 3: 5-13. © 2006 by American Thoracic Society. Reprinted with permission from Proceedings of the American Thoracic Society.

CIl V

v

Control

140

~

.~

@~ 130 - ro c:3~ 120 :,

£'I

.-:::

110

~ 100

'-

v

8

.......

90 80'

b~ln

I~

2~

• AQPI-null

3~

Normal saline challenge water, while the related glyceroaquaporins conduct small organic molecues like glycerol, urea, DL-glyceraldehyde and glycine, in addition to water. The stereoselectivity of glyceroaquaporins is indicated by the J O-fold higher rates of transport of ribitol than of D-arabitol, both of which are simple five-carbon reduced sugars. The high-sequence homology among the AQPs suggests that they use a common architecture to accomplish selective watel- transport. Although they are fairly small (AQPI is only 28 kDa), the high similarity between their N- and C-terminal segments indicates that the

10.23. Decreased pulmonary vascular permeability in AOP1-null humans. Computed tomography scans of the lung before and after intravenous infusion of saline revealed differences in water permeability of lung tissue. AOP1-null individuals and normal controls received infusions of up to 3 L of physiologic saline, and images of their bronchioles and adjacent venules were recorded and quantified. In both groups, the vessel wall of the bronchiole became thickened due to the accumulation of fluid (Al; however, the surrounding area did not accumulate fluid in the AP01-null individuals (B) since they lack the AOP to secrete water. Redrawn from Agre, P., Proc Am Thorac Soc. 2006, 3:5-13. © 2006 by American Thoracic Society. Reprinted with permission from Proceedings of the American Thoracic Society.

Channels

255

Extracellular

Membrane

Intracellular N terminus 10.24. X-ray structure of AOP1. Each TM helix (colored differently) tilts about 30" off the bilayer normal. Two half-helices (HB and HE) form one of the seven TM segments. The helices are twisted into a right-handed bundle. From Fujiyoshi, Y, et al., Curr Opin Struct Bioi. 2002, 12:509-515. (() 2002 by Elsevier. Reprinted with permission from Elsevier.

were quickly followed by that of the bovine AQPI at 2.2 A and AqpZ, the E. coli aquaporin, at 2.5 A ITSO]Ution. More recently, the structures of sheep and bovine AQPO have become available. The structures all share a unique AQP fold that consists of six TM (X-helices with two additional half-helices, so called because they each span half the bilayer. The TM helices cross i.n a righthanded twist at a 30° tilt to form an hourglass-shaped pore, with the two half-helices meeting in the center of the membrane (Figure 10.24). Both Nand C termini are cytoplasmic, with the loop connecting the N- and C-terminal segments on the extracellular side and varying in length. The NPA signature sequence for the AQPs is repeated in each of the half-helices and makes the interface between them. Purified AQPs assemble into

tetramers both in the crystal lattice (Figure 10.25) and when reconstituted into lipid bilayers, but clearly each protein has a pore. Since there is no evidence for cooperation between subunits, the functional unit of AQP is a monomer. New insights into channel selectivity have come from details of the structure of GlpF, the first highresolution AQP structure.

Glyceroaquaporins: GlpF The GlpF pmtein provides a channel for passive diffusion of glycerol into E. coli. In the cytosol glyceml is immediately converted to G3P, ensuring that the inside concentration of glycerol is low, which drives its uptake. The topology diagram for GlpF shows the symmetry between the N- and C-terminal segments (residues 6-108 and 144-254), with similar TM segments on each side of the two half-helices (M3 and M7) (Figure 10.26). In three dimensions the two segments form two inverted halves of the channel and are linked by a protruding periplasmic region (Figure 10.27). The two half-helices form an important junction in the center of the membrane, held by van der Waals interactions between the proline residues of the NPA signature motifs. The NPA motifs cup each other between the pmline and alanine side chains of the opposite helix, with their orientation stabilized by other very highly conserved residues. In addition, conserved glycines allow close contact between the (X-helices where they cmss near the center of the bilayer, stabilized by CH-O hydmgen bonds. The channel in GlpF starts on the outer surface with a wide vestibule and then constricts to form a selective channel that is 28 A long. The crystal structure sho\vs three glycerol molecules in transit (see Figure 10.27). The narmwest point in the channel defines the selectivity filter, with a very close fit for glycerol that involves hydmphobic interactions at the corner between Trp48 and Phe200 and hydrogen bonds between the hydmxyl

10.25. Tetramers of AOP1. Each monomer contains a pore, and hydrophobic interactions between monomers stabilize the tetrameric assembly, which is viewed from the top (A) and the side (8), with each monomer colored differently. From DeGroot, B. L., and H. Grubmuller, Curr Opin Struct BioI. 2005,15:176-183. © 2005 by Elsevier. Reprinted with permission from Elsevier.

Transporters and Channels

256

~L ~

:.

.E©J76

© ®®®

®

CD®©@ A

Lumen 10.42. X-ray structures of SERCA1 in the presence and absence of calcium ions. The Ca 2+ -free form of the enzyme was crystallized bound to an inhibitor, thapsigargin (TG). In the structure of E12Ca 2 + (violet), the three cytoplasmic domains are splayed open, with the ATP-binding site available. The two cyan circles point out the location of bound Ca 2+ ions. The red arrows indicate the movements of the cytoplasmic domains that would produce the Ca 2+ -free form. In the absence of Ca 2+, the E2(TG) structure (green) shows the N, P, and A domains gathered into a compact headpiece. The black dashed line illustrates the angular movement of the N domain. The cyan arrow shows the proposed entry path for Ca 2+. The bilayer shown is an MD simulation of DOPe. From Toyoshima, e., et al., FEBS Lett. 2003, 555: 106-11 O. © 2003 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

A.

B. 10.43. Rearrangement of TM helices when SERCA 1 binds calcium. The TM helices of SERCA 1 (numbered) are superimposed in positions observed in E1·2Ca 2+ (violet) and E2(TG) (green). Both A and B are viewed from the side, with a 90° rotation in the viewpoints (such that the view in B is the back of the view in Figure 10.42). Helices M8 and M9 are removed in B to show the movements of M5, M2, and M4. The double circles (red and white) show pivot positions of M2 and MS. The red arrows indicate the direction of movements during the change from E1·2Ca 2+ to E2(TG), and the cyan arrow shows the proposed pathway for entry of the first Ca 2 + ion. From Toyoshima, e., et aI., Annu Rev Biochem. 2004, 73:269-292. © 2004 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

Channels

267

B.

A.

10.44. Close contact between TM helices M5 and M7 in SERCA 1. The GxxxG motif (green space-filling models) in M5 interacts with Gly770 (blue space-filling model) at the pivot point in M7. The view in B is rotated 90" from the view in A. From Lee, A. G., Biochim Biophys Acta 2002, 1565:246-266. © 2002 by Elsevier. Reprinted with permission from Elsevier.

The two Ca 2 + ions bind in a cooperative manner to sites located side by side in the TM domain (Figure 10.45). Site I is surrounded by Ms, M6, and M8 helices at the center of the TM domain, while site II is closer to the cytoplasmic surface on the M4 helix 'where M4 is partly unwound. Asp800 on an unwound part of M6 contributes to both sites. Most mutations of the specific residues involved in binding Ca 2 + at sites 1 and rr abolish binding, as expected. An entrance to the Ca2+ -binding sites located just below the phospholipid interface is suggested in the EJ -Ca 2+ crystal stmcture (Figure 10.46). For vectorial transport, Ihis site should be closed off upon phosphorylation; however, the intermediates EIP·ADP·2Ca2+ and E1P·2Ca2+ are short-lived. To see the occlusion of this site in phosphorylated intermediates, SERCA has been crystallized

10.45. The calcium-binding sites in SERCA 1. There are two high-affinity sites that bind Ca 2 + (blue spheres), each with seven oxygens from amino acid residues along with two waters (red spheres). From Toyoshima, C, et aI., Annu Rev Biochem. 2004, 73:269-292. © 2004 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

with analogs of Pi and ATP present. The structure of EI-AlF4"·ADP·2Ca2+ (in which AIF4" is a phosphate analog) reveals movements ofTM helices MI, which makes a kink, and M2, which is lifted up about 10 A. These changes close the proposed entrance gate to site TI. The structure of yet another form or SERCA, the E2AIF.j(TG) complex, gives information about the dephosphorylation step. The N domain is moved away from the P domain by a rotation due to the bending of MS. The interactions of AIF4" with Asp3S1 and the catalytic Glul83 are consistent with an SN2 mechanism for dephosphorylation. In the next step protons musl enter the structure, although they are not seen at this

A.

10.46. Entrance gateway for Ca 2 + in SERCA1. A. The region in E1·2Ca 2 + at M 1 (gold) and M2 (lavender) makes a gateway lined by Glu109, Glu55. Asp59, and Glu58 (behind Asp59 at this angle) leading directly to Glu309 and the Ca 2 +ion (green) at site II. B. The conformational change occurring with phosphorylation blocks this entrance pathway as indicated in this region of the structure of E1AIF4" ·ADP·2Ca 2 +. From M011er, J. et aI., CurrOpin Struct BioI. 2005, 15:387-393.~) 2005 by Elsevier. Reprinted with permission from Elsevier.

v.,

Transporters and Channels

268

E1·2Ca 2 +

.....

E1·AIF ol-' ADP

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~

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Lumen

Lx 10.47. Structural transitions during the transport cycle in SERCA 1. The conformational changes during the Ca2+ -ATPase transport cycle are shown with structures from the major stages of the E1-E2 cycle (shown in center). Motions of the headgroup domains are indicated by dashed arrows. Color changes gradually from the N terminus (blue) to the C terminus (red). Key residues, including R560, F487, E183, D351, and D703, are shown in ball-and-stick representation. When present, ATP, ADP, AIF4, and TG are shown in space-filling molecules and Ca 2 + is circled. From Inesi, G., et aI., Biochemistry. 2006, 4513769-13788. © 2006 by American Chemical Society. Adapted with permission from American Chemical Society.

For Further Reading resolution. Overall, this form of the enzyme has few structural differences from the E2(TG) structure except for the mobile A domain. Thus four structures of SERCA can portray most of its conformational changes during the EI-E2 reaction cycle to give remarkable insight into the function of this complex ion pump (Figure 10.47). Binding of two Ca 2+ ions opens the headpiece su fficiently for ATP to bind, which leads to large movements as the headpiece domains close. Bending of the A and P domains bring the ATP close to Asp351, allO\.ving transfer of the phosphoryl group to generate the phosphoenzyme intermediate. Another conformational change allows ADP to leave and presumably opens a channel for extrusion of the calcium ions into the lumen. Structural biology has come a long way toward elucidating the transport mechanism of the Ca 2+-ATPase and, by extension, other members of the family of Ptype ATPases, including the Na+K+-ATPases in animal cells and the H+ -ATPases in plant cells, which are both essential for the formation of the membrane potential. While the mechanism is more complex than those of the other transporters and channels described in this chapter, it is accomplished by a relatively simple protein structure when compared with the F 1 Fa-ATPase described in Chapter 11.

FOR FURTHER READING

269

Mitochondrial ADP/ATP Carrier Nury, H., et aI., Relations between structure and function of the mitochondrial ADP/ATP carrier. Armu Rev Biochem. 2006,75:713-741. 'Pebay-Peyroula, E., et al.. Structure of the mitochond.·ial ADP/ATP carrier in complex with carboxyatractvloside. Nature. 2003,426:39-44. Pebay-Peyroula, E., and G. Brandolin, Nucleotide exchange in mitochondria: insight at a molecular level. Curr Opin Slruct Bioi. 2004, 14:420-425. Aquaporins Agre, P.. Aquaporin water channels (Nobel lecture). Angew Chem Int Ed. 2004,43:4278-4290. DeGroot. B. 1.., and H. Grubmuller, The dynamics and energetics of water permeation and proton exclusion in aquapar·ins. Curl' Opin Struct Bioi. 2005, 15: 176-183. 'Fu, D. X., et aI., Structure of a glycerol-conducting channel and the basis for its selectivitv. Science. 2000,407:599605. Fujiyoshi, Y, et aI., Structure and function ot water channels. Curl' Opin Struct Bioi. 2002,12:509-515. 'Murata. K., et aI., Structural determinants of water permeation through aquaporin-l. Nature. 2000, 407:599605. Stroud. R. M., et aI., Glycerol facilitator GlpF and the associated aquaporin family of channels. Curl' Opin Struct Bioi. 2003, 12:424-431. 'Sui, H., et aJ.., Structural basis of water-specific transport through the AQP 1 water channel. Nature. 200 I, 414:872878.

LacY and GlpT 'Abramson, J., 1. Smirnova, V. Kasha, G. Verner, H. R. Kaback, and S. Iwata, Structure and mechanism of the lactose permease of Escherichia coli. Science. 2003, 301 :61 0615. Abramson, J., H. R. Kaback, and S. Iwata, Structtmll comparison of lactose permease and the glycerol-3-phosphate antiporter: members of the major facilitator superfamily. CurrOpin Struct Bioi. 2004,14:413-419. Guan, L., and H. R. Kaback, Lessons from lactose permease. Annu Rev Biophys Bi011101 Struct. 2006, 35:67-91. "Huang, Y, M. J. Lemieux, J. Song, M. Auer, and D.-N. Wang, Structure and mechanism of the glycerol-3-phosphate transporter from Escherichia coli. Science. 2003,301:616620. Lemiuex, M. J., Y Huang, and D.-N. Wang, The structural basis of substrate translocation by the Escherichia coli glycerol-3-phosphate tranSpOJ1er: a member of the major facilitatorsuperfamily. CurrOpinStruct Bioi. 2004,14:405412. Lemieux, M. J., Y. Huang, and D.-N. Wang, Glycerol-3phosphate transporter of Eschen:chia coli: structure, function and regulation. Res i\llicrobiol. 2004, 155:623-629.

• Paper presents original structure.

Potassium Channels 'Doyle, D. A., el aI., The structure of the potassium channel: molecular basis of K+ conduction and selectivity. Science. 1998,280:69-77. Gouaux, E., and R. MacKinnon, Principles of selective ion transport in channels and pumps. Science. 2005, 310: 14611465. •Jiang, Y, et aI., Crystal structure and mechanism of a calcium-gated potassium channel. Nature. 2002, 417:523526. •Jiang, Y.. et aI., X-ray structure of a voltage-dependent K+ channel. Nature. 2003,423:33-41. MacKinnon, R., Potassium channels and the atomic basis of selective ion conduction (Nobel lecture). Angew Chern Int Ed. 2004,43:4265-4277. Roux, B., Ion conduction and selectivity in K+ channels. Annu Rev Biophys Biomol Struct. 2005, 34: 153-171. Tombola, F, et aI., How does voltage open an ion channeP A,1I7U ReI' Cell Dev Bioi. 2006, 22:23-52. 'Zhou, Y. et aI., Chemistry of ion coordination and hydration revealed by a K+ channel-Fab complex at 2.0 A resolution. NalUre. 200 I, 414:43-48 .

Transporters and Channels

270

Calcium ATPase 2

Lee, A. G., Ca +-ATPase stmcture in the EI and E2 conformations: mechanism, helix:helix and helix:lipid interactions. Biochim Biophys Acta. 2002, 1565:246266. ·'Toyoshima, c., et aI., Crystal structur·e of the calcium pump of sarcoplasmic reticulum at 2.6 Aresolution. Nature. 2000, 405:647-655.

'Toyoshima, c., et aI., Structural changes in the calcium pump accompanying the dissociation of calcium. Nature. 2002, 418:605-611. Toyoshima, C., and G. Inesi, Structural basis of ion pumping by Ca 2 + -ATPase Ol the sarcolasmic reticulum. Al1rlU Rev Biochem. 2004, 73:269-292. Toyoshima, c., [on pumping by calcium ATPase of sar·coplasmic reticulum. Adv Exp Med Bioi. 2007, 592:295-303.

11

Membrane Protein Assemblies

Extracellular

OM

Assemblies oj membrane proteins can span two membranes, exempli ied 111 the dynamic coupling netween inner membrane transporters and a hannel-tlli1n€e1 ,KrOSS the periplasm and outer membrane or rug efflux In Gram negative bacteria TI15 model of the AcrABfTolC systeln, which can export drugs S ch as novobiocin from insidt> rhe cell and ampicillin from he penplasm, is composed 01 the x-ray s ucture of TolC modeled to be the open s ate (red], he x-ray structure 01 AcrB (green), and a ,'epresenta Ion of AcrA based on the x ray struc LJre of ItS close homolog, MexA (blue). rrom Eswaran. J., el ill, Curr Opin Struct Bioi. 2004, 14:741 747 7004 by Elsevi r Reprinted With permission from Elsevier

Periplasm

1M

Cytosol

Most of the membrane proteins described in the previous chapters can carry out their tasks without partners, although some form homo-oligomers and others are involved in transient interactions, for example, with signaling proteins. Because they can function on their own, their high-resolution structures reveal a great deal about their mechanisms. In contrast, many membrane proteins function in large complexes and can be understood only when the other protein components in these multi protein assemblies are characterized as well. This chapter describes structures of some multi-

component complexes that can be viewed as molecular machines, or nanomachines, in the membrane. These vary from large enzymes composed of many subunits, such as ATP synthase, to dimers of protomers* that each have many subunits, such as cytochrome-bel oxidase, to structures formed when separate proteins interact, A protomer is the minimal structure from which a larger structure is buill. Although the term prolomer can refer to monomers. it is used here to denote subunits which themselves contain subunits. Similarly, mullimers are the larger units resulting from the assembly of protomers.

271

272

11.1. ATP synthase molecules observed on inside-out vesicles of bovine heart mitochondria. The knob-on-stalk appearance of the ATP synthase is seen in negative staining EM. From Walker, J. E, Angew Chern Int Ed. 1998, 37:2308-2319. © 1998 by Gesellschaft Deutscher Chemiker. Used by permission of WileyVCH Verlag GmbH.

Membrane Protein Assemblies was recognized by the award of the 1997 Nobel Prize in Chemistry to Paul Boyer and John E. Walker for "the elucidation of the enzymatic mechanism underlying the synthesis of ATP." Further experimentation and structural biology have contributed to the present understanding of this marvelous mechanism. Unlike the calcium pump described in Chapter 10, which as a single polypeptide couples ion flux to the hydrolysis of ATP, lhe ATP synthase is a molecular machine with many different polypeptide components. With eight (prokaryotic) to 18 (mammalian) different subunits, the complex has a molecular mass of 550 to 650 kDa. The F 1 domain is the catalytic domain that carl-ies out the synthesis and hydrolysis of ATP. Forming the knobs in Figure 11.1, F 1 is extrinsic to the membrane, so it can be removed by mild treatmenls and function as a soluble ATPase. E. coli has the simplest F 1 domain, with the composition cx3f3,yb£. The F o domain is lypically composed of ab 2c(lo_12J, although some complexes have two different b subunits instead of a homodimer, and the number of c subunits varies in differenl

sometimes across more than one membrane as seen in the proteins involved in drug efflux in Gram-negative bacteria (Frontispiece).

F,Fo-ATPASE/ATP SYNTHASE Familiar fOJ'decades for its knob-on-a-stalk appearance, the FIFo-ATPase is named for its two major structural domains, F 1 and Fo (Figure 11.1). This fascinating complex is also called the ATP synthase because it couples the flow of protons across the membrane to the synthesis of ATP as well as its hydrolysis, depending on the direction of proton flux. Found in the plasma membrane of bacteria, the mitochondrial inner membrane in eukaryotes, and the chloroplasllhylakoid membrane in plants, the ATP synthase is the major producer of ATP in cells using either oxidative phosphorylation or photosynthesis to generate a proton motive fOl-ce (pmf, the proton electrochemical gradient across the membrane that stores energy). The importance of ATP generation is clear to humans, who each use around 40 kg of ATP each day of a sedentary life (an athlete uses much more!). With around 100 mmol in the pool of adenosyl nucleotides in the body, this rate of consumption means thal each molecule of ADP must be phosphorylated about a thousand times a day to provide the needed ATP. The reverse reaction of the F 1 Fo-ATPase is also important under conditions when the utilization of ATP is needed to replenish the proton gradienl. How lhe flow of protons across the membrane is coupled to the catalylic reaction is a question thal is fundamental to life, Biochemical data on the catalytic mechanism of the F 1 Fo-ATPase gained beautiful support from the first high-resolution structures of its components. This

11.2. Overall structures of the F1 Fo-ATPase from E. coli. The arrangement of the subunits making up Fo and F, is diagrammed with one ex subunit removed to reveal the y subunit down the center. The three pairs of exf3 subunits of F, (magenta/pink, dark blue/light blue, and green; the third ex subunit is removed) surround the y (red) of the stalk, with [ (yellow) at its base and & (orange) along the back. The c subunit ring (black) of Fo is linked to the central stalk (yt:) as well as to the peripheral stalk composed of b2 (green) and b. The five predicted TM helices of the a subunit are represented (gold). Note: the bows show some of the crosslinks that either have little or no effect on (green) or inhibit (red) the activity, which demonstrated that c, y, and [ rotate together. From Capaldi, R. A., and R. Aggeler, Trends Biochem Sci. 2002, 27: 154-160. © 2002 by Elsevier. Reprinted with permission from Elsevier.

F, Fa-ATPase/ATP Synthase

sources and even under different growth conditions. When the F I domains are stripped off, Fa by itself forms a passive proton pore. The Fa domain is sensitive to the inhibitor oligomycin, for which it was originally named. In the complex, these two structural domains are connected by two stalks: a central stalk made up of y and £ subunits (yb£ in mitochondria) and a peripheral stalk made up of the band b subunits (Figure 11.2). The additional subunits in mammalian F I Fa-ATPases are mostly in the stalk regions (see Table I 1.1). The oligomycin-sensitivity conferring protein (OSCP subunit) in the mitochondrial peripheral stalk is equivalent to the bacterial D subunit. The b subunit in mitochondria has the role of the £ subunit in bacteria, and the small mitochondrial £ subunit (only 50 amino acids) at the foot of the central stalk has no counterpart in bacteria or chloroplasts. The overall shape of F I assembled with the c subunits of Fa is shown beautifully in the low-resolution images of the yeast mitochondrial FI-c complex published in 1999 (Figure 11.3). Subunit Structure and Function

F, Domain In all F I Fo-ATPases, the F I domain has alternating <X and j3 subunits forming a spherical hexamer, like sections of an orange, 'with a cavity in the center (Figure 11.4). The <X and j3 subunits of E. coli have 20% sequence identity and very similar folds. Both can bind nucleotides, but the catalytic sites are located on the j3 subunits at the <x/j3 interfaces. As described below, the three active sites are in three different conformations,

273

TABLE 11.1. Equivalent subunits in ATP synthases in bacteria, chloroplasts, and bovine mitochondria Type

Bacteria

Chloroplasts

Mitochondria a

F,

ex

ex

ex

(3

(3

(3

y

y

6

6

y OSCpb

€

€

6

a b c

a (or x) band b' (or I and II) c (or III)

a (or ATPase 6) b c

€

Fa

, Mitochondrial ATP synthase has additional subunits that have no equivalents in bacteria or chloroplasts. b Oligomycin-sensitivity conferring protein. From Walker, J. E, ATP synthesis by rotary catalysis (Nobel lecture). Angew Chem Int Ed. 1998,37:2308-2319.

depending on whether they bind ATP (the loosebinding TP conformation, partly open), or ADP (the tight-binding DP conformation, closed), or are empty (the E conformation, open). The y subunit has very long N- and C-terminal helices, 'which form an antiparallel <x-helical coiled coil that traverses the central cavity of the <Xj3 hexamer and continues as the central stalk to the Fa domain. The two long helical domains of yare connected by a globular domain that protrudes at the base of the stalk. The globular domain of y, visible in the higher-resolution structure of the complete bovine mitochondrial F,-ATPase obtained in 2000, has an <X!j3 structure that wraps around the other two helices

11.3. Low-resolution x-ray structure of the F, Fa-ATPase from yeast mitochondria. The electron density map of the F,-ClO complex from Saccharomyces cerevisiae at 3.9 A resolution shows the architecture of the complex from the side (A) and the cytoplasmic end (B), with insets identifying the subunits in A and numbering the c subunits in B. Note the asymmetry in B. From Stock, D., et al., Curr Opin Struct Bioi. 2000, 10:672-679. © 2000 by Elsevier. Reprinted with permission from Elsevier.

Membrane Protein Assemblies

274

A.

B.

11.4. The high-resolution structures of the ex, 13, and y subunits of the F,-ATPase from bovine mitochondria. The ribbon diagrams show ex (red), 13 (yellow), and y (blue) subunits as identified in the schematic drawing accompanying each figure. A. The entire F, particle shows the coiled coil of the y subunit in the central cavity between the ex and 13 subunits, with bound nucleotides (black ball-and-stick representation). The exl3 pairs are labeled E for empty, TP for binding ATP, and DP for binding ADP, as described in the text. B. A cutaway view showing only three subunits: CXTP, y, and I3DP (shaded in the diagram above the structure). From Walker, J. E., Angew Chern tnt Ed. 1998,37:2308-2319. © 1998 by Gesellschah Deutscher Chemiker. Used by permission of Wiley-VCH Verlag GmbH.

and contacts 8 and [ subunits (Figure 11.5). The position of the y subunit is asymmetric with respect to the cxl3 hexamer, which is important in the mechanism. Rotation of y, which unwinds the lower part of the coiled coil (see Figure 11.6A), is driven in one direction by hydrolysis of ATP and is driven in the opposite direction by the pmE. At the base of the y subunit where it contacts Fa is the [ subunit (8 in mitochondria), which has two domains, an N-terminal l3-sandwich and a helix-turnhelix C terminal. The N-terminal domain of [ makes contact with Fa and is essential for coupling. The [ subunit appears in very different positions in the E. coli and mitochondrial structures (Figure 11.6), and it also exhibits different conformations when crystallized as a pure protein and in complex with a portion of the y subunit. The closed conformation observed in isolated [ is similar to the conformation seen in the mitochondrial structure, but the open, partially unfolded conformation observed in complex with y is unlike that seen previously and suggests considerable flexibility in the structure (Figure 11.7). Cross-linking studies pro-

vide evidence for interactions between [ and 13 subunits that differdependingon which nucleotides are bound to F I , suggesting the flexible [subunit might have a regulatory function (see below). Data from cross-linking studies indicate that [ rotates with y and c, because both ATP synthesis and ATP hydrolysis coupled to proton pumping occur \",hen all three are covalently linked (see Figure 1J .2). The 8 subunit is required to bind F, to Fa, hence its historic designation in mitochondria as the oligomycin sensitivity-conferring protein. With the b 2 dimer from Fa ,8 forms the peripheral stalk that appears to hold the CXJ I3J knob in place while y rotates. For this reason b 2 8 has been called a stator, the stationary part of a machine in which a rotor revolves. As will be seen below, the rotor includes the central stalk and the ring of c subunits in the Fa domain of the enzyme.

Fa Domain The Fa structural domain consists of a ring of c subunits connected via the a subunit to the b dimer that forms the peripheral stalk. NMR studies indicate that

F, Fo-ATPase/ATP Synthase

275

11.5. The complete structure of the F,-ATPase from bovine mitochondria. The structure of the central stalk is visible in the x-ray structure of the complete F, domain obtained at 2.4 A resolution. A. The ribbon diagram shows the C( and 13 subunits (red and yellow, respectively) alternating around the y subunit (blue) with the y (green) and £ (magenta) at the base of the central stalk. B. Model of the mitochondrial ATP synthase based on EM data for bovine and yeast structures. Mitochondrial subunit names are different from those in the E. coli structure (see Table 11.1). In particular, the IS subunit is equivalent to the £. subunit in E. coli, and d along with six copies of F interact with the b subunits. From Stock, D., et al., Curr Opin Struct BioI. 2000, 10:672-679. © 2000 by Elsevier. Reprinted with permission from Elsevier.

the sm"l1 (8000 Da) c subunit forms two hydrophobic TM helices that make a coiled coil connected by a short polar loop. The number of c subunits varies, with J 2 in E. coli, 14 in chloroplasts, and 10 in bovine mitochondria. The c subunits form two concentric rings, with their N-terminal helices forming the inner ring and their C-terminal helices forming the outer ring (see Figure 11.3). The ring of c subunits is contacted by [ and y at the central stalk from F I , and half the polar loops contact the b subunit. An essential residue involved in proton uptake through Fo is cAsp61 (E. coli numbering) in the center of the C-terminal helix. When Asp61 binds " proton, the C-termin,,1 helix undergoes a conformational change that is critical to rotation of the ring. The very hydrophobic a subunit, predicted to form five TM helices, connects the ring of c subunits to the b subunit of the st"tor. A critical ,-esidue in the a subunit is Arg21 0, which is essential for ATP-driven proton translocation but not for passive proton transport when F I has been removed. The interaction of aArg210 with cAsp61 indicates there is a direct connection between the a subunit and the ring of c subunits. Therefore sub-

unit a is thought to contribute to the proton channel, with protons entering the translocation pathway either at an interface between the a and c subunits or via the a subunit, foJJowed by transfer to the c subunit. The a subunit has other basic and acidic residues essential for proton translocation, which are His245, GJu196, and Glu219. The b subunit is usually a homodimer, although some bacterial species have a heterodimer of band b' instead. It is very elongated and anchored in the membrane by its hydrophobic N-terminal region. The rest of the protein is hydrophilic and highly charged, except for a short stretch of hydrophobic amino acids near the C terminus. The C terminus is required for assembly of the ATP synthase. According to its circular dichroism spectra, the b subunit is highly helical. In agreement with this result, NMR data indicate the N terminus is helic"l, with residues 4 to 22 predicted as a TM segment. The two b subunits are in such close proximity that they are linked by disulfide bonds when cysteine residues are engineered at many different positions in the b subunit.

Membrane Protein Assemblies

276

A.

C

B.

11.6. Different conformations observed in mitochondrial and bacterial structures of the central stalk. The arrangement of the y (red) N-terminal helix (thicker), the C-terminal helix (thinner), and the [ subunit (yellow) varies between the mitochondrial ATPase (A) and the E. coli enzyme (B). The blue arrow shows the large movement of c involving rotation of 810 and translation of 23 A. From Capaldi, R. A., and R. Aggeler, Trends Biochem Sci. 2002, 27: 154-160. © 2002 by Elsevier. Reprinted with permission from Elsevier.

A.

Regulation of the F 1 Fo-ATPase

The FIFo-ATPase is a highly efficient motol~ Attempts to determine its thermodynamic efficiency (the ratio of free energy gained by pumping protons to the free energy expended in the phosphorylation of ADP to make ATP) from the measured torque of the F 1 domain suggest an efficiency near 100%. What then is the purpose of regulation? Cells would benefit by alteration of the efficiency of F I Fa-ATPase because if ATP concentrations in respiring cells are high, decreasing the net rate of ATP synthesis is beneficial. whereas if ADP concentrations are high, the cell needs rapid ATP synthesis. Crosslinking studies suggest that regulation of the activity of FIFo-ATPase could (partly) depend on the flexibility of the £ subunit described above (see Figure 11.6). The rate of cross-linking between £ and f3 subunits is affected by ATP/ADP concentrations: in the presence of ATP this rate increases, whereas in the presence of ADP it decreases. Thus the position of the £ subunit varies depending on the needed net rate of ATP synthesis. Recent studies show that £ is able to sample alternate conformations on a timescale of seconds. Do the conformations themselves suggest a possible mechanism? The conformation of the £ subunit seen in the E. coli structure allows its C-terminal helices to wind around the y subunit and extend toward the <X3f33 hexamer, with the central axis of the f3-sandwich roughly parallel to the N- and C-terminal helices of the y subunit. The conformation observed in the mitochondrial

B.

11.7. Conformational changes in the £ subunit. A. The closed conformation observed for the isolated £ subunit of E. coli is docked onto a model of the c subunit ring (below) and the end of the y subunit above. B. The open conformation observed when [ is crystallized with a portion of the y subunit has the C-terminal helices separated from each other as well as from the ~-sandwich and wrapped around portions of y. From Bulygin, V. v., et al., J Bioi Chem. 2004, 279:35616-35621. © 2004 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

F, Fa-ATPase/ATP Synthase

277

ATP

l

ADP + Pi

T

1

11.8. Sequential stages in the binding-change mechanism. Each catalytic site of the ATP synthase can have one of three conformations, designated 0 for open (empty) state, T for tight (ATP-bound) state, and L for loose (ADP + P; occupied) state. The figure shows one iteration through the states, which is coupled to a 120" rotation of the yc central stalk. This is also called an alternating sites hypothesis as it involves cycling of each of the three catalytic sites through three states. From Capaldi, R. A., and R. Aggeler, Trends Biochem Sci. 2002,27: 154-160. © 2002 by Elsevier. Reprinted with permission from Elsevier.

structure has shifted the central axis of the l3-barrel to be roughly perpendicular to the N- and C-terminal helices of the y subunit, ,vith the C-terminal helices flattened against the side of the l3-sandwich and away from Fl. These two states have been proposed to work as a rachet to affect the catalytic efficiency of F I , and this is supported by cross-linking studies showing different results when nucleotides are bound to Fl. Cross-linking studies also show that diFferential effects on hydrolysis and synthesis activities are possible. For example, formation of a cross-link between the C terminus of subunit £ and they subunit inhibited ATP hydrolysis by 75% and ATP synthesis by 25%, indicating that in some conformations, ATP synthesis is allowed when ATP hydrolysis is not. Finally, cross-linking results indicate that subunit £ can span the region of the central stalk and interact simultaneously with a 13 subunit of F, and subunit c of Fa. Catalytic Mechanism of a Rotary Motor

Kinetic studies of the F, Fa-ATPase established a number of characteristics of its mechanism: (1) Isotope exchange data are consistent with the idea that the enzyme does not release ATP at one active site until substrate is available to bind at another active site. (2) The exchange of '80 showed that all three catalytic sites on the three 13 subunits are equally capable of carrying out the reaction. (3) Because release of products is rate Ii 01iting, the Pi and ADP that remain bound can reversibly resynthesize ATP, resulting in the incorporation of 18 0 in different positions when the reaction is carried out in H 2 '80. (4) The catalytic sites on each of the 13 subunits in F 1 differ in affinity for ATP (when measured in excess ATP so ATP binds to all three sites): Kd for the first is < 1 nM, for the second is ~ 1 ~lM, and for the third is 30 IJ.M. Clearly, substrate binding shows negative cooperativity. However, when more than one site is occupied, the rate of ATP hydrolysis goes up 104 _ to lOS-fold, so catalysis shows positive cooperativity. Whether this cooperativ-

ity requires nucleotide binding to two or three sites is still controversial. Alternating Site Mechanism The kinetic results are all consistent with the Boyer binding-change mechanism that states that each of the three binding sites in F, is in a different conformation, either closed (tight), partly open (loose), or open (and empty; Figure J 1.8). The site in the open conformation is ready to bind substrate. When it binds nucleotide it closes, triggering conformational changes in the other two so that the closed one becomes partly open and the partly open one becomes fully open. In the mechanism, each site alternates between the three states in a cycle for both forward and reverse reactions (see Figure 11.8). The reaction for ATP synthesis involves (J) binding of ADP and Pi to the partly open site; (2) conformational change at that site that converts it to the closed site, where catalysis occurs (while changing the other two sites to the open and loose sites); and (3) a second conformational change to convert that site to the open site that allows dissociation of the product. The rotating y subunit interacts with one of the 13 subunits to drive its conformational change from closed (tight) to open (empty), which in tum triggers the conformational changes in the other two subunits. In the full F I Fa-ATPase, these conformational changes in F 1 are coupled to proton translocation through Fa. For ATP synthesis, the rotation is generated by the passage of protons, driven by the pmf. through Fa to cross the membrane; in the reverse direction, energy released by hydrolysis of ATP drives the rotation in the opposite direction and reverses the flow of protons (Figure 11.9). In this rotary motor, the central rotor formed by the y and £ subunits of F I rotates by 120 for each ATP synthesized or hydrolyzed, bringing subunit y into contact with a different 13 subunit with each rotation. The central stalk connects to the ring of c subunits and rotates with them to couple catalysis to proton transport. 0

278

Membrane Protein Assemblies

~

2

3

e;/'~

~0">

\0'

'i:''\.~

.

Co

Catalytic £1wpll

S')

/ \ rotation

....

4

5

-- /

\

ATP

,,''0,.

i~

u;.\

~.~..

i

\-"

2Q

cylci

'i:::~ Q-sile I

\

--

QH:,mb""'

'. ·······:::::::.r.- anlll1lycl/1 _Q

~

)~--2H+

N·phase

11.13. The mechanism of the 0 cycle illustrated on a functional unit of the chicken cytochrome-bcl complex. The cytochromeb (light green) and cytochrome-cl (yellow) subunits are shown with transparent surface representation, while the structure of the Rieske Fe/5 protein is represented by ribbon diagrams showing the two conformations (blue and red). The position of the membrane is indicated by dashed lines, with the intermembrane space labeled P-phase (for positive) and the matrix labeled N-phase (for negative). The [2Fe-25] center is represented by space-filling models in four positions, two showing the two conformations of the Rieske protein (blue and red) with two intermediate positions (yellow) to show the trajectory of its movement. The b-type hemes (blue) are near the quinone-binding sites, and the c-type heme (black) is not far from the docking site for cytochrome c. A quinone (green) binds to 0" and a quinone has been modeled to replace stigmatellin at 00 at the inhibitor binding sites (dotted light blue arrows). Electron transfers (small green arrows) and proton release and uptake (curved blue arrows) are indicated. In the reaction summarized in the text, two quinol molecules are oxidized to quinone at the 00 site, with two electron pairs following the divergent pathways (green arrows) to the [2Fe25] center, cytochrome C1 and cytochrome c and to the b·type hemes to reduce the quinone at the OJ site. From Crofts, A. R., Rev Physiol. 2004, 66:689-733. © 2004 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

b-type hemes, called b H and b L , and cytochrome CI has a c-type heme (a-, b-, and c-type hemes differ in their substituents off the tetrapyrrole ring and their linkages to the proteins). In addition to the two cytochromes, the catalytic unit has an iron-sulfur protein of the Rieske type, which is a [2Fe-2S] cluster in which one Fe is coordinated by two histidine residues and the other by two cysteine residues. The redox sites in these three subunits are buried in the membrane interior (see Figure 11.12B) because their quinone substrates are very hydrophobic, with isoprenoid side chains 30 to 50 carbons in length. In addi tion, redox reactions of quinones involve a semiquinone intermediate that needs to be shielded from the aqueous environment to avoid formation of damaging reactive oxygen species.

The Q Cycle Unlike other proteins that pump protons, such as bacteriorhodopsin (see Chapter 5) or cytochrome oxidase (see next section), the cytochrome-bcl complex does not provide a direct proton path across the membrane. In the overall reaction, two electrons from ubiquinol reduce two molecules of cytochrome C with uptake of two protons from the matrix (or cytosol in bacteria) and release of four protons to the intermembrane space (or outside in bacteria). This is accomplished by a special mechanism called the Q cycle, which uses two quinonebinding sites. The Qi site is closer to the inner surface and with heme b H forms the N center (for negative side), and the Qo site is closer to the outer surface and with heme b L forms the P center (for positive side). Quinone reduction takes place at Qj, quinol oxidation takes place at Qo , protons are taken up at Q i and exit from Qo , and electrons are cycled between quinoJ and the hemes before delivery to cytochrome c at the outer surface (Figure 11.13). In brief, the first ubiquinol binds to the P center and is oxidized, with its two electrons taking divergent paths. One electron is transferred to the Rieske Fe/S center, and [Tom there to cytochrome c\ and then to cytochrome c. The other is transferred to the b L heme, and from it to the b H heme and then to a ubiquinone that binds to the Qi site at the N center, making ubisemiquinone. A second ubiquinol binds to the P center, and again the two electrons follow these different paths, resulting in the reduction of a second cytochrome c and the reduction of the semiquinone to ubiquinol at the N center. Each oxidation of ubiquinol releases two protons, and two protons are taken up from the matrix for reduction of ubiquinone at the N center. Evidence for the Q cycle includes the ability of the inhibitor antimycin, which binds to the Q i site, to inhibit oxidation of heme b H and stop all electron transfer to cytochrome c. High-Resolution Structures The x-ray structures of bovine, chicken, and yeast cytochrome-bcl complexes all show a symmetric, pearshaped dimer that protrudes from the membrane in both directions, ~75 A into the matrix on the inside and ~35 A into the intramembrane space on the outside (see Figure 11.12). The position of the lipid bilayer is clear because of the bound phospholipids (see Chapter 8). More than half the mass of the complex is in the matrix portion, including the misnamed subunits Core I and Core 2 that function in mitochondrial peptide processing. Each protomer has eight TM helices from cytochrome b and one each from cytochrome CJ and the Fe/S protein, along with two or three additional TM helices from small subunits that surround the functiona I unit (Figure 11.14). The Fe/S protein has a hinge between an extrinsic domnin nnd the TM domain containing the metal center, allowing the [2Fe-2S] center

Membrane Protein Assemblies

282

p'

H"

QCRS·"

A.

~B~i

F 11.14. TM helices of cytochrome-bcl homodimer. Segments of one subunit are designated with asterisks. The TM helices from cytochrome b (red) are labeled A to H, those from cytochrome c, (yellow) are labeled CYT1, and those from the Rieske Fe/S protein (green) are labeled RIP1. The additional TM helices from small subunits OCR8 and 9 (blue and gray) are also shown. The dimer interface has two large hydrophobic clefts, labeled CFT. The hemes and the headgroups of the quinone (behind helix A) and stigmatellin (between B and C) are shown in ball-and-stick models. From Hunte, C, et aI., Structure. 2000, 8:669-684. @ 2000 by Elsevier. Reprinted with permission from Elsevier.

to be shuttled bet,,veen cytochrome b and cytochrome c (see Figure 11.13). The conformational change around this hinge rotates the extrinsic domain 60" and moves the [2Fe-2S] cluster 16 A. Mutations that limit the hinge movement lower the activity of the complex. The crystallization of the yeast cytochrome-bel complex in tbe presence of substrate and inhibitors has provided insight into the mechanisms of electron transfer and proton conduction. In the crystal structures one molecule of ubiquinone is bound to Q; at the N center. At the Q o site, the inhibitor stigmatellin binds in the same position that ubiquinone is expected to bind, while the inhibitor 5-n-heptyl-6-hydroxy-4,7-dioxobenzothiazole (HHDBT) is a hydroxyquinone anion that resembles an intelmediate step of ubiquinol oxidation (Figure 11.15). Critical residues at the P center include Hisl81 on the Rieske Fe/S protein and Glu272 of cytochrome b. In the first step of the Q cycle ubiquinol is hydrogen bonded to both of these residues to form an electron donor complex, which allows essentially simultaneous electron transfer to the Rieske cluster and the b L heme. Oxidation of ubiquinol to ubiquinone breaks this complex, allowing the ubiquinone to diffuse out to the medium or possibly to the N center of the opposite protomer. 1n addition, formation of the complex increases the pKa on the imidazole nitrogen of Hisl81, allowing it to take up a proton from ubiquinol; when the complex dissipates and the Rieske Fe/S center moves away, the pKa is lowered and the proton is released. The second proton [Tom the ubiquinol is transferred to Glu272 after electron transfer to the b L heme (Figure 11.16).

B.

11.15. Binding of different inhibitors to the 0 0 site of yeast cytochrome bc,. A. Electron density fitted to the structure of stig· matellin at the 0 0 binding site between Glu272 and the 12Fe-2SI center of the Rieske protein coordinated by His181 and His161. Hydrogen bonds are shown to the carbonyl oxygen (04) and hydroxyl group (08) of stigmatellin. From Hunte, C, et aI., Structure. 2000, 8:669-684. © 2000 by Elsevier. Reprinted with permission from Elsevier. B. Electron density fitted to the structure of the inhibitor 5-n-heptyl-6-hydroxy-4, 7-dioxobenzothiazole (HHDBT), whose structure is given in the inset. Residues that stabilize the binding are labeled, and hydrogen bonds to the carbonyl oxygen (04) and the deprotonated hydroxyl oxygen (06) are shown. From Palsdottir, H., et aI., j Bioi Chem. 2003, 278:31303-31311. © 2003 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

Complexes of the Respiratory Chain

283

11.16. A model of electron and proton transfer at the P center of the yeast cytochrome-bcl complex viewed from the membrane interior A. Ubiquinol is hydrogen bonded to His 181 of the Rieske Fe/S protein (residues 160-175 and 178-182 are represented as yellow strands) and to Glu272 of cytochrome b (residues 75-85 and 265-275 are represented as cyan ribbons) with the lengths of the hydrogen bonds given in angstroms. The side chains of His181, Glu272, and Arg79 are shown as stick models (carbon, green; oxygen, red; nitrogen, blue), along with the ubiquinol and the b L heme. This conformation is observed in stigmatellin-bound cytochrome bCI, with ubiquinol modeled to replace stigmatellin. B. The structure in cytochrome bC1 complexed with the inhibitor HHDBT, a hydroxyquinone anion inhibitor that resembles an intermediate step of ubiquinol oxidation, shows movement of the Glu272 toward the b L heme. From Hunte, c., et aI., FEB$ Lett. 2003, 54539-46.

11.17. The suggested pathways for proton uptake at the N center of the yeast cytochrome-bcl complex. Two distinct pathways connect the solvent at the matrix side with the quinone-binding pocket at the N center. The E/R pathway (right side of figure) has an entrance at Glu52 of the Ocr7 subunit and is gated by Arg218 (yellow side chain with blue nitrogen atoms). The cardiolipin/K pathway has a cardiolipin (CL, green) positioned at its entrance and is gated by Lys228 (yellow side chain with blue nitrogen atom). Water molecules (red spheres) are associated with charged residues, and hydrogen bond interactions (dotted and dashed lines) are indicated. The arrows indicate the access sites from the bulk solvent, and double-headed arrows indicate proton transfer between the residues and the ubiquinone (U06, cyan). From Hunte, c., et al., FEBS Lett. 2003, 545:39-46. © 2003 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

Membrane Protein Assemblies

284 cyt. c

N-side

4W

I I

4W

Iran~localion ~ub~tratc

11.18. Schematic diagram showing the paths of proton and electron transfer in cytochrome-c oxidase. The chemical reaction of 02 reduction to water (blue arrows) is coupled with the translocation of four protons (red arrows). Electrons flow from reduced cytochrome c in the intermembrane space (P-side) to the CUA center, and from there to heme a and then to the heme a3-CuB center, where they reduce oxygen. Protons from the matrix are either shuttled to the heme a3-CuB site and consumed in the production of water (substrate protons) or are translocated across the membrane. From Wikstrom, M., Biochim Biophys Acta. 2004, 1655:241-247. © 2004 by Elsevier. Reprinted with permission from Elsevier.

Rotation of protonated Glu272 allows it to hydrogen bond with a water molecule that is hydrogen bonded to a heme propionate side chain. From there the proton is released to the surface via a hydrogen-bonded water chain associated with four charged residues of cytochrome b, Arg79, Asn256, Glu66, and Arg70. While protons are released from the P center of cytochrome bCI, they are taken up at the N centel~ The x-ray structure of the yeast complex reveals two distinct pathways for proton uptake, called the E/R pathway and the cardiolipin/K pathway (Figure J 1.17). The two pathways are located at either end of the substrate-binding site, indicating a quinone can be reduced on both ends of the molecule without changing positions. The E/R path runs from Glu52 of one of the minor subunits and is gated by Arg218 of cytochrome b. A molecule of cardiolipin is at the entrance of the other path, which is gated by Lys228 of cytochrome b. Upon reduction of ubiquinone, a proton is abstracted from each of the gating residues (Arg218 and Lys228) and is replenished with a proton [Tom the malLix via hydrogen-bonded networks to those residues. The structures have provided an elegant explanation for the coupling of proton movements to electron transfer.

Cytochrome-c Oxidase Complex IV of the respiratory chain, cytochrome-c oxidase, carries out the reduction of O 2 to H 2 0 using four electrons coming from four molecules of reduced

B.

11.19. x-ray structure of cytochrome-c oxidase. The crystal structure of cytochrome-c oxidase from Rb. sphaeroides was determined at 2.3 A resolution. The subunits are closely packed: subunit I (green), subunit II (light gray), subunit III (dark gray), and subunit IV (magenta). The redox centers include heme a (light blue), heme a3 (red), and CUA and CUB (dark blue), and have Mg (light green), calcium (pink). lipids (orange). and water molecules (red spheres) A. Ribbon diagram shows the structure viewed from the side. B. The TM helices in cytochrome-c oxidase are viewed from the P-side. From Svensson-Ek, M., et aI., J Mol BioI. 2002, 321 ;329-339. © 2002 by Elsevier. Reprinted with permission from Elsevier.

Complexes of the Respiratory Chain

285

cytochrome c while pumping a total of eight protons. four "substrate" protons that combine with the oxygen atoms and four "pumped" or "vectorial" protons that contribute to the electrochemical gradient. Like complex III. complex IV is a dimeric multimer with the prokaryotic protomercontaining three to foursubunits, while in eukaryotes it has eight to 13 subunits. AJI the redox centers are contained in subunits I and II: two a-type hemes called a and a3. and two Cu-containing centers, CUA, which has two copper ions. and CUB' In addition, the complex has a Mg 2+ ion that is not involved in redox and a site for binding Ca 2+ or Na+. Spectroscopic studies revealed the path of electron transfer is from cytochrome c to CUA to heme a, then to a binuclear complex of heme a3 and CUB. and finally to O 2 (Figure 11.18). The four-electron reduction of O 2 must occur by accumulating reduced intermediates without their release as reactive oxygen species. High-Resolution Structures

Crystal structures have been obtained for cytochrome-c oxidase [Tom Paracoccus denitrifrcans, from Rhodobactersphaeroides, and from bovine mitochondria. In these stmctures. both bacterial complexes have four subunits and the bovine complex has 13 subunits, of which subunits I. If, and III are very similar to the corresponding subunits in the other two. The dimeric complex has an ellipsoid shape that protrudes beyond the lipid bilayer 32 A into the intermembrane space and 37 A into the matrix (Figure I \.19). The proteins cross the membrane with 21 to 28 TM helices: 12 [Tom subunit I. two h-om subunit II. and seven from subunit III. The seven additional subunits in the bovine complex are type U membrane proteins (each with a single TM helix and the N terminal inside) thaI together surround subunits I. II. and HT. Six phospholipid molecules are resolved in the yeast structure. Both heme a and the bimetallic heme a3/CuB center are buried in the membrane interior of subunit I, ~13 A from the outer surface. The CUA center has two cystei ne residues as ligands to the two copper ions (analogous to [2Fe-2SJ) and binds to a globular domain of subunit II on the outside surface. This globular domain meets a corner of subunit I on the surface where cytochrome c likely binds, as there are 10 acidic residues that can interact with the ring of lysine residues on cytochrome c (see Figure 4.2).

11.20. Proton uptake pathways in yeast cytochrome-c oxidase. The ribbon diagram shows subunits I, II, and III with heme a and a3 (green) as stick structures, with Ca and Mg metals (green spheres) and Cu metals (orange spheres). The two paths for water molecules are the D path (red) and the K path (blue). with water molecules colored accordingly. From Hosler, J. P., et al.. Annu Rev Biochem. 2006, 75:165-187. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

[Tom time-resolved spectroscopic studies. indicate that O 2 binds first to the Fe 2 + of heme a3 and quickly picks up two electrons from the Fe 2+ and the CUB 1+ to form a peroxide bridge between Fe 3+ and CUB 2+. The next intermediate is an unstable hydroperoxo compound - Fe 3+OOH - that is quickly cleaved to the oxoferro state Fe4+ = 0 with CUB 2+ -OH-. By now, four electrons have been transferred. three from the bimetal center and the fourth probably from Tyr288, creating a tyrosyl radical. The next two steps are slower, limited by the time it takes the protons to reach the binuclear center. A proton reacts with the OH- on CUB to release the first water; then two electrons and two protons react with the 0 2 -- on Fe 4 + of heme a to release the second water. The electrons are supplied by additional molecules of cytochrome c operating through CUA and heme a.

Oxygen Reduction

The reaction can be viewed as starting with a metal reduction phase, when each cytochrome c molecule docks on the surface of cytochrome-c oxidase and transfers its electron to CUA. The electrons are then transferred to heme a and then to heme a3/CuB, where the reduction of O 2 takes place. The characteristics of the O2 reduction site with heme a3 and CUB have been determined in crystals of both fully oxidized and fully reduced enzyme. These structures, along with results

Proton Pathways

Protons from the matrix utilize two uptake pathways to the bimetallic center that have been identified in the structures of cytochrome-c oxidase. As observed in bacteriorhodopsin (see Chapter 5), a proton pathway consists of a series of hydrogen-bonded water molecules linked to residues that are essential for pumping protons. The D pathway goes from Asp132 on the surface to Glu286 between heme a and heme a3, a distance of

Membrane Protein Assemblies

286

Mg.

11.21. Molecular dynamics simulation of a proton exit pathway in yeast cytochrome-c oxidase. The simulaton of the Rb. sphaeroides cytochrome-c oxidase structure in added water lasted over a nanosecond and revealed a chain of hydrogenbonded water molecules from Glu286 to the M g 2+ ion. From Hosler, J. P., et al., Annu Rev Biochem. 2006, 75: 165-187. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

~26 A, and the K path\vay begins at GlulOI on subunit II and passes in sequence Ser299, Lys362, Thr359, a farnesyl side group of heme a3, and Tyr288 of subunitl (Figure 11.20). Either the D or K pathway is used to conduct "substrate" protons to the heme a3/CuS site. Since genetic alteration of the D pathway eliminates proton pumping, the "pumped" protons use only the D pathway. The role of subunit III appears to be a proton antenna, as it has many chal-ged residues on the surface. When subunit III is Jacking, the rate of proton uptake into the D pathway is I-educed 50%. While a great deal of evidence supports the roles of the D and K pathways for proton uptake, the exit pathway for "pumped" protons is not clear from GJu286 to the outside. However, MD simulations of a singlefile column of water molecules suggest a path from Glu286 through a hydrophobic cavity to the Mg ion (Figure 11.2 J ). Add itional experimental and theoretical work is needed to define the exit pathway. A number of othel- questions are being explored, such as what controls the directionality of proton pumping) Are there mechanisms to regulate the efficiency of cytochrome-c oxidase? Yet tremendous progress has been made in understanding the mechanism of this vital enzyme complex and in general in addressing the nature of electron transfer coupled with proton pumping.

THE TRANSLOCON

The translocon (also called the translocase and the protein-conducting channel [PCC]) is a complex of

membrane proteins that functions to transport proteins across membranes as well as to incorporate proteins into the membrane (see Chapter 7). It has a wellconserved core of integral membrane proteins that partner with various peripheral proteins and chaperones. Since many proteins are translocated as they are synthesized, it also partners with the ribosome. The structures of the translocons from canine ER, yeast, and E. coli have been imaged with cryo-EM of twodimensional crystals. After a great deal of work, the first high-resolution x-ray structure for a translocon, that from the archaea Methanococcus jal1l1.Qschii, was solved at 3.2 A resolution. With this detailed structure as a model, the lower-resolu tion structure of the E. coli translocon with an actively translating ribosome could be analyzed. The translocon in bacteria, eukaryotes, and archaea are heterotrimers of similar composition. Two of the three proteins (corresponding to SecYE in bacteria) are essential for viability, while the third (SecG) stimulates translocation as well as the ATPase activity of SecA but is not essential under normal conditions of cell growth. Additional protein partners in E. coli include SecA, the motor for posttranslational translocation; YidC, a protein involved in insertion of hydrophobic TM segments into the bilayer; and the SecDFYajC heterotrimer that facilitates protein translocation in an unknown mechanism (see Chapter 7). The M. jannaschii Translocon Structure

The crystal structure of M. jal111a.schii SecY (X(3y greatly advanced understanding of the structure of the heterotrimeric translocon and suggested how it could move proteins both across the membrane and lateraUy into the bilayer. The (X subunit has JO helices positioned in the membrane to form a sort of balTel with a (-ectanguJar shape when viewed from the cytosol and a pseudo-symmetry between two halves formed by TM IS and TM6-10 (Figure 11.22). The loop between TM5 and TM6 connecting the two halves is proposed to act as a hinge at the back side of the rectangle. Many of the helices al-e tilted up to 35° from the bilayer normal, conU-ibuting to the funnel shape of the central cavity. Not all the helices span the bilayer fully, and in particular TM2a extends only halfway through the membrane and is only partially hydrophobic. Since it is bordered on three sides by the (3 and y subunits, the only lateral opening in the rectangular SecY complex is on the side opposite the proposed hinge, gated by TM2 and TM7. The nonessential (3 subu nit, at the side near the N terminus of the (X subunit, makes only limited contact with the (X subunit; its single TM he.lix is close to the C terminus of the y subunit at the external side of the membrane. The y subunit makes a girdle, or band, along two sides of the rectangle.It consists of two helices: an amphipathic helix that

The Translocon

287

A.

11.22. Structure of the M. jannaschii translocon viewed from the cytosol. The "front" is on the left, across from the hinge at the back made by TMS and TM6 on the right. A. The ex subunit is colored blue to red from the N to the C terminus, with the TM segments numbered; the 13 subunit is shown in pink, and the y subunit in purple. B. The structure is now colored to highlight the symmetry in the ex subunit, with the N-terminal half in blue and the C-terminal half in red. From van den Berg, B., et aI., Nature. 2004, 427:36-44. © 2004. Reprinted by permission of Macmillan Publishers Ltd.

lies on the cytoplasmic surface and a long, curved helix that runs along the back side of the ex subunit. With the f3 subunit on the third side, the two halves of the ex subunit can open only on the front like a clam shell. The M. jannaschii structure can be superimposed on the EM structure of E. coli translocon (SecYE) at 8 Aresolution with a good match of the TM ex-helices (Figure 11.23). This first high-resolution structure provided a number of insights into the function of the translocon and suggested it could function as a monomer (see below) with a singJe channel down the center. The channel is shaped like an hourglass, with funnel shapes above and below its central constriction (Figure 11.24). The opening of the constriction is only 3 Ain diameter and is lined by a ring of hydrophobic and inflexible lIe side chains. When cysteine residues were engineered at 30 positions throughout SecY, only the cysteines in the region of the central constriction made a disulfide bridge with a cysteine on a translocating polypeptide, giving strong evidence that this is the pore. However, the smalJ diameter of the constriction means that other TM helices would have to shift to open the channel enough to alJow passage of an extended peptide having a maximal width of around 12 A or an (X-helical peptide having a width of 14 A; this may happen dynamically as a peptide is passing through the channel to accommodate the peptide without a leak. TM2a is proposed to be a plug that closes the channel. Indeed, when the TM2a segment in E. coli is locked by an engineered disulfide bond to the y (SecE) subunit, the channel stays open. This result implies that TM2a moves 22 A to open the channel (see Figure 11.248).

11.23. Superposition of the x-ray structure of the M. jannaschii translocon on the cryo-EM structure of the E. coli translocon. The M. jannaschii x-ray structure (numbered helices, colored as in previous figure) was visually docked onto the electron density map of the E. coli SecY complex from cryo-EM (light blue). The labeled gray cylinders represent TM helices in E. coli that have no correspondence in M. jannaschii. The diamond indicates the axis of twofold symmetry in the E. coli complex. From van den Berg, B., et aI., Nature. 2004, 427:36-44. © 2004. Reprinted by permission of Macmillan Publishers Ltd.

Membrane Protein Assemblies

288

A.

11.24. The channel pore in the M. jannaschii translocon. A. The channel is in the center of the translocon when viewed from the top. as in Figure 11.22. Half-helix TM2a (green) acts as the channel plug. B. The side view shows the hourglass shape of the pore with constrictions at the pore ring formed by three lie residues (gold). The arrow shows the modeled movement of the plug (green) toward the y subunit (magenta). From van den Berg, B., et aI., Nature. 2004, 427:36-44. © 2004. Reprinted by permission of Macmillan Publishers Ltd.

TM Insertion

The x-ray structure also suggests how a TM segment of the nascent protein could be released into the lipid bilayer. The "front" of the lX subunit is the only possible lateral opening, since the other three sides are closed by the 13 and y subunits. A hinge movement between TM5 and TM6 at the back would open the [Tont to allow the peptide access to the bilayer (Figure 11.25). A dynamic fluctuation of this hinge movement would

allow the peptide inside the channel to partition into the lipid if it were sufficiently nonpolal~ In addition to overall hydrophobicity, the positions of some residues influence this lateral partitioning, according to studies with model peptides (see Chapters 4 and 7). While the x-ray stl-ucture shows a central pore in the translocon from M. jQIlI1Qschii, suggesting it is active as a single copy of the SecY heterotrimer, there is substantial evidence for higher-order mul timers of the Sec translocons. The SecYEG complex purified h'om E. coli exists as oligomers. The cryo-EM images from twodimensional crystals formed by slow detergent removal in the presence of phospholipids clearly show dimers of SecYEG (see Figure 11.23). Translocon assembly into dimers, or even oligomers, could be a dynamic process that is triggered by interactions with other partners, such as the ribosome or SecA in E. coli. Cryo-EM has now been used to investigate how the SecYEG translocon might interact with the ribosome.

The Translocon-Ribosome Complex

11.25. Proposed lateral gate of the M. jannaschii translocon. The TM segment of the nascent protein is depicted as a magenta cylinder, and its movement into the lipid bilayer is indicated by the arrow to the left. This is postulated to involve an opening between TM helices 7 and 8 on one side and TM2b and TM3 on the other, resulting from the hinge at TM5 and 6 as shown by the arrow on the right. From van den Berg, B., et aI., Nature. 2004,427:36-44. © 2004. Reprinted by permission of Macmillan Publishers Ltd.

Images of the canine translocon bound to a ribosome achieved with cryo-EM show a larger assembly of translocons with a diameter of ~ 100 'A, which fits a tetra mer of Sec61 heterotrimers. The ribosometranslocon assembly shows a central depression between the four heterotrimers that is believed to be filled with lipids, in contradiction to the proposed aqueous pore. The ribosome exit tunnel is nearly centered over the tetra mer, but the contacts between the ribosome and the translocon indicate that only two of the Sec61 heterotrimers have access to the tunnel.

The Translocon

289

3' mRNA entrance

11.26. Cryo-EM image of the complex of ribosome and translocons from E. coli. The small (30S, yellow) and large (SOS, blue) ribosomes are shown with the A, P, and E sites (magenta, green, and orange), and the L7/L12 stalk is labeled. The mRNA (cyan) is visible and so is the nascent chain (gold) in the exit tunnel. The translocating SecYEG dimer (dark blue) is at the exit tunnel, while the nontranslocating SecYEG (red) is at the S'mRNA exit. From Mitra, K., et aI., Nature. 2005. 438:318-324. © 2005. Reprinted by permission of Macmillan Publishers Ltd.

High-resolution EM studies of a translocating SecYEG complex from E. coli with a translating ribosome show a complex of two Sec heterotrimers in a front-to-front orientation and also suggest different conformations for translocating and nontranslocating complexes. These studies were performed with detergent-solubilized SecYEG added to a cell-free system of E. coli ribosomes translating an mRNA that encodes a chimeric protein with the signal-anchor of

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Contacts with the Ribosome The contacts between the translocating SecYEG dimer and the ribosome occur at three regions, called Cl, C2, and C3. They involve both ribosomal proteins and rRNA from the ribosome and two large cytoplasmic loops (between TM6 and 7 and between TM8 and 9) of SecY

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FtsQ fused to a stalling domain of SecM, which slowed it sufficiently to allow a snapshot of the translocation process by cryo-EM. Computer modeling was used to fit a homology-based atomic model of the E. coli translocon derived from the 1vl. jannaschii structure, along with the detailed structure of the ribosome, to the 11 A resolution EM images. This complex shows two assemblies of translocons (PCCs) associated with the ribosome (Figure 11.26). One is associated with the exit pore of the large subunit and is apparently involved with the movement of the nascent peptide. The other one is nonphysiologically bound to mRNA near its exit site in the small subunit and is thus considered to be in a nontranslocating state. The atomic coordinates of two SecYEG translocons in a front-to-front arrangement could be fitted to the nontranslocating structure. consistent with the inactive/closed state of the x-ray structure, and enabled analysis of the structural changes required to obtain the activeltranslocating state for the other one. The model of the translocating state shows sufficient opening of the SecY halves within the membrane plane to allow formation of a wider pore (Figure 11.27), However, the pore is not shared between the two monomers, and the two pores of the SecYEG heterotrimers seem to be in different states, one open to the bulk lipid and the other inaccessible to lipids,

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11.27. Nontranslocating and translocating SecYEG dimers in front-to-front orientation. The van der Waals surface representations obtained by fitting to the nontranslocating (A) and translocating (B) electron density observed by EM are shown with the SecY C-terminal halves transparent. The green arrow indicates the change in the heterotrimer interface at the front, and the yellow arrows point out the changes in the opening of SecY. One heterotrimer is blue/green and the other is in shades of red. The ribosomal side is behind the plane of the membrane. From Mitra. K., et aI., Nature. 2005, 438:318-324. © 2005. Reprinted by permission of Macmillan Publishers Ltd .

Membrane Protein Assemblies

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290

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11.28. Path of the nascent chain through the ribosome and translocon. A. The ribosome bound to a front-to-front SecYEG dimer is viewed from the side with a nascent chain (yellow) in the exit channel. Only the molecules from the ribosome that contact the translocon are shown. B. A schematic version of the molecules involved, with the rest of the ribosome and the membrane added. The nascent chain is yellow in the ribosome exit tunnel and the translocon pore is green. From Mitra, K., et aI., Nature. 2005,438:318-324. © 2005. Reprinted by permission of Macmillan Publishers Ltd.

(Figure 11.28). At Cl the rRNA helix 59 contacts one SecY, and at C2 the rRNA helix 24 contacts the other SecY. At C3 the proteins L29 and L23 from the ribosome make contact with the cytoplasmic region of SecG and possibly the N-terminaJ part of SecE in a nonessential but stabilizing connection. Since C3 is at the back, there is a large opening at the front, providing space between the translocon and the ribosome that is accessible to the cytoplasm. This access to the cytoplasm means the ribosome does not plug the translocon pore to prevent leakage, as earlier envisioned; thus the translocon itself must be capable of providing a tight seal to maintain the permeability barrier. Structures of the translocon with and without the presence of a ribosome-nascent chain complex answer many questions about protein export and leave many others unanswered (see Chapter 7). It is still not clear how the N-terminal portion of the nascent chain inserts into the translocon, presumably as a hairpin in the initial step. How does a TM segment reorient inside the transJocon, as indicated in studies of topogenesis? The dynamic interaction with SecA, presumed to include its insertion into the translocon, is not understood. What is the relation to other proteins that assemble at the translocon, such as SeeD, SecF, and YajC? StilJ other proteins, such as signal or leader peptidase and oligosaccharide transferase (in eukaryotes), are present on the outside of the membrane as part of the export

process. Clearly, the translocon is at the center of an amazing molecular machine that carries out dynamic and complex processes.

ABC TRANSPORTERS AND BEYOND

ABC transporters carry out the uptake or efflux of a wide variety of substances at the expense of hydrolysis of ATP (see Chapter 6). All ABC transporters have four domains, two TM domains and two nudeotidebinding domains (NBDs) that are synthesized as one to four polypeptides (see Figure 6.6). A molecular understanding of this important class of transporters is provided by the x-ray structures of two ABC transporters, the Sav1866 protein and the BtuCD complex. Many ABC transporters work in tandem with otber proteins to facilitate uptake or efflux across the two membranes and the space between them in the cell envelope of Gram-negative bacteria. The other components working with BtuCD in E. coli vitamin B I2 uptake include two other specific proteins 'whose structu res have been solved (BtuB and BtuF) and three less understood proteins (TonB, ExbB, and ExbD) required for energy coupling from the inner to the outer membranes. After a description of this complex system, the chapter ends with the structure and function of the Sav 1866 protein as well as others involved in drug efflux.

ABC Transporters and Beyond

291

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lizes two kinds of energy coupling and involves seven proteins to span the two membranes of the cell envelope. Interestingly, the genes encoding specific B I2 transport proteins are not in an operon but are scattered as bluE, blueD, and bluF loci in the E. coli chromosome. The recent acquisi tion of structures of all four B,rspecific transport components provides a structural basis to begin to understand the complexities of this system, which provides a good model for other ABC transport systems. Transport across the Inner Membrane Delivery by BtuF

5'-Deoxyadenosylcobalamin (coenzyme B l2 ) 11.29. S'-Deoxyadenosylcobalamin, vitamin B12. The Co(1I1} ion is liganded by four pyrrole N atoms of the corrin ring and the N atom of S,6-dimethylbenzimidazole (DMB), which is covalently linked through its 3'-phosphate group to a side chain of the corrin ring. The sixth ligand is a S'-deoxyadenosine in most physiological conditions, as shown, that is replaced during purification by a cyano group to produce cyanocobalamin.

Vitamin B '2 is transported across the inner membrane by an ABC transport system consisting of BtuCD and BtuF. BtuF is the soluble substrate-binding protein thaI avidly binds the cofactor (Kt ~ 15 nM) as it enters the periplasm through the BtuB channel in the outer membrane (discussed below). The x-ray structure of BtuF with bound vitamin B I2 shows two lobes that each consist of a central five-stranded l3-sheet surrounded by helices, a Rossmann-like fold (Figure 11.31). Betvveen the two lobes is a deep cleft with the substrate-binding site. The CN-cbl is bound with its DMB ligand present, and it contacts six aromatic residues, three from each lobe of BtuF. Unlike most of the ABC substrate-binding proteins in E coli, BtuF has a backbone ex-helix spanning the two domains that makes it unlikely to undergo a large hinge motion to the unliganded state. That such a large movement is not required to release the su bstrate is indicated by a zinc-binding protein from Treponema BtuB

The Vitamin 8 12 Uptake System

Vitamin B ,2 , or cyanocobalamin (CN-cbl), is a cofactor produced by some bacteria and archaea and is required by a variety of enzymes in most cells. Specific transporl systems enable cells to import this large, inflexible molecule, which consists of a corrin ring (a tetrapyrrole with a cobalt metal) plus two axial ligands, cyanide and 2,3-dimethyl-benzimidazole (DMB), covalently linked to the ring via aminopropanol-phosphate-ribose (Figure J 1.29). Transport of vitamin B '2 can be fully induced in E coli by growth on ethanolamine (because it is needed for the ethanolamine ammonia lyase reaction) and monitored by uptake of radiolabeled [57Co]CN-cbJ. Extensive biochemical and genetic studies have characterized two energized phases of vitamin B '2 transport: uptake across the outer membrane utilizing the specific receptor BtuB coupled with the TonB/ExbBD system for energy input, and uptake across the inner membrane via the ABC transporters BtuCD and BtuF (Figure J 1.30). Thus this system uti-

BtuCD

TonB-ExbBD complex 11.30. Components of the transport system for vitamin B12. Structures for BtuCD, BtuF, and BtuB have been solved, along with the C terminus of TonB, while the structures of ExbB and ExbD are not known. The general porin is included in the outer membrane because it may allow passive diffusion of vitamin B12 into the periplasm. From Kadner, R. J., et aI., in R. Benz (ed.), Bacterial and Eukaryotic Porins, Wiley-VCH, 2004, pp. 237-2S8. © 2004 by Wiley-VCH. Used by permission of Wiley-VCH Verlag GmbH.

Membrane Protein Assemblies

292

N

with gates at each end. Therefore to transport vitamin BIz to the cytoplasm requires a conformational change, which is likely to change the tilts of the TM helices analogous to the opening of LacY and GlpT (see Chapter 10). This conformational change is triggered by the binding and/or hydrolysis of ATP at the BtuD NBDs, likely A.

11.31. X-ray structure of BtuF, the periplasmic vitamin B1Zbinding protein. The ribbon diagram shows l3-sheets (blue) at the substrate-binding lobes and <x-helices (green) in the lobes and the backbone, with an asterisk denoting the helices that form the backbone. The substrate, vitamin B1Z, is shown as a ball-and-stick model. From Borths, E. L., et aI., Proc Natl Acad Sci USA. 2002, 99: 16642-16647. © 2002 by National Academy of Sciences, USA. Reprinted by permission of PNAS.

pallidum with a similar backbone, which requires a 4° tilt of the C-terminal domain to collapse the substratebinding site. BtuF carrying vitamin B 12 docks on the peri plasmic side of the BtuC dimer. Conserved glutamate residues on the surface of BtuF can be aligned with arginine residues on BtuC to form the stable complex described below. BtuCD in the Inner Membrane The BtuC and BtuD proteins transport vitamin BIz across the inner membrane at the expense of ATP hydrolysis. They both dimerize to form a heterotetramer, BtuCzD z, thus providing the two TM domains and the two NBDs that are standard for ABC transporters (see Figure 6.6). Crystals of the heterotetramer in the absence of both vitamin BIz and ATP allowed the structure to be refined at 3.2 A resolution, giving definition of the entire structure except for 17 C-terminal residues of BtuD and a few residues of the periplasmic loops of BtuC (Figure 11.32). Each BtuC subunit has 10 tilted TM helices, giving the assembled transporter significantly more TM segments than observed in most ABC transporters (which typically have 12). Between the BtuC subunits is a wide hydrophobic cavity forming a translocation pathway that is large enough to accommodate vitamin BIz but lacks a specific binding site for it. In the crystal structure the channel opens to the peri plasm. Two loops at the ends of TM helices 4 and 5 form a gate that closes the channel to the cytoplasm. The channel is proposed to work like an airlock

B.

11.32. X-ray structure of the ABC transporter BtuCzDz. A. Viewed from the side, the two BtuC subunits (purple and red) span the membrane with their L1 and L2 helices (gold) at the cytoplasmic surface. The TM helices are numbered. The two BtuD subunits (green and blue) associate with BtuC at the cytoplasmic side and have cyclotetravanadate molecules (ball-and-stick models) at their ATP-binding sites. B. The BtuD NBD domains (blue and green) are viewed from the cytoplasmic side, with the P-looplWalker-A motif (red), Walker-B motif (pink), and ABC signature motif (yellow) labeled (compare with Figure 6.7). From Locher, K. P, et aI., Science. 2002, 296:1091-1098. © 2002. Reprinted with permission from AAAS.

ABC Transporters and Beyond A.

293

B.

C.

11.33. MD simulation of the effect of ATP binding to BtuC2D2. Computer simulations based on the crystal structure of the BtuC2D2 heterotetramer were run for 15 ns in the absence of nucleotide (A). Then ATP was positioned at each of the two binding sites. In B, ATP is bound to binding site I (red) and in C, it is bound to binding site II (green). The cavities are colored to highlight the shifts in the transport channel (yellow) and the gap separating the NBDs (gray). From 0100, E. 0., and D. P. Tielelman, J BioI Chem. 2004, 279:45013-45019. © 2004 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

mediated by two short helices, L1 and L2, that make an L-shape at the interface between BtuC and BtuD (see Figure 1 1.32A). Both the overall fold and the nucleotide-binding sites of BtuD resemble those of other ABC NBDs described in Chapter 6 (see Figure 6.7). BtuD contains a six-stranded l3-sheet surrounded by nine a-helices, with a peripheral three-stranded l3-sheet. The nucleotidebinding sites, which contain critical conserved residues in the P and Q loops and the Walker-B motif, are occupied by vanadate salts in the crystals. Because the BtuD subunits face opposite directions in the dimel~ the ABC signature from one subunit is opposite the P-Joop of the other, creating two ATP-binding sites at the interface between BtuD subunits (see Figure 11.32B). In the structure lacking ATP, this interface is not extensive, so it is likely that the BtuD dimer is stabilized by the interfaces between each BtuCD pair, where conformational changes due to ATP binding and hydrolysis must be transmitted to the TM domains. Key residues for this transmission have been identified by analogy to other ABC transporters; for example, Leu96 in BtuD COlTesponds to PheS08 of CFTR, the si te of mutations in 70% of the cases of cystic fibrosis (see Chapter 6). Binding of ATP appears to have a significant effect on the BtuCD conformation. MD simulations used to dynamically probe the ATP-binding process suggest that docking of ATP draws the two NBDs closer to each other, which then alters the translocation pathway of the TM domain by reorienting TM helix 5 (Figure 11.33). Other effects that are likely to trigger further

BtuF-B I2

Membrane

Cytoplasm

11.34. Model of the BtuC2D2F heteropentamer. Ribbon diagrams of the BtuC, BtuD, and BtuF proteins are shown in the heteropentamer that is suggested from complementary groups on BtuC and BtuF. Two cyclotetravanadate molecules occupy the ATP-binding sites in BtuD, and vitamin B12 occupies the binding site of BtuF. From Locher, K. P., et al. FEBS Lett. 2004, 564:264268. © 2004 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

Membrane Protein Assemblies

294

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11.35. Assay systems for the activities of the BtuC2D2F heteropentamer. The stable heteropentamer is reconstituted into proteoliposomes in random orientation, which allows hydrolysis of external ATP (A) as well as uptake of vitamin B12 with preloaded ATP (B). ATP hydrolysis can also be measured in detergent micelles (C). From Borths, E. L., et aI., Biochemistry. 2005, 44: 16301-16309. @ 2005 by American Chemical Society. Reprinted with permission from American Chemical Society.

confm'mational changes and allo\,v delivery of vitamin B 12 to the cytoplasm include the hydrolysis of ATP and the binding of substrate-loaded BtuF. The specificity of the inner membrane transport system resides in the high-affinity binding site of BtuF, since the channel in BtuCD does not appear to specifically recognize vitamin B 12 . The complementary groups of BtuF and Btue allow a model to be made of the BtuC 2 D2 F heteropentamer, which shows the substrate is positioned directly over the translocation channel (Figure 11.34). Mixing purified Btu F with the purified BtuCD proteins in a 5: 1 ratio, followed by removal of excess BtuF, produces a stable BtuC 2 D 2 F complex that hydrolyzes ATP in detergent micelles and in reconstituted proteoJiposomes and also transports vitamin B I2 (Figure 11.35). Given the protein-protein interactions in the complexes, models have been proposed for the

coupling of ATP to vitamin B 12 transport, with conformational changes occurring in both TM and NBD domains (Figure 11.36). Crystallization of the proteins at other stages in the transport process as well as achievement of higher-resol ution structures will further the understanding of this process. Transport across the Outer Membrane

BtuB in the Outer Membrane The specific outer membrane ('eceptor for vitamin B I2 is the BlUB protein, a f3-barreJ protein with striking similarities to the FepA and FhuA iron receptors described in Chapter 5, These specific outer membrane transporters carry out active lI-ansport only in the presence of the TonB protein (see below), so they are called TonB-dependent transporters (TBDTs). BtuB has a high

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11.36. A model for the transport of vitamin B,2. BtuF (green) brings vitamin B'2 (red) to the BtuC2D2 complex (black), sending a signal (blue dashed arrows) across the membrane to trigger ATP binding and hydrolysis, Hydrolysis of two ATP molecules (yellow) promotes conformational changes that release vitamin B 2 to the cytoplasm and also release BtuF. From Locher, K. P., Curl' Opin Struct Bioi. 2004, ' 14:426-431. © 2004 by Elsevier. Reprinted with permission from Elsevier.

ABC Transporters and Beyond

11.37. Ribbon diagram of BtuB, the outer membrane receptor for vitamin B12. The 22-stranded j3-barrel surrounds the globular hatch domain in the interior, similar to the FepA and FhuA proteins also in the outer membrane (see Figure 5.23). From Chimento, D. P., et aI., Proteins. 2005, 59:240-251. © 2005. Reprinted with permission from John Wiley & Sons, Inc.

affinity for vitamin B 12 , and its crystal structure has been solved in the presence and absence of bound substrate. The BtuB protein has two domains: a C-terminal domain that makes the f3-barrel, and an N-terminal globular domain thaI folds into the barrel to make a hatch (Figure 11.37). The barrel consists of 22 tilted antiparallel amphipathic f3-strands connected by short turns on the periplasmic end and somewhat longer loops of nine to 19 residues on the outside. In the structure without bound substrate, the loop joining strands 9 and lO is ordered while loops 2, 3, and 4 are disordered; in the structure with vitamin B I2 bound, the reverse is the case, suggesting that the loops are mobile and close to trap the bound substrate. The hatch domain is highly conserved among TBDTs. It has a polar exterior (compatible with the polar interior walls of the barrel) and a hydrophobic core. The core is a four-stranded f3-sheet, with connecting loops that have distinct roles (Figure 11.38). The two helices are amphipathic and interact with the core 13sheet through their hydrophobic sides. The three apical loops are involved in binding vitamin B 12 , and the loop between the third and fourth strand is called a "latch" because it interacts with f3-strands 13 through 15 of the barrel that fold slightly into the lumen. Near the N terminus, seven residues (Asp6-Thr-Leu-Val-Val-Thr-Ala) form the "Ton box," defined as the major site of interaction with the TonB protein by data from genetic and cross-linking studies. The Ton box is inside the barrel on the peri plasmic side ofBtuB, and its exposure to the periplasm increases somewhat when substrate binds.

295

The BtuB protein binds eN-cbl with its axial ligand, DMB. The substrate-binding site on BtuB is on one side of the external face of the molecule and involves residues from both the barrel and hatch domains. Binding of two calcium ions is prerequisite to binding the vitamin B I2 and helps to order barrel loops 2, 3, and 4 that clamp around the bound substrate. Two apical loops from the hatch domain pack against the DMB, and equatorial side chains of the corrin ring fit into small pockets formed by both domains. The tight binding (nM KI) is not surprising in view of all the polar and van der Waals interactions between the receptor and the substrate, including 11 hydrogen bonds. The high affinity of BtuB for vitamin B I2 suggests a conformational change is required for the receptor to carry out transport. Indeed, no channel is evident in the available structures of BtuB or any other TBDT. However, the hatch domain in the internal cavity does not have a close fit to the barrel walls, and the many water molecules in the interface between them mediate half of the hydrogen bonds between the two domains. Yet two thirds of the water molecules present are hydrogen bonded to one domain or the other, not both, which is characteristic of a transient protein-protein interaction (Figure 11.39). Thus it is likely that the hatch moves out of the barrel to allow passage of vitamin B 12 , as BtuB

Substrate binding Loop / Loop 3 \ Loop 2 \

r'~

iRG"r

I

f\,,--, TDG

11.38. Conserved features of the hatch domain of TBDTs. The four j3-strands (labeled hj31-4, arrows) are connected by loops and <x-helices (cylinders) with particular roles: the apical loops 1 through 3 are involved in substrate binding (red), the loop between hj33 and hj34 is the latch (black). The site of interaction with TonB, called the Ton box, is connected to a "switch helix" (gold) in two iron receptors but not in BtuB. The connection of the hatch domain to the barrel domain is the linker (black). Residues of conserved motifs whose functions are not known are labeled (blue) From Chimento, D. P., et al., Proteins. 2005,59:240-251. © 2005. Reprinted with permission from John Wiley & Sons, Inc.

296

11.39. Water molecules at the interface between the barrel and hatch domains of BtuB. The polar interfacial regions are filled with many waters. Bridging waters (green) make hydrogen bonds to both domains, while nonbridging waters (blue) make hydrogen bonds to a single domain or to other water molecules. From Chimento, D. P., et al., Proteins. 2005, 59:240-251. © 2005. Reprinted with permission from John Wiley & Sons, Inc.

undergoes a significant conformational change to make room for this very large and inflexible substrate to pass.

TonB and Energy Coupling Such a major conformational change requires energy, and surprisingly BtuB and the other TBDTs in the outer membrane utilize the energy of the pmI' (proton motive force) across the inner membrane. The energy coupling is carried out by TonB and its accessory proteins, ExbB and ExbD, which are anchored in the inner membrane. TonB and ExbD are both predicted to have single TM segments near their N termini, with large periplasmic domains. ExbB is predicted to have three TM segments with a large cytoplasmic loop between the first two. In the cell, these proteins form a complex of ~260 kDa with the stoichiometry 1 TonB: 2 ExbD: 7 ExbB, which may dimerize. TonB has 239 amino acids in three domains: the N-terminal TM domain, a proline-rich periplasmic domain, and a C-terminal domain of 48 residues that contacts outer membrane receptors. The uncleaved signal sequence is its TM domain, which contains a highly conserved sequence along one face of the ex-helix. The TM domain of TonB interacts with the TM segment of ExbD. Little is known about ExbB and ExbD, but they seem to be similar to MotA/B, 'which use the pmI' to drive the motion of bacterial flagella. A high-resolution structure of the C terminus (residues 155-239) of TonB reveals a highly cylindrical dimer, in which each monomer has three l3-strands. one ex-helix, and a short 3 10 helix (Figure 11.40). If this structure has not been perturbed significantly by the lack of the rest of the protein, it indicates that TonB forms a dimer that extends at least 65 A, halFway across the periplasm. Models proposed for the energy coupling between TonB/ExbB/D and the TBDTs suggest that an

Membrane Protein Assemblies energized state of TonB contacts a TBDT in the outer membrane and alters the conformation of the transporter. Evidence for such a mechanism is provided by a high-resolution structure of BtuB complexed with the C terminus of TonB, which shows the Ton box of BtuB has become a l3-strand recruited by the l3-sheet in TonB (Figure 11.41). The interaction between the l3-strands is predicted to be sufficient to allowTonB to pull the hatch domain out of the barrel, opening a transport channel for vitamin B 12 • The function ofBtuB protein is the outer membrane transport of vitamin B 12 ; however, like other outer membrane proteins. it is also used as a receptor for colicins and bacteriophage. specifically colicins E and A and phage BF23. Entry of these agents into the cell, which is also TonB-dependent. is poorly understood. DRUG EFFLUX SYSTEMS In contrast to the mysterious TonB/ExbB/D complex that couples the inner and outer membranes for uptake, much more is known about how molecules on their way out of the cell cross the periplasmic space due to the high-resolution structure for the amazing channeltunnel made by ToIC, along with structures of several other proteins involved in efflux. Some of these proteins serve to export lipids from the inner membrane, and some are related to proteins involved in the secretion of proteins such as hemolysin and colicins. But they are best known for their role in drug efflux because the problem of multidrug resistance (MDR) is now limiting

11.40. The C-terminal domain of TonB. The ribbon diagram of residues 155 to 239 of TonB shows two molecules (red and blue) are intertwined along the entire 65 A length of the structure, with [3-strands alternating from each monomer. From Chang, C, et al., J Bioi Chem. 2001,27535-27540. © 2001 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

Drug Efflux Systems

297

ondary transporter AcrB belongs to the RND (resistance nodulation cell division) superfamily. AcrB is part of a well-characterized tripartite drug efflux system that includes the membrane fusion protein, AcrA, and utilizes the remarkable TolC tunnel to export drugs directly to the extracellular space. Representing the diverse mechanisms for drug efflux, these five proteins provide important models for the mechanisms that rid cells of unwanted toxic compounds. Sav1866, an ABC Multidrug Transporter

11.41. The ribbon diagram of the complex of BtuB and the C-terminal domain (residues 147-239) of TonB. The front of the BtuB protein, shown with the outer l3-barrel (copper) and the inner hatch domain (green). has been removed to show the lumen and the BtuB-TonB interaction. The purified proteins were combined in a molar ratio of 1:5 in the presence of vitamin B 12 and calcium. In the x-ray structure. the Ton box from BtuB (blue) has been recruited by the l3-sheet of the TonB C terminus (magenta). The substrate, vitamin B12 (red spheres). is bound in the extracellular region above the hatch domain of BtuB. From Shultis, D. D., et aI., Science. 2006, 312:1396-1399.

treatment options for many cases of cholera, pneumonia, gonorrhea, and tuberculosis. Today, pathogens resistant to almost any antibiotic can arise due to multidrug efflux pumps. Genome analyses predict a role in drug efflux for 6% to 18% of all transporters in bacterial membranes. A few of the bacterial drug transporters are related to human MDR proteins that are making many tumors highly drug resistant. In Gram-negative bacteria, many MDR proteins are organized into tripartite systems, each consisting of an inner membrane transporter and a peri plasmic lipoprotein called a membrane fusion protein, in addition to the ou ter membrane channel. The transporters use either ATP hydrolysis or the proton motive force as the source of energy and belong to at least six families in the transporter classification scheme described in Chapter 6 (ww,...,.tcdb.org). X-ray structures are now available for prokaryotic transporters involved in drug efflux from three of these classes. The Sav 1866 protein is an ABC transporter related to mammalian MDR proteins, including P-glycoprotein (see Chapter 6). The EmrE protein is a member of the small MDR family that exports toxic hydrophobic compounds. The sec-

Sav 1866 is a multidrug transporter from Staphylococcusaureus with homology to several human MDR transporters, including P-glycoprotein. It uses the hydrolysis of ATP to drive efflux of numerous drugs, including the cancer drugs doxorubicin and vinblastine. lL is a homodimeric ABC transporter, with each dimer contributingoneTM domain and one NBD. While the structure of the BtuCD transporter for vitamin B I2 uptake has been solved in the absence of bound nucleotide, the structure of Sav 1866 with bound nucleotide gives additional insight into the coupling of ATP hydrolysis with transport. Each Sav1866 subunit has an N-terminal TM domain (residues 1-320) and a C-tenninal nucleotidebinding domain (residues 337-578) with a short linking peptide (residues 321-336). The x-ray structure of Sav1866, obtained at 3.0 A resolution, shows an elongated molecule (120 A long) that extends well into the cytoplasm (Figure 11.42). The NBD structure is similar to those of other well-characterized ABC transporters, with the nucleotide-binding sites at the interface between subunits where the P-loop of one NBD is across from the ABC signature motif of the other (see Figure 6.7). In the x-ray structure, Sav1866 has two molecules of ADP tightly sandwiched at this interface. Like many ABC transporters, the TM domain of the Say 1866 dimer has twelve TM a-helices; their symmetry in the structure suggests the occurrence of a gene duplication event as postulated for AQPs and major facilitators (see Chapter 9), but not previously observed in an ABC transporter. Helices from the two subunits are intertwined at the center of the membrane, where they bend outward to form two wings, with TM 1-TM2 from one subunit aligning with TM3-TM6from the other (see Figure 11.42). This means the two subunits do not form separate lobes of the transporter, as postulated for many ABC transporters. Constrained by these interactions, the two subunits are not likely to act independently and the NBDs probably do not completely separate during the reaction cycle. Like other multidrug transporters (see below), Sav1866 has a large internal cavity and lacks a welldenned drug-binding site. Rather, drugs appear to bind nonspecifically in the cavity, whose affinity for the drugs is altered by conformational changes. In the x-ray

Membrane Protein Assemblies

298 A.

u

·s CJJ

Ol

TMDs

!i

2 u>.

NBDs

_1 C-ter 11.42. The ribbon diagram of the x-ray structure of Sav1866, an ABC drug efflux transporter in S. aureus. The views in A and B are rotated by 90° The subunits (yellow and green) are intertwined, especially as they cross the membrane (gray shading). Two molecules of bound ADP are visible between the NBDs. The TM segments (numbered in B) are connected by short loops on the extracellular side (labeled ECll, 2, and 3 in B) and long loops on the cytoplasmic side (labeled ICl). Based on Dawson, R. J. P., and K. P. locher, Nature. 2006, 443: 180-185. © 2005. Adapted by permission of Macmillan Publishers Ltd.

structure, the translocation pathway is accessible to the outside and the outward-facing cavity is polar. It has charged residues along its surface in the inner leaflet primarily from TM2-TMS, while at the outer leaflet it is lined by TMl, TM3, and TM6. In this conformation, extrusion of hydrophobic drugs occurs with simple diffusion from the low-affinity cavity. Biochemical studies of similar MDR proteins suggest that when the cavity is open to the cytoplasm, it has a high affinity for the drugs. Thus the transport mechanism appears to utilize an alternating access mechanism, like that observed with the major facilitators (described in Chapter 9) except using the hydrolysis of ATP to drive the conformational change. At the interface between the NBD and the TM domain lie portions of the long intracellular loops, ICLI and ICL2. Each of these loops contains a short helix running nearly parallel to the plane of the membrane; they are called "coupling helices" since both genetic and structural data implicate them in communication between domains. Interestingly, only one of the coupling helices contacts the NBDs of both subunits; the

11.43. Model for the interaction of the TM domains and the NBDs in Sav1866. Due to the extensive interactions between TM domains at the "wings" as well as the contact between one intracellular loop with both NBDs, small conformational changes rather than large domain movements are expected to result from ATP-binding. From Schuldiner, S., Nature. 2006, 443: 156-157. © 2006. Adapted by permission of Macmillan Publishers Ltd.

Drug Efflux Systems

A.

299

B.

11.44. X-ray structure of the EmrE dimer. Ribbon diagrams of the EmrE structure viewed from the plane of the membrane (A) and perpendicular to the plane of the membrane (8, top view). Each monomer is shown in rainbow color (N terminus, blue; C terminus, red), with a space-filling model of TPP+ (magenta). From Tate, C. G., Curr Opin Struct Bioi. 2006, 16:457-464. © 2006 by Elsevier. Reprinted with permission from Elsevier.

other one contacts only the opposite NBD. The portion of the NBDs that contacts the TM domain is primarily residues around the Q-Ioop. In addition, a newly recognized conserved motif called the "x-loop" near the ABC signature motif of the NBD is proposed to form and break a cross-link to the ICLs and thereby to transmit conformational changes upon ATP binding and hydrolysis. In short, the coupling between the NBDs and the TM domains appears to utilize communication of moderate conformational rearrangements [Tom the NBDs uponATP binding (Figure 11.43), rather than their association and dissociation as described for BtuCD (see above). EmrE, Small but Powerful

EmrE from E. coli is only 12 kDa (110 amino acids), and yet when overexpressed it causes bacteria to be resistant to tetracycline, tetraphenylphosphonium (TPP+), ethidium bromide, and other antiseptics and intercalating dyes. Found in both Gram-positive and Gramnegative bacteria but not in eukaryotes, it is a proton! drug antiporter that uses the energy of the proton motive force to drive transport of drugs out of the cytoplasm. Predicted to have four <x-helices, EmrE is very hydrophobic and has only eight charged residues. The most important of these is the only buried charged residue, Glu14, which is involved in coupling substrate transport and proton translocation. The pKa of Glu 14 is elevated to 8.5 due to nearby hydrophobic residues (Trp63, Tyr40, and Tyr60). When aspartate is substituted for Glu 14, its pK\ is only around 6.5 and it releases protons without using them to pump out drugs. Cysteine replacements to study accessibility of residues to alkylating agents identified other hydrophobic residues likely to be in the substrate-binding site. These residues

(Leu7, AlalO, Ilell, Tyr40, and Trp63) are from TM helices 1 (near GluI4), 2, and 3. While there is agreement that EmrE functions as a dimeI', the nature of the dimer is controversial. The structure of EmrE solved by both cryo-EM and x-ray crystallography appears to be asymmetric dimers (Figure 11.44). Each monomer contributes fourTM helices, as expected, with the substrate bound in a cleft formed by two helices [Tom one monomer and one from the other. An axis of pseudo-symmetry relates the first three helices of one monomer to the first three helices of the other. This dual topology was detected in a global topology genomic analysis of E. coli membrane proteins (see Chapter 6) and is supported by data from mutational analysis of EmrE fusion proteins. However, it is inconsistent with biochemical and cross-linking data. In the most compelling cross-linking result, the n08C mutant of EmrE was purified after cross-linking with 0phenylenedimaleimide, a rigid cross-linker about loA in length. In asymmetric dimers, the positions of T1 08 in the two monomers would be over 35 Aapart, strongly suggesting the cross-linked species, which has full activity, is a symmetric dimeI'. Some EmrE mutants have a C-out topology, while others have a C-in topology. Furthermore, different topologies are observed with different constructs, for example, if a histidine tag and epitope are placed at the N or C terminus. If the EmrE sequence is posed such that it is readily pushed into one topology or another, it may represent an interesting point in evolution. Gene duplication events that produced larger transporters have produced some with domains in parallel orientation, such as LacY, GlpT, and AcrB, and others with domains in antiparallel orientation, such as AQPs. The controversy over the EmrE dimer has intensified with the recent acknowledgment of a software

Membrane Protein Assemblies

300

~

Drug

Medium

/ Cytoplasm

11.45. Schematic representation of the MDR system composed of AcrABffolC. Drugs can enter the AcrB (yellow) in the membrane for export from either the cytoplasm or the peri plasm. AcrA {green} is thought to stabilize the docking of TolC (blue) on AcrB and allow drugs to be transported outside the cell. From Murakami, S., and A. Yamaguchi, Curr Opin Struct Bioi. 2003, 13:443-452. © 2003 by Elsevier. Reprinted with permission from Elsevier.

glitch that affected the solution of the EmrE crystal structure. Further structural and biochemical studies are needed to determine whether two EmrE monomers, identical in sequence, can insert into the membrane with opposite orientations. If this does occur in cells, it raises interesting questions regarding the determinants of membrane protein topogenesis. Tripartite Drug Efflux via a Membrane Vacuum Cleaner

Drug efflux in Gram-negative bacteria often employs three-component systems that span the cell envelope to extrude drugs directly into the outside medium. A well-characterized tripartite system in E. coli is composed of AcrB, an inner membrane transporter; AcrA, a periplasmic lipoprotein; and ToIC, the channel-tunnel protein (Figure 11.45 and Frontispiece). An analogous system is made up of the secondary transporter EmrB, the peri plasmic lipoprotein EmrA, and Tole. Similarly, the system for exporting hemolysin is made up of HlyB, HlyD, and Tole. In E. coli several systems for drug and protein export are known to use TolC, whereas drug efflux systems in other bacteria, such as Pseudomonas aeruginosa, have different tunnel components for the different efflux systems (see below). Assay of different transporters in vitro demonstrates that substrate specificity of these MDR systems resides in the inner membrane transporters. The AcrAB/ToIC system exports a broad variety of mostly amphiphilic compounds that

may be positively or negatively charged, zwitterionic, or neutral, including bile salts, erythromycin, and 13lactams such as ampicillin. Such MDR systems have been called "membrane vacuum cleaners" because they can remove a large number of unwanted substances h-om the inner leaflet of the cytoplasmic membrane and pump them out of the cell, preventing their accumulation in either the cytoplasm or the periplasm. AcrB, a Peristaltic Pump

The inner membrane transporter of a tripartite drug efflux system may belong to one of several different classes of transport proteins. As a member of the RND superfamily, AcrB is a proton/drug anti porter that utilizes the proton motive force to export many clinically important drugs. When purified and reconstituted into proteoliposomes, AcrB carries out proton-dependent transport of these drugs. Like numerous other transpNters, including other MDR proteins, the sequence of AcrB can be divided into two halves that are homologous, suggesting they evolved h-om gene duplication events. AcrB Structure

The first crystal structure of AcrB revealed a homotrimer in the shape of a jellyfish, with each 110-kDa monomer (1049 residues) providing 12 TM a-helices and also contributing to the large peri plasmic headpiece (Figure 11.46). In the bilayer-spanning domain the TM segments are fairly loosely organized around a central cavity of 35 A diameter, which must be filled wi th mem brane Iipids in vivo to avoid loss of the permeability barrier. The central helices, TM4 and TM 10, contain the only three charged residues in tbe hydrophobic TM segments of each monomer, Asp407, Asp40S, and Lys940, which can form salt bridges and are assumed to be involved in proton translocation. The headpiece is divided into two layers: the upper TolC-docking domain and the lower pore domain. The TolC-docking domain opens like a funnel to a diameter of ~30 A, matching the diameter of the TolC tunnel (see below). Below it, the pore domain has a large central pore lined by three helices, one £1-om each subunit. In the pore domain each subuni t has four l3-a-13 subdomains that together form a large substrate-binding pocket. The central cavity of AcrB is enormous, measuring in total around 5000 A3 Between the subunits are three openings to the peri plasm, called vestibules. Substrates can thel-efore access the central cavity either from the periplasm or the bilayer for transport out the central pore (Figure 11.47). An Alternating Site Mechanism

Because the first crystals of AcrB were of the trigonal form, they confined the trimer to exact threefold

Drug Efflux Systems

11.46. X-ray structure of the drug efflux pump AerB. A. View of the AerB trimer from the side, oriented with the TM helices on the bottom and the peri plasmic headpiece on the top. Two of the three subunits (colored purple, green, and blue) are seen clearly from this angle. B. The AcrB trimer, viewed from the periplasm, reveals a peptide strand from each subunit that reaches far into the neighboring subunit. C. The view from the cytoplasm of the AerB homotrimer shows the large central cavity. From Murakami, S., and A. Yamaguchi, Curr Opin Struct Bioi. 2003, 13:443-452. © 2003 by Elsevier. Reprinted with permission from Elsevier.

301

Membrane Protein Assemblies

302

-}, MCIPC OCAC TC

_ 1'1' 11.47. Solvent-accessible surface of AcrB in a cutaway model. The front subunit has been removed, allowing a view into the central cavity and revealing pore helices (yellow) and TM helices that form grooves (green). Broken lines indicate the putative membrane boundaries and the framework of the funnel and cavity. Broken arrows indicate the postulated translocation pathways for substrates from the membrane, such as deoxycholate (DOC). acriflavine (AC), and tetracycline (TC). as well as substrates from the periplasm/outer leaflet, such as cloxacillin (MCIPC). From Murakami, S, and A. Yamaguchi, Curr Opin Struct Bioi. 2003, 13:443-452. © 2003 by Elsevier. Reprinted with permission from Elsevier.

to O. Drug extrusion is accomplished by the diffusion of substrate along a pathway that bulges and occludes as it migrates toward the funnel, much like the action of a peristaltic pump. The funnel at the top of AcrB fits the dimensions of the bottom of the TolC tunnel. and their interaction can be modeled to involve six hairpins at the top of AcrB docking with six helix-tum-helix structures at the bottom of ToIC. Although cross-links can be inserted between the two molecules, both with chemical crosslinking and with disulfide Formation between inserted Cys residues, unmodified TolC does not bind AcrB in vitro. In contrast, purified AcrA exhibits micromolar affinity for both AcrB and ToIC, suggesting that the docking between AcrB and TolC is reinforced by AcrA. The role of AcrA is likely to be quite dynamic given that the complex Formation is transient, aJlowing TolC to partner with other efflux transporters. AcrA. a Membrane Fusion Protein

In the tripartite drug efflux systems, membrane fusion proteins do not contribute part of the transport pores but are required to stabilize the interactions of the inner membrane transporter with the channel protein.

L Access

symmetry. Solution of the AcrB structure from crystals of different space groups that allowed asymmetry of the monomers revealed three different conformations affecting the access in the pore region. The three conformations are called loose (L, or "access"), tight (T, or "binding"), and open (0, or "extrusion"), and they vary in the binding pocket and pore regions (Figure 11.48). A variation in the position of the central pore helices is key to the conformational differences. The central helix From the 0 monomer is inclined nearly 15° to\-vard the T monomer, which results in opening the pore from the 0 monomer to the exit funnel while contracting the T monomer' to Form the drug-binding pocket (Figure 11.49). The lining of the pocket has eight phenylalanine residues (see Figure 11.48), aJlowing hydrophobic and aromatic interactions with drugs of diverse sizes and structures, as seen in the crystal structures of AcrB with several different drugs. The observation of three subunits in three conformations suggests an alLernating site, Functionalrotation mechanism analogous to that of the F1-ATPase (Figure 11.50). In the rotation, a monomer first binds substrate in the L conformation, binds it more tightly in the T conformation, and then extrudes it to the funnel in the 0 conformation. The subunits communicate via the central helices. The conformational changes are coupled to proton translocation in the TM domain: proton uptake is postulated to drive the change from 0 to L and proton release to accompany the change from T

T Binding 11.48. Illustration of the three conformations of subunits of the AcrB trimer. Each subunit is in a different conformation: the access state (L, green). extrusion state (0, red). and binding state (T, blue) shown in a cut view of the pore domain from the exterior of the trimer. The vestibules are clefts that are open in the Land T conformations and closed in the 0 conformation, which is open to the exit pore. Phenylalanine residues are shown in ball-andstick representation, and the binding pocket of the T conformation contains a molecule of the drug minocycline (orange contour labeled "Drug"). The arrows indicate the relative movements of the subdomains, which are labeled PC1, PC2, PN1, and PN2. From Murakami, S., et aI., Nature. 2006, 443: 173-179. © 2006. Reprinted by permission of Macmillan Publishers Ltd.

Drug Efflux Systems

303

11.49. Visualization of the tunnels in the pore domain of the AcrB peristaltic pump. In a ribbon diagram of the AcrB trimer viewed from the side, the tunnels are highlighted (green surfaces). A. The tunnel in the L monomer (blue) goes from the cleft halfway toward the central pore. B. The tunnel in the T monomer (yellow) extends across the pore domain toward the pore. C. The tunnel in the monomer (red) is closed at the lateral edge and opens into the exit funnel. From Seeger, M. A., et aI., Science. 2006,313:1295-1298. © 2006. Reprinted with permission from AAAS.

°

Members of the membrane fusion protein family attach to the inner membrane either via lipid acylation ofa cysteine residue or with an N-terminal TM segment. However, studies of mutants lacking these regions reveal that membrane attachment is not essential for the drug efflux function of these proteins. MexA, from the

B.

%

11.50. Schematic representation of the AcrB alternating site functional rotation transport mechanism. The three conformational states are loose (L, access; blue), tight (1, binding; yellow), and open (0, extrusion; red). They are shown as viewed from the side (A, with dotted lines denoting the membrane) and from the top (B). The side chains presumed to be involved in proton translocation (Asp407, Asp408, and Lys940) are indicated in the TM part of each monomer in A. An acridine molecule is depicted as substrate, which first binds to the L state, then binds tightly to the aromatic pocket in the T state, and then is extruded toward the funnel in the state. From Seeger, M. A., et aI., Science. 2006, 313:1295-1298. © 2006. Reprinted with permission from AAAS.

°

MexAB/OprM tripartite system of P. aeruginosa, was the first membrane fusion protein to have a high-resolution structure. AcrA is highly homologous to MexA, so it is not surprising that its structure proved to be very similar. AcrA Structure AcrA consists of 397 amino acids including a cleavable N-terminal signal, residues 1 to 24. Its C-terminal ~90 residues are very sensitive to proteolysis and correspond to the region of MexA that did not give clear electron density in its x-ray structure, indicating it is highly flexible. Therefore, this C-terminal domain, which is essential for interaction with AcrB and ToIC, was cleaved fTom the AcrA molecule for structural analysis of its 28-kDa stable core. To solve the x-ray structure, four additional methionines were substituted in the AcrA fragment, allowing incorporation ofselenometbionine for phase determination. The effect of the four methionine residues on the activity of intact AcrA is not entirely clear: mutants with two of the substitutions function to give drug resistance in a 6AcrA mutant, but with all four substitutions the intact protein is not sufficiently expressed to give the resistance phenotype. The 2.7 A resolution structure of the AcrA fragment shows residues 53 to 299 in an elongated sickle that consists of three domains: a I)-barrel domain, a central lipoyl domain, and a coiled coil (X-helical hairpin (Figure 11.51). Nearest the membrane, the I)-barrel domain includes both the Nand C termini of the fragment. It has six antiparalJeJ I)-strands, two of which close off the barrel near the C-terminal end (where the proteolyticaUy sensitive domain would follow in the wildtype protein). The lipoyl domain contains a I)-sandwich

Membrane Protein Assemblies

304

cd

a1 u-helil:barrel domain to a position alongside the peri plasmic domain of AcrE. This puts the coiled coil well into the peri plasm, where it can interact with TolC (see Frontispiece). While both AcrE and TolC are trimers, the oligomeric state of AcrA is not clear. In vilro both AcrA

and MexA, lacking membl-ane associations, are soluble monomers. In crystals, MexA formed six- and sevenmember rings while the AcrA Fragment Formed a dimer of dimers, which clearly does not represent physiological association. However, the crystal structure of AcrA is very informative because each of the Four monomers in the crystal has a different conformation due to a hinge at the base of the helical hairpin. The largest difference among the four shows a rotation of 15° (Figure 11.52). Interestingly, this hinge movement is due to variation in unwinding of the helices. It can be correlated with the proposed iris-like opening of the TolC channel (see below), but it is not clear whether it actively opens TolC or passively maintains the intermolecular contacts. TolC, the Channel-Tunnel

Drug efflux via the tripartite AcrAB/ToIC results in expelling the unwanted compounds into the outside medium because TolC provides a pathway acmss both the periplasm and the outer membrane. This is evident in the remarkable high-resolu tion structure of TolC (2.1 A resolution) that shows a homotrimer with a (:'>-barrel channel domain in the outer membrane that connects to a unique a-barrel in the peri plasm (Figure 11.53). Since the oc-barrel is 100 A in length, the sum of the

Drug Efflux Systems

305

_ 15° ~ Mol DMol A M () I B Mol C

a2

a1

al

ep

90"

11.52. Four conformations of AcrA observed in the crystal structure. The crystal form captured each monomer (labeled MolA, MoIB, MoIC, and MolD) in a different conformation due to the flexible hinge between the helical hairpin and the lipoyl domain. Superposition of the four conformations reveals a 15° rotation between the most different structures. From Mikolosko, J., et aI., Structure. 2006, 14:577-587. © 2006 by Elsevier. Reprinted with permission from Elsevier.

peri plasmic length of TolC and AcrB is 170 A, which is long enough to span the periplasm. TolC provides an uninterrupted pathway with an inner diameter of at least 20 A fTom the inner membrane transporter to the medium. Passage through the TolC channel-tunnel is passive, as the inner membrane transport partners carry out the energized step.

Extracellular

-40

A

-100

A

A Unique Structure

The TalC channel is an anti parallel 12-stranded I)-barrel with a right-handed twist. Aromatic residues delineate the regions located in the lipid bilayer interface (see Chapter 5). As the case for other outer membrane proteins, the external loops of TalC are sites for colicin and bacteriophage attachment. However, TolC differs from other outer membrane I)-barrels in significant ways. Rather than having each polypeptide form a barrel, each TolC subunit conttibutes four I)-strands to its single pore. With neither a constriction formed by inward folded loops nor a plug formed by a separate domain, the pore is constitutively open. Finally, the I)-barrel is connected directly to the ex-barrel, 'which has a lefthanded twist, requiring interdomain linkers with conserved proline residues that make abrupt turns (Figure 11.54). The peri plasmic tunnel of TolC is also 12-stranded, with ex-helices packed uniformly in an anti parallel lefthanded superhelical twist. Each subunit contributes two long helices (H3 and H7) and two pairs of shorter helices (H2/H4 and H6/H8) stacking end-to-end to span the length of the barrel (Figure 11.54B). Three short

Periplasm

Entrance 11.53. High-resolution structure of the TolC channel-tunnel. TolC is a homotrimer (each subunit is a different color) that spans the outer membrane with a l3-barrel channel and extends into the peri plasm with an ex-barrel. See text for details. From Koronakis, v., et ai, Annu Rev Biochem. 2004, 73:467-489. © 2004 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

Membrane Protein Assemblies

306

Closed

Open

---J domain of the F 1 Fa-ATPase. The recent high-resolution structures of AcrB in crystal forms that allowed asymmetry in the trimer also show the subunits in different conFormations, pointing to an alternating-site rotation model analogous to that of the F\-ATPase. A typical approach to obtain different conformations is to carry out crystallization in the presence of substrate analogs or inhibitors. This approach has been especially successful with the sarcoplasmic reticulum calcium ATPase. Comparison of the structures of Ca 2+ -ATPase with and without calcium ions shows large movements of its three domains. Additional conformations have been obtained with AIF 4 -ADP as an ATP analog and with AlF4" and the inhibitor thapsigargin, which stabilizes the calcium-Free forms. As described in Chapter J 0, a nearly complete reaction cycle can be envisioned For this complex ATP-dependent pump based on high-resolution structures capturing its different conformations. Detailed comparisons of structures in di [ferent conformations clarify the specific sources of the structural changes. For example, AcrA is hinged at varying angles between the lipoyl domain and the long coiled coil, and the hinge angle depends on the unwinding of an

Motifs and Patterns

ex-helix at the base of the coiled coil. For the helicalbundle membrane proteins in general, conformational changes arise from bending of helices, rotation of helices, unwinding of helices, and/or loss of secondary structure creating short loops. These can result in quite large structural changes, such as hinging between domains and opening and closing of channels. Some membrane proteins have been crystallized only in the presence of a substrate analog or inhibitor to push the equilibrium to favor one conformation. This was the case for the transporters LacY and mitochondrial AAC, which are expected to undergo large conformational changes to accomplish substrate translocation. In contrast, addition of substrate or inhibitor to other membrane proteins has little effect on their overall conformation. These proteins often lack ,",veil-defined substrate-binding sites, having instead large internal pockets or cavities. The large binding pocket of AcrB can accommodate a variety of drugs that interact with different subsets of residues without triggering large stnlctural changes. Another example is the large cavity of the BtuC transporter that lacks a specific binding site and relies on BtuF for the specific delivery of their substrate, vitamin 8 12 . Also binding of inhibitors need not always cause large conformational changes but can instead give important insight into mechanisms of inhibition, as observed with cytochrome-bel complex and prostaglandin H 2 synthase. Even when structures of different conformations are lacking, the information from x-ray structures can contribute to understanding conformational changes. The B values (temperature factors) of the crystal structures show regions with high Aexibility that implies conformational freedom. Simulations based on one highresolution structure can also pick out flexible regions and then predict other conformations. Often features of a high-resolution structure lead to predictions for how the conformation is likely to change. For example, the bends in the basket helices of mitochondrial MC suggest these are Aexible sites where alternate conformations enable the carrier to open to the other side of the membrane. Another example is the proposal that the loop in the backbone or the SecY translocon functions as a hinge between the two halves of the heterotrimer, suggesting how the opposite side might open and allow peptides to move laterally into the bilayer. A different basis for prediction of conformational changes is the comparison of structures of related proteins, such as the potassium channels. The KcsA, KirBacl.J, and KirBac1.3 structures show the closed channels, whereas MthK, KvAP, and KvJ.2 show the open channels. In other cases EM images can indicate what a different conformational state might look like. For example, both LacY and GlpT x-ray structures show the transporters open to the cytoplasm, while a lower-resolution EM structure shows the related car-

311

rier OxlT in a state open to the peri plasm. In addition, evidence for the structure of LacY in the conformation open to the peri plasm comes from studies of Cys replacement mutants that determined reactivity to N-ethylmaleimide as well as cross-linking. Future progress in membrane structural biology will certainly include elucidation of x-ray structures in different conformations from those available today. In some cases these will be key to understanding the mechanism of the proteins, such as the TonBdependent outer membrane transporters whose channels are blocked in the structures now available. A significant achievement will be determination of the specific conformations of activated states, for example, of visual rhodopsin whose available x-ray structure portrays the inactive ground state. The solved structures of intermediates in the bacteriorhodopsin photocycle, trapped by different conditions at very low temperatures and/or by mutations, provide a hint of what can be achieved in future work on rhodopsin and other GPCRs.

MOTIFS AND PATTERNS

Recurring patterns in the structures of membrane proteins can involve general features or fine details. Only a few membrane protein structures with resolution around 2 A are now available. For these structures the ability to determine positions of amino acid side chains and of water molecules reveals the importance of hydrogen-bonding networks to both structure and f-unction. Interhelical hydrogen-bonding networks in rhodopsin include many highly conserved residues. Since genetic data indicate that some of these residues are essential for activation of rhodopsin, it is likely that conformational changes that accompany activation will produce different hydrogen-bonding networks. In the mitochondrial MC the hydrogen bond networks link all but one of the TM helices and thus must be altered to allow the outward-facing conformation to change to the inward-facing conformation. The hypothesized trigger for this change is perturbation by the negative charges on ATP of a particular salt bridge at the base of the hydrogen-bonding network. Important networks of interhelical and even intersubunit hydrogen bonds can be expected to be revealed in other structures as they are obtained at higher resolutions. Some general structural features that can be observed at lower resolution show that many helical membrane proteins deviate [Tom the now-classical model of bacteriorhodopsin with its fairly uniform serpentine structure. Helix irregularity would seem to be more the rule than the exception, as almost every description of the structure of a helical-bundle membrane protein reports on variations in helix lengths, tilt

Themes and Future Directions

312

angles, and bends. For example, four of the TM helices of the sarcoplasmic reticulum Ca 2+ -ATPase extend well past the membrane bilayer into the cytoplasmic region of the protein. Even visuali-hodopsin has a 33-residuelong TM helix. In addition to some very long helices, the presence of half-helices, as seen in aquaporins, certainly makes hydropathy predictions inadequate. The recently solved structure of the CIC chloride channel has been called "a jumble of helices." In view of such irregularities, specific helix-helix interactions are important in stabilizing helix packing. These are typically knob-into-hole nonpolar interactions, which account for the frequent occurrence of the GxxxG motif in the helices. While the character of most TM helices is predominantly hydrophobic, a surprising number of charged residues occur in some membrane domains. Notably the KvAP potassium channel has four arginine residues in its otherwise nonpolar voltage-sensor paddle. Another deviation h-om the serpentine structure is the protrusion of some mem brane proteins well beyond the membrane. Examples h-om the respiratory membrane include the FIFo-ATPase, cytochrome-bcl complex and cytochrome-c oxidase, all of which protrude significantly (h-om 32 to 75 A) on both sides of the membrane. Formate dehydrogenase in the E. coli nitrate respit-atory pathway has peripheral subunits that extend 90 A past the bilayer. The peri plasmic head of the AcrB drug efflux transporter is 70 Along, and the tube of the TolC channel-tunnel protrudes 100 Ainto the peri plasm. These extensions beyond the bilayer enlarge the depiction of membrane proteins as described at the end of Chapter 1. Symmetry in helical membrane proteins has been detected both in their amino acid sequences and, in cases of lower homology at the primary level, in their overall folds. While some have three repeats and threefold symmetry, as seen in the mitochondrial carriers, it is more common for membrane proteins to exhibit twofold symmetry. The sequences of aquaporins contain two homologous halves, and the twofold symmetry is apparent in their structure, in which each half folds into three and a half TM helices with a junction between the two half-helices in the center of the membrane. Twofold pseudo-symmetry is .-evealed in the xray structures of LacY and GlpT, with each consisting of two linked bundles of six ex-helices. Although the two halves of these transporters have low-sequence homology, bioinfo.-matics programs detect this pattern in most other members of the Major Facilitator Superfamily. The first ABC transporter to have symmetry noted in the two halves of its structure is the Savl866 drug exporter. Yet another example is the Sec translocon, whose major channel is formed by two pseudosymmetric halves of the SecY subunit (Sec61p in yeast and SecYex in archaea). Apparently a common occur-

rence III the evolution of helical membrane proteins has been gene duplication followed by fusion of the two genes. Interestingly, in some cases (e.g., the aquaporins) the repeated domains fused in inverted orientations. A precursor to this evolutionary step would be the association of two monomers with opposite orientations, as observed in the crystal structure of the drug efflux protein EmrE. Even though the physiological relevance of the asymmetric dimer of EmrE is controversial today, the fact that mutations can encourage this asymmetry suggests that EmrE could represent an early stage in the evolution of the proteins that contain inverted repeats. Symmetric folds and structural motifs often contain characteristic signature sequences that al-e usually first identified as highly consented residues among related or homologous proteins. The powerful tools of genetics and bioinformatics can be expected to identify more motifs in the many membrane proteins about which very little is known, and this in turn will provide critical information about their functions.

CONCLUSIONS

In spite of the rich diversity of their functions, it is not surprising that membrane proteins exhibit some structural themes, given the const.-aints of their lipid bilayer environment and of their mechanisms of biogenesis. As the number of high-resolution structures grows, more shared traits will be recognized. In addition, more specific interactions with lipids will likely be revealed, serving as a reminder of the complexity of the molecules that surround membrane proteins. Much more than a simple hydrophobic barrier, the heterogeneity of the lipid bilayer means that even the relative position of an amino acid in a TM helix affects the ability of a peptide to be a stable TM segment. Thus much more will be learned from ['urther studies of those proteins able to insert into the lipid bilayer, such as diphtheria toxin. Progress in the characterization of assemblies of proteins that form complexes, even nanomachines, in the membrane allows even more elaborate processes to be tackled at the molecular level. Topics such as intracellular trafficking, chemotaxis and flagellar movement, virus infection, and membrane fusion have long provided fertile areas for research. Now it is feasible to target the membrane proteins mediating such phenomenon for study by the methods of structural biology. Of course while membrane structural biology is making great advances, it continues to benefit fTom advances in membrane biochemistry, biophysics, genetics, bioinformatics, and computational biology. These productive methodologies are working together to solve a major riddle of life, as the very definition of a cell depends on how the membrane

Conclusions determines what it takes in and sends out and how it responds to the outside environment. And while many of the membrane proteins that comprise up to one third of the human genome are not well characterized, already there are many examples of membrane proteins involved in the etiology of diseases and a large majority of current drugs target membrane proteins. Therefore the benefits of increased knowledge of the structural biology of membranes will undoubtedly reach beyond advances in basic research to have an impact on human medicine and health.

313

FOR FURTHER STUDY

(multi-author) Nobel symposium on membrane proteins: structure, hmction and assembly. FEES Leu. 2003, 555. (multi-author) Insight on membranes. Nature. 2005, 438:587. (multi-author) Special section on crossing the bilayer. Science. 2005, 310: 1451. Pornillos, 0., and G. Chang, Inverted repeat domains in membrane proteins. FEBS Leu. 2006. 580:358-362. White S., et aI., Lipid bilayers, translocons and the shaping of polypeptide structure, in L. K. Tamm (ed.), Prolein-Lipid Interactions. New York: Wiley-VCH, 2005, pp. 3-25.

Appendix I

ABBREVIATIONS

AAC: ABC LnJnSporters: AChR: AFM: ANK: AQP: ATR: BA: BCl: BO: BPh: BR: CATR: CFTR: CHAPS: CHAPSO: CL:

CMC: CMT: CN-cbl: CT: CTAB: DAG: DAGK: DDM: DEPC: DI-IA: DHPC: DLPC: DLPE: DMB:

ex-hemolysi n f'-ee energy change for the transfer of a substance From one solvent to another ADP/ATP carrier a large class of transporters named for their ~TP-~inding

i::assettes acetyl choline receptor atomic Force microscopy repeated domain of ankyrins aquaporin atl-actyloside, an inhibitor of the mitochondriol ATP/ADP carrielbongkrekic acid, an inhibitor of the mitochondrial ATP/ADP carrier bacteriochlorophyll bacterio-opsin. which locks the retinal cofactor bacteriopheopyt in bacteriodlodopsi n carboxyatractyloside, an inhibitor of the mitochondrial ATP/ADP carrier cystic fibrosis tronsmembrane conductance regulator, a chloride channel 3-[3-(chola midopropvl) dimethyl-ammonioJ-l-propanesulfonate 3-([3-cholamidopropyl]dimethylommonio)-2hvdroxy-I-propanesulfonate cardiolipin critical micelbr concentration critical micellar temperature cyanocobalamin, or vitamin B I 2 cytidyl transferase cetyltrimethylammonium bromide diocylglycerol diacylglycerol kinase - but DGK in chapter 6 dodecyl I3-D-maltoside dieleidoyl phosphatidyJcholine docosahexaenoic ocid dihexanoyl phosphatidycholine dilauroyl phosphotidylcholine dilauroyl phosphatidylethanolamine 2,3-d imethvl-benzim idazole

DMPC: DMPE: DMPG: DMPS: DOPC: DOPE: DPPC: DPPE: DRMs: DSC: DSPC: DT: EDTA: EF: EGF: EM: EPR: ER: FDI-1: FRAP: FRET: FTIR: G3P: GdmCl: GOF: GPCR: GPl: GUVs: HDL: HHDBT:

HiPIP: HMM: HPr: ICL:

IMS: Ko: KcI:

dimyristoyl dimy,-istoyl dimyristoyl dimyristoyl

phosphatidylcholine phosphatidylethanolamine phosphatidylglycerol phosphatidylserine

dioleoyl phosphatidylcholine dioleoyl phosphatidylethanolamine dipalmitoyl phosphatidylcholine dipalmitoyl phosphatidyJethanolamine detel-gent-resistant membranes differential scanning calorimeu-y d iSlearovl phospha tidylchol ine diphtheria toxin ethylenediamine tetraacetic acid edema Factor. a component of anthrax toxin epidermal growth factor electron microscopy electron pa,-amagnet ic resonance endoplasmic reticulum Formate dehydrogenase fluorescence recovery aFter photobleaching fluorescence resonance energy transfer Fourie,- t1-onsForm infTared spectroscopy glycerol-3-phosphate guanidinium chloride gain of Function G-protein coupled receptor glycosylphosphatidylinositol giant unilamellar vesicles high-density lipoprotein 5-/7- heptyl-6- hydroxy-4, 7-diozobenwth iowle, an inhibitor of the cytochrome-be I complex hexagonal phase with nonpolar centers and polar groups and water outside inverted hexagonal phose (with polar centers and nonpolar exteriors) high-potential iron-sulfur protein hidden Markov model histidine-containing phosphocarrier protein intracellular loop intermembrane space of mitochondria binding constant dissociation constant

315

Appendix I

316

Kr:

lipid association constant describing selective

PEP PTS:

phosphoenolpyruvate-dependent

PG:

phosphotransferase system phosphatidylglycerol

PG H 2: PGHS:

prostaglandin H 2 prostaglandin H 2 synthase (also called

retention of lipids by pmteins)

L13,: LI3:

lamellar gel in which the chains are tilted lamellm- gel, also called So (ordered solid)

L~:

lamellar liquid crystalline, also called Lei (or Id,

Lc : LC:

lamellar crystalline

PH:

liquid condensed, a condensed phase in a lipid monolayer

pleckstrin homology domain that binds phosphoi nosi tol

PI:

phosphatidylinositol

LDAO:

lauryldimethyJamine oxide, or

pKa :

pH at which an acidic or basic hmctional group

LE:

dodecyldimethylamineoxide liquid expanded, an expanded phase in a lipid

PKC:

is 50% protonated protein kinase C

monolayer

PL:

phospholipid, glycerophospholipid

Lep:

leader peptidase

phospholipase A2

LF: LHJ, LH2:

PL A2: POPC:

light-hal\lesting complexes

liquid-disordered)

Lo :

lethal factor, a component of anthrax toxin liquid-ordered lipid state that occurs when PLs, sterols, and/or sphingomylins are present

cyclooxygenase, COX)

POPG: POX: PrP:

I-palmitoyl-2-0Ieoyl phosphatidylchoJine I-palmitoyl-2-0Ieoyl phosphatidylglycerol peroxidase prion protein

loss of function

PS:

phosphatidylserine

R:

MBD:

large unilamellar vesicles membrane-binding domain

the radius of curvature of the lipid/water interface

MC:

Monte Carlo

RC:

reaction center (photosynthetic)

MCF:

mitochondrial carrier family

RCK:

regulators of conductance of K+

MD: MDOs:

molecular dynamics

RND:

Resistance Nodulation cell Division, a

MDR:

membrane-derived oligosacchal-ides multidrug resistance

Ro :

superfamily of drug efnux proteins the inu-insic value of R for each lipid species

MFS: MLVs:

major facilitatOl- superfamily

S:

multilamellar vesicles

LOF: LUVs:

MPoPS:

I-myristyl 2-palmitoleoyl phosphatidylserine menaquinone

shape parameter for lipids, lipid volume/(cross-sectional area of polar head-group x lipid length)

SDS:

sodium dodecylsul fate, also called sodium

membrane scaffold proteins

SOSL:

site-directed spin labeling

N:

Newton, a unit of force

SOS-PAGE: SDS polyacrylamide electrophoresis

NBD: NBD-

nucleotide-binding domain cholestemllabeled with the

SERCA: SLUVs:

sarcoplasmic reticulum Ca2+ -ATPase short-chain/long-chain unilamellar vesicles

SM: SMS:

sphingomyelin an analog of somatostatin that is an

sn:

amphiphilic peptide with a positive charge stereochemical numbering

MQ:

MS: MSPs:

mechanosensitive

cholesterol: nitrobenzoxadia7.0lyl group NBD-DLPE: DLPE (a PL) labeled with the

laurylsulfate

nitrobenzoxadiazolyl group NMR:

nuclear magnetic resonance nuclear Overhauser effect (in NMR)

SOPC: SOPE:

J-steaml-2-0Ieoyl phosphatidylcholine

octyl-polyoxyethylene octyl f)-D-glucoside (sometimes put as f)OG)

SRP:

signal recognition particle

STGA:

outer membrane phospholipase A

3-HSOrGal pf)I-6Manpal-2Glcpal-archaeol small unilamellar vesicles

oligomycin-sensitivity conferring protein, a

SUVs: TBDTs:

TonB-dependent transporters

subunit of ATP synthase

TC:

transport classification

PA:

phosphatidic acid

TOG:

f)-D-galactopyranosyl

PM:

protective antigen, a component of anthrax TEMPO:

PC:

toxin phosphatidylcholine

a small lipid-soluble spin probe whose nitroxide group has an unpaired electron

PCC:

protein-conducting channel, also called the

TIM:

transJocatase across the inner mitochondrial

TU-I:

transition temperature for lipid transitions

NOE: NSAlDs: octyl-POE: OG: OMPLA: OSCP:

nonsleroidal anti-inflammatory drugs

I-thio-f)-D-galactopyranoside

translocon PCR:

polymerase chain reaction

PDB: PE:

Protein Data Bank phosphatidylethanolamine

J -stearol-2-oleoyl phosphatidylethanolamine

membrane from lamellar to hexagonal phases, typically L~-HII

Appendix I TO':

TM: TNBS:

317

melting temperature for fatty acids and pure lipids; also lIsed as transition temperature for lipid transitions from Lp to L" transmembrane tr-initrobenzenesulfonic acid

TOM: TRAM: TROSY:

translocase across the outer mitochondrial membrane translocating chain-associated membrane protein transverse relaxation optimized spectroscopy (in NMR)

Appendix II

SINGLE LETTER CODES FOR AMINO ACIDS A, C. D. E. F, G,

alanine cvsteine asparLate glutamate phenylalanine glycine H, histidine I. isoleucine K. lysine L, leucine

318

M. methionine N, asparagine p. proline Q. glutamine R, arginine S. serine T. threonine V. valine W. tryptophan Y. tyrosine

Index

I-palm itoyl-2-oJeoyl phosphatidylcholine (POPC) in lipid rafts, 34, 35 MD simulation of, 200, 220 2-heptyIA-hydroxyquinoline-N-oxide (HOQNO), 226 AAe. See ADP/ATP carrier (AAC) AB toxin model, 84 ABC transporters, 133-135, 136, 290-291. See also specific conformational changes in, 292-293, 294 nucleotide-binding domains of, 135, 136,292,293,297 structure of, 290 symmetry in, 312 acetylcholine, 139 acetylcholine receptors (ACchRs) muscarinic, 139 nicotinic, 139-140 patch clamp recordings of. 57, 58 AcrA, 302-304, 307 conformational Oexibility, 304, 305 slruclure of. 300, 303-304, 310 AcrB, 297, 300-302 alternating site mechanism, 300-302, 303 protrusion beyond membrane, 312 struclure of. 300, 301, 302, 310 active transporlers, 131, 132 primary, 133-134 secondary, 137-138 acyl chains, 13-17 addition or removal of, protein binding regulated by, 83 composition of, 16 in MD simulations, 200, 202 length of, in lipid-protein interactions, 98 names for, IS, 17 oleoyl-, 15 polyunsaturated, 15 stearyl-, 15,200 adhesion protein, 114 ADP/ATP carrier (AAC), 138,242, 249-250

ADP/ATP carrier (AAC) basket, 250, 251 binding in, 250-251, 252 conformational changes in, 252, 311 dimer formation in, 252, 253, 310 hydrogen bonding in, 251, 311 MCF motif in, 250, 252 structure of, 250 transport mechanism, 249, 252-253 adsorption, detergent removal by, 50 aerolysin, 85 Aeromonas, 85 Aeropyrul11 pemix, 259 aggregation number (N), 46, 48 Agre, Peter, 207, 241 helical proteins channels, 132-133 folding studies of, 162-164 helices, 102 distortion of, in transmembrane segments, 92-93 in MS channels, 239 location of. 113 membrane-spanning, amino acids in, 92 hemolysin (aHL), 85, 86, US alamethicin, 87, 88, 132-133 alkaline phosphatase (PhoA), 144, 145, 146 AMBER,198 amino acid distribution genomic analysis of, 153 in integral proteins, 145 amino acid(s) codes for, 78 hydrophobicity scales for, 142, 143, 182 in transmembrane domains, 92 positions of, 154 snorkeling of, 184 amphipathic helix in fJ-barrels, 114 insertion into interfacial region, 76, 82 amphiphile shape hypothesis, 29-30, 31 amphiphile(s) spontaneous assembly of, 4-5 tripod, 45

amphiphilicity,4 micelle formation and, 47 of fJ-barrels, 157 amphipols,45 amphitropic proteins, 68, 72-73 membrane binding by, 76 anaerobic respirat'ion, 222, 224 angiotensin receptor, 235 anion exchangers, 138 ANK repeats, structure of, 70 ankyrin, 70 deficiency, 70 almexins, 72, 83 annulus, 8, 94. See also boundary (annular) lipids antennae, 111-112,310 anthrax toxin, 85 anti torsion angle, 15, 16 antibiotics microbial resistance to, 296 porins and, 118 antibody fragments, proteins bound to, for crystallization, 208, 209 antimicrobial peptides, 87 antimycin, 281 anti port, 138 antiporters, 138 ApoAI, nanodiscs, 66 apolipoproteins,73 apparent free energy (DGapp), 182 aquaglyceroporins, 255-258 selectivity of, 254, 258 aquaporin I-null humans, 253, 254 aquaporins (AQPs), 253-254 physiological roles of, 253, 254 proton transfer in, 256-258 selectivity of, 253-254, 258 structure of, 254-255 symmetry in, 312 arachidonic acid, 220, 221 archaebacteria phospholipids in, 18, 20 translocons in, 286 archaeol, 20, 104 area vs. surface pressure isotherm, in monolayer formation, 51 aromatic amino acids, 92, 114

Index

320

arrestin, 230 artifacts, in x-ray crystallography, 209 Astral compendium, 142 atomic force microscopy, of lipid rafts, 34,35 AT? generation, 272 catalytic mechanism, 277, 278, 279 regulation of, 276 ATP synthases, 272. See a/so F 1 Fa-ATPase/ATP synthase catalytic sites, 277 composition of, 271-272, 273 equivalent subunits, 273 MD simulations of, 278 role in respiratory chain, 279 ATPase subdomain, of ABC transporters, 135 ATPases, 133-134 A-type, 134 F-type, 134 superfamilies of, 134 V-type, 134 ATP-binding cassettes. See ABC transporters ATP-sandwich, 135 atractyloside (ATR), 249 avenacins, 55 avidin-biotin binding, in potassium channels, 263 AxxxA motif, 154 B values (temperature factor), 207, 311 Bacillus subti/is, 170 bacteria. See also specific species anaerobic respiration, 222, 224 cell envelope structure, 113 drug efflux, 300 group translocation, 135, 137 multidrug resistance, 297 protein export, 170, 172, 176 topogenesis, 185 translocation apparatus, 175,286 bacterial membranes, I, 2 blebs in, 64, 65 fatty acid composition of, 17 lipid diversity in, 37, 39 lipid-anchored proteins in, 75 phase transitions in, 16,40 phospholipids in, 18, 20, 39 protein-rich domains in, 8 stnlctural support in, 10 bacteriochlorophyll (BChl), 109, Ill, 112 bacteriocins, 85 bacterio-opsin (BO), 166 bacteriopheopytins (BPh), 109 bacteriorhodopsin (BR), 8,45, [03, 166

bacteriorhodopsin (BR) crystallization of, 209, 210 helical bundles in, 102-107 helix-helix interactions in, 154 hydmphobicity plots fo.·, 144 integral proteins in, 92, 93 mutant proteins from, 105 nanodiscs,66 photoreaction cycle in, 105, 107 protein-folding studies in, 164, 166-167 proton path in, 106, 285 structure of, 103, 104,105,106,203, 204,210 Band 3 protein, 138, 186 barrel-stave model, 88 Basic Local Alignment Search Tool (BLAST), 142, 157 fl-barrels, 102, 113-118,213,214 amino acid residues in, 114, 119, 121 composition of, 114 enzymes with, 214 families of, 157 genomic analysis of, 157-158 hydrogen bonding, 114 hydrophobic mismatching in, 99 in porins, 113, 119, 120, 121, 132-133,157 in toxins, 85 location of, 113,167-168 number of fl-strands in, 114 pmtein folding in, 167-169 structure of, 115, 116, 210 transmembrane segments in, 96 BBF (Beta-Barrel Finder), 157 BChl (bacteriochlorophyll), 109, Ill, 112 fl-cyclodextrin, 34, 36 fl-D-galactopyranosyl I-thio-fl-o-galactopyranoside (TDG),243 bee venom, 87, 88 Beerendsen, H. J. c., 197 Benson, A. A., 5, 6 Beta-Barrel Finder (BBF), 157 fl-galactosidase (LacZ), 144 fl-hydroxybutyrate dehydrogenase, 50 bicelles,63 crystallization and, 209, 210 bile salts, 43, 48 binding ligands, 79, 83, 234 binding sites ATP synthase, 277 BtuB,295 calcium ATPase, 264-265, 267 conformational changes and, 311 cytochrome-bcl complex, 281,282 GlpT, 245, 246

GPCRs, 234 KcsA,260 LacY, 243 reaction centers, 112 binding-change mechanism, 277 bioinformatics tools for protein families, 141, 142 motifs and patterns studied with, 312 biotin-avidin binding, in potassium channels, 263 bitopic proteins, 69, 90, 184 Bla (fl-lactamase), 144 black films. See planar bilayers fl-Iactamase (Bla), 144 BLAST (Basic Local Alignment Search Tool), 142, 157 B/aslOch/oris viridis antennae, 111 reaction centers, 107, 108, 109, 110 blebs, 63-64, 66 blisters, 63-65, 66 blood clotting factors, 73 fJ-octylglucoside, 45, 50 bond angle energy, 199 bond length, function for, 198 botulinum neurotoxin, 85 boundary (annular) lipids, 50, 94, 204 EPR of, 94, 95, 96, 98 exchange with bulk layer; 95, 98 numbers of, 205 structure of, 205 Boyer binding-change mechanism, 277 Boyer, Paul, 272 BPh (bacteriopheopytins), 109 BR. See bacteriorhodopsin (BR) Bragg's law, 193, 194, 195 BtuB conformational change, 295 function of, 296 in outer membrane, 294-296 structure of, 295, 296 BtuBC (vitamin Bil transporter), 135 BtuBCDF, 272 BtuC, 311 BtuC2D2F complex, 292, 293, 294 BtuCD complex, 290 conformational changes, 293-294 in inner membrane, 292-294 BtuD,293 BtuF, 291, 292 bulk lipids, 94, 96 exchange with annular· layer, 95, 98 CI domain, 80, 81 C2 domain, 80, 81

Index Caenorhabdilis elegans, J 53 calcium ATPase, 264 binding sites, 264-265, 267 conformational change in, 264-267, 268, 269 E 1-E2 reaction scheme, 264, 269 structure of, 3 J0 x-ray crystallography of, 209-210 calcium binding, 83, 84 by calcium ATPase, 264-265, 267 dephosphorylation step in, 267 to OMPlA, 214, 216, 217 calcium channels, voltage-gated, 264 calcium pump, 264 calcium, in signaling, 264 carboxyatractyloside (CATR), 250 cardiolipin (CL), 17,38,40 in reaction centers, 109 mixed micelles containing, 63 structure of, 205. 206 cardiolipin/K pathway, 283, 284-286 carotenoids. in reaction centers, 109 Carpet mechanism, 87, 88 carriers, 133 catalytic domain. of prostaglandin Hz synthase, 217, 220, 221 catalytic mechanism, FiFo-ATPase, 277-278. 279 catalytic triad. in OMPlA, 2J4, 215 CATH (Class, Architecture, Topology and Homologus superfamily), 142 CATR (carboxyatractyloside), 250 caveolae, 36 formation of. 37, 38,90 caveolins,90 cell envelope, bacterial, 113 cell surface receptors, in supported bilayers, 57 cerebrosides, 20 CF (cystic fibrosis). 187, 188 CFTR (cystic fibrosis transmembrane conductance regulator), 135, 186 misfoJding of, 187, 188 channel(s). 253. See also specific channel in translocons, 287. 288 channel-forming toxins, 132-133 chaperones, J73. 176 CHAPS (3-[3-(cholamidopropyl) dimethyl-ammonioJ-lpropanesulfonate), 43 charge mutations, 186 CHARMM,198

321

cholate. 50 cholera toxin, 85 cholesterol, 20, 21 in lipid raf1s, 34, 35 in supported bilayers. 57. 59 in ternary phase diagrams. 32, 33 interactions with otber lipids, 22, 23, 33 MD simulation of, 201,202 proportion of, 8 solubility of, 22-23 thickening by, 22, 27, 100 chromophores, III circular dichroism. 161, 166 cis double bonds, 15,200 CL. See cardiolipin (Cl) Class, Architecture, Topology and Homologus superfamily (CATH), 142 CIC chloride channel. 312 cleavable signal sequences. 171. 172, 184, 185 cloud point. See Krafu point CMC (critical micellar concentration), 46, 47 CMT (critical micellar temperature), 47 cobra venom phospholipase Az, 128 cofactors, in photosynthetic reaction centers, 109-110 colic ins, 85-86, 87, 132-133 conduction pores, 253 cone cells, 228, 229 conformational changes, 310-311. See a.lso specific protein CONPRED, 152 cotranslational translocation, 170, 176 coupling helices. in Sav 1866, 298-299 COX (cycJooxygenase), 216, 218, 220, 221 critical micellar concentration (CMC), 46,47 critical micellar temperature (CMT), 47 cross-linking studies of ATP synthase, 277 of protein insertion, J 78, J 79, 182 crystal packing, 207, 209 crystallography. See a.lso x-ray crystallography process of, 207-210 CT (cytidyltransferase), 82 cubic phase, 28, 31, 207 crystallization in, 207, 209, 210 types of, 31 curvature frustration, 30, 31,40 curvature stress, in protein folding, 167,169 cyanocobalamin. See vitamin BIZ

cyclooxygenase (COX), 216, 218, 220, 221 cystic fibrosis (CF), 187, 188 cystic fibrosis transmembrane conductance regulator (CFTR), 135,186 cystic fibrosis transmembrane conductance regulator (CFTR) misfolding of, 187,188 cytidyltransferase (CT), 82 cytochrome b, 279 cytochrome-bc) complex, 154, 279-284 integral lipids in, 206, 208 proton pathways. 283, 284-285, 286 protrusion beyond membrane, 312 Q cycle, 281, 282 stTl.lctLll-e of. 279, 280, 281-282, 284 cytochrome-bcl oxidase, 271 cytochrome-c oxidase, 154, 284-286 integral lipids in, 206 oxygen reduction, 285 proton pathways, 283, 284, 285-286 protrusion beyond membrane, 312 structure of, 284, 285 cytochrome c, binding by, 69, 76, 77, 78 cytochromeci,281 cytochrome cz, 112 cytochrome oxidase, crystallization of, 208, 209 cytochrome P450, crystallization of, 209 cytochrome P450 reductase, nanodiscs, 66 cytochrome subunit, of reaction center. 110 cytoplasm. protein export from, 170-172.174,175,177 cytoplasmic domain, in rhodopsin, 234 cytoskeleton, of plasma membrane, 10 DAG (diacylglycerol), 73 DGK. See diacylglycerol kinase (DGK) Danielli, James,S Database of Interacting Proteins (DIP), 142 Davson, Hugh,S DDM (dodecyl ,B-D-maltoside), 43 Debye-HuckeJ theory, 79 Debye-Waller factor. See B values (temperature factor) decyl maltoside, 63 Deisenhofer, Johann, 107, 207 denaturation of proteins, 161 deoxycholate. 50 DEPC, phase diagram for, 32 detergent(s), 42,43. See also specific detergent

Index detergent(s) alternatives to, 45 bicelle formation, 63, 209, 210 crystallization and, 206, 208 definition of, 43 impurities in, 43 in protein-folding studies, 170 ionic, 43 lipid removal by, 50,206 mechanism of action, 45-48 membrane solubilization by, 48-50, 129 concentration required for, 48, 49 nonionic,43 porins solubilized with, 119 removal of, 50 nanodiscs produced by, 66 solubility of, 43 types of, 43-44, 45, 46 zwitterionic,43 detergent-resistant membranes (ORMs), 36-37, 73 GPI-anchored proteins in, 75 proteins in, 36, 37 DHA (docosahexaenoic acid), 201, 202 DHPC (dihexanoyl PC), 167 diacylglycerol (DAG), 73 diacylglycerol kinase (DGK), 129-130 kinetic analysis of, 63 mutations, 129 protein folding in, 169 structure of, 129, 130 differential scanning calorimetry (OSC) of fatty acid phase transitions, 15, 17, 29 of lipid phase transitions, IS, 17,29, 76, 77 of nanodiscs, 66 diffraction pattern, 193 diffusion of lipid bilayer, 24-25 rate of, 25 of membrane components, 7, 8 digitalis, 134 digitonin, 55 dihexanoyl PC (DHPC), 167 dilauroyl phosphatidylcholine (DLPC), 100 dimer formation, 271,309-310 mitochondrial carriers, 252, 253, 310 OMPLA, 215, 216, 310 dlodopsin, 234 translocons, 288 dimyristoyl phosphatidylcholine. See DMPC (dimyristoyl phosphatidylcholine)

dioleoyl phosphatidylchoJine (DO PC), 192,194,195 dioleoyl phosphatidylchoJine (DOPC), 196 DIP (Database of Interacting Proteins), 142 dipalmitoleoyl PC (DPoPC), 167 dipalmitoyl phosphatidylcholine (DPPC) MD simulation of, 198, 199,200, 201 phase diagram for, 32 diphosphatidyl glycerol. See cardiolipin (Cl)

diphtheria toxin (DT), 84,132-133 distearoyl phosphatidylcholine (DSPC), 32,100 disulfide bond oxidoreductases, 134 DLPC (dilauroyl phosphatidylcholine), 100 DMPC (dimyristoyl phosphatidylchol ine) hydrophobic mismatching in, 100 in protein-folding studies, )67 molecular dynamic simulation of, 201 Monte Carlo simulation of, 202, 203 NMR spectra of, 24 docosahexaenoic acid (DHA), 20 I, 202 dodecyl tl-D-maltoside (DDM), 43 dodecylmaltoside, 235 dodecyl-phosphocol ine (DPC), I 18 dolichols, 20 domains, protein-binding, 80-83 OOPC (dioleoyl phosphatidylchoJine), 196 DOPG,78 DPC (dodecyl-phosphocoline), I J 8 OPoPC (dipalmitoleoyl PC), 167 DPPC (dipalmitoyl phosphatidylcholine) MD simulation of, 198, 199,200, 201 phase diagram for, 32 DRMs. See detergent-resistant membranes (DRMs) drug efflux systems, 296-297 ToIC-dependent, 306 tripartite, 300 drug targeting cyclooxygenase, 216, 222 G-protein coupled receptors, 140, 227 Dsb system, 176 OSc. See differential scanning calorimetl)' (DSC)

DSPC (distearoyl phosphatidylcholine), 32, 100 DT (diphtheria toxin), 84 E2-AIF4-(TG) complex, 267 EBF (epidermal growth factor) receptors, 37 edema factor (EF), 85 Edidin, M., 7 EGF-domain, of prostaglandin H2 synthase, 219, 220 EI (enzyme I), 135 Ell, 135 elaidic acid, 15 electrical potential-driven transporters, 133 electrical properties patch clamp recordings of, 55-56, 57, 58 planar biJayers used to study, 53-55 electrochemical gradient, across membranes, 53 electron carriers, 134 in nitrate respiratory pathway, 223, 224 in reaction cycle, 112, 113 electron density distribution, 193 electron paramagnetic resonance (EPR), 32, 33, 97 boundary lipids, 94, 95, 96, 98 MS channels, 238 myelin basic protein, 70 photosynthetic reaction centers, 108, 109 electron spin resonance (ESR). See electron paramagnetic resonance (EPR) electron transfer, in reaction cycle, 112 electrophysiology, 53 electrostatic interactions in MD simulations, 199 in reaction centers, I J 2 lipid selectivity and, 98 protein binding by, 69, 76, 78, 79, 80, 83 x-ray crystallography and, 203 electrostatic switch, 76, 83 eleidoyl, 200 emission spectrum, 26 EmrE, 297, 299-300 asymmetry in, 312 mutations, 299 structure of, 210, 299 endoplasmic reticulum (ER) P450 cytochromes in, 130 protein export to, 170, 172, 176 energy coupling, in vitamin 812 transport system, 296

Index energy surFaces, 196. 197 ensemble average. 197 enterobactin, 124 EnvZ, 119 enzymes, 127-129.214. See also specific enzyme lipid requirements of. 94, 95. 127 lipolytic. 214 epidermal growth factor (EBF) receptors. 37 epitopes. antibody recognition of. 143 EPR. See electl'on paramagnetic I-esonance (EPR) equilibration phase. 199 equilibrium measurements, of protein folding. 161 ER (endoplasmic reticulum) P450 cytochromes in, 130 protein export to, 170. 172, 176 ER pathway, 283. 284-286 ergosterol. 21, 22 eIJ,throcytes, glucose transporters in. 131.132 Escherichia coli acyl chains. 16 adhesion proteins. ] 14 ATP synthase, 272. 276, 309 chaperones. 176 colicins, 85 DGK.129 fatty acid composition, J 7 glyceroaquaporins. 254. 255-258 group translocators. 137 integral pl·oteins. 92 iron receptors. 124 mechanosensitive channels, 235. 236.237 OmpA, 118 OMPLA.214-2]5 OmpX, 118 phase transitions in. 40 phospholipid composition. 37, 39 porins. ]] 9,122 protein Folding. 100, 145 protein odentation. J 43, 146 SRP, 172.173.174 TolC-dependent drug efflux systems. 306 translocation, 175 translocon. 175,287 transporters, 241. 242, 248 tripartite drug efflux. 300 eukaryotes. See also humans; yeast; specific species aquaporins, 253 cell membranes. 1, 2 ion channels. 138-139 lipid-anchored proteins. 75 mechanosensitive channels. 235

323

P450 cylochromes, 130 plasma membrane cytoskeleton. 10 protein export, ]70.176 protein orientation. 144 receptors. 139 signaling enzymes, 129 sterols, 20-21. 22 topogenesis. 185 translocation, 160. 176 translocation apparatus. 175 translocons. 175, 286 tl"ansporters. 248 ExbB,296 ExbD.296 excitation wavelength. 26 excitatory glutamate receptors, 139 extracellular proteins. 72 eye, photoreceptors in, 228. 229

Fa domain. 274-275 F 1 Fa-ATPase catalytic mechanism, 277-278, 279 conFormational changes in. 276. 277 protrusion beyond membrane. 312 regulation of. 276-277 F 1 Fa-ATPase/ATP synthase, 272-273 structure of. 272, 273 subunit structure and fLlnction, 273-274.275,276.302 F 1 domain. 273-274, 275, 276, 302 Farnesyl, 20 FASTA,142 Fatty acid amide hydrolase, 90 fatty acid(s). 73 cis, 15,200 names of, 14. 15 phase transitions. 15, 17.29 polyunsaturated, 15 MD simulations of, 200-20 1.202 species of. 14 frans.15,200

FDH-N (formate dehydrogenase-N) functional unit. 225, 226 structure of. 223. 224 FepA protein, J 23, 124 ferdchrome. J 24 ferrous ion, in reaction centers, 109 FhuA protein, 123. 124 flap movement, in phosphoJipases, 71 flip-flop diffusion. See transverse diffLlsion flippases. 25. 26 flotation experiments. 192 Fluid Mosaic Model, 3. 7. 8, 68 lateral diffusion in, 24 lateral domains in, 33 fluidity. membrane. 24 measurement of. 8-9

fluorescence depoJarization, 26 fluorescence recovery after photobleaching (FRAP), 24, 26, 32,64 fluorescence resonance energy transfer (FRET). 26. 33 fluorescence techniques, 24, 26 F 1 Fa-ATPase studied with, 278. 279 giant unilamellar vesicles studied using, 63 lipid rafts studied with, 34 supported bilayers studied with, 59 formate dehydrogenase. 222-226. 312 formate dehydrogenase-N (FDH-N) ftlllctional unit, 225. 226 structure of, 223, 224 FosFomycin.245 Fourier transForm, 193 Fourier transform infrared (FTIR) spectroscopy. 161 FrankJin. Ben. 5 FRAP (fluorescence recovery after photobleaching), 24. 26, 32. 64 free fally acids. 14 freeze-fracture techniques, 6, 62 FRET (fluorescence resonance energy transfer). 26, 33 Frye. L. D.. 7 FTIR (Fourier transfOim infrared) spectroscopy. 161 fumarate reductase, 214 FYVE zinc-binding domains, 80. 81. 82 G protein (guanine nucleotide binding). 140 gain-of-function (GOF) mutants, 238 y-aminobutyric acid. 139 ganglioside GM z . 17, 18 ganglioside(s). 20. 98 gated pores, 131, 132. 288 gating mechanisms, 253 KcsA, 260-264 MS channels. 238-239 voltage, 262-263 gauche torsion angle. 15, 16 gel filtration, detergent removal by, 50 gene fusion. 143. 145, 146 general import pore (GIP). 178 genetic approaches to motifs and pa ttems, 312 to protein orientation, 143 genomic analysis of GPCRs, 234 of proteins, ]41. ]45-154 geranylgeranyJ, 20 GES (Goldman-Engelman-Steitz) scale. 139, 143 giant unilamellar vesicles (GUVs). 62 Gibbs absorption isotherm. 79

Index GIP (general import pore). 178 GlpF, 254, 255-258 channel interior, 256, 257, 258 hydrogen bonding, 256, 258 selectivity of, 255-256 structure oC 255, 256 GlpT. See glycerol-3-phosphate transporter (GlpT) glucose transporters in erythrocytes, 131, 132 topology of, 186 glucose-3-phosphate transporters. 241 glyceroaquaporins, 255-258 selectivity of, 254, 258 glycerol, 17 glycerol-3-phosphate (G3P). 17, 244, 255 glycerol-3-phosphate transporter (GlpT).242,244-245 glycerol-3-phosphate transporter (GlpT) binding sites, 245, 246 compared to LacY, 245-248 structure oC 245. 246, 247, 311 symmetl'y in, 312 translocation mechanisms. 242, 245, 247 glycerophospholipids. See phospholipids (PLs) glycolipids, 17, 18 glycophorin A, 93, 94.164,165 glycosylation sites, 186 giycosyJphosphatidylinositol (GPO anchors. 73, 74, 75 glyceroaquaporins, 253-254 GOF (gain-of-function) mutants, 238 Goldman-Engelman-Steitz (GES) scale, 139, 143 Gorter, E., 5 Gouy-Chapman theory, 79 GPCRs. See G-protein coupled receptors (GPCRs) GPI (glycosylphosphatidylinositol) anchors. 73, 74, 75 G-protein coupled receptors (GPCRs). 140-141,226 G-protein coupled receptors (GPCRs) activation mechanisms. 234 function of, 227 genomic and proteomic studies of. 234 gramicidins, 52, 87, 88, 89, 98, 132-133 greasy slide, 122, 123 green fluorescent protein, 144, 145 Green, David E .. 5,6 Grendel. F, 5 GROMOS.198 Gronhuss mechanism, 256-258

group translocators, 134, 135-137 GTPases, 173 guanine nucleotide binding proteins (G proteins), 140 GUVs (giant unilamellar vesicles), 62 GuxG motif, 154,265,267,312 half-helices. 93,312 Halobacter salinarum, 102, 103 halorhodopsin, 154 HDL (high-density lipoproteins), model bilayer system of, 66 head groups, 4 binding sites For, 76 heat capacity, of lipid bilayers, effects of peripheral proteins on, 76, 77 helical bundles, 102,213,214 conFormational changes in. 311 in bacteriorhodopsin, 102-107 in photosynthetic reaction center, 107-108 topology of, prediction of, 147 Helical Hairpin Hypothesis, 162 helical symmetry, 312 helix distortion, in integral proteins. 92-93,311-312 helix packing, 154, 164, 165,312 helix pair motifs, genomic analysis of, 154 helix tilting, 94, 99 in MS channels, 239 helix unwinding, 93 helix-helix interactions, 93,154,156 hydrogen bonding in, 154. ISS. 156. 164 in helical bundles, 102,312 in TolC tunnel. 306 triplets in, 154, 155 helix-tum-helix structures, in voltage paddles, 262 heme binding site, prostaglandin H 2 synthase. 218, 221 heme subunits cytochrome-bcl complex. 279-280. 281 reaction center, 110 hemifluorinated surfactants, 45 hetero-oligomers, 3\ 0 hexagonal phase. 28, 29-31 NMR spectrum of, 39, 40 types of, 28, 29 HHDBT (5-n-heptyl-6-hydroxy-4, 7-diozobenzothiazole),282 hidden Markov models (HMM), 150 high-sensitivity titration calorimetry, 61 high-density lipoproteins (HDL), model bilayer system of, 66 high-potential iron-sulfur protein (HiPIP), 1 \2, 113

histidine, 135 histidine-containing phosphocarrier protein (HPr), 135 HMM (hidden Markov models), 150 HMMTOP, 145, 150 holins, 132-133 homo-oligomers, 310 hop diffusion, 25, 27 HOQNO (2-heptyl-4-hydroxyquinolineN-oxide), 226, 229 HPr (histidine-containing phosphocan'ier protein), 135 Huber, Robert, 107,207 humans ABC transporters, 135 aquaporins, 253,254 ATP generation, 272 G-protein coupled receptors. 140, 226,227 mechanosensitive channels. 226-227 P450 cytochromes, 130 hydrocarbon chains, torsion angles in, 15,16 hydrocarbon-packing energies, 30 hydrogen bonding. See also specific protein importance of, 311 interhelical, 154. 155, 156, 164, 311 MD simulations, 200 protein Folding, 162, 164 transmembrane secondary structures. 92, 102 water, 4 hydrophobic effect, 3, 4-5 in lipid phase transitions, 30, 31 in protein folding. 162 hydrophobic interactions, protein binding by, 69, 78, 80, 82 hydrophobic mismatch, 98-99, 100 hydrophobicity measurement of, 4, 5 of ,ti-barrels, 157 positive-inside rule overriden by, 185 hydrophobicity plots, 141, 143, 144 combined with positive-inside rule, 145 generation of, 144 hydrophobicity scales, 142, 143, 144 for protein insertion studies, 182, 183 IMS (intermembrane space), 250 inclusion bodies, protein refolding from, 170 inhibitors. crystallization in presence of,310 insulin receptor, 139 integral lipids, 204, 206

Index integral proteins, 68, 90-94. See also specific protein amino acid distribution in, 145 classification of, 90, 91, 204 crystallization oF, 207 environmental factors affecting, 91 hydrophobicity plots of, 143 lipid interactions with, 94-98, 99 lipids associated with, 204 numbers of lipids in, 205 structure of, predicting, 141-143 interaction domains, in annexins, 72 interdigitation,27 INTERFACE-3, 154 interfacial region, 10 insertion of amphipathic helix into, 76, 82 inten:nembrane space (IMS), 250 inward-rectifying channels, 258 ion, 138-139 ion carriers, non-ribosomally synthesized, 133 ion channels, 138-139 patch clamp recordings of, 55-56, 57, 58 planar bilayers used to study, 53-55 ion exchange chromatography, 69 ionic strength, effect on micelle formation, 46, 47 ionophores, 87, 133 ionotropic ATP receptors, 139 iron receptors, 123-124 structure of, 123, 124 iron-sulfur proteins, in cytochrome-bel complex, 281 isoforms (isozymes), 129 isoprene, 20 KcsA,259 binding sites, 260 gating and conformational changes, 260-263,264,311 potassium conduction mechanism, 260,261 protein folding in, 169, 170 structure and selectivity, 259-260, 261 kinetic crystallography, 107 of F l Fa-ATPase, 277 of protein folding, 161 KirBacl.1, 259, 311 KirBac1.3, 259, 3J J knob-into-hole interactions, 93, 154, 164,165,306,312 Krafft point, 47 KvAP, 259, 3JJ, 3J2 voltage paddle, 262, 263 Kyte-Doolittle algorithm, 143

lactose permease (Lac carrier; LacY), 1-3,137,241,242-244 compared to GlpT, 245-248 conformational changes in, 244, 311 crystallization of, 207, 208 protein folding in, 100,169,170 structure of, 242, 243 symmetry in, 312 symport in, 242, 243, 244 transport mechanism, 132, 133 LacZ (tl-galactosidase), 144 LamB (maltoporin), 61, 118, 122-123 lamellar phases, 28 La (liquid crystalline), 28,191,193 Lb (gel), 28 Lc (crystalline), 23, 28, 191 NMR spectrum of, 39, 40 Langmuir isotherm, 79 Langmuir trough, 50 large unilamellar vesicles (LUVs), 60, 61 lateral diffusion, 24 measurement oF, 24, 26 rate of, 25 lateral domains, 33-34, 36 lateral mobility, of membrane components, 7, 8 lateral pressures effect on MS channels, 238 in protein folding, 167, 169 lauryldimethylamine oxide (LDAO), 43, 49,109 LCIC (ligand gated ion channel) da tabase, 139 LDAO (lauryldimethylamine oxide), 43, 49,109 leader peptidase (Lep), 145, 146. See also signal peptidase in topogenesis, 183, 184 length scales, in simulations, 202 lethal factor (LF), 85 LH1, 111, 112 LH2, 111, 112 ligand binding, 79, 83, 209-210, 234 ligand gated ion channel (LCIC) database, 139 light-harvesting complexes, in reaction centers, 111,310 light-induced signal transduction, efficiency of, 229 line tension, 34 linear isoprenoids, 17, 18,20-23 lipid anchor, 76 lipid bi/ayers, 7 components of, 13 diffusion of, 24-25 rate of, 25

heat capacity of, effects of per"ipheral proteins on, 76, 77 matrix, 23-24 models, 10, 196,212 molecular, 24, 25, 196 organization of, 31-33 planar, 53-55 formation of, 53-54, 55, 56 reversible interactions of peripheral proteins with, 76 simulations of, 210-212 stresses in, protein folding and, 167, 169 structure of, 24,191-192 supported, 57, 59 thickness of, 22, 26-28, 98 x-ray crystallography of, 23, 24, 28, 191 lipid chain, insertion of, protein binding by, 80 lipid clamp, 71 lipid cofactors. See nonannular lipids lipid mono layers, 50-53 curvature of, 30, 40 formation of, 50, 51 interaction with signal peptides, 52 phase changes in, 51 roleof,31,40 lipid phases, categories of, 28 lipid raf-ts, 8-9, 10,33-36 clustering of smaller domains in, 36 domain formation in, 57 effects of protein binding on, 78 GPI-anchored proteins associated with,75 location of, 34 models of, 34 physical properties of, 34, 35 proteins in, 9, 34, 90 used as markers, 36 size oF, 34 lipid shells, 36 lipid(s). See also specific lipid asymmetry, 26-27, 28 boundary layer of, 8 classes of, 17, 18,94,204 complex, 17 configuration of, 204, 205 crystallization of, 207 definition of, 4 diversity of, 13, 37-40 effects of peripheral protein binding on, 76-78 headgroups, 4 interactions between, 22, 23, 33 phase transitions, 28, 29,191,192 polymerized, 57 polymorphism, 28, 30

Index

326

lipid(s) (COI1/") proportion of, 7, 8, 17 removal of, 50, 206 requirements fOl~ 94, 95, 127 selectivity for, 95, 98, 99 stoichiometry of, 95, 99 structu res of, 4 surface concentration of, 128 trafficking patterns, J0 volume of, determination of, 192 lipid-anchored pt'oteins, 68, 73-74, 75, 76, 90 lipid-protein interactions, 127 integral, 94-98, 99 peripheral, 77, 78-80 x-ray crystallography and, 203-207 lipolytic enzymes, 214 liposome swelling assay, 60, 6J, 122 Iiposomes, 42, 60-63 types of, 60 uses of, 60 liquid crystal theory, 193-194, 195 liquid crystallography, 192 liquid-disordered state (Ld), 33 liquid-ordered state (L o ), 33, 34, 35 extraction of, 36 loss-of-function (LOF) mutants, 238 LSGGO motif, 135,136 lungs aquaporinsin,253,254 pulmonary surfactant, 52 LUVs (large unilamellar vesicles), 60, 61 lysine, 105 Iysophosphatidylcholine, 238 lysophospholipids,47 MJ 3 coat protein, 98 MacKinnon, Roderick, 207, 241 major facilitator superfamily (MFS), 133,242 paradigm fOI~ 248, 249 symmetry in, 3 J 2 MalK dimer, 136 maltoporin (LamB), 61, 118, 122-123 maltose, 135 mass spectrometry, 64 Maxwell-Boltzmann distributions, 198 MBD (membrane-binding domain), of prostaglandi n H2 syn thase, 219 MCF (mitochondrial carrier family), 138, 249. See a/50 ADP/ATP carrier motif,252

MD simulation. See molecular dynamics (MD) MDOs (membrane-derived oligosaccharides), 129 MDR (multidrug resistance) system, 296, 300 MDRI (multidrug resistance protein 1), 135, 186 mechanosensitive (MS) ion channels, 226, 235 channel opening models, 238, 239 classification of, 235 gating mechanisms, 238-239 large (MscL), 235-236, 237, 238, 239 mini (MscM), 235 small (MscS), 235, 237-238 melillin, 87, 88 membrane binding, See peripheral protein binding membrane damp, 76 membrane components, 1, 3, 8. See a/50 lipid(s); protein(s) lateral mobility of, 7, 8 proportions of, 7, 8,17 tools for studying, 42 membrane fluidity, 24 measurement of, 8-9 membrane fusion proteins, 302-303 Membrane Integrating Sequence for Translation of Integral Membrane Protein Constructs (Mistic), J70 membrane permeability, 1-3 membrane rafts, 3 membrane receptors, 139 membrane scaffold proteins (MSPs), 66 membrane solubilization, by detergents, 48-49, 50,129 membrane structure early models of. 5, 6 paradigms of, 3 shift in, 8-10 membrane thickness, 22, 26-28, 98 membrane(s) basic functions of, 1 dynamic nature of, 3 electrochemical gradient across, 53 membrane-binding domain (MBD), of prostaglandin H 2 synthase, 219 mem brane-derived oligosaccharides (MDOs),129 membrane-spanning proteins, in supported biJayers, 57 MEMSAT, 145,147 menaquinol (MOH2), 224 menaquinone (MOl. 224, 226 Metall, 234

MelhanobaCleriu 111 lhennoaulotrophicus, 259 Melhanococcus janllaschii translocon, 175,286-287,288 MexA, 303, 304 MFS. See major facilitator superfamily (MFS) micelles crystallization and, 207, 208 formation of, 45, 46, 47 mixed, 63 size of, 48 surface dilution kinetics 01',128 Michaelis-Menten kinetics, 129, 132 Michel, Hartmut, 107,207 microdomains, in blebs, 64 misfolding diseases, 187-189, 190 Mistic (Membrane Integrating Sequence for Translation of Integral Membrane Protein Constructs), 170 mitochondrial carrier Familv (MCF), 138,249. See a/50 ADP/ATP carrier mitochondrial carrier family (MCF) motif,252 mitochondrial inner membrane, composition of, 7, 8 mitochondrial membranes, 249 mitochondrial proteins, import of, 170-175,177 mixed micelles, 63 MLV (mulLilamellar vesicles), 60 protein binding to, 83 model membranes, 42, 50 for folding studies, 162 of lipid bilayers, 10, 196 peptide insertion into, 87 molecular dynamics (MD), 197-202 aquaporins, 258 ATP synthase, 278, 286 BtuCD complex, 293-294 calculations, 198 examples 01',198,199,200,212, 220 phases of, 199 steps in, J97-198 molecular models, of lipid bilayers, 24, 25, 196 monoamine oxidase, 90 monotopic proteins, 68, 69, 90 Montal, M., 53-55, 56 Monte Carlo (MC) simulations, 202-203 configurational-bias, 203 example of, 203 motifs, 311-312. See a/50 specific motif MPTopo, 144,157

Index MQ (menaquinone), 224, 226 MS channels. See mechanosensitive (MS) ion channels MsbA,210 MscL channels, 235-236, 237, 238, 239 MscM channels, 235 MscS channels, 235, 237-238 MSPs (membrane scaFfold proteins), 66 MthK, 259, 311 structure of, 260. 261, 262 Mueller, P., 53-55, 56 multidrug resistance (MDR) system, 296,300 multidrug resistance protein I (MDRI), 135,297 multilamellar vesicles (MLV), 60 protein binding to, 83 multiple sclerosis, 70 multiple sequence alignments, 148 multiprotein assemblies. See protein assemblies multi-spanning membrane proteins, topology of, 186 muscarinic acetylcholine receptor, 139 lVlycobaclerium luberculosis, 235, 236, 237 myelin basic protein, 69, 78 myelin proteolipid, 95, 98 myristate, 73, 74 myristoyl switch, 83, 84 Na+ channels, voltage-gated, 264 Na+K+ -ATPase, 134, 138 transport mechanism, 134 NADPH oxidase, 134 nanodiscs, 66, 67 nanomachines, 307, 312 native gel electrophoresis, 156 NBDs. See nucleotide-binding domains (NBDs) Neher, Erwin, 55, 57 neonatal respiratory distress syndrome, 52 neural networks, J 48 neurotransmitter receptor superfamily, 139-140 neutron diffraction, 191 combined with x-ray crystallography, 194-196 of protein-detergent complexes, 49, 50 techniques, 193 Newton's laws of motions, 197, 198 Nicolson, G. L., 6, 7, 33 nicotinic acetylcholine receptor, 139-140,239 nigericin, 133 nitrate reductase, 222-224

327

nitrate respiratory pathway, 222, 224 NMR. See nuclear magnetic resonance (NMR) Nobel Symposium on Membrane Proteins (2003), 309 nonannular lipids, 94, 204, 205 numbers of, 205 nonpolar domain, 10 non-ribosomally synthesized channels, 132-133 non-ribosomally synthesized ion carriers, 133 nonsteroidal anti-inflammatory drugs (NSAIDs), 216, 222 NPT ensemble, 197, 200 NSAIDs (nonsteroidal anti-inflammatory drugs), 216, 222 NTP ensemble, 202 nuclear magnetic resonance (NMR), 116 fJ-barrels, 116. 168 comparison of crystal structures with,210,211 Fo domain. 274-275 hydrophobic mismatch, 98 lipid bilayer, 24 lipid phase transitions, 39, 40 Mistic, 170 OmpA, 118, 168 OmpX, 118 pore formation, 88, 89 nucleoside channel porin (Tsx), 118 nucleotide(s), 83 nucleotide-binding domains (NBDs) AAC, 251 ABC transporters, 135, 136,292,293, 297,298-299 NVE production phase, 200 NVT ensemble, 197,202 OG (octyl fJ-D-glucoside), 43, 50 oleic acid, 15 oligomerization, 309-310. See also dimer formation protein-protein interactions and, 310 purification process and, 310 x-ray crystallography of, 310 oligomycin, 273 oligomycin-sensitivity conferring protein (OSCP subunit), 273 OmpA porin, 118 protein folding in, 168, 169 structure of, 210 OmpC pol'in, J 18, 119-J20 OmpF porin, 49, 50, 53, 118, 119-120 black films of, 54 blisters and, 65, 66

channel properties of, 121 crystallization of, 208 structure of, 120 OMPLA. See outer membrane phospholipase A (OMPLA) OmpR porin, 119 OmpX porin, 114, 115, 118 structure of. 210 OprM,307 optical spectroscopy, 108 organophosphate phosphate antiporter family, 247, 309 orphan proteins, 156 OSCP subunit (oligomycin-sensitivity conferring protein), 273 osmoporin (OmpC porin), 118, 119-120 osmotic pressure, mechanosensi tive responses to, 235 octyl fJ-D-glucoside (OG), 43,50 outer membrane phospholipase A (OMPLA), 115,214-216 calcium binding sites, 214, 216, 217 catalytic triad, 214, 215 dimer formation. 215, 216, 310 physiological role of, 214-215 protein folding in, 170 structure of, 215 ovispirin, 88 oxidoreductases, 134 OxIT, 248, 31 I oxygen reduction, by cytochrome-c oxidase, 285 p bulge, 93 P450 cytochromes, 129, 130-131 PagP, 210, 21 I pairwise sequence alignments, 142 palmitoyl, 73, 74 palmitoyJ-sphingomyelin (PSpM), 34, 35 Paracoccus denilrificans, 285 paradigms of membrane structure, 3. See aL50 Fluid Mosaic Model; hydrophobic effect shift in, 8-10 passive transporters, 131, 253 patch clamps, 55-56, 57, 58 blisters, 64 mechanosensitive channels, 235, 238 patterns, 311-312. See also specific pattern Pc. See phosphatidylcholine (PC) PCC (protein-conducting channel). See translocon PE (phosphatidylethanolamine), J 7, 38,39,109

Index

328

PEP PTS (phosphoenolpyruvatedependent phosphotransferase system), 135 peptides, transmembrane, 87-88. See also transmembrane (TM) domains; specific peptide peptidoglycan, 113, 1 19 peripheral pl'Otein binding, 76 domains in, 80-83 effects on membrane lipids, 76-78 ligands in, 79, 83 modulation of, 83-84 sites of, 76, 80 peripheral protein(s), 10,68,69-72 amphitropic, 72-73 embedded in membrane, 90 interactions between lipids and, 77, 78-80 reversible interactions with lipid bilayer, 76 permeability barrier, 1-3 blebs and, 64 changes in, pressure-induced, 226 lipid-protein interactions and, 94 porins and, 118 translocon and, 175 permeases, 127 peroxidase (POX), 218, 220, 221 pertussis toxin, 85 PFam-A database, 153 PG (phosphatidylglycerol), 17,38 PGH 2 (prostaglandin H 2), 216 PGHS. See prostaglandin H 2 synthase (PGHS) P-glycoprotein (multidrug resistance protein 1), 135, 297 PH (pleckstrin homology) domains, 80, 81,82 phase diagrams, 31,32 detergent-composition, 47 ternary, 32, 33, 35 phase transitions bacterial membranes, 16, 40 blebs, 64 fallyacids, 15, 17,29 lipid, 28, 29,191,192 lipid monolayers, 51 nanodiscs, 66, 67 PHD, 145, 147, 148 PhoE (phosphoporin), 118, 121, 122 PhoE (phosphoporin) channel properties of, 121 protein folding in, 170 phosphatidic acid, 17, 129 phosphatidyl choline (PC), 15, 17 structure of, 205

phosphatidylethanolamine (PE), 17, 38,39, 109 phosphatidylglycerol (PG), 17,38 phosphatidylinositol (PI), 17 phosphatidylinositol3-phosphate (PIP3),82 phosphoenolpynJvate-dependent phosphotransferase system (PEP PTS), 135 phosphoglycerate kinase, 161 phosphoinositides, 80 phospholipase A2 (PLA2), 71, 128 binding sites, 76 phospholipase C, 71, 72 phosphoJipases, 70 Hap movement in, 71 x-ray crystallography of, 71 phospholipids (PLs), 17-18,20 anionic, 18 interactions with cholesterol, 7, 22, 33 MD simulation of, 199 mixed micelles of, 63 phase changes, 51, 67 properties of, 18 simulations of, 198, 202 species of, 37 structures of, 15, 19, 192, 195 zwitterionic, 18 phosphoporin (PhoE), 118, 121, 122 channel properties of, 121 protein folding in, 170 phosphotidylserine (PS), 17 photoreaction cycle, in bacteriorhodopsin, 105, 107 photoreceptors, retinal, 228, 229 photosynthesis, pmcess of, 107 photosynthetic reaction center antennae in, 111-112,310 cardiolipin in, 205 cofactors in, 109-110 helical bundles in, 102, 107-108 helix-helix interactions in, J 54 hydrophobic mismatching in, 100 light-hal\/esting complexes in, III lipids in, 109 oligomerization in, 310 proteins in, 108-109, 144 reaction cycle in, 112-113 structure of, 108, 109 surface of, 109,206 transport classification, 133-134 photosystems, types of, 108, 112 phototransduction cycle, in rod ceJJs, 229-230 PI (phosphatidylinositol), 17 PIP3 (phosphatidylinositol 3-phosphate), 82 PKC. See protein kinase C (PKC)

PLA2 (phospholipase A2), 71, 128 binding sites, 76 planar bilayers, 53-55 formation of, 53-54, 55, 56 plants P450 cytochromes in, 130 photosystems in, 108 pldA gene, 215 pleckstrin homology (PH) domains, 80, 81,82 PLs. See phospholipids (PLs) pmf (proton motive force), 272, 279, 296 polar clamp, 155, 156 polyethylene glycol, 57 polymerized lipids, 57 polyphosphorylated inositol, 80 polystyrene beads, detergent adsorption to, 50 poly topic proteins, 68, 69, 90 classification of, 152 genomic analysis of, 152, 153 pmtein insertion in, 184 topogenesis in, 184, 186 polyunsaturated fatly acids, 15 MD simulations of, 200-201,202 POPC (l-palmitoyl-2-0Ieoyl phosphatidylcholine) in lipid rafts, 34, 35 MD simulation of, 200, 220 pore architecture, in potassium channels, 259, 261,262 pore formation cholesteml, 55 colicins, 86, 87 peptides, 88, 89 pore(s) conduction, 253 gated, 131, 132, 288 general import, 178 porins, 8, 118-119,241. See also specific porin Ii-barrels in, 113, 119, 120, 121, 132-133,157 channel properties of, 119, 121 classification of, 118 detergent-solubilized, 119 mutations, 119 number of, 119 purification of, I 19 specific, 120-122 structure of, 114, 119, 122 trimeric, 157 porters, 133 positive-inside rule, 144-145, 146, 184, 185 combined with hydrophobicity plots, 145 factors overr'iding, 185, 186

Index posttranslational translocation, J70 potassium channels, 258-259 conformational changes in, 3 J I diversity of, 258 gating mechanisms, 259 inward-rectifying, 258 selectivity of, 258 stnJcture of, 259 potential energy, 197, 198,202 POX (peroxidase), 218, 220, 221 precipitation, detergent removal by, 50 pressure in phase diagrams, 31 lateral. in protein folding, 167, 169 permeability changes caused by, 226 primary active transporters, 133-134 prion protein (PrP), 186, 187 GPI anchor of, 75 probability distributions, 192, 193, 194, 197 probes, in phase diagrams, 31-32 production phase, 199, 200 prokaryotes. See also bacteria; specific species anaerobic respiration in, 222, 224 tJ-barrels in, 114 cell membranes in, 1,2 lipid diversity in, 21, 22, 37 mechanosensitive channels in, 235 protein export in, 170 translocation systems in, 160, 175 transJocons in, 175 transporters in, 248 prostaglandin H 2 (PGH2), 216 prostaglandin H 2 synthase (PGHS), 90, 214,216-222 catalytic domain, 2 J7, 220, 221 EGF-domain, 219, 220 MD simulation of, 220 mechanism of action, 218 structure 01',218,219,220 protease protection assay, 171 protease sensitivity, 143 protective antigen (PA), 85 protein assemblies, 10,271-272. See also specific complex as nanomachines, 307, 312 genomic analysis of, 156 respiratory chain, 279 Protein Data Bank, 141, 142 protein folding, 160, 161 a-helical,162-164 bacteriorhodopsin, 164, 166-167 tJ-barrels, 167-168, 169 bilayer stresses and, 167, 169 environmental factors in, 91 helix packing in, 164,165 hydrophobic mismatch in, 100

329 in vilro studies 01',161-162,170

stages in, 161, 162, 163, 164 thermodynamics of, 160, J63,I69 protein insertion, 160, 169, 178-184 biological scale for, J83 cross-linking studies of, 178, 179, 182 protein kinase C (PKC), 72 activation of, 73 binding sites, 80 surface dilution kinetics of, 128 protein misfolding diseases, 187-189, 190 protein sequences databanks of, 141, 142, 154 genomic information on, 141 protein sorting, hydrophobic mismatch in, 100 protein toxins. See toxins protein translocation, 171,175,178 apparatus, 160,175 colicins, 85 cotranslational, 170, 176 GlpT, 242, 245, 247 LacY, 242, 243 posttranslational, 170 process of, 170-175,177 proteins involved in, 171, 175, 178 toxins, 84, 85 translocon-ribosome complexes, 288-289 proteinO, trafficking, in rafts, 36, 37 protein(s), 1,7. See also specific protein protein(s) biogenesis of, 160, 170, 182 bioinformatics tools 1'01',141,142 classification of, 68-69,127, 14J. 213 denaturation of, 161 export from cytoplasm, 170-172, 174,175,177 functions of, 127 genomic analysis 01',145-154 in blebs, 64 in detergent-resistance membranes, 36,37,49 lipid requirements of, 94, 95 orientation of, 143-144, 145 orphan, 156 proportion of, 7, 8 protrusion beyond membrane, 312 raft, 9, 34 used as markers, 36 reconstituted into nanodiscs, 66 topogenesis in, 170, 184-187 transport, 131-132 protein-conducting channel (PCC). See translocon protein-lipid interactions, 94-98, 127 x-ray crystallography and, 203-207

protein-protein interactions, 7, 9 disruption of, during purification process, 310 in AAC, 253 oligomerization and, 310 proteoliposomes, 60, 62 proteomics, 156, 157, 234 protomers, 271 proton motive force (pmf), 272, 279, 296 proton pathways aquaporins, 256-258 bacteriorhodopsin, 106,285 cytochrome-bcl complex, 283, 284-286 F 1 Fo-ATPase, 277 LacY, 243, 244 proton symporters, bacterial. 137 proton/drug antiporters, 299, 300 PrP (prion protein), 186,187 GPI anchor of, 75 PS (phosphotidylserine), 17 Pseudomonas aeruginosa, 303, 306, 307 PSI, 108 PSIBLAST, 142 PSI!, 108 PSpM (palmitoyl-sphingomyelin), 34, 35 PufX,112 pulmonary surfactant, 52 purple bacteria antennae system in, 111-112 photosynthetic reaction centers of, 108, 109, 112 purple membranes, 103 composition of, 104 Q cycle, 281, 282 quenching processes, 26 quinone(s), reaction centers, 109 quinone-binding sites, cytochrome-bcl complex, 281, 282

RCK domain, 261, 262 reaction center (RC), 107. See also photosynthetic reaction center reaction cycle, photosynthetic, 112-113 reconstitution, 42, 50 detergent removal for, 50 goal of, 309 recoverin, 83, 84 redox loop, 214, 218, 224 redox potentials, nitrate respiratory pathway, 223, 224 redox sites, cytochrome-bcl complex, 280, 281 reduction potentials, heme subunit of reaction center, 110 reporter enzymes, 144

Index

330

Resistance Nodulation cell Division (RND) superfamily, 297 respiratory chain complexes, 279, 280 classification of, 279 protrusion beyond membrane, 312 respiratory distress syndrome, neonatal, 52 retinal, 104, lOS, 106 analogs, 105 in pmtein folding, 166 in rhodopsin, 231,232,233 isomeriz.ation of, 105. 107 retinal photoreceptors, 228. 229 retinitis pigmentosa. 187, 189. 190, 232 reverse signal-anchors. 184, 185 Rhodobacter capsulatus, 113, 114. 119, 145 RJ1Odobacter sphaeroides cytochrome-c oxidase, 284, 285 integral lipids. 206 reaction centers, 108,109,110,111, 112 Rhodopseudornonas acidophila, I I I Rhodopseudomonas blastica, 119 Rhodopseudomonas sphaeroides. See RJ1Odobacter sphaeroides RJ1Odopseudomonas viridis. See Blastochloris viridis rhodopsin, 95, 226, 227 activation of, 229 cytoplasmic domain, 234 DHA in, 201 dimer formation, 234 ex tracell ula r- transmem brane interface in, 232 function of, 228-230 hydmgen bonding in, 232, 311 hydrophobic mismatching in, 99, 100 mutations, 189, 190 01 igomerization state of, 234 polar clamp in, 156 (-etinal in, 231, 232, 233 structure of, 103, 140,209,211. 227, 228, 230-235 TM helices in, 232. 243 ribosome, translocon bound to, 288-289,290 ripple phase, 28, 29 RND (Resistance Nodulation ceJl Division) superfamily, 297 RnfA, 145, 146 RnfE, 145, J46 Robertson, J. David, 6 rocker-switch type mechanism, in translocation, 242, 245, 247 rod cells, 228, 229 hyperpolarization in, 228. 229 phototransduction cycle in. 229-230

mtational diffusion, 24 RxxRR motif, 248 SAl protein. 182 Sakmann, Bert, 55, 57 salt bridge, in AAC, 251, 252 saponins, 43. 55, 56 sarcoplasmic reticulum calcium ATPase (SERCAI) conformational changes, 265, 266, 267, 268 structure of, 264, 265, 266, 267 Sav 1866, 135, 290, 297-299, 312 coupling helices, 298-299 drug binding to, 297-298 structure of, 297, 298 scattering length distribution, 193 Schiff base, 104, 105, 106 Schindler, H., 55 SCQP (structural classification of proteins), 142 ScrY (sucrose pOlin). 118 SDS. See sodium dodecyJsuJfate (SDS) SDS polyacrylamide electrophoresis (SDS-PAGE), 171, 172 SDSL (site-directed spin labeling), 97 sea anemone toxins, 87 Sec (secretion) proteins, 174, l75, 176, 312 SecA, 88-90, 173, 174, 286, 290 SecB, 173, 174 SeeD, 174, 175,290 SecDFYajC, 286 SecE, 175 SecF, 175, 290 SecG, 175,286 secondary active transporters, 137-138 SecY, 175,311 structure of, 286-287, 288 tl'anslating versus nontranslating states, 289 SecYEG, 156, 174, 175,286 ribosome binding, 289-290 structure of, 288, 289 sensory rhodopsin II, 154 SERCA 1. See sarcoplasmic reticulum calcium ATPase (SERCAI) serine z.ipper motif, 155 serotonin, 139 serpentine receptors, 140 short-chain/long-chain unilameJlar vesicles (SLUVs), 62 signal peptidase, 176, 184. See also leader peptidase (Lep) signal peptides, 52 interaction with monoJayers, 52 signal recognition particle (SRP), 172, 173,174

signal transduction in rafts, 36 integral proteins in, 92 light-induced, efficiency of, 229 signal(s) initiating protein export, 170, 171, 172. 173, 177 stop transfer, 176, 178, 184, 186 that govern topogenesis, 184, 185, 186, 187 signal-anchors, 173, 176, 182, 184, 185 reverse, 184, 185 signaling proteins calcium, 264 in raf-ts, 36 isoz.ymes as, 129 membrane receptors as, 139 simulation,191,210-212.Seealso molecular dynamics (MD); Monte Carlo (MC) combined with x-ray crystallography, 210-212 lipid bilayers, 196-197 Singer. S. J., 6, 7, 33 site-dil-ected spin labeling (SDSL), 97 sitosterol, 2 J, 22 small unilamellar vesicles (SUVs), 60, 61 smectic liquid crystals, diffraction studies of, 193, 195 SMS,61 sn-glycerol-3-phosphate, 17 snorkeling by transmembrane domains, 92 of amino acid residues, 184 sodium channels, voltage-gated, 264 sodium cholate, 43 sodium deoxycholate, 43 sodium dodecylsulfate (SDS), 43 precipitation of, 50 protein unfolding in, 166, 169 solubility of, 43 sonication, raft isolation with, 36 special pair, J 10, J J2 specific porins, 120-122 spectroscopic rulers, 168 sphingolipids, 17, 18,20 sphingomyelins, 20, 21, 98 sphingosine, 20, 2 J spider toxins, 87 spin probes, 97, 238 squalene-hopene cyclase, 90 Src peptide, binding by, 78, 80 SRP (signal recognition particle), 172, 173.174 Staphylococcus aureus. 85. See also Savl866 stator, 274 stearic acid, 14, 15, 200, 202

Index sterols, 17, 18, 20-2 I, 23 STGA, 206, 207 stigmasterol, 2 I, 22 stop transfer signals, 176, 178, 184, 186 stopped-flow absorption spectroscopy, 166 Streptomyces lividans, 259 stll.lctural classification of proteins (SCOP),142 substr·ate concentration, calculation of, 129 substrate-analogs, crystallization in presence of, 3 J 0 sucrose porin (ScrY), I 18 sugars, group translocation of, 135 supported bilayers, 57, 59 surface concentration terms, 23 surface dilution effects, 128 surface pressure vs. area isotheml, in monolayer formation,S I surface tension measurement of, 43 micelle formation and, 47 surfactants amphiphilicity of, micelle formation and,47 definition of, 43 hemiAuorinated,45 pulmonary, 52 SUVs (small unilamellar vesicles), 60, 61 Swiss-Prot, 141, 142, 154 switch helix, in TonB-dependent transporters, 295 symport, J 31, 137, 138 in LacY, 243, 244 symporters, 137-138 targeting complexes, in protein export, 176 TAT (twin arginine translocation) pathway, 176 Tay-Sachs disease, 17, 18 TBDTs (TonB-dependent transporters), 294 structure of, 295, 311 Tb-MscL,235,236,237 TC (transport classification) system, 132-133,134,158 TOG (t!-D-galactopyranosyl I -thio-{l-D-galactopyranoside), 243 temperature in phase diagrams, 3 I micelle formation and, 47 temperature factor (B values), 207, 3 II TEMPO, 32 terpenes, 73, 74

331

tetanus neurotoxin, 85 tetraheme cytochrome subunit, reaction center, I 10 TF (trigger factor), 174 thapsigargin (TG), 264, 266, 310 Thermochromatium tepidum, 108, 110, 112 thermodynamics in lipid phase transitions, 30 of F 1 Fo-ATPase, 276 of protein folding, 160,163, 169 TIM complexes, 178 time average (Aave), 197 time scales, in simulations, 202 time-averaged probability distributions, 192,193,194,197 TM domains. See transmembrane (TM) domains TMHMM, 145, 147, ISO, 156 TNBS (trinitrobenzenesulfonic acid), 27 TolC tunnel, 296, 297, 300, 302, 304-307 drug efflux systems dependent on, 306 opening mechanism, 306-307 partners of, 307 protrusion beyond membrane, 312 structure of, 300, 304-305, 306, 307, 310 TOM complex, 178 TonB box, 124, 295 TonB protein, 133,294 energy coupling, 296 structure of, 296, 297 TonB-dependent transporters (TBDTs), 294 structure of, 295, 3 11 topogenesis, 170, 184- 185, 187 topologv models helical bundles, 147 protein orientation, 144,145,147, 156 TopPred, 145 toroidal modeJ, 88 torsion angle, 15, 16 torsional potential, 199 toxins, 84-85. See also specific toxin channel-forming, 132-133 peptide, 87 TRAM (translocating chain-associated membrane protein), 174, 175, 176 tra11s fatty acids, 15,200 trans torsion angle, IS, 16 transcription translation systems, ;1'1 vitro, 170 transducers, 226-227 transducin, 228, 229, 234

transfer free energy change (Dc.Gtl')' 142,143 transient confinement zones, 76 translocase. See translocon translocating chain-associated membrane protein (TRAM), 174, 175,176 translocon(s), 170, 175, 176,286 dimer formation, 288 function of, 287 pathways through, 178, 179,288 structure of, 286-287,288, 289-290 TM insertion, 288 translocon-ribosome complex, 288-289, 290 ribosome contacts, 289-290 transmembrane (TM) domains, 69, 84, 90 amino acids in, 92 deletion of, for crystallization, 209 formation of secondary structure in, 92, J02 helical bundles, 102 prediction of, 141-143, 148 snorkeling by, 92 thickness of, 37 translocon, 288 types of, 81 transmembrane (TM) electr-on transfer carriers, 134 transmembrane (TM) helices in rhodopsin, 232, 243 numbering of, 243 transmembrane (TM) peptide segment bitopic proteins, 68 transport classification (TC) system, 132-133,134,158,297 tr·ansport mechanisms, 131, 132,134 AOPfATP carTier, 252-253 lac permease, 132 Na+K+-ATPase, 134 transport proteins, 131-132 classification of, 132-133, 134, 158 genomic analysis of, 157 transporters, 241-242 classification of, 242 role of, 241 transverse diffusion, 24 rate of. 25 transverse relaxation optimized spectroscopy (TROSY), 116 transverse view of membrane, 9 trigger faclor (TF), 174 trimeric pOi-ins, 157 trinitrobenzenesulfonic acid (TNBS), 27 tripartite drug efAux, 300 triplets, in helix-helix interactions, 154, 155

Index

332

tripod amphiphiles, 45 Triton X-I 00, 36, 43, 50 in protein-folding studies, 170 micelles, 63 surface dilution kinetics oE, 128 Trojan peptides, 87 TROSY (transverse relaxation optimized spectroscopy), 116 Tsx (nucleoside channel) porin, 118 twin arginine translocation (TAT) pathway, 176 ubiquinone, I"eduction by cytochmme C2, 112 uncoupling protein, 138 unilamellar vesicles, 60 giant (GUVs), 62 large (LUVs), 60, 61 short-chain/long-chain (SLUVs), 62 small (SUVs), 60, 61 uniport, 131 unit membrane, 5, 6 valinomycin, 133 van der Ploeg, P., J 97 van der Waals forces, 199 in helix packing, 164 x-ray crystallography and, 203 VceC, 307 vestibules, 253 vibrational energy, 199

Vibrio cholerae, 307 viral enveloped formation, domain segregation fOl~ 37 vitamin 8 12, J35, 291 structure of, 291 vitamin B l2 transporter (BtuBC), 135 vitamin B l2 uptake system, 291 energized phases of. 291 transport across inner membrane, 291-293,294 transport across outer membrane, 294-296 voltage clamp, 53 voltage gating, 262-263 voltage paddle, 262, 263 volume of lipid, determination of, 192 von Heijne positive-inside rule. See positive-inside rule

Walker A and B motifs, 135, 136 WalkeJ~ John E., 272 water channels. See aquaporins; glyceroaquaporins water solubility, of detergents, 43 water, hydrogen bonding in, 4 Wimley-White (WW) scale, 139, 143 Wuthrich, Kurt, 116 x-ray crystallography artifacts in, 209

combined with neutron diffraction data, 194-196 combined with simulations, 210-212 compared with NMR studies, 210, 211 conformational changes and, 310-311 crystallization pmcess in, 207-210 history of, 207, 309 ligand binding in, 209-210 lipid-protein interactions, 203-207 of bilayer thickness, 100 of helical bundles, 102 of integral proteins, 90 of lipid bilayer, 23, 24, 28, 191 oE oligomers, 310 of pore formation, 88, 89 resolution, 204, 207, 209 techniques, 193 uses of. 191 YajC, 175,290 yeast cytochrome-bcl complex, 280, 282, 283, 284, 285 inlegrallipids, 206, 208 lipid-anchored proteins, 75 mitochondrial Fl-c complex, 273 translocons, 286 yeast two-hybrid analysis, 156 YidC, 174, 175, 176, 286