Ion Channels: From Atomic Resolution Physiology to Functional Genomics -No. 245 (Novartis Foundation Symposia)

ION CHANNELS: FROM ATOMIC RESOLUTION PHYSIOLOGY TO FUNCTIONAL GENOMICS

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Novartis Foundation Symposium 245

ION CHANNELS: FROM ATOMIC RESOLUTION PHYSIOLOGY TO FUNCTIONAL GENOMICS

2002

JOHN WILEY & SONS, LTD

Copyright © Novartis Foundation 2002 Published in 2002 by John Wiley & Sons Ltd, Baffins Lane, Chichester, West Sussex PO19 1UD, England National 01243 779777 International (+44) 1243779777 e-mail (for orders and customer service enquiries): cs-books@ wiley.co.uk Visit our Home Page on http://www.wiley.co.uk or http://www.wiley.com All Rights Reserved. No part of this book may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording, scanning or otherwise, except under the terms of the Copyright, Designs and Patents Act 1988 or under the terms of a licence issued by the Copyright Licensing Agency, 90 Tottenham Court Road, London, W1P 9HE, UK, without the permission in writing of the publisher. Other Wiley Editorial Offices John Wiley & Sons, Inc., 605 Third Avenue, New York, NY 10158-0012, USA WILEY-VCH Verlag GmbH, Pappelallee 3, D-69469 Weinheim, Germany John Wiley & Sons Australia, Ltd, 33 Park Road, Milton, Queensland 4064, Australia John Wiley & Sons (Asia) Pte Ltd, 2 Clementi Loop #02-01, Jin Xing Distripark, Singapore 129809 John Wiley & Sons (Canada) Ltd, 22 Worcester Road, Rexdale, Ontario M9W1L1, Canada Novartis Foundation Symposium 245 ix+273 pages, 67 figures, 11 tables

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Contents Symposium on Ion channels: from atomic resolution physiology tofunctional genomes, held at the Novartis Foundation, London, 12-14 June 2001 Editors: Gregory Bock (Organizer) and Jamie A. Goode This symposium is based on a proposal made by Mark, Sansom Mark S. P. Sansom Introduction: stretching the envelope in structure —function studies of ion channels 1 Nigel Unwin Structure of the acetylcholine-gated channel Discussion 15

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Katjusa Brejc, Willem J. van Dijk, August B. Smit and Titia K. Sixma The 2.7 A structure of AChBP, homologue of the ligand-binding domain of the nicotinic acetylcholine receptor 22 Discussion 29 Alok K. Mitra, Gang Ren, Vijay S. Reddy, Anchi Cheng and Alexandrine Froger The architecture of a water-selective pore in the lipid bilayer visuali2ed by electron crystallography in vitreous ice 33 Discussion 46 Dax Fu, Andrew Libson and Robert Stroud conducting channel 51 Discussion 61

The structure of GlpF, a glycerol

Mark S. P. Sansom, Peter Bond, Oliver Beckstein, Philip C. Biggin, Jose Faraldo-Gomez, Richard J. Law, George Patargias and D. Peter Tieleman Water in ion channels and pores — simulation studies 66 Discussion 79 Benoit Roux What can be deduced about the structure of Shaker from available data? 84 Discussion 101

CONTENTS Stefano Garofoli, Gennady Miloshevsky, Vladimir L. Dorman and Peter C. Jordan Permeation energetics in a model potassium channel Discussion 122 Jacqueline M. Gulbis safety switch? 127 Discussion 142

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The b subunit of Kvl channels: voltage-gated enzyme or

E. Perozo, L. G. Cuello, D. M. Cortes, Y.- S. Liu and P. Sompornpisut EPR approaches to ion channel structure and function 146 Discussion 158 General discussion I

From structure to channel physiology 165

Senyon Choe, Susan Cushman, Kent A. Baker and Paul Pfaffinger Excitability is mediated by theTl domain of the voltage-gated potassium channel 169 Discussion 175 Diane M. Papazian, William R. Silverman, Meng-chin A. Lin, Seema K. Tiwari-Woodruff and Chih-YungTang Structural organization of the voltage sensor in voltage-dependent potassium channels 178 Discussion 190 Michelle M. Pirruccello, Nikolaus Grigorieff and Joseph A. Mindell Electron diffraction of a bacterial CIC-type chloride channel 193 Discussion 203 John B. C. Findlay and Michael A. Harrison A protein chemical approach to channel structure and function: the proton channel of the vacuolar H+-ATPase 207 Discussion 218 Anthony Auerbach Discussion 234

Acetylcholine receptors, between closed and open

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David B. Sattelle, Emmanuel Culetto, Marta Grauso, Valerie Raymond, Christopher Franks and Paula lowers Functional genomics of ionotropic acetylcholine receptors in Caenorhabditiselegans and Drosophila melanogaster 240 Discussion 257 Final general discussion Index of contributors Subject index

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Participants Frances Ashcroft (Chair) University of Oxford, University Laboratory of Physiology, Parks Road, Oxford OX1 3PT, UK Anthony Auerbach Department of Physiology and Biophysics, State University of New York, 324 Gary Hall, South Campus, Buffalo, NY 14214, USA Senyon Choe The Salk Institute for Biological Studies, 10010 North Torrey Pines Road, Lajolla, CA 92037-1099, USA Pierre-Jean Corringer Neurobiologie Moleculaire, Institut Pasteur, 25, rue du Dr Roux, F-75015 Paris, France Declan Doyle Laboratory of Molecular Biophysics, The Rex Richards Building, Department of Biochemistry, University of Oxford, South Parks Road, Oxford 0X1 3QU, UK John Findlay School of Biochemistry & Molecular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK Dax Fu Biology Department, Building 463, 50 Bell Avenue, Brookhaven National Laboratory, Upton, NY 11973, USA Jacqueline Gulbis The Walter and Eliza Hall Institute of Medical Research, Post Office, The Royal Melbourne Hospital, Melbourne, VIC 3050, Australia Peter Jordan Department of Chemistry MS-015, Brandeis University, PO Box 549110, Waltham, MA 02454-9110, USA Jeremy Lambert Department of Pharmacology and Neuroscience, University of Dundee, Dundee DD1 4HN, UK Alistair Mathie Biophysics Group, Blackett Laboratory, Imperial College, Prince Consort Road, London SW7 2BZ, UK

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Keith Miller Harvard Medical School, Massachusetts General Hospital, Department of Anaesthesia, Fruit Street, Edwards 505, Boston, MA 02114, USA Joseph Mindell Department of Biochemistry, Brandeis University, 415 South Street,Waltham, MA 02454-9110, USA Alok Mitra Department of Cell Biology, The Scripps Research Institute, Mail Stop MB21,10550 North Torrey Pines Rd, Lajolla, CA 92037, USA Diane Papazian Department of Physiology, UCLA School of Medicine, 405 Hilgard Avenue, Box 951751, Los Angeles, CA 90095-1361, USA Eduardo Perozo Department of Molecular Physiology and Biological Physics, University of Virginia, POBox 800736, Charlottesville,VA 22908-0736, USA Chris Poll Novartis Horsham Research Centre, Wimblehurst Road, Horsham RH12 SAB, UK Benoit Roux Department of Biochemistry, Weill Medical College of Cornell University, 1300 York Ave, Box #63, Room W-220, New York, NY 10021, USA Mark Sansom Laboratory of Molecular Biophysics, Department of Biochemistry, Rex Richards Building, University of Oxford, South Parks Road, Oxford OX1 3QU, UK David Sattelle MRC Functional Genetics Unit, Department of Human Anatomy and Genetics, University of Oxford, South Parks Road, Oxford 0X1 3QX, UK Tilman Schirmer University of Basel Biozentrum, Abteilung Strukturbiologie, Klingelbergstrasse 70, Basel, CH-4056, Switzerland Hildgund Schrempf Universitat Osnabriick, Fachbereich Biologie/Chemie, Barbarastr. 11, D-49069 Osnabriick, Germany Titia Sixma Netherlands Cancer Institute, Division of Molecular Carcinogenesis, Plesmanlaan 121,1066 CX Amsterdam,The Netherlands Kenton Swartz Molecular Physiology and Biophysics Unit, NINDS Building 36, Room 2C19,36 Convent Drive, MSC 4066, Bethesda, MD 20892-4066, USA

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Peter Tieleman Department of Biological Sciences, University of Calgary, 2500 University Drive NW, Calgary, Alberta, CanadaT2N 1N4 Nigel Unwin MRC Laboratory of Molecular Biology, Hills Road, Cambridge CB2 2QH, UK Bonnie A.Wallace School of Crystallography, Birkbeck College, University of London, Malet Street, London WC1E 7HX, UK Su Li Novartis Horsham Research Centre, Wimblehurst Road, Horsham RH12 SAB, UK John Westwick Novartis Horsham Research Centre, Wimblehurst Road, Horsham RH12 5AB, UK

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Introduction: stretching the envelope in structure-function studies of ion channels Mark S. P. Sansom Laboratory of Molecular Biophysics, Department of Biochemistry, Rex Richards Building, University of Oxford, South Parks Road, Oxford OX1 3Q U, UK

This meeting has provided a timely opportunity to review progress in understanding the relationship between structure and function of ion channels. However, before considering this progress in more detail, it is important to consider why ion channels are of interest to investigators ranging from structural biologists and biophysicists to pharmacologists and physiologists. It is now recognized that mutations in ion channels can result in a wide range of diseases (Ashcroft 2000). Furthermore, it has been suggested that ion channels may make up some 25% of the potential targets for new drugs (Terstappen & Reggiani 2001). In addition to their biomedical importance, ion channels have for a long time been at the cutting edge of biophysical studies of membranes (Hille 1992). They now provide us with a real chance of linking up descriptions of membrane proteins at the genomic, structural and physiological levels. For the past 30 years or so, there have been almost continuous methodological advances for studying the function and structure of ion channel proteins. The development of the patch clamp technique (Neher & Sakmann 1976) allowed ionic currents flowing through single channel molecules to be measured with unique precision. This enabled more fine-grained dissection of the functional properties of different species of channel than had previously been possible, and also allowed detailed kinetic modelling of the conformational changes underlying channel gating (Colquhoun & Hawkes 1981). The 1980s saw the emergence of the first amino acid sequences of channel proteins (Noda et al 1982). The combination of these molecular insights with patch clamp recording techniques and in vitro mutation of channels resulted in a flourishing of molecular physiology studies that enabled assignment of physiological function to distinct regions within a channel sequence (Miller 1991). However, in the absence of three-dimensional structures for channel proteins, the precise relationships between structure and function often remained tantalisingly out of reach. The

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solution to this problem lay in the discovery of a bacterial homologue (KcsA) of mammalian K+ channels (Schrempf et al 1995). The breakthrough structure of KcsA (Doyle et al 1998) provided a paradigm for how a high resolution X-ray structure can inform functional studies. We are now witnessing a new phase of channel research, combining the search for new channel structures with poststructural studies using new approaches from spectroscopy, simulation, bioinformatics and genomics. We hope that this book will capture some of the excitement of these developments. A number of themes run through this meeting. The first is that of the continuing progress in determining structures of ion channels. Ongoing structural advances include: the nicotinic acetylcholine receptor (Unwin 2000); an acetylcholine binding protein (Brejc et al 2001); Cl- channels (Mindell et al 2001); regulatory subunits of voltage-gated K+ channels (Gulbis et al 1999); and also two channellike transporter proteins from the aquaporin family, namely GlpF (Fu et al 2000) and AQP1 (Ren et al 2001). In addition to such (essentially static) structures from X-ray and electron microscopy methods, significant advances are being made in understanding the conformational dynamics of channel proteins, using methods such as electroparamagnetic resonance (EPR) spectroscopy (Perozo et al 1999), molecular simulations and related computational approaches (Roux et al 2000, Sansom et al 2000) and detailed analysis of the effects of mutations on channel gating (Grosman et al 2000, Tiwari-Woodruff et al 2000). All of these approaches are moving us away from the picture of a rigid static pore through which ions flow, towards a more dynamic model of channel structure and function. A second theme is that of studying function at the microscopic (i.e. atomic resolution) level. This is an area where physiological and biophysical chemistry studies of ion channels converge. Obtaining a precise description of how selected ions flow through a channel at near diffusion-limited rates is an challenging problem in biophysical chemistry (Jordan 1999), and may provide a paradigm for theoretical studies of other, more complex, membrane transport proteins. In some respects it is chastening that we still do not possess a complete quantitative theory of why a K+ channel is K+ selective. However, at least we have now an X-ray structure to help us make substantial progress towards the Holy Grail of describing single-channel function in atomistic detail (Fig. 1). Similar approaches will have to be adopted for other pore-like transporters such as AQP1 and GlpF. A third theme is that of the challenges posed to ion channel research by genomics. For example, the genome sequence of a 'simple' animal, Caenorhabditis e/egans, has revealed a surprisingly large number of e.g. K+ channel genes (Bargmann 1998). It will be of considerable interest to see how these are related to the physiology of the animal as a whole (Davis et al 1999). And so, coming full circle, as genomics reveals a diversity of channel genes, a major challenge for the

INTRODUCTION: STRETCHING THE ENVELOPE

FIG. 1. Diagram of the central challenge facing research on channels: how can we relate channel structure (left) to channel physiology (right)? The channel structure is a model of the pore domain of Kir6,2 (Capener et al 2000); the single channel recording is also from Kir6.2 (diagram courtesy of F. M. Ashcroft).

pharmaceutical industry will be to devise compounds that exploit this diversity in terms of drugs exhibiting selective interactions with specific ion channel proteins. What of the challenges of the future? There are many. From a biomedical perspective, a pressing challenge is to arrive at a more integrated view of channel structure and function ('from atom to animal...'). In particular, one would like to see better integration of data from genomics, bioinformatics, mutation and structural studies of channels. From a fundamental biophysical point of view, it should be possible to extend our emergent understanding of channel structurefunction relationships both to a wider range of ion channels and to other more complex transporters (e.g. AQP1, GlpF and H+-translocating ATPases).

References Ashcroft FM 2000 Ion channels and disease. Academic Press, San Diego Bargmann CI 1998 Neurobiology of the Caenorhabditis elegans genome. Science 282:2028-2033 Brejc K, van Dijk WJ, Klaassen RV et al 2001 Crystal structure of an ACh-binding protein reveals the ligand-binding domain of nicotinic receptors. Nature 411:269-276 Capener CE, Shrivastava IH, Ranatunga KM, Forrest LR, Smith GR, Sansom MSP 2000 Homology modeling and molecular dynamics simulation studies of an inward rectifier potassium channel. Biophys J 78:2929-2942 Colquhoun D, Hawkes AG 1981 On the stochastic properties of single ion channels. Proc R Soc Lond B Biol Sci 211:205-235 Davis MW, Fleischhauer R, Dent LA, Joho RH, Avery L 1999 A mutation in the C. elegans EXP2 potassium channel that alters feeding behavior. Science 286:2501-2504

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Doyle DA, Cabral JM, Pfuetzner RA et al 1998 The structure of the potassium channel: molecular basis of K+ conduction and selectivity. Science 280:69-77 Fu D, Libson A, Miercke LJW et al 2000 Structure of a glycerol-conducting channel and the basis of its selectivity. Science 290:481-486 Grosman C, Zhou M, Auerbach A 2000 Mapping the conformational wave of acetylcholine receptor channel gating. Nature 403:773-776 Gulbis J, Mann S, MacKinnon R 1999 Structure of a voltage-dependent K* channel b subunit. Cell 97:943-952 Hille B 1992 Ionic Channels of Excitable Membranes, 2nd edn. Sinauer Associates, Sunderland, MA Jordan PC 1999 Ion permeation and chemical kinetics. J Gen Physiol 114:601-604 Miller C 1991 1990: annus mirabilis of potassium channels. Science 252:1092-1096 Mindell J A, Maduke M, Miller C, GrigorieffN 2001 Projection structure of a CIC-type chloride channel at 6.5 A resolution. Nature 409:219-223 Neher E, Sakmann B 1976 Single-channel currents recorded from membrane of denervated frog muscle fibres. Nature 260:799-802 Noda M, Takahashi H, Tanabe T et al 1982 Primary structure of alpha-subunit precursor of Torpedo calif arnica acetylcholine receptor deduced from cDNA sequence. Nature 299:793-797 Perozo E, Cortes DM, Cuello LG 1999 Structural rearrangements underlying K + -channel activation gating. Science 285:73-78 Ren G, Reddy VS, Cheng A, Melnyk P, Mitra AK 2001 Visualization of a water-selective pore by electron crystallography in vitreous ice. Proc Natl Acad Sci USA 98:1398-1403 Roux B, Berneche S, Im W 2000 Ion channels, permeation and electrostatics: insight into the function of KcsA. Biochemistry 39:13295-13306 Sansom MSP, Shrivastava IH, Ranatunga KM, Smith GR 2000 Simulations of ion channels — watching ions and water move. Trends Biochem Sci 25:368-374 Schrempf H, Schmidt O, Kummerlen R et al 1995 A prokaryotic potassium ion channel with two predicted transmembrane segments from Steptomyces lividans. EMBO J 14:5170-5178 Terstappen GC, Reggiani A 2001 In silico research in drug discovery. Trends Pharmacol Sci 22:23-26 Tiwari-Woodruff SK, Lin MC, Schulteis CT, Papazian DM 2000 Voltage-dependent structural interactions in the Shaker K+ channel. J Gen Physiol 115:123-138 Unwin N 2000 The Croonian Lecture 2000. Nicotinic acetylcholine receptor and the structural basis of fast synaptic transmission. Phil Trans R Soc Lond B Biol Sci 355:1813-1829

Structure of the acetylcholine-gated channel Nigel Unwin MRC Laboratory of Molecular Biology, Hills Road, Cambridge CB2 2QH, UK

A bstract. The structure of the acetylcholine-gated channel, trapped in open as well as closed states, has been analysed by electron crystallographic methods. The channel has large vestibules extending from the membrane which shape the acetylcholine-binding pockets and facilitate selective transport of cations across a narrow membrane-spanning pore. When acetylcholine enters these pockets it triggers a concerted conformational change that opens the pore by destabilizing a gate in the middle of the membrane made by a ring of pore-lining a-helical segments. The alternative 'open' configuration of porelining segments reshapes the lumen and creates new surfaces, allowing the ions to pass through. Recent results, at nearly 4 A resolution, have defined more precisely the structure of the pore and the design of the vestibular entrances. 2002 Ion channels—from atomic resolution physiology to functional genomics. Wiley, Chichester ( Novartis Foundation Symposium 245) p 5-21

The acetylcholine-gated channel, or ACh receptor, is a model neurotransmittergated ion channel found at the nerve/muscle synapse and in the electric organs of fish. Its structure has been explored extensively by electron crystallographic experiments, using postsynaptic membranes from the Torpedo ray. Based on this work, we now know what it looks like to nearly 4 A resolution, and have an elementary picture of how the channel opens. Descriptions of the acetylcholine-gated channel, in both the closed and open forms, have been given in recent publications (Unwin 1995, 2000, Miyazawa et al 1999). My intention here is to focus on the structure of the ion-conducting pathway. I will discuss our present understanding of the gate and the gating mechanism, and of how the channel is designed to achieve efficient, yet highly selective transport of cations across the membrane. The structural principles of gating and ion selectivity, as they apply to this channel, may be pertinent also to other examples discussed in this volume. Basic architecture The acetylcholine-gated channel is a ~ 300 kDa glycoprotein made from a ring of five rod-shaped membrane-spanning subunits (Fig. 1). These tilt by ~ 10° around

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the central axis, so that the whole complex has a slightly right-handed, coiled configuration. Each subunit is comprised of a large extracellular N-terminal domain, four predicted transmembrane regions (Ml—M4), and an extended M3M4 cytoplasmic loop. There are two ACh-binding a subunits, which have the same amino acid sequence, and three others, b,g and d, which are similar to the as. The ACh-binding pockets lie about midway between the membrane surface and the extracellular end of the complex (Fig. 1, asterisks). The five subunits together delineate a central, ~120A long ion-conducting pathway, which is divided into three parts: a narrow membrane-spanning pore and, on either side, large water-filled vestibules (Fig. 1). Two important functional regions lie within the pore: the gate, which is located near the middle of the membrane (see Fig. 3); and the constriction zone of the open pore, which is close to the cytoplasmic membrane surface (see Fig. 4). The two vestibules are quite

FIG. 1. Outline of whole channel, emphasizing features of the central ion-conducting pathway within, and on either side of the narrow membrane-spanning pore. The extracellular vestibule has a cylindrical shape whereas the cytoplasmic vestibule is nearly spherical, with narrow openings, or 'windows' lying directly under the membrane surface. The positions of the AChbinding pockets are indicated by asterisks; the bent helices in the membrane represent the M2 segments in the closed configuration; the gate is where they come closest to the axis of the pore.

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different, the extracellular one being essentially a ~ 65 A long by ~ 20 A diameter cylinder, and the cytoplasmic one a ~ 20 A diameter sphere. A special feature of the cytoplasmic vestibule is that it does not open directly into the cell interior. Instead, the ions have to pass through narrow (<8-9A wide) lateral openings lying directly under the membrane surface. These lateral 'windows' are shaped by ahelical segments projecting from the individual subunits and coming together at the base of the receptor to form an inverted pentagonal cone.

Role of the vestibules The walls of the vestibules together make up more than 70% of the mass of the channel. Much of this mass may be involved in shaping the ACh-binding pockets and sites of attachment for regulatory molecules and other proteins (e.g. rapsyn) that are concentrated at the synapse. However another likely physiological function of the vestibules is to serve as pre-selectivity filters, making use of charged groups at their entrances and on their inner walls to electrostatically guide and concentrate the ions which are selected for while screening out ions which are discriminated against. Rings of negative charge located at the mouths of the narrow membrane-spanning pore have been shown by mutation studies to be significant determinants of channel conductance (Imoto et al 1988). The modification of the ionic environment achieved by the vestibules may be thought of as providing an additional means for increasing the efficiency of cation transport. The extracellular vestibule provides a route which is narrow enough for the charged groups on the inner wall to influence ions at the centre, yet not too narrow to restrict their diffusion — and so is designed such that weak electrostatic interactions, distributed over a considerable distance, could combine to achieve a strong overall effect. In fact, the extracellular wall has a high net negative charge: on average — lle/subunit, based on amino acid composition (Unwin 1989). This charge is the same as that for the fast-acting enzyme, acetylcholinesterase (Nolte et al 1980, Ripoll et al 1993), where the whole protein surface plays a role in producing an electrostatic field that guides the positively charged ACh substrate to the active site. It is easy to imagine that the channel uses its predominantly negatively charged extracellular surface to guide, in a similar manner, inorganic cations to the pore (see also Adcock et al 1998). Similar principles are likely to apply to the cytoplasmic vestibule, since electrophysiological experiments have shown that there is no marked preference for cations to go in one direction across the membrane. Indeed, the a-helical segments shaping the 'windows' can be identified with the stretch of sequence containing heptad repeats of negatively charged residues, denoted MA (Miyazawa et al 1999), and once thought to be lining the membrane-spanning

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FIG. 2. Sequence comparison of aligned MA stretches involved in shaping the cytoplasmic windows. Presumed framing residues are boxed, and charged residues (E,D: negative; R,K: positive) are in bold. The acetylcholine- and 5-HT 3 gated channels are cation selective; the glycine- and GABAA-gated channels are anion selective.

pore (Finer-Moore & Stroud 1984). This identification, in turn, suggests that the windows do not merely function as molecular sieves, preventing impermeant cytoplasmic molecules from reaching the vicinity of the pore, but also make use of the negative charge to help to concentrate cations in the vestibule. Passing ions would be strongly influenced by negatively charged side-chains framing the windows, which are narrower for the most part than the diameter of an ion including its first hydration shell. Figure 2 shows the stretches along MA which, from tentative fitting to the threedimensional densities, appear to be involved in shaping the windows. This figure also shows aligned sequences from other channels in the ACh receptor family. There are several features consistent with the idea that these stretches may play a general role in pre-selecting ions. First, the presumed framing residues (boxed regions) are grouped in pairs, spaced seven residues apart: they therefore lie along lines on opposite faces of each helix, making an included angle of 100C, which closely matches the angle made by adjacent windows (ideally 108c). Second, there is a clear distinction between the cation-selective (AChR, 5-HT3) and the anion-selective (Gly, GABAA) channels in terms of the framing residues being predominantly negatively charged in the former case and positively charged in the latter. Third, it is only the a subunit which uses both faces to shape the three main windows of the acetylcholine-gated channel (Miyazawa et al 1999) and, appropriately, this helix is the most negatively charged. Closed pore The pore of the acetylcholine-gated channel is shaped mainly by the membranespanning segments, M2, one contributed by each of the encircling subunits (for a review, see Lester 1992). These segments are a helical, but bend inwards near the

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middle of the membrane to form the gate of the closed channel (Unwin 1993, Miyazawa et al 1999; see Fig. 1). The precise nature of the gate — whether it forms a physical occlusion, or is better described as a zone of low polarizability that reinforces the dielectric barrier to ion permeation — has not yet been clarified. Earlier studies had emphasised the likely importance of a ring of conserved leucine residues (aLeu251), and had suggested they might project inwards from the M2 segments, associating side-to-side to form an occluding gate (Unwin 1993). But the side-chain densities in the most recent ~ 4 A map do not come close enough to the pore axis to account for such a simple arrangement. Apparently there is a hole of ~ 3.5 A radius along the central axis of the pore where the gate is located. This is only slightly smaller than the radius of a Na+ or K+ ion having a single hydration shell. Therefore, to be effective in preventing ion permeation, the gate should counter any opportunity for electrostatic stabilization, which could liberate ions from their hydration shells, by forming a completely hydrophobic girdle around the hole. Two rings of hydrophobic residues are likely to be part of this girdle: one including the conserved leucine and the other including a valine (aVal255; also highly conserved in ACh receptors), one turn of a helix away. Residues in both these rings can be labelled by small uncharged photoactivable compounds when the channel is closed (White & Cohen 1992, Blanton et al 1998). The higher resolution map suggests that the leucines may interact side-to-side with alanine/ serine residues on neighbouring M2 segments, rather than with each other, to form the girdle (Fig. 3). Since the M2 segments are not closely packed and draw away from the pore axis (and hence from each other on either side of the bend), the interaction implicating aLeu251 may be the only one between these segments. The resulting limited stability of the structure would ensure its easy disruption and reassembly by conformational changes initiated at the binding sites. Other pentameric structures composed of bundles of a helices, such as phospholamban (Adams et al 1995), COMP (Malashkevich et al 1996) and the mechanosensitive channel (Chang et al 1998), all contain rings of hydrophobic side chains lying closer to the pore axis than those of the receptor. These side chains could function as occluding gates. However, the a-helical bundle structures involve more extensive side-to-side interactions, increasing their stability and making them less suited for rapid conversion between two widely distinct and precisely defined states. Open pore The structure of the open pore has been determined to 9 A resolution by spraying ACh onto postsynaptic membranes on the microscope grid and freezing the grid

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FIG. 3. The higher resolution maps suggest that the gate may be made by leucine and valine side-chains creating a hydrophobic girdle around the pore. The hydrophobic surface, in providing no electrostatic stabilization, would counter liberation of the ion from its hydration shell, and the central hole would be too narrow for a hydrated ion to pass through.

within 5 ms to trap the reaction before significant desensitization can take place (Berriman & Unwin 1994, Unwin 1995). The M2 segments encircling the open pore have a different configuration from those encircling the closed pore, being further apart near the middle of the membrane and coming together instead at the cytoplasmic membrane surface (Fig. 4A). The open pore is consequently narrowest at the cytoplasmic membrane surface, forming a constriction zone — where direct interaction between the hydrated ions and encircling protein determines most critically whether or not the ions can go through. A tentative alignment of the three-dimensional densities with the cytoplasmic portion of the M2 sequence (Unwin 1995) suggested that a line of small polar (serine or threonine) residues would lie almost parallel to the axis of the pore when the channel opens (Fig. 4B), an orientation that should stabilize the passing ions by providing an environment of high polarizability. The gating action

A simple mechanistic picture of the gating action, suggested by comparison of the ACh-activated structure with the structure obtained in the absence of ACh, would

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FIG. 4. Transient configuration of M2 rods around the open pore, and interpretation at 9 A (Unwin 1995). (a) A barrel of a-helical segments, having a pronounced twist, forms in the cytoplasmic leaflet of the bilayer, constricting the pore maximally at the cytoplasmic membrane surface. The bend in the rods is at the same level as for the closed pore, but instead of pointing inwards has rotated over to the side, opening up the pore in the middle of the membrane, (b) Schematic representation of the most distant three rods. A tentative alignment of the amino acid sequence with the densities suggests that a line of polar residues should be facing the open pore (from Unwin 1995).

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be as follows. First, ACh triggers localized disturbances at the two binding sites. Second, the effects of these disturbances are communicated, through 'twisting' rotations of the a subunits, to the structure in the membrane. Third, the M2 segments in the membrane transmit the rotations to the gate-forming side chains, drawing them away from the central axis; the mode of association near the middle of the membrane (Fig. 3) is thereby disfavoured, and the segments switch to the alternative side-to-side mode of association (Fig. 4), creating an open pore. The bend near the middle of the M2 segment, where the helix is partially unfolded, may play a crucial role in facilitating the rapid switch between the closed and open pore. In acting as a point of flexure, it would allow this critical part of the receptor to move economically within the rest of the membranespanning structure (which does not change significantly), so that there is no direct influence on the motion by the surrounding lipids. Interestingly, molecular dynamics simulations of isolated a helices in water show that those composed of the M2 stretch of amino acids have a propensity for localized flexure in their middle portion. The M2 helix unfolds in this region, in the simulations, allowing the bend to act as a molecular swivel, or hinge (Law et al 2000). Relation to mutagenesis experiments Mutagenesis combined with electrophysiological study of function has been a valuable approach for exploring the nature of the membrane-spanning pore and identifying the likely roles played by individual M2 side chains in affecting ion permeation. Several such experiments have highlighted the uniqueness of the conserved leucine residue in relation to the gating mechanism. For example, replacement of successive leucines by serines (Labarca et al 1995), or by threonines (Filatov & White 1995) markedly increased, by uniform increments, the opening sensitivity of the channel. An analysis of the influence of mutations on the binding of the open channel blocker, QX222, is consistent with the structural evidence that the open pore tapers towards the cytoplasmic end of M2 and that the line of polar pore-facing residues shown in Fig. 4B is likely to play an important role (Charnet et al 1990). The threonine at the point of maximum constriction (aThr244), when substituted by other residues of different volume, has a pronounced effect on ion flow, as if it is in the narrowest part of the open pore (Imoto et al 1991, Villarroel et al 1991). The critical nature of the cytoplasmic end of M2, including the preceding loop region, has been reinforced more recently by experiments investigating anion versus cation selectivity, carried out on other channels in the ACh receptor family (Corringer et al 1999, Keramidas et al 2000, Gunthorpe & Lummis 2001).

ACETYLCHOLINE-GATED CHANNEL

13

Cysteine scanning mutagenesis has also been applied extensively to the ACh receptor, and other members of the family, to probe the organization of residues composing M2, and other membrane-spanning segments, around the pore. Some of these experiments lead to a similar picture of the pore to the structural results, while others suggest there could be discrepancies, for example in the exact location of the gate (Wilson & Karlin 2001). One disadvantage of the mutagenesis approach is that amino acid substitutions often alter the local structure in an unknown and unpredictable way. This makes interpretation difficult, particularly when it comes to the key functional components. Mutation experiments also tend to over-emphasize the importance of a small number of identified amino acid residues, whereas weak interactions by many, widely distributed residues can combine to produce strong overall effects. The structure, as I have described, strongly suggests that this latter principle is an important one in determining the physiological properties achieved by the acetylcholine-gated channel.

Conclusions The electron crystallographic experiments have provided an elementary picture of the ion-conducting pathway of the acetylcholine-gated channel, showing how it is designed and suggesting how it functions. The vestibules appear to play a significant role in pre-selecting for cations and concentrating them at the mouth of the narrow membrane-spanning pore, thereby enhancing the overall efficiency of cation transport. The gate of the channel, near the middle of the membrane, appears to be made by hydrophobic side-chains creating a girdle around the pore. In the absence of polar groups to provide electrostatic stabilization, the ion will retain its hydration shell and then be too large to pass through the central hole. The close-to-open transition of the channel is mediated by a twisting of the a subunits between the binding pockets and the membrane. This weakens the stability of the gate, favouring the alternative 'open' configuration of M2 segments, where the pore is wider near the middle of the membrane and most constricted at the cytoplasmic membrane surface. The constriction zone, at the cytoplasmic membrane surface, appears to be shaped largely by polar residues and/or backbone groups. The polar surfaces in this region, and nearby charged groups, provide a polarizing environment that should facilitate selective transport by stabilizing passing cations.

14

UNWIN

Acknorvledgements The staff of the Marine Station at Roscoff, France are warmly thanked for their continued interest and for the frequent supply of Torpedo marmorata electric rays. The research has been supported in part by grants from the National Institutes of Health (GM-61941) and the European Commission (BIO4-CT98-0110), and by funds from the Louis Jeantet Foundation.

References Adams PD, Arkin IT, Engelman DM, Brunger AT 1995 Computational searching and mutagenesis suggest a structure for the pentameric transmembrane domain of phospholamban. Nat Struct Biol 2:154-162 Adcock C, Smith GR, Sansom MSP 1998 Electrostatics and ion selectivity of ligand-gated channels. Biophys J 75:1211-1222 Berriman J, Unwin N 1994 Analysis of transient structures by cryo-microscopy combined with rapid mixing of spray droplets. Ultramicroscopy 56:241-252 Blanton MP, Dangott LJ, Raja SK, Lala AK, Cohen JB 1998 Probing the structure of the nicotinic acetylcholine receptor ion channel with the uncharged photoactivable compound [3H]diazofluorene. J Biol Chem 273:8659-8668 Chang G, Spencer RH, Lee AT, Barclay MT, Rees DC 1998 Structure of the MscL homolog from Mycobacterium tuberculosis: a gated mechanosensitive ion channel. Science 282:2220-2226 Charnet P, Labarca C, Leonard RJ et al 1990 An open-channel blocker interacts with adjacent turns of a-helices in the nicotinic acetylcholine receptor. Neuron 4:87-95 Corringer PJ, Bertrand S, Galzi JL, Devillers-Thiery A, Changeux JP, Bertrand D 1999 Mutational analysis of the charge selectivity filter of the a7 nicotinic acetylcholine receptor. Neuron 22:831-843 Filatov GN, White MM 1995 The role of conserved leucines in the M2 domain of the acetylcholine receptor in channel gating. Mol Pharmacol 48:379-384 Finer-Moore J, Stroud RM 1984 Amphipathic analysis and possible formation of the ion channel in an acetylcholine receptor. Proc Natl Acad Sci USA 81:155-159 Gunthorpe MJ, Lummis SC 2001 Conversion of the ion selectivity of the 5-HT3A receptor from cationic to anionic reveals a conserved feature of the ligand-gated ion channel superfamily. J Biol Chem 276:10977-10983 Imoto K, Busch C, Sakmann B et al 1988 Rings of negatively charged amino acids determine the acetylcholine receptor channel conductance. Nature 335:645-648 Imoto K, Konno T, Nakai J, Wang F, Mishina M, Numa S 1991 A ring of uncharged polar amino acids as a component of channel constriction in the nicotinic acetylcholine receptor. FEES Lett 289:193-200 Keramidas A, Moorhouse AJ, French CR, Schofield PR, Barry PH 2000 M2 pore mutations convert the glycine receptor channel from being anion- to cation-selective. Biophys J 79:247-259 Labarca C, Nowak MW, Zhang H, Tang L, Deshpande P, Lester HA 1995 Channel gating governed symmetrically by conserved leucine residues in the M2 domain of nicotinic receptors. Nature 376:514-516 Law RJ, Forrest LR, Ranatunga KM, La Rocca P, Tieleman P, Sansom SP 2000 Structure and dynamics of the pore-lining helix of the nicotinic receptor: MD simulations in water, lipid bilayers and transbilayer bundles. Proteins 39:47-55 Lester HA 1992 The permeation pathway of neurotransmitter-gated ion channels. Annu Rey Biophys Biomol Struct 21:267-292 Malashkevich VN, Kammerer R A, Efimov VP, Schulthess T, Engel J 1996 The crystal structure of a five-stranded coiled coil in COMP: a prototype ion channel? Science 274:761-765

ACETYLCHOLINE-GATED CHANNEL

15

Miyazawa A, Fujiyoshi Y, Stowell M, Unwin N 1999 Nicotinic acetylcholine receptor at 4.6 A resolution: transverse tunnels in the channel wall. J Mol Biol 288:765-786 Nolte HJ, Rosenberry TL, Neumann E1980 Effective charge on acetylcholinesterase active sites determined from the ionic strength dependence of association rate constants with cationic ligands. Biochemistry 19:3705-3711 Ripoll DR, Faerman CH, Axelsen PH, Silman I, Sussman JL 1993 An electrostatic mechanism for substrate guidance down the aromatic gorge of acetylcholinesterase. Proc Natl Acad Sci USA 90:5128-5132 Unwin N 1989 The structure of ion channels in membranes of excitable cells. Neuron 3:665-676 Unwin N 1993 Nicotinic acetylcholine receptor at 9A resolution. J Mol Biol 229:1101-1124 Unwin N 1995 Acetylcholine receptor channel imaged in the open state. Nature 373:37-43 Unwin N 2000 The Croonian Lecture 2000. Nicotinic acetylcholine receptor and the structural basis of fast synaptic transmission. Philos Trans R Soc Lond B Biol Sci 355:1813-1829 Villarroel, A, Herlitze S, Koenen M, Sakmann B 1991 Location of a threonine residue in the asubunit M2 transmembrane segment that determines the ion flow through the acetylcholine receptor channel. Proc R Soc Lond B Biol Sci 243:69-74 White BH, Cohen JB 1992 Agonist-induced changes in the structure of the acetylcholine receptor M2 regions revealed by photoincorporation of an uncharged nicotinic competitive antagonist. J Biol Chem 267:15770-15783 Wilson GG, Karlin A 2001 Acetylcholine receptor channel structure in the resting, open, and desensitized states probed with the substituted-cysteine-accessibility method. Proc Natl Acad Sci USA 98:1241 1248

DISCUSSION Roux: I am a little puzzled about the relationship between the charge distribution in the pore and how this channel is functioning. I would expect it to carry an inward current of cations normally. So why would negative charge far on the intracellular side be functionally useful? Unwin: Ions do go through in both directions. When the channel first opens, the driving force is mainly for Na+ to go towards the interior of the cell. But as the voltage changes across the membrane, there is an increasingly strong driving force for K+ to go in the opposite direction. The negative charge is relevant for encouraging K+ to go more efficiently through the channel. I suspect that there is an analogous process operating in the extracellular domain. Of course, you notice that it is a long cylinder 20 A in diameter; this means that residues lining the extracellular wall will have an effect on ions going through. One knows from looking at the sequences that each of the subunits contains on average about the same net negative charge on their extracellular surfaces as acetylcholinesterase. The negative charges on acetylcholinesterase are very important for bringing the acetylcholine cation into the active site. One could imagine the negative charges on the extracellular surface of the receptor, doing a similar thing to encourage cations to go in that direction as well. Lambert: The M3/M4 loop in the GABA receptor usually has some putative phosphorylation sites. Could these also interfere with cation conduction?

16

DISCUSSION

Unwin: I don't know the GABA receptor well enough to know exactly where they are. If the phosphorylation sites are located close to those windows, you would imagine they could have a pronounced effect. Corringer: I have a question about the location of the ACh binding site. If I understood you correctly, you see the binding site within the subunits, whereas in the ACh binding protein (AChBP) structure the binding site appeared to be at the interface between the subunits. Could the cavities seen in your images correspond to the cavities that are within the subunits in the AChBP structure, but which are not actually the acetylcholine binding sites? Unwin: Certainly, in Titia Sixma's structure, the binding site appears to be close to the interface, although we haven't seen ACh in the binding site yet. We assumed that the cavities in a subunits were the binding sites. In the light of Titia's structure, this may not be entirely accurate. There are definitely pronounced cavities in the a subunits, but it looks like the actual binding site is closer to the interface than we thought. Corringer: Perhaps they are not. We just compared the pictures, and the cavities within the AChBP subunits seem to fit nicely with the cavities you see in the a subunits by electron microscopy. Unwin: The cavities in the a subunits of the receptor are opened relative to those in AChBP. We know this because when we fit the polypeptide chains on one side of the cavity, those on the other side start moving into the cavity. However, when you look at the subunit found between the a subunits the fit is better, and this hasn't got much of a cavity at all. Sixma: Do you think this could be because we have HEPES bound and you are looking at something without anything bound, so we are looking at different states? Or is this an intrinsic difference between the nicotinic receptor and AChBP? Unwin: One could say that you are looking at the desensitized state and we are looking at the closed state. This is the analogy we would use. Perozo: Perhaps it looks different to the trained eye, but I can also see cavities in the other subunits, not only in the a subunits. Unwin: It is all a matter of degree. The a subunits have cavities with water in. They are about as prominent as the windows into the cytoplasmic vestibule, and we know that ions have to go through the windows otherwise the channel wouldn't work. The a subunit cavities are about 8 A across, and they probably have at least 10 water molecules in them. The cavity on the subunit that runs between the a subunits may have no water at all. The other two subunits do not really have cavities; they are more like grooves. We think that the cavities may be used for allowing ACh to access the binding site, because it will be drawn into the extracellular vestibule along with the other cations. Sixma: I think that is unlikely.

ACETYLCHOLINE-GATED CHANNEL

17

Unwin: I don't think you can say it is unlikely, because the structure is different in this region from that of the AChBP. Matbie: In the channel you are looking at, the muscle-type nicotinic receptors, the ions go equally well into the channel as they do out of the channel. But this is not true of some of the neuronal nicotinic receptors, which rectify and therefore don't carry ions outwards as easily. Do you think the cytoplasmic vestibules would be the best place to look for regions of the channel where there is this differential effect? Unwin: Yes, because individual residues can have a big influence here. Mindell: Related to that question, you hinted that there are not many sitedirected mutagenesis data in these areas, in the basket strands. Are there any mutations that have an effect? One would predict that there should be individual mutations of charged residues that have profound effects on ion conduction, and this may make a more strongly rectifying channel. Miller: Dr Sine's group (Bouzat et al 1994) have shown that changing the MA region from the gamma subunit sequence into that of the epsilon subunit changes channel lifetime. Unwin: If you don't see any effect, you could argue that there may be a lot of weak interactions adding up to produce a big overall effect. Mutation of just one residue may not do much. You may need to do a number of mutations at once, but this could of course change the structure of the channel and make the results uninterpretable. Sansom: Comparing your closed-state structure with your open model, what sort of change in radius do you see? Is it about 7 A in diameter in the narrowest region of the closed model? It looked a little bit larger in the open-state structure. Unwin: I reckon that it increases its diameter by about 4 A in the middle of the membrane when this channel opens. This makes it similar to the cavity in the K+ channel. Roux: If the closed channel has a pore of 7 A lined by hydrophobic residues, this wouldn't create an insurmountable barrier to permeation. This means that if a large potential was applied, there should be some detectable current. Has this been observed? Sansom: At least in simple models, I'm not even sure that water would enter, if the channel is purely hydrophobic and 8 A long. Roux: But even if the pore is 7 A wide? Sansom: Yes.

Perozo: Wouldn't you see streaming potentials? Mindell: If ions aren't going through, then perhaps you wouldn't see streaming potentials, but you should see water permeability. Jordan: Going back to the windows, are you saying that you think that when the ion actually passes through the window, it loses part of its hydration shell, or is

18

DISCUSSION

there concerted movement of the residues so that it can maintain its hydration shell? Unwin: When we fit the helices there, we know that the windows are very narrow. The widest part of the widest one is about 9 A. The other two are 8 A or less, so it seems that the ions do have to lose some of their hydration shell. Choe: I have a question about the outer rim. If we imagine we have b-stranded sheets here, the length of the transmembrane b sheets will require you to have more crossings, unless you tilt your strands sideways. How do you reconcile this with the fact that you have only three crossings to make based on sequences? If these are b strands how would they fit with your structure? Also, you mentioned that two molecules are disulfide-bonded in the tube sample: are these natural disulfide bonds? Unwin: Yes, in Torpedo disulfide bonds are naturally formed between the d subunit of neighbouring receptors — I don't think human muscle has them. The strands going through Ml probably fold about three times. This fits very well with the enterotoxin structure that Titia Sixma solved a long time ago in the bottom part of the membrane. If you fit this into the densities, the a helices in the two structures superimpose to within an angstrom or so. Corringer: From Coen's work it seems that M3 and M4 are predominantly coiled in an a helix. Can you see any rod-shaped structures in the transmembrane regions? Ijnwin: In the top part of the membrane there may be a bit of a helix. We don't really know. Miller: It is not just photolabelling that suggests that M4 is an a helix. LasaldeDominicci and McNamee (Tamamizu et al 2000) have recently performed tryptophan scanning of M4 and their results fit pretty well with the structure being an a helix. I don't recall anyone reporting an alternating pattern such as one might expect from a b sheet. Lambert: You mentioned that the a subunits appeared to have rotated clockwise in the open conformation. Do the other subunits also move? Are all of them going clockwise? I ask because Myles Akabas interpreted his results, in a model of the GABAA receptor, as indicating that one of the subunits moved clockwise and the other four subunits moved anticlockwise (Horenstein et al 2001). Do you have any information on the movement of the other subunits in the open conformation? Unwin: What we observed was that all the subunits change at the level of the binding sites, and then from the binding sites down to the membrane, the two a subunits seem to behave differently to the others. They seem to have rotations in the order of 4° clockwise. The other showed no significant change. Perozo: Do you detect any accommodation from the surrounding transmembrane segments? In the b barrel are you proposing that the structure also changes around the M2 helices?

ACETYLCHOLINE-GATED CHANNEL

19

Unwin: We couldn't detect any changes in the b barrel between the two. We had poor resolution and we took a fivefold average. I haven't looked closely at the structure before it was averaged, so there could have been some localized changes that got smeared out. Corringer: 1 have a question about the transition of activation and desensitization. Do you see any change in the structure of the cytoplasmic domain upon desensitization or activation? Unwin: Not on activation. We haven't looked in great detail, though. Corringer: So it could be a gating mechanism as well. There could therefore be two selectivity filters. Unwin: That is possible. Papazian: This question goes back to one that was asked earlier. If, as you propose, the outer rim of the transmembrane segment of the receptor is b structure, given what you know about the internal structure would you care to speculate about the number of strands that might be required to surround that structure? Unwin: It fits about the same number as enterotoxin. Choe: You mentioned the disulfide-bonded pentamers. Where would the disulfide bond be? Is it in the transmembrane domain? Unwin: No. Jean-Luc Popot and Don Engelman showed it to be on the extracellular side. It is at the cytoplasmic end of the d subunits. It is a very floppy linkage: we are often looking at dimers of receptors, and they can be 100 A apart, yet they are still dimers. Choe: Have you tried putting the entire pentamer into your electron density map? Unwin: No, we haven't tried this. Miller: Could you comment on Karlin's cysteine scanning and your model of how the helices swing and open? Unwin: There have been a number of different interpretations of the cysteine scanning mutagenesis experiments. One that fits our results very well is Paul Slesinger's analysis of the 5-HT3 receptor. As far as I can see this is 100% consistent, except they seem to have a helix going on a bit beyond the threonine 244 in their case. Mitra: According to Karlin's model the position of the gate is somewhat different from the location of the critical leucine in your model. Unwin: I think he agrees that this is a flexible part of the structure. He wouldn't agree that it is where the gate is located. They think that the gate for the closed channel is at the cytoplasmic surface. There was a recent paper in which they have done the cysteine scanning on the desensitized structure (Wilson & Karlin 2001). In the desensitized structure they have another gate that extends from the cytoplasmic surface up to that leucine. This is a critical region of the protein;

20

DISCUSSION

whatever you do to it will change things more than just simply substituting the residue. That leucine is the only part in our structure where the M2 helices come close to each other. It is a sensitive part of the structure. Miller: There is lots of space around the M2 helices in your model. If you were photolabelling with a hydrophobic agent like TID (3-[trifluoromethyl]-3-[miodophenyl]diazirine), would you expect to get labelling in the regions behind M2 or, as is observed, just the residues in the channel lumen? Unwin: Parts of the M2 segment have got water behind them, which does complicate things. Fu: I have a hard time understanding the channel gate. Does the channel gating process require bending a helices to open up an aqueous pore? As we know, the geometries of a helices are determined by a hydrogen-bonding network made by main chain carbonyl oxygens and nitrogens. When you distort a helices by disputing hydrogen bonds, you may have to disrupt the entire H-bonding network, not just a few local bonds. This means that channel gating may need a huge amount of activation energy in order to rearrange all H bonds in the distorted helices. Unwin: It is like a swivel. Sansom: Firstly, you might not have to actually break the H bonds to give a distortion of the helix. You could have a cumulative small change in several residues, which would at least give a banana-shaped helix. Also, if there is some water floating around there, then the cost of breaking a hydrogen bond in the helix is not going to be so large, as an alternative H-bond to a water may be formed. Choe: That gating does not involve breaking or reforming the helical H bonds. The helix is rotating in and out of the pore, so the helical H bonds have nothing to do with the gating energies. Unwin: I think it is a swivel. I would say that the residues are probably facing in the same direction in either case. Miller: You are drawing that essentially as a V-shaped helix in the cartoon. Is the axis of rotation along one of the edges of the V? Unwin: It is complicated. The top half moves backwards slightly, and the bottom half swivels inwards. Sansom: The other thing that might be consistent with the hinge is the profound effect of putting a proline in, as is seen in the slow channel syndrome. One of these mutations is a proline, just above where the leucine is (Ohno et al 1995). This has a pronounced effect on gating that would be consistent with having a hinge in that region. Tieleman: Do you have any idea which part of the protein interacts with the M2 helix? Unwin: We don't know. Presumably Ml, M3 and M4!

ACETYLCHOLINE-GATED CHANNEL

21

References Bouzat C, Bren N, Sine SM 1994 Structural basis of the different gating kinetics of fetal and adult acetylcholine receptors. Neuron 13:1395-1402 Horenstein J, Wagner DA, Czajkowski C, Akabas MH 2001 Protein mobility and GABAinduced conformational changes in GABAA receptor pore-lining M2 segments. Nat Neurosci 4:477-485 Ohno K, Hutchinson DO, Milone M et al 1995 Congenital myasthenic syndrome caused by prolonged acetylcholine receptor channel opening due to a mutation in the M2 domain of the e subunit. Proc Natl Acad Sci USA 92:758-762 Tamamizu S, Guzman GR, Santiago J, Rojas LV, McNamee MG, Lasalde-Dominicci JA 2000 Functional effects of periodic tryptophan substitutions in the alpha M4 transmembrane domain of the Torpedo californica nicotinic acetylcholine receptor. Biochemistry 39:4666-4673 Wilson G, Karlin A 2001 Acetylcholine receptor channel structure in the resting, open, and desensitized states probed with the substituted-cysteine-accessibility method. Proc Natl Acad Sci USA 98:1241 4248

The 2.7 A structure of AChBP, homologue of the ligand-binding domain of the nicotinic acetylcholine receptor Katjusa Brejc*, Willem J. van Dijk*, August B. Smitt and Titia K. Sixma*1 * Division of Molecular Carcinogenesis, Netherlands Cancer Institute, Plesmanlaan 121, 1066 CXA msterdam, and' Department of Molecular and Cellular Neurobiology, Research Institute Neurosciences Vrije Universiteit, Faculty of Biology, De Boelelaan 1087, 1081 HV A msterdam, The Netherlands

A bstract. Acetylcholine binding protein (AChBP) is a novel protein with high similarity to the extracellular domain of the nicotinic acetylcholine receptor. It is secreted from glia cells in the freshwater snail, Lymnaeastagna/is, where it modulates neuronal transmission. AChBP forms homopentamers with pharmacology that resembles the a7 nicotinic receptors. In the crystal structure of AChBP at 2.7 A, each protomer has a modified immunoglobulin fold. Almost all residues shown to be involved in ligand binding in the nicotinic receptor are found in a pocket at the subunit interface. This pocket is lined with aromatic residues, and filled with a HEPES buffer molecule. The AChBP crystal structure explains many of the biochemical studies on the nicotinic acetylcholine receptors. Surprisingly the interface between protomers is relatively weakly conserved between family members in the superfamily of pentameric ligand-gated ion channels. The lack of conservation has implications for the mechanism of gating of the ion channels. 2002 Ion channels—from atomic resolution physiology to functionalgenomics. Wiley, Chichester (Novartis Foundation Symposium 245) p 22-32

The nicotinic acetylcholine receptor (nAChR) has long been the prototype of ligand-gated ion channels, also known as Cys-loop receptors. This superfamily also includes the GABAA and GABAC, the glycine and the 5-HT3 serotonin receptors (Corringer et al 2000). These transmembrane receptors form pentamers of related subunits, in which each subunit consists of an N-terminal ligand-binding domain, four membrane spanning regions (M1-M4) and a short intracellular 1 This chapter was presented at the symposium by Titia Sixma, to whom correspondence should be addressed.

22

THE 2.7 A STRUCTURE OF AChBP

23

region (Corringer et al 2000). Extensive biochemical studies have defined the complicated ligand-binding sites that are found at the subunit interface. The binding of ligands goes through a complex cycle of activation and desensitization (Karlin & Akabas 1995, Changeux & Edelstein 2001). Structurally the proteins are well defined by the work of Nigel Unwin and colleagues, who have studied the Torpedo receptor up to a resolution of 4.6 A (Miyazawa et al 1999, Unwin 2000), but an atomic model has been lacking. In Guus Smit's group, at the Free University in Amsterdam, the fresh water snail Lymnaea stagnates is studied as a model system. It has large neurons that have been characterized very extensively. This precise knowledge allows the in vitro culture of specific, cholinergic synapses (Feng et al 1997). Upon electrophysiological analysis of such an in vitro system an interesting effect was observed when the synapse was cultured in the presence of glia cells. It was found that glia cells suppress the synaptic transmission (Smit et al 2001). After an extensive series of studies Smit et al (2001) showed that this is mediated by secretion of a novel protein, called acetylcholine binding protein (AChBP). The AChBP protomer contains 210 residues and shows high homology (~ 24% sequence identity) with the N-terminal domain of the a subunits of the nAChRs, but it lacks the transmembrane region entirely (Smit et al 2001). The conserved regions are the hallmarks of the superfamily of ligand-gated ion channels (15-20% sequence identity with e.g. GABA and 5-HT3 serotonin receptors). The recombinant AChBP protein, expressed in yeast Pichia pasforis, assembles into stable homopentamers. Furthermore, AChBP contains all the arnino acid residues needed for the ligand binding, showing pharmacology that is similar to a homopentameric nAChR, such as the neuronal a7 receptor (Anand et al 1993). Thus it binds better to nicotine than to acetylcholine (ACh) and binds very well to epibatidine. Nicotinic antagonists are binding as expected, but it also binds to some of the muscarinic antagonists (Smit et al 2001), an effect that has also been observed for the homopentameric a9 receptor (Elgoyhen et al 1994). We can therefore conclude that AChBP is a soluble homologue of a ligand-binding domain of the nicotinic receptors. The recombinant AChBP has been crystallized and its structure was determined with phases from multi-wavelength anomalous diffraction on a Pb-derivative in two different crystal forms (Brejc et al 2001). The electron density was improved by non-crystallographic and multi-crystal averaging across three different crystal forms. The AChBP model was refined in the P422}2 crystal form with one pentamer per asymmetric unit at 2.7 A resolution. The AChBP homopentamer, composed of five protomers, forms a doughnutlike structure with a radius of 80 A and height of 62 A (Fig. 1), conforming to the electron microscopy data for the nAChR (Miyazawa et al 1999). Each AChBP protomer has a modified imrnunoglobulin (Ig) topology (Fig. 2A) with relatively

24

BREJC ET AL

FIG. 1. Pentamer of AChBP, viewed along the fivefold axis. Possible subunit organization for the Torpedo nAChR is indicated.

long and twisted strands. Both b sheets have at least one extra b strand compared to the standard Ig fold (Fig. 2B). An Ig fold was predicted by Le Novere et al (1999) and Corringer et al (2000), but the additional strands were not predicted and the binding site is in a different location. Where the Ig fold has a so-called 'tyrosine cornerstone', a residue that is conserved across different Ig families (Bork et al 1994), a disulfide bridge (Cysl23-Cysl36) is found in AChBP. This disulfide bond is absolutely conserved in the superfamily of pentameric ligand-gated ion channels and has importance for the stabilization of the fold as shown by Blount & Merlie (1990) and Green & Wanamaker (1997). Its structural similarity to the Ig 'tyrosine cornerstone' may indicate that this disulfide bond has a structural role in keeping the two b sheets together. The residues involved in ligand binding in the muscle and Torpedo nAChR subtypes have been extensively investigated, and are found at the interface between an a subunit and a neighbouring subunit, forming primary and

THE 2.7 A STRUCTURE OF AChBP

FIG. 2. (A) Protomer fold of AChBP. The protein has an N-terminal helix followed by a b sandwich. The C-terminal residue would be the start of the transmembrane receptor in nAChRs. (B) Similar orientation of the immunoglobulin chain. Note the location of the 'tyrosine cornerstone'. This residue is not present in AChBP, but is replaced by the conserved disulfide bridge.

complementary parts of the binding pocket, respectively (Corringer et al 2000). The a-subunit residues important for binding are clustered in 'loop' regions, A (Galzi et al 1990), B (Dennis et al 1988) and C (Dennis et al 1988, Kao et al 1984, Sine et al 1994). The a subunit contributes the primary binding site, consisting of loop A, B and C. The neighbouring subunit, g or d, contributes 'loops' D (Chiara & Cohen 1997), E (Chiara & Cohen 1997, Chiara et al 1999, Sine 1997) and F (Czajkowski & Karlin 1995) (Table 1); in homopentamers the equivalent loops will be contributed by a subunits. In the AChBP structure all these 'loop' regions are indeed found to form a single pocket region at the subunit interface, with 'loops' A—C contributed from one subunit and D-F from another (Fig. 3). 'Loops' A, B, C and F are indeed

26

BREJCET AL TABLE 1 Numbering of ligand binding residues in AChBP, compared to Torpedo and muscle a1, g and d and human a7-subunits Loop

ACbBP

a7

A

Tyr89 Trpl43 Tyrl85 Cysl87

Tyr93 Trpl49 Tyrl90 Cysl92

Cysl88 Tyrl92

Cysl93 Tyrl98

B C

D

Trp53

E

Gln55 Argl04 Vall 06 Leull2 Metll4

F

7

d

a-

Tyr93 Trpl49 Tyrl88 Cysl90

Trp55 Glu57 Leu 109

Tyrlll Tyrll7

Trp57 Thr59 Leulll

Cysl91 Tyrl95 Trp55 Gln57 Leu 109 Asn111

Leull9

Argll 3 Thrl19 Leu 121

Gln117 Leull9

Aspl74

Asp 180

Asp 164

Tyrl64

structural loops in the structure, but the residues, contributed from 'loops' D and E are found on b strands. The pocket is lined by aromatic residues of loop A-D, while the hydrophobic components of the residues in loop E form the lid of the pocket. The Tyrl64 that is contributed from the loop F region is not conserved, and the variation in length and sequence in this area indicates that the region may form a somewhat different fold in other family members, contributing another residue to the binding site. In the experimental electron density extra features were observed in the ligandbinding site. This density was attributed to a HEPES buffer molecule that was used in crystallization. HEPES was found to bind with an IC50 of 100 mM, which could cause partial occupancy at the concentrations used. The HEPES buffer has a positive charged nitrogen atom, similar to acetylcholine (ACh). This atom stacks on Trpl43, in a similar fashion as expected for acetylcholine stacking on the equivalent Trpl49 in nAChR (Zhong et al 1998). The position of the HEPES molecule could not be unambiguously refined. It is likely that the lack of specific hydrogen bonds to this weakly binding molecule causes variability in position, while the site is not fully occupied as well. However, its general position presents a reasonable preliminary model for the ACh binding. The HEPES molecule is almost entirely buried in the structure, with both subunits contributing equal interface areas.

THE 2.7 A STRUCTURE OF AChBP

FIG. 3. Stereo representation of the ligand binding site of AChBP, principal side in light grey, complementary side in dark grey. For the equivalent residues in other receptors see Table 1.

The crystal structure of AChBP adds some additional data to the ongoing debate on the relative arrangement of subunits in the muscle and Torpedo form of the nAChR (Umvin 2000). According to our structure, the location of the binding site is such that any principal binding site has its complementary binding site in the anticlockwise direction (Fig. 1). There is considerable sequence conservation in the superfamily of ligand-gated ion channels (LGICs). The conserved residues are mostly found within the protomer, contributing hydrophobic residues to the core and hydrophilic residues to structurally important sites. This conserved core structure indicates that the overall fold of the other family members will greatly resemble the AChBP fold. The only conserved regions that are lacking in AChBP are the residues that form the so-called Cys loop. This region, connecting the absolutely conserved cysteines (123—136 in AChBP, 128—142 in Torpedo receptor) is a conserved hydrophobic region in the LGIC superfamily, but in AChBP it is one residue shorter and mostly hydrophilic. The loop is found on the membrane facing side (see Fig. 2) of AChBP, and probably interacts with the transrnembrane domain in the receptor, a region that is absent in AChBP. A major surprise in the structure is the lack of conservation of the subunit interface across the superfamily of LGICs. The residues that are forming the subunit interface in AChBP vary between superfamily members, including changes in character from charged to hydrophobic and vice versa, without the necessary complementary changes. Thus it can be concluded that the precise interactions in the interface are not very important for preserving the pentamer

28

BREJC ET AL

formation. Moreover they are apparently not essential for maintaining the mechanism of opening a channel in response to a ligand binding. This has implications for the possible mechanism of gating. Once a ligand is bound to the extracellular domain of the receptor, this will have to cause a signal that is relayed to the transmembrane domain. One possible mechanism is a pivoting motion about the ligand-binding site of the entire protomer. Such a rigid body motion would presumably cause a rearrangement in the transmembrane domain. Other mechanisms can be imagined, but the lack of conservation of the subunit interface would seem to imply that they may well be found in the protomer. Possibilities involve a movement of the 'loop' C b hairpin and a rearrangement of the b sandwich structure. It will be interesting to see whether AChBP itself will be useful as a model to study such ligand-dependent movements. AChBP does not have a transmembrane domain, and presumably would not need such a movement for its function. Further studies are needed to show whether this will be the case. However, the crystal structure of AChBP will undoubtedly be a starting point for studying such movements since it shows which regions of the sequence are found at certain sites in the structure and thus will provide new ideas for experiments. References Anand R, Peng X, Lindstrom J 1993 Homomeric and native a7 acetylcholine receptors exhibit remarkably similar but non-identical pharmacological properties, suggesting that the native receptor is a heteromeric protein complex. FEES Lett 327:241-246 Blount P, Merlie JP 1990 Mutational analysis of muscle nicotinic acetylcholine receptor subunit assembly. J Cell Biol 111:2613-2622 Bork P, Holm L, Sander C 1994 The immunoglobulin fold. Structural classification, sequence patterns and common core. J Mol Biol 242:309-320 Brejc K, van Dijk WJ, Klaassen RV et al 2001 Crystal structure of an ACh-binding protein reveals the ligand-binding domain of nicotinic receptors. Nature 411:269-276 Changeux JP, Edestein SJ 2001 Allosteric mechanism in normal and pathological nicotinic acetylcholine receptors. Curr Opin Neurobiol 11:369-377 Chiara DC, Cohen JB 1997 Identification of amino acids contributing to high and low affinity dtubocurarine sites in Torpedo nicotinic acetylcholine receptor. J Biol Chem 272:32940-32950 Chiara DC, Xie Y, Cohen JB 1999 Structure of the agonist-binding sites of the Torpedo nicotinic acetylcholine receptor: affinity-labeling and mutational analyses identify gTyr-111 dArg-113 as antagonist affinity determinants. Biochemistry 38:6689-6698 Corringer PJ, Le Novere N, Changeux JP 2000 Nicotinic receptors at the amino-acid level. Annu Rev Pharmacol Toxicol 40:431-458 Czajkowski C, Karlin A 1995 Structure of the nicotinic receptor acetylcholine-binding site. Identification of acidic residues in the d subunit within 0.9 nm of the a subunit-binding site disulfide. J Biol Chem 270:3160-3164 Dennis M, Giraudat J, Kotzyba-Hibert F et al 1988 Amino acids of the Torpedo marmorata acetylcholine receptor alpha subunit labeled by a photoaffinity ligand for the acetylcholine binding site. Biochemistry 27:2346-2357

THE 2.7 A STRUCTURE OF AChBP

29

Elgoyhen AB, Johnson DS, Boulter J, Vetter DE, Heinemann S 1994 Alpha 9: an acetylcholine receptor with novel pharmacological properties expressed in rat cochlear hair cells. Cell 79:705-715 Feng ZP, Klumperman J, Lukowiak K, Syed NI 1997 In vitro synaptogenesis between the somata of identified Lymnaea neurons requires protein synthesis but not extrinsic growth factors or substrate adhesion molecules. J Neurosci 17:7839-7849 Galzi JL, Revah F, Black D et al 1990 Identification of a novel amino acid alpha-tyrosine 93 within the cholinergic ligands-binding sites of the acetylcholine receptor by photoaffinity labeling. Additional evidence for a three-loop model of the cholinergic ligands-binding sites. J Biol Chem 265:10430-10437 Green WN, Wanamaker CP 1997 The role of the cysteine loop in acetylcholine receptor assembly. J Biol Chem 272:20945-20953 Kao PN, Dwork AJ, Kaldany RR et al 1984 Identification of the alpha subunit half-cystine specifically labeled by an affinity reagent for the acetylcholine receptor binding site. J Biol Chem 259: 11662-11665 Karlin A, Akabas MH 1995 Toward a structural basis for the function of nicotinic acetylcholine receptors and their cousins. Neuron 15:1231—1244 Le Novere N, Corringer PJ, Changeux JP 1999 Improved secondary structure predictions for a nicotinic receptor subunit: incorporation of solvent accessibility and experimental data into a two-dimensional representation. Biophys J 76:2329-2345 Miyazawa A, Fujiyoshi Y, Stowell M, Unwin N 1999 Nicotinic acetylcholine receptor at 4.6 A resolution: transverse tunnels in the channel wall. J Mol Biol 288: 765-786 Sine SM 1997 Identification of equivalent residues in the 7,g,dand e subunits of the nicotinic receptor that contribute to a-bungarotoxin binding. J Biol Chem 38:23521—23527 Sine SM, Quiram P, Papanikolaou F, Kreienkarnp HJ, Taylor P 1994 Conserved tyrosines in the alpha subunit of the nicotinic acetylcholine receptor stabilize quaternary ammonium groups of agonists and curariform antagonists. J Biol Chem 269:8808-8816 Smit AB, Syed NI, Schaap D et al 2001 A glia-derived acetylcholine-binding protein that modulates synaptic transmission. Nature 411:261-268 Unwin N 2000 The Croonian Lecture 2000. Nicotinic acetylcholine receptor and the structural basis of fast synaptic transmission. Phil Trans R Soc Lond B 355:1813-1829 Zhong W, Gallivan JP, Zhang Y et al 1998 From ab initio quantum mechanics to molecular neurobiology: a cation-pi binding site in the nicotinic receptor. Proc Natl Acad Sci USA 90:90319035

DISCUSSION Wallace: Does AChBP show the toxin binding specificity of the neuronal or the neuromuscular channel, and have you tried to look at toxin binding? Sixma: We haven't looked very carefully with all the different conotoxins yet. We are collaborating with a group in Israel on bungarotoxin binding, and they have a nice peptide structure of the C-loop contacts with this toxin (Harel et al 2001). It seems to fit well with our structure. Mindell: Now that you know about the AChBP in snails, when you look back at the genomes that have been solved can you find homologues in other phyla? Sixma: My collaborator, Guus Smit, has looked at this and tells me that it is difficult because the databases are not always curated particularly carefully. One is

30

DISCUSSION

never sure whether something is a bona fide N-terminal domain alone. So far we haven't found it. Schrempf: Have you any results that prove an interaction with another nicotinic receptor in the membrane? Sixma: No, we have not. It is possible, but we don't need to invoke such a mechanism to see the results we have obtained. There are nicotinic receptors in the glial cells that secrete AChBP in Guus Smit's in vitro assays. In addition, the glial cells themselves have nicotinic receptors. FM: How do the binding sites of your AChBP compare with acetylcholinesterase? This binding site also has many side chains. Sixma: The size of the cavity is roughly similar, but all the residues are different. I had a hard time deciding how to superimpose it. I decided on the tryptophan that stacks the positive ion. Then you see that the size of the cavity is the same and all the residues come from different sides. Choe: I have difficulty in understanding how you decide the subunit assembly arrangements, in order to predict the ACh receptor subunits. The ACh receptors are not homopentameric, and you mentioned that there is no high sequence conservation at the interface, yet your model is relying on the a subunit sequences in segregating principal sites and complementary sites. This doesn't exist in a common receptor channel, so you have to decide which one you want to keep as the interface. Sixma: In our structure, every subunit has both a principal and a complementary site. In modelling the arrangement, I tried to show the two most commonly debated models. The only thing we can do is exclude models that have the position of the ligand binding site clockwise with respect to its a subunit. I am not trying to predict anything about where a g or a d subunit should be; we are just using literature data. The only thing we are saying is that any model that has the binding site clockwise to an a subunit can not be true. Choe: That preserves the principal site as the ligand binding site. Sixma: Yes, that is the definition of the principal site. Sattelle: It appears that the secreted AChBP is acting as some sort of ACh sponge, mopping up extracellular ACh. Is much known about the dissociation kinetics of the protein compared with native receptors? Sixma: We don't have very detailed kinetic data yet. Choe: Thinking about the AChBP versus the ACh receptor, the physiological role of the binding protein is not necessarily transmitting conformational change, it is just soaking up the ligand. If you soak out HEPES and replace it with some other buffers, would you expect to see conformational changes? Sixma: With the Hill coefficients being less than 1, you would expect it to be different from the receptor, because there is no cooperativity (at least in this region). On the other hand, there is no transmembrane region, and this could

THE 2.7 A STRUCTURE OF AChBP

31

transmit the cooperativity in the receptor. We don't know whether AChBP will show similar changes. Choe: Have you tried soaking out HEPES or replacing it with other buffers? Six ma: We can grow the crystals in the absence of HEPES and in the absence of Ca2+. They grow in a similar way. Perozo: How good a ligand is HEPES for the ACh receptor? Sixma: I don't know. Umvin: What is the occupancy of the HEPES? Sixma: I think it is not fully occupied. The density is variable around the peritamer. We are trying crystallization experiments with ligands, but we are stuck with terrible twinning problems of the crystals. Unwin: The reason I ask is because if it is only partly there, and the structure changes when it is not there, you might expect to see some disordered regions in the electron density maps. Sixma: We do see a little bit of disorder in the C-loop that is not fully resolved. But not a lot. On the other hand, in this 20+10+5-fold averaged structure, we can follow the run of the main chain pretty well. There is not a lot of disorder in this structure. C3n the other hand, if 20-30% of the molecules were disordered, I am not sure you would lose that density entirely. It is hard to tell. Miller: I don't recollect that anyone has reported that HEPES activates ACh receptors. The known structure-activity relationships would make it unlikely to be an agonist or competitive antagonist. Have you tried to put something as large as an N-methyl group into the putative acetylcholine-binding pocket, and if so, can you make any conclusions about the orientation of the binding? Sixma: I have avoided this because I thought everyone else would do it. I think it is a similar size to HEPES, so it should fit. For bigger ligands, I think things would have to open up, which requires conformational changes. Miller: There is a prediction from NMR experiments that the angle formed between the CH2-NMe3 bond of acetylcholine and the membrane is 40° in the desensitized form of the receptor (Williamson et al 2001). This might provide a useful constraint for modelling. Roux: I am intrigued by the evolutionary origin of AChBP. How did it occur? Did a part of the gene for the ACh receptor just separate and assume an independent role? Sixma: Guus Smit and his group are trying to address this. They have cloned a whole series of ACh receptors from Lymnaeastagualis. It seems that AChBP is more similar to Lymnaea receptors than to other receptors, which would make sense. It is very similar to the superfamily, and most similar to the nicotinic receptors. Roux: If this is just an AChBP binding domain that arose de novo, it would just have to bind ACh; in other respects it could be completely different.

32

DISCUSSION

Sixma: It is definitely a family member because it is also well related to the 5-HT3 serotonin, the GABA and the glycine receptors. Sansom: On the percentage identities to different types of nicotinic subunits, Nigel Unwin was suggesting in the structure that the AChBP seemed to match the non-a more than the a. At the sequence level, does it show a higher percentage identity to a or to non-a? Sixma: It is closer to a, particularly because it has the principal ligand binding residues. Gulbis: Is MetlH at the top of the ligand-binding pocket in the protein present in the ACh receptor, and is there any evidence that it oxidises? Sixma: It is not necessarily a methionine. It is just contacting the HEPES by the hydrophobic part of the residue. Gulbis: In view of its similarity to choline, the bound HEPES molecule could be taken to indicate the position and orientation adopted by acetylcholine. Sixma: It is certainly not preserved as a methionine throughout the family, so I don't think it is likely to be that essential. Mathie: Is there an ion permeation pathway through the AChBP? Can you get information about how ions are passed through the central pore? Sixma: It is a wide pore of 18 A, so anything should pass through. It is lined with charged residues. If you compare it with other members of the superfamily, and if you just look at residues that are actually lining the pore, their position and order changes a lot. There is a slight excess of charges that fits with the type of ions that go through between the different channels, but I am not so sure that this is meaningful, I haven't really looked at the width and where they are located. The channel isn't equally wide all the time, there is a bit of an indentation in the middle. References Harel M, Kasher R, Nicolas A et al 2001 The binding site of acetylcholine receptor as visualized in the X-ray structure of a complex between a-bungarotoxin and a mimotope peptide. Neuron 32:265-275 Williamson PTF, Watts JA, Addona GH, Miller KW, Watts A 2001 Dynamics and orientation of N+ (CD3)3-bromoacetylcholine bound to its binding site on the acetylcholine receptor. Proc Nad Acad Sci USA 98:2346-2351

The architecture of a water-selective pore in the lipid bilayer visualized by electron crystallography in vitreous ice Alok K. Mitra*, Gang Ren*, Vijay S. Reddyt, Anchi Cheng* and Alexandrine Froger* *Department of Cell Biology and Department of Molecular Biology, The Scripps Research Institute, 10550 North Torrey Pines Road, Lajolla, CA 92037, USA

Abstract. The water-selective pathway through aquaporin 1 (AQP1) membrane channel has been visualized by fitting an atomic model to a 3.7 A resolution three-dimensional density map. This map was determined by analysing images and electron diffraction patterns of lipid-reconstituted two-dimensional crystals of AQP1 preserved in vitrified buffer in the absence of any additive. The aqueous pathway in a monomer is characterized by a size-selective, ~ 4.0 ±0.5 A wide pore that spans a length of ~ 18 A and bends by ~ 25° as it traverses the bilayer. This narrow pore is connected by wide, funnel-shaped openings at the extracellular and cytoplasmic faces, and is outlined mostly by hydrophobic residues interspersed with short stretches of polar amino acids, which results in relatively inert pathway conducive to diffusion-limited water flow. Although not visible at the current resolution, the 3D structure suggests putative binding sites for water molecules in the size-selective pore. 2002 Ion channels—from atomic resolution physiology to functional genomics. Wiley, Chichester (Novartis Foundation Symposium 245) p 33 50

In all living organisms, the entry and exit of water across the lipid biiayer membrane is a fundamental physiological process. Several observations over many decades (see Agre et al 1993, Verkman et al 1996, for review) suggested the presence of water-specific channels or pores in certain membranes in order to account for very high osmotic permeability for water that could not be explained based on ubiquitous passive diffusion. Agre and colleagues (Denker et al 1988) first made the biochemical identification of such a water channel upon isolation and characterization of a very abundant 28 kDa integral membrane protein (CHIP28) from human red blood cell membranes. CHIP28 was later renamed AQP1 (Agre 1997) to signify it as an archetypal member of the rapidly growing aquaporin family of proteins, whose members have been identified both in eukaryotes and prokaryotes. Aquaporins belong to the larger MIP (major intrinsic protein of eye lens) superfamily (Pao et al 1991) of integral membrane proteins. MIP members 33

34

MITRA ET AL

display diversity in the nature of the osmotic-gradient driven diffusion of molecules which include apart from water (in aquaporins), glycerol, purines, pyrimidines and other neutral solutes (in the aquaglycerporins) (Verkman & Mitra 2000, for review). The polypeptide sequences of MIP members are characterized by six major hydrophobic, membrane-spanning domains and homologous N- and C-terminal halves, each containing a highly conserved AsnPro-Ala tripeptide motif. The internal sequence homology probably arose by a tandem, intragenic duplication event (Wistow et al 1991). There is a general consensus that the AQP1 channel is permeated only by water, and excludes, for instance, small chemical species such as H+ and NH3 (Zeidel et al 1992,1994, van Hoek & Verkman 1992). However recent reports have implicated permeability to CO2 (Prasad et al 1998) and cation conductance (Yool et al 1996) that await detailed scrutiny. In vivo, AQP1 organizes as tetramers (Verbavatz et al 1993); but, several lines of evidence (see Verkman et al 1996) indicate that each monomer is a functional channel. The osmo-regulated, bi-directional passage of water through AQP1 is diffusion limited and corresponds to the flow of about 2-5 x 109 water molecules/second per monomer (Zeidel et al 1992, Engel et al 1994). Residues adjacent to the two NPA motifs have been linked to function in AQP1 (as also in other aquaporins) based on site-directed mutagenesis experiments (Jung et al 1994). For instance, Cysl89 in the second, extracellular NPA loop is the site of binding of mercurial reagents that leads to reversible blockage of water transport (Preston et al 1993). We describe here the 3.7 A resolution atomic model of AQP1 water channel in the membrane bilayer determined by electron cryo-crystallography. For this purpose, fully hydrated, vitrified two-dimensional crystals of AQP1, which were generated in synthetic lipid bilayers, were used to acquire crystallographic data. The three-dimensional structure reveals the architecture of the channel and provides insight into elements in the structure and the polypeptide sequence that may confer its exquisite selectivity for water. An atomic-level understanding of the rapid, trans-bilayer water flow through AQP1 not only elucidates a basic mechanism of membrane transport at the structural level but also provides a paradigm for understanding more complex channels such as the voltage-gated and ligand-gated channels. Materials and methods Fully hydrated, 2D crystals of deglycosylated, purified AQP1, while embedded in vitrified, crystallization buffer (20 mM NaH2PO4/Na2HPO4, 100 mM NaCl, 0.1 mM EDTA, and 0.025% NaN3, pH 7.1) (Mitra et al 1995) were examined at -180 °C in Philips CM12 and CM200FEG microscopes. 3D electron crystallographic data (amplitudes and phases) to 3.7 A resolution were extracted

ATOMIC MODEL OF HUMAN AQP1 WATER CHANNEL

35

by processing images and diffraction patterns of many tilted (up to 60°) views of the 2D crystal (Ren et al 2000a, 2001). Uniformly sampled values of structure-factor amplitudes and phases along least-squares fitted curves for the continuous transform normal to the 2D crystal plane were used to synthesize a 3D density map after weighting the amplitudes by the figure of merit (cosine of the estimated error in phase). The statistics of the electron crystallographic data are provided in Table 1. Based on the topology reported in Ren et al (200()a), an initial model was fitted to the experimentally determined 3D density map using the program O (Jones et al 1991). The model was refined against the 3D diffraction amplitudes (Grigorieff et al 1996) using the positional refinement protocol in XPLOR (Brunger et al 1987). The Rfree (Brunger et al 1987) and the free-phase residual (j free ) (Grigorieff et al 1996) were used to monitor the course of the refinement. In order to improve the map, a SIGMA A (Read 1986)-weighted 2FO-FC map was calculated that was expected to be devoid of the model bias in the side chain positions of the original best-fitted model (Ren et al 2001) and was used for optimization of side chain orientations. Results and discussion Based on the processed image areas, the phases used for calculating the density map were determined from an averaged view of ~ 7 x 105 tetramers (~ 3 million AQP1 monomers). The observed, overall, low phase residual of 25.7° and the good statistics of the diffraction data (Table 1) led to a density map whose quality was sufficiently reliable to allow the delineation of an atomic model (Fig. 1). Hitherto, all investigations to derive atomic or near-atomic resolution structures using electron crystallography have exclusively employed specimens preserved in sugars, such as tannic acid, trehalose or glucose, including the recent study of AQP1 structure reported by Murata et al (2000). Therefore, our 3.7 A resolution electron-crystallographic analysis represents the first investigation in which fully hyclrated specimens embedded in vitreous ice were used to arrive at an atomicresolution model allowing a delineation of the unperturbed, close to in vivo structure. The atomic model presented includes residues L9 to A232. No continuous densities for the rest of the residues were visible in the map. The 2Fo-Fc map calculated at the final stage (see Methods section) helped to improve locations of some of the side chains (Fig. 1A,B). Elaborate refinements of smoothed B-factor along the helices and along individual residues (Grigorieff et al 1996) or lipid positions seen in projection (Ren et al 2000b) were not attempted. The tandemly repeating N- and C-terminal halves of AQP1 are each comprised of three tilted transmembrane a helices and a short a helix adjacent to the NPA

MITRA ET A I.

36

TABLE 1 Electron crystallographic data Two-side plane group

P4212

Unit cell dimension

a = b = 99.58 ± 0.50 A, c = 100 A

Number of observed amplitudes

48037 19839

Number of observed phases Number of structure factors and overall completeness (100-3.7 A) In-plane resolution and estimated resolution normal to the bilaverb

3

2947, 63%

3.7 A, 6 A

Electron diffraction Resolution limit Maximum tilt Number of patterns

3.3 A 57.5 44 6.7-28.9%(15.8%>) 15.4-47.3% (27.5%)

Image

Resolution limit Maximum tilt Number of images Phase residuale in resolution Zones Overall

3.7 A 60.2C 72 (100-7.0 A) 20.0c (7-4.0 A) 39.7c (4.0 3.7 A) 557c (100-3.7A) 25.7c

Refinement statistics Rfactorf

35.0%,

Rfree f,g

45.1%

jfree

62.4C

a

With figure of merit >0.27. Maximum completeness for data up to 60" tilt is 83%. Based on calculation of point-spread function (Unger & Schertler 1995) for the experimental map.

b

e

Weighting based on amplitudes. Using electron diffraction amplitudes (> 2s) between 24—3.7 A. Calculation based on 5% of significant reflections not used in refinement. h Averaged difference between phases calculated from the model and the observed phases with figure of merit > 0.50. f

g

ATOMIC MODEL OF HUMAN AQP1 WATER CHANNEL

FIG. 1. A representative section of the 3D density map defining the density for the transmembrane helix TM6 in the (A) experimental and the corresponding (B) calculated, 2Fo-Fc map. (C) Stereo view of another section of the 2Fo-Fc map showing the region around the cytoplasmic edges of the transmembrane helix TM3 and the short a helix H1 in the first NPA loop. Some of the amino acid residues are marked. The maps in this and in the other figures were rendered at 1.0s of the mean density and were generated using the program AVS (Upson et al 1989).

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MITRA ET AL

motif (TM1, TM2, TM3, H1 and TM4, TM5, TM6, H3, Fig. 2A). This is similar to the fold described in the two other reported, atomic-resolution structural studies of aquaporins (Murata et al 2000, Fu et al 2000). The long 3-4 inter-helix linker (Fig. 3A) containing a short a helix (H2) at the extracellular edge of TM3 connects the two halves. The TM3-TM4 linker dips in from the extracellular side towards the second NPA loop and at this location, highly conserved R195 (in all MIP sequence NPAR, except NPAV in g-TIP) is stabilized by a hydrogen bond with conserved N127 and a salt-bridge with D128 of the 3-4 loop (Fig. 3B). The observation of clear and continuous density for this trans-bilayer loop, possibly due to enhanced contrast (water versus protein in our case as against sugar versus protein in traditional high-resolution electron crystallography) proved to be very important in correctly defining the polypeptide topology based on our earlier, lower 4 A resolution, density map (Ren et al 2000a). The AQP1 monomers organize as tetramers as in vivo, consistent with the p4212 two-sided plane group of symmetry of the 2D crystal (Fig. 4A). The transmembrane helices in a monomer pack with an atypical right-handed twist (Walz et al 1997, Cheng et al 1997), whereas those at the monomer interface (TM4, TM5 of one monomer and TM1, TM2 of an adjacent monomer) in the tetramer are arranged with the more commonly observed left-handed packing. The spatial dispositions of the two halves of the monomer are according to an approximate, in-plane twofold axis of symmetry (Cheng et al 1997, Ren et al 2000a). The symmetry places the two short, transmembrane a helices TM2 (N-terminal half) and TM5 (C-terminal half) proximal near the fourfold axis and the two NPA loops, which are approximately, mutually orthogonal, vertically apposed and on the two sides of the symmetry axis. TM1, TM2, TM4 and TM5 define the interior of the monomer while TM6 and TM3, define the lipid-exposed exterior face (Fig. 2A). Adjacent helices in a monomer display elaborate hydrophobic packing interactions involving in some cases residues that are conserved in the aquaporin family. These include F18 (highly conserved) of TM1 with V107 of TM3; V50of TM2 with LI81 (conserved) of TM5; L139,1143 and L147 of TM4 with 1211, G215 (conserved), and G219, and A223 of TM6, respectively. The aqueous pathway through an AQP1 monomer is outlined by residues from TM1, TM2, TM4, TM5 and the short a helices H1 and H3. Figure 3B identifies schematically, residues that line or point into the channel. In Fig. 4B, the architecture of the monomeric channel is illustrated thorough a surface-rendered view of a transverse section through an AQP1 tetramer (Fig. 4A). The aqueous pathway is wide on both the extracellular and the cytoplasmic edges and tapers down like a funnel from both sides to form a narrow curved pore with an approximately uniform diameter of ~4.0db0.5 A and spans a distance of ~ 18 A through the interior of a monomer. The cytoplasmic and extracellular constrictions defining the two edges of this pore are ~ 9 A away from the

ATOMIC MODEL OF HUMAN AQP1 WATER C H A N N E L

FIG. 2. (A) Ribbon diagram for the polypeptide fold of an AQP1 monomer viewed from the extracellular face. The six transmembrane helices TM1 to TM6 and the short cytoplasmic (H1) and extracellular (H3) a helices in the two NPA loops and that (H2) at the extracellular edge of TM3 are indicated. (B) Demarcation of the polypeptide sequence of AQP1 into the observed a helices. Amino acids, whose side chains either line or point into the monomeric channel, are shaded. The mercurial-sensitive C189 and the analogous A73 are indicated.

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MITRA FT AL

FIG. 3. (A) The extracellular density in the experimental map corresponding to the segment connecting the two halves of the monomer—the linker between helices TM3 and TM4. (B) Stereo view of the pocket around the highly conserved R195 in helix H3 indicating stabilizing interactions with residues N127 (conserved) and D128 in the TM3-TM4 linker.

locations of the NPA motifs, an observation that disagrees with results reported by Murata et al (2000). This narrow pore, which acts as the size-selective filter to exclude small solutes, bends by ~25° as it traverses the bilayer (Fig. 4B). The apex of the curvilinear pathway is close to the middle of the bilayer where the inplane pseudo twofold axis of symmetry is also located. Thus, the ~ 18 A long, narrow part of the channel includes a region of the molecule where the approximate twofold symmetry is the strongest (Cheng et al 1997, Ren et al 2000a), leading to a symmetrical environment that is consistent with observed bidirectional water flow (Meinild et al 1998). Such a curved pathway can also promote an increase in strain and/or disruption of hydrogen-bonded network of permeating water molecules. This in effect enhances the possibility that

ATOMIC MODEL OF HUMAN AQP1 WATER CHANNEL

41

FIG. 4. (A) A ribbon diagram for the quaternary organization of the AQP1 monomer viewed from the extracellular side. (B) Surface-rendered illustration of the AQP1 tetramer sliced through the middle (through the fourfold axis) revealing the curved water-selective pathway in the two adjacent monomers and the region around the fourfold axis as viewed parallel to the bilayer. Top is the extracellular side. The ~25° bend of the size-selective pore as it traverses the bilayer is indicated by arrows. The MSMS (Sanner et al 1996) program was used for surface rendering of the atomic model.

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neighbouring residues in the surrounding protein, e.g. N76 and N192, could compete for hydrogen bonds (Murata et al 2000) with adjacent water molecules in the pore, thereby aborting H+ (Stilinger 1998) or OH~ conduction. Whereas mostly polar or charged residues line the entrances to the AQPl channel, the majority of the residues (~75%) outlining the ~ 18 A narrow part of the channel are hydrophobic (Fig. 2B) interspersed with polar residues, namely, N76, N192, HislSO, His74 and Ser28. Therefore, the surface presented to permeant water molecules in the interior, size-selective part of the channel is largely hydrophobic but with small polar regions to overcome the energy barrier. Such an arrangement results in a relatively non-interacting pathway for rapid diffusion in the absence of attractive energy minima that accompany an overall, polar environment. The dispositions of the two short a-helices (H1 and H3 in the two NPA loops, Fig. 2A) in the narrow pore are reminiscent of the pore helices in the structure of the K+ channel from Streptomyceslividans (Doyle et al 1998). In that channel, these helices are oriented so that the helix dipole moments act to attract K* near the centre of the bilayer. However, in AQPl, the orientation of each of the short helices is opposite to that in the K+ channel and will repel a cation near the bilayer centre. Also, the absence of an extensive polar lining in the AQPl channel means that it will be energetically expensive to locate a desolvated ion at the channel entrance while a hydrated ion including H3O+ and anions, which are almost always hydrated, are too big to pass through. In the cell milieu, water is the most concentrated species — the physiological concentration of water is ~ 55 M, many times higher than any other component, including common ions which occur in milli- to micromolar range. This fact alone will lead to a preference for osmotically driven flow of water through a size-selective pore that is designed for the passage of uncharged small solutes restricted by its diameter (~4.0±0.5A). Apart from the pores in each monomer, the region around the fourfold axis (i.e. at the centre of the tetramer) also represents a putative trans-bilayer pathway (Fig. 4B). In fact, on the basis of a 2.2 A resolution X-ray crystallographic structure determination of an aquaglyceroporin — the bacterial glycerol transporter GlpF — Fu et al (2000) have suggested that the pore at the centre of the tetramer transporter could act as a cation channel. They observed density peaks along the fourfold axis, which were assigned to two Mg2+ ions, one of which (at the extracellular face) was chelated by four symmetry-related glutamate (Glu43) residues. This region, bounded by symmetry-related mates of TM2 and TM5, has an irregular shape in the AQPl structure and, because of different inclinations of these two helices, has the largest diameter near the centre of the bilayer (Fig. 4B). The mouth of this pore is wider on the cytoplasmic side than on the extracellular side. At the extracellular side four symmetry-related Gln47 residues of TM2, which are in close apposition, constrict the diameter to

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< ~3 A. The interior of this pore has a very hydrophobic environment, but includes LysSl of TM2 (analogous to Glu43 of GlpF), presumably protonated (raised pKa) or chelating an anion not resolved in the 3.7 A density map. Interestingly, Lys5l has been implicated in proper folding of AQP1 during maturation in the Golgi (Foster et al 2000). Currently, the exact functional role of the pathway around the fourfold axis in AQP1 awaits further, detailed analysis. Although, at 3.7 A resolution we cannot visualize permeant water molecules in the AQP1 channel, we can speculate on possible binding sites. In analogy with the situation in the 2.2 A resolution structure of GlpF (Fu et al 2000) one putative site is shown in Fig. 5A. At the extracellular face, Arg206, Trp48 and Phe200 form a glycerol-binding pocket in GlpF (Fig. 5B) and were suggested to comprise the selectivity filter (Fu et al 2000). The analogous pocket in AQP1 located at the extracellular constriction of the channel (Fig. 4B) contains similar highly conserved Argl95, and the aromatic Phe56, but now includes Hisl80, and Phe2()0 is replaced by mercurial sensitive Cysl89. We note that Hisl80 is conserved in mammalian homologues AQP0 and primarily water-transporting members AQP2, AQP4, AQP5 and AQP6 but is a variable non-polar residue in aquaglyceroporins (e.g. Gly in GlpF, AQP3 and AQP7, Val in Nod26). We suggest that Hisl 80 is an important residue which contributes to the selection of water over glycerol at this putative binding site in AQP1 that is slightly larger and relatively more polar than the aforementioned glycerol binding pocket in GlpF. Near the centre of the bilayer, a cluster of hydrophobic residues surrounding the absolutely conserved, polar Asn76 and Asnl92 characterize the middle of the sizeselective pore (Fig. 4B). Interestingly, a similar hydrophobic cavity in the retroGCN4 leucine-zipper structure (Mittl et al 2000) has been observed. Here, a wall of Leu and Val residues define the hydrophobic interior and two symmetry-related Asn residues are the only polar side chains that point into the core sequestering a pair of water molecules. It is possible that Asn76 and Asnl92 in AQP1 also participate in forming a similar binding site for water molecules (Murata et al 2000). Conclusions An atomic model of the human AQP1 water channel in the membrane bilayer was determined at 3.7 A resolution by electron cryo-crystallography of 2D crystals preserved in vitreous ice. The fully hydrated, unperturbed, close to in vivo structure reveals the architecture of the water-selective pathway through an AQP1 monomer. The aqueous pathway is characterized by a ~4.0+0.5 A wide, size-selective pore that spans a length of ~ 18 A and bends by ~ 25° as it traverses the bilayer. This narrow pore, outlined mostly by hydrophobic residues with short stretches of polar amino acids widens into funnel-shaped openings at both the

44

MITRA FT AL

FIG. 5. (A) Putative binding site for water molecules in AQP1 at the extracellular edge of the size-selective pore in the monomer. His 180 conserved in primarily, water-transporting mammalian aquaporins is suggested to have an important role in preferential binding of water. (B) The corresponding glycerol-binding site observed by Fu et al (2000) in the X-ray structure of bacterial glycerol transporter GlpF.

extracellular and cytoplasmic faces. Such an arrangement results in a relatively inert pathway conducive to diffusion-limited water flow. The atomic model suggests putative, water-selective binding sites at the extracellular constriction of the aqueous pathway and near the middle of the bilayer. The atomic-resolution structure of AQP1 provides a rational framework for not only exploring sitedirected, mutagenesis-driven experiments to fully understand selectivity but also for embarking on identification of pharmacologically useful modulators of

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function though structure-based drug design and high-throughput screening of combinatorial drug libraries.

Acknowledgements We are indebted to M. Pique for generating the figures depicting pertinent features of the 3D density map. The research was supported by a grant from the National Institutes of Health, in part by a grant from the National Science Foundation and a special fellowship from the Cystic Fibrosis Research Foundation to AKM. AC and GR were supported by a post-doctoral fellowship from the American Heart Association Western Affiliate. VSR was supported by the MMTSB research resource RR12255 of NIH to TSRI. AKM is an Established Investigator with the American Heart Association.

References Agre P 1997 Molecular physiology of water transport: aquaporin nomenclature workshop. Mammalian aquaporins. Biol Cell 89:255—257 Agre P, Preston GM, Smith BL et al 1993 Aquaporin CHIP: the archetypal molecular water channel Am J Physiol 265:F463-F476 Brunger AT, Kuriyan J, Karplus M 1987 Crystallographic R factor refinement by molecular dynamics. Science 235:458-460 Cheng A, van Hoek AN, Yeager M, Verkman AS, Mitra AK 1997 Three-dimensional organization of a human water channel. Nature 387:627-630 Denker BM, Smith BL, Kiihajda FP, Agre P 1988 Identification, purification, and partial characterization of a novel Mr 28,000 integral membrane protein from erythrocytes and renal tubules. J Biol Chem 263:15634-15642 Doyle DA, Morais Cabral J, Pfuentzer RA et al 1998 The structure of the potassium channel: molecular basis of K+ conduction and selectivity. Science 280:69-77 Engel A, Walz T, Agre P 1994 The aquaporin family of membrane water channels. Curr Opin Struct Biol 4:545-553 Foster W, Helm A, Turnbull I et al 2000 Identification of sequence determinants that direct different intracellular folding pathways for aquaporin-1 and aquaporin-4. J Biol Chem 275: 34157-34165 Fu D, Libson A, Miercke LJW et al 2000 Structure of a glycerol-conducting channel and the basis for its selectivity. Science 290:481-486 Grigorieff N, Ceska TA, Downing KH, Baldwin JM, Henderson R 1996 Electroncry stallographic refinement of the structure of bacteriorhodopsin. J Mol Biol 259:393-421 Jones TA, Zou J-Y, Cowans S W, Kjeldgaard M 1991 Improved methods for building protein models in electron density maps and the location of errors in these models. Acta Crystallogr A 47:110 119 Jung JS, Preston GM, Smith BL, Guggino WB, Agre P 1994 Molecular structure of the water channel through aquaporin CHIP. The hourglass model. J Biol Chem 269:14648-14654 Meinild DA, Klaerke DA, Zeuthen T 1998 Bidirectional water fluxes and specificity for small hydrophilic molecules in aquaporins 0-5. J Biol Chem 273:32446-32451 Mitra AK, van Hoek AN, Wiener MC, Verkman AS, Yeager M 1995 The CHIP28 water channel visualized in ice by electron crystallography. Nat Struct Biol 2:726-729 Mittl PRE, Deillon C, Sargent D et al 2000 The retro-GCN4 leucine zipper sequence forms a stable three-dimensional structure. Proc Natl Acad Sci USA 97:2562-2566

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DISCUSSION

Murata K, Mitsuoka K, Hirai T et al 2000 Structural determinants of water permeation through aquaporin-1. Nature 407:599-605 Pao GM, Wu LF, Johnson KD et al 1991 Evolution of the MIP family of integral membrane transport proteins. Mol Microbiol 5:33-37 Prasad GV, Coury LA, Finn F, Zeidel ML 1998 Reconstituted aquaporin 1 water channels transport co2 across membranes. J Biol Chem 273:33123-33126 Read RJ 1986 Improved fourier coefficients for maps using phases from partial structures with errors. Acta Crystallogr A 42:140-149 Ren G, Cheng A, Reddy V, Melnyk P, Mitra, AK 2000a Three-dimensional fold of the human AQP1 water channel determined at 4 A resolution by electron crystallography of twodimensional crystals embedded in ice. J Mol Biol 301:369 387 Ren G, Cheng A, Melnyk P, Mitra AK 2000b Polymorphism in the packing of aquaporin-1 tetramers in 2D crystals. J Struct Biol 130:45-53 Ren G, Reddy VS, Cheng A, Melnyk P, Mitra AK 2001 Visualization of a water-selective pore by electron crystallography in vitreous ice. Proc Nad Acad Sci USA 98:1398-1403 Sanner M, Olson AJ, Spehner JC 1996 Reduced surface: an efficient way to compute molecular surfaces. Biopolymers 38:305-320 Stillinger FH 1980 Water revisited. Science 209:451-457 Unger VM, Schertler GF 1995 Low resolution structure of bovine rhodopsin determined by electron cryo-microscopy. Biophys J 68:1776-1786 Upson C, Faulheber TJr, Kamins D et al 1989 The application visualization system: A computational environment for scientific visualization. IFIEE Comput Graphic Appl 9:30-42 van Hoek AN, Verkman AS 1992 Functional reconstitution of the isolated erythrocyte water channel CHIP28. J Biol Chem 267:18267-18269 Verbavatz JM, Brown D, Sabolic I et al 1993 Tetrameric assembly of CHIP28 water channels in liposomes and cell membranes: a freeze-fracture study. J Cell Biol 123:605 618 Verkman AS, Mitra AK 2000 Structure and function of aquaporin water channels. Am J Physiol 278:F13-F28 Verkman AS, van Hoek AN, MaTetal 1996 Water transport across mammalian cell membranes. Am J Physiol 270-.C12-C30 Walz T, Hirai T, Murata K et al 1997 The three-dimensional structure of aquaporin-1. Nature 387:624-627 Wistow GJ, Pisano MM, Chepelinsky AB 1991 Tandem sequence repeats in transmembrane channel proteins. Trends Biochem Sci 16:170-171 Yool AJ, Stamer WD, Regan JW 1996 Forskolin stimulation of water and cation permeability in aquaporin 1 water channels. Science 273:1216 1218 Zeidel ML, Ambudkar SV, Smith BL, Agre P 1992 Reconstitution of functional water channels in liposomes containing purified red cell CHIP28 protein. Biochemistry 31:7436- 7440 Zeidel ML, Nielsen S, Smith BL, Ambudkar SV, Maunsbach AB, Agre P 1994 Ultrastructure, pharmacologic inhibition, and transport selectivity of aquaporin channel-forming integral protein in proteoliposomes. Biochemistry 33:1606—1615

DISCUSSION Tieleman: You gave a very accurate picture of the side-chains, but your resolution is only 3.7 A. How can you justify the detail in your pictures? Mitra: I wouldn't say that the side-chain positions are equally clear throughout the map. In the regions that I showed, the density is pretty clear and defines the

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side-chain orientations well. At 3.7 A we can't be absolutely sure about the orientations, but we can make optimal models. Doyle: Structurally this channel is a tetramer, but functionally it is a monomer. Is there a reason why this is actually tetrameric? Mitra: It is not uncommon to have monomeric functional units in an oligomeric membrane protein. Bacteriorhodopsin is an example; it organizes as a trimer but individual monomers photocycle. GlpF has been shown to be functional even in a monomeric state. It could be that because the lengths of TM2 and TM5 in AQP1 are particularly short and insufficient to span the bilayer it is necessary to form an oligomer in order to sequester these helices away from the lipid. Also the tetramer formation may have an indirect structural role in the correct architecture of the monomeric pores. Perozo: I have always been fascinated by the size of the cavity in the fourfold axis symmetry, and this is blocked by this residue 47. An obvious experiment would be to mutate it and see whether the cavity then conducts either water or ions. Mitra: The role of this region is a matter of considerable interest in the MIP field. Unpublished experiments (A. van Hoek, personal communication) on AQP1 suggested that after inhibition by mercurials there is an onset of leaky pathway which, for instance, could be the cavity along the fourfold axis. Recently, Agre and colleagues (Saparov et al 2001) showed that in planar lipid bilayers containing AQP1, cGMP-induced ion conductance is elicited that is not blocked by pCMBS, presumably implicating an alternate pathway. Actually, ion conductance properties were first reported in 1996 by Yool and colleagues from the University of Arizona, work that was unfortunately received with a lot of suspicion. The conclusion is that the region around the fourfold axis may serve as another solute or ion pathway; however the exact role awaits careful investigation. Umvin: I have Dr Yoshi Fujiyoshi's structure in the back of my mind. I got the impression that their pore that the water goes through is shorter. You seem to have a long, narrow tube. Could there be a difference because of the glucose in their case and the water in yours? Mitra: I am not quite sure about the full, transverse nature of the pore in their structure. Unwin: I felt happy with the shorter pore because the protein is designed to be as efficient a water transporter as possible. You have this long tube which the water goes down. Mitra: In their model, the critical region is the narrow part near the bilayer centre. Unwin: The tube is quite wide further up, and presumably there is no problem with water transport there because the surfaces have polar groups on them.

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DISCUSSION

Mitra: At least in the mouth of the pore, definitely. Auerbach: I am trying to relate this to Nigel Unwin's idea of a hydrophobic ring being a permeation barrier in the ACh receptor. You have a more extensive structure in the hydrophobic region. What are the dimensions of this region? Mitra: As I mentioned, there is uncertainty in the orientation of some of the side chains and therefore we present the pore diameter to be ~ 4 ±0.5 A. Mindell: Is there a continuous line of polar residues all the way down your pore which would be in contrast to Nigel Unwin's structure? Mitra: No, a majority of the side chains lining the narrow pore are hydrophobic, interspersed with some polar residues. For instance, Asn76 and Asnl92 lie in the middle of the bilayer surrounded by hydrophobic residues and analogous to a very similar arrangement seen in the retro-GCN4 structure (Mittl et al 2000). It is a fourhelix bundle with a very hydrophobic core but has two symmetry-related Asn residues binding two water molecules. Mindell: Outside that critical point, are there places where there are only hydrophobic residues all the way around the circumference of the pore? Mitra: Yes. Mindell: Roughly what percentage of the distance that a water molecule would have to traverse do they cover? Mitra: The total length on either side is about 9 A, so I would say 5—6 A. Miller: The other way to look at this is that you say that ions do not go through this pore, so it has to be hydrophobic in certain places, simply to act as a gate. Was this 4 A a radius or diameter? Mitra: Diameter. I would like to add that the way we were looking at the reasons behind selectivity was wrong. We were only thinking about what it could be that was making the channel selective for water. Instead, it appears that we should have been asking what it is that is making it deselective for other solutes. For instance, the two short NPA loop helices are oriented with their N-terminal pointing towards the middle, so that the centre of the bilayer is repulsive to cations. Anions are almost always hydrated, so they are too big to go through. In the physiological milieu, the highest concentration of anything present is water. It is almost a reverse way of looking at it. Jordan: How long is the narrow part of the pore? Mitra: It is narrow for about 18 A. Schrempf: How do you envisage the transport of ions in those members of the family that can transport water, glycerol and ammonium ions? Mitra: We carried out modest homology modelling for Nodulin 26, which is the subset of channels you mention. Nod26 is not quite a glycerol facilitator and not quite a water channel, but somewhere in the middle. In the model, there are some interesting differences, although overall, the nature of the residues lining

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the selectivity pore is very similar. However there are hints as to residues that might be responsible for differences in transport properties. Schrempf: Many organisms have a set of several channel genes in parallel. Could you exclude the possibility that, as each monomer seems to act individually, there might be a heteromeric composition of several types of these subfamily members? Mitra: I suppose you are referring to the situation as in the case of gap junctions. It is an interesting idea, but I haven't thought about it. Choe: What do you think is filling up the cavity at the fourfold axis? In nature there cannot be a vacuum in the middle of a membrane. Mitra: Presumably, but we don't know: there could be bound detergent, or lipids, or even vacuum. Choe: It could be a space-buffering zone. Mitra: I can't say what the situation is in vivo, Mindell: Peter's ion channel aquaporins seem to be anion channels. I am intrigued that you see this lysine right in the central pore. Is that lysine conserved among the whole family? Mitra: No, it is not. For instance, it is a glutamate in GlpF, which in the X-ray structure binds to a putative Mg2+ ion. It has been shown that Lys5l, at least in the context of AQP1, might be important for proper folding. Foster et al (2000) showed that in the endoplasmic reticulurn (ER), AQP1 does not have the traditional four-transmembrane topology. When they swapped a small analogous segment from AQP4, where this residue is leucine, AQP4 displayed similar abnormal topology in ER. Mindell: Does AQP6 also have the lysine? Mitra: No, this residue is glutamine: interestingly AQP6 was shown to be an anion channel from experiments in oocytes. Fu: In your electron density map, did you see channel-like cavity in the centre of the tetramer, which would suggest AQP1 may have a tetrameric channel? Mitra: At the current resolution we did not see any resolved density for candidate ions. Ashcroft: We heard earlier that the acetylcholine channel is closed by a hydrophobic gate. In the discussion there was an argument about whether water could go through this gate or not, and the feeling was that it didn't. Here we have another hydrophobic section: why is the water going through in this case? Sansom: It is not as hydrophobic. I don't want to disagree with Alok, but it would seem that the water channel and the GlpF channel are rather more amphipathic, in that if you cut sections through the channel, in most regions there is a polar side chain somewhere along it. Unwin: The acetylcholine has an extensive region of hydrophobicity that is about 8 or 9 A in length.

50

DISCUSSION

Mitra: Ours is not that long. We expected that as water goes through, there should be polar residues. From this point of view we were quite surprised to find the hydrophobic regions. Sansom: One hesitates to extrapolate from simulations to the real world, but maybe if you have something that is highly polar it is more likely to bind water. As you said, in this pore the water is barely pausing on its way through. If you only do a calculation of the geometry of the pore with water going in at diffusion-limited rates, it can barely match the rate of water going through AQP. Mathie: Is the channel gated at all? Mitra: Anthony et al (2000) have indicated in oocyte experiments that cloned AQP1 displays gating by cGMP, although the result has not been widely accepted. There are other members that have been shown to be regulated by phosphorylation or by pH. Mathie: In that case, would you imagine that the hydrophobicity of the pore region should be changed when it is gated? Mitra: I doubt that. For example, in Nod26, the site of phosphorylation is on the very flexible C-terminus. It is intriguing how it affects the channel. Sattelle: In the aquaporins, are there any naturally occurring mutations in the channel-lining residues? In the nicotinic receptors such mutations have been very instructive; they include mutations responsible for congenital myasthenias and one form of epilepsy. Mitra: AQP2 is one aquaporin that is directly implicated in the disesase nephrogenic diabetes insipidus. Different single-site mutations have been considered to be responsible for this disease. One such residue is Argl87 analogous to Argl95 which is at the extracellular mouth of the selectivity pore in AQP1. References Anthony TL, Brooks HL, Boassa D et al 2000 Cloned human aquaporin-1 is a cyclic GMP-gated ion channel. Mol Pharmacol 57:576-588 Foster W, Helm A, Turnbull et al 2000 Identification of sequence determinants that direct different intracellular folding pathways for aquaporin-1 and aquaporin-4. J Biol Chem 275:34157-34165 Mittl PR, Deillon C, Sargent D et al 2000 The retro-GCN4 leucine zipper sequence forms a stable three-dimensional structure. Proc Natl Acad Sci USA 97:2562-2566 Saparov SM, Kozono D, Rothe U, Agre P, Pohl P 2001 Water and ion permeation of aquaporin1 in planar lipid bilayers. Major differences in structural determinants and stoichiometry. I Biol Chem 276:31515-31520

The structure of GlpF, a glycerol conducting channel Dax Fu, Andrew Libson* and Robert Stroud* Biology Department, Bmlding463, 50 Bell Avenue, Brookhaven National Laboratory, Upton, NY 11973 and *Macromolecular Structure Group, Department of Biochemistry and Biophysics, Room S-964, Box 0448, University of California San Francisco, San Francisco, CA 94143-0448, USA

Abstract. The passage of water or small neutral solutes across the cell membrane in animals, plants and bacteria is facilitated by a family of homologous membrane channels, variously known as aquaporins though perhaps more correctly as aquaglyceroporins. The glycerol facilitator (GlpF) is a 28 kDa aquaglyceroporin that catalyses transmembrane diffusion of glycerol and certain linear polyhydric alcohols in Escherichia coli. X-ray crystallographic analysis of GlpF to 2.2 A resolution revealed an a-barrel structure, surrounded by six full-length transmembrane helices and two halfspanning helices that are joined head-to-head in the middle of the membrane. These helices are arranged to a quasi twofold manner relative to the central membrane plane, where highly conserved residues make helix-to-helix contacts that stabilize the relative position and orientation of the helices in the structure. This sequence-structure correlation suggests that the evolutionary divergence of aquaporins and aquaglyceroporins is constrained by a conserved structural framework within which specialized function may be developed. Three glycerol molecules were resolved in the central channel through the GlpF monomer, thereby defining a transmembrane channel for glycerol permeation. The structure of glycerol—GlpF complex provides insight into the chemical basis for transmembrane selective permeability. 2002 Ion channels — from atomic resolution physiology to functional genomics. Wiley, Cbtchester ( Novartis Foundation Symposium 245) p 51-65

Fluid balance is a ubiquitous biological process in all living organisms. The molecular basis of the rapid transmembrane flux of water and small nonelectrolytes remained elusive until the recent discovery of a superfamily of fluidtransporting membrane channels (Preston & Agre 1991, Preston etal 1992). On the basis of their permeability properties, these channels are subdivided into aquaporins — a subfamily of strict water-selective channels, and aquaglyceroporins — a homologous subfamily of less selective channels with permeability for small neutral solutes such as glycerol (Borgnia et al 1999a). 51

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The fundamental importance of aquaporins and aquaglyceroporins is attested by their strong genetic conservation from bacteria to plants and humans (Park & Saier 1996, Heymann & Engel 2000). More than 180 aquaporins and aquaglyceroporins have been identified in organisms ranging from bacteria to mammals, including 10 human isoforms (AQPO-9). The Escherichia coli aquaporin AQPZ and aquaglyceroporin GlpF share 20-25% pair-wise sequence identity with each other, and with all human isoforms identified thus far (Unger 2000, Beitz & Schultz 1999). This extensive sequence similarity indicates that AQPZ and GlpF are part of the development that leads to the human evolutionary tree. Functional studies of human aquaporins and aquaglyceroporins have been facilitated by the use of homologous E. coli isoforms. AQPZ and GlpF have been overexpressed, extracted from the membrane using non-ionic detergents and then purified to homogeneity in detergent micelles (Borgnia & Agre 2001). At the protein level, reconstitution of pure AQPZ or GlpF into liposomes makes it possible to examine transmembrane fluxes of water or glycerol by monitoring swelling or shrinkage of membrane vesicles in response to a transmembrane osmotic gradient (Agre et al 1999, Verkman 2000a). The rates of solute fluxes through these channels are diffusion limited, suggesting that AQPZ and GlpF function as channels rather than transporters. The channel selectivity was also examined, showing that AQPZ specifically increases water osmotic permeability (Calamita et al 1995) whereas GlpF increases glycerol (Maurel et al 1994) and water (Borgnia & Agre 2001) permeability. In addition, GlpF is permeable to certain polyols and antimonite (Heller et al 1980, Sanders et al 1997). At the organ level, the physiological significance of human aquaporins was implicated by phenotype analysis of aquaporin knockout mice and by pathophysiological studies of human aquaporin deficient diseases such as nephrogenic diabetes insipidus (Verkman et al 2000, Verkman 1999, Agre 1998). It is evident that human aquaporins can facilitate near iso-osmolar transepithelial fluid absorption and secretion, as well as rapid water movement driven by osmotic gradients (Verkman 2000b). Recent cloning of AQP8 from adipose tissue suggests that aquaglyceroporin is involved in glycerol release and body energy balance that may be relevant to obesity (Kishida et al 2000). Structural analysis has led to the determination of the general channel architectures of AQP1 (Cheng et al 1997, Walz et al 1997, Murata et al 2000) and GlpF (Braun et al 2000, Fu et al 2000). The structure of human red cell AQP1 was determined to 3.8x6.5 A resolution by electron crystallography (Murata et al 2000). More recently, we determined the structure of GlpF by X-ray crystallography to 2.2 A resolution (Fu et al 2000). Arrangements of amino acid side chains and chemical groups along the GlpF transmembrane channel are revealed at this higher resolution, together with three bound glycerol molecules. AQP1 and GlpF are strikingly similar structures. Both crystallize as tetramers, with

STR UCTURE OF GlpF

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FIG. 1. Ribbon representation of GlpF monomer (from a perspective viewed in parallel to the membrane plane) with two glycerol molecules bound in the middle of the channel.

each monomeric subunit as an asymmetric unit. The monomers are folded as an a-barrel structure, with a central channel surrounded by six full-length transmembrane helices and two half-spanning helices that are joined head-tohead in the middle of the membrane to give the appearance of a kinked transmembrane span (Fig. 1). In both AQP1 and GlpF, an internal quasi-twofold symmetry is evident. Superimposition of main chains of two halves of the GlpF

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FIG. 2. xylitol.

FU ET A L

Stereo arrangements of OH groups with respect to the alkyl backbones of ribitol and

showed a close match with a root-mean-square deviation of 1.4 A. The arrangement of the two halves in the structure creates an overall symmetrical relationship with respect to a quasi-twofold axis in the central membrane plane. This quasi-twofold structure corresponds to the two tandem repeats in the linear sequence that may have arisen from an intragenic gene duplication event (Wistow etal1991).

Channel permeability and selectivity Fluxes of water and polyols through AQPZ or GlpF channels have been monitored in real time using stopped-flow light scattering measurements (Borgnia et al 1999b, Borgnia & Agre 2001, Fu et al 2000). Permeability through the AQPZ channel was only observed for water, but not for any other neutral solutes. By comparison, GlpF was found to be permeable to a range of linear polyols of different chain lengths and to water at a slower rate compared with the rate through AQPZ (Borgnia & Agre 2001). In light of this flexibility for different sizes of polyols, it came as a surprise to find that the rate of ribitol flux was sevenfold faster than that of its stereoisomer xylitol (Heller et al 1980). The only difference between ribitol and xylitol is the stereo configuration of one OH group with respect to the alkyl backbone (Fig. 2). This result showed that GlpF is selective for one stereoisomer over another. The GlpF structure explains how permeability to multiple solutes can be reconciled with stringent stereo-selectivity (Fu et al 2000). The channel lining is strongly amphipathic, with oxygens and nitrogens lined up on one side and carbons on the opposite side of the lumen surface. This amphipathic channel uniquely matches the chemical structure of glycerol and other polyols, which are composites of the polar hydroxyl group arranged on a non-polar alkyl backbone. Of particular interest is the exquisite hydrogen bonding network seen at the narrowest constriction of the channel, where two consecutive OH groups of a polyol molecule are oriented with ideal geometry with respect to the donors and acceptors from within the channel, thus positioning the donor-acceptor pairs at

STRUCTURE OF GlpF

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FIG. 3. Schematic diagram showing the process of stereo selective permeation. A polyol molecule passes through the narrowest constriction of the GlpF channel where each of two consecutive OH groups satisfies the double hydrogen bonding geometry imposed by the hydrogen donor and acceptor groups located on the hydrophilic side of the GlpF channel.

about 120° in relation to each of two successive OH groups of the polyol (Fig. 3). As a polyol proceeds through the channel, each pair of successive OH groups has to satisfy this double H-bonding geometry. Thus, the narrowest constriction imposes a stereo selection upon the relative orientation of OHs, regardless of the chain length. This structure satisfactorily explains why ribitol is more permeable than xylitol, because all OHs in ribitol are on one side of the alkyl backbone (Fig. 3), matching with the arriphipathic channel, whereas in xylitol one OH is on the opposite side of the backbone. Common to AQPZ and GlpF is the absolute exclusion of ions (Fu et al 2000, Murata et al 2000). The first barrier against ion entry is made by electrostatic repulsion between NHs of the guanidimurn group of R206 and the carbonyl oxygens of G199 and F200 at the narrowest constriction of the channel. The negative charges on carbonyls are enhanced by a buried carboxyl of El 52 that interacts with main chain nitrogens of G199 and F200. These charge pairs are lined up at a 4 A-wide constriction, thus only allowing passage of dipoles that can interact with positive and negative charges simultaneously. Neither a cation nor an anion can fulfil this dual role of polarization. Both R206 and El 52 are

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invariant, suggesting that these residues are conserved across all aquaporins and aquaglyceroporins to exclude ions. The second barrier of ionic exclusion is imposed by an insurmountable activation energy for removing water of hydration from a passing ion in the hydrophobic sector of the membrane, where the cross-section of GlpF channel is only large enough to fit one fully dehydrated ion, but the hydrophobic side of the lumen provides no replacement for water of hydration on the entire hydrophobic face. By comparison, the K + -conducting channel (KcsA) contains a stack of carbonyl rings at its narrowest constriction, thus affording negatively charged oxygens to coordinate with a passing potassium ion and replace its water of hydration (Doyle et al 1998, Roux et al 1999). The GlpF structure at 2.2 A resolution provides the first structural and chemical clues as to the mechanism of channel-selective permeability. This crystal structure represents a snapshot in the permeation process that could be hypothesized as follows. Upon entry to the channel on the extracellular side, a glycerol molecule first meets with three highly constrained carbonyl oxygens and two nitrogens at the narrowest constriction, followed by two nitrogens in the centre of the channel, and then another set of three carbonyl oxygens on the cytoplasmic side. These two sets of carbonyl triplets and two central nitrogens are arranged in a closely spaced sequential order, serving as surrogate waters to form H-bonds with OH groups of a passing glycerol (Fig. 4). It is not clear how these H-bond acceptors and donors interact with waters before a glycerol molecule enters the channel. Furthermore, we do not know the sequential order of transmembrane events that occur in the permeation process. A glycerol molecule might move down the channel in a stepwise fashion by exchanging one set of H-bonds with the next in line. Alternatively, permeation may be carried out as a series of thermally activated jumps along several distinct binding sites in the channel (Lauger & Apell 1982). These hypotheses could be clarified by further crystallographic analysis of GlpF in complex with water and other polyols. Conserved structure framework The heritage of sequence conservation in the aquaporins and aquaglyceroporins is well preserved in protein structure from bacteria to mammals. Superposing peptide backbones of GlpF and human AQP1 shows a close alignment with a root-mean-square deviation of 1.8 A, indicating that GlpF and AQP1 are essentially built upon a common structural framework. The GlpF structure reveals how tertiary interactions hold together the structural framework. The building block for GlpF is made of a helices with regular geometry as seen in many other proteins. What makes GlpF unique is the inter-helix crossing-over packing that determines the positions and orientations of helices in the structure. Crossing contacts in GlpF occur within helical bundles of M2-M1-M4—M3 and

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FIG. 4. Atomic interactions in the GlpF channel reveal the structural basis of selective permeation.

M6-M5-M8-M7 (Fu et al 2000). These two bundles are joined by an anti-parallel interaction between M2 and M6 on one side of the channel. On the other side, the two half-spanning helices M3 and M7 close up the channel by a head-to-head interaction at the conjunction of two NPA motifs (Fig. 1). Furthermore, constrained by the quasi-twofold internal symmetry, all crossing contacts in GlpF are coplanar with the quasi-twofold axis in the central membrane plane

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FIG. 5. The conserved structure framework in aquaporin and aquaglyceroporin superfamily. All interhelical contacts in GlpF are localized to the central membrane plane. Highly conserved residues involved in helix-helix contacts are shown in CPK representation.

(Fig. 5). Outward from this plane, helices diverge to form the two vestibules on each side of the membrane (Fu et al 2000). The GlpF structure suggests that the transmembrane architecture of aquaporins and aquaglyceroporins are strongly constrained by a few key tertiary interactions that are localized to the central membrane plane. Structure—sequence correlation shows that most inter-helix contacts in GlpF are made by invariant or highly conserved resides, including four glycines, G49, G96, Gl 84 and G243. Far from the usual role in disrupting a helices, these glycines allow the closest approach of helices in the central membrane plane. Perpendicular to the membrane plane is the transmembrane channel, which contains seven functional

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determinant residues, including G199, G64, N68, N203 and three charged residues, R206, El 4 and El 52. All these residues are extremely conserved from bacteria to mammals. These locations of highly conserved residues suggest that aquaporins and aquaglyceroporins are related and probably descend from a common evolutionary origin. The evolutionary divergence has been constrained by an almost absolute conservation in helix-to-helix contacts and in positions that are principal determinants for transmembrane function. Therefore, human isoforms may be built upon a conserved structural framework. Specialized functions may be developed through natural selection of mutational drifts of non-structural components. The structure of GlpF could be used as a template to model human isoforms with adequate treatments of side chain rotamers.

Future studies The crystal structure of GlpF provides the first snapshot of the transmembrane permeation process in aquaporins, and unveiled a conserved structural framework within which specialized functions may be developed in human isoforms. Molecular dynamic simulations may reveal a detailed picture of atomic motions in the process of transmembrane selective permeation. Structure—function studies may lead to identification of functional determinants in human isoforms. The assignment of functional roles to key residues in sequences of human channels is crucial to understand the genetic consequences of human polymorphisms in aquaporins and aquaglyceroporins, thereby providing the structural basis for genetic diagnosis of disease or disease risk caused by genetic variations of certain human determinant residues.

Acknowledgements We thank T. Earnest for help and support at the Advanced Light Source (ALS), Lawrence Berkley National Library, J. Krucinski for assistance in crystallization, L. Miercke for assistance in protein purification, and Drs P. Maloney, S. Sine and P. Freimuth for discussion. This work was supported by an NIH grant GM24485 to RMSDF and AL received postdoctoral support from NIH.

References Agre P 1998 Aquaporin null phenotypes: the importance of classical physiology. Proc Natl Acad Sci USA 95:9061-9063 Agre P, Mathai JC, Smith BL, Preston GM 1999 Functional analyses of aquaporin water channel proteins. Methods Enzymol 294:550-572 Beitz E, Schultz JE1999 The mammalian aquaporin water channel family: a promising new drug target. Curr Med Chem 6:457-467

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Borgnia MJ, Agre P 2001 Reconstitution and functional comparison of purified GlpF and AqpZ, the glycerol and water channels from Escherichia coli. Proc Natl Acad Sci USA 98:2888-2893 Borgnia MJ, Nielsen S, Engel A, Agre P 1999a Cellular and molecular biology of the aquaporin water channels. Annu Rev Biochem 68:425-458 Borgnia MJ, Kozono D, Calamita G, Maloney PC, Agre P 1999b Functional reconstitution and characterization of AqpZ, the E. coli water channel protein. J Mol Biol 291:1169-1179 Braun T, Philippsen A, Wirtz S et al 2000 The 3.7A projection map of the glycerol facilitator GlpF: a variant of the aquaporin tetramer. EMBO Rep 1:183-189 Calamita G, Bishai WR, Preston GM, Guggino WB, Agre P 1995 Molecular cloning and characterization of AqpZ, a water channel from Escherichia coli. J Biol Chem 270:29063-29066 Cheng A, van Hoek AN, Yeager M, Verkman AS, Mitra AK 1997 Three-dimensional organization of a human water channel. Nature 387:627-630 Doyle DA, Morais Cabral J, Pfuetzner RA et al 1998 The structure of the potassium channel: molecular basis of K+ conduction and selectivity. Science 280:69-77 Fu D, Libson A, Miercke LJ et al 2000 Structure of a glycerol-conducting channel and the basis for its selectivity. Science 290:481-486 Heller KB, Lin EC, Wilson TH 1980 Substrate specificity and transport properties of the glycerol facilitator of Escherichia coli. J Bacteriol 144:274-278 Hermann JB, Engel A 2000 Structural clues in the sequences of the aquaporins. J Mol Biol 295:1039-1053 Kishida K, Kuriyama H, Funahashi T et al 2000 Aquaporin adipose, a putative glycerol channel in adipocytes. J Biol Chem 275:20896-20902 Lauger P, Apell HJ 1982 Jumping frequencies in membrane channels. Comparison between stochastic molecular dynamics simulation and rate theory. Biophys Chem 16:209-221 Maurel C, Reizer J, Schroeder JI, Chrispeels MJ, Saier MH Jr 1994 Functional characterization of the Escherichia coli glycerol facilitator, GlpF, in Xenopus oocytes. J Biol Chem 269:1186911872 Murata K, Mitsuoka K, Hirai T et al 2000 Structural determinants of water permeation through aquaporin-1. Nature 407:599-605 Park JH, Saier MH Jr 1996 Phylogenetic characterization of the MIP family of transmembrane channel proteins. J Membr Biol 153:171-180 Preston GM, Agre P 1991 Isolation of the cDNA for erythrocyte integral membrane protein of 28 kilodaltons: member of an ancient channel family. Proc Natl Acad Sci USA 88: 11110-11114 Preston GM, Carroll TP, Guggino WB, Agre P 1992 Appearance of water channels in Xenopus oocytes expressing red cell CHIP28 protein. Science 256:385-387 Roux B, MacKinnon R 1999 The cavity and pore helices in the KcsA K+ channel: electrostatic stabilization of monovalent cations. Science 285:100-102 Sanders OI, Rensing C, Kuroda M, Mitra B, Rosen BP 1997 Antimonite is accumulated by the glycerol facilitator GlpF in Escherichia coli. J Bacteriol 179:3365-3367 Unger VM 2000 Fraternal twins: AQP1 and GlpF. Nat Struct Biol 7:1082-1084 Verkman AS 1999 Lessons on renal physiology from transgenic mice lacking aquaporin water channels. J Am Soc Nephrol 10:1126-1135 Verkman AS 2000b Physiological importance of aquaporins: lessons from knockout mice. Curr Opin Nephrol Hypertens 9:517-522 Verkman AS 2000a Water permeability measurement in living cells and complex tissues. J Membr Biol 173:73-87 Verkman AS, Yang B, Song Y, Manley GT, Ma T 2000 Role of water channels in fluid transport studied by phenotype analysis of aquaporin knockout mice. Exp Physiol 85:233S-241S

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Walz T, Hirai T, Murata K et al 1997 The three-dimensional structure of aquaporin-1. Nature 387:624-627 Wistow GJ, Pisano MM, Chepelinsky AB 1991 Tandem sequence repeats in transmembrane channel proteins. Trends Biochem Sci 16:170-171

DISCUSSION Mindell: You talked about the conservation of these channels. Are there specific sequence differences that would explain this dramatically different selectivity? Fu: AQPs and GlpF have about 25% sequence similarity. But if we look at the channel-lining residues, they are highly conserved with the only exception at position 200. The GlpF has a Phe while AQP1 has a Cys at this position. Despite this strong similarity, there is no clear indication of the source of the water/glycerol selectivity. Mitra: This is debatable. Different labs do experiments differently. Some labs have claimed that GlpF does not conduct water very well at all. If you compare it with the sequence of AQP3 in mammals (which is a glycerol transporter), it does not have a phenylalanine in that place, but a slightly polar tyrosine. In fact, at this position only GlpF has a phenylalanine. All the others have tyrosine there. We think Hisl80 in AQP1 at the analogous site is important, but at that position, 180, it is glycine/valine for aquaglyceroporins. The additional polar residue there, in addition to phenylalanine versus cysteine, might act together to provide that occlusion of glycerol or transport of water. Sansom: You said that it might be the extracellular mouth that contributes to the selectivity. Where in the sequence does this occur? Fu: It is the only part that is not conserved. The selectivity is largely contributed by the linking group between the two halves of the protein. Mitra: This is between helices 3 and 4. Sansom: Has anyone tried making chimeras in which they switch these? Mitra: Bai et al (1996) have done switching between AQP2 and AQP1 in the hydrophilic loops. These resulted in chimeras with permeability similar to the wild-types. On the other hand an AQP2 chimera in which a fragment of 3—4 loop of GlpF replaced a corresponding segment of AQP2 displayed significantly reduced water permeability. Doyle: GlpF also has the conserved NP A residues. This probably has a role in the water movement, but does it have any role in the movement of glycerol? Fu: Yes, the two NPA tripeptide sequences are located in the middle of the channel. The Asp206 and Asp68 form H-bonds with two OH groups of a glycerol molecule that is bound in the middle of the channel. Choe: I would predict that the main chain phi, psi angles of glycine 199 are positive.

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Fu: I have checked the geometries. All the main chain, with the exception of F200, is located in the allowed areas of the Ramachandran plot. Sansom: So is your suggestion that the El 52 is aiding that distortion? Or do you think that it might be polarizing the carbonyl oxygen? Fu: The entire side chain of El52 is embedded in a highly hydrophobic environment. The only way that El52 could gain access to the aqueous channel is through hydrogen bonding interaction with the main chain nitrogen. This side chain—main chain interaction could strongly polarize carbonyl oxygens in the vicinity. Jordan: I am having a hard time seeing what it is about the interior of this channel that makes it difficult for water to get inside. Otherwise I worry about glycerol getting in there and just being stuck. What is it that keeps the water out? Fu: This is also a problem I have! People used to think that GlpF was only permeable to glycerol. I kept asking myself how this channel could be only permeable to glycerol but not water. Andy Libson and I came up with the following explanation. When a water molecule moves down the channel, the water itself has to be dehydrated because waters in aqueous solution are linked together by hydrogen bonds. Moving a water molecule through a narrow channel therefore requires removal of its hydration shell first. Further, to satisfy hydrogen donor and acceptor groups lining the hydrophilic face of the channel, at least two water molecules are required to be in a line. However, a string of water molecules does not have a non-polar backbone to satisfy the hydrophobic face of the channel. Jordan: So what you are saying is that you think that there might be a genuine free energy barrier for water to enter, because you have to break up the water structure. The glycerol is nicely designed to be solvated by the channel. Fu: Yes. Sansom: We should remember that structure is solved at low temperature (100 K). Things might move a little bit. Choe: I thought that the polarity of the molecule with respect to the membrane was strikingly clear. Is this the case for all aquaporin channels? Fu: Like other membrane proteins in general, GlpF follows the positive inside rule. There are 16 excess positive charges on the cytoplasmic face. But I do not think that all these charges are conserved. Choe: In other membrane proteins the polarity is not as distinct as in yours. Fu: There are two rings of charged side chains, forming two layers of charges which sandwich two layers of aromatic side chains that are located on the polarnon-polar interface of the membrane. These aromatic rings seem to be a common feature in membrane proteins. Choe: What is the membrane potential of the inner membrane of E. coli?

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Fu: I would guess it is — 30 to — 70 mV. Mindell: I have usually heard it quoted as —200 mV. It is a very high potential. Choe: It could therefore be more pronounced for E. coll water channels to have that stability by creating higher polarity. Roux: People say 'stability', but you don't expect a membrane protein to tumble over (i.e. with the intracellular side pointing outward). The presence of positively charged amino acids on the intracellular side may have more to do with the folding and trafficking than the stability of the folded protein itself. Fu: I think this dipole probably helps to orientate GlpF in the membrane. The protein is formed by two halves that have strong sequence similarities, and the dipole means that they can only organize in the membrane in the correct way. Mitra: In many cases the charge on one leaflet of the bilayer is different from the other. This also acts to orient the membrane protein in a particular way. Choe: Do you also have a similar level of polarity in your model? Mitra: I have not checked it in detail. Ashcroft: How long a molecule can go through the channel? Fu: Probably 20 A. A shcroft: Is it possible for something to span the channel? Fu: Yes, if you have a polyol of 5 or 6 carbons, this will be much longer than the pore. Choe: But these sugars are rings, not long chains. If you build that stereochemistry, the hydroxyl groups are all on one side. In one of your stereotypical variants, some are on the other side, but it had only sevenfold less permeability. Fu: This is because we were depicting things in two dimensions. If you view a three-dimensional molecule, the stereo arrangement of the OH groups is not so much different. Choe: It occurs to me that your sugar may linearize in order to cross over. As a ring you can't force it through the pore. Fu: People have used ring sugars to examine the permeability of the GlpF channel. Mindell: My impression was that those weren't sugars, they were polyalcohols. Fu: Yes. They don't form rings. Wallace: Physiologically, what is the main role of this channel? Is it to transport polyalcohols, glycerol, water or something else? Fu: These proteins can be induced in the presence of glycerol. This is an inducible glycerol-permeable channel. People believe that this channel is involved in glycerol uptake. When the channel is knocked out the bacteria grow much slower. Ashcroft: At the beginning of your presentation you showed four pores, one in each monomer, and you also had a channel in the centre, which you haven't said anything about.

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Fu: That putative channel was surprisingly empty along the fourfold axis. We do not know the physiological function of this fourfold pore. This channel has a small hydrophobic constriction on the cytoplasmic side, which is too small to allow permeation of any type of ion or water. Choe: When you isolate your protein, is it a stable tetramer? Does it ever monomerize or dimerize? Fu: When I purify the protein it is not stable. We have spent a long time trying to work out why. Perozo: What do you mean when you say that it isn't stable? Fu: This is a typical problem with membrane proteins. Then one day, by accident we found that if we added Ca2+ or Mg2+, it made the GlpF more stable. After the structure was solved, it became clear that Ca2+ or Mg2+ could help in stabilizing the formation of tetramer. In the absence of divalent cations, GlpF may be in either a tetrameric or monomeric form. Choe: At low concentrations do you ever see monomers? They have to go to monomers at some point if you keep diluting. Fu: But how can we determine this? The protein is in the detergent and it is very hard to determine the precise molecular weight. Choe: It is hard, but it is probably doable if you know how much detergent is bound in the complex. Schrempf: Has anyone done mutagenesis to prevent the contacts among the monomers? This would address the question. Mttra: There are some unpublished reports from the French group in Rennes that the amount of detergent in the purification cocktail can also modulate the oligomeric state. At low concentration of octyl glucoside (~ 1%), they claim that more of GlpF is in monomeric state, and as the detergent concentration is increased (~10%) the proportion of tetrameric structure increases. This is somewhat counterintuitive. Fu: I suspect that tetramer formation may be essential for the stability of the protein. Two transmembrane-spanning helices in GlpF are located close to the tetramer fourfold axis, each is approximately 20 A in length. They are apparently not long enough to cross the membrane if not forming the tetramer. By forming the tetramer, GlpF shields the short helices from the hydrophobic core of the membrane. Mitra: Couldn't this be done by dimers? Fu: That is possible. Papazian: What about the results of something simple such as sucrose gradients? Does the protein migrate as a single species? Fu: This will depend on how many detergent molecules are binding to a protein. The protein can go just about anywhere. One way to estimate detergent number per micelle would be to use radioactive detergent. This will allow one to calculate

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how many detergent molecules are bound per protein, and then it is possible to estimate their contribution to the apparent molecular weight of the proteindetergent micelle. Choe: Even with the analytical centrifuge you need to account for how much detergent is present in the complex. The lipid density is significantly different from protein density. Reference Bai L, Fushimi K, Sasaki S, Marumo F1996 Structure of aquaporin-2 vasopressin water channel. J Biol Chem 271:5171-5176

Water in ion channels and pores — simulation studies Mark S.P. Sansom, Peter Bond, Oliver Beckstein, Philip C. Biggin, Jose Faraldo-Gomez, Richard J. Law, George Patargias and D. Peter Tieleman* Laboratory of Molecular Biophysics, Department of Biochemistry, Rex Richards Building, University of Oxford, South Parks Road, Oxford, OX1 3QU, UK and * Department of Biological Sciences, University of Calgary, 2500 University Drive NW, Calgary, Alberta, Canada T2N 1N4

Abstract. The microscopic properties of water in narrow pores are relevant to the function of ion channels and related membrane transport proteins. The emergence of several high-resolution structures allows one to perform molecular dynamics simulation studies of water in such pores. Simulations of bundles of parallel a-helical peptides (e.g. alamethicin) have enabled development of methodologies and concepts appropriate to such investigations. In the narrow channels formed by such bundles, water molecules exhibit reduced rotational and translation motion. This reduction in water mobility may be a general property of narrow pores. We have used simplified channel models to explore the role of hydrophobicity/hydrophilicity in the entry of water into pores. Narrow pores with a hydrophobic lining, although physically open, may not admit water molecules, acting as a 'hydrophobic gate' that prevents water and ion permeation. Such a gate can be opened either by widening the pore or making its lining more polar. Simulations have been used to explore the behaviour of water in GlpF, a member of the aquaporin family of water pores, and OmpA, a bacterial outer membrane protein. Preliminary results suggest that a continuous water wire is not formed within the amphipathic GlpF pore. Simulations of OmpA, in which polar residues line the channel, indicate that a small conformational change in one of the channel lining side chains may open the channel. In summary, comparison of the behaviour of water in different narrow transmembrane pores suggests that an amphipathic pore is ideal for water permeation, and that either a highly hydrophobic pore lining or a charged pore-lining region can act as a gate. 2002 Ion channels—from atomic resolution physiology to functional genomics. Wiley, Chichester ( Novartis Foundation Symposium 245) p 66-83

Ion channels and related integral membrane proteins form transbilayer pores through which ions and/or other small molecules (e.g. water) may pass in a rapid (c. 108 s-1) but selective fashion. In order to understand the physical basis of rapid permeation of ions and/or water through such pores (Tieleman et al 2001) it is important that we fully understand the properties of solvent (i.e. water) 66

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FIG. 1. (A) Schematic diagram of a pore/channel formed by a transmembrane protein (dark grey) in a lipid bilayer membrane (cross-hatched). The water properties within the pore (p) are perturbed relative to those of bulk water (b), the degree of perturbation being indicated by the depth of the shading. (B) An example of a transbilayer pore, namely that formed by the bacterial protein GlpF (Fu et al 2000). The protein is shown in 'ribbons' format, with the approximate extent of the lipid bilayer indicated by the horizontal lines. The inner of surface of the pore is shown. Calculated using HOLE (Smart et al 1997,1993).

molecules within the pore, and how they may differ from those of water in its bulk state (i.e. on either side of the cell membrane). This is especially true for water pores (aquaporins). For ion channels, an understanding of pore water is an essential first step to understanding the dynamics and electrostatics of ions in the pore. Consider the structure of a 'typical' channel protein (Fig. 1A). The structures of a number of membrane proteins are known from X-ray crystallographic and electron microscopic studies of membrane proteins (see http:|| indigo1.biop.ox.ac.uk/membraneproteins.html for a summary). These include: (i) several porins from bacterial outer membranes (Schirmer 1998); (ii) bacterial potassium (KcsA; Doyle et al 1998) and mechanosensitive (MscL; Chang et al 1998) channels; (iii) the nicotinic acetylcholine receptor channel (nAChR; Unwin 2000); and (iv) two members (GlpF [Fig. 1B] and AQP1; Fu et al 2000, Murata et al 2000, Ren et al 2001) of the aquaporin family of water pores. From these we may generalize that a 'typical' pore is c, 3 nm long and has a radius which varies along the length of the pore, but which may be as low as c. 0.2nm in places (n.b. the radius of a water molecule is c. 0.l5nm, and that of a K+ ion is c. 0.13nm). From this it is evident that the pore dimensions are comparable to the size of the transported molecules and/or ions. It has been known for some time, from both experimental and

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theoretical studies, that liquids behave differently when confined in pores, slits and cavities of molecular dimensions relative to the same liquids in the bulk state (Granick 1991). Thus we would expect non-bulk properties of water within transbilayer pores. Of particular importance to channel and related pore proteins are the dynamic behaviour of water (e.g. diffusion coefficient) and also its electrostatic properties (e.g. local dielectric constant) (Roux et al 2000, Tieleman et al 2001). It is somewhat difficult to address these local microscopic properties experimentally, especially for biological pores. However, simulations provide ready access to information on the structural dynamics of water in association with proteins (Daggett & Levitt 1993). There is a considerable literature on the use of molecular dynamics (MD) simulations to explore the structure and dynamics of water, and a number of water models have been developed for use in such simulations (Jorgensen et al 1983, Levitt et al 1997). Although none of these models is perfect, comparison of the simulated properties of water in pores with water in bulk enables one to explore how the molecular environment provided by a pore alters the behaviour of the solvent. Recent years have witnessed an increase in the level of detail included in models of pores used in simulations. Initial studies employed either highly idealized models (e.g. smooth cylindrical pores [LyndenBell & Rasaiah 1996] or cavities [Sansom et al 1996]) or model peptides that capture some of the essential features of more complex ion channels (Chiu et al 1999, Roux & Karplus 1994, Tieleman et al 1999). More recently it has been possible to simulate complex channel and pore proteins, embedded in either an explicit lipid bilayer (Berneche & Roux 2000, Shrivastava & Sansom 2000) or a bilayer mimetic octane slab (Guidoni et al 2000). An example of a study of a model system is provided by simulations of pores formed by the peptide alamethicin (Sansom 1993). Water molecules diffused more slowly within the pore, and were non-randomly oriented relative to the pore axis. Initial studies used models of solvated alamethicin helix bundles in vacuo (Breed et al 1996). These were extended to alamethicin channels embedded in a fully solvated phospholipid bilayer (Tieleman et al 1998). Encouragingly, similar results were obtained from both sets of simulations, and from simulations on a slightly different peptide channel model (Mitton & Sansom 1996, Randa et al 1999). By combining results from simulations on a variety of systems performed by a number of research groups (reviewed in Tieleman et al 2001) it has been possible to analyse how DPORE /D B U L K (i.e.the ratio of the diffusion coefficient in the pore to that in bulk solution) varies as a function of (local) pore radius. For both water molecules and for simple monovalent ions a similar pattern emerges. Namely, that for radii above c. 0.8—1.0 nm the behaviour of water in the pore matches that of bulk water, whereas for narrower pores the value ofD P O R E / D B U L Kdecreases

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progressively until, at a radius of c. 0.2nm or less, it is not clear whether one is justified in describing water and/or ion motions in term of 'diffusion'. In this paper we examine three specific aspects of the behaviour of water in transbilayer pores: (i) water in a narrow hydrophobic 'gate'; (ii) water in the amphipathic pores formed by a member of the aquaporin family; and (iii) water within a highly charged pore-like cavity in the bacterial outer membrane protein OmpA.

Methods MD simulations were run using Gromacs v2.0 (http://www,gromacs.org/). The water model used was simple point charge (SPC) (Hermans et al 1984, van Gunsteren et al 1996). The time step used in all simulations was 2fs. NPT (i.e. constant number of particles, temperature and pressure) conditions were applied, with a constant pressure of 1 bar maintained independently in all three directions via a coupling constant of Tp = l.0ps (Berendsenet al 1984). Where present, water, lipid or octane, and protein were coupled separately to a temperature bath at 300 K with a coupling constant of tT = 0.1 ps. The LINCS algorithm (Hess et al 1997) was used to constrain bond lengths. Octane parameters were from GROMOS96 (van Gunsteren et al 1996). Long-range interactions were treated via PME (Darden et al 1993) using a cut-off of 0.9 nm for the direct space non-bonded calculations. Other simulation details were as described in Tieleman et al (2000). Analysis of simulation results was performed using GROMACS and/or locally written code. Pore radius profiles were analysed using HOLE (Smart et al 1997, 1993). Diagrams were drawn using RasMol (Sayle & Milner-White 1995), VMD (Humphrey et al 1996), MOLSCRIPT (Kraulis 1991) and Raster3d (Merritt & Bacon 1997).

Results Ahydrophobic gate Most ion channels are gated, i.e. they switch between an open state and a closed state. Gating can occur on a wide range of timescales and can be intrinsic (i.e. presumed to be due to random thermal motions of a channel protein) or can be coupled to allosteric modulation of channel protein conformation by ligand binding and/or voltage driven conformational changes. Structural studies on a number of channels suggest that a gate may be formed by a ring of side chains that form a narrow, hydrophobic region within a channel through which ions cannot pass. For example, in the nAChR it is suggested that a ring of conserved leucine side chains forms a hydrophobic region of length c. 0.8nm and radius c. 0.35 nm (Unwin 2000). In MscL there are two rings of hydrophobic side chains

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FIG. 2. (A) Diagram of a simplified model of a pore in a membrane mimetic. The pore (p) is shown as semi-transparent space-filling spheres, and the water molecules (w) as 'sticks'. The membrane mimetic (m) is omitted for clarity. The pore radius is 4.5 A, the mouth radius is 18 A, and two parallel dipoles (each of magnitude 2 x 10 -29 Cm) were present in the pore lining. (B) Average degree of openness (FO) vs. pore radius (Rp) for hydrophobic pores with various radius 18 A (solid line) and for equivalent pores with two parallel dipoles in the pore lining (broken line). Simulation times were 1 ns per data point.

that narrow the pore in the closed state of the channel, whereas in KcsA a ring of valine side chains forms a hydrophobic gate at the intracellular mouth of the pore. In this context, it is of interest to explore the pore dimensions and degree of hydrophobicity that are consistent with formation of a (closed) gate. In the longer term one should look at gating in terms of the energetics of ion permeation (Biggin et al 2001). Here, we explore gating of water entry into a pore. We assume that if water is excluded from a gate region then so will ions. A simplified model of a pore in a membrane was used in these simulations (Fig. 2A). Both the pore and the membrane were constructed from methane-like atoms, subject to positional restraints during the simulations. The dimensions of the pore region were adjusted to explore the effect of different pore and mouth radii on water behaviour. In the first set of simulations the pore lining atoms were apolar, i.e. their partial atomic charges were zero. In a second set of simulations, small dipoles were added to the pore lining by adding suitable partial charges to the pore-lining atoms. For the various pore configurations, 1 ns MD simulations were performed in the presence of SPC waters on either side of the pore. Water molecules were free to enter/exit the pore, and the penetration of water molecules into the pore (i.e. 'wetting') was measured. The openness of a given pore was defined as the time averaged density of water within the pore relative to the bulk density of water.

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The openness of simple hydrophobic pores depends strongly on the pore geometry. As the pore radius is increased there is a clear transition between a closed pore (i.e. one from which water is excluded) and an open pore (i.e. filled with water; Fig. 2B). The exact pore radius for the transition is influenced by other aspects of the pore geometry (Beckstein et al 2001), e.g. the mouth radius, but typically is c. 0.45 nm. A radius of 0.45 nm is in excess of that necessary to accommodate water molecules from a simple steric perspective. Thus a geometrically open (i.e. unoccluded) pore can remain functionally closed (i.e. exclude water molecules) if it is sufficiently hydrophobic. Adding two parallel dipoles (magnitude 2 x 10 - 2 9 Cm) to the pore lining was sufficient to open a pore that was otherwise closed (Fig. 2B). Again, the exact behaviour depended on the geometry of the pore but the trend was the same in all simulations, namely that making a narrow pore more polar can open it to entry of water molecules. Thus these simplified model channel simulations provide a proof of the principle of hydrophobic gating, i.e. a hydrophobic region in a pore can completely exclude water molecules even if it is not sterically occluded. This 'gate' can then be opened either by increasing the local pore radius and/or increasing the polarity of the pore lining. Aquaporins The aquaporins are a ubiquitous family of water permeable pores, found in organisms ranging from bacteria to animals and plants. Aquaporins per se are selective for water permeation whereas glyceroaquaporins also permit glycerol permeation (Borgnia & Agre 2001). Structures are known for AQP1 (at medium resolution by electron microscopy; Murata et al 2000, Ren et al 2001) and for GlpF (at 0.22 nm resolution, by X-ray diffraction; Fu et al 2000). In both cases the protein forms a six-helix bundle spanning the membrane. The bundle contains two porelining NPA-motif containing loops that help to line a central pore. The pore has an average radius of c. 0.2nm (Fig. 1B) and is amphipathic, i.e. it is lined by both hydrophobic and polar amino acids. A 5 ns duration simulation has been run for a GlpF monomer embedded in a bilayer mimetic octane slab in order to study the behaviour of water molecules within pores of this family. The protein conformation does not change significantly during the course of this simulation (Fig. 3A,B). Comparing the structures at the start and end of the simulation, the Ca RMSD is c. 0.2nm, which is typical for a simulation of a membrane protein based on an X-ray structure. Furthermore, the change in structure over the course of the simulation is substantially less for the TM a helices and for the NPA loops than is suggested by this overall figure, as structural fluctuations are greatest for the surface-exposed inter-helical loops. Based on simple considerations of the single molecule water

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FIG. 3. (A,B) Superimposed Ca traces of the GlpF molecule at the start (thin black line) and end (thick grey line) of a 5 ns simulation, viewed perpendicular (A) and down (B) the pore axis. (C) Snapshots from a simulation of GlpF in a membrane mimetic octane slab. The diagrams show just the water molecules within and at either mouth of the pore. The orientation is the same as that in Fig. 1B, i.e. with the extracellular mouth of the pore uppermost. Snapshots are shown at t = 0-5ns.

permeability of aquaporin (c. 3 x 109 waters s -1; Murata et al 2000) and the relative permeability of GlpF and aquaporin to water (c. 1:6; Borgnia & Agre 2001) one arrives at a mean time spent by a water molecule in GlpF of c. 3 ns. Thus one might expect to see one or two water permeation events in a 5 ns simulation. Some care was taken to ensure that the GlpF pore was fully solvated at the start of the simulation. Thus, at the start of the simulation the pore accommodated c. 26 water molecules. Snapshots of the water molecules within the pore taken during the simulation (Fig. 3C) indicate that the initially continuous column of waters is

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not stable but breaks during the course of the simulation. Analysis of the number density of water along the pore axis, averaged in 0.5 ns blocks indicates that during the simulation the water density falls to almost zero at various positions along the pore despite the absence of any significant changes in pore radius profile during the course of the simulation. At the end of the 5 ns the number of waters within the pore is c. 17. Diffusion coefficients of water molecules within the pore are lowered (c. threefold) relative to bulk water—this is perhaps a smaller decrease than might have been expected on the basis of the pore dimensions. By following the trajectories of those water molecules that spend at least 30% of their time within the pore one can see the water molecules that enter and leave the pore. One water molecule undergoes a 'permeation event' i.e. it enters one mouth of the pore, remains within for c, 2 ns and then leaves via the opposite mouth. Thus simulations of this duration begin to probe water permeation. Water movement within the pore does not appear to occur in a strict single-filing fashion. On the basis of these simulations we may conclude that water molecules penetrate the GlpF pore but do not form a continuous H-bonded chain throughout the simulation. Rather, a fluctuating and discontinuous chain of water molecules appears to be formed. This is in some respects similar to the behaviour of water in the simple model pore described above when the latter has a polar pore lining such that the average degree of openness FO c. 0.5. In this case the time-dependent density of water within the pore fluctuates between c. 0 and c. 1 relative to bulk density (Beckstein et al 2001). Interestingly, in long (>50ns) simulations of water in a carbon nanotube, Hummer et al (2001) observe water flux through the pore to occur in a pulsatory fashion, with fluctuations on a timescale of c. 4 ns. Thus, it may be a general property of water permeable pores that the water density within such pores fluctuates. Indeed, recent theoretical studies have emphasised the relationship of fluctuations in number of particles within a pore to its geometry (Bezrukov et al 2000). OmpA OmpA is a small outer membrane protein from Escherichiacoli. The structure of the N-terminal transmembrane domain (residues 1-171) of this protein has been solved by X-ray diffraction (Pautsch & Schulz 2000) and by NMR (Arora et al 2001). The protein forms an eight-stranded anti-parallel b barrel. There is some debate as to whether or not the OmpA protein can form a channel. On the one hand, the X-ray structure failed to reveal a continuous pore through the centre of the barrel. Instead there are three internal clusters of bound water molecules, containing four (extracellular cluster), eight (central cluster) and eight (periplasmic cluster) waters (Fig. 4A), separated from one another by salt bridges formed between ionic side chains in the protein interior. On the other hand,

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FIG. 4. (A) Structure of OmpA (shown as a trace of the polypeptide backbone) with the internal water molecules shown as spheres. The horizontal lines indicate the approximate location of the bilayer, and the region of the E52/R138 'gate' is labelled 'g'. (B) Trajectories, projected onto the pore (z) axis, of water molecules within an OmpA molecule embedded in a DMPC bilayer over the course of a 5 ns MD simulation. The location of the aromatic bands (presumed to correspond to the approximate extent of the bilayer) are shown by the arrows and the extent of the 'gate' region is labelled 'g'.

functional measurements have consistently revealed channel formation by OmpA (Arora et al 2000) and by related proteins (Brinkman et al 2000, Saint et al 2000) reconstituted into artificial lipid bilayers. For refolded OmpA, two different conductance levels are observed (Arora et al 2000). The lower conductance level (c. 60 pS in 1 M KC1) is consistent with a pore formed by an eight-stranded barrel. Thus, it is of some interest to examine the simulated dynamics of water molecules within OmpA, with the protein embedded in either an octane slab or a phospholipid bilayer (Bond et al 2001). The results of a 5 ns simulation of the 'closed' form of OmpA (i.e. that seen in the crystal structure) embedded in a DMPC bilayer are summarised in Fig. 4B in the form of trajectories of the internal water coordinates projected onto the barrel (i.e. z) axis. From this it is evident that water molecules show some movement within a cluster on a 5 ns timescale. This and other simulations of OmpA have revealed waters to exchange between the bulk and the two outer clusters, i.e. through the

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FIG. 5. Schematic of three classes of pore. (A) Hydrophobic pore (dark grey) in a hydrophobic membrane (dark grey). Water molecules (stippled grey) are present on either side of the pore but fail to enter. (B) Amphipathic pore (light grey) with dipolar side chains (arrows), and filled by water. (C) Polar pore (light grey) penetrated by waters but gated by a side chain ion pair in the centre of the pore.

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inner and outer 'mouths' of the pore. However, water was not observed to exchange across the 'gate' region (formed by side chains E52 and R138), i.e. between the central cluster and the periplasmic cluster. Hence, at least on a 5 ns timescale, the OmpA protein remains in a 'closed' conformation. By combining model building and further simulation studies (data not shown; Bond et al 2002) we have shown that a minor rearrangement of salt bridges between side chains within the interior of the molecule can 'open' the pore, at least to water molecules. Thus, it is tempting to identify the E52/R138 side chain pair with the gate of the OmpA channel.

Conclusions What may one conclude from these studies? Firstly it is evident that simulations can provide detailed insights into the dynamic behaviour of water within transbilayer pores. Of course there are limitations to such simulations, including their relative short duration (up to c. 10ns) so that statistical sampling is poor for longer timescale (e.g. permeation) events. Furthermore, there are concerns about the approximations implicit in the simulation forcefields employed. However, by combining simulations with suitable reality checks wherever possible one may start to realise some atomistic level detail of the dynamics of pores. On the basis of the results presented in this paper one may draw a tentative conclusion concerning the relationship between hydrophobicity/polarity of the pore lining and water permeability (Fig. 5). Note that these conclusions apply to pores of radius comparable to that of one or two water molecules. A completely hydrophobic pore (region) can act as a gate by resisting 'wetting'. A highly charged pore lining can also act as a gate by providing regions of the pore that form salt bridges and so exclude waters. In between these two extremes an amphipathic pore appears to be optimal for water permeation, as found in the AQP family of water pore proteins. Acknowledgements Research in MSPS's laboratory is supported by the Wellcome Trust. Studentships were provided by the Wellcome Trust (OB), MRC (RL), EPSRC & La Caixa Foundation (JFG), and State Scholarship Foundation (Greece) (GP). DPT is an AHMFR Scholar. We thank the Oxford Supercomputing Centre for computing support. Our thanks to Dax Fu and Alok Mitra for their interest in this work and for stimulating discussions concerning aquaporins.

References Arora A, Rinehart D, Szabo G, Tamm LK 2000 Refolded outer membrane protein A of Escherichia coli forms ion channels with two conductance states in planar lipid bilayers. J BiolChem 275:1594-1600

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Arora A, Abildgaard F, Bushweller JH, Tamm LK 2001 Structure of outer membrane protein A transmembrane domain by NMR spectroscopy. Nat Struct Biol 8:334-338 Beckstein O, Biggin PC, Sansom MSP 2001 A hydrophobic gating mechanism for nanopores. J Phys Chem B 105:12902-12905 Berendsen HJC, Postma JPM, van Gunsteren WF, DiNola A, Haak JR 1984 Molecular dynamics with coupling to an external bath. J Chem Phys 81:3684—3690 Berneche S, Roux B 2000 Molecular dynamics of the KcsA K+ channel in a bilayer membrane. BiophysJ 78:2900-2917 Bezrukov SM, Berezhkovskii AM, Pustovoit MA, Szabo A 2000 Particle number fluctuations in a membrane channel. J Chem Phys 113:8206-8211 Biggin PC, Smith GR, Shrivastava 1H, Choe S, Sansom MS 2001 Potassium and sodium ions in a potassium channel studied by molecular dynamics simulations. Biochim Biophys Acta 1510:1 9 Bond P, Faraldo-Gomez J, Sansom MSP 2002 OmpA— A pore or not a pore? Simulation and modelling studies. Biophys J, in press Borgnia MJ, Agre P 2001 Reconstitution and functional comparison of purified GlpF and AqpZ, the glycerol and water channels from Escherichia coli. Proc Nat Acad Sci USA 98:2888-2893 ' Breed J, Sankararamakrishnan R, Kerr ID, Sansom MS 1996 Molecular dynamics simulations of water within models of ion channels. Biophys J 70:1643—1661 Brinkman FSL, Bains M, Hancock RE, 2000 The amino terminus of P seudomonas aeruginosa outer membrane protein OprF forms channels in lipid bilayer membranes: correlation with a three dimensional model. J Bacteriol 182:5251-5255 Chang G, Spencer RH, Lee AT, Barclay MT, Rees DC 1998 Structure of the MscL homolog from Mjcobacterium tuberculosis: a gated mechanosensitive ion channel. Science 282:2220-2226 Chiu SW, Subramaniam S, Jakobsson E 1999 Simulation study of a gramicidin/lipid bilayer system in excess water and lipid. I. Structure of the molecular complex. Biophys J 76:19291938 Daggett V, Levitt M 1993 Realistic simulations of native-protein dynamics in solution and beyond. Annu Rev Biophys Biomol Struct 22:353-380 Darden T, York D, Pedersen L 1993 Particle mesh Ewald — an N.log(N) method for Ewald sums in large systems. J Chem Phys 98:10089-10092 Doyle DA, Morais Cabral J, Pfuetzner RA et al 1998 The structure of the potassium channel: molecular basis of K+ conduction and selectivity. Science 280:69—77 Fu D, Libson A, Miercke LJ et al 2000 Structure of a glycerol-conducting channel and the basis for its selectivity. Science 290:481-486 Granick S 1991 Motions and relaxations of confined liquids. Science 253:1374-1379 Guidoni L, Torre V, Carloni P 2000 Water and potassium dynamics in the KcsA K+ channel. FEES Lett 477:37-42 Hermans J, Berendsen HJC, van Gunsteren WF, Postma JPM 1984 A consistent empirical potential for water-protein interactions. Biopolymers 23:1513-1518 Hess B, Bekker H, Berendsen HJC, Fraaije JGEM 1997 LINGS: a linear constraint solver for molecular simulations. J Comp Chem 18:1463-1472 Hummer G, Rasaiah JC, Noworyta JP 2001 Water conduction through the hydrophobic channel of carbon nanotube. Nature 414:188-190 Humphrey W, Dalke A, Schulten K 1996 VMD: visual molecular dynamics. J Mol Graph 14:33 38 Jorgensen WL, Chandresekhar J, Madura JD, Impey RW, Klein ML 1983 Comparison of simple potential functions for simulating liquid water. J Chem Phys 79:926—935 Kraulis PJ 1991 MOLSCRIPT: a program to produce both detailed and schematic plots of protein structures. J Appl Cryst 24:946-950

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Levitt M, Hirshberg M, Sharon R, Laidig KE, Daggett V 1997 Calibration and testing of a water model for simulation of the molecular dynamics of proteins and nucleic acids in solution. ] Phys Chem B 101:5051-5061 Lynden-Bell R, Rasaiah JC 1996 Mobility and solvation of ions in channels. J Chem Phys 105:9266-9280 Merritt EA, Bacon DJ 1997 Raster3D: Photorealistic molecular graphics. Methods Enzymol 277:505-524 Mitton P, Sansom MSP 1996 Molecular dynamics simulations of ion channels formed by bundles of amphipathic a-helical peptides. Eur Biophys J 25:139-150 Murata K, Mitsuoaka K, Hirai T et al 2000 Structural determinants of water permeation through aquaporin-1. Nature 407:599-605 Pautsch A, Schulz GE 2000 High-resolution structure of the OmpA membrane domain. J Mol Biol 298:273-282 Randa HS, Forrest LR, Voth GA, Sansom MSP 1999 Molecular dynamics of synthetic leucineserine ion channels in a phospholipid membrane. Biophys J 77:2400-2410 Ren G, Reddy VS, Cheng A, Melnyk P, Mitra AK 2001 Visualization of a water-selective pore by electron crystallography in vitreous ice. Proc Natl Acad Sci USA 98:1398-1403 Roux B, Karplus M 1994 Molecular dynamics simulations of the gramicidin channel. Ann Rev Biophys Biomol Struct 23:731-761 Roux B, Berneche S, Im W 2000 Ion channels, permeation and electrostatics: insight into the function of KcsA. Biochemistry 39:13295-13306 Saint N, El Hamel C, De E, Molle G 2000 Ion channel formation by N-terminal domain: a common feature of OprFs of P seudomonas and OmpA of Escherichia coli. FEMS Microbiol Lett 190:261-265 Sansom MS 1993 Structure and function of channel-forming peptaibols. Quart Rev Biophys 26:365-421 Sansom MSP, Kerr ID, Breed J, Sankararamakrishnan R 1996 Water in channel-like cavities: structure and dynamics. Biophys J 70:693-702 Sayle RA, Milner-White EJ 1995 RASMOL: biomolecular graphics for all. Trends Biochem Sci 20:374-376 Schirmer T 1998 General and specific porins from bacterial outer membranes. J Struct Biol 121:101-109 Shrivastava IH, Sansom MSP 2000 Simulations of ion permeation through a potassium channel: molecular dynamics of KcsA in a phospholipid bilayer. Biophys J 78:557-570 Smart OS, Goodfellow JM, Wallace BA 1993 The pore dimensions of gramicidin A. Biophys J 65:2455-2460 Smart OS, Breed J, Smith GR, Sansom MSP 1997 A novel method for structure-based prediction of ion channel conductance properties. Biophys J 72:1109-1126 Tieleman DP, Breed J, Berendsen HJC, Sansom MSP 1998 Alamethicin channels in a membrane: molecular dynamics simulations. Faraday Discuss 111:209-223 Tieleman DP, Berendsen HJC, Sansom MSP 1999 An alamethicin channel in a lipid bilayer:molecular dynamics simulations. Biophys J 76:1757-1769 Tieleman DP, Berendsen HJC, Sansom MSP 2000 Voltage-dependent insertion of alamethicin at phospholipid/water and octane/water interfaces. Biophys J 80:331-346 Tieleman DP, Biggin PC, Smith GR, Sansom MSP 2001 Simulation approaches to ion channel structure-function relationships. Quart Rev Biophys 34:473-561 Unwin N 2000 The Croonian Lecture 2000. Nicotinic acetylcholine receptor and the structural basis of fast synaptic transmission. Philos Trans R Soc Lond B Biol Sci 355:1813-1829 van Gunsteren WF, Kruger P, Billeter SR et al 1996 Biomolecular Simulation: The GROMOS96 Manual and User Guide. Biomos & Hochschulverlag AG an der ETH Zurich, Groningen & Zurich

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DISCUSSION Roux: A hydrophobic pore, made of completely non-polar side chains could be different. The backbone of proteins is quite polar. Sansom: I guess it depends on the exact geometry of the side chains. Roux: But the backbone is polar: water molecules inside cavities are almost never more than 3 or 4 A away from the backbone. Sixma: But in a helix it is not polar all along the helix. Roux: That doesn't matter. A water molecule inside a non-polar cavity in the interior of a protein will usually find a way to hydrogen bond on the side of a backbone carbonyl group of a helix. It doesn't perturb the helix structure very much to allow such a hydrogen bond (Roux et al 1996). Sansom: That is going to depend on how densely packed the rings of hydrophobia side chains are. I am trying to visualize Nigel Unwin's acetylcholine receptor structure. Unwin: Looking at it, I can't see how water could get close to those polar groups. There is too much protection from the hydrophobic side chains. Roux: How many layers of side chains are there? Sansom: Three: leucine, valine and another leucine. I guess the thing to do is to calculate a solvent-accessible surface. Roux: As an example, we have tried simulations of KcsA in the 'open' state. If you take KcsA and slightly open the inner helices at the bottom, there are residues 101, 106 and 109 making three layers of non-polar amino acids. You just have to push these helices by about 3 A and then you get this stable tube of water (Roux et al 2000). Sansom: I agree; it is going to be very sensitive to the exact structure. Hummer et al (2001) have been doing some simulations on nanotubes, and depending on how they tune the van der Waals interactions water either enters or leaves the pore. Perozo: If you are going to assign a gain you want it to be very sensitive. You want a polar barrier that is easily lowered so that ions and water can go through. Sansom: When one tries to build an open model of KcsA the gate is increased to c.4 A . Roux: I think it is a 3—4 A radius. Perozo: It depends where you look, because it is a scissor-like motion. The distance change decreases as you approach residue 107, which we think forms a 'pivot' for the gate. Roux: It may be 6 A diameter in the narrowest part. Sansom: That seems a bit narrow. Roux: Even at 6 A there is penetration of water. Unwin: What length? Roux: Three amino acids, which is about 10 A.

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Sansom: Even in the simple model calculations there is some sensitivity to the geometry at the mouth of the channel. Roux: Protein material itself has a dielectric constant of 4 or 5, whereas your system is a slab of non-polar spheres with a dielectric constant of 1. From an electrostatic point of view this is equivalent to vacuum. Sansom: No, it is not the slab of vacuum; it has non-polar atoms for the slab as well. Roux: If one thinks in term of the Born (1920) model of solvation, the free energy of stabilization of an ion depends sensitively on the dielectric constant of the surrounding. Already for a dielectric constant around 4, one gets a large stabilization free energy, nearly 75% of the free energy that one gets for a dielectric constant of 80. Mindell: I am intrigued by the stripes of order that you see. It is not clear to me, however, that this would have a dramatic effect on the diffusion coefficient. Does it affect the diffusion in those models? Sansom: We were a bit nervous about trying to calculate the diffusion coefficient when we can see something moving in a stepwise fashion, so we haven't done this. I think people have suggested such arrangements of water in a number of calculations, e.g. of water in idealized narrow tubes (Allen et al 1999) where water clusters form distorted pentameric rings. Choe: You mentioned that the waters that were placed in the middle of the core tend to drift away only in one direction. Sansom: There does seem to be some asymmetry. I'm not sure why this is. It may reflect an asymmetry in the exact pattern of hydrophobicity in the surface lining the pore. We haven't looked at this properly yet. We would like to compare this effect for aquaporin, for example, where there are subtle differences in the nature of the residues lining the pore. Choe: How much does the electric field contribute to this behaviour? Sansom: We haven't yet estimated the field strength. In principle we could cancel it out by using an asymmetric bilayer. I am fairly certain that there is not such a marked dipole across AQP1 as there is across GlpF. Schrempf: Taking the biological data into consideration, I am surprised you are not using glycerol for your simulations for GlpF. Sansom: We are. If one believes the data in the Borgnia et al (2001) paper (and I know there is some discussion about it), the argument is not that GlpF has massive selectivity against water, so much as AQP has a massive selectivity against glycerol. Schrempf: The biological data clearly indicate that this is a glycerol-transporting protein. If you make Escberichia coli mutants and disrupt the gene it can also be tested. Sansom: But it lets water through. It allows water through at near its diffusionlimited rate (Borgnia et al 2001).

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Mathie: You showed one water molecule diffusing right across the membrane and you said that it took about 2ns. The other ones that you showed here are staying in the pore much longer than that and are settling half way through the pore. Is this just random? Sansom: I think so. To make this a bit simpler to present, we have chosen four waters that spend the longest time in the pore. We see a large number of waters entering and coming back in. This happens to be the one we have seen, in the absence of any concentration gradient of water, that goes all the way across. At equilibrium you would expect a large number to enter and then to exit again and this is what we see. Mathie: If you were to change the osmolarity in your models you would presumably push water through. Sansom: That is non-trivial to do, but in principle, yes. Mindell: Why is it so hard to make an osmolar gradient? Can't you just exclude water from part of the volume? Sansom: We have a finite simulation box with periodic boundaries at the edges, so the computational trick used is that if a water wanders out at the bottom it reappears at the top. Putting any asymmetry across the system is a bit challenging. There are ways to do it. Roux: A possible approach is to apply an external perturbative force (not necessarily having the form of an electric field though). According to KuboZwanzig Linear Response Theory (Hansen & McDonald 1986), the equilibrium response function can be extracted using any small perturbative force. For example, one could apply a generic force inducing a flux of water across the membrane. Sansom: Or you could drag one or more water molecules through the pore. Mindell: What if you just fill one box with some volume-occupying spheres? This would reduce the net number of water molecules on that side. Roux: You would have to restrain them in space somehow, because they would diffuse. Or you could have two membranes and have the gradient between them. Sansom: It is similar to the problem of how to put a voltage across the membrane. Asbcroft: You showed single-channel currents for OmpA, but you didn't tell us which ion was carrying the current. You also talked about water moving through this channel, but you didn't say that your simulation predicted whether the water could move. Sansom: We haven't done that yet. As to which ion goes through, I don't think this was mentioned. The experiment was in KC1 (Arora et al 2000). Miller: A simple question. Water has a dipole and can hydrogen bond. How is this allowed for in the molecular dynamics calculations?

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Sansom: You have got partial charges on the water. This is just a three-point charge model, so there aren't any lone pairs to put on the oxygen. Miller: You have your rings of water which satisfy the hydrogen bonding requirement: why are they all parallel to the bilayer and not tilted? Sansom: I am still unsure about these. It is peculiar, and I wouldn't have shown it if other people hadn't seen it as well. It may just be an artefact, but it is a consistent one, and it fits in with the general scheme that even if you have water penetrating a narrow hydrophobic pore it may behave in a very different fashion from bulk water. Miller: Presumably you don't see hydrodynamic effects in something that small. Ashcroft: Can I go back to the toy model of the acetylcholine receptor pore? You said that at 3.5 A radius, if it was lined by hydrophobic residues, the pore was essentially closed. However, if these are replaced with charged residues the pore opens. The critical leucine has been mutated a lot in the ACh receptor. What do you see in your model if you substitute this leucine with other residues? Do you get the same effect as is found for the channel? Sansom: With this toy model it would be difficult to do exactly the same. Corringer: It doesn't open. If you mutate the leucine to a threonine, you don't get spontaneous opening. Unwin: It depends on how many mutations you do. Corringer: If you do all five you see a few percent of channels opening spontaneously, but most are closed. Unwin: If we are talking about water going through this narrow hole, that is different. You are talking about the channel opening via a conformational change. Corringer: What I am saying is that the closed conformation is maintained. Unwin: I would expect that. If you are changing a leucine you probably should change the valines on the next ring up at the same time. Corringer: I should do both together. We have a very complex phenotype that I don't understand yet. We have increments that we interpret as spontaneous opening, but perhaps we have impaired the gate. Sansom: How would you distinguish between a spontaneous opening and an impaired gate? Corringer: A spontaneous opening can be reversed using an antagonist. Perhaps we have an impaired gate. In the double mutant the phenotype is very complicated. Unwin: I don't think people have ever measured how much water goes through the acetylcholine receptor. It may be reasonably impermeable to water. Sansom: The coupling between water and ions has been measured using streaming potentials. This is shown in some older work by Dani & Levitt (1990). Choe: As I was looking at this graph I was thinking of the bottom half of the KcsA pore, which is quite hydrophobic in the narrow part. The calculation

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seems to suggest that the conformational change should be from 2 A to about 4-5 A. This is quite a significant change. Perozo: It is not a linear change. The helices move away at the base, so the distance change that we see is about 10 A at the base, and this narrows out to about 2 A in the narrowest portion. Sansom: So what you are saying is that this would expand to about 5 A radius. Choe: That is more than the diameter of the helix. References Allen TW, Kuyucak S, Chung SH 1999 The effect of hydrophobia and hydrophilic channel walls on the structure and diffusion of water and ions. J Chern Phys 111:7985-7999 Arora A, Rinehart D, Szabo G, Tamm LK 2000 Refolded outer membrane protein A of Escherichia coli forms ion channels with two conductance states in planar lipid bilayers. J BiolChem 275:1594-1600 Borgnia MJ, Agre P 2001 Reconstitution and functional comparison of purified GlpF and AqpZ, the glycerol and water channels from Escherichia coli. Proc Nat Acad Sci USA 98:2888-2893 Born M 1920 Volumen und Hydratationswarme der lonen. Z Phys 1:45-48 Dani JA, Levitt DG 1990 Diffusion and kinetic approaches to describe permeation in ionic channels. J Theor Biol 146:289-301 Hansen JP, McDonald IR 1986 Theory of simple liquids, 2nd edn. Academic Press, London Hummer G, Rasaiah JC, Noworyta JP 2001 Water conduction through the hydrophobic channel of a carbon nanotube. Nature 414:188-190 Roux B, Nina M, Pomes R, Smith JC 1996 Thermodynamic stability of water molecules in the bacteriorhodopsin proton channel: a molecular dynamics free energy perturbation study. Biophys J 71:670-681 Roux B, Berneche S, Im W 2000 Ion channels, permeation, and electrostatics: insight into the function of KcsA. Biochemistry 39:13295-13306

What can be deduced about the structure of Shaker from available data? Benoit Roux Weill Medical College of Cornell University, 1300 York An, New York, NY 10021, USA

Abstract. Voltage-gated K+ channels are transmembrane proteins that control and regulate the flow of K+ ions across cell membranes in response to changes in membrane potential and are essential for the propagation of action potentials in the nervous system. One of the most studied voltage-gated channels is Shaker. Available experimental results clearly provide specific constraints on the structure of the channel, even though the direct translation of the available information into 3D structures is not trivial. The goal of this work is to develop a computational approach to construct and refine 3D models of Shaker by incorporating and integrating available experimental data. Our approach is based on comparative modelization and global conformational optimization using energy restraints extracted from experimental data. 2002 Ion channels—from atomic resolution physiology to functional genomics. Wiley, Chichester (Novartis Foundation Symposium 245) p 84-108

The activity of voltage-gated ion channels is the basic molecular mechanism underlying the electrical excitability of nerves and muscles (Hodgkin & Huxley 1952). These channels are specialized transmembrane proteins, which control and regulate the flow of ions across cell membranes by opening and closing ('gating') in response to changes in membrane potential (Hille 1992). The first identified and best-studied voltage-gated channel is the Shaker K+ channel from the fruitfly Drosophilia melanogaster (Tempel et al 1987); the corresponding voltage-gated K+ (Kv) channels in mammals are KV1.1-KV1.7 (Jan & Jan 1997). Normally closed at hyperpolarized resting potentials, Shaker K+ channels undergo a conformational transition from a closed to an open state at depolarization potentials (Cha et al 1999, Glauneretall999). Studies have shown that Shaker and all the channels in the Kv family are structurally and functionally similar. They are formed by four identical or homologous domains, or subunits (MacKinnon 1991). Analysis of the amino acid sequence suggests that each subunit contains six putative transmembrane (TM) segments, S1 to S6 (Jan & Jan 1997, Tempel et al 1987). The second (S2) 84

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and fourth (S4) segments contain several charged residues, which are affected by changes in membrane potential and form part of the voltage sensor that controls the gating of the channel (Aggarwal & MacKinnon 1996, Bezanilla 2000, Liman et al 1991, Logothetis et al 1992, Papazian et al 1991, Seoh et al 1996, Yellen 1998). The part of the protein forming the pore region responsible for the selectivity and conduction of K+ ions is located between segments S5 and S6 of a subunit in the region containing the essential amino acid 'signature sequence' TTVGYGD common to all K+ channels (Heginbotham et al 1992,1994). Although a growing body of information is available for Shaker and other voltage-activated channels in the Kv family, the only ion channel for which a structure at atomic resolution is currently available is the KcsA channel from Streptomyces lividans (Doyle et al 1998). The main features of the crystallographic structure are shown in Fig. 1. The channel is made of four identical subunits disposed symmetrically around a common axis corresponding to the pore (only two are shown in Fig. 1). Although the monomer of KcsA is formed by only two transmembrane helices, the amino acid sequence is, in fact, very similar to that of segment S5—S6, which is conserved in eukaryotic voltage-gated channels such as Shaker (Cortes & Perozo 1997, Doyle et al 1998, Schrempf et al 1995). Furthermore, a combination of structural and functional data with neurotoxin from scorpion indicates the extracellular vestibule of KcsA is structurally very similar to Shaker (MacKinnon et al 1998). In the absence of a detailed atomic structure, extensive studies using a variety of experimental approaches including electrophysiology, site-directed mutagenesis, resonance energy transfer and electron microscopy, have been used to probe the structure and function of the Shaker K+ channel. Undoubtedly, many of the experimental results obtained so far put very specific constraints on the structure of Shaker, though often indirectly. Nonetheless, the direct translation of all the available information into a 3D structure is not straightforward. The purpose of the present work is to develop a computational approach to construct and refine 3D models of Shaker by incorporating and integrating all available experimental data. The approach that we use is based on comparative modelization and global conformational optimization using energy restraints extracted from experimental data. Given the limited amount of information presently available, we do not expect to converge towards a unique 'best' model of Shaker. Instead, we seek to delineate and clarify, as objectively as possible, the current state of the knowledge about Shaker by generating an ensemble of plausible models which are consistent with the available data. Our hope is that such an ensemble of 3D models can play a useful role in the design of future experiments by indicating the areas of greatest uncertainty in the structure, by helping to examine the spatial relationship between functionally important residues, and by revealing inconsistencies between different experimental results.

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FIG. 1. Schematic view of the KcsA channel (only two of the four monomers are shown). The extracellular side is at the top and the intracellular side is at the bottom. The main structural elements are: the outer helix corresponding to S5 in voltage-gated K+ channels (residues by A391 to E418); the P loop formed by the pore helix and the selectivity filter which contains the signature sequence TTVGYGD (residues 1429 to T441); and the inner helix corresponding to S6 (residues G452 to N482).

Assumptions

The general topology of a Shaker subunit is illustrated schematically in Fig. 2. In particular, it is assumed that the segments S1-S4 are in an a-helical conformation and that the central pore formed by S5-P-S6 is structurally very similar to the crystallographic structure of the KcsA K+ channel. Although these assumptions are reasonable and currently supported by experimental evidence, some of them might turn out to be incorrect in the future. Nonetheless, such simplifications are necessary at this point.

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FIG. 2. Schematic view of one subunit of Shaker with its six transmembrane segments. It is assumed that S1 to S4 are in an a-helical conformation and that the central pore formed by S5—P-S6 is structurally similar to the crystallographic structure of the KcsA K+ channel shown in Fig. 1.

Helical conformation of the transmembrane segments S1-S4 It is assumed that the TM segments S1—S4, S1 (226-247): ARVVAIISVFVILLSIVIFCLE S2 (279-300): FFLIETLCIIWFTFELTVRFLA S3 (311-332): VMNVIDIIAIIPYFITLATVVA S4 (358-380): LAILRVIRLVRVFRIFKLSRHSK are in an a-helical conformation and are roughly perpendicular to the membrane plane, though some tilting of the helix axis is possible. This hypothesis is supported by a number of observations. Experimental studies have established that an isolated fragment corresponding to S4 adopts a predominantly a-helical conformation in methanol and in lipid membranes (Halsall & Dempsey 1999, Haris et al 1994, Mulvey et al 1989). The structure is a random coil in aqueous solution (Haris et al 1994). Similar studies with TM segments of the Na+ channel have shown that they adopt a-helical structures in detergent micelles (Doak et al 1996). There are also strong indications that the SI to S4 segments are a helical in the channel structure. Ala- (Li-Smerin et al 2000a) and Trp-scanning (Hong & Miller 2000, Monks et al 1999) mutagenesis studies suggest that S1 and S2 are amphipathic membrane spanning a helices that

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interface directly with the lipid membrane. Helical periodicity of functional alteration in the voltage-activation curves and gating kinetics were observed throughout S1 and S2. Trp-tolerant positions in the Shaker K+ channel are clustered on approximately half the a helix surface, as if the side chains are exposed to the hydrocarbon region of the lipid bilayer (Hong & Miller 2000, Monks et all 999). Similarly, Ala-scanning mutagenesis in the drkl K+ channel of S1 and S2 suggest that these segments are relatively simple amphipathic helices that span the full width of the membrane and make extensive contacts with the lipid membrane (Li-Smerin et al 2000a). The observations for the S3 and S4 segments are more complex. In the case of S3, the distribution of Trp-tolerant position is roughly consistent with a helical secondary structure, although the results are not as clear towards the extracellular side (Hong & Miller 2000). Results from Ala(Li-Smerin et al 2000a) and Lys-scanning (Li-Smerin & Swartz 2001) with the drkl K+ channel suggest that the S3 segment is entirely helical, but that the N-terminal region interfaces with both lipid and protein, whereas the C-terminal region interfaces with water. It has been speculated that a conserved proline at position 322 might induce a kink in the helical segment. Ala scanning of S4 reveals helical periodicity in only the C-terminal region (Li-Smerin et al 2000a). However, it seems likely that the absence of helical character in the N-terminal portion, which is exposed to the intracellular side, results from complexities in the aqueous and protein environment surrounding the segment. Structure of the centralpore S5-P-S6

It is assumed that the conformation of the central pore of Shaker, formed by the S5-P-S6 segment, is very similar to the crystallographic structure of KcsA (Doyle et al 1998). This is a very reasonable assumption given the high sequence similarity of the core of Shaker and the KcsA bacterial channel (Fig. 3). For 93 residues, the sequence identity is 31 for a global sequence similarity of 49%. This high similarity makes Shaker an excellent candidate for successful comparative modelling using the KcsA structure as a template (Fiser et al 2000). The structural similarity of Shaker relative to KcsA is also supported by experiments: AgTx2 binds to the KcsA channel, demonstrating that this prokaryotic K+ channel has the same pore structure as that of Shaker (MacKinnon et al 1998), and the chemical modifications of cysteines located along S6 by soluble thiol agents are generally consistent with the accessibility of the corresponding residues in KcsA (Liu et al 1997). Because the KcsA K+ channel is in a closed conformation (Roux et al 2000), the crystallographic structure is probably a better model for the closed state of Shaker. Nonetheless, although it appears to be very reasonable, the assumption that

THE STRUCTURE OF SHAKER

FIG. 3.

89

Sequence similarity between the KcsA and Shaker channels.

the S5—P—S6 core of Shaker is structurally similar to KcsA can be wrong for some regions of the structure. In particular, the sequence similarity is much smaller near the end of S6, which may indicate that the structure of this region differs from that of KcsA. In particular, S6 contains the motif Pro-ValPro at position 473-474-475, which is known to perturb a helices considerably (Barlow & Thornton 1988, von Heijne 1991). In addition, recent results from blocker protection in the pore of a voltage-gated K+ channel suggests that the structure of Kv channels differs from that of the KcsA by the introduction of a sharp bend in the S6 helices (del Camino et al 2000). Experimental data Intragenic suppression

Using an intragenic suppression strategy, Tiwari-Woodruff et al (1997) showed that charge reversal mutations of E283 in S2 and K374 in S4 disrupt maturation of the protein. Maturation was specifically and efficiently rescued by second-site charge reversal mutations, indicating that electrostatic interactions exist between E283 in S2 and R368 and R371 in S4, and between K374 in S4, E293 in S2, and D316 in S3. Further investigation of those mutants showed that the interaction between K374 in S4 and E293 in S3 and D316 in S3 is important for the closed state whereas the interaction between E283 in S2 and R368 and R371 in S4 is probably important for the open state (Tiwari-Woodruff et al 2000). Resonance energy transfer

Using lanthanide-based resonance energy transfer (LRET) to measure distances between Shaker K+ channel subunits at specific residues, Cha et al (1999) determined the distance between site-specific labelled monomers. Specific sites in the channel were fluorescently labelled by substituting cysteine for particular residues and attaching a cysteine-reactive compound of either a donor (a terbium-chelate maleimide, TbM) or an acceptor (fluorescein maleimide, FM). The intersubunit distances were evaluated using the relaxation time constant of acceptor-sensitized emission and donor emission without an acceptor. The

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distances estimated with this method are accurate because the lanthanide donor is in fast orientational averaging. For example, the distance measured across the pore at F425 is in excellent accord with the distance between the carbon a for the corresponding residue in the X-ray structure of KcsA. Furthermore, it was shown that the distances deduced independently from the energy transfer between the nearest and opposite subunits in the tetramer were consistent with Pythagoras' theorem. The most important distances measured between two monomers on opposite sides of the tetramer were 46 A for D273, which is located near the extracellular end of S2, 41 A for N353 near the extracellular end of S4, and 45 A for V363 in the middle of S4. Tethered pore blackers

Blaustein et al (2000) have used a clever method based on a series of compounds of varying length ending with a quaternary ammonium (QA) pore blocker. To locate the S1—S4 segments relative to the central pore these compounds were tethered to specific test residues using site-directed cysteine mutations. By examining the probability of blockade as a function of the length of the compounds, it was shown that the extracellular ends of S1 and S3 are approximately 30 A from the external opening of the pore while the S3-S4 linker is at 17-18 A from the pore. Scanning mutagenesis

Miller and co-workers have performed site-directed substitution of all residues in S1, S2 and S3, probing for significant functional changes (Hong & Miller 2000, Monks et al 1999). The function of the channel was significantly affected after substitution of Trp at the positions: A226, R227, V229, A230, S233, V234,1237, L238, S240,1241,1243, F244 and C245 for S1; E283, C286,1287, F290, E293, L294, R297 and A300 for S2; and M312, N313, 1315, D316, A319, 1320, P322, Y323, F324, L327, V330 and V331 for S3. The function of the channel was not affected by similar substitutions at other positions. Helical periodicity of functional alteration in the voltage activation curves and gating kinetics were observed throughout S1 and S2. A similar result was obtained with S3, although the distribution of Trp-tolerant position is not as clear towards the extracellular side. The Trp-tolerant positions are clustered roughly on one face of the putative a-helical segments, as if those side chains were exposed to the hydrocarbon region of the lipid bilayer. The results are consistent with Ala scanning mutagenesis studies on a related Kv channel (Li-Smerin et al 2000a, Li-Smerin & Swartz 2001). Interestingly, comparison of the amino acid sequence of several channels in the Kv family showed that the Trp-tolerant sites exhibit a high level of sequence variability, whereas the Trp-intolerant sites are highly conserved. This

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observation further reinforces the suggestion that Trp-tolerant residues are exposed to the lipid hydrocarbons, while the Trp-intolerant residues are involved in protein—protein contacts. Single-particle electron microscopy

Recently Sokolova et al (2001) obtained the 3D structure of the complete Shaker channel at 25 A resolution using single-particle electron microscopy. The 3D map clearly shows that the Shaker channel has a two-domain architecture with a transmembrane domain, corresponding to S1—S6, and a soluble domain corresponding to the Tl domain attached to the transmembrane domain by thin connectors. This two-domain architecture is consistent with the 'hanging gondola' model proposed by Kobertz et al (2000). Although the image is still at very low resolution, it shows that the transmembrane section of the channel has the shape of a compact square with sides of roughly 70 A. Modelization procedure The TM segments S1-S4 The S1—S4 segments were constructed as ideal a helices and then further refined by energy minimization. A rather conservative definition of helical segments S1 to S3, with only 21—22 residues, was used in the modelization to avoid 'over' constraining the system with long a-helical TM segments. In the case of S4, the helical segment was constructed as an a helix from residue 350 to 380 based on the mutagenesis results of Gonzalez et al (2000). Planar energy restraints were applied to keep the ends of the TM segments outside a 30 A thick hydrophobic slab and maintain the orientation of the helical segments. It is implemented as a half-harmonic potential with a force constant of 10kcal/mol-A 2 . This energy restraint keeps the helices roughly perpendicular to the membrane while allowing some tilting of the helices, in agreement with the orientation of TM helices in membrane proteins of known structure (Bowie 1997). For the sake of simplicity, the S1-S2 (31 residues), S2—S3 (10 residues), S3-S4 (17 residues), and S4— S5 (10 residues) linkers between the TM segments were not included explicitly. Instead, a simple half-quadratic restraint of 10 kcal/mol-A2 was used to limit the maximum distance between the C- and N-terminals of the ends of adjacent TM segments (S1—S2, S2— S3, S3—S4, and S4— S5). A maximum distance of 25 A was used for the S1—S2 linker while a distance of 10 A used for the other loops. Experimental studies have shown that the S3—S4 linker can be shortened from 17 to the five amino acids closest to S4 without disrupting the function and the gating of the channel (Sorensen et al 2000), indicating that the C- and N-terminals of the helices are probably located close to each other.

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The central pore

The 3D model of the central pore of Shaker was constructed on the basis of its sequence similarity with KcsA for which there is a crystallographic structure. The program MODELLLER v4 (Sali & Blundell 1993) was used to produce 3D homology-based models by satisfaction of spatial restraints. Its ability to produce high quality 3D atomic structures based on sequence similarity has been evaluated (Sali et al 1995). The segment S6 was modelled as a straight helix, even though there are indications of a sharp bend (del Camino et al 2000). The ideal fourfold symmetry of the tetrameric channel model was imposed with energy restraints. Fifty 3D models were generated. The best model was kept and then further refined with energy minimization of the side chains. Distance energy restraints

Several pieces of information can be translated as constraints on the range of allowed distances between different parts of the structure. This information is incorporated into the model as artificial distance-energy restraints which take the form of a quadratic function. All the restraints that were used are given in Table 1 . Energy restraints corresponding to the salt bridge between E293 in S2, D316 in S3 and K374 in S4 were applied (Tiwari- Woodruff et al 1997, 2000). A maximum distance of 3 A between the side chain atoms having the ability to form the salt bridge was applied with a harmonic force constant of 20kcal/mol-A2. Restraints corresponding to the salt bridge between E283 in S2 and R368 and R371 in S4 were also included. Because those salt bridges formed only in the open state (TiwariWoodruff et al 2000), weaker restraints with a force constant of 5 kcal/mol-A 2 and allowing for a maximum distance of 6 A were used. The LRET restraints are all applied between subunits A and D, and between subunits B and C. Because the linker residues between the TM segments are not included for the sake of simplicity, the LRET restraint measured for D273 which is part of the S1-S2 linker was transferred to F279 at the end of S2. Lipid-exposed residues

A directional energy restraint was applied to force the orientation of S1— S3 such that the Trp-tolerant residue points away from the core of the structure towards the lipid hydrocarbon region. The energy restraint takes the form of a pseudo dihedral angle, E expo (f) = Kexpocos(f-180) where f is the angle between the centre of mass of the lipid-exposed residues and the protein-exposed residues projected onto the XY plane. The restraint was

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93

Distance energy restraints

Inter-segment loops (for all submits) E247 CA

F247 CA

<25A

A300 CA A332 CA

V311 CA E358 CA

K380 CA

A391 CA

<10A <10A <10A

RE T measurements (all between submit A andD, and between submit B and C) V363 CB

V363 CB

F279 CB

F279 CB

Within 44-46 A

N353 CB

R371 CB

Within 36-40 A

Within 44-46 A

Salt bridge (distance takenfrom the nearest pair of any subunit) E293 OE*

K374NZ

<3A

D3160D

K374 NZ

<3A

E283 OE*

R368 NH*

E283 OE*

R371 NH*

<6 A <6 A

Planar restraints on ends of helices (for all subunits) A226 all atoms

z

< -15 A

E247 all atoms

z

> +15 A

F279 all atoms

z

>+15A z<-15A

A300 all atoms V311 all atoms

z

A332 all atoms

z

< - 15 A > +15 A

L358 all atoms

z>+15A

K380 all atoms

z<-15A

applied to all side chain atoms of the buried and exposed residues to ensure proper orientation with respect to the core structure S5—S6. The force constant of the pseudo-dihedral energy restraint Kexpo was 10 kcal/mol/rad2. An energy restraint based on a rigorous calculation of the solvent-accessible surface area (SASA) could be used in the later stage of the structural refinement. However, the calculation of the exact SASA (Richards 1977) is computationally much more expensive than the approximation used here, and is unnecessary at this stage.

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Conformational sampling and simulated annealing procedure

Initially, S1-S4 are disposed around the central pore in arbitrary positions and orientations to insure a thorough exploration of all conformational possibilities. To obtain a starting position, each TM segment is rotated around its axis using a randomly picked angle. The orientation of S1-S3 is chosen within a 10° window centred approximately to direct the Trp-tolerant residues away from the core of the structure whereas the orientation of S4 is completely randomized over 360o. The initial position of S1—S4 is then chosen randomly within a distance of 20-55 A away from the axis of the pore (with S4 between 20-25 A, S3 between 30-35 A, S2 between 40-45 A, and S1 between 50-55 A) as well as distributed within a 180 quadrant centred on their respective monomer. The random initial models were refined with cycles of molecular dynamics (MD) simulations at 600 K during 100 ps. To maintain constant temperature, the systems were simulated with Langevin dynamics with a friction constant corresponding to a velocity relation time of 1 ps - 1 . Electrostatic interactions were not included and van der Waals interactions were truncated at 5 A to reduce the computational cost. To maintain the helical conformation of the S1 to S4 segments an energy restraint based on the root-mean-square deviation was applied on all the backbone atoms (RMSD restraint). The backbone of the central core S5-P-S6 was fixed in space, but the side chains were allowed to move dynamically. The fourfold symmetry of the tetrameric channel was maintained during the simulated annealing procedure using energy restraints. To reduce the steric clashes and accelerate the convergence, the atoms were allowed to move along an unphysical 4th dimension w, i.e. the interatomic distances were calculated as r= x2 + y2 + z2 + w2 (van Schaik et al 1993). A harmonic restraint potential with the form 1/2K4Dw2 was used to progressively project the coordinates back onto the physical three-dimensional space. Lastly, the configuration was refined in 3D for l0ps at a reduced temperature of 300 K followed by 200 steps of energy minimization. Generating and refining one Shaker model took approximately 2 h on a Pentium III 850 MHz.

Results and discussion A general pattern rapidly emerged from preliminary conformational searches. In particular, it was observed that the S4 segment was localized near the groove between adjacent subunits to satisfy the distance of 45 A between V363 across the tetramer obtained by LRET (Cha et al 1999). The LRET restraints distance on F279 and N353 also contributed to this trend. In addition, it was observed that S2 and S3 were generally adjacent to one another, covering S4, and that the position of S1 relative to the rest of the structure was largely undetermined due to the long S1-S2 linker and the lack of distance constraints. The models have

THE STRUCTURE OF SHAKER

FIG. 4. search.

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Example of the random initial configurations of S1 to S4 used in the conformational

extended structures, and S1 is often quite distant from the rest, which is most certainly incorrect. According to a low-resolution electron microscopy map (Sokolova et al 2001), Shaker should have a compact square structure of roughly 70 A on the side. To form a compact structure, S1 should be in proximity to the other segments S2— S4. Furthermore, all the charged residues of S4 cannot be shielded from the lipid environment by only S2 and S3. Because it is unlikely that charged residues would be exposed directly to the hydrocarbon of the membrane, S1 must also be involved in shielding the charges of S4. Lastly, although the interhelical linkers are sufficiently long to allow a 'scrambling' of the TM segments from adjacent subunits, surveys of membrane proteins show that TM helices generally pack against neighbours in the sequence (Bowie 1997). In order to generate a family of plausible models, we introduced one artificial restraint imposing that the centre of S1 is no more than 10 A away from the centre of S4. On the basis of these preliminary results, 100 models were generated according to three plausible topologies shown in Fig. 4. The initial position of S4 was chosen within a +10° window centred along the groove between adjacent subunits of the

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FIG. 5. search.

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Example of the structural models of Fig. 4 after refinement with the conformational

central core to accelerate the conformational search. The position of S1-S3 is designed to cover S4 from the lipid hydrocarbon: S1-S2-S3, S2-S3-S1, S3-S2Sl and S1-S3-S2. The 100 models were refined with a simulated annealing procedure. One typical model of each type is shown in shown in Fig. 5. The 100 models are all equally plausible on the grounds of available information. Clearly, the spatial arrangement of S1—S3 cannot be determined without further information from further experiments. In the case of the S4 segment, the situation is slightly better and some general trends are emerging. Fig. 6 shows a superposition of the S4 segments from all 100 models with the central pore. The overall root-mean-square deviation on the backbone of all S4 is around 4—5 A. Although the orientation of the helix along its main axis is undefined at such resolution, the overall position of S4 is more or less determined by the modelization. In order to propose useful directions in mutagenesis studies of Shaker, we searched for pairs of residues between S4 and S5-P-S6 which are in frequent

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FIG. 6.

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Superposition of the S4 segments from the 100 models with the central pore.

contact in the 100 generated models (a contact is defined as an interatomic distance less than 5 A). The results are given in Table 2. More contacts were observed near the extracellular side than on the intracellular side. This is, in part, due to the conical (inverted tepee) shape of the pore domain. This pattern of contact suggests that experiments searching for correlated interactions between S4 and the central core might have more success by focusing on the region near the extracellular side. Remarkably, several of the residues observed to be in contact with S4 were previously noted in the mutagenesis study of Li-Smerin et al (2000b). In their study, they examined the residue at the 'surface' of the central core S5--P-S6 constructed according to the KcsA crystallographic structure. The mutants producing large changes in gating were found to cluster near the interface between adjacent pore domain subunits. It was concluded that the voltage sensor machinery must be in contact with this region of the central pore. This is consistent with the present finding that S4 is located in this region. The positively charged residues in S4 (R362, R365, R368, and R371) are separated by two residues and would not all face the same side of an a helix. This suggests that some arginine residues must face the central core formed by S5-P—S6. Interestingly, a large number of possible contacts between S4 and aromatic side chains are observed in Table 2, such as R362 with F433, W434 and W454. It is known that positively charged side chains (Arg, Lys, His) can associate very favourably with an aromatic side chains (Phe, Tyr or Trp) (Gallivan & Dougherty 1999). The interaction between isolated arginine and phenylalanine side chains (without the backbone) is about —10 kcal/mol, a value similar to that of simple cations with benzene in the gas phase (Sunner et al 1981). Interestingly, mutation of F433 significantly affects gating (Li-Smerin et al 2000b). One may also

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TABLE 2 Residue contacts between S4 and S5-P-S6 from the Shaker models

S5 and S6

S4

L358 A359 I360 L361

P430 P430

D431

F433 F433 W434

W434 F453

F453 W454

W454 I457

K456

I457

W434 F453

F453 W454

W454 I457

K456

I457

I457

V458

S460

L461

S460 L461 L461 L461 F404

W454 L461 L461 L461 L461 I464 I464 I464 I464

F373 L375 R377

P430 F433 F433 F433 1457 F404 F404 F404 F404 F404 I400 F404 L468 L396 M393

R394

G397

H378 S379 K380

M393 M393 M393

N480 E395

N480

R362 V363 I364 R365 L366 V367 R368 V369 F370 R371

W434 I457 S460 I457

I464

note that R362 is the only position in S4 which, upon substitution by a histidine, transforms the voltage sensor into a proton channel when Shaker is in its closed state (F. Bezanilla, personal communication). Other important aromatic residues in the interface between adjacent subunits could be F404, W434, F453 and W454. Substitution of W434 by a phenylalanine transforms the channel into a nonconducting mutant (Perozo et al 1993). Because of the uncertainty in the 3D models, any of the residues R365, R368 and R371 could be in proximity of F404. Summary We have developed a general protocol based on a restrained molecular dynamics conformational search procedure for incorporating various kind of experimental information in the modelization of the Shaker K+ channel. We have generated 100 atomic models of Shaker in the closed inactivated state using the currently available data. The main result of the restrained conformational search is that S4 is probably

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located at the interface between adjacent subunits of the central pore. This is largely consistent with the mutagenesis results of Li-Smerin et al (2000b). Specific contacts between S4 and S5-P-S6 were found using the models generated by the conformational search which could be used to design future mutagenesis experiments. It is hoped that the present approach will help better define the 3D structure of Shaker as more experimental information becomes available. References Aggarwal SK, MacKinnon R 1996 Contribution of the S4 segment to gating charge in the Shaker K+ channel. Neuron 16:1169-1177 Barlow DJ, Thornton JM 1988 Helix geometry in proteins. J Mol Biol 201:601-619 Bezanilla F 2000 The voltage sensor in voltage-dependent ion channels. Physiol Rev 80:555-592 Blaustein RO, Cole PA, Williams C, Miller C 2000 Tethered blockers as molecular 'tape measures' for a voltage-gated K+ channel. Nat Struct Biol 7:309-311 Bowie JU 1997 Helix packing in membrane proteins. J Mol Biol 272:780-789 Cha A, Snyder GE, Selvin PR, Bezanilla F 1999 Atomic scale movement of the voltage-sensing region in a potassium channel measured via spectroscopy. Nature 402:809-813 Cortes DM, Perozo E 1997 Structural dynamics of the Streptomyces lividans K+ channel (SKC1): oligomeric stochiometry and stability. Biochemistry 36:10343—10352 del Camino D, Holmgren M, Liu Y, Yellen G 2000 Blocker protection in the pore of a voltagegated K+ channel and its structural implications. Nature 403:321-325 Doak DG, Mulvey D, Kawaguchi K, Villalain J, Campbell ID 1996 Structural studies of synthetic peptides dissected from the voltage-gated sodium channel. J Mol Biol 258:672-687 Doyle DA, Morais Cabral J, Pfuetzner RA et al 1998 The structure of the potassium channel: molecular basis of K+ conduction and selectivity. Science 280:69-77 Fiser A, Sanchez R, Melo F, Sali A 2000 Comparative protein structure modeling. In: MacKerrel AD, Roux B, Watanabe M (eds) Biological membranes: a molecular perspective from computation and experiment. Marcel Dekker, New York p 275—312 Gallivan JP, Dougherty DA 1999 Cation-pi interactions in structural biology. Proc Natl Acad Sci USA 96:9459-9464 Glauner KS, Mannuzzu LM, Gandhi CS, Isacoff EY 1999 Spectroscopic mapping of voltage sensor movement in the Shaker potassium channel. Nature 402:813-817 Gonzalez C, Rosenman E, Bezanilla F, Alvarez O, Latorre R 2000 Modulation of the Sbaker K+ channel gating kinetics by the S3-S4 linker. J Gen Physiol 115:193-208 Halsall A, Dempsey CE 1999 Intrinsic helical propensities and stable secondary structure in a membrane-bound fragment (S4) of the shaker potassium channel. J Mol Biol 293:901-915 Haris PI, Ramesh B, Brazier S, Chapman D 1994 The conformational analysis of a synthetic S4 peptide corresponding to a voltage-gated potassium ion channel protein. FEBS Lett 349:371-374 Heginbotham L, Abramson T, MacKinnon R 1992 A functional connection between the pores of distantly related ion channels as revealed by mutant K+ channels. Science 258:1152-1155 Heginbotham L, Lu Z, Abramson T, Mackinnon R 1994 Mutations in the K+ channel signature sequence. Biophys J 66:1061-1067 Hille B 1992 Ionic Channels of Excitable Membranes, 2nd edn. Sinauer, Sunderland, MA Hodgkin AL, Huxley AF 1952 A quantitative description of membrane current and its application to conduction and excitation of nerve. J Physiol 117:500—544 Hong KH, Miller C 2000 The lipid-protein interface of a Shaker K+ channel. J Gen Physio! 115:51-58

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Jan LY, Jan YN 1997 Cloned potassium channels from eukaryotes and prokaryotes. Annu Rev Neurosci 20:91-123 Kobertz WR, Williams C, Miller C 2000 Hanging gondola structure of the Tl domain in a voltage-gated K+ channel. Biochemistry 39:10347-10352 Li-Smerin Y, Swartz KJ 2001 Helical structure of the COOH terminus of S3 and its contribution to the gating modifier toxin receptor in voltage-gated ion channels. ] Gen Physiol 117:205-218 Li-Smerin Y, Hackos DH, Swartz KJ 2000a Alpha-helical structural elements within the voltage-sensing domains of a K* channel. J Gen Physiol 115:33-50 Li-Smerin Y, Hackos DH, Swartz KJ 2000b A localized interaction surface for voltage-sensing domains on the pore domain of a K+ channel. Neuron 25:411-423 Liman ER, Hess P, Weaver F, Koren G 1991 Voltage-sensing residues in the S4 region of a mammalian K+ channel. Nature 353:752-756 Liu Y, Holmgren M, Jurman ME, Yellen G 1997 Gated access to the pore of a voltagedependent K+ channel. Neuron 19:175-184 Logothetis DE, Movahedi S, Satler C, Lindpaintner K, Nadal-Ginard B 1992 Incremental reductions of positive charge within the S4 region of a voltage-gated K+ channel result in corresponding decreases in gating charge. Neuron 8:531-540 MacKinnon R 1991 Determination of the subunit stoichiometry of a voltage-activated potassium channel. Nature 350:232-235 MacKinnon R, Cohen SL, Kuo A, Lee A, Chait BT 1998 Structural conservation in prokaryotic and eukariotic potassium channels. Science 280:106-109 Monks SA, Needleman DJ, Miller C 1999 Helical structure and packing orientation of the S2 segment in the Shaker K+ channel. J Gen Physiol 113:415-423 Mulvey D, King GF, Cooke RM, Doak DG, Harvey TS, Campbell ID 1989 High resolution 1H NMR study of the solution structure of the S4 segment of the sodium channel protein. FEBS Lett 257:113-117 Papazian DM, Timpe LC, Jan YN, Jan LY 1991 Alteration of voltage-dependence of Shaker potassium channel by mutations in the S4 sequence. Nature 349:305-310 Perozo E, MacKinnon R, Bezanilla F, Stefani E 1993 Gating currents from a nonconducting mutant reveal open-closed conformations in Shaker K+ channels. Neuron 11:353-358 Richards FM 1977 Areas, volumes, packing and protein structure. Annu Rev Biophys Bioeng 6:151-176 Roux B, Berneche S, Im W 2000 Ion channels, permeation and electrostatics: insight into the function of KcsA. Biochemistry 39:13295-13306 Sali A, Blundell TL 1993 Comparative protein modelling by satisfaction of spatial restraints. JMolBiol 234:779-815 Sali A, Potterton L, Yuan F, van Vlijmen H, Karplus M 1995 Evaluation of comparative protein modeling by MODELLER. Proteins 23:318-326 Schrempf H, Schmidt O, Kummerlen R et al 1995 A prokaryotic potassium ion channel with two predicted transmembrane segments from streptomyces lividans. EMBO J 14: 5170-5178 Seoh SA, Sigg D, Papazian DM, Bezanilla F 1996 Voltage-sensing residues in the S2 and S4 segments of the Shaker K+ channel. Neuron 16:1159-1167 Sokolova O, Kolmakova-Partensky L, Grigorieff N 2001 Three-dimensional structure of a voltage-gated potassium channel at 2.5 nm resolution. Structure 9:215-220 Sorensen JB, Cha A, Latorre R, Rosenman E, Bezanilla F 2000 Deletion of the S3-S4 linker in the Shaker potassium channel reveals two quenching groups near the outside of S4. J Gen Physiol 115:209-222 Sunner J, Nishizawa K, Kebarle P 1981 Ion-solvent molecule interactions in the gas phase. The potassium ion and benzene. J Phys Chem 85:1814-1820

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Tempel BL, Papazian DM, Schwarz TL, Jan YN, Jan LY 1987 Sequence of a probable potassium channel component encoded at Shaker locus of Drosophila. Science 237:770-775 Tiwari-Woodruff SK, Schulteis CT, Mock AF, Papazian DM 1997 Electrostatic interactions between transmembrane segments mediate folding of Shaker K+ channel subunits. Biophys J 72:1489 1500 Tiwari-Woodruff SK, Lin MA, Schulteis CT, Papazian DM 2000 Voltage-dependent structural interactions in the Shaker K+ channel. J Gen Physiol 115:123-138 van Schaik RC, Berendsen HJC, Torda AE, van Gunsteren WF 1993 A structure refinement method based on molecular dynamics in four spatial dimensions. J Mol Biol 234:751-762 von Heijne G 1991 Proline kinks in transmembrane alpha-helices. J Mol Biol 218:499-503 Yellen G 1998 The moving parts of voltage-gated ion channels. Q Rev Biophys 31:239-295

DISCUSSION Swartz: In terms of the location of S4 you propose, it seems that you are putting a tremendous amount of weight on the LRET (luminescence resonance energy transfer) distances from Bezanilla's group. The LRET distances they have from labelling a range of locations are all 45 A. The numbers were all the same whether they tagged the outer part of S2, the outer part of S3 or the outer part of S4. The question is, do these distances really constrain the location of S4? Roux: Cha et al (1999) reported several distances, ranging from 26-46 A. The measurements were done using LRET with lanthanides as donors and organic dyes as acceptors. The method yields distance estimates that are quite accurate because the influence of the anisotropy factors, normally important in conventional FRET (fluorescence energy transfer), are greatly reduced in lanthanide-based LRET. In fact, the main source of uncertainty in the measurements is the variability of the fluorophore's position with respect to the labelled residue because of the length of the cysteine-maleimide linker. The measurements are validated in the following way: by design, the channels that generate sensitized emission have three terbium-labelled subunits and one fluorescein-labelled subunit. The donor and acceptor can be between subunits that are either on opposite sides of the tetramer or are near neighbours, yielding two possible distances obeying the Pythagorean equation, and correspondingly, two possible decay rates for the energy transfer. Independent fitting of the decay rates shows that the two possible donor—acceptor distances obey the Pythagorean relation, which demonstrates the precision of the measured distances. For example, a distance of 30 A was measured across the tetramer between subunits labelled at Phe425, in excellent accord with the distance of 29 A between the corresponding residues (Gln58) in the KcsA crystallographic structure. Swartz: My initial reaction was that we all know it has to be 45 A or greater, on the basis of the structure of KcsA. Roux: Yes, if we measured S4 with a distance of say 39 A, we would feel this is too close.

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I should have mentioned that with the tethered quaternary ammonium of Blaustein et al (2000), the top of S4 was labelled. The estimated distances are highly consistent. In the paper they said that the loop of S4 is probably leaning on top of the core, as if there is no groove. But such a distance is possible if S4 lies in the groove. Papazian: There is now an additional constraint between S2 and S3 that comes from a different channel, ether-a-go-go (eag). eag has a metal ion binding site formed by two acidic residues, one in S2 and one in S3 (Silverman et al 2000). This constraint should be very helpful for modelling the Shaker channel. Ken Swartz' lab have put forward a convincing argument that the voltage sensor must pack in the groove. When he plotted his data from perturbation analysis of the Shaker pore domain onto the KcsA structure, the highest impact residues mapped right into that stripe between the subunits. Roux: I looked very carefully at this paper, and you did not conclude with certainty that S4 was there. Swartz: That's correct. I don't think we know where S4 is located. S3 could be there. If I interpret what you are saying correctly, you are betting even-thing on LRET distances. I am not quite convinced that this is such a solid constraint. Roux: S5/S6 looks a bit like a clover-leaf with the four subunits. If you start to build S4 anywhere but in the groove, there is no way that you can shield such a polar helix with S1, S2 and S3. The only way to do this is to put S4 in the groove with the three other helices on top, covering it. Swartz: Yes, but S3 has a lot of complexities to it also. What is unique about S1 and S2 is that they have really well defined faces where you can make radical substitutions to hydrophobic residues, and they are well tolerated. Both S3 and S4 have much more complex patterns, as if they are more buried. I believe S1 and S2 are located in the periphery of the complex. If you think S4 is a helix, you mutate every residue to alanine, and look at the energetic perturbation in gating, there are significant patches where those mutations are very well tolerated. So S4 could also have some lipid-exposed surfaces. Roux: It could be exposed to an aqueous pore. There is quite a bit of space around S4, and this is because of the tepee shape of the KcsA-like S5-S6 (Doyle et al 1998). On the extracellular side it is packed tight, but on the intracellular side there is quite a bit of space around S4. If you mutate amino acids to alanine there, you may find that they are tolerated. This doesn't imply that they are exposed to the lipid. Mindell: Rod MacKinnon's work on cysteine accessibility in S4 (Aggarwal & MacKinnon 1996) and Pancho Bezanilla's histidine mutagenesis (Starace & Bezanilla 2001) also suggest that water is getting quite a significant way up or down S4. Sansom: It might be interesting to step back and think a bit about the methodology here. One of the problems in trying to think about future

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experiments is the ratio of observations to parameters. Essentially, there are a lot of restraint data from different experimental approaches. We want to know how many restraints we need in order to hit the right target. Comparable situations that have been explored in the past, where we know what the target is, are bacteriorhodopsin and rhodopsin (Herzyk & Hubbard 1995). Could one take a cluster of three helices from either of these structures and then let the other four pack in around them? Roux: That's a good idea. One could take the bacteriorhodopsin structure and just come up with a small number of restraints. Sansom: I think that for rhodopsin, for example, the sort of data already exist comparable to the soft data that the physiology generates. You could see how many data you would have to include to get within a certain root-mean-square deviation of the known crystal structure. Roux: You could simulate the entire thing: you could take the bacteriorhodopsin structure, find which residues are exposed to the lipid, and then you could have some distance restraints between a few amino acids. You could even insert some wrong distances, just to see how robust it is. Sansom: This is going to be a problem that will come up time and again with different proteins. One wants a feel for what the radius of convergence is. A uerbach: What exactly have we learned? You started with S4 in the groove; there is a face of S4 that you can pack into the protein. Do we know where the S1/S2/S3 ordering is? Roux: No, not really. Auerbach: So S1/S2/S3 could be in any order on the outside. Roux: Pretty much, although I don't think that S1 is between S2 and S3. Perozo: So there are three key constraints: the overall dimensions of the molecule, Diane Papazian's electrostatics, and five distances. Roux: There is also lipid exposure from the tryptophan scanning. Perozo: What specific data do you need to converge into a more reliable model? Roux: If I had as few as five distances between S4 and S5/S6, you would see something coming out. We built the AgTx2/Shaker complex with nine constraints. Ashcroft: This goes back to what Mark was saying: if you have some indication of how many restraints you need to produce a reliable model of something for which the structure is already known, then you can come back to the experimentalists with a list. Roux: On the basis of the AgTx2/Shaker complex it is clear that if we have about a dozen constraints, we will be OK. With two restraints you can already be in the right ballpark. Sansom: But, of course, we don't know the answer for AgTx2, as we don't have the crystal structure of the channel—toxin complex.

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Roux: We did the barstar—barnase complex with the same method. With six or seven distances we got to about 1.7 A relative to the crystallograpic structure. Sansom: Essentially you are doing a jack-knifing test: can you take out data from the barstar—barnase, and see as you take more and more out, how much worse it gets? You need some sort of calibration. Roux: There are so many ways this can be done. In the current methodology the problem is not how accurate the model becomes with a certain number of restraints; rather, it is a matter of which restraints are fooling us because they were deduced from functional effects that we misunderstand. Sansom: In secondary structure prediction based on any pattern-learning approach, it is standard to take out the data and test how they match several times. Roux: It is clear that if I try to fold bacteriorhodopsin with too few restraints, I won't get the structure of bacteriorhodopsin. I am basically resolving the rough position of one helix here: I am saying that S4 must be at the interface between the subunits. Sixma: Compared to the toxin binding, how many variables do you have here, and how many did you have in the toxin? Now you have four independent units moving, whereas in the toxin there was just one. Roux: That is true, although in S1 to S4 they only have the angular rotation, whereas in the case of AgTx2 there is a full three-body rotation in three dimensions. Sixma: But the AgTx2 is moving as one element, whereas the subunits are moving independently, you may have many more variables. Roux: It is hard to tell, because the side chains are not rigid. Perozo: Diane Papazian, you also have a model of how S4 fits relative to the other subunits. Your proposal is that S4 is tilted relative to the membrane, whereas in Benoit Roux's model it is quite perpendicular. How do you explain this difference? Papazian: The tilt on our model came from trying to fit the S2-S4 constraints simultaneously in Shaker. However, we used extended side chains in our model. If you let the side chains rotate, the tilt can be much less. Mindell: But Benoit also constrained his helices to be perpendicular to the membrane. Roux: To a degree. The hydrophilic amino acids must be outside a slab of 30 A thick, which permits some tilting of the helices. Perozo: But you used the same set of data and your results are quite different. Roux: I am trying to generate a representative ensemble of models, which satisfy a number of established constraints about the structure of Shaker. The results should not be viewed as 'my model of Shaker', but rather as reflecting the present state of available structural information about Shaker. I believe that this approach will get us further in terms of defining what is known about the structure of Shaker. In my opinion, constructing and considering a single model (which satisfies known

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constraints) can be quite misleading because it can be very biased. Generating 100 models more or less randomly which have to satisfy a number of experimentally determined constraints is a sobering exercise. In particular, they show where the 'holes' are in the available information. That may not be a pleasant fact, but it makes it harder to delude oneself with one's favourite model. Also, it points in the direction where experimental data are most needed. On the other hand, when some structural feature appears to be recurrent, then one knows with confidence that it represents meaningful and robust knowledge about Shaker. Mindell: Once you are doing this kind of thing, is there a rule for trying to incorporate constraints based on sequence homologies? Roux: Before the tryptophan scan of Miller and his co-workers (Monks et al 1999, Hong & Miller 2000), people were tempted to just look for hypervariable regions and conclude that these must be exposed to the lipid. Once you have the tryptophan scan correlating to this, you don't really need anything more. What else would you use? Mindell: I would be looking more for things that are conserved, or that co-vary. Roux: Lockless & Ranganathan (1999) are doing this. If they find correlation between S5/S6 and S4, they will let me know. Swartz: We have done this, but we were unable to find any patterns. Perozo: Several studies in the mid-1990s looked for structural correlations within families of related proteins. With the exception of a few salt bridges, the general consensus is that this approach is not sensitive enough to pick up most compensating mutations. Papazian: If you look at divergent channels, covariance analysis would not necessarily pick out the structural constraints that we have identified in the Shaker voltage sensor. Sansom: If we relax the restraints on the S6 helix and let it bend a bit, how much would S4 then jiggle about? Roux: It is complicated to bend it, because S5 is in the way. I wasn't able to do this. In Gary Yellen's model, S6 is quite bent, and of course S4 will be influenced by this (del Camino et al 2000). Sansom: When we have tried to bend S6 at all, it is very difficult to maintain the KcsA-like packing. Roux: The S4 seems to be mostly packed against the core at the top, in the upper third of the structure. Choe: It would probably cost a lot more if you want to tilt it, because of the tepee shape of the core domain. Here's a hypothetical model: if you have KcsA sequences for the core domain S5/S6, but the rest is a Shaker sequence like you have, would you expect S4 to be in a similar conformation? Roux: 1 don't think it would make a large difference at the present level of modelizations.

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Auerbach: I know that you still have a lot of wiggle room in terms of the position of S4, but do you have a sense of whether residues of S4 are equally likely to interact with S5 or S6? Roux: Because S1 /S2/S3 is so underdetermined, by the time you have considered all these possibilities, S4 has almost gone round full circle. In the end you get a large number of amino acids of S4 that could possibly touch S5 or S6. Auerbach: Almost all the numbers in S5/S6 that you are picking out are in S5. Roux: One observation that could be a little discomforting is that some of the contacts suggested by the models were residues that did not have a large impact on the gating of Shaker upon mutation (Li-Smerin et al 2000). Swartz: There is an important qualification worth mentioning. The measurements you refer to are sensitive to perturbations at any step in the activation pathway. That is they don't distinguish between effects on the voltagesensor, the gate or the interaction between the two. We have now looked at the effects of pore domain shell mutations on charge translocation within the voltage-sensors by measuring perturbations in the charge versus voltage relations. These patterns are quite different than those we previously observed for perturbation of the conductance-voltage relations. My view is that, at least from our work, the position of S4 in relation to the core region (S5-S6) is a completely open question. Roux: I am not proposing any particular orientation of S4. I believe that somewhere among these pairs of amino acids, there are a few that could be experimentally verified. Auerbach: In the AgTx2, one of the double-mutant cycle analyses was well off. Do you have an explanation for this? Roux: We haven't finished this work, so what I can say is limited. Apparently, there is an effective coupling between Serll (of AgTx2) and Asp431 (of Shaker) (Ranganathan et al 1996) despite the fact that the two residues are not close in space. The coupling might arise because the toxin is very rigid and a coupling can be transmitted through the structure. Nonetheless, this case is an exception. Generally, most of the double mutants that appear to be coupled are indeed close in space (Schreiber & Fersht 1995, Buckle et al 1994). This is the basis for using these couplings as distance restraints in trying to derive a structural model. Mindell: Between yours and Rod's data on the toxin, it looks great that the outsides of the channels are the same, but how much constraint does that put on the other side? Gary's data argue that there are differences. Do you have a lot of freedom in there? Roux: In fact, even the turrets of Shaker are probably not much like KcsA. When we finally had our structure, we did an alanine scan of AgTx2 in the computer. We predicted that a few amino acids should affect the binding. In fact, they were

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mutated in the past. Some of them don't have a big effect, and they are involved with the turrets. Mindell: It seems that a lot of arguments of detail come from the interactions on the cytoplasmic end of this little notch. Roux: The precise structure near the tip of KcsA does not influence very much, except if there was this big kink in S6, then S4 might want to pack slightly differently. Miller: What is the sensitivity of the model to the size of the constraint? Let's say that the LRET is not very accurate, and the real constraint is 50 A and not 43 A. What happens to the predictions of your model? Roux: Then things would get quite undetermined. Instead of being +2 A I put my constraints as being +10 A, everything goes away. But the accuracy of LRET is better than that (Burmeister Getz et al 1998). Samom: In the context of hard versus soft restraints, you said you wanted to put a distance restraint for closed state versus open state. One was about 3 A and one was about 6 A. My back of the envelope calculation suggests that this might be a bit optimistic. If the LRET data suggesting quite a substantial rotation in S4 are accurate, then 6 A is a bit on the short side. I think the change is more like 20 A, if there are two side chains pointing towards one another and then you rotate both of them. Roux: Only one segment is thought to rotate, the S4. There is no evidence of S1, S2 or S3 undergoing significant rotations. References Aggarwal SK, MacKinnon R 1996 Contribution of the S4 segment to gating charge in the Shaker K+ channel. Neuron 16:1169-1177 Blaustein RO, Cole PA, Williams C, Miller C 2000 Tethered blockers as molecular 'tape measures' for a voltage-gated K+ channel. Nat Struct Biol 7:309—311 Buckle AM, Schreiber G, Fersht AR 1994 Protein-protein recognition: crystal structural analysis of a barnase— barstar complex at 2.0 A resolution. Biochemistry 33:8878-8889 Burmeister Getz E, Cooke R, Selvin PR 1998 Luminescence resonance energy transfer measurements in myosin. Biophys J 74:2451—2458 Cha A, Snyder GE, Selvin PR, Bezanilla F 1999 Atomic scale movement of the voltage-sensing region in a potassium channel measured via spectroscopy. Nature 402:809-813 del Camino D, Holmgren M, Liu Y, Yellen G 2000 Blocker protection in the pore of a voltagegated K+ channel and its structural implications. Nature 403:321-325 Doyle DA, Morais Cabral J, Pfuetzner RA et al 1998 The structure of the potassium channel: molecular basis of K+ conduction and selectivity. Science 280:69-77 Herzyk P, Hubbard RE 1995 Automated method for modeling 7-helix transmembrane receptors from experimental-data. Biophys J 69:2419-2442 Hong KH, Miller C 2000 The lipid-protein interface of a Shaker K+ channel. J Gen Physiol 115:51-58 Li-Smerin Y, Hackos DH, Swartz KJ 2000 A localized interaction surface for voltage-sensing domains on the pore domain of a K+ channel. Neuron 25:411-423

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Lockless SW, Ranganathan R 1999 Evolutionarily conserved pathways of energetic connectivity in protein families. Science 286:295-299 Monks SA, Needleman DJ, Miller C 1999 Helical structure and packing orientation of the S2 segment in the Shaker K+ channel. J Gen Physiol 113:415-423 Ranganathan R, Lewis JH, MacKinnon R 1996 Spatial localization of the K+ channel selectivity filter by mutant cycle-based structure analysis. Neuron 16:131 139 Schreiber G, Fersht AR 1995 Energetics of protein-protein interactions: analysis of the barnase barstar interface by single mutations and double mutant cycles. J Mol Biol 248:478 486 Silverman WR, Tang CY, Mock AF, Huh KB, Papazian DM 2000 Mg2+ modulates voltagedependent activation in ether-a-go-go potassium channels by binding between transmembrane segments S2 and S3. J Gen Physiol 116:663-678 Starace DM, Bezanilla F 2001 Histidine scanning mutagenesis of basic residues of the S4 segment of the shaker K+ channel. J Gen Physiol 117:469-490

Permeation energetics in a model potassium channel Stefano Garofoli, Gennady Miloshevsky, Vladimir L. Dorman and Peter C. Jordan 1 Department of Chemistry, MS-015, Brandeis University, PO Box 549110, Waltham, MA 02454-9110,USA

Abstract. Known structures of selective ion channels share a common property: a narrow constriction, presumably crucial for ionic discrimination. This region can be fairly long, imposing single file motion on waters and ion(s). We apply the semi-microscopic Monte Carlo approach to study permeation in the KcsA channel, decomposing energetics into a three-step process: cation dehydration; ion transfer into a uniform low £ dielectric; and transfer from the uniform dielectric into the channel. The influence of individual channel structural features is separately assessed. The aqueous cavity has only a modest stabilizing effect on nearby ions in the filter. Ionic solvation in the filter reflects the combined influence of the single file waters, the binding pockets' carbonyls, the a helices directed at the cavity and the negative residues near the extracellular surface of the channel; no one feature dominates. At all sites along the permeation pathway there is substantial discrimination favouring K+ over Na + ; conversely, there is little discrimination among the larger alkali cations. Selectivity for K+ over Na+ appears due to the inability of the filter's carbonyl oxygens to ideally coordinate Na + . 2002 Ion channels—from atomic resolution physiology to functional genomics. Wiley, Chichester (Novartis Foundation Symposium 245) p 109-126

Until recently theoretical study of ionic interaction with ion channel proteins was either based on structural speculations or limited to considering the model system gramicidin (see Roux & Karplus 1994). The situation is now dramatically different. Four distinct selective channel systems have been solved to atomic level resolution: a K+ channel from Streptomyceslividans (KcsA; Doyle et al 1998), a stretch-activated channel from Mycobacterium tuberculosis (Tb-McsL; Chang et al 1998), human red cell aquaporin 1 (AQP1; Murata et al 2000) and the Escherichia coli glycerol facilitator (GlpF; Fu et al 2000). All share a common feature, a constricted region where the transported species must lose much of its surrounding water and pass in close proximity to the channel protein. In KcsA the constriction is This paper was presented at the symposium by Peter Jordan, to whom correspondence should be addressed.

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visible from the X-ray structure and is associated with a single file domain like that in gramicidin (Wallace 1999), but substantially shorter, some 10-15 A long. A similar feature may be integral to the function of AQP1 (Murata et al 2000). The structure of KcsA confirmed many electrophysiological inferences, investing kinetic models of the multi-ion permeation pathway (Hille & Schwartz 1978, Neyton & Miller 1988) with structural reality (Doyle et al 1998). It also revealed some unexpected architectural details: the carbonyl binding pockets, the mid-channel aqueous cavity and the a helices aimed at the cavity. The AQP1 structure, with a helices pointed at the constriction, rationalizes how this protein forms water channels and simultaneously blocks proton transport. Nonetheless, questions remain. In KcsA, what creates essentially barrier-less free energy profiles for permeant ion transport, i.e. why are K+ channel conductances so high? What accounts for essentially insurmountable energetic obstructions to the flow of similar competing species, i.e. why is the K + /Na + permeability ratio so high? How do individual structural features affect permeation energetics? Which features simply lead to superposable static fields and which induce major dielectric reorientation? This chapter examines these issues, describing the effect of individual structural features on the permeation free energy profile, and suggesting reasons for certain aspects of channel design. We extend the approach to compare permeation energetics among the alkali cations, emphasizing the importance of hydration energetics. Modelling ion channels Many theoretical approaches illuminate structure-function relationships in ion channels. Gramicidin has been their proving ground (Roux & Karplus 1994, Dorman et al 1996, Woolf & Roux 1997, Chiu et al 1991, Jakobsson & Chiu 1987). Using the X-ray structure of KcsA as a guide, insight has been gained from electrostatic analysis (Roux & MacKinnon 1999), Brownian dynamics (BD) (Chung et al 1999) and molecular dynamics (MD) (Aqvist & Luzhkov 2000, Shrivastava & Sansom 2000, Berneche & Roux 2000, Guidoni et al 2000, Biggin et al 2001). In MD, the computational models hew closely to the known structure and provide a wealth of information. Among the simulational results are: a model for the permeation duty cycle (Aqvist & Luzhkov 2000); evidence for the permeant ions' role in structurally stabilizing the channel (Shrivastava & Sansom 2000); a detailed picture for the functional permeating assembly (Berneche & Roux 2000); identification of a possible secondary influence of the oriented a helices (Guidoni et al 2000); and a novel hypothesis for the origin of K + / N a + selectivity (Biggin et al 2001). In contrast, both BD and electrostatic studies are mesoscopic in nature, necessarily partially idealized. The transmembrane aqueous pathway is a

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continuum fluid with high permittivity, approaching or equal to that of bulk water, even though water in narrow constrictions must be ordered and nonpermittive (Partenskii & Jordan 1992, Partenskii et al 1994). BD successfully reproduced gross aspects of transport kinetics (Chung et al 1999). The Roux & MacKinnon (1999) study provided a basis for the cavity's ability to preferentially solvate monovalent cations. Our perspective on KcsA is somewhat different. We treat prescribed structural features that we believe critical for the energetics of ion transfer from water (Dorman et al 1996, 1999). These define an exactly soluble, computationally efficient statistical mechanical problem. The model, illustrated in Fig. 1, incorporates a few mobile, reorientable features (the ion[s], the single file waters in the channel, and the carbonyls forming the binding pockets); the remainder (the cavity, the oriented a helices and the negative residues) are, for computational convenience, treated as fixed background charges, although this restriction can be lifted. The bulk water domains are continua with high dielectric constants, for computational simplicity chosen as infinite. The cavity is treated in two ways: as a high £ continuum or by incorporating explicit cavity waters, 20 additional

FIG. 1. Semi-microscopic model geometry for the KcsA selectivity filter. It includes solvating CO groups (residues 75—78 of each tetramer strand), single file ions and waters, peptide dipoles, the 80Asp carboxylates, the aqueous cavity and its included ion. Bulk electrolyte and the cavity are treated as dielectric continua, e = . The Helmholtz layer (accounting for water immobilized by interaction with polar surfaces) separating the explicit sources in the filter from extracellular bulk water has a width of 2 A; that between the filter and the mid-channel water pool is 1.5 A. The pool radius is 5.0 A and it accommodates ~20 waters. The crystallographic occupancy sites (2 and 4) are ~ 18.5 A and ~ 11.0 A from the cavity centre.

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mobile, reorientable moieties in the low £ dielectric background. The surrounding membrane and those parts of the channel not explicitly modelled form a background continuum dielectric with eback ~2. The picture is semiquantitative, designed to deconstruct individual structural features' influences on ion transfer and to facilitate comparison of ion-channel interactions among the alkali cations. This approach permits description of the proximate structural reorganizations associated with ionic solvation in a channel environment; exactly treating charge induced dielectric relaxation of 'solvent', i.e. the parts of the system nearest the transported ion(s). The choice of eback ~2 derives from the index of refraction (the high frequency dielectric constant); it is the electronic contribution to £. This approach circumvents drawbacks of both Poisson-Boltzmann and Brownian treatments where the constriction is treated as a high e continuum, even though it contains ordered, non-permittive single-file waters (Partenskii & Jordan 1992, Partenskii et al 1994) and where all protein charges are immobile, with stabilization arising from (real) structural reorganization dealt with by assigning the protein an elevated effective e, between 4 and 20 (Antosiewicz et al 1994, Gibas & Subramaniam 1996). In our treatment, structural reorganization of the single file waters and the binding pocket carbonyls in the ions' immediate vicinity is treated exactly; other electrical features are stationary. The model is fundamentally electrostatic; channel solvation involves transfer of an ion into a cavity accompanied by dielectric reorganization of the immediate surroundings. The approach is very efficient computationally. With ~80-100 mobile sources, statistically reliable free energy perturbation calculations for any point on the permeation pathway are achieved in < 10 h on a personal computer. Model parameters roughly reproduce alkali metal hydration energies and gross aspects of both ion-water and water-water pair correlation functions. Permeation is a composite process. The ion is dehydrated, and exchanged for water in the gas phase; the ion is transferred to a cavity in the background dielectric and exchanged for water; it is then stabilized by exchange for a water molecule in the channel: lon(aq) + Water(vac) —> lon(vac) + Water(aq) lon(vac) + Water(aback) —> Ion(eback) + Water(vac)

Dehydration (la) Cavity

(1b)

Ion(eback) + Water(channel) —> lon(channel) + Water(eback) Stabilization (1c) The stabilization energy is computed by perturbation methods described previously (Dorman et al 1996); the dehydration energy is experimentally

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1

accessible ; the cavity term is a Born energy, determined by the ion's cavity radius in the channel. The model of Fig. 1 has five sites. Four have crystallographic correspondences (Doyle et al 1998) and/or rough electrophysiological identifications (Neyton & Miller 1988): sites 2 and 4 are the crystallographic sites (outer lock-in and enhancement respectively); site 3 is the interionic water site; and site 5 approximates the Ba block site (Jiang & MacKinnon 2000). The extracellular boundary site 1, identified from simulational studies, more or less accounts for extracellular vestibular water's influence on filter energetics. Default geometry is a strand-averaged symmetrization of the crystallographic coordinates. For computational efficiency carbonyl carbons are immobilized, a restriction that can be lifted. Oxygens rotate about the carbons, weakly tethered to equilibrium orientations determined by minimizing the crystallographic structure with an unoccupied filter (hypothetical). Waters and ions are not constrained. In default geometry the cavity, of ~ 5 A radius, is centred 26 A and 23 A from the extraand intracellular boundaries, respectively. The temperature is 300 K.

Role of the cavity The cavity's major role is to stabilize an ion near the centre of the membrane (Doyle et al 1998, Roux & MacKinnon 1999). However, does it also help stabilize ions in the selectivity filter? Or might it have some other secondary influence on permeation? Is proximity to the cavity as effective as bulk water in stabilizing filter ions; if not, how large is the penalty? Does the cavity isolate the filter from the low e domain on the channel's intracellular side? Are filter energetics very different in the open channel? Is the cavity as effective in stabilizing filter ions as additional single file waters? Tables 1—3 provide answers to these questions, by limiting consideration to ionwater interaction. Five basic variants from default geometry of Fig. 1 are treated: (1) replace the cavity by explicit waters; (2) vary cavity radius, at constant overall system width; (3) approximate an open state, shrinking overall system width until the cavity contacts the intracellular region or (4) deforming the cavity to a tube contacting the intracellular region and filled with explicit waters; and (5) eliminate the cavity, replacing it by additional single file waters. 1 The process of Eq. la is hypothetical, but energies can be estimated with a fair degree of confidence. What is needed is the absolute potential of the standard hydrogen electrode. The most recent experimental and theoretical determinations differ by ~ 7 kT (Reiss & Heller 1985; Tissandier et al 1998); dehydration free energies are thus uncertain to + 3.5 kT.

GAROFOLI ET AL

114

TABLE 1 Effect of cavity size and cavity occupancy (cases 1 and 2, see text) on monovalent ion stabilization free energies (Eq. 1c, in kT) for single occupancy of the model filter (Fig. 1, the default geometry)

Width/A

4 /A Explicit waters Rcavity

Default

49

5

File-5, Cavity-0

Case 1

49

5

File-5, Cavity-20

Case 2

49

8.5

File-5, Cavity-0

Case 2

49

0

File-5, Cavity-0

Site 1

Site 2

Site 3

Site 4

Site 5

-28.3 -28.1 -28.4 -28.0

-25.8

-20.5

-20.4

-19.2

-25.4

-20.4

-20.7

-20.6

-26.0

-21.0

-21.9

-25.2

-19.8

-21.6 -18.7

-14.7

Case 1 replaces the continuum cavity by explicit cavity water (see text).

TABLE 2 Effect of varying intracellular channel geometry (cases 3 and 4, see text) on monovalent ion stabilization free energies (Eq. 1c, in kT) for single occupancy of the model filter (Fig. 1, the default geometry)

Default

Width/A

Rcavity/A

Explicit Waters

Site 1

Site 2

Site 3

Site 4

Site 5

49

5

File-5, Cavity-0

-28. .3

-25.8

-20.5

-20.4

-19.2

-26.2 -21.2 - 25.7 -20.8

Case 3

33

5

File-5, Cavity-0

-28. .6

Case 4

49

Tube

File-5, Tube-72

-28. ,4

-21 .5 -21.1 -21 7 -22.3

Case 4 incorporates explicit 'tube' water (see text).

TABLE 3 Effect of replacing the continuum cavity by single file waters (case 5, see text) on monovalent ion stabilization free energies (Eq. 1c, in kT) for single occupancy of the model filter (Fig. 1, the default geometry) Width/A

Rcavity/A

Explicit waters

Site 1

Site 2

Site 3

Site 4

Site 5

File-5, Cavity-0 File-5, Cavity-0

-28. .3 -28.0

-25.8 -25.2

-20.5 -19.8

-20.4 18.7

19.2 -14.7

-28.1 -28.1

-25.6

-20.5

-20.5

-28. .2

-20.8

-21.4 -22.0

-21.8 23.6

-28..3

-25.8 -25.9

Default

49

5

Case 5

49

0

Case 5

49

0

Case 5

49

0

File-6, Cavity-0 File-7, Cavity-0

Case 5

49

0

File-8, Cavity-0

,1

-21.1

-24.5

Stabilization energies at site 1, furthest from the cavity, are, as expected, essentially independent of intracellular structure, suggesting an overall statistical uncertainty of +0.3 kT. Table 1 describes different modifications of the cavity: filling it with explicit waters (case 1) or varying its radius (case 2). The inclusion of explicit cavity waters tests the high £, continuum cavity approximation. But for site 5, adjacent

PERMEATION ENERGETICS

115 2

to the cavity, there are no major differences . Altering the cavity radius noticeably perturbs ionic stability at the two inner sites; the effect is moderate at site 4 (~ kT) and large (> 2 kT) only at the innermost site, 5. Table 2 illustrates the effect that transition to an open state may have on filter energetics. Whether the continuum cavity contacts the intracellular space (case 3) or the water-filled cavity is extended and connected to the intracellular space (case 4), outer site energetics is unaltered. Again changes at site 4 are small, but real (~ kT) and those at site 5 are moderate (~ 2—3 kT). Table 3 contrasts the influence of the cavity on filter ion stabilization with that of hypothetical additional single file waters. One single file water would be fully compensatory. A clear picture emerges. Not surprisingly, the cavity isn't designed with an eye toward filter energetics; additional single file waters would be more effective. It clearly helps stabilize ions at site 5; since ion—cavity interactions arise from image forces, quadratically dependent on valence, they may contribute to Ba stabilization near the channel—cavity boundary. The cavity electrically isolates filter ions from the non-permittive intracellular side of the channel assembly. Without a change in filter geometry, filter energetics could only be marginally altered in the transition from closed to open state. Ion transfer energetics — individual electrical features and stabilization As an ion enters the channel, ion—protein interaction must offset the ion's dehydration energy. The solvation environment along the interior of the permeation pathway is dramatically different from that in bulk water. The dehydration energy (Eq. la) ranges from 115 (Cs+) to 160 (Na+) kT. This is balanced by cavity (Eq. 1b) and stabilization (Eq. 1c) components. We first focus on stabilization, the process occurring within the uniform dielectric background; at each site it is roughly the same (to within ~ 15 kT) for the four alkali cations and compensates for 5CM)5% of the dehydration energy. Deconvolution of ionic interaction with individual structural features provides insight into how each helps make the filter ionophilic. We consider (hypothetical) single occupancy and separately assess the influence of bulk and cavity water, of the single file waters, of the binding pocket carbonyls, of the oriented a helices, and of the 80Asp near the extracellular mouth. Figures 2 and 3 decompose the stabilization free energy for one K + -like ion in the filter. They illustrate each feature's 2

This may overestimate the cavity's stabilizing ability. MD simulation of water in ~ 20 A cavities suggests an e of ~5 (Zhang etal 1995), like that in single file channels (Partenskii & Jordan 1992, Partenskii et al 1994). However, work on large cylindrical channels (radius ~8 A) is consistent with a larger e, ~ 30 (Sansom et al 1997).

116

GAROFOLIETAL

FIG. 2. Individual contributions of the various electrical features (continuum background, ; single file waters, ; binding pocket carbonyls, ; oriented a helices, ; negative residues, •) to monovalent ion stabilization energy at various occupancy sites in the model KcsA filter. The cavity is unoccupied.

contribution to the stabilization free energy (Eq. 1c), for scenarios where the cavity is ion free (Fig. 2) or occupied (Fig. 3) by a monovalent cation. For K+, the total stabilization energy at the physiological sites (2—5) ranges between 75 and 85 kT. Consider ionic stabilization with the cavity ion free (Fig. 2). Attraction to the dielectric background (bulk and cavity water) is strongest near the two boundaries; the cavity is half as effective as bulk water. Stabilization by the single file waters is complementary, weakest at the boundary sites where the ion has but one single file neighbour. Net interaction with waters of all kinds (bulk, cavity and single file) varies from ~ 20 to ~ 30 kT. The remaining dielectric stabilization mainly reflects ioncarbonyl and ion—helix attraction. Near the cavity the filter sites are very near the a helices'C-terminiandhelixinteractiondominates.TheC-terminiare ~ 6 Afromsites 4 and 5 but ~ 14 A from site 2; in all cases the amino termini are ~ 20 A away. The 80 Asp at the peptide—water interface are strongly shielded by nearby bulk solvent; their ability to stabilize filter cations is consequently much reduced3.

3

In the dielectric picture, each of these charges induces an electrical image (of opposite polarity) in the solvent, creating a dipole. Near the aqueous interface these charge separations are small; their influence on ions in the selectivity filter is much less than that of effective dipoles created by charged groups in the low £ interior.

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117

FIG. 3. Individual contributions of the various electrical features (continuum background, ; single file waters, ; binding pocket carbonyls, A; oriented a helices, ; negative residues, ) to monovalent ion stabilization energy at various occupancy sites in the model KcsA filter. The cavity is occupied by a monovalent cation.

Introducing an ion into the cavity (Fig. 3) has little effect on ion—peptide energetics. Interaction with the background (here including the cavity ion) destabilizes the filter ion; interionic repulsion decreases much more rapidly than R-1 due to solvent (image) effects. Ions at site 2 or 3 are much closer to the cavity ion's (negatively charged) image than are ions at site 4 or 5; net filter ion-cavity ion repulsion at site 2 is ~35% that at site 4, even though their direct coulombic interaction is ~60% as large. At site 5 there is significant compensation. Interaction with the single file water becomes relatively favourable since the cavity ion reinforces a site 5 ion's tendency to align channel water.

Ion transfer energetics — the ionic Born cavity Processes (a) and (b) of Eq. 1 require charge transfer between different dielectrics. Experimental data provide reliable estimates for step (a), dehydration (see footnote 1). Step (b) is a Born transfer from vacuum to the uniform background (eback = 2); the associated energy is

118

GAROFOLIETAL

TABLE 4 Ion-oxygen distances (in A) in an energy minimized KcsA K+ channel for alkali cations occupying the crystallographic sites (2 and 4). Bold face entries are for oxygens identified as forming part of the binding cavity boundary Binding site 4

Binding site 2

Ion

W-1

O-78

O-77

W-3

W -3

O-76

O-75

W-5

Na+

2.40

3.69, 4.24, 5.00, 4.75

2.38, 2.47, 2.36, 2.47

2.29

5.63

4.26, 3.83, 4.53, 4.64

2.28.2.31.

2.34

2.32, 2.38

K+

2.60

4.86, 3.77, 4.78, 3.14

2.64, 2.69, 2.66, 2.68

2.72

3.48

2.70, 2.75, 2.91, 2.82

2.67, 2.76, 2.75, 2.67

3.98

Rb+

2.86

3.95, 3.33, 5.07, 3.29

2.75, 2.85, 2.80, 2.73

2.71

3.62

2.83, 2.88, 2.79, 2.73

2.86, 2.90, 2.78, 2.81

3.60

Cs+

2.98

3.51, 3.72, 3.37, 5.16

2.92, 2.93, 3.02, 2.91

2.86

3.05

2.95, 3.06, 2.97, 3.02

3.07, 2.99, 3.04, 2.96

3.69

W-1 is the distance from the ion to the water oxygen at site 1, etc. O-78 is the distance from the ion to the carbonyl oxygens of residues 78, etc.

The cavity radius, Rcavity is determined by establishing a dividing surface separating ion from 'solvent'. This distance is not an intrinsic ionic property; it is also solvent dependent (Grunwald 1997). This is especially true in the nonuniform, inherently non-symmetric medium (even when time averaged) of a channel interior (Jordan 2002). In principle, the ion could ideally associate unconstrained single file waters. However, the binding pocket carbonyl oxygens are significantly constrained, their range of motion limited by the peptide backbone's rigidity (Doyle et al 1998, Berneche & Roux 2000). At each of sites 2-4 the oxygens from eight carbonyl and two single file waters form the binding environment. However, not all coordinate the permeant ion equally well. In addition, different ions are more or less effectively coordinated. Table 4 presents ion—oxygen distances for energy-minimized channels containing a single alkali cation at sites 2 and 4. The rest of the filter, the cavity and the extracellular vestibule are filled with water. The local environments are far from symmetric; only some ligands form part of the first solvation shell. To estimate the size of the cavity, we proceed somewhat arbitrarily, assuming (1) that at least one carbonyl from each set of ligands must form a part of the boundary and (2) that waters are not necessarily bounding ligands. Thus, at site 2 mean distances between ions and their bounding ligands are 2.58 A for Na+, 2.73 A for K+, 2.83 A for Rb+ and 3.00 A for Cs+. While mean ion-oxygen distances for the larger alkali cations are nearly optimal (similar to those in water), that for Na+ is very large. The Na+K+ difference is only 0.15 A, much less than in water, 0.38 A; the channel does not adjust as well to Na+ as to the other alkali cations. Consequently the Born energy,

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119

TABLE 5 Decomposition of ion transfer free energy (in kT) for alkali cations occupying the crystallographic sites (2 and 4) of a singly occupied channel (with ionfree cavity) Binding site 2

Binding site 4

Ion

Na+

K+

Rb+

Cs+

Na+

Dehydration

163.1 -98.8 -76.1 -11.8

134.3 -86.9 -73.9 -26.5

125.2 -74.8 -73.0 -22.5

115.9 -70.3 -69.7 -24.1

163.1 134.3 -100.5 -84.3 -86.2 -81.7 -22.7 -31.7

Cavity Stabilization Total

K+

Rb+

Cs+

125.2 -78.1 -81.8 -34.7

115.9 -69.7 -76.7 -30.5

Eq. 2, is relatively small for Na+. To quantitate the cavity energy requires an estimate of the size of the O atom; as we are using energy minimized structures to estimate cavity size, we choose a rather small value for the O atom radius, 1.3 A, consistent with recent ab initio studies (Roux & Karplus 1995).

Ion transfer energetics — summary Table 5 presents the individual contributions to the free energy of ion transfer from water to the channel interior for alkali cation occupancy of sites 2 and 4. Due to our approximations, total free energies are highly approximate. The results are most useful for contrasting filter interaction among the alkali cations. Cavity and stabilization components are comparable. Both the electrostatic interaction (stabilization) and the cavity contribution are largest for Na+. This ion interacts extremely well with the peptide, better than its larger congeners, consistent with the suggestion that internal Na+ blocks K+ permeation (Heginbotham et al 1999). However, as the peptide is insufficiently flexible, the Na+ cavity remains too large; the net interaction is inadequate to fully compensate for Na+'s much larger dehydration energy. Figure 4 presents estimates of the ion transfer free energies, relative to K+ at site 2, for alkali cation occupancy of the crystallographic sites4. Na+ occupancy is always unfavourable. On average it is ~15kT less stable than K+, comparable with observed permeability ratios that imply a AG ~10kT (Hille 1992). While the larger alkali cations are energetically similar, Cs+ clearly interacts less well 4

Sites 1 and 5 are both eliminated from consideration, the former because it really is not part of the single file, the latter because the explicit solvation environment is incomplete. States with ions occupying neighbouring sites are energetically inaccessible and thus ignored.

120

GAROFOLIETAL

FIG. 4. Permeation free energy, relative to K+ occupancy of site 2, for single and double alkali cation occupancy of four states of the selectivity filter of the model KcsA channel (Na+, ; K+, ; Rb+, ; Cs+, ). All energies have been displaced by +52.6 kT; the shifted free energies are independent of the cavity's ionic occupancy state.

than either K+ or Rb+; there appears to be a slight preference for K+ over Rb+. Both observations are consistent with experiments. Model limitations This approach to permeation energetics is highly approximate. Significant terms, like the energy required to create an uncharged Born cavity, have been ignored (Roux et al 1990). Dielectric relaxation has been limited to the ions' first solvation shell; other structural features were immobilized. The outer vestibule and the cavity are viewed as low and high e dielectric continua respectively. These are all restrictions that can be lifted. Conclusions Even in its limited form, this model provides a way to separately assess how individual architectural features of the channel affect permeation. It demonstrates that the cavity effectively isolates the filter from the intracellular domain and that it is especially effective in aiding stabilization of divalent ions at the cavity-filter boundary (the Ba-block site). It demonstrates that the oriented a helices, in addition to stabilizing an ion in the cavity, also contribute importantly to ionic stabilization at the cavity-filter boundary. It corroborates the idea that peptide

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121

rigidity may be at the heart of K+ channel selectivity (Doyle et al 1998). It demonstrates that discrimination among the larger alkali cations requires delicate energetic trade-offs involving Born stabilization in the dielectric cavity and dielectric stabilization by the surrounding charge distribution. Acknowledgements This work supported by the National Center for Supercomputing Applications and by the National Institutes of Health, grant GM-28643. We thank M. B. Partenskii for helpful comments. D. Patangia determined the data of Table 4.

References Antosiewicz J, McCammon JA, Gilson MK 1994 Prediction of pH-dependent properties of proteins. J Mol Biol 238:415-436 Aqvist ], Luzhkov V 2000 Ion permeation mechanism of the potassium channel. Nature 404:881-884 Berneche S, Roux B 2000 Molecular dynamics of the Kcs A K+ channel in a bilayer membrane. BiophysJ 78:2900-2917 Biggin PC, Smith GR, Shrivastava I, Choe S, Sansom MSP 2001 Potassium and sodium ions in a potassium channel studied by molecular dynamics simulations. Biochim Biophys Acta 1510:1 9 Chang G, Spencer RH, Lee AT, Barclay MT, Rees DC 1998 Structure of the McsL homolog from Mycobacterium tuberculosis: A gated mechanosensitive ion channel. Science 282:2220— 2226 Chiu SW, Jakobsson E, Subramaniam S, McCammon JA 1991 Time-correlation analysis of simulated water motion in flexible and rigid gramicidin channels. Biophys J 60:273—285 Chung S-H, Allen TW, Hoyles M, Kuyucak S 1999 Permeation of ions across the potassium channel: Brownian dynamics studies. Biophys J 77:2517—2533 Dorman VL, Partenskii MB, Jordan PC 1996 A semi-microscopic Monte Carlo study of permeation energetics in a gramicidin-like channel: the origin of cation selectivity. Biophys J 70:121-134 Dorman VL, Garofoli S, Jordan PC 1999 Ionic interactions in multiply occupied channels. In: Gramicidin and related ion-channel forming peptides. Wiley, Chichester, (Novartis Found Symp225)pl53-169 Doyle DA, Morais-Cabral J, Pfuetzner RA et al 1998 The structure of the potassium channel: Molecular basis of K+ conduction and selectivity. Science 280:69—77 Fu D, Libson A, Miercke LJ et al 2000 Structure of a glycerol-conducting channel and the basis of its selectivity. Science 290:481-486 Gibas CJ, Subramaniam S 1996 Explicit models in protein pKa calculations. Biophys J 71:138147

Grunwald E 1997 Thermodynamics of molecular species. Wiley-Interscience, New York Guidoni L, Torre V, Carloni P 2000 Water and potassium dynamics inside the KcsA K+ channel. FEBS Lett 477:37-42 Heginbotham L, LeMasurier L, Kolmakova-Partensky L, Miller C 1999 Single Streptomyces lividans K+ channels: functional asymmetries and sidedness of proton activation. J Gen Physiol 114:551-560 Hille B 1992 Ionic channels of excitable membranes, 2nd edn. Sinauer Associates, Sunderland, MA.

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Hille B, Schwartz W 1978 Potassium channels as multi-ion single-file pores. J Gen Physiol 72:409-442 Jakobsson E, Chiu SW 1987 Stochastic theory of ion movement in channels with single-ion occupancy. Application to sodium permeation of gramicidin channels. Biophys J 52:33-45 Jiang Y, MacKinnon R 2000 The barium site in a potassium channel by X-ray crystallography. J Gen Physiol 115:269-272 Jordan P 2002 Ionic energetics in narrow channels. In: Layton H (ed) Proceedings of the IMA Workshop on Membrane Transport and Renal Physiology. Springer-Verlag, New York, in press Murata K, Mitsuoka K, Hirai T et al 2000 Structural determinants of water permeation through aquaporin-1. Nature 407:599-605 Neyton J, Miller C1988 Discrete Ba2+ block as a probe of ion occupancy and pore structure in the high conductance Ca2+- activated K+ channel. J Gen Physiol 92:569-586 Partenskii MB, Jordan PC 1992 Theoretical perspectives on ion channel electrostatics. Continuum and microscopic approaches. Q Rev Biophys 24:477-510 Partenskii MB, Dorman V, Jordan PC 1994 Influence of a channel-forming peptide on energy barriers to ion permeation, viewed from a continuum dielectric perspective. Biophys 1 67:1429-1438 Reiss H, Heller A 1985 The absolute potential of the standard hydrogen electrode: A new estimate. J Phys Chem 89:4207-4213 Roux B, Yu H-A, Karplus M 1990 Molecular basis for the Born model of ion solvation. J Phys Chem 94:4683-4688 Roux B, Karplus M 1994 Molecular dynamics simulations of the gramicidin channel. Annu Rev Biophys Biomol Struct 23:731-761 Roux B, Karplus M 1995 Potential energy function for cation-peptide interactions: An ab initio study. J Comp Chem 16:690-704 Roux B, MacKinnon R 1999 The cavity and pore helices in the KcsA K+ channel: electrostatic stabilization of monovalent cations. Science 285:100-102 Sansom MS, Smith GR, Adcock C, Biggin PC 1997 The dielectric properties of water within model transbilayer pores. Biophys J 73:2404-2415 Shrivastava IH, Sansom MS 2000 Simulations of ion permeation through a potassium channel: molecular dynamics of KcsA in a phospholipid bilayer. Biophys J 78:557-570 Tissandier MD, Cowan KA, Feng WY et al 1998 The proton's absolute enthalpy and Gibbs free energy of solvation from cluster-ion solvation data. J Phys Chem A 102:7787-7794 Wallace B 1999 X-ray crystallographic structures of gramicidin and their relation to the Streptomyces lividans potassium channel structure. In: Gramicidin and related ion-channel forming peptides. Wiley, Chichester (Novartis Found Symp 225) p 23-37 Woolf TB, Roux B 1997 The binding of sodium in the gramicidin A channel: comparison of molecular dynamics with solid-state NMR data. Biophys J 72:1930-1945 Zhang L, Davis HT, Kroll DM, White HS 1995 Molecular dynamics simulation of water in a spherical cavity. J Chem Phys 99:2878-2884

DISCUSSION Tieleman: How do you treat the energy differences between the ions in vacuum and the ions in water? Jordan: We have a background dielectric constant of two. We have a three point model for water and we have adjusted things so that in terms of our thermodynamic cycle we can come within 3-5 kT of the dehydration energies.

PERMEATION ENERGETICS

123 +

+

+

The radii are reasonably well tuned. My Na is 1 A, K is 1.3 A, Rb is 1.5 A and Cs+ is 1.7 A: with these I come within about 3kT of the dehydration energies determined experimentally. Roux: How do you calculate the dehydration energy? Jordan: We make up a sphere about 15 A in diameter that has a bunch of our model waters in it. We go through the whole equivalent cycle and do the calculations. Roux: Do you correct for what is beyond 15 A? Jordan: Yes, but it is an imperfect approach, even for establishing parameters. We must estimate the size of the cavity that surrounds the ions. This is even more of a problem in our modelling of KcsA since the carbons of the carbonyls are immobile, so we can't in any consistent way determine the size of the cavities surrounding the ions. The cavity sizes we used have therefore been derived rather differently. We have just minimized the structures using molecular mechanics and looked at how the immediate surroundings of the ions at the various sites differ as the ion is varied. It is really cobbled together at that point, which is why the numbers are screwy. This is one of the reasons we really want to let those carbons move so that we have an internally consistent picture. Roux: With regard to the ion solvation problem, I don't doubt that it is possible to calibrate a potential function by doing the free energy calculation with the ions. Nonetheless, when you start to look at the literature, it is very disconcerting that the solvation free energy of these simple cations and chloride is uncertain, in fact well beyond the selectivity of biological channels. A recent discussion of the large variations in ion solvation can be found in Pliego & Riveros (2000). The estimate for the free energy of Na+ ranges from — 92 to — l00kcal/mol, and for Cl- it ranges from — 78 kcal/mol to — 86 kcal/mol. Then all of a sudden Cl- is very close to Na + , whereas I always thought it was close to K+. There is no way the computational people will be able to do anything meaningful until those numbers are tied down. Jordan: People have been working on this problem for a long time. The problem comes down to getting an absolute electrode potential for the hydrogen electrode. Rottx: Isn't it possible to choose something else as a reference? Jordan: It doesn't make any difference. If you had an absolute measurement of the hydrogen electrode it would be fixed. Every time this unknown parameter is changed, you move all the monovalent cations up an amount and all the monovalent anions down by the same amount. With the divalents whatever happens is doubled. Sansom: Does that change the relative position within the monovalent cations? Jordan: No, but it drastically alters cation and anion values. Among the more recent values, there was an experimental determination of the standard hydrogen electrode potential by Reiss & Heller (1985) that was about 7 kT different from an

124

DISCUSSION

extrapolated procedure done very recently, based on the idea that if you think about adding waters to the ion microclusters, you will eventually get a solvated ion (Tissandier et al 1998). This is for free energy. If you are interested in enthalpy of solvation, it is even worse. Roux: With improvements in technology is there any hope that a better measurement could be made? Jordan: No, because ultimately you are doing a measurement that requires you to come up with some way of approximating what is happening in the interface. One of the ways of dealing with this is to try to find how much of a potential change occurs when you bring the ion directly across the interface. This is very hard to do. People have worked on this and come up with different ways of doing it. If I had to guess I would say that there is an uncertainty of about + 3.5kT. These numbers differ from Marcus's estimates (Marcus 1991) by about 50 K j m o l - 1 , mainly because we use a different value for the absolute potential of the standard hydrogen electrode. But there is also another problem. If you look at the numbers he quotes in his tables and then go back to your freshman chemistry texts and work through the relative free energies for the alkali cations, the numbers aren't completely internally consistent. He has cobbled things together and made some approximations of his own. It is a real problem. Sansom: If you had a slightly more deformable cavity, do you think one could switch the exact value of the K+ versus Na+ selectivity? That is, would there be smaller DG if you allowed more deformability in the cavity? Jordan: I would imagine that if things were more deformable, the DGs of permeation would mush together. It depends on the root-mean-square displacements that I assign to the carbon motion. In our experiments the cavity radii were determined in a way that makes things roughly comparable to what people have determined using molecular dynamics. Sansom: At the back of my mind are channels such as Kir6.2, where the tyrosine of the filter is replaced by a phenylalanine and the Na + /K + selectivity is lower. Jordan: We can certainly adjust the model to give more mobility to the waters in the cavity. Until now we have been dealing with a spherical cavity, because this is a historical artefact. We introduced this because when one does continuum electrostatics it is easy to compete with a spherical cavity. If we are putting explicit waters in there (which doesn't cost us very much), we could change the shape of the cavity. We could probably also give the cavity walls some flexibility. Perozo: I am trying to come to terms with your numbers and calculations in the sense of a working channel. You calculate ion flux. What kind of single-charge currents do you get? Jordan: I can't do it. If I look at what I have here, if I solve for conductance using these free energies I would find values of absolutely zero.

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125

Perozo: Then, in any calculation based on the KcsA structure, do people get energies compatible with the single-charge currents that have been mentioned? That is, do people get something comparable to what would make sense given the actual conductances? Roux: To get the correct magnitude of flux, you don't expect free energy barrier of more than 3 or 4 kcal/mol. Perozo: So what are the values that people have obtained based on the structure? Roux: It depends whether you allow the structure to be flexible or not. If you keep the structure very rigid, you will get one kind of answer. If you allow the full dynamics of the structure, you get a fairly different kind of answer. Perozo: What I am really getting at is that you try to come to ways such that your terms make sense from the physical character point of view. Could it be that the structure used in the calculations is not the right one? Jordan: That is possible. I am stuck with what the experimentalists give me. Roux: According to our calculations, the selectivity filter of the X-ray structure can sustain a flux. There is no big free energy barrier with that structure, and as far as we can tell the activity filter is not in a 'closed' state. Sansom: You should be able to test this. You could apply different levels of restraint to the X-ray structure, from rigid to soft restraints allowing a very large distortion. We know that if it is held rigid the barriers will be too great to sustain the observed flux, so imagine letting it soften more and more, and then determine at what stage the free energies become compatible with the experimental flux. I don't think you have to allow it to change greatly, but some deformability is needed. Perozo: It is clear that it doesn't need to be a great change. It cannot change that much because it is surrounded by all the transmembrane helices. The question is, to what extent do we let it change? Sansom: Despite the reservations one might have about simulations, we have seen a degree of flexibility in terms of things such as a flip round the valine of one of the carbonyls, for example (Shrivastava & Sansom 2000). We would expect some deformability of those carbonyls to track the ions as they move through the channel. Benoit Roux, have you calculated PMFs (potentials of mean force) with different degrees of rigidity of the filter? Roux: That is not quite how we did it, but there are certainly indications that the flexibility of the selectivity filter is important. For example, the Tyr78 (of the GYG signature sequence) is forming a hydrogen bond with Trp68 in the crystallographic structure. If the hydrogen bond is made artificially stronger by applying an energy restraint, then the free energy barriers for ion conduction are increased. What is remarkable is that this hydrogen bond is nearly 12 A away from where the ions are located, even though it has an impact on ion conduction. This is an example of a very delocalized and long-range effect from flexibility. It is certain that the

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flexibility of the protein will affect the free energy barriers controlling ion conduction. Ashcroft: What happens in your simulations if you stick an F in there rather than aY? Roux: We haven't done this, since it would weaken the hydrogen bond; it would probably flatten the free energy profile. Unwin: My understanding is that the flux of K+ through the KcsA channel isn't nearly as high as with some K+ channels, and also the selectivity is not as good. Has anyone got any insights as to what to do to the channels to make them more selective and flux at a higher rate? Perozo: Depending on the permeant ion concentration, Kcs A has a conductance as high as most channels, and it is also quite selective. It is just like any other K+ channel. Unwin: So the design is absolutely optimized for flux and selectivity. Perozo: Yes. There are many K+ channels that conduct poorly compared with KcsA. Sansom: Peter Jordan, going back to your breakdown of terms at the different sites, and accepting all these reservations about absolute magnitudes, if we add all those terms up, does it look flat or do you see preference for certain sites within the filter? Jordan: If it is singly occupied, I see a preference for ion occupancy of the water site. This is my site 3.1 dropped consideration of the two external sites because, in both those cases, the environment is part explicit solvent and part continuum solvent. If I have two ions in the filter, there is only one occupancy possibility because you are not going to have the two ions at neighbouring sites. Sansom: Can you break down DG to DH and TDS? Could you look at how things would change from room temperature to crystallographic temperature? Jordan: Yes.

References Marcus Y 1991 Thermodynamics of solvation of ions. Part 5. Gibbs free energy of hydration at 298.15 K. Faraday Soc Trans I 87:2995-2999 Pliego JR, Riveros JM 2000 New values for the absolute solvation free energy of univalent ions in aqueous solution. Chem Phys Lett 332:597-602 Reiss H, Heller A 1985 The absolute potential of the standard hydrogen electrode: a new estimate. J Phys Chem 89:4207-4213 Shrivastava IH, Sansom MSP 2000 Simulations of ion permeation through a potassium channel: molecular dynamics of KcsA in a phospholipid bilayer. Biophys J 78:557-570 Tissandier MD, Cowan KA, Feng WY et al 1998 The proton's absolute enthalpy and Gibbs free energy of solvation from cluster-ion solvation data. J Phys Chem 102:7787-7794

The b subunit of Kvl channels: voltage-gated enzyme or safety switch? Jacqueline M. Gulbis The Walter and Eliza Hall Institute of Medical Research, Post Office, The Royal Melbourne Hospital, Melbourne, VIC 3050, Australia

Abstruct. The b subunit of Kvl channels at first appears to be a quirk of mother nature — a redox protein permanently co-opted into a K+ channel assembly in the central nervous system. The N-terminal cytoplasmic domain of the channel, Tl, mediates its assimilation into the complex. Recent structural and biophysical characterization of the protein components of this assembly has been enlightening, but its principal physiological office is still in question. Here we re-examine the structural data with a view to providing a biological rationale for this unlikely partnership. A fresh interpretation of key structural features of b and Tl provides incidental evidence that the main channel gate in the transmembrane region can be subverted by the cytosolic assembly as part of a cellular response to oxidative stress. A hypothetical model in which the Tl-b interaction modulates the channel by controlling a secondary gate in the cytosol is offered as a plausible means by which feedback regulation of Kvl channels might be achieved. 2002 Ion channels—from atomic resolution physiology to functional genomics. Wiley, Chichester ( Novartis Foundation Symposium 245) p 127-145

Voltage-gated K+ channels are classified into families (Kvx) and subfamilies (Kvx.y) according to amino acid sequence. Higher organisms such as mammals possess a large multiplicity of Kv channels exhibiting differing patterns of expression. Although the number and type of Kv channels in cells might be expected to vary between discrete tissues or stages of development, the existence of so many members of this genre is nonetheless perplexing. Kv channels release K+ ions in response to membrane depolarization (or in some instances hyperpolarization), with little to discriminate between their individual conductivity profiles. An explanation for their natural diversity, therefore, is likely to reside in other aspects of their function. The cytosolic assemblies of Kv channels are optimally located to provide an interface with the cellular microenvironment. Cytosolic domains can differ markedly between families, and in some cases are tailored to assimilate cellular 127

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proteins into complexes at the membrane surface. An intriguing possibility is that interdomain contacts may mediate feedback responses between channel activity and cellular status, the fine-tuning perhaps occurring by differential channel expression. This paper reviews the current state of play with respect to the cytoplasmic assembly of Kvl (Shaker) channels, and seeks to provide an architectural framework for the interpretation of biophysical data. Kvl channels have a substantial cytosolic complex comprising the N- and C-terminal domains of a, an ancillary subunit (b), and short S2—S3 and S4— S5 linker segments. The b subunit associates exclusively with Kvl channels and is highly abundant in the mammalian central nervous system. Despite being long since acknowledged as an integral part of the assembly, no principal physiological role has yet been ascribed to b, although important auxiliary functions have been recognized. Two recent crystal structures of the b subunit revealed salient and unexpected features. Firstly, the tertiary fold of the b subunit exposed a novel member of the aldo-keto reductase (AKR) enzyme superfamily, in keeping with an emerging theme of redox-family proteins associated with ion channels. A later study, b in stoichiometric complex with the N-terminal domain (Tl) of K v la, clarified the nature of the interprotein interface and indicated how the cytosolic assembly might be juxtaposed relative to the membrane. These studies complement other recently published structures of the pore-forming unit (Doyle et al 1998) and the Tl domain (Kreusch et al 1998) in placing our knowledge of K+ channels onto a firm structural footing. Historical context Only three variants of b are presently known (b1, b2, b3). All have a conserved C-terminal core region of about 330 amino acids related in sequence to aldo-keto reductases (McCormack & McCormack 1994, Chouinard et al 1995). The N-termini are unrelated in either form or function. b subunits first drew serious attention in the mid-1990s for their capacity to inactivate open Kvl channels (Rettig et al 1994). This activity, however, is not a general feature of b subunits, but restricted to K v bl isoforms. A stretch of thirty or so residues at the N-terminus of b1 (an inactivation-particle or gate) directly contacts the channel in the vicinity of the pore, stemming the flow of K+ ions. The process is analogous to that in Shaker and K V 1.4 channels, which have intrinsic inactivation gates. It has also been proposed that b plays a role in promoting cell surface expression of Kvl channels (Shi et al 1996). In the normal course of events Kv channels are folded and processed in the endoplasmic reticulum, from whence they are transported to the plasma membrane (Nagaya & Papazian 1997). Co-assembly of K v la and b subunits has clear and quantifiable effects on surface expression. This is

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likely to be an indirect consequence of enhanced stability, or export competence, of the co-operatively folded a/? heteromer, although a specialised mechanism cannot be discounted (Manganas & Trimmer 2000). Although these phenomena are likely to represent select physiological advantages to the cell, neither is sufficiently general to be considered the primary role of j? in the absence of other data. The hypotheses also fail to account for some of the more interesting and conserved structural features of ft, which, unless it is simply an opportunistic protein hitching a ride with the channel to the cell membrane, must be integral to its function.

The fi subunit is an oxidoreductase The crystal structure reveals that the ft subunit (i) shares a common symmetry with the channel, and (ii) is a redox protein. The structure of the conserved core of rat Kv/?2 (residues 36-367) was solved by multiple isomorphous replacement methods and refined to a resolution of 2.8 A (Gulbis et al 1999). Each asymmetric unit of the crystal contains a ft tetramer with C4 point symmetry, congruent with the K + channel structure (Doyle et al 1998), in which four subunits are symmetrically disposed about a molecular fourfold. The two faces of the ft tetramer are manifestly different — one being extremely flat, whilst a large concavity defines the other. The flat face of each ft tetramer lies flush against that of a second, forming a ft octamer (Fig. 1A). Several methionine residues support the formation of a strong 'dimer' interface. Consistent with sequence homology, each ft monomer adopts the form of an eight-stranded a/?-barrel (TIM barrel) (Fig. IB) similar to the tertiary fold of aldo-keto reductases. A nicotinamide adenine dinucleotide phosphate (NADP) cofactor is bound to the protein with high occupancy. An unusually extensive array of hydrogen bonds and steric contacts constrain the NADP in an extended configuration within a long and narrow binding pocket. Finally, three catalytic residues (Tyr90, LysllS, Asp95) which, by analogy with AKR enzymes are deemed responsible for hydride transfer, are appropriately positioned and oriented relative to the cofactor (Fig. 1C). Together these points form a strong case for validation of ft as a redox-active enzyme.

Does ft have catalytic activity? To investigate this possibility we monitored the absorbance spectrum of ft to assess the oxidation state of the cofactor. Bound NADPH (reduced form) peaks at 363 nm, a slight red shift from the textbook value of 340 nm. Whereas the purified protein binds NADP in the reduced state, all crystalline forms of ft fail to absorb at 363 nm, indicating that the nicotinamide ring of the cofactor is oxidized

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FIG. 1. b subunit structure. (A) The b octamer forms by dimerization of b tetramers. The two flat faces pack closely together to yield a tight complex with 422 point-group symmetry. (B) Ribbon representation of a single K v b2 subunit, in similar orientation to the one denoted by an asterisk in Fig. 1 A. The central strands are shaded to accentuate the TIM barrel architecture. An NADP ligand (black) is positioned at the C-terminal edge of the barrel. (C) Close-up of the NADP binding site, illustrating the proximity of three catalytic residues (D85, Y90, K118) to the nicotinamide ring. This figure was generated with the program MOLSCRIPT.

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to NADP+ in the crystals. We were unable to crystallize b in the presence of reduced NADPH or to successfully soak the ligand into intact crystals. Whilst this may be due to an insufficient number of trials, it is equally likely to signify disruption of crystal packing contacts. This could arise from a rearrangement of protein elements about the NADP binding site to accommodate the reduced form of the cofactor, leading to local differences between the structures of b-NADP+ and b-NAPH. Although oxidation and subsequent re-reduction of bound cofactor (using free NADPH) in situ was observed by spectrophotometric means, it was not possible to verify that this was due to the catalytic activity of b rather than of a trace contaminant in the preparation. In these experiments, an array of small aldehydes and alcohols were used as potential substrates to assess the enzymatic turnover of b. Surprisingly few reagents were successful in accomplishing oxidation of NADPH, and even in those instances (e.g. benzaldehyde) the reaction occurred over a prolonged period (hours). UV spectra (280-400 nm) were almost indistinguishable from those of a control experiment where a putative non-catalytic mutant, b-Y90F was substituted for b, suggesting that a minor preparative contaminant and not b was likely to be responsible for catalysis. No reaction was detected in the absence of purified b (J. Gulbis, S. Mann & R. MacKinnon, unpublished work 1999). In summary, whilst the data indicate that NADP switches between oxidised and reduced states whilst bound in its pocket, and indeed appears permanently affixed therein, the experiments failed to demonstrate substrate reduction by b. In view of the tight association of b and channel in vivo, one explanation is that b is inactive as a free octamer. A second explanation is that it has a low turnover rate. In fact, the presence of an asparagine residue (N158) in mammalian b subunits suggests that a high local concentration of substrate is required for catalysis to occur. In AKRs a conserved histidine at the same position is involved in substrate binding and selectivity. Mutations of this histidine in aldose reductase decrease the catalytic efficiency by several orders of magnitude, lowering the rate constant and markedly increasing the Km (Bohren et al 1994). Tertiary fold — relationship of the b subunit to AKRs The tertiary fold of b bears remarkable resemblance to that of some AKR enzymes. A comparison of the backbone traces of b and a representative AKR, 3a-hydroxysteroid-dihydrodiol dehydrogenase (HSD; Bennett et al 1997) shows only two conspicuous differences (Fig. 2A). In each case this involves a serious perturbation of surface loop structure, but affects neither the overall topology of the protein nor its core catalytic features. Located in highly accessible regions of the b tetramer, the loops are of particular interest, being optimally positioned for

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FIG. 2. (A) The b subunit is structurally related to AKRs. (A) b (left) and HSD (right) are viewed in similar orientation from the C-terminal side of the barrel. The strands are shaded as in Fig. 1. Regions of structure that markedly differ between the proteins are distinguished in dark grey. There are two major features of interest. Specifically: (i) a protruding loop in HSD (*) interfaces with the C-terminus, and (ii) a second loop located almost directly above the active site has been extended and reconfigured in b (**) to cap the NADP binding pocket. (B) Suggestive homology in two critical regions: (i) the T1-S1 linker to the 'active site loop' of HSD, and (ii) C-termini of b and HSD.

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interaction with other proteins. One such disparity almost entirely submerges the cofactor of b beneath the molecular surface, whereas in HSD the NADP is comparatively exposed. Here the elaboration of a short connecting loop in HSD to an extended and convoluted version in b forms a large contusion in the surface that encases the NADP binding site. This 'capping' loop has by far the highest mobility, based on experimentally observed thermal parameters, of any in b. A second, and perhaps more striking, distinction between the two structures occurs in the vicinity of the enzyme active site. The catalytic centre of HSD is protectively shielded from surrounding solvent as is usual for oxidoreductases, whilst that of ft atypically gapes open. This change reflects a difference in the relative positioning of the C-terrninal tail of each protein. A protruding elongated loop having no counterpart in ft interfaces with the terminus of HSD to encircle the reaction centre. Both loop and C-terminus contribute aromatic side chains to the enclosed steroid-binding site of HSD. In b such an interface cannot occur as its C-terminus has retracted too far to reach the active site. Consequently the equivalent surface of ft resembles a large open concavity, and has a correspondingly sparse population of substrate-binding residues. How might these structural anomalies, conserved in b subunits, be translated into functional attributes? In a recent paper (Gulbis et al 1999) the authors propose that ft activity is voltage-dependent, or, in other words, that its activity is intrinsically coupled to that of the voltage-gated a subunit, or channel. In fact these two proteins form a close and permanent partnership, insinuating a mutual raison d'etre. The ion-conducting path of a resting channel is poised to open in response to membrane depolarization. Movement of the voltage sensor in the electric field triggers a conformational change in the transmembrane domains, gating the pore. The proposal entails harnessing this gating movement to drive cytosolic redox chemistry or, conversely, exploiting local conformational rearrangements of b to transduce sensory information to the channel. The two structural differences outlined above are appealing because they immediately suggest means by which conformational switches in the proteins might be activated. Referring to the first example, the prominent 'capping' loop has no counterpart in HSD, rendering it unlikely that it is required for redox activity. It is, however, intimately connected with the NADP molecule by several conserved amino acids in the region 253-264. It is not implausible that small effects on cofactor conformation upon reduction of the nicotinamide moiety could be amplified and transmitted to a conjoined protein via this mobile loop. The redox status of NADP would thus act as a barometer, utilising ft to relay cellular signals to the membrane. A similar structural effect was recently proposed for rhodopsin, in which a change in the state of a retinal ligand drives the movement of a proximal loop (Subramaniam & Henderson 2000). Secondly, the b subunit appears to have an incomplete scaffold sheltering its active site. In generic

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TIM barrel enzymes the reaction centre is bounded by the eight loops at the carboxy edge of the barrel connecting each strand to the ensuing helix. Some exceptions occur, in which an adjacent domain or subunit contributes a loop to complete the active site. One might reasonably speculate that the elongated, protruding loop interacting with the C-terminus of HSD to enclose its active site could in practice be substituted in b by a loop proffered from elsewhere in the channel assembly. This implicitly argues for a functional interconnection between a and b. In support of this, it is interesting that in the C-terminal region, sequence alignments of HSD and b are inconsistent with three-dimensional structural superpositions. Although in the crystal structures ft appears to have a diminished C-terminal extension relative to HSD, conventional line-ups show that they are not only of similar length, but have significant homology at the extremity — especially at those few residues that facilitate interaction with the active site loop in HSD. Suggestive homology is also evident between the linker connecting Tl to the first transmembrane segment (S1) and the protruding loop of HSD (Fig. 2B). Furthermore, three arginines (R129, R159, R189) line a crevice at the periphery of the b tetramer that leads directly to the active site. They impart a substantial positive charge to the molecular surface, perhaps targeting acidic residues of the T1-S1 linker. Hence there are a number of indicators that the open scaffolding is a feature of ft structure only when it is expressed in the absence of its natural partner. The Tl-b assembly Kvl channels in mammalian brain tissue are invariably found associated with b subunits. The Tl domain of a mediates the interaction, and together with $ forms the major part of the cytoplasmic complex. In order to explore the physical nature of the assembly, constructs of Tl (1-135) and b (36-367) were coexpressed in SF9 cells, purified as a stoichiometric complex and crystallized. The structure was solved by molecular replacement methods employing a b tetramer as the search model and refined against data complete to 2.1 A (Gulbis et al 2000). In the Tl4-b4 hetero-octamer, four prominent loops of Tl employ surface hydrophobic residues to tether the Tl domain to the flat face of the b tetramer (Fig. 3A). The interface involves a number of the same residues mediating octamerization of free b (Fig. 3B), confirming the exclusive nature of a b-binding partnership. Tl— b has a central conduit running along its entire length that is sufficiently wide at the apex to permit the entry of ions. The lining of this passage consists predominantly of charged and polar amino acid side chains. An opening at the distal face of Tl widens to form a concealed chamber that sustains a negative electrostatic potential due to the presence of four equivalent buried aspartate

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FIG. 3. Comparative anatomy of (A) the T14-b4 complex, and (B) the b octamer (side view). Four conserved loops tether Tl (dark grey) to the flat face of a b tetramer.

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residues, apparently without ion-pairing partners. Four symmetrically disposed apertures in the surface of Tl, sufficiently wide to allow the insertion of large side-chains or divalent ions, tunnel directly into this curious chamber which presumably acts as a sink for positively charged species. A comparable cavity is found in the structure of a KV3 Tl domain (Bixby et al 1999), indicating that it may serve a general function in voltage-gated ion channels. The remainder of the central passage maintains a positive surface potential. Initial comparison of the Tl—b complex with crystal structures of Tl and b revealed only minor differences. To illustrate, a backbone trace of the free b subunit is virtually superimposable upon its counterpart in the complex. At first glance Tl also seems essentially unchanged, but closer inspection exposes small but significant departures in its tertiary and quaternary structure which may have a bearing on the electrophysiological attributes of Kvl channels. The a helix of Tl immediately preceding the T1-S1 linker, for example, deviates from helical periodicity significantly earlier than in published structures of uncomplexed Tl (Protein Data Bank entry 1T1D). Furthermore, relative to the free state, all four Tl protomers in the complex are slightly inclined with respect to a central fourfold. The net effect is to separate individual subunits on the (putative) membrane side, thereby enlarging the cavity entrance by approximately 2.5 A (Fig. 4A). This tilted conformer is reminiscent of crystal structures of Tl mutants that occasion a small leftward shift in the voltage-activation curves of Kvl channels (Cushman et al 2000). In fact b subunits themselves moderately affect the gating properties of Kvl channels, inducing a comparable negative shift in activation voltage (Heinemann et al 1996). It may be pertinent to add that actual current level is not significantly affected by mutations in, or indeed deletion of (Kobertz & Miller 1999), the Tl domain, or by the presence of b. The resemblance of the destabilizing Tl mutants to the conformer observed in the Tl—b assembly provides an explanation for the observed effects of ab co-expression on channel gating. Whereas the Tl-b complex explicitly defines the relative orientations of its protein components, it does not discriminate between alternate models (invertomers) for juxtaposition of cytosolic assembly and membrane (Fig. 4B). The confusion arises because the structured Tl domain is linked to the first transmembrane segment (S1) by a sufficiently long chain of amino acids to allow for two possible contingencies, both of which align the tetrad axis with the pore. The result of an electrophysiological assay employed to resolve this issue makes a persuasive argument for an arrangement in which Tl is sandwiched between and the channel (Gulbis et al 2000). In this model T1-S1 linkers form spacers, ensuring that the cytosolic structure does not impede the flow of cations to the open channel. It is consistent with a Tl—b 'hanging basket' suspended beneath the membrane (Fig. 5A), in similar fashion to the cytosolic vestibule of the nicotinic

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FIG. 4. Assembling the channel. (A) The Tl domain undergoes a conformational change as a consequence of binding b. Each Tl subunit is tilted with respect to a central fourfold axis, widening the main aperture to the inner cavity. The structure of the terminal helix is also disrupted. (B) The two possible ways in which a Tl—b assembly may be oriented with respect to the membrane are depicted schematically.

acetylcholine receptor (AChR) from Torpedo (Miyazawa et al 1999). A preponderance of acidic residues in the Tl—S1 linker may also be akin to AChR, where negatively-charged protein spacers appear to act as a prefilter, repulsing anions from entering lateral openings that surround the pore entrance. The T1-S1 linker: entrance portal or redox active-site? We are left, seemingly, with a paradox. Biophysical evidence indicates a rigid structural relationship between the Tl and transmembrane domains in which Tl—S1 spacers play a key part, enabling the flow of K+ ions to the pore entrance. Structural considerations, conversely, would appear to favour an intimate connection between b and the channel. This again involves the Tl—S1 linker, but in this instance as a functional component if the active site (Fig. 5A). Modelling T1-S1 into the assembly demonstrates that the important features of this arrangement mimic the active site of HSD. There is a proviso however; the

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K V 1 b SUBUNIT

139

FIG. 5. (A) Two possible roles for T1-S1. (B) A hypothetical model portraying a two-gate mode of Kvl channel regulation. When the Kvl pore is closed the T1 domain lies against the membrane, the entrance to its inner vestibule aligned with the pore. Four orifices in the T1 surface descend into the cavity, where unpaired aspartate residues impart a negative electrostatic potential to the inner surface. Each is occupied by a positively charged side chain arising from the S4-S5 loop separating the pore-forming and voltage-sensing domains, tightly securing T1 to the membrane surface. This effectively closes a cytosolic 'gate'. The b subunit is poised for catalysis, its reaction centre protected by interaction of its C-terminus with the T1-S1 linker. In this model, hydride transfer from NADP to a substrate initiates a cascade of events that instigate channel opening. A change in the configuration of NADP is transmuted to a large conformational adjustment of the 'capping' loop. This disrupts the interaction between b and T1-S1, incapacitating b. Subsequent retraction of each T1—S1 linker into structured 'spacers' distancing T1 from the membrane and causing the basic 'plugs' to disengage from T1 opens a peripheral cytosolic gate to K+ ions. This triggers four S4 voltage sensors to move cooperatively outwards within the electric field of the membrane, opening the main gate of the channel. When the main gate closes the same four residues reinsert into T1, disassembling the spacers. The T1 S1 loop is again available to form a functional active site in b, although this is contingent upon reversal of the oxidation state of captive NADP molecules — presumably as an outcome of the refreshed cellular environment.

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mechanism, this process triggers a subsequent S4 movement cannot be ascertained from this study. The model does, however, account for a number of key observations, (i) Redox activity of b in isolation is minimal or absent, (ii) NADP oxidation appears to invoke local conformational changes in b. (iii) The terminal helix of Tl is no longer a structured element in the T1-b complex, (iv) Short but significant sequence homology occurs between active-site elements of HSD and b/Tl-Sl segments, (v) The negative surface of Tl's inner vestibule is conserved in Kv families. (vi) S4 gating involves a stable intermediate state between the independent and cooperative transitions.

Opening Pandora's box Does this represent an efficacious bypass to open Kv channels in oxidative stress conditions? Is a two-gate mechanism feasible? Does it concur with biophysical experiments? And can these ideas be equally applied to other families of Kv channels? Perhaps the major contribution of this review is to provide an informed backdrop to future experimentation.

Acknowledgements I thank Roderick MacKinnon, in whose laboratory the structural work was carried out, for his generous patronage. My appreciation extends to the members of his group: to Ming Zhou for collaborative efforts, and especially to Sabine Mann for her tireless assistance and companionship. Many faculty, postdocs, students and technicians at the Rockefeller University provided assistance and support during my term there, including John Kuriyan, Brian Chait and members of their respective laboratories. James Trimmer provided the b subunit DNA, and staff at the BNLS and CHESS synchotrons willingly gave of their time. Particular thanks go to Declan Doyle, Elena Conti, Xiao Dan Li and Tony Auerbach for their help and insight, and to Peter Colman, Tom Garrett and Brian Smith for constructive advice on the manuscript.

References Auerbach A, Akk G 1998 Desensitization of mouse nicotinic acetylcholine receptor channels. A two-gate mechanism. J Gen Physiol 112:181-197 Baker OS, Larsson HP, Mannuzzu LM, Isacoff EY 1998 Three transmembrane conformations and sequence-dependent displacement of the S4 domain in shaker K+ channel gating. Neuron 20:1283-1294 Bennett MJ, Albert RH, Jez JM, Ma H, Penning TM, Lewis M 1997 Steroid recognition and regulation of hormone action: crystal structure of testosterone and NADP+ bound to 3 alphahydroxysteroid/dihydrodiol dehydrogenase. Structure 5:799—812

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Bixby KA, Nanao MH, Shen NV et al 1999 Zn2+-binding and molecular determinants of tetramerization in voltage-gated K+ channels. Nat Struct Biol 6:38-43 Bohren KM, Grimshaw CE, Lai CJ et al 1994 Tyrosine-48 is the proton donor and histidine-110 directs substrate stereochemical selectivity in the reduction reaction of human aldose reductase: enzyme kinetics and crystal structure of the Y48H mutant enzyme. Biochemistry 33:2021--2032' Chouinard SW, Wilson GF, Schlimgen AK, Ganetzky B 1995 A potassium channel b subunit related to the aldo-keto reductase superfamily is encoded by the Drosophila hyperkinetic locus. Proc Natl Acad Sci USA 92:6763-6767 Cushman SJ, Nanao MH, Jahng AW, DeRubeis D, Choe S, Pfaffinger PJ 2000 Voltage dependent activation of potassium channels is coupled to Tl domain structure. Nat Struct Biol 7:403-407 Doyle DA, Morais Cabral J, Pfuetzner RA et al 1998 The structure of the potassium channel: molecular basis of K+ conduction and selectivity. Science 280:69-77 Gulbis JM, Mann S, MacKinnon R 1999 Structure of a voltage-dependent K+ channel b subunit. Cell 97:943-952 Gulbis JM, Zhou M, Mann S, MacKinnon R 2000 Structure of the cytoplasmic b subunit- Tl assembly of voltage-dependent K+ channels. Science 289:123-127 Heinemann SH, Rettig J, Graack HR, Pongs O 1996 Functional characterization of Kv channel b-subunits from rat brain. J Physiol 493:625-633 IsacoffEY, Jan YN, Jan LY 1991 Putative receptor for the cytoplasmic inactivation gate in the Shaker K+ channel. Nature 353:86-90 Kobertz WR, Miller C 1999 K+ channels lacking the 'tetramerization' domain: implications for pore structure. Nat Struct Biol 6:1122-1125 Kreusch A, Pfaffinger PJ, Stevens CF, Choe S 1998 Crystal structure of the tetramerization domain of the Shaker potassium channel. Nature 392:945-948 Manganas LN, Trimmer JS 2000 Subunit composition determines Kvl potassium channel surface expression. J Biol Chem 275:29685-29693 Mannuzzu LM, Isacoff EY 2000 Independence and cooperativity in rearrangements of a potassium channel voltage sensor revealed by single subunit fluorescence. J Gen Physiol 115:257-268 Miyazawa A, Fujiyoshi Y, Stowell M, Unwin N 1999 Nicotinic acetylcholine receptor at 4.6 A resolution: transverse tunnels in the channel wall. J Mol Biol 288:765-786 McCormack T, McCormack K 1994 Shaker K+ channel bsubunits belong to an NAD(P)Hdependent oxidoreductase superfamily. Cell 79:1133-1135 Nagaya N, Papazian DM 1997 Potassium channel oc and b subunits assemble in the endoplasmic reticulum. J Biol Chem 272:3022-3027 Rettig J, Heinemann SH, Wunder F et al 1994 Inactivation properties of voltage-gated K+ channels altered by presence of b-subunit. Nature 369:289-294 Shi G, Nakahira K, Hammond S, Rhodes KJ, Schechter LE, Trimmer JS 1996 b subunits promote K+ channel surface expression through effects early in biosynthesis. Neuron 16:843 852 Subramaniam S, Henderson R 2000 Molecular mechanism of vectorial proton translocation by bacteriorhodopsin. Nature 406:653-657

DISCUSSION Ashcroft: My understanding is that not all of the b subunits confer redoxsensitive inactivation on voltage-sensitive K+ channels. I thought that some b

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subunits were not redox sensitive. What is the difference between those that are and those that aren't? Do they have a binding site? Gulbis: I imagine that all b subunits would be redox sensitive. They have all the attributes. There was some controversy over b3, which has some sequence differences. I don't think enough is really known about this. Ashcroft: There is always the possibility that there is a difference between binding and transduction. They might all have the same structure but just not transduce it in the same way. Gulbis: I guess they could do. I would imagine they all have a similar structure. I don't think the NADP ever comes out of that binding site. We couldn't wash it out, and we tried hard. The cofactor is nrobably an integral part of b subunit structure. As I understand it, recombinant material produced in bacteria fails to fold correctly — possibly because the NADP fails to incorporate. Papazian: I am curious about the interface between the subunit and Tl. Is this very extensive? How difficult would it be to engineer an interface between one of the a keto reductases that is very enzymatically active and the T1 domain? This would enable you to see whether enzymatic activity has some gating effect. Gulbis: Firstly, Olaf Pongs (Bahring et al 2001) recently reported effects of the enzymatic activity of b on gating. His results did suggest a relationship, although I found it a little difficult to interpret his data as he used chimeras rather than site mutations. Secondly, whilst the interface between ft and Tl is limited to a single loop on each T1 subunit that contacts the flat face of the b tetramer, an extensive array of polar and water-mediated contacts aids in cementing the two proteins together. This might prove difficult to replicate in a T1-AKR complex. Incidentally, the interacting loops of T1 do seem to be conserved throughout Kvl families. It is likely that they serve a general role as a protein-binding interface in Kv channel biology. Swartz: Are those regions of the b subunit that interact with T1 conserved? Gulbis: No, the surfaces are different. Choe: I thought that the model of the C-terminal segment of Tl reaching down was very refreshing. Have you tried to extend your T1 construct a little and see whether it forms a more stable complex? Gulbis: I hadn't developed the gating ideas when I was in Rod MacKinnon's lab, although I did note the sequence homology between the T1-S1 linker and the protruding loop of 3a-hydroxysteroid-dihydrol dehydrogenase (HSD). At that time I had a T1-S1 peptide synthesized and tried to show interaction with the free b subunit. It failed dismally. Perhaps in the free b octamer, residues that are accessible in the channel assembly are unavailable for interaction. I never tried this experiment on the complex.

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Mindell: Your model predicts that you can have exquisite length dependence of that linker. Getting rid of any length of that linker would have big effects. Gulbis: It depends on what part of the linker is removed. Mindell: Has anyone tried this? Gulbis: Not as far as I know. Swartz: There is work in Chris Miller's lab in which they deleted T1 (Kobertz & Miller 1999). There is nothing really deficient in those channels. Gulbis: I realise that. I don't think this is inconsistent with my model. I am not saying that this is how the channel always gates. I'm suggesting that the channel undergoes normal depolarization-elicited channel opening. However, in the event that the cell is redox-stressed, b provides a switch to open the channel in the absence of a depolarizing stimulus. Swartz: So you are talking about a modulation of the fundamental gating properties? Gulbis: Yes. Schrempf: You cannot exclude that there are other proteins in the cascade, not directly interacting with your b subunit. Gulbis: That's right. For example the C-terminal domain of the channel must be in the immediate vicinity, in all probability contacting the T1—S1 linker. The model I proposed in my talk is simply one way of putting our structural data into a context that is consistent with the known electrophysiological behaviour of Kvl channels. And it does provide a rationale for having redox proteins attached to Kv channels. Mathie: Perhaps I have misunderstood you, but in your model, when you have active b there was no ion permeation pathway and you couldn't then get normal gating of the channel. Gulbis: What I am suggesting is that when b is active the channel is closed. If you have evidence to the contrary then you should say so. Mathie: Not at all! But the only way that the channel could work with its normal voltage gating is if b wasn't active. Is that correct? Gulbis: No. What I mean is that whilst voltage-gating could occur when b is in an 'active' configuration, the actual process of gating would disrupt this and render b inactive. Choe: I have a question about the octameric packing. It is kind of mind boggling. Does this imply that the Kb exists as an octamer in the cytoplasm? Gulbis: In the normal course of events nascent b polypeptides translocate, along with the channel a preproteins, into the endoplasmic reticulum (ER), where folding and processing occur. Octamer formation may be a means of 'removing' excess b from circulation, and possibly only happens during overexpression. At first we thought the b-octamer was a crystallographic artefact,

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but changed our minds after two or three crystal structures of free b revealed an identical octamer. Choe: If Kb stays as an octamer as an isolated protein, this suggests that it won't be able to interact with the channel unless they are co-complexed in the ER lumen during synthesis. Gulbis: Yes. The inter-protein interface is, as stated earlier, of a somewhat tenuous nature. We were unable to make a T1—b complex from purified component proteins, except in buffers lacking salt. The complex we made in this fashion appeared stoichiometric, but disintegrated with the addition of 25 mM salt. To obtain a stable complex we had to co-express the b and T1 domains in SF9 cells. This is because each b tetramer has a higher affinity for another b tetramer than for T1 —which shares a common binding interface. Choe: What if you inject isolated Kb protein into Xenopus oocytes? Gulbis: I don't know. Papazian: Since we are talking about the possible states of b in the cell, I'll just mention an observation we made some time ago. We were looking at the issue of assembly between as and bs. There is nice work from Jim Trimmer's lab that shows that for some of the mammalian Kv channels that don't get to the cell surface very well, their co-expression with b increases the cell surface expression. We observed something on the flip side of this, too. We saw in mammalian cells that Kb was expressed at high levels but was quite unstable. It seemed to be only the fraction that associated with the a subunit (in this case Shaker), that was stable over significant periods. It is an extrapolation, but one might propose from this kind of result that in fact association with a is the main thing that b does, and that in the absence of this interaction it is not a very stable protein. Sansom: Are you implying that the Kb is a chaperone for the a subunits, rather than a modulatory protein? Papazian: There is evidence, from Trimmer's lab, that some Kv a subunits don't get to the surface well unless b is expressed. Whether bs are chaperones or not depends on your definition of a chaperone. I am suggesting that the issue of whether Kvb is a tetramer or an octamer in cells may be biologically irrelevant. It doesn't seem to be very stable unless it assembles with a. Ashcroft: The same thing is seen with KATP channel. Neither the Kir6.2 subunit nor the SUR subunit will get to the surface in the absence of the other one (Zerangue et al 1999).

References Bahring R, Milligan CJ, Vardanyan V et al 2001 Coupling of voltage-dependent potassium channel inactivation and oxidoreductase active site of KVb subunits. J Biol Chem 276:22923-22929

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Kobertz WR, Miller C 1999 K+ channels lacking the 'tetramerization' domain: implications for pore structure. Nat Struct Biol 6:1122-1125 Zerangue N, Schwappach B, Jan YN, Jan LY 1999 A new ER trafficking signal regulates the subunit stoichiometry of plasma membrane K ATP channels. Neuron 22: 537-548

EPR approaches to ion channel structure and function E. Perozo, L. G. Cuello, D. M. Cortes, Y.-S. Liu and P. Sompornpisut Department of Molecular Physiology and Biological Physics, and Center for Structural Biology, University of Virginia, Charlottesville, VA 22908, USA

Abstract. The fundamental processes that underlie ion channel function are permeation, selectivity and gating. In an effort to understand ion channel gating, we have used an approach that combines reporter-group spectroscopic techniques (spin labelling/ electron paramagnetic resonance, EPR) and electrophysiological methods with classical biochemical and molecular biological procedures. As an ideal test channel, we have focused our attention on the K+ channel from Streptomyces lividans, KcsA. Through sitedirected spin labelling, cysteine chemistry was used to introduce nitroxide radicals into specific sites within KcsA with high reactivity and specificity. EPR spectroscopy analysis of the spin labelled mutants yields two types of structural information: (1) mobility and solvent accessibility of the attached nitroxide through collisional relaxation methods and (2) distances between pairs of nitroxides through dipole-dipole interactions. Using this approach, we analysed the correlation between KcsA crystal structure and the EPR data, extend it to derive low-resolution folds of full-length KcsA and apply it in the determination of the molecular rearrangements responsible for pH-dependent gating. 2002 Ion channels—from atomic resolution physiology to functional genomics. Wiley, Chichester (Novartis Foundation Symposium 245) p 146-164

In recent years, use of electron paramagnetic resonance (EPR) spectroscopy together with site-directed spin labelling (SDSL) has opened up the possibility of pursuing real structural studies in systems that, because of size or physical-chemical properties (or both, like membrane proteins), have remained refractory to more traditional methods (Hubbell & Altenbach 1994, Hubbell et al 2000, 1998, Mchaourab & Perozo 2000). Using traditional site-directed mutagenesis methods, it is possible to introduce cysteines at practically any position within a protein sequence. These engineered cysteines are the targets to which a nitroxide spin-label is subsequently attached through a covalent thio-ether bridge. Figure 1A shows the most common thiol-specific spin label that has made possible these advances. Spin label I is a methanethiosulfonate derivative, a highly reactive chemical that has been successfully used in numerous spin labelling experiments 146

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FIG. 1. The nitroxide side chain and its spectrum. (A) Reaction of the methanethiosulfonate spin label (MSTSL) with a free cysteine. (B) Typical EPR spectrum in the intermediate motional regime. The small horizontal bar represents the width of the central resonant line (m = 0), the long vertical bar, its amplitude.

(Berliner et al 1982). Compared with fluorescent probes, nitroxides are quite small: the estimated volume of a nitroxide is slightly larger than a tryptophan. Here we will summarize recent progress on the use of SDSL and EPR spectroscopy in the analysis of structure—function relations in K+ channels. We

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shall briefly discuss the basic principles of the approach, its use in deriving structural information and the structural rearrangements responsible for channel gating.

EPR principles and data Figure 1B shows a typical EPR spectrum from a slow tumbling protein. These spectra are represented as the first derivative of the microwave absorption versus the magnetic field strength, and derive from the reorientation of unpaired electrons within an external magnetic field upon absorption of radiation. Analysis of spinlabelled molecules yields three types of structural information: (1) nitroxide dynamics from spectral line-shapes, (2) solvent accessibility of the attached nitroxide through collisional relaxation methods and (3) interspin distances between pairs of nitroxides through dipole—dipole interactions. The first two are local parameters interpreted in terms of structure in the vicinity of the probe, while inter-nitroxide proximities reflect global features of the backbone fold (see Hubbell et al 1998, Mchaourab & Perozo 2000). In a spin-labelled protein, the relation between the spectral line-shape, local structure and side chain dynamics is extremely complex, given the number of possible rotations around the label's attachment bonds to the protein backbone. Still, spin label dynamics (mobility) can be quantified through the lineshape parameter DHo, or the spectral second moment (Mchaourab et al 1996). Both are semi-empirical measures derived from the spectral line-shape, determined by the degree of averaging of the anisotropic g and hyperfine tensors respectively (their value increases as the frequency and amplitude of nitroxide motions is reduced). Mobility measurements are particularly useful, since they allow the classification of a spin-labelled site into one of three structural classes: exposed, buried or in tertiary contacts (Columbus et al 2001, Mchaourab et al 1996). Collision of nitroxides with paramagnetic fast relaxing reagents, such as oxygen and metal ion complexes, causes Heisenberg spin exchange that in effect, shortens the spin-lattice relaxation time T1 of the nitroxide (Hyde & Subczynski 1989). This phenomenon can be measured either by continuous wave power saturation techniques or by saturation recovery methods. Figure 2A illustrates the principle of the power saturation approach. Using T4 lysozyme embedded in a saturated O2 solution as an example (left panel), we placed spin labels either at the core of the molecule (position 1) or in its surface (position 2), and spectra were recorded at increasing microwave powers. If O2 molecules are unable to directly collide with the spin label (position 1), signal saturation occurs at a similar power as in the absence of the relaxing radicals (as in an N2-saturated solution). Conversely, a fully accessible spin label requires significantly higher microwave power to saturate (Fig. 2A, bottom right panels). Combination of accessibility data with

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FIG. 2. Structural parameters derived from spin-labelled systems. Examples are given from spin-labelled T4 lysozyme. (A) Solvent accessibility is obtained from power saturation experiments. Two positions chosen for their solvent exposure. Position 1 is buried in the protein core and shows saturation at low powers. Position 2 is fully exposed and shows saturation at high powers. In each case the central resonant line is obtained in the presence of a paramagnetic collision probe (i.e. O2), and in a N2 background. (B) Inter-spin distance determinations. Examples are given under dynamic conditions (room temp.) or in the rigid limit (~140K). The top spectra represent the sum of individually labelled protein (no spinspin interaction). The lower spectra were obtained from the double mutant (full spin-spin interaction). Mutants kindly provided by M. Mchaourab.

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spin label dynamics can be used effectively to determine the overall location of a specific residue in a soluble protein fold, or within the membrane protein-water interface for membrane proteins. Quantitative information on inter-spin distances can be obtained from dipole— dipole interactions between nitroxide groups in a system of two spin labels (either in a symmetric system or through the use of double mutants). In general, the dipole-dipole interaction is a function of the spatial separation of the two nitroxides, their relative orientation, and the frequency and amplitude of the motion averaging the orientation of the spin vector relative to the magnetic field. As a general rule, the broadening of the EPR spectrum when compared to the reference state of the spectral sum of the single mutants can be interpreted semi quantitatively to rank distances into three broad categories: close (<10A), intermediate (10-15 A) and far (15-20 A). Direct estimation of distances is typically achieved under rigid limit conditions (low temperature), as has been shown by a number of groups (Hustedt et al 1997, Mchaourab et al 1997, Rabenstein & Shin 1995, Steinhoff et al 1997). These dipolar interactions are reflected as a general broadening of the spectrum (shown in Fig. 2B under dynamic and rigid-limit conditions), and the extent of this broadening can be used to estimate actual distances with some precision (Rabenstein & Shin 1995). In general, any structural conclusion originating from the determination of interspin distances should be based on the presence of patterns of proximities rather than single distances. This way, the impact of the uncertainty in the position of the nitroxide ring relative to the a carbon is minimized. Site-directed spin label analysis of KcsA The Streptomyces lividans K+ channel (KcsA) is a 160-residue protein that forms homotetrameric channels closely related to the pore domain of larger voltagedependent channels (Schrempf et al 1995). When purified and reconstituted in lipid bilayers, KcsA catalyses single channel activities with selectivity properties identical to those of other eukaryotic K+ channels (Cuello et al 1998, Heginbotham et al 1998, Schrempf et al 1995). The fact that KcsA is easily expressed in Escherichia coli at milligram levels made this protein an ideal target for structural analysis. Crystallographic studies led to the high-resolution structure of KcsA by Doyle et al (1998), an achievement that lent a firm structural foundation to more than three decades of functional work on K+ channels. The crystal structure revealed that KcsA is formed by the association of four subunits, contributing equally to form a water-filled pore. Each subunit has two transmembrane segments, TM1 in the periphery of the complex, and TM2, lining the permeation path. Towards the extracellular face of the channel is the selectivity filter, where intimate contact with the permeant ions takes place.

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With the idea of obtaining structural dynamic information of the two transmembrane segments, we undertook an extensive SDSL study of KcsA in its native environment (Perozo et al 1998). Sixty-six single cysteine mutants were prepared and analysed for residues 22—52 and residues 86-120 of KcsA, where the two putative transmembrane segments are thought to be located. Using discrete Fourier transform methods (Cornette et al 1987, Donnelly et al 1994), angular frequency information was extracted from residue environmental profiles from each of the scanned protein segments and used in the assignment of secondary structure elements (Perozo et al 1998). Solvent accessibility information derived from either O2 or NiEdda collision frequencies was in agreement with the crystal structure, suggesting that TM1 has a large lipid-exposed surface, while TM2 is more protected from membrane lipids. An excellent example of the use of solvent accessibilities to describe unknown structures is found in the recent SDSL analysis of KcsA cytoplasmic domains (Cortes et al 2001). Because of the requirements specified by the successful crystallization conditions, no structural information exists on these regions of KcsA (Doyle et al 1998). Thus, we probed regions in the N- and C-termini of KcsA predicted to be soluble and exposed to the cytoplasm by site-directed spin labelling. The results can be viewed in the form of a residue environmental phase diagram by plotting Ni accessibility (ONiEdda) versus O2 accessibility (IIO2, Fig. 3A). The structural phases can be classified as aqueous (high ONiEdda and low IIO2), lipid exposed (high IIO2 and low IINiEdda) and buried (low IINiEdda and low IIO2). From these diagrams it is clear that the N-terminus region of KcsA resides at the membrane interface (straddling both membrane exposed and water exposed regions, Fig. 3A, top), while the C-terminus is wholly exposed to the aqueous environment (Fig. 3A, bottom). Using this data set, together with the known coordinates from the crystal structure, we were able to develop a backbone three-dimensional model of full-length KcsA (Fig. 3B), to complete the structural description of KcsA with the membrane as a reference point, and to provide a structural basis for understanding its mechanism of pHdependent activation gating. Spin-spin interactions have played a key role in the structural analysis of KcsA and in the molecular description of its activation mechanism (see below). Due the tetrameric stoichiometry of KcsA, spin—spin coupling can originate either from interactions between next-neighbours or from diagonally related subunits (Fig. 4A, only the diagonal subunits are shown). Thus, in an ideally labelled KcsA, four spin labels would be contributing to the spectral broadening (Fig. 4B). Under these conditions, a qualitative estimate of the magnitude of spin—spin interactions can be obtained from the ratio of amplitudes of the central resonance line (M = 0) between the under-labelled and fully labelled mutant, both normalized to the total number of spins in the sample (Perozo et al 1998). This

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interaction parameter, W, was used to survey patterns of spin coupling along the length of TM2 and to deduce possible inter-helical orientations. Strong broadening was in fact observed every three or four residues on one face of the C-terminal half of TM2 (in residues Al08, T112, VI15 and G116), as shown from the comparison between their spectra at underlabelling conditions (Fig. 4B, left) and those obtained from fully labelled channels (Fig. 4B, centre). In agreement with the relative inter-subunit angle between TM2 segments (Fig. 4A), significantly weaker spin—spin coupling (as measured from the W parameter) was detected on the N-terminal half of the helix (Fig. 4C). Similar results have been obtained in a parallel SDSL analysis of KcsA TM2 (Gross et al 1999). KcsA gating mechanism The structure of KcsA can be used as a remarkable blueprint to explain the principles of ion permeation (Doyle et al 1998). The gating mechanism, however, is harder to deduce from direct examination of the KcsA structure. The discovery that KcsA can be activated by changes in proton concentration (Cuello et al 1998) set the stage for the analysis of the types of protein motions that underlie activation gating in K+ channels. It was shown, using both radiotracer flux experiments and single channel current measurements in planar lipid bilayers, that the equilibrium between open and closed states in KcsA could be shifted towards the open state by lowering pH. Recent studies clearly show that channel opening is induced by intracellular changes in pH, and that a pH gradient is not necessary for channel activation (Heginbotham et al 1999). Earlier results from voltage dependent channels suggested that channel opening was associated to global conformational changes in the intracellular face of the channel (Armstrong 1971, Liu et al 1997). The first direct demonstration of the types of movements that may be associated with channel opening came from our lab (Perozo et al 1998) as large pH-dependent changes in inter-subunit spin-spin coupling were observed at selected residues along the C-terminal end of TM2 (Fig. 4B,C). From the crystal structure, these residues were shown to be located at the narrowest section of the internal helix bundle, close to the fourfold axis of symmetry (See Fig. 4A). In each case, spectra obtained at low pH showed a reduction in the magnitude of the inter-subunit dipolar coupling (Fig. 4B, right), leading to the suggestion that channel opening was a consequence of a widening in the inner vestibule of the channel. A more thorough analysis of these movements was subsequently carried out by studying changes in mobility and dipolar coupling along the main structural elements in the channel: the two transmembrane segments TM1 and TM2, as well as the regions flanking the selectivity (Perozo et al 1999). The changes in spectral line shape were analysed using discrete Fourier transform methods to

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detect possible patterns that could indicate specific types of movements in each of the TM segments originating from pH acidification. The majority of the residues along TM2 showed a reduction in the extent of spin—spin coupling as measured by the W parameter, indicating a translational movement or helical tilt away from the symmetry axis that would indicate and increase in the diameter of the internal vestibule of the channel. Vector analysis of these changes in the context of a helical wheel projection indicated that the pattern of change in inter-subunit proximity could be explained, in addition to the helical tilt, by a rotation of the individual TM2 helices in an anticlockwise direction (Perozo et al 1999). Similar analysis of the spectral changes in the 'external' transmembrane segment TM1 also revealed helix rotations and tilts compatible with a rigid-body movement of the entire helix. However, because TM1 is located further away from the axis of symmetry, changes in inter-subunit residue proximity are more subtle and harder to detect than in TM2. For this reason, and given that less than half of the TM1 residues are involved in any extensive tertiary contacts, changes in probe mobility (DDHo) are a very sensitive indicator of helix tilting and rotation. As in TM2, these data suggested an anti-clockwise rotation of TM1 upon channel gating. Limited data on the types of conformational changes occurring in the vicinity of the selectivity filter showed subtle changes in both probe mobility and spin-spin interaction (Perozo et al 1999). Interestingly, spin labels located at the C-terminal end of the pore helix reported larger changes in spectral line shape as those in the extracellular vestibule of the channel. Taken together, these spectral changes might be suggestive of possible conformational rearrangements that could affect the energetics of ion coordination at the selectivity filter. As stated above, distance determinations in KcsA are complicated by the geometric relationship among subunits, as a fully spin-labelled channel will show dipolar interactions between both neighbouring and diagonally related subunits. This problem was recently solved by the use of tandem dimer constructs to determine changes in inter-subunit distances at the inner helical bundle of KcsA (Liu et al 2001). These inter-subunit distances were then used to derive a backbone structure for the KcsA inner helical bundle in the open state with the aid of a novel computational algorithm (Fig. 5A). This computational approach (restraintdriven Cartesian transformation, ReDCaT), is based on an exhaustive sampling of rigid-body movement in Cartesian space, and was used to determine the type, direction and magnitude of the conformational changes in TM2 using these limited distance information (Sompornpisut et al 2001). The overall picture of KcsA gating emerging from these studies is summarized in Fig. 5B. Our gating model proposes that upon lowering the pH, TM1 and TM2 undergo an anti-clockwise rotation and a simultaneous movement away from the symmetry axis of the tetramer, thus opening the intracellular side of the permeation path. Each TM2 helix supports a scissoring-type motion with a pivot point near the

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FIG. 5. A molecular mechanism of KcsA gating. (A) Average structure of the inner TM2 helical bundle from the crystal structure, and the calculated open structure derived from intersubunit distances and the assumption of rigid-body conformational changes. ReDCaT refers to the computational approach used to calculate the open TM2 bundle (restraint-driven Cartesian transformations) (B) Schematic model of gating in KcsA.

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centre of the helix. Although the data are currently limited, small movements in or around the selectivity filter cannot be ruled out as having a role in activation gating.

References Armstrong CM 1971 Interaction of tetraethylammonium ion derivatives with the potassium channels of giant axons. J Gen Physiol 58:413-437 Berliner LJ, Grunwald J, Hankovszky HO, Hideg K 1982 A novel reversible thiol-specific spin label: papain active site labeling and inhibition. Anal Biochem 119:450-455 Columbus L, Kalai T, Jeko J, Hideg K, Hubbell WL 2001 Molecular motion of spin labeled side chains in a-helices: analysis by variation of side chain structure. Biochemistry 40:3828-3846 Cornette JL, Cease KB, Margalit H, Spouge JL, Berzofsky JA, DeLisi C 1987 Hydrophobicity scales and computational techniques for detecting amphipathic structures in proteins. J MoJ Biol 195:659-685 Cortes DM, Cuello LG, Perozo E 2001 Molecular architecture of full-length KcsA: role of cytoplasmic domains in ion permeation and activation gating. J Gen Physiol 117:165-180 Cuello LG, Romero JG, Cortes DM, Perozo E 1998 pH-dependent gating in the Streptomyces lividans K+ channel. Biochemistry 37:3229—3236 Donnelly D, Overington JP, Blundell TL 1994 The prediction and orientation of alpha-helices from sequence alignments: the combined use of environment-dependent substitution tables, Fourier transform methods and helix capping rules. Protein Eng 7:645-653 Doyle DA, Cabral JM, Pfuetzner RA et al 1998 The structure of the potassium channel: molecular basis of K+ conduction and selectivity. Science 280:69 77 Gross A, Columbus L, Hideg K, Altenbach C, Hubbell WL 1999 Structure of the KcsA potassium channel from Streptomyces lividans: a site-directed spin labeling study of the second transmembrane segment. Biochemistry 38:10324 10335 Heginbotham L, Kolmakova-Partensky L, Miller C 1998 Functional reconstitution of a prokaryotic K+ channel. J Gen Physiol 111:741-749 Heginbotham L, LeMasurier M, Kolmakova-Partensky L, Miller C 1999 Single Streptomyces lividans K+ channels: functional asymmetries and sidedness of proton activation. ) Gen Physiol 114:551-560 Hubbell WL, Altenbach C 1994 Site-directed spin labeling of membrane proteins. In: White SH (ed) Membrane protein structure, experimental approaches. Oxford University Press, New York, p 224-248 Hubbell WL, Cafiso DS, Altenbach C 2000 Identifying conformational changes with sitedirected spin labeling. Nat Struct Biol 7:735-739 Hubbell WL, Gross A, Langen R, Lietzow MA 1998 Recent advances in site-directed spin labeling of proteins. Curr Opin Struct Biol 8:649-656 Hustedt EJ, Smirnov AI, Laub CF, Cobb CE, Beth AH 1997 Molecular distances from dipolar coupled spin-labels: the global analysis of multifrequency continuous wave electron paramagnetic resonance data. Biophys J 72:1861 1877 Hyde JS, Subczynski WK 1989 Spin label oxymetry. In Berliner LJ, Ruben J (eds) Biological magnetic resonance, vol 8: spin labelling. Klewer—Plenum, New York, p 399-425 Liu Y, Holmgren M, Jurman ME, Yellen G 1997 Gated access to the pore of a voltagedependent K+ channel. Neuron 19:175—184 Liu YS, Sompornpisut P, Perozo E 2001 Structure of the KcsA channel intracellular gate in the open state. Nat Struct Biol 8:883-887 Mchaourab H, Perozo E 2000 Determination of protein folds and conformational dynamics using spin-labeling EPR spectroscopy. In: Berliner LJ, Eaton SS, Eaton GR (eds)

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Biological Magnetic Resonance, vol 19: distance measurements in biological systems by EPR. Kluwer-Plenum, New York, p 155-218 Mchaourab HS, Berengian AR, Koteiche HA 1997 Site-directed spin-labeling study of the structure and subunit interactions along a conserved sequence in the a-crystallin domain of heat-shock protein 27. Evidence of a conserved subunit interface. Biochemistry 36:1462714634 Mchaourab HS, Lietzow MA, Hideg K, Hubbell WL 1996 Motion of spin-labeled side chains in T4 lysozyme. Correlation with protein structure and dynamics. Biochemistry 35:7692-7704 Perozo E, Cortes DM, Cuello LG 1998 Three-dimensional architecture and gating mechanism of a K+ channel studied by EPR spectroscopy. Nat Struct Biol 5:459-469 Perozo E, Cortes DM, Cuello LG 1999 Structural rearrangements underlying K + -channel activation gating. Science 285:73-78 Rabenstein MD, Shin YK 1995 Determination of the distance between two spin labels attached to a macromolecule. Proc Natl Acad Sci USA 92:8239-8243 Schrempf H, Schmidt O, Kummerlen R et al 1995 A prokaryotic potassium ion channel with two predicted transmembrane segments from Streptomyces lividans. EMBO J 14:5170-5178 Sompornpisut P, Liu YS, Perozo E 2001 Calculation of rigid body conformational changes using restraint-driven Cartesian transformations. Biophys J 81:2530-2546 Steinhoff HJ, Radzwill N, Thevis W et al 1997 Determination of interspin distances between spin labels attached to insulin: comparison of electron paramagnetic resonance data with the X-ray structure. Biophys J 73:3287-3298

DISCUSSION Mindell: My understanding of KcsA gating, especially in Na+ and K+, is that even at low pH the open probability is still quite low. How does this affect things? Perozo: All the experiments we did were in low intracellular salt concentrations. All the experiments used to calculate open probability are done in a bilayer. In the bilayer conditions the surrounding solutions have a high ionic strength of about 200 mM K+. Under these conditions, we see very little flux. It is still pH dependent, but flux throughput is very low. If the ionic strength is lowered only on the inside of the channel, there is a vast increase in the size of the flux. This tells us that the gating mechanism probably has a strong electrostatic component. Choe: How low is the low ionic strength? Perozo: It is about 5 mM. Mindell: The current is from the 200 mM side, presumably. Perozo: Yes.

Mindell: Do you know how many channels are in that membrane? Perozo: No, and this is a bit of a problem. The open probability increases dramatically, but the open times don't increase much. Since the open times are so short, it is hard to know how many channels we have. This does not change the fact that the mechanism of opening is based on tilting rotations of the helices. These movements are probably there and they may be larger than we think. Ashcroft: What is the ionic strength inside the bacterium? Schrempf: It is about 150 mM.

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1 59

Asbcroft: Does this mean that the channel will be closed all the time? Perozo: Under the conditions that we have studied, probably. However, the volume of a bacterium is minuscule and the single charge conductance is fairly large. You need very few ions to cross the membrane to change the potential. However, this doesn't eliminate the possibility that the lipid composition in the bacterium membrane is different to that which we use in our experiments. There might also be associated subunits that we don't know about. Swartz: It seems to me that even if the open probability is much higher than under other conditions, all you are showing is a correlation. The question is, what can we do to go beyond this and show a causal relationship between the pH, the structural changes you are seeing by electron paramagnetic resonance (EPR) and the open state? Perozo: Here is the problem. If you want to know from EPR the absolute open state structures, given the uncertainties of the open probabilities, I probably cannot give them to you. All I can do is attach the gating mechanism to the structure. It is clearly a more complicated system than we think. One proposal that I like is to suggest that there are actually two gates. The lower gate is responsible for determining the length of the single channel burst. Once it opens, it allows the pore to become dynamic. This dynamic behaviour then determines the structure inside the burst: what we call the 'flicker'. This is actually a very flickery channel. If you do more of these experiments at low ionic strength, you see that the burst duration actually increases significantly, but the mean open time does not increase. Thus there could be two open states, one not conducting and one conducting. Corringer: If you propose a conformation for the open state, you can try to introduce two cysteines which, when bridged together, constrain this plausible conformation. This would be a nice demonstration. Such a strategy was used in the case of the P2X receptor. Swartz: If you find the titratable residue, then you would potentially have a route to show that the pH-dependent structural changes are in fact related to the opening and closing of the gate. Perozo: It depends what you want to do. If you want to do electrophysiology, this might be feasible. If you want to do spin label, you will not be able to use the cysteines for cross-linking, since they will react. Sansom: I have a couple of questions about your methodology, and particularly about mobility. With KcsA in the closed state, do you see any evidence of a gradient in mobility from extracellular to intracellular? Secondly, given that some X-ray structures for membrane proteins reveal tightly bound lipid (Fyfe et al 2001), if you had a tightly bound lipid would that result in a change in mobility due to interaction with lipid rather than protein? Perozo: We wouldn't be able to distinguish whether there was a tightly bound lipid versus another part of the protein that is interacting with the label. It all

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depends on the dynamic range of the label. EPR has a very specific window of dynamics that goes from hundreds of picoseconds to hundreds of microseconds. This is ideal for looking at conformational changes in proteins. However, lipid motion typically occurs within a very fast regime, beyond the window of dynamics that we can determine. In all the membrane proteins that have been spin labelled, as we go from the water-exposed residues to the lipid-exposed ones, the observed changes in mobility are not that significant. They are quite similar. From the point of view of spin labels, water and lipids are just two different types of solvents and they are both moving very fast. Sansom: Has anyone put a spin label on one of these sites where one can see a bound lipid in an X-ray structure? Perozo: As far as I am aware, the only membrane proteins that have been spin labelled and analysed in any systematic way are KcsA, bacteriorhodopsin, Lac permease, rhodopsin, colicin, diptheria toxin and some porins. We have just finished MscL as well. Mindell: MscL does have some bound detergent in the crystal structure, so who knows what this is in a bilayer. Sansom: I was thinking about bound lipid. For example, FhuA in the Ferguson et al (1998) structure has tightly bound lipopolysaccharide. Perozo: Within the time window in which we can look, it doesn't seem to affect the mobility of exposed residues very much. Sansom: In the closed state of KcsA, is mobility along Ml constant? Perozo: There is slight variation from end to end. We do see a clear decrease in mobility from the beginning of the N-terminus to the point at which it inserts into TM1. There is a clear gradient there, which is compatible with an extended helix that is floating around in the plane of the membrane. Mitra: With regard to the robustness of your method, if you were to do an experiment where you deliberately distorted a helix somewhat and then go through with your analysis, how different a result would you get? Perozo: It would depend very much on whatever surrounds the helices. One of the clear limitations of our technique is that the resolution is limited at the level of the backbone. Let's take the most basic scenario of a peptide oriented perpendicular to the membrane. If you spin label all of the peptides and look at the mobility, they are all going to be the same because they are all facing the same environment, unless there is some local interaction. If you have the same helix and it is bent, only the bent part with the semi-cavity that is formed restricts the mobility. The same applies for accessibility. The picture improves dramatically for us when we have multi-segment proteins, because their TM segments have to associate somehow. This association gives clear differences in environments, so now the environment that is oriented towards the rest of the molecule is clearly more motion-restricted than the side that is facing the lipid.

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Unwin: Are your experiments done on the truncated protein? Perozo: The EPR experiments are done with full length KcsA that has a His tag at the N-terminus. Everything is done at room temperature and in low ionic strength. Unwin: As far as you can tell, is the structure of the gate with the full-length protein the same as the one solved by X-rays? Perozo: We have done experiments to answer that question. We cut the C-terminus and watched what happened to the mobility parameter and spin—spin coupling. There were some changes in the mobility, as you would expect, when you get closer and closer to the point that is cut (residue 125). But the changes are not that significant and it gates almost as well. Unwin: That C-terminal helix is lying on a membrane and the gating rotates it around in some way. Perozo: That doesn't appear to change through the gating. Unwin: The helix that connects the two does a bit. Surely the link must be flexible? Perozo: That is what we think. It is hard to measure the conformational changes at the N-terminus because if TM1 moves slightly, it is still going to be independent of the membrane. In fact, the remarkable result is that if you profile the environmental parameters of this region, the mobility is actually flat. By looking at the mobility on its own we could not draw any conclusions about its structure. Yet if you look at the oxygen profile, you see amazing periodicity. If you look at the nickel data, you see the same periodicity, but almost 180° out of phase. This means that whenever there is a minimum in oxygen, there is a maximum in nickel. The only way we can explain this is by assuming that it lies flat on the membrane surface, and that there is a linker here. Unwin: Why is it there? Perozo: I have no idea. Roux: When people wonder about what can be structurally deduced out of a technique such as EPR, one must distinguish the different steps in the procedure. If you make a biophysical measurement, the first question you should ask concerns how reliable the distance or property measured is: what is the uncertainty? When you test EPR with known structures and you compare the distances, there is quite a good agreement. Perozo: There is some agreement, but it is not perfect. Roux: At least this gives an assessment of the significance of the measurements that are made and also more information about the window of uncertainty. Some of the uncertainty is related to the intrinsic perturbation that is introduced by the technique. The same is true with the thermodynamic double mutant cycle analysis. This technique was tested on barstar—barnase, for which there is a crystallographic structure of the protein—protein complex. These tests showed that the strong

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coupling between a pair of amino acids is more or less indicative of a short distance between them in the protein—protein complex (Schreiber & Fersht 1995, Buckle et al 1994). This type of validation, or control, is the first thing that must be known from any method if one is going to start deluding structural constraints. Second, one clearly needs to have enough data, or enough restraints. This would apply to any other technique. Third, one needs a computational procedure to incorporate the available data into a coherent structural model. For example, in the early 1980s people were struggling to deduce the three dimensional structure of proteins from nuclear magnetic resonance (NMR) data. They had trouble because they did not know how to integrate all the available data. Then, methods such as distance geometry (Havel & Wuthrich 1985) and restrained molecular dynamics (Brunger et al 1986) came along, which solved these problems. Beside the computational procedure, the crucial first step in structural studies of Shaker is to know how reliable the biophysical measurements are, and what is the intrinsic uncertainty. Perozo: The absolute distance information is probably the most complicated and unreliable. This is the same for all spectroscopically related data. My approach is to say that in any specific case, the precision of the distance is low, but the pattern of data obtained is far more important than any individual data point. The more data you have, the better. Clearly, this is a long-term process, and we hope to gradually improve the quality of the data in order to refine the models. In our case, there is a strict limitation on how well we can do, because of the size of the probe and the uncertainties about its conformation. But despite this, our data are fairly solid and our conclusions can be supported as long as we limit ourselves at the backbone fold level. Schrempf: I have a comment in support of some of your notions. We have analysed our mutants that have changes in the outer vestibule. If you look at the ratio of openings and the ratio of subconducting states, i.e. if you have exchanged the tyrosine residue in position 82, in the end you have increased the openings and different ratios of the different opening stages. This suggests that there are conformational stages that can be blocked in different positions, and also that there is another gating position. Perozo: Over the last two months we have started looking at single channel analysis of cysteine mutants, within and outside the pore region. There are clearly gating-related effects of cysteine mutants. Sixma: I was wondering about your model of the N-terminal helix. Is there anything special about the sequence there? Perozo: It is very amphipathic. Sixma: Are there a lot of tryptophans and tyrosines? These are often seen at the interface. Perozo: KcsA has a ring of tryptophans, and there are a couple here.

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Fu: Do you think it is possible to determine the pH dependency of the open profile? Perozo: It depends on how you do it. We have done this many times using 86Rb+ fluxes. In the wild-type KcsA it is fully closed at alkaline pH. At pH 7 there is an open probability of 15-20%. This rises sharply up to pH 5, and then remains high before decreasing at lower pH. We think this may have to do with the ability of the system to measure the fluxes. The logical experiment would be to do this at a single channel level. Fortunately for the EPR but unfortunately for the physiology, > 95% of the channels reconstitute with the intracellular side looking outside of the vesicle. This simplifies the pH and solution changes, but when we do patch clamp, the pH sensor, which is intracellular, will always be in the pipette. This makes it hard to do patch—clamp experiments. We have been reluctant to do what many others have done, which is to exchange the pipette solution. Fu: You determined the profile of those two distinct states, which would suggest a conformational movement that is associated with channel opening. Based on the pH dependency of the open profile, channel opening is presumably a result of pH titration of certain residues in the channel gate. When you mutated the channel lining residues, you may also have altered the pKa values of these side chains. Did you confirm that such mutations did not alter pH dependency of the channel open profile? Perozo: Every time we do a mutant, we check three things: that it is a tetramer, the temperature of stability of that tetramer, and its ability to support 86Rb+ flux. In all cases we see tetramers and at least some level of activity. Swartz: What is the status of knowing what groups you are titrating? Is it one, or many? Perozo: We have been working at this for almost two years. We started looking in the wrong place, because we thought it was on the outside. After Heginbotham and Miller showed that it was intracellular, we shifted gears and started mutating all the negative charges inside. There is a cluster of charges right at the base of KcsA at the exit point of TM2: these are prime candidates. When we mutate these there are shifts in the apparent pKa, but in no case did we see a dramatic decrease in the slope. And with a single mutation it wasn't possible to make a pH-independent KcsA. We then started looking at combinations of charges, which produced increasing shifts. But in all cases the slopes change only slightly, not dramatically. This would suggest that we are touching something that affects the pH sensor but is not the pH sensor itself. I think that there might be a contribution from a number of charges plus perhaps the lipid. Instead of being a point charge, this is an array of charges in delicate equilibrium. Under physiological conditions a change in pH balances the charges and local potentials, which I think will result in repulsion of the helices.

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Cboe: Can you imagine doing EPR experiments in the solid state with more oriented samples? Perozo: We have not done that experiment, but Wayne Hubbell's group has successfully oriented membranes in a special cavity. Those are very nice experiments.

References Brunger AT, Clore GM, Gronenborn AM, Karplus M 1986 Three-dimensional structure of proteins determined by molecular dynamics with interproton distance restraints: application to crambin. Proc Natl Acad Sci USA 83:3801-3805 Buckle AM, Schreiber G, Fersht AR 1994 Protein-protein recognition: crystal structural analysis of a barnase-barstar complex at 2.0 A resolution. Biochemistry 33:8878-8889 Ferguson AD, Hofmann E, Coulton JW, Diederichs K, Welte W 1998 Siderophore-mediated iron transport: crystal structure of FhuA with bound lipopolysaccharide. Science 282:22152220 Fyfe PK, McAuley KE, Roszak AW, Isaacs NW, Codgell RJ, Jones MR 2001 Probing the interface between membrane proteins and membrane lipids by X-ray crystallography. Trends Biochem Sci 26:106-112 Havel TF, Wuthrich K 1985 An evaluation of the combined use of nuclear magnetic resonance and distance geometry for the determination of protein conformation in solution. J Mol Biol 182:281-294 Schreiber G, Fersht AR 1995 Energetics of protein-protein interactions: analysis of the barnasebarstar interface by single mutations and double mutant cycles. J Mol Biol 248:478-486

General discussion I

From structure to channel physiology Ashcroft: How do we go from the crystal structure — or even something which is intermediate, like the electroparamagnetic resonance (EPR) data — to the singlechannel currents? It has been very noticeable at this meeting that although we are discussing channels, there has been an almost total lack of single-channel recording data. How do we interpret the single-channel kinetics in terms of structure, and structure in terms of the gating and permeation properties? Choe: I think we should derive DG between two conformations. This will give us a starting point from which to see whether it correlates with the open channel probability. Perozo: It is a complicated problem because of the way that ion channels function. They have been designed over the course of evolution to have regions that undergo conformational changes within microseconds. They are therefore metastable in many cases. In structural biology you need a defined structure in order to solve it. I think the answer lies in a combination of crystal structure which gives high resolution, spectroscopy which gives something intermediate and dynamic, physiology and theory. Sansom: In the F-ATPase field, Wang & Oster (1998) have performed mechanical calculations to interpolate between the different snapshots seen in John Walker's X-ray structures. They are attempting to reproduce computationally the nature of the motions that may occur between structural endpoints. Perozo: The problem in channels is that getting snapshots of the intermediates is the hard part. Let's say that we are able to standardize what we call the open state. It actually becomes more dynamic once the gates are open, and therefore less determined from the point of view of the structure. Miller: The other thing that is important, and Eduardo Perozo has been hinting at, is that it isn't just the equilibrium states that are of interest. We are also interested in gating. I study things like ethanol, which are mild peturbants. Ethanol shifts the acetylcholine concentration—response curve for activation to the left. The effect increases in magnitude along the kinetic pathway. Relative to the closed state, the free energy of the closed to open state transition-state decreases a little, the free energy of the open state decreases more, and the free energy of the open to 165

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desensitized state transition-state decreases most of all. The whole potential energysurface is influenced by ethanol. Non-equilibrium states are not amenable to structural studies, so this observation is quite a challenge. A shcroft: I guess the other problem is that there will be conformational changes associated with transitions between different closed states. Mindell: Some of the methods that Nigel Unwin has developed, using cryoelectron microscopy, offer some hope in this direction. He is able to isolate intermediate states that are at least metastable. Solving the electron microscopy structures of these states may help get the structures that facilitate modelling of the intermediates. Pero^p: Is that true? What is the occupancy of these intermediates when you freeze them? Unwin: This channel is very efficient at opening. With an excess of acetylcholine about 95% of the channels are supposed to be open at any given time. This is consistent with what we see. It may be a lucky case. Fu: In the crystal structures we can only see one snapshot of many conformational states. What does one crystal structure really tell us about the protein function? One of the key factors that will allow us to determine a meaningful structure is to find a ligand that will lock the protein conformation into one particular state. If we can do this, we may have a better chance of crystallizing and solving the structure. The next challenge is how we can extend our knowledge from one static picture to a dynamic model. This is where molecular dynamics studies and free energy calculations can help. One of the things I am still quite not sure about is the level of accuracy of molecular simulations. The free energy barriers between various conformational states could be small, perhaps only a few kCal/mol. Sansom: It is no secret that the accuracy of the calculations is a problem at the moment. It will get better, but not overnight. From looking at other fields, one of the ways to get structural information in the time scale at which physiologists measure gating, is to go to single-molecule spectroscopy. If you look at some of the single-molecule fluorescence studies on single enzyme turnover (Bai et al 1999), they have reinvented channel gating kinetics and see the sorts of patterns in their turnover kinetics that you see in channel gating kinetics. Perhaps the answer is to perform spectroscopic studies (e.g. using fluorescence) at the single channel level. Then you have to interpret them in structural terms, which is non-trivial. Miller: It is possible to do kinetics with EPR down to about a millisecond. Sansom: But not with a single molecule. Pero^p: For a single molecule you have to go to fluorescence, although there have been initial attempts to do single molecule EPR. Sansom: I think I have seen single-molecule infrared, as well.

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Choe: From a structural point of view, we all operate under the assumption that there are distinct conformations that structural biologists can offer between open and closed states. One has to remember this may not happen. Conformation averaged over your experimental measurement time may not change at all. One analogy is that if you have a door that is moving at a constant speed, on average it will have half occupancy of one conformation no matter what speed it is moving at. Instead, we may need to know how fast it switches between the two conformations. This might be the key that underlies the mechanism of selecting Na + versus K + . We operate under this traditional idea that the size is the key determinant and hydration matters, but these things have to come on and off in certain kinetic terms. This, coupled with the kinetic motion of a couple of distinct conformational states, will be impossible to trace by crystallographic methods. We should keep this as an open possibility. Miller: That is an interesting idea and EPR techniques might be useful in testing it. They can provide some structural information in the time dimension as well as the space dimension. Picture a protein flipping between two conformations and bearing a spin label on an amino acid residue that is mobile on the EPR time scale in one conformation but not the other. Then the spectra contributed by each conformation will be so different that the experimental spectrum may be deconvoluted into two components. This allows the ratio of the two conformations to be estimated. Choe: In my proposed scenario the ratio stays the same, but the rate of the conversion between the two changes. Miller: Rate information is available from EPR because of the phenomenon of exchange broadening of spectra. In the example above, the experimental spectrum is only the sum of the two-component spectra when the interchange between the two conformations is slow on the EPR time scale (> 1 //s). When it is fast on the EPR time scale (< 1 ns) a time average spectrum is obtained. In the intermediate time frame the spectra are sensitive to the rate of interchange. The same phenomenon is observed with NMR but on a slower time scale. Choe: I'll give you another analogy. As you go towards a revolving door, it revolves at a constant speed. If it spins very fast you can never get in; if it spins slowly it still spins with the same ratio between closed and open states, yet you can go through. Miller: That is right, and you can only get a picture of the half open door if you use a camera with a fast enough shutter speed. I would like to turn to another inherent methodological problem. We are interested in both silent and conducting conformational states in order to obtain a complete kinetic description of channel regulation and permeability. Electrophysiological methods only measure open conformational states. If you look at the experiments necessary to solve complex schemes in classical enzyme

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kinetics, complete descriptions were only achieved with a number of probes each reporting on a different conformational state. While electrophysiological measurements on channels are very powerful for analysing the exits and entries to the open state, it becomes increasingly difficult to tease out information as states become progressively more remote from the open state. A. uerbach: You can often tell the number of states even if they have the same conductance. It is not as if you just see one state. Miller: For example, if the system has three silent states before it opens, it is more difficult to get information about the first two then the third from the open channel kinetics measurements. If you are clever you can devise experiments that help, but it may be difficult to be unambiguous. References Bai C, Wang C, Xie XS, Wolynes PG 1999 Single molecule physics and chemistry. Proc Nad Acad Sci USA 96:11075-11076 Wang H, Oster G 1998 Energy transduction in the F, motor of ATP synthase. Nature 396:279-282

Excitability is mediated by the Tl domain of the voltage-gated potassium channel Senyon Choc*, Susan Cushmanf, Kent A. Baker* and Paul Pfaffingerf *Structural Biology Laboratory, The Salk Institute, l^a Jolla, CA 92037 and ^Division of Neuroscience, 5613, Baj/or College of Medicine, 1 Baylor Pla^a, Houston, TX 77030, USA

Abstract. The Tl domain of voltage-gated K + (Kv) channel is the N-terminal cytoplasmic part of the channel preceding the transmembrane pore domain of the channel. Several crystal structures of the Tl domain show that the four Tl subunits are arranged in a rotationally symmetric tetramer. The subunit interface of the Tl domain encodes the assembly specificity of intact functional K v channels. Along the fourfold symmetry axis of the Tl tetramer, a water-filled cavity exists. K + ions, however, do not pass through this Tl cavity. Instead, they are believed to enter the transmembrane pore through four identical inter-subunit spaces created between the membrane-facing C-terminal side of the Tl tetramer and the inner leaflet of the membrane. Several point mutations have been introduced into the putative membrane-facing region of the Tl tetramer. These mutations led to a systematic change of the channel's voltage sensitivity. Such functional change was accompanied by a distinct structural change in the C-terminal membrane-facing side of the Tl tetramer. Interestingly, a similar structural alteration that renders the channel more excitable is also induced by the binding of a cytoplasmic protein K v /? subunit. Within this conformationally flexible part of the Tl tetramer, non-Shaker type K v channel subunits invariably contain one Zn2+ per subunit. With the KV4.2 Tl, we demonstrated that the tetramer can be reversibly converted to monomers by chelating zinc away from the protein. The rate of removal of Zn2+ is pH-dependent. The structural ability of the Tl tetramer to alter conformation could be an essential property to mediate and process protein—protein interaction events in the cytoplasm to control excitability of intact full-length K v channels. 2002 Ion channels—from atomic resolution physiology to functional genomics. Wiley, Chi Chester ( Novartis Foundation Symposium 245) p 169-177

Voltage-gated, K + ion-selective (Kv) channels are involved at all levels of basic physiological functions. Some are role-specific such as those with endocrine and exocrine functions, and others serve fundamental cellular functions of excitable membranes such as cardiac excitability, learning and memory, as well as the control of synaptic efficacy. Functional, pore-forming Kv channels consist of four a subunits. The a subunit is encoded as a transmembrane protein chain with 169

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each a subunit containing six transmembrane helices (referred to as SI—S6) immediately following a highly conserved N-terminal domain on the cytoplasmic side. Four subunits are thought to cluster around a water-filled ion permeation pathway (Doyle et al 1998). The intracellular N-terminal domain (referred to as Tl domain) displays a high degree of sequence conservation within four major gene families (Shaker, Shab, Shaw and Shal}. These four distinct subfamilies of Kv channels are evolutionary conserved. They homo- and heterotetramerize exclusively between subunits that belong to the same subfamily, even from different species. Previous studies showed that the highly conserved cytoplasmic Tl domain is sufficient to encode specificity that governs the tetramerization process (Shen & Pfaflfinger 1995) in intact functional pore-forming K v channels. In the crystal structures of the truncated Tl tetramers from Shaker (Kreusch et al 1998) and Shaw(Bixby et al 1999), four Tl subunits form rotationally symmetric tetramers. The interfacial interaction is highly polar and the interface residues show a high degree of amino acid conservation in a subfamily-specific manner. The overall architecture of the Tl tetramer consists of four layer-like segments folded along the symmetry axis around the water-filled cavity, termed the Tl cavity. In Fig. 1A, layers 1-4 denote different parts of the Tl chain to signify their relative position to the rest of the molecule including the putative membrane, starting from the N-terminus of layer 1 to the C-terminus of layer 4, respectively. Cushman et al (2000) demonstrated that the Tl cavity is not part of the ion permeation pathway. Therefore, we concluded that the cytoplasmic entryway for ions should be found between the membrane-facing C-terminal side of the Tl teramer and the inner leaflet of the membrane (Choe et al 1999). The structurally underdefined large grey box in Fig. 1 symbolizes the putative cytoplasmic vestibule formed between the Tl tetramer and the membrane. This structural model is further supported by the crystal structure of the Tl/ Kv/? complex (Gulbis et al 2000). In the structure of the complex, the N-terminal side of the Tl tetramer (layer 1) interfaces the flat side of the KV/J tetramer coaxially. The side openings of K v channels are bounded by the linker segments between the C-terminus of the layer 4 and the N-terminus of the first transmembrane helix SI (linker B in Fig. 1 A) and also between the C-terminus of the S6and the entire Cterminal domain of the channel. The framework of this side opening may include the linker loop connecting S4 and S5 (linker C), which is thought to play a crucial role in responding to voltage activation or channel inactivation. Based on the mutational data Zhou et al (2001) recently concluded that the N-terminal end of the channel unfolds and extends its long flexible linker A to reach the central pore cavity of the channel through this side opening to inactivate the channel. A direct visualization for such conformational change may not be readily attainable by Xray crystallographic methods, but perhaps by NMR techniques.

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The subunit interfaces of the Tl tetramer are highly polar despite the fact that they are buried.. We have previously shown that the putative membrane-facing side of the Tl tetramer is conformationally flexible as compared to other parts of Tl. We further showed that site-specific mutations introduced into the flexible region of the molecule (layer 3) result in conformational alterations in the same region (Fig. IB), which then shifts the channel activation potential (Cushman et al 2000). A similar conformational change occurs within the Tl tetramer when it is complexed with Kv/? tetramer (Gulbis et al 2000). These experimental data combined together imply that the conformational changes occurring in the Tl tetramer, most notably in the membrane-facing region of Tl near the putative side openings, can be faithfully sensed by the voltage-sensing and ionconducting modules of the channel in the membrane. This energetic link would then require a tight conformational junction between Tl tetramer and the rest of the channel to be able to propagate conformational changes throughout the channel. These are not inconsistent with the idea of a hanging gondola with four connecting cables (Korberz et al 2000), however rigid or flexible they are. Within this relatively flexible membrane-facing part of the Tl tetramer (layers 3 and 4), all Tl domains of non-Shaker subfamilies bind Zn2+ at the tetrameric subunit interface (Bixby et al 1999). Zn2+ is tetrahedrally coordinated by four amino acids on the subunit interface: one histidine and two cysteine residues from one subunit, and a cysteine residue from the facing subunit. Since four Zn2+ atoms within the tetramer are deeply buried inside the protein, access to the zinc is limited by the protein conformation. It was found that the removal of zinc from the protein interface was sufficient to disassemble the Tl tetramer into monomers. At lower pH, the rate of monomerization is accelerated, probably depending on the conformational plasticity of the Tl domain (Jahng etal.2001). The Tl domain can be replaced by an artificial tetramerization module (Zerangue et al 2000) with only limited functional changes and diminished stability of the channel assembly. This implies that Zn2+ does not directly affect the channel property, but instead may influence it indirectly through the protein domain. Therefore, the conformational plasticity can enable Tl to mediate signalling through various protein—protein interactions with the cytoplasmic components such as Ca2+-binding KchIP to regulate K v channels. In the case of the interaction between Tl and Kv/? tetramers, the direct interaction involves the N-terminal side of the Tl tetramer, yet the Tl tetramer in the complex undergoes a structural change on the C-terminal membrane-facing side opposite from the K v/?contacting interface (Gulbis et al 2000). Interestingly, the altered conformation is closer to that of the previously reported two cavity-widening, left-shifting mutants of Tl tetramer (Cushman et al 2000). From these studies, we conclude that an important physiological role of Tl is to act as a structural platform

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FIG. 1(A). A composite structure of a K v channel. Side view of Tl tetramer (bottom half) is combined with the transmembrane ion-permeating part modelled by the structure of KcsA (top half). There is only one N-terminal inactivation domain shown for clarity. The fourfold symmetry axis is vertical in this view. Only three subunits are shown for clarity, in three different shades. Numbers 1 through 4 on the left side denote layers 1 through 4 from the Nterminus to the C-terminus of the chain, respectively. Connecting loops A through C are structurally undefined, but probably are important functional components. The large grey rectangular box in the middle indicates the putative cytoplasmic vestibule and probably includes elements involved in controlling channel properties. Through four sides of this box, ions are expected to enter and merge to the central pore to cross the membrane. It is structurally unknown how a single inactivation peptide ball would reach into the vestibule to block the ionic flow through the central pore region. The smaller box on the top denotes the narrowest part of the ion permeation pathway modelled on the basis of the transmembrane domain of the KcsA channel (Doyle et al 1998).

for interactions with cytoplasmic proteins in order to modulate the channel excitability. Conformational flexibility of the Tl tetramer induced by solution pH provides a plausible explanation by which binding of cytoplasmic proteins to Tl tetramer can trigger a conformational change perhaps by protonating a specific set of amino acids. Such conformational propagation through the Tl tetramer (Lockless & Ranganathan 1999) may mediate the protein-protein interactions in the

T1 - M ED I AT ED EX CIT A BI LIT Y

E1G. 1(B). Surface representation of conformational change of the Tl tetramer as a result of cavity-widening mutations. The level of shading ranges from 0.01 A to 1.8 A as given in the bar. The orientation of the Tl tetramer is the same as in the panel (A). Only two diagonally symmetric subunits are shown. The putative entry ways for ions will be between the top part of the tetramer and the membrane, which is the most significantly altered area as a result of mutational changes.

cytoplasm to affect channel function. Zn2+ is found abundantly at presynaptic vesicles, well stained by Zn2+-specific dyes, and is known to potentiate A-type currents. Localized tissue distribution of some K v channels and mechanisms of controlling cell surface expression will enable cells to pattern the biological activity by physical changes at genetic and cellular levels. These are some exciting research avenues for yielding a better understanding of channel regulation in response to cellular states and neuronal signals. New insights are beginning to emerge from discoveries of several cytoplasmic proteins: KchIP (K+ channel interacting protein) (An et al 2000) or auxiliary regulator proteins such as K v j8 subunit. These cytoplasmic proteins are known to make physical interactions directly with the N-terminal domain of K v channels. Understanding the molecular interactions between these proteins and biological controls for

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Retention Time (minutes) FIG. 2. Elution profile of the rKv4.2 Tl domain in Superdex75, run at 1 ml/min. The sample elutes at 64.5 ml in the presence of 50-60 j/M Zn2+ at pH 8.0 (solid line), and at 85.4 ml in the presence of 2mM EDTA (dotted line). The conversion is reversible by Zn2"1" or 2mM EDTA, respectively. Rates of conversion from tetramer to monomers are accelerated by slightly acidic pH (6.5-7.0) (Jahng et al 2002). The molecular weight estimates in the two different states are consistent with the results from analytical centrifugation. T and M denote tetramer and monomer peaks, respectively.

tissue-specific distribution will be instrumental to elucidate the mechanisms of channel excitability.

References An WE, Bowlby MR, Betty M 2000 Modulation of A-type potassium channels by a family of calcium sensors. Nature 403:553-556 Bixby KA, Nanao MH, Shen NV et al 1999 Zn 2+ -binding and molecular determinants of tetramerization in voltage-gated K + channels. Nat Struct Biol 6:38-43 Choe S, Kreusch A, Pfaffinger PJ 1999 Towards the three-dimensional structure of voltagegated potassium channels. Trends Biochem Sci 24:345-349 Cushman S, Nanao MH, Jahng AW, DeRubeis D, Choe S, Pfaffinger PJ 2000 Voltage dependent activation of potassium channels is coupled to Tl domain structure. Nat Struct Biol 7:403-407 Doyle DA, Morais-Cabral J, Pfuetzner et al 1998 The structure of the potassium channel: molecular basis of K + conduction and selectivity. Science 280:69-77 Gulbis J, Zhou M, Mann S, MacKinnon R 2000 Structure of the cytoplasmic /? subunit — Tl assembly of voltage-dependent K + channels. Science 289:123-127 Jahng AW, Strang C, Kaiser D, Pollard T, Pfaffinger P, Choe S 2002 Zinc mediates assembly of Tl domain of the voltage-gated K channel 4,2. Submitted Kobertz WR, Williams C, Miller C 2000 Hanging goldola structure of the Tl domain in a voltage-gated K + channel. Biochemistry 39:10347-10352 Kreusch A, Pfaffinger PJ, Stevens CF, Choe S 1998 Crystal structure of the tetramerization domain of the Shaker potassium channel. Nature 392:945-948

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Lockless SW, Ranganathan R1999 Evolutionarily conserved pathways of energetic connectivity in protein families. Science 286:295—299 Shen NV, Pfaffinger PJ 1995 Molecular recognition and assembly sequences involved in the subfamily-specific assembly of voltage-gated K + channel subunit proteins. Neuron 14:625633 Zerangue N, Jan YN, Jan LY 2000 An artificial tetramerization domain restores efficient assembly of functional Shaker channels lacking Tl. Proc Natl Acad Sci USA 97:3591—3595 Zhou M, Morais-Cabral JH, Mann S, MacKinnon R 2001 Potassium channel receptor site for the inactivation gate and quaternary amine inhibitors. Nature 411:657-661

DISCUSSION Miller: Is it possible sterically just to thread that chain A through the middle of the tetramer, and have the ball much closer to the channel? Choe: Rick Aldrich and others clipped this linker A down before we had the structure. When we had the structure we were able to see the domain boundary of Tl in their sequence, so we were able to count how many residues were left between their ball boundary and Tl. It was as few as 19 in one of their constructs. Unwin: Is there any possibility that the ball could be sitting on the fourfold axis of the Tl domain, such that Tl acts as a sort of presenter for the ball just underneath the channel? Choe: I wish I could answer that; it is a great idea. Obviously, that requires the channel not to have a fourfold symmetric arrangement. One guy has to sit close enough to the centre so that one is enough to do the job. Wallace: If you completely remove the Tl domain, what effect does it have on ion permeability and selectivity? Choe: This was done back in the mid-1990s in Carol Deutsch's lab, and was most recently revisited by Miller's group after the structure had been found. In all cases the channels they dealt with were from different species. In the case ofAp/ysia, we could not produce an assembled channel, so we could not properly measure any ion conductance. Miller's group truncated off in the middle of the B chain. The inactivation properties were altered: the on rate was four times slower but the off rate was unchanged. Inactivation properties can be eliminated without much influence on the overall structure by eliminating this part. The on rate was decreased, which is counterintuitive if the pocket was somewhere in the membrane. If the on rate was increased we might imagine that the pocket is somewhere inside, so we don't have anything in the way and consequently the access is faster. I don't remember what happened to the ion selectivity, but overall the key difference is that the channel is not stable, although this doesn't say much about the actual physical properties of the channels that were formed. Papa^ian: Those of us who are interested in how these things fold and assemble would really like to know the relative contributions of interactions within and

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without the plane of the membrane in terms of stabilizing the tetramer. Not everyone has KcsA, which stays together in SDS to a very high temperature. For other channels, it seems like your zinc situation is perfect. If you take the full length channel for one of these families where the Tl domains are held together with zinc, and you coordinate the zinc, what happens to the stability of the tetramer overall? Choe: Removing zinc is very hard. It takes hours. The zinc is buried 10 A deep from outside and 16 A from inside. The rate is changed as a function of pH, and this is not because the chelating affinity is changing; it is because of the conformational change of the protein. The reason that the rate of monomerization is altered as a function of pH and yet tetramerization isn't, is not because the extra protonation influences the affinity between zinc and the coordinating side chains, but because the access to that site, which was monitored by the tryptophan just outside, is changing so that at acidic pH the conformation allows chelation faster. Even so, the actual rate of removal is very slow. In practice, it has been technically very difficult. Schrempf: I am surprised that you didn't try to oxidise. It is known from chaperones, and also the RPA protein which has some binding activity for S-S, that activity is modulated by oxidation. The zinc basically keeps those residues in the vicinity where they can later be oxidised. What happens if you form S-S bridges? This would also be a reversible process. Choe: That is a good suggestion. Gulbis: Proteins such as the chaperone Hsp33 are directly activated by cellular oxidative stress. These become functional upon disulfide formation. If the zinc atom is removed from K V 3 Tl via oxidation, forming disulfides, the new topology would more closely resemble that of K v l (Shaker) Tl. Perhaps chelating the zinc modifies the local protein structure in such a way as to protect the pore entrance. In this way it could act as a physical obstruction. Choe: Also, Tl may disassemble in some cases. Gulbis: It may. However if an oxidizing agent were used in place of a chelating agent the cysteines would oxidize, which would undoubtedly affect this somehow. And it may have a less disruptive effect on the quaternary structure of Tl. Choe: The discovery of zinc was serendipitous. Nothing in the literature even hinted at its existence, but when we were tracing the map we ran into this bulky element. It took us a long time to identify this as zinc. Later of course it was confirmed that crystals grew better if we added zinc, which meant that zinc was captured during a protein preparation from whole cell lysate, and was never removed during purification. It was that stable. Gulbis: It would be interesting to know whether the Tl structure is different under oxidative conditions. Choe: KV4.2 has been documented to be sensing the oxygen state.

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Mathie: In terms of neuronal excitability, some K V 3 channels have positive thresholds for activation. Could you explain any of these positive thresholds in terms of differences in the Tl region? Or do you think this resides somewhere else? Choe: It's speculative at present. I reckon this excitability modulated by cytoplasmic protrusion of Kv channels to be a very similar scenario in general. There are at least three proteins that intimately regulate the excitability. Some are shifting the activation potential by as much as 35 mV. It is found in Caenorhabditis ekgans by a point mutation identified by genetic screening that is about two amino acids from the C-terminus of Tl. This point mutation shifts the sensitivity to the right by 35 mV. There are three examples that may be helpful. First, it has been known since 1994 that K/? shifts the sensitivity up and down and now binds here to influence the action over there. Second, Src kinase phosphorylates a few residues preceding the N-terminus of Tl, and this phosphorylation almost shuts down the channel. The third and most interesting example is a Ca2+ binding protein called Kchip. This protein is rnonomeric in cytoplasm, but when it binds to Tl, four molecules bind to the tetrameric Tl. This indicates that somehow there is room for four molecules binding together. Does it need all four? This is not known. One of the key ways of regulating this channel's assembly is by modulating the affinity at the interface. The way a squid axon does this is through RN A editing, as a result of cytoplasmic signals, which has been shown by Pancho Bezanilla. By looking at mRNAs, he was able to pick a dozen or so sites that undergo RNA editing changes which result in amino acid changes. These data were compared against the structure, and the majority of these amino acids were at the interface of Tl. It seems reasonable to think that all these cytoplasmic signals that control RNA editing enzymes can influence the subtlety of excitability, by affecting the affinity of the Tl assembly.

Structural organization of the voltage sensor in voltage-dependent potassium channels Diane M. Papazian, William R. Silverman, Meng-chin A. Lin, Seema K. Tiwari-Woodruff and Chih-Yung Tang Department of Physiology, UCLA School of Medicine, Los Angeles, CA 90095-1751, L'SA

Abstract. The structural organization of the voltage sensor in K + channels has been investigated by second site suppressor analysis in Shaker and by identification of a metal ion binding site in ether-a-go-go (eag). In Shaker, two groups of interacting charged residues have been identified. K374 in the S4 segment interacts with E293 in S2 and D316 in S3, whereas E283 in S2 interacts with R368 and R371, two voltage-sensing residues in S4. Interactions of E283 with R368 and R371 are voltage dependent. The results suggest that E283 is located in a water-filled pocket near the extracellular surface of the protein. During voltage-dependent activation of Shaker channels, R368 and R3^1 move into this pocket and come into proximity with E283. In eag channels, extracellular Mg2+ directly modulates the activation process by binding to two acidic residues that are located in an analogous pocket. These acidic residues are found only in eag family members, accounting for the specificity of Mg2+ modulation to that family. These compatible results from Shaker and eag suggest a model for the packing and conformational changes of transmembrane segments in the voltage sensor of K* channels. 2002 Ion channels—from atomic resolution physiology to functional genomics. Wiley, Chichesttr (Novartis Foundation Symposium 245) p 178-192

In voltage-dependent ion channels, the control of channel activity by voltage is due to the presence of an intrinsic charged voltage sensor that undergoes conformational changes in response to changes in the membrane potential, thereby regulating the probability of pore opening (Papazian & Bezanilla 1997). Many voltage-dependent channels are closed at the resting potential, but open upon depolarization of the membrane. This chapter will review aspects of voltage sensor structure and function in K + channels. Voltage-dependent K + channels include four subunits that surround the central ion selective pore, and may also contain auxiliary subunits that regulate function or cell surface expression (MacKinnon 1991, Rettig et al 1994, Shi et al 1996). Each 178

ORGANIZATION OF VOLTAGE SENSOR IN K + CHANNELS

FIG. 1. Membrane topology of the Shaker K + channel protein. The functional channel contains four subunits that assemble to form the pore. The first four transmembrane segments, S1-S4, contain the voltage sensor, whereas the last two transmembrane segments, S5 and S6, and the re-entrant P loop form the pore. The approximate positions of voltage-sensing residues in S2 and S4 are shown (Aggarwal & MacKinnon 1996, Seoh et al 1996). The locations of two glycosylated asparagine residues in the extracellular S1--S2 loop are also indicated (trees) (Santacruz-Toloza et al 1994).

subunit contains six transmembrane segments and a re-entrant P loop that forms the selectivity filter (Fig. 1). The mechanisms underlying selectivity and permeation are understood in molecular detail due to extensive electrophysiological analysis and the recently solved structure of the Streptomyces lividans KcsA channel, a bacterial K + channel protein with significant similarity in sequence and function to the pore domain of eukaryotic voltage-dependent K + channels (Doyle et al 1998, MacKinnon et al 1998). In contrast to the molecular mechanism of permeation, much less is known about the structure and function of the voltage sensor in voltage-gated channels. The mechanism of voltage-dependent activation has been studied in a number of channels, but perhaps most extensively in Shaker K + channels (Bezanilla 2000). Shaker channels are steeply dependent on voltage. It has been estimated that the equivalent of 12—13 elementary charges (CQ) cross the transmembrane field during the activation of one Shaker channel (Schoppa et al 1992). This represents the gating charge transferred by conformational changes of the voltage sensor during the activation process. Charged residues that contribute to the single

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channel gating charge, including four positively charged arginine residues in the S4 segment, and a negatively charged glutamate residue in S2, have been identified since mutations that neutralize these residues dramatically decrease the single channel gating charge (Fig. 1) (Aggarwal & MacKinnon 1996, Seoh et al 1996). Substituting the charge moving residues with cysteine or histidine and then probing their conformation-dependent reactivity with hydrophilic probes from opposite sides of the membrane has provided compelling evidence that the S4 residues traverse virtually the entire transmembrane electric field upon activation (Larsson et al 1996, Baker et al 1998, Starace et al 1997, Starace & Bezanilla 2001). In contrast, it is not known whether the acidic residue in S2 moves relative to the transmembrane field during activation or, instead, helps to shape the profile of the electric field detected by the moving S4 residues (Papazian & Bezanilla 1997). Voltage sensor function can be revealed by the analysis of gating currents, which are electrical measurements of the conformational changes of the voltage sensor (Armstrong & Bezanilla 1973). A comparison of the voltage dependencies of voltage sensor conformational change (Q-V relationship) and pore opening (PoV relationship) reveals that the voltage sensor begins to undergo conformational changes at potentials more hyperpolarized than those that lead to channel opening (Bezanilla 2000). These results suggest that voltage-dependent channels transit through several closed conformations, which differ in the position of the voltage sensor, before entering the conducting (open) state (Bezanilla 2000). Interestingly, detailed analysis of gating currents suggests that voltage sensor movement can be divided into a minimum of two phases (Bezanilla et al 1994). Mutations in the S4 segment can alter the properties of both phases of gating charge movement, suggesting that the S4 segment is involved in both types of conformational changes (Perozo et al 1994). This does not preclude the involvement of other segments of the protein in voltage sensor movement. Evidence that S2 may also be involved has been presented based on tagging the voltage sensor with fluorescent probes and biophysical analysis (Cha & Bezanilla 1997). To understand the molecular basis of voltage-dependent activation, it will be essential to determine the structure of the voltage sensor in several conformations along the activation pathway and to resolve how voltage sensor movement is coupled to pore opening. How are the transmembrane segments of the voltage sensor packed together in voltage-dependent K + channels? In our laboratory, we have used two approaches to identify likely structural interactions in the voltage sensor of K + channels, and to investigate how these interactions rearrange during activation. These approaches are second site suppressor analysis in Shaker K + channels and identification of a binding site for divalent cations that is present in the voltage sensor of ether-ago-go (eag) K + channels (Papazian et al 1995, Tiwari-Woodruff et al 1997, Silverman et al 2000). These studies have led to compatible results that suggest a

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packing arrangement and some of the conformational changes of the voltage sensor in K + channels. Experimental procedures Site-directed mutations were made using PCR methods (Tiwari-Woodruff et al 1997). For electrophysiological or biochemical analysis, cRNA encoding wildtype and mutant Shaker or eag proteins was injected into Xenopus oocytes (Papazian et al 1991). Ionic currents were recorded using a two-electrode voltage clamp as previously described (Tiwari-Woodruff et al 1997). For biochemical analysis, in vitro translation grade [35S]-methionine was co-injected with cRNA. Oocyte membranes were isolated, and Shaker protein was solubilized, immunoprecipitated and analysed by fluorography as previously described (Santacruz-Toloza et al 1994, Schulteis et al 1998). Results and discussion Second site suppressor analysis reveals voltage-dependent structural interactions in the voltage sensor of Shaker channels

For second site suppressor analysis, mutations that disrupt the normal folding or assembly of a protein are identified and then rescued by introducing a specific second site mutation. The interpretation is that the positions of the disruptive mutation and its specific second site suppressor are in proximity in the native three-dimensional structure of the protein. In Shaker channels, mutations that disrupt folding or assembly are easily identified because they prevent protein maturation. Like many other membrane proteins, the Shaker K + channel protein is detected immediately after synthesis as an immature, core-glycosylated protein in the endoplasmic reticulum (ER) (Schulteis et al 1995, Nagaya & Papazian 1997). The Shaker protein folds and assembles in the ER and is then transferred to the Golgi apparatus where the carbohydrate chains are modified to the complex type (Nagaya & Papazian 1997, Schulteis et al 1998). Conversion of the immature precursor to the mature product is accompanied by a significant change in the electrophoretic mobility of the protein, making it easy to distinguish these two forms on a denaturing acrylamide gel (Santacruz-Toloza et al 1994, Schulteis et al 1995). Biogenesis of the Shaker channel is strictly subject to the quality control system of the ER. Whereas the wild-type protein folds, assembles and leaves the ER very efficiently, mutant proteins that disrupt folding and assembly are recognized and retained in the ER, and may be targeted for degradation by cytoplasmic proteasomes (Schulteis et al 1998, Myers et al 2001). Therefore, misfolded

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proteins are readily identified by their failure to mature (Papazian et al 1995, Schulteisetall998). Position K374 will be used to illustrate the strategy of second site suppressor analysis. This highly conserved positively charged residue in the S4 segment does not contribute significantly to the single-channel gating charge (Seoh et al 1996), but instead plays a pivotal role in determining the structure of the voltage sensor. Neutralization or charge reversal mutations of K374 eliminate maturation, abolish functional expression, and lead to intracellular retention of the Shaker protein (Papazian et al 1995, Tiwari-Woodruff et al 1997). These results led us to hypothesize that the positively charged residue, K374, normally interacts with a negatively charged residue, and that this interaction is essential for formation of the voltage sensor. To test this hypothesis, charge reversal mutations were made at a variety of acidic residues located in and near the transmembrane segments of the protein (Tiwari-Woodruff et al 1997). These mutations were then paired individually with K374E, a charge reversal mutation that disrupts folding of the voltage sensor. Double mutant combinations were assessed biochemically to determine the efficiency of protein maturation, and electrophysiologically to determine whether functional channels were produced (Papazian et al 1995, Tiwari-Woodruff et al 1997). Of those positions assessed, only two, E293 in S2 and D316 in S3, could be mutated to suppress the defective maturation found in K374 mutants (Fig. 2A) (Papazian et al 1995, Tiwari-Woodruff et al 1997). Double-mutant proteins that matured efficiently were able to incorporate into functional channels, confirming that these proteins adopt a native three-dimensional structure (Fig. 2B) (Papazian et al 1995, Tiwari-Woodruff et al 1997). From these results, we conclude that K374 in S4 is a close neighbour of E293 in S2 and D316 in S3 in the native structure of the Shaker K + channel (Papazian et al 1995, Tiwari-Woodruff et al 1997). Figure 2C summarizes the results obtained from second site suppressor analysis of the Shaker voltage sensor (Papazian et al 1995, Tiwari-Woodruff et al 1997). In addition to the interactions of K374 with E293 and D316, proximity was inferred between E283 in S2 and R368 and R371 in S4. Because R368 and R371 are two of the charge-moving residues in S4 that undergo voltage-dependent conformational changes during activation (see Fig. 1) (Aggarwal & MacKinnon 1996, Seoh et al 1996), the results suggest that structural proximity with E283 may exist in some but not all of the conformational states of the protein. To test this hypothesis, we investigated the location of E283 in different conformations, using the approach of cysteine mutagenesis followed by extracellular exposure to hydrophilic sulfhydryl reagents (Tiwari-Woodruff et al 2000). To obtain a protein that expressed robustly, we performed these experiments using a double-mutant construct, E283C+R371Q, in a non-inactivating (IR) background.

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FIG. 2. Second-site suppressor analysis of K374 mutations. (A) Protein products of Shaker-IR (IR), the single mutant K374E, and several double mutant combinations are shown. The Shaker protein was immunoprecipitated and subjected to denaturing electrophoresis and fluorography. The upper and lower arrows indicate the positions of the mature and immature forms of the Shaker protein, respectively. This figure was originally published in Tiwari-Woodruff et al (1997). (B) Functional expression of the double mutant combination E293K+K374E in Xenopus oocytes. Using a two electrode voltage clamp, currents were evoked from E293K+K374E-IR channels by pulsing from a holding potential of — lOOtnV to potentials ranging from — 80 to +60 mV in 20 mV increments. This figure was originally published in Tiwari-Woodruff et al (1997). (C) Summary of structural interactions in the Shaker voltage sensor inferred from second-site suppressor analysis. E283 in S2 is shown interacting with R368 and R371 in S4, and K374 in S4 is shown interacting with E293 in S2 and D316 in S3 (dotted lines) (Papazian et al 1995, Tiwari-Woodruff et al 1997).

In E283C+R371Q-IR channels, extracellularly applied methanethiosulfonate reagents (MTSES and MTSET) dramatically accelerate deactivation (TiwariWoodrufFet al 2000). This effect is not observed in control channels. Changes in deactivation kinetics were used to monitor the reactivity of E283C in different protein conformations. Application of the sulfhydryl reagents at either — lOOmV or +40 mV, potentials at which the channel is in resting or activated conformations, respectively, had identical effects on deactivation kinetics (Fig. 3). Furthermore, the rates of reaction of the reagents were similar in both resting and

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FIG. 3. Extracellular MTS reagents accelerate deactivation in E283C+R371Q-1R channels. MTSET (5 mM) or MTSES (1 mM) was applied for 2min while clamping the membrane at — 100 or +40 mV, as indicated. Tail currents were fitted with a single exponential function to obtain a time constant for deactivation (Tdeact). A box plot of rdeact values obtained before and after treatment is shown. These results were originally published in Tiwari-\\"oodruff et a! (2000).

activated conformations. Reaction rates were low compared to those of MTS reagents with sulfhydryl groups in solution (Stauffer & Karlin 1994), suggesting that position 283 is located in a pocket somewhat protected from the extracellular solution. We concluded that position 283 can be accessed from the extracellular solution in both resting and activated conformations (Tiwari-Woodruff et al 2000), consistent with the finding that position 283 does not contribute to the single-channel gating charge, and therefore does not move relative to the transmembrane field during voltage-dependent activation (Seoh et al 1996). Taken together, our results and those obtained in other laboratories indicate that in the resting conformation, R368 and R371 are near the intracellular surface of the Shaker protein, whereas E283 is near the extracellular surface (Larsson et al 1996, Starace et al 1997, Baker et al 1998, Tiwari-Woodruff et al 2000, Starace & Bezamlla 2001). This strongly suggests that the interactions between these positions, which we have inferred from second site suppressor analysis, do not occur in the resting conformation. Analysis of the voltage-dependence of the double mutant combinations E283R+R368E-IR and E283R+R371E-IR indicate that these mutations stabilize the protein in different conformations (Tiwari-Woodruff et al 2000). Whereas E283R+R368E-IR increases the relative stability of a nonconducting state, E283R+R371E-IR increases the relative stability of a fully activated conformation. From these results, we have proposed that E283 and R368 interact transiently in an intermediate closed state along the activation

ORGANIZATION OF VOLTAGE SENSOR IN K + CHANNELS

FIG. 4. Tentative model of voltage sensor conformational changes during activation of Shaker channels. At rest, E283 is near the extracellular surface of the protein, whereas R368 and R371 are near the intracellular surface (Tiwari-Woodruff et al 2000, Larsson et al 1996, Starace & Bezanilla 2001). Upon activation, first R368 and then R371 move into proximity with E283. In the intermediate conformation, R371 is shown in parentheses because the data do not address its position in this state. This model is adapted from one originally published in Tiwari-Woodruff et al (2000).

pathway, whereas E283 and R371 achieve their closest proximity in the open conformation (Fig. 4) (Tiwari-Woodruff et al 2000).

Identification of a metal ion binding site in the voltage sensor ofeag K+ channels

We have also identified structural constraints on the packing of transmembrane segments in the voltage sensor by locating a binding site for metal ions in eag K"1" channels. Extracellular Mg2+ dramatically slows eag activation kinetics (Terlau et al 1996, Tang et al 2000). Analysis of gating and ionic currents indicates that Mg2+ directly modulates the process of activation gating, suggesting that the binding site is located in or near the eag voltage sensor (Tang et al 2000). We hypothesized that the acidic side chains of aspartate and glutamate residues coordinate bound Mg2+ in the eag voltage sensor, as found in other metal ion binding sites in proteins (Williams 1993). Sequence comparison indicates that members of the eag subfamily contain two acidic residues, one each in S2 and S3, which are not present in other subfamilies of voltage-dependent K + channels (Fig. 5 A) (Warmke & Ganetzky 1994). These are good candidates to contribute to the ion binding site because modulation of voltage-dependent activation by extracellular Mg2+ occurs in many members of the eag subfamily, but not in other voltage-dependent K + channels (Terlau et al 1996). We tested the hypothesis that these eag-specific acidic residues coordinate the bound ion by mutating them and determining the consequences for Mg2+ modulation of activation gating kinetics (Silverman et al 2000). Mutations D278V and D278E of the eag-specific acidic residue in S2 abolish Mg2+ sensitivity of activation kinetics (Fig. 5B) (Silverman et al 2000).

ORGANIZATION OF VOLTAGE SENSOR IN K + CHANNELS

\8"?

Interestingly, the two mutations have differential effects on activation kinetics. The D278V channel activates slowly with kinetics that resemble the wild-type channel in the presence of Mg2+. In contrast, the D278E channel activates rapidly with kinetics that resemble the wild-type channel in the absence of Mg2+. These results strongly suggest that bound Mg2+ shields the charge at D278, and that this contributes to its effect on activation kinetics. Because Mg2+ sensitivity is abolished by the conservative mutation D278E, which conserves the negative charge but lengthens the side chain by one methylene group, the binding site is likely to be sterically constrained near D278. We conclude that D278 in S2 contributes to the Mg2+ binding site (Silverman et al 2000). A similar analysis was performed after mutating D327, the eag-specific acidic residue in S3. D327F dramatically reduces the Mg2+ sensitivity of eag channels (Silverman et al 2000). Like D278V, neutralization mutations at D327 slow activation kinetics so that they are similar to the wild-type in the presence of Mg2+. We conclude that D327 also coordinates the Mg2+ ion. Our results indicate that extracellular Mg2+ binds in a water-filled pocket between S2 and S3, where it is coordinated by the two eag-specific acidic residues (Silverman et al 2000). Does this pocket correspond to the one in Shaker that contains E283? To address this question, we investigated the effect of mutating D274, the eag counterpart of E283 in S2, on Mg2+ sensitivity. Mg2+ modulates

FIG. 5. Identification of the Mg2+ binding site in eag channels. (A) Aligned sequences of S2 and S3 segments from eag family members and other voltage-dependent K + channels are shown. Arrows indicate acidic residues (in bold), numbered according to the Drosophila eag sequence. Acidic residues corresponding to positions 274, 284, and 319 are conserved among all voltagedependent K + channels, whereas those corresponding to positions 278 and 327 are found only in members of the eag subfamily (Warmke & Ganetzky 1994). Abbreviations used: d, Drosophila; t, rat; b, bovine; h, human; m, mouse; erg, eag-related gene; elk, eag-like K + channel; ShB, Shaker. (B) Currents from wild-type eag and the mutants D278V and D278E were recorded at +60 mV in the presence (dashed traces) and absence (solid traces) of lOmM extracellular Mg2+. Scale bars represent 2^A, O.S^A and 2^(A for wild-type, D278V, and D278E channels, respectively, and 25 ms. A single exponential component has been fitted to the late phase of wild-type current activation (bold curves). D278V and D278E channels are insensitive to Mg~+. Note that the mutations have a differential effect on activation kinetics, with D278V mimicking the slow activation kinetics of the wild-type in the presence of Mg2+, and D278E mimicking the fast activation kinetics of the wild-type in the absence of Mg2+. These results were originally published in Silverman et al (2000). (C) A model for the packing of the voltage sensor in K + channels based on second site suppressor analysis in Shaker channels and identification of a metal ion binding site in eag (Papazian et al 1995, Tiwari-Woodruff et al 1997, Silverman et al 2000). Inferred structural constraints are shown as dotted lines. The black circle denotes a bound Mg2+ ion coordinated by the side chains of D278 and D327 in eag (underlined italics). All other residues are numbered according to the Shaker sequence. This model represents the activated conformation of the voltage sensor (Tiwari-Woodruff et al 2000) and has been adapted from a figure originally published in Silverman et al (2000).

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activation gating in both D274A and D274E, indicating that D274 is not required for Mg2+ binding. In fact, Mg2+ slows activation kinetics more in D274E than it does in wild-type eag. Importantly, Mg2+ washes off more slowly from D274E than from wild-type channels. These results suggest that D274 does not interact with the bound ion in the wild-type channel. However, a longer acidic side chain is able to interact with the ion, increasing the apparent affinity for Mg2+. From these results we conclude that the Mg2+ binding site is located in an extracellular-facing pocket between S2 and S3, analogous to the one containing E283 in the Shaker voltage sensor (Fig. 5C) (Silverman et al 2000). In eag, bound Mg2"1" is thus in an optimal position to modulate the movement of S4 residues into the pocket, and to constrain any movements of S2 relative to S3 during activation gating. Voltage sensor structural constraints identified in eag and Shaker are quite compatible, consistent with structural conservation between these two divergent channel proteins. Our results suggest a model for the packing of the voltage sensor in an activated conformation (Fig. 5C). Tilting the S4 segment in this model provides a location for solvent-filled pockets that have been suggested to exist at both the extracellular and intracellular faces of the membrane (Tiwari-Woodruff et al 2000, Islas & Sigworth 2001). Interestingly, the S4 tilt angle is similar to that proposed by Kenton Swartz and his colleagues for a voltage sensor/pore interaction surface on the basis of perturbation analysis of the pore domain in Shaker channels (Li-Smerin et al 2000). This raises the possibility that the S4 segment interacts directly with the pore. Although speculative, this is an easily testable proposal. Knowledge of voltage sensor-pore interactions will be required to understand how conformational changes of the voltage sensor result in pore opening. This remains a key unanswered question in ion channel structure and function.

A cknowledgements This work was supported by grants from the National Institutes of Health (GM43459) and the American Heart Association, Western States Affiliate.

References Aggarwal SK, MacKinnon R 1996 Contribution of the S4 segment to gating charge in the Shaker K + channel. Neuron 16:1169-1177 Armstrong CM, Bezanilla F 1973 Currents related to movement of the gating particles of the sodium channels. Nature 242:459-461 Baker OS, Larsson HP, Mannuzzu LM, IsacoffEY 1998 Three transmembrane conformations and sequence-dependent displacement of the S4 domain in Shaker K* channel gating. Neuron 20:1283-1294 Bezanilla F 2000 The voltage sensor in voltage-dependent ion channels. Physiol Rev 80:555 592

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Bezanilla F, Perozo E, Stefani E 1994 Gating of Shaker K channels: II. The components of gating currents and a model of channel activation. Biophys J 66:1011—1021 Cha A, Bezanilla F 1997 Characterizing voltage-dependent conformational changes in the Shaker K + channel withfluorescence.Neuron 19:1127-1140 Doyle DA, Morais Cabral J, Pfuetzner RA et al 1998 The structure of the potassium channel: molecular basis of K + conduction and selectivity. Science 280:69—77 Mas LD, Sigworth FJ 2000 Electrostatics and the gating pore of Shaker potassium channels. ) Gen Physiol 117:69-90 Larsson HP, Baker OS, Dhillon DS, IsacoffEY 1996 Transmembrane movement of the Shaker K f channel S4. Neuron 16:387-397 Li-Smerin Y, Hackos DH, Swartz KJ 2000 A localized interaction surface for voltage-sensing domains on the pore domain of a K 4 channel. Neuron 25:411-423 MacKinnon R 1991 Determination of the subunit stoichiometry of a voltage-activated potassium channel. Nature 350:232-235 MacKinnon R, Cohen SL, Kuo A, Lee A, Chait BT 1998 Structural conservation in prokaryotic and eukaryotic potassium channels. Science 280:106-109 Myers MP, Khanna R, Papazian DM 2001 Differential targeting of Shaker mutant proteins to cytoplasmic proteasomes. Biophys J 80:219a Nagaya N, Papazian DM 1997 Potassium channel a and ft subunits assemble in the endoplasmic reticulum. J Biol Chem 272:3022-3027 Papazian DM, Bezanilla F 1997 How does an ion channel sense voltage? News Physiol Sci 12:203-210 Papazian DM, Timpe LC, Jan YN, Jan LY 1991 Alteration of voltage-dependence of Shaker potassium channel by mutations in the S4 sequence. Nature 349:305-310 Papazian DM, Shao XM, Seoh SA, Mock AF, Huang Y, Wainstock DH 1995 Electrostatic interactions of S4 voltage sensor in Shaker K + channel. Neuron 14:1293—1301 Perozo E, Santacruz-Toloza L, Stefani E, Bezanilla F, Papazian DM 1994 S4 mutations alter gating currents of Shaker K channels. Biophys J 66:345-354 Rettig J, Heinemann SH, Wunder F et al 1994 Inactivation properties of voltage-gated K + channels altered by presence of beta-subunit. Nature 369:289—294 Santacruz-Toloza L, Huang Y, John S A, Papazian DM 1994 Glycosylation of Shaker potassium channel protein expressed in insect cell culture and in Xenopusoocytes. Biochemistry 33:56075613 Schoppa NE, McCormack K, Tanouye MA, Sigworth FJ 1992 The size of gating charge in wildtype and mutant Shaker potassium channels. Science 255:1712—1715 Schulteis CT, John SA, Huang Y, Tang C-Y, Papazian DM 1995 Conserved cysteine residues in the Shaker K + channel are not linked by a disulfide bond. Biochemistry 34:1725-1733 Schulteis CT, Nagaya N, Papazian DM 1998 Subunit folding and assembly steps are interspersed during Shaker potassium channel biogenesis. J Biol Chem 273:26210-26217 Seoh S-A, Sigg D, Papazian DM, Bezanilla F 1996 Voltage-sensing residues in the S2 and S4 segments of the ShakerY^ channel. Neuron 16:1159—1167 Shi G, Nakahira K, Hammond S, Rhodes KJ, Schechter LE, Trimmer ]S 1996 ft subunits promote K + channel surface expression through effects early in biosynthesis. Neuron 16:843-852 Silverman WR, Tang C-Y, Mock AF, Huh K-B, Papazian DM 2000 Mg2+ modulates voltagedependent activation in ether-a-go-go potassium channels by binding between transmembrane segments S2 and S3. J Gen Physiol 116:663-678 Starace DM, Bezanilla F 2001 Histidine scanning mutagenesis of basic residues of the S4 segment of the Shaker K + channel. J Gen Physiol 117:469-490 Starace DM, Stefani E, Bezanilla F 1997 Voltage-dependent proton transport by the voltage sensor of the S baker K? channel. Neuron 19:1319 1327

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Stauffer DA, Karlin A 1994 Electrostatic potential of the acetylcholine binding sites in the nicotinic receptor probed by reactions of binding site cysteines with charged methanethiosulfonates. Biochemistry 33:6840-6849 Tang C-Y, Bezanilla, F, Papazian DM 2000 Extracellular Mg2+ modulates slow gating transitions and the opening of Drosophila ether-a-go-go potassium channels. } Gen Physiol 115:319-338 Terlau H, Ludwig J, Steffan R, Pongs O, Stuhmer W, Heinemann SH 1996 Extracellular Mg2* regulates activation of rat eag potassium channel. Pfliigers Arch 432:301-312 Tiwari-Woodruff SK, Schulteis CT, Mock AF, Papazian DM 1997 Electrostatic interactions between transmembrane segments mediate folding of Shaker K + channel subunits. Biophys J 72:1489-1500 Tiwari-Woodruff SK, Lin MA, Schulteis CT, Papazian DM 2000 Voltage-dependent structural interactions in the Shaker K + channel. J Gen Physiol 115:123-138 Warmke JW, Ganetzky B 1994 A family of potassium channel genes related to eag in Drosopbila and mammals. Proc Natl Acad Sci USA 91:3438-3442 Williams RJP 1993 Magnesium: an introduction to its biochemistry. In: Birch KJ (ed) Magnesium and the cell. Academic Press, New York, p 15-29

DISCUSSION Pero^o: What do you think Mg2+ is doing? Is it fair to say that it limits the relative motion of the helices, and that is why it slows down the whole process? Papazian: That is one possibility. For aficionados in the voltage sensing field, it is known that the voltage sensor undergoes more than one conformational change, and that there may well be conformational changes outside of S4. The one place that people have focused on is S2. It is possible that Mg2+ could be restricting the movement of S2 relative to S3, and in addition affecting the movement of S4 into the vicinity. The effect of Mg2+ on the electrophysiology is quite complicated, and it is clearly working at different phases of the activation mechanism. It is possible that it is affecting different conformational changes. Pero^p: Is it possible to measure the affinities for Mg2+ at different stages in the activation? Papazian: That has been hard, but I am starting to suspect that Mg2+ may bind much better to the closed state. It does not get forced out of the channel at depolarized potentials, but we think that the affinity might be higher in the closed state. Sansom: How do we know that the second-site suppressor mutations are in the same subunit, and not rescuing between subunits? Papazian: That is a good question. For some of them we don't know. For the bottom set, the data that we have suggest that they are within one subunit, that is, that the two mutations have to be on the same subunit in order to effectively rescue each other. We have also done some electrophysiological analysis that suggests that the residues have to be on the same subunit.

+

ORGANIZATION OF VOLTAGE SENSOR IN K CHANNELS

191

Mindell: You said that the 283 methanethiosulfonate (MTS) reaction is very slow, but the Mg2+ wash-in is very fast. Papa^ian: Yes. Mindell: If these are in the same pocket, how are those alterations compatible with one another? Is it to do with the size of the pocket? Papa^ian: Perhaps. Of course, the MTS reagents are much larger than Mg2+. In addition, the size and shape of the pockets in Shaker and eag may not be identical. We haven't really measured the wash-in rate for Mg2+, which would be nice to know. As a result, I can't really compare this with the MTS reaction rate. Swart^: Have you looked at the effect of Mg2+ on the voltage dependence of opening? Papa^ian: It has no effect on the steady-state voltage dependence. It is all in the kinetics. Swart^i Is there any effect on the Q-V relationship? Papa^ian: No. We looked at this in collaboration with Pancho Bezzanila. We wanted to look at the effect of Mg2+ on gating currents, eag is a weird channel: the conformational changes that occur at hyperpolarized potentials are ratelimiting for channel opening, which is not the case in Shaker. It makes it very hard to study gating currents, especially when Mg2+ is added and they get even slower. Snmrt^: So with Mg2+ the gating currents are slowed, but the equilibrium Q - V relationship is not affected. Ashcroft: Your experiments with Mg2+ suggest that changes in the plasma concentration of the ion might have an effect on the cardiac action potential. Does this happen? Papa^ian: Channels of the eag family are expressed in the nervous system and cardiovascular systems. These are two systems where decreases in physiological Mg2+ lead to increases in excitability. Although you might hypothesize that the changes in excitability are due to Mg2+ modulation of eag channels, at least this doesn't seem to be the case in the heart. HERG is an eag-type channel that is expressed in the heart. HERG channels help to repolarize the cardiac action potential, primarily as the channel slowly deactivates. Reduced Mg24' would slow down HERG deactivation even more, increasing outward K + current, which of course would be anti-excitable. So, the increase in excitability is not mediated by HERG. Perhaps Mg2+ modulation of HERG is protective in hypomagnesaemia. A shcroft: I didn't understand why your mutations disrupted folding. You said that the 371 mutation only interacted with 283 in the activated conformation. Is the channel open in the endoplasmic reticulum? Papa^ian: We have now looked at many combinations of mutations that either mature or don't. The ones that mature, with 2 exceptions, don't shift activation at all or they shift activation to the left. This suggests that they stabilize the protein in

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an activated conformation. Our data would suggest that the protein folds in an activated conformation. ROHX: You have shown a very specific interaction between charged residues, but there are also others in S2, S3 and S4. Would you expect that all the charges in this very low dielectric region should be paired with some other charge, or is it known that they are not? Papa^ian: I would be surprised if they are. I accept the idea that there are substantial pockets of solvent that intrude into the plane of the bilayer in the region of the voltage sensor. A lot of these charged ionic residues are probably solvated. I think that the part that is buried is probably short, and includes 374. This is a highly conserved lysine residue that we think is important for determining the fold, and it doesn't seem to move relative to the field. It isn't accessible to solvent by any of the standard approaches. ROMX: That is pretty low along S4. It is almost in the intracellular side. Is this salt bridge lost in the activated conformation? Papa^ian: I don't know. Mitra: What happens when you chelate Mg2+ out, using EGTA for example? Papa^ian: eag takes on rapid activation kinetics.

Electron diffraction of a bacterial CIC-type chloride channel Michelle M. Pirruccello*f, Nikolaus GrigoriefPt and Joseph A. Mindellt 1 *Rosensteil Basic Medical Science Research Center and W.M.. Keck Institute for Cellular Visualisation and ^Department of Biochemistry and Howard Hughes Medical Institute, Brandeis University, Walt ham, MA, USA 02478

Abstract. CIC-type Cl channels, as two-pore homodimers, display an architecture unprecedented in selective ion channels, yet little is known regarding their mechanisms of selectivity and gating. In contrast to the great successes with K + channels, a decade of mutagenic analysis has revealed little about the structure and function of the CICs: even the number of ion-conducting pores per complex is controversial. Thus, for these proteins direct structural information is particularly important. We have formed twodimensional crystals of a bacterial C1C homologue, and are analysing their structure using cryo-electron microscopy of glucose-embedded specimens. Here we report the measurement of electron diffraction patterns from these crystals. Unfettered by the imaging limitations of the electron microscope, the diffraction patterns reveal ordering of the crystals to at least 3.8 A resolution, suggesting that they can be used to generate an atomic model of the protein. We present an improved projection structure of the channel at 6.5 A using amplitude data derived from four electron diffraction patterns, with crystallographic statistics comparable to those reported for other high-quality twodimensional crystals. 2002 Ion channels—-from atomic resolution physiology to functional genomics. Wiley, Chichester (Novartts Foundation Symposium 245) p 193-206

CIC-type chloride channels serve a host of biological functions: from maintaining the resting membrane potential of skeletal muscle cells to mediating acid secretion in bone-reabsorbing osteoclasts. Only a few CIC-type channels have been studied in detail to date: these are unique in both form and function when compared to the better characterized gap junctions, neurotransmitter-activated channels, and S4type voltage-gated channels. Relatively little is known in detail about the C1C channels: indeed the number of ion-conducting pores per channel complex has This paper was presented at the symposium by Joseph A. Mindell, to whom correspondence should be addressed.

193

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PIRRUCCELLO ET AL

been controversial (Maduke et al 2000). The combination of unique function and ignorance motivates direct structure determination on the C1C channels. We recently produced two-dimensional (2D) crystals of a prokaryotic C1C channel homologue, EriC (Mindell et al 2001). The crystals are flat sheets approximately 4^m on one side, and contain 104-105 channels, probably in a single layer. These crystals are excellent candidates for structural analysis by electron crystallography and we recently obtained a projection structure of the EriC channel at 6.5 A resolution with image data obtained using cryo-electron microscopy (Mindell et al 2001). The structure reveals an orthorhombic unit cell with dimensions of 137 x 76 A and p22!2j symmetry. Because the image-derived amplitudes fall off quickly at high resolutions, we amplified the high-resolution data in this map using a procedure which assumes that the attenuation for the EriC crystals is similar to that observed for bacteriorhodopsin, for which accurate amplitude data are known (Havelka et al 1995). This procedure is similar to applying a negative temperature factor to boost the high-resolution data. However, more accurate amplitudes may be derived from electron diffraction patterns of EriC crystals. Use of these amplitudes will facilitate the determination of a high-resolution structure of the EriC channel. If feasible, measuring amplitudes from electron diffraction patterns has significant advantages over measuring them from image data. Because the diffraction pattern is insensitive to specimen drift, very long exposures at very low electron doses are possible. This allows the use of illumination conditions which maximize the coherence of the electron beam and therefore the sharpness of the spots. Furthermore, whereas images are modulated by the oscillating contrast transfer function (CTF), diffraction patterns are not so affected; thus the amplitudes need no CTF correction and are significantly more accurate. Finally, since a larger area of crystal contributes to a diffraction pattern than to an image, more unit cells are averaged, yielding intensities that are also more precise. Of course, since electron diffraction yields only intensity data, phases must still be obtained from images. Here, we demonstrate the feasibility of electron diffraction on the EriC crystals, and present an improved projection map which uses the amplitudes from electron diffraction in combination with the previouslyreported phases. The new map preserves the overall appearance of the channel protein, but displays some minor changes. Materials and methods Specimen preparation and electron diffraction

Crystals of EriC were prepared as described previously (Mindell et al 2001). 4 pi of the purified crystal solution was applied to a carbon-coated copper grid. The

C1C-TYPE CHLORIDE CHANNELS

195

crystals were then embedded in 50 jul 1% glucose with 0.25 mg/ml bacitracin. The grid was mounted into a Gatan cryo-holder, transferred into the microscope, and cooled to liquid nitrogen temperature. Diffraction patterns were collected on a Philips CM12 Transmission Electron Microscope located at Brandeis University in Waltham, Massachusetts, with an accelerating voltage of 120 kV. The patterns were recorded on a Gatan Slow Scan CCD camera with a 1024 X 1024 pixel CCD chip using the gain-normalization option of the Gatan software. Data analysis Background subtraction, indexing of the patterns, integration of peak intensities, and merging of the reflections was performed using the MRC image processing programs as described previously (Baldwin & Henderson 1984, Ceska & Henderson 1990). The scale factor (S) and the isotropic temperature factor (B) relative to the best pattern were calculated as follows (Ceska & Henderson 1990):

B 2,d

^(1 scaled 11 observed) = r^T H~ InJ

This equation was fit using a linear least squares routine to yield the parameters S and B for each new pattern. Image phases to 6.5 A, averaged from seven merged projection images and rounded to 0° and 180° (Mindell et al 2001), were combined with the diffraction data using a small program written for this purpose. In addition, the program used image amplitudes for very low-resolution reflections not measured due to CCD saturation. Results Two-dimensional crystals of EriC belong to the space group p22j2i, with an orthorhombic unit cell possessing dimensions of 137 x 76 A. Diffraction patterns were recorded at an electron dose of approximately 2 e~/A 2 . A total of 57 untilted diffraction patterns were collected at varying camera lengths and exposure times. Four images, recorded at a camera length of 1.1 m and an exposure of 40 s, were of suitable quality for analysis, based on crystallographic statistics. Reflections were measured to a resolution of 4—6 A depending upon the pattern. However, there were distinguishable reflections to a resolution of 3.8 A in the best patterns. An example of a background-subtracted diffraction pattern is shown in Fig. 1. After background subtraction, indexing and spot integration, we used several different R factors to assess the quality of the images. R^- reflects variations in intensity between Friedel-related reflection pairs. Figure 2 shows a graph of R^ values in resolution ranges for the patterns used in the projection structure. Each

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FIG. 1. Electron diffraction from a two-dimensional crystal of EriC, an E. ro//CIC-type Cl channel. Background-subtracted electron diffraction pattern of a glucose-embedded crystal, recoded on a CCD camera using 120 kV accelerating voltage. Note the mirror symmetry about the h and k axes, and the systematic absences along those axes, as well as the 'systematic presences' of the 13,0 spot (arrow).

pattern was initially measured to 4 A, and the sharp rise in R factor at a specific resolution for each pattern represents the limit for measuring reflections distinguishable from the background. Rf generally increased at about 6.5 A, and then decreased again at about 5 A for those patterns that possessed discernible reflections at high resolution, corresponding to a band of weak intensities in the diffraction patterns in that resolution range. R^m measures differences between intensities related by the predicted p22]2j symmetry of the crystal; its values are presented in Table 1. Measured values of R t) , w are always larger than those of R,-, to a varying degree for each image.

C1C-TYPR CHLORIDE CHANNELS

197

FIG. 2. Friedel R factor (Rv) versus resolution for the electron diffraction patterns used in the improved projection structure. R/ was calculated from unsealed diffraction intensities to a resolution of 4 A for each pattern. In all patterns, the R/ rises sharply at a resolution of ~6.5 A. In patterns with reflections that extend to 4 A, the R/ begins to decrease at about 5.0 A.

We also used R factors to assess the quality of the merged diffraction data set. An overall R,, was calculated using the scaled image amplitudes measured from all patterns. This R factor provides an estimate of differences between Freidelrelated intensities for the entire data set. The merging R factor (R.meroe) provides an assessment of the variations in intensities between diffraction patterns. This value was calculated after merging the intensities. The overall merging R factor is 21.6%, and the overall R^is 16.6% (Table 1). Relative scale and isotropic temperature factors were determined for each pattern, by using the best pattern (-^"20) as a reference. The scale factors were determined by plotting ^(Iscaiedllobs) versus d~~ 2 (A~ 2 ) as discussed under Materials and methods. The patterns all have relative temperature factors close to zero (Table 1), reflecting the consistency of the crystals. Due to the intense central beam, 20 low-resolution spots were missing from the merged diffraction amplitude data set. Therefore, amplitudes previously measured from images provided this data. Temperature and scale factors were calculated for the low-resolution image data (with the merged diffraction data as a reference), and the scaled results were substituted for the missing diffraction amplitudes.

PIRRUCCELLO ET AL

198

TABLE 1 Crystallographic data from EriC diffraction patterns Pattern b(1,0) number (pixels)

ft (0,1) (pixels)

R

28

(-0.850, - 14.600)

(-22.650, 0.975)

0.0563 0.1950 0.0805* 0.2033*

20

(6.500, 13.000)

(22.300, -10.625)

0.0784

39

(8.925, 11.775)

(21.100, -15.925)

0.1031

40

(8.950, 11.700)

(21.100, -15.875)

Overall

/

R

.rjw

^mt•ry

dumber Resolution Scale factor

Of SpOtS (-4)

Temperature factor ( A2 1

254 544*

6.00 4.39*

0.22"?9

0.00986

0.08764

216

6.""

1.00

0

0.1176

233

6.61

1.1396

0.0737

0.0815 0.1729

234

6.61

1.3580

0.07334

937

6.5

0.16 55

0.21 65

* indicates a value calculated with data of higher resolution than used in the data set. All other values are calculated from the reflections used in the structure. Pattern #20 was the reference pattern, and thus had a scale factor of 1 and a temperature factor of 0.1^ is the intensity of reflection (h,k). TheFriedel relationship states that //,* = /_;,_£. The plane group of the crystal imposes an additional symmetry such that Ihk — /*-* = /-At — f-h-k-

ff

l,

ni,k = number of spots merged for that reflection.

A final projection structure was calculated to 6.5 A resolution using the phases obtained from image data (as previously published, Mindell et al 2001) with the merged amplitudes (Fig. 3). This map is grossly similar to the projection map derived from image data, but differs in detail (Fig. 4). The overall shape of the protein is little changed, but the surrounding lipid areas are flatter, with fewer ripples than before. Within the mass of the protein, there are several more clearlydefined circular areas of high scattering density, though the average density of the protein between these circular regions remains higher than that of the surrounding lipid. The four off-axis low-density areas observed in the original projection remain, and are somewhat more prominent in the new map. The density well on the twofold symmetry axis is deeper than in the image-derived projection.

C1C-TYPE CHLORIDE CHANNELS

FIG. 3. 6.5 A projection map of EriC. The map uses amplitudes calculated from electron diffraction patterns and phases calculated from image data. Four 137x76 A unit cells are shown. The zero contour line is roughly the average density of the lipid area; negative and positive contours are shown as dotted and solid lines, respectively.

Discussion We have collected electron diffraction patterns from two-dimensional crystals of EriC, the EscherichiacoliClC-tj^e. Cl~ channel protein, and used them to determine an improved 6,5 A projection map of the protein. These experiments demonstrate the feasibility of electron diffraction with these crystals and reveal their order to high resolution. The projection map derived using these data, while grossly similar to the image-derived projection, is different from the previous map in several details.

The diffraction patterns Electron diffraction patterns collected from EriC crystals display detectable reflections consistently to ~3.8 A. For this analysis, we chose to focus on data to

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PIRRUCCELLO ET AL

FIG. 4. Comparison between the previously calculated projection structure (using only image data) and the improved map. (a) Projection map calculated from scaled image amplitudes. This is the previously published structure (Mindell et al 2001). (b) Projection structure of EriC using diffraction amplitudes, same data as in Fig. 3. In this figure, lighter shades of grey correspond to higher scattering densities and darker shades correspond to lower scattering densities.

CIC-TYPE CHLORIDE CHANNELS

201

6.5 A resolution, which is the extent of our image-derived phase information. The orthorhombic unit cell dimensions are consistent with those measured from imaged crystals. These observations suggest that the crystals are sufficiently ordered to ultimately determine an atomic model of the protein. The R factors for this projection structure compare favourably with that of other structures in the literature. Electron diffraction data on the LHC (light harvesting complex, Wang & Kuhlbrandt 1991), for example, show values of R/~22% and R-merv ~35%. The average Ry for EriC to 6.5 A is 8% (calculated using data from Table 1), with 10% corresponding to the highest R^ for an individual pattern. The R*merge value for EriC is also comparable to that observed for LHC with a value of 22%. The frequent presence of strong reflections at resolutions greater than 4 A implies that the crystals are ordered to near atomic resolution and suggests that an atomic model may ultimately result from these studies. The symmetry of the diffraction patterns is generally consistent with the previously reported p22^2^ symmetry of the crystals. The ratio (p) of the square of the average amplitude ({|F|}2) and the average intensity ({/}) reflects the symmetry of a crystal (Wilson 1949). If the (h,k$) projection is centrosymmetric, as expected by the observation of p22j2i symmetry of imaged EriC crystals, p = 2/71 = 0,637, If the crystals are non-centrosymmetric, p = 0.785. For the diffraction intensities used in the structure, p = 0.588. This result strengthens the previous rinding of a twofold symmetry axis in the projection. In addition, systematic absences of the odd reflections on the h and k axes and apparent mirror planes in the untilted patterns along those axes both predict screw axes along h and k. Thus, alternating dimers are inserted in the membrane in opposite orientations. However, two reflections (the 13,0 and 17,0), expected to be absent, are present on every pattern. The meaning of these 'systematic presences' is unclear; perhaps they reflect a minor breakage of symmetry due to the interaction of the carbon support film on one side of the crystals. Consistent with this hypothesis is the observation that Rsym>R/ for all the patterns — in this case the increased Rfvm would reflect actual deviations from the expected identity of upward and downward facing protein dimers. Alternatively, both the systematic presence and the increased R sym values could reflect small crystal tilts, leading to slightly different values of ^* for nominally equivalent spots. In sum, the diffraction data support the p22j2j symmetry of the crystal, but perhaps with small deviations.

The projection structure

Figure 4 compares the projection map generated with boosted high-resolution image amplitudes with the new map using diffraction-derived amplitudes (Mindell et al 2001) and demonstrates the improvements in the new map.

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PIRRUCCELLO ET AL

A notable feature of the new map is the increased flatness of the lipid areas between the proteins. In the image-derived structure, the lipid areas show regularly spaced ripples, suggesting too much boosting of some high-resolution Fourier components. In the new projection, the lipid areas are smoother, as would be expected for a disordered lipid bilayer. The outline of the protein itself is similar to that in the previous structure, but several of the circular high-density areas, expected to be a helices, are better defined. Also, elongated high-density features are more linear, as would be expected for tilted helices, and the areas between helices are lower in density, giving an appearance similar to previously reported projection structures at this resolution. The four off-axis very low-density regions, previously interpreted as aqueous cavities in the protein, are still present in the new structure, and are indeed more prominent, supporting the speculation that these cavities form portions of the ionconducting pore. The new low-density area on the symmetry axis of the dimer rekindles the possibility of a single conducting pore, originally raised by Fahlke et al (1998). However, multiple recent electrophysiolgic studies on the eukaryotic CICs argue against this possibility (Saviane et al 1999, Lin & Chey 2000, Weinreich & Jentsch 2000). In addition, a low mass density is required on a protein symmetry axis; this is not sufficient evidence to demonstrate a conduction pathway (see, for example the structures of GlpF and aquaporin, Fu et al 2000, Murata et al 2000). In conclusion, we have demonstrated the feasibility of using electron diffraction to measure reflection intensities in 2D crystals of EriC. We find that the crystals are ordered to at least 3.8 A and that the diffraction intensities are reproducible from crystal to crystal. Using the intensities measured for untilted EriC crystals in combination with previously determined phases, we calculated an improved projection map at 6.5 A resolution. The new map is substantially unchanged, though some details seem slightly improved.

A. cknorvledgements This work was supported in part by NIH grant DK2801 to JAM.

References Baldwin J, Henderson R 1984 Measurement and evaluation of electron diffraction patterns from two-dimensional crystals. Ultramicroscopy 14:319-336 CeskaT, Henderson R 1990 Analysis of high-resolution electron diffraction patterns from purple membrane labeled with heavy-atoms. J Mol Biol 213:539-560 Fahlke C, Rhodes TH, Desai RR, George AL 1998 Pore stoichiometry of a voltage-gated chloride channel. Nature 394:687-690 Fu D, Libson A, Miercke LJ et al 2000 Structure of a glycerol-conducting channel and the basis for its selectivity. Science 290:481-486

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Havelka W A, Henderson R, Oesterhelt D1995 Three-dimensional structure of halorhodopsin at 7 A resolution. J Mol Biol 247:726-738 Lin CW, Chen TY 2000 Cysteine modification of a putative pore residue in C1C-0: implication for the pore stoichiometry of C1C chloride channels. J Gen Physiol 116:535-546 Maduke M, Miller C, Mindell JA 2000 Decade of C1C chloride channels: structure, mechanism., and many unsettled questions. Annu Rev Biophys Biomol Struct 29:411-438 Mindell J, Maduke M, Miller C, GrigoriefFN 2001 Projection Structure of a CIC-type chloride ion channel at 6.5 A resolution Nature 409:219-223 Murata K, Mitsuoka K, Hirai T et al 2000 Structural determinants of water permeation through aquaporin-1. Nature 407:599-605 Saviane C, Conti F, Pusch M 1999 The muscle chloride channel C1C-1 has a double-barreled appearance that is differentially affected in dominant and recessive myotonia. J Gen Physiol 113:457-468 Wang DN, Kuhlbrandt W 1991 High-resolution electron crystallography of light-harvesting chlorophyll a/b-protein complex in three different media. J Mol Biol 217:691-699 Weinreich F, Jentsch TJ 2001 Pores formed by single subunits in mixed dimers of different CLC chloride channels. J Biol Chem 276:2347-2353 Wilson AJC 1949 The probability distribution of X-ray intensities. Acta Cryst 2:318-321

DISCUSSION A shcroft: If your hypothesis is right and each pair of holes corresponds to a single pore with a bent path, would you not expect to see a reduced density between the holes? Mindell: If it is mostly protein on both sides, then it is a relatively narrow path and it isn't surprising that we don't see a very reduced density. I don't think that in this structure it argues against the kink. It is not a high density there. A shcroft: I find it fascinating that the channel has double-barrelled kinetics, as if each pore has its own gate, but there is also a common gate that shuts both pores. How do you imagine that the common gate operates, especially since the two pores seem to be so far apart? Mindell: That is a great question; I don't know. Presumably it is some kind of cooperative interaction across the subunits. But this doesn't say anything. One model we could consider is that there are two openings on one side of the membrane, converging to one opening on the other side. But there is clear evidence that this is not what is happening, because you can modify each pore independently from either the extracellular side or the intracellular side in C1CO. This argues strongly against there being a common opening on one side. Pero^o: Have you considered the possibility that, as with the aquaporins, this happens to be oligomeric just by chance, but is actually a monomer that exhibits this double barrel? Mindell: It is a possibility. However, even in the projection structure these two things in the monomer look quite different, and you wouldn't expect them to behave as identically as they do in any electrophysiology experiment. I think it is

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hard to argue that the two pores are in a monomer. You wouldn't see two identical pores with this kind of architecture, if that were to be the case. Wallace: How big are these pores? In your map, they look quite large relative to the size of water. Mindell: It is an issue I try to avoid, because I can't really measure the sizes of the pores. The density, especially in projection, is a continuous function, and I don't know where the side chains end and where the hole begins. The pores are in the order of 8 and 6 A, but this is not a meaningful number because there is no clear cutoff. Sansom: How selective are Cl ~ channels to different monovalent anions? Mindell: They are actually rather promiscuous. Almost any monovalent anion will go through. Sansom: Can large organic anions permeate? Mindell: Thiocyanate blocks, and I have some ideas of some heavy metal blockers that we are currently working on. Univin: What are the other monovalent anions that it has to worry about? Mindell: In physiological situations, there aren't really any. This is probably why it has never evolved an exquisite selectivity mechanism like the K + channels. Umvin: Couldn't there just be a 9 A hole, then? A Cl~ with its hydration shell should pass through this, whereas the other relevant anions would be too big. Mindell: Absolutely, and I think this is likely to be what is happening. There is reason to believe from the minimal selectivity that there is not a lot of dehydration. In fact, nitrate goes very7 well through many of these channels. A lot of them have higher conductances in nitrate than in Cl~. Nitrate looks a bit like a hydrated chloride. I think this is a reasonable model. Fu: The Cl~ channel has 12 putative transmembrane spanning regions. Is there any similarity between the Cl"" channel and other membrane transporters that contain 12 transmembrane spans? Mindell: There is no sequence similarity at all. There was recently a projection structure published of a 12 transmembrane domain transporter, and it doesn't look anything like this one. Swart^: Is there evidence from the dependence of gating on Cl" concentration for specific Cl ~ binding sites? Mindell: You are referring to fact that the voltage dependence of C1CO is carried by a Cl~ ion, and not by S4 or any charge on the protein. This implies that Cl~ is binding somehow and moving through the field on the process of opening. We don't know what this binding process has to do with permeation at all. In fact, another model that you could propose is that perhaps one of these holes is the binding pocket that opens the gate and the other hole is the pore. People have tried to look at this, and it is complicated. It looks like the two sites may have slightly different selectivity profiles, but not different enough to really tease this apart.

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Swart^j: Is it clear in your structure where the subunit interface is? Mindell: No. Sivart^: I think there are a couple of different ways you could approach this. Mindell: Nine months ago there were a lot of different ways we could go. Since then, Thomas Jentsch has published a paper in which he made tandem dimers of C1CO and C1C1 (Weinreich & Jentsch 2001). These have two very different pores. He sees a double-barrelled channel with one very large and one very small pore. This makes us think that the channel has to be entirely enclosed within a subunit. This eliminates many possible subunit interfaces. Choe: Is there the possibility of extending the phase beyond 6.5 A? Mindell: Yes. I already have some data to 5 A. Alok Mitra has 3.7 A, and I have every expectation that with the best images 1 will get that. The crystals are ordered to that high resolution. Mitra: You said that you had collected some low tilt images. At a certain viewing angle, the two sides of the kinked pore should appear approximately in register in a projected reconstruction along that direction. Mindell: I have seen tempting hints of things like this, but I don't have a good enough reconstruction yet to be able to say for sure. I am hoping that in the near future I'll be able to address this with a low tilt 3D structure. Pero^o: I have a question about your Cl ~ flux assay. Why does it stabilize around 1.3? Mindell: Every time we do these assays there seems to be a fraction of vesicles that never reconstitute with protein. Pero^o: Could it be that you have 30% dead channels? Mindell: If that was true, you should get to a point by adding more protein where you overwhelm this, and you have multiple channels per vesicle. We never saw that. The fact that it levels off suggests that this is just an inaccessible population. There are reasonable explanations for this observation, but in the fit we just deal with it by means of a fudge factor. Pero^o: How do you reconstitute these channels? Mindell: We make multilamellar vesicles and sonicate them. We then mix them with detergent solubilized channels. It is exactly the same as I showed with the 2D crystal, only with much less protein. Miller: Do you use synthetic lipid? Mindell: We can do it in synthetic lipids or Escberichiacoli lipids. The crystals are in synthetic lipid and they don't form in E. coli lipid. Sansom: It almost looks like there is a pseudo-twofold symmetry within a subunit. If you go to your high-resolution image, is that still there? Mindell: I haven't seen anything that makes me think of a pseudo-twofold symmetry. Unwin: Did you do this in glucose? Mindell: Yes.

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Unwin: Glucose has a similar density to the protein. Those holes are really quite pronounced. Mindell: It is quite dramatic. There shouldn't be much water here. We were initially a little puzzled as to why we could see the holes at all. Either the water is quite tightly bound there, or when drying in the glucose enough water, is trapped in for you to really see the holes. It will be interesting to look at these in ice, but we haven't done this yet because the glucose has worked so well. Ashcroft: What is known from mutagenesis studies about which residues get closest to the pore? You have an a helix really close to one of the pores. Mindell: There are a number. One of the important things to realize is that the pore will have contributions from many different helices. Because it is not in the symmetry axis, it is going to take a lot of different parts of protein to contribute to the pore. There is really only one set of residues in what we would call the D3/D4 loop that has been reasonably well shown to be lining the pore. It is on the basis of the cross-linking of those residues that the single pore model arose. This one-pore model came from an interesting experiment that I think was poorly interpreted. Christoph Fahlke in Al George's lab had a cysteine mutant of C1C1 that he showed quite nicely was lining the pore (Fahlke et al 1997). Then he found that with copper phenanthroline he could cross-link that cysteine and block the pore (Fahlke et al 1998). He couldn't imagine how that could happen if the cysteines were in two different pores, so he proposed a one-pore model. But he did all his experiments in macroscopic recordings. I look at the structure and think, what if this were D4, and the pore-lining cysteine was sitting on this helix? Then this could be facing the pore, but if part of gating involves a conformational change of this helix that then exposes it to the central axis, perhaps we are getting cross-linking when the channel is closed and this thing is facing the central axis. The advantage of this hypothesis is that it still has two pores and also that it is very testable. It predicts that the channel should only be cross-linkable in the closed state. We are in the process of trying to test this by looking at the state dependence of cysteine cross-linking. Swart^: This would also be a nice explanation for the inactivated state, too. Mindell: Absolutely. This is a nice way for there to be interaction across the dimer interface. If we can do this experiment then a range of other things become possible. References Fahlke C, Yu HT, Beck CL, Rhodes TH, George AL Jr 1997 Pore-forming segments in voltagegated chloride channels. Nature 390:529-532 Fahlke C, Rhodes TH, Desai RR, George AL Jr 1998 Pore stoichiometry of a voltage-gated chloride channel. Nature 394:687-690 Weinreich F, Jentsch TJ 2001 Pores formed by single subunits in mixed dimers of different C1C chloride channels. J Biol Chem 276:2347-2353

A protein chemical approach to channel structure and function: the proton channel of the vacuolar H+-ATPase John B. C. Findlay and Michael A. Harrison School of Biochemistry and Molecular Biology, University of Leeds, Leeds LS2 9JT, UK

Abstract. The vacuolar H+-ATPase provides a channel through which protons can be pumped across the bilayer. It is a complex assembly of about 20 subunits made up from 13 different polypeptide chains. The proton channel is located in the bilayer and therefore must be formed from one or both of the two intramembraneous subunits, called in yeast Vphlp (100 kDa) and Vma3p (16 kDa). Electron microscopy and the use of water soluble and hydrophobic chemical probes in conjunction with mutagenesis to cysteine or glutamic acid residues, suggest that the membrane sector consists of a single Vphlp subunit in association with a hexameric complex of the four-helical bundle Vma3p subunit. This hexamer encloses a large central pore lined by the first transmembrane helix. This pore appears to be impermeable, however; instead, a glutamic acid residue critical to transport function is located on the outside of the hexamer, deeply buried in the membrane and accessible to probes and inhibitors resident in the hydrophobic phase of the bilayer. The stoichiometry and chemistry of inhibitor binding supports the postulate that the mechanism of action involves rotation of the hexamer in the plane of the bilayer. Mutagenesis data on the Vphlp subunit indicate that it is vital to proton transport. It is likely, therefore, that the proton channel is formed at the interface of the Vphlp and Vma3p subunits, the protons moving via a network of interacting charged amino acid side-chains. 2002 Ion channels—-from atomic resolution physiology to functional genomics. Wiley, Chichester (Novartis Foundation Symposium 245) p 207-222

The vacuolar H+-ATPase (V-ATPase) is a multi-subunit complex whose function is to pump protons into intracellular compartments, or in some cases into the extracellular medium (Finbow & Harrison 1997, Stevens & Forgac 1997). It is an obligatory component of most eukaryotic cells, playing a central role in endocytic/exocytic processes, pH regulation, acid/base balance, bone resorption and so on. In overall organization and in the evolutionary relationship between individual subunits (Fig. 1), the V-ATPase resembles the mitochondrial 207

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FIG. 1. Domain organisation in the proton-motive ATPases. Both V-type (left) and F-type (right) ATPases comprise soluble nucleotide triphosphate-binding domains connected via stalk structures to a transmembrane proton translocating domain. Unlike the F-ATPase, which can operate in either ATP-generating or proton pumping modes, the V- ATPase is unidirectional and operates exclusively as a proton pump. The V-ATPase contains as many as thirteen different species of subunit, whereas the bacterial F^O-ATPase contains as few as eight. For the V-ATPase model, numbers indicate VMA gene designations in Saccharomyces. The principal ATP-binding subunits of the two complexes (V-ATPase subunits 1 and 2, F-ATPase subunits a and ft) are related at the sequence level, as are the proton-translocating subunit c proteolipids of the respective membrane domains. The relative positions of VMA4, VMA8, \'MA6, \ 7MA5 and KAL4 13 gene products within the V-ATPase complex are hypothetical.

F^FO-ATPase, consisting of an ATPase head group complex (Vj) joined to an intramembranous sector (Vo) by two stalk structures (Boekema et al 1997, Stock et al 1999). The movement of protons across the lipid bilayer is facilitated by the intramembranous sector, which consists of two species of integral membrane protein: The lOOkDa subunit a and 16kDa subunit r, or 16kDa proteolipid (Fig. 1). In Saccharomyces, a system widely used in the study of the V-ATPase because of its genetic pliability, these polypeptides are represented by the Vphlp and Vma3p subunits respectively. This contribution will outline the protein chemical studies which have given a low-resolution picture of the proteolipid component of the VQ membrane sector, and how it might interact with Vphlp to provide the permeation pathway of protons. Methodology The methodology used in this approach to channel structure and function consisted of generating site-directed mutant forms of the 16kDa proteolipid which could be expressed in the Saccharomyces system. The modified proteolipids could then be addressed by a variety of chemical agents whose interaction or lack of interaction with the polypeptides provides information on the organisation and

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disposition of the proteolipids within the membrane. These data, in conjunction with biophysical analysis and functional characterisation, have provided constraints for a low-resolution model of the proteolipid complex. This model in turn has allowed the mapping of residues central to activity, and therefore provided insights into the mechanisms by which protons are channelled across the bilayer. Results Outlines of the various experimental approaches used, the results obtained from them and the interpretation of these results in the context of the organization, structure and mechanism of action of the V-ATPase are given below. The basic strategy underpinning each experiment was to introduce residues with modifiable side chains (cysteines or glutamates) into a form of the 16 kDa proteolipid which is equivalent to the yeast Vma3p proteolipid. These modified proteolipids were then expressed in a yeast strain which is deleted for the native VMA3 gene and therefore unable to support V-ATPase function. The mutant proteolipids become incorporated into a hybrid V-ATPase, complementing the VMA3 mutation and restoring V-ATPase function (Harrison et al 1994). The modifiable side chains carried by the mutant proteolipids act as targets for sitespecific labelling or cross-linking experiments, providing information on membrane disposition and protein-protein contacts. Overall organization of the transmembrane sector Sequence analysis of the 16 kDa proteolipid family shows that there is a very high degree of conservation between species. Hydrophobicity plots and protease susceptibility studies support the presence of four transmembrane segments, consistent with a duplication event from the related 8 kDa subunit c of the F^FQATPase (Mandel et al 1988). FTIR spectroscopy analysis of the proteolipid from the arthropod Nephrops indicates that the protein is greater than 60% a helical (Holzenburg et al 1993), suggesting that these transmembrane regions were a helices, the N- and C-termini being in the intravesicular space. This proposed topography places a single potentially charged glutamate residue into the fourth transmembrane segment and is consistent with the two transmembrane helix structure proposed for F-ATPase subunit c. The proteolipid from Nephrops hepatopancreas can be prepared in substantial quantities, occurring as protein-dense membrane arrays with a degree of order. Electron microscopy studies on this negatively stained preparation have suggested a hexameric arrangement of these four-helix bundle subunits surrounding a central pore that is more than 15 A in diameter (Holzenburg et al

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1993). In the native state, this hexamer could be of six identical subunits. When expressed in Saccharomyces, the Nephrops proteolipid may interact with additional forms of yeast 16kDa proteolipid present as homologous subunits (Hirata et al 1997). Examination of the staining density after SDS-PAGE of isolated V0 domains from yeast indicate that the V-ATPase contains one Vphlp subunit per hexamer of 16 kDa proteolipids. From hydropathy plots, one suggestion is that Vphlp contains nine transmembrane regions (Leng et al 1999) though our analysis indicates a 7TM topography. Thus, to produce ATP driven movement of protons across the bilayer, the V-ATPase complex utilised about 13 different subunits (over 20 polypeptide chains in all), with the proton pathway arising from the assembly of up to 33 transmembrane segments. Clearly this must be a vital and highly regulated activity! Organization of the 16 kDa proteolipid Since recombinant 16 kDa proteolipids can incorporate functionally into a yeast VMA3 knockout strain, mutagenesis in combination with modification with a variety of chemical probes could be utilized to determine the location and orientation of potentially any residue side-chain in the intact V0 complex. Accessibility of introduced thiol side chains to water-soluble fluorescein maleimide can be assayed from mass shift of the isolated proteolipid on SDSPAGE caused by covalent attachment of the probe. Thus, cysteines on one face of helix 1 were freely available to fluorescein maleimide, which in contrast was shown not to react with any other transmembrane segments (Jones et al 1995). Conversely, the very hydrophobic pyrenyl maleimide (Fig. 2) was employed with

FIG. 2. Site-directed labelling of mutant 16kDa proteolipids. Lipid-soluble probes N-(lpyrenyl) maleimide (PM) and N-(l-pyrenyl)cyclohexylcarbodiimide (PCD) modify cysteinyl thiol and protonated carboxyl residues respectively. Fluorescence emission spectrum for pyrene-conjugated 16 kDa proteolipid is shown (right). Double labelling of pairs of sites on the proteolipid can result in long-wavelength excimer emission in the 470 nm region (broken line) only when fluorophores are closely aligned, indicating proximity between labelled residues.

PROTON CHANNEL OF THE V-ATPase

FIG. 3. Site-directed disulfide cross-linking of the 16 kDa proteolipid. The 16 kDa proteolipid is proposed to assemble as a hexamer of four helical bundles surrounding a central pore (left). A dimer of 16 kDa proteolipids was modelled (right) on the NMR-derived structure of the related 8kDa subunit c proteolipid from F-ATPase (Girvin et al 1998) using the programme Sybyl (Tripos Associates). Helices were positioned in accordance with experimentally observed disulfide cross-links. The derived position of the DCCD-reactive glutamate (El40) is consistent with accessibility to the lipid phase.

cysteine mutants to identify discrete loci on helices 2, 3 and 4 that were accessible from the lipid phase (Harrison et al 1999). The study was extended to include labelling of both the native glutamate residue of helix 4, and several glutamate mutant proteolipids. Modification of specific sites by the lipid-soluble and fluorescent pyrenyl analogue of dicyclohexylcarbodiimide (PCD; Fig. 2) confirmed the proximity of defined faces of helices 2, 3 and 4 (but not 1) to the lipid phase (Harrison et al 2000). Significantly, PCD modifies only protonated carboxyl groups, and forms a stable end product only in the absence of water. The final approaches to elucidating the helical packing arrangement come from disulfide-bond formation between introduced cysteine residues, and from complementary secondary mutations which restore function to disruptive mutations (Harrison et al 1999). Although such studies can be hard to interpret, what did emerge clearly was dimer formation with cysteines at positions on helix 1 predicted to be at the lipid/aqueous phase interface. Dimer formation also occurred involving a cross-link from cysteines positioned on opposite faces of helix 1 (Fig. 3: Residues Ala25 and Ala27). All these data strongly and consistently indicate that helix 1 from each proteolipid subunit is situated at the heart of the VQ complex, where they interact with one another lining a large channel accessible to water soluble reagents (Jones et al 1995). The most striking and highest yield intermolecular cross-links were obtained between cysteine residues positioned on helix 2 and helix 3 of adjacent proteolipids (Fig. 3: Glyl0l/104 and Gly62), indicating that these helices (in

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conjunction with helix 1) form the principal intermolecular contacts within the proteolipid hexamer (Harrison et al 1999). Interestingly, in all cases proteolipid dimers predominated, rather than higher order oligomers. This infers a degree of non-equivalence in the packing, giving rise to a 'trimer of dinners'. A model of the proteolipid dimer, in which the orientation of each helix is constrained only by the disulfide cross-linking data, is fully consistent with the pattern of lipid accessibility obtained from the labelling experiments. Thus, data obtained via a number of independent experimental approaches are mutually consistent and converge to produce a low-resolution model of the proteolipid complex (Fig. 3). Functionally sensitive sites Extensive cysteine mutagenesis indicated that the helices tended to have faces that were functionally sensitive (i.e. mutations adversely affected activity) and others that were much less so (Jones et al 1995, Harrison et al 1999). This was interpreted as revealing regions that are involved in helical interactions and packing on the one hand and exposed to the environment of the protein on the other. This interpretation highlighted a particularly interesting 'glycine stripe' region on one face of helix 3 of the proteolipid. This helical face appears to be critical for intermolecular helical contacts (see Fig. 3), and any modifications in this region profoundly perturbed activity. Recent work examining structural features that drive dimerization of integral membrane proteins has identified the motif GxxxG as a key influence (Ubarretxena-Belandia & Engelman 2001, Senes et al 2001). Helix 3 of the 16 kDa proteolipid contains a multiple form of this motif (GxxxGxxGxxxGxxxGxxG; Fig. 4), underlining its key role in driving assembly of the proteolipid complex. Cross-linking studies (Fig. 3) indicate that helix 3 makes contact with helix 2 of a neighbouring proteolipid. The presence of the GxxxG motif is proposed to allow multiple weak hydrogen bonds between the Cx protons of the glycine residues and oxygens of the neighbouring transmembrane helix (Senes et al 2001). Clearly, this form of interaction is important in driving assembly of the 16 kDa proteolipid complex. As will be discussed below, however, some residues/regions potentially accessible from the hydrophobic milieu and hence not thought to be liable to structural and functional restraint turn out to be vital to activity and reveal much about the mechanism of action of the molecular complex. The most significant of these is the glutamate residue in helix 4 of the proteolipid (Glul40 in the Xepbrops polypeptide). Removal of the negative charge abolishes all activity (Supek et al 1994). Furthermore, when the native and Glul40Gly mutant proteolipids are co-expressed in yeast, the proton-pumping ability of the resultant complex is reduced to such a level as to suggest that if as few as one of the six subunits in the hexamer is non-functional, then the whole complex is inactive (Hughes et al 1996).

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FIG. 4. The 'glycine stripe' region of helix 3 is crucial in helix-helix interactions. The helix 3 helix 4 region of the 16 kDa proteolipid, corresponding to the second subunit c-derived tandem repeat element, contains a glycine repeat motif with a-helical periodicity (left). This motif gives rise to a groove on helix 3 (right), presumed to allow close intermolecular packing and weak hydrogen bonding between the glycine Ca proton and oxygen of the neighbouring chain. The helical hairpin was modelled using Sybyl (Tripos Associates) and visualized using Rasmol.

It seems clear, therefore, that, like a corresponding aspartic acid in the F0-ATPase equivalent, this residue is an obligatory part of the mechanism of proton translocation. Situated near the middle of a hydrophobic transmembrane segment, it is reasonable to assume that this side chain is sequestered in the hexamer away from an hydrophobic environment. The protein chemical data, however, say otherwise. Dicyclohexylcarbodiimide (DCCD), an extremely hydrophobic reagent which covalently associates with protonated carboxylates, is a very potent inhibitor of proton pumping. The same is true of a bulky tributyl tin derivative, another potent inhibitor of both F- and V-ATPases, but one that does not bind covalently to the proteolipid (Hughes et al 1996). The interaction between this compound and the protein produces a substantial enhancement in the fluorescence yield of the probe. This can be used to demonstrate that the tin derivative can be displaced from the native complex (but not from the E140G mutant), by DCCD. The inference drawn from the results is that both hydrophobic probes are binding to approximately the same site on the protein,

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accessible from the lipid environment, and that the alkyl tin does not (unlike DCCD) require a carboxyl moiety for this interaction. A further line of study confirms that this glutamate is accessible from the lipid bilayer (and coincidentally that other glutamates can be placed in lipid exposed regions). After modification of these residues by the fluorescent PCD, quenching studies on reconstituted, pyrene-labelled V0 complexes show accessibility of the fluorophore only to lipid-soluble quenchers such as TEMPO and spin-labelled stearic acids (Harrison et al 2000). Related experiments using the parallax method (Chattopadhyay & London 1987) to determine distance of the pyrene fluorophore from the centre of the bilayer gave an approximate distance of 9A between the modified glutamate side chain and the centre of the bilayer (Harrison et al 2000). Measurement by ESR methods of spin-spin interactions between the paramagnetic DCCD analogue Ar-(2,2,6,6-tetramethyl piperidineoxy)-N'-cyclohexylcarbodiimide attached to Glul40 of the Nephrops protein and spin-labelled stearic acids also indicated that the glutamic acid was about 10 A from the hydrophobic/hydrophilic boundary at the bilayer surface (Pali et al 1999). In summary, therefore, all these data point to a lipid-accessible protonated residue located approximately midway into the cytoplasmic half of the lipid bilayer. Protein modelling studies which amalgamate the 'low resolution' data provided by this protein chemical approach would place this glutamate near the helix 2/helix 4 interface (Fig. 3), a location consistent with the position proposed for the equivalent aspartate residue in the F-ATPase F.0 domain (Rastogi & Girvin 1999). Implications for the proton channel The structural studies outlined above clearly establish that the key glutamate residue in the fourth transmembrane helix of the 16 kDa proteolipid is not sequestered within the structure of the V0 complex, but is on an outer surface exposed to lipid. The high sensitivity of this surface to functionality is further attested by the inhibitory potency of the hydrophobic single transmembrane papilloma virus protein E5 (Thomsen et al 1999). Binding of this oncoprotein is specifically to the proteolipid component (Goldstein et al 1991), and requires the presence of the transmembrane glutamate residue (Andresson et al 1995). This has profound implications for the mechanism of proton translocation, which must be unlike any process carried out by channels and transporters. Moreover, the fact that one dysfunctional unit inserted into the V-ATPase either by mutagenesis or the attachment of a single DCCD moiety is sufficient to disable the whole complex, implies a very high degree of cooperativity. The third element to be considered is the role of the other integral membrane protein present, Vphlp. Hydropathy plots project that this polypeptide could possess up to nine transmembrane segments, a

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FIG. 5. Operation of the vacuolar ATPase as a molecular motor. The ATP hydrolysing unit comprising (1) Vmalp and (2) Vma2p generates torque on a central stalk structure postulated to comprise subunits (4) Vma4p and/or (8) VmaSp. This structure is fixed relative to the 16 kDa proteolipid hexamer (3), which is then in turn rotated relative to the ATP hydrolysing domain. Membrane subunit Vphlp is proposed to form part of a stator element, fixed relative to the ATPase domain via a secondary stalk. Rotation generated by the turnover of ATP therefore generates movement of the proteolipid complex relative to the single copy Vphlp. The pathway of proton translocation is proposed to form transiently at the interface between the two subunits.

few of which intriguingly contain potentially charged residues. Mutagenesis studies have shown that removal of acidic (Glu789), histidine (His743) or basic (Arg735) residues abolishes proton pumping (Leng et al 1999, Kawasaki-Nishi et al 2001). It is clear, therefore, that at least one of the transmembrane segments (residues within the region 720-750) is deeply implicated in the mechanisms of proton translocation. The most straightforward interpretation is to postulate an interaction between this segment of Vphlp and the helix 2/helix 4 surface of the 16 kDa proteolipid (see Fig. 3). The interface between these two regions would thus provide the route by which protons migrate from one side of the membrane to the other, perhaps via a network of interacting, transiently protonated side chains.

Mechanism of action A static picture such as this, however, does not align completely with the effects of the various mutations and inhibitor studies. These are more consistent with a speculative mechanism proposed earlier (Harrison et al 1997), based on analogies with the FoF1-ATPase (Sambongi et al 1999, Fillingame 1999), involving rotation of hexameric 16 kDa proteolipid complex against the Vphlp subunit (Fig. 5). The latter is held static by interaction with components in the cytoplasmic sector of the

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V-ATPase assembly. Protons are picked up from the cytosol and funnelled to Glul40 on a single proteolipid subunit through the Vphlp-Vma3p interface. This facilitates rotation such that the now protonated glutamate moves out of the Vphlp-Vma3p interface and into a lipid accessible environment. The process can then be repeated until the protonated glutamate then migrates through 360°, reenters the interface and loses the proton to the intravesicular surface of the membrane. This mechanism would account for complete inhibition of proton translocation caused by a single dysfunctional proteolipid subunit and inhibitor binding to lipid-exposed surfaces of the V0 complex. The energy required to pump protons against their concentration gradient by such a mechanism would come from the hydrolysis of ATP by the head group region. Changes at the nucleotide binding site here would be translated to the top of the central axle (or stalk) and thence to the proteolipid hexamer tightly bound to the bottom of the axle. This is a rather simplistic view of a form of molecular motor, but one that accommodates most of the data collected so far. There are likely, of course, to be other components particularly in the cytosolic compartment that may play a role, at least in regulating this process, but what we have so far provides a basis for interpreting structural details from other structures.

Residual enigmas The low-resolution structural model that emerges from these studies suggests a large central channel which at first sight should facilitate the movement of quite large substances across the membrane. But it is clear that the V-ATPase or even the V0 transmembrane sector is not freely permeable. It follows that this pore is not freely accessible at least to compounds that cannot pass across the lipid bilayer. Yet side chains in this central region can be modified by probes which are to some extent water soluble but not by those which partition exclusively into the hydrophobic milieu. There are two possible explanations for these observations: the first is that the channel is tightly sealed, at least at one end, by protein elements either from the proteolipid itself, or more likely from other subunits in the complex. The cytoplasmic 'loops' connecting transmembrane helices 1 and 2 and helices 3 and 4 of the 16kDa proteolipid are certainly the most sensitive to mutational change, which can be interpreted as suggesting the presence of sites of interaction with cytoplasmic phase subunits. The other, not necessarily mutually exclusive, explanation is that the channel has within it an environment that is substantially hydrophobic thereby presenting a barrier to the diffusion of hydrophilic material. Only further detailed structural studies will resolve this puzzle.

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A cknowledgements We gratefully acknowledge financial support from the Wellcome Trust and the BBSRC for the studies reported here and the contributions of many colleagues, including Phil Jones, Malcolm Finbow, Derek Marsh, Andreas Holzenburg, Glen Hughes, Yong-In Kim, James Murray and Ben Powell.

References Andresson T, Sparkowski J, Goldstein DJ, Schlegel R 1995 Vacuolar H+-ATPase mutants transform cells and define a binding site for the papillomavirus E5 oncoprotein. J Biol Chem 270:6830-6837 Boekema EJ, Ubbink-Kok T, Lolkema JS, Brisson A, Konings WN 1997 Visualisation of a peripheral stalk in V-type ATPases: evidence for the stator structure essential to rotational catalysis. Proc Natl Acad Sci USA 94:14291-14293 Chattopadhyay A, London E 1987 Parallax method for direct measurement of membrane penetration depth utilizing fluorescence quenching by spin-labeled phospholipids. Biochemistry 26:39-45 Fiilingame RH 1999 Molecular rotary motors. Science 286:1687-1688 Finbow ME, Harrison MA 1997 The vacuolar H+-ATPase: a universal proton pump of eukaryotes. Biochem J 324:697-712 Girvin ME, Rastogi VK, Abildgaard F, Markley JL, Filingame RH 1998 Solution structure of the transmembrane H+-transporting subunit c of the F1F0 ATP synthase. Biochemistry 37:8817-8824 Goldstein DJ, Finbow ME, Andresson T et al 1991 Bovine papillomavirus E5 oncoprotein binds to the 16K component of vacuolar H+-ATPases. Nature 352:347-349 Harrison MA, Finbow ME, Findlay JBC 1997 Postulate for the molecular mechanism of the vacuolar H+-ATPase (hypothesis). Mol Memb Biol 14:1-3 Harrison MA, Jones PC, Kim Y-I, Finbow ME, Findlay JBC 1994 Functional properties of a hybrid vacuolar H+-ATPase in Saccharomyces cells expressing the Nephrops 16 kDa proteolipid. Eur J Biochem 221:111-120 Harrison MA, Murray J, Powell B, Kim YI, Finbow ME, Findlay JBC 1999 Helical interactions and membrane disposition of the 16-kDa proteolipid subunit of the vacuolar H+-ATPase analyzed by cysteine replacement mutagenesis. J Biol Chem 274:25461-25470 Harrison MA, Powell B, Finbow ME, Findlay JBC 2000 Identification of lipid-accessible sites on the Nephrops 16 kDa proteolipid incorporated into a hybrid vacuolar H+-ATPase: site directed labeling with N-l-(pyrenyl)cyclohexylcarbodiimide and fluorescence quenching analysis. Biochemistry 39:7531-7537 Hirata R, Graham LA, Takatsuki A, Stevens TH, Anraku Y 1997 VMA 11 and VMA 16 encode second and third proteolipid subunits of the Saccharomyces cerevisiae vacuolar membrane H+ATPase. J Biol Chem 272:4795-4803 Holzenburg A, Jones PC, Franklin T et al 1993 Evidence for a common structure for a class of membrane channels. Eur J Biochem 213:21-30 Hughes G, Harrison MA, Kim YI, Griffiths DE, Finbow ME, Findlay JBC 1996 Interaction of dibutyltin-3-hydroxyflavone bromide with the 16 kDa proteolipid indicates the disposition of proton translocation sites of the vacuolar ATPase. Biochem J 317:425-431 Jones PC, Harrison MA, Kim YI, Finbow ME, Findlay JBC 1995 The first putative transmembrane helix of the 16 kDa proteolipid lines a pore in the V0 sector of the vacuolar H+-ATPase. Biochem J 312:739-747 Kawasaki-Nishi S, Nishi T, Forgac M 2001 Arg-735 of the 100-kDa subunit a of the yeast VATPase is essential for proton translocation. Proc Natl Acad Sci USA 98:12397-12402

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Leng XH, Nishi T, Forgac M 1999 Transmembrane topography of the 100 kDa a subunit (Vphlp) of the yeast vacuolar proton translocating ATPase. J Biol Chem 274: 14655-14661 Mandel M, Moriyama Y, Hulmes J D, Pan Y-C E, Nelson H, Nelson N 1988 cDNA sequence encoding the 16 kDa proteolipid of chromaffin granules implies gene duplication in the evolution of FT-ATPases. Proc Natl Acad Sci USA 85:5521-5524 Pali T, Finbow ME, Marsh D 1999 Membrane assembly of the 16 kDa proteolipid channel from Nephrops norvegicus studied by relaxation enhancements in spin-label ESR. Biochemistry 38:14311-14319 Rastogi VK, Girvin ME 1999 Structural changes linked to proton translocation by subunit c in the ATP synthase. Nature 402:263-268 Sambongi Y et al 1999 Mechanical rotation of the c subunit oligomer in ATP synthase (F0F1): direct observation. Science 286:1722-1724 Senes A, Ubarretxena-Belandia I, Engelman DM 2001 The Ca — H • • • O hydrogen bond: a determinant of stability and specificity in transmembrane helix interactions. Proc Natl Acad Sci USA 98:9056-9061 Stevens TH, Forgac M 1997 Structure, function and regulation of the vacuolar H+-ATPase. Annu Rev Cell Dev Biol 13:779-808 Stock D, Leslie AGW, Walker JE 1999 Molecular architecture of the rotary motor in ATP synthase. Science 286:1700-1705 Supek F, Supekova L, Nelson N 1994 Features of vacuolar H+-ATPase revealed by yeast suppressor mutants. J Biol Chem 269:26479-26485 Thomsen P, Rudenko O, Berezin V, Norrild B 1999 The HPV-16 E5 oncogene and bafilomycin A1 influence cell motility. Biochim Biophys Acta 1452:285-295 Ubarretxena-Belandia I, Engelman DM 2001 Helical membrane proteins: diversity of functions in the context of simple architecture. Curr Opin Struct Biol 11:370-376

DISCUSSION Ashcroft: How similar is your V-ATPase to the F1/F0 ATPase? Findlay: It has elements of overall similarity in that some of the machinery for dealing with ATP is obviously closely related, both in sequence and in structure. The channel protein looks to be a dimer of the F0 one. It is an internally duplicated event, so instead of being two transmembrane, which it is in F0, it is four transmembrane, but both only have one transmembrane negative charge. In the F0 there is still a controversy as to how many subunits there are in the intact complex. There is debate as to whether it is 10 or 12. The stoichiometry may or may not be similar in terms of the total number of transmembrane segments. The V0 seems clearly to have 24. A shcroft: Does the F 1 /F o ATPase have this glutamate that you suggest functions as a proton shuttle? Findlay: It has an equivalent aspartate but we only have one glutamate for the four transmembrane segments of V0.

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Miller: You have presented one part of the channel in detail here. It looks as though we need another protein next to it. Do we know anything about that protein? Findlay: This protein in yeast is called Vphlp. We don't know much about it at the moment, but it does seem to have a number of charges in one of the transmembrane segments in particular. It is a monomer that looks to have six to nine transmembrane segments. There are data suggesting that mutagenesis of some of these charges kills proton pumping. It is a big step from this observation to saying that it is part of the proton channel, but there is a suggestion that it is functionally involved. Asbcroft: I have a problem understanding what is going on at the top. You suggest that the channel rotates against a fixed protein that runs the length of the channel in the membrane. From the cryoelectron microscopy pictures it looks like they are connected. What happens when this channel rotates? Findlay: In the cryoelectron microscopy pictures the top is knocked off. The supposition is that the ATP-utilizing top of the mushroom and the stalk that goes down into the membrane are fixed. The axle and the membrane component rotate with respect to that complex. ATP hydrolysis drives the axle that runs down from the head group into the transmembrane segment, and it drives the latter round. A. shcroft: So is the axle what is plugging the pore? Findlay: Yes, we think the axle may be plugging the pore, but we don't know which subunits are involved. Sansom: Going back in the light of the model that you now have to the mutagenesis experiments, do you think this suggests that glutamate residues can be tolerated in a relatively hydrophobic environment? Findlay. Yes, that was a surprise. We have a good assay for whether the protein folds properly in the membrane. If it goes into the bilayer and folds properly, the whole complex assembles. If in any way the fold is dysfunctional, then that head group is not attached. In many of these mutants, where there was a glutamate, the head group was assembled, so by and large the complex was going into the membrane correctly. The supposition from this is that it is possible to tolerate a protonated residue in the bilayer. Miller: In the membrane where the lipid carboxyls and head groups line up, what potential do they produce in the middle of the bilayer? Roux: The electrostatic potential inside the bilayer is slightly positive relative to the surroundings (McLaughlin 1977, Hladky & Haydon 1973). Umvin: In the structure of the F0, is the position in the membrane similar? Is it on the outside?

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Findlay: It looks to be roughly similar. Our guess before the FO structure came out was that it is wedged at an interface between two helices, but accessible from the lipid bilayers. I think this is still true. Scbrempf: Could you comment on the different number of subunits making the spring-like structures in the membrane. It seems to depend on the type of ATPase you are looking at. Findlay: Our impression is that this one always seems to be hexameric. I know there is a lot of controversy about the F1/F0 one. Some people get 10 and others get 12.1 have never worked on that subunit. Schrempf: There are systems such as Manduca, where it is known that if you take the head off, the main structure seems to survive. In such a situation these rings should be closed. Findlay: When we take the head off and then try to do transport studies, we get a complex that is impermeable to protons. We have always assumed that one of these small subunits is in some way associated with the top and doesn't allow anything to go through. But there is a recent paper that suggests that there is a Ca2+ calmodulin-dependent opening up of this pore. This structure is thought to be a dimer of hexamers formed head-to-head between two vesicles. Presumably, where the headgroups would be, the hexamers are associating. Under these circumstances, they are gap junction-like and if you add Ca2+ the central entity seems to open up. The mechanism is hard to envisage, because this is a large central pore. Schrempf: Could you exclude a kind of inhibitory protein? Findlay: My instinct is that there are proteins in there that regulate it, whether by blocking the pore or some other mechanism. Schirmer: I have a question concerning FO- It is a round-ish, circular structure in contrast to your star-like structure. How does this happen? Findlay: F0 is an even larger structure still, in which the helices are almost side-byside. I tried to follow what the protein chemistry is telling us. The cross-linking, juxtaposition and labelling work all suggest that no other one of those helices is exposed to the central pore. There is a suggestion (and this is part of the folklore of the field) that depending on the form of the protein you isolate, it can be the more compact structure or an extended one. Perhaps this is where we are seeing these two systems operating. The big channel one is a much expanded, slightly reorganized structure, which is due to self association. In contrast, the ATPase structure is associated with a different set of subunits. Schirmer: There are no electron micrographs of the full complex? Findlay: The work from Alan Brisson indicates the complexed structure. We have some early pictures of what it looks like. What I am hinting at is that in isolation it may look broader than it looks in intact complexes. Sansom: Does TM3 have a glycine-rich face?

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Findlay: Yes, there is a glycine stripe. It is very sensitive. If you mutate residues in that stripe, it seriously affects the structure and function of the protein. Sansom: Are there any other glycine rich patches on other helices? Findlay: Not obviously. A shcroft: How big is the hole in the central channel? Findlay: In our case, it is at least 15 A, so it is quite large, but is not as big as the F0, one. A shcroft: When they did the experiments in which the two vesicles were fused together, how did they measure the conductance through the two? Findlay: They measured the migration of choline from one to another. In fact, there is some older work from Israel's group that was not understood at the time (Birman et al 1990). He carried out a very similar experiment and found that there was Ca2+-dependent movement of acetylcholine from one compartment to the next in membrane preparations from Torpedo electric organs. This protein was implicated in the process. A shcroft: Is your ATPase modulated by Ca2+? Findlay: Not as far as we know. It is very pH sensitive. There are many interesting observations now emerging. I have just shown one lobster protein, but we have also used the native yeast protein and also ones from various other species that will substitute in yeast. But yeast actually has three of these vacuolartype channels, one of which seems to have five transmembrane domains. The suggestion is that the fifth one may be in the middle of the pore, implying that the pore may be big enough to take another ring of helices. Miller: Returning to that glutamate which you showed us was in the hydrophobic region of the membrane. A fatty acid in water has a pKa of ~ 2—4, but one sitting in a bilayer at the aqueous-lipid interface can have a pKa of ~ 6-7. So what would you expect of that carboxyl in the middle of a bilayer? If it is protonated, then it is not so unlikely to be in the bilayer. Findlay: It substantially reduces its polarity. Miller: It might be a lot better off in water, that's for sure. But is the energetic cost really so high as one would intuitively think? R.OUX: Once it is protonated, the free energy of this group is not much worse than that of a water molecule. The thing is, how likely is it to be protonated? It would be quite pH sensitive if it were. Findlay: That's what the environment between those two proteins is actually all about: the protonation process. A shcroft: Why is it so unusual to expect a glutamate to be in the middle of the bilayer, when the S4 domain of Kv channels has arginines and lysines? Findlay: The difference is that we are prejudiced from history to believe that in a hydrophobic (lipid) environment there would not be a charged residue. In the

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middle of the protein it is more easy to envisage that it would be tolerated because there could be compensating buried charges. Sansom: If you look at those membrane proteins for which we have crystal structures, then the frequency of finding a glutamate or arginine in a transmembrane helix is very low (Ulmschneider & Sansom 2001).

References Birman S, Meunier FM, Lesbats B, Le Caer JP, Rossier J, Israel M 1990 A 15 kDa proteolipid found in mediatophore preparations from Torpedo electric organ presents high sequence homology with the bovine chromaffin granule protonophore. FEBS Lett 261:303-306 Hladky SB, Haydon DA 1973 Membrane conductance and surface potentials. Biochim Biophys Acta 318:464-468 McLaughlin S 1977 Electrostatic potentials at membrane-solution interfaces Current Topic Membrane Transport 9:71-144 Ulmschneider MB, Sansom MSP 2001 Amino acid distributions in integral membrane protein structures. Biochim Biophys Acta 1512:1-14

Acetylcholine receptors, between closed and open Anthony Auerbach Center for Single-Molecule Biophysics and Department of Physiology and Biophysics, State University of New York, Buffalo, NY 14214,USA

Abstract. Muscle acetylcholine receptors switch between conformations that either allow ('open') or prohibit ('closed') ion permeation. We probed the dynamics of this structural transition using linear free-energy relationships. Specific regions of the protein were perturbed (mutations, voltage or agonists) and the opening and closing rate constants were estimated from single-channel currents. Usually, a log—log plot of rate constant versus equilibrium constant was linear, with the slope indicating the sensitivity of the transition state to the perturbation as being between that of the open and closed conformation. The spatial gradient in this slope, from open-like at the transmitter binding sites to closed-like at the middle of the membrane, suggests that gating is a wave that propagates between the binding sites and the membrane domain. 2002 Ion channels—from atomic resolution physiology to functional genomics. Wiley, Chichester ( Novartis Foundation symposium 245) p 223—239

The muscle acetylcholine receptor (AChR) is an allosteric protein that gates — it adopts alternative structures in which its ion permeation pathway is either closed or open. When the two transmitter binding sites are empty, the AChR gating equilibrium constant is exceedingly low (10- 5 —10- 7 ). When the transmitter binding sites are occupied by acetylcholine, the gating equilibrium constant increases by about six or seven orders of magnitude. The driving force for this increase favouring the open conformation is its ~2500-fold higher affinity for the transmitter. Once the transmitter binding sites are occupied by acetylcholine, the channel opens within ~20ms. The AChR is a low-noise, high-gain, high-speed molecular switch. The structural components of this switch are known only at 4—5 A resolution (Unwin 2000), although the structure of AChBP, a homologue of the AChR extracellular domain, is known at atomic resolution (2.7 A; Brejc et al 2001). The AChR is composed of five homologous subunits and is approximately cylindrical, with a long axis of ~150 A and a diameter of 50-80 A. Both electron crystallography and the folding topology predicted from the primary sequence indicate that ~50% of the amino acids form the extracellular domain, ~20% are within the membrane 223

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and ~30% form the intracellular domain. Each transmitter binding site lies at a subunit interface situated about midway up the extracellular domain, ~45 A above the middle of the membrane. About a third of the protein, or >750 residues, separates these locations. A comparison of the closed- and open-channel AChR structures at a resolution of 9 A shows that gating involves substantial movements of regions of the protein that are between the binding sites and the gate (Unwin 1995). This observation is supported by functional studies showing that mutations of many of the residues that are thought to lie in this region influence the gating equilibrium constant. AChR gating is a massive chemical reaction that is likely to involve the movement of thousands of atoms and changes in thousands of non-covalent bonds. In single-channel recordings, the transition from a non-conducting to a conducting conformation appears as an instantaneous jump in the current. The energetics of the transitions between stable states of the protein can be inferred from the molecular rate constants for gating, which in turn can be estimated from the durations of the closed and open intervals in a single-channel record. Estimating the gating rate constants is not always straightforward because there are many open and closed stable states and some have lifetimes that are brief and difficult to measure. However, there is a consensus that for the adult mouse AChR activated by acetylcholine (—100 mV, 22 °C), the diliganded opening rate constant (b2) is ~50000s~ 1 and the diliganded closing rate constant (x 2 ) is ~2000s- ! , yielding a diliganded gating equilibrium constant (b 2 /x 2 = Q2) of ~25. The channel opening and agonist dissociation rate constants are fast and the lifetime of the diliganded-closed state is close to the time-resolution of the instrumentation (~10ms, on a good day). It is therefore difficult to obtain accurate estimates of b2 using wild-type AChR activated by acetylcholine. Fortunately, a number of perturbations reduce this rate constant so that it can be more readily measured. For example, b2 depends on the agonist used to activate the channel. Choline, a breakdown product of acetylcholine, supports an opening rate constant of only ~200 s~ 1 . At high choline concentrations (where binding is saturated), the diliganded-closed state of the AChR has a lifetime of ~5 ms, which is well within the instrumental limits. A notable feature of the durations of open and closed intervals associated with gating is that they are each distributed as a single exponential. That is, there is no indication of stable (> 10 ms) intermediates in the gating reaction. This observation allows us to define diliganded AChR gating as a concerted, single-step reaction:

A2C

y.-.

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where A2C is the diliganded-closed receptor and A2O is the diliganded-open receptor. There are several ways in which to rationalize this apparently simple behaviour with the enormity of the gating reaction. One possibility is that one of the many molecular events that constitute gating is much slower than the others and thus limits the overall rate of the reaction. Data are presented below that argue against this hypothesis. Another possibility is that no single molecular motion is ratelimiting, but that all of the intermediates of the reaction are brief. In this case, the apparent gating rate constants do not reflect any single microscopic molecular transition, but rather the aggregate lifetimes of ensembles of short-lived, structurally distinct conformations (microstates) in which the channel is either closed or open. By analogy, the decay of the endplate current appears as a single exponential even though it reflects the aggregate lifetime of all diliganded states, including the brief intermediate state A2C. The single exponential behaviour of AChR gating allows us to invoke a simple scheme for the reaction: one barrier, two wells. Such a reaction is usually drawn as two intersecting parabolas that represent the lowest energy profile across the energy landscape connecting the endpoints of the reaction, A2C and A2O. Kinetics, however, does not provide direct information on the shapes of the wells or the barrier, or on the absolute free energies of the stable states. Rather, the rate constants yield only two points on the pathway relative to the starting point, the free energy of the endpoint and the free energy of the transition state. Both of these energies reflect the properties of ensembles of microstates. The free energy of the transition state ensemble pertains to the whole system — ligands, protein, water, ions and membrane. This aggregate energy is spread over many microstates and probably involves many of the thousands of atoms and bonds that move during gating. It therefore is likely to reflect structures that are spatially dispersed. Perturbations can be used to probe the character of the gating pathway in a restricted region of the protein. Different agonists specifically perturb the vicinity of the transmitter binding sites. The membrane potential specifically perturbs the charges and/or dipoles that govern the voltage sensitivity of the reaction (which are, presumably, in the membrane domain). Mutations perturb the local environment near the altered side chain. There is always the possibility that each of these experimental manipulations could have a long-range effect on the system, but for the moment we will assume that there is only a localized perturbation. With this assumption, it is possible to extract information about the position of the perturbed region along the reaction coordinate. The method is rateequilibrium linear free energy (LFER) analysis, first used in physical organic chemistry by J. Bronsted (see Leffler & Grunwald 1989) and more recently by A. Fersht and others to probe protein folding pathways (see Fersht 1999).

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The concept is simple. Suppose that a small perturbation of the system differentially affects the free energies of the closed (C = DG°losed) and open (O =DG°open) ensembles. This, of course, will result in a change in the gating equilibrium constant. The key LFER assumption is that the effect of the perturbation on the transition state free energy (T = G++) is a linear combination of its effects on the endpoints. That is, the sensitivity of the transition state to the perturbation is a fraction (F) like that of the open state, and (1 — <1>) like that of the closed state: T* - T = F(O* - O) + (1 - F)(C* - C)

where * indicates the perturbed condition and F is a number between zero and one. F is an interesting parameter because is describes the character of the transition state at the site of the perturbation. However, from the form of this equation it does not appear to be readily accessible from the experiments. A simple rearrangement of the terms leads to a formulation that can be directly related to the experimentally derived rate constants: (T* - C*) = F(O* - C*) - F(O - C) - (C + T)

(T* — C*) is proportional to the log of the opening rate constant and (O* — C*) is proportional to the log of the gating equilibrium constant. The other terms in the equation do not vary with the perturbation and can be incorporated into a single constant. The equation can now be written as log ( b 2 ) = F log (Q2*) + constant. If the linear combination assumption holds true, for a series of localized perturbations the plot of the log of the opening rate constant versus the log of the gating equilibrium constant should be a straight line. The slope of this line is Q (the fractional sensitivity of the transition state compared to the open state), and the ordinal intercept represents the intrinsic barrier to gating, i.e. the free energy of the transition state when the gating equilibrium constant is unity. For a one-step gating reaction the slope of the relationship between the log of the closing rate constant and the log of the equilibrium constant is, by definition, F — 1: log (x2*) = (F - 1) log (F 2 *) 4- constant According to these formulations, b2 ocQ2F*and y.2 oc Q2F-1, It is more direct, and more statistically correct, to quantify the slope of the relationship between the logarithms of the forward and reverse rate constants (Estell 1987). However, in this form the slope is equal to (F — 1)F and the

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sought-after value of F is less intuitively gained from an examination of the plot, Note that for the extreme values of F (0 and 1) the transition state energy will track that of the closed and open state, respectively, so there will be no correlation between the rate and equilibrium constants. The LFER method is familiar to electrophysiologists who study the voltagedependence of channel block by a charged ligand. Often, the kinetics of a singlestep block reaction are influenced by the membrane potential, and voltage sensitivities of the blocking and unblocking rate constants are used to estimate the position of the barrier or well as an 'electrical distance'. The standard assumptions are that the voltage influences only the blocker and not the protein (i.e. that the perturbation is local) and that the electric field changes linearly with distance across the membrane. This is strictly analogous to the LFER approach, in which F provides an estimate of the position of the barrier along the reaction coordinate in the region of the perturbation. We applied LFER analysis to several positions in the AChR (Grosman et al 2000a). In one experiment, 11 different side chains were substituted for a serine at position 12' of the d subunit. This residue is situated in the M2 membrane-spanning segment (which lines the pore) and is three residues removed (towards the extracellular side) from a leucine that lies near the middle of the membrane. Remarkably, the LFER at d12' is indeed linear over a range of equilibrium constants that spans more than three orders of magnitude (Fig. 1). This supports the assumption that there is, at this position, a linear relationship between the sensitivity of the transition-state free energy and those of the end states. This relationship, which could have been anything, is a straight line because the perturbations were sufficiently small that the linear approximation works. The lack of curvature over such a wide range of equilibrium constants suggests that the reaction mechanism is similar for all of the perturbed structures. A simple interpretation of this result is that there is a restricted set of pathways, or perhaps a single reaction pathway, for the gating of diliganded AChR. The wild-type point falls on the same line as the mutants. In this case the mutations did not disrupt the overall characteristics of the gating reaction, insofar as the relative sensitivity of the transition state is concerned. Grosman et al (2000a) found that the linearity and slope of the relationship was the same with two different weak agonists, choline and acetylthiocholine. The F value for the mutation series at d12' is 0.28. At the transition state, which is determined by all of the atoms and bonds in the entire system, this location is ~30% like the open state and ~70% like the closed state, with regard to its sensitivity to the perturbation. The relationship between free energy and structure is not known and is complex, but cannot be arbitrary. Accordingly, we interpret the slope of the LFER at d12' to indicate that this residue has a structure that is only 30% open-like when the system is at the transition state.

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FIG. 1. Analyses of a mutation series of a single AChR residue. The serine at the 12'position of the d subunit (in the second membrane-spanning segment) was mutated to 11 different amino acids. Each circle reflects a different side chain (left panel, left to right: A, S, G, W, C, T, I, Y, V, N, Q, K). Left: Log rate-equilibrium LFERs (Bronsted plots). The relationships are linear and the slopes for the opening and closing rate constants are complimentary, consistent with a single-step reaction and a single reaction pathway. The values of F are 0.28 + 0.02 (opening) and 0.27 + 0.02 (closing). The opening and closing rate constants are equal (Q = 1) at 645 s-1. The arrow marks the wt (serine). Right: Log rate-rate relationship. The slope of this line is -2.438 ±0.267 and yields Q = 0.30 (correlation coefficient =0.945; dotted lines are 95% confidence limits). These analyses indicate that the d12' side chain is ~30% like that of the open state and 70% like that of the closed state (with regard to its sensitivity to perturbations) at the gating transition state.

Other regions of the protein have been explored using LFER analysis (Grosman et al 2000a). The transmitter binding sites were probed both by mutations (eight different residues) and by changing the agonist (13 different ligands). In this region, the F values were always in the range 0.9-1.0. That is, these perturbations altered the opening rate constant (in some cases, more than 1000-fold), but left the closing rate constant essentially unchanged. According to the above interpretation, the structures of the liganded binding sites at the transition state closely resemble those of the open conformation. At the transition state, when the pore is mostly closed, the agonists and the residues at the binding sites are in the high affinity configuration. The status of the membrane domain at the transition state was investigated further both by changing the voltage and by mutations. For diliganded AChR, the F values of the voltage sensor (which is in the membrane) and for six residues below the centre of the d subunit M2 segment (i.e. the intracellular limb) were all approximately 0. These perturbations altered the closing rate constant, but left the opening rate constant essentially unchanged. At the transition state for diliganded gating, the cytoplasmic limb of M2 is in the closed configuration.

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FIG. 2. The spatial map of F values in the AChR. Centre: AChR topology, as predicated from the sequence. The transmitter binding sites (TBS) and the gate (in M2) are separated by ~45 A. Left: there appears to be a gradient in F values along the long axis of the protein, from ~1 at the TBS to ~0 below the middle of M2. The results are from mutation series of 27 different residues. Right: a high-resolution map ofF values in the d subunit M2 segment. For the upper half of this segment, 0.42 < 0.28 (grey circles) and for the lower half,
Several locations presumed to be between the binding site and the gate have been probed using LFER analysis. Three residues in the extracellular linker of the a subunit are mostly open at the transition state (F 0.8; Grosman et al 2000b). Two residues near the extracellular margin of M3 and M4 in the a subunit are approximately midway between open and closed (F ~ 0.45; Wang et al 1999, Bouzat et al 2000). Four residues in the extracellular limb of M2 of the d subunit are mostly closed (F ~ 0.3) while those in the intracellular limb are completely closed (F ~ 0) (G. Cymes et al 2001). From these few measurements we can construct a low-resolution map of F values (Fig. 2). There appears to be a gradient of F values along the long axis of

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the AChR, from 1 (open-like) at the binding site, to 0 (closed-like) at the gate. The map, however, is quite fuzzy. Only a handful of positions have been probed. Moreover, the estimate of F at each position is imprecise and depends both on the number of mutations and the extent of the change in the equilibrium constant. Finally, without atomic resolution, the placement of a specific amino acid within the AChR structure is inexact. Thus the granularity of the gradient in F, if it indeed exists, is uncertain. F may change in a continuous manner, or may change in a block-wise fashion with discrete domains of the protein each having a single, characteristic F value. The spatial distribution of F values, which in general could have been anything, suggests that there is some underlying spatial organization to the gating reaction. One interpretation is that AChR gating is concerted (there are no stable intermediates) and asynchronous (all of the atoms do not move at the same time). If the residues near the binding site were the first to move and those near the gate the last, then the .F values at the binding site should be larger than those at the gate, as observed. For a single-step reaction the principle of detailed balance constrains closing to be the reverse of opening. Thus, according to this interpretation, when the AChR closes the movement of residues at the gate precedes those at the binding site. The gradient in F values suggests that the structural motions that constitute gating proceed as a reversible 'conformational wave', which has a locus of initiation (the binding site or the gate), and propagates along the long axis of the protein. The spatial gradient in F does not illuminate the mechanism of propagation. One possibility is that a conformational wave occurs because the perturbations in one residue (or domain) are coupled to those of its neighbours. In this 'knock-on' mechanism, one side chain influences the disposition of its neighbours, as in a chain of dominoes. Another possibility is that the protein motion starts synchronously, but that there is an environmental gradient (e.g. in mechanical compliance) that determines the spatial distribution of F. (Detailed balance requires the gradient to invert upon gating.) Either mechanism could produce the conformational wave. Kinetic measurements also do not address the nature of the forces that drive propagation, which are likely to involve everything that governs water, lipid and protein structure. F values only suggest a sequence of events, and say nothing about their timing. A lower limit for the propagation time of a conformational change might be derived from the speed of sound in water (1500ms"1), which can cover 50 A in ~3 ps. A reasonable upper limit is the inverse of the opening rate constant which for acetylcholine is ~20^s. This time range covers more than six orders of magnitude. The wave could be even slower, as single-channel measurements only record the moment in the reaction where the rate of ionic flux changes dramatically. It would be extremely valuable, although probably difficult, to

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obtain a more direct experimental measurement of a propagated conformational change during gating, perhaps using fluorescence. The presence of a gradient inF argues against the possibility that one of the microstate transitions that constitutes gating is slower than the others and serves to limit the overall reaction rate. If this were the case we would expect the map of F values to be all ones or zeros, depending on whether the perturbed region changed before or after the rate-limiting rnoiety. The large number of positions exhibiting fractional F values argues against a single, rate-limiting step in the AChR gating reaction. A long-range goal is to relate the map of F values to the sequence of microstate structural transitions that constitutes gating. The structures of the closed and open AChR (Unwin 2000) and AChBP (Brejc et al 2001) are the starting points. In the closed AChR, gating begins at the transmitter binding sites. In the AChR and the AChBP, this region is replete with aromatic residues, which form an energetically favourable environment for cationic agonists (Fig. 3A). The position of a HEPES molecule fortuitously bound in the pocket of AChBP is consistent with functional data indicating that the indole ring of W143 (AChBP numbering) interacts with the quaternary amine of acetylcholine (Zhong et al 1998). Mutations of four of the residues in the mouse AChR a subunit (Y89, Y185, Y192 and D194) significantly reduce diliganded gating, specifically by lowering the opening rate constant. That is, for these residues F & I. Mutations of the tyrosine residues have multiple consequences. In the adult mouse AChR, the order of the effect of a phenylalanine substitution is 185 > 89:»192 (Chen et al 1995, Auerbach et al 1996, Akk et al 1999). In addition to decreasing the diliganded gating equilibrium constant, these mutations also decrease the affinity of the closed conformation and the unliganded equilibrium gating constant (Grosman & Auerbach 2000). D194 is unique among these four because an asparagine substitution has almost no effect on the affinity of the closed conformation of the binding site (Akk et al 1996). Two other mutations, aS268I (Grosman et al 2000b) and eD175N (Akk et al 1999), also reduce diliganded gating without perturbing the net stability of the closed-liganded AChR, but these residues are not present in AChBP. In the open AChR, gating begins near the middle of the membrane domain. Unwin (1995, 2000) has suggested that a change in the disposition of a hydrophobic girdle formed by the 9' and 10' residues of the M2 segment (see Fig. 2) renders the channel permeable to ions. The F-value analysis (of the d subunit M2 segment) is consistent with this hypothesis. The intracellular limb of M2, from 10' to 2', is all fully closed-like (F = 0) at the transition state. Mutations of the 11' side chain do not influence the gating equilibrium constant, suggesting that this residue is in a similar environment in the closed and open conformations. The remaining resides that have been probed in the extracellular limb of M2 have a

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FIG. 3. (A) Close-up of the transmitter binding site of AChBP (Brejc et al 2001). The view is from the minus (non-a, in the AChR) side of the binding pocket. The residue numbers are for AChBP. The vicinal cysteines (Cl 87-188) that signify an AChR a subunit are at the tip of loop C (left foreground). The ligand is HEPES. W143 is on loop B, which follows the b7 strand. \X"53 is on the minus side of the pocket. Mutations of the three Tyr residues reduce both ACh binding (to the closed conformation) and channel gating, while mutations of D194 only reduce gating. These four residues all have F % 1. Distances to the plane of W143 are, approximately, 10 A (D194 carboxyl; Y185 hydroxyl), 8.2 A (Y192 hydroxyl) and 6.4 A (Y89 hydroxyl). (B) Transverse sections at the level of the ligand binding site. (Left) The unliganded Torpedo AChR oty subunit (from Fig. 10, Unwin 2000). Tunnels appear to link a central cavity with the extracellular (left arrow) and vestibule (lower arrow) compartments. (Right) The liganded Limnea AChBP (Brejc et al 2001). The vicinal cysteines on loop C and the binding pocket are at the upper left (\X 143; Y192, Y89 and Y185 are in grey, the start indicates the location of the HEPES ligand). W55, which contributes to the minus side of the pocket of the adjacent interfacial binding site, is in grey at the lower right. The distance between W143 and W55 is approximately 24 A. The arrow indicates the b7 strand. The cavity region of the AChR (left) is occupied by hydrophobic residues in the AChBP (black dots, drawn to a radius of 1.4 A around each atom), thus forming the core of a ft sandwich. A testable hypothesis is that ligand binding triggers the formation of this hydrophobic core, which contributes to the stability of the open-channel (or desensitized) AChR conformation.

similar F value of ~0.3, which suggests that during opening they move before the 10' position. Conformational changes to the extracellular domain that accompany gating of the AChR can be inferred from a comparison of unliganded Torpedo AChR (Unwin

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2000) and liganded Limnea AChBP (Brejc et al 2001). Structural elements of the AChBP correspond to electron density in a map of the homologous AChR a subunit in the closed state (Fig. 3B). These include loop C and a number of b strands on the molecular perimeter. In the AChBP structure, however, these elements appear to be positioned so that a central cavity, present in the a subunit of the closed AChR, has collapsed inwards to form a b sandwich. This central cavity is less apparent in the other AChR subunits, as well as in the open-channel structures of the a subunits (Unwin 1995). This suggests that the tertiary fold of the AChBP may correspond to the open (or desensitized) AChR. The strands of the AChBP b sandwich (in particular, b7) appear to lie closer to the centre of the subunit than in the closed AChR. The disposition of the b strands in AChBP is consistent with an anti-clockwise twist in the a subunits at the level of the binding sites that is observed to occur during the closed-to-open gating transition (Unwin 1995). Trpl43 arises from loop B, forms the base of the ligand-binding pocket and is immediately adjacent to b7'. It therefore seems reasonable to speculate that agonist binding triggers a conformational change that results in an inward, radial motion of pi and perhaps other strands. Accordingly, the reversible formation of a hydrophobic core between b sheets could, in part, determine the relative stability of the closed versus the open AChR conformations. This hypothesis can be tested by a combination of mutagenesis, cysteine-accessibility measurements, kinetic analysis techniques, and F value measurements in the region of b7 and elsewhere. With regard to future kinetic experiments, the first priority is to refine the map of F values. Many more residues, in each of the different subunits, need to be probed for their positions in the reaction. The structure of AChBP will serve as a guide to locate specific residues in the extracellular domain of the AChR. It will also be interesting to probe the gating trajectories of unliganded and monoliganded AChR, as there is evidence that AChRs without agonists open via a different or altered path (Grosman et al 2000a). Testing for correlations in F values (i.e. measuring F on mutant backgrounds) may provide information regarding the shape of the transition state ensemble. Finally, it is important to probe the 0 map for gating in other ion channels, if not other non-channel allosteric proteins. Reactions in which the transition state occurs early or late in the reaction would not be expected to show a gradient in F values even if a wave were present. Reactions in which a F gradient is apparent, however, may illuminate more general mechanisms of conformational changes in proteins. A cknow ledge ments I thank Gisela Cymes, Claudio Grosman, Jacqueline Gulbis, Meyer Jackson, Stuart Licht and Fred Sachs for comments on the manuscript. I am particularly indebted to Nigel Unwin for allowing a joint inspection of his unpublished AChR density maps, and for active discussions regarding the comparative structures of the AChR and the AChBP. Supported by NS-23513.

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References Akk G, Sine S, Auerbach A 1996 Binding sites contribute unequally to the gating of mouse nicotinic alpha D200N acetylcholine receptors. J Physiol 496:185-196 Akk G, Zhou M, Auerbach A 1999 A mutational analysis of the acetylcholine receptor channel transmitter binding site. Biophys J 76:207-218 Auerbach A, Sigurdson W, Chen J, Akk G 1996 Voltage dependence of acetylcholine receptor gating: different charge movements in di-, mono-, and unliganded receptors, j Physiol 494:155-170 Brejc K, van Dijk WJ, Klaassen R et al 2001 Crystal structure of Ach-binding protein reveals the ligand-binding domain of nicotinic receptors. Nature 411:269-276 Bouzat C, Barrantes F, Sine S 2000 Nicotinic receptor fourth transmembrane domain: hydrogen bonding by conserved threonine contributes to channel gating kinetics. J Gen Physiol 115:663-672 Chen J, Zhang Y, Akk G, Sine S, Auerbach A 1995 Activation kinetics of recombinant mouse nicotinic acetylcholine receptors: mutations of alpha-subunit tyrosine 190 effect both binding and gating. Biophys J 69:849-859 Cymes G, Grosman C, Auerbach A 2002 Structure of the transition state of gating in the acetylcholine receptor channel pore: F -value analysis. Biochemistry, in press Grosman C, Auerbach A 2000 Kinetic, mechanistic, and structural aspects of unliganded gating of acetylcholine receptor channels: a single-channel study of second transmembrane segment 12' mutants. J Gen Physiol 115:621-635 Grosman C, Zhou M, Auerbach A 2000a Mapping the conformational wave of acetylcholine receptor channel gating. Nature 403:773-776 Grosman C, Salamone FN, Sine SM, Auerbach A 2000b The extracellular linker of muscle acetylcholine receptor channels is a gating control element. J Gen Physiol 116:327-340 Estell DA 1987 Artifacts in the application of linear free energy analysis. Protein Eng 1:441-442 Fersht A 1999 Structure and mechanism in protein science: a guide to enzyme catalysis and protein folding. WH Freeman, New York Leffler JE, Grunwald E 1989 Rates and equilibria of organic reactions: as treated by statistical, thermodynamic, and extrathermodynamic methods. Dover Publications, New York Unwin N 1995 Acetylcholine receptor channel imaged in the open state. Nature 373:37-43 Unwin N 2000 The Croonian Lecture 2000. Nicotinic acetylcholine receptor and the structural basis of fast synaptic transmission. Philos Trans R Soc Lond B Biol Sci 355:1813-1829 Wang HL, Milone M, Ohno K et al 1999 Acetylcholine receptor M3 domain: stereochemical and volume contributions to channel gating. Nat Neurosci 2:226-233 Zhong W, Gallivan JP, Zhang Y, Li L, Lester HA, Dougherty DA 1998 From ab initio quantum mechanics to molecular neurobiology: a cation-pi binding site in the nicotinic receptor. Proc Nad Acad Sci USA 95:12088-12093

DISCUSSION Jordan: What temperature do you work at? Auerbach: Room temperature: 22-24 °C. Jordan: Can you do it at a different temperature? You had a short break at the end between the F of 0.3 and a F of about 0, and you give this a structural interpretation. Would you expect this to alter much if you changed the temperature? Rates of reaction are pretty temperature dependent.

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A uerbach'. We could change the temperature. We could probably work within a 20-30 °C range of temperature, but I'm not sure how much we would learn from such experiments. Jordan: You are breaking up the rate constant, and you have entropic and enthalpic effects in here. You are trying to get structural inference out of this, and you would like to see that your F isn't terribly temperature dependent. Fu: I think the temperature could be important. In this linear function, two parameters determine the slope and the intercept of the line in your free energy coordinates. The intercept is determined by the activation energy of the gating reaction, which is a function of temperature. The intercept can only be held as a constant when all mutations have the same activation energy. But if mutations change the activation energy, you have to deal with two variables in your regression analysis. A uerbach: If the constant was different with different mutations, they would show up off the line, and it would cause these points to look like outliers. The constant is the height of the transition state barrier when the equilibrium constant is 1, where the two lines cross. We have this height for a number of residues, but we haven't figured out what this information is providing. I think that in this context temperature experiments would be useful. Right now we are focusing on just trying to get more F values. ]ordan: But if you are trying to infer structure and you haven't completely deconvoluted what you are looking at, you are making assumptions. The whole linear free energy idea assumes that the free energy landscape moves in a very concerted fashion. It is important to see that this concerted motion is the same at two different temperatures. Roux: You could imagine that the free energy varies linearly but the entropy and enthalpy are non-linear. It is just that they conspire to give you linear free energy. Then you would have real problems. What really acts on the system is the free energy. A uerbach: It might reveal something interesting if the F values changed with different temperature. Jordan: I don't think it would address the issue of structure versus free energy, though. Koux: How much has been done on allosteric proteins where the structure is known? A uerbach: With haemoglobin and GroEL, single point mutations have been made and calculations of an average F for the whole protein have been done. This gives the position of the transition state and the reaction coordinate for the whole system. What hasn't been done extensively is probing of multiple substitutions at one position, and also trying to map this onto the structure. I think that this is the first time that has been done. This is partially

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because the ACh receptor is so long, which gives us a nice spatial extent to probe. It is also partially because single-channel kinetics makes it relatively easy to get the rate constants. Roux: It would be interesting to understand whether the linear free energy relationship is obeyed for many residues. However, the interpretation of why it is linear is interesting. Why would you get the different F values at different positions? It would be interesting to probe this in a system for which the structure is known. Auerbach: A linear approximation works when the perturbation is small. Roux: Even in the example you gave you can think of things that are hard to understand. Let's say you have a state that is like a cavity, and you have a residue there, and then you want to go to other states and other cavities. There is a tunnel, which you have to go through. Then when you get to the other side you are happy again. If you make something bigger, you can imagine that you would perturb the equilibrium in a completely different direction from the rate constant, because the transition state may have a completely different nature. Electrostatically, it is possible that if I have a positive charge, and I have to move from here to there, and I make a mutant that is like a negative change that would put a linear electric field, then you would see as I move in the direction I have a constant change of free energy. But for collisions it is not to easy to understand how you can get a linear free energy relationship. Auerbach: I think the multiple trajectory hypothesis is unlikely. It has been proposed that Fs are all zeros or ones, and the apparent F value reflects the probability of taking one pathway or another. The linearity of our Bronsted plot suggests that this is not true. However, there are more complicated possibilities. There may be a million trajectories that span the range of 0-1. Or there may be two trajectories between, say, 0.4 and 0.5, that give you something that looks fairly linear. Some trajectories indeed have fractional F values. What does a fractional F value mean? An example of multiple pathways occurs with mutations at the delta M2 10 prime position. Mutations that perturb the equilibrium constant to a small extent seem to follow a .F of zero. With bigger perturbations, that is, larger changes in the equilibrium constant, the slope changed. Such a non-linear Bronsted plot indicates that when there is a big enough perturbation to the system, the reaction mechanism changes. One possibility is that the protein can take alternate pathways. At low energies it takes one pathway, and with big perturbations it tends to take another. Alternatively, there may be a 'catalytic' effect with big perturbations. Roux: The chemistry of these amino acid substitutions is not that obvious. A. uerbach: We don't know what the detailed chemistry is; one of the beautiful aspects of this analysis is that we don't have to worry about the chemistry, just the free energy.

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Sansom: I was interested by your outlier at the 12' position. You said that this outlier indicates that you are getting extra catalysis. Awrbach: Yes. The proline substitution had an independent effect on the transition state. Sansom: And the transition state energy is lowered more than you'd expect? It is interesting that of the mutations you make, the only one that affects the backbone (which is also one that if you project it on a helical wheel is just above the 9') is the one that has additional catalytic effects. This would be directly consistent with a hinge or a swivel (Sansom & Weinstein 2000). Auerbach: We haven't looked in this much detail. I think this residue might be projecting towards the internal part of the protein and not the lumen. Sansom: 12' is one turn above 9', which is where the proposed hinge is. The proline at 12' would lead to a disruption of the 12'—9' hydrogen bond, so it will give a greater hinging ability. This correlates with your lower free energy for the transition state. This would correlate directly with a hinging model for the transition. Fu: You showed that most of the spots will be falling in a line, but you do see some exceptions. Because you only have a few spots to make a line, if you make enough observations will the spots off the line also make another parallel line, with the same F? A uerbach: Eiach point is four different patches. There is error in the rate constants in each patch, and some variation between patches. The points that are not exactly on the line may reflect noise, or some additional catalytic effect or anticatalytic effect on the transition state, or some other reaction mechanism. I think we would be hard pressed to fit those data by anything other than a straight line. Fu: My point here is that when you introduce a point mutation you could equally possibly affect F or the constant. We have data showing a set of mutants that produced two separate lines, but they are clearly parallel to each other. This observation suggested that our mutants do not affect F, but the constant. Roax: You seem to be referring to a specific example. Have you done F analysis, and if so, on what system? Fu: Yes, on the ACh receptor. Auerbach: Did you study receptors having multiple mutations? If so, you measured F on a background of another F. This is quite complicated. F on a background may or may not be the same as F with the wild-type. I haven't seen your data, so I'm not sure. But if you had two sets of lines that were clearly separated, with the same slopes, you might speculate that there are two possible intrinsic barriers, each with the same F. Sixma: It is not completely obvious what you would expect to see if there was an intermediate.

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Auerbach: We would expect to see an extra component in our closed interval duration histograms. The histogram I showed at the beginning was an exponential. An exponential on a log x axis looks a bit like a warped Gaussian. We would expect to see one warped Gaussian and another bump coming off the end, to indicate the presence of another short-lived state. This is not to say that there isn't something shorter than 10/^s or so. But this is the limit of our instruments, and down to this limit we see just one component. An intermediate would show up as a second bump in that histogram. Sixma: Is it possible that you will hit on other residues that would cause an intermediate, and you are not yet looking at the right regions? Auerbach: That is possible. We have been concentrating mostly in the M2 region. Sixma: You are now either in the ligand binding site or the actual gate region. A uerbacb: No, we have also measured residues in the M2/M3 loop and at the top of M4, above the gate. We have explored more than the M2 9-10' region. We haven't sampled the extracellular domain very much, because we didn't know where things were until very recently. I think that if it is possible that the transmission is through the whole protein. The protein's job is to transmit the signal between the binding site and the gate. We have to think of the protein as a dynamic object, and that this conformational wave is really a fundamental aspect of its function. It is not going to be a simple matter of only a few residues being involved. The transition state is probably distributed over a large spatial area. I should mention that the fact that we see so many fractional values indicates that it is not a single microstate transition that is rate limiting. If this were the case, everything before that transition should be a of 1 and everything after should be a O of zero. The fact that we see this nice distribution of Os suggests that the transition state is spread over a large number of residues. Corringer: What about the opening of non-liganded or monoliganded receptors: do you have any idea of the transition state structure of these? Auerbach: That is a good question. We actually have some data on that. The trajectory of unliganded receptors is different from that of liganded receptors. We have measured unliganded receptor gating, by changing the voltage. It is hard to measure the number of channels in the patch when you have unliganded receptors, so getting the rate constants is difficult. If you change the voltage this is another perturbation: we are perturbing the dipoles or the charges that sense the voltage, which are somewhere in the membrane. We find that for diliganded receptors they are closed-like at the transition state. We don't know where they are, but they are probably below the 9' and 10' residues. If we look at this in unliganded gating, we see the exact opposite behaviour. The value for those dipoles is 1 instead of 0. It is presumably the same dipoles and the same structure, because the equilibrium constant of gating is sensitive to voltage to the same

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extent. Presumably, the structures are the same, but their positions in the reaction coordinate are different. Unliganded gating may take a different trajectory than diliganded gating, even though the endpoints in the reaction are likely to be the same. It is quite hard to make a F map for perturbations of unliganded gating, because of the problem of not knowing the number of channels per patch. Reference Sansom MSP, Weinstein H 2000 Hinges, swivels & switches: the role of prolines in signalling via transmembrane a-helices. Trends Pharmacol Sci 21:445—451

Functional genomics of ionotropic acetylcholine receptors in Caenorhabditis elegans and Drosophila melanogaster David B. Sattelle, Emmanuel Culetto, Marta Grauso, Valerie Raymond, Christopher J. Franks and Paula Towers AiRC Functional Genetics Unit, Department of Human A natomy and Genetics, University of Oxford, South Parks Rd, Oxford OX1 3QX, UK

Abstract. Genetics, genomics and electrophysiology are transforming our understanding of the nicotinic acetylcholine receptors (nAChRs). Caenorhabditis elegans contains the largest known family of nAChR subunit genes (27 members), while Drosophila melanogaster contains an exclusively neuronal nAChR gene family (10 members). In C. elegans, several genetic screens have enabled the identification of nAChR subunits, along with novel proteins that act upstream and downstream of functional nAChRs. The C. elegans genome project has identified many new candidate nAChR subunits and the calculated electrostatic potential energy profiles for the M2 channel-lining regions predict considerable functional diversity. The respective roles of subunits are under investigation using forward and reverse genetics. Electrophysiological and reporter gene studies have demonstrated roles for particular subunits in levamisole-sensitive muscle nAChRs and a role for nAChRs in pharyngeal pumping. Recombinant homomeric and heteromeric C.. elegans nAChRs have been expressed in Xenopus laeris oocytes. In D. melanogaster, three new nAChR a subunits have been cloned, one of which shows multiple variant transcripts arising from alternative splicing and A-to-I pre-mRNA editing. Thus, studies on the genetic model organisms C. elegans and D. melanogaster have revealed different routes to generating molecular and functional diversity in the nAChR gene family and are providing new insights into the in rho functions of individual family members. 2002 Ion channels—from atomic resolution physiology to functional genomics. Wiley, Chichtster (Xovartis Foundation Symposium 245) p 240-260

Nicotinic acetylcholine receptors (nAChRs) mediate the fast actions of the neurotransmitter acetylcholine (ACh) at nerve-muscle junctions (Taylor et al 2000) and in the nervous system (Lindstrom 2000). Distinct native nAChR subtypes with different physiological and pharmacological properties have been

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FIG. 1. Schematic representations of homomeric and heteromeric nicotinic acetylcholine receptor (nAChR) molecules and the loops (A—F) of the extracellular, N-terminal region that contribute to the ACh binding site.

recognized, and are formed from different combinations of polypeptide subunits (vertebrate muscle: a l, b l, yl [or el], (51; vertebrate nervous system: a2-10, b 2-~4). The binding of ACh to an nAChR molecule results in the transient opening of the integral ion channel. Muscle and neuronal nAChRs of vertebrates are pentameric membrane molecules composed of a and non-a subunits (Fig. 1), the a subunits being defined by a pair of adjacent cysteines in the extracellular N-terminal domain that contribute to the ACh binding site (for review see Galzi & Changeux 1995). Mutations in nAChR subunits result in several kinds of congenital human myasthenias (Engel et al 1998) and in one type of epilepsy (Steinlein et al 1995). Also, there is growing interest in the development of subtype-selective drugs that act on nAChRs for use in the management of disorders such as Alzheimer's disease, Tourette's syndrome and schizophrenia (Lena & Changeux 1998). The recently sequenced genomes of Caenorhabditis elegans, the first metazoan to have its genome fully sequenced (The C. elegans Sequencing Consortium 1998), Drosophila melanogaster (Adams et al 2000) and human (Lander et al 2001) have revealed a rich diversity of nAChR subunits. Nevertheless, a major limit to our current knowledge of the functions of nAChRs is that for no single organism is the subunit composition of all nAChR subtypes known. The ready application of genetics to these nematode and insect model organisms (Baylis et al 1996) is

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likely to assist in determining the in vivo functions of the numerous nAChR isoforms. C. elegans: a genetic model for the study of receptor gene families The free-living nematode worm, C. elegans, is 1 mm long at the adult stage, has 302 neurons, and its complete cell lineage and nervous system synaptic connectivity have been described. A detailed genetic map is available, together with an overlapping physical map of cosmids and yeast artificial chromosomes (YACs). Its genome is 97 megabases encoding at least 19 100 proteins. The genome has revealed that genes encoding receptors important in synaptic functions, rather like the old king's sorrows in Shakespeare's Hamlet, Act 5 Sc. V, 'Come not single spies but in battalions'. For example, there are 90 genes encoding ionotropic receptor subunits, the gene 'superfamily' which includes the nAChR subunit genes (Fig. 2) and 1000 G protein-coupled receptors (Bargmann 1998). So a major challenge now is to make functional sense of the molecular diversity in complex receptor gene families (Sattelle 1998). C. elegans has an advanced genetic 'toolkit' to add to the wealth of new sequence data, which facilitates analysis, by forward and reverse genetics, of the functions of individual receptor gene family members. Genetic screens identify novel nicotinic receptor subunits that are drug targets In 1974, Sydney Brenner first isolated C. elegans mutants resistant to the cholinergic anthelmintic drug levamisole, which is used to control nematode parasites. Later, the genetics of levamisole resistance was characterized (Lewis et al 1980,1987) and it was shown that three of the genes associated with resistance (Fig. 3) lev- /, unc-29&nd unc-38 encode nAChR subunits: LEV-1 (a), UNC-29 (non-a) and UNC-38 (a) (Fleming et al 1993,1996,1997). In the same study it was shown that expression of these three subunits in Xenopus laevis oocytes results in low amplitude, inward currents in response to levamisole. The levamisole-induced currents were reduced in amplitude by nicotinic antagonists mecamylamine and neosurugatoxin (Fleming et al 1997). Although the pharmacology of this recombinant receptor was similar in some respects to that of native nematode muscle nAChRs (Fleming et al 1993,1997, Tornoe et al 1995, 1996) the very small amplitude currents pointed to a missing subunit. Two dominant mutations of lev-1 in the second transmembrane (M2) region resulted in particularly high resistance to levamisole (Fleming et al 1997). Thus, the identification of viable nAChR mutants in C. elegans permits manipulation of receptor expression, synaptic targeting in vivo, and may add to our understanding of drug resistance mechanisms.

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0.05,

FIG. 2. Phylogenetic relationships obtained with the neighbour-joining protein distance method of analysis on a ClustalX alignment of the nAChR subunit amino acid sequences of C. elegans. The signal peptide sequence and the hyper-variable TM3-TM4 intracellular loop have been removed. Bootstrap values are indicated as a percentage at each node. The product of the C. elegans unc-49A gene, a GABAA-type receptor subunit gene, was selected as the outgroup.

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A genetic approach that screens for reduced pharyngeal pumping (Raizen et al 1995) has identified other mutants resulting in the identification of two interacting loci of interest in the present context (eat~2and eat-18}. Electromyogram recordings are consistent with these genes encoding either nAChR subunits, or associated proteins, although further work is needed to resolve this. Another genetic screen for mutants with a neuronal degeneration phenotype has resulted in the isolation of a small number of genes, one of which, deg-3, encodes an nAChR a subunit (Treinin & Chalfie 1995). Moreover, a suppressor screen for the deg-3 dominant mutation has identified des-2 (=acr-4\ which encodes another a subunit closely related to deg-3 (Treinin et al 1998). These two functionally dependent a subunits are encoded in a single operon (Treinin et al 1998). Genomics reveals an extensive and diverse gene family of nicotinic receptor subunits — the largest currently known for any organism A cross-hybridization strategy, using either D. melanogaster nAChR cDNA, previously cloned C. elegans nAChRs, or vertebrate nAChR subunit cDNAs, has permitted the cloning of four subunit cDNAs: acr-2 (Squire et al 1995), acr-3 (Baylis et al 1997) (non-as), acr-16 (a) (Ballivet et al 1996) zndjtf-38 (a), which maps to a levamisole-resistant locus (E. Culetto, personal communication). However, the majority of novel putative nAChR subunits have been identified via The C. elegans Genome Consortium. Using the reverse transcription (RT)PCR, the transcription of 18 novel putative nAChR subunit genes has been demonstrated in C. elegans. When combined with the nine other a subunit genes described earlier by other workers (deg-3, acr-4 [ = des-2], Ce21 [ = acr-16], acr-6] and ourselves (unc-38, unc-29, lev-1, acr-2, acr-3}, this is now the largest known family (27 members to date) of nAChR subunit genes in a single species. Four major groups of mainly a subunits can be recognized, their names deriving from the first reported member (Mongan et al 1998): DEG-3-like; ACR-16 [Ce21]like; UNC-38-like and ACR-8-like (Fig. 2). The DEG-3-like group has no counterpart in vertebrates, and neither have the three members of the ACR-8-like group. A separate group containing only non-a subunits, the UNC-29-like group, has also been identified (Fig. 2). Five C. elegans nAChR a subunits contain a modification in loop C of the ACh binding site in which the normally conserved Tyr-x-Cys-Cys is replaced by a distinct motif (Tyr-x-x-Cys-Cys). The members of the C. elegans ACR-16-like group show homology to the vertebrate a 7 subunits. We have found two new non-a relatives of the ACR-16-like a subunits, ACR-9 and ACR-14. As is the case for the vertebratea 7 subunit, ACR-16 can form functional homomeric receptors when expressed in Xenopus/aevis oocytes (Ballivet et al 1996, Raymond et al 2000). The question of whether such subunits form homomeric nAChRs in vivo remains controversial but the discovery in

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TABLE 1 Expression in neurons and muscle cells of selected C. elegans nicotinic acetylcholine receptor (nAChR) subunit genes

nAChR. subunitgene deg-3, des-2 unc-29 acr-5 acr-2

Location of expression

PVC, PVD, FLP (touch neurons) Head neurons body wall muscle B-type motor neurons Motor neurons (multiple classes)

Reporter used in localisation studies

LacZ GFP GFP GFP

References

Treinin & Chalfie 1995, Treininetal 1998 Fleming et al 1997 Winnier et al 1999 Nurrishetal 1999

C. elegans oi ACR-16-like non-a subunits and DEG-3-like non-a subunits raises the possibility that some heteromeric receptors may exist. Most of the C. elegans nAChR subunits appear to be expressed at very low levels in vivo since only two, ACR-7 and UNC-29, have been isolated as expressed sequence tags (ESTs).

Subunit spatial expression patterns that cross tissue boundaries are common in C. elegans Detailed studies of vertebrate nAChRs have resulted in the concept of distinct families of receptor subunits expressed in muscle and in the nervous system, although the discovery that the vertebrate a9 subunit is expressed in epithelial cells as well as certain brain regions has raised the possibility that more complex patterns of expression might be found (Elgoyhen et al 1994). Also, the finding that the vertebrate 'neuronal' a7 subunit can be expressed in developing muscle, though only prior to synapse formation, suggests a possible role for this subunit in synaptogenesis (Fischer et al 1999). As shown in Table 1, several of the nAChR subunits studied to date in C. elegans are expressed in both neurons and body wall muscle cells.

Genetics identifies genes functionally linked to nAChRs The major levamisole resistance loci include genes whose functions remain to be resolved but which are not nAChR subunits. The unc-50 gene encodes a product that may be involved in assembly or transcriptional control of receptor subunits (Fig. 3). Although not studied extensively in C. elegans, a vertebrate orthologue of unc-50, UNCL, a candidate RNA binding protein, appears to be localized in the inner nuclear membrane and enhances the amplitude of currents recorded from

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recombinant vertebrate nAChRs when co-expressed in Xenopus oocytes with either a4/?2 or a3/?4 subunits (Fitzgerald et al 2000). The lev-11 and unc-22 genes (Fig. 3) are involved in muscle contraction, encoding respectively tropomyosin and twitchin. Identifying all the remaining unresolved loci will be of interest. A separate genetic screen has identified aldicarb-resistant mutants (Nguygen et al 1995). Aldicarb is an acetylcholinesterase (AChE) inhibitor. This screen resulted in isolation of 18 new loci (ric genes, resistant to inhibitors of cholinesterase) and identified components that were both pre- and postsynaptic. Of these, ric-3 appears to play a role in nAChR assembly in C. e/egans, as it also confers levamisole resistance. The ability to identify, in screens for resistance to cholinergic drugs and chemicals, novel candidate genes that act either upstream or downstream of nAChRs is of considerable interest (Sattelle & Culetto 1999, see Fig. 3). Mutant suppressor screens based on nAChR subunit mutants also offer the prospect of identifying genes functionally linked to particular nAChR subtypes (Treinin et al 1998). This approach involves screening for mutations, which reduce or suppress the particular nAChR mutant under investigation. This will be increasingly important in understanding how the diverse spatial and temporal patterns of nAChR subunit expression are controlled.

Molecular modelling predicts novel functions for subsets of nicotinic receptor subunits Restrained molecular dynamics simulations have been used to generate homopentameric M2 helix bundle models for C. elegans nAChR a subunits (Mongan et al 1998). The calculated electrostatic potential energy profiles are displayed as the potential energy profile of a single positive charge moved along the length of the pore (z) axis (Fig. 4). The profile calculated for the M2 region of ACR-8 differs strikingly from that of ACR-16, largely due to the presence of a histidine residue at the — 1' position in M2, predicting an apparent electrostatic barrier to cation permeation (Fig. 4). Thus, the presence of this histidine is expected to result in a major change in cation permeability of the channel, raising the possibility that it may represent an ACh-gated anion channel subunit. The difference between the electrostatic energy profiles of ACR-8 and ACR-16 resemble qualitatively the differences observed between the vertebrate glycine receptor a subunit and the a7 nAChR subunit (Sankararamakrishnan et al 1996), which are respectively able to form anionic and cationic channels when expressed in Xenopus oocytes. Examples of native ACh-gated anion channels have been reported in invertebrates. Two pharmacologically distinct ionotropic receptors mediate ACh-induced Cl~ currents in pleural ganglion neurons of the mollusc Aplysia call/arnica (Kehoe & Mclntosh 1998). One is a rapidly desensitizing response, the

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FIG. 4. (A) A 'ribbons' diagram of the C. elegans ACR-16 pentameric M2 helix bundle model with the channel axis horizontal. In (B) the strikingly different electrostatic potential energy profiles are shown for ACR-16 (favourable for cation permeation) and C. elegans ACR-8 (less favourable for cation permeation). The overall profile for ACR-16 resembles that of the y7, whereas that of ACR-8 is more reminiscent of that observed for the glycine receptor a subunit. Based on Mongan et al (1998).

other a more sustained response. The molecular entities responsible for these AChmediated anion channels are not known for any organism, so it will be of interest to pursue heterologous expression studies to see whether recombinant nAChRs containing ACR-8-like subunits (or other subunits) result in ACh-gated anion channels.

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(A) Voltage-clamp recording of ACh-induced currents from expressed vertebrate and C. elegans neuronal nicotinic receptors

(B) Voltage recording of C. elegans pharynx muscle membrane potential in response to ACh

FIG. 5. Responses to acetylcholine (ACh) of (A) recombinant homomeric C. elegans ACR-16 and vertebrate a7 nAChRs expressed in Xenopus laevis oocytes and (B) native C. elegans nAChRs of the pharynx.

Expressed recombinant nicotinic receptors are used to explore functional and pharmacological diversity in the nAChR gene family In 1997, Fleming et al showed that a recombinant nAChR formed when C. elegans nAChR subunits UNC-38, UNC-29 and LEV-1 are co-expressed in Xenopus laevis oocytes was activated by ACh and levamisole but blocked by mecamylamine. Of all the C. elegans nAChR subunits sequenced to date, the UNC-38-like and UNC-29like groups most closely resemble the vertebrate muscle subunits. The recent finding t\\&tjtf-38, a member of the UNC-38-like group, encodes a novel nAChR a subunit offers the prospect of further recombinant expression studies that will help clarify the subunit composition of the levamisole-sensitive body wall muscle nAChRs. Ballivet and colleagues (1996) cloned the ACR-16 subunit. Its primary structure is 47% identical to the chicken neuronal a7 subunit and it forms functional homomeric nAChRs in heterologous expression studies. The recombinant nAChR generated by the ACR-16 homomer-forming a subunit (Fig. 5) differs in

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aspects of its pharmacology from UNC-38-containing nAChRs. Oocytes expressing either ACR-16 or a.7 nAChRs generate currents in response to ACh or nicotine. In both cases nicotine is more potent than ACh. Nicotine is a full agonist on a.7 (Couturier et al 1990), but only a partial agonist on ACR-16 (Ballivet et al 1996). Both receptors desensitize rapidly and are blocked by d-tubocurarine and dihydro-/J-erythroidine (Ballivet et al 1996). However, ACR-16 is relatively insensitive to N-methyllycaconitine (MLA) and the snake toxin a-bungarotoxin, both potent antagonists of a.7 nAChRs (Ballivet et al 1996). Of particular interest was the finding that levamisole, an agonist of the UNC-38, UNC-29, LEV-1 receptor, showed no agonist action on ACR-16 but instead was an antagonist (Ballivet et all 996). Recently, we compared the actions of several anthelmintics on ACR-16 and chicken a.7. Morantel, pyrantel and butamisole were antagonists of both nAChRs. Oxantel, a partial agonist of a.7, showed no agonist action on ACR-16; it was, however, an antagonist (Raymond et al 2000). The endectocide ivermectin, an agonist at L-glutamate-gated Cl ~ channels, potentiates the actions of ACh on chicken a.7 (Krause et al 1998, Raymond et al 2000) but has no such effect on ACR16 (Raymond et al 2000). Thus, studies on oxantel and ivermectin have identified new pharmacological differences between the chicken a.7 subunit and its C. e/egans homologue ACR-16. Since the discovery of the DEG-3 subunit (Treinin & Chalfie 1995), there has been interest in the prospect of studying functional expression of members of this class of nAChR subunits. Recently, Treinin and colleagues have shown that, although not able to function as homomers, a functional recombinant heteromeric receptor composed of two types of a subunit (DEG-3, DES-2 [ = ACR-4]) could be generated (Yassin et al 2001). Of particular note was the finding that such receptors showed a higher affinity for choline than for ACh. There have been suggestions that choline can function as a neurotransmitter at certain stages of neuronal maturation (Albuquerque et al 2000) and it will be of interest to address this further in C. e/egans, where there is a large family of DEG3 like receptors. In vertebrates, a subunit heteromers of a9 and alO also yield functional receptors. These a9alO receptors more closely resemble the native nAChR in the cochlear epithelium than the a9 homomeric receptor (Elgoyhen et all 994, 2001). Studies on native nicotinic receptors in C. elegans The successful application of electrophysiology to the study of (a) synaptic transmission at nerve-muscle junctions; (b) pharynx muscle physiology; and (c) neuronal cell bodies in C. elegans has enabled, for the first time, the study of native C. elegans nAChRs. The work of Richmond & Jorgensen (1999), using

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patch-clamp recordings obtained from body wall muscles, has shown that two ACh receptors are present, one of which is levamisole sensitive. A role for unc-38 and unc-29 in the levamisole-sensitive muscle receptor has been demonstrated, a finding consistent with results of genetic experiments. Data from the Avery lab (Raizen et al 1995) and our own recent studies (Fig. 5) point to a role for nicotinic receptors in regulating pharyngeal muscle activity.

Role of nicotinic receptor genes in egg-laying behaviour The Schafer laboratory has deployed genetics to analyse the genes responsible for the egg-laying behaviour of the nematode and the effects of various drugs on this behaviour. The egg-laying behaviour is biphasic and the interval between successive layings is under the control of the neurotransmitter ACh, whereas the delay between bursts of laying is under the control of 5-hydroxytryptamine (5-HT). The rinding that the ACh-controlled phase is also sensitive to nicotine stimulation provided these workers with a genetic model with which to investigate the molecular events associated with long-term exposure to nicotine (Waggoner et al 1998, 2000), events which in human and other vertebrates can lead to addiction. Recently, Kim et al (2001) found that seven levamisole-resistance genes, including the nAChR subunit genes unc-38, unc-29 and lev-1', are essential for stimulation of egg laying by levamisole, though in the absence of the drug their effects on egg laying are less pronounced. As egg laying mutants also responded to other nAChR ligands, it seems likely that other nAChRs are involved in the egglaying neural circuitry. It is of interest that mutations in unc-38 also resulted in altered sensitivity of egg laying to the neurotransmitter 5-HT. Thus, nAChRs and 5-HT receptors feature in neural pathways regulating egg-laying. These studies show that a combination of genetic and behavioural studies can add to the spectrum of physiological roles of nAChR subunits.

The role of nAChRs in nervous system development Vertebrate muscle and neuronal nAChR subunits are, to varying degrees, developmentally regulated and there is growing evidence that nAChRs play a role in maturation of neurons and neuronal circuits (Zoli 2000). Also, a marked decrease in nAChRs is characteristic of ageing pathologies such as dementia (Zoli 2000). However, little is known of the regulatory mechanisms involved. Studies on C. elegans have shown that in the unc-29(e1072) mutant, which lacks this non-a subunit, there is reduced neuronal sprouting and this is a postsynaptic effect, as it is seen when the expression of the subunit is directed only to muscle cells (Zhao & Nonet 2000). The finding that disruption of nAChRs generates sprouting of the

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neuron indicates that the decision to sprout involves communication between nerve and muscle and additional experiments support the view that muscle inactivity is sensed by the motor axon resulting in modulation of synaptic contacts (Zhao & Nonet 2000).

Drosophila melanogaster: a genetic model for the study of receptor gene families The fruit fly, D. melanogaster, is well suited for studies of neurotransmitter receptor gene families. It has a larger nervous system (~250 000 neurons) and more complex behaviours than C. elegans. Its complete genome is ~120 megabases, encoding at least 13 600 genes. There are about 50 ligand-gated ion channels and 160 G proteincoupled receptors (Littleton & Ganetzky 2000).

The nAChR gene family of D. melanogaster— new insights from genomics In the recently completed genome of D. melanogaster four new nAChR subunits have been identified, resulting in a gene family of 10 members to date (Fig. 6). There are seven a subunits and three non-a subunits. The physiology and pharmacology of nAChRs containing SAD, ALS and/or Da3 (a subunits) have been addressed in co-expression studies with the vertebrate j?2 subunit in Xenopus laevis oocytes (Bertrand et al 1994, Schulz et al 2000). However, to date there has been no successful functional expression using only D. melanogaster nAChR subunits. Three newly identified a subunits (Da5, Da6 and Da7) most closely resemble the mammalian a? subunits and the C. elegans ACR-16-like subunits. The remaining fly subunits are more closely related to one another than to known mammalian a or /? subunits. The D. melanogaster nAChR subunits are likely to be purely neuronal (see Breer & Sattelle 1987, Gundelfinger & Shulz 2000) as L-glutamate is the classical excitatory neurotransmitter at insect nerve-muscle junctions. Alternative splicing adds to the diversity of the fly nAChR gene family in the case of Da4 (Landsell & Millar 2000), Da5 and Da6 (M. Grauso, personal communication). In the case of Da6 pre-mRNA A-I editing has been detected (M. Grauso, personal communication).

Conclusions The largest and most diverse nAChR gene family known is that of C. elegans with 27 subunits and there are a wealth of mutants available for study- Using this model organism, we have observed nAChR subunits that are neuron-specific, musclespecific and others found in both neurons and muscle cells. Functional roles for unc-38, unc-29 and lev-1 have been demonstrated at motorneuron-body wall muscle synapses. In D. melanogaster a smaller, and apparently exclusively neuronal

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FIG. 6. A phylogenetic tree for the Drosophila melanogaster nAChR subunits has been constructed using the method described to generate the C. elegans nAChR tree (Fig. 2). The product of the D. melanogaster rdl gene, a GABAA type receptor subunit, was used as the outgroup. Accession numbers of published subunits are: ALS, CAA30172; SAD, CAA37652; Da3, Caa75688; D<x4, CAB77445; ARD, CAA27641; SBD, CAA39211; RDL, Aaa28556; Da5, D«6 and Da7 have been cloned in our laboratory and their Genbank submission is in progress. We recently sequenced D/?3, the new putative nAChR ft subunit cDNA.

nAChR gene family has been identified. Functional heterologous expression studies on SAD, ALS and Da3 subunits have been carried out. Considerable additions to nAChR molecular diversity in D. melanogaster result from alternative splicing and RNA editing. Thus, different molecular strategies for generating functional diversity among nicotinic acetylcholine receptors have been identified using these invertebrate model organisms. Future challenges The application of electrophysiology (Richmond & Jorgensen 1999) and Ca2+ imaging (Kerr et al 2000) to native excitable cells of C. elegans has been a major

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step forward and the increasing use of primary cultures will provide further impetus to receptor/channel studies in both C. elegans and D. melanogaster. Detailed study of the temporal and spatial patterns of expression for all members of a particular nicotinic receptor gene family and the identification of their functional roles are a realistic prospect. Recently a range of antisense approaches have been applied to neural signalling in C. elegans (im review see Culetto & Sattelle 1999). Comprehensive gene silencing experiments, using double-stranded RNA interference (dsRNAi), have been used to investigate the entire complement of genes on chromosomes I (Fraser et al 2000) and III (Gonczy et al 2000) of C. elegans. This approach has proved extremely useful and is being developed into large-scale, high-throughput procedures (Maeda et al 2001) but some neuronal genes are found to be somewhat resistant to silencing by this method. Nevertheless, with improvements in dsRNAi technology, it is likely to prove extremely effective when used in physiological studies on cultured, excitable cells. The use of mutant suppressor screens and microarrays is already beginning to yield useful data and will be a rich source of new information on genes interacting with nAChRs as well as the effects on subunit gene expression of long-term exposure to nicotinic receptor ligands. Both genetic model organisms allow complex behaviours to be analysed (Davis 1996, Rankin 2000) using the tools of molecular and cell biology, genetics, genomics, and physiology to which are now being added the powerful new post-genome technologies, such as DNA microarrays and proteomics.

References Adams MD, Celniker SE, Holt RA et al 2000 The genome sequence of Drosopbila melanogaster. Science 287:2185-2195 Albuquerque EX, Pereira EFR, Alkondon M, Eisenberg HM, Maelicke A 2000 Neuronal nicotinic receptors and synaptic transmission in the mammalian central nervous system. In: Clementi F, Fornarsi D, Gotti C (eds) Neuronal nicotinic receptors. Springer-Verlag, Berlin, p337-358 Ballivet M, Allied C, Bertrand S, Bertrand D 1996 Nicotinic acetylcholine receptors in the nematode Caenorhabditis elegans. } Mol Biol 258:261-269 Bargmann CI 1998 Neurobiology of the Caenorhabditis elegans genome. Science 282:2028-2033 Baylis HA, Matsuda K, Squire M et al 1997 ACR-3, a Caenorhabditis elegans nicotinic acetylcholine receptor subunit. Molecular cloning and functional expression. Receptors Channels 5:149-158 Baylis HA, Sattelle DB, Lane NJ 1996 Genetic analysis of cholinergic nerve terminal function in invertebrates. J Neurocytol 25:747-762 Bertrand D, Ballivet M, Gomez M, Bertrand S, Phannavong B, Gundelfinger ED 1994 Physiological properties of neuronal nicotinic recepors reconstituted from the vertebrate /?2 subunit and Drosophila a subunits. Eur J Neurosci 6:869-875 Breer H, Sattelle DB 1987 Molecular properties and functions of insect acetylcholine receptors. 1 Insect Physiol 33:771-790 Brenner S 1974 The Genetics of Caenorhabditis elegans. Genetics 77:71~94

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Couturier S, Bertrand D, Matter JM et al 1990 A neuronal nicotinic acetylcholine receptor subunit (alpha 7) is developmentally regulated and forms a homo-oligomeric channel blocked by a-BTX. Neuron 5:847-856 Culetto E, Sattelle DB 1999 Antisense approaches to neural signalling in C. elegans. In: Leslie RA, Hunter A J, Robertson HA (eds) Antisense technology in the central nervous system. Oxford University Press, Oxford, p 23O-243 Davis R 1996 Physiology and biochemistry of Drosophila learning mutants. Physiol Rev 76:299-317 Elgoyhen AB, Johnson DS, Boulter J, Vetter DE, Heinemann S 1994 a9: acetylcholine receptor with novel pharmacological properties expressed in rat cochlear hair cells. Cell 79:705—715 Elgoyhen AB, Vetter DE, Katz E, Rothlin CV, Heinemann SF, Boulter J 2001 alO: a determinant of nicotinic cholinergic receptor function in mammalian vestibular and cochlear mechanosensory hair cells. Proc Natl Acad Sci USA 98:3501-3506 Engel AG, Ohno K, Sine SM 1998 Congenital myasthenic syndromes: experiments of nature. ] Physiol Paris 92:113-117 Fischer U, Reinhardt S, Albuquerque EX, Maelicke A 1999 Expression of functional x7 nicotinic acetylcholine receptor during mammalian muscle development and denervation. Eur J Neurosci 11:2856-2864 Fitzgerald J, Kennedy D, Viseshakul N et al 2000 UNCL, the mammalian homologue of UNC50, is an inner nuclear membrane RNA-binding protein. Brain Res 877:110-123 Fleming JT, Tornoe C, Riina HA, Coadwell J, Lewis JA, Sattelle DB 1993 Acetylcholine receptor molecules of the nematode Caenorhabditis elegans. In: Pichon Y (ed) Comparative molecular neurobiology. Springer-Verlag, Berlin, p 65-80 Fleming JT, Baylis HA, Sattelle DB, Lewis J A 1996 Molecular cloning and in vitro expression of C. elegans and parasitic nematode ionotropic receptors. Parasitology 113:S175-S190 Fleming JT, Squire MD, Barnes TM et al 1997 Caenorhabditis elegans levamisole resistance genes lev-1, unc-29 and unc-38 encode functional nicotinic acetylcholine receptor subunits. ] Neurosci 17:5843-5857 Fraser A, Kamath RS, Zipperlen P, Martinez-Campos M, Sohrmann M, Ahringer J 2000 Functional genomic analysis of C. elegans chromosome I by systematic RNA interference. Nature 408:325-330 Galzi JL, Changeux JP 1995 Neuronal nicotinic receptors: molecular organization and regulations. Neuropharmacology 34:563-582 Gonczy P, Echeverri C, Oegema K et al 2000 Functional genomic analysis of cell division in C. elegans using RNAi of genes on chromosome III. Nature 408:331—336 Gundelfinger ED, Schulz R 2000 Insect nicotinic acetylcholine receptors: genes, structure, physiological and pharmacological properties. In: Clementi F, Fornasari D, Gotti C (eds) Neuronal nicotinic receptors. Springer-Verlag, Berlin, p 497-521 Kehoe JS, Mclntosh JM 1998 Two distinct nicotinic receptors, one pharmacologically similar to the vertebrate a7-containing receptor, mediate Cl currents in Aplysia neurons. J Neurosci 18:8198-8213 Kerr R, Lev-Ram V, Baird G, Vincent P, Tsien RY, Schafer WR 2000 Optical imaging of calcium transients in neurons and pharyngeal muscle of C. elegans. Neuron 26:583-594 Kim J, Poole DS, Waggoner LE et al 2001 Genes affecting the activity of nicotinic receptors involved in Caenorhabditis elegans egg-laying behavior. Genetics 157:1599-1610 Krause RM, Buisson B, Bertrand S et al 1998 Ivermectin: a positive allosteric effector of the a 7 neuronal nicotinic acetylcholine receptor. Mol Pharmacol 53:283—294 Lander ES, Linton LM, Birren B et al 2001 Initial sequencing and analysis of the human genome. Nature 409:860 -921

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Lansdell SJ, Millar NS 2000 Cloning and heterologous expression of Da4, a Drosophila neuronal nicotinic acetylcholine receptor subunit: identification of an alternative exon influencing the efficiency of subunit assembly. Neuropharmacology 39:2604-2614 Lena C, Changeux JP 1998 Allosteric nicotinic receptors, human pathologies. J Physiol Paris 92:63-74 Lewis JA, Wu CH, Berg H, Levine JH 1980 The genetics of levamisole resistance in the nematode Caenorhabditiselegans. Genetics 95:905-928 Lewis JA, Elmer JS, Skimming J, McLafferty S, Fleming J, McGee T 1987 Cholinergic receptor mutants of the nematode Caenorhabditiselegans. } Neurosci 7:3059-3071 Lindstrom J 2000 The structure of neuronal nicotinic receptors. In: Clementi F, Fornasari D, Gotti C (eds) Neuronal nicotinic receptors. Springer-Verlag, Berlin, p 101-162 Littleton JT, Ganetzky B 2000 Ion channels and synaptic organization: analysis of the Drosophila genome. Neuron 26:35-43 Maeda I, Kohara Y, Yamamoto M, Sugimoto A 2001 Large-scale analysis of gene function in Caenorhabditis elegans\>v high throughout RNAi. Curr Biol 11:171-176 Mongan NP, Baylis HA, Adock C, Smith GR, Sansom MSP, Sattelle DB 1998 An extensive and diverse gene family of nicotinic acetylcholine receptor alpha subunits in Caenorhabditiselegans. Receptors Channels 6:213-223 Nguyen M, Alfonso A, Johnson CD, Rand JB 1995 Caenorhabditis elegans mutants resistant to inhibitors of acetylcholinesterase. Genetics 140:527-535 Nurrish S, Segalat L, Kaplan JM 1999 Serotonin inhibition of synaptic transmission: Gzo decreases the abundance of UNC-13 at release sites. Neuron 24:231-242 Raizen DM, Lee RY, Avery L 1995 Interacting genes required for pharyngeal excitation by motor neuron MC in Caenorhabditiselegans. Genetics 141:1365-1382 Rankin CH 2000 Context conditioning in habituation in the nematode Caenorhabditis elegans. Behav Neurosci 114:496-505 Raymond V, Mongan NP, Sattelle DB 2000 Anthelmintic actions on homomer-forming nicotinic acetylcholine receptor subunits: chicken a"7 and ACR-16 from the nematode Caenorhabditiselegans. Neuroscience 101:785-791 Richmond J, Jorgensen EM 1999 One GABA and two acetylcholine receptors function at the C. elegans neuromuscular junction. Nat Neurosci 2:791-797 Sankararamakrishnan R, Adcock C, Sansom MS 1996 The pore domain of the nicotinic acetylcholine receptor: molecular modelling, pore dimensions, and electrostatics. Biophys J 71:1659-1671 Sattelle DB 1998 Genetic, genomic and functional studies on the acetylcholine receptor gene family of Caenorhabditiselegans. ] Physiol 513:18S Sattelle DB, Culetto E 1999 Functional genetics of cholinergic synaptic transmission in Caenorhabditis elegans. In: Beadle DJ (ed) Progress in neuropharmacology and neurotoxicology of pesticides and drugs. Royal Society of Chemistry, Cambridge, p 146- 160 Schulz R, Bertrand S, Chamaon K, Smalla KH, Gundelfinger ED, Bertrand D 2000 Neuronal nicotinic acetylcholine receptors from Drosophila: t\vo different types of alpha subunits coassemble within the same receptor complex. J Neurochem "74:253~7 2546 Squire MD, Tornoe C, Baylis HA, Fleming JT, Barnard EA, Sattelle DB 1995 Molecular cloning and functional co-expression of a Caenorhabditis elegans nicotinic acetylcholine receptor subunit (acr-2). Receptors Channels 3:107- 115 Steinlein OK, Mully JC, Propping P et al 1995 A missense mutation in the neuronal nicotinic acetylcholine receptor oc4 subunit is associated with autosomal dominant nocturnal frontal lobe epilepsy. Nat Genet 11:201-203 Taylor P, Osaka H, Molles B, Keller SH, Malany S 2000 Contributions of studies of the nicotinic receptor from muscle to denning structural and functional properties of ligand-gated ion

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channels. In: dementi F, Fornasari D, Gotti C (eds) Neuronal nicotinic receptors. SpringerVerlag, Berlin, p 79-100 The C. ekgans Sequencing Consortium 1998 Genome sequence of the nematode C. elegans: a platform for investigating biology. Science 282: 2012-2018 Tornoe C, Bai D, Holden-Dye L, Abramson SN, Sattelle DB 1995 Actions of neurotoxins (bungarotoxins, neosurugatoxin and lophotoxins) on insect and nematode nicotinic acetylcholine receptors. Toxicon 33:411-424 Tornoe C, Holden-Dye L, Garland C, Abramson SN, Fleming JT, Sattelle DB 1996 Lophotoxin-insensitive nematode nicotinic acetylcholine receptors. J Exp Biol 199:2161-2168 Treinin T, Chalfie M 1995 A mutated acetylcholine receptor subunit causes neuronal degeneration in C. elegans. Neuron 14:871—877 Treinin M, Gillo B, Liebman L, Chalfie M 1998 Two functionally dependent acetylcholine subunits are encoded in a single Caenorhabditis elegans operon. Proc Natl Acad Sci USA 95:15492-15495 Waggoner LE, Zhou GT, Schafer RW, Schafer WR 1998 Control of alternative behavioral states by serotonin in Caenorhabditis elegans. Neuron 21:203-214 Waggoner LE, Dickinson KA, Poole DS, Tabuse Y, Miwa J, Schafter WR 2000 Long-term nicotine adaptation in Caenorhabditis elegans involves PKC-dependent changes in nicotinic receptor abundance. J Neurosci 20:8802-8811 Winnier AR, Meir JY, Ross JM et al 1999 UNC-4/UNC-37-dependent repression of motor neuron-specific genes controls synaptic choice in Caenorhabditis elegans. Genes Dev 13:2774-2786 Yassin L, Gillo B, Kahan T, Halevi S, Eshel M, Treinin M 2001 Characterization of the deg-3/ des-2 receptor: a nicotinic acetylcholine receptor that mutates to cause neuronal degeneration. Mol Cell Neurosci 17:589-599 Zhao H, Nonet ML 2000 A retrograde signal is involved in activity-dependent remodelling at a C. elegans neuromuscular junction. Development 127:1253-1266 Zoli M 2000 Neuronal nicotinic acetylcholine receptors in development and aging. In: Clementi F, Fornasari D, Gotti C (eds) Handbook of experimental pharmacology, vol 144: neuronal nicotinic receptors. Springer-Verlag, Berlin, p 213-246

DISCUSSION Sansom: Why does C. elegans have so many different channels? Although it has only 5000 synapses, it has 27 nicotinic receptors and 40-odd TWIK-like K + channels. Also, there are two subgroups of nicotinic receptors in C. elegans with no known counterparts in vertebrates. Sattelle: It is one route to functional diversity; in the nicotinic receptor gene families of other organisms different strategies are evident such as alternative splicing and RNA editing. Further examples of the considerable diversity in invertebrate ligand-gated channels are to be found among ligand-gated anion channels. For example, we now have a 5-HT-gated Cl~ channel and a histaminegated Cl~ channel, as well as ACh-gated, glutamate-gated and GABA-gated Cl~ channels. Fu: In the pattern of expression of the different isoforms, is there an overlap between them?

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Sattelle: The subunits UNC-29, LEV-1 and UNC-63 all overlap to a degree in that they are expressed in the muscles of the body wall, but they appear to have nonoverlapping expression patterns in terms of some of the neurons in which they are expressed. That is why we deployed these three subunits for coexpression, in the hope of mimicking, at least in part, the muscle receptor. Miller: Returning to Mark Sansom's question of why C. elegans has so many channels and so few synapses, one way to modulate neuronal behaviour in a nervous system with so few synaptic connections per cell would be to switch receptor (or subunit) subtypes to change the properties of each synapse appropriately. Sattelle: Studies using green fluorescent protein (GFP) show that some subunits are expressed in very few neurons. I think many of the C. elegans nicotinic receptor described subunits are expressed in neurons and probably all the D. melanogaster subunits are components of neuronal nAChRs. Many of those C. elegans subunits that are expressed in muscle are also expressed in different subsets of neurons. It will be interesting to explore further receptor expression patterns at particular synapses throughout development. Sillier: Do you think a single synapse can express different receptors, depending on what is happening in its environment? Sattelle: It appears that there are two distinct nicotinic receptors expressed at the nerve-muscle junction, where knocking out a single subunit (UNC-63) completely abolishes the levamisole response but not the ACh response. We know that there is differential sensitivity to (and actions of) levamisole when recombinant muscle and neuronal receptors are compared. As to how environmental factors might impact, this remains to be investigated. Schrempf: Has anyone attempted to use one of the genes as a way of studying in detail splicing and editing, and how this is regulated? Sattelle: Not yet. The splicing and editing reported to date is all in D. melanogaster\ and we haven't yet reconstituted a fully functional D. melanogaster nicotinic receptor from only Drosophila subunits. To date there are no nicotinic receptor mutants in D. melanogaster. The diversity seems to be particularly high in the embryo. Corringer: You identified one a7-like subunit in C. elegans (ACR-16); are there other such subunits? Sattelle: The largest group of all in C. elegans is the a7-like group, with at least 9 members. We haven't yet looked at the other a7-like subunits, but plan to do this in the future. Corringer: Just from sequence alignment, are there similarities between the mammalian subunits and the C. elegans subunits? Sattelle: The ACR-16-like subunits resemble a? subunits of vertebrates. Interestingly, there are two ACR-16-like subunits that demand to be clustered

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in that gfoup which are in fact non as. This is a novel finding. It will be interesting to see whether there are non-oc partners of the a7-like subunits, at least in C. elegans. Corringer: One has a histidine in the — 1' position. Would you expect this to be an anionic receptor? Sattelk: Perhaps — there are two prolines close to the histidine residue (PPxxH) in ACR-8. We must try the expression studies, because if it is an ACh-gated anion channel, that would be of considerable interest. There are three changes in and around M2 of c<7 that are required to convert a cation channel into an anion channel. At this stage we cannot rule out that it is an unusual subunit that forms part of a cation channel. Miller: How common are anion channels in ACh receptors? There's one in snail that I know of. Sattelk: There are two that Jacsue Kehoe has identified on one cell in Aplysia; two ACh-gated Cl~ channels with different time-courses of response to ACh and each with a distinct pharmacology. We have also found an insect neuron where two glutamate-gated Cl~~ channels co-exist on the same cell. Ashcroft: You said that when you express the Drosophila ACh channels in a heterologous system, you are unable to see any current. What happens in the fly itself? Do they actually have ACh-gated channels? Sattelk: Yes. The insect nervous system is one of the richest nervous tissues known with respect to nAChR density. If you monitor radiolabelled a bungaratoxin binding, receptor concentration (mg"1 membrane protein) compares well with those of the electric organs of certain electric fish. Values comparable with those found in the electric organ of Electrophorus have been reported in insect nervous tissue. Receptor densitites in insect nervous system are not as high as those reported from the electric organs of Torpedo species. Patch-clamp studies on insect neurons have shown at least three different nicotinic receptor subtypes. A shcroft: From looking at the whole Drosophila genome, do you think you have all of them? Sattelk: You can never be quite sure about the non-as, but we are reasonably sure we have most of the as. Lambert: Have you tagged the non-functional ones? Do they make it to the cell surface? Sattelk: We have not done that, but Neil Miller at UCL has done some nice work with epitope-tagged subunits in various combinations. He has expressed these in S2 cell lines, and some do appear to reach the surface. Binding of nicotinic receptor ligands can sometimes be detected, but to date we are unable to observe functional receptors by electrophysiology. There may be some assembly problems and we cannot rule out that some subunits may have quite novel roles. The cell line

260

DISCUSSION

could perhaps present problems, but we have expressed other ionotropic and metabotropic receptors in these D. melanogaster S2 cells without difficulty. Sansom: Going back to the diversity question, if you look at non-channel receptors or non-receptor proteins in C. elegans, have people seen a similar large diversity there? Sattelle: Overall, there are estimated to be 19 100 protein-encoding genes, larger than the estimated 13 600 protein-encoding genes of D. melanogaster. In C. elegans there are ~1000 G protein-coupled receptors, ~80K + channels and ~90 neurotransmitter-gated ion channels, but apparently no Na + channels. So care is needed in making generalizations. However, multiple isoforms are often seen in C. elegans receptor gene families. Alternative splicing and RNA editing may result in comparable (possibly greater) diversity in the sometimes smaller gene families (e.g. nAChRs) of D. melanogaster.

Final general discussion A shcroft: Because we are never going to get the 3D structure of all the ion channels we want to know about in the foreseeable future, we will have to do quite a bit of modelling and inference from other channels. Knowing the 3D structures of just one or two channels, how do we go from there to get the others? What do the modellers need the electrophysiologists to come up with in order that we can put together a huge amount of data obtained from mutagenesis studies to produce models of the ones we haven't got structures of? R.OUX: As a modeller, one thing that is important to me would be to agree on the significance of the information that is extracted from experiments. For example, in the case of Shaker, experimental observations are translated into some constraint or inference about the structure. Then, important questions are: what is the uncertainty about such a constraint? What are the upper and lower bounds? Is there a consensus about the significance of this constraint? Does the scientific community agree? Is there an alternative interpretation of the experimental observation? It would be highly desirable to clear the air about this sort of issue. Mindell: The problem is that not everyone agrees on what the errors are in the measurements. When you are looking at your own data and it has taken two years to get them, you will see them in the rosiest light. This issue is frequently raised in discussions. I don't think it will be possible to give you a golden number. Papa^ian: There is some work from Don Engleman in which he combined mutagenesis of glycophorin with modelling and structural analysis (MacKenzie et al 1997). I think this is very informative. He had a lot of mutagenesis data, and the modelling did not converge to one lowest-energy model, but predicted several possibilities with similar energetics. But only one of these models was compatible with the body of experimental data. When the structure was eventually solved by NMR, this model turned out to be accurate. Modelling has to be a reiterative process of taking available data, making a preliminary model, and then testing the model by experimentation. Pero^o: We have to get into the perspective that a model can always be improved. There are always new experimental data that either agree or disagree with the model. A shcroft: This needs a lot more close interaction between the modellers and the experimentalists. Roux: The work on glycophorins is a special case. The experiments and the modelling were done by the same group (Lemmon & Engelman 1994). 261

262

FINAL GENERAL DISCUSSION

Sansom: One of the cases where we might be able to do it is for rhodopsin, now that we have the X-ray structure (Palczewski et al 2000). Before this there was a large body of mutational data on G protein-coupled receptors (GPCRs), which in many ways was analogous to what we have done for ion channels. Perhaps there is a case for someone to go back and analyse these data retrospectively to try to rank different types of observation in terms of their fit to the known structures. We might get the dull answer that they are all about the same, but we might be able to highlight the methods that pin things down more precisely. Mindell: We have actually been doing this in a small way. In Chris Miller's lab we have been doing a systematic review of the site-directed mutagenesis on the membrane protein structures that have been solved, looking at how good they were in retrospect. In general, they have been surprisingly good. The constraints that come out of the mutagenic studies when mapped onto the structure do pretty well. What is turning out to be the biggest variable is not the method, but how carefully done the experiments are. We all know that you can do cysteine mutagenesis carefully and you can do it poorly. The people who do it carefully get more reliable results. I don't think it will be possible to tie a number to a method. Mitra: I would like to make a comment here. It is not uncommon that papers published from reputable labs with regard to the effects of a given mutation differ from the results published earlier. It was not that the experiments were not done carefully; it was more due to parameters and the system being optimized better. It is difficult to rank the experiments. Sansom: In terms of formal restraints in the modelling procedure, there are numerical analysis ways of doing robust refinements. After one passes through the data you reduce the weights on the outliers, and then iterate until you no longer find any outliers. In this respect you may be able to filter out the occasional rogue measurements in an objective fashion, if you have sufficient data to determine the solution. Miller: I don't think it is only a matter of assessing rogue measurements or doing statistical filtering to remove outliers. Many of the states examined are transient. They exist only briefly on the time scale of the cysteine accessibility experiment, so that more than one state is necessarily sampled in the result. To get cleaner data uncontaminated by a second conformation requires conducting the cysteine accessibility experiment on a faster time scale. Unfortunately, this is technically very demanding and would limit the number of mutants that could be studied in detail. Perhaps the best strategy in practice is to base the initial modelling on a survey of many mutants at low time resolution (including contamination from other conformations). Then, this model can be refined by making specific critical predictions which can be experimentally tested at the higher resolution offered by time-resolved accessibility measurements.

FINAL GENERAL DISCUSSION

263

Similar considerations apply to studies of ligand-gated channels where the time of exposure to agonist is critical. Mindell: Benoit Roux's example of the AgTx2 was very enlightening. For the one mutant that didn't work from the modelling, he had a different explanation for a long-distance effect. We have no way experimentally of picking up those effects. As careful as you can be, there will still be outliers. You can't predict which measurements have higher errors than others in this way. Koux: The AgTx2 was fairly easy, because it matches the outer surface of Shaker so well. You could almost find that Serl 1 cannot be close to the Asp431 at the same time as Arg24 by simple examination. Miller: So are you saying it is a two-way street? You can tell him that he should mutate something that is not intuitively obvious as he looks at the primary structure. Koux: So far it has not been. The communication between the modellers and experimentalists is not easy, probably because what is 'easy' for one group is difficult for the other and vice versa. Papa^ian: One problem with a lot of the methods used experimentally is that if you get a negative result you can conclude nothing. Sansom: In terms of improving communication, at the moment we have a structure and highlight residues where we think there are important contacts, but in building models we are taking a statistical average of all these data to drive a particular conformation. In the display and analysis of this we throw away this statistical approach. I wonder whether it might be worth adopting a rather more sophisticated use of bioinformatics-type procedures to illustrate the natural sequence variability or mutational data superimposed upon the proposed structure, and show how this changes as you change the proposed model. This might give greater insight. At the moment we are starting to sink in a sea of data. If there are 100 or so restraints, how can one retain a mental picture of which ones are satisfied and which ones fail as a function of changes in the model? One needs better tools to rapidly assess what is failing and what is succeeding. Koux: At the moment there are so few constraints it is pretty easy to satisfy all of them, unless they are really inconsistent. Sansom: I was thinking of bringing in softer constraint data from bioinformatics or analysis of sequence variability. With the aquaporins, Heymann & Engel (2000) used correlated mutational analysis to aid in folding the sequence into the electronmicroscopy derived density. The information available will get richer as we get more high-resolution structures. Coupling data mining on such structures to bring in soft restraints will not position things precisely, but may improve our radius of convergence. Koux: But such correlated mutation analysis has been tried on K + channels, hasn't it?

264

FINAL GENERAL DISCUSSION

Swartz: In specific regions. Between the core domain and the voltage sensor a number of groups have tried to look at the sequence variability and pick out places where a residue is on one variant and there is a compensation somewhere else. No one has done it systematically across the protein as far as I know, although I think Eduardo Perozo's group might have tried. Pero^p: The problem is that in systems that were more easily understood at the time, the correlation analysis showed that it wasn't very sensitive. It only works with very specific, strongly energetically coupled groups (like salt bridges). The idea is very interesting, anyway. Ashcroft: If I look at a model, I don't know which mutations would be the most useful for testing it. Some of them must be much stronger tests than others, because they will provide greater restraint on the way that the model is constructed. Roux: I showed 100 models in my paper (Roux 2002, this volume). Given the restraints and the window of uncertainty I assign to them, one can see how things are located in space. There are assumptions, but if one tries to remove everything that is debatable, then one still has some sort of rough position for S4. Pero^o: That's where your misgivings might be misplaced. You develop an approach and there is a set of data that you have to work with. That's all you have. The model will be as good as the amount of reliable data you have. As the experimentalists keep pumping out reliable data, you will improve your models. Swartz: I would vote for the first generation of models being very picky in terms of the constraints used. People will then have more faith in the predictions. Mindell: I have been struck at this meeting by how few structures there will be. It is astounding how good KcsA is as a model for the Shaker K+ channel. The acetylcholine (ACh) binding protein looks just like the ACh receptor. To a certain degree you would expect proteins that are related to have the same fold, but it really looks like there is a lot of room for sequence variation but a similar overall structure at the same time. This will make all our lives easier. References Heymann JB, Engel A 2000 Structural clues in the sequences of the aquaporins. J Mol Biol 295:1039-1053 Lemmon MA, Engelman DM 1994 Specificity and promiscuity in membrane helix interactions. Q Rev Biophys 27:157-218 MacKenzie KR, Prestegard JH, Engelman DM 1997 A transmembrane helix dimer: structure and implications. Science 276:131-133 Palczewski K, Kumasaka T, Hori T et al 2000 Crystal structure of rhodopsin: a G proteincoupled receptor. Science 289:739-745 Roux B 2002 What can be deduced about the structure of shaker from the available data? In: Ion channels: from atomic resolution physiology to functional genomics. Wiley, Chichester (Novartis Found Sym 245) p 84-108

Index of contributors Non-participatingco-authors are indicated by asterisks. Entries in bold indicatepapers; other entries refer to discussion contributions.

\

Findlay, J. B. C. 207, 218, 219, 220, 221 *Franks, C. 240 *Froger, A. 33 Fu, D. 20, 30, 49,51, 61, 62, 63, 64, 162, 163,166,204,235,237,257

Ashcroft, F. 49, 63, 81, 82,103,126,142, 144,158,159,165,166,191,192, 203, 206, 218,219, 220, 221, 259, 261, 264 Auerbach, A. 48, 103, 106, 168, 223, 234, 235, 236, 237, 238

G B

*Garofoli, S. 109 *Grauso, M. 240 *Grigorieff, N. 193 Gulbis, J. M. 32,127, 142,143, 144, 176

*Baker, K. A. 169 *Beckstein, O. 66 *Biggin, P. C. 66 *Bond, P. 66 *Brejc, K. 22

H *Harrison, M. A. 207

C *Cheng, A. 33 Choe, S. 18,19,20,30,31,49,61,62,63,64, 65, 80, 82, 83,105,142,143,144, 158, 163, 165,166,167,169,175,176,177, 205 Corringer, P.-J. 16, 18, 19, 82, 159, 238, 258, 259 "Cortes, D. M. 146 *Cuello, L. G. 146 *Culetto, E. 240 *Cushman, S. 169

J Jordan, P. C. 18, 48, 62,109,122, 123, 124, 125,126,234,235

L Lambert,]. 15,18,259 *Law, R. J. 66 *Libson, A. 51 *Lin, M. A. 178 *Liu, Y.-S. 146

D

M

*Dorman, V. L. 109 Doyle, D. 47,61

Mathie, A. 17, 32, 50, 81, 143,177 Miller, K. 17,18,19, 20, 31, 48, 81, 82,107, 165, 166, 167,168, 175, 205, 218, 219, 221,258,259,262,263 *Miloshevsky, G. 109

F *Faraldo-G6mez, J. 66 265

INDEX OF CONTRIBUTORS

266

Mindell, J. A. 17, 29, 48, 49, 61, 63, 80, 81, 102,104,105,106,107,143,158,160, 166,191,193, 203, 204, 205, 206, 261, 262, 263, 264 Mitra, A. K. 19, 33, 46, 47, 48, 49, 50, 61, 63, 64,160,192, 205, 262

p Papazian, D. M. 19, 64,102, 104, 105, 142, 144,175,178,190,191,192,261,263 *Patargias, G. 66 Perozo, E. 16,17,19,31,47,64,79,83,103, 104,105, 124, 125, 126,146,158, 159, 160,161,162,163,165,166,190,203, 205,261,264 *Pfaffinger, P. 169 *Pirruccello, M. M. 193 R

*Raymond, V. 240 *Reddy,V. S. 33 *Ren, G. 33 Roux, B. 15, 17, 31, 63, 79, 80, 81, 84, 101, 102,103,104,105,106,107,123,124, 125,126,161,192, 219, 221, 235, 236, 237,261,263,264 S

Sansom, M. S. P. 1, 17, 20, 32, 49, 50, 61, 62, 66, 79, 80, 81, 82, 83,102,103,104, 105,107, 123,124,125,126, 144, 159, 160,165,166,190, 204, 205, 219, 220, 221,237,257,260,262,263

Sattelle, D. B. 30, 50, 240, 257, 258, 259, 260 Schirmer, T. 220 Schrempf, H. 30, 48, 49, 64, 80, 143, 158, 162,176,219,220,258 *Silverman, W. R. 178 Sixma, T. K. 16, 22, 29, 30, 31, 32, 79, 104, 162,238 *Smit, A. B. 22 *Sompornpisut, P. 146 *Stroud, R. 51 Swartz, K. 101, 102, 105, 106, 142, 143, 159, 163, 191, 204, 205, 206, 264 T

*Tang, C. Y. 178 Tieleman, D. P. 21, 46,66,122 *Tiwara-Woodruff, S. K. 178 *Towers, P. 240 U

Unwin, N. 5, 15, 16, 17, 18, 19, 20, 21, 31, 47, 49, 79, 82, 126, 160, 161, 166, 175, 204,205,206,219 204, 205, 206, 21 V

*vanDijk, W. J. 22 W

Wallace, B. A. 29, 63, 175, 204

Subject index acetylcholine receptors (AChRs) 223-234 gating 223,224-225 spatial organization 230 genomics 240-257 propagation 230-231 rate—equilibrium linear free energy 227-233,234-235,23 structure 223-224 ACh-binding pockets 6, 16 acr-2 245 acr-3 245 acr-4 245 ACR-7 246 ACR-8-like group 245 acr-16 245 ACR-16-like group 245-246, 258-259 ACR-16 subunit 249-250 AgTx2 103,106, 263 alamethicin 68 aldicarb 247 aldo-keto reductase (AKR) superfamily 128 ft subunit, relationship to 131-134 a barrel structures 53 a helical segments 7, 8, 9 a subunits 6,16, 24-25 rotation 12, 18 Alzheimer's disease, drug management 241 amino acid sequencing 1 amphipathic channels 54, 71 antimonite permeability 52 antisense approach, neural signalling 254 AQP1 see aquaporin 1 AQP2 50 AQPZ 52 channel permeability and selectivity 52, 54 ion exclusion 55 aquaglyceroporins 51, 52 conservation 56-59, 61 obesity and 52 aquaporin 1 (AQP1) 33-46, 109 « barrels 53 aqueous passage 38, 40, 42 diffusion-limited 34

A

acetylcholine-binding protein (AChBP) 16, 22-29,223,231,233 a subunit 24-25 binding sites 30 8 subunit 25 disulfide bonds 24 evolutionary origin 31—32 i// subunit 25 homopentamer 23 immunoglobulin fold 23-24 loop regions 25-26 toxin binding 29 tyrosine cornerstone 24 acetylcholine-gated channel 5-15 ACh-binding pockets 6, 16 a helical segment 7, 8, 9 y. subunits 6, 16 rotation 12, 18 anion channels 247—248 basic architecture 5-7 charge distribution 15 disulfide bonds 18,19 framing residues 8 gate 6, 9 gating action 10, 12, 20 hydrophobic residues 9 ion-conducting pathway 6 leucine residues 9, 12 M2 8, 9, 10, 12, 20 M4 18 MA 7-8 mutagenesis experiments 12-13,17 pore closed 8-9 constriction zone 6 open 9-10, 17 vestibules 6-7 cytoplasmic 7—8, 19 extracellular 7 role 7-8 windows 7—8, 18 267

SUBJECT INDEX

268

aquaporin 1 (AQP1) (cont.) binding sites 43 carbon dioxide permeability 34 cation conductance 34, 42 conservation 56, 61 constricted region 110 density map 35 electron crystallographic data 36 fourfold axis region 42-43, 47, 49 gating 50 hydrophobic residues 42, 43, 48, 49 monomer halves, spatial disposition 38 mutagenesis experiments 34 N- and C-terminals 35, 38 NPAloop 34,38 polar residues 42, 48 pores 38,40,42,47 quasi-twofold symmetry 53 structure 52-53, 71 aquaporins 51, 52 channel-lining residue mutations 50 conservation 56-59, 61 correlated mutation analysis 263 physiological significance 52 water molecule behaviour in 71—73 Argl87 50 Asn76 43,48 Asnl92 43,48

B bacteriorhodopsin 47 0subunit 127-142,173,177 catalytic activity 129,131 cell surface expression 128 channel inactivation 128 historical context 128-129 octamer 129, 134, 143-144 redox activity 129, 142 relationship to AKRs 131-134 Tl-j8 assembly 134-137,142 T1-S1 linker ' 137-140 tertiary fold 128,131-134 Born energy 113,117,118-119 Brownian dynamics 110-111 bungarotoxin 250 binding 29 butamisole 250

c Caenorhabditis elegans 241

antisense approach, neural signalling 254 calcium imaging 253 channel diversity, reasons for 257-258 electrophysiology 253 gene silencing 254 as a genetic model 242 nicotinic receptors 250-251 protein diversity 260 subunit spatial expression patterns 246 calcium imaging 253 carbon dioxide permeability 34 cardiac action potential 191 cation conductance 34, 42 cavity 111-112,170 permeation energetics 117-119 radius 118 role 113-115 charge distribution 15 chloride channels, monovalent anion selectivity 204 see also CIC-type chloride channels choline 224,250 CIC-type chloride channels, electron diffraction 193-203 data analysis 195 diffraction patterns 199,201 projection structure 201-202 specimen preparation 194-195 COMP 9 conformational sampling 94 conservation 27-28, 56-59, 61, 170, 209 correlated mutation analysis 263-264 Cysl89 34 Cys-loop 27 Cys-loop receptors 22-23 cysteine scanning mutagenesis 13, 19-20 cytoplasmic vestibules 7-8, 19, 170 D £W/EW 68-69 D194 231 D274 188 D278 185, 187 D327 187 DEG-3 245,250 DEG-3-like group 245,246 dehydration energy 112,123

SUBJECT INDEX dementia, nAChR reduction 251 DES-2 245,250 diabetes insipidus, nephrogenic 50, 52 dicydrohexylcarbodiimide 213 dipole—dipole interaction 148, 150 distance energy restraints 92 disulfide bonds 18,19,24 double-stranded RNA interference 254 Drosophila melanogaster 241 as a genetic model 252 nAChR gene family 252 drug development/targets 1, 241, 242, 245 drug resistance 242

E E5 214 El 52 55-56,62 E283 182-184 eat-2 245

eat-18 245 egg-laying behaviour, nicotinic receptor genes 251 electron crystallographic data 36 electron diffraction 193-203 data analysis 195 specimen preparation 194-195 electron paramagnetic resonance (EPR) spectroscopy 146-158, 159-160, 167 principles and data 148—150 electrophysiology 253 electrostatics 68, 110-111 epilepsy 241 EriC crystals 194 electron diffraction 194-202 data analysis 195 diffraction patterns 199,201 projection structure 201—202 specimen preparation 194—195 ether-a-go-go (eag) 191 acidic residues 185,187-188 expression 191 magnesium binding 185, 187-188, 190, 191 metal ion binding 102 voltage sensor 180, 184-188 evolution 31-32,59 extracellular vestibules 7

2.69

F

F200 55 F0 domain 214,220 F0FrATPase 215,218 flicker 159 framing residues 8 Friedel R factors 195-197, 201

G G199 55 gate/gating 28, 84 acetylcholine-gated channels 6, 9, 10, 12, '20 AChR 223, 224-225, 230 AQP1 50 current analysis 180 equilibrium constant 223 hydrophobic 69—71 KcsA 154-157, 158-159, 161 unliganded 238-239. GCN4 leucine zipper 43 gene silencing 254 genomics 2-3,240-257 glia cells 23 GlpF 42,43,49,51-61, 109 a barrels 53 amphipathic channel lining 54, 71 antimonite permeability 52 channel permeability and selectivity 52, 54-56 conservation 56-59, 61 crossing contacts 56-57 hydrogen bonding 54-55 ion exclusion 55—56 NPA motif 57,61 physiological role 63 polarity 62 polyol permeability 52, 54 quasi-twofold symmetry 53-54 stability 64 structure 52-54, 71 vestibules 58 water in 62, 71-73 Glul40 212 glycerol release 52 glycine stripe 212, 220 glycophorin mutagenesis 261 gramicidin 110 green fluorescent protein 258

270

SUBJECT INDEX

Gromacs 69 GROMOS96 69 GxxxG motif 212

H hanging basket 137 hanging gondola 91,171 Heisenberg spin exchange 148 HEPES buffer 26,30-31 HERG 191 HislSO 43 HOLE 69 hydrogen bonding 54—55 hydrophobia gate 69-71 hydrophobia residues 9, 42, 43, 48, 49 3a-hydroxysteroid-dihydrodiol dehydrogenase (HSD) 131, 133, 134

I immunoglobulin fold 23-24 interspin distances 148,150 intragenic suppression 89 ion exclusion 55-56 ion solvation 123 ivermectin 250

L

J jtf-38

relationship to AKRs 131 -134 Tl-/? assembly 134-137,142 T1-S1 linker ' 137-140 tertiary fold 128,131-134 KchIP 173,177 KcsA 2, 109 cavity 111-112 permeation energetics 117-119 radius 118 role 113-115 constricted region 109-110 crystallographic structure 85 gating mechanism 154-15"7, 158-159, 161 hydrophobia gate 70 mobility 159-160 permeation energetics 111, 115-120 pH dependency 154, 162-163 simulation 79 site-directed spin labelling 150-154 structural similarity to Shaker 88-89 structure 85 kinetic modelling 1 Kubo-Zwanzig Linear Response Theory 81

245,249

K K374 181-182 K v channels 84-85,127 conservation 170 cytoplasmic vestibule 170 proteins regulating excitability 177 side openings 170 Tl domain and excitability 169-175 tetramerization 170 voltage sensor 171,178-190 function 179 S2/S4 involvement 180 two-phase movement 180 K v l channel, 0 subunit 127-142 catalytic activity 129,131 cell surface expression 128 channel inactivation 128 historical context 128-129 octamer 129,134,143-144 redox activity 129,142

lanthanide-based resonance energy transfer (LRET) 89-90, 92, 94, 101,' 107 leucine residues 9, 12 leucine side chains 69-70 LEV-1 251,258 lev-11 247 levamisole resistance 242, 250 ligand binding 23, 24-25 ligand-gated ion channel superfamily, conservation 27-28 light harvesting complex (LHC) 201 LINGS algorithm 69 lipid-exposed residues 92 94 LysSl 43,49

M M2 8,9,10,12,20 M4 18 MA 7-8 magnesium binding 185, 187-188, 190, 191 McsL 109 hydrophobic side chains 69-70

271

SUBJECT INDEX mechanosensitive channels 9 membrane proteins 1, 67 N-methyllycaconitine (MLA) 250 MIP members 33-34 MODELLER 92 molecular dynamics 68, 69, 110 Molscript 69 morantel 250 muscle 246,247 mutagenesis experiments 12-13, 17, 34, 90-91,261 myasthenias 241

N nephrogenic diabetes insipidus 50, 52 nervous system development, nAChRs in 246, 251-252 neuronal sprouting 251-252 nicotinamide adenine dinucleotide phosphate (NADP) 129 nicotinic acetylcholine receptor (nAChRs), genomics 240-257 diverse gene family 245-246 drug targets 242, 245 egg laying behaviour 251 functional and pharmacological diversity 249-250 genes functionally linked to 246-247 mutations 241 nervous system development 246, 251-252 novel functions 247-248 nitroxide dynamics 148 Nod26 48,50 non-liganded receptor opening 238-239 O

O program 35 obesity 52 OmpA 73-76 oxantel 250 P

patch clamping 1 permeation energetics 111, 115-120 perturbations 225 pH dependency 154,162-163 pharyngeal muscle activity 245, 251

F 226, 228, 229-232, 233, 235, 236, 237, 238-239 phospholamban 9 PME 69 Po-V relationship 180 polar residues 42, 48 polyol permeability 52, 54 pores

AQP1 38,40,42,47 blocking 90 closed 8-9 constriction zone 6 models 68,92 open 9-10, 17 openness 70-71 Shaker 88-89,92 typical 67 power saturation 148 proline 20 proteolipid 16kDa 209,210-212 conservation 209 proton channel see vacuolar H+-ATPase pyrantel 250

Q Q-V relationship 180, 191 quaternary ammonium pore blocker 90 QX222 12 R

R206 5S--56 R factors 195-197,201 RasMol 69 Raster3d 69 rate-equilibrium linear free energy (LFER) 225-233, 234-235, 236 redox activity 129,142 residue environmental phase diagram 151 restraint-driven Cartesian transformation (ReDCaT) 155 rhodopsin 262 ribitol 54,55 rie-3 247 RN A editing 177 S

S1-S4 87-88,91 S2, voltage sensing 84-85, 180

SUBJECT INDEX

272

S4 94, 95-96, 97,101,102,104,188 voltage sensing 85,180 scanning mutagenesis 90-91 schizophrenia, drug management 241 second site suppressor analysis 181-184, 190-191 Shaker 84-101 central pore 88-89,92 conformational sampling 94 distance energy restraints 92 intragenic suppression 89 lanthanide-based resonance energy transfer (LRET) 89-90, 92, 94, 101,'107 lipid-exposed residues 92-94 modelization 91-92 pore blocking 90 S1-S4 87-88,91 54 94,95-96, 97, 101, 102, 104, 188 scanning mutagenesis 90-91 simulated annealing 94 structural similarity to KcsA 88-89 two-domain architecture 91 voltage sensor 85, 179, 181-184 side chain associations 97 SIGMAA 35 simulated annealing 94 single-molecule spectroscopy 166 site-directed spin labelling 146 KcsA 150-154 solvent accessibility 148 solvent-accessible surface area (SASA) 93 spectral second moment 148 spin label I 146-147 spin label dynamics (mobility) 148, 159-160 spin labelled molecules, structural information from 148 Src kinase 177 stabilization energy 112 synaptogenesis 246 T

Tl 128, 169-175 cavity 170 conformational changes 171-172 physiological role 171-172 removal 175 voltage-sensing 171 zinc binding 171, 173, 176 Tl-0 assembly 134-137, 142, 170, 171 T1-S1 linker " 137-140

tandem dimer constraints 155 temperature 234-235 threonine 12 TIM barrel 129, 133 tin derivative 213-214 Tourette's syndrome, drug management 241 toxin binding 29 tropomyosin 247 tryptophan scan 105 twitchin 247 tyrosine cornerstone 24

U unc-22 247 UNC-29 246,251,258 UNC-29-like group 245, 249 unc-38 251 UNC-38-like group 249 unc-50 246 UNC-63 258 UNCL 246-247 unliganded gating 238-239

V vacuolar H+-ATPase 207-218 16kDaproteolipid 209,210-212 functionally sensitive sites 212-214 mechanism of action 215-216 transmembrane organization 209-210 valines 9 vestibules 6-7, 58 cytoplasmic 7-8, 19, 170 extracellular ~ role 7-8 VMD 69 voltage gating 139-140 voltage sensor I"7!, l^S-^O function 179 S2/S4 involvement 84-85,180 two-phase movement 180 Vphlp 210,214215,218

W water 66-78 behaviour AQP1 34,38,40,42 GlpF 62,71-73 binding, AQP1 43 dynamic behaviour 68

SUBJECT INDEX

273

electrostatic properties 68 models 68, 69 windows 7—8, 18

XPLOR 35 xylitol 54, 55

X

Z

X-ray crystallography 52

zinc binding 171,173,176