Biomembrane Transport

Biomembrane

Transport

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Biomembrane Transport Lon J. Van Winkle Midwestern University

With contributions by Ovidio Bussolati, Gian Gazzola, and John McGiven Bryan Mackenzie, Milton H. Saier, Jr., Peter M. Taylor, Michael J. Rennie, and Sylvia Y. Low

A C A D E M I C PRESS San Diego

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This book is printed on acid-free paper. @ Copyright 9 1999 by ACADEMIC PRESS All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher. Academic Press a division of Harcourt Brace & Company 525 B Street, Suite 1900, San Diego, California 92101-4495, USA http://www, apnet, com Academic Press 24-28 Oval Road, London NW1 7DX, UK http ://www.hbuk.co.uk/ap/ Library of Congress Catalog Card Number: 98-89087 International Standard Book Number: 0-12-714510-9 PRINTED IN THE UNITED STATES OF AMERICA 99 00 01 02 03 04 EB 9 8 7 6

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To my wife, Mikki, who taught me to learn by resolving differences.

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Contents

W. The Gibbs-Donnan Effect Also Generates

Foreword xi Preface xiii

Osmotic Pressure 47 VI. Chemical Reactions Drive Primary Active Transport 49 VII. Reversal of Transport May Drive Chemical Reactions 55 VIII. How Do Fluctuations in the Local Hydrogen Ion Potential Facilitate Formation of Phosphoric Acid Anhydride Bonds by the Mitochondrial FoFI_IATP Synthase? 56 IX. Conversion of Solute Total Chemical Potential Gradients to Gradients of Other Solutes during Co- and Countertransport 57 X. Dissipation of Solute Gradients through Mediated Transport Processes May Also Perform Work 61 XI. Application of Thermodynamic Principles to the Solution of Practical Transport Problems 63 XII. Summary 63

1. I m p o r t a n c e of B i o m e m b r a n e T r a n s p o r t I. Introduction 1 II. Solute and Solvent Fluxes Are Determined by Barriers and Propelling Forces 3 III. Biomembrane Transport in Context 7 IV. Summary 10

2. Biomembrane Composition, S t r u c t u r e , and Turnover I. Introduction 13 II. Is the Fluid Mosaic Model of Membrane Structure Still Adequate? 13 III. Some Components of the Biomembrane Can Be Reconstituted 29 IV. How Are Biomembrane Composition and Structure Regulated? 30 V. Summary 38

4. T r a n s p o r t Kinetics I. Introduction 65 II. Kinetics of Diffusion 66 III. How Do Measurements of both the Diffusional and the Osmotic Permeability Coefficient for Water Inform Us about the Mechanism of Water Transport across a Plasma Membrane? 70 IV. Do Lipophilic Substances Migrate across Biomembrane Phospholipid Bilayers by Simple Diffusion? 73 W. Lipid-Soluble Substances Are Used to Attempt to Measure the Width of Unstirred Water Layers on Either Side of Biomembranes 74

3. T h e r m o d y n a m i c s a n d T r a n s p o r t I. Introduction 39 II. Similar Mathematical Expressions Serve for the Free Energy Change in a Chemical Reaction and in the Migration of a Solute or Solvent 39 III. Changes in Enthalpy and Entropy May Contribute Differently to the Free Energy Changes Associated with a Biochemical Reaction and Migration of a Solute 43 IV. The Total Chemical Potential Change for a Transport Process Also May Have an Electrical Component 44

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VI. Do Such Determinations of the Apparent Widths of Unstirred Water Layers Reflect the Intended Physical Phenomenon or Our Ignorance of How Lipid-Soluble Substances Cross Biomembranes? 76 VII. Protein versus Lipid-Mediated Mechanisms of Fatty Acid Migration across Biomembranes 79 VIII. Protein-Mediated Biomembrane Transport Is Probably Always Substrate Saturable 81 IX. Kinetics of Saturable Transport 83 X. Identification and Minimization or Deduction of Processes That May Obscure a Transport Process of Interest 98 XI. Kinetic Differences among Substrate-Saturable Transport Processes That Form, Propagate, or Dissipate Solute Gradients 116 XII. Summary 124 Appendix 126 5. Structure a n d Function of Transport Proteins That Form Solute Gradients I. II. III. IV.

Introduction 133 P-Type ATPases 135 FoFI-ATP Synthases (F-Type ATPases) Summary 166

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6. Transport Proteins That P r o p a g a t e Solute Gradients

II. III.

IV. V~

Introduction to Symporters and Antiporters 169 Both Erythroid and Nonerythroid Tissues Express Anion Exchangers 170 ASC and Excitatory (Anionic) Amino Acid Transporters Comprise One of Two Known Families of Mammalian Na+/Amino Acid Symporters 208 Both AE and EAAT/ASC Proteins Have Additional Functions 233 Summary 237

8. A P r o p o s e d S y s t e m for t h e Classification of T r a n s m e m b r a n e Transport Proteins in Living O r g a n i s m s Io Introduction 265 II. Work of the Enzyme Commission as a Basis for the Systematic Classification of Transport Proteins 265 III. Phylogeny as a Basis for Protein Classification: Criteria for Family Assignment 266 IV. Proposed Transport Protein Classification System 267 go Representative Examples of Classified Families 272 VI. Cross-Classification of Transport Proteins 272 VII. The Two Largest Superfamilies of Transporters: The MF and ABC Superfamilies 275 VIII. Macromolecular Transport Proteins in Bacteria 275 IX. Conclusions and Perspectives 276

9. Regulation of Plasma M e m b r a n e Transport I~ Introduction 277 II. Regulation of Transport by Changes in Driving Force: The Role of Plasma Membrane Potential 277 III. Regulation of the Activity of Existing Transporters through Modifications of Transporter Molecules 278 IV. Regulation of Transport by Changes in the Repertoire of Transport Proteins in the Plasma Membrane 284 V~ Coordinated Regulation of Transport Systems 287 VI. Derangements in Transport Regulation 287 VII. Summary 293

10. B i o m e m b r a n e Transport a n d I n t e r o r g a n Nutrient Flows: The A m i n o Acids I~ Interorgan Nutrition

II. 7. Channel Proteins Usually Dissipate Solute Gradients I. Introduction 239 II. Structure, Function, and Evolution of Channel Proteins 240 III. Kinetics of Transport via K + and Other Channels 254 IV. Summary 262

III. IV. V~

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295 Interorgan Amino Acid Nutrition: General Principles and Key Issues 295 Control of Interorgan Amino Acid Metabolism: Metabolic Control Theory and Safety Factors 308 Physiologically Important Flows of Amino Acids and Related Compounds 311 Amino Acid Nutrition under Special Circumstances 319 Summary 325

Contents

1 1. S e l e c t e d T e c h n i q u e s in M e m b r a n e Transport I. Introduction 327 II. Purification and Reconstitution of Transport Proteins 327 III. Methods for Isolating cDNAs Coding for Transport Proteins 328 IV. Heterologous Expression Systems for Transport Proteins 329 V. Voltage-Clamp Techniques in Xenopus Oocytes 332 VI. Probing Transport with Ion-Selective Microelectrodes 338

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VII. Optical Methods for Measuring Membrane Transport 339 VIII. Structure-Function Studies of Transport Proteins 339 IX. Genetic Approaches to Understanding Transporter Function 341 X. Summary of Preparations Used to Study Native Membrane Transport 341 XI. Commentary Epilogue 343 References 345 Index 387

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Foreword

Originally conceived as an update and expansion of my 1975 edition of Biological Transport, Van Winkle's Biomembrane Transport integrates recent advances in this broad field with several historically important concepts. Van Winkle argues convincingly that each of the transport proteins functions by interacting intimately with its specific substrate to provide a pathway for the movement of the substrate across a biomembrane. He points out that all such proteins need to move in order to catalyze transport, although the extent of the conformational changes varies greatly among the proteins. This perspective departs significantly from the view that the proteins that form, propagate, and dissipate solute gradients across biomembranes function by a variety of distinct mechanisms. My good impression of Van Winkle's efforts at this integration of transport is strongly heightened by his attention to detail. In separate chapters of this book, Dr. Van Winkle describes what is known about the structures and catalytic mechanisms of several examples of each category of transport protein. In this process he exposes differences as well as similarities in the structures and mechanisms of action of proteins of the same and different categories. What stands out in each of these chapters is how frequently the actual thermodynamics and kinetics of substrate transport appear to differ from currently accepted formulations for the transport. These revelations add up to an important contribution to a field in which numerous investigators are pressing to discover details of transporter structure and action, even though the characteristics of transport itself may still remain inadequately described and appreciated. With these caveats in mind, several guest authors integrate the actions of various types of transport pro-

teins in chapters on transporter regulation and the resulting interorgan flows of their substrates. I call attention especially to the remarkable, current development of the subject of competition of amino acids for transport across the blood-brain barrier presented in Chapter 10, particularly in phenylketonuria, where phenylalanine in excess is the dangerous competitor, and in maple syrup disease, where it is instead leucine, a leucinosis. Learning of such physiological and pathophysiological functioning of the transporters is of course the purpose for studying them, although this goal may sometimes be obscured in experiments using powerful new molecular procedures. In a guest chapter on some of these techniques, Dr. Bryan Mackenzie makes the important observation that we are likely to return to greater use of conventional preparations to study biomembrane transport as an appropriate emphasis of their overall physiology is restored. This concern urges the modern investigator to understand and be prepared to use a wide array of procedures available for studying transport at the subcellular, cellular, tissue, and organismal levels of biological organization. Additionally, we can comprehend fully the breadth of our field of biomembrane transport by examining carefully for its bounds, for example, among enzymes whose characteristic actions differ from those of transporters in destabilizing their substrates rather than in simply moving them from one phase to another. In short, Lon Van Winkle's effort helps us very much in describing what we know and what we do not know about biomembrane transport. I believe he has done an outstanding job. He has ranged thoughtfully in his invitation of guest authors to broaden his already good perspective. His book asks many questions and provides good answers. For myself, a person who has faced questions on membrane transport for about a half-century,

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I find this book both insightful and provocative. Watching it evolve has been very satisfying and rewarding to me, I having until now privately included Lon Van Winkle among my own students, even though no such formal arrangement was ever made. I encourage other stu-

dents of membrane transport to study this book to seek the benefits of the continued development of the field.

Halvor Christensen

Preface

Some new investigators may find themselves in the field of biomembrane transport in part serendipitiously because an interesting cDNA clone happened to encode a transport protein. Others may have been led to the field through their investigations relatively late in their careers, well after their formal training was complete. It is hoped that this book will help such individuals fill deficiencies that may exist in their knowledge of biomembrane transport. Beyond this more limited goal, the book is intended to give any interested student of biochemistry and molecular biology insight into what is as well as what is not yet known about biomembrane transport and its importance to the physiological functions of cells. The book is divided into three main parts. The first part (Chapters 2 to 4) covers fundamental principles of biomembrane structure and transport. In the second part (Chapters 5 to 7) we discuss the structures and functions of transport proteins that form, propagate, and dissipate solute total chemical potential gradients. Finally, three chapters (8 to 10), written by prominent guest authors, span the topics of classification, regulation, and integration of the functions of biomembrane transport proteins. Modern techniques for the study of biomembrane transport are discussed briefly in several sections of various chapters and in Chapter 11. Chapters 8 to 11 add not only important dimensions to the book, but also the unique perspectives of the guest authors. I leave it to the guest authors themselves to reveal their sometimes novel views on transport in their individual chapters and do not speak for them in this preface except coincidentally. Transport proteins have evolved on numerous occasions to catalyze migration of a solute or the solvent across biomembranes. Such evolution has been necessary because membrane lipid bilayers otherwise present virtually impenetrable barriers to most hydrophilic sol-

utes. Hence, it became possible to regulate the composition of intracellular and extracellular fluids with the advent of biomembrane transport proteins. Moreover, such regulation was made progressively more sophisticated as more types of proteins evolved to transport the same as well as different solute species. Modern organisms appear now to need such diversity of biomembrane transport processes to compete successfully with other species. Such circumstances also mean, however, that the biomembrane transport proteins that evolved in apparently unrelated families and superfamilies nevertheless evolved under similar constraints; new transport processes have had to improve the ability of the organism to fit into a successful niche in the biological community by influencing a single main function of their cell or cells. Consequently, virtually all such biomembrane transport proteins function in two fundamentally similar ways. 1 First, they provide pathways for the migration of their substrates across biomembranes. Such pathways involve temporary association of the substrate with one or more sites along the pathway, thus rendering the pathways selective for one or a few chemically and physically similar solutes. Moreover, such mediated transport is substrate saturable apparently because the interactions between substrate and transport protein necessarily slow migration of the solute relative to the rate at which it could migrate over the same distance by ordinary diffusion. Nevertheless, the rate of biomembrane transport varies among proteins over nearly 10 orders of magnitude, apparently owing to a need for 1 We are discussing here the majority of transport proteins that are produced by organisms for their own uses. Not included in this summary are transport proteins, such as cz-hemolysin, that are produced by an organism in order to cause the death of the cells of another. The latter proteins function by insertion of the transport protein molecules into the plasma membranes of cells of the target organism.

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such differences in rate under various physiological conditions. While it is conceivable that differences in the way in which transport proteins interact with their substrates could alone account for their wide range of known transport rates, we think this wide range in rates also depends on differences in the magnitude of the other fundamentally similar way in which virtually all transport proteins function. 2 As we shall see, virtually all biomembrane transport proteins need to move in order to catalyze transport. In the cases of transport proteins historically known as carriers or primary active transporters, such conformational changes may be relatively large and easy to document, although none of the proteins completely reverses its conformation across the membrane. In contrast, the more rapid migration of solutes and the solvent across the membrane via channels appears to require only the small movements that all macromolecules normally undergo. Some readers may question whether we have accepted prematurely data showing that channel proteins must be able to move normally in order to catalyze transport. These data are based primarily on computer simulations of protein structure and on similarities between channel proteins and enzymes in their interactions with substrates. We maintain, however, that the principal reason the movements of channel proteins during transport are not as well documented as the movements of other types of transport proteins is that channel proteins are not expected to move except to open, close, or inactivate. However, as for all proteins and other molecules at temperatures well above absolute zero, channel proteins and other membrane constituents do exhibit predictable motion, and their movements as well as that of the substrate are needed for transport to occur.

In a similar vein, we challenge the common notion that transport in some cases occurs by a process that resembles ordinary diffusion. The notion of transport by diffusion of course contradicts the theory that transport via transmembrane pathways formed by proteins requires the proteins to move during transport. As we shall see, however, it is also our position that even lipophilic solutes do not appear to migrate across the highly ordered lipid bilayers of biomembranes by processes that resemble ordinary diffusion. If we are right, one consequence would be that the widths of the unstirred 2 Use of the word "we" to refer to the primary author here or in other sections of this book should not be taken to mean that guest authors share all of his opinions about biomembrane transport. The opinions expressed in each chapter are those of the author or authors of it and may or may not be shared by the others.

water layers on either side of the lipid bilayer have been vastly overestimated. We hope that readers will accept our good intention of such challenges to common theories and beliefs about the mechanisms of biomembrane transport. We accept at the outset that many of our notions may be incorrect, but we think that accepted paradigms may themselves also not be well supported by experimental data. Our purpose then is to provoke thought and further study in these instances. It is after all such an inquisitive spirit, as well as our disagreements, that inspires us to develop and test creative new theories about the functions of biomembrane transport proteins. The field of biomembrane transport also has become too broad for a detailed discussion of all important instances of such transport. Consequently, we discuss many principles that are pertinent to all transport processes, but the examples selected to illustrate these principles are only a very few of the numerous wellstudied examples that could have been chosen. Similarly, to discuss the relationship of protein structure to function in enough detail to present a full view of the state of the art, only some of the many important examples of transport proteins had to be selected. If, however, we are correct in our assertion that virtually all transport proteins function according to fundamentally similar principles and mechanisms, then selection of these examples should indeed give the reader the necessary insight into the broad field of biomembrane transport. Many people contributed to the production of this book, and I will not attempt to mention each one by name lest I forget someone more deserving than those I remember. Most people who helped to prepare the book are members of various departments at Midwestern University, including Biochemistry, Library Services, Media Resources, and Research Affairs. Individuals in these departments who must be mentioned by name because of the quantity of work they performed include Allan Campione, Barbara Le Breton, Michael Moore, and Eileen Suarez. Moreover, several colleagues provided constructive criticisms of more than one chapter, sometimes exposing differences in our opinions. Although many of these differences were constructively resolved, some still remain, so my colleagues are not to blame for my ideas about transport that may turn out to be wrong. Readers should also see the acknowledgments in individual guest chapters for persons who contributed to production of those parts of the book. Colleagues who reviewed several or all of the first seven chapters include Stefan Br6er, Halvor Christensen, Jacquelyn Smith, Susan Viselli, Douglas Webster, and James Young.

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1 Importance of Biomembrane Transport

1. INTRODUCTION Asexual, sexual, and cellular reproduction and the functioning of cells and organisms may be viewed in relation to various modifications of the central dogma. The dogma historically places nucleic acids and in particular D N A at a central position in biology. D N A is of course needed for organisms to reproduce and for them to pass their genes to the next generation. The only measure of an organism's biological productivity is the relative quantity of heritable D N A that it, and in some cases, its relatives contribute to subsequent generations. Despite the importance of nucleic acids to our comprehension of how living things function, other cellular constituents are of course required for cells and organisms to reproduce and remain alive. In particular, biocatalysts are needed both to interpret the information in nucleic acids and to propagate the cells and organisms that contain them (Fig. 1.1). Biocatalysts also convert free energy into biochemical and biophysical forms useful in performing the work of living and reproducing. Any biological molecule or combination of such molecules that increases the rate of a process in vivo qualifies here as a biocatalyst. Familiar forms of biocatalysts include enzymes, ribozymes, chaperones, and biomembrane transport proteins. The same biocatalyst molecule may also increase the rate of more than one process, as we will come to expect in this volume when we consider the multiple functions of many biomembrane transport proteins (especially in Chapter 6). Moreover, these multiple processes may be of the same type, such as multiple independent biomembrane transport processes, or they may be of different types, such as a transport process

that is coupled to a chemical change. For example, the F-type ATPases (or ATP synthases) of chloroplasts convert the free energy of the proton gradient formed by light-driven active transporters into the free energy normally realized in ATP when the ATPases also catalyze transport of protons along their total chemical potential gradient. Biocatalysts that function in biomembrane transport constitute a quantitatively significant portion of all proteins. As pointed out in Chapter 8 of this volume, recent complete genome analysis revealed that about 10% of all genes in microorganisms encode transport proteins. Moreover, catalysts are needed to insert these transport proteins asymmetrically into biomembranes. In the case of photosynthesis, proton gradients can be formed only if transport is asymmetric, and subsequent use of the gradients for ATP synthesis requires that the ATP synthases also function asymmetrically. As for the membranes in chloroplasts and other intracellular organelles, the plasma membrane also is asymmetric, and this asymmetric structure helps to organize important biological processes. For example, watersoluble signaling molecules bind to receptors on the outside of cells. As a result of such binding, a cascade of events often is produced within cells to change their metabolism (Fig. 1.2). These changes frequently involve net transport of solutes asymmetrically in one direction or the other across biomembranes. Clearly, the symmetric functioning of such a system would be of little value to cells. Hence, the asymmetry of the barriers that biomembranes form is as critical to their normal functioning as the transport proteins and other catalysts that are associated with them. The importance of this asymmetry to the normal functioning of the transport biocatalysts may some-

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1. Importance of Biomembrane Transport

FIGURE 1.1 Extension of the central dogma of molecular biology to include reverse transcription and RNA replication. Each of the processes depicted is needed by some or all organisms in order for them to function and survive. Solid arrows in the dogma and its extensions are meant to indicate the directions of information transfer. In addition, proteins and ribozymes are shown as containing information needed to catalyze the processes. Hence, the information needed to sustain life is contained both in nucleic acids and in biocatalysts.

times be more subtle than in the conspicuous instances just discussed. For example, the asymmetric functioning of inwardly rectifying K + channels appears to allow the channels to transport K + ions into cells against their total chemical potential gradient. As we shall see in Chapter 7, this asymmetric functioning depends primarily on the underlying asymmetry of a polyamine gradient with which the channels interact. If verified, this surprising transport would occur critically at the resting membrane electrical potential. Consequently, it makes the membrane more sensitive to depolarizing stimuli. The existence of K + channels in membranes was postulated in the first place because this ion appeared to traverse the hydrophobic interior regions of biomembranes more rapidly than anticipated from the hydrophilic character of K +. In the absence of a protein mediator, the rates at which many solutes permeate biomembranes appear to depend on their molecular masses and lipid solubilities. More hydrophobic substances are sometimes viewed as being able to permeate the phospholipid bilayers of biomembranes more easily than hydrophilic ones, owing in part to their ability to dissolve in and subsequently diffuse through the hydrophobic region at the center of such bilayers. Diffusion of smaller solutes is of course more rapid than larger

FIGURE 1.2 Binding of hydrophilic signaling molecules to their receptors on the outside of a cell frequently activates a cascade of events in the plasma membrane and cytosol. In the case depicted, the norepinephrine-bound receptor actually can stimulate numerous Gprotein molecules (shown as a single o~-subunit that has separated from the 3'- and/3-subunits) each to activate an adenylate cyclase molecule. One result of the signaling in this case is the asymmetric net transport of Ca 2+ into cells along its total chemical potential gradient. The whole system must, of course, also operate asymmetrically across the membrane to be effective (adapted from Opie, 1991, with permission from Lippincott-Raven Publishers).

ones, so better correlations between permeability and hydrophobicity are obtained when the permeabilities are corrected for the size of the solute. Hence, when a substance appears to permeate a biomembrane more rapidly than anticipated from these properties, it becomes reasonable to look for a transport process that may mediate migration of the solute across the membrane. For example, the paradoxically very rapid transport of the solvent water across biomembranes may now be understood largely owing to the presence of water channel proteins in the membrane (see Sections II and III of Chapter 4 for further discussion). Moreover, other membrane proteins, such as the Na+-dependent glucose transporter (Loike et al., 1996; Loo et al., 1996), appear to catalyze transport of significant amounts of water in addition to that catalyzed by water-specific channels. Nevertheless, the migration of water across artificial phospholipid bilayers is still, in our view, paradoxically rapid, and special ways of accommodating water molecules in the bilayer structure have been proposed to account for this migration (e.g., Haines, 1994).

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Solute and Solvent Fluxes

Similarly, other substances may pass across biomembranes more rapidly than anticipated from their molecular size and structure. As for such migration of water, the migration of these solutes across the phospholipid bilayer may be catalyzed by proteins. Alternatively, the solutes may migrate more rapidly because of asyet poorly appreciated properties that appear to permit more rapid permeation of the lipid bilayer than anticipated from molecular size and hydrophobicity alone. For example, c~-tocopherol is a highly lipid-soluble substance whose membrane permeability can be increased by converting it to the larger and less lipid soluble substance tocopherol succinate (Bonina et al., 1996). Hence, protein-mediated transport may not always be present when migration of a solute is more rapid than anticipated. 1 Conversely, protein-mediated transport cannot always be ruled out solely because a solute migrates across the membrane at a rate anticipated from its physical properties alone. Nevertheless, the ability of any molecular or ionic species to move across a biomembrane depends only on the degree to which the membrane serves as barrier to that migration.

evolved partially to overcome the barriers. 3 Rather than using cellular free energy to make biomembrane transport faster than the rate of migration that could be achieved by ordinary diffusion, this free energy is used instead in combination with biomembrane barriers to produce total chemical potential gradients of solutes across biomembranes. Transport along these gradients then serves to perform additional work such as ATP synthesis, signal transduction, and regulation of cellular volume. A. Unidirectional Solute or Solvent Flux D e p e n d s on the D e g r e e to Which a B i o m e m b r a n e Serves as a Barrier to That Migration

The unidirectional flux of a solute or the solvent across a biomembrane proceeds much more slowly than could occur if free diffusion were possible over the same distance. Even the fastest transport via channels has been estimated to proceed no more rapidly than about 8% of the rate that could be achieved owing to free diffusion (calculated by Stein, 1986; p. 202). 2 While it is an interesting theoretical question whether a system could be constructed to catalyze biomembrane transport at a rate exceeding that which would occur if ordinary diffusion were possible, it is difficult to imagine a need for such a system except perhaps in the case of macromolecules. Consequently, the barrier functions of biomembranes can be seen to be at least as important to the lives of cells as the transport processes that have

In transport that is not saturable by substrate, the rate at which the substrate traverses the membrane depends only on the total chemical potential of the substrate, the total surface area of the membrane, and the permeability of the membrane to the substrate. The rate of nonsaturable unidirectional transport is not usually coupled to an obvious source of cellular free energy, nor does it depend on the concentration of the substrate on the other side of the membrane. For example, the unidirectional flux of a solute at a concentration of, say, 1.0 mM will occur at the same rate regardless of whether the solute concentration on the other side of the membrane is 0.1 or 10 mM. The rate of protein-mediated, substrate-saturable transport also need not be influenced by the concentration of the same substrate on the other side of the membrane, although unidirectional flux in the reverse direction will, of course, depend on this concentration. When the rate of mediated unidirectional transport is not influenced by the presence of the same substance or ion on the other side of the membrane, the transport is believed to be catalyzed by uniporters. Such transport is also sometimes imprecisely attributed to facilitated diffusion of the solute across the membrane via a carrier as discussed further in Section VIII of Chapter 4. As for biomembrane barriers, propelling forces influence protein-mediated unidirectional solute and solvent fluxes. The simplest of these forces is the total chemical potential gradient of the substrate. While a mathemati-

1Undetected mediation of transport by a protein is, of course, nearly impossible to rule out formally for any biomembrane. 2A possible exception to the results of this calculation may be transport via nonselective channels formed by some toxins such as a-hemolysin. These toxins form relatively wide pathways for the migration of water and most solutes. To our knowledge, the transport rates via these toxin channels has not been determined and compared to the rate that could occur if ordinary diffusion were possible.

3Similarly, other transport processes increase the rate of migration of inorganic and organic solutes in the cytosol by helping the solutes partially to overcome barriers to their free diffusion (e.g., Bronner, 1996; Luxon, 1996; Weisiger, 1996). However, the transport processes do n o t help the solutes to exceed their rates of ordinary diffusion. Terms, such as "self-diffusion", that are sometimes applied to the uncatalyzed migration of solutes in the cytosol are not equivalent to free diffusion of the solutes in the absence of cytosolic barriers.

II. SOLUTE AND SOLVENT FLUXES ARE DETERMINED BY BARRIERS AND PROPELLING FORCES

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1. Importance of Biomembrane Transport

cal expression for this gradient will be derived in Chapter 3, the reader's existing concept of total chemical potential should more than suffice for the present discussion. B. A Propelling Force is N e e d e d to P r o d u c e N e t Flux of a Solute or t h e Solvent in O n e Direction across a B i o m e m b r a n e Since propelling forces influence unidirectional flux, they also produce net flux when the propelling force is greater in one direction across the membrane than in the other direction. In the simplest case, a substance or ion migrates more rapidly along its total chemical potential gradient toward thermodynamic equilibrium than it moves in the reverse direction. As we discuss in several chapters, however (e.g., Chapters 4 to 7), this simple phenomenon does not account fully for the net transport catalyzed by most proteins. An exception appears to be the transport catalyzed by proteins, such as c~-hemolysin, that do not select among low-molecularweight solutes. All transport proteins that are substrate selective and saturable can be made to catalyze transport of a substrate against a total chemical potential gradient by coupling the transport to another source of free energy. C. The E n d e r g o n i c Migration of a Solute a g a i n s t Its Total Chemical Potential G r a d i e n t Can O c c u r O n l y W h e n It Is C o u p l e d to an E x e r g o n i c Process of G r e a t e r M a g n i t u d e Even uniporters (Section XI,G of Chapter 4) and channels (Section II,C of Chapter 7) may in some cases use the total chemical potential gradient of one substrate (or inhibitor) to generate a gradient of another. Such interconversions of gradients by uniporters and channels are, however, usually much less conspicuous and efficient than the propagation of one solute gradient into that of another by symporters and antiporters. The latter proteins couple the migration of one or more solutes to the co- or countermigration, respectively, of one or more other solutes. Hence, they are sometimes also termed cotransporters, countertransporters, exchange-transporters, or even secondary active transporters. The tightness of the coupling (i.e., the degree to which transport occurs only in the presence of all co- or countersubstrates) helps to determine how efficiently the free energy of one solute gradient is converted into that of another. When slippage or tunneling (i.e., uncoupled transport) is relatively frequent, the free energy transfer is relatively inefficient, whereas tightly coupled transport results

in the nearly complete conversion of the free energy in the gradient of one solute into that of another. Interestingly, transport in which coupling between co- (or counter-) substrates is not obligatory could lead to transport of a large amount of one cosubstrate relative to the other. This high ratio could be mistaken for the stoichiometry of comigration rather than the stoichiometry of cotransport of the substrates. 4 Such a high apparent stoichiometry of comigration of one substrate relative to the other would, however, actually reflect a high degree of uncoupling rather than the actual stoichiometry of comigration. Consequently, a total chemical potential gradient of the cosubstrate transported in greater amount would be dissipated without much transport of the other cosubstrate against its gradient. In contrast, the stoichiometry of cotransport of one cosubstrate relative to the other may be high, owing to the obligatory comigration of several ions or molecules of this first substrate to the transport of a single ion or molecule of a second kind. In this case, a total chemical potential gradient of the first cosubstrate across a membrane could produce a much steeper gradient of the second one, assuming only that a mechanism exists to maintain a steady-state gradient of the first cosubstrate (to be discussed further in Section IX,B of Chapter 3). Many solute gradients are maintained across biomembranes by coupling them to yet another source of free energy such as that realized from A T P hydrolysis. Conversely, transport along these gradients may drive A T P synthesis. When transport along a gradient normally coupled to A T P synthesis occurs without such coupling, however, additional free energy must be expended to maintain the gradient. In fact, when thermodynamically coupled processes are uncoupled for the purpose of generating thermal energy in mammals, uncoupled transport rather than uncoupled A T P hydrolysis results in thermogenesis. Rapid hydrolysis of A T P for thermogenesis might put at risk the numerous other cellular processes that rely on a well regulated A T P 4We define the stoichiometry of co- (or counter-) transport as the ratio of the number of ions or molecules of one substrate actually transported with a particular number of ions or molecules of the other substrate in the average transport cycle. This definition is contrasted here with our definition of the stoichiometry of co- (or counter-) migration, which is the number of ions or molecules of each species of substrate that are transported together in a single catalytic cycle of a transport protein. In the case of nonobligatory symport or antiport, the stoichiometry of comigration cannot be measured in every transport cycle since comigration does not occur in every cycle. Consequently, the stoichiometryof comigration may be difficult to determine experimentally, whereas the stoichiometry of cotransport can almost always be determined. Similarly, the stoichiometry of comigration may be difficult to determine experimentallywhen obligatory symport and antiport occur together, and different species of substrate have different probabilities of dissociating from the transport protein during its cycle (see Section III,B of Chapter 6 for further discussion).

Solute and Solvent Fluxes

supply. These processes are distributed throughout the cell, so the total volume of cytoplasm in which the ATP supply must be regulated is relatively large. In contrast, the ATP synthesis that is driven by a proton gradient in animals is restricted to the inner mitochondrial membrane. D. The Thermal Energy Released to Maintain a Solute Total Chemical Potential Gradient Provides Conspicuous Evidence of the Free Energy Content of the Gradient It is well established that an F-type ATPase catalyzes conversion of the free energy in the proton gradient across the inner mitochondrial membrane into the free energy realized in the phosphoric acid anhydride bonds of ATP (Chapter 5). The considerable free energy associated with this proton gradient becomes even more conspicuous in mammals when the proton gradient is disconnected from ATP synthesis by mitochondrial uncoupling proteins (UCPs). The thermal energy that is generated in opposing the action of UCPs serves both to warm the animal under cold stress and as a device to rid the animal of excess dietary free energy intake. While the first of these proteins to be discovered (UCP1) is expressed exclusively in mitochondria of brown adipose tissue (Ricquier et al., 1991), a second protein (UCP2) is widely distributed in the tissues of mammals including humans (Harper, 1997; Wolf, 1997). Twenty to 40% of mammalian mitochondrial oxygen consumption is needed to support the proton transport catalyzed by proteins such as UCP2 (Harper, 1997; Rolf and Brown, 1997). UCPs catalyze uncoupled H + transport across the inner mitochondrial membrane by a process that is distinct from the H + transport catalyzed by F-type ATPases. Hence, UCPs do not act to uncouple H + transport from ATP synthesis by F-type ATPases. Rather, they compete with F-type ATPases to transport protons and thus reduce the quantity of protons that could otherwise be used for ATP synthesis by 20 to 40%. The proton transport catalyzed by UCPs is associated with transport of a variety of inorganic and organic anions (Garlid, 1990), and physiologically important ones appear to be ionized fatty acids (Garlid et al., 1996; Jezek et al., 1997). Proton transport appears, however, not actually to be coupled to mediated fatty acid transport. Rather, UCPs are believed to catalyze uniport of fatty acids and other anions (Garlid, 1990). In the case of fatty acids, uniport of their anionic form out of mitochondria could be followed by their uncatalyzed migration into mitochondria in association with protons (Gar-

5

lid et aL, 1996; Jezek et aL, 1997). The latter migration of uncharged fatty acid molecules across the membrane may be relatively rapid, whereas transport of fatty acids in their normally anionic form appears always to be transport protein mediated (see Section VII of Chapter 4). Consequently, UCPs appear to catalyze uncoupled proton transport indirectly by enabling fatty acids to behave as cycling protonophores (Skulachev, 1991; Garlid et aL, 1996; Wojtczak et aL, 1998) (Fig. 1.3). The presence of possible proton-conducting groups on the side chains of some functionally important amino acid residues in UCP1 has, however, militated against universal acceptance of this protonophore theory (e.g., Bienengraeber et aL, 1998). Moreover, other investigators have concluded that fatty acids do not increase the rate of proton transport as a result of their own transport by UCPs (Gonzalez-Barroso et al., 1998). Regardless of the mechanism of proton transport owing to UCPs, the thermal energy generated in opposing the action of UCPs exposes the free energy content of proton gradients. Interestingly, UCPs are homologous to several other mitochondrial transport proteins including the ATp4-/ ADP 3- and H2POa-/OH- antiporters (Aquila et al., 1987; Klingenberg, 1990). Although the phosphate transporter was originally believed to catalyze H2PO4-/ H § cotransport, more recent evidence indicates that it

O

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Mitochondrial Matrix

FIGURE 1.3 Scheme showing how fatty acids may act as cycling protonophores to facilitate proton transport across the inner mitochondrial membrane. In this model it is proposed (Skulachev, 1991; Garlid et al., 1996) that uncoupling proteins (UCPs) catalyze uniport of fatty acid anions out of the mitochondrial matrix. The fatty acid anions are proposed, then, to associate with protons at the outer surface of the inner mitochondrial membrane, owing to the relatively high concentration of protons there. The undissociated fatty acids migrate relatively rapidly across the lipid bilayer without the help of a biocatalyst, whereas the fatty acid anions require a transport protein (in this case a UCP) to catalyze their migration. Once inside the mitochondrial matrix, the fatty acids dissociate from protons, owing to the relatively low proton concentration. While other authors (Skulachev, 1991; Garlid et al., 1996) show the UCP-catalyzed transport of the anionic forms of fatty acids as a "flippase" (to be discussed in Chapter 2), the actual mechanism by which these forms of fatty acids may migrate across the membrane via UCPs remains to be determined.

6

1. Importance of Biomembrane Transport

catalyzes H2PO4-/OH- exchange (Stappen and Kr~imer, 1994). Extrusion of O H - would of course accomplish the same end as H + uptake. Hence, although UCPs do not appear to catalyze H + transport directly, they could conceivably be modified effectively to do so. Such mutability of both substrate selectivity and the combinations of co- and countersubstrates received by transport proteins appears to have resulted frequently in the evolution of important new physiological functions in many families of such proteins (see the summary of families that contain homologous members in Chapter 8).

ies is found in the E A A T / A S C family of amino acid transporters (Chapter 6). Proteins in the E A A T subfamily catalyze the concentrative uptake of anionic amino acids in neurons and other tissues at the expense of both the Na+and K § gradients across the plasma membrane (Fig. 1.4). Consequently, many of these proteins help to reduce the glutamate concentration in the vicinity of glutamate receptors in the central nervous system to a level below the values of the dissociation constants of these receptors. Interestingly, however, several members of the E A A T subfamily appear to have evolved to express primarily a related but quite different additional function of the proteins. The latter members of the E A A T subfamily are postsynaptic proteins that catalyze mainly glutamate-stimulated C1- transport (Fig. 1.4) and relatively little glutamate transport (e.g., Fairman et al., 1995; Sonders and Amara, 1996; Arriza et al., 1997). Hence, some E A A T proteins may have a central rather than an auxiliary role in signal transduction (Sonders and Amara, 1996). Likewise, members of the ASC subfamily apparently evolved in yet another context to catalyze Na+-dependent exchange of zwitterionic amino acids (Fig. 1.4). Each of these different transport functions is important to the ability of different cells to perform their specialized functions. Another way in which changes in substrate selectivity may contribute to the evolution of important new functions among related proteins is for the stoichiometry of transport but not the substrate species themselves to change. We discuss in Chapter 5 the importance of

E. M e d i a t e d Transport Is Substrate Selective Biomembranes function as barriers to form compartments and consequently to organize metabolism among tissues and organs as well as among subcellular organelles. These functions of various biomembranes also depend, however, on the different substrate selectivities of their transport proteins. Hence, for example, the improper sorting of the homologous H+K +- and Na+K +selective ATPases to the basolateral and apical membranes, respectively, of an acid-secreting epithelium (instead of the other way around) would have disastrous consequences for the organism. In these cases, protons would be secreted inappropriately into interstitial spaces, whereas Na § would be extruded incorrectly into the lumens of pertinent organs such as the stomach (see also Section II,B,5 of Chapter 5). Another example of the importance of the evolution of transport proteins with different substrate selectivit-

CI-

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C

_

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FIGURE 1.4 Schemeto emphasize the various transport functions and relative substrate selectivities of different members of the EAAT/ASC protein family. The sizes of the abbreviations of the substrates are meant to indicate the relative amounts of transport by each group of transport proteins. EAAT1 to EAAT3 catalyze concentrative uptake of anionic amino acids such as glutamate (Glu-) to a greater extent than they catalyze glutamate-stimulated C1- transport as channels. In contrast, EAAT4 and EAAT5 catalyze more glutamate-stimulated C1- transport than they do glutamate uptake. Members of the other subfamilyof transport proteins in the EAAT/ASC family (i.e., the ASC subfamily) catalyze Na+-dependent exchange of zwitterionic amino acids (AA -~) as well as channel-like C1- transport. The relative amounts of these transport activities remain, however, to be determined for different ASC proteins. For this reason, they are shown approximately to be equal for ASC proteins, although such may not be the case for different members of this subfamily.

7

Biomembrane Transport in Context

V-type ATPases in acidification of intracellular compartments at the expense of ATP hydrolysis. On the other hand, ATP synthesis is usually accomplished in oxidative tissues by the related F-type ATPases in mitochondria. Part of the explanation of how these two homologous families of ATPases evolved to perform opposite functions is that the stoichiometry of H + ions transported per ATP molecule hydrolyzed or synthesized is lower by one or two protons in V-type than in F-type ATPases. For this reason, a much larger and usually unattained proton gradient would be required for V-type ATPases to carry out net ATP synthesis. Vtype ATPases may also catalyze some uncoupled proton transport in the reverse direction out of intracellular compartments (i.e., they may leak), which would help to make ATP hydrolysis by the enzyme irreversible. Similarly, F-type ATPases may also catalyze proton transport in either direction. Unlike V-type ATPases, however, proton transport remains coupled to ATP synthesis or hydrolysis in F-type ATPases. While reversal of the function of F-type ATPases is unusual in mitochondria, extrusion of protons at the expense of ATP hydrolysis under anaerobic conditions is a normal adaption of F-type ATPases is some bacteria. Hence, we see that differences in the reversibility of solute migration as well as in substrate selectivity combine with barrier action to determine a variety of functions of biomembranes in different cells and organelles. F. Reversibility of Solute Transport That solute transport must be reversible for optimum physiological functioning is no better exemplified than in the case of C1-/HCO3- exchange in the red blood cell (Chapter 6). The anion exchanger (AE1) catalyzes release of HCO3- from erythrocytes in exchange for C1in capillaries of respiring tissues (Fig. 1.5). The HCO3is produced in red blood cells by carbonic anhydrase, owing to their uptake of the CO2 produced in nearby cells. This process helps blood carry more total CO2 (CO2 plus HCO3-) than would otherwise be possible. A greater capacity for bulk flow of CO2 from peripheral tissues to the lungs appears to be particularly important during aerobic exercise. Anion exchange must be fully reversible, however, in order for erythrocytes to take up most efficiently HCO3- in exchange for C1- and convert it to CO2 for excretion by the lungs (Fig. 1.5). Similarly, we shall see that reversible solute transport into and out of cells is essential for normal nutrient flows among tissues and organs in other cases, such as in the fed and fasted states. These nutrient flows among tissues and organs will be discussed in Chapter 10 using amino acids as examples.

Lungsl

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FIGURE 1.5 Schemeto show why reversal of C1-/HCO3- exchange in erythrocytesis needed to help to carry CO2 from peripheral tissues to the lungs. Solid arrows show the net migration of the carbon in CO2 in the blood capillaries of peripheral tissues, whereas the dashed arrows show the net migration of the carbon within capillaries of lungs. Abbreviation: AE1, anion exchanger 1.

Transport may also be made reversible by using different transport processes to catalyze solute migration in one direction or the other across biomembranes. For example, Na+K+ATPase catalyzes K + uptake and Na + extrusion against their total chemical potential gradients across the plasma membrane of most animal cells. The reverse net transport of both cations is catalyzed by Na + and K + channels. In general, channels allow substrates to migrate along their total chemical potential gradients. Transport of Na + and K + in both directions across the membrane helps to produce and dissipate transmembrane electrical potentials in excitable cells, and it results in other types of cellular work, such as regulatory cellular volume increases and decreases. The activities of the transport processes themselves must also be regulated in these cases to produce physiologically desirable results (see Chapter 9 for further discussion of transport regulation).

III. BIOMEMBRANE TRANSPORT IN CONTEXT Most students of biomembrane transport eventually consider their findings in the broader context of the environments of cells in situ (e.g., see Chapter 10). Nevertheless, relatively few scientists actually study transport into the cells of perfused tissues, organs, and even whole multicellular organisms. Isolation and characterization of transport activities in a given cell type is particularly difficult in the latter context. In order to isolate and study a single biomembrane transport activity for a substrate, one frequently needs precisely to control

8

1. Importance of Biomembrane Transport

the concentrations of inhibitors of other processes that compete with the activity to transport the substrate. This control is difficult to achieve in intact organs where the inhibitors and substrate may need to migrate relatively long distances to reach the cell membranes. Moreover, the uptake measured in whole organs may represent a composite of several cell types only one of which is the type of interest. For this reason, investigators frequently chose first to isolate cells or even biomembrane vesicles from the cells and then to characterize their transport in a controlled environment in vitro. The possible physiological significance of the transport processes is then usually discussed in the context of what is known about substrate concentrations in extracellular fluids in vivo. Also considered is how the transport is influenced by signaling molecules and other signaling processes such as changes in membrane electrical potential often measured in isolated cells. In light of these attempts to understand transport in its physiological context, surprisingly little attention has been paid so far to the effects on transport of the physical environment of cells in situ. For example, what immediate effects on biomembrane transport are introduced during isolation and purification of a particular cell type or their biomembranes? Do the characteristics of transport change immediately in some or all types of cells when they are isolated? Or do these characteristics remain relatively stable regardless of what may need to be done to the surrounding environment in order to isolate the cells of interest for further investigation? A. H o w Much Does the Cellular Environment in Vivo Influence B i o m e m b r a n e Transport? A partial answer to the preceding questions comes from the study of amino acid transport in early mouse embryos. Cleavage-stage conceptuses and blastocysts lie in close association with the reproductive tract during development. They are, however, quickly and easily separated from the reproductive tract for a period of about 5 days after conception, at which time blastocysts implant in the uterus. When blastocysts are removed from the uterus a day before implantation, their plasma membrane system B ~ transport activity remains constant for several hours in culture (Van Winkle and Campione, 1987; Van Winkle et al. 1990d). In contrast, blastocysts removed from the uterus a few hours prior to implantation experience a dramatic increase in their system B ~ X-AG and/~-transport activities, whereas system b+2 decreases in activity (e.g., Fig. 1.6). The activities of these transport systems change within a few minutes after embryo isolation, and the changes are complete within about half an hour (Van Winkle and Campione, 1987).

FIGURE 1.6 Changes in the transport activities of systems B ~ and b+2 but not system b ~ upon removal of blastocysts from the uterus just prior to implantation (A). In contrast, no change in system B ~ transport activity is observed when blastocysts are removed from the uterus 24 hr before implantation (B). Changes in activity are statistically significant when they are marked with a double asterisk (p < 0.01) (data from Van Winkle et al., 1990d).

Interestingly, the changes also occur on the same time course in blastocysts within the uterus when it is simply massaged gently with a blunt instrument, whereas no such changes in transport system B ~ activity are observed when the uterus is massaged 24 hr prior to blastocyst implantation. Since transport cannot be measured easily in preimplantation embryos within the reproductive tract, it is unclear how the activities of their transport systems may change when they are removed form the uterus relative to their activities in this initial condition. There is, however, little doubt that some of the activities do change as a result of isolation at least in blastocysts nearing implantation. While the possible physiological implications of these changes in blastocysts nearing implantation is of interest primarily to those of us who study early development, the fact that the changes occur at all should evoke broader interest. We currently study transport primarily by isolating the pertinent cells, biomembranes, or even the transport proteins themselves, and the transport proteins may be expressed in other cells or in proteoliposomes. While such studies produce new insights into the functions of transport proteins, the proteins may not function as they normally do in vivo. For this reason, it is anticipated that new investigations will more frequently involve whole tissues, organs and even intact organisms. Numerous examples of such studies are discussed by Taylor and associates in Chapter 10 of this volume. Here we discuss a few examples of the sometimes surprising place of biomembrane transport in the context of multicellular organisms.

Biomembrane Transport in Context

9

B. Transepitheliai Nutrient Transport May Not Be Equivalent Simply to a Composite of All Pertinent Biomembrane Transport Processes for the Nutrient Most of us attribute a central importance to nutrient transporters in the placenta since in few cases is a need for relatively massive transfer of nutrients so conspicuous. While the importance of biomembrane transport to normal placental functioning is difficult to deny, findings with intact animals have shown that the mechanism of transfer of nutrients across the placental trophoblast can be much more complex than anticipated. The simplest way for organic and inorganic solutes to traverse the biomembrane barriers between mother and fetus appears to be for the nutrients to be taken up against their gradients by transporters in the microvillous membrane, for them to then diffuse across the cytosol of the placental trophoblast, and finally for them to migrate out of the cells via transport proteins in the basal membrane. Consequently, much study has focused on identifying and characterizing transport systems and proteins in the two membranes and attempting to envision how the transport processes could be coordinated to catalyze net flux toward the fetus. When this flux is studied in intact animals, however, we quickly learn that we must understand how biomembrane transport fits into a much broader biochemical context, if we are to understand how vectorial nutrient transfer actually Occurs.

For example, only about 38% of the leucine, 11% of the glycine, and none of the serine appearing in the blood plasma of fetal sheep gets there through direct transfer from mother to fetus across the placenta. Rather, amino acids released during placental and fetal protein degradation and nonessential amino acids synthesized in placental and fetal tissues provide most of the amino acids appearing in fetal blood (Geddie et al., 1996). Glycine is synthesized primarily from serine in the placenta for transfer to the fetal circulation (Fig. 1.7), whereas serine is synthesized from glycine and other substrates in the fetal liver (Thureen et aL, 1995). A little over half of the leucine released from the placenta to the fetus appears to arise from placental protein degradation (Ross et aL, 1996). The possibility that the sheep placental trophoblast may also take up and degrade maternal plasma proteins apparently has not been ruled out formally, although the trophoblast in the chorioallantoic placenta appears to have a relatively low endocytic capacity at least in the rodent (Pratten and Lloyd, 1997). In contrast, epithelial cells of the rodent visceral yolk sac placenta display prominent endocytosis and could

FIGURE 1.7 Glycine and serine transport and metabolism in the ovine placenta. While some glycine is transferred directly to the fetus from the mother, most of the glycine appearing in fetal blood plasma from the placenta is produced from serine. Two separate serine pools in the placenta appear to be derived from maternal and fetal sources, and both of these pools are used to produce glycine (adapted from Geddie et aL, 1996 with permission from W. B. Saunders Company Ltd.)

conceivably take up proteins in order to supply amino acids to the embryo/fetus beginning just after implantation and continuing until parturition. 5 Uptake of nutrients by this route clearly permits postimplantation rat embryos to grow at their normally rapid rate even in culture when the epithelium is in direct contact with proteins in the medium (e.g., Beckman et al., 1990, 1991, 1994, 1996, and 1997). It is less clear, however, whether plasma proteins actually reach the yolk sac epithelium in large enough quantities in vivo to contribute significantly to the amino acids reaching the embryo/fetus. If plasma proteins are the principal source of amino acids for the embryo/fetus in vivo, however, then the mechanism of such nutrition in the rodent is significantly more complex than previously anticipated. From about the time of implantation until organogenesis is nearly complete, the epithelium of the yolk sac placenta would take up maternal plasma proteins, degrade them in lysosomes, and then release the resultant amino acids to the embryo. For the amino acids to reach the embryo they would first be transported out of the lysosomes and epithelial cells via amino acid transport systems apparently expressed selectively in the lysosomal and plasma membranes (e.g., Pisoni and Schneider, 1992). Even after the chorioallantoic placenta becomes functional during the latter half of gesta5Two prominent placentas (i.e., the yolk sac and the chorioallantoic placentas) appear to transfer nutrients to the embryo/fetus of several rodent species during a major portion of their gestation, whereas most other eutherian species transfer nutrients primarily or exclusively via the chorioallantoic placenta.

|0

1. Importance of Biomembrane Transport

tion in the rodent, the major source of amino acids to the fetus may still be via the yolk sac placenta (Beckman et al., 1994, 1997). If such is the case, then amino acid transport via the chorioallantoic placenta may not serve primarily for net transport of amino acids to the fetus. Rather, this organ may be viewed better as regulating amino acid levels in the fetus. It may even serve for the net flux of some amino acids from the fetal to the maternal circulation depending on the quantity of amino acids supplied to the fetus via the yolk sac placenta, the maturity of fetal organs, and the nutritional requirements of the fetus. Regardless of what conclusions are finally drawn concerning the role of biomembrane transport in the nutrition of fetuses of various species, two fundamental tenets emerge here. First, biomembrane transport is critical to such nutrition as well as to other processes that are required in order to supply nutrients to the tissues and organs of multicellular plants and animals. Nevertheless, the role of biomembrane transport in these processes may not be as simple or as direct as we first envision. Second, the study of biomembrane transport leads us naturally away from excess reductionism, thus helping to insure that its function will eventually be understood in its physiological context. As we have seen, the ways in which biomembrane transport contributes to the normal functioning of multicellular organisms may not be obvious, although its contribution is obviously real. Biocatalysts usually are needed in order for transport to proceed at rates compatible with life. The discovery and characterization of these transport proteins and systems has thus helped to explain the otherwise unexpectedly rapid rates of migration of some hydrophilic substances across biomembranes.

IV. SUMMARY We have seen that the study of biomembrane transport requires us to consider its biophysics and physical chemistry as well as its biology. Moreover, biomembrane transport is central to the functioning of all multicellular organisms regardless of whether it is considered at the subcellular, tissue, or systemic levels of their organization. Similarly, the study of biomembrane transport is as legitimate a component of investigations into mechanisms of development and differentiation as it is into the functioning of fully formed tissues and organs. Hence, there is scarcely a biological journal or a subsection within such journals from the biophysical to the evolutionary levels of investigation that does not contain articles on the subject of biomembrane transport. Such was, however,

not necessarily the case before biomembrane transport became a fully legitimate field of investigation in these academic disciplines. Partly as a result of the establishment of provisional boundaries to the various academic disciplines of biological sciences in the middle half of this century, some students of biomembrane transport saw the opportunity to cross these artificial subdivisions in highly productive ways. Thanks to the efforts of these pioneers, most modern scientists view their own research as pertinent to a wide range of biological disciplines. Research groups that are focused on certain aspects of biology may of course develop within or between institutions as a consequence of common interests. Most of the time, however, these groups are composed of individuals with broad training, only some of which may have been considered part of the academic discipline historically defined for the department in which they happen to find themselves. The recent dissolution of the Physiology Study Section of the U.S. National Institutes of Health (Ehrenfeld, 1998) is but one consequence of this evolution toward multidisciplinary scientific investigations. By analogy, numerous proteins have evolved over billions of years to catalyze transport of solutes and the solvent across the barriers formed by biomembranes. As the demand for different types of transport increased owing to evolution of more numerous as well as more complex species, the needed transport processes also of course evolved (e.g., see Chapter 8). Similarly, as our investigations have led us to a fuller understanding of biology, the disciplinary boundaries that once helped us to define ourselves now help us more easily to recognize various aspects of biology to which our unfolding work may apply. Viewed as part of our cultural evolution, academic disciplines are destined to become extinct because new paradigms better fitted to the scientific milieu are replacing them. One wonders what scientific approach will evolve to render current interdisciplinary and multidisciplinary approaches obsolete. We continue our exposition of the central position of biomembrane transport in biology in Chapter 2. There we consider the physical and chemical natures of biomembrane barriers, their origins, and their fates. It is impossible fully to understand how various transport proteins form, propagate, or dissipate solute and solvent gradients (Chapters 3 to 7) without understanding the nature of the barriers across which they catalyze transport. Moreover the contributions of these biomembrane transport processes to the physiological functioning of cells and organisms is rooted in the physical and chemical nature of biomembrane barriers and how the barriers may change in various physiological and pathophysio-

Summary logical conditions. Hence, an understanding of the physical and chemical nature of membranes is needed to understand both cellular physiology and the physiology of whole multicellular organisms such as those described in Chapter 10. Similarly, the regulatory mechanisms

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needed to coordinate biomembrane transport in cells and in multicellular organisms (Chapter 9) can only be understood fully if one understands the nature of the barrier for which regulated transport is needed in the first place.

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2 I Biomembrane Composition, Structure, and Turnover

1. INTRODUCTION

process of unknown origin may actually be 20 times more rapid than the more obviously directed processes of endocytosis and exocytosis (Farge, 1995). Endocytosis and exocytosis depend on the cytoskeleton for movement of vesicles to and from the plasma membrane, and the cytoskeletal infrastructure influences the motion of membrane constituents. As for the plasma membrane, however, the function of the cytoskeleton should be viewed neither as passive nor simply structural.

For many years it was unclear how phospholipid bilayers only about 5 nm thick could nevertheless be strong enough to withstand the stresses on the plasma membranes of most cells. True, the sizes of most animal cells are small enough (i.e., --~20/zm in diameter) for adhesive forces between water molecules to maintain a more or less spherical cell shape inside a lipid bilayer surface. For this reason, plasma membranes might, as first approximations, be able to lie relatively passively as unreinforced thin lipid barriers at the surface's of cells. A few moments reflection on the requirements of the membranes of most cells in their natural environments leads, however, immediately away from any such notion of a placid existence for most cell membranes. For example, monocytes surrounded by thin membranes are greatly deformed as they migrate between vascular endothelial cells in response to injury or infection (Fig. 2.1). The endothelial cells, on the other hand, must withstand powerful hemodynamic sheer stresses on their lumenal surfaces that would occur at arterial branch points. In fact, when such stresses are excessive, as in hypertension, they seem to initiate or contribute to development of atherosclerosis (Fig. 2.1). While each cell type may have its own specific requirements for movement, reinforcement, and signaling, all eukarocytic cells benefit from the normally inconspicuous cytoskeletal components that support their membrane structures and functions (Fig. 2.2). Cell membranes also face continuous challenge to their integrity from within. An area of membrane about equal to the area of the entire cell surface turns over about every 30 min in many cells due to endocytosis and exocytosis. A less conspicuous vesiculation and refusion

!I. IS THE FLUID MOSAIC MODEL OF MEMBRANE STRUCTURE STILL ADEQUATE? A. The Lipid Bilayer Hypothesis The lipid bilayer is now well established as the fundamental structure of most biomembranes (Fig. 2.3A). Nevertheless, nonlamellar lipid structures also form important components of biomembranes. These structures and their influences on membrane function have been reviewed recently in a volume edited by Richard Epand (1998). For this reason and to conserve space, we focus principally on the lipid bilayer structure of membranes in this chapter. Many phospholipids will assume the bilayer structure spontaneously in membranelike structures known as liposomes under the right experimental conditions either alone or in combination with other lipids (Gregoriadis, 1993). More realistic "snapshots" of fully hydrated artificial phospholipid bilayers (e.g., Fig. 2.3B) have been produced recently through computer simulations (e.g., Jakobsson, 1997). The phospholipids that are present in biomembranes are highly amphipathic; they have phosphate-containing groups esterified to molecules (usually glycerol) that also have long-chain fatty acyl

13

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2. Biomembrane Composition, Structure and Turnover

FIGURE 2.1 Involvement of monocytes and endothelial cells in development of the fatty streak and, eventually, atherosclerosis. Note in particular the physical stresses that monocytes need to place on their own plasma membranes in order for them to pass between endothelial cells and the hemodynamic sheer stresses to which endothelial cells are exposed at arterial branch points. Such sheer stresses would, of course, increase as blood pressure increases.

groups (Fig. 2.3C). The phosphate-containing groups are electrically charged and, hence, highly hydrophilic, whereas the hydrocarbon side chains of the fatty acyl groups are quite hydrophobic. In the phospholipid bilayer, these hydrophobic side chains extend within each leaflet of the bilayer toward each other to form the hydrophobic interior of biomembranes (Fig. 2.3A). In contrast, the hydrophilic phosphate-containing groups seek positions in the bilayer between the hydrophobic interior and either of two external aqueous phases. One surface of the bilayer faces the cytosol of cells, whereas the other surface of the bilayer borders the lumen of

organelles, the interior of membrane vesicles, or the exterior of cells (Fig. 2.3D). The formation of bilayers is primarily an entropydriven process because water exists in an ice-like rather than a liquid state when the water is associated with the hydrocarbon side chains of the fatty acyl groups of the phospholipids. When the hydrocarbon side chains associate with each other in the interior of the lipid bilayer instead of with water, the water can become liquid rather than remain ice-like. It is this greater freedom of movement of water in the liquid state that drives formation of these so-called hydrophobic bonds or, more properly,

Fluid Mosaic Model of Membrane Structure

| 5

FIGURE 2.2 Cell in culture fixed and stained to expose proteins that form the filamentous cytoskeleton (adapted from Alberts et al., 1994, with permission from Garland Publishing, Inc.).

hydrophobic interactions. In the present case, these hydrophobic interactions refer to the sequestration of the hydrocarbon side chains in the interior of the lipid bilayer of biomembranes away from most of the water. Animal biomembranes also contain other lipids, such as cholesterol, in addition to phospholipids. Cholesterol is less amphipathic than are phospholipids because the hydrophilic portion of cholesterol is due primarily to its uncharged and relatively small hydroxyl group rather than to an electrically charged and much larger phosphate-containing group (Fig. 2.3C). For this reason, cholesterol appears in many instances not to be as confined as phospholipids are to one leaflet or the other of the membrane bilayer. It is this greater ability of cholesterol to flip from one leaflet to the other that is

believed by some investigators to permit cells to undergo rapid shape changes without causing one leaflet of their plasma membrane to "wrinkle" and the other leaflet to "gap" (Fig. 2.3E). Cholesterol also migrates from the outer to the inner leaflet of the platelet plasma membrane when these cells are activated apparently owing to migration of phosphatidylethanolamine in the reverse direction (Boesze-Battaglia and Schimmel, 1997). Although clearly valid, the lipid bilayer hypothesis remains an active field of investigation. For example, we are still attempting fully to understand the consequences of the asymmetric distribution of lipids across the bilayer (see below). In addition, we are only beginning to appreciate how the existence of most membrane

16

z. Biomembrane Composition, Structure and Turnover

lipid bilayers in a liquid-crystalline state (defined here as the state of transition between the wholly liquid and wholly crystalline phases) contributes to their function. 1 The complex composition and asymmetric distribution of lipids in the leaflets of the membrane bilayer probably contributes to the relatively wide range of temperatures over which the membrane "melts" (Fig. 2.4). The continuous transition of lipid in membranes between the liquid and crystalline states creates in membranes transient domains that undoubtedly influence not only the structure but also the function of membrane constituents. Because membrane constituents may be more concentrated in one type of domain than in the other, the influence of these transient domains on membrane function may be quite different from the effects on function of wholly liquid or wholly crystalline bilayers. For example, what is the consequence to, say, a glutamate transport protein molecule when it is present in a crystalline vs a liquid domain of different lipid compositions? Could existence in one or the other domain influence whether the protein functions in some instances as a glutamate transporter and in other instances as a C1- channel? (See Sections III and IV of Chapter 6 for further discussion of such multiple transport functions of these proteins.) In addition, many integral membrane proteins appear to be associated preferentially with either liquid or crystalline domains depending on the domains for which the protein molecule has greater affinity (Marsh, 1995). The sizes of crystalline domains are larger than are liquid ones, at least in artificial bilayers, apparently owing to a smaller number of nucleation sites in the former case (Sankaram et al., 1992). Such differences in the sizes of the liquid and crystalline domains as well as the preferential association of different proteins with one or the other of the domains likely influences the interactions among protein molecules in the bilayer. 1The terminology in the literature is somewhat unclear in regard to what is meant by the liquid-crystalline state of the lipid bilayer, perhaps because the existence of phase separation in the bilayer has only recently gained wider acceptance (e.g., Brown and London, 1997). Here we define the wholly solid, gel, or crystalline state as the state of the bilayer before it begins to melt, whereas the wholly liquid or fluid state is defined as the state of the bilayer after it has melted. At physiologically normal temperatures the membrane lipid bilayer exists between these two states in what is termed here the liquid-crystalline state. Phospholipids in the liquid-crystalline state of membranes may be viewed as highly ordered, as in a crystal, and yet highly mobile, as in a liquid. The same is to some extent also the case, however, for the lipid in bilayers just above or just below their melting temperatures. The lipid is simply more ordered in the crystalline state and more mobile in the liquid state. Hence, what is perhaps more important to appreciate about the lipid in biomembranes is that the order and

mobility of the lipid varies with location in the membrane; some such transient domains appear to be wholly crystalline while others appear to be fully liquid. The possible importance of melting and freezing of these transient domains to the physiologicalfunctioning of biomembranes is discussed in this and subsequent chapters.

FIGURE 2.3 Lipid bilayer structure ofbiomembranes. (A) The lipid bilayer in which circles represent charged phosphoryl-containing portions of the lipid molecules and hydrophobic hydrocarbon chains extend toward the center of the bilayer. (B) Computer simulation of a fully hydrated artificial lipid bilayer in which the phosphoryl-containing portions of the lipid molecules can be seen to mix on a molecular level with water molecules. Hydrogen atoms in water molecules are shown in white, and the oxygen atoms in water are somewhat lighter in color than atoms in phospholipid head group. The sizes of the water molecules and phospholipid head group atoms have been reduced in order to see into the structure (adapted from Chiu et aL, 1995, with permission from The Biophysical Society). (C) Details of the structures of several membrane lipid molecules as they would be aligned in one layer of the bilayer. The zigzag lines represent hydrocarbon chains of various lengths (adapted from Finean and Miche 11,1981, with permission from Else vier Science). (D) One surface of the bilayer always faces the cytosol, whereas the other surface may face the lumen of intracellular organelles

(except mitochondria and peroxisomes), the inside of membranebound vesiclesorthe extracellular environment (adapted from van Helvoort and van Meer, 1995,with permission from Elsevier Science). (E) Cholesterol molecules in the bilayer may flip from one layer to the other when a shape change is needed in the membrane, thus helping to prevent formation of gaps and wrinkles in either layer.

It might, at first, seem to be a simple matter to study the effects of crystalline and liquid domains in membranes on their function by studying the membranes below or above their melting temperatures. It can be shown, however, that the fluid (i.e., entirely liquid) and gel (i.e., entirely crystalline) states of membrane lipids are probably not equivalent to liquid and crystalline lipid domains in membranes in phase transition. For example, triiodothyronine decreases the freedom of lipid motion in artificial membranes in the fluid state but increases it in the liquid-crystalline and gel states (Farias et aL, 1995). A possible explanation for this phenomenon is that the hormone is partitioned mainly to the crystalline phase of the liquid-crystalline state, whereas no such partitioning is possible in the fluid state. If, however, partitioning occurs in the liquid-crystalline state but it cannot by definition occur in the fluid or gel states, then this phenomenon becomes itself proof that the transient crystalline and liquid lipid domains in membranes in phase transition are unlikely to have a composition and structure that are identical to those of membranes

Fluid Mosaic Model of Membrane Structure

FIGURE 2.3

wholly in the gel or fluid state. If triiodothyronine is partitioned to the crystalline domains then it is not evenly distributed in bilayers in the liquid-crystalline state, whereas it would be evenly distributed in the wholly fluid or wholly gel states. Preferential association of some lipid species with particular integral membrane proteins may also result in partitioning of the lipid into a liquid or crystalline domain since the lipid species attracted to a protein molecule may itself be more likely to be in one or the other of these states (Marsh, 1995). The lipid compositions of liquid and crystalline domains in mem-

|7

(Continued)

branes can be studied using X-ray microanalysis (Hui, 1995), and electron microscopy helps to determine the geometries of the domains (e.g., Fig. 2.5). As just discussed, however, it becomes a difficult problem indeed to study the effects of different domains on the functions of proteins that intrude into the lipid bilayer. In fact, it is even conceivable that some proteins require the crystalline or liquid lipid domain in which they exist at a given moment to "melt" or "freeze," respectively, in order for the proteins to complete their functions (e.g., see Section VI of Chapter 3 regarding the functioning of Na+K+ATPase).

|8

2. Biomembrane Composition, Structure and Turnover

/

C

> > > > >

OH

c=o 9.o O O CH2-CH 2 CH2

C.Oo C.Oo

~.o ~.o O 9 CH=-CH 2 CH2

(~"O ~ C(=O (~ 9 (~=O 9 'C=O ,O !~O ~C= CH2- CH2 CI~ CH2-cH 2 CH2- CH2 Cholesterol CH2 HO ,CH2 ~H2 H C~ ~H2 C,H= ,CH= ,CH~ 0 9. O ,O 9 9 O=P-O O=P-O" O~P-O" o.,P-O o'-f=oo=,P-O" O-P-O'o H O O H ~ ~ HOOH2-~~OH~ ,o o 0 ~H2 H C]~2 /CH2 CH~ c.o ,c CH~ H~OO'O ~'O'O'O'O'O'O H 'O' 0 CH C,H2 CH O O CH3 (i; nln3 NH3* CH~",I~-CH3 CH~'N(~i)~CH3 6H HOCH " .0 CH3 CH3 Diphosphaditylglycerol ( DPG ) P PhosphatidylSphingomyelin Galactosyl O" ",r-NHC~, ( SM ) ceramide HOCH~o ~ CH3 inositol( PI ) serine( PS ) ethanolamine choline ( PC ) ( cerebroside) 0 (PE) !

!

!

.o o. I

!

o !

~--~'%L,=,

+

HOCH~o ~OOH Monosialoganglioside ( GMj )

Fluid Mosaic Model of Membrane Structure

FIGURE 2.4

19

Phase transition of lipids in the membranes of living

Acholeplasma laidlawii cells (circles) and in the membranes isolated

FIGURE 2.3 (Continued)

from these cells (triangles). According to the terminology used here, the bilayers are in the gel state at temperatures along the lower plateaus and they are in the fluid state at temperatures along the upper ones. At temperatures between the plateaus the bilayers are in the liquid-crystalline state. The broad range of temperatures over which the membrane "melts" and, hence, at which both liquid and crystalline lipid domains are present appears to be due in large part to the presence of integral membrane proteins in the bilayer and that these proteins associate preferentially with some lipid species (e.g., Marsh, 1995). Asymmetric distribution of lipids across the bilayer and heterogeneity of fatty acid residues that are saturated, monounsaturated, or polyunsaturated also probably contributes to the characteristics of the membrane lipid phase transition (adapted from Mantsch and McElhaney, 1991, with permission from Elsevier Science).

B. Protein Intrusion into the Bilayer The protein content of biomembranes varies from about 20% to more than 70% depending on the membrane. Proteins can also be associated with membranes peripherally as well as being integral components of them (Fig. 2.6). Biomembrane transport is generally believed to be catalyzed by integral membrane proteins, although their activities may be profoundly influenced by peripheral proteins. Integral proteins have one or more membrane-spanning segments. Most such segments are believed to be c~-helices of about 21 consecutive mainly hydrophobic residues oriented more or less perpendicularly to the plane of the membrane (von Heijne, 1994). Some of the helices may, however, be tilted, and they may even be parallel to the membrane in some cases (Persson and Argos, 1996). In addition, B-barrel components of some integral membrane proteins have been proposed to traverse the plasma membranes of eukaryotic cells (Fischbarg and Vera, 1995) as well as the outer membranes of Gram-negative bacteria and mitochondria (see summary of outer membrane porins in Chapter 8 of this volume). While the trans-

membrane component of the GLUT1 glucose transport protein is shown to be all /3-barrel in Fig. 2.7, more recent data are consistent with the possibility that it is composed of a mixed structure (Ducarme et al., 1996). In this mixed model, 10 c~-helices and 4/3-strands of GLUT1 are proposed to traverse the plasma membrane. The tertiary and even quaternary structures of several integral membrane proteins have also been studied in some detail. For example, the aquaporin-1 (CHIP28) monomer, a member of the protein family that forms water channels in numerous epithelial and nonepithelial tissues (Verkman et al., 1996), has been observed using cryoelectron crystallography to form tetrameres of four water channel pathways through the plasma membrane (Fig. 2.8). Each monomer forms a water channel apparently surrounded by six transmembrane c~-helices. In the case of acetylcholine-gated inorganic ion channels, the channels have even been imaged in different closed and open conformations (Unwin, 1995). In spite of these impressive advances, the structures of most integral membrane proteins are still only slowly being described in detail. Moreover, while our ability both to predict

20

z. Biomembrane Composition, Structure and Turnover

FIGURE 2.5 Freeze-fracture electron micrograph of one face of a phospholipid bilayer. Crystalline and liquid domains appear to be associated with characteristic surface undulations that help to distinguish them (Bar = 500 nm) (adapted from Hui, 1995, with permission from Taylor & Francis, London, UK).

topologies of membrane transport proteins and to test these predictions is improving (e.g., Jones et al., 1996; Persson and Argos, 1996), the detailed biochemical and biophysical mechanisms by which the proteins catalyze biomembrane transport, and the mechanisms of bioenergetic coupling of transport processes to each other or

to chemical change, remain, in most instances, very active areas of research. In spite of our still incomplete understanding of these transport mechanisms, many investigators assume some fundamental knowledge of the laws that govern them. For example, although transport proteins have asymmetric orientations in biomembranes, most investigators believe that the proteins catalyze thermodynamically symmetric transport in instances where the transport is not coupled to a conspicuous source of free energy. However, such may not always be the case, as discussed in several of the following chapters (e.g., Section II,C,2 of Chapter 7). C. B i o m e m b r a n e Structure Is A s y m m e t r i c as Well as H e t e r o g e n o u s 1. Proteins

FIGURE 2.6 Fluid mosaic model of biomembrane structure. Both integral and peripheral protein molecules are associated with the lipid bilayer, and the protein as well as the lipid molecules themselves diffuse laterally in the bilayer. In addition, the lipids near proteins are shown to be slightly disturbed relative to the more regular order in the rest of the bilayer. Actually, the lipid should be composed of more ordered "crystalline" domains and less ordered "liquid" domains, and some proteins should be shown as increasing order while

others decrease it (see text).

The mechanism of insertion of protein molecules into eukaroytic biomembranes appears to insure that each copy of a particular protein will have the same asymmetric orientation in a membrane (see also Section IV,B below). Most integral membrane proteins are inserted into membranes in the endoplasmic reticulum by a process that involves amino acid residue signaling sequences (Fig. 2.9). Although much is known about the topological information in the protein to be inserted, we are still learning how the cell decodes this informa-

Fluid Mosaic Model of Membrane Structure

2 |

2. Lipids

FIGURE 2.7 Possible structure of the GLUT1 glucose transport protein molecule. Note the considerable B-barrel structure (arrow) that has been proposed to span the plasma membrane. This protein appears to catalyze the transport of water as well as of glucose (adapted from Fischbarg and Vera, 1995, with permission from the American Physiological Society).

tion (von Heijne, 1994). Moreover, the number of transmembrane segments may change after initial insertion of proteins into the membrane as is the case for aquaporin-1 (CHIP28) monomers during trafficking from the endoplasmic reticulum to the plasma membrane (Verkman et aL, 1996). Hence some membrane transport proteins may undergo relatively large conformational changes during processing and even while functioning. (See detailed discussion of specific examples of transport protein function in Chapters 5 to 7.) Nevertheless, most transport and other integral membrane proteins likely retain their asymmetric orientations after insertion into biomembranes.

While integral membrane proteins have asymmetric orientations, lipids have asymmetric concentrations across biomembranes. For example, among the four most abundant categories of phospholipids in plasma membranes, the anionic one (phosphatidylserine) and a zwitterionic one (phosphatidylethanolamine) are usually more concentrated in the inner than in the outer leaflet of the bilayer, whereas the converse is true for the other zwitterionic phospholipids (sphingomyelin and phosphatidylcholine) (Table 2.1). More rapid movement of phosphatidylserine and phosphatidylethanolamine from the outer to the inner leaflet than the reverse is catalyzed by a membrane-bound MgZ+ATPase (Zachowski, 1993; Auland et al., 1994). Free energy is required to move these phospholipids against their concentration gradients and, in the case of the anionic phosphatidylserine, against the inside negative membrane electrical potential. Similarly, the outward movement of phosphatidylcholine is two to three times more rapid than inward migration (Zachowski, 1993), and stimulation of this difference by cytosolic ATP indicates that the outward transport may also be catalyzed by an ATPase. In contrast, the location of sphingomyelin in the outer leaflet appears to result from its synthesis in the lumen of the cis-Golgi through transfer of a phosphocholine residue from phosphatidylcholine (leaving diacylglycerol) to ceramide (see also Section IV,C below). Since little or no movement of sphingomyelin from the outer to the inner leaflet has been observed in healthy cells, this phospholipid appears to remain in the outer leaflet after it is synthesized. Although more sphingomyelin may be synthesized from ceramide and phosphatidylcholine in recycling endosomes than in the cis-Golgi in some cells (Fig. 2.10), its exclusive synthesis in the outer leaflet still appears to account for its asymmetric distribution in the plasma membrane. The asymmetric distribution of lipids across the plasma membrane is scrambled by several normal as well as artificial processes. Cellular activation by a variety of stimuli is associated with an increase in the cytosolic free C a 2+ concentration. Since C a 2+ in the cytosol inhibits MgZ+ATPase, it has been proposed that inhibition of this enzyme also leads to the increase in the concentrations of phosphatidylserine and phosphatidylethanolamine in the outer leaflet of platelet cell membranes during activation. Inhibition of MgZ+ATPase by N-ethylmaleimide does not, however, result in redistribution of phosphatidylserine to the outer leaflet of the platelet plasma membrane (Bas~e et aL, 1993). Hence, scrambling of lipid asymmetry during cell activation may involve more than simply inhibition of

22

2. Biomembrane Composition, Structure and Turnover

FIGURE 2.8 Projection structure of the aquaporin-1 channel-forming integral membrane protein molecule of 28 kDa (CHIP28) in the membrane at 6 A resolution by cryoelectron crystallography. The protein appears to be a tetramer of four 28-kDa monomers, each of which forms a water channel. The putative, channelforming transmembrane a-helices are numbered 1 to 6 in one monomer (Bar = 10A) (adapted from Mitra et al., 1995, with permission from Nature Structural Biology).

Mg2+ATPase by Ca 2+, at least in platelets. In this regard, membrane fusion events such as endocytosis and exocytosis also probably lead to local transient scrambling of lipid asymmetries (Zachowski, 1993). The initial movement of phosphatidylserine from the inner to the outer leaflet appears to precede vesicle shedding by platelets (Basge et al., 1993). Hence, it is more likely that cortical granule exocytosis rather than subsequent

vesicular budding contributes to the scrambling of phospholipid asymmetry during platelet activation. This mechanism also would account for the movement of sphingomyelin from the outer to the inner leaflet, a movement which does not appear to occur by other biochemical means in most biomembranes. In addition, a Ca2+-dependent "scramblase" appears to catalyze the degradation of the phosphatidylserine and phosphati-

Fluid Mosaic Model of Membrane Structure

23

FIGURE 2.9 Synthesis and insertion of protein molecules into the lumen or membrane of the endoplasmic reticulum (ER). (A) Although the simplified diagram is for insertion of a protein into the lumen of the ER, the presence of multiple, uncleaved start- or stop-transfer signal peptides in the primary structure of a protein presumably can lead to insertion of a multipass, integral membrane protein molecule as in B. (B) Hypothetical model for insertion of a double-pass protein molecule in the ER membrane (adapted from Alberts et al., 1994, with permission from Garland Publishing, Inc.).

dylcholine concentration gradients across the platelet plasma membrane (Comfurius et al., 1996) as well as the plasma membranes of other human cells (e.g., Zhou et al., 1997). Regardless of the mechanism, the movement of phosphatidylserine to the outer leaflet has important physiological and pathophysiological consequences (reviewed more extensively by Zachowski, 1993). In platelets, phosphatidylserine in the outer leaflet favors conversion of coagulation factor X to Xa and the association of factor Xa with factor Va. These changes help to generate a catalytic surface that promotes coagulation. Similarly, abnormal red cells, such as ones that are sickled, adhere more strongly to vascular endothelial cells perhaps as a consequence of the greater concentrations of phosphatidylserine in their outer leaflets relative to normal cells. Also as a consequence of greater external phosphatidylserine exposure, apoptotic lymphocytes and some tumorigenic cells may be destroyed more readily by monocytes or macrophages. Differences in phospholipid composition of the inner and outer leaflets of erythrocytes and quiescent platelets has been used successfully to design hemocompatible surfaces (Chapman, 1993). Phosphatidylcholine coating reduces the adsorption of fibrinogen and platelets to artificial surfaces and prevents platelet activation by the surfaces. These surfaces should be useful in the production of better artificial blood-contacting devices including catheters, indwelling biosensors, extracorporeal circuits, and filtration membranes. It has also been proposed that the process of forming asymmetric distributions of phospholipids in the plasma membrane has itself a function independent of the func-

tions of the asymmetrically distributed phospholipids themselves or subsequent scrambling of their asymmetric distribution (reviewed by Williamson and Schlegel, 1994). In this view, the transport of the phospholipids from the outer to the inner leaflet would create a force in the inner leaflet which bends the membrane inward. Such is almost certainly the case in erythrocytes where excessive transport of phosphatidylserine and phosphatidylethanolamine from the outer to the inner leaflets results in the formation of stomatocytes (i.e., mouthlike cells) (reviewed by Zachowski, 1993). In cells with a less restrictive cytoskeleton, the transport of phospholipids from the outer to the inner leaflet could conceivably contribute to invaginations of the membrane such as those that occur during endocytosis. Whether phospholipid transport p e r se contributes significantly to formation of endocytic vesicles or to other membrane processes remains to be established experimentally.

3. Carbohydrates Unlike proteins and lipids, carbohydrates are not considered to be an integral component of membranes. Their peripheral association with biomembranes is, however, highly asymmetric. Oligosaccharides are covalently bound to membrane lipids and integral proteins only on their noncytosolic sides. Oligosaccharides are assembled, transferred to membrane proteins or lipids, and subsequently modified in the lumen of the endoplasmic reticulum and Golgi apparatus (Fig. 2.11). No mechanism is known for the assembly and attachment of these molecules to segments of the proteins or lipids at the cytosolic surfaces of membranes. The resultant

24

2. Biomembrane Composition, Structure and Turnover

TABLE 2.1 Percentage of Each Main Phospholipid Class Present in the Outer Leaflet of Various Animal Plasma Membranes a Cell

Human erythrocyte Mouse erythrocyte Rat erythrocyte Monkey erythrocyte Human platelet Pig platelet Mouse erythroleukaemic cell LM cell Mouse synaptosome Rabbit intestinal brush border Rabbit kidney brush border Trout intestinal brush border Middle Posterior Rat cardiac sarcolemna Krebs ascites cell Rat hepatocyte Bile canalicular surface Contiguous surface Sinusoidal surface Chick embyro fibroblast Chick embryo myoblast Quail embyro myoblast

Sphingomyelin

PC

PE

PS

80 85 100 100 82 93 91 80

77 50 62 63 67 45 40 45 48

<4 0 6

63

32 30 35

20 20 20 0 13 20 34 50 24 10-15 34 28 23

93 47

43 52

46 36 25 45

32 31 0 19

65

85 80 85

50 0 55 35 65 73

0 20 0 20 45 44

80

65

0 9 6 15 20 44 15

aAbbreviations: PC, phosphatidylcholine; PE, phosphatidylethanolamine; PS, phosphatidylserine (from Zachowski, 1993with permission from the Biochemical Society and Portland Press).

impressive structural difference between the inner and outer surfaces of the plasma m e m b r a n e (Fig. 2.12) appears to have surprisingly little general effect on solute or solvent transport in either direction across the membrane. This carbohydrate-rich zone on the surfaces of cells is known as the glycocalyx. In spite of its likely influence on the properties of the layer of extracellular water with which it is associated, the glycocalyx appears to have little or no general effect on b i o m e m b r a n e transport. Nevertheless, specific glycosylation of some integral m e m b r a n e proteins appears to be essential for their transport activities (e.g., Asano et al., 1991; Ntifiez and Arag6n, 1994), whereas glycosylation of other proteins is not needed at least for transport (e.g., Conradt et al., 1995). The detailed functions of the glycocalyx seem likely to emerge as we learn more about the importance of this complex structure. Currently, components of the

FIGURE 2.10 Synthesisof sphingomyelin in BHK cells. Although transfer of the phosphocholine residue (represented by "+") from phosphatidylcholine to ceramide to form sphingomyelin is shown to occur in both Golgi and recycling endosomes, the latter compartment appears to be the major site of this process in BHK cells. Sphingomyelinase was used experimentally to convert sphingomyelin in the plasma membrane to ceramide plus phosphocholine (adapted from Kallen et al., 1994, with permission from Elsevier Science).

glycocalyx are believed to function mainly in interactions of cells with their environments including other cells. For example, selectins are CaZ+-dependent integral m e m b r a n e proteins that recognize specific oligosaccharides expressed on the surfaces of other cells. Initial recognition by neutrophils destined to migrate out of a blood vessel at sites of inflammation appears to be a selectin-mediated process (McEver, 1992). Interestingly, the binding of carbohydrate by selectins appears to reduce the motion of lipids and proteins in the plasma m e m b r a n e s of the selectin-containing cells (Hajela et al., 1996). Since the binding of ligands by selectins also appears to increase cytosolic viscosity, we suggest that these effects of ligand binding result from an influence of ligand-bound selectins on the cytoskeleton. Changes in the cytoskeleton not only influence the viscosity of the cytosol, but such changes could also alter the motion of proteins and lipids in the plasma membrane. D. M o t i o n of Lipids a n d Proteins The classical fluid mosaic model of b i o m e m b r a n e structure states that the cell m e m b r a n e is "a two-dimensional oriented solution of integral p r o t e i n s . . , in the viscous phospholipid bilayer solvent" (Singer and Nicolson, 1972). In this well-known model (Fig. 2.6) lipids apparently diffuse and rotate freely in each leaflet of the bilayer in which they are present, but they are able to flip from one leaflet to the other relatively rarely. Moreover, as discussed above for cholesterol vs phospholipids, the probability that a lipid will move from one leaflet to the other decreases as the extent to which

Fluid Mosaic Model of Membrane Structure

25

FIGURE 2.1 1 Assembly (not shown), transfer (A), and modification (B) of oligosaccharides to form glycoproteins in the endoplasmic reticulum (ER) (A) and Golgi (B). Glycosylation occurs almost as soon as a polypeptide chain containing a glycosylation site enters the lumen of the ER. Oligosaccharides are also transferred to lipids to form glycolipids in a similar process (adapted from Alberts et aL, 1994, with permission from Garland Publishing, Inc.).

they are amphipathic increases. Apparently, the free energy barrier is too high in the absence of a catalyst to allow the hydrophilic heads of phospholipids frequently to pass from one surface of the membrane to the other through its hydrophobic interior. Similarly, many integral membrane proteins diffuse laterally and rotate in the bilayer but they rarely if ever reverse their orientation in it. Protein motion may also be influenced differently by phospholipids in the inner and the outer leaflets. The asymmetric distributions of phospholipid polar heads across biomembranes could conceivably influence protein diffusion and rotation asymmetrically, although the asymmetric fluidities of the inner and outer leaflets of the lipid bilayer seem more likely so to influence protein motion. The physiological significance of the asymmetric fluidities of the bilayer leaflets remains largely to be determined, although its regulation seems likely to be

involved in numerous normal and pathological processes. For example, some of the effects of movement of phosphatidylserine from the inner to the outer leaflet of the plasma membrane bilayer (discussed in Section II,C,2 above) may be attributable to changes in the relative fluidity of the leaflets rather than (or in addition to) the increased concentration of the polar head of phosphatidylserine in the outer leaflet. In addition, the pore-forming protein, perforin, is inserted into membranes more easily when their lipids are less tightly packed. This property along with others, such as the nature of the glycocalyx, may help cytotoxic T lymphocytes, which secrete perforin, to kill target cells but not themselves; the lipids in the outer leaflet of the T lymphocyte plasma membrane are more tightly packed than those in the target cells (Williamson and Schlegel, 1994). The asymmetric fluidities of the two leaflets of the plasma membrane lipid bilayer appear to result mainly

26

2. Biomembrane Composition, Structure and Turnover

FIGURE 2.12 The glycocalyx of the lymphocyte plasma membrane. The glycocalyx is more than five times as thick as the lipid bilayer of the plasma membrane. Although the lipid bilayer of the plasma is not clearly delineated, it is known to be about 5 nm wide (adapted from Alberts et al., 1994, with permission by Garland Publishing, Inc.).

from the different fatty acyl compositions of membrane phospholipids. For example, the phosphatidylserine in human erythrocyte plasma membranes has a greater proportion of unsaturated fatty acyl chains than does the phosphatidylcholine in membranes of these cells (Myher et al., 1989). Hence, the inner leaflet of the lipid bilayer, which normally contains a higher concentration of phosphatidylserine, melts over a lower temperature range than the outer leaflet of the bilayer, which contains a higher concentration of phosphatidylcholine. A similar explanation also probably accounts for differences in the fluidities of the inner and outer leaflets of biomembranes of other cells such as fibroblasts (E1Hage Chahine et al., 1993). From our discussion thus far, it can be concluded that the fluid mosaic model of membranes is still largely valid, although the first experiments (e.g., Kornberg and McConnell, 1971) did not detect the full complexity of phospholipid motion. In fact, studies on lipid mobility still frequently do not discriminate between the motion of lipids in the liquid and crystalline domains that are present in biomembanes at physiological temperatures. Hence, while the liquid-crystalline state of lipids has been studied extensively in both artificial and natural membranes, the liquid and crystalline domains within such membranes (Fig. 2.5) are only now being characterized (e.g., Hui, 1995).

It is also a remarkable characteristic of metabolism in cells at different temperatures that they can produce membrane lipid compositions that are, nevertheless, at their melting temperatures. Even more remarkably, the membranes of at least some living cells undergo phase transitions at different temperatures than the same membranes after isolation from the cells (Fig. 2.4). Hence, phase transitions of lipids in membranes appear surprisingly to depend on the structure and metabolism of whole cells as well as on the composition of biomembranes p e r se. As for the emerging complexity of lipid motion in membranes, the motion of integral membrane proteins is more variable than indicated by the surface antigens selected initially for study (e.g., Frye and Edidin, 1970). The latter proteins are clearly much more mobile than are the membrane proteins that are tethered to the cytoskeleton. In addition to these nearly immobile proteins, many membrane proteins appear to experience transient confinement to domains in membranes that are not the result of a liquid-crystalline phase transition. Still other proteins appear to undergo much more rapid movement in particular directions than can be accounted for even by their completely unrestricted lateral diffusion in the lipid bilayer. These nonrandom migrations of some proteins and the limitations on the motion of others appear to result mainly from the now well

Fluid Mosaic Model of Membrane Structure

established interactions of the cytoskeleton with constituents of the plasma membrane. E. The Cytoskeleton and Other Cellular Structures Influence Protein and Lipid Motion in Biomembranes Numerous integral membrane proteins are now known to be tethered to the cytoskeleton and, hence, are not free to diffuse laterally in the lipid bilayer (Jacobsson et aL, 1995). The structures and functions of some such proteins have been studied in detail as have the ways in which they attach to the cytoskeleton. For example, 1/3of the band 3 protein, which catalyzes anion exchange in erythrocytes (discussed in greater detail in Chapter 6), is attached to the cytoskeletal protein, spectrin, via ankyrin (Fig. 2.13). The band 4.2 protein appears to stabilize the association between band 3 and ankyrin (Inoue et aL, 1994; Golan et al., 1996). Interestingly, the lateral diffusion of the band 3 protein is restricted by this attachment although its rotational movement is not restricted. Apparently the cytosolic and membrane domains of the band 3 protein are connected to each other by a flexible hinge (Wang, 1994). A mutation in the hinge area (i.e., a nine-residue deletion in the band 3 protein) results in ovalocytic erythrocytes (reviewed by Wang, 1994). The ovalocytic red cells in this condition, known as Southeast Asian ovalocytosis, also are very rigid. Apparently the hinge is needed to maintain the erythrocyte shape, strength, and flexibility

FIGURE 2.13 In erythrocytes, some of the band 3 protein is attached to the cytoskeletal protein spectrin via ankyrin. Although this attachment limits the lateral diffusion of the band 3 protein molecule within the lipid bilayer, it somewhat surprisingly does not appear to limit rotation of the molecule. The zig-zag lines represent the hinge region of the band 3 protein molecule, which connects its cytosolic and membrane domains (adapted from Wang, 1994, with permission from Elsevier Science).

27

attributed to the interaction between the band 3 protein and the cytoskeleton. A model completely explaining the needed interactions and the function of the hinge apparently as a swivel is still being developed. Similarly, models are only now emerging for the structures connecting the cytoskeleton to membrane proteins destined to form clusters, such as those formed by acetylcholine receptor protein molecules at neuromuscular junctions (Gomparts, 1996). In the case of clustering of postsynaptic glycine receptor protein molecules, the receptorassociated protein gephyrin appears to anchor the receptor proteins to the cytoskeleton (Kirsch and Betz, 1995). In contrast to relatively immobile membrane proteins that are tethered to the cytoskeleton, sometimes the association of an integral membrane protein with the cytoskeleton may lead to their rapid nondiffusional movement in a particular direction. For example, integrins undergo directed excursions toward the edge of migrating fibroblasts (Schmidt et al., 1994). Moreover, integrins with mutations in their cytoskeletal binding regions display different degrees of directed transport. Hence, their directed excursions appear to require binding of the integrins to the cytoskeleton presumably also in association with cytoskeletal motor proteins. These directed movements of integrins apparently are needed in migrating fibroblasts in order for them to traverse the length of an extending lamellipod fast enough to supply adhesion receptors to points of new cellsubstratum contact (Schmidt et al., 1994). In addition to the restricted or directed motion of integral plasma membrane proteins that results from attachment to the cytoskeleton, the cytoskeleton may also limit the motion of integral membrane proteins by temporarily confining them to a region of the membrane skeleton meshwork (reviewed by Jacobson et aL, 1995). In this "membrane skeleton fence model" (Kusumi et aL, 1993; Tomishige et al., 1998) the membrane is divided into small domains or compartments with areas of 0.04 to 3.0/~m 2 by cytoskeletal components just under the membrane (Fig. 2.14). Because the distance between the membrane and the cytoskeletal meshwork may vary, proteins that are not bound to the cytoskeleton may escape to adjacent compartments at frequent intervals. Nevertheless, it is likely that all integral membrane proteins are confined in their lateral diffusion in the lipid bilayer at least some of the time by the cytoskeletal meshwork just beneath the plasma membrane. Hence, the fluid mosaic membrane model (Fig. 2.6) must be modified to take into account all of these effects of the cytoskeleton on plasma membrane protein molecules. Finally, there appear to be at least two ways in which membrane lipids and proteins are confined

28

2. Biomembrane Composition, Structure and Turnover

FIGURE 2.14 The cytoskeletal meshwork just under the plasma membrane appears transiently to limit the lateral diffusion of integral membrane protein molecules in the lipid bilayer. Because the distance between the meshwork and membrane vary, protein molecules, such as the transferrin and EGF receptors, can escape at regular intervals to different domains delineated by the meshwork (adapted from Kusumi et aL, 1993, with permission from the Biophysical Society).

transiently to small domains that do not necessarily involve the cytoskeleton. First, and as discussed above, numerous crystalline and liquid domains may exist temporarily in lipid bilayers of biomembranes and the presence of protein or lipid molecules in crystalline domains restricts their motion more than when they are in liquid ones. Similarly, the cytoskeleton would not be needed for confinement of protein molecules in clusters formed through interactions among their oligosaccharide moieties. Association of molecules of the rat form of the glutamate transport protein EAAT1 (GLAST-1) with each other or with other proteins in the plasma membrane could conceivably occur in this manner (Conradt et al., 1995). Such interactions have also been proposed to form glycosphingolipid rafts that may be associated with glycosyl-phosphatidylinositol-linked proteins (Fig. 2.15) in the Golgi and at the cell surface (Simons, 1995). Similar types of confinement of proteins in membranes have also been postulated based on data such as that for a lipid linked isoform of neural cell adhesion molecules which, because they are not integral membrane proteins, cannot be confined through physical interaction with the cytoskeleton (Simson et al., 1995). These proteins may be confined owing to interactions among themselves (the proteins lie outside the membrane lipid bilayer, Fig. 2.15) or with other proteins that are associated with the cytoskeleton.

N-terminal

"'i...... m PresentIon GPI anchors

(' In ~~) CH2-CH-CH2~ S

Membrane bilayer

I Fatty acids

FIGURE 2.15 Major components of glycosyl phosphatidyl-inositollinked proteins in membrane bilayers. Protein molecules of this type may associate with one another or with glycolipid rafts through intermolecular hydrogen bonds and other types of interactions. Such associations could limit diffusion of the proteins and lipids in the lipid bilayer by a mechanism that does not require involvement of the cytoskeleton. Residue abbreviations: Eth, ethanolamine; P, phosphate; M, mannose; GI, glucosamine; In, inositol; GPI, glycosyl phosphatidyl-inositol (adapted from Brown and Waneck, 1992, with permission from Williams and Wilkins, Baltimore, MD).

Reconstitution of Biomembrane Components

I11. SOME COMPONENTS OF THE BIOMEMBRANE CAN BE RECONSTITUTED Attempts to reconstitute proteins in artificial membranes range from fully successful to entirely unsuccessful. In some cases, such as reconstitution of a glutamate transporter (Danbolt et al., 1990), there is a reproducible and apparently obligatory loss of about 80% of its transport activity upon isolation, purification, and reconstitution of the protein in proteoliposomes. Although many technical as well as more physiological explanations beyond the scope of this chapter may contribute to the variable success in reconstituting different proteins, our current discussion already informs us of what some of these explanations may be. First, attachment to other proteins, such as those in the cytoskeleton, may be more important to the functions of some integral membrane proteins than others. For example, isolation of the system A amino acid transport protein and reconstitution of its transport activity in proteoliposomes appears to require coisolation and association with the ~3/~1 integrin (McCormick and Johnstone, 1995). This integrin could mediate interaction of the system A transport protein with the cytoskeleton. Moreover, it has been suggested that a system A regulatory protein may be a component of the cytoskeleton (Ruiz-Montasell et al., 1994). In contrast, anion exchange via band 3 does not require its connection to the cytoskeleton (e.g., Motais et al., 1997 and see Chapter 6). In this regard, whether a protein molecule requires association with another, different protein molecule in order to function (and whether its function can be reconstituted without the other protein) may depend on which of its functions is under consideration. For example, while the anion exchange function of the band 3 protein does not require attachment to the cytoskeleton, its function in maintaining erythrocyte shape and mechanical strength almost certainly does require this association. Similarly, the association of some membrane proteins through, say, their oligosaccharide moieties may be required for the proteins to function normally in all situations in vivo, whereas their known biochemical function, such as transport, may not be influenced detectably by the oligosaccharides upon heterologous expression of the proteins in other cells (e.g., Conradt et aL, 1995). In addition, the transport and other functions of some integral membrane proteins may depend on the asymmetric lipid distribution or on the resultant asymmetry in fluidity between membrane bilayer leaflets, whereas the functions of other proteins may not depend on either of these asymmetries. Likewise, the functions of some proteins may depend on whether they are in a liquid

29

lipid or crystalline lipid domain and even on whether the domain undergoes a local phase transition while the protein is in the domain. Other proteins may have no such complex requirements of the lipid milieu in order to function. Similarly, the requirements of particular lipid species or changes in species for protein function are likely different for various proteins. It is, however, difficult currently to measure the effects of different lipid species on protein function under more or less physiological conditions, especially since some lipid components of the plasma membrane are partitioned preferentially to the liquid or crystalline phase (see Section II,A above). Nevertheless, our ability to perform such studies may improve now that liquid and crystalline domains can be observed (Fig. 2.5) and their lipid composition determined (Hui, 1995). Similarly, it appears already to be possible to produce proteoliposomes containing both the pertinent major lipids and MgZ+ATPase (e.g., Auland et al., 1994). As discussed in Section II,C,2 above, Mg 2+ ATPase catalyzes migration of phospholipids from the outer to the inner leaflet of the plasma membrane. Hence such liposomes should form a more or less normal asymmetric distributions of phosphatidylserine and phosphatidylethanolamine in the presence of Mg 2+ ATP. A technical question would be whether proteoliposomes need to be formed with an asymmetric distribution of MgZ+ATPase even though simply supplying MgZ+ATP in either the inside or the outside compartment would appear to be enough to produce lipid asymmetry across the artificial bilayer. Even if asymmetric reconstitution of the enzyme is necessary, however, success at asymmetrically reconstituting band 3 protein molecules in proteoliposomes (Boulter et al., 1996) shows that asymmetric reconstitution of membrane proteins is feasible. Moreover, the asymmetric reconstitution of some proteins in lipid bilayers appears to occur automatically (Dierks and Kr~imer, 1988). General acquisition of the ability to control whether proteins are asymmetrically oriented should allow more general determination of the importance of an asymmetric orientation to protein function when the bilayer is or is not also asymmetric in regard to lipid concentrations. Asymmetric reconstitution of a protein for phosphatidylcholine transport in the reverse direction of phosphatidylserine and phosphatidylethanolamine could conceivably permit formation of liposomes with the correct asymmetries of all three of these major membrane phospholipids. Additional measures would be required to produce an asymmetric distribution of sphingomyelin across artificial liposome bilayers since the asymmetric distribution of this phospholipid in the outer leaflet of the bilayer is produced at the time of its synthesis in the Golgi. Ultimately we may need to learn how to

30

2. Biomembrane Composition, Structure and Turnover

reconstitute in proteoliposomes all of the ways that living cells influence the fluidity and other characteristics of their membranes (e.g., Fig. 2.4). These measures are necessary both to produce artificial proteoliposomes in which the structure of the lipid bilayer corresponds to the actual structure of the bilayer in biomembranes and to test the effects of bilayer structure and composition on integral membrane protein function.

IV. H O W IS BIOMEMBRANE COMPOSITION AND STRUCTURE REGULATED? A. Vesicular Traffic The phospholipid bilayer is synthesized primarily in the endoplasmic reticulum. Lipids in the newly synthesized bilayer could conceivably be transferred to each of the other membranes in the cell via exchange of the lipid molecules among the membranes. This transfer process, termed monomeric exchange, has been shown to be catalyzed by water soluble phospholipid transfer

(or exchange) proteins in in vitro experiments (e. g., Dowhan, 1991). Monomeric exchange appears, however, mainly to transfer newly synthesized phospholipids from the membrane of the endoplasmic reticulum to mitochondrial and peroxisomal membranes (see Section IV,C below). Additions to other membranes occurs principally through vesicular trafficking from the endoplasmic reticulum through the Golgi and subsequently to lysosomes, endosomes, or the plasma membrane (Fig. 2.16). Vesicular trafficking also occurs among these different compartments, and proteins and even the proportions of different lipids may vary among vesicles with different origins and destinations. Such trafficking involves formation of membrane vesicles from each of the membrane compartments and subsequent fusion of the vesicles with the target compartments. The formation of vesicles and their subsequent fusion with the proper target membrane involves specific targeting signals and docking mechanisms (A1berts et aL, 1994). Movement and fusion of the vesicles also require various components of the cytoskeleton

FIGURE2.16 Vesiculartrafficking between and among the endoplasmic reticulum (ER), Golgi apparatus, endosomes, lysosomes, and the plasma membrane. New membrane is synthesized in the ER and may be modified in other compartments. The new membrane is directed to the appropriate compartment by mechanisms that are still being elucidated (see text). Thick arrows indicate compartments that are connected by vesicular trafficking and in most cases vesicles move in both directions. Dotted lines are used to represent membranes of organelles that receive lipid through monomer exchange rather than vesicular trafficking. Also shown (thin arrows) is the presumed development of some endosomes into endolysosomes and subsequently into lysosomes after addition of acid hydrolases via vesicular trafficking from the trans-Golgi network (adapted from van Helvoort and van Meer, 1995, with permission from Elsevier Science).

How is Biomembrane Composition and Structure Regulated?

and their respective motor proteins (see Section IV,D below). Such trafficking remains a very active field of investigation, and the details of the process vary to some extent from cell type to cell type. If a cell and its organelles are to remain more or less the same size, trafficking of vesicles to target membranes must of course be balanced by removal of membrane constituents mainly through endocytosis and budding, respectively. Vesicular trafficking is also the means by which most integral membrane proteins are sorted and sent to the biomembranes in which they function. B. Protein Sorting Integral membrane proteins contain information in their primary structures for insertion into lipid bilayers. Most such proteins are inserted into bilayers in the endoplasmic reticulum (Fig. 2.9). These proteins also contain information in their primary and conformational structures which permits them to be retained in a particular compartment or sent to another one (e.g., Fig. 2.16). It is commonly believed that proteins are sent from the endoplasmic reticulum to the Golgi and from the Golgi to the plasma membrane via default pathways that do not require signals in nonpolarized cells. Signals probably are required, however, to direct proteins to either the apical or basolateral plasma membranes of polarized cells (see Section D below). In fact, the conclusion that any protein sorting occurs by default probably should remain provisional even for apparently nonpolarized cells. The number of cell types that are considered to be polarized has increased as we have learned more about their biochemical nature and the details of their physical appearance. Similarly, we have only begun to learn the mechanisms of protein sorting and vesicular trafficking even in well-studied cells. For example, site-directed mutagenesis of a plasma membrane Ca2+ATPase led in the cases of two different single amino acid residue substitutions to retention of the protein in the endoplasmic reticulum (Guerini et aL, 1996). Such results for two separate amino acid residue substitutions seem unlikely to occur if a signal is needed for retention of proteins in the endoplasmic reticulum but not for transport of proteins to the cell surface. In this case, it might be argued that proper folding is needed for proteins to follow the default pathway. If, however, proteins are monitored for proper folding or even for the absence of a signal on their way to the surface, then this folding or the apparent absence of a signal are themselves signals. Similarly, maintenance of some proteins in the trans-Golgi network by retrieving them from the cell membrane (Stanley, 1996) is inconsistent with the simple model that proteins are either retained in intracellular compartments due to signaling or

31

directed to the plasma membrane by default. For these and other reasons, it has become apparent that bulk flow out of the endoplasmic reticulum to the plasma membrane via default pathways is unlikely to be a major mechanism of protein sorting (reviewed by Rothman and Wieland, 1996). C. Phospholipid Synthesis, Transbilayer Transport, and M o n o m e r i c Exchange Most membrane phospholipids are synthesized on the cytosolic side of the endoplasmic reticulum (Fig. 2.17). Hence they would accumulate in excess in the inner leaflet of the bilayer were it not for hydrophilichead-group-specific phospholipid translocators, or "flippases", that appear collectively to catalyze equilibration of most phospholipids across the membrane of the endoplasmic reticulum (Fig. 2.18). 2 These apparently equilibrating flippases are clearly unlike the Mg 2+ ATPase of the plasma membrane (Section II,C,2 above) which appears to transport phospholipids in one direction across the bilayer against their total chemical potential gradients. Subsequent to synthesis of most phospholipids in the endoplasmic reticulum, sphingomyelin is synthesized in the outer leaflet of the lipid bilayer of the cis-Golgi through transfer of a phosphocholine residue from phosphatidylcholine to ceramide (Fig. 2.19). Because phospholipids (particularly sphingomyelin) flip from one leaflet of the bilayer to the other only very slowly in the absence of a catalyst, sphingomyelin tends to remain in the outer leaflet during subsequent trafficking of vesicles to the plasma membrane (see also Section II,C above). Nevertheless, since membrane fusion is believed at least partially to scramble phospholipid asymmetry in the bilayer and because membrane fusion occurs repeatedly during vesicular trafficking other mechanisms may exist to produce and maintain an asymmetric distribution of sphingomyelin in the plasma membrane. In theory, phospholipids in the inner, cytosolic leaflet could move through monomeric exchange directly between and among the endoplasmic reticulum, Golgi, endosomes, lysosomes, and plasma membrane. Monomeric exchange appears, however, to occur mainly between the endoplasmic reticulum and mitochondrial and peroxisomal membranes possibly as a result of the action of phospholipid exchange proteins or because the cytosolic leaflets of the lipid bilayers of these organelles can in some cases establish areas of close apposition (reviewed by van Helvoort and van Meer, 1995). Such transfer of phospholipids to 2We use the widely used term "flippase" with reluctance. The "-ase" suffix should be reserved for proteins that catalyze chemical change.

32

2. Biomembrane Composition, Structure and Turnover

FIGURE 2.17 Phospholipid synthesis is catalyzed by enzymes in the endoplasmic reticulum membrane. Synthesis occurs exclusively in the cytosolic leaflet of the lipid bilayer, since the active sites of the enzymes face the cytosolic compartment (adapted from Alberts et al., 1994, with permission from Garland Publishing, Inc.).

mitochondrial and peroxisomal membranes is, of course, needed because these organelles are not connected to the others via vesicular trafficking (Fig. 2.16). Similarly, these organelles must receive and insert integral membrane proteins via the cytosol rather than through vesicular trafficking. Signal sequences are involved in the insertion of proteins into the inner mitochondrial and peroxisomal membranes. Similar processes also are required for insertion of proteins into the thylakoid membrane of chloroplasts and the plasma membrane of prokaryotes. Interestingly, phosphatidylethanolamine acts as a chaperone in assembly of the lactose transporter into the Escherichia coli plasma membrane (Bogdanov et al., 1996). Because we need to conserve space for our main topic of biomembrane transport, we consider further here only formation and maintenance of epithelial cell plasma membranes as an example of how proteins are incorporated into the appropriate membrane.

D. Biogenesis of Epithelial Cell Polarity as an Example of Regulation of M e m b r a n e Composition and Structure Many of the descriptions of mechanisms by which membrane composition is regulated (other than regulation of expression of genes for specific proteins; e.g., Chapter 9) have arisen from studies of the biogenesis of epithelial cell membrane polarity. The basolateral and apical plasma membrane domains are maintained by tight junctions in epithelia (Fig. 2.20). These structures prevent mixing of the components of the apical and basolateral membranes by preventing lateral diffusion of lipids and proteins between the two membranes. Since tight junctions prevent mixing in the outer but not the inner leaflet of the bilayer, however, lipids in the inner leaflet and peripheral proteins bound covalently or noncovalently to lipids in the inner leaflet are apparently free to diffuse laterally between the apical and basolat-

How is Biomembrane Composition and Structure Regulated?

33

FIGURE 2.18 Flippases appears to catalyze the movement of phosphatidylcholine and probably other phospholipids from the inner leaflet where they are synthesized to the lumenal leaflet of the endoplasmic reticulum membrane lipid bilayer. Also indicated (A) is the movement of lipids from the endoplasmic reticulum, through the Golgi apparatus (not shown) to the plasma membrane via vesicular trafficking. The details of the flipping indicated in A are shown in B (adapted from van Helvoort and van Meer, 1995 (A) and Alberts et al., 1994 (B) with permission from Elsevier Science and Garland Publishing, Inc.).

eral membranes. Consistent with this model, glycosphingolipids, which are in the outer leaflet, remain apically enriched. Similarly, phosphatidylcholine, which may also be asymmetrically distributed in the outer leaflet, is kept predominantly in the basolateral membrane. Phosphatidylserine and phosphatidylethenolamine are, however, apparently free to move in the inner leaflet between the apical and basolateral membranes. In addition to tight junctions, a second way in which integral membrane proteins can be retained in either the basolateral or the apical membrane is through attachment to the cytoskeleton. For example, by binding to Na+K+ATPase, the membrane skeleton restricts the access of this enzyme to endocytic vesicles and hence may help to preserve its polarized location in the plasma membrane (Hammerton et aL, 1991). The different compositions of the basolateral and apical membranes are produced through preferential sorting of lipids and proteins to one membrane or the other via membrane vesicles. For example, in Madin-

Darby canine kidney (MDCK) cells, at least two populations of plasma membrane-directed vesicles form in the trans-Golgi network. One population of membrane vesicles carries Semliki Forest virus spike glycoprotein selectively to the basolateral membrane. In contrast, another population of vesicles carries influenza virus hemagglutinin to the apical surface (Fig. 2.20). The different biochemical information provided in each of the two populations of vesicles that distinguishes whether they are sent to the apical or basolateral membrane is only beginning to emerge. 1. The Cytoskeleton and Motor Proteins Help to Move Membrane Vesicles from the Golgi to Their Apical or Basolateral Destinations

Both microtubules and actin filaments and their motor proteins appear to be involved in the selective targeting of membrane vesicles to the apical membrane (Fig. 2.21). Some evidence also has accumulated to

34

2. Biomembrane Composition, Structure and Turnover

FIGURE 2.19 Synthesis of sphingomyelin in the lumenal leaflet of the cis-Golgi membrane. Since there appear to be no flippases for sphingomyelin or other phospholipids in the cis-Golgi, sphingomyelin is present primarily in the outer (noncytosolic) leaflet after trafficking in membrane vesicles to the plasma membrane (see Fig. 2.16).

support the theory that microtubules are needed for movement of vesicles to the basolateral membrane (reviewed by Brown and Stow, 1996 and by Weimbs et aL, 1997). Although movement along components of the cytoskeleton is probably a general mechanism by which vesicular trafficking occurs, it is not yet clear whether it is the only way to produce directed vesicular movement. The details of how motor proteins recognize vesicles for transport to either the basolateral or the apical membrane is also being investigated. Similarly, the manner in which the cytoskeleton may be involved in the docking and fusion of vesicles with each of these membranes is an active area of research, and the docking and fusion proteins that may serve in each of these cases should soon be identified and compared. Different cytoskeletal components and a growing array of motor proteins (e.g., Goodson et al.,

1997) provide several fundamentally similar ways in which directed vesicular movement can be achieved. Vesicular traffic that depended on more or less random vesicular movement would, of course, be much less efficient than directed movement.

2. Lipid and Protein Sorting to the Basolateral Membrane The mechanism by which some lipids, such as phosphatidycholine, are delivered preferentially to the basolateral membrane in MDCK and other polarized cells remains an active area of research. Some components of the process by which proteins are sorted to this membrane are, however, already beginning to emerge. Sorting to the basolateral membrane appears to involve signals in the cytosolic domains of proteins, as opposed

How is Biomembrane Composition and Structure Regulated?

FIGURE 2.20 Formation of at least two populations of plasma membrane-directed vesicles in the trans-Golgi network of MDCK cells. For example, one population of membrane vesicles (open circles) carries Semliki Forest virus spike glycoprotein to the basolateral but not to the apical membrane, whereas another population of vesicles (solid circles) carries influenza virus hemagglutinin to the apical but not to the basolateral membrane (adapted from Simons, 1995, with permission from John Wiley & Sons, Inc.).

to apical signals which seem to be in either transmembrane or lumenal protein domains (Fig. 2.22). As expected, removal of the cytoplasmic "tails" of the apically sorted proteins does not alter their destination, whereas such treatment redirects some basolateral proteins to the apical membrane. For this reason, the basolateral signal appears to be dominant to the apical one. In addition, several different signals for basolateral sorting appear to exist. One goal of current research is to determine how these signals are decoded by one or more proteins that are presumed somehow to cluster cargo proteins into membrane patches destined for the basolateral membrane. Such patches presumably form vesicles in the trans-Golgi network, and the vesicles are subsequently transported preferentially to the basolateral membrane (reviewed by Simons, 1995). Proteins termed NSFs, SNAPs, and SNAREs are known to be involved in vesicular docking and fusion in intra-Golgi transport, and these proteins also appear to be involved in transport of vesicles to both the basolateral and the apical membranes (Low et al., 1998). Investigators in this field are, however, still attempting to learn the details of how vesicles from the trans-Golgi network are directed specifically to the basolateral membrane. Much effort continues to be devoted to determining the protein composition of such vesicles and

35

FIGURE 2.21 Motor proteins move membrane vesicles from the trans-Golgi network to the apical plasma membrane. In this scheme, the vesicles are first carried by the microtubule (MT)-associated motor protein dynein along microtubules and then by myosin-I along actin filaments (adapted from Fath et al., 1993, with permission from Company of Biologists Ltd.). In contrast (and not shown) kinesinlike motor proteins may carry vesicles to the basolateral membrane in the reverse direction along microtubules (Weimbs et al., 1997).

their lipid composition may hold as yet unexplored clues to the mechanism of their selective trafficking. Interestingly, the intracellular parasite Chlamydia trachomatis interrupts vesicular trafficking from the trans-Golgi network to the plasma membrane (Fig. 2.23) apparently in order for it to acquire newly synthesized sphingomyelin (Hackstadt et al., 1996). Perhaps study of these or similar pathogens and the nature of the vesicles they do or do not attract in polarized cells will yield insight into the nature of the vesicles and the ways in which they are normally directed to the basolateral or apical membranes. 3. Vesicular Traffic to the Apical Membrane

Transport of membrane lipids and proteins to the apical membrane may be direct or indirect. In the indirect pathway, vesicles with apical cargo are first sent to the basolateral membrane. Subsequently, proteins and lipids to be sent to the apical membrane are incorporated into endocytic vesicles of the basolateral membrane. The latter process presumably results in formation of basolateral endosomes from which apically

36

2. Biomembrane Composition, Structure and Turnover

FIGURE 2.22 Signals for sorting of proteins to the basolateral membrane appear to be located in their cytosolic domains, whereas the transmembrane or lumenal domains of proteins appear to contain signals that direct them to the apical membrane. As expected, sorting to the apical membrane still occurs when the cytosolic "tails" of the pertinent proteins are removed. Somewhat surprisingly, however, several basolateral proteins are sent to the apical membrane after removal of their cytosolic tails. Hence, the basolateral signal appears in some instances to be "dominant" to the apical one (see text) (adapted from Simons, 1995, with permission from Wiley-Liss, Inc.).

Plasma

SM / Endoytic . Compartment ",,

",,

f

i

Chlamydial usion

(~)

T

Golgi Apparatus Ceramide From Endoplasmic~ Reticulum / ' FIGURE 2.23 The intracellular parasite Chlamydia trachomat& interrupts vesicular trafficking from the transGolgi network to the plasma membrane. This interruption of normal cellular functioning apparently allows the parasite to acquire needed sphingomyelin (SM). The parasite apparently cannot intercept endocytic vesicles nor vesicles emanating from recycling endosomes. In some cells the latter compartment may be the major site of sphingomyelin synthesis (shown in greater detail in Fig. 2.10) (adapted from Hackstadt et al., 1996, with permission from Oxford University Press).

How is Biomembrane Composition and Structure Regulated?

directed vesicles are then formed (Mostov et aL, 1992). The constituents of these vesicles then pass through an apical recycling compartment before the proteins and lipids are sent finally to the apical membrane (Weimbs et al., 1997). It is currently unclear where and by what process apical proteins are segregated from basolateral ones in the indirect pathway. Partly because the parasite Chlamydia discussed above does not intercept endocytic vesicles (Fig. 2.23), it should be interesting to compare the composition of the vesicles involved in trancytosis from the basolateral to the apical membrane with the composition of vesicles moving from the trans-Golgi to the basolateral membrane. Some polarized cells, such as hepatocytes, do not appear to have a direct route for trafficking vesicles from the Golgi to the apical plasma membrane and hence rely exclusively on the indirect route to sort proteins and lipids first to the basolateral surface and from there to the apical membrane. Other cells, such as Caco-2 cells, use both the direct and indirect routes, whereas MDCK cells appear to use the direct route exclusively (Simons, 1995). Elucidation of the direct route of protein sorting to the apical plasma membrane involves an interesting question. Since the signals for apical sorting appear to be either in the transmembrane or lumenal domains of proteins (Fig. 2.22), how are such signals recognized in the cytosol for transport to the apical membrane? For example, glycosyl-phosphatidyl-inositol-linked proteins are attached covalently to lipid in the lumenal leaflet of the membrane bilayer, but the protein molecules do not penetrate the membrane. Hence, their signals for transport to the apical membrane appear to lie entirely within the lumens of the pertinent organelles and membrane vesicles. Although mechanisms for recognition of such signals can, of course, be envisioned (e.g., Le Gall et aL, 1995 and see paragraph below), an actual mechanism of recognition has not as yet been demonstrated for any such instance. It is also perplexing that a particular protein may be sorted to the apical membrane in some cells and to the basolateral membrane in others. For example, the transferrin receptor is located in the basolateral membrane of most cells but it is in the apical membrane of the placental trophoblast (Cerneus et aL, 1993). In the latter case, the receptor presumably contributes to the uptake of iron from the maternal circulation by the trophoblast in order to deliver it to the fetus. It remains to be determined, however, how transcytosis of the receptor occurs in the trophoblast while it does not occur in other epithelia. Similarly, unlike most epithelia, LDL receptor is apically rather than basolaterally located in the renal epithelium (Pathak et aL, 1990). Na+K+ATPase is also usually located in the basolateral membrane although it is sent preferentially (and appro-

37

priately) to the apical membrane of the secretory epithelium in choroid plexus apparently via a microtubuledependent process (Alpert et al., 1994). It is of course relatively easy to envision simple mechanisms, such as phosphorylation, by which cytosolic basolateral signals might be inactivated. Proteins so phosphorylated conceivably at tyrosine residues (Roush et al., 1998) might then be redirected from the basolateral to the apical membrane. It is, however, curious that a simple mechanism such as phosphorylation has not as yet been shown to apply to rerouting. In fact, phosphorylation may sometimes be involved in transcytosis of proteins expected to lie in the apical membrane (Casanova et al., 1990). If phosphorylation were also involved in rerouting, however, one would still need to learn the mechanism by which the normally basolateral proteins are selected for phosphorylation in order to redirect them to the apical membranes. Further modulation of the already complicated sorting machinery could also account for the rerouting of some proteins to the apical instead of the basolateral membrane in some epithelia, although this mechanism is considered by some investigators to be less likely than phosphorylation or a similar covalent modification of the proteins to be sorted. It has also been suggested that interaction of membrane proteins, such as Na+K+ATPase, with the cytoskeleton may retard their degradation (Nelson, 1994). Preferential stabilization in this manner could also result in concentration of a protein in one membrane or the other. While much remains to be learned (Weimbs et al., 1997), a working hypothesis has emerged for the sorting of lipids and proteins directly to the apical membranes of cells in polarized tissues (Simons, 1995; Simons and Ikonen, 1997). This model is based primarily on several features that appear to distinguish apical transport from other instances of vesicular trafficking and on the lack of detection of involvement in apical trafficking of proteins known to be involved in other vesicular routes. Sorting to the apical membrane is proposed to occur via the formation of glycosphingolipid clusters or rafts in the Golgi complex (Fig. 2.24). These clusters are believed to form owing to the ability of the oligosaccharide moities to form intermolecular hydrogen bonds (Simons, 1995) and possibly the tendency of the long saturated hydrophobic acyl chains of these glycolipids to pack into crystalline lipid domains (Schroeder et al., 1994). The ability of the oligosaccharides to form hydrogen bonds should probably increase during modification of their structures in the Golgi. Glycosyl-posphatidyl-inositollinked proteins appear to become able to form a complex with glycosphingolipids when Golgi enzymes render them endoglycosidase H-resistant owing to the processing of their oligosaccharide moities. The glycosphingolipid raft with its cargo is envisioned to move via

38

2. Biomembrane Composition, Structure and Turnover

cytosol

FIGURE 2.24 Formation of glycosphingolipid rafts in the Golgi through hydrogen bonding between oligosaccharide moities. The rafts should also associate with glycosyl-phosphatidyl-inositol-linked proteins (Fig. 2.15) and they could conceivably also associate with integral membrane proteins. Several rafts containing proteins are envisioned to associate into sorting platforms in the trans-Golgi network from which apically directed vesicles could form with the aid of as-yetunidentified putative coat proteins (adapted from Simons, 1995, with permission from John Wiley & Sons, Inc.).

vesicular trafficking to the trans-Golgi network where integral membrane proteins destined for the apical membrane could conceivably become associated with it. At least one apically directed integral membrane protein has, however, been found not to associate with such rafts (Alonso et al., 1997), so at least one other mechanism appears to exist for apical trafficking. When rafts are involved, they are envisioned to associate into sorting platforms from which apical transport vesicles could form with the help of as yet unidentified apically directed putative coat proteins. Separate docking and fusion mechanisms that are unique either to the apical or basal membranes may provide a final proofreading process by which proper targeting is achieved (reviewed by Weimbs et al., 1997). V. SUMMARY Much is being learned about how plasma membrane composition and structure are regulated. The anatomy of vesicular trafficking has been described reasonably well, and the biochemical processes that achieve proper targeting of lipids and proteins are beginning to be elucidated. Information within the structures of proteins is being discovered that helps to target them to or retain them in the endoplasmic reticulum, various portions of the Golgi, lysosomes, endosomes, or the plasma membrane. Moreover, investigators in this field are beginning to study how the cell deciphers this information. The mechanisms by which the cytoskeleton and its associated motor proteins help to move vesicles to the correct destinations are also being described as are the coat, docking, and fusion proteins that result in formation,

recognition, and fusion of vesicles with the correct target compartments. One of the most challenging problems in the field appears to be to describe the processes by which lipids and proteins are sorted to apical membranes of polarized cells, since sorting to this compartment appears to differ mechanistically from much of what is known about sorting and vesicular trafficking. Testable new hypotheses, such as the one involving glycolipid rafts in apical sorting, should help us to learn the mechanism of sorting to apical membranes. In addition, description and solution of new problems should inform us about how differential sorting occurs. For example, it will be interesting to learn how highly homologous subtypes of G A B A transport proteins are sorted to different regions of the plasma membranes of epithelial cells and neurons (Ahn et aL, 1996). After proteins reach their target membranes, their functions may be influenced both by the asymmetric distribution of phospholipids across the lipid bilayer and by the asymmetric fluidity of each leaflet in the bilayer. Similarly, whether a protein is in a liquid or a crystalline domain of the lipid bilayer could influence its function, which could conceivably be different in each of these lipid environments. It is, however, only now being determined whether these physically and chemically different environments have physiologically significant effects on the functions of integral membrane proteins. More appears to be known about the effects of the cytoskeleton on the functions of membrane proteins; the cytoskeleton not only influences the mobility and function of proteins that are tethered to it, but its meshwork below the membrane surface also appears to limit the ability of all other integral membrane proteins to migrate laterally in the lipid bilayer. Hence, progress in each of these areas of research continues to lead to revision of the original fluid mosaic model of membrane structure and to knowledge of the implications of the revisions to membrane function. In contrast to the emerging nature of our knowledge concerning modulation of the function of membrane proteins by membrane lipids and the cytoskeleton, much is already known about the biochemical actions of the proteins. The action of biomembrane transport proteins depends, of course, on the formation of a barrier to the free mixing of intracellular and extracellular constituents by the membrane lipid bilayer. Moreover, transport across the membrane barrier frequently results in or requires work. Since work can be viewed conveniently as the coupling between work-performing exergonic processes and work requiring endergonic ones, the next chapter appropriately concerns the thermodynamics of biomembrane transport.

C

H

A

P

T

E

R

3 I Thermodynamics and Transport

or molecules from a more ordered or confined to a less ordered or confined condition within a system. If the system is also closed to the passage of energy, then, of course, only entropy-driven processes are possible; the second law requires that entropy remain the same or increase in a closed system. When the universe, rather than a living cell or an organism, is considered to be the system, the distinction between entropy- and enthalpydriven processes is obscured. In this case, the enthalpydriven processes described above are instead entropydriven processes in which more condensed chemical potential energy in organic matter is converted to less condensed thermal energy. Nevertheless, it is sometimes useful to measure the enthalpy as well as the entropy changes associated with processes in living cells. These processes may have differences in the details of their mechanisms, and some of these differences are exposed when they are considered in relation to whether they are primarily enthalpy or primarily entropy driven. For these reasons, we will use both of these concepts to discuss the thermodynamics of membrane transport. In addition, even solely entropy-driven or entirely enthalpy-driven processes usually occur in association with an opposing change in enthalpy or entropy, respectively. Again, these concepts can be combined into one thermodynamic concept of the driving force of a process, the change in free energy (AG).

1. INTRODUCTION In order to live, cells must perform work and some of this work involves biomembrane transport. If the cells work to produce solute and solvent gradients across biomembranes, then the gradients may be dissipated to perform other work. The capacity of a solute or solvent gradient to perform work is, of course, greater when the gradient is further removed from equilibrium. For example, the degree to which taurine can contribute to a regulatory volume decrease by being released from a cell depends on the degree to which it has been concentrated in the cell against its chemical potential gradient. The amount of work that a solute or solvent gradient can perform can also be expressed in terms of the thermodynamic parameter free energy. Free energy changes usually involve changes in both enthalpy (heat content) and entropy (degree of disorder). Hence, a given chemical or physical process also may be described as either or both entropy and enthalpy driven. In living cells, most processes are primarily enthalpy driven, although some important processes, such as dissipation of ion gradients, are primarily entropy driven. The enthalpy-driven component of processes in cells of heterotrophic organisms usually results from the conversion of chemical potential energy present initially in ingested organic matter into thermal energy that is transferred to the environment. Such energy conversions are, of course, consistent with the first law of thermodynamics, which requires only that energy be conserved. The second law of thermodynamics underlies the component of spontaneous processes that is entropy driven. In the case of processes driven by solute or solvent gradients across biomembranes, the increase in entropy may be viewed primarily as the passage of ions

I!. SIMILAR MATHEMATICAL EXPRESSIONS SERVE FOR THE FREE ENERGY CHANGE IN A CHEMICAL REACTION AND IN THE MIGRATION OF A SOLUTE OR SOLVENT Detailed thermodynamic and kinetic assessments have been applied to biomembrane transport data in numerous excellent papers and books (e.g., Stein, 1986;

39

40

3. Thermodynamics and Transport

Hille, 1992). These assessments become, of course, increasingly complex mathematically as the number of different substances that are transported in the complete process increases. Additional complexity is introduced when transport is coupled to chemical change. Hence, the assessments are organized in some of these writings in an increasing order of mathematical complexity, a In the present volume, however, we discuss these processes in reverse order of mathematical complexity in several chapters including this one. Although presentation of transport processes in reverse order of their mathematical complexity may seem in some ways perverse, they are organized in this manner to reflect important relationships among the transport processes. Much biomembrane transport in living organisms occurs because primary active transport produces and maintains solute and solvent gradients. These gradients serve subsequently either to produce and maintain gradients of other solutes or, when the gradients are dissipated, to do other work such as propagation of a signal along a biomembrane. Different types of transport can be distinguished by the details of the thermodynamic expressions that apply to them. Nevertheless, the thermodynamic expressions for overall transport processes are conceptually quite simple even for the most complex cases of primary active transport. Since we will discuss primary active transport ahead of several other types of transport in later sections, it is useful to consider how the mathematical expressions are similar for biomembrane transport and chemical reactions. Each of these processes occurs during primary active transport. As stated above, the driving force of any process is reflected by the change in free energy that is associated with it. When the free energy of a system decreases, the process producing the decrease in free energy proceeds spontaneously. In contrast, when a process would produce an increase in free energy it proceeds instead in the reverse direction. All components of a system contribute to its free energy content, and these components can undergo changes within the system as long as the changes result in a decrease in the free energy of the system. As we shall see below, the free energy of a component of the system frequently may even increase as long as it occurs in association with a larger decrease in the free energy of another component. The free energy of a component, A, of a system is known as its chemical potential. The chemical potential aThe mathematical complexity of the thermodynamic or kinetic representation of a transport process need not, however, reflect the relative complexity of the mechanism by which a solute or the solvent actually migrates across the membrane via a transport protein. Too little is as yet known about these mechanisms of migration to draw conclusions about the relative degrees of their complexity.

of A is defined as the amount by which the free energy of a system is increased or decreased by the addition or loss, respectively, of one mole of this component. 2 Since the chemical potential of A is both a measure of how much A a system contains and how much work may be associated with the loss or gain of it, the chemical potential of A is expressed as calories or joules per mole. In the cases of biomembrane transport and chemical reactions in living cells, component A is frequently a solute dissolved in water. For this reason, it is usually more convenient to express quantities of solutes in relation to the volume of solvent in which they are dissolved (i.e., their concentrations). The relationship between the chemical potential of A and its concentration can be stated approximately as ~1,A ~ /Li,~ -}- R T

ln[A],

(3.1)

where/-/,A and/X~ are the chemical potential and standard chemical potential of A, respectively, R is the gas constant and T is the temperature in ~ Although molal rather than molar concentrations are preferable because the former are independent of temperature, the common use of molar concentrations does not introduce appreciable error. In this case, the standard chemical potential becomes the chemical potential of a 1 M solution of A. While the standard chemical potential is defined as 1 M, the reader is reminded that the chemical potentials of much less than one mole of a solute would actually be measured in the relatively dilute solutions and at the volumes of most pertinent biophysical experiments. Now the statement of the approximate chemical potential of A in Eq. (3.1) must be corrected for any factor that causes its behavior to deviate from ideal. When the cohesive forces among substances in a solution are not completely uniform, another expression of concentration, the activity of substance A, can be calculated from its molar concentration as aA = 3/ [A],

(3.2)

where aA is the activity of A in the same units as its concentration and y is the activity coefficient. As the concentration of A decreases, its behavior approaches the ideal and y approaches a value of unity. The more precise expression of the chemical potential of A is written /d,A = /d,~

nt- R T

In aA,

(3.3)

where/X~ now represents the chemical potential of A at 1 M activity. From this simple equation, it should be 2The free energy of a component also depends on temperature and pressure, which are frequently nearly constant in biological systems. The importance of varying pressure in biological systems is considered in Section V of this chapter.

41

Mathematical Expressions of AG

clear that changes in chemical potential and free energy will be the same regardless of the process by which A is lost from (or gained by) the system. If A is lost from the system, its chemical potential and, hence, its free energy contribution to the system will decrease, whereas these parameters will increase if A is gained by the system. Moreover, the chemical potential of A in the system would change by the same amount regardless of whether its activity were to, say, decrease due to conversion of A to substance B in a chemical reaction or due to transport of A out of the system across a biomembrane (Fig. 3.1). The change in the chemical potential of the system can be calculated by subtracting the final chemical potential of A in the system, ~ A 2 , from its initial chemical potential, ~ A a . The mathematical expression for the change is /J'A1 -- /-~A2 =

(/-l'~

-'[-

RT

A G In aAa )

-[- R T

-- (/.l,~

In

aA2),

In (aA1/aA2).

(3.5)

~-~ (/A, B 1 -

In the case where A is consumed in a chemical reaction there is, of course, a concomitant increase in the contribution of B to the chemical potential of the system. A <-->B For a chemical reaction that occurs in solution at a constant volume and under isothermal conditions, the free energy change results, however, entirely from the loss in chemical potential of the reactants and gain in chemical potential of the products. The change in chemi-

/A,A2 )

(3.6)

It can be seen by inspection of Eq. (3.6) that A G would, indeed, have its most negative value when/zm is at its minimum and/ZA1 is at a maximum. This equation can be rearranged to give A G = R T In

A. Derivation of a T h e r m o d y n a m i c Expression for a Chemical Reaction

/A,B2) -- (/d, A a -

= R T In (aBa aA2/aB2 aA1).

(3.4)

where aAa is the initial activity of A in the system and aA2 is its final activity. Equation (3.4) can be simplified to /J, Aa -- 1%A2 ~-- R T

cal potential for the reaction in which A is converted to B can be represented by combining the contributions to the change in chemical potential of A and B. The expression for the change in chemical potential that is attributable to B is identical in form to that for A in Eq. (3.5). Furthermore, the total change in chemical potential is equal to the change in free energy, A G, when the changes in chemical potential are combined in a way that, by convention, produces a negative A G when the reaction proceeds spontaneously from A to B. Of course, A is most inclined to proceed to B when /d, A1 is at a maximum and/d, Ba is at a minimum. Equation (3.5) for A and an identical expression for B can, hence, be combined to produce the total change in free energy as follows

(aA2/aB2) + R T In (aBa/aA1)

or

A G = - R T In (aB2/aA2) + R T In (aB1/aA1).

(3.7)

Now the standard free energy change of a reaction, AG ~, is defined as the free energy change that occurs in association with the reaction as it proceeds from the standard state. Equation (3.7) can thus be made to represent AG ~ by setting the initial activities of A and B, aAa and am, equal to their standard states (i.e., 1 M each). In addition, if the reaction is permitted to proceed to equilibrium, then the final activities of A and B will, of course, represent this equilibrium Keq--

aB2/aA2.

Hence, AG = AG ~ = - R T

In (aB2/aa2) + R T In (1 M/1 M),

which simplifies to the familiar form AG ~ =

- R T In Keq.

(3.8)

Likewise, the thermodynamic Eq. (3.7), for the chemical reaction, can now be combined with Eq. (3.8) to produce the familiar form AG = AG ~ + R T In (aB1/aA1). FIGURE 3.1 The decrease in the contribution of A to the free energy content of a system is the same regardless of whether it is lost through conversion to B in a chemical reaction (1) or due to transport out of the system (2).

(3.9)

The same equation can be derived less explicitly but more directly beginning with Eq. (3.3) and an expression of the same form for substance B

42

3. Thermodynamics and Transport /J'A = ~ ~

-{- R T

In

(3.3)

aA

MEMBRANE

and COMPARTMENT A

/xB = IZ~ + R T I n aB.

-" COMPARTMENT B

In this case, the change in free energy is equal to the difference between the chemical potentials of the product and the reactant AG

= /%B-

/~A = ~tJ~~

/-J~~

+ R T In aB -- R T In aA.

CA

(3.10)

..d

:

CB

9

Since AG ~ =

/.~~B -

].j~~A

Eq. (3.10) simplifies to the familiar form AG = AG ~ + R T In (aB/an),

(3.11)

which is the same as Eq. (3.9) where a~ and aA in Eq. (3.11) are equivalent to aB1 and aA1 in Eq. (3.9). It should also be noted that there is no need to indicate in the chemical reaction A ~--~B

FIGURE 3 . 2 Movement of substance C between compartments A and B of a system through a membrane. In order for the example to be precisely parallel to the example for a chemical reaction in which the volume of the system remains constant (Fig. 3.1), the volumes of compartments A and B must be equal (see text for full discussion).

that other products, such as energy, are lost or gained. Specifically, in the reaction A ~ B + heat the term "heat" is unnecessary. The heat is accounted for by the enthalpy change, AH. Change in enthalpy is, however, a component of the free energy change that is used to represent the thermodynamics of the reaction. That is, the change in free energy is fully represented by AG

= AH-

TAg

(3.12)

tion makes it unnecessary in practice to be concerned with the volume of compartments in most biophysical experiments.) In the present example, substance C is designated CA when it is in compartment A and CB when it is in compartment B in order to remind us that we are comparing the chemical potential of C in compartment A to its chemical potential in compartment B. In order to change Eq. (3.7) for the reaction A +--~B

as well as by AG = AG ~ + R T In (aB1/aA1),

(3.9)

where AS is the entropy change and Tis the temperature in ~ B. Parallel T h e r m o d y n a m i c Expression for Migration of a Solute Expressions entirely parallel to Eq. (3.9) and (3.11) can also be derived for a system in which the substance C migrates from compartment A of the system past or through a barrier, possibly a membrane, into another compartment, B, of the system (Fig. 3.2). To make the present derivation entirely parallel to that above for a chemical reaction at a constant volume, the volumes of compartments A and B must, for the present, also be equal. (Activities with units of concentration are, of course, quantities of solutes normalized in regard to volume. Expression of activities in units of concentra-

to an equation for the migration of C from compartment A to B (Fig. 3.2), one need only substitute CA for A and CB for B as follows AG

= - R T In (aCBZ/aCA2) + R T In (aCB1/aCA1).

(3.13)

In the case of simple migration of a solute that is not coupled to another process, the activities of C in compartments A and B are equal when the system is permitted to reach equilibrium. Since at equilibrium aCA2 -- aCB2

the standard free energy change is zero AG ~ = -RT

In (aCB2/aCA2) = O.

Hence, the thermodynamic expression for migration of C between compartments A and B in Eq. (3.13) simplifies to AG

= R T In

(aCB1/aCA1).

(3.14)

43

Contributions of Enthalpy and Entropy Change to AG

Additional considerations frequently must be made when developing thermodynamic expressions for migration of real molecules and ions across biomembranes. For example, the electrical potential across the membrane that results from the combined influences of permeant and nonpermeant ions on both sides of the membrane influences the bioenergetics of transport of all permeant ions. Before we discuss in Section IV below the effects of transmembrane electrical potential on transport, however, it will be useful to consider how the similar thermodynamic expressions for transport and a chemical reaction may nevertheless differ. As discussed above, we will consider the thermodynamics of various types of transport in decreasing order of mathematical complexity, mainly because the bioenergetically most complex of these transport processes, primary active transport, creates solute gradients that make most other forms of transport possible. Although the thermodynamic expressions take the same form for the chemical reaction and the transport that occur during primary active transport, the relative magnitude of the enthalpy and entropy changes associated with chemical change and migration of a solute may differ.

i!I. CHANGES IN ENTHALPY AND ENTROPY MAY CONTRIBUTE DIFFERENTLY TO THE FREE ENERGY CHANGES ASSOCIATED WITH A BIOCHEMICAL REACTION AND MIGRATION OF A SOLUTE Simple qualitative consideration of the two systems discussed above for a chemical reaction and a migration of a solute leads, perhaps first, to the notion that they are energetically different rather than that they can be represented by similar thermodynamic expressions. This insight arises, of course, from the simple realization that reactions and transport are clearly different processes. Unlike chemical reactions, almost all transport processes do not appear even to involve transient formation

TABLE 3.1

of chemical bonds between the transported solute and another substance or the transport protein. For these reasons, chemical reactions and solute or solvent transport may also differ in the relative contributions which enthalpy and entropy changes usually make to the thermodynamic expression for either process AG

:

AH-

(3.12)

TAS.

Chemical reactions in living cells are more frequently primarily enthalpy driven (Table 3.1). In contrast, migration of a solute from a place of greater to a place of lower chemical potential is usually mainly entropy driven. Hence, reversal of this primarily entropy-driven process is typically coupled to the mainly enthalpydriven process of hydrolysis of phosphoric acid anhydride bonds in primary active transport. In similarly coupled processes involving hydrolysis of high-energy phosphate bonds, microtubules and actin filaments are made to depolymerize. Since polymerization is, in each case, an entropy-driven process (e.g., Lauffer, 1983), hydrolysis of GTP (microtubules) or ATP (actin filaments) to their corresponding nucleoside diphosphates and Pi reverses entropy-driven processes. This reversal of entropy-driven processing is analogous to the spontaneous migration of a solute with its chemical potential gradient which is reversed in primary active transport (e.g., Table 3.1). Conversely, the exchange of GTP or ATP for the GDP or ADP bound to tubulin or actin monomers, respectively, reverses the conformational change that results from GTP or ATP hydrolysis and, hence, favors polymerization. Such conformational changes appear also to be involved in the primary active transport catalyzed by P-type ATPases. In these cases, however, hydrolysis of the nucleotide results in temporary phosphorylation of the enzyme during the transport cycle. Nevertheless, ATP hydrolysis is associated with a conformational change in the enzyme that results in the transport of solutes against their chemical potential gradients (Vasilets and Schwarz, 1993; Chow and Forte, 1995; Andersen

Principal Contribution to the Decrease in Free Energy ( - A G ) of Several Types of S p o n t a n e o u s Processes in Living Cells W h e r e A G = A H - T A S Contribution to AG (+ or - )

Type of spontaneous process

Example a

1. Enthalpy driven 2. Primarily enthalpy driven 3. Primarily entropy driven

Asn + H20 <--->Asp- + NH+4 ATP 4- + H20 <--->ADP 3- + Pi 2- -~- H + Migration of a solute along its total chemical potential gradient (e.g., see Fig. 3.3) Polymerization of tubulin to form microtubules

4. Entropy driven

aAt physiological pH, temperature, and pressure.

AH

- TAS

z~/vs -

TAS

+

AH

<

- TAS

-

AH

<

- TAS

-

-TAS

< AH

-TAS

< AH

44

3. Thermodynamics and Transport

and Vilsen, 1995) in a manner analogous to reversal of the entropy-driven processes of tubulin and actin polymerization. In contrast, the binding of ATP to the enzyme favors the reversal of the conformational change that occurs on nucleotide hydrolysis, again analogous to conformational changes associated with GTP and ATP binding to tubulin and actin dimers and monomers, respectively. In some cases, the reversal of the conformational change resulting from ATP binding to P-type ATPases also is associated with transport of K + into the cytosol against its chemical potential gradient. In the case of the active K + transport by P-type ATPases, however, the free energy increase for K + transport is considerably smaller than the free energy increase associated with transport of other solutes in the reverse direction (Table 3.2). Hence, nucleotide hydrolysis is, in each case, associated with reversal of the otherwise most exergonic entropy-driven component of the overall process of microtubule polymerization, polymerization of actin filaments and migration of solutes transported by P-type ATPases. The amino acid residue sequences of the tubulin, actin, and P-type ATPase proteins show no homology. Hence, they appear to represent convergent evolution of homologous mechanisms for the reversal of entropydriven or primarily entropy-driven processes. It should be interesting to learn whether evolution of processes involving hydrolysis of phosphoric acid anhydride bonds by homologous proteins will all tend to be associated with reversal of primarily entropy-driven processes when one of them is so associated. In this regard, the amino acid residue sequence of tubulin shows some homology to that of hexokinase, which undergoes conformational changes while it catalyzes coupling of nucle-

TABLE 3.2 The A G for Extrusion of Inorganic Cations Is M u c h Higher Than the A G for U p t a k e of Cations in Cases W h e r e P-type ATPases Catalyze both U p t a k e and Extrusion of Cations a A G for the indicated process (kJ mo1-1) Enzyme b

Na+or H + extrusion

K + uptake

Total

Na+K+ATPase H+K+ATPase

34.2 42

7.6 2.5

41.8 44

aExamples calculated in Chow and Forte, 1995). bin the case of Na+K+ATPase, 3 Na + are extruded and 2K + are taken up by cells. In contrast, only 1 H + is extruded and 1 K + taken up in each cycle catalyzed by H+K+ATPase. The number of ions moved per cycle and per ATP hydrolyzed is lower in the case of H+K+ATPase primarily because the activity gradient against which H + is extruded is about 105-fold greater than the activity gradient against which Na + is extruded, whereas the free energy available to do work as a result of to ATP hydrolysis is about the same in each case.

otide hydrolysis to reversal of a reaction that is, however, entirely enthalpy driven (i.e., category 1 in Table 3.1). Nevertheless, the current example indicates that the association between hydrolysis of high-energy phosphate bonds and the reversal of primarily entropydriven processes has evolved on several apparently independent occasions.

IV. THE TOTAL CHEMICAL POTENTIAL CHANGE FOR A TRANSPORT PROCESS ALSO MAY HAVE AN ELECTRICAL COMPONENT Primary active transport processes frequently move ions across electrically polarized membranes. For this reason, we must derive a thermodynamic expression that includes the effect of membrane electrical potential on ion transport before considering the thermodynamics of coupling between ion transport and hydrolysis of ATP. Even for the simplest possible type of migration of electrolytes with their chemical potential gradients, the thermodynamic expression derived above must be expanded because electrical potential differences develop prior to equilibrium. A. The Migration of Ions Has an Electrical C o m p o n e n t The migration of cations and anions in water involves both chemical potential differences and electrical potential differences. For example, the movement of Na + ions and C1- ions from the 0.15 M NaC1 solution depicted in compartment A of Fig. 3.3 to pure water in compartment B proceeds, at first, unequally. Because the mobility of C1- in water is about 50% greater than the mobility of Na+, more C1- than Na + moves initially from compartment A to compartment B. Hence, compartment B takes on a net negative charge which accelerates the movement of Na + and slows the movement of C1- into compartment B (Hille, 1984). The resultant electrical potential can be added to the chemical potential (Eq. (3.3)) to give the total chemical potential of Na+,/ZtNa+A, in compartment A after slightly more C1- than Na + has :moved to compartment B. In this case, the total chemical potential for Na + in compartment A is ~ZtNa+A -- j(LNa+A nt- ZNa+ u~'%IFA,

(3.15)

where /ZtNa+Ais the total chemical potential of Na + in compartment A,/ZNa+A is the chemical potential of Na + in A, Zya+is the number of electrical charges on a sodium ion (i.e., + 1), S i s the Faraday constant (96.5 kJ volt -1 equivalent-l), and ~A is the electrical potential of com-

45

Electrical Component of Total Chemical Potential

to be reduced somewhat from its chemical potential in this compartment owing to the - 1 value of z o /J, tC1-A = /d,~ _ -[" R T

FIGURE 3 . 3 Migration of Na + ions and C1- ions across any interface between a 0.15 M NaC1 solution (compartment A) and water (compartment B). In this case, a m e m b r a n e is positioned at the interface and hence the compartments are separated by a finite distance. The presence of the m e m b r a n e does not, however, alter the thermodynamic expression for migration of Na § and C1- from A to B as long as the m e m b r a n e is equally permeable to both ions. Since the mobility of a C1- ion is about 50% greater than the mobility of a Na § ion, compartment B immediately takes on a transient net negative charge and compartment A acquires a transient net positive one.

partment A relative to an arbitrary zero point. Since, according to Eq. (3.3) /d,Na+ A = /d,~

-}- R T

In aNa+A ,

(3.16)

where aNa+A is the activity of Na + in compartment A and/X~ is the standard chemical potential of Na+, Eq. (3.15) can be expanded to /d,tNa+A = /d,~

-Jr- R T l n aNa+A -}- ZNa+ ~"a~/'A.

(3.17)

To simplify our consideration of the present case, the arbitrary zero to which XFAis compared may be set equal to the charge in compartment B (which is actually negative). Now ZNa+ is, of course, positive and the electrical potential due to the greater mobility of C1- makes compartment B slightly negative and compartment A slightly positive. Hence, it can be seen from Eq. (3.17) that at the time we are observing the system, the total chemical potential for Na + in compartment A will be somewhat higher than its chemical potential in this compartment. Conversely, the total chemical potential of C1- in compartment A will be somewhat less than its chemical potential in this compartment. This somewhat lower total chemical potential for C1- in compartment A can be seen from an expression similar to Eq. (3.17)

In a c l _ A + ZC1-~'XI/'A.

(3.18)

When the two ions reach equilibrium across the barrier represented in Fig. 3.3 then there will, of course, be no electrical or chemical potential difference between the compartments. The total chemical potential of each ion on each side of the barrier will be equal to its total chemical potential on the other, assuming that the volumes of compartments A and B are equal. In addition, there will be no electrical potential contribution to the total chemical potential of either ion on either side of the barrier at equilibrium. A more interesting equilibrium can, however, be established if a permanent electrical potential difference is established across the membrane. In this case, the activity and the chemical potential but not the total chemical potential will differ for each ion on each side of the membrane at equilibrium. These differences in the activities of Na + and C1- from the condition in which there is no permanent electrical potential difference across the barrier can be produced simply by introducing impermeant ions of one charge or the other into one of the compartments depicted in Fig. 3.3. Introduction of an impermeant anion or cation means, of course, that the barrier is now more permeable to some solutes than to others, as is the case for most biomembranes. The changes in the equilibrium activities of Na + and C1- that are produced by introducing an impermeant cation or anion into one compartment is known as the GibbsDonnan effect or, in some texts, simply the Donnan effect. B. The G i b b s - D o n n a n Effect Produces M e m b r a n e Electrical Potential The Gibbs-Donnan effect can be illustrated by introducing into compartment B of the two-compartment system shown in Fig. 3.3 a second sodium salt, the anion of which is unable to cross a membrane separating the two compartments (Fig. 3.4). The impermeant anion in Fig. 3.4 was made multivalent in order to resemble cellular constituents, such as nucleic acids and proteins. As we shall see, the effects of the impermeant anion on the distributions of Na + and C1- across the membrane are not influenced by whether the impermeant anion is monovalent or multivalent. The valence of the impermeant ion will, however, significantly influence the osmotic pressure across the membrane once the Gibbs-Donnan equilibrium is reached (see Section V below). Assuming that the volumes of compartments A and B remain equal, the equilibrium that is established by migration of Na + and C1- across the membrane in Fig.

46

3. Thermodynamics and Transport

In (aNa+AlaNa+B)= ZNa+

uJ-('tI/B --

gkrA)lRT

(3.23)

and for C1In (aCl-A/aCl-B) = ZCl-~J(~B -- ~ A ) / R T

(3.24)

Now, since ZNa+ = +1 and zcl- = - 1 , we can write the equation 5 ( ~ B -- ~A)/RT = In

FIGURE 3 . 4 A system in a vessel composed of two compartments of equal volume, a semipermeable m e m b r a n e between the compartments, and the indicated solutions in each compartment. The membrane is permeable to Na +, CI-, and water but not to A 15-. The NalsA dissolved in compartment B could be the sodium salt of a protein or another highly soluble polyvalent anion. For purposes of illustration, 15 Na + or 15 C1- can be taken as equivalent to an activity of 0.15 M of each of these ions.

3.4 can be represented by the equations for the total chemical potential of Na § and C1- in each of the compartments. These equations are for compartment A /./,tNa+A = /.~~ + -1/./,tCl_ A :

~t/,~ _ "at-

R T In

RT

aNa+A + ZNa+ 5 a l r A

I n aC1-A + Z C 1 - g a t r A

(3.17) (3.18)

and for compartment B /.s

= /.s176 + +

/xtCl_B = /x~

R T In

aNa+B +

ZNa+OJ"aI$B

+ R T In aCl-B + Z C l - g ~ B

(3.19) (3.20)

At equilibrium, however, and /./,tCl_ A = /./,tCI_B.

Hence + nt-

R T In = /X~

aNa+A + ZNa+ u--~"alCA

+ R T In aNa+B + ZNa+ 5altB

(3.21)

and /z~

+ R T In aC1-A + ZC1-5at)'A = /x~ + R T In aCl-B + Z O - f ~ B ,

(3.22)

which simplify to forms of the familiar Nernst equation for Na §

In

(aCl-B/aCl-A).

(3.25)

Using Eq. (3.25) several important consequences of the Gibbs-Donnan effect can be explained. For purposes of illustration, it will be assumed that the numbers of mobile Na § and C1- ions depicted in Fig. 3.4 actually represent their activities. (For readers who prefer to use actual units of concentration, 15 Na + or C1- in Fig. 3.4 can be equated to 0.15 M activity of either ion.) It is also assumed that the migration of water has been maintained at equilibrium owing to application of enough pressure to compartment B to keep its volume constant (see Section V below). The distribution of Na + across the membrane in Fig. 3.4 that nearly fitS the requirements of Eq. (3.25) is 27 Na + in B and 18 Na § in A (Fig. 3.5). This distribution would mean, from Eq. (3.25), that compartments A and B would contain 18 and 12 CI-, respectively. In this case, however, the total positive charge in A(+18) equals the total negative charge in this compartment (-18). Likewise, the total positive and negative charges in compartment B are both 27. Thus, the membrane potential would be zero, which contradicts Eq. (3.25). If the activity gradients for Na § and C1- were greater than the values listed above, then there would be a net positive charge in compartment B and a net negative one in A. There is, of course, no mechanism by which introduction of the impermeant anion into compartment B could, by itself, establish a net positive charge on that side of the membrane. Hence, in the present example aNa+AlaNa+B =

/.ltNa+A --- /d,tNa+B

/~~

(aNa+A/aNa+B) ---

aCl-B/aCl-A >-- 2/3.

(3.26)

Note, however, that the more these ratios exceed 2/3, the more the membrane electrical potential calculated from Eq. (3.25) decreases in magnitude (compartment B is negative). In contrast, the membrane electrical potential as assessed by the actual numbers of charged ions present in each compartment increases in magnitude (compartment B is still negative) as the ratios exceed 2/3. For example, if, say, 24 instead of 27 Na § ions were present in compartment B, then, of course, 21 Na § would be in A, 16 C1- would be in A and 14 C1- would be in B. In this case, the total negative charge in B would be - 2 9 while the positive charge in this compartment would be + 24. Hence, the magnitude of the membrane electrical potential based on assessment of the

Gibbs-Donnan Effect Generates Osmotic Pressure

47

m e m b r a n e potential that is p r o d u c e d in living cells by only a slight gradient in total charge (roughly 1 part in 1000; Hille, 1992) nevertheless contributes a b o u t as much to the total chemical potential difference of N a § ions or K § ions across the m e m b r a n e as do the m o r e than 10-fold differences in their activities and concentrations.

V. T h e G I B B S - D O N N A N EFFECT A L S O GENERATES O S M O T I C PRESSURE

FIGURE 3.5 The system in Fig. 3.4 after it reaches the equilibrium defined by Eq. (3.25). Although the total numbers of negative and positive charges are shown to be equal on both sides of the membrane, compartment A is actually slightly more positive than negative, whereas the reverse is true for compartment B. For example, it can be assumed that the compartments each have a volume of 1 ml, the membrane has an area of 1 cm2 and the 18 Na + in A, 27 Na + in B, 12 C1- in B, and 18 CI- in A actually represent activities of 0.18, 0.27, 0.12, and 0.18 M, respectively. In this case, the ratios of 0.18 M/0.27 M for Na + and 0.12 M/0.18 M for C1- between the two compartments need to be offset by about one billionth in order to produce the transmembrane electrical potential of -10 mV that must be present according to Eq. (3.25). The pressure on the solution in compartment B also must be higher than that on the solution in A to produce equilibrium (see Section V of the text).

actual charged ions increases from zero as the ratio exceeds 2/3, w h e r e a s the m a g n i t u d e of this electrical potential should decrease according to Eq. (3.25). T h e solution to this a p p a r e n t p a r a d o x is that the ratios of the N a + and C1- activities in Eq. (3.26) exceed 2/3, but only by a very small amount. Ratios of 2/3 can be used in Eq. (3.25) to calculate an electrical m e m b r a n e potential of - 1 0 m V at 298~ ( S = 23,061 cal V -1 equiv -1 and R = 1.987 cal deg -1 mol-1). It can be calculated for the m e m b r a n e depicted in Fig. 3.5 (assuming that 15 N a + or C1- = 0.15 M activity of these ions) that the 2/3 ratio would n e e d to be offset by only about one billionth ( 1 0 -9 ) t o p r o d u c e an electrical m e m b r a n e potential of 10 m V if the m e m b r a n e has an area of 1 cm 2 and the v o l u m e of each c o m p a r t m e n t is 1 ml (see similar calculation for living cells in Hille, 1984). Conversely, the r e a d e r may also now begin to appreciate that very small total ion activity gradients p r o d u c e electrical m e m b r a n e potentials that have relatively large effects on the total chemical potentials of ions. For example, we show in Section V I , A below that the electrical

In addition to the generation of an electrical potential difference across m e m b r a n e s , a n o t h e r c o n s e q u e n c e of the G i b b s - D o n n a n effect is to p r o d u c e osmotic pressure. Osmosis is defined here as the migration of water from a place of higher total chemical potential of water to a place of lower total chemical potential of water through a m e m b r a n e p e r m e a b l e to water but not all solutes. 3 As for solutes, the free energy of water in a c o m p a r t m e n t can be expressed as its chemical potential, /Xs, w h e r e /Xs =/X~ + R T l n as.

(3.27)

Since water molecules are u n c h a r g e d (z = 0), the m e m b r a n e potential does not directly influence its total chemical potential. Nevertheless, by influencing the total chemical potential of anions and cations in the system, the m e m b r a n e potential can have a significant indirect effect on the total chemical potential of the solvent. F o r the example illustrated i n Figs. 3.4 and 3.5, it can be seen that at equilibrium of the solutes, the total n u m b e r (or activity) of dissolved solute particles in c o m p a r t m e n t B is 27 Na + + 12 C1- + 1 A 15- = 40, whereas the total in compartm e n t A is 18 Na + + 18 C1- = 36 (Fig. 3.5). This difference is due both to the i m p e r m e a n t anion and to its consequences for the distributions of Na + and C1- across the m e m b r a n e . T h e difference in the total activities of solutes in c o m p a r t m e n t s A and B also m e a n s that the system r e p r e s e n t e d in Fig. 3.5 is not yet at equilibrium in regard to the solvent. Let's assume that no force is at work to maintain equal volumes in c o m p a r t m e n t s A and B. In this case, we could a t t e m p t to allow the system to reach equilibrium by allowing water to m o v e from c o m p a r t m e n t A to c o m p a r t m e n t B, thus making the volume of B g r e a t e r than A. T h e r e is, however, an i m p e r m e a n t ion in com-

3The reader may notice that we are careful not to state or imply that water or relatively small hydrophilic solutes actually migrate through most biomembranes by ordinary diffusion. Nevertheless, the thermodynamic expression for the overall process is not altered if it is assumed that water moves between compartments A and B by that simple process. Similarly, our convenient assumption here that a membrane may be permeable to water or solutes should not be taken to mean that permeation occurs by diffusion.

48

3. Thermodynamics and Transport

partment B, and its presence there also results in unequal distributions of permeant ions between compartments A and B (see above). If water moved from compartment A to compartment B, it would tend to concentrate the solutes in A and dilute them in B (Fig. 3.6). To reestablish the appropriate ratios of the activities of Na + and C1- according to Eq. (3.25), each of these ions would also move from A to B. Hence, if water were allowed to move freely from A to B until it reached equilibrium, it would continue to do so until all of the water and permeant ions had moved from A to B. To maintain constant volumes of compartments A and B, it is necessary to establish an equilibrium by allowing pressure, rather than volume, to change. Pressure (P) and volume (V) can be introduced into the expression for the total chemical potential of water, txts, by expanding Eq. (3.27) as follows

constant pressure that has been assumed until now to be adequate for our considerations of total chemical potential. For the system depicted in Fig. 3.5, it is possible to stop the net migration of water molecules from A to B and establish equilibrium by applying enough pressure to the solution in compartment B to stop the net migration of water. At equilibrium

where /.LtSA and lxtsB are the total chemical potentials of water in compartments A and B, respectively. Similarly, we can write from Eq. (3.28) and (3.29) /X~ + R T l n aSA + V s ( P A - P o ) = ~~ + R T l n aSB + Vs (PB - Po), which simplifies to I

PA-

PB = ( R T / V s ) ( l n

I

~ts = ~t/'S -+" Vs (P - Po)

and I

/.Lts = /.L~ -t- R T l n as + Vs (P - Po),

(3.28)

where Vs is the volume occupied by one mole of water, P is the pressure applied to the solution, and Po is the

(3.29)

/.LtSA = ~tSB ,

(asB/aSA)),

(3.30)

where the subscripts A and B indicate that the parameters are for the solutions in compartments A and B, respectively. The difference between PA and PB is the amount by which the pressure on compartment B must exceed that on A to stop the net migration of water. More generally, equations such as Eq. (3.30) can be developed for any water solution in comparison to pure water to define the osmotic pressure, 7r, of the solution. The osmotic pressure of a solution is defined here as the amount of pressure that must be applied to the solution in order to stop the net movement of water into it from a compartment of pure water through a semipermeable membrane. For the solutions in compartments A and B of Fig. 3.5, respectively ~'A : ( R T / V s ) ( l n

(aw/asA)),

and 7rB = (RT/Vs)(ln (aw/asB)), where aw is the activity of pure water. Hence, for the system in Fig. 3.5 involving compartments A and B 7rA -- 7rB = ( R T / V s ) ( l n

FIGURE 3.6 Effect of the equilibrium depicted in Fig. 3.5 on the net migration of water (represented by the solid arrows) between the compartments. Because the total activities of solutes in compartment

B are greater than in compartment A, the activity (and total chemical potential) of water is greater in A than in B. Hence, water and, as a result of the water movement, Na+ and C1- would all tend to move from A to B. The movement of water and ions from A to B would continue until compartment A disappears, unless enough pressure is applied to the solution in B (in excess to that applied to the solution in A) to stop the net migration of water.

(asB/aSA)).

(3.31)

The difference between the osmotic pressures of the two solutions, ~'A -- 7rB, is, o f course, the degree to which the pressure that is applied to the solution in compartment B must exceed the pressure on the solution in compartment A in order to stop the net migration of water molecules from A to B, which is the same as the difference between P A and PB in Eq. (3.30). Another way to stop the net m o v e m e n t of water from A to B without applying different pressures to the two solutions is to establish a steady state rather than an equilibrium. In this case, pressure and volume can again remain constant while the excess of permeant ions in

49

Chemical Reactions Drive Primary Active Transport

compartment B is reduced by their active transport to compartment A. Active transport of C1- from B to A would not only tend to equalize the osmotic pressures of the two solutions, but it would also reduce or reverse the membrane electrical potential. In contrast, the active transport of Na + from B to A would both counteract the difference in osmotic pressure and increase the existing membrane electrical potential. Active extrusion of Na + from compartment B is analogous to the action of Na + K+ATPase and other such primary active transport processes in the plasma membranes of most cells. In the case of Na+K+ATPase, the extrusion of more monovalent cations from the cell than are taken into the cell from the extracellular fluid increases the inside negative electrical potential across the membrane. In addition, this transport by Na+K+ATPase helps to reduce the cellular swelling that would otherwise occur as a result of the GibbsDonnan and other effects (see also Section X below).

VI. CHEMICAL REACTIONS DRIVE PRIMARY ACTIVE TRANSPORT

A. C o m p a r i s o n s of the Free Energies of Cation Transport and ATP Hydrolysis That Are Catalyzed by Na+K+ATPase In contrast to the hypothetical example just described, the concentration and activity gradients of most solutes across the biomembranes of living cells usually lie in the same direction as their total chemical potential gradients across these membranes. Hence, in most real examples of primary active transport, solutes are moved against their concentration and activity gradients as well as against their total chemical potential gradients. In the case of Na+K+ATPase, the endergonic processes of Na § extrusion from cells and K + uptake by them are driven by the exergonic process of ATP hydrolysis to ADP and Pi. Separate thermodynamic expressions can be derived for each of these coupled processes. To derive an expression for the transport of Na +, the intracellular and extracellular compartments may be distinguished with "i" and "o," respectively. Equation (3.17) previously derived for a compartment that was designated A /d,tNa+A ---- /.s176 + nt- R T In aNa+A + ZNa+ ~ " ~I'tA

We have seen that the total chemical potential of a solute may depend on its charge, the electrical membrane potential, and other permeant and nonpermeant ions in the system. In the hypothetical example just described, the active transport of two Na + ions from compartment B to compartment A in Fig. 3.5 would reestablish osmotic balance. This endergonic process actually decreases the Na § ion activity gradient in this case. The total chemical potential of Na + is, however, also dependent on the membrane potential against which Na + must be moved in going from B to A. Hence, the active movement of Na + from B to A in Fig. 3.5 represents formation of a gradient of the total chemical potential of Na § such that

From Eq. (3.17) it can be seen that the movement of two Na + ions from B to A would increase /./,tNa+A by increasing both the R T l n aNa+A and the Zya+ S ~A terms -'1- R T In aNa+A + ZNa+ f

'(I)'A,

(3.17)

where aNa+A would increase as would the magnitude of ~A. Similarly, from Eq. (3.19) it is clear that p, tNa+B would decrease due to a decrease in both aNa+B and ~B (i.e., ~B would become more negative) /ZtNa+B "-- /d,~

may then be rewritten /.LtNa+i- /.s176 + nt- R T l n aNa+i + ZNa+f ~I/'i

(3.32)

for the total chemical potential of Na § inside cells and /d,t N a + o -

/d,~

q- R T In aNa+o -}- ZNa+ ~"XI~o

(3.33)

for the total chemical potential of Na § outside them. The total chemical potential gradient against which Na § must be moved and, hence, the free energy change for extrusion of Na § from cells can now be written as the difference between the total chemical potential in each compartment AGNa+ = A/d, tNa+o-i-- /[s

/d,tNa+i .

When combined in this way, Eqs. (3.32) and (3.33) simplify to

/d't Na+A ~ /d,tNa+B 9

/.LtNa+A -- fiI,~

(3.17)

q- R T In aNa+B q-- ZNa+ ~ - Xt)'B.

(3.19)

In primary active transport, such movement of solutes is driven by hydrolysis of phosphoric acid anhydride bonds in ATP.

AGNa+ : A/.s = R T l n (aNa+o/aNa+i) nt- ZNa+ ~ - (~Ito -- ~Ifi).

(3.34)

A similar expression can be derived for the total chemical potential gradient of K § across the plasma membrane. In this case, however, the equation is written to reflect the total chemical potential difference against which K § must be moved in order to be taken up by cells. Therefore, the equation for K § transport by Na+K+ATPase is AGK+ = A/d, tK+i_o--

RTln (aK+i/aK+o)

-k- ZK+ ~ (~Iti -- aI*o)

(3.35)

to reflect its movement in the opposite direction of Na + (compare Eqs. (3.34) and (3.35).

50

3. Thermodynamics and Transport

Finally, the thermodynamic expression for intracellular ATP hydrolysis must take into account the reactants and products of the following chemical equation ATP 4- + H 2 0 ~ ADP 3- + Pi 2- + H +

(3.36)

The thermodynamic expression takes the general form of Eq. (3.11) for a chemical reaction A G = AG ~ + R T l n (aB/aA)

(3.11)

AG~

A G = - R T In K e q nt- R T In (as/aA).

Most intracellular biochemical reactions occur, however, in the presence of buffers that maintain the pH value near 7. Moreover, these reactions occur in solutions that are dilute enough in regard to the solutes to allow us to assume that the activity and concentration of the solvent, water, has a large constant value of 55.5 M. Hence, neither the activity of the reactant, H20, nor that of the product, H +, in Eq. (3.36) is usually included in calculations of the values of the observed equilibrium constant, K'eq, the standard free energy change, AG ~ or the total free energy change, A G. While K'eq and AG ~ are marked with a prime to indicate that their values may differ from those that would be obtained when the activities of H20 and H + are included in the calculations, the total free energy change, A G, needs no such designation. If they have not previously done so, readers should satisfy themselves that omission of these activities in the calculation of AG ~ from K ' e q is balanced when the activities of water and protons are also omitted from the general expression, R T ln(aB/aA), thus rendering AG unaltered. Several other factors, such as ionic strength and the activities of metal ions, also influence the free energy of hydrolysis of ATP. One such factor, the Mg 2+ activity, is especially important since Mg 2+ forms complexes with ATP and ADP to form the substrates and products actually involved in intracellular biochemical reactions. The thermodynamic effects of the Mg 2+ activity on several such reactions has been studied in detail (Alberty, 1969). All such activities are, however, also assumed to remain constant for most calculations of the free energy of hydrolysis of ATP. The thermodynamic expression for ATP hydrolysis under physiological conditions can therefore be written d- R T l n

(aADpapi/aATP).

K'eq = [ADPleq

[Pileq/[ATPleq =

2.22 X 105M.

Moreover, AG~ can be calculated from an equation that resembles Eq. (3.8) to be

or when combined with Eq. (3.8)

AGATP = A G ~

the convenient assumption that the values of solute activities are near enough to the values of solute concentrations to allow us to use the latter values in calculations without significant error. Under these assumptions for the ATP phosphohydrolase reaction, K'eq can be calculated from the equilibrium concentrations of ATP, ADP, and Pi to be

(3.37)

Eqs. (3.34), (3.35), and (3.37) can now be used to calculate the free energy changes associated with each of the processes that are coupled bioenergetically by Na+K+ATPase. Since the solutions of solutes both inside and outside cells are relatively dilute, we can also make

= -RTln

g ' e q = - 31.7 kJ mo1-1

at 310~ (R = 8.314 J deg-lmol-1). From this value of AG ~ the actual free energy change at pH 7 and 310~ can be calculated from Eq. (3.37) for any cell in which the cytosolic concentrations of ATP, ADP, and Pi are known. For rat hepatocytes, these concentrations have been estimated to be 3.38, 1.32, and 4.80 mM, respectively. In this case Z~GATP = -31.7 kJ mo1-1

+ R r l n ((0.00132 M)(0.00480 M)/(0.00338 M))

AGATP = -31.7kJ mo1-1 + -16.2 kJ mo1-1 AGATP = -47.9 kJ mo1-1.

More generally, the free energy of hydrolysis under typical intracellular conditions can be calculated to range from about - 4 2 to about - 5 4 kJ mo1-1 (Chow and Forte, 1995). Similarly, Eq. (3.34) can be used to calculate the free energy of Na + extrusion from cells. Typically, the intracellular and extracellular concentrations of Na + are about 11 and 140 mM, respectively (Chow and Forte, 1995), and we are assuming that these concentrations are nearly equivalent to the activities of Na+. Moreover, the membrane electrical potential of cells is frequently about -0.05 V (inside negative). Under these conditions and assumptions AGNa+ = R T l n ([Na+]o/[Na+]i) "q- ZNa+ ~ ('tI'to -- 'tI)'i)

(3.34)

AGNa+ = R T In (0.140 M/0.011 M) -t-9 ZNa+ ~ " (-]-0.05 V )

AGNa+ = 6.56 kJ mo1-1 + 4.82 kJ mo1-1 AGNa+ = 11.4 kJ mo1-1,

where f is 96.5 kJ V -1 equiv -1, R is 8.314 J deg -1 mo1-1, and T is 310~ The stoichiometry of transport is, however, extrusion of 3 Na + and uptake of 2 K + for each ATP hydrolyzed under physiological conditions. Hence, the minimum free energy needed (or the work that must be done) to move 3 moles of Na + out of a cell per mole of ATP hydrolyzed is

Chemical Reactions Drive Primary Active Transport AGNa+T-- 3 X 11.4 kJ(mol of ATP hydrolyzed) -1

51

all types of transport ATPases, including the P-type, V-type, and F-type ATPases (see below and Chapter 5).

AGNa+T = 34.2 kJ(mol of ATP hydrolyzed) -1.

Finally, the minimum free energy needed to move 2 K + into the cell can be calculated using Eq. (3.35) and intracellular and extracellular K + concentrations of 140 and 5 mM, respectively (Chow and Forte, 1995). In this case AGK+ =

R T l n ([K+]i/[K+]o) -Jr- ZK+ ~ (XIri -- xI~ro)

(3.35)

AGK+ = R T l n (0.140 M/0.005 M) -}- ZK+ ~" (--0.05 V )

AGK+ = 8.59 kJ mo1-1 - 4.82 kJ mo1-1 AGK+ = 3.8 kJ mo1-1. Since 2 K § are moved per ATP hydrolyzed, the total free energy needed for uptake of 2 moles of K + per mole of ATP hydrolyzed is AGK+ w =

2 • 3.8 kJ(mol of ATP hydrolyzed) -1

A G K + w --

7.6 kJ(mol of ATP hydrolyzed) -1.

We can now compare in several ways the free energy changes associated with each aspect of the overall process catalyzed by Na+K+ATPase. First, compare the free energy needed to move a mole of K § ions and a mole of Na + ions against their concentration (and activity) gradient across the plasma membrane of a typical cell. Threefold more energy is needed for the movement of Na + in spite of the fact that its concentration gradient is not as steep as that for K § This difference between Na + and K + is, of course, due to the inside negative membrane electrical potential against which Na + must also move, whereas K + is pulled by the inside negative electrical potential against its concentration gradient. In fact, we will show in calculations made in Section X below that K § may in many cases be very near to its equilibrium predicted by the Oibbs-Donnan effect. Hence, we shall see that Na+K+ATPase is frequently more of a Na § pump than a Na+K + pump under physiological conditions. From the above calculations it can also be seen that the total free energy available from ATP hydrolysis (42 to 54 kJ tool -1) is somewhat greater than the energy needed for K § uptake and Na § extrusion (34.2 + 7.6 = 41.8 kJ(mol of ATP hydrolyzed)-l). One may, at first, gain comfort from such calculations performed here and by numerous other authors. Enough free energy to drive the endergonic transport processes is indeed available from the hydrolysis of ATP. The details of coupling of the free energy changes associated with transport and chemical reactions are, however, virtually unknown for

B. C o m p a r i s o n of H o w Well W e U n d e r s t a n d Coupling b e t w e e n Multiple Transport or Multiple Chemical Processes to H o w Well W e U n d e r s t a n d Coupling of Transport to a Chemical Reaction The earlier example of the coupling of movement of Na + with its activity (and concentration) gradient while C1- moved against such a gradient toward the GibbsDonnan equilibrium (difference between Figs. 3.4 and 3.5) makes intuitive sense. It is possible to reason, for example, that an electrical membrane potential develops when Na + begins to move across the semipermeable membrane with its concentration gradient. In this case, C1- also moves in the same direction toward the positive side of the membrane and in so doing creates an activity gradient of this ion. Similarly, we will find that the bioenergetic coupling of protein-mediated transport of two or more solutes frequently seems relatively easy to comprehend (e.g., see Sections IX and X below). We may also assume that we comprehend relatively well the bioenergetics of coupling between many chemical reactions, such as the two that may be viewed as constituting the reaction catalyzed by adenylate kinase ATP 4- + AMp2-<---~ 2 ADP 3-.

(3.38)

This reaction can, of course, be written as the combination of two reactions, one exergonic ATP 4- + H20 <--9A D P 3- + Pi 2- q- H +

(3.39)

and the other endergonic AMP 2- + Pi 2- + H + <---->ADP 3- + H 2 0

(3.40)

under usual physiological conditions. The overall reaction may be viewed as the hydrolysis of one phosphoric acid anhydride bond and the formation of another. In addition to these thermodynamic considerations under conditions that prevail in vivo, many of us comprehend the free energy contained in the phosphoric acid anhydride bonds of ATP as resulting from electrostatic repulsion among negative charges, resonance stabilization that is possible when the bond is hydrolyzed, consumption of a rare proton at pH 7, and the greater solvation that is possible after the bond is hydrolyzed. Hence, it is not believed to be a conceptual stretch for most of us to reason that roughly equal amounts of free energy might become available and then be consumed by the transfer of the phosphoryl group from ATP to AMP to produce 2 ADP. Formal calculations of the actual free energy changes for the overall reaction in Eq. (3.38) or

52.

3. Thermodynamics and Transport

for the combination of the two half reactions in Eqs. (3.39) and (3.40) again comfort us that the reaction would occur in the direction written in Eq. (3.38) under many physiological conditions. The bioenergetic coupling between transport and chemical reactions is, however, much more difficult to comprehend than the coupling between two transport processes or between two chemical processes. Our struggles to comprehend the bioenergetics of coupling between chemical and transport processes also probably exposes our failure to comprehend on a fundamental level all bioenergetically coupled processes. Of course, thermodynamics does not concern itself with how processes interact, so thermodynamics should not be expected to tell us how chemical reactions or transport may drive each other, only that they d o drive each other (Christensen, 1975). If, however, we attempt to assess the thermodynamics of the biochemical and biophysical steps of an overall process, such as primary active transport, we may gain some insight into the bioenergetics of how transport and a chemical reaction are coupled. C. Possible Mechanism for the Bioenergetic Coupling of ATP Hydrolysis and Cation Transport by Na+K+ATPase and Other P-type ATPases While the bioenergetics of coupling may still be mysterious, the chemical and physical processes by which ATP is hydrolyzed and ions are moved across biomembranes can, in many instances, be described in some detail. These detailed descriptions have generally been well accepted, although some investigators still challenge them (e.g., Myung and Jencks, 1995). We will continue here with a description of the chemical and physical changes catalyzed by Na+K+ATPase as an example of the processes catalyzed by P-type ATPases. The possible mechanisms of bioenergetic coupling catalyzed by F-type ATPases (e.g., mitochondrial ATP synthase) and V-type ATPases (e.g., those that acidify lysosomes), are discussed in Section VIII below, and the structures and functions of all of these types of ATPases are described further in Chapter 5. According to the extended reaction scheme for Na +K+ATPase of Albers (1967) and Post et al. (1969), the inward-facing E1 form of the enzyme binds both ATP and 3 Na + ions relatively tightly (Fig. 3.7). Hydrolysis of the bound ATP by the enzyme and the associated autophosphorylation leads first to occlusion of the 3 Na + ions and subsequently to a conformational change in the enzyme from the inward facing, E1 P form to the outward-facing EzP form (Vasilets and Schwarz, 1993). The still phosphorylated EzP form has a relatively low

/.~,,.~p. (2K)E2.,,

2KE2ATP~

/2/EK2PADP--/2KE2P I /T

I

/

14

3NaE2AT~I , 3NaE2/ l

2K/E1ATP 2~~1PADP /2KE, E1A]TP

III

/2KE, P

//

- E1P,~DP /I /

////

- 3NaE1----------(3Na)EIP

3Na!2P(DP 3N!E!P S E2conformations

i

q

E~conformations

i

FIGURE 3.7 Extended Albers-Post reaction scheme for Na+K + ATPase. Thick arrows depict steps in the transport-reaction cycle that appear to occur during normal functioning of the enzyme. Occluded cations are shown in parentheses (adapted from Vasilets and Schwarz, 1993 with permission from Elsevier Science).

affinity for the 3 Na + ions and, hence, releases them to the extracellular fluid. Binding of 2 K + ions to the phosphorylated EzP form occurs, however, with relatively high affinity, and dephosphorylation of the E2 form occludes the 2 K + ions from the external milieu (Fig. 3.7). Binding of ATP to the E2 form then favors a conformational change in the enzyme from the E2 to the E1 form. The K + ion has a relatively low affinity for the E1 form and, hence, is released to the cytosol, thus completing the transport-reaction cycle (Fig. 3.7). While all P-type ATPases undergo a similar transportreaction cycle, V- and F-type ATPases appear to function via physical and chemical mechanisms which are in detail quite different from P-type ATPases (see Chapters 5). Nevertheless, the same thermodynamic concepts are needed to attempt to understand all instances of bioenergetic coupling between a chemical reaction and a transport. In the case of Na+K+ATPase, what may be the free energy changes that occur during ATP hydrolysis, phosphorylation of the enzyme, occlusion of 3 Na + ions, and transition from the E1 P to the EzP conformation (Fig. 3.7)? One bioenergetically pertinent clue may come from a surprising observation by Klodos et al. (1994). They found that upward jumps in the NaC1 concentration in the medium greatly and transiently increases the rate of conversion of EIP to EzP and vice versa. These transient kinetic behaviors were not simulated well by relatively simple models, but they did fit a kinetic model that involved a phase change in the lipid surrounding

53

Chemical Reactions Drive Primary Active Transport

Na+K+ATPase proteins in the membrane. The authors suggested that one phase may stabilize and contain mostly E1P while the other phase may stabilize and contain mostly E z P (Klodos et al., 1994). Since biomembranes function at their melting temperatures and, hence, contain transient liquid and crystalline domains (see Chapter 2), we suggest that the conversion from the E~P to the E z P conformation results from melting of the crystalline lipid domain containing the enzyme. Conversely, freezing of the liquid domain surrounding the E2 conformation would in this model favor its conversion to the Ea conformation. Klodos and associates (1994) did not distinguish which Na+K+ATPase conformation might be favored in the liquid or crystalline phase, and their kinetic model did not appear to require such a distinction. We, however, favor the above model because lower temperatures favor the E~P conformation of Na+K+ATPase reconstituted in proteoliposomes (Fig. 3.8), whereas higher temperatures favor the E2P conformation (Yoda and Yoda, 1987). Yoda and Yoda also found that cholesterol in the membrane appears to interfere with the E~P to E 2 P conformational change (Fig. 3.9), and this cholesterol makes the lipid in the bilayer less fluid (e.g., Raynard and Cossins, 1991). In fact, it has been concluded from additional studies (summarized in Vilch6ze et aL, 1996)

80-

o

"ou" 60 _C

~O

~

E2 P

1

40 -

o

E*P 20

0

. . . . . . . .

)

I 5

I 10

.

I ..... 15

E1P

1 20

Temperature (oc)

FIGURE 3.8 Effect of temperature on the conformation of Na+K+ATPase. Percentages of the enzyme in each conformation are shown. E1P and E2P correspond to the inward- and outward-facing conformations of the enzymes that normally occur in biomembranes (see Fig. 3.7). E*P is an intermediate conformation that occurs in the reaction sequence EIP ~ E*P ~ E2P. Egg phospholipid containing 28 mole percent cholesterol was used to reconstitute the enzyme in lipid bilayers of proteoliposomes. (data from Yoda and Yoda, 1987 are replotted here).

E2P (15 ~

) /

( \\

E 60

~ _

E1P (3 ~) ,.%;

_

E*P (1.5~ ......

} 4o E2P

~

IL

/~/ //"-..~

\

...----'V ~,/

z_--- ~ - - 2 20

30 40 Mole % Cholesterol

E*P

x 50

FIGURE 3.9 Effect of the cholesterol content of the lipid bilayer on the conformation of Na+K+ATPase. Percentages of the enzyme in each conformation at 3~ and at 15~ are shown. See legend of Fig. 3.8 for further details. (data from Yoda and Yoda, 1987 are replotted here).

that cholesterol specifically eliminates cooperative phase transition of phospholipid in the bilayer. Hence, the EaP (and El) conformation appears to be favored when more of the membrane is in the crystalline phase, whereas the E z P (and E2) conformation seems to be favored when more of the membrane is in the liquid phase. Consistent with this model, n-alkanols may inhibit Na+K+ATPase (Hegyvary, 1973) because they increase membrane fluidity (e.g., Kutchai et al., 1980), lower the phase transition temperature of the lipid in the membrane (e.g., Hill, 1974), and, hence, cause the membrane bilayer to contain too much liquid phase lipid. Similarly, cholesterol (Raynard and Cossins, 1991) and monoketone anesthetics (Tanii, 1996) may reduce Na+K+ATPase activity because they shift the membrane composition to too high a proportion of the crystalline phase. At least two types of forces may stabilize crystalline lipid domains in membrane lipid bilayers. The lipid molecules in membranes may be attracted to each other both by hydrogen bonds between, say, oligosaccharide moities of glycolipids at the bilayer surfaces and by hydrophobic interactions in the interior of the bilayer (see Chapter 2). A comparison of the latent heat of

54

3. Thermodynamics and Transport

fusion (i.e., the enthalpy change associated with melting or freezing) of hydrogen bonded substances, such as water, or hydrophobic substances, such as benzene, and the free energy of hydrolysis of ATP indicates, however, that only on the order of about 10 lipid or glycolipid molecules in a crystalline domain might undergo a phase change as a result of hydrolysis of one ATP molecule. Since crystalline lipid domains appear to contain on the order of 1000 lipid molecules (Sankaram et al., 1992), it seems unlikely that ATP hydrolysis could cause net melting of a significant proportion of a crystalline domain. Rather, ATP hydrolysis seems more likely to result in a change in the positions of liquid and crystalline domains within the membrane relative to the position of Na+K+ATPase. This migration of a phase boundary (Klodos et al., 1994) would, we think, put Na+K +ATPase in a liquid domain after ATP hydrolysis (thus favoring the E2P conformation) and in a crystalline domain after ATP binding (thus favoring the E1 conformation). 4 As discussed in Chapter 2, different proteins may associate preferentially with different lipids in the bilayers of membranes and the lipids with which they associate might form liquid or crystalline domains. It is conceivable that binding of ATP results in a Na+K+ATPase molecule that associates preferentially with crystalline-forming lipids, whereas hydrolysis of the ATP and possibly autophosphorylation of the enzyme attracts liquid-forming lipids to it. In this regard, the interactions of Na+K+ATPase with amphophilic styrol dyes appear to depend on the conformational state of the enzyme (Fedosova et al., 1995). Hence, the El(P) and Ez(P) conformations of Na+K+ATPase could, indeed, present different surfaces to the membrane lipid bilayer, which might be more attractive to and favored by lipid in the crystalline or liquid states, respectively. Missing, of course, are data that indicate whether ATP binding and hydrolysis attract crystalline- and liquid-forming lipid domains, respectively. Regardless of whether induction of phase transitions cause El(P)/ Ez(P) conformational changes, rather than simply being associated with the changes, it would nevertheless be 4It should also be noted that both a decrease in enthalpy and an increase in entropy contribute to the free energy decrease associated with ATP hydrolysis (Table 3.1). Components of the transport process catalyzed by Na+K+ATPase may, however, be solely enthalpy or entropy driven. In these cases, the magnitudes of the individual enthalpy and entropy (times absolute temperature) changes may each greatly exceed the magnitude of the overall free energy change (i.e., AG will be negative regardless of whether the magnitudes of AH and TAS are large or small relative to the magnitude of AG). Hence, much more thermal energy may be released transiently to or absorbed temporarily from the environment in association with a component of the transport process than is apparent from the free energy available from ATP hydrolysis (see example at the end of this section).

significant if different phases merely stabilize different conformations once they form. This model could be important in describing the details of how the forward transport operation of Na+K+ATPase is favored rather than its reverse ATP-synthetic operation. Such should, of course, be the case for whatever form the free energy takes in its conversion from the largely chemical form in ATP to the form in cation gradients under the physiological conditions of most cells. If the phosphorylated E2P form of the enzyme is stabilized by the liquid domain that it attracts, then it would be difficult for it, once so stabilized, to be converted back to the E1P conformation that would presumably be needed to donate the phosphoryl group in ATP synthesis (Fig. 3.7). Similarly, if the resultant ATP-bound E1 form of the enzyme attracts and is stabilized by a crystalline domain, then the free energy barrier to its conversion to the E2 form needed for phosphorylation by inorganic phosphate would be opposed by its preference for and stabilization by a crystalline domain. No additional free energy for melting or, of course, for freezing of membrane lipid would be needed in order for ATP hydrolysis to drive Na + and K + transport. Rather, the enzyme would take advantage of liquid and crystalline domains of lipid that exist at equilibrium during phase transitions to favor transport and ATP hydrolysis at physiological levels of intracellular ATP, ADP, and Pi. Presumably, most of the relatively large changes in enthalpy and entropy associated with "melting" or "freezing" of a crystalline or liquid lipid domain around Na+K+ATPase could occur as a result of "freezing" or "melting," respectively, of similar amounts of lipid in domains elsewhere in the membrane. In addition and as discussed above, the relative concentrations of different lipid species are different in the liquid and crystalline domains of the membrane lipid bilayer. In this model, the location at any given moment of the free energy resulting from ATP hydrolysis becomes obscured. As discussed above, the free energy of hydrolysis of ATP is only about 1% of the enthalpy and entropy (times absolute temperature) changes proposed here to be involved in the phase transitions of membrane lipid domains. Consequently, Na+K+ATPase may operate by interconverting relatively large but more or less constant amounts of free energy, while at the same time using the free energy of ATP hydrolysis to change the locations of liquid and crystalline lipid domains in the membrane rather than permitting these locations all potentially to be random. That is, ATP hydrolysis might be used in part to decrease the entropy of the membrane in regard to the locations of liquid and crystalline domains in order to help to produce the conformational changes in the Na+K+ATPase molecule that are needed for it to catalyze cation migration against the pertinent

Reversal of Transport May Drive Chemical Reactions total chemical potential gradients. To our knowledge, the measurement of such changes in the entropy of the lipid bilayer have not, as yet, been attempted. Nevertheless, these or other similarly obscure types of changes in entropy, enthalpy, and free energy may need to be measured if we are to comprehend, at a fundamental level, the thermodynamics of how chemical reactions and transport can be coupled. For example, it was shown recently that the binding of Mg 2+ to the sarco(endo)plasmic reticulum CaZ+ATPases is a wholly entropy-driven process (recall definition in Table 3.1). Binding of Mg 2+ (which is involved in reversal of ATP hydrolysis by the ATPase) results in an enthalpy change of about +180 kJ mo1-1, whereas the change in entropy multiplied by absolute temperature is about 192 kJ mo1-1 (Schwarz and Inesi, 1997). Hence, much more thermal energy may be extracted from or added to the environment at a given moment during the catalytic cycle than appears to be available from the overall free energy change or even from the free energy change associated with ATP hydrolysis (i.e., 42 to 54 kJ mol-1). Consideration of these components of the overall free energy change may eventually expose mechanisms for the cyclic freezing and melting of whole membrane lipid domains during the transport-reaction cycle.

VII. REVERSAL OF TRANSPORT MAY DRIVE CHEMICAL REACTIONS In the case of Na+K+ATPase, we learned in Section VI,A that the free energy of ATP hydrolysis may be only slightly larger in magnitude than the free energy of Na + extrusion plus K + uptake. Hence if the intracellular ATP level were to drop somewhat and ADP and Pi levels were to increase, then the free energy of ATP hydrolysis would no longer be sufficient to drive transport. Rather, the reversal of transport could conceivably drive synthesis of ATP from ADP and Pi 9Such is, in fact, the case for the processes catalyzed by Na+K+ATPase. According to the model presented above for this enzyme (Fig. 3.7), the reduced level of ATP and concomitant increase in the Pi concentration would favor dissociation of the nucleotide from the enzyme, stabilization of the E2 conformation by a liquid lipid domain, and conversion of the E2 form of the enzyme to the E2P form. As for the forward process described above, the free energy for the "melting" and "freezing" of a lipid domain that may be needed for the complete cycle would be supplied at the expense of "freezing" and "melting," respectively, of other lipid domains in the membrane bilayer.

55

In contrast to phase transitions of lipid in the plasma membrane bilayer, the F-type and V-type ATPases in mitochondrial and other membranes may couple transport and chemical reactions by a different mechanism. Moreover, the normal direction of operation of the Ftype ATPases in mitochondria is the formation of ATP at the expense of a total chemical potential gradient of H § ions across the inner membrane. 5 For this reason, we will focus our discussion in this section on the thermodynamics of coupling between H § transport and ATP synthesis. Let us also derive a thermodynamic expression for the combined processes of phosphorylation and H § transport, since we did not do so for the processes catalyzed by Na+K+ATPase. Such a thermodynamic expression could, of course, also be derived in the same way for a combination of the latter processes simply by taking the sum of the expressions for the individual processes (each derived in Section VI,A above). The F-type and V-type ATPases are composed of catalytic and membrane sectors, the structures of which will be discussed in greater detail in Chapter 5. For now, we need only to know that the membrane Fo sector of the mitochondrial F-type ATPase conducts protons across the inner mitochondrial membrane, whereas the catalytic F1 sector hydrolyzes or synthesizes ATP. The reader will now recognize that each sector actually has its own catalytic activity and that the F1 sector is said to be catalytic specifically because it catalyzes a chemical change. The F1 sector of the mitochondrial enzyme is attached to the inner membrane on the matrix side, while the membrane sector spans the membrane (Fig. 3.10). Possible mechanisms for the bioenergetic coupling of transport and ATP synthesis or hydrolysis are discussed in the next section and in Chapter 5. Here we will derive the thermodynamic expressions for the processes. The thermodynamic expression for migration of protons through the inner mitochondrial membrane resembles Eq. (3.35) for the uptake of K § by cells via the plasma membrane. The equation for protons is AGH+

=

A/d, tH+i-o

+ zH+ ~ i

--

RT In (aH+i/aH+o)

(3.41)

- q'o),

where ' T ' and "o" now refer to H + activities and membrane electrical potentials inside and outside the inner mitochondrial membrane, respectively. If we assume that H + concentrations are nearly equivalent to their activities then Eq. (3.41) becomes 5The proton gradient is, of course, created across the inner mitochondrial membraneby the oxidation-reductionreactions of the electron transport chain. The details of the bioenergeticcouplingbetween these reactions and proton transport is also only now coming to be understood at a fundamental level (e.g., see summaryof redox-driven proton pumps in Tables 8.1 and 8.2 of Chapter 8).

56

3. Thermodynamics and Transport

F-ATPase

V-ATPase

SubunitM.W.kDa 55 J3 50 .y 31 ~ . ~ a

b c

B ;

SubunitM.W.kDa A 68 B 57 C 44

1517~ i ~

E a c

30

F1

IN MEMBRA O'~

V1

26 20 16

CYTOPLASM

Vo MEMBRANE ~

LUMEN

FIGURE 3.10 Structures of F- and V-type ATPases. The F-type ATPase diagrammed is for the enzyme in E. coli but all F- and Vtype ATPases have similar structures (adapted from Nelson, 1995 with permission from Landes Biochemical).

AGH+ = 2.303 RT(pHo - pHi) -I- ZH+ O~- (~Ir __ ~Ir

(3.42)

Similarly, the thermodynamic expression for ATP synthesis resembles Eq. (3.37) for ATP hydrolysis. If we assume that concentrations are nearly the same as activities in the matrix of the mitochondrion then the free energy of ADP phosphorylation in the reaction ADP 3- + P i 2- + H + ~-~ ATP 4- + H 2 0

Vlll. HOW DO FLUCTUATIONS IN THE LOCAL HYDROGEN ION POTENTIAL FACILITATE FORMATION OF PHOSPHORIC ACID ANHYDRIDE BONDS BY THE MITOCHONDRIAL FoF1ATP SYNTHASE? Recall that Eqs. (3.37) and (3.44) were derived under the assumption that Reactions (3.36) and (3.43) occurred at pH 7. In contrast, the total chemical potential difference associated with a pH gradient is believed to provide the free energy for ATP synthesis by the F-type ATPase in mitochondria (Fig. 3.10). Initial insight into part of the mechanism by which a proton total chemical potential gradient could help to drive phosphorylation, thus, comes simply from observing that the reaction depicted by Eq. (3.43) would be shifted toward ATP synthesis by a decrease in pH. Perhaps more importantly, much of the increase in free energy required for phosphorylation of ADP results from the free energy needed to bring together the A D P 3- and Pi2- anions (Fig. 3.11). If, however, the protons entering the mitochondrion through the Fo component of the ATPase could somehow be directed to ADP 3- or to Pi 2at their binding sites on the F1 component temporarily to neutralize their charges, then the free energy of ATP

(3.43)

@ AGp = AG~

+ R T l n ([ATP]/([ADP] [Pi])).

(3.44)

As we have seen on several occasions, the total free energy change for coupled processes is equal simply to the sum of the free energy changes for the individual processes. Hence, the thermodynamic expression for the total free energy change associated with the processes catalyzed by the mitochondrial FoF1 ATP synthase is AGT

=

AGH+ + AGp,

(3.45)

where A GT is the free energy change for the overall process, AGH+ is the free energy decrease associated with migration of protons into the mitochondria, and AGp is the free energy increase required for ADP phosphorylation. When A GT is negative, ATP is synthesized and when A GT is positive, protons are pumped out of the mitochondrial matrix at the expense of ATP hydrolysis. Our first hint concerning how the free energy in proton gradients might be converted to free energy in phosphoric acid anhydride bonds of ATP requires that we reconsider the assumptions under which Eqs. (3.37) and (3.44) were derived.

"O- ~=-O- IP- O- ~ O- I Rib I-l Adenine I O" ATP 4O ' J O'. H~O ihydrolysis, with

"O- ~-OH OPi

.....

"r~lief of ~arge repulsion

+ H O - I ~ ) - ~ - O q Rib H Adenine I O" OADP 2-

I resonance stabilization

;~PI~

L'?

H+

Li 1 onization

H. +-o-

9 9 ,P-O- ,p- o-I O"

ATP 4- + H20

AG ~

O"

Rib mAdenine I ADP 3-

= ADP 3- + Pi 2- + H + -31.7 kJ/mol

FIGURE 3.1 1 Most of the work of ATP 4- synthesis from ADP 3and Pi 2 at physiological pH values is needed to appose the negatively charged substrates. These substrates are unlikely to approach each other spontaneously because such a process alone would require an increase in free energy at physiological concentrations of the substrates. Conversely, separation of these charges is the principal reason for the decrease in free energy associated with ATP hydrolysis, although other factors also contribute to these bioenergetic considerations. Nevertheless, a transient increase in the total chemical potential of H+, as could conceivably occur near the F1 sector of ATP synthase due to the flow of H + through the Fo sector (Fig. 3.10), would greatly reduce the negative charges on ADP 3- and Pi e- and hence the work needed to oppose them (see discussion in text) (adapted from Lehninger, 1993 with permission from Worth Publishers).

Conversions of Solute Gradients

synthesis would be considerably reduced. In this model, the free energy decrease associated with the flow of protons with their concentration gradient is coupled to a reduction in the free energy needed for ADP phosphorylation. This view has been challenged in part because the equilibrium constant for synthesis or hydrolysis of ATP is not influenced by the surrounding pH while ATP is bound to the FI component of the ATPase (see Chapter 5 for a description of the more generally accepted binding change model; Boyer, 1993). The pH of the surrounding bulk water solution may, however, not influence the effective H + concentration at the nucleotide binding site. Rather, ATP synthesis and hydrolysis appear to occur at a hydrophobic binding site that is thus sequestered from surrounding bulk water. Under such conditions, the water concentration would also be below that of water solutions and hence would also favor the formation of ATP (Reactions (3.36) and (3.43)). As for the H + concentration, it can no longer be assumed that the water concentration is constant and nearly equal to that of pure water. Under such conditions it is not surprising that synthesis and hydrolysis of ATP appears to occur several times during occupancy of the ATP (or ADP + Pi) binding site and that the equilibrium constant is nearly equal to one (reviewed by Nakamoto, 1996). While the ATP (or ADT + Pi) binding site may be constrained and optimized to catalyze chemical change (e.g., Senior et aL, 1995), sequestration of ADP and Pi with migrating protons and away from water should minimize the free energy change associated with ATP synthesis and hydrolysis. Whether the migrating protons participate directly in the chemistry of ATP synthesis seems to depend on whether the protons bind only to amino acid residues during transport or whether they may also be associated with the nucleotides or Pi. In this regard, it was proposed in a somewhat different context that protons might be transferred temporarily to a dihydrogen phosphate at the end of the pathway in which they are transferred across the Fo complex to FI (Bartl et aL, 1995). The temporary formation of such a neutral phosphoric acid species owing to fluctuations in the local H + potential would decrease considerably the work needed to appose the phosphoric acid and ADP 3- (Fig. 3.11). Formation of a phosphoric acid anhydride bond prior to dissociation of the transiently associated protons would insure that much of the free energy initially associated with the proton total chemical potential gradient is now associated instead with a high-energy phosphate bond of ATP. This simple model also is consistent with the observation that ATP hydrolysis drives H + transport when the forward process is reversed due to a reduction of the proton total chemical potential gradient and an increase in the ratio of the ATP to ADP

57

and Pi concentrations. In this case, one of the dissociable protons produced during ATP hydrolysis could remain transiently associated with Pi 2- long enough to be transferred back to the aspartyl residue on Fo from which a proton appears to emanate in the forward process. Actually, protons are thought to move across sites on several amino acid residues in the Fo sector when they move through it across mitochondrial or bacterial membranes (e.g., Bartl et aL, 1995). Primary active transport processes similar to that catalyzed by reversal of mitochondrial FoFaATP synthase may serve numerous functions, such as acidification of endosomal and lysosomal compartments by a V-type ATPase. The components of V-type ATPases are homologous to the components of F-type ATPases, although the former cannot use a proton total chemical potential gradient to synthesize ATP. This irreversibility appears to result from transport of a portion of the actively transported protons in the reverse direction along their total chemical potential gradient via V-type ATPases without concomitant chemical change (Nelson, 1992a,b, 1995). 6 Acidification of intracellular compartments by V-type ATPases serves useful functions in its own right, such as facilitation of the dissociation of ligands from their receptors after fusion of endocytic vesicles with endosomes (Fig. 3.12). We are, however, interested here primarily in the ability of solute gradients to drive other transport processes or to perform other transport related work. Hence, we return now to the thermodynamics of coupling between multiple transport processes.

IX. CONVERSION OF SOLUTE TOTAL CHEMICAL POTENTIAL GRADIENTS TO GRADIENTS OF OTHER SOLUTES DURING CO- AND COUNTERTRANSPORT A. General Considerations The biochemically simplest form of cotransport resulting from the Gibbs-Donnan effect requires a semipermeable membrane, but it does not require a transport protein. Hence, it is substrate selective only in regard to membrane permeability. Nevertheless, the Gibbs-Donnan effect (described above) led to the movement of Na + with its total chemical potential gradi6From a thermodynamic standpoint the irreversibility of the primary active transport catalyzed by V-type ATPases cannot be attributed to the transport of 2 rather than the 4 protons transported per cycle by F-type ATPases (see Chapter 5). The transport of 2 protons per cycle would permit production of a steeper proton gradient than the transport of 4 protons per cycle, and the steeper gradient could be used for ATP synthesis were it not for transport in the reverse direction in the absence of chemical change.

58

3. Thermodynamics and Transport

FIGLIRE 3.12 The functions of acidification of membrane-bound compartments in eukaryotic cells include dissociation of ligands from their receptors in endosomes so that the receptors may be returned to the plasma membrane. This process is shown in the upper-right portion of the figure (i.e., processes 3 and 4) (adapted from Lukacs et al., 1995 with permission from Landes Bioscience).

ent in association with C1- along its initial chemical potential gradient (Figs. 3.4 and 3.5). Granted, in the case of the G i b b s - D o n n a n effect, it appears to be the small membrane potential that would develop when some Na + has moved from compartment B to compartment A that may then "pull" C1- along its new total chemical potential gradient from B to A. Nevertheless, the movements of Na + and C1- can be viewed to be as closely coupled in regard to free energy changes as any symport or antiport process that requires a transport protein for coupling. In fact, thermodynamic expressions that are very similar to the one for the G i b b s Donnan effect (Eq. (3.25)) can be derived for the coupled symport or antiport of two solutes. For example, consider a hypothetical Na§ - symport protein in the plasma membrane of a cell. The free energy needed to move Na § out of the cell against its

total chemical potential gradient was derived in Section VI above to be A GNa + = R T In

(aNa+o/aNa+i)

-'1- ZNa+ S (tI/"o -- tI)'i).

(3.34)

Equation (3.34) can, of course, be rearranged to give the decrease in free energy when Na + moves into the cell with rather than against its total chemical potential gradient AGNa+ = R T In (aNa+i/aNa+o) -I- ZNa+ u-~"(tIYi -- tIto).

(3.46)

A similar expression for the movement of C1- into the cell via the hypothetical carrier is given by AGcl_ = R T In (acl-i/acl-o) "F ZC1-u~-'(tIfi- tIfo).

(3.47)

Conversions of Solute Gradients

Hence, the total free energy change for cotransport is A G T = AGNa+ nt- A G c 1 or

AGT = R T In (aNa+i/aNa+o) 4- R T In (acl-i/ac~-o) (3.48) -t- Z C I - ~ z ' ( a I J i - aIto) -1- ZNa+ ~ - ( a I t i -

~Ito) 9

Now the only difference between the terms involving membrane potential is that z for Na § is positive and z for C1- is negative. Hence, Eq. (3.48) simplifies to AGT = R T In (aNa+ilaNa+o) + R T In (acl-i/acl-o).

(3.49)

Moreover, at equilibrium in regard to the solutes (and steady state in regard to the system) R T In (aNa+i/aNa+o) = - R T In (acl-i/acl-o), which can be rearranged and simplified to

59

intuitive sense, since the free energy conversions from a total chemical potential gradient of one solute to a gradient of another are between similar types of changes in the total chemical potential of each solute across the membrane. As discussed above, a major challenge to biophysicists is to comprehend in a similar way the bioenergetic coupling between chemical reactions and biomembrane transport. It is also now instructive to make the additional considerations of membrane electrical potential and solute charges that are unnecessary for symport or antiport of uncharged solutes or for the hypothetical Na+/C1- cotransport just discussed. In the latter case, membrane electrical potential did not influence transport because the two ions were assumed to be transported in a stoichiometry of 1:1 and because they are of equal but opposite charge. Such conditions and assumptions frequently do not apply to symport and antiport of cations and anions.

In (aNa+i/aNa+o) = In (acl-o/acl-i) or simply

B. The Thermodynamics of Na+/Ca z+ aNa+i/aNa+o = acl-o/acl-i.

(3.50)

The latter relationship is, of course, the same as that derived in Section IV above (Eq. (3.26)) for the GibbsDonnan equilibrium. In fact, Eq. (3.50) applies well to cotransport in cells at a steady state (and at equilibrium in regard to the solutes) when the symporter is 100% efficient and, thus, has no leaks or slippage (i.e., for the hypothetical cotransport just described, entry of Na + can only occur in association with C1- and vice versa). For Eq. (3.50) to apply perfectly, it must also be assumed that the transport occurs in a closed system where the activities (and concentrations) of Na § and C1- outside the cell cannot vary independently (Naftalin, 1984). Although each of these assumptions is rarely if ever entirely correct, in many cases the fact that the two assumptions are not fully correct does not greatly change the result. The general expressions for symport and antiport of any uncharged solutes, A and B (or for charged solutes when the transmembrane electrical potential difference is zero), take the anticipated forms for symport

aAi/aAo = aBo/aBi

(3.51)

aAi/aAo = aBi/aBo.

(3.52)

and antiport

As we saw above, Eq. (3.51) also applies even in the presence of a membrane electrical potential difference to solutes with opposite charges of equal magnitude. Similarly, Eq. (3.52) applies to solutes with equal charges. The similarities of Eqs. (3.50), (3.51), and 3(.52) to Eq. (3.26), derived for the Gibbs-Donnan effect, make

Exchange as an Example of the Bioenergetics of Coupling b e t w e e n the Migration of Solutes Calcium ions can be extruded from the cytosol of cells in primary active transport via P-type, Ca2+ATPases in the plasma membrane or in the membranes of intracellular, membrane-bound compartments. In addition, Ca 2§ may be extruded from cells in exchange for extracellular Na § that had itselfbeen extruded by Na+K+ATP ase. Each of these processes contributes to maintaining the relatively low cytosolic total chemical potential of Ca 2+ in most cells in what is sometimes termed their resting states. Such a steady, resting state is needed in order for transient increases in the cytosolic total chemical potential of Ca 2+ to be able to function in initiating signaling processes. In the present case we wish to derive a thermodynamic expression for the coupling of Na + uptake with Ca 2+ extrusion. This exchange is catalyzed by an antiporter that is selective for these cations and has the stoichiometry 3Na+o + Ca2+i ~ 3 Na+i + Ca2+ o.

(3.53)

An equation for the free energy of uptake of Na + across the plasma membrane was derived above (Eq. (3.46)). In the case of Na+/Ca 2+ exchange, however, 3 Na + ions enter the cell for each Ca 2+ ion extruded. Hence, the free energy available from Na + uptake to move 1 mole of Ca 2+ out of the cell against its total chemical potential gradient is three times more than would be calculated from Eq. (3.46). Equation (3.46) can be multiplied by three to produce

60

3. Thermodynamics and Transport

In ([Na+]i/[Na+]o)3 + 3 ZNa+ 5 ( ~ i ~o),

/~G3Na+ -" R T

(3.54)

where the activities of Na + inside and outside cells are, again, assumed for our purposes to be nearly equal to their molar concentrations. Similarly we can write for the extrusion of C a 2+ AGca2+ - R T ln([CaZ+]o/[Ca2+]i)l

(3.55)

-q- ZCa2+ ~-'(~Ifo -- ~I/i) 9

If it is assumed that the system is at a steady state, that the cations are at equilibrium, and that virtually all of the free energy in the total chemical potential gradient of Na § can be used to maintain a similar gradient of Ca 2+ (and vice versa) then

and AG3Na+ -- - A G c a 2 + or

(3.56)

qt_ ZCa2+ ~'(~I)'i -- "kilo)"

Equation (3.56) can be rearranged and simplified to ln([CaZ+]i/[Ca2+]o) = In ([Na+]i/[Na+]o)3

(3.57)

nt- (3 ZNa+ -- ZCa2+)u-~-"(~Ifi- "tIl'o)/RZ,

which takes the general form for antiport ln(aAi/aA~

ln(aBi/aB~ + (mZB -- n Z A ) ~ ( t I f i -

(3.58)

~o)IRT,

where m and n represent the stoichiometries for uptake and extrusion, respectively, of solutes B and A and "a" represents their activities inside (i) and outside (o) cells. Similarly, the general form of the equation for symport is In

In (aBi/aBo) m + (mZB + nZA)~(ats'i- ~o)/RT,

(aAo/aAi) n --

In ([CaZ+]i/[Ca2+]o) = In (0.011 M/0.140 M) 3 + ((3 x 1) - (1 x 2 ) ) f ( - O . O 5 V ) / R T In ([CaZ+]i/[Ca2+]o) - -7.631 - 1.872 - -9.503 and [Ca2+]i/[Ca2+]o = 0 . 0 0 0 0 7 4 6 2

A G T = AG3Na+ + A G c a 2 + = 0

R T In ([Na+]i/[Na+]o)3 + 3 ZNa+ J-'(tIfi- a~)'o) = R T In ([CaZ+i/[CaZ+]o)a

For the specific case of Na+/Ca 2+ exchange, we can now calculate the steady-state concentration gradient of C a 2+ that could be maintained by the total chemical potential gradient of Na + across a typical cell membrane. If we assume, as we did previously, that a typical cell has a membrane potential of about - 5 0 mV and an intracellular Na + concentration of 11 mM and that the extracellular Na + concentration is about 140 mM, then from Eq. (3.57)

(3.59)

where the ratio of the activities is, of course, reversed relative to antiport for one of the solutes and the charges on the ions are added instead of calculating their difference. 7 7Equations (3.58) and (3.59) can also be used to determine apparent stoichiometries of counter- or cotransport, respectively (e.g., Schroeder, 1995; Chen et aL, 1995; Zerangue and Kavanaugh, 1996a), when these stoichiometries are not already known. We do not, however, recommend use of these equations for this purpose without adequate justification. In most instances it has not been demonstrated that the stoichiometry calculated from these equations matches the actual stoichiometry of transport. As we discuss further in Section III of Chapter 6, such indirect determinations of stoichiometry appear frequently to be inadequate substitutes for measuring the actual unidirectional fluxes of each co- or countersubstrate.

at 310~ This concentration gradient of more than 13,000-fold more free C a 2+ outside cells than in the cytosol is about the gradient that is maintained by a typical cell. For a typical cell in the resting state, C a 2+ leaks appear to be small enough to make unnecessary extrusion of C a 2+ by means other than this exchange. Nevertheless, it has been determined that the C a 2+ leaks are, likewise, small enough also to permit C a 2+ ATPase in the plasma membrane of steady state, smooth muscle cells to maintain the s a m e C a 2+ concentration gradient (Lucchesi et aL, 1988). The reason for this apparently redundant function is probably that smooth muscle and other cells frequently are not at steady state in regard to their cytosolic C a 2+ concentration, which may fluctuate intracellularly by several orders of magnitude to accomplish signaling. While either Na+K+ATPase plus the Na+/Ca 2+ exchanger o r C a 2+ ATPase alone could maintain cytosolic C a 2+ at its relatively low, resting state concentration, the rate of recovery of this state after initiation of C a 2+ signaling would be faster if multiple processes for decreasing cytosolic C a 2+ operate. In addition, there are other demands on the Na + total chemical potential gradient including symport of several solutes, such as amino acids, and the opening of Na+-selective channels (see the following section). In fact, it is probably best to view the numerous transport processes as contributing together to the final distributions of various solutes across the plasma membrane. In this view, for example, both Na+K+ATPase and C a 2+ ATPase should be viewed as contributing to the final distributions of both C a 2+ and Na + across the plasma membrane. Changes in the activities of either or both of these primary active transport processes would influence the movement of both ions across the plasma membrane since they would tend to equilibrate via the Na+/Ca 2+ exchanger. A model for the simultaneous operation of these three transport

Dissipation of Solute Gradients

processes in smooth muscle cells is presented in greater detail elsewhere (Skinner et aL, 1993). X. DISSIPATION OF SOLUTE GRADIENTS THROUGH MEDIATED TRANSPORT PROCESSES MAY ALSO PERFORM WORK The thermodynamic expressions for dissipation of solute gradients via channels are the same as those derived above [e.g., Eq. (3.46) for Na + and Eq. (3.47) for C1-]. These gradients are, of course, produced as a result of primary active transport and, in some cases, to its coupling to symport or antiport. For example, taurine exodus from cells via channels may contribute to a regulatory volume decrease in the cells when the channels open only because Na+/taurine symport had previously accumulated taurine in cells against the total chemical potential gradient of taurine. The concomitant exodus of water with taurine also via channels or via other processes that are apparently not mediated by proteins (e.g., see next chapter) results in a decrease in cell volume. Hence, we can think of these separate processes of transport via channels, symport, antiport, and primary active transport as bioenergetically coupled instead of separate processes. In the present example, ATP hydrolysis drives the exodus of taurine via channels as surely as it drives Na + and K + pumping. We have already derived thermodynamic expressions for each component of these coupled processes, and there is little need to write additional equations for these processes in this section. Rather, we will focus instead on physiologically significant aspects of the regulation of channel transport that, when driven by primary active transport, permits cells to produce and use ion gradients and membrane electrical potential in their functioning. A. Importance of the M e m b r a n e as a Barrier to Ordinary Diffusion In order to produce the highest resultant free energy during primary active transport of inorganic ions, it is important that the solute total chemical potential gradients not deteriorate greatly owing to migration of the ions apparently through leaks in the plasma membrane. It is unlikely that such leaks result from simple diffusion of inorganic ions across the lipid bilayer as discussed more thoroughly in Chapter 4. Although the leaks might still be lipid mediated, we prefer the theory that they are primarily the consequence of unwanted protein mediated transport. As discussed further in Chapter 6, many transport proteins have multiple transport functions, but it is possible that not all of these functions are simultaneously useful to cells. Regardless of their

61

cause, wasteful Na + and Ca 2+ leaks through the smooth muscle cell membrane appear to be small enough to allow the plasma membrane CaZ+ATPase alone to maintain an approximately 13,000-fold excess of Ca 2+ outside vs inside these cells at steady state (e.g., Lucchesi et al., 1988). It is also possible to show that Na+K+ATPase plus Na+/Ca 2+ exchange are able to produce and maintain this Ca 2+ gradient. As discussed above, operation of these mechanisms together would more rapidly remove Ca 2+ from the cytosol than either mechanism alone when deviations from the steady state Ca 2+ gradient occur during normal cell functioning. For the present discussion, it is important to appreciate that the cell membrane can be maintained in a relatively leak-free condition in regard to most inorganic ions when this condition is needed. The remarkable nature of this leakfree condition can be better appreciated when one remembers not only that ordinary diffusion of inorganic ions across biomembranes is virtually prohibited, but also that if ordinary diffusion did occur over this distance, it would be more than 10 times faster than the fastest known transport catalyzed by channels. B. Contribution of Na+K+ATPase to the M e m b r a n e Electrical Potential Since the plasma membrane can be maintained in a relatively leak-free condition in regard to inorganic ions, Na+K+ATPase contributes directly to production of the membrane electrical potential. By catalyzing extrusion of 3 Na + ions and uptake of 2 K + ions in each cycle, the pump extrudes net positive charge, which contributes to the inside negative membrane potential. It can also be calculated, however, that the direct contribution of this transport imbalance to the membrane potential is relatively small in most cells (e.g., Stein, 1990). The latter calculations are not intended to be taken alone since it is, of course, the hydrolysis of ATP by Na+K+ATPase that may provide much of the free energy needed to produce and maintain the membrane potential. This coupling of ATP hydrolysis to production of membrane potential is accomplished by also coupling part of the transport catalyzed by Na+K+ATPase to regulated migration of one of the cations with its total chemical potential gradient. C. Na+K+ATPase, K+-selective Channels and the G i b b s - D o n n a n Effect Allow Most Cells to Convert S o m e of the Free Energy of ATP Hydrolysis into the Free Energy of the Plasma M e m b r a n e Electrical Potential The free energy increase associated with the uptake of K + by Na+K+ATPase against its total chemical potential gradient is relatively small because the gradient is

62

3. Thermodynamics and Transport

small. The gradient is small because most cells at rest have open K + channels that allow K + to migrate more or less freely with its total chemical potential gradient from the cytosol to the extracellular fluid. 8 Although this process might, at first, seem wasteful and futile, the m o v e m e n t of K + out of the cytosol greatly increases the inside negative m e m b r a n e potential. Na+K+ATPase is primarily a Na + pump, and K + might even be viewed as an enzyme cofactor in most cells at rest. Consequently, most of the free energy of A T P hydrolysis is devoted to moving Na + out of the cell rather than to uptake of K + by it (Table 3.2). That the enzyme can be viewed approximately as a Na + pump also means that it contributes much more significantly to the membrane potential than it would otherwise. In this case, Na+K+ATPase effectively extrudes nearly three rather than one positive charge per cycle. In fact, the resultant m e m b r a n e potential and the m o v e m e n t of K + relatively freely toward G i b b s - D o n n a n equilibrium via channels in the plasma m e m b r a n e of most cells at rest agrees well with the distribution of this ion across the plasma membrane. For example, in skeletal muscle cells, C1- ions as well as K + ions are relatively free to migrate across the sarcolemma via channels that select for them. Using a m e m b r a n e potential of about - 9 0 m V for this tissue, the G i b b s - D o n n a n equilibrium concentration ratios of K + and C1- can be calculated using an equation similar to 3.25 in which K + is substituted for Na + and assuming that the concentrations of these ions are low enough to be nearly equal to their activities. In this case

~ - ~ i - q%)/RT = In ([K+]o/[K+]i) = In ([C1-]i/[C1-]o), where "i" and " o " designate the intracellular and extracellular compartments, respectively. Substituting the appropriate values for the constants and for the membrane potential (96, 500 J V -~ equiv-~)(-0.090 V)/(8.314 J deg -~ mo1-1) (310~ - In ([K+]o/[K+]i) = In ([C1-]i/[C1-]o) -3.370 = In ([K+]o/[K+]i) = In ([C1-]i/[C1-]o) +3.370 = In ([K+]i/[K+]o) = In ([C1-]o/[C1-]0 [K+]i/[K+]o = [C1-]o/[C1-]i = 29.1 The actual concentration ratios of K + and C1- inside and outside m a m m a l i a n skeletal muscle cells can be calculated from their concentrations (Hille, 1984) to be about [K+]i/[K+]o

--

0.155 m/0.004 m - 38.8

and 8See Section II of Chapter 9 regarding hepatocytes for an important exception to this generalization.

[C1-]o/[C1-]i---

0.123 M/0.0042 M = 29.3

Thus it can be seen that the steady-state concentration ratios of intracellular-to-extracellular K + and extracellular-to-intracellular C1- exceed their ratios expected from the G i b b s - D o n n a n equilibrium by only a small amount. In fact, the intracellular-to-extracellular K + concentration ratio is so near equilibrium that inhibition of Na+K+ATPase results in almost no increase in membrane electrical potential because K + may now achieve this equilibrium. Actually, the m e m b r a n e potential decreases somewhat just after inhibition of Na+K+ATPase apparently as a result of inhibition of the direct, electrogenic contribution of the Na+K + p u m p activity to membrane potential (Alberts et al., 1994). In addition, it should be clear from these considerations that the inside negative m e m b r a n e potential results both from a small contribution to the G i b b s - D o n n a n effect by impermeant intracellular anions and a large contribution to this effect by the relatively i m p e r m e a n t extracellular cations Na + and Ca 2+. The extrusion of the cations via P-type ATPases and Na+/Ca 2+ exchange and the relative absence of Na + and Ca 2+ leaks in the plasma m e m b r a n e generate most of the m e m b r a n e potential because K + and C1- frequently approach a G i b b s - D o n n a n equilibrium set up by the i m p e r m e a n t cations. 9

D. H o w M u c h W o r k C o u l d Be P e r f o r m e d by t h e R e s t i n g P l a s m a M e m b r a n e Electrical Potential? A n equivalent of electrical charge represents about 4.8 kJ of free energy at a m e m b r a n e electrical potential of 50 m V ( f x (~o - ~i) - 96.5 kJ V -1 equiv -1 x 0.05 V - 4.8 kJ equiv-1). This quantity of free energy is, of course, the same as the contribution of an inside - 5 0 m V electrical m e m b r a n e potential to the total chemical potential of an equivalent of monovalent intracellular anions or extracellular cations. While 4.8 kJ equiv -a may, at first, seem small relative to the free energy of hydrolysis of A T P (42-54 kJ mo1-1 in most cells), it should also be appreciated that the concentrations of m a n y ions is greater than the concentration of ATP. For example, the extracellular Na + concentration is more than 40-fold higher than the intracellular A T P concentration. Of course only a small proportion of Na + ions would need to move to produce the m e m b r a n e depolarization that is associated with an action potential. [Recall the example in Section IV above regarding 9Surprisingly, some types of K+ channels may catalyze net migration of K+ against its total chemical potential gradient (see Chapter 7). Hence, the distribution of K + ions across the plasma membrane appears to depend on sometimes unanticipated K +transport by several different processes.

Summary the proportion of the ions that need to move to produce a membrane potential of - 1 0 mV and see Hille (1984) for such examples for actual cells.] Nevertheless, this small amount of ion movement represents a substantial temporary diminution of the free energy stored by total chemical potential gradients of Na + and Ca 2+ and, likewise, an increase in the free energy stored by gradients of K + and C1- across the plasma membrane.

XI. APPLICATION OF THERMODYNAMIC PRINCIPLES TO THE SOLUTION OF PRACTICAL TRANSPORT PROBLEMS A kinetic approach frequently is used to distinguish among multiple transport activities for the same substrate (see Chapter 4). Thermodynamic concepts have, however, been used recently to help investigators to recognize multiple transport activities apparently attributable to a single protein. Uptake of the excitatory amino acid, glutamate (Glu-), by glial cells is driven mainly by the Na + total chemical potential gradient across the cell membrane. Although Glu- carries a net negative charge at neutral pH, its cotransport with 2 (Bouvier et al., 1992) or 3 (Zerangue and Kavanaugh, 1996a) Na + ions results in a positively charged current across the plasma membrane. The magnitude of this inward positive current is, of course, greater at more negative inside electrical membrane potentials, and it should approach zero as the magnitude of the inside negative membrane electrical potential is decreased and finally reversed. In addition to Na + and Glu- uptake, each transport cycle also appears to include extrusion of 1 K + ion and uptake of 1 proton. Now when each member of a family of Glu-/Na + cotransport proteins was expressed in Xenopus oocytes, somewhat different than anticipated results were obtained (e.g.; Wadiche et al., 1995). Expression of these proteins in oocytes led in most instances to reversal of the current at sufficiently high inside positive electrical membrane potential (Fig. 3.13). The reverse currents were not attributable to flow of Glu- and Na + in the reverse direction since all currents appeared on first application of the amino acid to the solution bathing oocytes. Furthermore, the reverse currents were eliminated when C1- was removed from the medium (Fig. 3.13). Hence, it was concluded based on this and other evidence that this family of excitatory amino acid transporters ( E A A T ) catalyze both Glu-/Na + cotransport and channel-like transport of C1-. Since transport was studied by expressing the protein in Xenopus oocytes, it is, of course, conceivable that the E A A T proteins activate one or more transport activities endogenous to the oocyte rather than serving for both Glu-/Na +

63 membranepotential(mV)

-150

I

-1(00 I

.soI

.~0

130

C +500 104~ M cl'~

ooooo ,

~ 1_11.117mO

t-^

-500 0 r-

o

+100 I

0 m M CI-ou t

--lOOO

0

- -1500 - -2000

FIGURE 3.13 Glutamate (Glu-)/Na + cotransporters produce the anticipated currents at relatively low inside-negative membrane electrical potential differences. The current is, however, reversed on addition of the amino acid at sufficiently high, inside positive membrane electrical potential. In contrast, the current is not reversed on addition of Glu- in the absence of CI-. Hence, it is concluded that the Glu-/ Na+ cotransporter also acts as a C1- channel. The experiment shown is for expression of excitatory amino acid transporter (EAAT)I after injection of its cRNA into Xenopus oocytes (adapted from Wadiche et al., 1995 with permission from Cell Press).

cotransport and an independent C1- migration. Nevertheless, an understanding of the thermodynamic principles of biomembrane transport particularly as they relate to membrane potential permitted these authors to recognize that several transport activities probably exist where only one had been anticipated (Fig. 3.13).

XII. SUMMARY Several factors may contribute to the total chemical potential of a solute or the solvent and hence to their free energies of biomembrane transport. Moreover, primary active transport is coupled to a chemical change, and it may drive subsequent symport, antiport, and uniport. Hence, the total free energy change of a transport process or a series of processes may have numerous obvious components. Knowledge of these components frequently has practical implications for the interpretation of transport experiments, such as that just described for data showing the apparent function of glutamate transporters also as C1- channels. In addition, knowledge of the free energy changes associated with each component of transport is required in order fully to understand how transport contributes to the work performed by cells.

64

3. Thermodynamics and Transport

One component of the total chemical potential of a solute or the solvent is its chemical potential. In addition, this total for a charged solute includes the effect of the membrane electrical potential difference on its tendency to migrate across a membrane. The migration of K + ions across the plasma membrane with their total chemical potential gradient through channels after the ions are first taken up by Na+K+ATPase contributes to the work of producing the membrane electrical potential in many cell types. Subsequent temporary migration of Na + ions with their total chemical potential upon activation of ligand- or voltage-gated channels helps to produce the action potentials needed for the propagation of nervous impulses and the signaling of muscle cells to contract. The total chemical potential gradients of these and other solutes may also be used for additional work, such as regulatory volume increases or decreases of cells, and these volume changes may also serve in signaling processes. Volume changes of course require water transport usually via water channels, and the tendency of water molecules to migrate into or from cells depends on the difference in the osmotic pressures of the intracellular and extracellular fluids. Moreover, the movement of nutrients, such as amino acids and monosaccharides, and their interorgan flows in multicellular organisms may depend on gradients of other solutes with which transport of the nutrients is coupled. Such total chemical potential gradients of solutes are first established through primary active transport and the hydrolysis of ATP. Despite some knowledge of the details of the processes through which hydrolysis (or synthesis) of ATP may be coupled to transport, an understanding of the bioenergetics of this coupling is only now emerging. We proposed in this chapter that the free energy changes associated with components of primary active transport processes may include much larger changes in the magnitude of enthalpy and entropy than are associated with ATP synthesis or hydrolysis. These larger changes in enthalpy and entropy were proposed to be associated with "melting" and "freezing" of crystalline and liquid domains in membrane lipid bilayers, which are in phase transition at physiological temperatures. In this context,

it may be possible to couple primarily enthalpy-driven processes such as the hydrolysis of ATP with the reversal of primarily entropy-driven processes such as the reversal of migration of solutes with their total chemical potential gradients. In this model it was proposed that hydrolysis of ATP helped to cause lipid phase transitions to occur near Na+K+ATPase and other P-type ATPase proteins in the membrane. These phase transitions were proposed to be needed to favor conformational changes of the proteins that lead to uptake or extrusion of solutes from cells. Hence it can be seen in this model that the free energy of hydrolysis of ATP may be converted to a free energy of the position of a solute by first converting part of the free energy to the free energy of position of liquid and crystalline membrane lipid domains. The magnitudes of the entropy (times absolute temperature) and enthalpy changes associated with the phase transitions of the entropy-driven melting and enthalpy-driven freezing of the lipid domains (but not the magnitudes of the free energy changes) can be calculated greatly to exceed the magnitudes of the free energy changes associated with ATP hydrolysis and biomembrane transport. In addition to the free energy available from solute transport, the cell depends on transport occurring rapidly enough to do work useful to it. Transport proteins are the catalyst that permit solutes and the solvent to migrate across biomembranes much more rapidly than they can by permeating phospholipid bilayers. Although ordinary diffusion of solutes and the solvent may be relatively rapid over short distances in intracellular and extracellular aqueous solutions, such diffusion is virtually halted across thin biomembranes. Ironically, many transport processes have been described elsewhere as facilitated diffusion, diffusion through channels, and even nonmediated simple diffusion through the lipid bilayer. We shall see, however, that while the concept of ordinary diffusion is important to understanding biomembrane transport, biomembrane transport hardly qualifies as that simple process. Since measurement of the rate of solute or solvent transport is by definition a kinetic rather than a thermodynamic problem, we turn in the following chapter to a consideration of biomembrane transport kinetics.

C

H

A

P

T

E

[4 1

R

Transport Kinetics

distinctions that are sometimes made between channels and carriers, such as differences in their turnover numbers or in the extent to which they are gated, as arbitrary and based primarily on the historical circumstances of their discovery and study. Channels catalyze transport by interacting intimately and selectively with their substrates, as do carriers (Chapters 5 to 7). In fact, the catalytic sites of channels, carriers, and enzymes exhibit homologous principles of structural design (Marban and Tomaselli, 1997). To emphasize the similarities between channels and carriers, we define transporters here as proteins that catalyze movement of solutes or the solvent from one side of a biomembrane to the other regardless of their categorization also as channel or carrier proteins (Chapter 8). Several authors have in fact expressed the notion that many and, as we understand them, perhaps all proteinmediated transport processes are built on a central transmembrane channel (e.g., Wilson, 1978; L~uger, 1980; Nikaido and Saier, 1992; Chen and Eisenberg, 1993). Symport, antiport, and the coupling of transport to chemical reactions appear to be features added to these channels (e.g., Nikaido and Saier, 1992). Since animals with a nervous system and the requisite gated channels evolved after other rather complex forms of life had already appeared, the actual order of evolution of specific transport processes may now be obscured. Nevertheless, it seems likely that channels, uniporters, antiporters, symporters, and primary active transport pumps have evolved from one another on numerous occasions. In fact, as discussed in the preceding chapter, the same transport protein may have thermodynamically distinct symporter activity for some substrates and channel-like activity for others (Wadiche et al., 1995a and

!. INTRODUCTION The kinetics of mediated biomembrane transport vary greatly among transport processes. For example, the turnover number (i.e., the maximum number of molecules that may pass through a transport pathway per second) of processes historically attributed to membrane carriers may range from about 1/sec (e.g., human erythrocyte phosphate exchanger) to nearly 105/sec (e.g., human erythrocyte chloride exchanger) (Hille, 1984; Stein, 1986). Membrane channels have even greater turnover numbers that typically exceed 106/sec (e.g., Andr6, 1995) and may, in some cases, seem to approach their theoretical maximum (Hille, 1984). Presumably, carriers must undergo a conformational change or interact physically in some way with their substrates to "slow" the rate of movement of the solute or solvent to less than the rate that occurs for channels. In fact, channels are sometimes viewed as forming aqueous diffusion pores that do not present much of a barrier to the migration of their substrates. Nevertheless, many channels appear to be capable of at least one type of conformational change since they are gated. That is, they are normally closed (or open), but they open (or close) relatively rapidly in response to ligands or changes in the electrical membrane potential. In contrast, regulation of the activities of carriers usually occurs more slowly (Chapter 9). Some gated channels are, however, also regulated by relatively slow processes, such as phosphorylation, and many types of channels appear not to be gated. Hence, gating should be viewed as one of many mechanisms of transport regulation rather than as a defining characteristic of the entire set of transport processes known as channels. We view the

65

66

4. Transport Kinetics

see also Chapter 6 for further discussion and additional examples). Although the kinetics of specific instances of each of these types of transport are discussed in this and other chapters, the reader should appreciate that their mechanisms and pertinent kinetic steps may have fundamental similarities which span the range of transporter types. A protein may also have functions in addition to biomembrane transport, and its functions may be modified and regulated by ligand binding or by enzymecatalyzed covalent modification. Accessory proteins may also regulate transporter activity and substrate selectivity, and a given accessory protein might have different effects on different transport proteins. I!. KINETICS OF DIFFUSION A. Low-Molecular-Weight Hydrophilic Ions and Molecules Are Unlikely to M o v e Across Biomembranes by Simple Diffusion Simple diffusion is necessary for virtually all biochemical processes including biomembrane transport. Diffusion is the process by which many substances reach biomembranes, and they must reach the membrane in order to be transported across it. It is, however, frequently assumed that even highly hydrophilic ions and molecules may also permeate biomembranes in significant amounts by a process that resembles simple diffusion. This unwarranted assumption seems to arise, in part, from the frequent discussion in the literature of diffusion and biomembrane transport in the same context. In fact, biomembrane transport is often discussed as if it occurred by simple diffusion. Certainly some of the same thermodynamic and kinetic principles apply to understanding both diffusion and biomembrane transport. Moreover, it has been known for many years that the movement of some substances through biomembranes correlates well with their lipid solubilities. In the cases of substances with high lipid solubility, it is conceivable that they might migrate by diffusion once they have entered the hydrophobic interior of the lipid bilayer (but also see Sections IV-VI below). For most small hydrophilic ions and molecules whose membrane transport is of biochemical interest, however, there is, as yet, no good evidence that they sometimes permeate biomembranes by simple diffusion. Simple diffusion must, by its nature, be associated with relatively low Qa0 values that exceed 1.0 only slightly. The Q~0 value is the degree to which the rate of a biological process increases in association with an increase in the ambient temperature of 10~ ~ For most aThe reader may recognize that Q10 values are proportional to the values of the free energy of activation (AGactivation).While more rigorous theoretical treatments may require the use of A Gactivation values, we think that the use of Q10 rather than AGactivation values makes the present discussion easier to understand.

enzyme-catalyzed reactions, the Qa0 value is above 2.0 as is the case also for most instances of carrier-mediated but not channel-mediated transport (e.g.; Christensen, 1975). In the case of possible biomembrane transport by simple diffusion, the Qa0 value might sometimes be above 1.0 because of other physical effects associated with the membrane, such as the unstirred water layers next to it (e.g., Verkman et al., 1996; Yano et al., 1996). Although the sizes of such layers appear frequently to be overestimated (see Section VI below), the Q10 values that could be associated with the migration of a solute (or the solvent) across such layers are in any case less than 1.5. The only known transport processes for small hydrophilic ions and molecules with Q~0 values below 2.0 are catalyzed by channel proteins. In the case of the water channels in rat cholangiocytes and other cells, temperature does not greatly influence water flux between about 10~ and 30~ (Verkman et al., 1996; Yano et al., 1996). Similarly, the Q10 values for ion channel proteins are about 1.3 (e.g., Frankenhauser and Moore, 1963; Latorre and Miller, 1983). Nevertheless, these proteins catalyze transport by interacting intimately and selectively with their substrates (Chapter 7). Such intimate and selective interactions are, of course, incompatible with the theory that the transport mechanism is, in each case, simple diffusion. Ion channels also show substrate saturation (Section VIII below), again, unlike the process of simple diffusion. In addition to water channels, water moves across biomembranes via a process (or processes) that does not appear to involve proteins. In this instance, however, the Q10 value appears to be greater than 2.0 (Verkman et al., 1996; Yano et al., 1996). This relatively high Qx0 value for a process that is not mediated by a protein appears to be due, at least in part, to the need to break hydrogen bonds between the water molecules that are to migrate across the membrane and other water molecules on either side of the membrane (Christensen, 1975; Verkman et al., 1996). Moreover, a rather precisely defined, temperature-sensitive process has been proposed to be the way in which individual water molecules may migrate through the lipid bilayer in the absence of water transport proteins (Haines, 1994). In the later case, temperature is thought to influence the rate of migration because the membrane must be fluid enough to form spaces for the migrating water molecules. Although this water migration may be called diffusion by some authors, it hardly qualifies as diffusion in the ordinary sense. Would not similarly complex processes also be needed for any other relatively small ion or hydrophilic molecule to cross the membrane without the aid of an integral membrane protein? In this regard, the bilayers of artificial phospholipid membrane vesicles have very low but measurable permeabilities to inorganic ions,

Kinetics of Diffusion such as Rb + and Na + (e.g., E1-Mashak and Tsong, 1985). The Q~0 values of these permeabilities are, however, about 3.0, so migration of the ions across the bilayers does not resemble ordinary diffusion. In spite of these considerations, we derive below mathematical expressions for the diffusion of water and other hydrophilic substances from one side of a membrane to the other. Components of the expressions are determined experimentally and relate to physiologically important concepts for the movement of substances across membranes. In our view, however, a more complex transport process always underlies transport that is treated as simple diffusion. In other words, the values of the resistance of the lipid bilayer as a barrier to diffusion, R, or the inverse of this resistance which is the diffusion coefficient, D, approach infinity or zero, respectively, for simple diffusion of low-molecularweight hydrophilic substances across biomembranes. The other values for these or related parameters that have been published in the literature appear actually to reflect protein-mediated or other complex transport processes, although the processes catalyzing transport may not, as yet, have been identified and studied experimentally.

B. M o s t L o w - M o l e c u l a r W e i g h t Hydrophilic S u b s t a n c e s Reach the Sites of Their B i o m e m b r a n e Transport by Simple Diffusion at the Cellular a n d Subcellular Levels Simple diffusion of small hydrophilic substances clearly is required in order for most biochemical processes including biomembrane transport to proceed. In order to be transported through a biomembrane, ions or molecules must make their way to the membrane through an aqueous solution. The random motion of ions or molecules due to thermal energy is an extremely effective means for their migration over short distances such as those needed to reach biomembranes at the cellular and subcellular levels (Christensen, 1975). In fact, it is unlikely that any membrane transport process can catalyze the migration of a low-molecular-weight solute at a rate faster than the rate of diffusion of the substance in water. Diffusion is a primarily entropy-driven process (Table 3.1). The random motion of a solute and the solvent in which it is dissolved help to determine the rate at which non homogenous mixtures of the substances approach homogeneity by diffusion. Thus, diffusion of a solute (depicted by dark circles in Fig. 4.1) is influenced not only by the random motion of the ions or molecules, but also by the nature and motion of the solvent molecules (light circles in Fig. 4.1). The solvent should also be viewed as diffusing into the region of high concentra-

67

FIGURE 4.1 Both solute (dark circles) and solvent (light circles) migrate by simple diffusion into regions in which the other is more concentrated. The rates of their migration depends both on their own velocity of motion and on the velocity and the resistance to motion of the other substance.

tion of the solute, and this diffusion is influenced by the nature and motion of the solute ions or molecules. Hence both of these cases of diffusion are limited by the resistance of the other substance as well as by the velocities of the ions or molecules of the diffusing substance. The kinetic expressions for diffusion of both solute and solvent take similar form. C. Derivation of M a t h e m a t i c a l Expressions for Simple Diffusion of a Solute The rate net of migration of a solute by diffusion is proportional to the difference in its concentration in two adjacent spaces, such as sides A and B of the container depicted in Fig. 4.2. This proportionality may, however,

FIGURE4.2 Diffusionof substance C will occur from side A to side B of a vessel through an opening in an otherwise impermeable barrier.

68

4. Transport Kinetics

not be the same for all instances of the same concentration difference, since the viscosities of the solutions may increase or decrease with changes in solute concentrations. The rate of solute migration is also inversely related to the resistance to migration of the solvent as well as the inherent limitations to movement of the solute due to its size, shape, and attraction to or repulsion from like solute ions or molecules and the solvent. Also represented in Fig. 4.2 is an impermeable barrier with an opening for the diffusion of the solute. Clearly such barriers and openings also influence the flux of the solute; the flux will, of course, be faster when the openings are larger or more numerous and slower when they are smaller or fewer in number. Here, the main purpose of the barrier is to define compartments A and B. We intentionally avoid stating or implying that the barrier may be a biomembrane with holes or pores large enough to permit simple diffusion. Based on the evidence discussed above (and below), we think that it is unlikely that biomembranes have such pathways for the migration of small hydrophilic ions and molecules. The net flux of the solute, C, from side A to side B of the container depicted in Fig. 4.2 can be represented by the equation vi = A (aCA -- aCB)/Rd,

(4.1)

where vi is the initial velocity of net migration of C, A is the area of the hole in the barrier, d is the width of the barrier, R is the resistance to flux of the solute, and aCA and aCB are the activities of C on sides A and B of the container, respectively. The resistance to flux, R, also is defined as the inverse of the diffusion coefficient. Hence, Eq. (4.1) may be modified to become a form of Fick's first law of diffusion 12 i :

DA

(aCA- acB)/d,

(4.2)

where D is the diffusion coefficient. Inspection of Eq. (4.2) will show that for vi to be the velocity of net movement of a quantity of substance C from compartment A to compartment B (e.g., moles sec -1), D must have the units of area/time (e.g., c m 2 sec-1). These units are also consistent with the relationship between the diffusion coefficient for ions or molecules and the time that the ions or molecules take to diffuse an average distance at a given temperature (Einstein's relationship). The relationship between D and this average distance is given by the equation D = b2/2t,

(4.3)

where b is the average distance traveled by such ions or molecules after time t. It is also sometimes useful to define the rates of diffusion of ions or molecules in terms of their permeability coefficients rather than their diffusion coefficients. If

we proceed carefully, it is possible to use permeability coefficients to describe not only the migration of ions or molecules in free solution, but also their migration across membranes, without implying that they cross it by simple diffusion. The term "permeability coefficient" is, however, misleading to the degree that it implies permeation of the lipid bilayer by solutes or the solvent. It is incorrect to view transport proteins as increasing the abilities of ions and molecules to permeate the lipid bilayers of membranes. Rather, these proteins each produce pathways for the migration of specific substrates across membranes. Permeability coefficients were, however, defined before we appreciated that migration of small hydrophilic ions and molecules across biomembranes occurs principally via relatively complex, protein-catalyzed processes rather than because the ions or molecules sometimes simply permeate the lipid bilayer. With these caveats in mind, reference to permeability coefficients may allow us to discuss the migration of solutes across membrane barriers without necessarily evoking the concept of simple diffusion which is, of course, evoked when we refer to diffusion coefficients. In addition, the use of permeability coefficients helps us to avoid the need to measure accurately the width of the biomembrane barrier, which may not have a precise or constant value. For example, the processes that are studied in physiological experiments frequently are the migration of solutes or the solvent via proteins that may vary in structure or conformation and hence in their effective widths. Permeability coefficients can be used to represent these complex transport processes, but again it must be remembered that none of the processes is simple diffusion. In this context, we may define the permeability coefficient, P, as P = D/d.

(4.4)

Equations (4.2) and (4.4) may now be combined to produce 1;i =

P A ( a C A - aCB).

(4.5)

D. Parallel Expressions for the Diffusion of Water and for Its Migration across B i o m e m b r a n e s While it is possible to write an expression similar to Eq. (4.2) for the simple diffusion of water in a purely aqueous environment, we are usually more interested in the migration of water across biomembranes. In the latter case, it is often useful to express the quantity of water as a volume rather than a number of moles. In order to express migration of water in this way, Eq. (4.5) can simply be multiplied on both sides by the

69

Kinetics of Diffusion

molar volume of water. The value of this parameter is approximately 18 ml mo1-1, although it depends somewhat on the ambient temperature and pressure. Since it is safe to assume that in most biological experiments our conclusions will not be greatly altered by the effects of temperature and pressure on the molar volume of water, we may modify Eq. (4.5) for the migration of water as follows V s pi = V s

P A (aWA -- awB),

(4.6)

where Vs is the molar volume of water and awA and awB are the activities of water in compartments A and B, respectively. It is also convenient to use solute rather than solvent activities and, as in the previous chapter, concentrations may be assumed to be nearly equal to the corresponding activities. Since the solutions on each side of the barrier in Fig. 4.2 are composed only of solutes and the solvent, the difference between the concentrations of solvent in the two compartments is inversely related to the difference between the total concentrations of dissolved ions and molecules in each compartment. The total concentrations of dissolved substances is, of course, the total osmolarity. Since total osmolality can, however, be easier to measure than total osmolarity, it is often used in place of total osmolarity without greatly changing our conclusions. Hence, Eq. (4.6) is frequently rewritten as V s vi :

Vs

P A (OSmB -- OSmA),

(4.7)

where OSmA and OsmB are the osmolalities of the solutions in compartments A and B, respectively. Finally, a single term, the initial transmembrane net volume flux of water, Jvi, is frequently substituted for Vs vi. Since Jvi = Vs vi,

Eq. (4.7) can be written Jvi = Vs P A (OSmB -- OSmA).

(4.8)

E. Derivation of Expressions for the Migration of Lipid-Soluble Substances across Biomembranes As discussed above, migration of low molecular weight hydrophilic substances across biomembranes does not appear to occur by simple diffusion. Nevertheless, it may sometimes be useful to measure the apparent diffusion and permeability coefficients associated with migration of hydrophilic solutes or water across the membrane (see Section III below). In addition, we will discuss in Section IV below why migration of lipidsoluble substances across biomembranes is thought by some investigators to occur by ordinary diffusion. In the latter cases, the permeability coefficient, P, is sometimes

redefined in terms of a new diffusion coefficient, Dmem, which is intended to represent the diffusion coefficient of the solute within the hydrophobic region of the membrane. One also needs to consider a third term, the partition coefficient, fully to derive P for the apparently simple diffusion of a solute through the hydrophobic region of a lipid bilayer. The pertinent partition coefficient is the ratio between the activity of a solute dissolved in the hydrophobic region of the plasma membrane divided by the activity of the same solute dissolved in the cytosol or extracellular fluid when the solute has reached equilibrium between dissolution in these places. Clearly, the pertinent partition coefficient is complex not only because the cytosol and extracellular fluid have different compositions, but also because the phospholipid bilayers of biomembranes are highly ordered, heterogenous structures (as described in Chapter 2) unlike oils or pure organic solvents. It is, however, easier to measure the partition coefficient of a solute between oil or a pure organic solvent and water than between the hydrophobic region of a membrane and water. Moreover, partition coefficients measured between water and some oils and organic solvents permit calculation of permeability coefficients that appear to apply reasonably well to the membrane transport of several nonionic substances (see Stein, 1986, for a more detailed account). The reader is, however, cautioned that when the partition coefficients between biomembranes and water solutions have been measured, they sometimes differ dramatically from those for oil and water. For example, the value of the partition coefficient of a-tocopherol between oil and water is greater than the value for the more hydrophilic derivative, tocopherol succinate, but the magnitude of these values is reversed for partitioning between the erythrocyte membrane and water (Bonina et aL, 1996). The relationship between the permeability coefficient, P, and the partition coefficient, Kp, can be derived by first considering the mathematical representation of Kp g p = aCAM/aCA = aCBM/aCB,

(4.9)

where aCA is the activity of substance C in compartment A (now more usefully defined as the extracellular fluid of a cell, Fig. 4.3), aCB is the activity of C in compartment B (i.e., the cytosol), aCAM is the activity of C in the membrane in equilibrium with the extracellular fluid, and aCBMis the activity of C in the membrane in equilibrium with the cytosol. Since aCB and aCA can be determined for the solutions of C in the cytosol and extracellular fluid, respectively, aCAM and aCBM can be calculated from Eq. (4.9) to be

70

4. Transport Kinetics

for migration of hydrophilic substances across lipid bilayers).

!II. H O W DO MEASUREMENTS OF BOTH THE DIFFUSIONAL AND THE OSMOTIC PERMEABILITY COEFFICIENT FOR WATER INFORM US ABOUT THE MECHANISM OF WATER TRANSPORT ACROSS A PLASMA MEMBRANE?

FIGURE 4.3 Migration of substance C from one side of a phospholipid bilayer to the other also involves migration of the substance through the hydrophobic region of the bilayer. In the latter case, CBM is in equilibrium with C in the cytosol and CAM is in equilibrium with C in the extracellular fluid. The details of the processes by which C crosses the surfaces of the bilayer or by which C traverses its hydrophobic interior are not meant to be reflected in the scheme shown.

aCAM = Kp aCA

(4.10)

aCBM = Kp aCB.

(4.11)

and

Equation (4.2) can now be rewritten for a membrane by substituting DME M for D and the activities of C in the membrane for the activities of C in the aqueous phases (and also replacing the barrier depicted in Fig. 4.2 with the membrane represented in Fig. 4.3) vi = Dmem A(Kp aCA -- Kp acs)/d or

vi = Dmem A Kp(aCA- acs)/d

(4.12)

From Eq. (4.12) it can be seen that, for substances to which organic solvent/water Kp values apply, P can be defined for diffusion of the substances in the hydrophobic region of biomembranes as P = Dmem Kp/d.

(4.13)

While Eqs. (4.12) and (4.13) are believed by some investigators to apply reasonably well to the migration of lipid-soluble substances across biomembranes, diffusion and permeability coefficients that are measured for hydrophilic solutes and for water clearly do not represent simple diffusion (e.g., recall that Q10 values exceed 2.0

As discussed above, water does not pass across lipid bilayers by a mechanism that resembles simple diffusion. It also does not move through the pathways created by water channel proteins in a manner that resembles ordinary diffusion. In this regard, some measurements of the values of the water permeability coefficients have been used to draw conclusions about the nature and the dimensions of the water pathways that are produced by channel proteins. If these conclusions are correct, then measurement of the so called diffusional and osmotic water permeability coefficients, while inaccurate in regard to the mechanisms of water transport that their names imply, may, nevertheless, constitute a useful means of assessing water transport. In numerous cell types, the osmotic water permeability coefficient has been observed to exceed the diffusional water permeability coefficient by severalfold. Such has also been found to be the case even for gramicidin channels incorporated into artificial lipid bilayers, although it does not appear to be true for the artificial bilayers alone. As for bilayers that do not contain proteins or peptides, the values of the diffusional and osmotic water permeability coefficients appear to become equal when the function of water channels in cell membranes is inhibited with mercurials. Such data have sometimes been interpreted to mean that water moves single file through channels and that the ratio of the values of the permeability coefficients is a measure of the number of water molecules that span the channel. The reasoning involved in this interpretation depends on knowledge of the differences in the ways in which the two permeability coefficients have been measured. The osmotic permeability coefficient, Pf, is usually determined by measuring net water flux into or out of cells under hypotonic or hypertonic conditions, respectively. The permeability coefficient, P, in Eq. (4.8) can be designated Pf for net osmotic water flux Jvi = Vs P f A ( O s m s

-- OSmA)

(4.8)

and this slightly modified form of Eq. (4.8) can be rearranged to calculate Pf

Pf = Jvi/(Vs A (Osms -- OSmA))

(4.14)

Diffusional and Osmotic Permeability Coefficient

where A is the plasma membrane surface area, Jvi is the transmembrane net volume flux of water, Vs is the molar volume of water (approximately 18ml/mol), and OSmB and OSmA are the osmolalities of the cytosol and extracellular fluid, respectively. OSmB can be measured by exposing cells to various solutions and determining which is isotonic. Similarly, OSmA can be established experimentally for the extracellular fluid. Jvi can then be determined by measuring the initial rate of change of cell volume on exposure of cells to hypotonic or hypertonic conditions. Similarly, the initial dimensions of the cell can be used to calculate the plasma membrane surface area, A. On the other hand, the diffusional water permeability coefficient, Pd, historically has been determined under isotonic conditions, so a different procedure must be used to measure water flux. Tritiated water may be used conveniently for such measurements, and its somewhat greater molecular weight relative to unlabeled water is not expected to alter greatly measurement of water flux. Pd can be calculated by measuring the initial rate at which tritiated water crosses the membrane. Equation (4.5) for water (instead of a solute) is 12i =

P A (aWA --

aws)

(4.15)

and this equation may be modified to read vi* = PdA [3H20],

(4.16)

where 1,'i* is the initial rate of movement of tritiated water across the membrane, A is the plasma membrane surface area, and [3H20 ] is the initial molar concentration of tritiated water on one side of the membrane or the other (activities are again assumed to be virtually identical to concentrations). In order to calculate P d , Eq. (4.16) may be rearranged to give Pd

=

vi*/A[3HzO]

9

It might be anticipated that the value of Pd would exceed the value of Pf somewhat, since the former represents unidirectional water flux, whereas the latter is for net flux. As just discussed, however, Pf appears to exceed Pd by severalfold in numerous types of cells. A frequent interpretation for the difference between Pf and Pd is that Pf exceeds Pd by an amount approximately equal to the number of water molecules inside water channels. It is reasoned that for, say, a single-file channel containing 10 water molecules, 10 such molecules would need to leave the channel before the first tritiated water molecule could pass across the membrane. In contrast, no such delay would be observed in detecting water movement owing to an osmotic gradient. Each water molecule would contribute to a change in cell volume as soon as the molecule appeared in or left the cytosol. Although this assessment may, at first, seem reasonable,

7 |

the real error in it appears to lie in the assumption that membrane channel proteins could contain enough water significantly to influence the volume of the cell in the absence of movement of water entirely from one compartment to the other. The volume of water contained in water channel proteins of course cannot exceed the volume of the membrane itself. It can be calculated that the membrane of a typical cell constitutes about 0.07% to its volume. The changes in cell volume that are actually measured in order to calculate Pf from Eq. (4.14) may, however, exceed the normal cell volume by 30% or more (e.g., Roberts et al., 1994; Yano et aL, 1996; Suleymanian and Baumgarten, 1996). Hence, even if the entire volume occupied by the plasma membrane were water, it could contribute no more than about 0.2% of the observed volume changes (0.07/30 = 0.2%). For this reason, we think it is unlikely that comparisons of Pf to Pd values lead to insight into the number of water molecules that may fill single-file channels. Our conclusion is supported by the rigorous theoretical treatment of Manning (1975). According to this author, the value of Pd should equal the value of Pf even for pores so narrow that they would permit only the single-file passage of water molecules. Hence, measurement of these values cannot be used to calculate the number of water molecules that channel proteins may contain. More recent theoretical assessments of the physics of water migration through channels appears to provide more sophisticated views of the possible dimensions of channels (e.g., Hill, 1995). Nevertheless, these views seem still not fully to consider that the nature of the interaction between a water molecule and its transport protein is probably more intimate and selective for water than would be predicted if friction were the principle impediment to migration. As discussed above, it appears unlikely that water or hydrophilic solutes migrate across biomembranes by processes that resemble ordinary diffusion through narrow pores regardless of whether the conditions are isotonic. While measurement of apparent Pd and P~ values may provide useful information, investigators should remember that neither value really represents simple diffusion of water across biomembranes. Nevertheless, if we can determine what the values of Pd and Pf really mean, then they may yield insight into how water channels function or how they are regulated. In this regard, we think that the differences in the osmotic and the diffusional water permeability coefficients may reflect a regulatory increase in the flux of water through channels under nonisotonic conditions. It is, in fact, quite reasonable that an osmotic gradient could increase the rate of migration of water molecules through channels by opening them further. Such gradi-

72

4. Transport Kinetics

ents for cells under physiological conditions are usually not as large as the gradients used to study channels experimentally (e.g., Roberts et aL, 1994; Yano et al., 1996; Suleymanian and Baumgarten, 1996). Hence, it seems unlikely that cells would usually benefit from attempting to resist osmotically induced water flow for self-protective purposes. In fact, important signaling processes that alter cell metabolism may result from cell swelling or cell shrinking (Fig. 4.4) (Graf and H~iussinger, 1996). Consequently, their ability to change volume rapidly could be important to normal cell functioning. Moreover, solute channels, such as those for taurine (e.g., Van Winkle et al., 1994), open in response to hypoosmotic stress presumably to produce a regulatory volume decrease (Fig. 4.4). In the latter case, water channels are expected to increase in activity more rapidly than solute channels, since the regulatory volume decrease follows an increase in cell volume. In contrast, volume changes in hepatocytes in response to insulin or glucagon (Graf and H~iussinger, 1996) likely result

Institution of Effector

RVD t-.

,,m

"anabolic" 09

O

Insulin

Glucagon "catabolic"

t~ t...

,,..~ I,.,,

r

CO

RVI

FIGURE4.4 The cell swelling or cell shrinking that occurs in hypotonic or hypertonic conditions is usually followed by a regulatory volume decrease (RVD) or a regulatory volume increase (RVI), respectively. Osmoticallyinduced volume changes may occur more rapidly than anticipated from the measured capacity to transport water under isotonic conditions (see discussion in test). We propose that the opening of water channels under other-than-isotonic conditions increases water flux. Subsequent opening of channels for solutes could produce an RVD, whereas active transport of solutes appears to be needed for an RVI. In contrast, increased solute transport may precede increased water fluxin order to produce the volume changes associated with insulin or glucagon treatment of hepatocytes. More importantly, the concomitant opening of water channels to permit more rapid cellular shrinkage or swelling could help to cause the metabolic changes associated with exposure to these hormones. While the presence of water channel proteins in hepatocytes is still controversial (e.g., Yano et al., 1996; Tsukaguchi et al., 1998), these proteins are present in most other cell types, and their transport of water contributes to changes in cellular volume (adapted from Graf and H~iussinger,1996,with permission from the European Association for the Study of the Liver).

from changes in solute transport followed by water transport. In either case, we propose here that water channels open in response to osmotic imbalances, possible as a result of protein phosphorylation (Johansson et al., 1998), and that the more rapid water flux serves important physiological functions, such as the signaling that results from changes in cellular volume (Fig. 4.4). It should be evident from the preceding discussion that we think the difference between the values determined for Pd and Pf depend on the different osmotic conditions under which they are measured rather than that they have inherently different values, which depend on the number of water molecules that span water channels. For these reasons, we predict that the Pd values determined by measuring migration of isotopically labeled water will exceed or equal Pf values determined by measuring cellular volume changes when the two permeability coefficients are determined under the same osmotic conditions. Somewhat surprisingly, we know of no published experiments in which these water permeability coefficients have intentionally been determined under identical osmotic conditions. The two values have, of course, been determined under the same conditions when the conditions were isotonic. In these cases, the values of osmotic water permeability coefficients are zero by definition, whereas the values of diffusional water permeability coefficients are above zero. We predict that the ratio of the two coefficients and the absolute values of the coefficients themselves will increase rapidly as the osmotic pressure difference across the membrane is increased in cells that have water channels. In our view, the ratio of Pf to Pd will approach but will not exceed a value of one when they are measured under the same osmotic conditions. Pd should always exceed Pf somewhat no matter what the osmotic difference across intact cell membranes, since the unidirectional flux of radiolabeled water molecules is measured to determine Pd , whereas Pf will always reflect the net migration of water (assuming that the amount of water initially in the m e m b r a n e is negligible relative to the amount of water that migrates). The requisite migration of some water in the reverse direction will always reduce somewhat the Pf value that is calculated from volume changes. For example, water moves into cells under hypotonic conditions to increase their volume, while some solutes, such as taurine, migrate out of the cells. Both water and taurine actually move in both directions, however, as can be demonstrated by measuring u p t a k e of radiolabeled taurine under hypotonic conditions (e.g., Van Winkle et al., 1994). Hence, P values determined by measuring the rates of net migration of water or a solute always are lower than the values that would be obtained from unidirectional flux. In the case of water transport, the

Lipophilic Substance Migration Pd value will always exceed the Pf value by a finite a m o u n t when these p a r a m e t e r s are m e a s u r e d under the same conditions. It remains to be d e t e r m i n e d w h e t h e r the Van Winkle theory or some other one provides the correct explanation for differences b e t w e e n the Pd and Pf values that have been reported for water transport. 2 More important is that the determination of the values of Pd and Pf has stimulated us to seek explanations for their relative values. Such explanations m a y give us insight into both the structure and the physiological functioning of water channels, especially if the two coefficients are m e a s u r e d simultaneously as described above, but under a variety of osmotic conditions. In order accurately to interpret these data, it is also necessary to r e m e m b e r that osmotic and diffusional permeability coefficients do not usually reflect permeation of the lipid bilayers of b i o m e m b r a n e s by water molecules. Moreover, the diffusional permeability coefficient never reflects simple diffusion of water across b i o m e m b r a n e s , contrary to what was believed when these coefficients were first formulated. Similarly, m e a s u r e m e n t of permeability coefficients for ionic and nonionic solutes can inform us about transport of the solutes and the significance of the transport to cellular physiology even though the solutes usually do not cross b i o m e m b r a n e s by permeating lipid bilayers. Accurate determination of apparent permeability coefficients requires, however, that we consider the effects 2 There is a third possible explanation for the observation that Pf exceeds Pd by about 10-fold because the former is measured in the presence of an osmotic gradient whereas the latter is not. In this third explanation it must be proposed that the number of water molecules transported together via water channels (i.e., their flux coupling) is an unexpectedly large number. In support of this possibility, Wright and associates (Loo et al., 1996b) have shown that about 260 water molecules appear to be transported with each glucose molecule by the Na+-dependent glucose transporter. If the stoichiometry of water transport via channels is also very large, then the velocity of water transport could be more rapid in one direction than in the reverse direction under anisosmotic conditions. For example, if 850 water molecules were transported together by a water channel, then a 10fold difference in the flux ratio (i.e., influx/efflux) would be observed when the transport of water into and out of cells in a solution of half the osmolarity of an isotonic solution is compared to the transport in an isotonic solution. The ratio of the water concentrations would be raised to a power of 850 to calculate the flux ratios of 10 and 1, respectively, for these two conditions (Eq. (7.1) in Chapter 7). If the unidirectional water flux from isotonic solutions is always the same, then the ratio of Pf to Pd would also be 10 in this case. In contrast, for the transport of only 10 water molecules at a time (or their occupation of a water channel together as discussed in the text), a flux ratio only slightly different than one is expected under anisosmotic conditions. Even the concentration of pure water is only slightly above that of water in an isotonic solution, so such a concentration ratio of intracellular to extracellular water would produce a flux ratio of only about 1.06 after the concentration ratio is raised to a power of 10 (see Section III,B of Chapter 7 for further discussion of the effect of flux coupling on the flux ratio.)

73

of other possible barriers to the m e a s u r e d migration of substances across biomembranes. In particular, unstirred water layers next to m e m b r a n e s are believed by some investigators to influence m e a s u r e d fluxes of solutes and the solvent, although we believe the importance of these layers may be greatly exaggerated (see Section VI below). Before we discuss estimation of the dimensions of the unstirred water layers and their putative effects on transport in Section V below, we must first consider the transport of lipid-soluble substances. Such lipophilic substances are used in experiments to estimate the width of unstirred water layers. Unlike water and highly hydrophilic ions and molecules, lipidsoluble substances appear more likely to p e r m e a t e biom e m b r a n e s by a mechanism that may resemble simple diffusion.

IV. D O LIPOPHILIC SUBSTANCES MIGRATE ACROSS BIOMEMBRANE PHOSPHOLIPID BILAYERS BY SIMPLE DIFFUSION? It is useful to think about the steps that may need to occur in order for relatively small, lipid-soluble molecules, such as those of n-butanol, to move from one side of a m e m b r a n e lipid bilayer to the other unaided by a protein catalyst. The molecules first diffuse through a bulk water solution and then through an unstirred water layer on one side of the m e m b r a n e . They then migrate across a w a t e r - p h o s p h o l i p i d interface and subsequently into the more hydrophobic interior of the lipid bilayer. By reversing these steps for the other leaflet of the bilayer, the molecules will appear in the bulk water solution on the other side of the m e m b r a n e . While one might envision that a barrier could be e n c o u n t e r e d at several of these steps, the barrier is usually assessed in terms of simple diffusion in the water phases, simple diffusion in the hydrophobic interior of the m e m b r a n e , and partitioning of the lipid-soluble molecules between the two. The preceding partitioning seems, however, less important to migration of a solute across the lipid bilayer than partitioning, say, b e t w e e n the area of the hydrophilic, phosphate-containing heads of the m e m b r a n e phospholipids and the hydrophobic interior of the membrane. To our knowledge, the latter partitioning has never been adequately modeled or studied. With such caveats in mind, let us consider why sufficiently lipophilic substances are considered by some investigators not to experience much of a barrier in moving from the water next to one side of a m e m b r a n e to the water next to the other side. In Section VI below we explain why we think that b i o m e m b r a n e s are significant barriers to the migration of virtually all substances. Interested read-

74

4. Transport Kinetics

ers are referred to more detailed experimental and theoretical treatments (e.g., Stein, 1986) of the proposed importance of diffusion and partitioning between water and the hydrophobic interior of the phospholipid bilayer in the overall process of membrane permeation by various substances. The classic studies of Collander and B~irlund (1933) with algal cell membranes (e.g., Fig. 4.5) have been repeated in several laboratories for a variety of other cells and even for artificial lipid bilayers. These experiments demonstrate more or less linear relationships between the log10 of the permeability coefficients (P) of numerous substances and the log10 of their partition coefficients (K1,) between water and nonpolar organic solvents. The latter solvents are selected to resemble the partitioning that is anticipated between water solutions and the hydrophobic regions of biomembranes because the actual partitioning between water solutions and these hydrophobic regions may be difficult or impossible to measure accurately. For these reasons, the original partitioning studies were performed with water and olive oil (Fig. 4.5), and hexadecane has been used more recently to represent biomembranes (e.g., Stein, 1986). When the size (i.e., the molecular volume) of the permeant is used to adjust the value of P, a very good

.i

BBBB

Methyl

i

.,=..001

Ethylalcohol~ Urethylan0

_AO

E

I

Trimtyl citrate

Methyl~)

t5

EthylumPM~

O

OThioure a

$

UreaO LadamideO Diethylmalonamide 0 Urotrophin 0.001

--9 Methylolurea

O (la~=,rol

0

"0

"" Dirnethylurea

~D

0.01

s

d~l)Glycerol mety~lurPa rl emer

0

E

Some substances, such as n-butanol molecules, are assumed to be lipid-soluble enough to pass through the membrane lipid bilayer unimpeded except by their rate of ordinary diffusion. That is, the phospholipid bilayer is not considered to be a barrier to the diffusion of these

Ur;ane Triethyl citrateO

Drol ethylether

Acetamide,~ Glyc~ '~

0

0

V. LIPID-SOLUBLE SUBSTANCES ARE USED TO ATTEMPT TO MEASURE THE WIDTH OF UNSTIRRED WATER LAYERS ON EITHER SIDE OF BIOMEMBRANES

Cyanimide~D ~Va~itl~ll~e Pmpi~ ~noc~lomhy~ Dm~tin

0.1

~=~

i

correlation is observed between the log10 of the sizecorrected P values and the log10 of the Ke values (Figs. 4.6 to 4.9). Such results for a variety of substances are taken as evidence that sufficiently lipid-soluble substances are not slowed during their migration across biomembranes owing to diffusion through the phospholipid bilayer. It is worth noting, however, that the loga0 of the strength of interaction of organic cations with a transporter in the proximal tubule of rat kidney is directly proportional to the log~0 of their hydrophobicity (Ullrich et al., 1992). Hence, some instances of apparent migration of hydrophobic substances through the lipid bilayer may actually result from their protein mediated transport.

-2

10,

._N (.9 O ._1

O Dieyandiamide

e'

.-4

Q.

e9

10 Malonamide

0.0001

e~o, 0.0001

I

I

0.001

0.01

-6 -7

I,i, 0.1

1.0

Partition coefficient

FIGURE 4.5 The permeability of algal cells increases with the lipid solubilities of various solutes. Lipid solubilities are considered to increase as the olive oil/water partition coefficients of permeants increase. The partition coefficient, Kp, is the ratio between the activity of a solute dissolved in olive oil to the activity of the solute in water when the solute has reached equilibrium between the two solvents (adapted from Collander, 1937 with permission from the Royal Society of Chemistry).

i

I

-5

-3

_,

, I, -1

,

I 1

L O G K hexadecane

FIGURE 4.6 Relationship between the lipid solubilities of several substances and their size-corrected permeability coefficients for the human erythrocyte plasma membrane. Partition coefficients (Kp) between hexadecane and water are taken as the pertinent measures of the relative lipid solubilities. See text for discussion of the method of correcting the permeability coefficients (P) for molecular size. The substances are: 2, ethanediol; 3, ethanol; 4, glycerol; 5, n-hexanol; 6, methanol; 7, n-propanol; 9, urea; and 10, water. (adapted from Stein, 1986, with permission from Academic Press).

Lipid-Soluble Substances

75

4

5

$6

s

40

"O O

(I,)

15 r O O

O O

6 N co v

6 N

v

L9 O

o

._J

._1

8e

e5

le -2 L -3

-2

-1

LOG K

0

1

hexadecane

-6

I

-7

-6

,

,

LOG K

I

-5

I

-4

,,

L

-3

FIGURE 4.7 Correlation between the size-corrected permeability coefficients of six n-alcohols for the dog red cell membrane and their hexadecane/water partition coefficients (Kp). The partition coefficients are considered to be a measure of the relative solubility of the alcohols in the interior of the phospholipid bilayer of the plasma membrane. The method used to correct the permeability coefficients (P) for size is discussed further in the text. The number of carbons in the n-alcohols is indicated by the number next to the data point (e.g.; 1 = methanol through 6 = n-hexanol) (adapted from Stein, 1986, with permission from Academic Press).

FIGURE 4.8 Permeability of the algal cell membrane (Chara ceratophylla) to several nonelectrolytes of varying lipophilicity. Lipid solubilities are measured as the partition coefficients (Kp) of substances between hexadecane and water. The method of size-correction of the permeability coefficients is discussed in the text. The substances are: 1, formamide; 2, acetamide; 3, n-propionamide; 4, n-butyramide; 5, urea; 6, ethanediol; 7,1,2-propanediol; 8, glycerol (adapted from Stein, 1986, with permission from Academic Press).

substances f r o m the bulk w a t e r solution on o n e side of the m e m b r a n e to the bulk w a t e r solution on the other. N e v e r t h e l e s s , the rates of a p p e a r a n c e or loss of the substances in o n e or the o t h e r of the bulk w a t e r phases is slower t h a n a n t i c i p a t e d for o r d i n a r y diffusion across a p h o s p h o l i p i d bilayer only a b o u t 5 n m thick. H e n c e it is a s s u m e d that this a p p a r e n t lack of m i g r a t i o n at the a n t i c i p a t e d rate is d u e to a n e e d for the substances also to diffuse t h r o u g h u n s t i r r e d w a t e r layers on each side of the b i o m e m b r a n e b e f o r e their m i g r a t i o n f r o m o n e b u l k w a t e r p h a s e to the o t h e r can be detected. F o r these reasons, it is also a s s u m e d that such substances as n - b u t a n o l can be used to d e t e r m i n e the width of the u n s t i r r e d w a t e r layers. If these a s s u m p t i o n s are correct, t h e n the thickness of the unstirred w a t e r layers are easy to d e t e r m i n e using Eq. (4.4), which is

substance a p p e a r s to diffuse. This f o r m u l a seems to apply b e c a u s e P is d e t e r m i n e d for the m i g r a t i o n of the substance f r o m the bulk w a t e r on o n e side of the m e m b r a n e to the bulk w a t e r on the o t h e r side. Since the m e m b r a n e is t h o u g h t n o t to be a barrier, we are really a t t e m p t i n g to e s t i m a t e the distance, d, o v e r which the substance on o n e side of the m e m b r a n e m u s t m o v e in o r d e r for us to detect it on the o t h e r side. This distance is t a k e n as the width of the u n s t i r r e d w a t e r layers the c o m p o s i t i o n s of which a p p a r e n t l y are not easily a m e n a ble to direct assay. F o r the h u m a n red b l o o d cell p l a s m a m e m b r a n e , P for b u t a n o l has b e e n f o u n d to be a b o u t 6 • 10 -2 cm sec -1. M o r e o v e r , D for b u t a n o l in w a t e r is a b o u t 1 • 10 -5 cm2sec -1. H e n c e the thickness of the m e m b r a n e plus its associated u n s t i r r e d layers can be calculated f r o m Eq. (4.4) to be

P = D/d,

(4.4)

w h e r e P is the p e r m e a b i l i t y coefficient m e a s u r e d for m i g r a t i o n of the substance f r o m the bulk w a t e r on o n e side of the m e m b r a n e to the bulk w a t e r on the o t h e r side, D is the diffusion coefficient of the substance in w a t e r (since the m e m b r a n e is a s s u m e d not to be a barrier), and d is the width of the space across which the

P-

D/d

d -

D/P

hexadecane

d - 10 -5 cm 2 sec-1/6 • 10 -2 cm sec -1 d = 1.7 • 10 .4 cm d = 1.7/xm.

(4.4)

76

4. Transport Kinetics 0

-

40

=,,=,

E" "O (D

=,=

13 -2

6

8 N

r,9 O

-

-3

10/e 2

/

...J

O5

-7

i -6

I -5

,

I ...... -4

I -3

,I -2

LOG K hexadecane

FIGURE 4.9 As for biomembranes, artificial phospholipid bilayers show a linear relationship between the log10 of the size-corrected permeability coefficients of several substances and the log10 of their hexadecane/water partition coefficients. The partition coefficients (Kp) are considered to be a measure of the relatively solubilities of the substances in the interior of the bilayer. Permeability coefficients were corrected for size as discussed in the text. The substances are: 1, formamide; 2, acetamide; 3, n-propionamide; 4, n-valeramide; 5, urea; 6, water (adapted from Stein, 1986, with permission from Academic Press).

Since the width of the plasma membrane even with the glycocalyx is less than 100 nm, most of this distance is attributed to unstirred water layers. Presumably, one would repeat such determinations for several lipidsoluble substances and obtain similar estimates of the width of the unstirred layers.

Vl. DO SUCH DETERMINATIONS OF THE APPARENT WIDTHS OF UNSTIRRED WATER LAYERS REFLECT THE INTENDED PHYSICAL P H E N O M E N O N OR O U R IGNORANCE OF H O W LIPID-SOLUBLE SUBSTANCES CROSS BIOMEMBRANES? The assumptions and calculations above for the human erythrocyte may at first seem quite reasonable. When, however, they are considered in the context of the dimensions of the erythrocyte, they begin to lose their appeal. Even with its glycocalyx (Fig. 2.12), the membrane could occupy no more than about 100 nm of the width attributed to unstirred layers. Hence it can be calculated from the result in the preceding Section that a layer of unstirred water about 0.8 txm thick is

present on each side of the erythrocyte plasma membrane ((1.7 tzm - 100 nm)/2 = 0.8/zm). Since the surface area of the average human erythrocyte is about 140 tzm 2, the volume of unstirred water inside the cell would be about 112 tzm 3 (0.8 txm x 140/xm2). The volume of the average human erythrocyte is, however, only about 90 txm3! Hence, measurements of butanol transport must involve detection of butanol within unstirred water inside cells, which contradicts the assumptions that have been made in determining P for butanol. That is, it makes little sense to envision dependence of the value of P on unstirred water layers that a solute must traverse during exodus from red blood cells when the solute starts out at various positions already within unstirred water inside the cell? It could be assumed that the permeability coefficient and unstirred water layers, such as one may determine for erythrocytes, apply only to the membrane and to an external water layer. In this case we would have to assume that migration of butanol occurs across an unstirred water layer, the volume of which is more than twice the average volume of erythrocytes (i.e., 1.6 txm • 140/xm 2 = 224/xm 3 around each cell). Others may feel that it is still possible to justify these or similar assumptions that are required for acceptance of the method of determining the width of unstirred water layers using butanol or a similar substance. We think, however, that the validity of these assumptions and method need to be reconsidered. In particular, we need to determine whether biomembranes are barriers to the migration of even highly lipid-soluble substances. Moreover, we need to determine more directly the dimensions of possible unstirred water layers next to the membrane. Few if any substances cross the barrier created by a biomembrane as rapidly as is anticipated for simple diffusion of the substance. These findings contradict the conclusion reached by many investigators (Section IV above) that the membrane lipid bilayer should not be a barrier to the migration of highly lipid-soluble substances. Hence, they have used unstirred water layers as a possible explanation for the apparent barrier. Certainly some impairment of stirring occurs in the water next to membrane lipid bilayers. The question is whether the nearly 2/xm thickness estimated above for human erythrocytes really contains mostly unstirred water. To our knowledge, no one has actually measured 3 The determination of P values have, in some cases, also been measured in a way that would not require quantification of the butanol within intracellular unstirred water p e r se (e.g., Garrick et al., 1980). Incorrect assumptions also appear to have been made in these cases, however, since the rates of migration of butanol and other substances in a bulk solution of 33% hemoglobin were used as models for the rates of migration of the substances through intracellular water.

77

Apparent Widths of Unstirred Water Layers

these layers next to a b i o m e m b r a n e directly and compared their dimensions to the dimensions estimated indirectly using transport of a lipophilic substance. If they occupy a volume greater than that of some m a m m a l i a n cells, however, then such m e a s u r e m e n t s should be feasible. In this regard, a recent study has shown that when microelectrodes are used to m e a s u r e ions near an artificial phospholipid bilayer, stirring motions next to the bilayer also must be considered in order to give a theoretically acceptable account of water migration across it (Pohl et aL, 1997). In this context, let us consider data that are already available for the p e r m e a t i o n of m e m b r a n e s by various substances. We noted above that good relationships exist between the log10 of the P values of various substances and the log10 of their Kp values (Figs. 4.5-4.9) but these relationships are consistently good only when P is corrected for the molecular volume or size of the p e r m e a n t (see Figs. 4.10-4.13 for the uncorrected relationships). The extent of this correction is, however, based on the degree to which a relationship exists between the volume of the p e r m e a n t and log10 of PA/ Kp (where A is a constant). The negative slope of this relationship, mv (Stein, 1986, pp. 74-76), is multiplied by molecular volume in order to correct P for size. Hence, the correction of P for molecular volume will almost certainly improve the correlation b e t w e e n logl0 P and log10 Kp. The question is w h e t h e r the correction

..=

-1

="

-2

-,-

O (b (/)

E v

O

O o

1

...J -3

-4

'--

I -2

-3

....

I -1

LOG K

,

,

,

I 0

,

I 1

hexadecane

FIGURE 4.1 1 The permeabilities of dog red cell membranes to various n-alcohols are unrelated to their lipid solubilities. Data are the same as those in Fig. 4.7 except that the permeability coefficients are not corrected for size. As for Figs. 4.6-4.10 and 4.12-4.13, the straight line is for unit slope, not the best fit of the data to a straight line (adapted from Stein, 1986,with permission from Academic Press).

-3

-2

3

5

u

-4

(3

(b

3

4e

O "~-4

I:L

Q..

O O

-5

..J

f05 14

-7

-5

-3

-1

1

LOG K hexa,:L=csr~

FIGURE 4.10 The permeability coefficients of various substances for the human erythrocyte plasma membrane does not exceed about 10-2 cm/sec regardless of their lipid solubilities. Data are the same as those in Fig. 4.6 except that the permeability coefficients (P) are not corrected for size (adapted from Stein, 1986, with permission from Academic Press).

08 | -5

,

LOG K

I. . . . . . . . . -4

I ....... -3 "

-2

I

hexadecane

FIGURE 4.12 Limited relationship between the lipid solubilities of several nonelectrolytes and their permeability coefficients for algal cell membranes. Data are the same as those in Fig. 4.8 except that the permeability coefficients (P) are not corrected for size (adapted from Stein, 1986, with permission from Academic Press).

78

4. Transport Kinetics -2

60 "6" v

40

E0

IO

05

-6

I

-5

I

-4

.

.I

-3

I

-2

LOG K hexadecane

FIGURE4.13 As for biomembranes, the uncorrected permeability coefficients of several substances for artificial phospholipid bilayers are rather poorly correlated with their lipid solubilities.The data are the same as those in Fig. 4.9 except that the permeability coefficients (P) are not corrected for size (adapted from Stein, 1986,with permission from Academic Press).

leads to greater insight into the mechanisms by which various substances permeate membrane lipid bilayers. In this regard, although good relationships exist between the size-corrected permeability coefficients and the partition coefficients of various substances (e.g., Figs. 4.6-4.9), the uncorrected values of P exceed 10-2cm sec -1 only very rarely (e.g., Figs. 4.5 and 4.104.13). Hence, there seems to be an upper limit to the permeability of membranes regardless of the lipid solubility of the migrating substance. To obtain greater lipid solubility investigators have frequently studied substances in a series with an increasing number of methylene groups which also increases their sizes (e.g.; n-alcohols, Figs. 4.7 and 4.11). We contend, however, that size and lipid solubility may not be the only important determinants of membrane permeability. For example, the permeability of the highly lipid-soluble substance, ~-tocopherol, is increased when a-tocopherol is converted to the larger and less lipid-soluble substance tocopherol succinate (Bonina et al., 1996). More importantly, undue emphasis on size and lipid solubility evokes the bias that a mechanism of migration of substances across membrane lipid bilayers is ordinary diffusion. In contrast, we suggest that the way in which the highly ordered lipid bilayer interacts with substances migrating across it may result in an upper limit to the

value of the permeability coefficient of approximately 10 -2 cm sec -~ for most membrane lipid bilayers. As discussed above for one of the most permeable substances studied, n-butanol (e.g., Fig. 4.11), the measured permeability is only about 0.3% of what it would be if the membrane did not constitute or somehow generate a barrier to its simple diffusion. Hence, it was concluded in Section V above that the actual distance over which diffusion occurred was much larger than the width of the membrane, and this additional width was attributed to unstirred water layers. Again, however, there has to our knowledge, been no independent verification of the existence of these unstirred layers of the dimensions determined using butanol, nor has it been shown that the layers are the reason why measured P values are much lower than anticipated for lipophilic substances. In our view, the layers need to be demonstrated in independent experiments to be as thick as they have been calculated to be and to not mix freely with bulk solutions containing the transported substances. Otherwise, additional experiments are needed to determine why lipid-soluble substances have much lower membrane permeabilities than anticipated for simple diffusion. In fact, we think that available data already support the conclusion that lipid bilayers are the principal barriers to migration of lipid-soluble substances across biomembranes. First, the Qa0 values for migration of n-hexanol, n-propanol, and methanol across the human erythrocyte membrane all exceed 2.0 (Brahm, 1983). As discussed above, Q10 values above 2.0 are inconsistent with a mechanism of simple diffusion. Even for simple diffusion limited by unstirred water layers, the Q10 value is not expected to exceed 1.5 (e.g., Verkman et aL, 1996; Yano et aL, 1996). Because the activation energies for migration of all three of these alcohols are equally high, it was concluded that they were transported via the same pathway. Moreover, the relative lipophilicities of the three alcohols (i.e., the ratios of the Kp values for partitioning between olive oil and water) are 750:16:1, so their activation energies do not appear to be related to their solubilities in the hydrophobic region of the lipid bilayer. It is also interesting that the activation energy for hexanol transport across the dog erythrocyte plasma membrane has been calculated to increase from 11 kJ mo1-1 above 20~ to 133 kJ mo1-1 between 15 ~ and 20~ (Brahm, 1983; Garrick et al., 1982). Temperature changes in this range may influence the physical characteristics of membrane phospholipids (e.g., Chapter 2), but such changes probably do not greatly change the properties of water, unstirred or otherwise. Hence, we conclude provisionally that the lower-than-anticipated permeability coefficients that are observed even for highly lipid-soluble substances are due to the barrier

Protein-Mediated Biomembrane Transport is Substrate Saturable

that the lipid bilayer itself presents to the substances. Because butanol and similar substances are unlikely to permeate biomembranes by a process that resembles simple diffusion (i.e., the lipid bilayer is likely to be a significant barrier to their migration), estimates of the width of unstirred water layers with these substances probably do not reflect a real physical phenomenon. That is, although unstirred water layers probably exist, it does not appear that their dimensions can be estimated by observing the transport of butanol or other lipid-soluble substances. Perhaps the viscosity of the membrane lipid bilayer is more pertinent than unstirred water layers to permeation of the bilayer by lipophilic substances. In this regard, it is possible to estimate apparent diffusion coefficients for the lateral migration of multispan proteins in the bilayer and compare them to the coefficients for similarly sized proteins in water (e.g., Edidin, 1996). The pertinent viscosity is, however, for the migration of lipophilic substances across the bilayer. As in the case of diffusion, it also seems inappropriate to attempt to apply our concept of viscosity of a bulk fluid, such as an organic solvent, to the highly ordered structure of the membrane phospholipid bilayer. Surely neither lipid-soluble nor water-soluble solutes experience the same physical forces during migration across ordered phospholipid bilayers as they experience during migration in bulk fluids. In this context, we turn now to a consideration of the mechanism of transport of the relatively large and physiologically important substances, the fatty acids. Long-chain fatty acids are thought by some investigators to traverse lipid bilayers by a process that resembles simple diffusion. While both lipid-based and proteincatalyzed processes have been proposed to mediate fatty acid migration across biomembranes, neither of these processes resembles ordinary diffusion. Vii. PROTEIN-VERSUS LIPID-MEDIATED MECHANISMS OF FATTY ACID MIGRATION ACROSS BIOMEMBRANES Important evidence has now accumulated to support the conclusion that long-chain nonesterified fatty acids are transported across animal (reviewed by Berk, 1996) and bacterial (e.g., Black and DiRusso, 1994) cell membranes by an integral membrane protein. It had been proposed earlier that binding of albumin to membrane receptor proteins could be involved in fatty acid uptake, primarily because most of the nonesterfied fatty acids in blood plasma are bound to albumin. High-affinity albumin receptor proteins have, however, not been detected, whereas such receptor proteins (now known to

79

be transport proteins, see below) have been identified for free fatty acids (reviewed by Fitscher et al., 1996). Albumin also could conceivably cause the kinetics of fatty acid transport to appear to be substrate saturable if albumin-bound and unbound fatty acids in plasma are not in equilibrium (i.e., if the rate of dissociation of fatty acids from albumin limits fatty acid uptake) (Weisiger, 1985). It is, however, widely accepted that this equilibrium prevails under most physiological and experimental conditions, even by proponents of the view that fatty acids enter cells by simple diffusion (e.g., Zakim, 1996). Moreover, several lines of evidence indicate that cell membranes contain transport proteins for free fatty acids (erythrocytes appear to be an exception; Kleinfeld et aL, 1998). First, it has been shown for several types of cells, including hepatocytes, adipocytes, cardiac myocytes, and skeletal muscle myocytes, that fatty acid transport occurs primarily by a substrate-saturable process at physiological concentrations (about 20-200 nM) of unbound fatty acids (e.g., Fig. 4.14). This process also shows trans-stimulation, counter-transport, and cisinhibition (Fig. 4.14), which are anticipated for many types of mediated biomembrane transport (Berk, 1996). (See Sections XI,B and XI,C below for further discussion of these characteristics.) Furthermore, initial rates of uptake are not correlated with intracellular fatty acid metabolism (Stremmel and Berk, 1986), although some investigators still assert on theoretical grounds that fatty acid metabolism limits their transport (e.g., Zakim, 1996). The fatty acid membrane transport protein, referred to in the literature as the plasma membrane fatty acidbinding protein, has now been identified and its cDNA has been cloned (reviewed by Berk, 1996). When expressed in X e n o p u s oocytes and other cells, the protein stimulates fatty acid uptake (Fig. 4.15) as anticipated for a fatty acid transporter. Furthermore, antibodies to the protein specifically inhibit long-chain fatty acid transport but not mediated glucose or medium chain fatty acid transport. The antibody does not, however, influence fatty acid uptake by fibroblasts, which fail to express the protein. Finally, fatty acid transport activity is increased in proteoliposomes also containing the putative fatty acid transport protein. Interestingly, the plasma membrane fatty acid-binding protein is identical to the mitochondrial isoform of aspartate amino transferase (see Berk, 1996 for review and references). Largely for this reason, some investigators think that other proteins are more likely to be the catalysts that transport long-chain fatty acids across biomembranes (Man et al., 1996). Actually, several proteins now appear to catalyze long-chain fatty acid transpOrt (Van Winkle, 1999), and up-regulation of at least three of them ap-

80

3.01

4. Transport Kinetics

A

B m

BSA = 600 A I

~

9 Oleate Oleate + Palmitate + S.E.M

2.0

u

CO

~d

r = 0.995

~ Q. 1.0

M

r = 0.997

0

200

Unbound Oleate (nM)

400

-0.01 0

0.01

0.05

0.1

1/S

FIGURE 4.14 Transport of long-chain free fatty acids across the rat hepatocyte plasma membrane is substrate saturable. In fact, nonsaturable fatty acid transport was such a small proportion of the total transport shown that it apparently did not need to be subtracted in order to characterize the large substrate-saturable component. (A) Relationship between the unbound oleate concentration and its initial velocity (vi) of uptake by isolated hepatocytes in the absence and presence of 55 nM unbound palmitate.(B) Lineweaver-Burk double-reciprocal plots of vi and the oleate concentration reveal an oleate concentration at half-maximum velocity (Km value) of 88 nM, a Vma x value for oleate of 2.6 pmol sec -1 (5 • 10 4 cells) -1, and a Ki value for palmitate of 27 nM (adapted from Sorrentino et al., 1996, with permission from The American Physiological Society).

A 240

== ~"

B Results: Day 3 Injected on Day 0 with:

50-1 tl mAspAT OH20

capped

/

~

:H2%S.pAT capped mRNA

-

~Jz

r= 0.99

160

mRNA -11..20 nl --" /

~r

~-~30

,.L,, Qe...

~

80

~

..''" ..........

10 .~~~'~' |'

0

~" !

'1

1

!

2

Oleate: BSA = 150 taM |

Hours

!

3

"!

' ! "

4

|

!

5

O-

!

1

0

"'%..........:.....

Oleate: BSA = 150 ~tM Ou: 120 nM ,

i

1

i

i

2

,

i

Days

FIGURE 4.15 Increase in oleate uptake by Xenopus oocytes after injection of mRNA encoding a rat plasma membrane fatty acid-binding protein (FABPpm). (A) Uptake of oleate by oocytes 3 days after injection of FABPpm mRNA (solid circles) or the vehicle alone (control, open circles). (B) The increase in oleate uptake is expressed in oocytes 2 days after FABPpm mRNA injection (adapted from Berk, 1996, with permission from Blackwell Science Inc.).

3

'i

~ -i

4

Protein-Mediated Biomembrane Transport is Substrate Saturable

pears to account for the increased saturable oleate transport in adipocytes of both genetically obese and genetically diabetic rats relative to their normal counterparts (Berk et aL, 1997). Since the aspartate amino transferase of the mitochondrial matrix appears also to be a plasma membrane fatty acid transporter, one must wonder whether the protein has a role also in the transport of fatty acids into mitochondria in addition to, or for a different purpose than, fatty acid uptake for oxidation via the carnitine shuttle. It has not to our knowledge been determined whether a portion of the total amount of mitochondrial aspartate amino transferase might reside in the inner mitochondrial membrane. Similarly, it is apparently not yet clear whether the protein also may catalyze the amino transferase reaction while associated with the plasma membrane and whether the position of the protein in the cell is regulated in association with alternate metabolic needs. The sometimes unexpected multiple functions of transport-related proteins have been noted in this and the preceding Chapters, and some of these multiple functions are discussed further in Chapter 6. While fatty acids may also be taken up by cells via a nonsaturable process, such transport appears not to be a significant component of total uptake at physiological concentrations of unbound fatty acids (Fig. 4.14). Even if cells that do not express a plasma membrane fatty acid transport protein take up physiologically significant amounts of fatty acids by another process, however, this multistep process is not simple diffusion. Instead, ionized fatty acid molecules are first incorporated physically into the ordered structure of one leaflet of the membrane lipid bilayer. The hydrophilic carboxyl groups of the fatty acids are oriented near the surface of one side of the membrane and their hydrophobic, hydrocarbon tails extend toward the membrane interior. The fatty acids appear then to flip from one leaflet to the other after which they must somehow leave the bilayer in order to complete their migration across it. While such movement of fatty acids from one leaflet to the other may occur more rapidly in membrane vesicles than appreciated previously (e.g., Kamp et aL, 1995) this so-called flip-flop is by no means equivalent to simple diffusion. Furthermore, fatty acid flip-flop is considerably slower in larger vesicles than in smaller ones, apparently owing to the lower degree of membrane curvature in larger vesicles (Kamp et al., 1995). It has, to our knowledge, not been determined whether the even lower degree of curvature normally present in the plasma membrane of most cells virtually prohibits flipflop of fatty acids at a rate that would have important physiological consequences. In addition, the possible physiological importance of more rapid fatty acid flip-

8 |

flop when the degree of curvature increases, as in endocytosis, apparently has not been investigated. Partly because the nonsaturable diffusion of lipophilic substances, such as fatty acids, seems so feasible, the substrate-saturable kinetics of fatty acid transport (e.g., Fig. 4.14) have gained acceptance relatively slowly. In contrast, substrate-saturable transport of lowmolecular-weight hydrophilic substances via carriers is a well-established phenomenon. Nevertheless, some investigators may still not appreciate that the very rapid migration of small ions and molecules through channels is also substrate saturable (e.g., see below). Because it may apply to the biomembrane transport of virtually all solutes and possibly even the solvent, we turn now to a discussion of the saturation phenomenon and, later, to kinetic formulations for it.

VIIi. PROTEIN-MEDIATED BIOMEMBRANE TRANSPORT IS PROBABLY ALWAYS SUBSTRATE SATURABLE

As discussed in the preceding Section, concentrations of unbound long-chain free fatty acids on the order of about 100 nM are needed to achieve half maximal velocity of their transport apparently via the plasma membrane fatty acid binding protein (Fig. 4.14). In contrast, the substrate concentrations needed to achieve halfmaximal velocity of ion transport via channels is on the order of about 100 mM (e.g., Fig. 4.16). Accordingly, protein-mediated transport appears virtually always to be saturable, although the substrate concentrations needed to achieve half-maximal velocities may vary over a range of at least six orders of magnitude. 4 More importantly, both of the transport processes illustrated in Figs. 4.14 and 4.16 begin to become saturated at nearphysiological concentrations of the substrates. In some other cases, such as arginine transport via system b ~ (Fig. 4.17), the substrate concentration appears sometimes to become saturating well below physiological concentrations (Van Winkle et aL, 1990a). In such cases, however, competing substrates are often present in extracellular and intracellular fluids, and these competing substrates may raise the concentration actually needed to achieve half-maximal velocity into the physiological range. This competition is achieved in the case of arginine transport via system b ~ by the presence of leucine, lysine, and tryptophan, also in physiological solutions 4 In addition, when studying transport processes that are saturated over a particular range of substrate concentrations, investigators frequently find it convenient to deduct a so-called nonsaturable component of transport. We discuss in several places (e.g., at the end of Section IX below) why we think that it is likely that protein-mediated processes also catalyze most apparently nonsaturable transport.

82

4. Transport Kinetics

A I max ~" o. v

7

Na *

6

t"

5 o ~,

4

"~: :D

3

I I

I I

2

Km = 100mM ! I

I 0

I

i

100

I 300

200

400

500

Na * activity (mM)

B

lOO ~max

~

80

i0o E"

pS

e-~-----"~ ----r-

100

K

-I-

El v

9

o

20 0

~6 E

oo

K m = 108 mM

0

C

E

i

~ 2o 0

92

Or)

-i

40

=

I 200

Na +

100

,m E

I 300

I 400

CI- activity (mM)

0

I

I 250

I 500

I

750

I

1000

K + or Na + activity (mM)

FIGURE 4.16 Transport of Na +, CI-, and K+ via channels is substrate saturable although the substrate concentrations at half-maximum velocities (Km values) are on the order of 100 mM. (A) A rat myotube acetylcholine receptor channel shows a Km value for Na + of about 100 mM (from Horn and Patlak, 1980, with permission from National Academy of Sciences (USA)). (B) A mouse glycine receptor channel has a Km value for C1- of about 108 mM (from Bormann et al., 1987, with permission from the Physiological Society). (C) A rabbit sarcoplasmic reticulum K+-preferring channel shows a Km value for K+ of 54 mM and a Km value for Na + of 34 mM (adapted from Coronado et al., 1980, with permission from Rockefeller University Press).

with arginine (Van Winkle et aL, 1988, 1990a,b). Similarly, the substrate concentrations needed to achieve half-maximal velocity via primary active transport processes, such as Na+K+ATPase, are of the order of magnitude of the physiological concentrations of their substrates (e.g., Fig. 4.18). Since Na+K+ATPase transports Na + and K + ions against their total chemical potential gradients, while transport of these ions via channels occurs in the direction of these gradients, it is reasonable that the substrate concentrations needed to halfsaturate the processes are 10- to 100-fold higher for channels than for Na+K+ATPase (Fig. 4.16 vs Fig. 4.18). If the substrate concentration needed to achieve halfmaximal velocity is usually near the physiological concentrations of the substances selected for transport by integral m e m b r a n e proteins, then it may be difficult or impossible to demonstrate that water transport via

channels is substrate saturable. As discussed in the preceding chapter, the physiological concentration of water usually is assumed to approximate that of pure water (i.e., 55.5 M ) , because most water solutions inside and outside living cells are relatively dilute. In order accurately to determine the water concentration at halfmaximal velocity of its transport, however, the water concentration should be varied from about 1/5 of the concentration at half-maximal velocity to about fivefold above this concentration. Such manipulations of the water concentration would, however, be impossible or highly toxic to cells, and even relatively small changes in the water concentration from that which is isotonic appear to cause water channels to open (see above). Nevertheless, as for ion channels, the close interaction between water and channel protein molecules during transport makes it highly likely that water channels

Kinetics of Saturable Transport 1.5

E

"7,

9

to perpetuate the myth that biomembrane transport may in some cases proceed primarily by that simple process. Unlike simple diffusion, virtually all proteinmediated transport processes are substrate saturable as is the case also for enzyme-catalyzed chemical reactions. In m o s t cases, the kinetics of this saturable transport closely resemble the kinetics described for enzymes by Michaelis and Menten (1913) almost a century ago.

!J,

'7" eo~

1.0-

r

tO tO tO

83

IX. KINETICS OF SATURABLE TRANSPORT

0

~ o.s-

A. Kinetic Formulations

0.0

0

I

I

I

I

1

I

I

I

20

40

60

80

100

120

140

160

[Arginine], I~M

FIGURE 4.17 L-Arginine transport via system b~ in fertilized mouse eggs shows a substrate concentration at half-maximumvelocity (Km value) on the order of about 1 ~M in the absence of other amino acids. Most physiological solutions contain other good substrates of system b~ however, such as L-leucine, L-lysine, and L-tryptophan. These other amino acids competitively inhibit arginine transport. Under such conditions, the effective Km value for arginine transport lies roughly in its range of physiological concentrations (Van Winkle et al., unpublished data and see Van Winkle et al., 1990a,b for additional evidence).

would be found to be substrate saturable if the necessary experimental conditions could be achieved. We describe above the relationship between the physiological concentrations of substrates and the substrate concentrations needed to achieve half-maximal transport velocity via channels, uniporters, symporters, antiporters, and primary-active transporters (e.g., Figs. 4.14 and 4.16-4.18). In spite of this relationship, transport via channels is frequently assumed to resemble simple diffusion and, unfortunately, uniport continues to be referred to as facilitated diffusion. While Christensen reminded us more than two decades ago (1975) that facilitated diffusion has more in common with primary active transport than with simple diffusion, the notion that simple diffusion underlies some forms of mediated transport appears difficult for some investigators to abandon. Similarly, some scholars have ignored the view that virtually all mediated transport processes may be built around simple transport pathways resembling channels (e.g., Nikaido and Saier, 1992). Use of the term "diffusion" to help to describe or name any proteinmediated membrane transport process (and probably also any lipid-mediated process) may therefore also help

Mediated transport across biomembranes usually is related to the substrate concentration (raised to the power of the number of like substrate ions or molecules transported together) by a simple rectangular hyperbola (Fig. 4.19). The shape of the curve in Fig. 4.19 implies that the substrate binds transiently to a mediating structure (now known in most instances to be a protein), the supply of which is limited. One may represent such transport process most simply as k~ k2 $1 + M ~- MS--~ M + $2, k -1

(4.17)

where S 1 and S 2 a r e the same substrate molecule on side 1 and side 2 of the membrane, respectively, and M is the mediating structure (i.e., the transport protein). When the concentration of S on side 1 is sufficiently high, M will become almost completely occupied or saturated (i.e., it will be almost entirely MS), and the transport process will approach a maximum velocity, Vmax. AS for other formulations in this chapter (see above), we are concerned with the initial velocities of transport. Hence, we do not need to consider reversal of the step marked k2 in Scheme (4.17). That is, the transported substance, S, is, initially, all on side 1 of the membrane. The initial velocity of appearance of S on side 2 of the membrane is determined before significant reversal of the process can occur due to accumulation of S on side 2. It should be noted that in practice, transport of a substance across a biomembrane may be measured when some of the substance is already on the other side of the membrane. In such cases, the initial velocity of transport is, however, still usually measured because the experimental conditions are manipulated so that transport in the reverse direction is not measured even though it occurs. For example, by including a radiolabeled form of S initially only on the outside of cells, the initial rate of S uptake can be measured despite the simultaneous exodus of unlabeled S. In fact, the values of the kinetic parameters for uptake of S by cells may

84

4. Transport Kinetics

A

B

120 100

:~

a0 60

.....

. . . . . . . . . . . . . . . .

i i'ti I .......... 9 /

.E

//:"

/

...

.....,...."" ...... ....!":"

,~ 4O

,<

<

20 0

80

"6 60

-~

40

".~

120

20 "-J 0

0

20

40

60

80

100

120

140

[Na*], m M

..... //"

~ 4-~

I

I

I

I

0

5

10

15

20

[K+], m M

FIGURE 4.18 Na + and K + transport by Na+K+ATPase shows substrate concentrations at half-maximum velocities (K0.s values) one to two orders of magnitude below the Ko.svalues for their transport via channels (e.g., Fig. 4.16). (A) Na + extrusion by a sheep Na+K+ATPase shows a Kmvalue of about 6.3 mM (filled circles), whereas a considerably higher Na + concentration is needed to achieve half-maximal velocity when the glutamyl residue normally at position 779 of the c~-subunit is replaced with a glutaminyl (open squares) or an alanyl (open diamonds) residue. (B) The K0.5value for K + uptake by a sheep Na+K+ATPase is about 0.78 mM (filled circles), but more K + is needed to achieve half-maximal velocity when an alanyl (open circles) but not a glutaminyl (open squares) residue is substituted for the glutamyl residue normally at position 779 of the ~subunit. (data from Feng and Lingrel, 1995, are replotted here).

c h a n g e d e p e n d i n g u p o n the type of t r a n s p o r t that is b e i n g investigated a n d the q u a n t i t y of u n l a b e l e d S that is a l r e a d y p r e s e n t in the cells (see also discussion of trans-stimulation in Section XI,B below). M o r e d e t a i l e d kinetic f o r m u l a t i o n s and the m e a n i n g of the values of kinetic p a r a m e t e r s for different types of t r a n s p o r t w h e n t h e r e is zero (e.g., zero trans e x p e r i m e n t s ) , very high (e.g., infinite trans e x p e r i m e n t s ) , or an i n t e r m e d i a t e a m o u n t of u n l a b e l e d S a l r e a d y on the o t h e r side of the m e m b r a n e m a y be f o u n d e l s e w h e r e (e.g., Stein, 1986). H e r e we will derive an e x p r e s s i o n for the kinetics of t r a n s p o r t d e p i c t e d in Fig. 4.19 by m a k i n g o n e additional, generally valid assumption. A t e a c h c o n c e n t r a t i o n of S at which the initial velocity of t r a n s p o r t is m e a s u r e d , it is a s s u m e d that the c o n c e n t r a t i o n of MS in S c h e m e (4.17) quickly r e a c h e s a constant, s t e a d y - s t a t e concentration. This steady-state a p p r o x i m a t i o n (Briggs a n d H a l d a n e , 1925) permits us to a s s u m e for e a c h c o n c e n t r a tion of S that

1000

.m e"

L

,.- 500 r

.m r L

...,,. >

0

0.1

0.2

0.3

0.4

0.5

[S], mM

FIGURE 4.19 Relationship between the concentration of substrate and the velocity of its saturable biomembrane transport. The curve forms a rectangular hyperbola described by the Michaelis-Menten equation (Eq. (4.26)). The horizontal line marked "Vmax"is the maximum velocity approached by the curve at relatively high substrate concentrations. At one-half of Vmax,the substrate concentration is by definition the Km value. In the present case, one-half Vmaxoccurs at a substrate concentration of about 50/xM.

12i =

k2 [MS],

(4.18)

w h e r e vi is the initial velocity, k2 is the rate c o n s t a n t for the step m a r k e d k2 in S c h e m e (4.17), a n d [MS] is the c o n c e n t r a t i o n of the MS c o m p l e x (it is again a s s u m e d that c o n c e n t r a t i o n s can be s u b s t i t u t e d for activities). E v e n t h o u g h a derivation of the Eq. for the h y p e r b o l a s h o w n in Fig. 4.19 (i.e., the M i c h a e l i s - M e n t e n e q u a t i o n ) is p r e s e n t e d in n u m e r o u s b i o c h e m i s t r y t e x t b o o k s , it is

Kinetics of Saturable Transport

also shown below because it can help to provide a foundation for understanding the meaning of the kinetic parameters Km, Vmax, and Ki. At steady state, the rate of formation of MS is equal to the rate of its breakdown. Moreover, these rates take the familiar chemical kinetic forms (4.19)

vf = k l ( [ M T ] -- [ M S ] ) [ S ]

and

85

logical concentrations of substrates, is the Km value (Fig. 4.19). That is, when Km = [S] Eq. (4.26) becomes Vi -- Vma x or Vi ~-~

Vd ~-~

k-1 [MS] + k2 [MS],

(4.20)

when ve and Vd are, respectively, the rates of formation and degradation of the MS complex; kl, kq and k2 are the rate constants for the steps shown in Scheme (4.17); and [MT], [MS], and [S] are the concentrations of the total M present (both substrate bound and unbound), the MS complex, and S, respectively. Since, at steady state

[S]/2[S]

Vmax/2.

For the more general case where more than one of the same ion or molecule may be transported together (e.g., Fig. 4.18), Eq. (4.26) can be modified to read Vi = Vma x [s]n/([S] n .-+- go.5 n)

(4.27)

where n is the number of identical ions or molecules transported together and K0.5 is the substrate concentration at half maximum velocity (Km when n = 1).

Pf = Vd

we may write kl

( [ M T ] - [MS])[S]

= k_ 1

[MS] +

k2

[MS]

(4.21)

Equation (4.21) can be rearranged in several algebraic steps to read

[MS] = [MT][S]/([S] +

( / 2 nu k - 1 ) / k l )

(4.22)

Now the relationships among the three kinetic constants in Eq. (4.22) is defined as the Michaelis-Menten constant, Km K m -- ( k 2 q-- k _ l ) / k I

(4.23)

and Eqs (4.18), (4.22), and (4.23) can be combined to read Vi -- k 2 [ M T ] [ S ] / ( [ S ]

+ Km).

(4.24)

Finally, since the maximum velocity of transport (Vmax) will occur when M is fully saturated (i.e., when [MS] = [MT] Vmax = k2 [ M T ] ,

(4.25)

Eq. (4.24) can be simplified to the familiar form of the Michaelis-Menten equation Vi--

gmax [ S ] / ( [ S ]

+ Kin)

(4.26).

Since the Vmaxvalue for a given process depends only on the total concentration of the catalyst, [MT], vi in Eq. (4.26) approaches the Vmax value as the substrate concentration, IS], is increased above the Km value. Moreover, it can be seen from Eq. (4.26) that the substrate concentration at half-maximal velocity, and the rate discussed in Section VIII above in regard to physio-

B. Fitting of Experimental Data to Kinetic Formulations and Statistical Comparisons of the Values of Kinetic Parameters D e t e r m i n e d from Such Fittings The best fit of transport data to a hyperbola defined by Eq. (4.26) can be determined with nonlinear regression analysis (Atkins and Nimmo, 1980; Gardner and Atkins, 1982; Ritchie and Prvan, 1996). Computer software for performing nonlinear regression analysis is now available commercially from several companies (e.g., Sigma Plot, Jandel Scientific Corp, San Rafael, CA 94901, USA), and the software also can be used to calculate kinetic parameters and to estimate the level of uncertainty in their values. The reader is cautioned, however, that the statistical uncertainties in Km and Vmax values that are determined using nonlinear regression analysis should not be used to compare the values to other such values in statistical tests. Rather, statistically significant differences among the values of Km and Vmax may need to be examined in the familiar way, by obtaining three or more replicate values for the parameters in each of the tissues or conditions under consideration. The mean values and their variances for each tissue or condition may then be calculated and the values compared with appropriate parametric or nonparametric statistical tests. Alternatively, transport experiments that were not designed explicitly to measure Km and Vmaxvalues may nevertheless support the theory that these values are different for different tissues or for the same tissue under different conditions. Such theories may then be tested by obtaining replicate values, as described above, or they can be tested by collecting single sets of kinetic

86

4. Transport Kinetics

data or by combining data into single sets for each tissue or condition. In the latter cases, the method of Eisenthal and Cornish-Bowden (1974), as later modified by these authors (Cornish-Bowden and Eisenthal, 1978), and by Porter and Trager (1977) may be used to compare Km or Vmaxvalues statistically. Briefly, this method involves determining values of Km and Vmaxbased on each pair of data points such as those depicted in Fig. 4.19 and then comparing ranges of values so determined for different tissues or conditions. From such assessments, medians and confidence intervals rather than means and standard errors are obtained. One may then use the finding that the confidence intervals for the values obtained for different tissues or under different conditions either do or do not overlap to conclude that the values are or are not statistically distinguishable from one another at a probability greater than the probability that the intervals contain the true values of the kinetic parameters (e.g., Table 4.1). One may also gain a visual appreciation of whether significant differences may exist between or among the values of kinetic parameters by examining linear transformations of the pertinent data. To produce such graphs, Eq. (4.26) Vi :

Vma x [S]/([S]-Jr- g m )

(4.26)

can be converted algebraically to several forms that represent straight lines for different values of [S] and vi. These three forms of Eq. (4.26) are 1/vi = Km/(Vmax[S]) + 1/Vmax, [S]/v i = [S]/Vma x -Jr- gm/Vma x

(4.28) (4.29)

and

TABLE 4.1 Kinetic Parameters for L-Lysine Transport by M o u s e Blastocysts in Different Isotonic Solutions a

Principal solute(s) in the isotonic solution in which uptake was measured

NaC1 LiC1 Sucrose

Median value of the parameter (92-94% confidence interval) b

Km

Vmax

61(48-90) 59(43-72) 5(1-8)

68(58-75) 42(39-51) 44(32-54)

aA modification (Cornish-Bowden and Eisenthal, 1978; Porter and Trager, 1977) of the nonparametric statistical method of Eisenthal and Cornish-Bowden (1974) was used to estimate the median values of kinetic parameters and their 92-94% confidence intervals (Van Winkle et al., 1990c). When the 92-94% confidence intervals do not overlap, it is more than 95% likely that neighboring values are different because even the 90% confidence intervals overlap somewhat when p = 0.05 in t tests (data from Van Winkle et al., 1990c with permission from Elsevier Science). bKm in/xM and Vmaxin fmol blastocyst -1 min -1.

lei/[S ]

-

-

Vmax/gm-

12i/g m

(4.30)

or

vi = Vmax -- Vi Km/[S].

(4.31)

When data are presented in graphs defined by Eqs. (4.28), (4.29), and (4.30), they are termed, LineweaverBurk, Hanes and Eadie-Hofstee (or simply Hofstee) plots, respectively (Fig. 4.20A-4.20C). The Hofstee plots shown in Fig. 4.21B correspond to Eq. 4.31. One can gain from these linear plots an immediate sense of the magnitude of the differences that may exist among values of Km and Vmax. For example, the differences in the values of the kinetic parameters reported in Table 4.1 are immediately obvious from the linear plots of the data in Fig. 4.21B. The magnitude of these differences may, however, not be as obvious from the nonlinear plots of the data shown in Fig. 4.21A. In Fig. 4.21B, the values of the y-intercepts correspond to Vmax values, whereas the negative values of the slopes correspond to Km values. In contrast, the Vmax values (i.e., the vi values at infinitely high substrate concentrations) are more difficult to estimate from the data as presented in Fig. 4.21A. Similarly, it is difficult to estimate the Km values visually (i.e., the substrate concentrations at onehalf the Vmax values) whenever the Vmax values are, themselves, difficult to judge accurately. While most readers are now familiar with the definitions of Km and Vmax, the relationship of their values to each other and to other kinetic parameters may not be as clear. Moreover, it is useful to consider why transport proteins have circumscribed values for these parameters and how the values may differ for transport in opposite directions across the membrane. For these reasons, we turn now to a more detailed consideration of the meaning of Km and Vmax. C. Meanings of the Kinetic Parameters K m a n d Vmax

1. Finite Values of Km and VmaxAre a Consequence of the Close Biophysical Interactions between Small Ions and Molecules and the Protein Molecules That Catalyze Their Biomembrane Transport

Because biomembranes appear to be barriers to the migration of even highly lipophilic substances, it is possible for the rates of migration of solutes and water across membranes to be increased by catalysts. Even the best catalysts cannot, however, increase the rate of biomembrane transport to near that which would occur if simple diffusion across the membrane were possible. For example, Stein (1986, p. 202) has calculated that gramicidin channels operate at no more than about 8 percent of the rate of migration of K + ions that occur by simple diffusion, and even the fastest K + channel protein mole-

87

Kinetics of Saturable Transport

A

B

C

L

13,/[s] [S]/ar Slope

= KJVM~

Slope

=

9

/

IlVM,~

1

~

SLOPE = - I / K M

VM~ /Ki

l l VMAx - -

-~/K. I

i

~/iS]

_KM

_ _

[S]

Vv~x

~

'

FIGURE 4.20 Lineweaver-Burk (A), Hanes (B), and Eadie-Hofstee (or Hofstee)(C) plots of transport data such as those shown in Fig. 4.19. The plots represent different linear transformations of the Michaelis-Menten equation (Eq. (4.26)) and correspond to Eqs. (4.28) (A), (4.29) (B), and (4.30) (C), respectively (adapted from Stein, 1986, with permission from Academic Press).

cules operate no faster than the channels formed by the gramicidin peptide. Since membranes appear to be barriers to diffusion even when they contain rapid channels, one can expect transport always to be saturable via the pathways through membranes created by integral membrane transport protein molecules. As discussed above, the Km values vary widely among protein cata-

lyzed transport processes. Vmax values also vary widely since their values frequently correlate at least roughly with Km values. These kinetic data also give us insight into how transport protein molecules may function. Since proteins catalyze transport, but the resultant rate never approaches that of free diffusion, even for the fastest transport via

7~

B 60

60

9

I

~

"T,r .n

/////Jl'-''-

I- 5 0 .........=

/

f / f /

E

9

/-

~.~

go

..

9

40 NaCI

m

3 o i / L -/

9

o

sucrose

~ 20

NaCI ........... Sucrose

buffered

LiCI

Phosphate-buffered

LiCI

0 0

i 200

J

400

i

600

[Lysine], I~M

I

800

1000

0

2

4

[L-lysine], ~tM

FIGURE 4.21 Nonlinear (A) and linear (B) plots of L-lysine uptake as a function of lysine concentration by mouse blastocysts under different isotonic conditions (the principal solutes are shown for each line). The lines shown in A represent the best fit for the data points obtained under each condition to a hyperbola, whereas the lines in B represent the best straight lines for the data points after linear transformation (Eq. (4.31)) of the Michaelis-Menten equation (Eq. (4.26)). (B adapted from Van Winkle et al., 1990c, with permission from Elsevier Science).

88

4. Transport Kinetics

channels, transport proteins must interact intimately with their substrates. These intimate interactions may include ionic and hydrophobic interactions and hydrogen bonding depending on the transport process and the chemical structure of its substrate. Such interactions slow the rate at which the substrate moves by free diffusion, even for the short width of biomembranes which otherwise are virtually impermeable to the migration of hydrophilic solutes. An understanding of the details of these close biophysical interactions is emerging for a number of transport proteins (see Chapters 5 to 8 for various examples). We are also only beginning to appreciate the significance of asymmetric insertion of transport protein molecules into biomembranes. Although this asymmetric structural orientation of transport proteins in biomembranes has been known for several years, it is still frequently assumed without much testing that many proteins function symmetrically in regard to their Km and Vmax values. For example, channels are frequently assumed to operate symmetrically in their open states, but their Km and Vmax values for uptake vs exodus have, to our knowledge, not usually been determined and compared. We anticipate that channels would operate symmetrically in regard to their Km and Vmax values about as frequently as do uniporters and antiporters. In this regard, nonaccumulating glucose transport has been shown to be directionally symmetric (i.e., Km ex~ -- Km uptake, Vmaxex~ = Vmaxuptake) in rabbit erythrocytes (Regen and Morgan, 1964) and rat hepatocytes (Craik and Elliot, 1979), but asymmetric in human erythrocytes (Wilbrandt, 1955; Bloch, 1974) and rat thymocytes (Whitesell et al., 1977). In our view, symmetric operation of transport proteins should be viewed as coincidental or possibly due to evolutionary selection for such function, since the precise mechanisms of transport are unlikely to be symmetric for asymmetrically oriented protein molecules in biomembranes. Obligatory exchange or antiport might, at first, seem to be an exception to this conclusion. A moments reflection should, however, lead us to drop that exception. Even for exchange, the values of the kinetic parameters need not be identical for uptake and exodus. Such has in fact been found to be the case for nonobligatory glucose exchange (Bloch, 1974) and obligatory anion exchange (Chapter 6), although the exchange itself is, by definition, symmetric. Ironically, many enzymes for physiologically reversible reactions may operate in a more symmetric manner than do transport proteins, even though enzymes catalyze chemical changes, whereas transporters do not destabilize their substrates. For example, amino transferase reactions are catalyzed by proteins with more or

less invariant sites for binding of amino acids and 2-oxo carboxylic acids (c~-keto carboxylic acids) regardless of whether they catalyze the forward or the reverse of reactions of this type

~C

_

R1

+NH 3

I

COO- + H m C m C O O -

I

R2 +NH3

- H m CI ~ C O O - +

I

R1

(4.32) O

'

I

c-coo

R2

In the above reaction, the enzyme molecule is structurally about the same whether it receives the substrates on the left or the substrates on the right. In contrast, a transport protein molecule presents a different orientation to identical substrate molecules on opposite sides of a biomembrane. For this reason, the effectiveness of transport proteins as catalysts, and hence their Km and Vma x values, could vary considerably for migration of substrates in one direction across biomembranes vs the other direction. Asymmetric transport should not be confused with an asymmetric biochemical reaction. A biochemical reaction may be asymmetric because the structures of the reactants and products are quite different, not because the enzyme catalyst is different for the forward and reverse reactions. Moreover, although the enzyme may receive substrates and products for the forward and reverse reactions differently, it treats identical substrate molecules in the same way. In contrast, identical substrate molecules on opposite sides of a biomembrane are received differently by the same transport protein molecule. The possibility that virtually all transport proteins exhibit structural and hence functional asymmetry for identical substrate molecules on opposite sides of biomembranes may have heretofore unappreciated thermodynamic as well as kinetic implications.

2. Can Transport Proteins Produce Solute Gradients Owing to Their Asymmetric Structures and Functions and the Free Energy Needed to Produce and Maintain Such Asymmetries? It has been concluded elsewhere that transport that is not coupled to a conspicuous source of free energy in ATP or a solute gradient may nevertheless have different values of the kinetic parameters for uptake and exodus across the plasma membrane without breaking the laws of thermodynamics as long as the ratios of the Km to Vmax values for uptake and exodus

Kinetics of Saturable Transport

are equal. Just such equal values for the ratios of these kinetic parameters have in fact been found in many studies. It is also conceivable, however, that the ratios of Km to Vmax as well as their absolute values need not be identical for uptake and exodus. Such transport should produce total chemical potential gradients, which for substrates with no net charge can be expressed entirely as gradients of activity or concentration. Although the sources of free energy needed to drive such transport processes remain to be identified, the processes could conceivably be driven simply by the free energy needed to construct and maintain asymmetric biomembranes. 5 In addition to their asymmetric orientations in biomembranes, most transport protein molecules are believed to undergo conformational changes in order to be able to receive substrates for transport on one side of the membrane or the other. While the differences in the free energies of the different conformations of transport proteins have not been measured in most instances, these differences in free energy would not need to be large to produce modest solute gradients. For example, a &G value for the different conformations of 1.8 kJ mo1-1 would be needed to maintain a twofold greater steady state concentration of an uncharged solute on one side of a biomembrane than on the other (see Eq. (3.34) and several similar equations in Chapter 3). Moreover, different conformations of allosteric proteins differ in their free energies by an average of about 22 kJ mo1-1 (Goldsmith, 1996). Hence, the free energies of different conformations of a transport protein would need to differ by less than 10% of the average such difference for allosteric proteins in order to produce a twofold difference in solute concentration across a biomembrane. In this regard, the affinity constants of various inhibitors (measured directly) or substrates (measured indirectly) for binding to the two conformations of the anion exchange protein, band 3 (Knauf et aL, 1992), can be used to calculate a free energy difference (Goldsmith, 1996) of about 3.4 to 6.0 kJ mol -~ for the two conformations, depending on the substance bound. It remains to be determined, however, whether band 3 or other transport proteins are themselves inherently able to produce gradients in the total chemical potentials of solutes across biomembranes (see also Chapter 6). 5 The proposed need for continuous free energy input to maintain constituents of biomembranes in asymmetric orientations warrants emphasis here. If free energy were not expended to maintain membranes and their constituents they would soon stop performing their functions. Maintenance of transport proteins in asymmetric conformations may be viewed as gradients that could conceivably be converted into solute gradients. Both gradients would be maintained only through a more or less continuous free energy expenditure by cells. Asymmetrically oriented transport proteins and their maintenance are, therefore, different from Maxwell's demon or a perpetual motion machine of the second kind.

89

It is, nevertheless, possible to imagine how the free energies of the different conformations of uniporters, symporters, and antiporters might be utilized to produce total chemical potential gradients of their substrates across biomembranes. These possible mechanisms for the production of gradients by the transport protein per se are described here only to demonstrate the conceptual feasibility of such processes. In the case of a uniporter, one conformation of the transport protein might be favored when substrate is bound, while the other is favored in the unbound state. A similar mechanism could produce gradients of two or more solutes via symport, while antiporters could produce gradients through obligatory exchange of two different solutes each of which favors a different conformation of the transport protein molecule. On a macroscopic level, structures with functions analogous to those proposed for asymmetrically organized transport proteins are easy to imagine. Cages or traps with doors that open in only one direction are an obvious example of how a barrier with an asymmetrically constructed entrance can provide greater concentrations of animals on one side of the barrier than on the other. In this case, the asymmetric door influences the thermodynamics as well as the kinetics of migration in one direction relative to the other. The asymmetric door also may be viewed as having different conformations when viewed from different sides of the barrier. The asymmetric distribution of these conformations across the barrier forms a gradient that can be propagated into a gradient of animals. The overall process is exergonic, owing to the conversion of the free energy expanded to construct the device into the free energy of the device itself. A continued input of free energy would also be required to repair the device and thus prevent the intended function of the device from deteriorating. In our view, current dogma sometimes equating migration of solutes through channels or uniporters to diffusion has militated against the design of studies to determine whether some of these proteins can serve to produce total chemical potential gradients of solutes or even the solvent. Similarly, the covert assumption most of us make in assessing the thermodynamics of transport entirely as a function of solute total chemical potential gradients (e.g., as we did in Chapter 3) is that transport protein molecules operate in a thermodynamically symmetric manner in the absence of coupling to an obvious source of free energy in ATP or in a previously formed solute gradient. Now that we know that the structures of most if not all transport protein molecules are asymmetric in biomembranes, however, we need to consider the possible thermody-

90

4. Transport Kinetics

namic as well as kinetic consequences of their asymmetric operation. 6 In this regard, we must also consider the converse. That is, some of the membrane-spanning helices of transport protein molecules may migrate within or even into and out of the lipid bilayer. The extreme (and we think unlikely) consequence of such migrations would be the complete reversal of the orientation of components of the protein molecule within the membrane. Transport protein molecules might lose asymmetry to the extent that they reverse their orientations in biomembranes. Investigation of these possibilities will, of course, require further study of the relationships between the structures of transport proteins and the values of their kinetic parameters for transport of substrates in both directions across biomembranes. It should also be useful to understand the relationship of the values of these kinetic parameters to the affinity (or dissociation) constants, since the latter values may be used to determine the differences in free energy of different conformations of transport protein molecules. Toward these ends, the structures and functions of several transport related proteins are considered in greater detail in Chapters 5 to 7.

3. Relationship of Km to the Substrate Dissociation Constant (Kd) and to Vmax Several additional interrelated concepts concerning the meanings of the Km and Vmax values also warrant consideration. Most of these concepts are intended to apply best to the simplest type of transport via channels or uniporters, although the concepts also may apply reasonably well to aspects of more complex types of transport. First, it is frequently and incorrectly asserted, even in the current literature, that the gm value is virtually equivalent to the dissociation constant (Kd) or to the inverse of the affinity of a substrate for its transporter. Such is the case, however, only in relatively rare instances where an equilibrium as well as a steady state assumption can be made in regard to Scheme (4.17). Since k2 usually contributes significantly to the value of Km in the simple representation of transport in Scheme (4.17), the Km value (Eq. 4.23) will also usually exceed the Kd value (Kd = k-Jka) when this Scheme applies. 6 While our discussion here has been primarily a theoretical one, we present evidence in Chapter 7 (Section II,C) that inwardly rectifying K + channels may indeed catalyze the migration of K + ions against their total chemical potential gradient. Such studies have, however, not been performed intentionally to test the present hypothesis that transport proteins may catalyze such migration owing to their asymmetric orientations in structurally asymmetric membranes. Experiments designed with the intent of determining whether channels and uniporters may sometime transport solutes against their total chemical potential gradients are, therefore, needed to test our hypothesis.

Historically, the fact that the Km value usually differs from the Kd value has been important in kinetic assessment of whether a transport process operates symmetrically or asymmetrically. In particular, the mobile carried model implied that the transport mediator might present the same face to substrates on either side of the membrane in some instances of passive transport. Hence, the value of Kd was expected to be the same at the inside and outside surfaces of the membrane, even when the transport process operated asymmetrically. In contrast, the Km values for uptake and exodus were expected to differ for asymmetric transport. As discussed in Section 2 above, it has until now been expected for asymmetric transport that the ratios of the Km to Vmax values would be equal for uptake and exodus via transport processes that are not coupled to an immediate source of free energy. Although it may now be anticipated that the values of these ratios need not be identical, we still expect a correlation between Km and Vmax values because the Km value is at least partially dependent on the values of rate constants that also help to determine the Vmax value. The Km and Vmax values should thus be viewed as measuring some of the same aspects of transport. For example, in the simplest formulations for transport under the steady state approximation, both the Vmax value (Eq. (4.25)) and the Km value (Eq. (4.23)) depend directly on the value of k2. It is therefore not surprising that the Km and Vmax values are correlated; transport processes with higher Km values also frequently have higher Vmaxvalues than processes with lower Km values. Nevertheless, the ratios of Km t o Vmax need not be precisely the same for exodus and uptake, owing to the asymmetric conformations of transport proteins in biomembranes. Now, as discussed above, in the simplest formulation for transport (Scheme (4.17)) Km~Kd because (4.23)

Km = (k-1 -k- k2)/kl, whereas

Kd = k-1/kl. For multistep transport models, however, the Km value may be either larger or smaller than Ko. For processes in which only one more step than in Scheme (4.17) is inserted kl ~ k3 S~ + M ~ (MS)I (MS2) ~ M + k -1 -2 the Km value can be shown to be

S2

(4.33)

Kinetics of Saturable Transport

Km = (k_flk~)(k3 + k-2 + k2(k3/k-1))/ (k3 + k-2 + k2),

(4.34)

where lowercase k's are the rate constants shown in Scheme (4.33). It can be seen from Eq. (4.34) that whether the Km value is larger or smaller than Ka depends on the ratio of the rate constants for the exit of S from the transporter on each side of the membrane. A procedure for deriving the expression for the Km value in Scheme (4.33) and in more complex cases is given in Appendix A of Stein (1986). Scheme (4.33) may apply best to channels, whereas additional considerations involving the reorientation of the protein to its original position without its substrate have historically been applied to carriers such as uniporters under zero trans conditions. In this regard, it should be noted that only modest success has been achieved in determining whether actual transport processes fit kinetically more complex models of transport, such as the somewhat more complex one shown in Scheme (4.33), or whether they fit the simple model in Scheme (4.17). Modern molecular studies indicate that many types of transport involve multiple steps (Chapters 5 to 7) which display only simple carrier kinetics (Hern~indez, 1998). Apparently these multiple steps either are not detected well by current kinetic procedures or they are not modeled well by known kinetic formulations (but see also Chapter 11). Finally, for the kinetically more complex process of symport, the relative values of Km and Vmax for one substrate at various concentrations of the other can, in theory, be used to determine the order of substrate binding. For example, when one substrate molecule must bind first, changes in its concentration are not expected to influence the Vmax value of the other substrate (Stein, 1986). While one may be inclined to question a kinetic model that makes such a counterintuitive prediction, several instances where the concentration of one substrate does not influence the Vmax value of the other substrate have been reported in the literature (summarized in Stein, 1986). The latter data could, of course, also result from undetected uncoupling or slippage of the cotransport process (i.e., its reversion to uniport of a single solute). Moreover, it has been concluded in more recent studies that one substrate molecule binds before the other even when a decrease in the concentration of the first substrate is observed to decrease the Vmax value of the other (e.g.; Boorer et al., 1996; Mackenzie et al., 1996a). For such reasons it is somewhat difficult to judge whether any currently emerging kinetic model of cotransport accurately reflects accumulating knowledge of the molecular structures and functions of symporters (see Chapter 6 for further discussion of the difficulties associated with studying the effects

91

of cosubstrates on each other's transport). Current procedures for studying the details of the kinetics of transport and the theoretical considerations needed to interpret the results of such experiments are discussed more thoroughly in Chapter 11. D. The Parameter, K~ in Inhibition Analysis 1. Determination of K~ Values The Ki value of a competitive inhibitor of a transport process frequently is equal to the Km value for transport of the inhibitor by the same process. For this reason, we shall see that determination of Ki values can be particularly useful in testing whether two or more solutes share a transport process (Section 3 below). Historically, competitive inhibitors are viewed as raising the Km value of a substrate without altering its Vmax value, whereas noncompetitive inhibitors decrease the Vmaxvalue of a substrate without changing its Km value. In Hofstee plots (Eq. (4.31)) competitive inhibition is reflected by an increase in the magnitude of the slope of the line defined by the data points (recall that slope = -Km), whereas noncompetitive inhibition results in a decrease in the y-intercept (Vmax value) (Fig. 4.22A). In practice, more complex types of inhibition are also observed in which both the Vmaxand Km values are affected (e.g., arginine inhibition of taurine transport shown in Fig. 4.22B). These other types of inhibition have been discussed extensively by other authors (e.g., Dixon and Webb, 1964). In cases of competitive inhibition, such as fl-alanine inhibition of taurine transport (Fig. 4.22B), the Ki value can be calculated from the equation. Ki = [I]/((Kmapp/Km)- 1),

(4.35)

where [I] is the inhibitor concentration, Km has its usual meaning, and gmapp is the apparent value of gm for substrate transport in the presence of the inhibitor. A more direct graphical method with which to determine the Ki value and whether inhibition is competitive or noncompetitive is to determine vi at multiple concentrations of both the inhibitor (I) and the substrate (S). When the resultant data are assessed in a Dixon plot (i.e., a plot of 1/vi vs the inhibitor concentration at several concentrations of the substrate) the resultant series of straight lines for different concentrations of the substrate intersect at a point where the inhibitor concentration on the x-axis is equal to the negative of the Ki value. Moreover, such lines intersect above the x-axis for competitive inhibition (Fig. 4.23A), whereas they intersect on the x-axis for noncompetitive inhibition (Fig. 4.23B). These results can be explained by considering that

92

4. Transport Kinetics

B

A

150 0

,_\,

I .o,n.,b,.o.

- \~ Competitive~ '\ In~bition ~

xr/

125

100

~

5O

9

?n

hibitor

./

Noncompetitive~

"~

~:

r \ I

\

%

25

%

\ \

I

%

9

I

lOmM . \

0

~.

, 0

2

a 4

I 6

/[Taurine], nl-conceptus -~. h -~

FIGURE 4.22 Competitive and noncompetitive inhibitors of transport produce decreases in the slope (more negative) and y-intercept, respectively, of Hofstee plots. (A) Changes in the Km (negative of the slope) and Vmax (y-intercept) values of transport owing to competitive and noncompetitive inhibition, respectively, as determined using Hofstee plots (Eq. (4.31)). (B) Inhibition of taurine transport by/3-alanine is competitive, whereas inhibition by L-arginine appears to be mixed (i.e., both competitive and noncompetitive) (B adapted from Van Winkle et al., 1994, with permission from Elsevier Science).

w h e n t h e s u b s t r a t e c o m p e t e s with t h e i n h i b i t o r , t h e inh i b i t o r s h o u l d h a v e no effect at infinite s u b s t r a t e conc e n t r a t i o n . H e n c e , t h e line at infinite s u b s t r a t e c o n c e n t r a t i o n m u s t be p a r a l l e l to y e t a b o v e t h e x-axis if t h e

A

s u b s t r a t e is to h a v e a finite Vmax v a l u e (Fig. 4.23A). In c o n t r a s t , n o n c o m p e t i t i v e i n h i b i t o r s a r e e x p e c t e d to slow t r a n s p o r t , e v e n at infinite s u b s t r a t e c o n c e n t r a t i o n (Fig. 4.23B).

B

=.

;i"

J,,

[i]

_

[I]

FIGURE 4.23 Dixon plots for competitive (A) and noncompetitive (B) inhibition of biomembrane transport. Competitive inhibitors become increasingly less effective as the substrate concentration is increased (A). In contrast, noncompetitive inhibitors are equally effective regardless of substrate concentration (B). The negative of the value of x at the point at which the lines intersect is equal to the Ki value.

Kinetics of Saturable Transport

2. The K~ Values for Competitive Inhibition of Transport Frequently Have a Different Meaning Than for Inhibition of Enzyme Catalysis Competitive enzyme inhibitors bind reversibly to enzymes, and they compete with the substrate for such binding. Competitive inhibitors of enzymes are, however, frequently not also substrates of the enzyme. In such cases, the Ki values for these inhibitors are equal to the values of their dissociation constants (Ka values; Fig. 4.24). In contrast, competitive inhibitors of catalysis by transport proteins usually vie with the substrate both for binding and for transport. For this reason, the Ki value of a substance for competitive inhibition of transport is usually identical to its Km value for transport (Fig. 4.25). For example, the Km values for transport of Lalanine, L-lysine, and the bicyclic amino acid analog 3-aminoendobicyclo [3,2,1 ] oct ane- 3- carb oxylic acid (BCO) are (within experimental variability) identical to their Ki values for inhibition of transport of each other via amino acid transport system B ~ (see Section 3 below). In some instances, however, competitive inhibitors may not be transported. For example, D-tryptophan competitively inhibits L-tryptophan transport by amino acid transport system T (L6pez-Burillo et al., 1985; Van Winkle et al., 1990b), but o-tryptophan does not appear to be a system T substrate. Similarly, cationic amino acids are competitive inhibitors of amino acid transport system ASC, but they are not transported by it (Thomas and Christensen, 1970). Moreover, maltose and phlorizin are reversible competitive inhibitors of Na+independent glucose uptake by G L U T transport proteins, but these substances are not transported by the proteins (Baldwin, 1993). In these cases, inhibition corresponds to the simpler Scheme where Ki = Kd (Fig. 4.24) rather than where Ki = Km (Fig. 4.25). Because it is possible, albeit infrequent, that competitive inhibitors of transport may not themselves be transported,

kI E +S

=

k. 1

k2 >

ES

> E +P

4-

I

k -1

1

K.= n

p

k'

-1

1

-K'

d

El

FIGURE 4.24 Competitive inhibitors of enzyme action frequently do not serve as enzyme substrates. In such c a s e s Ki values are equal to their dissociation constants (Kd values).

93 kI

M+S 1

,i

+

k2 MS

M+S 2

k. 1

11

MI

k'.l + k' 2 k"2

K. ~

n

~

~

,

kl

K

m (I)

M

FIGURE4.25 Competitiveinhibitors of biomembrane transport are also usually substrates for transport. Hence their Ki values are usually equal to their Km values for transport.

considerable evidence must be obtained to support the conclusion that a competitive inhibitor is transported by the same transport process that it inhibits. In part because Ki = Km when the inhibitor is also a substrate, measurement of Ki values is of considerable importance in the inductive process, known as A B C testing (see below), in which an investigator attempts to determine whether two or more solutes share a transport process. 7

3. Importance of Quantitative Inhibition Analysis in Determining Whether Two or More Solutes Compete for the Same Transport Process When two substances mutually inhibit each other's transport, it is occasionally and erroneously concluded that they share a transport process. Such conclusions may arise in part from the converse and correct conclusion that if two solutes do not inhibit each other's transport, then they do not share a transport process. When inhibition does occur, however, it may be noncompetitive, and even mutually competitive inhibitors are not necessarily substrates for the same transport process (see above). In order to demonstrate competition of 7A different procedure that does not rely on the values of these kinetic parameters may be used to characterize other types of transporters. For example, in the cases of channels, substrate-saturable transport may be more difficult to study or a given transporter may have only one known substrate. In many such cases, it has been possible to identify high-affinity inhibitors that act more or less selectively on particular transport proteins or sets of proteins (see summaries and consideration of such inhibitors of channels in Hille, 1992).

94

4. Transport Kinetics

two solutes for the same transport process, evidence must be gathered to show that each solute behaves as an inhibitor in the same way that it behaves as a substrate. 8 W h e n mutual inhibition of transport is observed between two or more solutes, it is first necessary to determine whether the inhibition is competitive or noncompetitive. In the case of Na+-dependent transport of L-alanine, L-lysine, and B C O by mouse blastocysts (system B ~ in Table 4.2), these determinations were m a d e using Dixon plots (e.g., Fig. 4.26). Since these amino acids inhibited each other's transport competitively, the Ki values d e t e r m i n e d from the Dixon plots could be used further to test the hypothesis that these three substances share the same transport process. In Section 2 above it was learned that when an inhibitor is also a substrate of a transport process, its Km value for transport is equal to its Ki value for inhibition of transport of another substrate. For the present example, since the Km value for L-alanine transport is the same as its Ki value for inhibition of B C O transport, and since the Km value for B C O transport is equal to its Ki value for inhibition of L-alanine transport (Table 4.3), these two substances appear to share the same transport process (AB portion of the A B C test). If either equivalency had not been observed, then it could be concluded that different unshared processes (or possibly multiple shared processes) are responsible for transport of each of the two substances (see Section X below for further discussion of such heterogeneity). Even if the values of Km and Ki for a given solute are the same, however, they may be equal coincidentally rather than because the solute interacts with the same process as an inhibitor and as a substrate. For this reason, it is frequently desirable to seek additional evidence that two solutes share a transport process. Such evidence can be gained by studying quantitatively the effects of additional, potential inhibitors of transport of the substrates thought to share a transport process. If the additional solutes are found to have the same Ki values for the competitive inhibition of transport of each putative substrate, then it becomes more likely that the substrates share a transport process (C portion of the A B C test). If, however, any of the additional solutes affects transport of putative substrates differently, then either the substrates do not share the same transport system or multiple shared or unshared 8 We cite amino acid transport experiments especially beginning at this point in the text principally because the potential for using the values of kinetic parameters to show competition for transport via the same agency (systems summarized in Table 4.2) is greatest among the amino acids (see discussion in later Sections). Examples of transport catalyzed by agencies with substrate selectivities other than amino acids are discussed in detail in Chapters 5 to 7. The author asks the readers indulgence for his selecting his own work for purposes of illustration rather than many fine studies of other investigators.

TABLE 4.2 Summary of Some of the Distinguishing Characteristics of Amino Acid Transport Systems a Na+-Dependent systems A. For zwitterionic amino acids System A: prefers less bulky substrates System ASC: can be distinguished from system A by lack of reactivity with N-methyl substrates System B: broad scope system in trophectoderm of mouse blastocysts and possibly other epithelia System B~ A system in renal and intestinal epithelia that may be identical to system B System Gly: has strong preference for glycine and sarcosine System N: prefers glutamine, histidine, and asparagine B. For cationic and zwitterionic and amino acids System B~ broad scope system in oocytes, early conceptuses, and possibly some adult tissues C. For anionic amino acids System X-AG: prefers glutamate, aspartate, and other relatively small anionic substrates II. Na+-Independent systems A. For zwitterionic amino acids System asc: appears to be analog of system ASC System L: prefers bulky substrates System T: selects for benzenoid substrates B. For cationic and zwitterionic amino acids System b~ prefers large substrates that do not branch at the a- or/3-positions C. For cationic amino acids System bx+: substrate selectively nearly limited to cationic substrates System b2+: similar to bl + but interacts differently with specific substrates System y+: prefers cationic substrates but also transports certain zwitterionic substrates with Na + System y+L: higher affinity than prototypic system y+ for bulky zwitterionic substrates in the presence of Na + (distinct for system L, which is Na + independent for transport of zwitterionic substrates) D. For anionic amino acids System Xc-: prefers glutamate and relatively large anionic substrates

I.

aAdapted from Van Winkle, 1993, with permission from Elsevier Science.

transport processes are present. In the case of Na +d e p e n d e n t amino acid transport by blastocysts, L-valine and 2-aminoendobicyclo(2,2,1)heptane-2-carboxylic acid ( B C H ) inhibit transport of both L-alanine and B C O competitively with the same K / v a l u e s (Table 4.3), although the Ki values of L-valine are, of course, not expected to equal those of B C H . These data were used to support the conclusion that a single Na+-dependent system, t e r m e d B ~ transports both L-alanine and B C O in mouse blastocysts. The results of A B testing b e t w e e n L-alanine and L-lysine indicate that L-lysine also is probably a substrate of system B ~ The r e a d e r may observe, however, that more data are presented in Table 4.3 to

Kinetics of Saturable Transport

A

0.40

B

95

0.20

.c_

E 6.4 ~

o .Q

L-alanine

0.20

6.4 pM L-alanine

0.10

o

E T--

16.4 pM L-alanine

16.4 I~M L-alanine

46.4 I~M L-alanine -1.25

2.50

5.00

[BCO] (mi)

46.4 ~M L-alanine -0.15

0.30

[L-lysine] (mM)

0.60

FIGURE 4.26 Dixon plots of BCO (A) and L-lysine (B) inhibition of Na+-dependent L-alanine transport by mouse blastocysts. Inhibition is competitive in both cases as indicated by intersection of the lines above the x-axis (see also Fig. 4.23). The process catalyzing the transport depicted was subsequently designated system B~ (adapted from Van Winkle et al., 1985, with permission from American Society for Biochemistry & Molecular Biology).

support the conclusion that B C O and L-alanine share the same system than that L-lysine and L-alanine do. Moreover, B C O and L-lysine are thought to share system B ~ because they both compete for transport with L-alanine. Mutual inhibition of transport b e t w e e n B C O and L-lysine was not studied quantitatively. Hence,

TABLE 4.3 ABC Testing Indicates That L-Alanine and BCO Share a Na+-Dependent Transport System in Mouse Blastocystsa

Inhibitor and Ki (or Kin) b value (pM) Substrate BCO L-Alanine L-Lysine

BCO 430 320 --

L-Alanine BCH 29 35 35

1000 990

L-Valine L-Lysine 110 101 m

110 140

aln addition, the AB portion of the ABC test indicates that L-alanine and L-lysine share the same system. That is, the Km and Ki values in each column for BCO, L-alanine, and L-lysine (AB portions of ABC tests) and the Ki values in each column for BCH and L-valine (C portion of the ABC test) are statistically indistinguishable from each other based on the known experimental variability in such values for blastocysts (Van Winkle et al., 1990a-c). Hence, it can be concluded that L-alanine, BCO, and L-lysine are all transported by the same Na+-dependent system in blastocysts (see text for details). This system is termed B~ (data from Van Winkle et al., 1985, with permission from American Society for Biochemistry & Molecular Biology). bKm values are reported when an amino acid is listed as its own "inhibitor."

when concluding that two or more substances share the same transport process, both deductive and inductive processes may be particularly evident. Such conclusions may gain additional support over time, since the potential for A B C testing is often much greater than the testing that is actually performed.

4. Further Importance of Understanding the Meanings of Ki and Km When Designing Experiments Involving Inhibition of Transport A serious conceptual error in experimental design still appears occasionally in the current literature. In such studies investigators assume that by using inhibitor concentrations 10- or lO0-fold above the substrate concentration, they will detect inhibition by solutes that compete with the substrate for transport. Fortunately, since Km and Ki values frequently fall in the range of concentrations of substrates and inhibitors used in such studies, the anticipated inhibition is often detected despite the inattention to experimental design. I m p o r t a n t inhibition may, however, go undetected if such experiments are designed without preliminary recognition of approximate Km and Ki values. This point can be m a d e more obvious by considering a simple hypothetical example. Let us assume that an investigator plans to study previously u n e x a m i n e d amino acid transport processes in one of n u m e r o u s cell types that need to take up arginine. The investigator

96

4. Transport Kinetics

selects 10/xM L-arginine as the substrate concentration for initial screening and decides to test a series of amino acids and amino acid analogs as possible competitive inhibitors of the transport process. The investigator reasons that since the substrate concentration is 10 txM, he will be able to detect inhibitors that may also be good substrates by setting the inhibitor concentrations at 100 txM. The experimental design will work very well indeed if the Km and Ki values are all near, say, 10 tzM. If, however, the Km and Ki values are near 500 tzM for the best substrates and inhibitors, then only about 16% inhibition is anticipated, and this amount of inhibition may not be found to be statistically significant. For a transport process with Km and Ki values near 500 p.M, the process will, of course, appear not to be substrate or inhibitor saturable if the concentrations of substrate and competitive inhibitors selected for study are well below 500/xM. Before beginning such inhibition studies, investigators should obtain estimates of Km and Ki values. Alternatively, relatively low substrate and high inhibitor concentrations may be selected based on knowledge of the range of Km and Ki values for similar transport processes. For amino acid transport, a substrate concentration of i /zM is below the Km (and Ki) values of substrates for most known systems, whereas inhibitor concentrations of 10 mM exceed most such Ki (and Km) values. Since the substrate concentration is probably below the Km value for its transport, while the inhibitor concentrations are most likely above their Ki values for inhibition of transport, studies designed in this manner should reveal the anticipated inhibition. It is also important to note that these studies are made possible by the high purity of most commercially available preparations of individual amino acids (or on our ability to purify them before use in experiments), at least in regard to contamination by other amino acids. Nevertheless, some contamination by substances that inhibit transport may occasionally be encountered. Moreover, some cationic or anionic amino acid analogs at 10 mM may be deprotonated or protonated at neutral pH to an extent large enough also to interact significantly with systems for zwitterionic amino acids that have Km values near 1/xM. For these reasons, investigators must be willing to repeat such inhibition studies with different substrate and inhibitor concentrations when more precise determinations of some Km and Ki values indicate that the results of the initial study may not accurately represent the characteristics of the transport system(s) present.

5. Possible Significance of Other Types of Inhibition Our focus in this section has been principally on competitive inhibition and the meaning of Ki (and Km) values. We noted above, however, that inhibition may be

complex as indicated by the mixed (both competitive and noncompetitive?) inhibition of taurine uptake by arginine (Fig. 4.22B). In a different example, inorganic and organic cations inhibit cationic amino acid transport via system b ~ competitively (e.g., Fig. 4.21B), but incompletely (e.g., Fig. 4.27). In this case, the Ki values for Na + and Li + can be calculated to be about 14 mM (see Fig. 4.21B and Eq. (4.35)) but these calculations obscure the fact that inhibition by the cations is inhomogeneous. For example, using the Ki value of 14 mM for monovalent cations, it can be calculated that the rate of lysine transport is somewhat slower than anticipated at 18.8 mM Na + (sodium acetate in Fig. 4.27), whereas it is nearly 70% more rapid than anticipated at 150 mM Na +. The latter data for the acetate salt of sodium can not be explained by activation of Na+-dependent transport at the higher Na § concentration because Na § dependent amino acid transport also requires C1- ions in blastocysts (Van Winkle et al., 1988b) and because the same effect is observed for other cations (Figs. 4.21B and 4.27). Furthermore, cations per se do not influence leucine transport by system b ~247Apparently, cations influence transport via system b ~ at a cation receptor subsite for the side chains of cationic amino acids. The substrate receptor site does not appear to include this cationspecific subsite when zwitterionic amino acids are transported. The resistance of lysine transport to complete

TMAO (I-I) NaCI (0) Taurine (A) Na Acetate (O) Choline CI ( I )

A

"7 C: ~

~E c~

=.o -i"

9

E 0

0 I 300

I

.....

milliOsmolarity I, , milliOsmolarity

I,,

150 of indicated I 150

I

substance I

I

3o0 i

o

of sucrose

FIGURE 4 . 2 7 Effect of cationic substances on L-lysine uptake via system b ~ When the milliosmolarity of the indicated substances was less than 300, sucrose was used to adjust the total milliosmolarity to this value. Inhibition can be shown to be attributable to cations rather than anions as described elsewhere (adapted from Van Winkle et al., 1990c, with permission from Elsevier Science).

Kinetics of Saturable Transport

inhibition by inorganic and organic cations (Fig. 4.27) may therefore occur because the lysine receptor site includes both the distinct cation receptor subsite and a part of the site that receives zwitterionic amino acids. Cations that are not a-amino acids would not compete with lysine for the latter site and, hence, may not be able completely to inhibit lysine transport (Fig. 4.27). It has been proposed that similar sites and subsites may also be present on proteins responsible for amino acid transport via systems y+, ASC, and asc and that these receptor sites or subsites may, therefore, have a common evolutionary origin (Van Winkle et aL, 1990c). The cation harmaline is an especially strong competitive inhibitor of cationic amino acid transport via systems b ~ asc, and y+, and it competitively inhibits the interaction of Na + with the Na+-dependent system ASC (Young et aL, 1988, 1991; Van Winkle et al., 1990c). Moreover, the Ki values for harmaline inhibition of systems b ~ and y+ are considerably higher in isotonic solutions of electrolytes than in solutions of nonelectrolytes. Hence, harmaline appears to interact with the putative cation receptor subsite described above. Harmaline is, however, also a noncompetitive inhibitor of zwitterionic amino acid transport via systems b ~ asc, and ASC. If Scheme (4.17) applies to these transport systems, then harmaline probably decreases the value of the rate constant k2 for zwitterionic substrates, whereas it most likely increases k_l/ka for cationic amino acids. Such an effect on k-1/kl could be due largely to competition between harmaline and cationic amino acids for binding at the putative cation receptor subsite. Selective reduction of the k2 value for zwitterionic but apparently not for cationic amino acids may also result from interaction of harmaline with the same subsite as cationic amino acids. Presumably, when a cationic amino acid molecule occupies the cation receptor subsite, harmaline cannot influence the value of k2 because harmaline is not bound to the subsite. In the case of Na+-dependent transport of zwitterionic amino acids via system ASC, harmaline competes with Na + for interaction with the putative cation receptor subsite. Thus it is likely to reduce the k2 value and, hence, vi, regardless of the concentration of the zwitterionic amino acid substrate. Similarly, zwitterionic amino acid transport via system y+ is Na+-dependent, although the effect of harmaline on transport has not been investigated in that case. Nevertheless, one might predict from these data that an as-yet unidentified cation is required for symport or antiport of zwitterionic amino acids via systems b ~ and ascl. Harmaline could produce noncompetitive inhibition of zwitterionic amino acid transport by competing with such a cation for its receptor subsite. In fact, it has been suggested that uptake of zwitterionic amino acids via system b ~ occurs by obliga-

97

tory exchange for intracellular K + or amino acids, some of which are cationic (see Section XI,E below). If harmaline is taken up by cells, then it could competitively inhibit exodus of cationic substrates and, hence, act as a noncompetitive inhibitor of zwitterionic amino acid uptake via system b ~ Alternatively, when harmaline is bound to cation receptor subsites, it may be bulky enough partially to obstruct the pathway through the membrane for zwitterionic amino acids. Zwitterionic amino acids following or attempting to follow the pathway through the membrane would, however, not be able to influence noncompetitive harmaline inhibition according to this alternate scenario. D6ves and Angelo (1996) proposed an entirely different theory to account for these complex kinetics. These authors suggested that inhibition of system y+ by cations could be attributed to an influence of the cations on the negative surface potential. That is, at low ionic strength, the negatively charged glycocalyx and possibly some phospholipid at the external surfaces of biomembranes would attract cationic amino acids and, hence, increase their concentrations near the surface. Such an effect would decrease the apparent Km value for transport, although the actual value would, of course, remain unaltered. Moreover, the effect would be masked by cations in the extracellular solution, which would, according to this theory, mimic inhibition of cationic amino acid transport. These authors also found that in the case of human erythrocytes, the effect is actually attributable to system y+L rather than to system y+. Their theory cannot, however, easily explain the much stronger inhibition by harmaline than by other cations. Moreover, in the case of system b ~ the zwitterionic amino acid taurine that is excluded as a substrate by system b ~ is, nevertheless, effective as an inhibitor of cationic amino acid transport (Fig. 4.27). To be consistent with the theory of D6ves and Angelo, taurine must, therefore, interact with the negative surface potential of cells in the same way as cations, including that both zwitterions and cations are concentrated at the surfaces of cells at low ionic strength. Zwitterionic amino acids that are also substrates of system b ~ should, therefore, also be concentrated at the membrane surface according to this theory. As discussed above, however, inhibition of system b ~ by cations is restricted to its cationic amino acid substrates; cations per se do not inhibit leucine transport by system b ~ Moreover, cations in the medium do not prevent the large effects of membrane surface potential on transport of cationic and anionic substances by CaCo-2 cells and brush border membrane vesicles (Iseki et aL, 1997). Hence, we think that it is unlikely that negative surface potential alone accounts for the complex kinetics of transport described above for systems b ~ y+, asc, and ASC.

98

4. Transport Kinetics

Molecular studies can be expected to help us to understand further the results of these kinetic studies, especially if investigators employing, say, site-directed mutagenesis, also remember to perform detailed kinetic analyses of pertinent mutant forms of transport proteins. The presence of multiple subsites to receive substrates for transport also opens the possibility that so called leaks for the migration of low-molecular-weight hydrophilic solutes may have their basis not only in slippage of normally coupled transport processes, but also in heretofore undetected transport of solutes not thought to be substrates of the process. For example, most members of the family of proteins that transport excitatory amino acids were found also to serve as channels for inorganic anions (e.g., Section XI of Chapter 3) so they presumably also have receptor sites for such anions (Zerangue and Kavanaugh, 1996a). The channels open for anion transport in the presence of excitatory anionic amino acids, which are themselves transported in a Na+dependent but C1--independent manner (Amara, 1992). In fact, some members of this family of proteins might be viewed more accurately as ligand-gated anion channels than as excitatory amino acid transporters (Sonders and Amara, 1996 and see also discussion of E A A T proteins in Chapter 6). The unappreciated channel activity of other systems and proteins thought currently to have only one transport function might yet be interpreted as leaks in instances where the basis for the mediated transport have not been identified. In the case of system b ~ the binding of cationic solutes to a subsite not required for the transport of zwitterionic amino acids may, nevertheless, lead to transport of the cationic solutes independent of whether zwitterionic amino acids are also present. By virtue of their binding to the cation receptor subsite, even cations that are not amino acids may thus be transported by system b ~ albeit with high Km values relative to the transport of amino acids. 9 While such apparently nonspecific transport of solutes by a single transport process may be relatively small and hence difficult to detect, the combined effects of such transport processes might come to require consideration for a given solute. In the cases of inorganic cations, amino acid transport systems b ~ ascl, y+, and ASC could be part of the physical basis of apparent "leaks" for these ions in biomembranes, although in our view such leaks must still involve equally complex mechanisms of transport that are characteristic 9 In fact, as discussed above, zwitterionic, cationic, and possibly even anionic amino acids should, by virtue of their positively charged amino groups, be transported by the high Km transport process postulated here. This mechanism could conceivably even account for the unanticipated high Km component of transport associated with expression of the amino acid transport related proteins BAT and CAT2a (see Section X,E below).

of all transport proteins (also see Section X,A of Chapter 3 regarding leaks for inorganic ions in biomembranes). In this regard, some Na+-dependent glutamate transporters also appear to transport several cations, including Na + in a glutamate-independent manner, and a similar result has been observed for other Na +dependent transport proteins (summarized in Vandenberg et al., 1995). Interestingly the Km value for Na + transport alone by the Na+/glucose symporter is lower than the value for its cotransport with glucose, although the velocity of Na+transport at physiological Na + concentrations is much higher for symport than for transport of Na + via this "leak" (Chen et al., 1997). Let us now turn our discussion from the sometimes complex characteristics of single transport systems and proteins to the practical problem of how the activity of a single process may be obscured by other transport and metabolic processes. In the following Section we discuss methods to minimize or deduct the effects of these other processes so that the characteristics of the transport process of interest may be examined.

X. IDENTIFICATION AND MINIMIZATION OR DEDUCTION OF PROCESSES THAT MAY OBSCURE A TRANSPORT PROCESS OF INTEREST The most frequent reason why a transport process for a solute or the solvent may be obscured or even overlooked is that the biomembrane contains several different transporters of the substrate. Transport of organic solutes may be further complicated by their metabolic alteration. Since some of the aspects of the procedure for studying transport of a given type of solute or the solvent may be specific to the substrate, and because our space is limited, we consider in this section mainly the transport of organic solutes (but see Chapter 11 for procedures for the study of the transport of other solutes). While the specific procedures for the study of transport may vary among substrate types, many of the principles for isolating and characterizing transport processes are the same regardless of the solute. One of these principles is the use of inhibitors of selected transport systems to isolate and study the activities of other transport processes. For example, ouabain can be used to inhibit Na+K+ATPase so that other transport processes for Na + and K + can be characterized separately. As we shall see, this approach also is used commonly in the study of multiple transport processes for an organic solute. In addition, it may be necessary regardless of the solute to measure transport via other processes and to deduct transport by these other processes from total transport to yield transport by the process of interest.

Identification of Obscuring Processes Each of these approaches may, however, introduce error. For example, in cases where transport that is m e a s u r e d under one condition must be deducted from transport m e a s u r e d under another, the difference is less precisely known than either of the original measurements. F u r t h e r m o r e , transport via the process of interest may be altered in ways not involving direct inhibition when experiments are p e r f o r m e d in the presence of inhibitors of alternate transport processes. We consider below several such methods for isolating the transport activity of a single process. In addition, we describe an example of how it may be reassuring to show that the characteristics of the transport process do not change when changes are applied to the experimental protocols that are used to isolate the transport activity in intact biomembranes. Since we will limit our discussion principally to organic solutes, we will begin our discussion by considering how metabolism of these solutes may influence characterization of their b i o m e m b r a n e transport.

frequently the case when accumulation of radiolabeled metabolites of the substrate occurs and can be measured. Nevertheless, investigators must remain alert to the possibility that at some time point in the uptake of an organic substrate, its metabolism may b e c o m e at least partially rate determining. If such a change does not result in a detectable decrease in the velocity of net uptake of the substrate, then errors about the characteristics of transport might be made. Such a circumstance could arise, for example, if the initial rate of accumulation of substrate owing to uptake were not easily perceived as different from the rate of accumulation of metabolites of the substrate that obtains at steady state intracellular substrate concentration (Fig. 4.28). It is also important that metabolism not become more rate determining as the external substrate concentration is

Net substrate taken up /

A. Effect of C o n c u r r e n t S u b s t r a t e M e t a b o l i s m Investigators of b i o m e m b r a n e transport must sometimes be concerned about the effects of solute metabolism on the kinetics by which a solute appears to be taken up by cells. For example, it was believed formerly that apparently saturable fatty acid transport was attributable instead to saturable metabolism of these substances and that the migration of fatty acids across biom e m b r a n e s actually was not substrate saturable. Subsequently it was shown that fatty acid transport is substrate saturable because uptake into cells could be m e a s u r e d over a time period short enough to preclude detectable fatty acid metabolism (Stremmel and Berk, 1986). Hence, saturable fatty acid metabolism cannot account for the observed saturation p h e n o m e n o n . In order for metabolism to help to limit uptake, enough substrate must accumulate inside cells to permit a significant portion of it also to be transported out of the cells via the same or other routes. W h e n the rates of uptake and exodus of a substrate reach a steady state, but net uptake of the substrate still occurs, then some other process, such as metabolism, must account for the observed net uptake of the substrate. Conversely, if metabolism is fast enough to maintain the intracellular concentration of the transported substance near zero, then such a process would actually aid in the accurate determination of kinetic parameters. Such rapid metabolism would virtually ensure the unidirectional flow of the transported substance and, hence, could help to m a k e the m e a s u r e m e n t of initial velocities relatively easy. In such cases, the metabolites must still be identifiable as substrate that has been taken up, as is

99

o

~' I--

/

/

/

~

/"

/ Substrate metabolized

~/"

Intracellular

g

0

t

2

3

time ( rain )

4

FIGURE 4.28 Illustration of how the characteristics of substrate metabolism might be mistaken for the characteristics of substrate transport into cells. In the schema shown, the intracellular substrate concentration ([S]) would increase initially owing to uptake with little exodus or metabolism. As the intracellular [S] increases, however, so will its exodus and metabolism. The intracellular [S] will reach a steady state when the rate of uptake is exactly equal to the rate of exodus plus metabolism. To reach an intracellular steady state IS], the initial velocity of substrate transport must exceed the rate of substrate metabolism. As the steady state [S] is approached, however, substrate exodus will become more significant and the net rate of uptake will approach the rate of metabolism. It is also assumed that metabolites of the substrate remain inside cells and are detected as representing substrate that has been taken up. In this case, the choice of a 2-min period over which to measure the initial velocity of transport would reflect both the rate of transport and the rate of metabolism. A choice of a shorter time period would, of course, reflect mainly the rate of transport, whereas a longer one would reflect mainly the rate of metabolism. In such circumstances, an inhibitor that decreases net uptake of substrate at longer but not at shorter periods of measurement probably influences substrate metabolism rather than transport. In contrast, an inhibitor of transport that is not itself transported should decrease net uptake regardless of the time period over which net uptake is measured.

| 00

4. Transport Kinetics

changed. Somewhat surprisingly, that latter phenomenon is, under some circumstances, more likely to occur as the substrate concentration is lowered rather than as it is raised (see Section X,B below). Such errors usually can be prevented if one is careful to show that, during the period over which the initial velocities are to be measured, uptake increases linearly with time across the range of extracellular substrate concentrations to be used in the study. A related practical problem arises when metabolic products of the substrate leave the cell. If formation and exodus of the product is rapid enough, then the useful metabolic "sink" described above will be lost unless the amount of product exiting the cell can be measured. This measurement would be relatively easy for some metabolic products, such as 14CO2, provided that the assay is designed to detect all such metabolic products. If the transported substrate is, however, not radiolabeled or not otherwise distinguishable from endogenous substrate, then metabolites produced specifically from transported substrate would become virtually impossible to identify. In the latter case, substrate taken up could not be distinguished from the same substance already present in the cell or produced endogenously by it. Such problems can be obviated by studying the transport of metabolism-resistant substrate analogs. While these analogs are useful in transport studies, it is still the central goal to characterize the transport of naturally occurring substrates. A principal reason for studying transport processes in the first place is to learn how solutes that are encountered by cells in situ migrate across their membranes. A final concern is that products of the transported substrate may influence substrate transport. Assuming that the products are not present initially, their effects on transport might be greater at higher exogenous substrate concentrations (depending on the Km value of the rate-limiting enzyme in the pathway producing the metabolite and the concentration of the metabolite needed to influence transport). In practice, however, such unintended influences of metabolic products appear to be absent or minimal in most studies, especially when transport is measured over time intervals short enough to reflect initial velocities. For example, if accumulation of a metabolite were to inhibit uptake, then its accumulation would result in transport that increased more slowly with time. Hence, a time interval would be chosen for subsequent studies that is short enough not to include this slowing of uptake. When such initial velocities are measured, one can calculate the initial velocity of transport for the process of interest by subtracting the initial velocities of transport via other processes from the initial velocity of total transport.

B. Deduction of Transport via A p p a r e n t l y Nonsaturable Routes Even prior to the development of modem computer software, deduction of the apparently nonsaturable component of transport depicted in Fig. 4.29 from total transport was in many instances relatively easy to accomplish. Particularly when the Km values for mediated transport are relatively low, the nonsaturable component can be so small as to be negligible at the substrate concentrations required for study of the saturable ones (e.g., Fig. 4.30). Somewhat surprisingly, however, when apparently nonsaturable transport is relatively conspicuous, it may be necessary to add a quantity to rather than subtract a quantity from total transport in order to remove the effect of the nonsaturable component of transport (Fig. 4.31). Perhaps even more surprisingly, an addition rather than a substraction is more likely to be needed at lower rather than at higher substrate concentrations (e.g., Fig. 4.31). When a saturable process is so rapid and able to accumulate substrate against its total chemical potential gradient that the intracellular total chemical potential greatly exceeds the extracellular total chemical potential after a very short time interval, it may become necessary to correct for nonsaturable transport with an addition to rather than a subtraction from measured total transport. The total chemical potential gradient formed through saturable transport would be proportionally more steep for lower substrate concentrations, since the kinetics of saturable transport are closer to first-order as the substrate concentration is reduced below the Km value. Recall also that the total chemical potential gradient of a substance that carries no net charge is proportional to the logarithm of the ratio of its intracellular to extracellular concentrations rather than to the magni-

Tt - 912 "T

E

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~r 8 O

E e ._o E

X'Saturable

~4

eW

o

Not saturabl,

4

8

12

16

20

[o~-Aminoisobutyrate] m M

FIGURE4.29 Deductionof nonsaturable transport from total transport yields transport owing to saturable processes. The rates of a-aminoisobutyrate (AIB) uptake into the cells of the rat diaphragm were measured over the indicated range of extracellular AIB concentrations (results of Akedo and Christensen, 1962). Reproduced, by permission, from Christensen (1975).

Identification of Obscuring Processes

y

101

Observed-,.,~,

.34

~L

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I'/ ~~L-le)ucine ( 9) I i= ~ I L-lysJne ( 9) 0 l 0

~

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component

I

I

I

10 20 Amino acid concentration, mM

FIGURE 4 . 3 0 Nonsaturable transport of L-lysine is relatively slow at lysine concentrations needed to study transport via system b ~247 in mouse blastocysts. The extracellular concentration of L-[3H]lysine was about 1/xM at all concentrations of nonradioactive amino acids studied. Based on these data, one would determine Km and Vmax values for lysine transport by component b ~ at total concentrations of lysine of about 300/xM and below. At these concentrations, the nonsaturable component of lysine transport can be seen to be relatively small (uninhibited 1/i is marked 0.54) (adapted from Van Winkle et al., 1988a, with permission from American Society for Biochemistry & Molecular Biology).

tude of its concentration difference across the membrane. (Concentrations are again assumed to approximate activities. It is also assumed for the present illustration that the intracellular substrate concentration is initially zero.) In this case, significant nonsaturable exodus of a substrate could begin to occur while uptake of the substrate via the saturable route continues to increase almost linearly with time. It must be determined on a case by case basis whether nonsaturable transport can simply be deducted from total transport to give transport via saturable routes (e.g., Fig. 4.29) or whether more complex adjustments, such as those shown in Fig. 4.31, are needed for the particular substrate and cell type under investigation. C. Use of Inorganic I o n - D e p e n d e n c e to Deduct or Otherwise Minimize Transport of Organic Solutes by Other M e d i a t e d Processes

1. Study of Inorganic Ion-Dependent Transport Processes Numerous transport processes for organic solutes are inorganic ion dependent, and most such processes probably involve symport or antiport of the organic and

"- ~'~

eeee

Z

~'"

,'I

~

Passive Nonsaturable Process 0

Substrate Concentration

in Suspending Solution

FIGURE 4.31 The correction for passive, nonsaturable uptake may be either positive or negative depending on various factors including the extracellular total chemical potential of the substrate. The total chemical potential is proportional to the logarithm of the concentration when the net charge on the substrate is zero (a similar plot has been made for uptake of cationic and anionic substrates; Christensen and Liang, 1966a). A time interval was chosen that was short enough to approximate the initial velocity of uptake for the mediated component, but such was not the case for the nonsaturable process. At low external substrate concentrations, the cellular substrate levels achieved via the saturable component exceeded the external levels. Hence, the nonsaturable component had a net outward direction, and the total measured uptake had to be increased to reflect actual mediated uptake. At high external substrate levels, the cellular substrate concentration remained below the external level, and the net direction of nonsaturable transport was inward. In the latter case, nonsaturable uptake could be deducted from total uptake to yield mediated uptake (data form Winter and Christensen, 1964). Reproduced, by permission, from Christensen (1975).

inorganic solutes. Such symport or antiport should, of course, be demonstrated rather than assumed to occur by studying the dependence of inorganic ion transport on the presence of the organic substrate. Similarly, the stoichiometry of transport of the cosubstrates should be measured directly rather than inferred from the effect of the inorganic ion concentration on transport of the organic solute (see also Section III,B of Chapter 6). These experiments may be technically difficult to perform since the inorganic ion flux associated with organic solute transport may be small relative to the total flux of the inorganic ion. For the practical purpose of studying organic solute transport, however, the inorganic iondependence of transport of an organic solute can be exploited to study the inorganic ion-dependent process regardless of whether the inorganic ion actually is also transported. Such an approach has been used particularly frequently for the study of Na§ transport processes although it is in theory equally feasible for any inorganic ion-dependent transport process.

102

4. Transport Kinetics

For example, Na+-dependent amino acid transport has been studied in many types of cells by performing experiments in the presence and absence of Na +. The Na+-dependent component can then be obtained by deducting transport measured in the absence of Na + from transport in its presence. Deduction of the Na+-independent component also usually results in deduction of apparently nonsaturable transport, so this approach is particularly appealing when it is possible. Nevertheless, it is important to remember that when measuring initial velocity, a linear increase with time of the quantity of substrate taken up must be measured both in the presence and in the absence of the inorganic ion. While investigators usually remember the latter requirement, other important considerations are more frequently overlooked. First, measurement of the inorganic ion-dependent and -independent components of transport usually requires that a control osmolyte be substituted for the inorganic ion. It must be shown, however, that the control osmolyte does not itself influence transport. It is perhaps obvious that if Li + can substitute even in a limited way for Na + in stimulating transport, as is the case for amino acid transport system A, then a different cation should be selected to replace Na + in order to measure transport that is not Na + stimulated. It is, however, less frequently appreciated that the osmolyte that is used as a substitute may inhibit either the iondependent transport of interest (perhaps an advantage) or an ion-independent component of transport. The latter phenomenon could produce misleading results because some portion of the component of transport assumed to be ion dependent would, of course, actually be ion-independent. For example, both the Na+-dependent amino acid transport system B ~ and the Na+-independent system b ~ in mouse blastocysts are inhibited by choline (Van Winkle et aL, 1988a,b), and choline is frequently substituted as an osmolyte for Na +. Hence the difference between amino acid uptake by blastocysts in Na +- vs choline-based media would be greater than uptake by Na+-dependent systems and would represent transport via the Na+-dependent systems plus a portion of system b ~ Actually, inhibition of system b ~ extends beyond choline to other cations including Na + (see Section IX,D,5 above). The inhibition by choline is, however, greater than that by most other cations, and, unlike that by most other cations, it extends to the transport of zwitterionic as well as cationic amino acids (Van Winkle et al., 1988b, 1990c). Fortunately, other cations, such as Li +, that are used as osmolites to replace Na + have the same effect as Na + on amino acid transport system b ~ but not system B ~ in blastocysts. These other cations can therefore be used instead of choline to replace Na + in order to study Na+-dependent trans-

port by deducting transport in Na+-free medium from transport in the presence of Na +. In a similar vein, one must remain aware that the inorganic ion upon which transport is dependent may itself influence apparently ion-independent transport in unanticipated ways. In the example just discussed, Na + not only stimulates transport via system B ~ but it also surprisingly inhibits transport of cationic amino acids via system b ~ (Van Winkle et. al., 1990c). For this reason, if only sucrose or mannitol had been tested as an osmolyte to replace Na + in studies with blastocysts (or with other cells that contain systems with some of these same characteristics), then it might have been concluded erroneously that blastocysts do not contain a Na+-dependent component of cationic amino acid transport. In this case, loss of Na+-dependent system B ~ transport activity in the absence of Na + could have been obscured by an increase in transport via system b ~ (Van Winkle et al., 1985, 1990c). While conceptually simple, attempts to study inorganic ion-dependent components of transport by substituting another osmolyte for the inorganic ion may be fraught with unanticipated difficulty. One must perform enough studies to insure that the osmolyte used to replace the ion of interest does not, itself, influence transport in uncontrolled ways. The ideal replacement would be one that has all the effects on transport by various processes as the inorganic ion for which it is substituted except for the ion-dependent process that is to be studied.

2. Transport Processes That Do Not Require the Presence of an Inorganic Ion for Activity As discussed above, some transport processes for organic solutes do not appear to require the presence of an inorganic ion. In fact, these processes are sometimes inhibited by one or more ions. Fortunately, it is usually possible to identify osmolytes that are physiologically inert in regard to the transport processes intended for study, so that ion-independent transport processes may be studied in isolation from ion-dependent ones. It may also be useful to perform studies under more physiological conditions in which the inorganic ion that had been replaced for convenience of study is added back. For example, it could be hypothesized that the amino acid transport ascribed to system b ~ is simply an altered functioning of the Na+-dependent system B ~ in the absence of Na +. To test this possibility, we used a proven inhibitor of system B ~ namely BCO, that does not alter significantly transport by system b ~ If systems B ~ and b ~ were indeed separate processes, then system b ~ should be observed largely unobscured by system B ~ even in Na+-containing medium when enough BCO is present to inhibit most transport by

Identification of Obscuring Processes

system B ~ As shown in Fig. 4.32, such was observed for leucine transport in mouse blastocysts. The excess of BCO used to inhibit system B ~ also inhibited transport of leucine by a second Na+-independent system L in blastocysts. Moreover, the competitive inhibition of leucine transport by L-lysine had a Ki value that was expected for system b ~ (Fig. 4.32). Such tests of whether ion-independent transport processes are really separate from ion-dependent ones and, if they are separate entities, whether they are influenced in some unanticipated manner by the ion should be performed when the tests are possible. D. Aside from Their Fundamental Metabolic Importance, Amino Acids Are a Fortunate Choice for Study of Structural Specificity and H e t e r o g e n e i t y of Transport Mediation As indicated in the foregoing discussion, the transport of a solute or the solvent may occur via numerous routes across a biomembrane. The activities of each of these transport processes for the same substrate can be difficult to isolate for detailed characterization, especially when there is little tolerance for variability in substrate structure. Amino acid transporters may, however, each accept a wide variety of substrates albeit with different selectivities. This circumstance can be exploited for amino acid transport (and usually to a

!

i

i

I

i

!

.c_

E

lO r J~

-

-

/

/

Km =580 I~M V.~x--31fmol blastocyst 9 -I ~min -I I~. =73 I~M

0

05 -

/

No inhibitor K~ =1301~M V.~x=36 frnol blastocyst 9 -~ min 9 -~

w=-

L~ o

I

i

5O

,

,

9

I

130

1/[ L - l e u c i n e ] , m M -1

FIGURE 4 . 3 2 Effect of 300/xM L-lysine on the Lineweaver-Burk plot for L-leucine transport via system b ~ in the presence of Na +. Ten millimolar BCO was used to inhibit the interaction of lysine and leucine with transport system B ~ and the interaction of leucine with system L. Nevertheless, the transport rates were corrected slightly for residual uptake via these other processes to produce the data shown (adapted from Van Winkle et al., 1988a, with permission from American Society for Biochemistry & Molecular Biology).

103

lesser extent for transport of other solutes) to isolate and characterize multiple transport system activities that have overlapping but different substrate specificities. Moreover, it is possible to learn about the structures of the substrate receptor sites of amino acid transport systems by examining the structures of the amino acids and the amino acid analogs that they accept or do not accept as substrates. In combination with molecular studies that are now possible for transport-related proteins (Chapters 5 to 8) it should become feasible to produce detailed descriptions of the mechanisms of protein-mediated biomembrane transport, especially in the case of amino acid transport. The molecular characterization of transport proteins for many substances began before such characterization of amino acid transport proteins. Nevertheless, it can be anticipated that the rich structural diversity of amino acids and amino acid analogs will continue to facilitate detailed characterization of the three-dimensional structures and functions of their transport-catalyzing proteins. E. Analysis of Heterogeneity of Amino Acid Transport Investigators must learn to recognize evidence that for a given instance more than one transport process may operate. They must also remember to test their proposals as to the number of mediated transport processes they anticipate are present. The number of components of transport may be exposed by examining the best statistical fit of transport data in nonlinear regression analysis or even simply by carefully inspecting data in Hofstee plots. Because of the spacing of the data points, Hofstee plots are usually more efficient at exposing multiple components of transport than are plots of other linear transformations. For example, linear transformations of transport data for the amino acid transport-related proteins BAT (related to b ~ amino acid transporter, Fig. 4.33) and CAT2a (cationic amino acid transporter 2a, Fig. 4.34) unexpectedly exposed at least two kinetically distinct components of transport. While it seems to us most likely that such expression of each of these transport-related proteins results in the expression of more than one transport activity in oocytes (see Section X,J below), it is also conceivable that one or both of these transport-related proteins alone catalyzes more than one transport process. Regardless of their origin, the presence of two or more components of transport for a given solute in the same biomembrane can obscure detection and complicate characterization of each component. Multiple components of transport for a particular substrate may be obscured both by the design of the experiments (see below) and by the nature of the transport pro-

104

4. Transport Kinetics

300

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~ V~,x=28 Pmol ooocyte.1, min.1 50 i,-.--._.Component 2

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~

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400

600

800

mCAT-2a

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~

1000

1200

1400

ar~/[ L-Leu], n1-1 oocyte 9 -1 (5 9 rain)-1 FIGURE 4.33 Expression of the system b~ transport protein BAT in Xenopus oocytes increases L-leucine transport via at

least two kinetically distinct components of transport. The concave curvature of the Hofstee plot reflects these multiple components of transport. The data represent net transport resulting from rabbit BAT expression (i.e., the difference between transport in oocytes injected with BAT cRNA and those injected with vehicle). Transport resulting from BAT expression was resolved into the two components shown using the method of Spears et al. (1971) (adapted from Van Winkle, 1993, with permission from Elsevier Science).

cesses. For example, it would be impossible to detect two transport systems for an amino acid by kinetic analyses, such as those shown in Figs. 4.33 and 4.34, if the Km values for the two systems were virtually identical. In cases where Km values for different processes are within about one order of magnitude of each other, the technique of detailed analog inhibition analysis may be needed to detect, isolate, and study each of multiple transport processes for the same solute. The term analog is used here in the broadest sense to mean solutes that seem structurally similar enough potentially to compete for transport. The solutes may be naturally occurring or synthetic analogs of the naturally occurring ones. Although analog inhibition studies are useful in distinguishing and characterizing each of several transport activities for a solute, conclusions as to the completeness of their exposition must be considered provisional. Further studies may show heterogeneity within what had appeared previously to be a homogeneous component of transport. Formerly, these further experiments involved mainly the study of the effect of additional competitive and noncompetitive inhibitors, but more recently, molecular studies have revealed multiple forms of transport proteins with very similar transport activities.

0.0

0

10 20 30 I / [L-arginine] (nA / mM)

40

50

FIGURE 4.34 Expressionof the cationic amino acid transport protein CAT2a in Xenopus oocytes increases L-arginine transport via at least two kinetically distinct components of transport. The data represent the current produced by L-arginine transport owing to mouse CAT2a expression (i.e., the difference between the arginine current in oocytes injected with CAT2a cRNA and those injected with vehicle). Transport resulting from CAT2a expression was resolved into two components with Km values of 37/xM and 6.6 mM using the method of Spears et al. (1971) (adapted from Van Winkle et al., 1995, with permission from Elsevier Science).

F. The N a t u r e of t h e Interactions a m o n g A m i n o Acid A n a l o g s That C o m p e t e for Transport M a y Reveal H e t e r o g e n e i t y of Transport Mediation Solutes that compete for some but not every one of a group of transport processes may be useful in isolating and characterizing each of the transport activities. As we shall see, inhibition between two competing amino acid analogs may become quite complex, even for a single cell type. If such complexity is detected, however, the means are available to isolate and to characterize thoroughly each of the transport systems that together produce the complexity. The means for isolating and studying each transport activity depend on the observation that when a given amino acid is paired with each of several of its analogs, the pairs compete for different but usually overlapping sets of transport processes. Christensen (1975, pp. 179-181) listed five classes of inhibition that might be observed between competing pairs of analogs, and we bring the list to seven by adding the last two listed below. While at least 10 such types of mutual inhibition might be listed formally, we find it

Identification of Obscuring Processes c o n v e n i e n t t o discuss 4 of t h e s e t y p e s of i n h i b i t i o n u n d e r o u r t r e a t m e n t of Class I V b e l o w . W h e n o n e o r m o r e s u b s t r a t e - s a t u r a b l e p r o c e s s e s for t h e t r a n s p o r t of e a c h of t w o a n a l o g s a r e p r e s e n t in a b i o m e m b r a n e , a n d t h e y c o m p e t e for t r a n s p o r t via at l e a s t o n e of t h e p r o c e s s e s , t h e f o l l o w i n g a r e p o s s i b l e d e s c r i p t i o n s of t h e i n t e r a c tions b e t w e e n the analogs.

"lJ a

nonsaturable

component

T

_(

0

9

"=

[b]

"lJ a

T 0

nonsaturable component

,,

~ [b] ~--

| 05

1. Class I: C o m p l e t e and H o m o g e n e o u s T h i s t y p e of m u t u a l i n h i b i t i o n w o u l d o c c u r b e t w e e n t w o a n a l o g s w h e n a single s a t u r a b l e p r o c e s s s e r v e s t o m e d i a t e t h e u p t a k e of each. T h e s h a p e of t h e c u r v e s h o w n in d i a g r a m 1 of Fig. 4.35 w o u l d o b t a i n b o t h for i n h i b i t i o n of t h e u p t a k e of a n a l o g a at i n c r e a s i n g c o n c e n t r a t i o n s of a n a l o g b a n d for i n h i b i t i o n of t h e u p t a k e of

"lJ a

nonsaturable component

T

(..

[b]

0

"lJ a

nonsaturable component

T 0

(.

....

[b]

FIGURE 4.35 Diagrams to illustrate the slowing of transport via a mediated process for solute, a, on raising the concentration of an analog, b. The inhibition depicted in diagram 1 is termed "complete and homogeneous" (class I in the text) because uptake approaches a constant value attributable to apparently nonsaturable uptake as the concentration of the inhibitory analog is increased and because the shape of the curve is a precise rectangular hyperbola. The inhibition in diagram 2 is also homogeneous but it is incomplete. In this case (2), at least one other mediated transport process for a, that is not inhibited by b, appears to be present. As in diagram 1, the inhibition depicted in diagram 3 is complete, but now the inhibition is termed "heterogenous" because the curve is the sum of more than one (in this case two) rectangular hyperbolas. Each hyperbola reflects the presence of a distinct, substrate-saturable transport process. Since the eye is not efficient at analyzing the shapes of such curves, other means may be needed to assess them (see text). Similarly, the inhibition in diagram 4 is heterogenous but it is incomplete so a is also transported by at least one component not inhibited by b (adapted from Christensen, 1969 with permission from Wiley and Sons).

106

4. Transport Kinetics

b at increasing concentrations of a. The inhibition is said to be homogenous because the shape of the curve is a rectangular hyperbola, such as is observed when only one transport system participates. (More than one transport system might, in rare instances, also produce a single rectangular hyperbola when the K~ values of a given analog for each system are equal.) Inhibition is termed complete because the velocity of uptake of one analog via the saturable process approaches a constant value attributable only to apparently nonsaturable uptake as the concentration of the inhibitory analog is raised.

2. Class II: Homogenous but Complete in Only One Direction In this case, a curve such as that shown in diagram 1 of Fig. 4.35 (but with the positions of a and b reversed) would be obtained for inhibition of uptake of analog b by analog a, but when the positions of the analogs are reversed to the position actually shown in diagram 1, the curve shown in diagram 2 would instead be observed. The curve in diagram 2 is a rectangular hyperbola so inhibition is probably for a single system, but the inhibition is incomplete. Hence, there appears to be at least two mediated processes for transport of a; one that is shared with b and one or more that is not shared (diagram 2). While more complex interpretations for incomplete inhibition are sometimes possible (e.g., see Section IX,D,5. above), our assumption here is that when two analogs compete for transport by a single mediating agency, they are capable of completely inhibiting each other's transport. 1~

3. Class III: Homogeneous but Complete in Neither Direction In this case, two curves each of the same form as that in diagram 2 of Fig. 4.35 would be observed for mutual inhibition between the analogs. The simplest and most frequently correct interpretation for such data is that the analogs share one system and that they each migrate by one or more additional systems not shared by the other analog. In this scenario, further inhibition would be observed in diagram 2 on addition of other analogs that compete for transport via the systems that are not shared by analogs a and b. 10 The reader is reminded that inhibition is described here for analogs known to compete for transport. If it is not already known that analogs compete for transport, the characteristics of inhibition that are observed can be used to determine whether the analogs compete for one or more transport processes (see Sections X,G and X,H below).

4. Class IV: Complete but Homogeneous in Only One Direction Inhibition by one analog is in this case heterogenous and reflects the presence of two or more mediated transport processes. Although perhaps not conspicuous to the eye, the transport and its inhibition depicted in diagram 3 of Fig. 4.35 is for the sum of two mediated transport processes. Consequently, the curve in Diagram 3 is the sum of two rectangular hyperbolas. The simplest interpretation for this type of interaction is that one analog, a, is transported well by two substrate saturable agencies, each of which can be inhibited by the other analog. The same interaction is not observed, however, when the positions of the analogs are reversed apparently because analog b is not a good substrate of the second system. Hence, at the relatively low substrate concentrations that would be used for such analog inhibition studies, the transport of b will occur primarily by one system, the inhibition of which by analog a will appear to be homogenous and complete (i.e., the same as in diagram i of Fig. 4.35 but with the positions of a and b reversed). Such a result would be obtained, for example, in the study of the transport of leucine and valine by the two most conspicuous systems in mouse blastocysts. Leucine interacts strongly with both systems b ~ and B ~ in blastocysts, whereas valine interacts relatively strongly only with system B ~ Nevertheless, valine would inhibit leucine transport via system b ~ as well as via system B ~ as the valine concentration is raised. In contrast, leucine inhibition of valine transport would reflect primarily system B ~ activity at the relatively low valine concentration that would be used to measure its transport. The latter inhibition would appear, therefore, to be homogenous and complete, although an obscured small portion of it actually would occur also via system b ~ A variation of this type of interaction that belongs technically in a different class is that the heterogenous inhibition is incomplete. If the example above for blastocysts is now made hypothetical, a third component of mediated leucine transport can be imagined that does not interact even weakly with valine. On the other hand, if we modify the original example for blastocysts above so that an additional valine-selective system rather than a leucine-selective system is imagined, then inhibition would appear to be homogenous but incomplete in one direction and heterogenous and complete in the other. Finally, if entirely leucine-specific and entirely valine-specific systems were both present, then the inhibition would be incomplete in both directions.

5. Class V: Complete but Heterogenous in Both Directions The simplest interpretation of such results is that the two analogs compete for at least two transport processes. The same number of systems may be exposed

Identification of Obscuring Processes

in each direction of the inhibition although the system with the lowest Km (and Ki) value for one analog may not be the system with the lowest Km (and Ki) value for the other analog. It is also possible for reasons discussed under Class I V above that more agencies contribute significantly to the transport observed in one direction than in the other.

6. Class VI: Heterogenous in Both Directions, but Complete in Only One In this case, the analogs probably share multiple systems, but one analog also is transported by one or more saturable processes not shared by the other analog (diagram 4 in Fig. 4.35). In such cases, the process or processes not shared by one analog can be studied conveniently by performing experiments in the presence of an excess of that analog (see Sections X,G and X,H below).

7. Class VII: Heterogenous and Incomplete in Both Directions Not only are multiple systems likely to be shared by the two analogs in this type of interaction, but each analog also appears to migrate by one or more systems not shared with the other analog. Again, the systems that are not shared by the analogs can be studied by performing experiments in the presence of an excess of the analog that is not transported by the system(s) to be studied. The two added classes VI and VII follow logically from the five classes listed originally by Christensen (1975). Adding them here, however, emphasizes the likelihood that a complexity introduced for one test substrate may well apply to both. That risk has become more appreciated perhaps than it was in 1975 due in part to the finding that some systems receive both zwitterionic and cationic amino acids as good substrates. For example, the interaction between leucine and lysine for transport by mouse blastocysts is categorized under class VII. Mutual inhibition occurs between these analogs for transport via systems B ~ and b ~ but lysine is transported selectively by system b+2 and leucine is transported selectively by systems B and L (Van Winkle et al., 1985, 1988a, 1990d; Van Winkle and Campione, 1990). G. H o w Can the Interactions A m o n g C o m p e t i n g Sets of Analogs be Used to Isolate and Study Apparently H o m o g e n o u s C o m p o n e n t s of Transport? It is frequently difficult to perceive whether inhibition is for one or more than one transport process. For example, the sum of the two rectangular hyperbolas in Dia-

107

gram 3 of Fig. 4.35 may not appear greatly different than the single hyperbola in Diagram 1. Multiple transport processes may, however, be exposed by performing further analog inhibition analyses. One goal of analog inhibition analysis is to identify strong inhibitors of all but one component of transport. The activity of that component can then be isolated and its substrate selectivity and other kinetic characteristics studied in the presence of the inhibitors of the other components. That is, the goal is to reduce each possible class of the interactions defined above to a series of the simplest type of interaction under class L Although this goal is rarely realized for every component of substrate-saturable transport, the number of transport activities may be reduced to a number small enough to permit their deduction from each other by additional means. It must be remembered, however, that sometimes apparently homogenous components of transport may actually be heterogeneous especially when a solute has nearly the same Km (and Ki) value for more than one system in the same membrane. Furthermore, the selection of relatively low or high substrate concentrations for study may obscure detection of high or low Km systems, respectively, by low and high Km systems. For example, the low Km and conspicuous Na+-dependent component of zwitterionic and cationic amino acid transport, termed system B ~ (Van Winkle et al., 1985), obscured for several years detection of the high Km component of Na+-dependent zwitterionic amino acid transport, termed B, in mouse blastocysts (Van Winkle et al., 1990d). For reasons such as these, investigators must remain open to the possibility that apparently homogeneous components of transport actually may be heterogeneous even for both of two structurally dissimilar analogs expected to share fewer systems then structurally similar ones. Since the identification, isolation, and characterization of each of several transport processes for the same substrate may be complex, we describe an example of this procedure in the next Section. Possible difficulties include not only those inherent in this complex problem, but also a continuing inclination on the part of investigators to terminate characterization of transport prematurely. Even worse, many investigators frequently assume erroneously that the transport processes in tissues not previously well studied will nevertheless turn out to be forms of those already known. This haste may spring from the laudable desire to accomplish apparently more important goals, such as to learn the physiological functions of solute or solvent transport in a tissue. Similarly, newer investigators who have more thorough training in molecular biology than in classical biochemical procedures and techniques may be less likely to perceive hints from their data that the transport they observe may be complex or novel. Scientists must re-

| 08

4. Transport Kinetics

t~9 100

(l)

r ~ o9 e'-

(-. o e-o

+

i_

0

Compound tested as inhibitor

I

S

(10raM) _

-r '-r" _J Compound tested as inhibitor

(10mM; 5mM BCH)

FIGURE 4 . 3 6 Effect of various amino acids on Na§ L-leucine (A) and L-lysine (B) uptake by mouse blastocysts. Note in particular that 10 mM L-lysine is a partial inhibitor of L-[3H-]leucine (1/zM) uptake and vice versa (adapted from Van Winkle et al., 1988a, with permission from The American Society for Biochemistry and Molecular Biology, Inc.).

member, however, that their ability to perceive the correct physiological functions of transport depends on their ability to interpret properly the transport data they plan to and subsequently do obtain.

"

transport in mouse blastocysts with consideration of a broad approach to studying leucine transport. Not unexpectedly, tryptophan shares some but not all of its transport agencies with leucine. As we shall see, if tryptophan and leucine had been studied as described in the preceding Section, theirs would be a class VI interaction. Na+-independent L-leucine transport was studied initially in regard to the ability of a wide variety of structurally different amino acid analogs to inhibit it. To insure that systems with low Km and low Vmax values would not be obscured by systems with higher Km and Vmax values, the initial substrate concentration studied was about 1 /zM (at or below the anticipated Km value), whereas inhibitor concentrations were 10 mM (speculated to be above the Ki values) (Fig. 4.36). These initial observations revealed L-lysine to be an incomplete inhibitor of leucine transport while, in similar studies, leucine incompletely inhibited lysine transport. However, unlike the incomplete inhibition discussed for four of the seven classes listed in Section X,F above, it was not known from these initial studies whether the inhibition was incomplete because it was weak or because it was partial. To determine whether the mutual inhibition was weak or partial in each instance, it was studied at several inhibitor concentrations. As shown in Figs. 4.30 and 4.37, the interaction between leucine and lysine

~'i

I~

'-

.=.

i

i

i

i

"

'

'

E

H. An Example of the Use of Analog Inhibition Analysis to Isolate and Characterize at Least Five C o m p o n e n t s of Mediate Tryptophan Transport Apparently in the S a m e Biomembrane ~ Part of the decision about how to begin to determine the routes of transport of a solute, such as tryptophan, should be based on what is already known about transport of tryptophan analogs (in this case, other natural or synthetic amino acids) in the tissue of interest. Such knowledge will permit us to start by determining whether tryptophan is also transported by these systems for other amino acids rather than beginning as we might have otherwise with a broad approach to the study of the systems that catalyze migration of tryptophan across the membrane. Nevertheless, the initial approach to the study of amino acid transport in the membrane should be broad. For these reasons, we will begin our discussion of the elucidation of five mediated routes of tryptophan 11A summary of the procedure used in such studies is outlined in the Appendix to this Chapter.

"-

0.4 ,i

o >" 0

\

t~ 0.3

9

= 0.2 --

L-alanine ( u ), L-valine ( A), and D/ leucine ( )9

~

~ =~

/

f Componentb~ Nonsaturable -

"',,,

~

""

c~176

"a 01 14-

~"

Zw,~rJ~S . . . . . . . .

=

I

prefemng component

x ~ ' ~ l " ~ ~ ( ~ Lhtsidme( ) ~L-!eucine( 9) / 0 -9 --0 10 20 A m i n o acid c o n c e n t r a t i o n , m M "

"

"

t

O

"

'

i 0 II

/

FIGURE 4.37 Inhibition of L-leucine uptake by L-lysine is incomplete because it is partial rather than weak, and the inhibition is apparently homogenous. Conversely, uptake of L-lysine by blastocysts is inhibited in a similar way by L-leucine (Fig. 4.30). Blastocysts appear to contain at least three components of Na+-independent L-leucine transport that are termed here, component b ~ the zwitterionpreferring component (system L), and the nonsaturable component (adapted from Van Winkle et al., 1988a, with permission from The American Society for Biochemistry and Molecular Biology, Inc.).

Identification of Obscuring Processes

and unshared components of saturable transport of each amino acid. At the time of its discovery, the observation that leucine and lysine shared a Na§ component of transport was somewhat surprising. Hence, ABC testing was needed to show that this agency, now known as system b ~ does indeed transport both cationic and zwitterionic amino acids (Table 4.4). Leucine transport via component b ~247 could be studied directly by performing experiments in the presence of a bicyclic amino acid analog (BCO or BCH), which inhibits the zwitterionpreferring component (system L) known not to be shared with lysine (Fig. 4.38). In contrast, it was necessary to subtract the leucine-insensitive component from total Na§ lysine transport to characterize lysine uptake via component b ~247Nevertheless, these and other studies permitted us to conclude provisionally that component b ~247 is a single system and that its good substrates include relatively large cationic and zwitterionic amino acids that do not branch at the/3-position. From these results, we anticipated that blastocysts contain at least two Na§ transport pro-

TABLE 4 . 4 ABC Testing Shows That L-Leucine and L-Lysine Share a Na+-lndependent Transport System in Blastocysts a

Inhibitor and Ki (or Kin) b value (pM) Substrate

L-Leucine

L-leucine L-lysine

140 130

L-Lysine 79 48

L-Valine

L-Alanine

5600 4100

3400 m

109

aThat is, the Km and Ki values in each column for L-leucine and L-lysine (AB portion of the ABC test) and the Ki values in the column for L-valine (C portion of the ABC test) are statistically indistinguishable from each other based on the known statistical variability in such values for blastocysts (Van Winkle et al., 1990a-c). The shared transport system is termed b ~ (data from Van Winkle et aL, 1988a, with permission from American Society for Biochemistry & Molecular Biology).

bKm values are reported when an amino acid is listed as its own "inhibitor."

appeared to be homogenous (although possibly not), and it was partial (rather than weak) in both directions (class III or possibly class VII above) with both shared

0.75

o) o

Na+-dependent component (~sterns B o,. and B)

0.50

~5 (D C :3

Non,saturable component

"i5

~..

. . . . J,

Cydoleucin; u.zo,/k t (Na*-independent; A

1

Zwiterionpreferring component

tCO (Na+-ir~dependent; o ) Threonine (Na*-independent cinee (Na+-independent;B)

O! 0

.

.

.

.

Compone~ bO,§

.

4 8 Amino acid concentration, mM

I

12

FIGURE 4 . 3 8 BCO inhibits the Na+-dependent and zwitterion-preferring components of Lleucine transport, but not component b ~ or the apparently nonsaturable component. Uptake of L-[3H]leucine from a 0.8 /zM solution was measured at various concentrations of several analogs including nonradiolabeled L-leucine in the presence of Na + or after its replacement with Li + (Na + independent) (adapted from Van Winkle, 1988a, with permission from The American Society for Biochemistry and Molecular Biology, Inc.).

1 10

4. Transport Kinetics

cesses for tryptophan. The lack of branching of tryptophan at the fl-carbon meant that it might be a good substrate of component b ~ (Van Winkle et al., 1988a). Moreover, the component of leucine transport not inhibited by lysine appears now to be a form of the wellknown system L for which tryptophan is a good substrate. Although tryptophan shows some interaction with the component of lysine transport not inhibited by leucine (now known as system b+2; Van Winkle and Campione, 1990), the interaction is weak and it seemed (and subsequently was found to be) unlikely that much, if any, tryptophan is transported by it. Hence, we focused on components b ~ and L as potential systems for the transport of tryptophan. We also knew that cationic amino acids could be used selectively to inhibit tryptophan transport by system b ~ whereas bicyclic amino acids inhibit system L. These selective inhibitors were important because we wondered whether additional Na+-independent systems might be present in mouse blastocysts. Tryptophan transport had not been thoroughly characterized in these embryos, and at least one other system that selects for benzenoid amino acids is known to be present in other types of cells. The most direct way to test whether components of tryptophan transport in addition to those known to receive leucine as a substrate were present in blastocysts was to determine whether a leucine-insensitive component of tryptophan transport could be detected in these cells. (i.e., would leucine inhibition of Na+-independent tryptophan transport most resemble diagram 3 or diagram 4 in Fig. 4.35?). The inhibition was expected to be heterogenous (diagrams 3 and 4) rather than homogenous (diagrams 1 and 2) because we anticipated that tryptophan would be transported both by system b ~ and by system L (Fig. 4.35). As shown in Fig. 4.39, a leucine-resistant component of tryptophan transport was detected although a tryptophan-insensitive component of leucine transport was not (i.e., the inhibition is under class VI described above). While these data are for cleavage-stage conceptuses, the same three Na+-in dependent components of transport persist into the blastocyst stage, although the relative contributions of each change as development proceeds (Van Winkle et al., 1990b). Moreover, the leucine-resistant component of tryptophan transport was shown to be sensitive to inhibition by the known inhibitors of system T, D-tryptophan and N-methyl-L-tryptophan (Fig. 4.40), throughout preimplantation development of conceptuses. From these and other data we concluded that early embryos contain three Na+-independent components of tryptophan transport; namely systems b ~ L, and T (Table 4.5). As discussed above, our conclusion must remain provisional since additional heterogeneity could be revealed

in future studies of any of these apparently homogeneous components of transport. Interestingly, system T in erythrocytes transports triiodothyronine with a much lower Km value than it shows for tryptophan (Zhou et al., 1990, 1992). Nevertheless, the physiological concentrations of all benzenoid amino acids combined and of all thyroid hormones combined are well below the Km values for each group. Hence, these substances probably do not interfere much with the transport of one another under most physiological conditions. The importance of this dual transport of related groups of substances to the functioning of erythrocytes and other cells is still being investigated. 12 In addition to these Na+-independent components of tryptophan transport, it can be seen from Table 4.5 that blastocysts are thought also to contain two Na +dependent components of transport. The more conspicuous component of Na+-dependent tryptophan transport was shown using the AB portion of the ABC test (Table 4.5) probably to be shared with L-alanine. Moreover, the most conspicuous component of L-alanine transport in blastocysts is system B ~ (Van Winkle et al., 1985). Hence this system is probably also the most conspicuous component of Na+-dependent tryptophan transport in blastocysts. In addition, a less conspicuous component of Na+-dependent transport was observed in the presence of enough L-lysine (20 mM) and BCH (10 mM) to inhibit virtually all system B ~ activity (Fig. 4.41). These amino acids also would have inhibited systems b ~ and L, respectively, and the experiments were performed in the presence of 2 mM N-methyl-Ltryptophan to inhibit system T. Since neither L-arginine nor trans-4-hydroxy-L-proline inhibited transport of radiolabeled tryptophan under these conditions, whereas nonradioactive L-tryptophan did inhibit it (Fig. 4.41), we concluded that this component of transport should be attributed to system B in blastocysts. This conclusion is supported by the observation that tryptophan strongly inhibits L-alanine transport by system B in blastocysts (Van Winkle et al., 1990d). For these reasons, we believe that alanine and tryptophan would show the class I interaction described above when the component of transport presumed to be system B is isolated in these ways. Since ABC testing has not been performed to establish the scope of substrates received by system B, however, it is conceivable although we think unlikely that another or an additional Na+-dependent process transports tryptophan in blastocysts. Because of its inconspicuous nature, the component of tryptophan transport attributed to system B (Fig. 4.41) was not characterized further. 12See Section IV,F on aromatic amino acids in Chapter 10 for a discussion of triiodothyronine transport in other cells.

Identification of Obscuring Processes

A

B

L-Leu -resistant compon,e,nt

120

120

120 lOO

100

"ID

L-Leu

80 e-

"

80

L-Harg

t

c t,-

N

40

0

80

~

20

a.

100

~

o C

40

-~ 60

No detectable L-Trp -

resistant component

100

.="_

60

o C

0

10

20

D-TrD

=.d

L-Trp

a. ,--m

X['3H] keu

0 =

60

40

t_ 0

-r

,--I

,_,1

.

"1" p....,

"T-

~-.

111

20

I 10

2O

30

0

0

I 30

41- L-Trp -e- L-Leu

L-Trp 10 20 Amino acid concentration, mM

,I 20

1 20 30 Amino acid concentration, mM

40

FIGURE 4.39 Early mouse embryos contain an L-leucine-resistant component of Na+-independent Ltryptophan transport (A), but not a tryptophan-resistant component of leucine transport (B). The leucineresistant component of [3H]tryptophan transport (i.e., experiments performed in the presence of 10 mM Lleucine) was inhibited by D-tryptophan, N-ethylmaleimide (NEM), and nonradiolabeled L-tryptophan,but not by L-homoarginineor additional L-leucine (inset of A). The inset of B is for [3H]leucine uptake in the presence of 10 mM L-tryptophan and further supports the conclusion that a tryptophan-resistant component of leucine transport is not detected in early embryos, (adapted from Van Winkle et al., 1990b, with permission from Elsevier Science).

I. Are T h e s e Transport S y s t e m s Really I n d e p e n d e n t Entities? Analog inhibition analysis has proved to be a useful tool in the detection, isolation, and characterization of different transport activities. In the case of amino acid transport systems, it appears to be fortunate for investigators that the natural substrates have considerable structural diversity. This structural diversity increases the potential for variety among transport systems and thus may have made it possible for scientists to isolate and study the systems using analog inhibition analysis. Data obtained in such studies, however, beg the question whether the different systems are independent agencies. That is, could the same agency, such as a single transport protein, display characteristics of more than one transport system? It may be difficult to rule out this possibility for results such as those discussed above until the transport protein for each agency has been characterized and shown to be distinct. Nevertheless, we think that it is unlikely in most instances that the same trans-

port protein catalyzes more than one component of transport, unless it is associated with different accessory proteins or is modified in some other way. In fact, we think that molecular studies are likely to reveal additional heterogeneity within components of transport thought currently to be catalyzed by a single agency. Biomembrane transport processes, including transport ATPases, carriers, transport systems, and channels, are composed of at least one transport protein (e.g., transporters) and in some cases one or more accessory proteins usually of more poorly defined function ((e.g., fl-subunit of Na+K+ATPase) (see also Chapters 5 to 7). Thus, different transport processes may result from expression of different transport proteins p e r s e or from expression of different accessory proteins in association with a single transport protein or set of proteins. Wellstudied transport processes, such as Na+K+ATPase, have not as yet revealed dramatic changes in characteristics, such as substrate selectivity, when the transport protein (e.g., an a-subunit of Na+ATPase) is expressed in association with different accessory proteins (e.g.,

1 12

4. Transport Kinetics 150 ,--

L-Leu -resistant component

"U

II

"~100 9 ~

100

80

BCH

80

L-Phe

~.

rp

'~ 20

"~

~' "" o

50 L-Harg

L-Leu

010

, N-methyI-L-~rp 0

10

9 1~6 Amino acid concentration, mM

.1.--

"7"D-Trp

I 20

"-_

27

FIGURE 4.40 Inhibition of the L-leucine-resistant component of Na+-independent benzenoid amino acid transport in mouse blastocysts by D-tryptophan and N-methyl-L-tryptophan. These two L-tryptophan analogs are known to be inhibitors of system T in other cells (adapted from Van Winkle et aL, 1990b, with permission from Elsevier Science).

different/3-subunits of Na+ATPase; Chapter 5). In the case of amino acid transport systems that are thought to be composed of more than one protein molecule, current research is directed at how a putative accessory protein molecule (e.g., BAT) may activate more than

TABLE 4.5

one transporter molecule (e.g. Peter, et aL, 1996; Yao et aL, 1999). Some amino acid transport systems have, of course, now been shown to be independent entities through isolation and characterization of their transport proteins and the cDNAs that encode them (e.g., Van Winkle, 1993; Malandro and Kilberg, 1996). The molecular approach will eventually allow us to determine which amino acid transport processes are separate entities and which are not. Meanwhile, other data strongly support the view that different amino acid transport systems have a different molecular composition. First, a particular transport system may be expressed in association with different systems in different tissues. For example, although some of the transport characteristics of amino acid transport systems ASC and A are similar, and hence they could conceivably result from expression of the same transport protein, system ASC occurs independently of system A in several cell types (e.g., rabbit reticulocytes and pigeon erythrocytes). For this reason, it was concluded over two decades ago that systems ASC and A are separate agencies (Christensen, 1975). This conclusion has now been verified through the cloning of cDNAs encoding two transport proteins, the characteristics of which correspond reasonably well to system ASC but not to system A (Arriza et aL, 1993; Shafqat et aL, 1993; Utsunomiya-Tate, et al., 1996). In addition, several similar amino acid transport processes appear and disappear during development on different schedules. For example, the Na+-independent

M o u s e Blastocysts Contain at Least Three Na+-Independent Systems (L, T, and b ~247 and Two Na+-Dependent systems (B~247and B) for L-Tryptophan Transport a Values of kinetic parameters (73-77% confidence interval) b

Vmax

Km

Ki (inhibitor)

One-cell (Trp substrate) L T bO,+

4.9 (4.6-5.5) 76 (75-112) c

24 (21-38) 3500 (3500-5500) c

63 (BCH) 870 (N-methyl-L-Trp)

Blastocyst (Trp substrate) L T b~ B B ~247 B ~ (Ala substrate)

1.6 (1.3-2.4) 47 (36-58) 25 (24-26) c -~10 25 (22-27)

24 (13-47) 1700 (1200-2400) 55 (48-64) c --~8 32 (28-37)

589 (BCH) 780 (N-methyl-LTrp) 3.8 (L-Arg)

System and stage of development

--~20 (L-Ala) 13 (L-Trp)

aAlso shown are kinetic parameters for tryptophan transport and inhibition of its transport via the three Na+-independent transport systems in one-cell embryos. The AB portion of the ABC test indicates that tryptophan competes with L-alanine for transport via system B ~ (from Van Winkle, 1990b, with permission from Elsevier Science). bUnits for values are as follows: Vmax in fmol conceptus -1 min-1; Km and Kg in ~M. CUptake via the indicated component of transport was too slow relative to uptake via other components accurately to estimate kinetic parameters.

Identification of Obscuring Processes

"1o .O ,m JE

100

trans-4-OH-L-Pro

80

L-Arg

1 13

molecular studies should reveal whether any of these distinct transport systems share a component, such as an accessory protein or, conceivably, a transporter.

.c_ o

J. Expression of Heterologous Transport-Related Proteins in X e n o p u s Oocytes and Other Cells Is a Useful Tool in Verifying That Transport Systems are Separate Agencies

d N

60 (No Na*)

e-e-

o. 40 L-Trp

e~

._i

---, 9 20 -r

eo

0

I

4

I

8

I

12

I

16

Amino acid concentration, mM

FIGURE 4.41 Na+-Dependent transport of L-tryptophan that resists inhibition by substrates of the conspicuous Na+-dependent system B ~ in blastocysts. System B ~ activity was inhibited by performing experiments in the presence of 20 mM L-lysine and 10 mM BCH, which also inhibited systems b ~ and L, respectively. At the same time, system T was inhibited with 2 mM N-methyl-L-tryptophan. The lack of detectable system B ~ or system b ~ activity was verified by the inability of L-arginine to inhibit uptake of L-[3H]tryptophan, whereas nonradiolabeled L-tryptophan inhibited Na+-dependent uptake of its own radiolabeled form. This component of tryptophan transport is attributed to system B (adapted from Van Winkle et al, 1990b, with permission from Elsevier Science).

system b+2 appears, whereas b+l disappears during preimplantation development of mouse embryos (Table 4.6). In contrast, systems L and T remain about the same or decrease in activity somewhat during this same period. Similarly, the Na+-dependent system Gly disappears at about the same time that system B ~ appears during preimplantation development. At the intermediate four- to eight-cell stage both system Gly and system B ~ appear to be detectable (Table 4.6). It is difficult to envision such a dramatic change from narrow- (system Gly) to broad-scope (system B ~ amino acid transport without at least some chemical changes in the transport agency. In fact, we now know that the disappearance of system Gly from embryos occurs in association with loss of an mRNA encoding a glycinespecific transport protein, and the appearance or disappearance of several other transport processes is associated with the gain or loss of mRNA encoding the appropriate protein (e.g., Van Winkle, 1993; Van Winkle et al., 1994; Van Winkle and Campione, 1996). Hence, these and other instances of regulation of expression of transport system activities on different developmental schedules support the theory that most of the activities are catalyzed by separate agencies. Further,

Numerous investigators are now confident enough in the technique of expressing cRNAs in Xenopus oocytes to use it to study the detailed kinetics of transport via proteins that the cRNAs encode (e.g., Hirayama et al., 1996, 1997 and see Chapter 11 for further discussion). This procedure was devised initially to identify transport related proteins through expression cloning. Transport resulting from expression of a protein in oocytes or other cells is reflected by the difference between transport in oocytes injected with the pertinent cRNA and those injected with vehicle (control oocytes). For electrogenic transport, uptake or exodus can be studied by measuring the effect of substrate on electrical current across the oocyte plasma membrane at a constant membrane electrical potential difference. Moreover, it may become interesting to study transport at different electrical potentials. When transport is measured as electrical current, the same oocyte may be used for numerous measurements, thus reducing experimental variability and the number of oocytes needed for such studies. Significant stimulation of transport after expression of a cRNA in oocytes usually means that the cRNA encodes a transport-related protein. This procedure has been quite useful in helping to verify that numerous amino acid transport systems are indeed independent entities (Van Winkle, 1993; Malandro and Kilberg, 1996). The possible effects of the expression of an injected cRNA on more than one transport process in oocytes may, however, complicate the use of this procedure to study the characteristics of a single transport process in detail. A major (and usually unaddressed) concern with such studies is that endogenous transport-related proteins may influence and may be influenced by expression of heterologous (i.e., cRNA-encoded) transportrelated proteins. For example, at least two amino acid transport systems appear to increase in activity when the system b~ amino acid transport protein BAT, from rabbit, is expressed in Xenopus oocytes (Fig. 4.33) (Van Winkle, 1993). This unexpected complexity has been thoroughly verified for the BAT homolog from rat (Peter et aL, 1996). More recently, Young and associates (Yao et al., 1999) showed that expression of the rat BAT protein and the related

1 14

4. Transport Kinetics TABLE 4.6

Developmental Regulation of Expression of Biomembrane Transport Processes in M o u s e Eggs and Preimplantation Conceptuses a,b Transport activity detected at each stage of development

Process

Egg c

1-cell

2-cell

4 to 8-cell d

Blastocyst

Na+-dependent processes for amino acid *System B . . . . . *System B ND System XAGND *System XA -e ND System Gly +++ System/3 ND

9 +++ +

? 9 +++ +

+(?) +(?) 9 ? +++ +

+++ + +++ +++ +

Na+-independent processes for amino acids *System b ~ + *System b+2 System T +++ System Xc+++ System L +++ *System b+l + *Tau/Gly channel1 ND *Tau/Gly channel2 ND

+ +++ +++ +++ + + -

+ +++ + +++ + ? 9

+ 9 ND + +++ 9 9 ?

+++ +++ +++ ++++ +

Processes for inorganic ions Na+/K+-ATPase *Novel K + transporter

+++ ?

+++ +++

+++ ?

+++ +++

+

+

ND

+++

+++ 9

Na+-independent process for organic ions System for choline ND

aprocesses marked with an asterisk were first characterized in conceptuses. /)Key: - , not detected; +, detected; + + + , relatively conspicuous activity detected; ?, insufficient or ambiguous data; ND, not determined. We report transport activity per conceptus both to be consistent with previous studies and because the surface area increases by only about twofold in conceptuses between the 1-cell and blastocyst stages. Data from Haghighat and Van Winkle (1990); Van Winkle (1988, 1992, 1993); Van Winkle and Campione (1987, 1990, 1991, 1992, 1996); Van Winkle et al. (1985, 1988a,b, 1990a-e, 1991, 1992, 1993, 1994). COvulated, unfertilized egg. dMixtures of compacted and noncompacted conceptuses have usually been studied at the 4 to 8-cell stage of development. eApparently novel system XA-- has not been named previously. It prefers L-and D-aspartate as substrates over L-glutamate and over other amino acids.

protein 4F2hc each stimulate Na+-dependent pyruvate a n d u r i d i n e u p t a k e as w e l l as a m i n o acid u p t a k e (Fig. 4.42). T h e t r a n s p o r t is e x p l a i n e d m o s t e a s i l y b y t h e t h e o r y t h a t B A T is a n a c c e s s o r y r a t h e r t h a n a t r a n s p o r t p r o t e i n a n d t h a t its e x p r e s s i o n in o o c y t e s a c t i v a t e s m o r e t h a n o n e e n d o g e n o u s t r a n s p o r t p r o c e s s . 13 T h e t h e o r y t h a t a c c e s s o r y p r o t e i n s a r e c o m p o n e n t s of t r a n s port systems raises the reverse possibility, however, 13 Alternatively or in addition, it is conceivable that expression of proteins in the BAT family cause transport proteins to be moved from intracellular locations to the plasma membrane. It is also possible to envision other less direct effects of these proteins on transport, although the simplest explantation is that they have direct effects on transport protein activities (Van Winkle, 1999).

t h a t t r a n s p o r t p r o t e i n s e x p r e s s e d in o o c y t e s m i g h t interact with more than one endogenous accessory protein. Such an event could explain the observation ( V a n W i n k l e et al., 1995) t h a t e x p r e s s i o n o f e a c h of the known cationic amino acid transporter (CAT) p r o t e i n s in X e n o p u s o o c y t e s r e s u l t s in d e t e c t i o n o f at l e a s t t w o k i n e t i c a l l y d i s t i n c t c o m p o n e n t s of t r a n s p o r t (e.g., Fig. 4.34). F u r t h e r m o r e , t h e e x p r e s s i o n of m a m m a l i a n t r a n s p o r t - r e l a t e d p r o t e i n s in a s s o c i a t i o n w i t h Xenopus proteins could produce different characteristics o f t r a n s p o r t t h a n t h e c h a r a c t e r i s t i c s n o r m a l l y associated with expression of the protein. Alternatively o r in a d d i t i o n , it is c o n c e i v a b l e t h a t t r a n s p o r t - r e l a t e d p r o t e i n s a r e a b l e t o f u n c t i o n in m u l t i p l e c o n f o r m a t i o n s

Identification of Obscuring Processes r

.r

"7

E lo.0 - A

Na* medium

o o3

~

E

7.5

o

o

o o

m

o

o

E

5.0

~.

2.5

1 15

ii

0.6

C

Na* medium Ch § medium

0.4

E o.

0.2 ,.r,,

II

t-

~o

=9

0.0 r4F2hc BAT

H20

r4F2hc BAT

H20

r4F2hc BAT

H20

A

"T

r-

~T t" .m

E

o 03

|

0.0

B

I

6

I Na*medium

Ch § medium

oE

03

d,

500

-D

r--]

4oo

Na + medium

Ch + medium

o

o E

4

~

300

~

2OO

"

100

E

(3.

~. 2

ID e~

~

L

o

0

|

0

..I

r4F2hc BAT

H20

r4F2hc BAT

H20

r4F2hc BAT

H20

r4F2hc BAT

H20

FIGURE4.42 Uptake of choline, pyruvate, uridine, and leucine in Xenopus oocytes expressing rat BAT and 4F2hc transport-related proteins. Oocytes were injected with 10 nl of water or water containing jejunal 4F2hc or renal BAT cRNA (1 ng/nl) and incubated for 5 days at 18~ in MBM. (A) Uptake of choline (0.2 mM, 20~ 30 min); (B) uptake of pyruvate (0.2 mM, 20~ 30 min); (C) uptake of uridine (10/zM, 20~ 30 min); (D) uptake of L-leucine (0.2 mM, 20~ 30 min). Fluxes were measured in NaC1transport buffer (open columns) or in choline chloride transport buffer (hatched columns). Each value represents the mean _ SEM of 10-12 oocytes (adapted from Yao et aL, 1999 with permission from the National Research Council of Canada).

each for which is associated with different Km and Vmax values. In the later case, one conformation may be favored more in Xenopus oocytes than in mammalian somatic cells. Because of these possibilities, conclusions about the physiological characteristics and functions of m a m m a lian transport-related proteins must remain provisional if they are based solely on expression of the proteins in Xenopus oocytes. Furthermore, investigators must r e m e m b e r to determine whether expression of a transport-related protein in Xenopus oocytes influences more than one transport activity for a given solute. Otherwise, emergent models that sometimes employ multiple kinetic steps to describe transport (e.g., Hirayama et al., 1996 and 1997) may depend on an erroneous

assumption that a single process catalyzes transport of the solute. K. Even Purified T r a n s p o r t Proteins M a y Display M u l t i p l e C o m p o n e n t s of T r a n s p o r t W h e n R e c o n s t i t u t e d in P r o t e o l i p o s o m e s A substrate may have multiple components of transport, owing to the presence in biomembranes of more than one protein involved in its transport (see above). Even purified transport proteins are, however, observed sometimes to display complex kinetics consistent with the presence of more than one c o m p o n e n t of transport when they are reconstituted in proteoliposomes (e.g., Fig. 4.43). Several explanations for such observations

1 16

4. Transport Kinetics 1.5

1.0

E

p0.5

0.2

0.4

0.6

0.8

v,/[S o] (sec-1)

1.0

1.2

FIGURE4.43 Biphasickinetics of lactose exchange by the purified E. coli/3-galactoside transporter in proteoliposomes. Liposomeswere loaded with 3 mM [1-14C]lactose, and the initial rates of exodus were measured in a pH buffered solution containing 0.1 to 25 mM unlabeled lactose. The kinetics of transport are consistent with the presence of multiple components of transport or negative cooperativity or both of these phenomena. See text for discussion of the mechanisms that may produce these kinetics (adapted from Lolkema et al., 1991, with permission fom The American Chemical Society).

are possible and considerable experimentation may be needed to distinguish among them. In the case of the E. coli/3-galactoside transporter (Fig. 4.43) formation of a dimeric transport protein molecule was considered less likely than the existence of protein monomers with two binding sites (Lolkema et al., 1991). Both sites could be catalytic or one could be a regulatory site. In the latter case, substrate binding to the regulatory site could raise the Km and Vmax values such that the protein would function reasonably well for transport over a wide range of substrate concentrations. Similarly, if the transporter has two catalytic sites, either inherent to the monomeric protein molecule or as a result of its dimerization, then the two sites could exhibit negative cooperativity for transport. Negative cooperativity as well as the presence of two transport sites with inherently different Km values could produce the curved Hofstee plots frequently interpreted to mean that multiple transport activities are present. Production of positive and negative cooperativity through oligomerization of transport protein monomers is considered further in

Chapters 5 and 6. In addition, it is possible that a transport protein monomer has different conformations such that one form has Km and Vm~x values different from the other. Even if high and low Km components of transport are inherent to a purified and reconstituted protein monomer, however, several explanations for the phenomenon must be considered. In addition to the possibility that each monomer alone may catalyze both components of transport, it is also possible that some of the monomers are chemically or physically different from the others. For example, some of them may have been phosphorylated or otherwise altered chemically to produce monomers with different transport characteristics. Similarly, the tertiary structures of some of the monomers may have been altered during their isolation, and such alterations could produce changes in the characteristics of transport. Finally, some monomers may be reconstituted in their reverse orientation in proteoliposomes, and the kinetics of transport are likely to differ for the two orientations. Artifactual causes of multiple transport activities can be ruled out by studying transporter function also in native membranes, although the underlying physiological causes and functions of multiple components of transport may still need to be identified.

XI. KINETIC DIFFERENCES A M O N G SUBSTRATE-SATURABLE TRANSPORT PROCESSES THAT FORM, PROPAGATE, OR DISSIPATE SOLUTE GRADIENTS The kinetics of transport attributed to single transport-related proteins and single transport processes can be studied in intact cells and after expression of the proteins in X e n o p u s oocytes and other cells. Based on data obtained from such studies, detailed kinetic models of transport are proposed and tested further. Because of interactions that may occur among transport-related proteins in the same cell, however, such detailed models for single proteins and single processes must be considered provisional until they can be verified using purified transport-related proteins in proteoliposomes. Nevertheless, several major differences in the kinetics of transport are known for processes that form, propagate, or dissipate solute gradients. We review here some of these differences and refer the reader to other sources for more detailed (and, in most instances, hypothetical) considerations of these kinetic differences. (For example, see Stein, 1986 for a good compilation of earlier work, and articles such as those by Parent et al., 1992 and Hirayama et al., 1996, 1997 for application of these kinetic models to transport

Kinetic Differences Among Transport Processes

proteins expressed in Xenopus oocytes.) Procedures for kinetic modeling are also described in greater detail in Chapter 11 of this volume. A. Processes That Form Solute Gradients The reader may recall from Chapter 3 (Section II) that we discuss transport processes that form, propagate, or dissipate solute gradients in this decreasing order of thermodynamic complexity because propagation and dissipation of solute gradients occur across biomembranes of living cells only because the gradients are formed in the first place by primary active transport (see also Chapter 3, Sections V I - X and Chapters 5 to 7). Hence, a conspicuous difference between primary active and other forms of transport is that only primary active transport is thought to move solutes against their total chemical potential gradients without dissipating gradients of other solutes or the solvent (but see also Section IX,C,2 above). Gradients formed through primary active transport can then be utilized to perform other work, such as to produce gradients of other solutes or physical or electrical changes in cells (see Sections XI,B to XI,G below). The kinetics of primary active transport also frequently involve the migration of two or more of the same ion or molecule together in the same direction across biomembranes. In the latter cases, the stoichiometry of transport appears to have evolved in association with constraints resulting from the steady state solute gradients that appear to be needed across biomembranes and the maximum free energy available from the hydrolysis of a phosphoric acid anhydride bond. For example, as discussed in Chapter 3, the free energy available from hydrolysis of ATP is only slightly higher than that required to extrude 3 Na + and take up 2 K + in one cycle of the process catalyzed by Na+K+ATPase. This and other ATPases appear to have evolved the capacity to occlude and then transport one or more ions depending on the free energy that is required to move them against their total chemical potential gradient and that is available from ATP hydrolysis. When two or more of the same ion or molecule are transported in the same direction in one transport cycle, the kinetics of their transport differ somewhat from simple Michaelis-Menten kinetics for transport of one ion or molecule at a time. When two or more ions or molecules are transported together, the plots of vi vs substrate concentration (IS]) are sigmoidal in shape (e.g., Fig. 4.18) rather than hyperbolic (Fig. 4.19). The sigmoidal curves can be rendered hyperbolic by plotting vi vs [S] n, where n is the number of identical ions or molecules that are transported in the same direction in a single transport cycle (Eq. (4.27)). It might also seem

1 17

appropriate to describe the uptake of 2 K + ions and extrusion of 3 Na + ions during the transport cycle as a type of antiport. As in antiport, Na + on the inside of the cell stimulates K + uptake and K + outside stimulates Na + extrusion. Similarly, the transport of two or more ions together in the same direction across a biomembrane could conceivably be termed symport. The terms antiport and symport are, however, usually reserved to describe transport that does not occur as a direct consequence of ATP hydrolysis. Rather, symporters and antiporters propagate solute gradients but they are not commonly believed to form them. B. Transport Processes That P r o p a g a t e Solute Gradients The solute gradients formed during primary active transport may be either dissipated (Section XI,G below) or used to form and maintain gradients of other solutes. In antiport, a gradient of one solute can be used to generate a similar gradient of another solute. In symport, a solute gradient in one direction across a biomembrane may be used to form and maintain a gradient of another solute in the reverse direction. The kinetics and thermodynamics of this transport differ fundamentally from primary active transport because at least one of the solutes must migrate in the direction of its lowest total chemical potential on either side of the membrane. Similarly, the kinetics of symport and antiport differ from uniport (including that occurring through channels). In symport and antiport, the movement of one solute along its gradient appears directly to be coupled to the migration of another solute against its total chemical potential gradient. No such direct coupling occurs in uniport, although uniporters may under some conditions use solute gradients to produce gradients of other solutes by a different mechanism (see Section XI,G below).

1. Symport In symport, each of two or more different ions or molecules appear to be transported together across a biomembrane. Consequently, each solute also stimulates transport of the other. For example, in the case of Na+/glucose cotransport across the intestinal epithelial plasma membrane, Na § and glucose each stimulate uptake of the other into cells. Since Na § is extruded from cells via Na+K+ATPase (and in some cells by Ca 2§ ATPase in combination with Na+/Ca § exchange), the movement of dietary glucose into epithelial cells against its total chemical potential gradient can be viewed as being driven by the migration of Na § with its gradient. Hence, the gradient of Na § produced in primary active transport

1 18

4. Transport Kinetics

is propagated as a glucose gradient. The gradient is then usually dissipated through exodus of glucose via uniport on the serosal side of the epithelial cells (Van Winkle, 1999). In other cases, however, the solute gradient formed by a symporter can be utilized to produce yet another solute gradient by a different symporter or antiporter.

2. Antiport Exchange transport (antiport) of a solute may or may not be obligatory. When exchange is nonobligatory, migration of the solute ion or molecule in one direction across the m e m b r a n e is stimulated by migration of the same (homoexchange) or another (heteroexchange) solute ion or molecule in the reverse direction, but migration may still occur in either direction, albeit at a slower rate, without exchange. Transport of cationic amino acids via system y§ is an example of this nonobligatory exchange (White and Christensen, 1982). The capacity to catalyze nonobligatory exchange (i.e., both antiport and uniport) may be viewed as slippage of an antiporter or futile cycling of a uniporter depending on the perceived primary function of the transport process. We discuss in Section XI,F below how regulation of whether a transporter catalyzes antiport or uniport may be used by erythrocytes to decrease or increase net glucose uptake depending on free energy needs. It is likely, however, that we are only beginning fully to appreciate how conversion of transporter function between antiport and uniport may be used to regulate interorgan nutrient

flows and metabolism. In the following sections we describe kinetic distinctions among obligatory exchange, nonobligatory exchange, and uniport after which we use the same kinetic model to explain cis- and transinhibition a n d - s t i m u l a t i o n phenomena. We then consider how the absorption of some nutrients across renal and intestinal epithelia may be driven owing to their uptake by obligatory exchange. As we shall see in Section XI,E below, the latter antiport appears to be driven by symport of other nutrients with Na § which in turn is driven by Na§247 C. Kinetic Distinctions A m o n g O b l i g a t o r y Exchange, Nonobligatory Exchange, and Uniport According to the model shown in Fig. 4.44A, a substrate ion or molecule, R, may be transported from side 1 to side 2 across a b i o m e m b r a n e only when the transport protein, M, is oriented in the m e m b r a n e to receive the substrate. In the ping-pong model shown, this orientation is depicted as the transport protein being near the side of the m e m b r a n e from which transport is to occur. We now know, however, that this mobile carrier model is probably incorrect in most instances. Nevertheless, the model is the same kinetically if the transport protein is viewed as having two conformations, one of which receives (or releases) solute only on side 1 of the m e m b r a n e and the other of which receives (or releases) solute only on side 2. It is also assumed that steps 4, - 4 , 2, and - 2 in Fig. 4.44A occur as a result of the

FIGURE 4.44 Ping-pong model to distinguish among obligatory exchange, nonobligatory exchange and uniport. (A) In obligatory exchange, the rate constants for steps 4 and -4 are zero. The rate constants for steps 4 and -4 have values greater than zero but less than the values of the rate constants for steps 2 and -2 in nonobligatory exchange. Hence, both obligatory and nonobligatory exchange exhibit trans-stimulation. In contrast, the values of the kinetic constants for steps 4 and -4 are equal to the values of the constants for steps 2 and -2 in uniport. Consequently, uniport does not exhibit trans-stimulation. In all cases, the values of the kinetic constants for steps 1, -1, 3, and -3 are assumed to be much larger than the values of the other kinetic constants. (B) The same model may also be applied to cis inhibition of transport of the substrate, R, by the competitive inhibitor G. In the case shown, G on side 1 of the membrane would slow the migration of R from side 1 to side 2 of the membrane owing to cis-inhibition. No such slowing of the migration of R from side 2 to side 1 would occur initially, however, owing to the absence of G on side 2. See text for further discussion.

Kinetic Differences Among Transport Processes

proposed conformational change of the transporter and that the values of the kinetic constants for these steps are much smaller than the values of kinetic constants for steps 1, - 1 , 3, and - 3 (i.e., the conformational changes of the transporter are the slow steps). Under these assumptions, obligatory exchange would occur when the values of the kinetic constants for steps 4 and - 4 a r e zero. In this case, the transporter would be trans-stimulated by the same or another substrate on the other side of the membrane because the transport protein undergoes its conformational change only when associated with a substrate molecule. A similar result would be obtained if obligatory exchange were viewed to occur as a result of transport of substrate simultaneously in each direction. Interestingly, antiporters usually are viewed to operate by transporting solute in one direction at a time, whereas symporters usually are assumed to transport two or more substrate ions or molecules simultaneously across the membrane. Although kinetic studies usually are designed with these models for symport and antiport in mind, few studies appear to be designed to distinguish between the possibilities of simultaneous and nonsimultaneous transport of substrate ions and molecules. Interestingly, when such tests have been performed for obligatory anion exchange proteins, the results support the conclusion that the proteins operate by a two-site simultaneous mechanism rather than a ping-pong model (see further discussion in Section II,B of Chapter 6). According to the ping-pong model usually applied to antiport, exchange will be nonobligatory but still transstimulated when the values of the kinetic constants for steps 4 and - 4 are greater than zero, but less than the values of the constants for steps 2 and - 2 (Fig. 4.44A). In this case, the transporter functions both for uniport and antiport, but uniport is slower than exchange, owing to a more rapid conformational change when the protein is associated with substrate. When, however, the values of the kinetic constants for steps 4 and - 4 are equal to those for steps 2 and - 2 , then the kinetic characteristics usually attributed to uniport are observed. In this case the transport protein can change conformation to receive substrate on either side of the membrane equally rapidly regardless of whether it is associated with substrate. Consequently no trans-stimulation is observed for uniporters. The other possibility not considered extensively in other Sections of this volume is that the values of the kinetic constants for steps 4 and - 4 exceed those of steps 2 and - 2 . In this case, the transporter would exhibit trans inhibition. Examples of trans-inhibition have been observed for amino acid uptake both by yeast and by Neurospora. In addition, trans-inhibition may in some instances account for the lower-than-anticipated

1 19

accumulation of one cosubstrate via symport, at least in bacteria (Wright, 1989). Some of the instances of trans-inhibition of amino acid uptake have, however, been attributed to special feedback loops rather than to the kinetic model described here (reviewed by Christensen, 1975, pp. 153-156). Alternatively, transinhibition might occur because steps 2 and - 2 for some amino acids exceed steps 2 and - 2 for others. In this case, the second group of amino acids would produce trans- as well as cis-inhibition of the first group as appears to be the case for obligatory exchange via system ASC (see Section III,B,3 of Chapter 6). D. The S a m e Kinetic Model May Be Used to Explain trans-Stimulation, trans-lnhibition, cis-lnhibition and c/s-Stimulation The kinetic model just used to explain how transstimulation and trans-inhibition may occur can also be applied to cis-inhibition and cis-stimulation. In cisinhibition, another substrate (e.g., G in Fig. 4.44B) simply competes with the substrate, R, for the transport protein, M. Under the conditions shown in Fig. 4.44B, however, G initially slows the migration of R from left to right across the membrane, whereas no such effect on the migration of R from right to left would occur initially. Under such conditions, G would cause a net migration of R from right to left. This phenomenon is considered further in Section XI,G below where we explain how uniporters and channels may, under some circumstances, form a total chemical potential gradient of one solute while they dissipate the gradient of another. The phenomenon of cis-stimulation may also be explained using the present kinetic model. This phenomenon appears to have been observed for a uniporter of lysine in the basolateral membrane of intestinal epithelia (e.g., Munck and Munck, 1997). In this case, intracellular leucine seems to stimulate exodus of lysine from enterocytes. Such an effect could conceivably occur for a uniporter if leucine somehow increases the rate constants of steps 2, - 2 , 4, and - 4 (Fig. 4.44A), possibly owing to an allosteric effect. Testing of this possible allosteric mechanism for the cis-stimulation of lysine transport by leucine must be preceded by unambiguous demonstration that the phenomenon actually occurs in these cells. Nevertheless, cis-stimulation of lysine exodus from enterocytes by leucine would help to explain the Na+-dependent net transport of cationic amino acids across the intestinal epithelium of species that have a Na+-dependent transport system for leucine but not lysine or arginine (Munck and Munck, 1997). Absorption of cationic amino acids from the lumen of the intestine

120

4. Transport Kinetics

also appears to involve their uptake via obligatory exchange across the apical membranes of enterocytes. E. Apparently Obligatory Exchange of Amino Acids as an Example of Propagation of Solute Gradients via Antiport The amino acid transport associated with expression of the b~ protein BAT in X e n o p u s oocytes appears to occur only through exchange of an extracellular amino acid molecule for an intracellular one. If this obligatory exchange is also associated with BAT expression in renal and intestinal epithelia, then it could account for the efficient absorption of cationic amino acids and cystine against their total chemical potential gradients (Chillar6n et al., 1996). In order to show that such gradients are formed by obligatory exchange, however, it is usually necessary to show that transport is transstimulated. It must be demonstrated rather than assumed that solute gradients are formed through transstimulation of an antiporter because even a strict uniporter (i.e., one that does not exhibit trans-stimulation) may also catalyze the migration of a substrate against its gradient owing to cis-inhibition by another substrate (Section XI,G below). Moreover, in the case of obligatory rather than nonobligatory exchange, the transport rate should be zero when substrate is not present on the other side of the membrane. In practice, it is difficult to demonstrate that the rate of amino acid transport by a system in X e n o p u s oocytes is zero when no substrate is on the other side of the membrane. Not only may rabbit BAT expression influence more than one transport activity in these cells (see Section X,J above), but the total intracellular concentration of amino acids is about 6 mM, and this total is difficult to reduce to zero (Chillar6n et al., 1996). In the case of cut-open oocytes, it was shown that the current, owing to alanine exchange for arginine, is reduced to zero, as expected, when arginine is not present on the other side of the membrane (Coady et al., 1994). Such experiments do not, however, preclude electrically silent alanine transport. Moreover, exodus of arginine, cystine, and leucine from oocytes can be measured in the absence of extracellular amino acids (Chillar6n et al., 1996). In the latter case, it is assumed that the exodus occurs via systems other than those influenced by BAT expression since exodus of arginine, cystine, and leucine does not increase when BAT is expressed. Although it remains conceivable that BAT expression in X e n o p u s oocytes is associated with a small amount of undetected uniport, it is clear that exchange accounts for most of the amino acid transport associated with it. Zwitterionic amino acid uptake owing to BAT expression is electrogenic because these amino acids

trans-stimulate exodus of cationic ones. Moreover, this current is greater when the proportion of intracellular amino acids that are cationic is higher. Finally, the amino acid transport activity attributed to BAT expression appears to catalyze a nearly 1:1 exchange under a variety of conditions in which both amino acid influx and efflux have been measured (Chillar6n et al., 1996). Obligatory exchange via BAT-associated transport system b ~ in the apical membranes of renal and intestinal epithelia would lead to uptake of cationic amino acids and cystine against their concentration gradients if they are exchanged for dipolar amino acids that had been accumulated against their total chemical potential gradients by another process (Fig. 4.45). The system for the latter transport in the brush borders of kidney and intestine has been proposed to be the Na+-dependent symport system B ~ (Chillar6n et al., 1996), although the Na+-dependent system B ~ also appears to be conspicuous in the epithelia of some species such as rabbit (Munck and Munck, 1994, 1995). In this Scheme, the

+

AA* or C s s C

AA o

o

,

I

~o

l

1

FIGURE 4 . 4 5 Model for the reabsorption of cationic amino acids (AA +) and the tetrapolar amino acid cystine (CssC) via system b ~ in the apical membrane of the renal epithelium. In the model, dipolar amino acids (AA ~ are taken up against their gradient via a Na*/AA ~ symporter termed B ~ The gradient of A A ~ is then used to drive uptake of AA + and CssC against their gradients via the b ~ antiporter. A A + and AA ~ (including Cys produced from CssC) are then proposed to leave epithelial cells via systems y+L and L, respectively, in the basolateral membrane (adapted from Chillar6n et al., 1996, with permission from the American Society for Biochemistry & Molecular Biology, Inc.).

Kinetic Differences Among Transport Processes Na+-dependent accumulation of dipolar amino acids is driven by the Na + gradient that is produced and maintained by Na+K+ATPase in the basolateral membrane (Fig. 4.45). Once cationic amino acids and cystine have been accumulated in this manner by the epithelial cells, the cationic amino acids could leave across the basolateral membrane via system y+L, perhaps in exchange for Na + and zwitterionic amino acids. Finally, zwitterionic amino acids including cysteine (produced from cystine) could leave epithelial cells by system L (Fig. 4.45). Lysine may also leave the cell via a system in the basolateral membrane (not shown) that appears to be cisstimulated by leucine (e.g., Munck and Munck, 1997). A more direct coupling to a gradient formed by Na+K+ATPase may also occur if exodus of K + occurs in exchange for amino acid substrates of system b ~ across the apical membrane (Ahmed et al., 1995). The apparent ability of some forms of system b ~ to exchange K + for amino acids would also help to explain how the current attributed to B A T expression can be reversed in hyperpolarized oocytes (i.e., those caused to have an inside negative electrical membrane potential difference of a greater magnitude than - 1 0 0 mV). In the later case (Fig. 4.46), the positive current that results from uptake of alanine in exchange for cationic amino acids or K + could be reversed, owing to exodus of endogenous alanin e (or other zwitterionic amino acids) in exchange for exogenous K +. Not all forms of B A T appear to be associated with K + transport, however (Section III,E,1 of Chapter 6). An alternative explanation for current reversal across the hyperpolarized membrane is that cationic substrates of system b ~ accumulate temporarily in the unstirred water layer on the outside of the plasma membrane owing to exchange for zwitterionic amino acids at higher (i.e., less negative inside) membrane electrical potential differences (Fig. 4.46). These cationic amino acids in the extracellular unstirred water layers might be taken up to reverse the current when the membrane electrical potential difference is suddenly made more negative (Coady et aL, 1994). If such is the case, it is unclear why zwitterionic amino acids do not accumulate in unstirred water to permit current reversal when arginine is present outside the oocyte (Fig. 4.46).

F. Reversible Regulatory U n c o u p l i n g of Linked Transport Is Well Illustrated by the N a + - I n d e p e n d e n t Glucose Transporter GLUTI If the obligatory exchange described above for system b ~ became less tightly coupled, then a dipolar amino acid molecule that had been taken up via system

121 200 I(nA)

100 -

-200 i

-

100J

100

i..

___~

.T_

i

V m (mV)

Ala

Arg -100

]

FIGURE4.46 The normally outward current of positive charge owing to exodus of cationic amino acids in exchange for alanine can be reversed at sufficiently high inside-negative membrane electrical potential in Xenopus oocytesexpressing the rabbit BAT protein. This reversal of current may result from exodus of zwitterionic amino acids in exchange for extracellular K+ when the magnitude of the inside negative membrane electrical potential becomes sufficiently large. Alternatively, cationic amino acid substrates, detained temporarily in an extracellular unstirred water layer, might be taken up and hence reverse the current when the magnitude of the inside-negative membrane potential is suddenly increased from the holding value of -50 mV to values between -100 and -175 mV (see text for further discussion). Also shown are currents for the uptake of arginine at various electrical potential differences presumably in exchange for intracellular zwitterionic amino acids (adapted from Coadyet aL, 1994, with permission from Elsevier Science).

B ~ would be able to leave the cell via system b ~ without concomitant uptake of another substrate molecule. Such slippages of coupled transport have been observed to occur in the primary active transport processes that form solute gradients as well as among the transport systems that propagate the gradients. For example, slow extrusion of Na + via Na+K+ATPase is observed in the absence of K +, although in this case most of the free energy in ATP may still be captured in the Na + gradient. When such slippages occur in antiporters or symporters, however, they lead to loss of some of the free energy stored in solute gradients. Since no process is 100% efficient, various symporters and antiporters probably fall on a continuum from those that catalyze highly coupled transport with minimal slippage to those that also catalyze uniport well. The apparently obligatory exchange associated with B A T expression is an example of highly coupled transport, whereas amino acid transport system y+ continues to operate quite well for uniport when substrate for exchange is not available on the other side of the plasma membrane (White and Christensen, 1982).

122

4. Transport Kinetics

Superimposed on this range of the possible degrees to which transport of one solute is coupled to another may be the capacity in some cases to regulate whether the transporter functions more for uniport or more for coupled transport. In this regard, uniport of glucose by the GLUT1 transport protein in human erythrocytes is favored when ATP synthesis is needed, whereas this protein appears to catalyze exchange of one glucose molecule for another when flux through glycolysis is adequate. While allosteric regulation appears to account for the ability of the GLUT1 in human erythrocytes to switch between uniport and antiport, regulation of GLUT1 in pigeon erythrocytes may involve covalent modification of this or another protein by an AMPdependent protein kinase. This kinase activity is increased under conditions that lower the intracellular ATP concentration, which then favors glucose utilization in glycolysis and consequently glucose uptake via uniport. When ATP is not depleted, the pigeon GLUT1 protein appears to catalyze primarily exchange which would of course, result in no net transport of glucose in either direction (e.g., Cloherty et al., 1996). These studies with the glucose transporter stimulate several questions. First, what proportion of all uniporters, antiporters, and symporters are actually regulated by switching between coupled and uncoupled transport under different physiological conditions? Moreover, does the switch involving phosphorylation of the glucose transporter result in conversion of the transport protein molecule from one which catalyzes obligatory antiport to one which catalyzes obligatory uniport? Similarly, does allosteric regulation involve shifting of an equilibrium between a form of the transport protein that is an uniporter and a form that is an antiporter or can each form catalyze both coupled and uncoupled transport albeit in different proportions? Finally, could these regulatory processes involve conversion of the transport protein between smaller and larger oligomeric forms? For example, the human GLUT1 protein appears to exist in the plasma membrane as a tetramer (e.g., Herbert and Carruthers, 1992). What if exchange occurs in some cases because subunits of oligomers function together to catalyze migration of solutes simultaneously in opposite directions across the membrane, whereas monomers or smaller oligomers can catalyze migration in only one direction at a time? Interestingly, formation of GLUT1 tetramers from dimers appears to require intramolecular disulfide bonds (Herbert and Carruthers, 1992), whereas uniport activity of pigeon GLUT1 appears to require reduced intracellular glutathione (Cloherty et al., 1996). Reduced glutathione could conceivably reduce disulfide bonds and, thus, convert GLUT1 tetramers to dimers. In this model, GLUT1 dimers may function as uniporters, whereas tetramers may be antipor-

ters. In support of this interpretation, GLUT1 tetramers have at least two glucose binding sites, whereas dimers have only one population of such sites (Herbert and Carruthers, 1992). The reader should note that the mechanism proposed here for the simultaneous exchange or two glucose molecules is in contrast to the model described in Section XI,C above for exchange (Fig. 4.44A) in which glucose would be proposed to migrate across the membrane in only one direction at a time (see also similar considerations for anion exchange proteins in Chapter 6). More broadly, one wonders whether many differences in reported stoichiometries of symport and antiport may actually represent the ability of cells to regulate them. Some transporters could conceivably undergo such regulation without necessarily sometimes becoming uniporters. For example, careful measurements sometimes have shown a glutamate molecule (Glu-) apparently to be cotransported with 2 Na § ions (e.g.; Bouvier et al., 1992) and sometimes with 3 Na § ions (e.g., Zerangue and Kavanaugh, 1996) by members of the excitatory amino acid (glutamate) transporter (EAAT) family of proteins. Regulation of the number of Na § ions that are cotransported with a glutamate molecule could influence the rate and extent of Gluuptake by cells against its gradient to terminate signal transduction in neurons. 14 For example, initial cotransport of 2 Na § with 1 Glu- might produce more massive uptake of Glu- at higher extracellular Glu- concentrations since extracellular Na § would not be as difficult for Na§247 to replenish as when 3 Na § were cotransported. After this more rapid initial uptake of Glu-, however, it might be useful to switch to cotransport of each Glu- with 3 Na § in order to reduce extracellular Glu- to a concentration that would virtually completely terminate association of this neurotransmitter with its receptor. In a similar vein, the stoichiometry of proton transport by a V-type ATPase from the red beet plant appears to decrease as the pH gradient is increased, although the absolute pH values on either side of the membrane also influences the stoichiometry (Davies et al., 1994). In both of these cases, greater input of free energy is needed to catalyze transport against a gradient as the gradient grows larger. The structures and functions of the Glu-/Na § cotransporters and other symporters and antiporters are discussed more thoroughly in Chapter 6. Here we must complete our kinetic comparison of transport processes that form, propagate, or dissipate solute gradients. As for symporters and antiporters, substrate saturable uniporters surprisingly may also sometimes use a gradient 14See Section III,B,3 of Chapter 6 for other possible reasons for such differences in experimental data.

Kinetic Differences Among Transport Processes

123

of one solute to generate a gradient of another even when they function strictly for uniport and are not transstimulated. Unlike symport and antiport, however, there is an obligatory loss of the free energy in a total chemical potential gradient when uniport is used to convert the gradient of one solute into that of another regardless of whether the efficiency of the conversion is at its theoretical maximum. G. Dissipation of Solute Gradients via Uniporters As noted above, cis-inhibition of transport via uniporters may, under some conditions, be expected to produce concentration gradients of transported substances analogous to the gradients produced by antiporters. In contrast to the gradients produced by antiport, however, the gradients produced by uniport form during dissipation rather than during propagation of the free energy in solute gradients. That is, the maximum total chemical potential gradient that can be achieved through uniport is always lower than that achievable through antiport (see below). In addition, the formation of a new gradient through trans-stimulation of an antiporter is expected to be much more rapid than formation of a gradient by cis inhibition of an otherwise kinetically similar uniporter. Obligatory antiport could produce a theoretical maximum total chemical potential gradient of the substance moving against its gradient equal to the gradient of the substance moving with its gradient (e.g; Eq. (3.52) in Chapter 3). Hence, at steady state, the primary active transport catalyzed by Na+K+ATPase could form and maintain gradients of equal sizes of several substances through propagation of the Na § and K § gradients via obligatory antiport and symport. The amount of ATP that would need to be hydrolyzed by Na§247 in order to maintain the necessary Na § and K + gradients would, of course, increase as the number of solute gradients resulting from the Na § and K § gradients increased (Fig. 4.47). In contrast to obligatory antiporters, the solute gradients that can be produced through cis-inhibition of uniporters have theoretical maxima that are always less than the gradients of the substances producing the gradients (Fig. 4.48). Hence, the gradient must be dissipated to some degree as a result of uniport. The extent of degradation or dissipation of the gradients is directly related to the extent to which the Ki (or Km ) value of the substance producing cis-inhibition (and, thus, a gradient of another substance) exceeds the absolute value of its concentration gradient. Since the physiological concentrations of many solutes lie within an order of magnitude of their Km values for transport across the

FIGURE 4 . 4 7 Accumulation of an amino acid (aa-2) in cells against its total chemical potential gradient owing to obligatory exchange for another amino acid (aa-1). Amino acid-1 is assumed to be accumulated 10-fold against its gradient owing to the combined action of Na+K + ATPase (process 1) and a symporter for aa-1 and Na + (process 2). According to Eq. (3.52) (Chapter 3), aa-2 can be accumulated against its total chemical potential gradient via the antiporter (process 3) to the same extent that aa-1 is accumulated against its gradient. Amino acid-1 and aa-2 are assumed to be zwitterionic amino acids, so the membrane electrical potential difference has no effect on their total chemical potential gradient.

plasma membrane, most instances of cis inhibition are unlikely to produce much of a gradient of another substance in vivo. In cases where the Ki (and Km) values are particularly low relative to substrate concentrations, however, significant gradients could conceivably be produced through cis inhibition of uniporters. For example, let us assume that an amino acid uniporter has a Km (and Ki) value for uptake and exodus of amino acid-1 (aa-1) of 2 tzM 15 and that the intracellular and extracellular concentrations of aa-1 are 20 and 2/xM, respectively. Let us also assume that amino acid 2 (aa-2) competes for transport with aa-1 via the uniporter and that the sum of the intracellular and extracellular concentrations of aa-2 is 22/xM. To keep our example simple, we also assume that the amino acids are zwitterions so that membrane electrical potential does not influence their total chemical potential gradients. Under 15We assume symmetric operation of the uniporter to simplify our discussion. As discussed in Section IX,C above, we think that such symmetric operation of uniporters and channels may occur less frequently than their asymmetric operation. Moreover, since the strict uniporter under discussion does not exhibit trans-stimulation, its Km and Vmax values are assumed not to be influenced by whether the substrate and the inhibitor are also present on the other side of the membrane.

124

4. Transport Kinetics

FIGURE 4 . 4 8 Accumulation of an amino acid (aa-2) in cells against its total chemical potential gradient owing to cis-inhibition by another amino acid (aa-1). Amino acid-1 is accumulated against its gradient as described in the legend of Fig. 4.47. Cis-inhibition of the exodus of aa-2 by aa-1 is 5.5-fold greater than inhibition of aa-2 uptake by aa-1 at a Ki value of 2 /zM for aa-1 owing to the 10-fold greater concentration of aa-1 inside the cell (20/zM) than outside it (2/zM). That is, the apparent K,n value for transport of aa-2 is 5.5-fold higher in the presence of 20/zM of aa-1 than it is in the presence of 2/zM of aa-1 (Eq. (4.35)). Hence, the intracellular concentration of aa-2 will be 5.5-fold greater than its extracellular concentration at steady state (i.e., when the rates of aa-2 uptake and exodus are equal as can be calculated using Eq. (4.26) where the apparent Km value for exodus is 5.5-fold larger than the apparent Km value for uptake).

these conditions and assumptions, the degree of inhibition of aa-2 transport by aa-1 would be greater for exodus of aa-2 than for its uptake. If the initial concentrations of aa-2 were ll/zM on each side of the membrane, then uptake of aa-2 would proceed more rapidly than exodus until the intracellular and extracellular concentrations of aa-2 are producing equal rates of aa-2 transport in both directions. If it is assumed that the Km values are equal for aa-2 uptake and exodus via the uniporter, 15 then it can be calculated that the rate of aa-2 exodus and uptake would be equal when the intracellular and extracellular concentrations of aa-2 are 18.6 and 3.4 /zM, respectively (Fig. 4.48). This theoretical maximum gradient is somewhat below the theoretical maximum of 20 to 2/zM that could be achieved through obligatory exchange (Fig. 4.47). The gradient produced through cis-inhibition of a uniporter must, therefore, result in dissipation of solute gradients to some extent even if the new gradient could achieve its theoretical maximum.

Gradients produced through c/s-inhibition may, however, be more resistant to change as a result of perturbations in the gradient of the substance producing the gradient than are gradients produced through obligatory exchange. That is, although the rate of production of a gradient through c/s-inhibition may be slower than the rate of production of a gradient through transstimulation, the former may be more stable once established. Furthermore, the relative stability of gradients formed through c/s-inhibition are higher when the Km value for transport of the accumulated substance is higher. For example, the gradient of aa-2 produced as described above through c/s-inhibition would be degraded more slowly after loss of the gradient of aa-1 when the Km value for aa-2 is higher. Similarly, the rate of formation of the gradient of aa-2 owing to cisinhibition of transport by aa-1 would also be slower when the Km value for aa-2 is higher. It has not, to our knowledge, been determined whether gradients of amino acids actually form in vivo owing to c/s inhibition and, if they do form, whether they are important to the normal physiological functioning of cells, a6 In pathological conditions such as phenylketonuria (PKU), accumulation of phenylalanine extracellularly appears to inhibit uptake of other amino acids by brain, whereas acute or chronic phenylalanine accumulation in other tissues may competitively inhibit amino acid exodus from these cells (Christensen, 1990). This dual effect of sequestration of amino acids in other tissue and inhibition of brain amino acid uptake is proposed to contribute to the negative effects of PKU on brain development.

XII. SUMMARY We have discussed in this chapter several reasons why neither hydrophilic nor lipophilic solutes or the solvent are likely to migrate across biomembranes by ordinary diffusion. Nevertheless, diffusion is an important mechanism by which these ions and molecules may reach the membrane for transport. Moreover, concepts developed under the assumption that diffusion occurs across biomembranes can be used to help us to understand mechanisms of transport regulation. For example, we proposed based on such data that water channels may open in response to osmotic gradients across the plasma membrane (see Section III above). Although it remains controversial whether hydrophobic solutes cross biomembranes primarily via lipidor protein-mediated processes, there appears to be little doubt that protein-mediated processes catalyze most of 16Physiologically significant cases of such transport may, however, occur for the inwardly rectifying K + channels discussed in Section II,C Chapter 7.

Summary

the migration of hydrophilic solutes. Even the relatively rapid migration of water through the lipid bilayers of biomembranes is usually several orders of magnitude slower than the migration that occurs via water channels. Based on the observation that protein-mediated transport has been found always to be substrate saturable when such saturability has been tested at concentrations of substrate that are near physiological, we suggested that transport of water via channels also is probably saturable. Such saturability would, however, be difficult to demonstrate for water since it is impossible to raise its concentration much above its physiologically normal one. Furthermore, raising or lowering the concentration of water may cause water channels to open, thus making substrate saturation of the channels difficult to detect. When the kinetics of transport can be studied fully, they usually resemble the kinetics described originally by Michaelis and Menten for enzyme-catalyzed chemical changes. Nevertheless, the kinetic parameters Kin, Vmax, and Ki may have different meanings for enzyme and transporter catalysis. In the case of Ki, the values for enzyme inhibition frequently are equal to the values of the dissociation constants for binding because the competitive inhibitors usually are not also enzyme substrates. In contrast, competitive inhibitors of transport also usually are substrates, so their Ki values are equal to their Km values in these cases. Largely for this reason, measurement of Km and Ki values for solutes can be used to help to determine whether they share transport processes with other solutes whose transport they inhibit. Analog inhibition analysis can also be used to identify, isolate, and characterize transport processes when these processes might otherwise obscure each other's transport of the same substrate. (See the Appendix to this Chapter for a summary of the use of analog inhibition studies for this purpose.) Finally, we described kinetic differences among transport processes that form, propagate, and dissipate solute gradients. In some cases of formation of solute gradients in primary active transport where the stoichiometry of cation transport is greater than 1, the kinetics of transport differ from the simplest Michaelis-Menten formulation that is based on the transport of one ion at a time. Similarly, two or more different species of solute ions or molecules appear to be transported together across the membrane when symporters propagate the

| 25

solute gradients formed during primary active transport. In contrast, solute exchange in most instances of antiport is thought to occur through transport of one ion or molecule at a time, as in the ping-pong model shown in Fig. 4.44A (where k4 and k-4 are zero). Nevertheless, obligatory exchange rather than net transport in a single direction occurs because the transport protein is believed to reorient itself across the membrane only when it is associated with substrate. In the case of nonobligatory exchange, reorientation of the transporter is also more rapid when it is associated with substrate, but reorientation of the transport protein alone also occurs. In both cases solute gradients are propagated, although when exchange is not obligatory the gradient is also dissipated to some extent. The maximum dissipation of solute gradients occurs when reorientation of the transporter is not influenced by whether it is associated with substrate. The resultant uniport nevertheless is capable of forming new solute gradients at the expense of others through cis-inhibition. The ability of each of these types of transport processes to produce new total chemical potential gradients of solutes by kinetically different mechanisms probably relates to their similar fundamental structures, each of which provides a saturable pathway for substrate migration across a biomembrane. Since even the fastest of such pathways operate at less than one-tenth of the rate of diffusion, all such transport proteins must interact with their substrates to slow their migration. Such interactions permit coupling of the processes to sources of free energy, such as that realized from hydrolysis of ATP or from migration of another solute along its total chemical potential gradient. Even without direct coupling to such conspicuous sources of free energy, however, transport protein molecules are organized asymmetrically across biomembranes that are themselves also asymmetric. Since the transport proteins also function asymmetrically, the proteins have the potential to produce gradients of their substrates without additional input of free energy beyond the asymmetric organization and maintenance of the protein molecules in the membrane. The reliance of the functions of transport proteins on their underlying asymetric structures leads us naturally to detailed considerations of the structures of transport proteins that form (Chapter 5), propagate (Chapter 6), or dissipate (Chapter 7) solute gradients.

126

4. Transport Kinetics

APPENDIX Use of Analog Inhibition Studies and ABC Testing to Detect, Isolate and Characterize Multiple Components of Transport of Amino Acids and Other Solutes

What are analog inhibition studies and A B C testing?

They are methods of isolating transport system activities and defining their substrate selectivities.

What may be concluded from analog inhibition studies in which no cross inhibition is observed?

The analogs (e.g., amino acids) probably do not share a saturable transport system (AB part of the qualitative ABC test).

What may be concluded from studies in which even the unlabeled form o f the substrate is not observed to inhibit transport?

Transport is nonsaturable (or Km value for transport is considerably higher than 10 mM).

Appendix

| 27

What if cross inhibition o f saturable transport is complete?

Need to know whether inhibition by analogs is competitive or noncompetitive and whether one or more than one system is involved (see later).

Cross inhibition may also be incomplete. What may be concluded from such results?

Inhibition may be incomplete because it is weak or because it is partial.

How can it be determined whether incomplete inhibition is weak or partial?

Measure transport at various concentrations of the inhibitor.

128

4. Transport Kinetics

What may be concluded from studies showing that Arg inhibition is incomplete because it is weak?

"- -,

-

[3H] L-Ile uptake (1 lxM)

j

System(s) selects the substrate for transport over the inhibitor.

L-Arg

L-Ile

~ ~ ~ ~

I

I

I

I

i

I

5

10

15

20

25

30

Inhibitor concentration (mM)

What may be concluded from studies showing that Arg inhibition is incomplete because it is partial?

\

L-Arg [3H] L-Ile uptake (1 IxM) L-Ile

i

I

I

i

I

5

10

15

20

25

Inhibitor concentration (mM)

More than one transport system for L-Ile may be present.

Appendix How may one isolate and characterize the component o f transport that may receive both lie and Arg as substrates?

\

[3H] L-Ile uptake (1 ~M)

\/~

~

L-Arg

Possible shared component

,. L-Ile I

5

| 29

First; identify inhibitors of the unshared component(s).

[3H]L-Ile , . uptake (1 ~M)

~Arg [3H]L-Arg " ~ I 11 (1uptake ~M)

1; I

5 10 -Inhibitor concentration (mM)

.~-Ile i 12

~ 1'0 Inhibitor concentration (mM)

And then; perform transport experiments in the presence of the inhibitors. I

I

I

10 15 20 25 Inhibitor concentration (mM)

..Epta.keath!g.h.tI 0 . . . . . .

[3H]L-Ile ~i uptake L-Arg ] ~'~Lille

.Ept~ea!h!g.h.tI2) . . . .

[3H]L-Arghl uptake L-Arg ] ~_L-ile

5 10 1'5 Inhibitor concentration(mM)

5 10 1'5 Inhibitorconcentration (mM)

130

4. Transport Kinetics

Do we know from these studies that L-Ile and L-Arg compete for the same transport system(s)? Uptake at high [I1] ........................ [3H] L-Ile uptake I | . LAr - g

I ~(.r I "~7

Uptake at high [12] k,"........................ [3H] L-Arg uptake ]l-~ L-Arg ( L-Ile

~

5 10 1~ Inhibitor concentration (mM)

To show competition for transport, it should be determined both whether the mutual inhibition between LArg and L-Ile is competitive;

[ ~ -

- -1- ~ - , -

In the presence ofI 1

In the presence of 12

vi

vi _

_

5 10 15 Inhibitor concentration (mM)

_

vi

_

_

[L-Ile]

vi _

[L-Arg]

And whether each amino acid has the same characteristics as a substrate as it has as an inhibitor (AB part of the quantitative ABC test). ( AB part ) Substrates Substrate (A) L-Ile (B) L-Arg

Km

Ki

Value ? Value Value ? Value

(C part ) Additional Amino Acid Ki(C1)

Ki(C2)

Value

Value

I17

Value

I17

Value

To further test whether L-Ile and L-Arg share the same system, determine whether other amino acid analogs (e.g., C1, C2, etc.) influence transport of each substrate in the same way (C part of the ABC test).

Appendix

| 31

Summary I. Mutual inhibition between two amino acid analogs may be; A. Absent (analogs do not share systems; AB part of the qualitative ABC test). B. Complete (analogs may share all systems). C. Incomplete. II. Inhibition may be incomplete because it is; A. Weak (system(s) selects substrate over inhibitor). B. Partial (more than one system may be present).

IV. To characterize the component of transport isolated in this manner one should determine whether; A. The two analogs of interest inhibit each others transport competitively. B. The two analogs behave as substrates in the same way as they behave as inhibitors (AB part of the quantitative ABC test). C. Other analogs influence transport of both substrates in the same way (C part of the ABC test).

III. To isolate and characterize the transport component of interest (e.g., a component that appears to be shared by two analogs) one can; A. Identify inhibitors of the other components of transport of analog A. B. Identify inhibitors of the other components of transport of analog B. C. Perform experiments in the presence of inhibitors of other components of transport. V. Study of the transport of more than two analogs results eventually in; A. Full assessment of the substrate selectivity of the isolated system or; B. Detection of unanticipated heterogeneity within what had appeared to be a single component of transport.

This Page Intentionally Left Blank

C

H

A

P

T

5]

E

R

Structure and Function of Transport Proteins That Form Solute Gradients

I. INTRODUCTION

Vacuolar (V)-type ATPases usually concentrate protons against their gradients in intracellular membranebound compartments such as endosomes and lysosomes (Fig. 3.12, Chapter 3). V-type ATPases are, however, incapable of the reverse process (i.e., coupling of ATP synthesis to transport of protons along their gradient) apparently because some transport occurs in the reverse direction without coupling to chemical change (Nelson, 1992b). In contrast, the normal physiological function of most eukaryotic energy coupling Factor (F)-type ATPases is to couple the transport of three or, more likely, four protons to the synthesis of one molecule of ATbp3P from ADP and Pi (Fillingame, 1997). Another difference between F- and V-type ATPases is that the latter transport only two protons per hydrolyzed ATP (Nelson, 1995) although this stoichiometry p e r se cannot account for the inability of V-type ATPases to catalyze the reverse reaction (see also Section III,A below). In spite of these differences, F- and V-type ATPases appear to function by the same general mechanism. For this reason and to conserve space, our detailed discussion of the structure and function of F-type ATPases in Section III below will be considered also to represent V-type ATPases. While virtually all F- and V-type ATPases, transport protons (Nelson, 1992b), various P-type ATPases transport a wide variety of different inorganic cations (MOiler et al., 1996). P-type ATPases are so named because one of their aspartyl residues is temporarily phosphorylated during the transport cycle, whereas no such phosphorylated intermediate has been detected for F- and V-type ATPases. P-type ATPases also have lower molecular masses and much less

Formation of solute gradients may be driven by several sources of free energy including (but not limited to) oxidation-reduction reactions, electromagnetic radiation, and ATP hydrolysis. In this Chapter we consider only the transport ATPases in detail. Solute gradients themselves may also be used to form new gradients, but in these cases the transport proteins propagate solute gradients rather than form them. In addition, the symporters, antiporters, and even uniporters that propagate or dissipate solute gradients may have an inherent ability to form solute gradients from some of the free energy needed to synthesize and maintain them asymmetrically in biomembranes (see Chapter 4). Since most transport protein molecules appear to undergo conformational changes during transport, the differences in the free energies of the different conformations may be the amount of free energy available to form solute gradients. When, however, such conformational changes in transport proteins are also coupled to a conspicuous source of free energy such as that in ATP, they are able to produce the steepest solute gradients. Transport ATPases form several categories (summarized in Chapter 8), and the three categories to be discussed here are termed P-, F-, and V-type ATPases. The F- and V-type ATPases share the same multimeric structure and a common evolutionary origin. Consequently, we consider them here to be subcategories of a single F-/V-type ATPase. Nevertheless, V- and Ftype ATPases perform physiologically different functions.

133

134

5. Structure and Function of Transport Proteins That Form Solute Gradients

extensive oligomeric structures than F- and V-type ATPases. Although P-type and F-/V-type ATPases appear to have evolved from unrelated ancestors, it is nevertheless useful to consider how their structures and functions are similar. The structural and functional differences between P-type and F-/V-type ATPases will become clear from their detailed descriptions later in this Chapter. A. Transport and Chemical Change Occur in the M e m b r a n e and Extramembrane Components, Respectively, of Both P- and F-/V-Type ATPases The F-/V-type ATPases have distinct membrane and extramembrane sectors that are each composed of multiple subunits (Fig. 3.10, Chapter 3). The two sectors function together to catalyze proton transport in the membrane (F0/V0) sector and ATP hydrolysis or (in the case of F-type ATPases) synthesis in the extramembrane (F1/V1) sector. Numerous impressive advances have been made over the past four decades in understanding the details of the mechanisms by which F-type ATPases contribute to photosynthesis and oxidative phosphorylation (e.g., see Section III below). Nevertheless, we are still learning how F-type ATPases transfer free energy between solute gradients and phosphoric acid anhydride bonds. Similarly, it is still largely a mystery how P-type ATPases couple transport to chemical change. But like the F-/V-type ATPases, P-type ATPases clearly have membrane and extramembrane components that function together to catalyze ion transport and ATP hydrolysis (or synthesis), respectively (Toyoshima et al., 1993) (Fig. 5.1). Moreover, both P-type and F-/V-type ATPases undergo conformational changes that appear to be involved in the bioenergetics of coupling of transport to chemical change. We describe much of what is known about these physical changes in Sections II and III below. While the details of the processes are clearly different for the two types of ATPases, it is interesting that some of the fundamental mechanisms underlying the transfer of free energy are the same. B. P- and F-/V-Type ATPases Appear to Function in a Fundamentally Similar Manner Both P- and F-/V-type ATPases couple inorganic ion transport through biomembranes to ATP synthesis or hydrolysis several nanometers away (e.g., Toyoshima et al., 1993; Fillingame, 1997). The spatial separation of these two processes may seem at first unnecessary since chemical catalysis appears in this case also to require a

FIGURE 5.1 Proposed three-dimensional structure of the Type II P-type ATPase molecule. The sarco(endo)plasmic reticulum Ca 2+ ATPase molecule shown consists of membrane and cytosolic components that are the sites of transport and ATP hydrolysis (or synthesis) respectively. The proposed positions of transmembrane segments 2 to 8 (M2 to M8) and the extracellular loop between M7 and M8 are also shown (adapted from Toyoshima et al., 1993, with permission from Macmillan Magazines Ltd.).

hydrophobic environment resembling that of the lipid bilayer. Indeed, since most of the membrane electrical potential appears to lie across the outer half of the membrane phospholipid bilayer (Stiarmer et al., 1989; Stein, 1990), the negatively charged nucleotides might not even need to enter the electric field to reach a hydrophobic environment. Nevertheless, neither P- nor F-/V-type ATPases evolved in a way that allowed the lipid bilayer to be exploited as the hydrophobic environment serving for ATP synthesis or hydrolysis. This result may be coincidental or perhaps using the membrane lipid bilayer as the site for chemical catalysis would disrupt the critical function of the bilayer as a barrier to the migration of hydrophilic substances. The free energy changes associated with ATP synthesis or hydrolysis at some distance from the membrane, on one hand, and ion transport across it, on the other, appear to be coupled through conformational changes in the various components of F-/V- and P-type ATPases. ATP hydrolysis results in conformational changes in the extramembrane components of both types of enzymes, which are transmitted to the membrane components to produce inorganic ion transport against a total chemical potential gradient (MOiler et al., 1996; Fillingame, 1997). Conversely, transport of the ions along their gradients produces conformational changes in the membrane

| 35

P-Type ATPases

components that, when transmitted to the extramembrane components, result in A T P synthesis by both Fand P-type ATPases. Both types of enzymes are also oriented in regard to the ion gradients they produce or utilize so that the higher total chemical potentials of the ions are on the side of the m e m b r a n e opposite to the extramembrane components. In spite of these similarities it is tempting to conclude that the catalytic mechanisms of P-type and F-/V-type ATPases are fundamentally different because the former are temporarily bound covalently to phosphate during their transport cycle while the latter bind phosphate noncovalently. Nevertheless, similar mechanisms appear to be used to accomplish both types of binding. For example, the positively charged amino group of an essential lysyl residue in F-type ATPases appears to be required for pi2-/pi 1- binding near A D P 3- during A T P synthesis (Weber and Senior, 1997). The amino group probably forms hydrogen bonds with the phosphate and thus contributes significantly to its free energy of binding. Similarly, an indispensable lysyl residue lies near the aspartyl residue to be phosphorylated in the P-type sarco(endo)plasmic reticulum Ca2+ATPase. In this case, the positively charged amino group on the lysyl residue appears to be part of a stereospecific acid-base catalyzed transfer of phosphate from A T P 4- to the negatively charged carboxyl group of the aspartyl residue (MOiler et al., 1996). Hence for both F- and P-type ATPases, part of the function of positively charged amino groups of essential lysyl residues appears to be to help to appose negatively charged substrates. The reader will undoubtedly notice additional similarities as well as differences in the mechanisms of bioenergetic coupling used by P- and F-/V-type ATPases, as the known details of their structures and functions are described in Sections II and III below.

Type II

IIA

~ /

~~cNscNANKA IHKA o0ern roao,es

Heavy Metal ATPases (Prokaryotes & Eukaryotes)

FIGURE 5.2 Proposed evolution of P-type ATPases. Evolution of Type I and Type II P-type ATPases from a common ancestor appears to have occurred in ancient prokaryotes. Subsequently, the Type II ATPases evolved into three types including the Type IIA and Type IIB ATPases of eukaryotes and the Type II ATPases of modern procaryotes. Further diversification of Type IIA ATPases led to the evolution of the H+K+ATPases (HKA), Na+K+ATPases (NKA), and sarco(endo)plasmic reticulum (SERCA) Ca2+ATPases(SCA). Diversification in the Type IIB line led to the evolution of the plasma membrane Caz+ ATPases (PMCA) and the P-type proton ATPases et al., (HA). See text for further discussion (adapted from Mr 1996, with permission from Elsevier Science).

II. P-TYPE ATPases A. Evolution a n d Classification of P-Type ATPases P-Type ATPases are divided into two classes that evolved from a common ancestor prior to the appearance of eukaryotes (Fig. 5.2). The most conspicuous difference between the two classes is in their N- and Cterminal membrane-associated domains (MOiler et al., 1996). The N-terminal domain is larger in Type I than in Type II P-type ATPases, whereas the reverse relationship applies to the C-terminal membrane-associated domain (Fig. 5.3). The differences in the sizes of these domains means that large sequences of amino acid residues must have been added to or lost from the N- and C-termini of P-type ATPases during evolution. These

FIGURE 5.3 Proposed structure of Type I(A) and Type II(B) Ptype ATPases. Type I ATPases are thought to have two putative transmembrane a-helices in their N-terminal membrane-associated domain that are not present in Type II ATPases, whereas Type II ATPases appear to have four such transmembrane a-helices in their C-terminal membrane-associated domain that are not present in Type I ATPases. Other components are described more thoroughly in the text and in other figures (adapted from MOileret al., 1996,with permission from Elsevier Science).

136

5. Structure and Function of Transport Proteins That Form Solute Gradients

evolutionary changes may have occurred through intermediate forms of the ATPases in which subunits were products of separate genes and hence not joined by peptide bonds. For example, the Kdp protein complex of E. coli (KdpK+ATPase) is composed of three subunits, each of which is required for its transport activity (e.g., Buurman et al., 1995). The largest of the subunits, KdpB, catalyzes ATP hydrolysis and it is homologous in amino acid residue sequence to other P-type ATPases. The other two subunits, and in particular the highly hydrophobic KdpA, form the pathway through which K + ions are transported. In fact, the K d p A subunit has features homologous to K+-channel proteins (Mr et al., 1996). Hence, the existence of the Kdp protein complex supports the notion that proteins capable of primary active transport may evolve through the association of proteins that catalyze chemical change with proteins that catalyze transport (e.g., Nikaidao and Saier, 1992). Except for a short sequence of 10 amino acid residues, the KdpA subunit is not homologous to either the N- or C-terminal membrane-associated domains of other P-type ATPases (Mr et al., 1996). Therefore, it appears from their structures that other P-type ATPases evolved through the combination of a catalytic subunit similar to KdpB with another channel-forming protein molecule. Combination of the genes encoding all of the subunits into one larger gene might have produced the ancestor to modern P-type ATPase genes. Alternatively, the KdpK+ATPase may represent evolution of an oligomeric structure from P-type ATPases through combination of a P-type ATPase molecule with another transport protein molecule. During evolution of a combined function of these two transport proteins, the membraneassociated domains of the P-type ATPase may have been lost. In the latter case, the genes encoding the extramembrane and membrane components are free to evolve separately. Such separate evolution of membrane and extramembrane components has occurred much more extensively among F-/V-type ATPases than among P-type ATPases (see Section III below). After evolution of Type II P-type ATPases apparently from an ancestral Type I ATPase, two subtypes of Type II ATPases evolved (Fig. 5.2). Type liB ATPases have larger C-terminal hydrophilic tails than do Type IIA ATPases (Mr et al., 1996). These larger tails permit regulation of Type IIB ATPases by calmodulin and other such effectors. Further diversification of Type IIA and Type IIB ATPases led to evolution of Na+K +-, H+K +-, K+H +-, and sarco(endo)plasmic reticulum Ca2+(SERCA) ATPases in the Type IIA line and plasma membrane Ca2+(PMCA) - and H+ATPases in the Type liB line (Fig. 5.2). The Type IIA line is more conspicuous in animals, whereas the Type IIB line is

more conspicuous in plants; only the two types of Ca2+ATPases (i.e., PMCA and SERCA ATPases) are found in both plants (or fungi) and animals. In addition to variation in substrate selectivity, the Type IIA ATPases have undergone interesting structural diversification. Both the Na+K+ATPases and the H+K+ATPases are heterodimers containing the catalytic a-subunit and a /3-subunit, the full function of which is still under investigation. The a-subunit of H+K+ATPase probably evolved through duplication of one of at least three genes encoding c~ subunits of Na+K+ATPase (Maeda, 1994). Each a-subunit of Na+K+ATPase may be associated with one of two isoforms of the/3-subunit (Chow and Forte, 1995), thus forming a total of at least six possible enzyme dimers. Alternative splicing is not known to occur in RNA transcripts encoding either subunit of Na+K § and H+K+ATP ases, but it does occur in transcripts encoding SERCA ATPases (see Section II,B,4 below). This splicing produces various subisoforms of the three known isoforms of SERCA ATPases. Since the SERCA ATPases do not form heterodimers, one wonders whether evolution of Na+K+ATPase heterodimers accomplished an end similar to that accomplished among SERCA ATPases by evolution of alternative splicing. For example, alternative splicing and a heterodimeric structure may both result in formation of a variety of different forms of the same enzyme. Somewhat different forms of enzymes appear to help different tissues meet their unique metabolic needs. As for the Type IIA SERCA ATPases, alternative splicing also produces subisoforms of Type IIB PMCA ATPases (see below). Moreover, the activities of the PMCA ATPases may be regulated not only by calmodulin, but also by nonrandom formation of homooligomers. In contrast, no such nonrandom homooligomerization has been described for Type IIA monomers (SERCA ATPases). Their a-subunits do not have the extended C-terminal tail apparently required for regulation both by calmodulin and by nonrandom oligomerization (Mr et al., 1996). Nevertheless, recent studies indicate that the C-terminal tails of Type IIA ATPases help to regulate their activities through K § binding (e.g., Ishii et al., 1997), and Na+K+ATPase appears to form (O/~) 2 t e t r a m e r e s (Linnertz et al., 1998). Moreover, SERCA ATPases are frequently so concentrated in the sarcoplasmic reticulum that they function as dimers or higher oligomers apparently formed through random association (but see also Section II,C,4 below). While P-type ATPases are classified conveniently according to all of these broad structural and functional differences, the similarities of the structures of these ATPases are also striking. Differences in structure account for differences among the ATPases in the speci-

P-Type ATPases

ficities of their transport and its regulation, but these differences are accomplished through relatively small variations on a fundamental structural theme that is the et al., 1996). Let us same for virtually all of them (Mr turn now to a more detailed description of the structural components of P-type ATPases that underlie the common mechanism by which they function. Since Type I and Type II P-type ATPases are structurally and functionally similar and because our space is limited, we focus here primarily on the Type II ATPases. B. Structure of Type II P-Type ATPases As for F-/V-type ATPases, P-type ATPases have membrane and extramembrane sectors where transport and chemical change, respectively, are catalyzed (Fig. 5.1). Unlike the F-/V-type ATPases, however, these sectors are part of the same polypeptide chain in P-type ATPases (Toyoshima et aL, 1993). The cytosolic sectors of the Type II P-type ATPases are composed of a large domain C, a region B, and a "hinge" region (Fig. 5.4). Also present in the cytosol are five "stalk" segments ($1 to $5) that emanate from the membrane-spanning helices of the N- and C-terminal membrane-associated domains (Mr et aL, 1996). Both the N- and Cterminal membrane-associated domains appear to contribute components to the pathway through which inorganic ions migrate across the membrane. Moreover, the cytosolic tails at the N- and C-termini of the enzyme may be involved in its regulation depending on the particular

FIGURE 5.4 Proposed structure of Type II ATPases including conserved amino acid residue sequence motifs. The specific motifs shown are for Na+K+ATPase. Also shown are the proposed 10 transmembrane a-helices (M1 to M10) and 5 a-helices (S1 to $5) that form the stalk in the three-dimensional model presented in Fig. 5.1. The sequence motifs and other components of the enzyme are discussed in the text. Key: NT domain, N-terminal membrane-associated domain; CT domain, C-terminal membrane-associated domain (Adapted from MOiler et al., 1996, with permission from Elsevier Science).

137

ATPase under consideration. Conserved amino acid residue sequence motifs in various portions of the enzyme molecule are also important to its function (Fig. 5.4). Let us now consider each of these components of Type II P-type ATPases in greater detail. Unless indicated otherwise, much of the information in the first three subSections below is presented in a review by Mr and associates (1996). 1. The N-Terminal Membrane-Associated Domain

A consensus model for Type II P-type ATPases indicates that these proteins have four transmembrane a-helices in the N-terminal membrane-associated domain (Fig. 5.4). Each of these transmembrane a-helices is associated with a cytosolic "S" sequence. These S sequences are so named because they are believed to comprise most of the "stalk" region of the proteins (Figs. 5.1 and 5.4). The four sequences, $1 to $4, will be discussed in conjunction with our discussion of the four transmembrane a-helices, M1 to M4, of the Nterminal membrane-associated domain. The first transmembrane a-helix penetrates the lipid bilayer from the cytosolic side and, hence, it is preceded by $1. While M1 has the anticipated low abundance of hydrophilic amino acid residues, $1 has an amphipathic nature. For example, highly hydrophilic amino acid residues tend to be segregated along one side of the $1 sequence of the rabbit la SERCA ATPase when the $1 sequence is arranged in its predicted a-helix conformation (Fig. 5.5). Presumably, associations between the more hydrophobic regions of the five stalk sequences (including one from the C-terminal membraneassociated domain, Fig. 5.4) help to stabilize their three-dimensional conformation. The C-terminal end of M1 is followed by a relatively short, hydrophilic extrasystolic loop from which the hydrophobic M2 emanates. M2 is followed on its cytosolic side by the second amphipathic stalk sequence $2. Between $2 and $3 is the cytosolic region B (see Section II,B,2 below). $2, $3 and region B together comprise the smaller of the two relatively large cytosolic looPs (Fig. 5.4). Like M1 and M2, the helix M3 contains mostly hydrophobic amino acid residues with a preponderance of aliphatic side chains. As Mr and associates point out (1996), however, M3 is estimated to be at least 22 residues in length, unlike M1 and M2, which may be too short to span the membrane as standard a-helices. As for $1 and $2, the cytosolic $3 sequence that precedes M3 has a relatively large number of hydrophilic amino acid residues that are predicted to be in a helical conformation, but unlike, $1 and $2, $3 does not display an amphipathic pattern.

138

5. Structure and Function of Transport Proteins That Form Solute Gradients

2. Cytosolic Regions and Domains

FIGURE 5.5 Helical wheel showing the locations of amino acid residue side chains around the S1 a-helix of the stalk region of rabbit la sarco(endo)plasmic reticulum Ca2+ATPase. Amino acid residues 40-58 of the protein are shown. Note in particular that six of the eight residues that have highly hydrophilic side-chains (shown as white on black) lie along one side of the S1 a-helix and that the other two highly hydrophilic residues also lie along one section of the helix. Some more recent models indicate that $1 may not actually be part of the stalk (e.g., Zhang, et al., 1998b). Abbreviations: A, alanine; E, glutamate; F, phenylalanine; G, glycine; I, isoleucine; K, lysine; L, leucine; P, proline; Q, glutamine; S, serine; V, valine; W, tryptophan.

Following another relatively short extrasystolic loop emanating from M3, the highly hydrophobic M4 sequence traverses the membrane. The C-terminal end of M4 is less precisely defined than in the cases of M1 to M3 since the $4 sequence is also highly hydrophobic. For this reason, the $4 a-helix is believed to be buried inside the stalk region of Type II ATPases. Both $4 and M4 may be involved in free energy transduction, as will be discussed further in Section II,C below. Moreover, the site of transient enzyme phosphorylation is only about 22 residues beyond $4, not only in all Type II ATPases, but also in homologous regions of Type I ATPases and even in a corresponding region of the Bsubunit of the Kdp protein complex (KdpK+ATPase). The amino acid residue sequence near the site of phosphorylation as well as the sequence of $4 are highly conserved among P-type ATPases. The residues form an interesting pattern of alternating hydrophobic and hydrophilic side chains following $4, and this region forms the beginning of the large cytosolic loop.

By far the most conspicuous of the cytosolic regions is the large cytosolic loop between M4 and M5, which has considerable secondary and tertiary structure. Consequently, the major portion of it is appropriately termed a domain (i.e., domain C; Fig. 5.4). Since ATP is invariably the substrate for chemical change in the extramembrane components of P-type ATPases, it is somewhat surprising that these components have widely variable lengths and amino acid residue compositions. In contrast, the membrane-associated domains are much less variable among P-type ATPases, although they receive a wide variety of inorganic ions as substrates. Apparently the conserved sequence motifs (Fig. 5.4) and other structurally similar Sections are enough to confer the same catalytic function on the extramembrane components of all P-type ATPases. For example, domain C appears to contain an alternating/3-a-/~ structure in all P-type ATPases, and this structure is predicted to form an essential tertiary organization with the ATP binding site at the C-terminal edge of the parallel/~-sheet structure (Branden and Tooze, 1991). While this tertiary structure may help to appose the ATP-binding site and the phosphoryl-accepting aspartyl residue, several other residues that are located in specific sequence motifs are needed for normal catalysis. For example, the phosphorylation site contains both the aspartyl (D) residue to be phosphorylated and a neighboring lysyl (K) residue (Fig. 5.4) that is essential for phosphorylation in SERCA ATPases (Maruyama and MaLennan, 1988). None of the other residues in the sequence motif surrounding the phosphorylation site (ICSDKTGTLT) is essential for phosphorylation of SERCA ATPases since conservative substitutions do not prevent phosphorylation. Nevertheless, semiconservative substitution of these residues greatly disrupts Ca 2+ transport. Mr and associates (1996) suggested that these neighboring residues may be needed to maintain the conformation of the moderately hydrophobic crevice, which may in turn be needed for phosphorylation. As anticipated, substitution of an asparagyl residue for the aspartyl residue at the phosphorylation site of the yeast H+ATPase destroys its normal function (Portillo et al., 1995; Harris et al., 1994). Interestingly, however, the latter mutation does not impair growth (Rao and Slayman, 1993), apparently because redundant mechanisms are sufficient for nutrient uptake when H+-dependent glucose transport is no longer driven by the H + gradient normally formed and maintained by H+ATPase in yeast. SERCA and other P-type ATPases also contain many other residues that are required for autophosphorylation. In the DPPR motif of SERCA ATPases (Fig.

P~Type ATPases

5.4) both the aspartyl residue and the second prolyl residue are needed. Similarly, both the glycyl and aspartyl residues of the TGD motif are required for normal phosphorylation and C a 2+ transport. The K G A P E motif (Fig. 5.4) appears to be close to the site of ATP binding and hydrolysis since chemical modification of the lysyl residue in this motif abolishes catalysis in SERCA and other ATPases. Nevertheless, site-directed mutagenesis of this lysyl residue of SERCA ATPases does not greatly alter C a 2+ transport unless the residue is converted to a very different one such as a glutamyl residue. Moreover, while substitution of an arginyl residue for this lysyl residue in the H+ATPase of yeast reduces H + transport by 90%, this decrease can be partially reversed by compensatory second-site mutations of other residues in cytosolic regions (Maldonado and Portillo, 1995). SERCA ATPases also need two additional, somewhat isolated, lysyl residues for normal ATPase and transport activities. The first conserved lysyl residue is located about 20 amino acid residues N-terminal to the KGAPE motif (Fig. 5.4), and it is associated with a conserved phenylalanyl residue 5 amino acid residues N-terminal to it (i.e., FXXXXK). The second residue is located 14 residues N-terminal to the "hinge" motif (Fig. 5.4) in both Type II and Type I P-type ATPases. The latter lysyl residue is associated with a conserved prolyl residue 4 amino acid residues N-terminal to it (i.e., PXXXK). Both the lysyl residue within the KGAPE motif and the one about 20 residues N-terminal to it can be chemically modified by several aminoreactive reagents. Hence, these lysyl residues are probably present at exposed positions near the surface of the ATPases. While domain C is the major component of the head region of P-type ATPases shown in Fig. 5.1, region B and the junctional region J (Fig. 5.4) may also project into the head. This globular head of the ATPases appears to extend about 6.0 nm from the cytosolic surface of the plasma membrane (Tohoshima et al., 1993). Region J is the sequence of amino acid residues between the end of domain C and M5. Hence, domain C and region J together comprise nearly all of the large cytosolic loop between the last transmembrane c~-helix of the N-terminal membrane-associated domain and the first helix of the C-terminal membrane-associated domain (Fig. 5.4). Region J contains both the hinge region present in all P-type ATPases and the fifth stalk sector, $5 (Figs. 5.1 and 5.4). It is the most conserved region of the polypeptide chain of P-type ATPases, as pointed out by Mr and associates (1996), probably because it is present in close proximity both to the ATP-binding region of these enzymes and the intramembrane ionbinding sites needed for transport. Region J is predicted to have a stiff helical structure that could conceivably

139

help to transmit small but well-defined conformational changes between domain C and the ion translocation sites. Only one other cytosolic loop contributes significantly to the cytosolic component of P-type ATPases. The loop is termed (or mistermed) the small cytosolic loop because it is the smaller of the two largest loops. The major portion of this loop is also known as region B (Fig. 5.4). Region B is located between $2 and $3 of the N-terminal membrane-associated domain, and it contains from about 65 to 175 amino acid residues depending on the ATPase. The region is particularly variable in length among plasma membrane C a 2+ ATPases as a result of alternative splicing (Section II,B,4 below). It is also likely to be the site of anionic phospholipid binding, and these phospholipids substantially increase the activity of plasma membrane C a 2+ ATPase (Penniston and Enyedi, 1994). Region B may have an antiparallel B-sheet conformation in most P-type ATPases, and an important although yet undefined role in energy transduction has been attributed to it. It appears to be located strategically between the stalk and head regions of P-type ATPases in their normal conformations, and hence it may help to link the catalytic sites for transport and chemical change. Region B may help to form this link by stabilizing the structures of $4 and $5, which are positioned to conduct conformational changes between domain C and the membraneassociated domains. 3. Putative Structure of the C-Terminal Membrane-Associated Domain

A well-defined 10-helix model has been developed for the membrane-associated N- and C-terminal domains of Type II P-type ATPases (Fig. 5.4). As for all such models, however, it must be remembered that the model will need to be modified as new data are obtained. Particularly in regard to the C-terminal membrane-associated domain, much uncertainty remains concerning the precise number and locations of its transmembrane sequences (Mr et aL, 1996). This domain has a very high content of hydrophobic amino acid residues with only short extramembrane loops (Fig. 5.4). Short loops are more resistant to protease action than longer ones, which makes the topologies deduced from hydropathic plots difficult to verify experimentally. Nevertheless, available evidence supports the conclusion that the Cterminal membrane associated domain contains six membrane traverses, the first two of which we will now see exemplify the difficulties encountered in establishing the structure of the C-terminal membrane-associated domain. The M5 and M6 transmembrane segments were pre-

| 40

5. Structure and Function of Transport Proteins That Form Solute Gradients

M5 NKA: - - S N I PESCA: - - S N V G E -

Turn

M6

PLP PEA

T V T ILC I D L G T D P.V,QL L W V N L V T D -

r-~HLRP

E KA E sDIM

~ lT~ BindingVestibule M5"--I~t~ ["~ "E~iFL(~822~---M6 -

Cytosoi

Membrane

I 813 FIGURE 5.6 Aminoacid residue sequence of the fifth and sixth putative transmembrane segments of Type IIA P-type ATPases (M5 and M6). The complete sequence shown is for a portion of the gastric H+K+ATPase. Note that the segments are barely long enough to traverse the membrane as a-helices in part because these helices are believed to form a cavity around bound cations (binding vestibule). Moreover, prolyl (P) residues (enclosed in diamonds) may form kinks that are conformationallymobile owing apparently to cis-trans isomerization. These conformational changes appear to contribute to cation translocation. Abbreviations: HKA, H+K+ATPase;NKA, Na+K+ATPase;SCA, sarco(endo)plasmicreticulum Ca2+ (SERCA) ATPase (adapted from Mr et al., 1996, with permission from Elsevier Science)

viously believed to comprise only one transmembrane segment. This conclusion was based primarily on immunological studies on the locations of particular peptidyl sequences on the cytosolic or noncytosolic sides of the membrane. Subsequent studies using proteolysis indicate, however, that this portion of the protein contains two transmembrane segments. In fact, it is believed that M5 and M6 may help to form cavities in which cations are bound for subsequent transport. Partly for this reason, the two segments appear to be just long enough to span the m e m b r a n e (Fig. 5.6). Moreover, the two segments appear to undergo conformation-dependent movements that are probably important to cation transport (Lutsenko et al., 1995; Mr et al., 1996; Beguin et al., 1998). Especially interesting are kinks formed by prolyl residues (Fig. 5.6) which may be subject to c i s - t r a n s isomerization. In such a case, the resultant conformationally mobile kinks could contribute to ion translocation (Brandl and Deber, 1986). As for M5 and M6, the proposed structure of the remainder of the C-terminal membrane-associated domain of the Type II ATPases (Fig. 5.4) is also provisional. While the preponderance of evidence seems to favor the presence of four more membrane traverses in

these ATPases (termed M7 through M10), it can be argued from some experimental data that M8 may not be within the lipid bilayer. Moreover, the proposed relatively large extracellular loop between M7 and M8 contains a sequence of nine hydrophobic amino acid residues following a putative turn that could conceivably traverse the membrane in a/3 conformation at least in Na+K § and H+K§ An alternative explanation for the conflicting data concerning the precise number and arrangement of m e m b r a n e traverses could be that the traverses do not all remain permanently within the lipid bilayer. We propose that some complete traverses may migrate into and out of the m e m b r a n e to regulate transport activity or as part of the transport cycle. 1 For example, proteolytic digestion of Na+K+ATPase results in release of a peptide containing M8 (Madyanov et al., 1992; JCrgensen, 1992; Shainskaya and Karlish, 1994). Moreover, some evidence favors the cytosolic location of the C-terminal portion of the relatively large putative extra1While crystallographers in general appear to feel that only relatively small conformational changes occur in proteins, it is also worth noting that only small changes may be possible for proteins in crystals, which is, of course, not a natural environment for most of them.

P~Type ATPases

systolic loop between M7 and M8, whereas other evidence favors the conventional conformation. Hence, the nine hydrophobic amino acid residues in this "loop" (Fig. 5.4) may migrate temporarily into the membrane as a/~-traverse while M8 moves out during inorganic ion transport by some P-type ATPases. While it is unconventional in the case of P-type ATPases to propose that some transmembrane segments enter or leave the membrane to perform or regulate transport, it is known that such segments of other transport proteins enter the membrane to produce their actively transporting forms. For example, the number of membrane traverses changes from four to six during trafficking of aquaporin-1 monomers from the endoplasmic reticulum to the plasma membrane, and one membrane traverse appears to reverse its orientation during such trafficking (Verkman et aL, 1996). Moreover, volume-sensitive anion channel proteins appear to be inserted spontaneously into the plasma membrane in response to cell swelling. They also appear to leave the membrane when no longer needed for transport (Strange et aL, 1996). Similarly, highly hydrophobic hairpin turns between helices in the channel-forming domains of the bacterial toxins, colicins, appear to insert themselves spontaneously into lipid bilayers (reviewed by Sansom, 1997). Such a hairpin appears to exist in some P-type ATPases between the transmembrane helices M5 and M6 (Fig. 5.6). Hence, both M5 and M6 could conceivably enter or leave the membrane rather than simply move somewhat within it. In regard to the possibility that M8 might enter and leave the membrane, research has been conducted in an attempt to understand the mechanism of insertion of such amphipathic amino acid residue sequences into membranes (e.g., Shai, 1995). Since insertion of such sequences appears to be facilitated by the presence of other helices in the membrane (Shai, 1995; Sansom, 1997), the other membrane-spanning segments of P-type ATPase protein molecules should facilitate insertion of M8. We consider further in Section II,C below the relationship of the possible structures and conformations of P-type ATPases to their functions. To complete our discussion of the structures of Type II ATPases, however, it is necessary to consider not only that different isoforms exist for transport of the same ions, but also that different subisoforms of some P-type ATPases are produced through alternative splicing of the primary transcripts that encode them. A wide variety of different P-type ATPases as well as their isoforms were compared in order to establish the conventional consensus structure described above. For this reason, we will focus now only on those P-type ATPases for which subisoforms are known to be produced through alternative splicing. These ATPases include the CaZ+ATPases of the plasma

141

membrane (PMCA ATPases) (Carafoli, 1992) and sarcoplasmic reticulum (SERCA ATPases) (Mr et aL, 1996). To avoid confusion, we refer in this volume to different forms of a protein that are produced through alternative splicing as subisoforms, whereas different forms of the protein that are produced from different but related genes are termed isoforms.

4. Subisoforms and Alternative Splicing Humans and other species contain at least four PMCA genes, and each gene transcript is subject to alternative splicing. While up to four sites of alternative splicing appear to exist in each transcript (Carafoli, 1992), only two of the sites have been shown to produce different subisoforms (Fig. 5.7). Splice site A is located adjacent to the nucleotide sequence encoding the putative phospholipid-sensitive site in region B of the enzyme (Figs. 5.4 and 5.7). Although the magnitude of the effect of this regulatory site o n C a 2+ transport activity is potentially greater than for calmodulin (Penniston and Enyedi, 1994), the effects of site A splicing on this regulation are still under investigation (Carafoli and Stauffer, 1994). Splice site C is in the region of the transcript encoding the calmodulin-binding domain of PMCA ATPases (Fig. 5.7). The reader may recall that the calmodulinbinding domain is in the C-terminal cytoplasmic tail of Type IIB ATPases beyond M10. It is this expanded regulatory region that distinguishes the Type IIB ATPases from those of Type IIA. The calmodulin-binding domain beyond M10 inhibits transport activity by interacting with the autoinhibitory site between M2 and M3 of PMCA ATPases. Hence, as for splice site A, the autoinhibitory site also lies in region B. Binding of calmodulin releases the enzyme from this inhibition. The subisoforms produced through alternative splicing at splice site C appear to have different affinities for calmodulin, and these affinities are in some cases pH dependent (Carafoli and Stauffer, 1994). Insight into the physiological significance, if any, of the different affinities of the subisoforms for calmodulin, or of other functions of alternative splicing at splice sites A and C, may emerge as the developmental regulation and tissue distribution of the subisoforms are investigated. The PMCA ATPase genes 1 and 4 are transcribed about equally in various tissues, whereas genes 2 and 3 are expressed in a more tissue-specific manner (Carafoli and Stauffer, 1994; Zacharias et al., 1995). Gene 2 is expressed principally in brain and muscle, and expression of gene 3 is restricted primarily to brain (Table 5.1). In the case of all four isoforms, however, alternative splicing leads to complex patterns of expression of different subisoforms in different regions of the brain (Table 5.2). In contrast to the PMCA ATPases, both the

142

5. Structure and Function of Transport Proteins That Form Solute Gradients

FIGURE 5.7 Locations of splice sites A and C of the plasma membrane C a 2§ (PMCA) ATPase gene transcript. (A) The splice sites are shown in relation to structurally important portions of the resultant protein (compare part A of the figure to Fig. 5.4). Abbreviations and explanations for part A: TM, putative transmembrane regions 1 to 10; CaM, Ca2§ domain, which has two subdomains labeled A and B; I-CaM, autoinhibitor region that interacts with CaM in the absence of Ca2§ PL, putative acidic phospholipidsensitive region; P(D), site of transient enzyme autophosphorylation that occurs during the transport cycle; F, part of the ATP-binding region. (B) The splice options for producing subisoforms of the enzyme from the isoforms encoded by genes 1 to 4. Also shown in the left columns of part B of the figure are the designations for each splice variant (from Zacharias et aL, 1995, with permission from Elsevier Science).

splicing of transcripts e n c o d i n g the s a r c o ( e n d o ) p l a s m i c reticulum Ca2+ATPases ( S E R C A A T P a s e s ) a n d their tissue distributions are c o n s i d e r a b l y less complex.

Tissue Distribution of Site C Alternative Splicing Products of Plasma Membrane Ca z§ (PMCA) ATPase Genes 1 to 4 a

TABLE 5.1

Tissue

Gene 1

Gene 2

Gene 3

Gene 4

Brain Skeletal muscle Heart muscle Smooth muscle Other Tissues

la, lb, lc, le lb, lc lb, lc lb lb

2a, 2b 2a, 2b 2a, 2b (2b) (2b)

3a, 3b, 3g

4a, 4b 4b 4a, 4b 4a, 4b 4b

m --m

aSee Fig. 5.7B for the differences among the splice variants. Isoform 2 is present only in small amounts in smooth muscle and other tissues (adapted from Carafoli and Stauffer, 1994, with permission from John Wiley & Sons, Inc.).

S E R C A A T P a s e s are T y p e I I A A T P a s e s and, hence, do n o t h a v e the larger C - t e r m i n a l tail of T y p e IIB A T P a s e s , which interacts with calmodulin. N e v e r t h e less, alternative splicing of T y p e I I A S E R C A A T P a s e g e n e transcripts takes place in the r e g i o n e n c o d i n g the C - t e r m i n a l tail ( r e v i e w e d by MOiler et al., 1996), as is the case for T y p e IIB P M C A A T P a s e s . M o r e o v e r , the activity of S E R C A A T P a s e is r e g u l a t e d by calmodulind e p e n d e n t p r o t e i n kinase, which p h o s p h o r y l a t e s the enz y m e at seryl residue 38 ( X u et al., 1993; T o y o f u k u et al., 1994). F o r these reasons, it will be interesting to learn w h e t h e r alternative splicing in the C - t e r m i n a l tail influences r e g u l a t i o n t h r o u g h p h o s p h o r y l a t i o n n e a r the N-terminus. T h r e e genes e n c o d e S E R C A A T P a s e s in h u m a n s a n d o t h e r species (Mr et al., 1996). G e n e 3 is exp r e s s e d only in e n d o t h e l i a , e p i t h e l i a , a n d b l o o d cells, w h e r e a s g e n e 1 is e x p r e s s e d only in fast-twitch muscle cells. T h e larger fetal s u b i s o f o r m l b is r e p l a c e d d u r i n g

TABLE 5.2 Distribution of PMCA ATPase G e n e Transcripts a n d Splice Variants in H u m a n Brain” Splice options Brain region Frontal cortex Parietal cortex Occipital cortex Temporal cortex Caudate Cerebellum Globus pallidus Hippocampus Hypothalamus Inferior olive Olifactory bulb Putamen Substantia nigra Thalamus

lx

++ +++

++ +++ ++ +++ +++

+++

+++ +++ +++ +++ ++ +++

la

+++ +++ +++

++++

+++ +++ +++ +++ + + ++ +++ +++ +++

lb

lc

++ ++ ++ ++ ++ ++ +++ + + ++ +++ ++ + ++

++ ++ ++ +++ ++ + +++ ++ -

+ +++ ++ -

++

le

++

+++ +++ ++ +++ ++ -

-

+++ +++

2w

2x

-

-

-

-

-

-

-

-

-

+ ++ + + + +++ + ++ +++ ++

++ +

+ + +++ + + -

+

22

2a

2b

3x

32

++ ++ ++ ++ +++ ++++ +++ ++++ + +++ +++ ++++

+ + + + + +

+++

+++ +++ +++ +++ +++

+ +

-

+++

+

+ -

++ + +

++

+

+++ +++

++ ++ +++ +++ +++ ++ +++ +++ +++ ++ +++

+++ +++ +++ +++ ++ ++ ++ ++ +++

++

++ ++ -

+ -

-

++ ++

3a

+++ +++ +++ ++++ +++ +++ +++

+++ +++ +++ +++ +++ ++ +++

3b

4x

4a

+++ +++ +++ +++ +++ +++

+++ +++ +++ +++ +++ +++ +++ +++ +++ +++ ++ +++ +++ ++

+++ +++ +++

+++

+++ +++ +++ +++ +++ ++ +++

4b ++T

+++ +++

++

++ +++

-

++

++++ ++

+++ -

-

-

++

++

+++ ++

++ ++ ++ ++ +++

aSee Fig. 5.7B for the differences among the splice variants. All designations of splice option abundance are strictly a comparison of relative band intensities within a single gel o r blot in order to emphasize the relative preponderance of particular mRNAs over others at a given splice site. They are not intended to represent absolute amounts and should not be interpreted as such. Key: - = not detectable; + = weak; + + = present; + + + = abundant; + + + + = very abundant. (Adapted from Zacharias et a[., 1995, with permission from Elsevier Science).

144

5. Structure and Function of Transport Proteins That Form Solute Gradients

development by the somewhat smaller adult subisoform la as a result of alternative splicing of the gene 1 transcript. Gene 2 also encodes two subisoforms but in this case the subisoforms are expressed in a tissue-specific manner. While the larger subisoform 2b appears to be a "household" enzyme with wide tissue distribution, the smaller subisoform 2a is expressed primarily in slowtwitch, cardiac, and smooth muscle cells. Interestingly, coronary artery smooth muscle cells appear to have at least two Ca2+-containing compartments in their sarco(endo)plasmic reticulum, and the transport activities that load these compartments also appear to be heterogenous (Elmselhi et al., 1996). The heterogeneity in SERCA ATPase activities responsible for filling different compartments in coronary artery smooth muscle cells may result from expression of subisoforms 2a and 2b in this tissue. Expression of these two subisoforms may also account for the different sensitivities of the two activities to superoxide. In addition, since SERCA ATPase molecules may be present in the membrane as homodimers, it will be interesting to learn whether dimerization protects one subisoform of the enzyme from this oxidation.

5. Different P-Type ATPases May Function as Monomers, Homodimers, Heterodimers, or Even Heterotrimers While most P-type ATPases appear to function as monomers, the SERCA ATPases most likely operate as dimers under physiological conditions (Martonosi, 1996). SERCA ATPase molecules catalyze highly cooperative C a 2+ transport in their native membranes, and the Hill coefficient for transport (n in Eq. 4.27) of Chapter 4) is --~4. In contrast, no cooperativity is observed when SERCA ATPase monomers catalyze C a 2+dependent ATP hydrolysis in detergent solutions. Since t w o C a 2+ ions appear to be sequestered and transported together for each molecule of ATP hydrolyzed, a Hill coefficient of 4 should obtain if SERCA ATPase molecules function as dimers. Functioning of these c~subunits as dimers would also explain why ATP hydrolysis and C a 2+ transport show cooperativity in relation to the ATP concentration (Martonosi, 1996) with a theoretical maximum for the Hill coefficient of 2. Dimerization may occur through random association of SERCA ATPase monomers owing to their relatively high concentration in the sarco(endoplasmic) reticulum. All such P-type ATPases show a concentration-dependent tendency to form oligomers, and the concentration of CaZ+ATPase within the lipid bilayer of the sarcoplasmic reticulum of fast-twitch skeletal muscle is about 30,000 monomeric enzyme molecules per square micrometer of membrane (Martinosi, 1996). Conse-

quently, if monomers were present in the membrane they would be on average only about 5 nm apart. This distance is less than the distance of the top of the "head" of the ATPase monomer from the surface of the lipid bilayer (Fig. 5.1). In contrast to the SERCA ATPases, PMCA ATPases may function as homodimers regardless of their concentration within the lipid bilayer of the plasma membrane (Carafoli, 1992). While SERCA ATPase constitutes about 80% of the total protein in skeletal muscle sarcoplasmic reticulum (Martonosi, 1996), PMCA ATPase is no more than about 0.1% of the total protein in the plasma membrane (Knauf et al., 1974). Nevertheless, the mass of a PMCA ATPase molecule in the erythrocyte plasma membrane appears to be nearly the size anticipated for its dimeric form (Cavieres, 1984). Moreover, dimerization of the enzyme apparently through the calmodulin-binding domain, results in its activation (Carafoli, 1992). The possible physiological significance of interconversions between PMCA ATPase monomers and homodimers is, however, still under investigation (see also Section II,C,4 below). Some P-type ATPases also require a noncatalytic /3-subunit in order to function normally (Fig. 5.8). The a-subunits of Na+K+ATPase and H+K+ATPase remain

FIGURE 5.8 Scheme showing the relationship between the a- and /3-subunits of Na+K+ATPase. The/3-subunit has one transmembrane segment, is highly glycosylated, and contains structurally important disulfide bonds. It may also help to form K+-binding sites as indicated in the diagram (K +). The extracellular ouabain binding site (O) and intracellular phosphorylation and nucleotide binding sites are also shown for the c~-subunit. The transmembrane scheme for the a-subunit does not, however, represent the currently accepted model for the protein molecule, which contains 10 putative transmembrane segments (adapted from Chow and Forte, 1995, with permission from Company of Biologists Ltd.).

P-Type ATPases

in the endoplasmic reticulum (Chow and Forte, 1995) or are degraded rapidly after nonselective sorting to the plasma membrane (Coupaye-Gerard et aL, 1997) unless they form heterodimers with their/~-subunits (Fig. 5.9). The/~-subunit of Na+K+ATPase appears to help direct the heterodimer to the basolateral membrane of polarized cells, whereas the/3-subunit of H+K+ATPase helps direct this enzyme to the apical membrane (Chow and Forte, 1995). It is also possible to express in LLC-PK cells chimeric H+K+ATPase a-subunit molecules containing the C-terminal half of the a-subunit of Na + K+ATPase. In this case the chimeric protein combines with the Na+K+ATPase /3-subunit, but the resultant complex is delivered to the apical membrane (Gottardi and Caplan, 1993). Similarly, a chimeric Na+K+ATPase a-subunit containing the C-terminal portion of H+K+ATPase assembles with the H+K+ATP ase/~-subunit and takes it to the basolateral membrane (Muth et al., 1998). Hence, both the a- and the /3subunits may carry signals for the delivery of these ATPases to their respective membranes. The signals in the N-terminal halves of the a-subunits of both ATPases appear, however, to be dominant to any signals in the /~-subunits. The location of this proposed dominant signal has recently been more precisely defined as a sequence of eight amino acid residues in the fourth puta-

, ~!, ~

Ii

i

Apical membrane

S

\\

Basolateral membrane

-~'"

FIGURE 5.9 Hypothetical model for assembly and trafficking of Na+K+ATPase (NaK) and H+K+ATPase (HK) heterodimeric protein molecules. The a- and/3-subunits are normally synthesized and assembled into heterodimers in the endoplasmic reticulum (ER). The heterodimers move through the Golgi network where the structure and composition of their oligosaccharide moities are modified. H+K+ATPase molecules are sent selectively to the apical membrane, whereas Na§ molecules are taken to the basolateral membrane. While overexpression of the/3-subunit of H+K § sometimes leads to its expression as a monomer on the apical surface, the other a- and/3-subunits appear rarely to move beyond the ER in their monomeric forms (adapted from Chow and Forte, 1995, with permission from Company of Biologists Ltd.).

| 45

tive transmembrane segment of the a-subunit (Caplan, 1997). Both this dominant signal and the one that may be present in the /3-subunit of Na+K+ATPase would presumably need somehow to be inactivated in order for this transporter to be sent in a few appropriate instances to the apical rather than to the basolateral membrane (see also Section IV,D,3 of Chapter 2). In contrast to the requirement of the/3-subunit for normal enzyme sorting, the role of the /3-subunit in ion transport is still under investigation. Nevertheless, considerable circumstantial evidence indicates that the /3-subunit is needed for the normal transport of K + ions. In particular, the/3-subunit may be needed for normal occlusion of K + (reviewed by Chow and Forte, 1995,). Moreover, the extracellular domain of the /3-subunit appears from studies with /~-subunit chimeras of the Na+K +- and H+K+ATPases to determine the strength of the interaction of the whole ATPase with K +. Multiple genes encoding multiple isoforms of the/3-subunit are also expressed with various isoforms of the a-subunit, and the various combinations of different isoforms of the a- and/3-subunits may have a range of physiological functions. Each isoform may also undergo varying degrees of glycosylation, thus further expanding the range of possible functions of the various forms. The high level of glycosylation of the /~-subunit of the gastric H+K+ATPase may help to protect the enzyme from the harsh extracellular environment that it helps to produce in the stomach (Chow and Forte, 1995), although this hypothesis needs to be tested further. The details of assembly of the a- and/3-subunits is also an active field of investigation. Specific regions of each polypeptide are required for formation of the complex but detailed study of the association is hampered because the monomers irreversibly dissociate upon detergent extraction. Coimmunoprecipitation of the subunits is, however, possible and, in the case of Na+K+ATP ase, a third, smaller y-polypeptide also coprecipitates (Mercer et aL, 1993). This putative y-subunit is highly conserved among species and most likely has an essential function. While the Na+K+ATPase molecule may for this reason be a heterotrimer, we are reminded also that the boundaries of the components of many protein complexes may be imprecise. For example, Na+K + ATPase a-subunit molecules also bind to ankyrin molecules, and ankyrin molecules appear to help mediate attachment of the enzyme molecules to the cytoskeleton (e.g., Nelson and Veshnock, 1987). For these reasons, it is not fully clear which protein molecules are part of one protein complex and which are part of another. While such physical boundaries of protein complexes may be imprecise, the functions of the associations among the components of the complexes remain amenable to experimental investigation.

| 46

5. Structure and Function of Transport Proteins That Form Solute Gradients

C. Relationships b e t w e e n the Structure and Function of P-Type ATPases As we have just discussed, important structural and functional relationships exist between the catalytic a-subunit of P-type ATPases and other protein molecules not usually considered to be components of the ATPases. Such longer-range effects of other protein molecules on transport remain an active and important area of research (e.g., see Section IV,A of Chapter 6). These studies are needed for us to comprehend fully the function of the ATPases in the context of living cells and organisms. Similarly, completion of the transport cycles of P-type ATPases could conceivably involve phase transitions in the membrane lipid bilayer as discussed in Chapter 3 (Section VI, C). It is, however, also important to focus on how the catalytic c~-subunit itself functions even though its interactions with other components of the cell contributes to this function. In this regard, much is now known about which amino acid residues of the a-subunit are needed for various steps in the active transport cycle (Fig. 3.7 in Chapter 3). The functions of these residues and the structures that they form in P-type ATPases are considered in detail below.

1. Components of Both the N- and the C-Terminal Membrane-Associated Domains May Be Involved in Ion Migration Among P-type ATPases, the structure and function of SERCA ATPases have been particularly well studied by various procedures including site-directed mutagenesis. For this reason, we will focus in this Section mainly on these transporters (Andersen and Vilsen, 1995). Nevertheless, the model for SERCA ATPases applies reasonably well to all P-type ATPases (MOiler et al., 1996). As discussed above, a three-dimensional model for the structure of P-type ATPases has been developed (Fig. 5.1). This structure corresponds to a proposed model for the arrangement of the 10 putative transmembrane a-helices (Stokes et al., 1994). Based in part on the amphipathic nature of some of the transmembrane helices, and on the clustering of lesswell-conserved residues on one side of them, a threedimensional arrangement of the helices has been proposed (Fig. 5.10A). 2 This arrangement is also based on the observation that helices M5, M6, and M8 in the Cterminal membrane-associated domain and M4 in the N-terminal one appear to form the pathway through which Ca 2+ ions pass (Stokes et al., 1994; Andersen and Vilsen, 1995). The same helices most likely also form 2 Results from a more recent study in the Stokes laboratory at about 8 ,~ resolution support a modified arrangement of the helices (Zhang et al., 1998).

the pathway for transport catalyzed by other P-type ATPases. Extensive site-directed mutagenesis studies show rather conclusively that these four helices contain all but one of the 19 residues known to be needed for normal Ca 2+ binding by SERCA ATPases (Ca2+-affinity mutants in Fig. 5.11). Note that known mutations of residues in other transmembrane helices (shown as solid circles in Fig. 5.11) do not greatly affect Ca 2+ affinity or transport (Andersen and Vilsen, 1995, 1998). Moreover, other evidence supports the conclusion that E908 in M8 (Fig. 5.11) contributes only peripherally to Ca 2+ binding (MacLennan et al., 1997), so Ca 2+ binding and transport may actually occur primarily via M4, M5, and M6 (Fig. 5.10B). Nevertheless, complete replacement of helix M3 in the SERCA ATPase molecule with M3 of Na+K + ATPase significantly reduces to 1% the Ca 2+ affinity as measured by its ability to activate enzyme autophosphorylation (NCrregaard et aL, 1993). Hence, transmembrane segments of other than M4 to M6 clearly have at least indirect effects on the pathways of ion migration formed by P-type ATPases. Some of the CaZ+-affinity mutants in M4, M5, and M6 are also EzP mutants, whereas other mutants in these helices show only the EzP phenotype (Andersen and Vilsen, 1995, 1998). Dephosphorylation of the EzP form of the enzyme is blocked in such mutants (diamonds in Fig. 5.11). Since the cytosolic component of the enzyme contains the site of autophosphorylation, we must now briefly describe how this component may function before we consider the interactions between the cytosolic and membrane-associated domains.

2. ATP Hydrolysis and Enzyme Autophosphorylation Are Catalyzed in the Large Cytosolic Loop Using fluorescence energy transfer between derivatives of selected amino acid residues in the SERCA ATPase, it is possible to estimate the distances between and among the residues, the lipid bilayer surface (in which are inserted phospholipids with fluorescent labels), and the enzyme autophosphorylation site (reviewed by MOiler et al., 1996). These covalent modifications of the ATPase and membrane lipid usually do not alter transport or ATPase activity. With this and other approaches, the locations of several residues in the large cytosolic loop can be arranged in relation to each other (Fig. 5.12). Also shown in Fig. 5.12 are the approximate locations of these residues in the head of the ATPase molecule (Fig. 5.1). With these landmark residues, one can begin to project the three-dimensional locations of many of the residues in the large cytosolic loop that influence various steps in the transport cycle of SERCA ATPases (landmark residues at the corners shown in Fig. 5.12 are marked by arrows in Fig. 5.11). In this

P-Type ATPases

147

A

6~

I F

C

~'A S ~ G -. .-r - - - - _ . : ~ ( ~ ) R ~ ' / ~ Y

/

G

L

L

&

v~::LP,x'~ '~ , P

Q F

L

Ca**

I W'i~]li~ ..~ ~i~i #t~ ~T

N

A L T

I

-

_@

L~ ~

y-

'

g

N

"=='~(~,,~

@ A ~

M~ L

l

N

M8 895

/

I

s-"'.ii;"i;:n~l~--~|

'

D(~I

#

/

@L A

- ~

V vvv T

@ H-~'-'~ e

G F@ G

v

S ~

L

0

M9 929

9 ttid,j;t,-.lr

FIGURE 5.10 Hypothetical packing of the putative 10 transmembrane helices of the sarco(endo)plasmic reticulum Ca 2+ (SERCA) ATPase (compare arrangement to that shown in Fig. 5.1). The arrangement shown in A is based in part on the amphipathic nature of some of the helices and on the clustering of less well conserved amino acid residues on one side of the helices. Such variable residues in transmembrane helices are more likely to be in contact with membrane lipid. The arrangement also reflects the hypothesis that M4, M5, M6, and M8 form the pathway across which Ca 2+ ions migrate. The helices are viewed from the cytoplasmic surface, so odd numbered ones run away from the viewer, whereas even numbered ones run toward the viewer. Note, however, that many of the helices are highly inclined especially according to a recent model (Zhang et al., 1998) so they do not usually run perpendicular to the membrane. (A modified packing arrangement of the helices is also presented in this new model.) Filled diamonds mark the N-terminal residues of each helix, and their position numbers in the protein molecule are shown in the centers of the helices. The numbers 2 to 7 mark the remainder of the first seven residues in each helix, and arrows emanating from them indicate residues in subsequent heptads. More variable residues are circled, and those thought to contribute to Ca 2+ binding are enclosed in triangles. C-terminal residues in each helix overlap with those at or near the N-terminus, so the C-terminal ones are shown at higher radii (adapted from Stokes et al., 1994, with permission from Elsevier Science). (B) Modified arrangement for the pathway shown in A based on evidence that M8 actually contributes only peripherally to Ca 2+ binding. In this case, the transport pathway is proposed to be formed from M4, M5, and M6 alone. Also shown in B are residues in these transmembrane helices that appear to contribute to Ca 2+ binding at sites I and II (adapted from MacLennan et al., 1997, with permission from Elsevier Science).

r e g a r d , t h e r e a d e r s h o u l d n o t e t h a t s o m e w h a t less strict

For example, conservative substitutions of most of the

criteria may have been used to decide whether mutants

r e s i d u e s in t h e s e q u e n c e m o t i f s u r r o u n d i n g

s h o w n in Fig. 5.11 i n f l u e n c e t h e a c t i v i t y o f t h e e n z y m e

p h o r y l a t i o n site (i.e., t h e a s p a r t y l ( D ) r e s i d u e 351 in t h e

as c o m p a r e d t o t h e c r i t e r i a o c c a s i o n a l l y u s e d in t h e t e x t .

sequence ICSDKTGTLT)

the phos-

do not prevent phosphoryla-

148

5. Structure and Function of Transport Proteins That Form Solute Gradients

3. Physical Relationships among the Sites of ATP Hydrolysis, Enzyme Autophosphorylation and Ion Transport

B

FIGURE 5 . 1 0 (Continued)

tion (discussed in Section II,B,2 above), although several semiconservative substitutions do prevent phosphorylation or the E~P-to-EzP transition (Fig. 5.11). (The reader may also wish to review the extended Albers-Post reaction Scheme for P-type ATPases in Fig. 3.7 of Chapter 3.) It has been proposed that the conserved lysyl residue 684 and the regions around the conserved lysyl residue 492 help to catalyze an acid-base destabilization of the y-phosphoryl group of ATP and its transfer to the carboxyl group of aspartyl residue 351 (Mr et al., 1996). (These two conserved lysyl residues are the same two described in Section II,B,2 above to precede the hinge and KGAPE motifs, respectively, Fig. 5.4.) Substitutions of residues at or near position 492 result, however, in ATPase molecules with near normal Ca 2+ affinities and turnover rates, whereas substitutions for lysyl residue 684 prevents both phosphorylation and the E1P-to-EzP transition. While the latter results may seem at first contradictory, it is indeed possible for different substitutions to produce these two phenotypes. For example, conservative substitution of an arginyl residue for lysyl residue 684 results in a phosphorylatable enzyme that is nevertheless incapable of the E1P-to-EzP transition, whereas other substitutions prevent phorphorylation in the first place (Vilsen et al., 1991). These results indicate that lysyl residue 684 is more important than 492 or its neighboring residues in helping to catalyze enzyme autophosphorylation during the transport cycle.

It is also possible to use the results of site-directed mutagenesis to begin to visualize the physical relationships that are required among amino acid residues in SERCA ATPase molecules for free energy transfer to occur (Fig. 5.11). The free energy transfer occurs over a distance of about 4.0 nm between the ATP and Ca 2+ binding sites (Toyoshima et al., 1993) and appears to involve conformational changes. Hence, mutations in which conversion from the E1P to the E2P conformation is blocked (squares in Fig. 5.11) seem particularly likely to be important to the process of free energy transfer. Such residues are prominent both between M4 and the phosphorylation site in the large cytosolic loop and in the smaller cytosolic loop between M2 and M3 (Fig. 5.11). These residues must somehow help to connect the site of ATP hydrolysis to that of Ca 2+ binding and transport. While this connection seems to be quite direct in the case of residues between M4 and the phosphorylation site, it is less clear precisely how the cytosolic loop between M2 and M3 weaves into the three dimensional structure connecting the sites of chemical change and ion transport. While M4 is believed to help form the pathway through which Ca 2+ ions pass, neither M2 nor M3 appear to do so (Fig. 5.10). MOiler and associates (1996) proposed that the loop between M2 and M3 (region B, Fig. 5.4) contributes to free energy transfer by helping to stabilize the $4/$5 link between domain C and the membrane c~-helices that form the transport pathway. As discussed in Chapter 3, it is also possible that the free energy transfer is less direct than implied from mutagenesis studies. In particular, we suggested that the free energy transfer may involve conversion of membrane lipid that surrounds an ATPase molecule from a crystalline to a liquid state. In this model, the mutations in the cytosolic loop between M2 and M3 would not necessarily need to be in a structure that directly connects the ATP and Ca 2+ binding sites in order to interrupt the conversion of free energy from that in ATP to that in a Ca 2+ gradient. In fact, even the residues between M4 and the site of phosphorylation could conceivably influence both the conversion from the E1P to the E2P conformation and the associated conversion of free energy indirectly through an influence on membrane lipid. If some of the free energy conversion occurs through an effect on the locations of membrane liquid and crystalline phospholipid domains, then the free energy change associated with conversion between different protein conformations may be smaller than for ATP hydrolysis or for Ca 2+ transport against its gradient.

P-Type ATPases

~L

| 49

402 492 t~515

684 365

601

703

712

N t ~ ~ ~rmf_.._.. ,d~,~(~ ~ " ~ " - ~ _ j ' f

M

1

2

3

4

5

6

7

_.._..~_

8

9

~

C

10

FIGURE 5.1 1 Topology of the sarco(endo)plasmic reticulum C a 2+ (SERCA) ATPase showing amino acid residues that have been studied by site-directed mutagenesis. The functionally important residues are indicated by their single-letter abbreviations inside a symbol showing the phenotype that results when the residue is replaced. Key: open circles, indicate mutants unable to occlude Ca2+or that display at least a threefold reduction in apparent Ca2+ affinity; triangles pointing up show mutants that do not undergo autophosphorylation of residue 351; squares indicate mutants in which the transition from EIP to E2P is blocked; diamonds indicate mutants in which the dephosphorylation of E2P is blocked; the triangle pointing down indicates the Tyr763to-Gly763 mutation where ATP hydrolysis is uncoupled from Ca2+transport. The filled circles indicate mutants displaying normal Ca2+ affinity and a maximum turnover rate above 20% of that of the wild-type enzyme. Framing of a residue abbreviation by two symbols indicates either that two steps in the transport cycle are affected by the same mutation or that different substitutions elicit different effects. Sites for binding of phospholamban (PL), regulatory phosphorylation of a seryl residue in isoform 2, and binding of thapsigargin (rectangle in M3) are also shown. 9 - Mutants retaining C a 2+ transport with normal affinity; O = Ca2+-affinity mutant; ~ = EaP to E2P mutants; P ~ regulatory serine phosphorylation site in isoform2; r---q -- thapsigargin binding site; O = E z P dephosphorylation mutants; A - phosphorylation negative mutants; V = uncoupled mutant; ~ 3 = phospholamban binding site. (Adapted from Andersen and Vilsen, 1995, with permission from Elsevier Science.)

In a survey of allosteric p r o t e i n s for which t h e r e are data, c o n f o r m a t i o n a l changes of the s a m e b i o e n e r g e t i c m a g n i t u d e as those of P-type A T P a s e s occur without hydrolysis of p h o s p h o r i c acid a n h y d r i d e b o n d s (Goldsmith, 1996). M o r e o v e r , the free e n e r g y changes associa t e d with all of these c o n f o r m a t i o n a l changes are generally smaller than those of A T P hydrolysis and active cation transport. F o r example, it has b e e n e s t i m a t e d that the differences in free e n e r g y of a l t e r n a t e conform a t i o n a l states of allosteric p r o t e i n s (--~22 kJ mol-1; G o l d s m i t h , 1996) are only a b o u t half of the free e n e r g y

change of A T P synthesis or hydrolysis (--~48 kJ mo1-1) and active cation t r a n s p o r t (--~42 kJ (mol of A T P hydrolyzed) -1) (see C h a p t e r 3). T h e r e f o r e conclusions a b o u t the p a t h w a y of free e n e r g y transfer in P-type A T P a s e s m u s t r e m a i n provisional until all of the p e r t i n e n t forms of free energy, i n t e r m e d i a t e to those i n h e r e n t in A T P and ion gradients, have b e e n identified. To d e v e l o p a m o d e l that is consistent with these considerations, let us first a s s u m e naively that all of the free e n e r g y is t r a n s f e r r e d b e t w e e n A T P and a cation g r a d i e n t by the p e r t i n e n t s e q u e n c e s of a m i n o acid residues in

| 50

5. Structure and Function of Transport Proteins That Form Solute Gradients

K51~,q / /

I I I l

\ \

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FIGURE 5.12

~

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,,,_, '~

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I,

-

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:

j

i

/

t

/

/

/

~-

.... I

'I

Locations of several residues in the large cytosolic loop of sarco(endo)plasmic reticulum Ca 2+

(SERCA) ATPase in relation to the surface of the membrane phospholipid bilayer, the site of ATP (A-RibP-P-P) binding, and the site of transient enzyme phosphorylation (D351). The locations of each of the numbered amino acid residues are indicated in Fig. 5.11 as well. Also shown are the relationships of all of these residues to the three-dimensional structure of the cytosolic component of the enzyme (outline drawn from structure shown in Fig. 5.1). Units of distance when not indicated are in ,~. Abbreviations; C, cysteine; D, aspartate; E, glutamate; K, lysine (adapted from MOiler et al., 1996, with permission from Elsevier Science).

the A T P a s e s and the structures that these s e q u e n c e s form. As anticipated f r o m the p r e c e d i n g discussion, the resultant m o d e l will be insufficient to a c c o u n t fully for the free e n e r g y transfer. W e will t h e n e x p a n d the m o d e l in the following Section in a way that could a c c o u n t for the free e n e r g y c h a n g e s that are k n o w n to occur during t r a n s p o r t by P-type A T P a s e s . This e x p a n d e d m o d e l is i n t e n d e d to p r o v o k e n e w theories and investigations into the actual m e c h a n i s m of free e n e r g y transduction. 3 3 It is also anticipated that new results will be forthcoming from additional site-directed mutagenesis studies. Although it is easy to form the impression that most such pertinent experiments have been performed (e.g., Fig. 5.11), new literature dissuades us from such a notion. For example, conversion of the conserved lysyl residue at position 758 in $5 of the SERCA ATPase to an isoleucyl residue renders the E2P form of the enzyme more rapidly dephosphorylated but less able to undergo the subsequent CaZ+-dependent E2 to Ea( Ca2+)2 transition (SCrensen et al., 1997).

4. Possible Mechanisms for Free Energy Transfer during the EIP-to-E2P Transition and Subsequent Conformational Changes P - T y p e A T P a s e m o l e c u l e s are v i e w e d as c h a n n e l s or p a t h w a y s t h r o u g h the m e m b r a n e that are c o u p l e d to an i m m e d i a t e source of free e n e r g y in A T P (e.g., A n d e r s e n a n d SCrenson, 1996). C o n s i s t e n t with this view, they are also k n o w n to u n d e r g o c o n f o r m a t i o n a l changes to catalyze t r a n s p o r t r a t h e r t h a n to carry their substrates across the m e m b r a n e (i.e., m o d e l A r a t h e r t h a n B in Fig. 5.13). In the case of P - t y p e A T P a s e s , such conform a t i o n a l changes n e e d s o m e h o w to link the inorganic ion binding site to the A T P binding site s o m e 4 n m away ( T o y o s h i m a et aL, 1993). It has b e e n c o n c l u d e d that a m i n o acid residue substitutions that p r e v e n t the E1P-to-EzP transition (squares in Fig. 5.11) m a y influence m o v e m e n t of the $4 stalk s e q u e n c e , which also

P-Type ATPases

9

9

o

FIGURE5.13 Two general mechanisms by which proteins may catalyze transport of solutes across biomembranes. In the first model (A) the protein undergoes a relatively small conformational change to permit the solute to migrate across the membrane phospholipidbilayer via the pathway formed by the protein. The second mechanism (B) requires the transport protein to undergo a conformational change large enough to carry the solute across the bilayer. The extreme of the latter case is complete reversal of the orientation of the protein in the membrane as shown in B. We favor the model shown in A as the most likely general mechanism by which virtually all transport proteins function. The degree to which the sizes of the conformational changes may vary among transport proteins is, however, still under investigation (adapted from Andersen and SCrensen, 1996, with permission from Elsevier Science)

may mediate rotation or tilting of M4 to release occluded Ca 2+ ions in the case of S E R C A ATPases. The latter structural changes also appear to lead to conformational changes at the autophosphorylation site such that the phosphorylated aspartyl residue becomes able to react hydrolytically with water (Andersen and SCrensen, 1996). In this regard, several amino acid residue substitutions in transmembrane helices of the S E R C A ATPase molecule result in E2P mutants that are unable to undergo dephosphorylation (Fig. 5.11). Presumably the latter mutants are unable to complete a final, relatively small portion of the E1P-to-E2P conformational change that is needed for reaction of the phosphorylated enzyme with water. Since the autophosphorylation site lies well outside the membrane, it is interesting that the only amino acid residue substitutions that have been shown to result in this dephosphorylation-negative E2P phenotype are located within the membrane (Fig. 5.11). The conformational change that permits dephosphorylation must therefore cross a considerable distance without demonstrated involvement of intervening amino acid residues of a type that can also prevent dephosphorylation when mutated. Hence, we see that clear gaps still

151

exist in the description of the structural changes that Ptype ATPases must undergo to complete their transport cycles. We propose that the coupling of ATP hydrolysis to cation transport can be understood only by considering mechanisms of free energy transfer in addition to those associated with protein conformational changes. Some of these additional mechanisms of free energy transfer may occur through an effect on membrane phospholipid, a For example, some of the amino acid residues in the membrane-associated domains may be needed for enzyme dephosphorylation (diamonds in Fig. 5.11) because they are needed for the transition of the phospholipid domains surrounding each enzyme molecule from a crystalline to a liquid state. Conversion of the lipid domains surrounding enzyme molecules from crystalline to liquid was proposed in Chapter 3 to occur in association with the E1P-to-EzP transition. Moreover, in the case of S E R C A ATPases, it has been proposed that conversion from the E1P to the EzP conformation is associated with dimerization of the enzyme monomers (Boldyrev and Quinn, 1994). In this case, the enzyme would not operate continuously in the dimeric form as discussed above. Instead, it would change back and forth between the monomeric and dimeric forms during the transport cycle. Perhaps the crystalline lipid domains that are proposed to surround the ATPase molecules in their E1P conformations prevent the monomers from forming dimers. If so, a change in state of the lipid to liquid could conceivably facilitate dimerization. 5 We suggest that oligomerization of S E R C A ATPase monomers may lead to the conformational changes that are needed at the sites of phosphorylation in order for them to react with water. Hence, dephosphorylation-negative EzP mutants may only occur in the membraneassociated c~-helices of S E R C A ATPases (Fig. 5.11) because only such mutations can influence the membrane phospholipid surrounding the enzyme molecules in this case to resist conversion from the crystalline to the liquid state. Also in this case, a membrane lipid phase transition would be needed only for the final stage of the E1P-to-E2P conformational change in which the enzyme is converted to a form that reacts with water rather than for the whole conformational change. The observation 4 The present discussion is not intended to deny that the free energy changes associated with each step of the transport cycle have been more or less accurately estimated for P-type ATPases (Jencks, 1989; Stein, 1990). Rather, we argue that the free energy differences among the forms and conformations of P-type ATPase proteins are insufficient alone to account fully for the known free energy conversions. 5 Alternatively, crystalline lipid domains could stabilize dimers, whereas the dimers could dissociate into monomers when the domain "melts." In either case, conversion between dimers and monomers could conceivably be needed for enzyme dephosphorylation.

| 52

5. Structure and Function of Transport Proteins That Form Solute Gradients

that PMCA as well as SERCA ATPases also may form dimers (e.g., Carafoli, 1992) is consistent with the conclusion that all P-type ATPases may form dimers at least transiently and that interconversion between dimeric and monomeric states may be necessary to complete the transport cycle. It has also been suggested that the P-type H+ATPase of Neurospora may undergo dynamic interconversions between the monomeric and dimeric forms (Bowman et al., 1997), and such may be the case also for cq~ and (a/~)2 dimers and tetrameres of Na+K+ATPase (Linnertz et al., 1998). In contrast to P-type ATPases, the conformational changes in F-type ATPases appear more directly to connect the sites of transport and ATP hydrolysis or synthesis (e.g., Fillingame, 1997). Moreover, it appears unlikely that these already highly oligomeric proteins form "superdimers" between two whole FoF1 complexes during the transport cycle (Fig. 5.14), although the three catalytic subunits within the F1 sector may so influence each other in F-type ATPases. There also seems to be

no support for the notion that the transport cycles of F-type ATPases involve phase transitions in membrane lipid domains, whereas data supporting this theory for P-type ATPases were presented in Chapter 3. Instead, the most widely accepted model for the action of F-type ATPases involves sequential changes in the free energy of binding of nucleotides at each of the three catalytic sites on the extramembrane F1 sector (e.g., Boyer, 1993). The changes in binding affinities in the F1 sector are believed to be driven by (or to drive) proton transport across the Fo sector. We turn now to a detailed consideration of the structure of these other transport ATPases and the mechanisms by which they appear to transfer free energy between the sites of proton transport and chemical change in ATP.

II!. FoF~-ATP SYNTHASES (F-TYPE ATPases) In contrast to the P-type ATPases, the extramembrane (F1) and membrane (Fo) sectors of F-type ATPases are each composed of several separate proteins (Fig. 5.14). Hence, it should in theory be somewhat easier to study the effects on function of structural alterations in distinct components of F-type ATPases. Nevertheless, as for P-type ATPases, the mechanisms of free energy coupling are still emerging for this class of primary active transport proteins. We begin our discussion of F-type ATPases with a description of their threedimensional structures and classification. In subsequent Sections we discuss the structures of the components of these enzymes in greater detail and we consider how they may catalyze the conversion of the free energy in a proton gradient into that in the phosphoric acid anhydride bonds of ATP. A. Evolution and Classification of F- and V-Type ATPases

FIGURE 5 . 1 4 Model of the structure of the F-type ATPase in E. coli. The subunits inside the hexamer formed by the a- and/3-subunits can be seen only because one of the three a-subunits is not shown. The horizontal lines represent the surfaces of the membrane phospholipid bilayer (adapted from Capaldi et al., 1994, with permission from Elsevier Science).

Evolution of F- and the related V-type ATPases appears to have occurred over 3 billion years ago. Nelson (1992b) proposed that these two types of ATPases evolved from a single ancestral proton pump composed of a homohexamer. Each of the subunits of the hexamer is believed to have had both an extramembrane and a membrane domain. Hence, the hexamer would have had a structure quite similar to that of modern F- and V-type ATPases, which have three catalytic and three homologous noncatalytic subunits in their extramembrane sectors (Fig. 5.14). The ancestral homohexamer would, however, have had either no additional subunits or additional subunits that are entirely different from the ones in modern F-/V-type ATPases. Such additional subunits in modern F-/V-type ATPases include the

153

FoF1-ATP Synthases (F-Type ATPases)

y-, 6-, e-, a- and/3-subunits of the E. coli enzyme that appear only once or twice in each enzyme molecule (Fig. 5.14). The stoichiometries of association of subunits in modern FoF1 and VoV1 protein complexes as well as similarities among amino acid residue sequences of the subunits present in the enzymes in different species were used to construct the following Scheme for evolution of the subunits retained in modern F- and V-type ATPases (Nelson, 1992b). At least 3 billion years ago the proposed ancestral gene separated into two genes, one of which encoded subunits in the membrane sector (as in Vo and Fo) and one of which encoded subunits in the extramembrane sector (as in F1 and V~) (Fig. 5.15). Early evolution of the genes that encoded extramembrane subunits is believed to have included a gene duplication with concomitant evolution of genes encoding two different but homologous subunits of an ancestral proton pump. The resultant heterohexamer of the extramembrane component presumably was composed of three subunits that resembled the three A-subunits of modern V-type ATPases and three subunits that resembled the three B-subunits of this enzyme.

The precise order of evolution of F- and V-type ATPases is obscured not only for the usual reason that it is difficult to reconstruct ancient events, but also because it is unclear what may have provided the selective impetus for the separate evolution of the two types of ATPases. Conceivably, a duplicate of a gene encoding subunits of a homohexameric V1 sector evolved first into a second pair of genes encoding two different subunits in an ancestral heterohexameric F1 sector capable of helping to perform photosynthesis. Subsequently, this new heterohexameric structure might have given rise to improved heterohexameric V-type ATPases. Alternatively, homohexameric V-type ATPases may have evolved into heterohexameric V-type ATPases as described in the preceding paragraph. A second round of gene duplications could have then given rise to subunits in an ancestral F1 sector that were capable of helping to perform photosynthesis. Elsewhere it has been proposed that accumulation of oxygen in the atmosphere and the advantages of oxidative metabolism as well as the disadvantages of oxidative stress led an F-type ATPase involved in oxidative phosphorylation to evolve from a V-type enzyme. In our view the latter possibility

Ancestral gene Memb. Extra.

• 1

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F-ATPase

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I

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or

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Arch.| Vacuoii

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r Subunit A Subunit B V-ATPase V-ATPase I

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Subunita Subunit13 F-ATPase

FIGURE 5.15 Proposed scheme for the evolution of F- and V-type ATPases. An ancestral gene is believed to have encoded both the membrane (Memb.) and extramembrane (Extra.) components of each of the six proposed subunits of the homohexameric ancestral protein. This gene is believed to have split into separate genes encoding the two components which then evolved separately but of course not independently. The gene encoding membrane components must have undergone duplication at some time during evolution in order to produce the genes encoding membrane components of both F- and V-type ATPases. From the ancestral membrane (Memb.) gene evolved genes encoding the modern subunit c (also known as proteolipid) of F-type ATPases and the genes encoding the same subunits of the V-type ATPases. In the case of the genes encoding the c-subunits in V-type ATPases (except those in Archaeobacteria), the ancestral gene is also believed to have undergone a duplication followed by a fusion to encode a c-subunit that is twice the size of the c-subunit in F-type ATPases. The ancestral gene encoding extramembrane (Extra.) subunits is also believed to have undergone duplication on more than one occasion. One duplication appears to have produced genes encoding different subunits in the ancestral heterohexameric extramembrane component. Each of these genes is believed then to have undergone a second duplication to produce genes encoding the catalytic subunits, A and/3, or noncatalytic subunits, B and a, in V- and F-type ATPases, respectively. Other abbreviations; Eubact., eubacterial; Chloro., chloroplast; Mito., mitochondrial; Arch., archaebacterial; and Vacuol., vacuolar forms of subunit c (proteolipid) (adapted from Nelson, 1995, with permission from Landes Bioscience).

154

5. Structure and Function of Transport Proteins That Form Solute Gradients

seems unlikely because evolution of photosynthesis utilizing F-type ATPases probably preceded oxygen accumulation. Hence, an F-type ATPase for oxidative phosphorylation probably evolved from or possibly in combination with the photosynthetic one. 6 In any case, additional gene duplications in ancestral genes (Fig. 5.15) would have been needed for the evolution of genes encoding the A-subunits of modern V-type ATPases (Fig. 3.10) and the homologous/3-subunits of modern F-type ATPases (Fig. 5.14). Similarly, duplication of the gene encoding the other major subunit of the extramembrane sector would have permitted evolution of the B- and a-subunits of V- and F-type ATPases, respectively. Subsequent evolution of the extramembrane sectors of the enzymes appears to have involved the addition of a variety of different subunits to different forms of the enzyme in order to serve the diverse needs of various organelles and cell types. Nelson (1992b) proposed a similar course of evolution for the c-subunit of the membrane sector of the enzymes. Each subunit of the ancestral membrane sector is assumed to have had three membrane-spanning helices because an F1 facing loop is needed in subunit c for interaction of the modern F0 sector with F1 and because the ancestral gene is believed to have evolved from a gene that encoded both membrane and extramembrane domains (Fig. 5.15). A gene encoding either the two most C-terminal or the two most N-terminal of these three helices appears therefore to have evolved into a gene encoding the ancestral c-subunit, which is also known as proteolipid. The genes encoding proteolipid have remained highly conserved in numerous species to the present day. The number of copies of this highly hydrophobic membrane c-subunit in the Fo complex (Fig. 5.14) has been determined experimentally to be 10 _+ 1 (Foster and Fillingame, 1982), so it is either present in a variable amount or, more likely, its stoichiometry is difficult to determine. Twelve copies of the c-subunit are proposed to be present in the most recent model of the E. coli F-type ATPase (Fillingame, 1997; see Section III,B below). Different genes encode the proteolipid in the F-type ATPases of mitochondria and chloroplasts, and each of these genes appears to have evolved from the same ancestral gene that gave rise to the gene encoding proteolipid in bacteria (Fig. 5.15). In contrast, a more distant ancestor of the latter group appears to have given rise to genes encoding the proteolipid in Archaeobacteria and a similar subunit in all Vtype ATPases. In the case of V-type ATPases, a gene duplication was followed by fusion to produce a 16 kDa 6 Actually photosynthesis using hydrogen donors other than water is believed to have evolved prior to evolution of photosynthesis that generates oxygen. Hence, F-type ATPases may have evolved before evolution of organisms that produce or utilize oxygen.

rather than an 8-kDa c-subunit in modern forms of the enzymes (Figs. 3.10 and 5.15). Both the inability of the membrane sector of V-type but not F-type ATPases to conduct protons when the cytosolic sector is removed and the lower stoichiometry of proton transport in V-type than in F-type ATPases have been attributed to the presence of the larger c-subunit in V-type than in F-type ATPases (Nelson, 1995). The reader may note that both of these effects should permit V-type ATPases to produce steeper total chemical potential proton gradients across the membrane than do F-type ATPases. Somewhat surprisingly, however, most V-type ATPases do not produce transmembrane proton gradients that approach their theoretical maximum (Nelson, 1992b). 7 Rather, the uncoupled transport of some protons across the membrane in the direction of their lowest total chemical potential without concomitant chemical change appears in most instances to prevent V-type ATPases from either producing near maximum proton gradients or catalyzing the reverse reaction (i.e., synthesis of ATP when the proton gradient is sufficiently steep). It is currently unclear whether this uncoupled transport of protons occurs via the intact VoV1 complex or some other process. For these reasons, it will be interesting to learn how the larger c-subunit in V-type ATPases helps to produce functional characteristics that are different from those of F-type ATPases. Most of the other subunits of the membrane sectors of F- and V-type ATPases appear to have been added after evolution of the c-subunit. Only in the case of subunit a (Figs. 3.10 and 5.14) is it believed that the original ancestral protein may have given rise to the modern protein, possibly from the third transmembrane segment thought to have been present in the ancestral protein (Nelson, 1992b). Alternatively, this third transmembrane helix may have been replaced by the modern subunit a, and a variety of other subunits have been added to different forms of the enzymes apparently to serve the specialized needs of different organelles and cell types. In addition, most of the genes that encode components of F-type ATPases and, of course, all of those that encode components of V-type ATPases in eukaryotic cells have been transferred to the nucleus. Nevertheless, some of the genes are still located in the v The V-type ATPases in archaebacteria are exceptions to this generalization. These enzymes catalyze proton extrusion against a much greater total chemical potential gradient than most other V-type ATPases (reviewed by Nelson, 1992b). More efficient proton transport permits some species to grow at an environmental pH of two. Moreover, the physiological function of V-type ATPases in some Archaeobacteria and, somewhat surprisingly, also in the eubacterium, Thermus thermophilus, appears to be ATP synthesis (Yokomyama et al., 1998). It will be interesting to learn whether the more efficient proton transport in Archaeobacteria results from the smaller size of the c-subunit in their V-type ATPases (Fig. 5.15).

FoF1-ATP Synthases (F-Type ATPases)

genomes of mitochondria and chloroplasts, and in some cases the locations of genes in either an organelle or the nucleus vary among species (Nelson, 1992b). While the evolution of F- and V-type ATPases is an interesting multidisciplinary topic, it promises ultimately to occupy a full volume. We return now to the main purpose of this Section, which is to describe the structure and function of F-type ATPases. We first describe the structure of the Fo membrane sector of these enzymes and the pathway by which protons may cross the membrane. Subsequently, we discuss the structure of the extramembrane F1 sector and then consider how ATP synthesis or hydrolysis in this sector might be coupled to proton transport in the Fo sector. B. The M e m b r a n e Sector Fo Forms the Pathway for Proton Transport

I. The Fo Sector Is Composed of Subunits The Fo complex of E. coli is composed of a single a-subunit, two b-subunits, and multiple copies of subunit c. While Fo in chloroplasts contains homologs of these three peptides, the mitochondrial form of the Fo sector is considerably more complex (Pedersen, 1996). Not only does the bovine heart mitochondrial Fo sector contain at least nine different polypeptides, but it also contains no clear homolog to the b-subunit of Fo in E. coli. Nevertheless, the Fo sectors of eubacteria, chloroplasts, and mitochondria appear to function in a similar manner. Moreover, most site-directed mutagenesis studies have been performed to modify the E. coli enzyme. For these reasons, we in general focus our discussion on Fo of E. coli. In spite of numerous site-directed mutagenesis studies, the precise arrangement of subunits in Fo is still under investigation. The two b-subunits in Fo were shown recently to interact as a dimer with F1 in the stalk region of the FoF1 complex (Fig. 5.14) (Rodgers et aL, 1997), but little else is known about their location among most of the other subunits in Fo. A newly formulated model for the Fo c-subunit oligomer is being tested using site-directed mutagenesis (Groth and Walker, 1997), although this method can provide only indirect information about protein quaternary structure. High resolution X-ray crystallographic studies would undoubtedly tell us the number and arrangement of subunits in Fo, but unlike the extramembrane F~ component, the lipophilic Fo sector has not as yet been crystallized in three dimensions. In earlier models, the two b-subunits and single a-subunit were centrally located in the sector (Fig. 3.10), whereas a more recent model based on electron microscopic imaging studies (Birkenh~iger et aL, 1995) places them on the periphery of the c-subunits in the Fo com-

155

plex (Fig. 5.16). The latter model also assumes that 12 c-subunits are present in the complex, since the transport of four protons is believed to be coupled to synthesis (or hydrolysis) of one ATP, and each c-subunit is viewed as helping to transport only one proton during a given transport cycle (Fillingame, 1997). Moreover, it is proposed in the binding change model for ATP synthesis (or proton pumping) that F-type ATPases contain three sites of ATP synthesis (or hydrolysis). Accordingly, the presence of 12 c-subunits in Fo would provide the requisite number of sites for proton transport. It is, however, somewhat difficult to envision how the single a-subunit could be peripherally located rather than centrally located among the 12 c-subunits. The a-subunit appears to interact with each of the c-subunits to catalyze proton translocation (see next Section); such interactions could occur most easily if the a-subunit were in a central rather than a peripheral location. It is conceivable that each c-subunit could rotate past the

FIGURE 5.16 Longitudinal section of the structure proposed for the E. coli FoFIATPase complex showing the locations of amino acid residues in various subunits in relation to each other. In the model, the a-subunit is shown to be located peripherally to 12 c-subunits in the Fo membrane sector of the protein complex, although some investigators still propose a more central location for the a-subunit. Also shown are the three extended a-helices of the y-subunit that are believed to be important to the transfer of conformational changes between the sites of proton transport across the a- and c-subunits and the sites of ATP synthesis (or hydrolysis) on the fl-subunits. During operation of the enzyme two "catches" or attachment points are proposed to form, the first between the side-chains of residues 268 and 269 of the y-subunit and the/3-subunit that has an empty (L) nucleotide binding site and the second between the horizontally inclined a-helix of the y-subunit (containing residue yC87) and the/3subunit that has a tight (H) nucleotide binding site (see text for discussion of other residues shown) (adapted from Fillingame, 1997, with permission from Company of Biologists Ltd.).

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5. Structure and Function of Transport Proteins That Form Solute Gradients

a-subunit on the periphery of the complex rather than, say, the a-subunit in the center of the complex rotating past each of the c-subunits surrounding it. Consideration of the free energy required to produce either of these types of rotations within the membrane may help to determine whether either or both of the proposed structures of the Fo complex are feasible. Although the proposed structure of the intact Fo sector is still highly speculative, progress is being made in regard to the relationship of the c-subunits to each other (Groth et al., 1998), and the structure of the c-subunit monomer in the membrane is now well established (Fillingame, 1997). The hairpin structure of the c-subunit is shown in Fig. 5.17 beside a conjectural arrangement for the portion of the a-subunit that is thought to be involved in proton translocation. Both the number of transmembrane segments in the a-subunit and its orientation in the membrane remain to be determined. The results of biochemical, genetic, nuclear magnetic resonance, and infrared spectroscopic studies have been used to construct a likely pathway of proton transport across the membrane via the a- and c-subunits of the Fo sector of F-type ATPases.

FIGURE 5.17 Functionally important amino acid residues in subunit c of the E. coli F-type ATPase. Also shown is the C-terminal region of subunit a, the structure of which is largely unknown. In fact, the conjectural orientations of the transmembrane helices of subunit a that are shown would need to be reversed to accommodate the model of proton transport described in the text. Residues shown in diamonds are believed to be essential for transport, and residues in boxes appear to be functionally important. Note also that the side-chain of Ala 24c is actually close enough to the side chain of Asp 61c to be in van der Waals contact with it (see also Fig. 5.18). Consequently, when the carboxyl group is moved from residue 61 of subunit c to residue 24 of this subunit through site directed mutagenesis the function of the enzyme is not lost (see text). Third site mutations at the residues shown in large circles enhance the activity of the enzyme in which the carboxyl group has been transferred from residue 61 to residue 24. Abbreviations: A, alanine; D, aspartate; E, glutamate; G, glyine; H, histidine; I, isoleucine; K, lysine; L, leucine; M, methionine; Q, glutamine; R, arginine; Y, tyrosine (adapted from Fraga et al., 1994, with permission from American Society for Biochemistry and Molecular Biology).

2. A Proposed Proton Pathway The carboxyl group of the aspartyl residue at position 61 in the second of the two transmembrane helices of subunit c (Asp61c) has for some time been viewed as the proton binding site of the F-type ATPase in E. coli (Fig. 5.17). The carboxyl group of a glutamyl residue at the same location in all other F-type ATPases serves a similar function (Nakamoto, 1996). Moreover, this proton binding site probably lies between packed pairs of adjacent c-subunits (Girvin et al., 1998). It now appears that a proton actually is transferred between the side chains of several residues before it reaches the anionic one on the residue at position 61. The proton is then transferred to the side chains of several more residues before its transport is complete. Nevertheless, the side chain carboxyl group of Asp61c of E. coli is a useful place to begin our discussion. Although Asp61c is a hydrophilic amino acid residue, it is present in the hydrophobic center of the membrane. Presumably, the otherwise highly hydrophobic transmembrane helices of the c-subunit stabilize Asp61 within the membrane as shown in Fig. 5.17. Apparently because it is present in a hydrophobic environment, the pK~ of the carboxyl group of Asp61c is about 7 rather than the lower value that it would be if it were in a more hydrophilic environment. In fact, it is proposed that the carboxy group actually takes on a lower pKa value as well as the one near 7 during the transport cycle of F-type ATPases (see Section III,D below). Interestingly, in the three-dimensional structure of the c-subunit (Fig. 5.18), Asp61 may be seen as close enough to the alanyl residue at position 24 to be in van der Waals contact with it (Girvin and Fillingame, 1995). As one might expect, the use of site-directed mutagenesis to replace Asp61c with a glycyl residue abolishes proton translocation. If, however, Asp61c is converted to a glycyl residue at the same time that Ala24c is replaced with an aspartyl one, the resultant FoF1 doublemutant retains some of its normal function (Miller et al., 1990). That is, the site of the carboxyl group can be shifted from residue 61 to residue 24 without loss of c-subunit function! Presumably the close apposition of the side chains of these resides accounts for the apparently more or less proper positioning of the essential carboxyl group in each case. The enzyme with an aspartyl residue both at position 61 and at position 24 is also functional, although it displays an unusual pH sensitivity (Zhang and Fillingame, 1994). In order for a proton to reach the position of the carboxyl group of Asp61c it appears first to need to be bound transiently at several other sites on the c- and a-subunits. As for Asp61, the carboxyl group of the alanine residue at the C-terminus of the c-subunit (posi-

FoF1-ATP Synthases (F-Type ATPases)

FIGURE 5.18 Proposed interaction between the two transmembrane c~-helices of the c-subunit of E. coli F-type ATPase. The arrowheads indicate well-defined regions of interaction between Tyr 10 to Ala 25 of helix-1 and Val 60 to Ala 77 of helix-2. The side-chain of the aspartyl (D) residue at position 61 is close enough to the sidechain of the alanyl residue at position 24 (not marked) to be in van der Waals contact with it. The c-subunit of the Fo sector interacts with the F1 sector via its loop between the two helices shown at the top of the diagram (adapted from Fillingame, 1997, with permission from Company of Biologists Ltd.).

tion 79) has a higher-than-anticipated pKa value. Since this conserved residue lies outside the hydrophobic region of the membrane and does not face Fa (Fig. 5.17), it is unlikely to fall in a hydrophobic environment. Hence, the greater-than-normal pKa value of its carboxyl group indicates that it is involved in hydrogen bonding (Brzezinski et al., 1992). The glutamyl residue at position 219 of the a-subunit is also known to be

157

essential for proton conduction. These results are consistent with the conclusion of Bartl et al. (1995) that the carboxyl group of Ala79c, the phenolic group of the conserved tyrosyl residue at position 10 of the c-subunit, and the carboxyl group of Glu219a form a hydrogenbonded chain that in infrared spectroscopic studies shows large proton polarizability owing to collective proton fluctuation. The investigators propose that these functional groups form the start of a hydrogen-bonded pathway along which protons are translocated. 8 The results of infrared spectroscopy also indicate that the next two hydrogen bonds in the pathway are between a carboxy group and the imidazole group of a histidyl residue (Bartl et al., 1995). The histidyl residue is probably at position 245 in the a-subunit, which is known from site-directed mutagenesis to be needed for normal proton translocation. Hence, the functional groups on His245a, Glu219a, Tyrl0c, and Ala79c appear to transfer protons to the carboxyl group of Asp61c via hydrogen bonding. The functional groups of three other residues and structural water are proposed then to receive the protons from the carboxyl group of Asp61c to complete the transport process. In the present model, an essential guanidino group of the arginyl residue at position 210 of the a-subunit is proposed to help to transfer protons from the carboxyl group of Asp61c to the F1 sector (Bartl et al., 1995). To correspond to this model, the putative transmembrane helix of the a-subunit containing this Arg210 residue and the Glu219 residue discussed above would probably need to be in the reverse orientation of the entirely conjectural one shown in Fig. 5.17. The side-chain functional groups of the conserved lysyl and arginyl residues at positions 34 and 41, respectively, of the c-subunit also appear to be involved through hydrogen bonding in the transfer of protons from the carboxyl group of Asp61c to the F1 sector (Bartl et al., 1995). Since the total number of hydrogen bonds formed by the side-chains of the residues described above are, however, insufficient to traverse the 6-nm-long-Fo complex, structural water is also proposed to comprise part of the transmembrane pathway. In addition, it has been proposed that dihydrogen phosphate may be involved in the translocation of protons instead of Lys34 of subunit c (Bartl et aL, 1995). As discussed in Chapter 3, we find the latter possibility particularly appealing since such a phosphate should, 8 Some authors have concluded that since Na + as well as H + gradients drive ATP synthesis in certain bacteria, hydrogen bonding cannot account for the pathway of proton transport. While it is clear that hydrogen bonding is not required for ATP synthesis in these bacteria, available data support the conclusion that hydrogen bonding does occur when protons are transported. In addition, Na + as well as H + ions could temporarily neutralize the negative charge on Pi 1- as we proposed in Chapter 3 and, thus, help to appose it to the charges on ADP 3-.

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5. Structure and Function of Transport Proteins That Form Solute Gradients

upon receiving a proton, be transiently easier to appose to the negatively charged A D P 3- for ATP synthesis. Our theory is in contrast to the proposal of Bartl and associates (1995) that the free energy that would be gained by dihydrogen phosphate is used for the translocation of subunit c along subunit a. Of course, these hypotheses are not mutually exclusive if the free energy needed for the proposed chemical and physical changes does not exceed the free energy of proton transport. In this regard, the movement of the c-subunit along the a-subunit could require considerable free energy if the a-subunit is peripherally located in Fo and the c-subunits rotate past it (Fig. 5.16). Recent evidence indicates, however, that the binding change model of thermodynamic coupling and the conformational changes proposed to be involved in it may need significant revision. In particular, the new data indicate that cooperativity may not exist among the three pertinent nucleotide binding sites in the F1 sector (Reynafarje and Pedersen, 1996) (see below). Since cooperativity usually involves communication through changes in protein conformation, the sizes of the conformational changes in the F1 sector may be smaller than if cooperativity were to occur. If the associated conformational changes have been exaggerated also for the Fo sector, then an a-subunit located centrally among the c-subunits rather than peripherally to them should minimize the extent of movement needed for the a-subunit to interact sequentially with each c-subunit. Before we consider further the binding change model and possible modifications of it for the coupling of ATP synthesis or hydrolysis to proton transport (Section III,D below), we describe the structure of the Fa sector and the chemical change that it catalyzes. In contrast to the Fo sector for which structure and movement are highly speculative, much data support the proposed structure of the Fa sector and the conformational changes that it appears to undergo. C. The Cytosolic Sector F1 Catalyzes Chemical Change 1. Subunit Structure of F1

The structures of the E. coli and the animal F-type ATPases are pictured in Fig. 5.19A. Viewed in cross section, the hexagonal subunit array of the F1 sector is particularly evident (Fig. 5.19B). While the three a- and the three/3-subunits comprise the bulk of the F1 sector, the single 3'- subunit is very important to the structure of F~ and its functional association with Fo (Pedersen, 1996). In this regard, the y-subunit is prominent in the "stalk region" connecting the "basepiece" of Fo and the "headpiece" of F1 (Fig. 5.19A). While the stalk

FIGURE 5 . 1 9 Similarities in the proposed structures of E. coli and mitochondrial (animal) F-type ATPases. (A) Subunits of the membrane-bound Fo sector comprise the basepiece, while the headpiece is composed entirely of subunits of the extramembrane F1 sector. Some subunits of both sectors extend into the stalk that connects the headpiece and basepiece. In these models, the entire 3,-subunit is believed to rotate and so the C-terminal end of its long a-helix must rotate within a hydrophobic sleeve formed in the headpiece by the Ot3/~3 heterohexamer. The positions and stoichiometries of the subunits listed at the bottom of A are not yet known. (B) Two proposed structures for the headpiece of the mitochondrial F-type ATPase shown in cross section. The structure on the left has been proposed based on X-ray crystallographic studies, whereas the structure on the right is predicted by Pedersen (1996) to occur under more physiological conditions than those used to crystallize the bovine heart F1 sector. The /3-subunits are believed to have different conformations (shown arbitrarily as different shapes) at a given moment according to the model on the left, whereas no such difference is proposed according to the model on the right (see text for further discussion) (adapted form Pedersen, 1996, with permission from Plenum Publishing Corp.).

regions of the E. coli and mitochondrial enzymes contain portions of additional subunits of the F1 and Fo sectors, our discussion of the function of the whole FoF~ complex will involve only one of these additional subunits, namely, the bacterial e-subunit to be discussed in Section III,D below. Before we consider catalysis by the F1 sector and the whole complex, let us examine briefly

FoF1-ATPSynthases (F-TypeATPases) important structural details of the catalytic/~-subunits and their associated ~-subunits in F1.9 The structure of the nucleotide-binding site on the /3-subunit and its catalytic function have been studied in great detail (Weber and Senior, 1997). The adenylbinding subdomain of the E. coli/3-subunit consists of the side chains of amino acid residues Tyr331, Phe404, Ala407, Phe410, and Thr411. Tyr331 helps to provide a nonpolar environment for the ring of the adenyl group with which it is stacked. All of these amino acid residues are in or near the third helix of the C-terminal domain of the/3-subunit (Weber and Senior, 1997). While there appear to be no specific interactions of the residues of the/3-subunit with the ribosyl group of A D P and ATP, we shall see that there are many important interactions with phosphoryl groups. As in numerous nucleotide-binding proteins, a P-loop surrounds the phosphoryl groups at the catalytic site of the /3-subunit (Weber and Senior, 1997). The loop in F1 of E. coli consists of residues 149 to 156 in the/3-subunit ( G G A G V G K T ) . The main-chain atoms of Gly152, Val153, and Gly154 and the side chains of Lys155 and Thr156 appear to make contact with the phosphoryl groups. Of these residues Gly154, Lys155, and Thr156 are essential. Even an arginyl residue cannot be substituted at position 155, while only a seryl residue is tolerated at position 156. Lys155 appears to lie close to the y-phosphoryl group of A T P or to Pi and to form specific hydrogen bonds with one or the other of these substrates, depending on the direction of the reaction. Lys155 contributes significantly to the free energy of binding of A T P or Pi and thus it probably contributes to completion of the catalytic cycle. The side-chain oxygen of Thr156 probably forms a coordination site for binding of the Mg 2+ of M g A T P or M g A D P (Weber and Senior, 1997). 1~ Among six other residues of the fl-subunit that are near the phosphoryl groups of the bound nucleotide (i.e., residues Glu181, Arg182, Glu185, Met209, Asp242, and Arg246), Glu181 is particularly important to the pertinent catalysis (Weber and Senior, 1997). The carboxyl group of Glu181 appears to lie about 4.4 A from the y-phosphoryl group of ATP. Consequently, this carboxyl group may activate a water molecule for nucleophilic attack on the phosphoryl group and therefore help to catalyze A T P hydrolysis or, conversely, A T P 9We may at times refer to the F1 sector and the/3-subunits in it as the sites of catalysis in FoF1 ATPases. We remind the reader, however, that proton transport also requires catalysis.Hence, we mean that the F1 sector and the/3-subunits are the sites of chemicalcatalysis. 10For simplicity here we refer in most instances simply to ATP or ADP rather than to the actual substrates which form complexes with Mg2+.

159

synthesis. 11 In this regard, replacement of Glul81 with an alanyl or a glutaminyl residue reduces the rate of the hydrolytic step of the catalytic cycle by two orders of magnitude, whereas the reverse step in A T P synthesis is slowed by three orders of magnitude (reviewed by Weber and Senior, 1997). Nevertheless, the mutant enzymes still accelerate hydrolysis of A T P by a factor of 106, so the side chains of other residues also contribute significantly to catalysis. Mutation of either Met209 or Arg246 also impairs catalysis, apparently owing to the proximity of their side chains to the side chain of Glu181. In addition to residues in the/~-subunit, several residues in the noncatalytic c~-subunit (Fig. 5.19) also are needed for normal catalytic activity (Weber and Senior, 1997). Site-directed mutation of each of the residues Ser347c~, Gly351~, Ser373c~, Ser375a, and Arg376c~ in the E. coli protein greatly reduces catalytic activity. Interestingly, these mutations each affect specifically multisite A T P hydrolysis by F1, whereas unisite hydrolysis, which occurs at substoichiometric M g A T P concentrations, is not greatly affected. These mutations of the c~-subunit appear therefore to interfere with the interactions between the c~- and /3-subunits at their mutual interfaces. As a consequence, the mutations are believed also to disrupt the interactions among the/~-subunits. The interactions between the c~- and/3-subunits have, therefore, been proposed to be required for positive catalytic cooperativity among the /3-subunits (Weber and Senior, 1997). This cooperativity is a fundamental tenet upon which is based the binding change mechanism for operation of F-type ATPases to be discussed in Section III,D below.

2. ATP Synthesis and Hydrolysis Nucleotides bind to each of three catalytic sites on the /3-subunits near one of their interfaces with c~-subunits of the F1 sector (Pedersen, 1996). Nucleotides also bind to noncatalytic sites on each of the three a-subunits, although binding of nucleotides to these noncatalytic sites is not needed for activity, at least in the case of the E. coli enzyme. In the binding change model for A T P synthesis (Boyer, 1993), nucleotide-binding sites on each of the three/3-subunits are envisioned to be different (left side of Fig. 5.19B). A T P (or the unbiological and nonhydrolyzable A T P analog AMP-PNP), occupies a tight binding site on one/3-subunit, A D P and Pi occupy a loose binding site on a second/3-subunit, and 11Because it is not yet certain whether the conformation of the F1 complex that has been studied in crystalsis a physiologicallynormal one, it is believed to be premature to conclude that the structure supports or does not support a particular mechanism of chemical catalysis (Weber and Senior, 1997).

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5. Structure and Function of Transport Proteins That Form Solute Gradients

the binding site on the third/3-subunit lies open. In the binding change model, the affinities of each of the three binding sites for nucleotides changes sequentially and cooperatively as a result of proton transport. Since all three nucleotide-binding sites appear actually to be occupied most of the time, their designations have been changed by some authors from "open," "loose," and "tight" to "L," "M," and "H," respectively (Weber and Senior, 1997). The different shapes of the /3-subunits depicted on the left side of Fig. 5.19B are meant to convey this difference among them in their conformations and in their relative affinities for nucleotides. For ATP synthesis to occur, ADP and Pi are believed to bind to an open or "L" site (Boyer, 1993; Weber and Senior, 1997). The loose or "M" binding site subsequently formed is proposed then to undergo transition to a tight or "H" site owing to proton translocation and associated enzyme conformational changes. The ratio of ADP to ATP at the tight binding site is near unity, and hence the change in affinity of the binding site for nucleotides is believed to drive the formation of ATP (Boyer, 1993; Weber and Senior, 1997). Subsequently, the ATP is released from the tight binding site apparently also as a result of proton transport and enzyme conformational changes. The cycle could, of course, be reversed at higher ATP concentrations or smaller proton gradients, movement being initiated by ATP binding to an open site. Subsequent hydrolysis of ATP would result in the reverse set of conformational changes, changes in the affinity of nucleotide binding, and translocation of protons in the opposite direction. The change in free energy associated with the change in binding affinity has been shown to be large enough to account fully for the transfer of free energy between a proton gradient and a phosphoric acid anhydride bond (Weber and Senior, 1997). It has not yet been determined, however, whether the associated protein conformational changes also involve large enough changes in free energy to be the only link needed in the process through which free energy is transferred. D. The Binding Change and Related Models of Thermodynamic Coupling b e t w e e n Proton Transport and ATP Synthesis/Hydrolysis Involve Coordinate Conformational Changes in the Fo and F1 Sectors There remains little doubt that F-type ATPases undergo conformational changes during ATP synthesis and during the reverse process in which a proton gradient is formed at the expense of ATP hydrolysis. It remains an open question, however, whether these conformational changes involve large enough changes in free

energy to provide the only direct link between the free energy in the proton total chemical potential gradient and that in phosphoric acid anhydride bonds. The model shown on the right of Fig. 5.19B is consistent with our theory that dihydrogen phosphate could receive transiently a transported proton and hence bind near ADP 3without experiencing electrostatic repulsion. As pointed out by Weber and Senior (1997), the presence of a proton gradient increases the affinity of the F-type ATPase for Pi by several orders of magnitude without affecting its affinity for ADP. Formation of a covalent bond between ADP 3- and phosphoric acid prior to dissociation of a proton from it could then be part of the mechanism by which free energy in a proton gradient is converted to that in phosphoric acid anhydride bonds. Although such additions to or modifications of the binding change hypothesis continue to arise, this hypothesis is still the most widely accepted model for ATP synthesis by F-type ATPases. For this reason, we present below a brief description of the binding change mechanism and a proposed structural basis for it. Additional information and discussion may be found in several excellent reviews (e.g., Boyer, 1993; Pedersen, 1996; Nakamoto, 1996; Fillingame, 1997; Weber and Senior, 1997; Kagawa and Hamamoto, 1997). According to numerous studies, nucleotides exhibit negative cooperativity when binding to the three catalytic sites on the/3-subunits of F-type ATPases. Conversely, catalysis is promoted upon binding of more than one nucleotide. Boyer (1993) discussed the possibility that this positive catalytic cooperativity and the associated negative cooperativity for binding may obscure each other and, hence, come falsely to resemble Michaelis-Menten kinetics. For these reasons, it is desirable in experiments with F-type ATPases to measure binding and catalysis independently. In a recentstudy, however, cooperative catalysis was not observed when the investigators were careful to measure actual rates of ATP hydrolysis rather than rates of nucleotide dissociation from the enzyme (Reynafarje and Pedersen, 1996). If these differences are confirmed, the binding change hypothesis may require considerable revision. In the binding change model, the previously observed cooperative catalysis is interpreted to mean that ATP is synthesized at all three of the catalytic sites, although the sites are proposed to be at different stages of the catalytic cycle at any given moment (Fillingame, 1997; Weber and Senior, 1997). For example, at the time that a nucleotide binds to an open (L) site, ADP is bound at a loose (M) site and ATP is bound at a tight (H) site. During ATP synthesis, each of the three sites is believed to be converted sequentially from an open to a loose and then to a tight nucleotide binding site. In this Scheme, the transport of H + ions along their total chemi-

FoL-ATP Synthases (F-Type ATPases)

cal potential gradient produces conformational changes in Fo and consequently in Fa that lead to tight ADP and Pi binding, ATP synthesis from the tightly bound substrates, and then release of ATP from what had been a tight binding site. The deviation from threefold symmetry that is predicted by the binding change model was actually observed for the/3-subunits in X-ray crystallographic studies of the bovine heart mitochondrial Fa sector (Abrahams et aL, 1994), although the physiological significance of this observation has been questioned by Pedersen (1997) among others (e.g., Weber and Senior, 1997). Each/3-subunit has a different conformation presumably because it comes into contact with the centrally located but asymmetric y-subunit differently (left side of Fig. 5.19B). The nucleotide binding site appears literally to be opened by movements of strands in a/3 pleated sheet of the/3-subunit that lie proximal to the binding site and by more global movement of up to 2 nm of the C-terminal third of the/3-subunit, which contains abundant c~-helical structure (Fillingame, 1997). Rearrangement of hydrogen-bonding is believed to form a "catch" or attachment point between a displaced loop of the/3-subunit that has the open nucleotide binding site and residues near the C-terminus of the y-subunit (Fig. 5.16). A second "catch" is believed to be formed between the horizontally inclined helix of the y-subunit (Fig. 5.16) and the C-terminal helical domain of the/3subunit that has a tight (H) binding site. It is proposed that the conformational changes in the/3-subunits result from rotation of the y-subunit in a hydrophobic sleeve surrounding its long C-terminal c~-helix (Abrahams et aL, 1994). The sleeve is formed by the 0/3/3 3 hexamer of F~ (Fig. 5.19). The rotation of the y-subunit is thought to be driven by proton transport, which results in the sequential changes in the nucleotide binding sites and ATP synthesis as described above. In this regard, nearly full (i.e., >280 degrees) (Sabbert et aL, 1996, 1997) or completely full (Noji et aL, 1997) unidirectional ATPdriven rotation of the y-subunit has been observed to occur within the rest of the isolated and immobilized F~ sector. Presumably, this rotation and the associated functioning of the two "catches" can occur in either direction depending on whether ATP is synthesized utilizing a proton gradient or hydrolyzed to produce one. Since the ~- and/3-subunits of F1 lie about 4.5 nm away from the surface of the membrane, whereas the ysubunit extends from the sleeve formed by c~3/33to the c-subunits of Fo (Fig. 5.19A), it is logical to propose that the free energy in a proton gradient may be converted to a mechanical change in 3' in order to connect proton transport across Fo to ATP synthesis on the/3-subunits of Fa. The rotation of y that is proposed to result from

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proton transport could lead to conformational changes in the/3-subunits and consequently ATP synthesis. Since proton transport occurs in association with the c-subunits in Fo, studies have been designed to determine how these subunits may interact with a proposed ye complex in the E. coli F-type ATPase. The polar extramembrane loops of the c-subunits face Fa and undoubtedly encompass the sites of their interaction with the ye complex (Fig. 5.17). Conversion of the glutaminyl residue at position 42 of the c-subunit (Gln42c) to a glutamyl residue results in an FIFo complex with normal ATPase activity, but that activity is uncoupled from proton transport (Mosher et al., 1985). Uncoupling may result from the inability of the F1 sector to block passive proton transport on binding the mutant Fo sector. Conversion of the arginyl residue at position 41 (Fig. 5.17) to a lysyl residue resulted in a similar phenotype, and varying degrees of uncoupling were also observed for other mutations at position 42 and for mutations at the conserved prolyl residue at position 43 (Fillingame, 1997). Mutations in the ye-complex that suppress the uncoupling caused by mutations of Gln42c are believed to be pertinent to understanding the connection between proton transport and ATP synthesis/hydrolysis in the binding change model. Fillingame and associates (Zhang et al., 1994) found only four such mutations upon screening bacteria for them, and all four mutations lay at position 31 of the e-subunit (Fig. 5.16). These same investigators also tested for the apposition of residue 31 of the e-subunit to residues in subunit c by converting residues in each subunit to cysteinyl residues and then testing whether they formed disulfide bonds upon oxidation. Such crosslinking occurs between position 31 of the e-subunit and positions 40, 42, and 43 (but not 39) of subunit c (Zhang and Fillingame, 1995). Similarly, subunit crosslinking can be produced between positions 42, 43, and 44 of subunit c and position 205 of the ysubunit (Fig. 5.16). That the y-subunit may function in association with the e-subunit in E. coli is supported by several additional studies (reviewed by Fillingame, 1997). First, formation of a "ye complex in vivo is supported by the observation that these subunits bind to each other in vitro. Moreover, their region of interaction has been defined by crosslinking studies similar to those just described. These proteins can be so linked in the region of position 205 of the y-subunit and the N-terminal domain of e. Although the C-terminal domain of subunit e interacts with the c~- and/3-subunits, deletion of the C-terminal domain of e does not result in loss of function. For these reasons, it is proposed that the ye-complex moves as a unit from one c-subunit to the next as protons are transported by them (Fillingame, 1997). This motion is thought to

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5. Structure and Function of Transport Proteins That Form Solute Gradients

generate torque on the y-subunit, causing it to rotate within the hydrophobic sleeve formed by the a3/~3 hexamer of F1. According to this proposed structural basis for the binding change hypothesis (Fillingame, 1997), the ysubunit inside the a3/~3 complex would rotate 120 ~ as each tightly bound ATP is released during ATP synthesis. Each such rotation would establish two new "catches" (attachment points described above) between y and the two/3-subunits that have just formed open and tight nucleotide binding sites (Fig. 5.16). Since it is assumed here that there are 12 c-subunits per FoF~ complex and because transport of four protons appears to be associated with formation of each phosphoric acid anhydride bond, each 120 ~ rotation is thought to be driven by the migration of four protons across four adjacent c-subunits. In this model (Fillingame, 1997), subunit c is proposed to form a complex with ye only when the carboxyl group of Asp61c is in its low pKa form. (The influence of the hydrophobic environment of this carboxyl group on its pKa value was discussed in Section III,B,2 above.) For ATP synthesis, the low pKa form of Asp61c would receive protons from the side of the membrane on which the total chemical potential of H + ions is the highest (i.e., the outside). Association of subunit c with the a-subunit could conceivably be needed for production of the low pKa form of Asp61c. Movement of subunit c away from the a-subunit (and another c-subunit into association with it) would then convert Asp61 on the first c-subunit to the high pK, form (and Asp61 on the incoming c-subunit to the low pKa form). This conversion of Asp61 to the high pK, form (or release of a proton from it to the side of the membrane on which the H + total chemical potential is lowest) is believed to result in a change in the loop structure of the c-subunit so that it dissociates from the ye-complex. Subsequent movement to the next c-subunit would be unidirectional presumably because of an asymmetric orientation of the ye-complex in relation to subunits a or b. After transport of the fourth proton and concomitant completion of rotation of 120 ~ around Fo, enough torque is presumed to have been generated also to have driven the 120 ~ rotation of the y-subunit within the c~3/33hexamer. As a consequence of the rotation and the associated ATP synthesis, "catches" between the y-subunit and the/3subunits with open (L) and tight (H) nucleotide binding sites would be broken and new catches formed with newly formed open (L) and tight (H) nucleotide binding sites. Concomitantly, ATP would be released from the previously tight-binding site. An alternative to the proposed smooth 360 ~ operation of the Fo sector is for it to produce torque on the ye complex 120 ~ at a time (Fillingame, 1997). To pro-

duce this torque, protons might not be released from high pKa forms of subunit c until the y-subunit is ready to form new catches. In this Scheme, transition of the carboxy group of Asp61c from the low to the high pKa form would occur as described above, but a proton would not be released immediately from it. Rather, the next three c-subunits would each move in turn into an association with the a-subunit. Because the protons would not dissociate from the carboxyl groups of four Asp61c residues until they are ready to rotate the yecomplex, the free energy of the four protons might be stored so that they could dissociate together to produce the 120 ~ rotation. The ye-complex would then experience no torque and hence remain more or less stationary until four more protons had been added and were ready to dissociate from carboxyl groups on the Asp61 residues of four adjacent c-subunits. A central rather than peripheral location of the a-subunit among the c-subunits shown in Fig. 5.16 would appear to be more compatible with such a model, since residues in the asubunit may be needed to complete proton transport (Fig. 5.17 and Section III,B,2 above). In part because the structure of the Fo sector is not yet known in detail, it is also not known whether part or all of it may rotate with the ye-complex. As just discussed, the c-subunits in Fo could move to produce torque on the ys-complex or a more centrally located a-subunit could rotate with ye while the c-subunits remain stationary. Alternatively, the a3/33-complex may actually rotate owing to conformational changes in Fo transmitted to it via the ~/e complex. Regardless of which subunits of the F1Fo complex form the stator and which form the rotor, one may still grasp the machinelike manner in which the whole complex appears to function. The reader may wish to review other scholarly accounts of the binding change model and the structural changes in F-type ATPases that appear to underlie it (e.g., Boyer, 1993; Pedersen, 1996; Nakamoto, 1996; Fillingame, 1997; Weber and Senior, 1997; Kagawa and Hamamoto, 1997). Central to the hypothesis is the proposal that the y-subunit, in association with other subunits of the F1 complex, helps to relay conformational changes from the c- subunits of Fo to the/~-subunits of F1. In the form of the model presented here, the transport of four protons across four of the 12 c-subunits proposed to be present in Fo results in rotation of by 120 ~ and in conformational changes in all three/3subunits. Nevertheless, only one newly synthesized ATP is released per turn of 120 ~. The same process might proceed with more subtle conformational changes in the/~-subunits if additional mechanisms help to account for the transfer of free energy. For example, direct but transient association of a transported proton with dihydrogen phosphate would make the resultant electrically

FoF1-ATPSynthases (F-Type ATPases)

neutral phosphoric acid molecule more likely to bind near the nucleotide binding site containing ADP 3-. Dissociation of the protons after formation of a phosphoric acid anhydride bond between ADP and Pi would help to complete the transfer of free energy to this bond. While all F-type ATPases are believed to function by the same general mechanism, different species have different metabolic needs. Presumably the form of the enzyme that evolved to function in the cells of different species helps the cells to perform best metabolically. Similarly, the form of the enzyme in different organelles in the same species may need to be different as is the case for chloroplasts and mitochondria in the same variety of plant. In fact, even the same organelle is known in some cases to express a slightly different F-type ATPase in different cells of the same organism. This difference in the mitochondrial F-type ATPase results from alternative splicing of a transcript encoding the y-subunit of the F1 sector. E. Isoforms, Subisoforms, and Alternative Splicing Metabolism varies among tissues and organs in part because they express different forms of the same enzymes. Such isoforms of proteins may be encoded by homologous genes or they may be produced in different cells through alternative splicing of the transcript from a single gene. In keeping with definitions used in this volume, we refer here to isoforms produced through alternative splicing as subisoforms even in cases where homologous genes encoding isoforms do not appear to exist. In this regard, there appears as yet to be no evidence for the expression of isoforms of any of the subunits of F-type ATPases. Nevertheless, alternative splicing has been observed to produce slightly different subisoforms of the critically important y-subunit of the F-type ATPase in human and bovine mitochondria (Matsuda et ai., 1993a,b). Similarly, such subisoforms have been detected for some of the subunits of V-type ATPases. We describe these subisoforms of F- and V-type ATPases in the following subsections. In the case of Vtype ATPases, we will also discuss briefly the expression of isoforms of subunits from different genes, since, in contrast to F-type ATPases, such isoforms have been observed for some of the subunits of V-type ATPases.

1. Subisoforms of the v-Subunit of F-Type ATPases Two subisoforms of the y-subunit of the mitochondrial F-type ATPase have been detected in several mammalian species (Matsuda et al., 1993a,b). The "liver" subisoform is expressed not only in hepatocytes but

| 63

also in other tissues and organs including brain, thyroid, spleen, pancreas, kidney, and testis. It is, however, not expressed in heart, skeletal muscle, or diaphragm, which express instead the "muscle" subisoform. Both subisoforms appear to be expressed in skin, stomach, intestine, and aorta, although it has apparently not been determined whether the subisoforms have similar or different tissue distributions within these organs. During processing of the gene transcript encoding the y-subunit, the 9th of 10 exons is deleted during production of the mRNA encoding the muscle isoform (Fig. 5.20). Interestingly, the only change in the protein that results from this deletion of exon 9 is loss of the C-terminal aspartyl residue which is present in the liver subisoform but not in the muscle one (Fig. 5.20). Both exon 9 and exon 10 contain a guanylate residue at their 5' ends, which completes a codon encoding a leucyl residue (Matsuda et al., 1993a,b). This codon is followed by a termination codon in exon 10, whereas the termination codon is preceded by one encoding an aspartyl residue in exon 9 (Fig. 5.20). Now, whether the C-terminal residue of the y-subunit has one (leucine) or two (aspartate) carboxyl groups is significant because the C-terminal domain forms an a-helix, which is believed to rotate inside the hydrophobic sleeve formed by the a3B3 hexamer of the F1 sector (Fig. 5.19). Moreover, the rotation of y inside this sleeve is believed to produce conformational changes needed for ATP synthesis. Proton transfer to dihydrogen phosphate also may occur in association with the rotation of y (see above). In any case, the number of carboxyl groups at the C-terminus of the y-subunit within the hydrophobic sleeve should have considerable consequences for the function of the ATPase and the degree to which its activity is influenced by pH. Perhaps for these reasons, low pH induces expression of the muscle subisoform of the y-subunit in cultured human sarcoma cells (Endo et al., 1994). The induction is blocked by cycloheximide and by protein kinase C inhibitors. Since increased needs for free energy frequently are associated with a decrease in pH, induction of expression of the muscle subisoform by a decrease in pH may permit a more rapid metabolic rate. The metabolic rate of muscle often must increase dramatically, and the FoF~ complex containing the muscle subisoform of the y-subunit responds to these increased metabolic needs. In contrast, the additional carboxyl group at the C-terminus of the liver subisoform of the y-subunit may limit the turnover number of the whole enzyme too much to accommodate the large changes in metabolic rate required in muscle. Nevertheless, a decrease in pH could conceivably increase the maximum activity of the liver enzyme more than the one in muscle, although the maximum activity of the muscle form

164

5. Structure and Function of Transport Proteins That Form Solute Gradients

Liver - type Heart/skeletal muscle-type

Liver - type

~ ~-~ 9~,,....~10

I

1 234

III

1 234

Heart I skeletal muscle - type

5 6 7 8 9 10

I I I i ,~ / TAA 56

7 8 10

I-P~

poly A

TAA

Liver- type

*NH3-~

DI_~J-COO-

298 aa

Heart I skeletal

+NHz-~

J-COO-

297 aa

muscle - type

presequence

conse"~ed region

FIGURE 5.20 Diagramof the introns and exons of the gene encoding the y-subunit of mammalian F-type ATPase (top) and the two different mRNA (middle) and protein (bottom) molecules that result from alternative splicing of the primary transcript. The presence of an extra codon before the termination codon in exon 9 as compared to exon 10 results in a liver-type protein with one more amino acid residue (Asp 298) than the muscle-type protein. Otherwise, the two subiosforms are identical, and the C-terminal residue in the muscle-type protein is leucine. Because the C-terminus of the y-subunit appears to rotate within a hydrophobic sleeve formed by the O~3~3 heterohexamer (Fig. 5.19), rotation of the subunit within the sleeve should be influenced by whether there are one (leucine) or two (aspartate) carboxyl groups on the Cterminal residue. A large portion of the C-terminus of the y-subunit is also highly conserved among species (see text for further discussion) (adapted from Endo et al., 1994, with permission from American Society for Biochemsitry and Molecular Biology).

would still be higher owing to titration of two rather than one carboxyl groups on the C-terminus of the y-subunit of the liver enzyme. Association of the carboxyl groups with protons would presumably make the C-terminus of the y-subunit easier to accommodate within the hydrophobic sleeve formed by the Og3fl3 hexamer. In support of this concept, the reader may recall from Section III,B,2 above that the c-subunit of the Fo sector is still functional when it has an extra carboxyl group at its proton "binding site" in the hydrophobic interior of the m e m b r a n e , but in this case the enzyme has an unusual sensitivity to inhibition by an increase in pH.

2. Isoforms and Subisoforms of Subunits of V-Type ATPases As discussed above, F-type ATPases are expected to vary somewhat from tissue to tissue because of their different requirements for A T P synthesis. V-Type ATPases are, however, expected to vary not only among different tissues, but even within the same cell, since differences probably exist among intracellular organ-

elles and vesicles in the nature and extent of proton pumping that is required in them (e.g., see Fig. 3.12 in Chapter 3). Such modest functional differences have, in fact, been demonstrated for A T P - d e p e n d e n t acidification of rat liver endosomes and lysosomes (Van Dyke, 1996). These functional differences among organelles and vesicles could result from expression of different isoforms and subisoforms of several of the subunits in the V1 and Vo sectors of V-type ATPases ( U m e m o t o et al., 1991; H e m k e n et al., 1992; van Hille et al., 1994, 1995; Berkelman et al., 1994; Peng et al., 1994; Bartkiewicz et al., 1995; H e r n a n d o et al., 1995; Nishigori et al., 1998). Because intracellular organelles and vesicles and their requirements for proton pumping are active areas of current research, it is likely that additional isoforms and subisoforms of the subunits in V-type ATPases are yet to be discovered (Van Dyke, 1996).

a. The V1 Sector Isoforms of both the A- and the B-subunits of the heterohexameric e x t r a m e m b r a n e V1 sector (Fig. 3.10 in Chapter 3) have been reported for the V-type ATPases in mammals and birds. Two isoforms of the catalytic

FoFI-ATP Synthases (F-Type ATPases)

A-subunit have been identified in human osteoclastoma cells (van Hille et al., 1995). Only one of these isoforms has been shown to be expressed in normal adult tissues, where the subunit is believed to be ubiquitous. Nucleotide sequences that encode the two isoforms are 73% homologous, and they are 63% homologous even in their most divergent segments (van Hille et aL, 1995). Moreover, both isoforms appear to function in V-type ATPases in osteoclastoma cells, and genes encoding both isoforms appear to be present in normal human cells. For these reasons, it seems likely that expression of both isoforms of the A-subunit eventually will be detected either during development or in normal differentiated cell types. In the ubiquitously expressed isoform of subunit A in the chicken, alternative splicing has been found to generate two subisoforms designated A1 and A2 (Hernando et al., 1995). The A1 and A2 mRNAs are identical except for one exon of either 90 or 72 nucleotide residues, respectively. The 90-residue exon that is unique to the A1 mRNA encodes one of three ATP-binding consensus sequences in A1, whereas the A2 mRNA encodes only the other two such sequences (Hernando et aL, 1995). These different but ubiquitously expressed subisoforms of subunit A are presumed to be important either for the different trafficking or for the different functioning of V-type ATPases that may be needed by different organelles and vesicles. The possible physiological significance of these two subisoforms in the trafficking and functioning of V-type ATPases remains, however, under investigation. Different isoforms of the noncatalytic B-subunit of V-type ATPases have also been identified in barley (Berkelman et al., 1994) as well as in birds and mammals. The different catalytic and pharmacological characteristics of the V-type ATPases of chicken kidney cells and osteoclasts have been attributed to the expression of different isoforms of the B-subunit in these tissues (Bartkiewicz et al., 1995). The isoform expressed in osteoclasts (and referred to as the brain type isoform) appears also to be expressed in a variety of human tissues, whereas the kidney type isoform is expressed in only a few types of cells and organs (van Hille et al., 1994). Interestingly, although alternative splicing has not been observed to generate subisoforms of the B-subunits, the yeast B-subunit appears to be generated by an unusual type of protein processing. In this protein splicing process, amino acid residues 284 to 737 appear to be removed from the precursor protein so that the mature protein is produced subsequently by joining the C-terminal residue 283 of the resultant N-terminal domain to the N-terminal residue 738 of the C-terminal domain (Hirata and Anraku, 1992). Evidence for such processing of the B-subunit in birds and mammals has to our knowledge not been sought. Nevertheless, if the

165

latter type of processing is found to vary for any of the subunits of the V1 sector, then this alternative splicing of proteins could contribute further to the variety of forms that V1 may take. Such further variety appears to result in mammals from the production of heterogenous forms of one other subunit of the V1 sector (Hemken et al., 1992). The brush border membrane of rat kidney epithelia has been found to have a cluster of apparently different forms of subunit E of the V-type ATPase, whereas microsomes do not show this heterogeneity. The heterogeneity cannot be attributed to different types or amounts of glycosylation, phosphorylation, or covalent lipid binding. Genomic screening indicates that two genes encoding the E-subunit may be present in the rat, although it remains to be determined whether both of these genes are functional. Regardless of the structural basis for the observed heterogeneity, the heterogeneity could contribute to variation in function of V-type ATPases. Such variation appears to be needed not only among membranes in different tissues, but also among different membranes in the same cell and even perhaps at different positions within the same membrane.

b. Isoforms and Subisoforms o f Subunits o f the Vo Sector Variety among V-type ATPases may also be generated from isoforms and subisoforms of the subunits in the Vo sector. In this regard, at least two genes appear to encode active c-subunits in the Vo sector of yeast (Umemoto et al., 1991). The two deduced amino acid sequences of the isoforms are only about 57% homologous, so utilization of one or the other or both subunits to form the Vo sector could conceivably result in considerable variation in function. A small multigene family also appears to encode several isoforms of the c-subunit in the oat (Lai et aL, 1991). In the latter case, the deduced amino acid residue sequences of the isoforms are 97 to 99% identical. Multiple isoforms of the Vo c-subunit have also been detected recently in humans (Nishigori et aL, 1998). V-type ATPases also contain subunits for which there is no analog in F-type ATPases, and the Acll5 protein in the Vo sector is one of these subunits. It is a glycosylated membrane protein with several transmembrane segments (Nelson, 1995). While its function is currently under investigation, it is of interest here because tissuespecific subisoforms of the protein are known to be produced through alternative splicing (Peng et al., 1994). One form of the protein is expressed in all five of the rat and bovine organs that have been tested, whereas the other form, which contains a six-amino-acid-residue insert, is expressed predominantly or exclusively in brain. The difference in amino acid sequence occurs in the two subisoforms in a region predicted to have a

166

5. Structure and Function of Transport Proteins That Form Solute Gradients

protease-sensitive motif. Hence, it has been suggested that the subisoforms could have different half-lives (Peng et al., 1994). All such differences in the structures of subunits of V-type ATPases may affect the function or location of the enzymes in cells, but description of the actual effects on function or location and elucidation of the mechanisms by which the effects occur is apt in each case to require much scientific effort.

IV. SUMMARY Many of the known details of the structure and function of P-type and F-type ATPases have been described in this chapter. While the models developed for the thermodynamic coupling between ATP synthesis (or hydrolysis) and ion transport have conjectural components, they are in general founded on a large amount of experimental data and on the resultant ingenious hypotheses of several investigators. All of us need to assess critically these impressive findings and insights in order to understand both what we do and what we do not know about how P- and F-type ATPases catalyze transport. A great deal remains to be learned about these catalytic mechanisms, and our models for the structures and functions of the ATPases will undoubtedly be modified as we learn more about them. In this spirit, the following summary is intended to be provocative of new ideas and not merely a summary of what we know about transport ATPases. To recapitulate from the introduction to this Chapter, striking similarities as well as differences abound in the functions of P-type and F-type ATPases. For example, both types of ATPases have membrane and extramembrane components that are connected by an intervening "stalk" region. Moreover, transfer of conformational changes between these components appears to be on the pathway of conversion between the free energy in phosphoric acid anhydride bonds and that realized in ion gradients. For both types of enzymes, however, it remains to be determined whether their conformational changes alone are large enough to be the only route of free energy transfer. While these general similarities in the structure and function of P- and F-type ATPases have undoubted importance, the differences in the details of their proposed structures and mechanisms of action are striking. The large FoF1 protein complex is proposed to operate as a molecular machine with continuous 360 ~ rotation of the central y-subunit. The rotation is proposed to be driven in one direction by proton migration along its total chemical potential gradient and in the reverse direction by ATP hydrolysis. In contrast, the a-subunits of P-type ATPases are composed of single polypeptide

chains capable of catalyzing both ATP hydrolysis (or synthesis) and ion transport. Hence, continuous rotation of a portion of a P-type ATPase in one direction during catalysis probably is not part of their mechanism of action, unless perhaps rotation occurs around a covalent bond. An observation that seems to have been fundamental to development of the machinelike binding change model of ATP synthesis by F-type ATPases is that the three catalytic subunits of these enzymes show positive catalytic cooperativity. Might not a similar model conceivably have been brought forth for the operation of the P-type ATPases, however, if they had also been found in general to be multiprotein complexes that display catalytic cooperativity? In fact, the reader may recall that some studies indicate that SERCA P-type ATPases function primarily as homodimers and show positive cooperativity between catalytic sites both for C a 2+ transport and for ATP hydrolysis. Were it not for the ability of these and other P-type ATPase molecules also to function apparently as monomers, a more machinelike model could also be proposed for their function. In this case, each a-subunit in the homodimer could operate in turn to catalyze ATP hydrolysis and ion transport rather than their operating, say, in unison or even in random order. Similarly, it is as easy to imagine that the three catalytic subunits of an F-type ATPase operate in unison or even in random order as it is to imagine their sequential operation. Hence, cooperative catalytic activity p e r se neither supports nor denies the sequential operation of the catalytic subunits in either F-type or in P-type ATPases. Furthermore, recent studies indicate that cooperativity may actually not occur among the catalytic subunits of F-type ATPases (Renafarje and Pedersen, 1996). For all of these reasons, the precise nature of the proposed dramatic conformational changes of the subunits of F-type ATPases, and the mechanism by which they function to couple transport to ATP synthesis or hydrolysis, remain a provocative field of investigation. The nature and size of the conformational changes that P- and F-type ATPases undergo are important because the conformational changes are believed to constitute part of the pathway of free energy transfer between phosphoric acid anhydride bonds and ion gradients. The free energy changes that are associated with conformational changes in proteins appear, however, to be no more than about half the size of free energy changes associated with ATP synthesis (or hydrolysis) and ion transport. We may therefore be wise to consider whether free energy may move from that in phosphoric acid anhydride bonds to that in ion gradients via pathways in addition to the free energy changes associated with changes in protein conformation. For example, it

Summary

was proposed in Chapter 3 that some of the free energy in phosphoric acid anhydride bonds may be converted into the free energy realized when the locations of liquid and crystalline lipid domains are organized in membranes. These different domains were proposed to be associated with different conformations of P-type ATPases and thus to add to the size of the free energy change associated with conversion of one conformation to the other. In addition, transient conversion of enzyme monomers to dimers during a transport cycle may be associated with conversion of the EIP conformation to the EzP conformation, at least in S E R C A ATPases. This conversion of monomers to dimers should, because of its organized nature and free energy of binding, also contribute to the size of the free energy change associated with conversion of EIP to EzP. To test such theories we need to devise methods to identify and measure each component of the free energy changes associated with conformational changes of P- and F-type ATPases as they function in their normal environments in biomem-

| 67

branes. In the process, we may need to think more creatively about the possible intermediate forms and pathways that free energy may take as it is converted from a form as obvious as that inherent in phosphoric acid anhydride bonds to another conspicuous form such as that realized in ion gradients. It may at first seem easier to envision how the free energy inherent in a gradient of one solute is converted into that of another than it is to conceive of conversion of this free energy into a chemical form. We turn in the next Chapter to a consideration of such apparently simpler free energy conversions which are catalyzed by symporters and antiporters. Detailed descriptions of the molecular structures of several such proteins in the next chapter help to expose both our impressive ability to explore the structures and functions of these transport proteins and the naivete (or perhaps even arrogance) implicit in the concomitant assumption that we may now soon understand on a fundamental level how they catalyze transport and free energy conversions.

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R

Transport Proteins That Propagate Solute Gradients 9

1. INTRODUCTION TO SYMPORTERS AND ANTIPORTERS

1

transfer of the free energy first to a high-flee-energy conformation of a transport protein molecule. The highflee-energy conformation may result from binding of the first solute and transport along its gradient. Subsequent conversion of the protein molecule back into its low free energy conformation with concomitant transport of the other solute against its gradient would complete the free energy transfer. The low-frequency conformation of such an antiporter would, however, need somehow to be determined by the original solute gradient, since antiporters frequently operate in either direction. The free energy needed to produce and maintain the asymmetric structures of membranes and their transport proteins might also be transferred into gradients of the substrates of the proteins (Section IX,C,2 of Chapter 4 and see also Section II,C of Chapter 7). That is, one conformation of a transport protein may have an inherently higher free energy than the other owing to its asymmetric orientation across an asymmetric membrane. In the absence of substrate, the protein would form the appropriate conformational gradient across the membrane. In the presence of substrate, however, we propose that this conformational gradient might be partially or wholly propagated into a gradient of the substrate. If this theory is correct, then symport and antiport may appear in some cases to be thermodynamically more efficient in one direction than in the other. In addition to these free energy contributions, free energy changes are associated with substrate selectivity and the organized migration of some but not all solutes. The free energy of this organized transport also must be contained somehow within the construction and functioning of transport protein molecules. An attractive

Since the free energies realized in different solute gradients take similar forms, it might at first seem to be a simple matter to propagate one from another. The detailed mechanisms of such conversions are, however, neither simple nor direct. If transport proteins migrated across membranes as carriers, solutes could conceivably drive them to one face of membranes with their total chemical potential gradients. In the case of, say, an antiporter, the protein could then follow its gradient back to the other face of the m e m b r a n e carrying a different solute against its gradient. Transport proteins do not, however, migrate across membranes to catalyze transport. Rather, they undergo conformational changes while embedded in them. Such conformational changes are believed in the case of transport ATPases to be part of the route of transfer of the free energy in phosphoric acid anhydride bonds to that of solute gradients (Chapter 5). Similarly, the free energy in a gradient of one solute might be transferred to that of another through 1We prefer the term "propagate" rather than the term "convert" to describe the production and maintenance of a total chemicalpotential gradient of a solute through symport or antiport with another solute. To constitute such production and maintenance of an additional gradient, the gradient of one solute must be propagated into that of another solute, while the gradient of the first solute is maintained by some other means such as primary active transport. Since the gradient of the first solute usually is maintained while it is propagated into a gradient of the second, it seemsinaccurate to use the term "converted" to describe the transport catalyzedby these transport proteins. Rather, free energy is transferred to the second solute gradient via the first. Moreover, numerous solute gradients may actually be propagated across cell membranes from a single gradient such as that of Na+.

169

| 70

6. Transport Proteins That Propagate Solute Gradients

hypothesis is that all forms of free energy are combined into a single form of a transition state. This high free energy of the transition state would again take multiple forms when the transport cycle is complete. The high free energy of the transition state could conceivably be contained transiently within the transport protein molecule and one or a few surrounding phospholipid domains. As discussed above, not only may symporters and antiporters take bioenergetically distinct conformations when facing different sides of the membrane, but substrate binding may also favor one conformation of the transport protein over the other. Similarly, different conformations of a transport protein may associate preferentially with a liquid or a crystalline phospholipid domain (recall related discussions in Chapters 3-5). Such associations would represent a component of the free energy change since they help to organize the locations of the lipid domains rather than permit them to be random. Consequently, a transport protein molecule and the phospholipid domains that surround it might be able to contain all of the free energy of the proposed high free energy transition state. In this Chapter we discuss the transfer of free energy from one or more solute gradients to that of one or more others by two apparently unrelated families of symporters and antiporters. We begin with the wellknown example of antiport by a family of erythroid and nonerythroid proteins termed anion exchangers 1, 2, and 3. We then discuss the mechanism of Na+/amino acid symport by another family of proteins expressed both within and outside the brain. Unlike apparently simpler symporters, the latter proteins also cotransport protons and some of the proteins exchange the three cosubstrates for K + (Zerangue and Kavanaugh, 1996a). Moreover, some of the transporters may catalyze K § homoexchange and exchange of the three cosubstrates for each other (Kavanaugh et aL, 1997; Millar et aL, 1997). In both the latter, obviously complex case of symport/antiport and in the case of anion exchange, investigations continue into the kinetic mechanisms by which transport occurs. For example, neither a ping-pong (e.g., Fig. 4.44 in Chapter 4) nor a simultaneous mechanism appears fully to explain all of the characteristics of anion exchange (e.g., Salhany, 1990, 1992, and 1996). By choosing these complex examples we are forced to consider in detail uncertainties and questions about the kinetics of symport and antiport. Consequently, we can more readily challenge frequent assumptions such as that most instances of antiport have a ping-pong mechanism, whereas most instances of symport have a multisite simultaneous one. It is somewhat difficult to challenge the assumption that symport occurs only by a simultaneous mechanism, perhaps because most experiments are

planned and their results interpreted assuming this model. Transport appropriately defined as symport may, nevertheless, occur without migration of all cosubstrate ions and molecules together in their final stoichiometric amounts (see Section III,B,3 below). Moreover, now that oligomerization of transport protein molecules is known to be fairly common, simultaneous mechanisms of antiport as well as symport seem more feasible and have been tested. Oligomerization of transport proteins also means that we must consider allosteric regulation of their function. Undetected or unappreciated instances of allosteric regulation may obscure accurate perception of symporter and antiporter function.

II. BOTH ERYTHROID AND NONERYTHROID TISSUES EXPRESS ANION EXCHANGERS Anion exchange (AE) proteins appear to occur ubiquitously in mammalian and avian tissues, although different tissues may express different isoforms. The erythroid isoform (AE1) was the first of the anion exchange proteins to be characterized, and it is also known as band 3 because it migrates as the third band from the top on SDS-polyacrylamide gel electrophoresis of the human red cell membrane proteins (summarized by Passow, 1992). Band 3 is a 100-kDa glycoprotein that is expressed in kidney as well as in cells of erythroid origin. It accounts for more than a quarter of the total mass of protein in the red cell membrane, which is equivalent to about 1.2 x 106 copies of the protein per cell. Since the protein appears actually to be present in the membrane as a homodimer (see below) and the surface area of the red cell is about 140/xm 2, each dimer is separated from its nearest neighbor by an average distance of only about 15 nm (Fig. 6.1). AE1 is believed to help to increase the total CO2 (CO2 plus HCO3-) that can be dissolved in and carried by blood from the vasculature of respiring tissues. It catalyzes net transport of HCO3- out of red blood cells in exchange for C1- after HCO3- is formed from CO2 and H20 by carbonic anhydrase (Fig. 6.2). CO2 and H20 are produced in greater amounts when cellular respiration is faster, and the whole process is reversed in the vasculature of the lungs as a result of excretion of CO2. As for other antiporters, the red cell anion exchanger is properly viewed as propagating one solute gradient into another. Unlike the gradients used by many other antiporters, however, the initial gradient of HCO3- is produced metabolically rather than through primary active transport. Because of the service of erythrocyte anion exchange for pH and CO2 homeostasis, it was somewhat surprising

Erythroid and Nonerythroid AE Expression

171

FIGURE 6.1 Freeze fracture electron micrographs of bovine red cell membranes. The concentration of normally abundant integral membrane protein molecules (A) is reduced by about 70% in cells from animals deficient in anion exchanger 1 (AE1) (B). As calculated in the text and shown here by the difference between A and B, AEl-dimers are normally separated from neighboring dimers by about 15 nm (bar, 100 nm; magnification, x200,000). (adapted from Inaba et al., 1996, with permission from the American Society for Clinical Investigation).

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to learn that some AEl-deficient mice (Peters et aL, 1996) and cattle (Inaba et al., 1996) survive into adulthood. Since these animals do not compensate for their AEl-deficiency by expressing the normally nonerythroid AE isoforms, AE2 and AE3, or another rapid anion exchange transport activity in their red cells, other mechanisms appear to serve well enough for pH and CO2 homeostasis to permit survival even in the absence of AE1. These mechanisms include but are not limited to normal expression of the AE2 and AE3 isoforms of these proteins in other tissues (e.g., Wang et aL, 1996). Nevertheless, AEl-deficient cattle experience mild acidosis, and this condition worsens during exercise or dietary acid load apparently owing to reduction in the total CO2 that can be carried by blood and hence excreted by these animals. AEl-deficient mice and cattle also suffer from chronic hemolytic anemia for which they partially compensate by increasing erythropoiesis (Peters et al., 1996; Inaba et aL, 1996). Premature red cell destruction appears to occur primarily as a result of the loss of AE1

172

6. Transport Proteins That Propagate Solute Gradients

as a structural component of erythrocytes (recall discussion in Chapter 2). Nevertheless, as for pH and CO2 homeostasis in the whole animal, the structure and viability of red blood cells does not depend on the band 3 protein alone. Consequently, red cells survive albeit for a shorter than normal time because other components of the membrane and cytoskeleton are able to maintain erythrocyte integrity well enough to support the lives of some animals. We discuss further the importance of AE1 to red cell structure in Section IV,A of this chapter. We also consider in this Chapter the importance of the structures of AE1, AE2, and AE3 to their functions both as antiporters and as channels. Furthermore, we discuss the kinetics of this transport and its possible regulation by allosteric effectors. First, however, we describe structural similarities and differences among the isoforms and subisoforms of this family of anion exchange proteins, and we consider a novel mechanism by which some portions of these proteins may have evolved.

A. Evolutionary C h a n g e s in the Structures of AE lsoforms H a v e O c c u r r e d Primarily in Their N-Terminal Cytosolic D o m a i n s A form of AE1 (band 3) appears to be expressed in all vertebrates including jawless fishes (e.g., Kay et al., 1995). In contrast, AE2 and AE3 have thus far been detected only in birds and mammals. If these findings are borne out by further investigations then it should be possible to conclude that duplication of an ancestral AE1 gene led to evolution of genes encoding AE2 and AE3 perhaps owing to selection for mechanisms of pH homeostasis that would better support life on land. The C-terminal and central portions of the three types of anion exchange proteins have been relatively well conserved during evolution (Espanol and Saier, 1995). In contrast, the short (i.e., about 60 amino acid residues) N-terminal segment of the AE1 molecule has no sequence similarity to the longer (about 320 residues)

N-terminal portions of AE2 and AE3 (Fig. 6.3). The N-termini of AE2 and AE3, are however, quite similar (Espanol and Saier, 1995), so the genes encoding these two types of proteins probably evolved from a common ancestor first derived from a gene encoding AE1. Moreover, some short sequences within the 320-residue-long N-termini of AE2 and AE3 appear to have evolved by a proposed new mechanism for protein evolution that involves trinucleotide residue expansion in D N A (see Sections II,A,2 and II,A,3 below). Instead of the N-terminal, central, and C-terminal segments of the A E proteins just discussed, these proteins are more often divided into N-terminal and Cterminal domains. The N-terminal cytosolic domain associates with specific protein molecules in the membrane, cytoskeleton, and cytosol, whereas the Cterminal membrane-associated domain forms the pathway for anion transport (Passow, 1992). The amino acid residue sequence of the C-terminal domain is, of course, highly conserved among A E proteins as indicated in the preceding paragraph, and it constitutes about 40% of the sequences of AE2 and AE3 and about 60% of the sequence of AE1 (Fig. 6.3). In contrast, the sequence of the N-terminal domain grows increasingly less well conserved among the proteins as the N-terminus is approached. Additional diversity is generated near the beginnings of the N-terminal domains of the proteins through alternative splicing. The N-terminal end is needed by AE1 and possibly AE2 and AE3 for their interactions with other protein molecules (Ding et al., 1994, 1996; Wang et al., 1995; Schneider and Post, 1995), and it is likely needed in AE2 and AE3 for functions that are yet to be determined.

1. Structural Similarities and Differences among the Isoforms and Subisoforms of A E Proteins

As we have seen, at least three related genes encode the anion exchange isoforms in mammals and birds, and subisoforms are produced from each gene. In contrast to the transport proteins discussed in Chapter 5, however,

FIGURE 6.3 Amino acid residue sequence alignments of 13 anion exchange (AE) proteins. The sequence of each protein (named in the top three lines) begins with the first residue (usually a methionyl one) and proceeds down the page and on to subsequent pages. Conserved residues can be identified by reading across the columns. Residues conserved in all AE1, AE2, AE2 plus AE3, and all AE proteins are marked by asterisks or one-letter amino acid abbreviations in the pertinent columns. Notes in the left column indicate portions of the sequences of the N-terminal cytoplasmic domains that are rich in particular amino acid residues. Also indicated in the left column are putative membrane spanning segments and extramembrane loops of the Cterminal membrane-associated domains. Some parts of the sequences are omitted (marked by the word, OMISSION) because they are not needed for the discussion in the text (adapted from Wood, 1992, with permission from Elsevier Science).

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Erythroid and Nonerythroid AE Expression

subisoforms of anion exchange proteins are produced through transcription from alternative promoters as well as through alternative splicing of the transcripts (Wang et aL, 1996). For example, a truncated version of AE1 is produced in mouse, rat, human, and chicken kidney apparently through transcription from an alternative promoter that is not utilized in mammalian erythroid tissue (Fig. 6.4). The AE1 transcript in mammalian kidney is missing the first three exons of the transcript that is present in erythroid tissue. Consequently, the renal AE1 protein molecule is shorter at its N-terminus by 65 amino acid residues in humans and by 79 residues in rats and mice (Fig. 6.3). This region of the AE1 molecule is needed in red blood cells for binding to ankyrin, the band 4.1 protein, several glycolytic enzymes, and hemoglobin (Ding et al., 1994, 1996; Wang et aL, 1995; Schneider and Post, 1995). The latter structural and metabolic functions of band 3 appear to be at least as important as its transport function in red blood cells

|8 |

(recall the AE1 deficient animals discussed above). Because the truncated AE1 molecule expressed in kidney probably cannot serve these structural and metabolic functions, the main function of the truncated AE1 molecule in kidney may be to help to regulate excretion of protons and other ions through anion exchange. Again, however, other mechanisms appear to regulate pH and excretion of ions well enough to support adult life since the AE1 deficiency described above would extend to the truncated as well as the longer form of AE1. In contrast to the single subisoform of AE1 expressed in erythroid tissue in mammals, this tissue in the chicken appears to express at least four AE1 subisoforms (Cox et al., 1995). Alternative splicing of transcripts produced from two alternative promoters results in the production of eight transcripts (four from each promoter) (Fig. 6.5). In addition, initiation of transcription at one of the promoters begins at two alternative sites, which are separated by 21 nucleotide residues (Cox et aL, 1995).

FIGURE 6.4 Use of alternative promoters and splicing to produce mRNAs encoding subisoforms of anion exchangers 1 (AE1), 2(AE2), and 3 (AE3) in mammals. Exons are numbered and shown as boxes that are untranslated (hatched) or serve to encode amino acid residues (open). Sites for the initiation of transcription and translation are indicated by arrows and ATG, respectively. The number of codons in the long open reading frame of each mRNA is shown in the last column. The tissue distribution of each mRNA is indicated in parenthesis in the first column (adapted from Wang et al., 1996, with permission from American Society for Biochemistry & Molecular Biology, Inc.).

182

6. Transport Proteins That Propagate Solute Gradients

FIGURE 6.5 Production of 12 mRNAs encoding four anion exchanger 1 (AE1) subisoforms (a to d) using alternative promoters and splicing in the chicken. The use of promoter 1 or 2 results in m R N A with untranslated exons 2 or 1, respectively, at the 5'ends of the molecules. Moreover, initiation at promoter 1 can occur at major and minor sites which are 21 nucleotides apart, as indicated by dark rectangles at the 5' ends of the AEI-1 m R N A variants. Thus, a total of 12 m R N A s are produced from the two promoters. The 12 mRNAs encode four protein subisoforms which increases in size from a to d and vary in structure only near their N-termini. Translation initiation sites are marked with A U G (adapted from Cox et al., 1995, with permission from American Society for Biochemistry & Molecular Biology, Inc.).

Consequently, a total of 12 different transcripts encoding four subisoforms of AE1 appear to be produced in chicken erythroid tissue. Since each set of four transcripts that is produced from a given initiation site encodes the same set of four protein subisoforms, the three different initiation sites may serve functions other than to increase the variety of proteins produced. Perhaps different sites of initiation are needed for proper regulation of expression of the subisoforms of AE1 in erythroid tissue. As for all AE subisoforms, the four erythroid-specific AE1 subisoforms of the chicken differ in length and in

some cases amino acid residue sequence at or near their N-termini (Cox et al., 1995). No more than about the 90 most N-terminal residues may be missing or different in different subisoforms, and even the shortest of these four subisoforms is still 45 residues longer than the truncated kidney-specific AE1 subisoform in the chicken. Since the smallest two of the four proteins are expressed in the plasma membranes of transfected human erythroleukemia cells, whereas the largest of the proteins is retained in a perinuclear compartment, it has been suggested that the different N-termini of the erythroidspecific AE1 proteins may help to target them to differ-

Erythroid and Nonerythroid AE Expression

ent membrane compartments (Cox et aL, 1995). To our knowledge, it has not been determined whether these different N-termini bind different intracellular proteins and hence might serve different structural and metabolic functions in various parts of the cell. As for regulation of AE1 expression in erythroid cells, regulation of its expression in kidney is more complex in the chicken than it is in mammals (Cox and Cox, 1995). Although transcription is initiated at a single nonerythroid specific promoter both in the chicken and in mammals, the resultant transcript is alternatively spliced in the chicken to produce m R N A molecules encoding two subisoforms. One of these chicken kidney subisoforms is the truncated AE1 protein also expressed in mammalian kidney. Somewhat surprisingly, the truncated chicken kidney subisoform of AE1 appears also to be expressed in erythroid tissue in addition to the four subisoforms described above. The other chicken kidney subisoform is, however, a somewhat larger protein (Cox and Cox, 1995) apparently not expressed in any other tissue. The larger chicken kidney subisoform has 63 N-terminal amino acid residues that are not present in the truncated AE1 protein. The first 21 of these residues are unique to this larger kidney subisoform, whereas the other 42 are identical to the corresponding residues in the nontruncated erythroid AE1 subisoforms (Cox and Cox, 1995). Perhaps this additional complexity in the pattern of expression of AE1 subisoforms in chickens is needed in part because the pattern of expression of AE2 subisoforms also differs between these birds and mammals. At least three subisoforms of AE2 are produced in mammals through transcription from alternative promoters and alternative splicing of the resultant RNA (Wang et al., 1996). AE2a, the largest of the AE2 proteins, is expressed in a wide variety of tissues. AE2b is produced primarily in stomach using a different promoter than for AE2a. The resultant transcript encoding AE2b is also spliced differently than the transcript encoding AE2a (Fig. 6.4). The first exon of the AE2b transcript corresponds to a region within the second intron of the AE2a transcript. This exon of the AE2b transcript contains the 5' untranslated region plus codons for the first three amino acid residues. Since the first exon of AE2b is spliced to an exon corresponding to exon 3 of AE2a, the 17 N-terminal residues of AE2a are not present in AE2b. In this regard, the hydroxyl group of the seryl residue at position 10 of AE2a may be a cAMP-dependent phosphorylation site. Consequently it was proposed that the transport activity of AE2a but not AE2b may be regulated in a cAMPdependent manner (Wang et aL, 1996). Alternatively or in addition, the different N-terminal sequences of AE2a

183

and AE2b may result in their sorting to different regions of the plasma membranes of polarized cells. The smallest mammalian subisoform of AE2 is produced exclusively in stomach through the use of a promoter in the region of the AE2 gene corresponding to intron 5 of the AE2a transcript (Wang et al., 1996). This first exon of the AE2c transcript is spliced to an exon corresponding to exon 6 of the AE2a transcript, and the translation initiation site is in this case near the 5' end of the exon corresponding to exon 6 (Fig. 6.4). Consequently, AE2c is a truncated version of AE2a that does not contain its first 199 residues. Interestingly the N-terminal regions of AE2a and AE2b contain clusters of histidyl and glutamyl residues that we propose may be involved in evolution of new functions in proteins (see below). To date, however, no function has been attributed to these clusters. 2 As for the AE1 gene, regulation of AE2 expression in the chicken is different from that in mammals (Cox et al., 1996). Both alternative sites of initiation of transcription and alternative splicing of the RNA transcript result in the production of two AE2 subisoforms. Unlike mammals, however, both of these relatively large AE2 proteins are expressed preferentially in stomach where they apparently contribute to acid secretion. They also contain the histidyl and glutamyl clusters present in larger subisoforms of mammalian AE2 and AE3. As for the truncated and nontrucated subisoforms of AE2, large and small subisoforms of AE3 are also expressed in mammals (summarized in Wang et al., 1996). The brain subisoform AE3b has nearly 200 more N-terminal amino acid residues than the cardiac subisoform AE3c (Fig. 6.4). In this case, however, the use of an alternative promoter and alternative splicing introduces a translation initiation site into the AE3c mRNA that is not present in the AE3b mRNA. Consequently, the 270 N-terminal amino acid residues of AE3b are replaced by 73 different residues in AE3c (Fig. 6.4). As for the truncated AE2c subisoform expressed exclusively in mammalian stomach, the AE3c subisoform does not contain the histidyl and glutamyl clusters present in larger subisoforms of these proteins. We propose that the histidyl- and glutamyl-rich sequences in AE2a, AE2b, and AE3b evolved through trinucleotide residue expansions in an ancestor to the AE2 and AE3 genes. More recently, further enlargement of the gene encoding AE3 proteins through trinucleotide residue expansion apparently has led to an increase in the size of a cluster of anionic amino acid residues in the proteins 2 Readers are cautioned that a paper reporting a regulatory function of the histidyl cluster (Sekler et al., 1996) has since been retracted

(Cell, 1997, 90(6)).

| 84

6. Transport Proteins That Propagate Solute Gradients

(i.e., addition of residues 141 to 143 of the AE3 proteins shown in Fig. 6.3). 2. Have Trinucleotide Residue Expansions Contributed to the Evolution of AE Proteins? Several regions within the N-terminal domains of anion exchange proteins contain clusters of identical amino acid residues. These clusters appear to have formed through trinucleotide residue expansions in the genes that encode the proteins. A similar mechanism of evolution may have led also to diversification of channel transport proteins (e.g., see Section II,B of Chapter 7). Trinucleotide expansions appear to be tolerated within the coding regions of genes often without harm, whereas tetranucleotide and other expansions seem more frequently to be deleterious even when they do not produce frame shifts (see Section II,A,3 below). Apparently expansions that add a series of nonidentical amino acid residues to the resultant proteins adversely affect their functions, whereas additions of clusters of identical amino acid residues are relatively innocuous. As such new regions in proteins evolve, other amino acid residues may replace some of the residues in the cluster through mutation. Consequently, what may have been a relatively innocuous change in a protein to add a cluster of identical residues may over time evolve a useful new function (Green and Wang, 1994). In Section II,A,3 below we discuss our evidence that trinucleotide residue expansions contribute to protein evolution by creating relatively harmless clusters of virtually every amino acid residue. First, however, let us consider possible instances in which trinucleotide residue expansions may have contributed to the evolution of some isoforms of the anion exchange proteins. Several examples of possible trinucleotide residue expansions are present in the genes encoding the mouse and rat AE3 protein molecules (Fig. 6.3). The cluster of glutamyl residues at positions 135 to 145 of these proteins is three residues longer than a similar cluster of anionic residues in AE2. These additional residues could have been added to the AE3 protein through expansion of the trinucleotide sequence, GAG, in the corresponding region of the gene from which the mouse and rat genes descended. An earlier expansion could conceivably have produced the other neighboring trinucleotides, G A A and GAG, that encode glutamate. In this case, expansion of one or the other of the trinucleotides is presumed to have occurred first followed by silent mutations to produce the other. If the earlier expansion occurred just prior to the time of gene duplication to produce separate AE2 and AE3 genes, then the original gene may have encoded only glutamyl residues in this region. The aspartyl residues currently in

this region of AE2 (Fig. 6.3) may have evolved later through point mutations, while the cluster of glutamyl residues was retained and further expanded in AE3. Even the glycyl and alanyl residues immediately preceding and following the glutamyl cluster of AE3 (Fig. 6.3) could conceivably have evolved from trinucleotides that initially encoded glutamate (GAA) but that evolved after an earlier expansion to encode alanine (GCA) and glycine (GGA). Similarly, expansion of the trinucleotide sequence, AAG, could have produced the region in the AE3 gene that now encodes a cluster of lysyl residues at positions 308 to 312 and possibly the arginyl and lysyl residues at positions 306 and 307 (Fig. 6.3). It remains to be determined whether these and similar Sections of AE3 and other anion exchange proteins contribute to their transport function or its regulation. One such section of some AE2 and AE3 subisoforms could, however, have undergone a series of evolutionary changes that seem unlikely to have occurred if the resultant protein did not have a function that was subject to natural selection. This histidyl-rich motif runs from residue 79 to residue 83 and is H H I H H in isoforms of AE2 and H H T H H in isoforms of AE3 (Fig. 6.3). We propose that the H H X H H motif arose initially as a relatively innocuous cluster of five histidyl residues through trinucleotide residue expansion in an ancestral gene encoding an anion exchange protein. A point mutation in the codon for the middle residue from CAC to CTC would have led to its replacement by a leucyl residue. If the resultant protein had a beneficial function, it would have been selected over the protein with a cluster of five histidyl residues. Subsequent selection for a second point mutation (i.e., from CTC to ATC) would have led to replacement of the leucyl residue by an isoleucyl one in the modern AE2 (Gehrig et al., 1992). This isoleucyl residue could have been converted to a threonyl residue in AE3 through a final point mutation from ATC to ACC. While admittedly speculative, the feasibility of such evolution of proteins initially through trinucleotide residue expansion is supported by our finding that clusters of virtually all amino acid residues are present in proteins at much greater than anticipated frequencies. 3. Proteins May Evolve Through Formation and Expansion of Amino Acid Residue Clusters Proteins evolve through numerous processes already known to most readers. Readers may, however, only now be aware that codon reiteration or trinucleotide residue expansion appears also to be a mechanism for the evolution of some segments of proteins (Green and Wang, 1994). In this process, relatively innocuous clus-

Erythroid and Nonerythroid AE Expression

ters of additional amino acid residues are proposed to be added to proteins through trinucleotide residue expansions in the genes that encode them. A mechanism for producing such expansions has been proposed (reviewed by Wells, 1996) and excessive expansions (i.e., more than 40 repeats) of a codon encoding glutamine results in at least six human neurodegenerative disorders (reviewed by Sutherland and Richards, 1995). It has been proposed that glutaminyl clusters may in some cases be beneficial (Stott et al., 1995) rather than simply harmless because such clusters appear to help proteins form oligomers. Glutaminyl clusters greater than 40 residues long may, however, cause a given species of protein molecules to have undesirably strong affinity for each other or for other proteins that have glutaminyl clusters (Stott et al., 1995). In addition to glutamine, seven other amino acid residues have been shown to have a statistically significant tendency to occur in clusters in proteins (White and Jacobs, 1993). In contrast, eight residues have been found by other investigators n o t to form clusters, and some of these amino acid residues are even reported to be anticlustering (White and Jacobs, 1993; Green and Wang, 1994). Several of these apparently nonclustering residues are, nevertheless, present in clusters in transport proteins. In contrast to other investigators, we have found that not just some but rather every amino acid residue appears in clusters in various proteins more frequently than anticipated owing to chance alone. Because of the possible importance of our data to the evolution of proteins in general and anion exchange and other transport proteins in particular, we consider them here in some detail. Each of the 20 amino acid residues appears in clusters of the same residue more frequently than anticipated owing to chance (Figs. 6.6 and 6.7). These data cannot be explained simply because clusters of some amino acid residues would be tolerated better in hydrophobic or hydrophilic regions of proteins. While clustering might occur more frequently than anticipated owing simply to chance in regions of proteins that are similarly hydrophobic or hydrophilic, they would occur less frequently than expected in dissimilar regions. Consequently, the overall frequency of clusters is expected to be nearly random regardless of the properties of the amino acid residues. Clusters of six identical residues occur 3 (valine) to 10,000 (glutamine and histidine) times more frequently than expected (Figs. 6.6 and 6.7). Moreover, the probability that a cluster of identical residues will be extended by at least one more residue increases as the cluster grows longer (Figs. 6.6 and 6.8). Hence, very large clusters of some amino acid residues appear in proteins against astronomically large odds.

185

For example, the mean frequency of appearance of glutaminyl (Q) residues in the National Biomedical Research Foundation-Protein Identification Resource (NBRF-PIR) database was determined to be 3.43%. 3 Hence, it can be calculated that a cluster of 7 Q residues should occur once in every 1.8 x 10 l~ heptapeptides within entries in the data base (1/0.03437). The June, 1994 NBRF-PIR database contains 70,848 entries with a total of 20,816,057 amino acid residues. Each sequence of an entry contains n - 6 heptapeptides where n is the number of residues in the sequence. The total number of heptapeptides in the database is 70,848 • ( N - 6) = 20,816,057 - 70,848 • 6 = 20,390,969 where N is the average number of residues in the 70,848 entries in the data base (i.e., N = 20,816,057/70,848 = 293.8129). The probability of there being a homogenous cluster of 7 Q residues in the database is, therefore, 0.114% (20,390,969 • 0.03437) if clusters of Q residues occur only owing to chance. A total of 1840 clusters of 7 Q residues were, however, detected in the database (total number of clusters of 7 Q residues = the number of sequences with a cluster of 7 Q residues + 2 • the number of sequences with a cluster of 8 Q r e s i d u e s . . . + 32 • the number of sequences with a cluster of 38 Q residues (the longest cluster of Q residues in the June, 1994 data base)). Moreover, the number of clusters of 7 Q residues detected in this manner is an underestimate, since multiple clusters of 7 Q residues separated by non-Q residues in the same entry would not have been detected by the method available for searching the database. Hence, clusters of 7 Q residues occur more than 1.61 • 106 times more frequently in proteins than anticipated owing to chance alone (1840/0.00114). Even under the conservative assumption that clusters of more than 7 Q residues should be counted as a cluster of 7 only once, clusters of 7 residues still occur 2.35 • 105 times more frequently than anticipated owing to chance (268/0.00114). Similar calculations for clusters of 32 and 38 Q residues indicate that they occur more than 1041 and 1048 times more frequently, respectively, than anticipated owing to chance. Calculations for clusters of 13 histidyl, 23 prolyl, 24 glycyl, 30 glutamyl, 32 asparagyl, and 35 seryl residues show that they appear in the data base more than 1013, 1014 , 10 21 , 10 31 , 10 32, and 10 34 times more frequently, respectively, than anticipated owing to chance. Similarly, when the mean frequency of appearance of Q residues in the database (3.43%) is increased by 2 SEM to make it 97.5% probable that the actual frequency of occurrence of Q in the database is below the frequency used for these calculations (i.e., mean + 2 • SEM = 3 The method used to determine the frequency of appearance of residues in the data base is described in the legend to Fig. 6.6.

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I

2

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3

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4

i

5

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6

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FIGURE 6.6 Clusters of three or more identical amino acid residues usually appear in proteins more frequently than anticipated owing to chance. We used DNASIS for Windows Sequence Analysis Software (Hitachi, San Bruno, CA) to determine the number of entries in the June 1994 National Biomedical Research Foundation-Protein Identification Resource (NBRF-PIR) data base that contain clusters of two to six identical amino acid residues. The same data base also was utilized to determine the number of proteins containing other specific short sequences of amino acid residues as indicated in Figs. 6.8 to 6.10, Tables 6.1 and 6.2, and the text. In analyses of protein primary structure performed elsewhere (White and Jacobs, 1990, 1993; White, 1994a,b; Klapper, 1977), single representative proteins were selected from families of related proteins. These selections were made to minimize the possibility that some short sequences or regularities in proteins might be detected more frequently simply because some families of proteins had more representatives in the data base than other families of proteins. In the present studies, however, we looked for a general tendency of every amino acid residue to form clusters of identical residues in proteins. Unlike previous studies, we did not use sequences of residues in proteins in the data base to form hypotheses about which sequences might occur more frequently. Hence, it was not necessary to select only one member of each family of proteins for our analysis or even to insure that duplicate entries were not present in the data base. Although some arbitrarily selected sequences might appear more frequently in the data base than anticipated owing to chance because some families of proteins are represented more frequently there, other arbitrarily selected sequences should, likewise, appear less frequently. Consequently, one would not expect to verify the hypothesis that clusters of each of 20 amino acid residues occur in proteins more frequently than anticipated owing to chance unless such were actually the case. In fact, redundancies and overrepresentation of some families should add variability to the analyses and perhaps prevent detection of clusters of some of the residues. In addition, clusters of identical amino acid residues were not found to be particularly well conserved among proteins in the same family (see text in this chapter and also Section II,B of Chapter 7 for examples). Hence, arbitrarily selecting single members of each family of proteins for the present investigation would have reduced the data base and the ability to detect clustering. The numbers of entries in the NBRF-PIR data base with clusters of the indicated lengths were divided by the numbers of such clusters expected owing to chance to calculate the extent to which clusters occur more or less frequently than expected (i.e., observed/ expected). These values for clusters of 2-6 residues are indicated for each residue by the one-letter abbreviations for the amino acids. The solid line connects the means of the logarithms of the 18 to 20 values for clusters of the indicated sizes (clusters of 4 W or 6 C residues were not observed). The number of entries expected to contain at least one

Erythroid and Nonerythroid AE Expression

3.43 + 2 x 0.23 = 3.89%), clusters of 7, 32, or 38 Q residues can still be calculated to occur about 6.69 x 1 0 5 , 1 0 39 , and 1 0 46 times more frequently, respectively, than anticipated owing to chance. A similar figure of about 3.89% has also been reported elsewhere as the average glutamine content of proteins (Dayhoff, 1978; White, 1992; Green and Wang, 1994). Moreover, when the amino acid residue compositions of proteins reported elsewhere (e.g., Dayhoff, 1978) are used for our calculations, it can still be concluded that virtually all residues are found in clusters in protein much more frequently than anticipated owing to chance (e.g., Fig. 6.8B). Rigorous statistical analyses using actual numbers of entries in the data base show that each amino acid residue is indeed found in clusters of identical residues more frequently than anticipated owing to chance (p < 0.01, Table 6.1). The observation that clusters of identical amino acid residues become more likely to be extended as the length of the cluster increases (Fig. 6.8) is consistent

187

with the theory that clusters may increase in size through amplification of trinucleotides that encode the residues. Such amplification may occur through D N A polymerase slippage (Streisinger et al., 1966) which, when accompanied by insufficient mismatch repair (Heale and Petes, 1995), appears to result in unstable expansion of trinucleotide repeats as in the cases of human neurodegenerative disorders (Sutherland and Richards, 1995). In addition, some clusters seem to result from expansion of longer nucleotide repeats such as the cluster of 20 glutaminyl residues in the yeast G A L l 1 protein that are encoded by four tandem repeats of the sequence CAGCAGCAACAACAA (Suzuki et al., 1988). The theory that larger clusters form from smaller ones is also supported by our observation that clusters of two identical residues usually occur in proteins about as frequently as anticipated owing to chance, whereas clusters of three or more residues usually occur more frequently than expected (Table 6.2 and Fig. 6.6). In contrast, randomly selected tri- or tetrapeptides occur in

instance of a given cluster was calculated by first calculating the probability that an entry of average length (293.8 residues per average entry = 20,816,057 total residues/70,848 entries) does n o t contain the cluster = (1 - frequency of the residue in the data basem)293"8-m+1,where m = number of residues in the cluster. The probability that an entry does contain the cluster = 1- the probability that it does not, and the number of entries expected to have the cluster = the probability that an average entry has the cluster x 70,848 (the number of entries in the data base). The frequencies of each residue in the data base were determined as described in the following paragraph. Similar conclusions can be drawn regardless of whether these frequencies (present figure) or previously published (e.g., Dayhoff, 1978) frequencies (Fig. 6.7) for the occurrences of amino acid residues in proteins are used for the calculations. The amino acid composition of the data base was measured by determining the frequencies at which each residue preceded six different randomly selected tripeptides within entries in the data base. The mean (___ SEM) percentage frequency was calculated from these six determinations for each of the 20 amino acid residues. Nearly identical results were obtained when the mean (and SEM) were calculated using the arcsine transformation (Woolf, 1968). The six randomly selected tripeptides used here occurred a mean number of 2535 times in the data base. This number is close to the expected average number of occurrences of randomly selected tripeptides, namely (frequency of occurrence of the average residue in the data base) 3 x the number of tripeptides in the data base = 0.053 x 20,674,361 = 2584. The number of tripeptides in the data base = 70,848 entries in the data base x (N - 2) = 20,674,361, where N is the number of residues in the average entry (20,816,057 residues/70,848 entries = 293.8129 residues/entry). Since we would not have detected multiple occurrences of the tripeptides in a single entry, but about 94 entries are expected to have two of a randomly selected tripeptide, we apparently detected the anticipated number of tripeptides. While most clusters of two residues occur no more frequently than anticipated owing to chance, clusters of three or more residues usually occur more frequently than expected when either the frequencies of amino acid residues determined here or elsewhere (Fig. 6.7) are used in the calculations (p < 0.01; see analysis in Table 6.2). In contrast, more complex sequences of amino acid residues occur no more frequently than expected owing to chance for up to a total of six residues in the sequences D R Q T D R (circles) and K Q T N K Q (triangles). (No instance of all six of these residues was observed as anticipated for the sequence K Q T N K Q (0.15 entry was expected to have it) and, perhaps, even for the sequence D R Q T D R (0.56 entry was expected to have it).) In these cases, the numbers of entries that were expected to have a particular sequence of two to six residues were calculated as above except that the frequencies of each of the pertinent residues in the data base were multiplied instead of raising the frequency of a single residue to the power of a cluster length, m. Similarly, in the rest of the calculations, m was equal to the lengths of the short sequences of interest. The amino acid sequences D R Q T D R and K Q T N K Q are encoded by the tetranucleotide repeats in microsatellite DNAs (GACA)n and (AAAC)n, respectively. For comparison, clusters of identical residues may, of course, be encoded by trinucleotide repeats, and alternating sequences of two amino acid residues may be encoded by dinucleotide or hexanucleotide repeats (see text and Fig. 6.10 for examples).

| 88

6. Transport Proteins That Propagate Solute Gradients

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FIGURE 6.7 The tendency to form clusters of three or more amino acid residues in proteins is so strong that it does not depend on highly accurate determination of the frequency of amino acid residues in the protein data base. In this case, the observed/expected ratios were calculated using the frequencies of amino acid residues in proteins published by Dayhoff (1978) rather than as determined by us for the NBRF-PIR data base (see the legend to Fig. 6.6). As can be seen by comparing this figure to Fig. 6.6, the relative positions of some of the amino acid residues change considerably although the general trend remains virtually the same.

proteins no more frequently than anticipated owing to chance (Table 6.2). Similarly, pairs of different amino acid residues also occur in proteins about as frequently as anticipated owing to chance, but alternating repeats totaling four or six residues occur more frequently than expected (Figs. 6.9 and 6.10). As for clusters of the same amino acid residue, this phenomenon for pairs of different residues could occur through expansion of the six nucleotides that encode the pair (Fig. 6.9). In contrast, expansion of at least some simple nucleotide sequences that would lead to more complex arrangements of unlike amino acid residues do not seem to occur or are strongly selected against. For example, sequences of amino acid residues encoded by the tetranucleotide repeats (GACA), and (AAAC)n are not found in protein more often than expected

owing to chance (Fig. 6.6). These tetranucleotide repeats are microsatellite DNAs known to be present in numerous species (Epplen, 1988; Beckmann and Weber, 1992). Since there is no known biochemical mechanism that should lead to tri- or hexanucleotide expansions more frequently than tetranucleotide expansions (Heale and Petes, 1995), we favor the theory that all of these expansions occur, but that tetranucleotide expansions are more strongly selected against than are tri- or hexanucleotide expansions when they occur in coding regions of genes. The fact that about two thirds of tetranucleotide expansions produce frame shifts probably accounts for much of this negative selection. However, most dinucleotide expansions also produce frame shifts, but dinucleotide expansions appear to occur more frequently than antic-

Erythroid and Nonerythroid AE Expression

| 89

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N U M B E R O F R E S I D U E S IN T H E C L U S T E R A F T E R IT H A S B E E N E X T E N D E D BY O N E R E S I D U E

FIGURE 6.8 As cluster length increases, the proportion of proteins in which the cluster is extended by at least one more residue also increases. The number of entries in the data base with at least one more residue identical to that in a cluster was divided by the number of entries with the initial cluster to produce the percentages shown. Calculation of percentages was discontinued arbitrarily at clusters of 14 residues or when the number of entries in the data base with the cluster fell below 20. Percentage values for each clustering amino acid residue are indicated by the one-letter abbreviation for the amino acid. The dashed line connects the mean percentages of clusters of the indicated lengths that are extended by at least one more residue while the solid line connects the percentages for glutamine (N).

i p a t e d owing simply to chance (Fig. 6.10). A s for m o s t h e x a n u c l e o t i d e expansions, d i n u c l e o t i d e expansions give rise to alternating pairs of two different a m i n o acid residues (e.g., Fig. 6.10). P e r h a p s for this reason, di- and h e x a n u c l e o t i d e e x p a n s i o n s a p p e a r to be ret a i n e d in coding regions of genes m o r e f r e q u e n t l y t h a n are t e t r a n u c l e o t i d e expansions. All of these expansions are, h o w e v e r , r e t a i n e d less f r e q u e n t l y t h a n are trin u c l e o t i d e e x p a n s i o n s (Figs. 6.6 to 6.10 and r e c e n t analysis of locations of various n u c l e o t i d e expansions in genes ( D e F o n z o e t aL, 1998)). It is, of course, possible that s o m e clusters of identical residues have b e e n selected during e v o l u t i o n for their beneficial attributes. A limited review of the literature indicates, h o w e v e r , that clusters of identical residues f r e q u e n t l y s e e m to have no obvious function. F o r example, clusters of S , L , T , E , G , H , P , N , and W residues sometimes are c o n s e r v e d in p r o t e i n s in the s a m e family but

f r e q u e n t l y they are n o t conserved. 4 T h e latter p h e n o m e n o n occurs e v e n w h e n proteins in the s a m e families also have the s a m e k n o w n functions ( M a j e a u and C o l e m a n , 1991; Schlesser e t a/.,1988; Bullard e t a L , 1988; TsengC r a n k e t al., 1990; F f r e n c h - C o n s t a n t e t a/.,1991; G r a b n e r e t al., 1991; K r i s t e n s e n e t aL, 1991; W o o d g e t t , 1990; 4 Compare amino acid residue sequences in Sakura et aL (1988); Woolford et al. (1988); Majeau and Coleman (1991); Piatkowski et aL (1990); Smythe et al. (1990); Ono and Means (1989); Slade et al. (1990); Guiltinan et al. (1987); Tanaka et al. (1992); Schlesser et al. (1988); Bullard et al. (1988); MacKrell et aL (1988); Tseng-Crank et al. (1990); Tanaka et al. (1990); Hohl et al. (1991); Ffrench-Constant et aL (1991); Plaza et al. (1991); Grabner et al. (1991); Feng et aL (1993); Kristensen et al. (1991); Dorer et al. (1990); Leid et al. (1992); KOster et al. (1991); Krolewski and Dalla-Favera (1991); Woodgett (1990); Gregory et aL (1987); Hsieh et al. (1992); Selski et al. (1993); Peverali et al. (1990); Kurowski and Ludwig (1987); Bae et al. (1989); Dangott et al. (1989); Grover et al. (1993); Celeste et al. (1990); Nakayama et al. (1992); and White et aL (1993).

| 90

6. Transport Proteins That Propagate Solute Gradients

TABLE 6.1

Frequency of Association (%) of t h e S a m e and Different A m i n o Acid Residues with a Cluster of Each Residue a

Frequency at which the specified residue(s) precedes the indicated cluster (%) Clustering amino acid residue

Cluster length

Numbers of entries in the data base with the cluster

Clustering residue

Ala (A) Arg (R) Asn (N) Asp (D) Cys (C) Gin (Q) Glu (E) Gly (G) His (H) lie (I) Leu (L) Lys (K) Met (M) Phe (F) Pro (P) Ser (S) Thr (T) Trp (W) Tyr (Y) Val (V)

6 5 5 4 3 7 7 7 7 4 7 5 3 4 7 7 7 2 4 5

433 288 208 690 449 268 188 237 45 186 119 199 475 147 194 205 72 4103 97 51

52.4 35.8 53.8 32.6 5.3 75.7 69.1 61.2 75.6 15.1 35.3 29.9 6.9 9.5 46.1 54.7 65.3 2.4 24.7 27.5

Nonclustering residues (mean _ SEM) 2.5 2.8 2.4 2.8 3.5 1.3 1.2 1.2 1.3

___ 0.6 _ 0.4 +__0.6 +_ 0.4 _ 0.5 ___ 1.1 ___ 0.3 +_ 0.4 ___ 0.5

Nonclustering residue that was a statistical outlier

13.2(P) 17.1(E) 30.7(S) 9.0(D) 16.5(S)

3.4 ___ 1.0 3.7 ___ 0.8 3.9 2.8 2.4 0.9

___ 0.9 +_ 0.8 ___ 0.6 ___ 0.5

25.5(L)

18.1(A)

4.0 ___ 0.8 2.9 ___ 0.8

19.6(D)

aEach amino acid residue was found to precede a cluster of the same residue more frequently than anticipated from its frequency in the data base (p < 0.01, two-by-two contingency table analyses of the actual numbers of entries with the residues in front of clusters vs in front of randomly selected sequences (Woolf, 1968)). In addition, several residues were associated with clusters of another residue more frequently than were other noncluster residues. Such statistical outliers are more than four standard deviations above the mean frequency of occurrence of the other noncluster residues (Shoemaker and Garland, 1967). Identical conclusions concerning statistical outliers were drawn after transforming the data to angles (arcsine transformation; Woolf, 1968).

Hsieh et aL, 1992; Kurowski and Ludwig, 1987; White et al., 1993). Similarly, the mouse homologs for the spinocerebellar ataxia 1 and Huntington disease genes have only 2 and 7 tandem trinucleotide residues, respectively, that encode glutamine (Lin et al., 1994; Banff et al., 1994). Hence, the longer clusters of 19 to 36 and 11 to 34 glutaminyl residues, respectively, encoded by normal alleles for the human forms of these genes (Huntington's Disease Collaborative Research Group, 1993; Orr et aL, 1993; Cha and Dure, 1994) seem unlikely to be needed for their functions. More dramatic examples of the apparent lack of function of glutamine clusters include the finding that removal of more than 200 amino terminal residues including a cluster of 37 glutaminyl residues had no conspicuous effect on the function of a glutamine- and proline-rich transcriptional activator in Saccharomyces cerevisiae (Laurent et al., 1990). In addition, deletion of the 27-residue polyglutamine region from the C-terminus of the Neurospora gene regulatory protein nit-4 did not appear to alter its function (Yuan et al., 1991). Hence, rather than clusters of identi-

cal residues having a beneficial function, we favor the view that insertion as well as deletion of clusters of up to about 40 identical residues often may have little or no effect on protein function. The number 40 applies to the maximum number of glutaminyl residues that appear to occur in proteins encoded by normal alleles of genes corresponding to several human neurodegenerative disorders (Huntington's Disease Collaborative Research Group, 1993; La Spada et al., 1991; Nagafuchi et al., 1994; Koide et al., 1994; Burke et al., 1994; Orr et al., 1993; Kawguchi et aL, 1994; Sutherland and Richards, 1995; Cha and Dure, 1994). Similarly, clusters of more but not less than 16 alanyl residues in the HOXD13 protein were shown to cause synpolydactyly (Muragaki et al., 1996). Hence, the maximum number of identical residues that may be relatively harmless in clusters appears to vary depending on the particular residue and probably also the position in the proteins in which the clusters occur. In this regard, the largest clusters in the June, 1994 NBRF-PIR database varied from 3 residues for tryptophan to 13 residues

Erythroid and Nonerythroid AE Expression TABLE 6.2 Clusters of Three or Four but Not Two Identical A m i n o Acid Residues Occur in Proteins M o r e Frequently Than Anticipated O w i n g to Chance

Type of sequences in proteins

n

Quotient of the observed/expected frequency in the data base (mean __. SEM) a

Clusters of two identical residues Two nonidentical adjacent residues

20 20

0.89 __+0.06] 0.84 _+ 0.03J~ n.s.

Clusters of three identical residues Three nonidentical adjacent residues

20

1.96

12

0.93 _+ 0.08J

Cluster of four identical residues Four nonidentical adjacent residues

20

8.01

14

0.93 ___0.06J

0.28~ p < 0.01 |

2.24~ P < 0.01 /

aThe observed/expected values were calculated as described in the legend to Fig. 6.6. The means ___SEM were calculated from these values for clusters of each of 20 identical residues or 12-20 randomly selected nonidentical adjacent residues. Clusters of three or four but not two identical residues occurred more frequently than anticipated owing to chance and more frequently than randomly selected sequences of nonidentical residues (p < 0.01, Kruskal-Wallis H tests, Woolf, 1968).

for histidine, 23 for proline, 32 for asparagine, and 38 for glutamine. It also seems likely that selection against clusters of all amino acid residues occurs at least to some extent because the normalized frequencies of clusters of the same length vary widely among the residues (Fig. 6.6). There is no reason to expect that the various trinucleotide expansions occur with different frequencies. Regardless of this possible selection against all clusters to some degree, and the existence of a maximum number of identical residues that can occur in clusters without almost certainly affecting protein function adversely, it is clear that clusters of every residue occur in proteins more frequently than anticipated owing to chance. Because insertions of clusters of 3 to nearly 40 identical amino acid residues may frequently be harmless, we propose that their production could result in the evolution of useful new regions in proteins through subsequent mutations within the trinucleotide expansions that produce clusters. Furthermore, the evolution of completely new and potentially useful regions in proteins might occur with qualitatively different results from DNA encoding functionally neutral clusters of identical amino acid residues than from DNA in duplicate genes that are relatively free to undergo rapid evolution throughout their nucleotide sequence. The constraints on evolution of new regions from functional

191

or once-functional regions could conceivably be quite different than evolution of new regions from functionally neutral clusters of identical amino acid residues within functional proteins. For example, perhaps functionally neutral clusters are retained once they are formed unless an amino acid substitution is beneficial. This theory is consistent with the observation that clusters of identical residues are more common than are sequences in which other amino acid residues intrude into the clusters (Figs. 6.8 to 6.10, Table 6.1 and data not shown). Such a process would also result in the production, through silent mutation, of different codons for the same amino acid residue in more ancient trinucleotide expansions. In this regard, amino acid residues in clusters frequently are encoded by the same codon, but often they are encoded by codons that differ by one nucleotide residue (Sutherland and Richards, 1995; Suzuki et aL, 1988; Sakura et aL, 1988; Woolford et al., 1988; Henthorn et al., 1986; Chaplin et al., 1986; Chen et aL, 1986; Fragoso and Goldenberg, 1992; Ohta et al., 1987; BOrki et al., 1991). A small proportion of clusters also probably evolved relatively slowly through selection of very different codons for the same amino acid residue. Such appears to be the case, for example, for the clusters of four seryl residues in the seventh putative transmembrane segment of most of the members of the Na+-dependent amino acid transport family discussed in Section III below (i.e., the EAAT/ASC family). Most clusters appear, however, to have formed relatively rapidly through trinucleotide expansion, and the more ancient of these expansions now contain silent mutations. Other observations also support the theory that trinucleotide expansions of up to about 40 repeats may be both relatively harmless and a mechanism for evolution. Among both cancer cells and bacteria, the greater the ability of the cells to produce trinucleotide and other expansions, the more rapidly they evolve under environmental stresses (Sutherland and Richards, 1995; Cairns and Foster, 1991; Ionov et al., 1993; Aaltonen et al., 1993; Thibodeau et al., 1993). This possible mechanism for increasing the rate of evolution during stress is reminiscent of the ability of another type of repeated sequence, the transposable elements, also able to increase the rate of evolution under stress (Gerasimova et al., 1984; McClintock, 1984; Walbot and Cullis, 1985). For all of these reasons, we suggest that formation of simple nucleotide residue repeats and, in particular, trinucleotide residue expansions within coding regions of genes may reflect a normal process by which all species evolve and by which their rates of evolution may increase in response to environmental stresses. It seems likely that useful additions to proteins evolve through relatively innocuous trinucleotide expansions in the genes that

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FIGURE 6 . 9 Tandem repeats of two nonidentical amino acid residues appear in proteins more frequently than anticipated owing to chance but not as frequently as clusters of identical residues of the same length. The quotients of observed/expected frequency ratios were determined as described in the legend to Fig. 6.6, and these values are the same as the values in Fig. 6.6 for clusters of identical amino acid residues. Also shown are the ratios for tandem repeats of two different amino acid residues (e.g., (GS),,) that may be encoded by hexanucleotide repeats. In these cases n = 1.5 and 2.5 for sequences of three and five

| 93

Erythroid and Nonerythroid AE Expression

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i

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I I I I 2 3 4 5 Length of Cluster or Sequence (Residues)

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FIGURE 6.10 Greater-than-expected frequency of occurrence of tandem repeats of two nonidentical amino acid residues encoded by the microsatellite D N A dinucleotide repeat sequences (AC)" and (AG)" (Epplen, 1988; Beckman and Weber, 1992). The encoded amino acid sequences are (TH)" and (RE),,, respectively. Also shown are the values for clusters of each of these individual residues (i.e., T', H,,, R', and E~ from Fig. 6.6). See the legends to Figs. 6.6 and 6.9 for further details of the analysis.

encode them followed by selection for beneficial mutations within the expansions. In contrast to the possible evolution of additions to some subisoforms of anion exchange proteins through formation of clusters of identical amino acid residues in their N-terminal domains, the membrane-associated C-terminal domains of these proteins have remained highly conserved in all AE isoforms and subisoforms. None of the C-terminal domains contains clusters of identical residues in their transmembrane segments, and

each appears to catalyze anion exchange in the same way. The ability of the proteins also to serve as channels is, however, influenced by the length of the extracellular loop between transmembrane segments five and six (Fi6vet et al., 1995), and this loop appears to have become longer in the mouse AE2 protein through trinucleotide expansion (see Section IV,B below). We begin our discussion of the membrane-associated domains of AE proteins with some interesting observations on similarities in their primary structures.

residues, respectively (e.g., GSG and GSGSG). It should also be noted that the alternating residues are the same residues as those for which data on clustering is shown. In each case, the degree to which the observed frequency exceeds the expected frequency is greater for clusters than for alternating pairs of residues, although in the case of G,,, S', and (GS)', these values are nearly identical except for sequences six residues long. The unusually high frequency of occurrence of (GS)n is consistent with data in Table 6.1, which show that seryl residues precede clusters of glycyl residues much more frequently than most other residues except glycine. (A) Clusters or alternating sequences of residues that are among the most likely of all residues to form clusters (i.e., Q', H,,, and P" in Fig. 6.6). (B) Examples of clusters or alternating sequences of residues that are less likely than Q,H, and P residues to form clusters (Gn, S', An, and D" in Fig. 6.6).

194

6. Transport Proteins That Propagate Solute Gradients

FIGURE 6.1 1 Model for the membrane-associated C-terminal domain of human erythroid anion exchanger 1 (AE1). The locations of the first eight putative transmembrane segments is relatively well established, whereas the presence and precise locations of the last six segments are less certain. One of the larger extracellular loops is N-glycosylated, and the other is susceptible to hydrolysis by chymotrypsin (C). In addition, cysteine 843 is fatty-acylated and an intracellular loop is subject to hydrolysis by trypsin (T). Ct = C-terminus Nt = N-terminus. (adapted from Wang et al., 1994, with permission from Oxford University Press).

B. Structure and Function of the C-Terminal M e m b r a n e - A s s o c i a t e d Domains of AEI and Related Anion Exchange Proteins The membrane-associated domains of AE proteins appear to be composed of 14 transmembrane c~-helices with intervening intracellular and extracellular loops of varying sizes (Fig. 6.11). 5 The sizes of the extracellular loops are particularly variable; a minimum number of residues form some of the loops, whereas many more than the minimum number form others (Wang et al., 1994). One of the larger extracellular loops is Nglycosylated and another is susceptible to cleavage by chymotrypsin (Fig. 6.11). In part because of the small size and, hence, lack of susceptibility to proteolysis of many extramembrane loops, the existence and precise 5 The results of more recent experiments have been used to propose a 12- and another 14-span model for AE1 (Popov et al., 1997; Tanner, 1997; Tang et al., 1998). As we shall see, however, the structure of the C-terminal membrane-associated domain actually may change enough to allow several such static structural models to be correct at one time or another during the transport cycle.

locations of the last six putative membrane transverses are less certain than the first eight. Nevertheless, these proteins are almost certain to span the membrane an even number of 10, 12, or 14 times because both the Cand the N-terminus are located in the cytosol (Fig. 6.11). Under the assumption that AE proteins contain 14 membrane traverses, it is interesting to compare the primary structures of the putative transmembrane segments both to each other and to the traverses of other membrane-associated proteins. In the latter case, the transmembrane segments of anion exchange and some other transport proteins have a greater frequency of residues with aliphatic side-chains in the centers of the spans, while both aliphatic and aromatic side-chains are present on residues nearer the membrane surface (Espanol and Saier, 1995). In contrast, other proteins with multiple spans have the reverse distribution (Fig. 6.12). It may be possible through site-directed mutagenesis to determine the importance of these different distributions of aliphatic and aromatic residues in the structures and functions of different proteins. Such studies should become more interesting after it has been determined

Erythroid and Nonerythroid AE Expression

200

195

t

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25

150

~

20

100

,--~'~ 15 ~n,

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50

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I

i

i

i

i

140 120

60

~

40

80

7= 20 o

20

C

---

5

cO

0

2

4 6 8 10 Transmembrane Helix

12

14

FIGURE 6.13 Distribution of amino acid residues (K, R, H, D, and E) that may carry a charge in the putative transmembrane segments of anion exchange (AE) proteins. The total number of these residues in each of the 14 putative transmembrane segments is shown for seven members of the AE protein family. While few or no such residues are found in most even-numbered transmembrane segments according to some models (e.g., Fig. 6.11), other models place a few charged residues in these segments (see text). The reader is reminded that although these residues may usually be charged, they are sometimes uncharged in hydrophobic environments such as that between the surfaces of a biomembrane (from Espanol and Saier, 1995, with permission from Taylor and Francis).

~' loo "~ I= "6

10

FWY

15

10 5

0

0

''

~

lb Position

1~

26

FIGURE 6.12 Distribution of aliphatic (I, V, L, M; circles) and aromatic (F, W, Y; squares) amino acid residues across (A) putative transmembrane segments of the anion exchangers (AE), (B) bacterial rhodopsins (BR), and (C) the three subunits of the reaction center complex from Rhodobacter spaeroides (RC). A sliding window of three residues was used in locating features of each plot. The numbers of residues are shown for all transmembrane segments in seven members of the AE-family (A), 11 members of the BR family (B), and the RC complex (C) (from Espanol and Saier, 1995, with permission from Taylor and Francis).

which other amino acid residues contribute to catalysis of anion exchange. The effects of reversing the distribution of these residues in membrane-spanning segments of the proteins alone as well as the effects of such reversals on proteins already altered using site-directed mutagenesis might be informative of the mechanisms by which the proteins function. Within the family of AE proteins, another somewhat surprising observation has been made concerning the distribution among the transmembrane segments of residues that may carry a charge (i.e.; D, E, K, R, and H).

According to some models for membrane traverses 3 to 12, only odd-numbered traverses may contain such residues (Fig. 6.13). Uncertainty in the number of traverses and their precise locations in different models makes this observation concerning the locations of potentially charged residues less strict, especially in regard to traverses 9 and 10. 6 Moreover, transmembrane helix 4 may have one such residue depending on the model used to locate transmembrane segments (Saier, personal communication) (Fig. 6.11). Nevertheless, the orientations of the odd-numbered transmembrane helices that have relatively large numbers of these residues is such that their C-termini are toward the outside (Espanol and Saier, 1995). These orientations may have significance to the mechanism by which the helices are inserted into the membrane. This pattern of the distribution of D, E, K, R, and H residues among the transmembrane helices also is likely to be pertinent to the structure of the pathway through which anions migrate. 1. Structure of the A E 1 Anion Pathway

Site-directed mutagenesis has been used to study the relationship between the structures of AE proteins and their function, although progress for these proteins has 6 The reader is reminded that although these residues may usually be charged at physiological pH values, they may not be charged in hydrophobic environments such as that in the interior regions of phospholipid bilayers.

196

6. Transport Proteins That Propagate Solute Gradients

been slower than for many other transport proteins (e.g., P-type ATPases described in Chapter 5). We may attribute this slower progress to the lack of fully satisfactory cell types in which to express mutant proteins (e.g., a null phenotype cell line) and to the difficulty in verifying that the mutant proteins are expressed in the plasma membrane rather than retained in the Golgi or endoplasmic reticulum (e.g., Alper, 1994; Mtiller-Berger et al., 1995a,b). Nevertheless, various mutagenesis and related procedures, such as the use of reagents selectively to modify the side-chains of particular amino acid residues, have been used to gain insight into the transmembrane segments and amino acid residues that may help to form the pathway for anion transport by AE1. The C-terminal membrane-associated domains of AE proteins still catalyze apparently normal anion exchange after removal of their N-terminal domains (reviewed by Alper, 1994). The finding that these truncated proteins transport anions does not, however, rule out a role of the N-terminal cytosolic domain in that transport. For example, the various N-terminal domains of the AE proteins may help to regulate their transport functions differently. Moreover, the N-terminal domain appears to coordinate anion transport with cellular metabolism and structure (Section IV,A below). In spite of the possible importance of the N-terminal domain to AE function, the apparent lack of its direct participation in transport seems to have militated against the frequent use of site-directed mutagenesis to study it. For these reasons, the present discussion of AE1 structure and transport function is limited primarily to its C-terminal membrane-associated domain. Mutation of each of three histidyl residues in three different transmembrane segments of the C-terminal domain results in reduced or undetectable anion exchange (Mtiller-Berger et al., 1995a,b). Moreover, one of the histidyl residues appears to interact with a glutamyl residue in another transmembrane segment through hydrogen bonding. Similarly, the effects of two of the mutations of histidyl residues on transport are partially or completely reversed by a second site mutation of a lysyl residue in a fifth transmembrane segment. These and other observations led Passow and associates (Mtiller-Berger et al., 1995a,b) to propose the following model for the pathway across which anions migrate during transport by AE1. Although admittedly speculative (M011er-Berger et al., 1995a,b), the model forms a framework for our further discussion. The pathway is proposed to be composed of membrane traverses 5,8,9,10 and 13 (Mtiller-Berger et aL, 1995a,b), which contain residues K558, E699, H721, H752, and H852, respectively, according to some models

of the mouse AE1 protein (Fig. 6.14). 7 Interestingly, the status of helices 9,10, and 13 as membrane traverses is less certain than that of helices 5 and 8 (e.g., Fig. 6.11 for human AE1 and see discussion above). The reader should also note for the present discussion that residues K558, E699, H721, H752, and H852 in the mouse AE1 (Fig. 6.14) correspond to residues K539, E681, H703, H734, and H834 in the human protein (Fig. 6.11). From these sequences it can be seen that histidyl residues 752 and 852 (734 and 834 in the human), which in some models are in helices 10 and 13 (e.g., Fig. 6.14), fall within cytosolic loops in more recent models (e.g., Fig. 6.11). We propose that the positions of these residues actually may change during the transport cycle so that both models may be correct to some degree. For example, histidyl residues 752 and 852 may lie in cytosolic loops when the AE1 protein takes a conformation that receives substrate in the cytosol, whereas the residues may lie in a protected hydrophobic environment in or near the membrane in the AE1 conformation that receives substrate outside the cell. In support of this theory, when AE1 is stabilized in its outward-facing conformation with the stilbene compounds DNDS or DIDS, histidyl residue 752 (734 in the human) is protected from the histidyl reactive agent DEPC (Fig. 6.15). In contrast, this residue is not protected from cytosolic exposure to D E P C when AE1 is in its inward-facing conformation (Mtiller-Berger et aL, 1995a). Similarly, other histidyl residues, possibly including 852 and 721 (834 and 703 in the human), are protected from cytosolic DEPC by DNDS (MtillerBerger et al., 1995a) or DIDS (Hamasaki et al., 1992). Another histidyl residue (i.e., mouse 837 or human 819) that is considered to be in the cytosolic loop between membrane traverses 12 and 13 in both models presented 7 Other models are consistent with the theory that charged amino acid residues occur only in odd-numbered membrane transverses (Fig. 6.13). The precise locations of the transmembrane helices as well as the nature of the anion pathway they form are uncertain enough to permit presentation of several possible models here without misrepresenting current knowledge. Moreover, residues and portions of the helices that contain them appear to enter or leave the hydrophobic environment of the membrane when the protein undergoes its conformational changes during the transport cycle (see text). Consequently, acceptance of any one static model (e.g., as in Fig. 6.11 or Fig. 6.14) as fully accurate may be unwarranted. In this regard, somewhat newer 12-span and 14-span models have been proposed for AE1 based in part on the results of more recent experiments (Popov et al., 1997; Tanner, 1997; Tang et al., 1998). Again, however, a variety of static structural models may have been generated in part because the actual structure of AE1 may change enough to resemble each of these models at different times during the transport cycle. Consistent with the proposal in the text, it was shown recently that spans 6 and 7 of the 14span model could be deleted without loss of transport activity (Groves et aL, 1998).

198

6. Transport Proteins That Propagate Solute Gradients

OutwardJ ~ ~

Inward

outside K.558

(

,

g~ DEPC

":::::

K869 ~

DNDS binding site

....

FIGURE 6.15 Mobile histidyl residues are protected from the histidyl reactive agent DEPC in the outward-but not in the inward facing conformation of the anion exchanger 1 (AE1). See text for discussion of the specific histidyl residues that are protected in this way when stilbene compounds such as DIDS or DNDS stabilize AE1 in its outward facing conformation (adapted from Hamasaki et al., 1992, with permission from Elsevier Science).

here (Figs. 6.11 and 6.14) also is protected from DEPC by stilbene compounds. These results show why it is difficult and perhaps even why it may not be possible to define precisely the positions of putative transmembrane segments 9 to 14 and particularly 9, 10, and 13. We propose that the residues that comprise the last six putative transmembrane segments, and perhaps even the number of transmembrane segments, change during the transport cycle. These changes occur as a result of the conformational changes in AE1 that are needed for it to catalyze anion exchange. In the model proposed by Mtiller-Berger and associates (Fig. 6.16), the side-chain of residue K558 in transmembrane helix 5 is proposed also to interact with the side-chains of residues H721, H837, and H852 in other helices or cytosolic loops of mouse AE1. K558 has been shown, however, to be some distance from the histidyl residues; it lies near the stilbene derivative binding site on the outside of the membrane (Passow et aL, 1992), whereas the histidyl residues react with DEPC on the inside (Hamasaki et al., 1992). Hence, it is tempting to speculate that these residues do not interact directly but rather indirectly through effects on AE1 conformation. For example, perhaps the second site mutation of K558 to N reverses the effects of site-directed mutagenesis of each of the histidyl residues to Q through an effect on the balance of positive charges at each face of the membrane. The proper balance of positively charged residues on one side of the transport pathway or the other could conceivably be more important to normal AE1 function than the absolute charges on each side of the pathway. Consistent with this possibility, the second site conversion of K558 to N reverses the inhibition of transport resulting from conversion of H837 to Q but not the inhibition resulting from conversion of H837 to R (Mtiller-Berger et al., 1995a). Perhaps the relative strength of the attraction of anionic substrates to the total positive charges at each end of the transport path-

H752 I DEPC (Helix 10) i binding site E699 (Helix 8)

to substrate binding site FIGURE 6.16 Amino acid residues and transmembrane helices of mouse anion exchanger 1 (AE1) that may help to form the pathway across which anions migrate. Binding of stilbene compounds, such as DNDS, blocks binding of the histidyl reactive agent diethyl pyrocarbonate (DEPC, also see Fig. 6.15). Transmembrane helix 9 (not shown) is also believed to help to form the pathway for anion migration (see text for discussion) (adapted from Mtiller-Berger et al., 1995b, with permission from the American Chemical Society).

way must be properly balanced in order for the anions to follow the pathway. The latter notion is consistent with the possibility that the histidyl residues become part of the pathway when they function to attract anions into the cell (i.e., when the conformation of AE1 is to receive external substrates), whereas the histidyl residues leave the pathway when anions are to be transported out of the cell (i.e., when the conformation Of AE1 is to receive internal substrate). Helix 8 is also believed to comprise part of the pathway across which anions migrate (e.g., Tang et al., 1998). Conversion of E699 in this helix to D through sitedirected mutagenesis shifts the pK value for the influence of pH on transport from 5.8 to 6.7 and reduces transport by about two-thirds (Mtiller-Berger et al., 1995b). E699 in helix 8 is presumed to interact with H752 in helix 10 because conversion of H752 to S has the same effect on transport as conversion of E699 to D. Both mutations are believed to disrupt the interaction between the side-chains of these two and perhaps residues in other helices and hence to shift the pK for the influence of pH on transport to a higher value. More recently, R748 in helix 10 (R730 in the human) has been shown also to be essential for transport (Gartner et aL,

Erythroid and Nonerythroid AE Expression

1997; Karbach et al., 1998). These investigators have proposed that a hydrogen-bonded series of functional groups on residues in helices 5,8,9,10, and 13 could be arranged such that they traverse the membrane from one side to the other (Mtiller-Berger et aL, 1995a,b). Full formation of this hydrogen-bonded chain by charged residues at low pH values is proposed by the authors to inhibit transport, whereas the breaking or absence of the bonds at higher pH values is thought to permit transport. As discussed above, however, the residues are located so that they could form such a chain in some models of AE1 structure (e.g., Fig. 6.14), but they do not fall at the proper locations according to other models (e.g., Fig. 6.11). It is unlikely that a completely definitive structure for the pathway across which anions migrate could be deduced from the results of studies involving only chemical modification and mutagenesis. Such is the case for more thoroughly studied transport proteins including the SERCA ATPases (Chapter 5), although several labor-intensive techniques have provided considerable insight into the tertiary structure of the fl-galactoside transporter (Kaback et al., 1997; Frillingos et al., 1998). What we need in conjunction with these newer approaches are high resolution X-ray crystallographic studies. The inability to form the physiologically meaningful three-dimensional crystals needed for such studies is a major impairment to understanding how most membrane transport proteins function. For example, definitive identification of the transmembrane helices that form the pathway for anion migration across AE proteins would greatly facilitate the planning of other experiments, such as site-directed cross-linking mutagenesis and second site suppression (e.g., Kaback et al., 1997), to determine more precisely how transport is catalyzed. In spite of these shortcomings, formation of two-dimensional crystals of the membrane-associated domain of AE1 has helped investigators gain insight into its structure and transport function (see Chapter 2 and Section II,B,3 below).

2. Kinetics of Anion Exchange Definition of the three dimensional structures of the AE proteins will eventually help to reveal how they exchange anions. Currently, a ping-pong model is believed to fit transport data for AE1 best (e.g., Brahm et aL, 1992), whereas a two-site simultaneous model seems to account best for the transport observed for AE2 (e.g., Knauf et al., 1992). Since the degree of protein oligomerization is unlikely to be higher for AE2 than for AE1, it is unlikely that the formation of dimers, tetramers, or higher oligomers alone leads to simultaneous rather than ping-pong exchange by AE proteins.

199

Oligomerization probably leads to allosteric effects, however, and such effects have not always been considered in assessing whether a ping-pong or simultaneous model best fits the transport data. Moreover, the results of kinetic experiments may be incorrectly interpreted because more than one anion exchange transport activity may operate in membranes simultaneously. In most kinetic studies of AE proteins there has been no attempt to isolate these different possible components and to characterize their transport activities independently. Historically, quantitatively less important substrates or nonphysiological temperatures have been used to study the kinetics of anion transport across the red cell membrane. Such approaches have been used in some experimental protocols because the rates of transport of C1- and HCO3- at physiological temperatures can be too rapid to measure accurately. The mammalian AE1 is an obligatory or nearly obligatory exchanger that nevertheless functions asymmetrically in the membrane, as anticipated from our discussion in Chapter 4 (Section IX,C,2). Its favored conformation is toward the inside of the cell by about 10 to 1 (Liu et al., 1996), but the transport that it catalyzes at O~ appears to fit Michaelis-Menten kinetics reasonably well both for uptake and exodus (Brahm et al., 1992). Moreover, the transport data are in general consistent with a pingpong model (Fig. 4.44 in Chapter 4) that also explains the observed cis-inhibition and trans-stimulation of the transporter. Data obtained at 38 ~ C are, however, inconsistent with a ping-pong model unless an intracellular self-inhibition site is included in the ping-pong model and in the modified Michaelis-Menten equation that describes it (Brahm et al., 1992). The latter selfinhibition is particularly conspicuous and complex for C1-self-exchange. Consequently, even the introduction of two sites of substrate inhibition into the ping-pong model and the Michaelis-Menten equation is insufficient to account for the self-inhibition that is observed for C1- homo-exchange.(Brahm et aL, 1992). 8 In spite of these observations, the now-classic study of Jennings (1982) still supports the notion that AE1 catalyzes transport by some sort of ping-pong mechanism. In these experiments with resealed red-cell ghosts, one cycle of C1- exodus was apparently observed after which anion transport nearly ceased because the transporter could not return to its inward-facing conformation. This test has never been applied to AE2 or AE3 because the concentration of these transport proteins in the plasma membrane of nonerythroid cells is too 8 We use the terms homoexchange and self-exchange interchangeably here to mean simply the exchange of identical solute particles. Some authors use these two terms also to imply differences in the conditions under which transport is measured (e.g., Schnell and Besl, 1984).

200

6. Transport Proteins That Propagate Solute Gradients

low to detect easily only a single cycle (or in this case a half cycle) of transport. Under the current circumstances in which the highly homologous AE1 and AE2 proteins are believed to operate by fundamentally different mechanisms, however, it would seem important to devise methods by which the Jennings (1982) study of AE1 could be repeated for AE2. In the case of AE2, some transport data fit the pingpong model so poorly that a two-site simultaneous mechanism has instead been proposed for its operation (Knauf et al., 1992). The ping-pong and two-site simultaneous models predict conspicuous differences between the interactions of the antiporter with competitive and noncompetitive inhibitors according to extensions of Michaelis-Menten kinetics. In the case of an extracellular competitive but nonpermeant inhibitor, the slope of the Dixon plot for inhibition of C1- exodus is predicted by the ping-pong model but not by the two site simultaneous model to be independent of the intracellular C1concentration ([Cli-])(Restrepo et al., 1991). In contrast, both the ping-pong and two-site simultaneous models predict that the slope of the Dixon plot for inhibition of C1- exodus will vary with [Cli-] for a dead-end reversible noncompetitive inhibitor (Restrepo et al., 1992). In the case of the ping-pong model, however, the linear relationship between the slopes of the Dixon plots and 1/ [Eli-] (i.e., the slope of the slopes of the Dixon plots vs 1/[CI~-]) is not expected to change when the extracellular C1- concentration is changed, whereas it is expected to change according to the two-site simultaneous model (Restrepo et al., 1992). By each of these criteria for competitive and noncompetitive inhibitors, AE2 catalyzes transport by a simultaneous rather than a pingpong mechanism (Restrepo et al., 1991, 1992). To our knowledge, however, the efficacies of these tests have not been verified by applying them to AE1 for which a ping-pong model is believed to operate. Moreover, it is unclear how our current concept of AE function may be influenced by erroneous assumptions about the conditions under which transport is measured. For example, it is frequently assumed that transport occurs in a thermodynamically symmetric manner in regard to the substrate under equilibrium exchange conditions. In this case, homoexchange is measured at equal substrate concentrations on both sides of the plasma membrane. Intact cells have an inside negative membrane electrical potential, however, so such conditions are thermodynamically asymmetric in the physiologically most relevant case. Thermodynamic symmetry in regard to substrate obviously will obtain only if the values of its total chemical potential are equal on both sides of the membrane. Moreover, the cation ionophore nystatin is sometimes used to allow cations and C1- ions to equilibrate and, thus, supposedly to make the C1- concentrations

equal on both sides of the red cell membrane (e.g., Liu et al., 1995). This compound would, however, reduce

the total chemical potential gradient rather than the concentration gradient of C1- to zero. Assuming that the red cell membrane electrical potential remains near its normal value of about - 1 0 mV (Zavodnik et al., 1996, 1997) during nystatin treatment, the extracellular C1- concentration would be about 50% greater than the intracellular concentration at its thermodynamic equilibrium (Eq. (3.25) in Chapter 3). The red cell membrane electrical potential may, however, vary with the intracellular C1- concentration (e.g., Gedde et al., 1997), thus further complicating interpretation of such experiments. Consequently, it is unclear in what way the results of the kinetic assessment of C1- transport by AE1 may be in error owing to the assumption that the C1- concentration is the same on both sides of the red cell membrane in the presence of nystatin. Perhaps most importantly, many detailed kinetic assessments of AE transport are performed under the assumption that a single transport activity is present. At least two kinetic components of C1-/HCO3- exchange appear, however, to be present in the erythrocyte membrane (Fuhrmann et al., 1992). Moreover, both positive and negative cooperativity apparently have been observed for AEl-catalyzed transport under some experimental conditions (e.g., Schnell and Besl, 1984) (see Section II,B,4 below). These complexities in the kinetics are, however, often not recognized in studies of anion exchange. Consequently, the impression that AE1 and AE2 function by ping-pong and simultaneous mechanisms, respectively, may have been established prematurely under the assumption that Michaelis-Menten kinetics for a single transport activity are nearly adequate to account for anion exchange. 3. Multimeric Structure of A E 1

Although several explanations are possible for the complex kinetics of AEl-catalyzed transport, it is likely that this complexity results at least in part from the formation of dimers and higher oligomers of the protein in vivo. Noncovalent interactions between the membrane-associated domains of monomeric band 3 molecules produce stable dimers (Reithmeier and Casey, 1992). Pairs of dimers then appear to form tetramers via their cytosolic domains, and ankyrin was shown recently to be needed for this further oligomerization (Yi et al., 1997; Che et al., 1997). The N-terminal cytosolic domain of AE1 also is needed for it to interact with other proteins (see Section IV,A below). The dimers and higher oligomers of the band 3 protein are, nevertheless, both structurally (Fig. 6.17; Wang et al., 1994) and functionally (see next section) highly asymmetric. As discussed above, the AE1 protein mo-

Erythroid and Nonerythroid AE Expression

FIGURE 6.17 Three-dimensional map of the C-terminal membraneassociated domain of the anion exchanger (AE1) homodimer. The map was constructed from electron microscopy and image reconstruction of two-dimensional crystals. The bulky basal region appears to be embedded in the lipid bilayer, and two other regions (top of the map) protrude into the cytosol. These protrusions appear to take different positions in two-dimensional sheets and tubular crystals (adapted from Wang et aL, 1994, with permission from Oxford University Press).

nomer has at least two conformations; one conformation receives substrates on the inside of the cell, and the other receives substrate on the outside. The inwardfacing conformation appears to have the lower free energy, and it is favored over the outward-facing conformation by about 10 to 1 (Liu et al., 1996). The only putative structural representations of these different conformations obtained so far (Wang et aL, 1993; Wang, 1994) may lie in the movement of membrane-associated subdomains C relative to two other membraneassociated subdomains, A and B (Fig. 6.18), and in the movement of regions of the membrane-associated domains that protrude into the cytosol (Fig. 6.17). Somewhat surprisingly, when AE1 monomers form dimers they also appear to form a new pathway through which anions have been proposed to migrate (e.g., Fig. 6.18), although each monomer alone appears to catalyze anion exchange. As discussed above, the detailed structures of each of these possible pathways may become apparent only after three-dimensional crystals of the band 3 protein have been formed and studied. Nevertheless, one wonders whether the formation of a new and possibly additional pathway for the transport of anions when monomers form dimers contributes to the complex kinetics of transport observed for anion exchange proteins.

4. AE1-Catalyzed Transport Exhibits Both Positive and Negative Cooperativity There is little doubt that allosteric effects help to regulate transport catalyzed by AE1. The nature of the allosteric effects that are observed depend, however, on

20 |

FIGURE 6.18 The flexible subdomains, C, of the anion exchanger 1 (AE1) membrane-associated domain may move in relation to other subdomains. (A) Projection map of a single AE1 dimer in a twodimensional sheet (dashed lines) superimposed on the map for a dimer in tubular crystals (solid lines). (B) Drawing of the proposed structure of the dimer showing a change in the positions of the two Csubdomains by about 30 A in the two maps. The two C-subdomains are proposed to move within the membrane (which is parallel to this page) possibly in association with conformational changes in the dimer during anion transport. The precise locations of the subdomains in the dimer are, however, still under investigation. Note also the presence of a central transport pathway apparently formed as a consequence of dimerization, although AE1 monomers appear also to catalyze anion exchange (adapted from Wang et aL, 1993, with permission from Oxford University Press).

whether the kinetics of uptake, exodus, or equilibrium exchange are studied (Schnell and Besl, 1984). Moreover, some of these allosteric effects are eliminated by stilbene disulfonates at concentrations that appear to inhibit one of the two monomers in dimers (Barzilay and Cabantchik, 1979). Hence, the allosteric effects depend as expected on the multimeric structure and function of the AE1 protein molecule. When AEl-catalyzed uptake is measured at constant intracellular substrate concentration, complex kinetics are observed that may reflect either multiple transport activities or negative cooperativity. At HCO3- concentrations of i m M or below, at least two kinetically distinct components of uptake appear to operate via heteroexchange (Fuhrmann et al., 1992) (Fig. 6.19). Similarly, curved Hofstee plots may reflect more than one saturable component of phosphate or sulfate uptake by homoexchange at concentrations between two and 100 m M (Fig. 6.20) (Schnell and Besl, 1984). As discussed by Salhany, however (1990), these kinetics are also consistent with the occurrence of negative cooperativity. In this regard, higher concentrations of substrates (Fig. 6.21B) have been observed actually to inhibit uptake (Barzilay and Cabantchik, 1979), thus supporting the notion that at least some of the observed effects are produced through negative cooperativity.

202

6. Transport Proteins That Propagate Solute Gradients

30 t.-

E tO x ;~

20

::L

0

200

400

600

800

1000

p,M NaHCO a

FIGURE 6.19 Anion exchanger 1 (AE1) appears to catalyze two kinetically distinct components of HCO3-/C1- transport in human red cells. The Km values for the putative high- (dotted line) and low(dashed line) capacity components of HCO3-/C1- exchange are about 1100 and 10 ~M, respectively, according to nonlinear regression analyses of the nearly identical results of the two experiments shown. These data are, however, also consistent with allosteric negative cooperativity for exchange by AE1 dimers (see text for further discussion) (adapted from Fuhrmann et al., 1992 with permission from Elsevier Science).

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Substrate inhibition has also been observed for exodus of sulfate, bicarbonate, and chloride at intracellular substrate concentrations above 200 mM (Figs. 6.21A and 6.22) (Barzilay and Cabantchik, 1979; Brahm et al., 1992). Hyperbolic kinetics obtain, however, for exodus of phosphate and sulfate (Schnell and Besl, 1984) at intracellular concentrations between about 5 and 120 mM and constant extracellular substrate concentration (Fig. 6.23). For uptake as well as exodus, the complex kinetics, apparently involving substrate inhibition, become hyperbolic in the presence of enough DNDS to inhibit one or the other of the subunits in AE1 dimers (Fig. 6.21). Consequently, these complex kinetics appear to depend on allosteric interaction between the subunits, at least at relatively high substrate concentrations (Salhany, 1990). Interestingly, the negative cooperativity for uptake and exodus of sulfate and phosphate becomes positive (Fig. 6.24) when both their intracellular and the extracellular concentrations are varied together between 2 and 100 mM in red cell ghosts (Schnell and Besl, 1984). Because observation of this positive cooperativity appears to depend on changing the substrate concentration

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FIGURE 6.20 Uptake of sulfate (A and B) and phosphate (C and D) by homoexchange at 100- and 110-mM intracellular substrate concentrations, respectively, appears to exhibit negative cooperativity. Alternatively, the human anion exchanger 1 (AE1) could catalyze two kinetically distinct components of uptake into red cell ghosts (see text for further discussion). Osmotic balance was maintained at different substrate concentrations by changing the potassium citrate or sorbitol concentration (adapted from Schnell and Besl, 1984, with permission from Springer-Verlag).

203

Erythroid and Nonerythroid AE Expression 0.75 _

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FIGURE 6.21 Exodus (A) and uptake (B) of sulfate by the erythrocyte anion exchanger 1 (AE1) are both inhibited at high substrate concentrations. Inhibition of exchange by the stilbene compound, DNDS, abolishes substrate inhibition apparently owing to inhibition of one or the other of the monomers in the AE1 dimer. Substrate inhibition is proposed to result from allosteric negative cooperativity, which is possible only when both monomers are active in the dimer (see text for further discussion) (data from Barzilay and Cabantchik, 1979, are replotted here).

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on both sides of the membrane simultaneously, it would seem also to depend on binding of substrate to AE1 subunits also facing the two different sides. While such positive cooperativity could conceivably be observed when two or more substrate molecules bind on the same side of the membrane and undergo transport together in the same direction, such a mechanism is made less likely by the observation that cooperativity does not occur (Fig. 6.23) or is negative (Fig. 6.20) when the substrate concentration is varied between about 2 and 120 m M on only one side of the membrane (Schnell and Besl, 1984). Hence, in one possible model (Fig. 6.25), positive cooperativity is observed when the substrate concentrations are about the same on each side of the membrane owing to two-site simultaneous exchange. On the other hand, the more the substrate concentrations on opposite sides of the membrane differ, the more likely it is that the subunits in the anion exchange dimer adapt to operate in the same direction

'

?'~ c'i' ...... 100

200

300

,

......

400

Concentration

500

600

700

(mM)

FIGURE 6.22 Substrate inhibition of HCO3- and C1- homoexchange in human red cell ghosts. Transport via anion exchanger 1 (AE1) was measured at various extracellular (A), intracellular (B), or both extracellular and intracellular (C) substrate concentrations. The lines shown were obtained by nonlinear least-squares fit of experimental data assuming Michaelis-Menten (A) or modified (B and C) Michaelis-Menten kinetics (see text for further discussion). Osmotic balance was maintained at different substrate concentrations by changing the sucrose concentration (adapted from Brahm et al., 1992, with permission from Elsevier Science).

simultaneously. In the latter case, electrostatic repulsion between anions may lead to the observed negative cooperativity for uptake and exodus. Presumably the single round of C1- exodus observed by Jennings (1982) to occur from red cell ghosts with a stoichiometry of about one C1- ion per AE1 monomer can occur when monomers in AE1 dimers operate together in the same direction. Anion exchange owing to transport of two

204

6. Transport Proteins That Propagate Solute Gradients

x10-7

~

A

8

7

B

~

'7, w

r

m u

o

c

=m

E

m

O

E

4-

4

A

f Jee 1.5

-

|

50 x104

\

2i

i

[s]=

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li

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C

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D

1.0 0.5 -

1

\o [

50 [p]~ 100

,

.t,,

1 J~-~f/

[P],n

I

3 x lO-a

FIGURE 6.23 Exodus of sulfate (A and B) and phosphate (C and D) by homoexchange at 100-mM extracellular substrate concentrations exhibit hyperbolic kinetics. Alternatively, positive cooperativity for the human anion exchanger 1 (AE1) may be obscured by changes in red cell ghost membrane electrical potential that occur in association with changes in substrate concentration (see text for further discussion). Osmotic balance was maintained at different substrate concentrations by changing the potassium citrate concentration (adapted from Schnell and Besl, 1984, with permission from Springer-Verlag).

anions in one direction at a time by an AEl-dimer is, nevertheless, a ping-pong-like mechanism, and even two-site simultaneous exchange by a dimer is ping-ponglike for each monomer considered separately. 5. Is a Combined Ping-Pong and Two-Site Simultaneous Model Feasible?

Band 3 dimers clearly function allosterically, and their allosteric function is asymmetric. Hence, it is not surprising that the kinetics of the transport that they catalyze do not reflect simple ping-pong or two-site simultaneous processes in which hyperbolic transport kinetics are expected to obtain. Nevertheless, one can begin to understand the mechanism by which transport may occur by considering these two types of transport mechanisms and the complex kinetic data for actual transport described above. More rigorous mathematical accounts of many of these data and their fitting to a transport model have been published elsewhere (Salhany 1990, 1992, 1996).

In the first model presented here, the alternative interpretations of either multiple transport activities or negative cooperativity (Figs. 6.19 to 6.21B) can be combined to account for anion uptake at constant intracellular anion concentration. In this model, the AE1 protein is proposed to progress from a low Km system of limited capacity at a low extracellular substrate concentration to a high Km and high-capacity system as the extracellular substrate concentration is raised. Consequently, the system would be able to catalyze anion exchange at a rate appropriate to existing extracellular anion concentrations. 9 A similar phenomenon could conceivably occur for exodus at least at relatively high intracellular sub9 Likewise, transport proteins that were discussed previously (e.g., CAT proteins, Section X,E of Chapter 4) could conceivably have curved Hofstee plots (Fig. 4.34) because they adapt to the substrate concentration to which they are exposed. We propose here that the Km and Vmax values may in some cases increase as the substrate concentration is raised. Such an increase in the values of the kinetic parameters would help to insure that transport would be responsive to changes in substrate concentrations and would continue to occur at proportionally significant rates regardless of the absolute concentration of substrate.

Erythroid and Nonerythroid AE Expression

A

B ~" X10-7

xlO-Z

//~

=.

.

4

m

./'

2

/

w m

'-,.,,,,

.

/

A

205

o c~

50

IS]

I

100

I

,,

I,

0.5

1.0

1.5xlt~ I

0.5

~p] 1.0

1.5x104

!

c

=l

E

m

0

E

:r

C

D

xl0.~

xl0 4

1.5

1.5-

1.0

0.5

t

/

./ ./ 50

o/"

1.0-

0.5-

|,,

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100

FIGURE 6.24 Exchange of sulfate (A and B) and phosphate (C and D) exhibit positive cooperativity when the intracellular and extracellular substrate concentrations are varied simultaneously between 2 and 100 mM. Models to reconcile these results with the negative cooperativity observed under other conditions are presented in Fig. 6.25 and in the text. Osmotic balance was maintained at different substrate concentrations by changing the sorbitol concentration (adapted from Schnell and Besl, 1984, with permission from Springer-Verlag).

strate concentrations (Fig. 6.21A). This negative cooperativity becomes positive, however, when the kinetics of both uptake and exodus are examined simultaneously by changing the intracellular and extracellular concentrations together in equilibrium exchange experiments (Fig. 6.24) Under equilibrium exchange conditions, one subunit of the AE1 dimer apparently faces the inside and the other the outside of cells such that association of an anion with each site leads to their cooperative simultaneous exchange by the dimer (Fig. 6.25B). Each monomer may, however, be viewed as operating via a pingpong-like mechanism in that it transports anions in only one direction at a time. In this model, the AE1 dimer may adapt to transport one or two anions in the same direction as the intracellular and extracellular substrate

concentrations become more dissimilar. When only the intracellular or extracellular substrate concentration is changed, positive cooperativity (Fig. 6.24) appears to be lost (Fig. 6.23) or even to become negative (Fig. 6.20). Negative cooperativity would occur if the binding of the second anion by the AE1 dimer favors dissociation of one or both of the anions before they are transported (Fig. 6.25A). An interesting alternative to this first model is to suppose that positive cooperativity always occurs, but that it is obscured when only the intracellular or the extracellular substrate concentration is varied. In this second model, we propose that positive cooperativity may be obscured by changes in the membrane electrical potential. It may be obvious to the reader that membrane electrical potential can influence the kinetics of electrogenic bio-

206

6. Transport Proteins That Propagate Solute Gradients

A

B outside

outside

inside

inside

outside

.........

inside

outside

.........

inside

FIGURE 6 . 2 5 Proposed models for anion exchange by the band 3 protein (AE1). When uptake is measured at various extracellular substrate concentrations ([Sout]) and a constant intracellular one ([Sin]), cooperativity appears to be negative. When, however, exodus is measured at various [Sin ] but constant [Sout], cooperativity becomes negative only at very high [Sin]. According to part A of the first model proposed here, this difference in the detection of negative cooperativity for uptake and exodus could result from the conformational change in the band 3 dimer diagramed in A. In the diagram, the pathways for anion migration through each monomer are shown to converge on the outside but to diverge on the inside of the membrane. Consequently, anions to be transported would repel each other more during uptake than during exodus. Transport by the dimer shown in A would occur when an anion is bound to one or both of the monomers, but simultaneous transport is less likely in part because anions bound at both sites foster their own dissociation before transport can occur. Part B of this same model is proposed to apply when the anion concentrations are set equal and varied together on both sides of the membrane. Under these conditions, the membrane electrical potential was found to be near zero, and positive substrate cooperativity was observed in the red cell ghosts used for the studies (Schnell and Besl, 1984). It is proposed in this first model that when the substrate concentration gradient is small, the band 3 dimer adopts a conformation in which the monomers face substrate on opposite sides of the membrane (B). In this case, transport is facilitated when substrate is bound to both subunits. Alternatively, it is proposed in a second model that the membrane electrical potential influences the effective anion concentration in addition to its known effect on anion total chemical potential. Although total chemical potential is a more precise and fundamental concept than effective concentration, effective concentration must be considered in plots of kinetic data because these plots are generated using concentrations rather than total chemical potentials (e.g., Figs. 6.19 to 6.24). Since the membrane electrical potential has been observed to increase in magnitude as the actual anion concentration gradient across the membrane is increased, the effective anion concentration would become greater than its actual concentration by a proportionally larger amount as the actual concentration is lowered below the constant anion concentration on the other side of the membrane (see text). Consequently, the relationship between transport velocity and effective anion concentration would according to this model be sigmoidal rather than hyperbolic (Fig. 6.23) and rather than the apparent combination of two or more rectangular hyperbolas (Fig. 6.20). Similarly, the Hofstee plots of these sets of data in which transport velocity divided by effective anion concentration is plotted on the abscisa would curve back to the left at their bottom owing to positive substrate cooperativity (e.g., as in Fig. 6.24). This second model makes no prediction about whether transport occurs primarily as shown in A, as shown in B, or in some other way. Regardless of the details of the transport process, however, transport would always exhibit positive cooperativity in relation to the effective substrate concentration.

membrane transport. We contend, however, that membrane electrical potential may also affect nonelectrogenic obligatory anion exchange. We suggest that changes in the membrane electrical potential influence the effective anion concentrations as well as the values of their total chemical potential. 1~ Consequently, transport kinetics deduced using the values of the actual anion concentra10 A further semiquantitative account of this theory is given in Section III,B of Chapter 7.

tions may not properly reflect the characteristics of transport. For example, increasing the magnitude of an insidenegative membrane electrical potential increases the values of the total chemical potential of anions on the inside of the membrane and decreases these values of anions on the outside. We propose that a change in membrane electrical potential may change the effective intracellular and extracellular anion concentrations in

Enythroid and Nonerythroid AE Expression

the same manner that it changes their total chemical potentials. 11 Consequently, if an inside-positive membrane electrical potential becomes larger as the extracellular anion concentration is lowered, then the effective extracellular anion concentration (or the tendency of anions to enter the cell) would decrease to a lesser extent than expected from the decrease in actual anion concentration alone. In this regard, Shnell and Besl (1984) showed that the membrane electrical potential ranged from about + 140 mV to about - 140 mV under experimental conditions in which the intracellular and extracellular anion concentrations, respectively, exceeded the anion concentration on the other side of the membrane by the greatest amount. In other words, when the external anion concentration was lowered to characterize anion uptake by red cell ghosts, the membrane electrical potential became increasingly more positive on the inside. Similarly, an increasingly larger outside-positive membrane electrical potential was generated under the reverse conditions in which the intracellular substrate concentration was lowered below that of extracellular substrate in order to determine the kinetics of anion exodus. According to these data and the model under discussion, the effective anion concentration would become proportionally higher as its actual concentration is lowered owing to an increase in the magnitude of the transmembrane electrical potential where the positive side of the electric field is on the other side of the membrane. Such changes in the effective substrate concentrations would, if large enough, render the relationships between the rate of transport and the effective substrate concentration sigmoidal for uptake (Fig. 6.20) and for exodus (Fig. 6.23) as well as for equilibrium exchange (Fig. 6.24). That is, the lower the actual substrate concentrations in Figs. 6.20 and 6.23, the greater the proportion by which they should be increased to represent the effective anion concentrations. If the substrate concentrations in Figs. 6.20 and 6.23 should actually be proportionally larger as they are decreased, then not only would the relationships between velocity and effective substrate concentration be sigmoidal, but the resultant Hofstee plots also would bend back to the left at their bottoms as is the case for positive substrate cooperativity (e.g., Fig. 6.24). Let us now summarize these two models for the observed AE1 transport data. In the second model just 11 In a similar vein, Stein (1990), while considering the bioenergetics of Na+K+ATPase-catalyzed transport, concluded that introduction of an inside-negative membrane electrical potential where none had been present would reduce the effective Na + concentration at the cytosolic face of the plasma membrane. Hence, although it may be unconventional to propose that membrane electrical potential may influence effective concentrations of cations and anions on both sides of the membrane, such a notion is not new.

207

discussed, the differing transport mechanisms shown in Figs. 6.25A and 6.25B are irrelevant since positive cooperativity is proposed to occur under all experimental conditions. Positive cooperativity could conceivably occur by simultaneous transport of two anions either in the same (Fig. 6.25A) or opposite (Fig 6.25B) direction. Positive cooperativity may be obscured, however, by changes in the membrane electrical potential in experiments in which the substrate concentration is varied on only one side of the membrane. In this model, binding of anions to the two substrate receptor sites as shown in Fig. 6.25 would not result in electrostatic repulsion between the anions. In the first model discussed above, however, transport of one or two anions in the same direction is proposed to exhibit negative cooperativity because of electrostatic repulsion between the two anions (Fig. 6.25A). Conversely, simultaneous transport of two anions in reverse directions is proposed to produce positive cooperativity (Fig. 6.25B). Thus, the mechanism of transport by AE1 dimers is proposed in this model to change from simultaneous transport of one or two anions in the same direction when the substrate concentrations on each side of the membrane are widely different (Fig. 6.25A) to one anion in each direction when these concentrations are nearly the same (Fig. 6.25B). In both models, the substrate concentrations are proposed to influence the observed kinetics of transport albeit by different mechanisms. The concept that the kinetics and stoichiometry of transport might change in response to substrate concentration was first presented in Chapter 4 for Na § dependent glutamate transporters (Section XI,F). The stoichiometry of Na+/glutamate cotransport was proposed to increase from two to three as the extracellular glutamate concentration is lowered in order first to accomplish bulk glutamate transport and then to reduce the glutamate concentration to a level that would not favor binding to its receptor. We return now to a more detailed consideration of the structure and function of these transport proteins. These transporters comprise a subfamily of proteins that catalyze uptake of Na § anionic amino acids, and probably H § together in exchange for K § The extrusion of K § is thought to be needed for reorientation of the transporters to receive the three cosubstrates for uptake (e.g., Kavanaugh et al., 1997). The proteins are therefore believed to operate by a ping-pong mechanism for antiport, while they also catalyze symport of Na § H § and anionic amino acids by a simultaneous mechanism. As we shall see, however, existing data are as yet insufficient to support final conclusions concerning the details of the transport mechanism or even the precise stoichiometry of transport.

208

6. Transport Proteins That Propagate Solute Gradients

!I1. ASC AND EXCITATORY (ANIONIC) AMINO ACID TRANSPORTERS COMPRISE ONE OF TWO KNOWN FAMILIES OF MAMMALIAN Na+/AMINO ACID SYMPORTERS The Na+-dependent amino acid transport proteins that are known to be expressed in humans and other mammals fall into two evolutionarily unrelated families. Members of the first family are Na + and C1- dependent (Amara, 1992), and they catalyze transport of a wide variety of known and putative neurotransmitters, some of which are amino acids (Van Winkle, 1993). Some members of this family are also expressed in tissues outside the nervous system where they perform additional functions such as cellular volume regulation. The stoichiometry of transport appears to be one or two Na + ions per organic solute molecule for this first family of transporters, whereas the stoichiometry for the second family may be 2 or 3 Na + ions per organic molecule (but see also Sections III,B and III,C below). This second family is the main topic of the present section. Transport proteins in the second family have a wide variety of names in different species owing to the timing of their discovery and their perceived physiological functions. To simplify the discussion here, we use the human designations excitatory amino acid transporter (EAAT) or system ASC-like transporter (ASC) for equivalent proteins in all mammalian species. 12The first mammalian members of the EAAT/ASC family were discovered in brain, although these proteins are now known to be expressed in a variety of tissues outside this organ. Members of the second family cotransport either anionic (EAAT subfamily) or zwitterionic (ASC subfamily) amino acids with Na + and probably H +, but none of the known mammalian members of this family prefers organic substrates that are not amino acids. Although the EAAT/ASC proteins also function as channels to transport CI- (see Section IV,C below), their transport of amino acids appears usually not to be C1dependent (reviewed by Amara, 1992 but see also Van Winkle, 1993). A. Evolution of the EAAT and ASC Subfamilies The E A A T and ASC subfamilies show about 40% similarity in their amino acid residue sequences (Fig. 6.26), but it is currently difficult to guess when they evolved from a common ancestor. Since the family is distantly related to H+/glutamate and H+/Na+/glutamate symporters in bacteria (Kanner, 1996), it is tempting 12 This family is termed the Glutamate : Na + symporter (ESS) family in Chapter 8 (number 2.27 in Table 8.2).

A 2

EAAT 3

5

ASC 1

4

1

2

I

3

I

93%

81% 72%

55%

16o% 42%

FIGURE 6 . 2 6 Amino acid residue sequence similarities among the human forms of five Na+-dependent anionic amino acid transporters (EAAT1 to EAAT5) and three Na+-dependent zwitterionic amino acid transporters (ASC1 to ASC3). The mouse form of ASC2 is shown here because the human version either has not yet been identified or is ASC3 (ASC3 is the B ~ transporter cloned by Kekuda et al., 1996; see text for discussion). The partial sequences shown contain 256 residues in regions of the proteins that were selected because they contain relatively high similarities and only one gap (EAAT3 has only 255 residues in this region). The first residues shown corresponds to residues 239 (EAAT1), 238 (EAAT2), 209 (EAAT3), 264

(EAAT4), 218 (EAAT5), 219 (ASC1), 239 (ASC2), and 227 (ASC3)

of the proteins. The proteins comprise a family composed of two subfamilies, termed E A A T and ASC, within each of which there is greater than 50% identity (A) for the partial sequences shown (B), even among the most distantly related members of each subfamily. (A) Percentage identities among the sequences (or average percentage identities in cases where more than two sequences are compared). (B) Sequence alignments (full sequences shown in Kekuda et al., 1996; Utsunomiya-Tate et al., 1996; and Arriza et al., 1997). Identical amino acid residues at the same positions in the proteins are usually (but not always) shown as white letters on a black background.

to speculate that the ancestor to the two subfamilies catalyzed H+-dependent and perhaps Na+-dependent anionic amino acid transport. Potassium ion countertransport may have evolved later in the E A A T line, since a critical glutamyl residue that is needed for K + transport (Kavanaugh et al., 1997) is instead a glutaminyl residue both in the ASC (e.g., Utsunomiya-Tata et al., 1996) and in the bacterial (Tolner et al., 1992 a,b, 1995a) transporters (but also see discussion below). The ASC subfamily is composed of at least two and probably three known members in mammals. We define ASC3 here as the B ~ transporter that was cloned by Ganapathy and associates from a human placental choriocarcinoma cell line (Kekuda et al., 1996). While these authors point out interesting differences in the relative substrate selectivities of ASC3 and other transport proteins in this subfamily, the amino acid residue sequence of ASC3 (Kekuda et al., 1996) is about 80% homologous to mouse ASC2 (Utsunomiya-Tate et al., 1996). More-

ASCand ExcitatoryAminoAcidTransporters

209

B ASC3 ASC1 ASC2 EAAT4 EAAT3 EAAT2 EAAT5 EAAT1

ASC3 ASC1 ASC2 EAAT4 EAAT3 EAAT2 EAAT5 EAAT1

' ~ ' ~

ASC3 ASC1 ASC2 EAAT4 EAAT3 EAAT2 EAAT5 EAAT1

ASC1 ASC2 EAAT4 EAAT3 EAAT2 EAAT5 EAAT1

20 i

!

R A I LFJ l i . l l aALl e l ~l ' i : l m K

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LI ~ A I n l G I

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nA vv rA4 K.mm~.N. ~P!Js

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vV~.~..~v !i~l vCl~il~' mvJkl[I V

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210

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~

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lq. 190

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10

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FIGURE6.26 (Continued)

over, we have found ASC2 and ASC3 to be 93% homologous across a continuous stretch of about half of their residues (Fig. 6.26). Hence, it is conceivable that ASC2 and ASC3 represent the product of the same gene in mice and humans, respectively (see also Kanai, 1997). If such is the case, however, the highly electrogenic alanine transport apparently catalyzed by the protein that we term here ASC3 (Kekuda et al., 1996, 1997), is in

stark contrast to the nonelectrogenic transport catalyzed by ASC2 (Utsunomiya-Tate et al., 1996 and see Section III,D below). The other member of the ASC subfamily (ASC1) was isolated from human brain nearly simultaneously by two groups of investigators (Shafqat et al., 1993; Arriza et al., 1993). It is about 60% homologous to ASC2 and ASC3 over their entire length, and these proteins

2 | 0

6. Transport Proteins That Propagate Solute Gradients

are 72% homologous in the selected stretch of 256 amino acid residues mentioned above (Fig. 6.26). Each member of the ASC subfamily exhibits transport characteristics that are similar to the zwitterionic amino acid transport system, ASC, first described by Christensen and associates (e.g.; Wheeler and Christensen, 1967). As for the EAAT subfamily (e.g.; Arriza et al., 1997), members of the ASC subfamily also serve as amino acidactivated C1- channels (Zerangue and Kavanaugh, 1996b). This C1- channel activity is particularly variable among members of the E A A T subfamily, and this variability may have evolutionary as well as functional implications. EAAT1,-2, and -3 show similarly low C1- channel activities relative to their abilities to transport anionic amino acids (Arriza et al., 1997). Moreover, these proteins show about 55% sequence similarity in the stretch of 256 amino acid residues considered in Fig. 6.26. In contrast, EAAT4 has predominately C1- channel activity (Fairman et al., 1995), and 81% homology to EAAT1 is detected in the region analyzed. The similarity between EAAT4 and EAAT1 is also higher over their entire sequences than among other combinations of EAAT proteins (summarized by Gegelashvili and Schousboe, 1997). Similarly, EAAT5 has relatively high CI- channel activity (Arriza et al., 1997), and the homologies detected among EAAT5,-1, and -4 are greater than those detected among EAAT5, -2, and -3 (Fig. 6.26A). While the latter differences in the percentages of similarity (i.e., 55 vs. 60% in Fig. 6.26A) are not statistically significant for the segments of the proteins analyzed, EAAT5 has 46% identity with EAAT1 and 43% identity with EAAT4 over their full lengths (Arriza et al., 1997). These values are significantly larger (p < 0.05, contingency tables) than the 36 and 37% identities of EAAT5 with EAAT2 and EAAT3, respectively, over their full lengths. For these reasons, we suggest that first EAAT5 and later EAAT4 evolved through a series of two duplications of the gene encoding an ancestral form of EAAT1. In both instances, the duplicate genes appear to have evolved to catalyze primarily C1- transport via channels rather than anionic amino acid transport. Nevertheless, these C1- channels are activated by anionic amino acids, and these amino acids serve as neurotransmitters in the central nervous system. Consequently, it has been proposed that EAAT4 and EAAT5 may serve primarily as ligand-gated C1- channels (Arriza et al., 1997). Although the common ancestor of EAAT1,-4, and-5 may have served primarily for amino acid transport, it may have had as-yet unappreciated structural characteristics that made it readily adapted to catalyzing mainly C1channel activity. Alternatively, the ancestor of EAAT1,-4, and-5 may have functioned principally as

a C1- channel, and this channel activity may have been reduced in favor of amino acid transport during evolution of EAAT1. Both of these possibilities are consistent with the observation that the potential for reversal of the C1- current owing to transport by EAAT1 is closer to potentials for reversal of the currents resulting from transport by EAAT4 and EAAT5 than are the potentials for reversal of the currents resulting from transport by EAAT2 and EAAT3 (Wadiche et al., 1995a; Fairman et al., 1995; Arriza et al., 1997). 13 As for the differences within the E A A T subfamily, differences between the primary structures of the E A A T and ASC subfamilies (Fig. 6.26) also must account for their distinctive substrate selectivities. The ASC proteins transport zwitterionic amino acids under physiological conditions, whereas members of the E A A T subfamily transport anionic ones. Moreover, the E A A T proteins extrude K § in exchange for anionic amino acids and their cosubstrates, whereas the ASC proteins apparently do not transport K § The ability of E A A T proteins to countertransport K § can, however, be lost through mutation of a single glutamyl residue (Kavanaugh et al., 1997). Consequently, the presence of a glutaminyl (Q) residue rather than a glutamyl (E) residue at the corresponding position in ASC proteins could account for their inability to transport K § (see Section III,D below). 14 Moreover, system ASC begins to select anionic amino acids over zwitterionic ones for transport when the pH approaches a value of 5 (Vadgama and Christensen, 1984; Christensen, 1984), and the ASC2 protein exhibits a similar characteristic (Utsunomiya-Tate et al., 1996). This change in substrate selectivity of system ASC at reduced pH appears to result from titration of its transport protein rather than its amino acid substrates (Vadgama and Christensen, 1984; Christensen, 1984). Hence, the structural differences producing different substrate selectivities in ASC and E A A T transport proteins may be small. 15 13 Additional functional diversity may be produced in the E A A T proteins through alternative splicing of gene transcripts at least in the case of EAAT2 (Meyer et al., 1998). Functional differences among three putative EAAT2 subisoforms is currently under investigation. 14 Conversion of an adjacent tyrosyl residue in E A A T proteins to a phenylalanyl residue also destroys K § transport (Zhang et al., 1998). Such a difference also could account for the inability of ASC proteins to transport K § (see Section III,D). 15 We do not intend to imply that such small differences have little importance to evolution of different protein functions. In fact, according to a current model of molecular evolution, significant changes in, say, the substrate selectivity of a biocatalyst are proposed to occur suddenly, often as a result of substitution of one or a few amino acid residues (Dean, 1998). The loss or gain of K § as a substrate for transport by members of the ASC and E A A T subfamilies, respectively, may be one such sudden evolutionary change in function. As discussed here, this change in substrate selectivity appears to occur as a result of a single amino acid residue substitution.

ASC and Excitatory Amino Acid Transporters

The ASC and E A A T proteins also differ functionally in that the ASC proteins appear to transport only 1 Na + ion per amino acid molecule (Utsunomiya-Tate et al., 1996; Zerangue and Kavanaugh, 1996b), whereas the E A A T proteins seem to transport 2 or 3 (e.g., Zerangue and Kavanaugh, 1996a). These conclusions are, however, based entirely on kinetic and thermodynamic data for the effect of the Na + concentration on amino acid transport. As pointed out in Chapter 4, this stoichiometry should be measured rather than inferred from such effects of one cosubstrate on transport of another. Similar types of effects are also used without adequate justification to deduce the order of substrate binding. We shall see in Section III,B below that the actual stoichiometry of transport may differ considerably from that which is predicted from the effect of one cosubstrate on the transport of another. Let us now consider the structure and function of transport proteins first in the ASC and then in the E A A T subfamilies. B. A Lesson on the Study of Transport from the ASC Systems and Transporters ~6 In this decade, molecular techniques have been used to revolutionize the study of amino acid transport. Owing perhaps to this rapid expansion of knowledge, each new discovery of an amino acid transporter has not led to its full functional characterization and comparison of its characteristics to those of a transport system that appears to employ it. Some characteristics are, of course, more important than others in determining whether a protein is likely to be the catalyst in a transport system, and unnecessary effort should not be expended in demonstrating such parallels. Much may still be learned about which characteristics to select for study, however, by consulting the earlier literature. One such lesson is described in this section for determining the stoichiometry of symport and antiport by the ASC systems and transporters. As we shall see, this stoichiometry can be determined reliably only by measuring solute fluxes directly. Indirect methods involving the measurement of induced current or apparent cooperativity will not suffice alone, contrary to conclusions drawn from such studies in many otherwise outstanding papers published in this decade. We begin our discussion of the transport catalyzed by the ASC systems and proteins with a description of their structure. Knowledge of the relationship of this structure to symport of Na + and amino acids is only 16 We use the term "transporter" when the transport under investigation can be attributed to a protein of known primary structure. When the protein has not been identified or for the results of transport studies that preceded isolation of the protein we use the term "system" to refer to the catalytic entity.

21 1

now emerging from molecular studies. While such studies may further inform us about the transport mechanism, they are best formulated with accurate knowledge of the transport process under investigation. For example, one becomes better able to plan studies to determine, say, the structure of the substrate receptor site when one knows from thorough transport experiments how many ions and molecules may be received at the site. Hence, the study of structure and function cooperatively advance each other and neither can be fully understood alone.

1. Structures of the ASC Proteins

Since the ASC transport proteins were identified only a few years prior to this writing (Arriza et al., 1993; Shafqat et al, 1993; Utsonomiya-Tate et aL, 1996; Kekuda et al., 1996), much remains to be learned about their secondary, tertiary and quaternary structures. As for the P-type ATPases (Chapter 5) and the anion exchange (AE) proteins (Section II above), the most uncertainty in the secondary structures of the ASC and E A A T proteins is in the C-terminal portions of their membrane associated domains (Kanai, 1997). The hydropathy plots for ASC proteins (e.g., Fig. 6.27) are very similar to those for the E A A T proteins (e.g., Fig. 6.28). While the first six putative membrane traverses stand out clearly in these plots, it is less certain whether the seventh peak represents a membrane spanning region. Moreover, the long hydrophobic stretch (LHS, Figs. 6.27 and 6.28) undoubtedly spans the mem-

5.00 4.00 3.00 1 2 3 4 5 6 7 LHS 2.00 1.00 0.00 -1.00 -2.00 -3.00 -4.00 -5.00 . ~ . . ~ 1 ~ ~ J ~ l ~ L ~ . . _ _ J ~ 1 101 201 301 401 501 n

-

553

FIGURE 6.27 Hydrophobicity plot of mouse ASC2. The plot was constructed according to the Kyte-Doolittle hydropathy analysis using a window of 21 amino acid residues. Numbers correspond to putative transmembrane segments of hydrophobic regions. Putative span 7 may, however, be too short actually to span the membrane as an a-helix and it is unclear how many spans are contained within the long hydrophobic stretch (LHS). Because of this uncertainty, proteins in this family have been proposed by different investigators to contain a total of 8 or 10 transmembrane segments (see Fig. 6.35) (adapted from Utsunomiya-Tate et al., 1996, with permission from American Society for Biochemistry & Molecular Biology, Inc.).

212

6. Transport Proteins That Propagate Solute Gradients 1

2

3

$113 (rat EAAT2)

4

5

6

7

LHS

H326 Y403 R479 (rat EAAT2) (rat EAAT2)(rat EAAT1) E404 (rat EAAT2)

FIGURE 6.28 Hydrophobicity plot of rabbit EAAT3. The plot was constructed according to the Kyte-Doolittle hydropathy analysis using a window of 21 amino acid residues. Seven putative transmembrane segments are numbered although the actual number of such segments may total 8 or 10 depending in some instances on whether the 7th segment is considered actually to transverse the membrane and whether the long hydrophobic stretch (LHS) is assumed to contain 2 or 3 transmembrane segments (e.g., Fig. 6.35). Positions corresponding to pertinent amino acid residues in other proteins in this family are also indicated in the figure (i.e., Sl13, H326, Y403, and E404 of rat EAAT2 and R479 of rat EAAT1) (adapted from Kanai, 1997, with permission from Current Biology Ltd.).

brane, although it is as yet unclear how many times (Gegelashvili and Schousboe, 1997). Two predicted N-glycosylation sites are present in the relatively large extracellular loops between the putative transmembrane segments 3 and 4 of all three ASC proteins. Moreover, each of these proteins contains several possible protein kinase-mediated phosphorylation sites (Shafqat et al., 1993; Utsunomiya-Tate et al., 1996; Kekuda et al., 1996). Interestingly, a putative leucine zipperlike motif in ASC1 is not present in ASC2 or ASC3. It has not yet been determined whether this motif in ASC1 allows it to form oligomers in biomembranes. As discussed in Section II,B above, the transport kinetics of oligomeric proteins are more likely to exhibit allosteric effects than are the transport kinetics of monomeric ones.

Alanine uptake by the ASC transporters exhibits hyperbolic kinetics (e.g., Fig. 6.29). Similarly, alanine uptake increases in a hyperbolic manner as the Na + concentration is raised (e.g., Fig. 6.30). By these criteria, the stoichiometry of Na+/amino acid cotransport appears to be one to one (Zerangue and Kavanaugh, 1996b; Utsunomiya et al., 1996). When the actual stoichiometry of Na + and amino acid cotransport is measured, however, the results may be quite different than anticipated from such kinetic analyses (Wheeler and Christensen, 1967). For example, as for the ASC transporters (Fig. 6.30), hyperbolic kinetics also obtain for the influence of the Na + concentration on alanine transport by system ASC in pigeon erythrocytes (Fig. 6.31). These data are consistent with the conclusion that the stoichiometry of alanine and Na § cotransport is 1 to 1. In contrast, the molar quantity of Na + actually transported is about 2.5-fold greater than the quantity of alanine transported (Table 6.3). Hence, 2 or 3 Na + ions are transported with every alanine molecule by system ASC (Wheeler and Christensen, 1967). On the other hand, Na+//3-alanine symport by system/3 occurs with a stoichiometry of about 1 to 1 (Table 6.3), although the relationship between/3alanine transport and the Na + concentration (Fig. 6.32) indicates that more than 1 Na § ion is transported with each /3-alanine molecule (Wheeler and Christensen, 1967). Only in the case of Na+/glycine symport by system Gly does the stoichiometry inferred from the influence

o o o

8 (~

2"

6 4

2. Kinetics of ASC-Catalyzed Symport and Antiport Neither system ASC (Wheeler and Christensen, 1967) nor the ASC1 transport protein (Zerangue and Kavanaugh, 1996b) can use a Na + total chemical potential gradient to drive accumulation of zwitterionic amino acids against a gradient. Rather, the symporter also appears to catalyze obligatory exchange of intracellular Na + and amino acids for these substrates outside cells. Consequently, a gradient of one amino acid could form a gradient of another through obligatory exchange, but the ASC-catalyzed Na § fluxes appear to be the same in both directions.

~

10

9

F= 8 --

o

6

>

2 0

2

'

0

0.2

V/S ( p m o l / m i n 0

I

50

I

100

I

150

I

200

i|

,

o.4

0.6 9IJ.M)

I

250

I

300

L-Alanine (~M)

FIGURE 6.29 L-Alanine transport exhibits hyperbolic kinetics in Xenopus oocytes expressing ASC2. Virtually all transport was Na + dependent and attributable to ASC2 expression. Either a Hofstee plot (inset) or nonlinear regression analysis can be used to calculate Km and Vmax values of about 20 /xM and 11 pmol oocyte -1 min -] respectively (adapted from Utsunomiya-Tate et al., 1996, with permission from American Society for Biochemistry & Molecular Biology, Inc.).

2.13

ASC and Excitatory Amino Acid Transporters

A .=_ E

§

__J 0 0 0 0

E 0

B

_

"E

0.8

o

0.6

Q.

"O

.c_

N

E L

o.4

e-

<,

c

0

0.2

N

I

20

I

i

I

40

i

60

I

I

i

80

'i

100

~.......~------~

0

0

-------------O

I

I

I

I

20

40

60

80

Na § (mM)

I

100

[Na +] mM

FIGURE 6.30 L-Alanine uptake by Xenopus oocytes expressing ASC2 (A) or ASC1 (B) increases in a hyperbolic manner as the Na + concentration is increased. Virtually all transport is Na + dependent and attributable to ASC2 or ASC1 expression. (A) ASC2 catalyzed alanine uptake from a 50-/zM solution (adapted from Utsunomiya-Tate et aL, 1996, with permission from American Society for Biochemistry & Molecular Biology, Inc.). (B) Normalized current induced by 300/xM alanine and the indicated concentration of Na + owing to the C1- channel activity of ASC1. The magnitude of the C1- current appears to be proportional to the amount of concomitant Na+/alanine cotransport (adopted from Zerangue and Kavanaugh, 1996b, with permission from American Society for Biochemistry & Molecular Biology, Inc.).

of the Na + concentration on glycine transport (Fig. 6.33) match the actual stoichiometry of transport reasonably well (Table 6.3). a7 Such results appear to require different kinetic models for the transport catalyzed by systems ASC,/3, and Gly. The transport catalyzed by system Gly appears, however, to fit currently accepted models in which the actual stoichiometry of glycine and Na § transport is consistent with the stoichiometry deduced from the influence of the Na § concentration on glycine transport (Table 6.3). Hence, we need to develop additional models here only for the cases of systems ASC and/3. The reader is reminded that the detailed mechanisms of transport are in all cases still under investigation. For Na+-dependent/3-alanine transport by system/3, more than 1 Na § ion as well as a/3-alanine molecule apparently must bind to the transport system before Na § and/3-alanine can migrate across the membrane. 17 While some transport activities that were at one time believed to be single systems have subsequently been found to be heterogeneous, no such evidence for such heterogeneity has, to our knowledge, been published for systems ASC, Gly, and/3 in pigeon erythrocytes. Moreover, the lack of correspondence between the actual stoichiometry of transport and the stoichiometry anticipated from the influence of the Na + concentration on transport of its cosubstrate has now been described for a transport protein (i.e., the Na+-dependent betaine and G A B A transporter, BGLT-1; BrOer and Lang, personal communication).

Each of these Na + ions appears to bind at least as well as the others since the relationship between/3-alanine transport and the Na + concentration is consistent with positive cooperativity of Na + binding (Fig. 6.32). It appears, however, that only 1 of the Na + ions is bound to a site from which it may be transported with the /3-

=

~ ~

0.14

I

'

'

[ala] mM

0.07

03

,

0

I

,

I

35

I

70

[Na+], m M

FIGURE 6.31 Effect of the Na + concentration on L-alanine uptake by system ASC in pigeon erythrocytes. The relationship between transport and Na + concentration appears to be hyperbolic at all alanine concentrations studied (adapted from Wheeler and Christensen, 1967, with permission from American Society for Biochemistry & Molecular Biology, Inc.).

214

6. Transport Proteins That Propagate Solute Gradients

TABLE 6.3

Inconsistencies between the Kinetic Dependencies of Amino Acid Transport on Na + Concentration (Column A) and the Measured Stoichiometries of Na+/Amino Acid Cotransport (Column B) in Pigeon Red Blood Cells a (A) Apparent kinetic dependency of amino transport on Na + concentration b

Transport system (amino acid substrate)

Assumption that can resolve inconsistency between findings summarized in columns A and B

(B) Mean +_ SEM stoichiometry of Na+/amino acid cotransport c

Gly (glycine)

Higher than first order (n > 1)

1.53 _+ 0.09 (N = 25)

None needed

ASC (L-Alanine)

First order (n = 1)

2.52 +_ 0.07 (N = 24)

Transport of only one of the Na + ions is detected kinetically

/3 (/3-Alanine)

Higher than first order (n > 1)

0.96 +_ 0.11 (N = 7)

Only one of the participating Na + ions is actually transported

aAdapted from Wheeler and Christensen (1967) with permission from American Society for Biochemistry & Molecular Biology. bValues of the Hill coefficient, n, in Michaelis-Menten type equations of the form: Vi(AA) = Vmax [Na+]n/([Na+] n 4- K0.5n) are sometimes unjustifiably concluded to equal the stoichiometry of Na + cotransport regardless of whether the measured transport velocities are for Na + or, as in this case, an amino acid (AA) cosubstrate. CMean +_ SEM stoichiometries (also known as coupling ratios) were calculated from multiple determinations (N) of the increase in initial velocity of Na + uptake when the indicated amino acid was added as compared to the increase in initial velocity of amino acid uptake when Na + was added. Tracers were 22Na+ and 14C-labeled amino acids.

a l a n i n e m o l e c u l e , p e r h a p s a f t e r e n o u g h N a + ions h a v e b o u n d e l s e w h e r e to t r i g g e r t r a n s p o r t . In c o n t r a s t , b i n d i n g of t h e last N a + ion for t r a n s p o r t by s y s t e m A S C s e e m s to b e c o n s i d e r a b l y w e a k e r t h a n t h e b i n d i n g of t h e 1 or 2 N a + ions t h a t m a y p r e c e d e it. This w e a k e r b i n d i n g of t h e last N a + ion c o u l d r e s u l t f r o m t h e e l e c t r o s t a t i c r e p u l s i o n t h a t is a n t i c i p a t e d to o c c u r a m o n g N a § ions t h a t n e e d to b i n d n e a r to e a c h o t h e r in o r d e r to b e t r a n s p o r t e d t o g e t h e r by s y s t e m A S C . This e l e c t r o s t a t i c r e p u l s i o n m a y b e a n a l o g o u s to

0.04

.

.

.

.

.

I

t h a t w h i c h a p p e a r s to o c c u r a m o n g K + ions d u r i n g transp o r t via a c h a n n e l ( J a n a n d Jan, 1997 a n d see also C h a p t e r 7). B i n d i n g of t h e last N a + i o n by s y s t e m A S C c o u l d also t r i g g e r its s y m p o r t with t h e o t h e r N a § ions a n d alanine. Similarly, it has b e e n p r o p o s e d t h a t b i n d i n g of a d d i t i o n a l K § ions to a c h a n n e l p r o t e i n facilitates d i s s o c i a t i o n a n d c o n s e q u e n t l y t r a n s p o r t of K § ions t h a t a r e a l r e a d y b o u n d to t h e p r o t e i n ( J a n a n d J a n , 1997). In a n y case, if t h e b i n d i n g of t h e last N a § to s y s t e m A S C is by far t h e w e a k e s t , t h e n h y p e r b o l i c r a t h e r t h a n s i g m o i d a l kinetics s h o u l d o b t a i n for t h e r e l a t i o n s h i p between alanine transport and the Na § concentration. It m i g h t b e n o t e d also t h a t a l t h o u g h i n o r g a n i c ions satu-

M, ~-al C

I

) "~

//~

o.o

~

0.2

1

" O.lO

o.,

~

"~

o.4 0.0

0.0

i

17.5 35

70

[Na*], mM

~

0.05

0.2

~

140

FIGURE 6.32 Relationship between the Na + concentration and/3alanine uptake by system/3 in the pigeon red blood cell. Nonsaturable uptake has been deducted from total uptake to produce the data presented. Since a hyperbolic relationship is first order in regard to the Na + concentration, the curves shown reflect a higher-than-first order dependence of /3-alanine uptake on the Na + concentration (adapted from Wheeler and Christensen, 1967, with permission from American Society for Biochemistry & Molecular Biology, Inc.).

~>~

0

35

70

[Na*], mM FIGURE 6.33 Sigmoidal relationship between the Na + concentration and glycine uptake by system Gly in pigeon erythrocytes. Since a hyperbolic relationship is first-order in regard to the Na + concentration, the curves presented reflect a higher-than-first-order dependence of glycine uptake on the Na + concentration (adapted from Wheeler and Christensen, 1967 with permission from American Society for Biochemistry & Molecular Biology, Inc.).

ASC and Excitatory Amino Acid Transporters

rate transport via channels in a hyperbolic manner (e.g., Fig. 4.16 in Chapter 4) it should not be assumed that only one ion is transported at a time (see also Section III,B,2 of Chapter 7). Weaker binding of the last ion to be cotransported could conceivably account for the hyperbolic shape of such curves as discussed here for Na + transport by system ASC.

215

translocation

inner surface

*Na*

/

(4~

\.

.q ~Na* Na* , ~/j~, -.E --*S / \ Na* E E

*Na* is preferentially

unloaded inside cells

*Na§

3. A Model for Symport That Does Not Require Simultaneous Entry of all Ions or Molecules in Their Final Stoichiometric Amounts Consideration of all existing data concerning the stoichiometry of substrate cotransport by system ASC provokes modification or replacement of the model for symport developed above for this system. In the case of system ASC, the stoichiometry varies from about 4 Na + ions with each amino acid molecule in the cases of homoserine and threonine to about 4 amino acid molecules with each Na § ion in the case of proline (Thomas and Christensen, 1971; Koser and Christensen, 1971). Consequently, the model for simultaneous cotransport of Na § ions and amino acid molecules developed above must be modified to allow the stoichiometry to vary widely depending on the amino acid species. Moreover, amino acid analogs with sufficiently long and hydrophobic side-chains serve as good competitive inhibitors but not substrates for transport by some forms of system ASC. These relatively large molecules may be inherently poor substrates or they may interfere with Na § binding and consequently its apparently requisite comigration with an amino acid molecule across the membrane. It becomes somewhat difficult, however, to reconcile this concept of a well-defined substrate receptor site with the widely variable stoichiometry that must be proposed for association of different amino acid species and Na § with the site. To account for these observations Koser and Christensen (1971) in collaboration with E.L. Thomas proposed the following model in which all cotransported ions and molecules need not migrate simultaneously. In case one of this model (Fig. 6.34), the radiolabeled Na + ion, is proposed to be more likely than the radiolabeled amino acid molecule (*S) to dissociate from the system, so several labeled *Na § ions are taken up with each labeled molecule. This mechanism could account for the observed stoichiometry of homoserine or threonine cotransport with Na+. In contrast, the second case shown in Fig. 6.34 appears to apply to amino acids, such as proline, several labeled molecules of which are taken up with each labeled Na + ion (Koser and Christensen, 1971). This model needs some modification, however, to account for all of the data of Koser and Christensen (1971). In particular, the unlabeled amino acid molecule

/

9 E @

\.

S *Na§ ~ * S _ . N a . S

E/

\ S

s

*S is preferentially

unloaded inside cells

*Na*

/

(~ E

\.

S . -,,~-Na* +*S Na* E / \ ~"

E

S ~ " Na* + S

*Na* and *S are equally likely to be

unloaded inside cells

FIGURE 6 . 3 4 Scheme to account for different Na+/amino coupling ratios among amino acids cotransported with Na + by system ASC. Only the steps at the internal surface of the membrane are illustrated here. E. L. Thomas participated with B. H. Koser and H. N. Christensen in the development of this Scheme. According to this model, radiolabeled *Na + is preferentially unloaded at the inner surface of the membrane in some cases (diagram 1) while the labeled amino acid (*S) is not unloaded, whereas the reverse is true in other cases (diagram 2). Both of these phenomena as well as the third case (diagram 3) in which both substrates are unloaded (or the substrates have equal probabilities of being unloaded) would account for differences in the coupling ratio that are observed for different amino acid species (see text) (adapted from Koser and Christensen, 1971 with permission from Elsevier Science).

(S) exiting the cell in case 2 is probably of a different species than the labeled one (*S) that enters. This condition must be applied to the model in case 2 because system ASC interacts differently with the amino acid species that is in this case shown as labeled, depending on whether it is on the inside or the outside of the cell. It can be concluded that system ASC treats some amino acids, such as asparagine and proline, asymmetrically because these amino acids influence the stoichiometry of transport of other amino acids with Na + in opposite ways depending upon whether they are on the inside or the outside of the cell. When the coupling ratio (stoichiometry) for Na+:amino acid coexodus is studied in the presence of various extracellular amino acids, proline and asparagine reduce the Na+:amino acid coupling ratio of exiting substrates (Table 6.4). In contrast, intracellular proline and asparagine increase the coupling ratio for couptake of Na + and other amino acids (Table 6.5). In the latter case, however, both asparagine and proline slow the absolute rates of both Na + and amino acid entry. Consequently, although intracellular proline

216

6. Transport Proteins That Propagate Solute Gradients

TABLE 6.4 N a + / A m i n o Acid Coupling Ratios during Their Exodus from Pigeon Erythrocytes via S y s t e m ASC in Exchange for the Indicated Entering A m i n o Acid" Coupling ratio (22Na§ acid) Entering amino acid

Ala exiting

Pro Asn Ala Ser Thr

1.2-1.6 b 1.3-1.6 2.2-2.6 2.2-2.5 2.5-3.2

Thr exiting

2.7, 2.4, 3.8 3.7, 4.3,

3.2 2.9 3.7 4.3

aCoupling ratios were determined by first loading cells with 22Na+ and 3H-labeled amino acids (about 15 and 5 mM, respectively) and then measuring their efflux in medium containing unlabeled Na + (140 mM) and the indicated nonradiolabeled amino acid. The resultant initial velocities of exodus were used to calculate the coupling ratios (adapted from Koser and Christensen, 1971, with permission from Elsevier Science). bWhen a range of values is given, four determinations are included within the range.

TABLE 6.5 Effect of Loading Cells with an A m i n o Acid on the U p t a k e Rates of Na + and t h e S a m e or A n o t h e r A m i n o Acid via S y s t e m ASC in Pigeon Erythrocytes a Coupled entry b Uptake of

Cells loaded with

Experiment 1 2.5 mM Asn

Asn

2.5 mM Asn

Thr

0.5 mM Thr

Asn

0.5 mM Thr

Thr

Na §

Amino acid

Coupling ratio

54 49 57 50 191 190 244 236

64 64 87 82 34 29 78 79

0.85 0.76 0.65 0.61 5.7 6.5 3.1 3.0

and asparagine increase the coupling ratio and thus appear to favor dissociation of *Na § rather than *S (case 1 in Fig. 6.34), they must do so by somehow interacting with the transport system. Perhaps the proline and asparagine molecules are able to displace *S but they are unlikely to exit with bound *Na +. Rather, they may hold the ASC transport protein in the inward-facing conformation and hence slow the overall transport process. Only when another amino acid molecule, such as the *S that had been transported in, displaces proline or asparagine do the cosubstrates become likely to migrate back to the outside of the membrane. By then *Na + may have been displaced by an unlabeled Na + ion as in case 1, thus increasing the Na+:amino acid coupling ratio for uptake of *S. Another mechanism by which the coupling ratio may increase is through an increase in the rate at which the transporter appears to cycle. For example, hydroxyproline has a coupling ratio of about 3.2 as compared to the ratio of about 0.22 for proline (Table 6.6). Moreover, the rate of hydroxyproline transport by system ASC is about sixfold greater than the rate of proline transport (Christensen et al., 1967; Koser and Christensen, 1971; Kilberg et aL, 1981). Hence, hydroxylation of the substrate, proline, appears to increase the maximum rate at which the ASC transporter cycles by more than 80fold (6.0 • 3.2/0.22)! Part of this increased cycling may be viewed as producing an increase in the rate of amino acid uptake, but the other part appears to result in a greater Na+:amino acid coupling ratio. The reader may now recognize that the Na + exchange rate appears also to increase by more than 80-fold when hydroxyproline

TABLE 6.6

N a + / A m i n o Acid Coupling Ratios for U p t a k e via S y s t e m ASC in Pigeon Erythrocytes a Coupling ratio Na§ Mean _ SEM b

Experiment 2 0.5 mM Ser

Pro

0.5 mM Ser

Thr

0.5 mM Thr

Pro

0.5 mM Thr

Thr

177 156 368 418 296 238 484 467

44 40 240 234 19.2 18.5 117 103

4.0 4.0 1.53 1.22 15.4 12.8 4.2 4.6

alllustrative results are shown. The cells were loaded with the amino acid indicated in the second column by incubating them in a 25 mM solution of it for 4.0 to 5.25 hr. After washing the cells twice, the uptake, of 22Na+ and a 3H-labeled amino acid were measured simultaneously. The resultant initial velocities were used to calculate the coupling ratios (adapted from Koser and Christensen, 1971, with permission from Elsevier Science). bin mmol (liter of cells) -1 min -1.

Alanine Serine Threonine Proline Hydroxproline Asparagine Cysteine c

2.52 3.94 4.50 0.22 3.16 1.66 4.5

___ 0.08 +_ 0.13 +_ 0.12 +_ 0.02 ___ 0.05 ___ 0.07

acid)

Vmax(Na+)fVmax(aa) 2.41 3.74 4.66 0.25 3.13 1.84 3.4-6

aCoupling ratios in the middle column were determined as described in the footnote of Table 6.5 (adapted from Koser and Christensen, 1971, with permission from Elsevier Science). bMeans and standard deviations were calculated from numerous determinations in the original paper, and SEM was calculated here from the standard deviation reported in the original paper under the conservative assumption that only 16 determinations were made in each case. Cln the presence of 10 mM dithiothreitol.

ASC and Excitatory Amino Acid Transporters

replaces proline as the cosubstrate. This result is in contrast to the decrease in the velocity of Na + transport associated with the increase in its coupling ratio described above for the influence of intracellular asparagine and proline on the transport of other amino acids. TM A model for the Na + and amino acid receptor subsites of system ASC and similar systems is presented below (Section III,E,3). Moreover, different models to account for the observed cotransport of amino acids and Na + by several systems are presented in this and the preceding subsections. There is, however, no model that can account adequately for the observed stoichiometries in every known instance of Na+/amino acid cotransport. Hence, it is essential always to measure unknown stoichiometries of cotransport directly rather than to attempt to deduce them from a model relating transport of one cosubstrate to the concentration of another. Moreover, such measurements should be repeated for a variety of known substrates since the stoichiometry of their transport with a cosubstrate may vary as in the case of amino acid transport by system ASC.

4. The Stoichiometry of Cotransport Should Be Measured Rather Than Inferred from Cooperative Kinetic Effects Since the stoichiometry of cotransport cannot be inferred reliably from cooperative kinetic effects, it must be determined experimentally in each instance. Likewise, such models in which stoichiometry is deduced from cooperative kinetics should not be extended also to deduce the order of cosubstrate binding. Although we present a model to account for most known characteristics of EAAT-catalyzed transport in Section III,C,3 below, the model cannot be extended to other instances of co- and countertransport. Moreover, we have attempted to construct the model so that it does not imply unproven events. 19 Our success will be measured by the extent to which only new details need to be added to the model. 18 It also appears that in order to be substrates for transport by system ASC, Na + and amino acids must be separable. The positively charged side chains of cationic amino acids appear to occupy the receptor subsite for Na + (see Section III,E,3 below) but unlike Na+, the side chains are covalently linked to the rest of the amino acid molecules. Consequently, cationic amino acids are competitive inhibitors but not substrates for transport by system ASC (Thomas and Christensen, 1971) apparently owing to the inability of the positive charge on their side chains to be separated from the remainder of the molecule. 19 Technically, our model does propose the occurrence of unproven events since the transport of neither Na + nor K + by the E A A T proteins has been measured directly. Enough circumstantial evidence appears to exist, however, to warrant this conclusion, although the stoichiometry of transport remains to be determined (see Sections III,C and III,D of the text).

7.1 7

C. EAAT S y m p o r t e r s P r o p a g a t e a Na + Gradient into Gradients of Anionic A m i n o Acids As for the ASC proteins, the E A A T proteins catalyze obligatory exchange. Unlike the ASC proteins, however, E A A T proteins use the transmembrane Na + total chemical potential gradient to concentrate amino acids against their gradients. This phenomenon is possible even though exchange is obligatory because K + is by itself also a substrate. Although the total chemical potential gradient of K + is much smaller than that of Na + (Chapter 3), it lies more importantly in the reverse direction of Na+. Consequently, Na + ions usually enter the cell with an amino acid molecule, whereas K + ions are more inclined to follow their gradient out of the cell. For these reasons, both the Na + and the K + gradients drive the uptake of anionic amino acids against their total chemical potential gradients. Before we consider the kinetics of this transport in greater detail, let us review briefly the structure of E A A T proteins.

1. Structure of the E A A T Proteins As discussed above, the structures of the E A A T and ASC proteins are very similar. While the locations of the first six putative transmembrane segments seems obvious from hydropathy plots alone (Figs. 6.27 and 6.28), the locations of the more C-terminal membrane traverses are much less conspicuous in these plots. Consequently, the exact number of membrane traverses in the C-terminal region of E A A T proteins has been controversial (Fig. 6.35A vs. Fig. 6.35B), especially in regard to the long hydrophobic stretch (Fig. 6.28). In particular, a highly conserved region of this stretch (LCHS in Fig. 6.36) was proposed to cross the membrane in some models (membrane traverse number 8 in Fig. 6.35A), whereas it is placed in the cytosol in other models (Fig. 6.35B). This highly conserved sequence is interesting both because conserved regions frequently are essential to transport and because hydrophobic regions may greatly influence the free energy of binding of hydrophilic substances to the proteins. Changes in the relative free energy of their binding may help to transfer the free energy of one solute gradient into that of another. Moreover, binding of substrate by the rat EAAT2 transporter appears to alter its conformation (Grunewald and Kanner, 1995), and conformational changes seem undoubtedly to be at least part of the mechanism by which free energy conversions occur during transport (see also Chapters 3-5). Hence, the more recent finding (Wahl and Stoffel, 1996) that the LCHS in rat EAAT1 (Fig. 6.36) actually appears to traverse the membrane

2 |8

6. Transport Proteins That Propagate Solute Gradients

A

B

7

(,,.,

4

AAXFIAQ

COOH ~ FIGURE 6.35 Proposed membrane topology of E A A T proteins based primarily on their hydrophobicity plots (A and B) or on this criterion plus the results of reporter glycosylation scanning studies (C). The model of Kanai and Hediger (1992)(A) differs from that of Pines et al. (1992)(B) primarily in whether the segments labeled 7 and 8 traverse the membrane. Putative segment 7 may be too short to cross the membrane as an c~-helix, whereas putative segment 8 is in a highly conserved portion of the long hydrophobic stretch (Fig. 6.28) which is labeled LCHS in Fig. 6.36. Also shown in A and B are the locations of the aminoand carboxyl-termini, conserved SSSS and A A X F I A Q amino acid residue sequence motifs, conserved possible protein kinase A (PKA) and C (PKC) phosphorylation sites, and putative N-glycosylation sites (symbols attached to the second extracellular loop) (adapted from Kanai et aL, 1994 with permission from Federation of American Societies for Experimental Biology). An apparently more precise definition of the probable locations of the membrane traverses beyond the sixth one is shown in C for rat EAAT1. Reporter glycosylation sites were introduced into the protein or fragments of it at sites indicated by filled circles and arrows. Native N-glycosylation sites are marked with arrowheads. Subsequent detection of the locations of the glycosylation sites was used to construct the proposed topology (adapted from Wahle and Stoffel, 1996, with permission from The Rockefeller University Press and see Grunewald et aL, 1998 for another recent model for E A A T proteins).

twice (Fig. 6.35C) has considerable implications for the mechanism by which these proteins catalyze transport. Several possible regulatory phosphorylation sites are also present in the intracellular loops of all EAAT proteins (Fig. 6.35) (Gegelashvili and Schousboe, 1997). Only the conserved intracellular site in the loop between transmembrane segments 2 and 3 (Fig. 6.35) has, however, been shown in at least one case (i.e., in rat EAAT2) actually to be phosphorylated and consequently to help to regulate transport (Casado et al., 1993). Interestingly, a protein kinase C-dependent process also regulates anionic amino acid transport in cells expressing rat EAAT1. In this case, however, phosphorylation of EAAT1 occurs at a site other than the sites expected to be substrates for protein kinase C (Conradt and Stoffel, 1997). Apparently protein kinase C is needed directly or indirectly to activate phosphorylation at other sites (discussed further in Section III,A,3 of Chapter 9). In regard to the loops on the other side of the membrane, the extracellular loop between transmembrane segments 3 and 4 contains two conserved consensus se-

quences for N-linked glycosylation (Fig. 6.35). While gly+ cosylation at these sites is unnecessary for Na /glutamate symport by rat EAAT1, glycosylation is needed for formation of EAAT1 dimers in vitro (Conradt et al., 1995). If dimerization also occurs in vivo, it could lead to as yet unappreciated allosteric effects on transport. Interestingly, the rat and mouse EAAT3 proteins also have a third glycosylation site in this loop, whereas the human and rabbit EAAT3 proteins have, instead, an additional site in the first extracellular loop (Bjcr~s et al., 1996). Ongoing investigations are designed to learn how these and other structural similarities and differences among the transport proteins in the E A A T / A S C family may influence their function. Important aspects of this function include the kinetics of the transport that they catalyze. 2. Kinetics of Anionic Amino Acid Transport Careful analysis of the current induced by substrate at constant membrane electrical potential indicates that EAAT3 catalyzes the exchange of 3 Na § ions, 1 proton,

ASC and Excitatory Amino Acid Transporters

2 | 9

C

464 206

368 425

77

313

445

487,

280

342|

398~tL,,

~EL~406

q~ 495 501

273

FIGURE 6 . 3 5

(Continued)

and 1 anionic amino acid ( A A - ) molecule for 1 K + ion (Zerangue and Kavanaugh, 1996a). Since unidirectional transport of each of these solutes alone is electrogenic, the current produced by their transport together must be the sum of their individual currents. Moreover, some transport produces current in the reverse direction or is electrically silent, since K + and AA-/H+/Na + may be transported in either direction and since E A A T 3 should catalyze homo- as well as heteroexchange. 2~ When the net current resulting from the sum of these components of total transport is normalized and measured as a function of the concentrations of the various ions, the stoichiometry of transport can be deduced from the cooperativity exhibited (e.g., Fig. 6.37). In addition, K0.5 values can be calculated from the relationships between normalized current and substrate concentration (Fig. 6.37). Since K + and AA-/H+/Na + would compete for transport in the same direction, however, the observed K0.5 values 20 W e use designations, such as A A - / H + / N a + to indicate that these solutes are cotransported. Moreover, K + is by itself a substrate of the E A A T proteins. In no case do we m e a n to imply that K + may substitute for Na + or H + for cotransport with A A - .

543

514

are higher than those that would be observed for uninhibited transport. Unfortunately, the mutual cis-inhibition between AA-/H+/Na+ and K + for transport cannot be characterized by measuring the current that their transport produces. For example, the current owing to uptake of both Glu- and K + from the extracellular medium is the sum of their currents in the absence of the other (Fig. 6.38). Such would, however, be the case regardless of whether AA-/H+/Na + and K + compete for the same pathway for transport. In the case of competition, the combined current reflects both mutual inhibition of the current produced by the other substrate and the fact that the competing substrate produces current in the reverse direction. In contrast, the combined current would represent the sum of two larger individual currents in the case where K + and AA-/H+/Na + do not compete, but the sum of these currents would be the same as in the case where they do compete. Even if a scheme were used to detect a reduction in current owing to competition, however, it would not detect potentially large components of the

220

6. Transport Proteins That Propagate Solute Gradients

EAATw ~ E ,A, ,A1T 2

E A AT3

EAAT4

M , ,, , ,', ', M ': A ': S,:T,,:EMG,,A O ,, M MP': K,,Q,=V,:,E,:, VP ,, D,,:S. H,L ,G ~ . ,LOCVD. ,K. ~L ,G. .K. ,NVL H NN V ,,R M S.E~E. .PVKOH, ,R H L G L,R M G K P A R K G C PSWK R F L K NNWV M S S H G N SL F L R E S G Q R L G R V G W L Q R L Q E S L Q Q R A L R T R L RLQTMT L EHVLRF LRRNAF I .

[

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vI

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v

C

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LI~

R~G D S~

",9c j i v - Ao oI = N l i l K

lib

V ,.I~G

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299 355

G I F / I / ~ R ~ F S L F A I V I llWml l i l I I V _ R U B ] ~ FR A GIVllm~IlImI-=ILiHRiFP I

3 s s

EBD F ~

42o

SBN , . . ~ A m . V B D ~ V ~ L i V DI~G I 1 ~ i I~ S / I EBN L ~ T ~ A ~ V

329

419 38 9 44 S

r~c L'LrceI'F F

TR . R L S P Q E

S A A Q~E T T E Q S G K..

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E K L A ; PD EA INMTJ s A E V S L L . . . . . . . K INETIVT VP ST R E RVK P P $ PEM ESFTAVMTTAI SKINKTIKE Y L Y T VVTRTMVRTEINGSIEPGASMPPPFSVE.INGTISF E~RAI. P~V S~C Q ~ S L ~ VR E N D DiN DiN

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GNSVIEENEMKKPYQLIAQDNETEKPIDSETKM RVHED ! EMTKTQS I YDDMKNHRESNSNQCVYAAHNSV EVN I VNPFALEST I LDNEDSDTKKSYVNGGFAVDKSDT T L PS LGKPYKS LMAQEKGASRGRGGNESAM 564

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21 The concept of effective concentration was introduced in Section II,B,5 of this chapter.

' '

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GV DL NY

ABI C R K D F A R D T G T E E--RHBKNRDVEM VY I i/K DT I D S Q H VE K EQMDVSS I E~Q ELQEAEL 561

574

FIGURE 6.36 Amino acid residue sequence alignment among members of the human EAAT subfamily of Na+-dependent amino acid transport proteins. Also shown are eight possible transmembrane segments (I to VIII) as well as the large conserved hydrophobic sequence (LCHS). If the LCHS does not traverse the membrane then it would lie outside the cell in the model presented. This extracellular location is in contrast to the membrane or cytosolic locations proposed in other models (e.g., Fig. 6.35). Possible N-linked glycosylation sites are shown in boxes (NXS or NXT) in a large extracellular loop between transmembrane segments III and IV (adapted from Arriza et al., 1997 with permission from the National Academy of Sciences, USA).

unidirectional fluxes of substrates owing to electrically silent h o m o e x c h a n g e . The contributions of h o m o e x change to the total unidirectional fluxes must be known in order to characterize accurately competition between A A - / H + / N a § and K § for transport. In this regard, since a given actual extracellular K § concentration is predicted to have a higher effective extracellular concentration at - 6 0 m V than at + 2 0 mV, the Km (and Ki) value for K + may be lower than the 17 m M determined at + 2 0 m V (Fig. 6.37). 21 For these reasons and for reasons discussed in the preceding Section (III,B), we strongly encourage investigators to charac-

I S

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terize transport by measuring unidirectional fluxes of radiolabeled substrates as well as by measuring the currents that their net transport produces. Data obtained in earlier studies of transport now known to be attributable to rat E A A T 2 (e.g., Kanner and Bendahan, 1982; Pines and Kanner, 1990) can be used also to draw conclusions similar to those above for E A A T 3 expressed in oocytes. In these earlier studies transport of radiolabeled anionic amino acids was measured in isolated m e m b r a n e vesicles and in proteoliposomes containing the purified and reconstituted transport protein. From all of these studies, one can formulate a possible m o d e l for the transport cycle of E A A T proteins.

221

ASC and Excitatory Amino Acid Transporters

A

x

~0.8

• jz~

E

.,...

"o9 0.6

N

40

.E

-20 ~ -40 -60

0.4

!,.,.,.

0

-8o ,i

0

1 := 0.8

50

,

..........

,i

i

10 100 [L-Glu] H.M ....

i

i,

i

0.4

E o 0.2 t~

Z

0

0

9

.

-6o iii! ......

. . . . . . [~1 M , 10 20 30 40 50 60 70 [K § mM

~

.

10

.

IH,]

100 nM

.

.

D

0.8 4O

~ 0.6

0,,0

o

50 100 150 200 250 300 350 [H § nM •

20

4

1SO ! .................

0

.c_

/

.~-

1 ~ 150 200 250 300 [L-Glu] ~M

E

(D N

40

.N "~ 0.4 E o 0.2 z

C

x

"o 0.6

B

= 0.8

"o9 0.6

E o 0.2

Z

1

z~

2O

0.2

Y

0

0

-. . . . . . .

25

50

75

10 _ 1 0 0 [Na'] mM , 100 125 150

[ N a § mM

FIGURE 6 . 3 7 Effects of extracellular substrate concentrations on the current induced by exchange of Glu-/H+/Na + for K + in Xenopus oocytes expressing EAAT3. Except when they were intentionally varied, external substrate concentrations were 10/zM Glu-, 32 nM H +, 60 mM Na +, and 40 mM K +. Currents owing principally to Glu-/H+/Na + uptake and K + exodus (A, B, and D) were measured at a membrane electrical potential of - 6 0 mV, whereas reverse currents (C) were measured at +20 mV. Normalized net currents (I/Imax) were fit by the least-squares method to the Equation I/Imax = [ion]n/([ion]" + Km'). According to some models, the values of n may reflect the stoichiometry of transport, and values of n greater than 1 indicate positive cooperativity. The values of n are 1.0 for Glu-, H +, and K +, and 2.3 for Na +. Apparent K0.5 values for Glu-, H +, K +, and Na + are 27 /~M, 26 nM, 17 mM, and 46 mM, respectively. The membrane electrical potentials that produce current reversal (Erev) are shown in the insets for various substrate concentrations. The relative magnitude of the slopes of the lines in the insets and the apparent stoichiometries of transport of the substrates according to this thermodynamic criterion are 1.0 Glu-, 0.79 H +, 3.17 Na +, and 0.98 K +. Both this thermodynamic method and the one involving measurement of relative cooperativity are indirect measures of stoichiometry and may be unreliable indicators of it (see text) (adapted from Zerangue and Kavanaugh, 1996a, with permission from Macmillan Magazines Ltd.). .

3. Possible Model for EAAT-Catalyzed Transport The actual fluxes of each of the co- and countersubstrates of anionic amino acid transport by E A A T proteins have not as yet been measured except as they may contribute to the net electrical current associated with transport (Fig. 6.37). Hence, the stoichiometries of transport deduced from these and other experiments need to be verified by measuring them directly (see Section III,B above). It is tempting to speculate that since the E A A T and ASC proteins are in the same family, they may have the same stoichiometries of transport. This inclination may be even more appealing since the stoichiometry deduced for Na + and anionic amino acid cotransport via E A A T proteins (e.g., Fig. 6.37) is about the same as that actually measured for transport of Na + with some amino acids via system ASC (e.g.,

alanine and hydroxyproline in Table 6.6). However, the different relationships between amino acid transport and the Na + concentration that have been observed for the E A A T (Figs. 6.37D and 6.39) and ASC (Fig. 6.30) proteins, as well as the lack of correspondence between such relationships and actual stoichiometries (e.g., Table 6.3), dissuade us from making premature judgements about the stoichiometry of transport catalyzed by E A A T proteins. Furthermore, we think that previous conclusions concerning the order of binding of Na + and anionic amino acids to E A A T proteins are premature. While existing kinetic data for these proteins and the systems that appear to contain them can be interpreted to mean that Na + binds first (e.g., Fig. 6.40), we are now reluctant to accept conclusions drawn from such studies. As discussed above, the conclusions depend on unwarranted

222

6. Transport Proteins That Propagate Solute Gradients

6~ r

-Io.

)

FIGURE 6.38 The sum of the currents produced by the simultaneous uptake of K + and Glu- (also with H + and Na +) by EAAT3 would be the same regardless of whether the substrates compete for uptake (see text for further discussion). The currents resulting from the application of 10/zM Glu-, 40 mM K +, or both of these solutes together were measured at the indicated values of the membrane electrical potential in Xenopus oocytes expressing EAAT3 ([Na +] = 60 mM) (adapted from Zerangue and Kavanaugh, 1996a, with permission from MacMillan Magazines Ltd.).

assumptions about the meaning of cooperative effects (or the lack of such effects) of one cosubstrate on the kinetics of transport of the other. Zerangue and Kavanaugh (1996a) actually used two indirect methods to estimate the stoichiometries of coand countertransport via EAAT3 (Fig. 6.37). The cooperative effects among cosubstrates described above were used to support their conclusion that 3 Na § ions are transported with each of the other co- and counter-

100

................ ' ......... ............

E

.E_ 8oX E 0

1:: 60O

/

e-

I--

---~ wr E404D

40-

1= t~ r

<,

a, 20-

0

'

0

I

30

'

'

I

60

'

'

I

"

'

90

I

120

'

'

150

Na concentration (raM)

FIGURE 6.39 Sigmoidal relationship between the Na + concentration and D-aspartate uptake by HeLa cells expressing either wild-type (WT) rat EAAT2 (open squares) or the E404D mutant (filled circles). Such results are obtained at D-aspartate concentrations of 200/zM or below (adapted from Kavanaugh et al., 1997, with permission from American Society for Biochemistry & Molecular Biology, Inc.).

substrates (Fig. 6.37). Their principal criterion was, however, a thermodynamic one (insets in Fig. 6.37) similar to the one mentioned briefly in Chapter 3 (footnote 7). While we think that the thermodynamic criterion may actually be a more reliable measure of stoichiometry than the one involving cooperativity, there are few if any data directly to support our view (see below). Hence, we still encourage investigators to measure and compare the actual unidirectional fluxes of all co- and countersubstrates rather than or in addition to deducing them using such thermodynamic criteria. This thermodynamic criterion is distinctly different from the one involving cooperativity, although the criteria share a theoretical foundation formed by transport kinetics and thermodynamics. 22 Moreover, both criteria are indirect measures of stoichiometry and should for this reason be verified before they are used for this purpose. Cooperativity has been found to reflect stoichiometry in only one of at least four cases of amino acid transport for which it has been studied (Table 6.3 and footnote 17 of this chapter). Furthermore, the stoichiometry of co-transport of amino acids with Na § varies widely among amino acid species at least in the case of system ASC (Table 6.6). Similarly, the stoichiometry of countertransport may vary greatly at least for some substrates of system b ~247(see below). Consequently, it may become necessary to verify the efficacy of indirect measures of stoichiometry not only for each transport system, but also for each alternative substrate of the system. In the case of the E A A T proteins, neither the cooperative nor the thermodynamic criterion has been shown in any instance accurately to reflect the actual stoichiometry of co- and countertransport. In fact, it is difficult to locate studies in the literature in which the efficacy of the thermodynamic criterion has been tested as a measure of the actual stoichiometry of transport. It is, of course, more tedious and costly to measure simultaneously the unidirectional fluxes of radiolabeled co- and countersubstrates across the biomembranes of cells than it is to measure the current reversal potentials at various substrate concentrations often utilizing the same single cell for numerous measurements (see also Chapter 11). Nevertheless, we can begin to gain some sense of how well the thermodynamic criterion may reflect stoichiometry by examining a study in which unidirectional flux, net current and current reversal potentials were measured for reasons other than to test this criterion. Electrogenic exchange of zwitterionic and cationic amino acids is catalyzed by a conspicuous transport ac22 We attempt further to combine the historically separate fields of transport kinetics and thermodynamics in Section III,B of Chapter 7. Moreover, the thermodynamic effect of membrane electrical potential on effective ion concentrations, which should be used for interpreting transport kinetics, was discussed in Section II,B,5 of the present Chapter.

223

ASC and Excitatory Amino Acid Transporters

A

[Na*]

B

[G~u-]

[] 20 mM

1.0 mM

10

o~

x

-;

940 mM 5

10

x

1.5 mM

>

60 mM

5

120 mM

II/

..~

2.5mM 10.0 mM

-0.3

0

03

0.6

0.9

1 / [Glu] (mM -1)

-3

-2

-1

0

1

2

I

3

(1 / [Na*] 2 (mM-2)) x 103

FIGURE 6.40 Kinetic data consistent with the interpretation that Na + binds before Lglutamate for symport by a system in rat intestinal brush border membrane vesicles that most likely contains an EAAT protein. The Na+-dependent component of total transport is shown where the units of the initial velocity (V) are pmol (mg protein) -1 min-1. According to classic enzyme kinetic theory (e.g.; Stein, 1986), when the double-reciprocal plots for Glu- uptake at various Glu- and Na § concentrations intersect exactly on the y-axis when 1/[Glu-] is plotted (A), but to the left of this axis when 1/[Na+]2 is plotted (B), then it may be concluded that the 2 Na§ ions bind before the glutamate molecule. Furthermore, the exponents of 1 for [Glu] and 2 for [Na§ reflect their stoichiometries of transport according to kinetic theory. These and related plots may, however, be unreliable indicators of the actual stoichiometry of cotransport (see text). For this reason, such studies should not be used either to deduce the stoichiometry of transport or the order of substrate binding until the proposed kinetic model has been shown to apply to the transport process under investigation (adapted from Prezioso and Scalera, 1996, with permission from Elsevier Science).

tivity associated with B A T expression in Xenopus oocytes. The r e a d e r may recall that this transport activity resembles system b ~ (see Sections X,J and XI,E of C h a p t e r 4 and see also Section III,E below). Moreover, L-alanine and its analog, 2-amino isobutyrate (AIB), c o m p e t e for transport by this activity with about equal Km (and Ki) values (Coady et aL, 1996). These authors also found that alanine and A I B induce similar currents and amounts of L-arginine countertransport at various m e m b r a n e electrical potentials. Most importantly, in regard to the t h e r m o d y n a m i c criterion, the current reversal (i.e., equilibrium) m e m b r a n e electrical potential was the same for alanine and AIB. 23 According to these findings, the stoichiometry of exchange of arginine for either A I B or alanine appears to be about 1:1. In contrast, unidirectional alanine and A I B transport indicated that their actual stoichiometries of exchange with arginine differ by about 30-fold (Coady et aL, 1996). Hence, the t h e r m o d y n a m i c criterion appears also to be an unreliable and incompletely u n d e r s t o o d m e a s u r e of stoichiometry, at least in this instance where adequate data appear to be available to assess it. In fact, most authors 23These data are complicated somewhat because the intracellular substrates are a mixture of amino acids. Nevertheless it seems clear that the stoichiometries of exchange that would be calculated from the substrate distribution ratios at the reversal potentials (i.e., apparently at equilibrium) would be the same for alanine and AIB.

have m e a s u r e d the stoichiometry of amino acid exchange via system b ~247 directly (e.g., A h m e d et al., 1995; Chillar6n et al., 1996), whereas indirect methods appear to be used more frequently to estimate stoichiometry when one or more of the cosubstrates is an inorganic ion. In the case of E A A T proteins, one such inorganic ion, K § probably should be viewed as a substrate in its own right, rather than as serving simply to reorient E A A T proteins to receive AA-/H+/Na § on the outside of cells. True, its physiologically most conspicuous transport may be exodus, but K + undoubtedly influences the function of the proteins from outside as well as inside the cell (e.g., see Section IV,C below). Moreover, it undoubtedly competes with the cosubstrates for transport as if it were A A - / H + / N a § although this competition has not been characterized (Section III,C,2 above). The reader is also reminded that since the proposed A A - - d e p e n d e n t fluxes of Na § and K + have not actually been measured, it is conceivable (although we think unlikely) that neither of these cations actually is transported. For these reasons, we present the conservative model for E A A T - c a t a l y z e d transport shown in Fig. 6.41 as one that still needs to be verified and then to be amplified through further investigation. In the model, it is proposed that A A - and Na § are cotransported in exchange for the same substrates or K + on the other side of the m e m b r a n e . Similarly, we

224

6. Transport Proteins That Propagate Solute Gradients

kI ~ k2 ? n A ~

T~~

AAhT ~rnNa

ke~kT?

tuNa+ AAhT

+

xK~_:Na::T

k~

nAA"

TRANSLOCATION

"- NatAnT

xK

J

TRANSLOCATION

k2

Na~/!~ T .., nAA"

.

-- Nal~1A/~ T nAA"

Na~lIT

Na~IT

OUTER

INNER

FIGURE 6.41 Model for transport by E A A T proteins. These transporters (T) appear to catalyze obligatory exchange, but because of the total chemical potential gradients of Na + and K + across the plasma membrane, the system usually functions for net exodus of K + and

net uptake of the cosubstrates, Na + and anionic amino acids (AA-). Only four of the many possible rate constants are shown in order to indicate that it is not yet known whether the binding or unbinding of the cosubstrates, Na + and AA-, is ordered or random. Similarly, four of the complexes, Na+m AAn- T, are shown in the figure for convenience of illustration and are not meant to imply that the complexes formed through different orders of substrate binding are different. It is also likely that protons are cosubstrates with Na + and AA-, but their transport is not shown because the kinetics of their transport and its relationship to the transport of Na + and AA- have not been studied as extensively as the other co- and countersubstrates. The Na + and K + have also not been shown actually to migrate across the membrane, although much circumstantial evidence indicates that they are transported. Determination of the stoichiometry of their transport depends, however, on direct measurement of their migration with AA-, so the stoichiometry of transport is not indicated in the model. The modified ping-pong model shown for obligatory exchange also accounts for the trans-stimulation and probably cis-inhibition that occur between and among K + and different anionic amino acids (AA-), the later always in association with Na +.

p r o p o s e that K + is t r a n s p o r t e d by E A A T p r o t e i n s by b o t h h o m o - or h e t e r o e x c h a n g e . C o n s e q u e n t l y , these substances exhibit cis inhibition and trans stimulation as e x p e c t e d for the modified ping-pong m o d e l shown. A two-site s i m u l t a n e o u s m o d e l also is consistent with m o s t available data for this e x c h a n g e and might be m o r e likely than a ping-pong m o d e l if E A A T p r o t e i n s f o r m dimers in vivo (recall Section II,B above c o n c e r n i n g the t r a n s p o r t catalyzed by A E 1 dimers). It should also be e m p h a s i z e d that, as far as we know, each step in this m o d e l is readily reversible (Fig. 6.41). C o n s e q u e n t l y , a l t h o u g h a m i n o acid e x c h a n g e r e q u i r e s N a +, it does not a p p e a r to m a t t e r on which side of the m e m b r a n e N a + is supplied ( K a v a n a u g h et al., 1997). A p p a r e n t l y anionic a m i n o acids m a y dissociate f r o m and t h e n reassociate with the t r a n s p o r t p r o t e i n while one or m o r e N a + ions r e m a i n b o u n d to it, a l t h o u g h it has not b e e n d e t e r m i n e d w h e t h e r N a + ions can u n d e r g o

a similar process while one or m o r e anionic a m i n o acid m o l e c u l e s r e m a i n bound. As discussed above, h o w e v e r , T h o m a s and C h r i s t e n s e n (1971) c o n c l u d e d that e i t h e r a N a § ion or a zwitterionic a m i n o acid m o l e c u l e could dissociate f r o m and t h e n r e a s s o c i a t e with system A S C for t r a n s p o r t without c o n c o m i t a n t dissociation of the c o s u b s t r a t e (Fig. 6.34). H e n c e , it is r e a s o n a b l e to suppose that the s a m e m a y be true for the A S C - r e l a t e d , E A A T proteins (Fig. 6.41). F u r t h e r studies are n e e d e d b o t h to show t h a t N a § and K § ions actually m i g r a t e across the m e m b r a n e via E A A T p r o t e i n s and to d e t e r m i n e the s t o i c h i o m e t r y of their co- and c o u n t e r t r a n s p o r t with anionic a m i n o acids. It also n e e d s to be d e t e r m i n e d w h e t h e r the a p p a r e n t l y multiple N a § ions bind with equal affinity and are transp o r t e d t o g e t h e r , since such a p p e a r s n o t to be the case for the r e l a t e d A S C t r a n s p o r t e r s (Section III,B above). Similarly, the t r a n s p o r t of a p r o t o n is not shown in

ASC and Excitatory Amino Acid Transporters

the model because it is not clear in what capacity or stoichiometry it may associate with the transporter and the cosubstrates for transport. D. Use of Site-Directed M u t a g e n e s i s to Gain Insight into the M e c h a n i s m of EAAT/ ASC-Catalyzed Transport Several charged amino acid residues in hydrophobic regions of the E A A T proteins are required for normal transport activity, whereas similar mutations in several other such residues have not been found to influence transport. For example, mutation of histidyl residue 326 of the rat EAAT2 transporter (Fig. 6.28) to a lysyl, arginyl, asparagyl, or threonyl residue abolishes amino acid transport activity (Zhang et aL, 1994). Since this histidyl residue in the sixth putative membrane traverse is conserved among E A A T and ASC transporters (Fig. 6.26B) it may be required by all of them for transport. In contrast, lysyl residue 298 in the fifth putative transmembrane segment of EAAT2 does not appear to be required for transport activity (Zhang et al., 1994). Even its conversion to the very different threonyl residue reduces amino acid transport only to about 40%,and more conservative substitutions do not appear to influence transport. Since this residue is also conserved among E A A T and ASC proteins, however, it probably is needed to maintain some more subtle function of the proteins than simply whether they can transport amino acids. Similarly, a conserved aspartyl residue at the beginning of the long hydrophobic stretch (Fig. 6.28) as well as one near the end of it (i.e., D398 and D470, respectively, in the rat EAAT2) may be required for transport by all members of this family. Mutation of these residues to asparagyl, glycyl, and even glutamyl residues in rat EAAT2 abolishes amino acid transport (Pines et al., 1995). Since these two aspartyl residues are also conserved in several related bacterial anionic amino acid (Tolner et al., 1992a,b, 1995a) and even dicarboxylate (Engelke et al., 1989; Jiang et al., 1989) transporters, they may be central to the functioning of all of these proteins. In contrast, two other anionic amino acid residues near the end of the long hydrophobic stretch (E461 and D462 in rat EAAT2) do not appear to be needed for transport (Pines et al., 1995). Since all five E A A T proteins have anionic amino acid residues at these two neighboring positions (Fig. 6.36), whereas the ASC proteins and bacterial anionic amino acid transporters have only one (Fig. 6.26B and Tolner et al., 1992a,b, 1995a), it might be interesting to learn the effects of a double mutation at these two positions in the E A A T proteins. It has also not been determined whether the single anionic amino acid residue at one or the other of these positions

225

in the ASC and bacterial anionic amino acid transport proteins is needed for transport activity. Interestingly, the two related dicarboxylate transport proteins do not contain either of these anionic amino acid residues ( Jiang et aL, 1989; Engelke et al., 1989). Hence, perhaps at least one of these anionic residues is needed for recognition of the positively charged c~-amino group of amino acid substrates. On the other hand, another residue may be needed for recognition of the second organic acid group of anionic amino acids and even that of the dicarboxylate transporters. The arginyl residue at position 479 of the rat EAAT1 protein (Fig. 6.28) is conserved in mammalian and bacterial anionic amino acid transport proteins (Fig. 6.26B and Tolner et aL, 1992a,b, 1995a) and even in bacterial dicarboxylate transporters (Engelke et al., 1989; Jiang et al., 1989). This residue is, however, not conserved in the ASC proteins (Fig. 6.26B). Mutation of this arginyl residue abolishes amino acid transport by the rat EAAT1 protein (Conradt and Stoffel, 1995). We propose that the side-chain of this residue may be needed for recognition of the organic acid group on the side-chains of anionic L-amino acid substrates (or the c~-carboxylate group on D-aspartate; Gazzola et aL, 1981). In contrast, the arginyl residues at positions 122 and 280 of the rat EAAT1 protein are not as well conserved as R479 among the E A A T (Fig. 6.36) and bacterial (Engelke et al, 1989; Jiang et al., 1989; Tolner et al., 1992a,b, 1995a) transport proteins. Moreover, R122 and R280 are not needed for amino acid transport activity, although their simultaneous mutation lowers the Km value for aspartate transport by rat EAAT1 (Conradt and Stoffel, 1995). Finally, the presence or absence of two amino acid residues has particularly important implications for the physiological functions of all proteins in this family. The tyrosyl and glutamyl residues at positions 403 and 404, respectively, of the rat EAAT2 protein (Fig. 6.28) are not only required for normal transport, but their conservative mutation alters the substrate selectivity of the protein (Pines et al., 1995; Kavanaugh et aL, 1997; Zhang et al., 1998). Perhaps most importantly in regard to physiological function, the ability of the Y403F and E404D mutants to transport K + appears to be lost entirely! The presence of phenylalanyl (F) and glutaminyl (Q) residues at these positions in the ASC proteins (Fig. 6.26B) may account for their inability to countertransport K § thus rendering them also unable to use a Na § gradient to concentrate amino acids (see below). It will also be interesting to learn whether any of the bacterial proteins discussed here are able to transport K § since they also carry a glutaminyl or another uncharged residue at the position corresponding to E404, whereas tyrosyl residues at positions corresponding to Y403 are

226

6. Transport Proteins That Propagate Solute Gradients

conserved among the bacterial proteins (Jiang et al., 1989; Engelke et al., 1989; Tolner et al., 1992a,b, 1995a).

1. The Y403F and E404D Mutant Forms of Rat EAAT2 Do Not Transport K + So They Catalyze Obligatory Nonelectrogenic Anionic Amino Acid Exchange The effects on transport of the Y403F mutation are very similar to those of the E404D mutation. The major difference between the effects of the two mutations on transport is that E404D influences selectivity for anionic amino acid substrates, whereas Y403F increases the affinity of the transporter for Na+, and it somewhat broadens the ability of the transporter to receive monovalent cations in place of Na + (Pines et al., 1995; Kavanaugh et al., 1997; Zhang et al., 1998). For these reasons, we describe only the effects of the E404D mutation in detail. Both the wild-type and the E404D mutant rat EAAT2 protein transport radiolabeled D-aspartate

A EAAT2 WT i

No EAAT protein

EAAT2 E404D j

i

well, but transport by the mutant is not electrogenic (Fig. 6.42). Moreover, the normal ability of substrate to activate an anion channel in E A A T proteins is lost in the rat EAAT2 E404D mutant when K + but not DAsp-/H+/Na § is introduced into the medium of oocytes expressing the protein (Fig. 6.43). Similarly, L-Asp-/H+/ Na + but not K + stimulates exchange uptake of L-Asp-/ H+/Na+ by the E404D mutant rat EAAT2 in proteoliposomes, whereas both K + and L-Asp-/H+/Na + stimulate exchange uptake by the wild-type protein (Fig. 6.44). Finally, both AA-/H+/Na + and K + stimulate exodus of D-Asp-/H+/Na + from proteoliposomes via wild-type rat EAAT2, but K + does not stimulate exodus via the E404D mutant protein (Fig. 6.45). For these reasons, it may be concluded that the glutamyl residue at position 404 of the rat EAAT2 and probably the homologous residues of other E A A T proteins are needed for them to transport K + (Kavanaugh et al., 1997). In contrast to loss of K + transport, the E404D mutation appears to decrease the Km value for transport of both D- and Laspartate by EAAT2. D-[3H]Aspartate uptake is inhibited by the unlabeled form of this amino acid more strongly in Hela cells

i

i

40nA

' ~ [3H]D-Aspartate

0oo ~e)

A

100 l.tM D-Asp I

I ~

,

50 mM K+

I

I

I

50 s ~

f

wt

<____ B

100 l.tM D-Asp I

50 mM K+

I

I'-'--]

I00 nAl

E404D

FIGURE 6.4Z Uptake of [3H]aspartate by Xenopus oocytes expressing the E404D mutant form of rat EAAT (B) is not electrogenic (A). Uptake by the mutant protein appears to occur in exchange for identical ionic species on the inside of the oocyte (see text). In contrast, transport by the wild-type protein produces current apparently owing to the symport of 2 or 3 Na + ions and 1 proton with each D-aspartate molecule in exchange for 1 K + ion. The membrane electrical potential and the D-aspartate concentration were -30 mV and 100 txM, respectively. (A) Horizontal bars represent times of D-aspartate application to oocytes expressing wild type (WT) rat EAAT2, the E404D mutant EAAT2, or no EAAT protein. (B) Vertical bars represent quantities of D-[3H]aspartate taken up by the same oocytes represented in A. Similar results were obtained with 12 other oocytes (adapted from Kavanaugh et al., 1997, with permission from American Society for Biochemsitry & Molecular Biology).

C

100 l.tM D-Asp

20 s

50 mM K+ I

I

No EAAT protein FIGURE 6.43 Potassium ion activation of the anion channel activity of the wild-type (wt) but not the E404D mutant form of rat EAAT2. Xenopus oocytes expressing wild-type (A), the E404D mutant (B), or neither of these EAAT2 proteins (C) were exposed to 100 tzM Daspartate or 50 mM K + at a membrane electrical potential of zero. Under the conditions of these experiments, uptake of NO3- (104 mM) produced most of the observed current (adapted from Kavanaugh et al., 1997, with permission from American Society for Biochemistry & Molecular Biology, Inc.).

227

ASC and Excitatory Amino Acid Transporters 200-

A

B

lOO A

e-

B

A

2

80

= 50-

40

-r

o

O.

E 30 m O

E

A

60 w

E

Q. v

~100-

~ a

,--., I

[]

40

50-

20

m Q.

/

El

a "1" '

I

1

'

I

2

'

I

3

Time . ...___

__.---o

...O""

0 0

'

I

5

'

I

10

-

0

'

A

I

5

'

v

I

10

time (rain)

0

'

I 1

'

I

2

'

I

3

(min)

FIGURE 6.45 Potassium ions trans-stimulate D-[3H]aspartate exodus via the rat EAAT2 protein (A) but not its E404D mutant form (B). Proteoliposomes were prepared from HeLa cells expressing the wild-type (A) or the E404D mutant form (B) of EAAT2. The liposomes contained D-[3H]aspartate and about 150 mM Na+. Exodus of the radiolabel was measured into 150 mM KCI (filled circles) or 150 mM NaC1 (open symbols) also containing 100/zM D-aspartate (squares), 100 /~M L-glutamate (triangles), or no addition (circles) (adapted from Kavanaugh et al., 1997, with permission American Society for Biochemistry & Molecular Biology, Inc.).

FIGURE 6.44 The E404D mutation of rat EAAT2 suppresses transstimulation of D-aspartate uptake by intravesicular K +. Proteoliposomes were prepared from HeLa cells expressing either the wild-type or the E404D mutant form of EAAT2. (A) Net D-aspartate uptake by proteoliposomes containing 120 mM K + is considerably slower when the glutamyl residue at position 404 of EAAT2 (triangles) is replaced with an aspartyl residue (circles). (B) Ten millimolar Laspartate on the inside of the proteoliposomes (open symbols) stimulated D-aspartate uptake by both the wild type (triangles) and the E404D mutant form (circles) of EAAT2 relative to uptake in the absence of intravesicular L-aspartate (filled symbols) (adapted from Kavanaugh et aL, 1997, with permission American Society for Biochemistry & Molecular Biology, Inc.).

2. Do Phenylalanyl and Glutaminyi Residues at Positions Corresponding to Y403 and E404 of Rat E A A T 2 Render All ASC Proteins Unable to Catalyze K § Transport?

expressing the E404D mutant rat EAAT2 than in cells expressing the wild-type protein (Fig. 6.46A). Similarly, inhibition of transport by dihydrokainate (Fig. 6.46B) and L-aspartate (Pines et al., 1995) is stronger for the E404D mutant than for wild-type EAAT2. In contrast, the strength of L-glutamate inhibition of D-aspartate transport is not influenced by the E404D mutation in rat EAAT2 (Fig 6.46C). Moreover, the E404D mutation reduces L-glutamate transport more than it reduces aspartate transport (Pines et al., 1995), probably as a result of nearly identical reductions in the Vmaxvalues of their transport. Hence, the E404D mutation appears both to increase the strength of the interaction of EAAT2 with L- and D-aspartate and to abolish the interaction of EAAT2 with K + (Kavanaugh et al., 1997). The absence of tyrosyl and glutamyl residues at positions in the ASC proteins that correspond to positions 403 and 404 in EAAT2 may explain why at least two of the ASC transporters appear not to receive K + as a substrate. As a consequence, these ASC proteins catalyze nonelectrogenic amino acid exchange but not net uptake.

Three hundred/xM L-alanine induces some current in X e n o p u s oocytes expressing ASC1 (Fig. 6.47A), but the current is much lower than the 29 nA expected if one net positive charge moves into the oocyte with each alanine molecule (Zerangue and Kavanaugh, 1996b). Moreover, the relatively small current that is induced appears to result entirely from the anion channel activity of this family of transport proteins (Fig. 6.47). Similarly, ASC2 catalyzes nonelectrogenic L-alanine transport (Utsunomiya-Tate et al., 1996). These results are consistent with the interpretation that the ASC transporters catalyze Na+-dependent exchange of zwitterionic amino acids for each other but not for K +. Since exchange of K + ions for an equal number of other cations and a zwitterionic amino acid molecule would also be electrically silent, however, Zerangue and Kavanaugh (1966b) performed additional studies to show that K + is not transported by ASC1. First, they found that L-alanine exodus from X e n o p u s oocytes expressing ASC1 is not stimulated by K + (data not shown), whereas exodus of anionic amino acids is greatly stimulated by K + in oocytes expressing an EAAT protein (e.g., Fig. 6.45A). Furthermore, Lalanine uptake is not influenced by raising the extracel-

228

6. Transport Proteins That Propagate Solute Gradients ~"

1201

looL A

2~

+ _-

..... 0

~)

. . . . . .

'WT . . . . . . . E~34D

,. . . . . . . . . . . . . .

20 40 60 80 CONCENTRATION OF D-ASPARTATE (I~M)

1 2 0 1. .0.0. .~. . . . . . . . . ,

~-'-. - w.T. E404D

iii

6o-i -\\

.,

i~40 200

. 0

,

, ,,

,

,,

100 200 CONCENTRATION OF DHK (I.IM)

1201' c O..

.,

.

...........

~'~ '-.

100

W.T. ' E404D

3()0

' ''

m~ 80

I-

~ 1 a

9'

40 20 0

'

. . . . .

I

''

v . . . . . .

I:'

"'

"

100 200 300 CONCENTRATION OF L, GLUTAMATE (p.M)

FIGURE 6.46 D-Aspartate (A) and dihydrokainate (DHK) (B) each inhibit D-[3H]aspartate uptake via i tile E404D mutant form of rat EAAT2 (filled triangles) more strongly than they each inhibit uptake via the wild-type (W.T.) protein (open triangles). In contrast, Lglutamate inhibits transport by both proteins about equally well (C). Proteoliposomes for transport studies were prepared from HeLa cells expressing the EAAT2 protein or its E404D mutant form (adapted from Pines et al., 1995, with permission American Society for Biochemistry & Molecular Biology, Inc.).

lular K + concentration from 0 to 50 mM under voltage clamp conditions (Zerangue and Kavanaugh, 1996b), indicating that extracellular K + does not compete with alanine for uptake and that intracellular K + is not exchanged for extracellular alanine. Finally, Na+-depen dent L-alanine uptake appears to equal alanine exodus in oocytes expressing ASC1 (Fig. 6.48) thus indicating that ASC1 catalyzes exchange of zwitterionic amino acids for each other rather than for K +.

In contrast to these data for ASC1 and ASC2, available data for ASC3 indicate that it may catalyze highly electrogenic amino acid transport (Kekuda et al., 1996). The current associated with L-alanine transport by X e n opus oocytes expressing ASC3 (Kekuda et al., 1996) is much larger than the current associated with alanine transport by ASC1 (Fig. 6.47) and ASC2 (UtsunomiyaTate et al., 1996). The current in the latter two cases is attributed to alanine-induced C1- conductance. Since the C1- channel activity varies considerably among E A A T proteins (Arriza et al., 1997), it is conceivable that such could be the case also for ASC proteins. It is also possible, however, that the electrogenic transport catalyzed by ASC3 results from K + countertransport. As of this writing, the transport producing alanine-induced current via ASC3 has not been identified. Since the ability of some if not all proteins in the EAAT/ASC family to transport K + may depend on the presence of critical tyrosyl and glutamyl residues (Y403 and E404 in rat EAAT2), the evolution of concentrative (K+-dependent) and nonconcentrative (K+-indepen dent) amino acid transporters from each other could conceivably occur relatively easily. Alternatively, the evolution of concentrative or nonconcentrative transport may depend on the loss or gain of Na + or H + symport with the amino acid. In this regard, some of the bacterial anionic amino acid transport proteins discussed in this chapter are Na + dependent (Tolner et al., 1992a) and some are not (Tolner et al., 1992b, 1995a). In fact, whether the bacterial proteins are Na+-dependent appears to depend on the local chemical and physical environment of the membrane in which they are expressed (Tolner et al., 1995b). Na+-Independent uptake of anionic amino acids by bacteria is driven by the transmembrane proton gradient and its associated electrical potential (Tolner et al., 1995a). The bacterial proteins are, however, also able to catalyze amino acid homoexchange (Tolner etal., 1992b), which could not be driven by a cosubstrate total chemical potential gradient or by membrane electrical potential. For this reason, studies are needed to determine what solute normally is exchanged for anionic amino acids and their cosubstrates in these bacteria and to determine whether the exchange is obligatory. While the solute that is exchanged could conceivably be K +, the bacterial proteins lack the glutamyl residue (but not the tyrosyl residue) apparently needed for K + transport by mammalian proteins in this family. As for the EAAT/ASC/bacterial family, other families of amino acid transporters also exhibit a variety of possible interactions with Na + and K +. Presumably transporters in different families can evolve to transport or not to transport a particular solute in a variety of ways. Nevertheless, some of the same constraints on this evolution appear to apply even to unrelated families of transport proteins.

ASC and ExcitatoryAmino Acid Transporters

A

, ,

B

C ~,+30 E +20 "~ +10 ~ 0

, L-alanine

+20 0-_I -20 ---~ -40 -60

~

,, 5nAL__ 10 sec

40 current (nA) ~-

229

~ -10 a~ -20 ~" -30

I

10

I

I

30 [Cl-]o

I

I

I

I

I

II

100

[CI-]omM 100

potential ( m V ) t 2 ~ -60 6 . -. 30 2 0

30 0

FIGURE 6.47 The current induced by application of 300/xM L-alanine to Xenopus oocytes expressing ASC1 is virtually all attributable to C1- migration and hence to activation of C1- channel activity. The current (A) is smaller than the 29 or more nA anticipated for the migration of at least one net charge with each L-alanine molecule (Zerangue and Kavanaugh, 1996b). Moreover, the current (B) and the potential at which it is reversed (reversal potential; B and C) depend on the C1- concentration (adapted from Zerangue and Kavanaugh, 1996b, with permission American Society for Biochemistry & Molecular Biology, Inc.).

E. The N a + - D e p e n d e n t EAAT/ASC Proteins a n d t h e N a + - l n d e p e n d e n t S y s t e m s asc, b ~247 a n d y§ M a y Interact in Similar W a y s w i t h Na § K § a n d A m i n o A c i d s 24 Several apparently unrelated families of Na+-dependent and Na+-independent amino acid transport systems and transporters have been described (reviewed by Van Winkle, 1993 and by Malandro and Kilberg, 1996). 25 While these proteins may have evolved independently, some of them show strikingly similar interactions with Na +, K +, and amino acids. Three broad themes are discussed here. First we consider briefly the finding that some but not all b~ transport systems appear able to transport a K + ion in place of an amino acid molecule and, hence, need not catalyze obligatory 24Some investigators consider system y+L to be a variant of system y+ and so system y+L is not considered separately here. Nevertheless, the Na+-dependent interaction of zwitterionic amino acids with system y+L is several orders of magnitude stronger than their interaction with system y+ (Dev6s et aL, 1997). 25Since the transporters in systems asc and b~ have not been identified and sequenced, it is not yet known whether they are structurally homologous either to the transporters in system y+ (CAT subfamily) or to those in system ASC (ASC subfamily). It remains formally possible that members of the BAT subfamily (system b~ proteins) are transport proteins rather than accessory proteins, although evidence is mounting against such a possibility.

amino acid exchange. These findings are reminiscent of a major distinction between proteins in the A S C and E A A T subfamilies only the latter of which transport K +. Next, we note that stoichiometry of amino acid exchange via at least one b~ transport system may vary as widely as the stoichiometry of Na+/amino acid cotransport via system ASC. Finally, we consider the finding that several Na+-independent transport systems and transporters nevertheless interact with cations including Na + in a manner that resembles the interaction of Na + with the ASC transport proteins. In addition, all of these systems appear to receive cationic amino acids at a binding site composed of the subsites for binding of Na + and zwitterionic amino acids. 1. Some Forms of System b ~247May Exchange Zwitterionic Amino Acids for K § As discussed in Chapter 4, system b ~ appears to catalyze obligatory amino acid exchange. This obligatory exchange is proposed by Chillar6n et al. (1996) to drive uptake of cationic amino acids in renal and intestinal epithelia through their exchange with zwitterionic substrates (Fig. 4.45 in Chapter 4). Although exchange of like charged substrates for each other may also occur, the exchange that occurs under physiological conditions appears to involve primarily heteroexchange of a cat-

230

6. Transport Proteins That Propagate Solute Gradients "T

o

"5 250 X 3

r

Na*

300m

200 150

E o 100 ~6 50 . ~~

"

TABLE 6.7 Charge Flux Equals Unidirectional Flux of 3H-Labeled Amino Acids in X e n o p o s O o c y t e s Expressing Rat BATa

I

0

choline + I

influx efflux

I

influx efflux

FIGURE 6.48 Na+-Dependent uptake and exodus of zwitterionic amino acids appears to occur by obligatory exchange in Xenopus oocytes expressing ASC1. Influx of L-[3H]alanine (300 tzM) into oocytes was Na + dependent and equal to the efflux of L-[3H]alanine (or its unlabeled equivalent) from other oocytes upon adding 300/xM Lalanine to their bathing solution. An unlabeled equivalent of alanine is defined here as an amount of an otherwise undefined mixture of amino acids that behaves for transport by ASC1 as a known amount of alanine would behave. Since oocytes contain a mixture of amino acids, many of which can be transported by ASC1, the behaviors of solutions of oocyte homogenates were compared to solutions of known alanine concentrations to determine alanine equivalent concentrations. The abilities of solutions to elicit C1- current in voltage-clamped oocytes expressing ASC1 were used to determine the concentrations of alanine equivalents in oocyte homogenates to be 1.2 ___ 0.1 nmol oocyte -~. Interpretation of the results of the present study are complicated because the interactions of the various zwitterionic amino acids with an ASC system or transporter may be dissimilar (e.g., Table 6.6). The conclusion drawn from the results in the figure, that the ASC1 transporter catalyzes obligatory exchange, is, however, corroborated by other studies (see text) (adapted from Zerangue and Kavanaugh, 1996b, with permission American Society for Biochemistry & Molecular Biology, Inc.).

ionic amino acid molecule for a zwitterionic one. Only under less physiological conditions, such as abnormally high concentrations of a cationic or zwitterionic amino acid alone on one side of the membrane, does exchange of like-charged amino acids comprise a significant part of total transport by system b ~247(Ahmed et al., 1995). Hence, when L-phenylanine or L-arginine uptake is measured in X e n o p u s oocytes expressing the rat (Tate, et al., 1992) b~ transport protein BAT, a 1:1 ratio is observed between the number of molecules and the number of charges that cross the membrane (Table 6.7). At an extracellular amino acid concentration of 50 IxM, cationic amino acids appear to be taken up in exchange for zwitterionic amino acids exclusively, whereas the reverse exchange seems to be true for uptake of cationic substrates. Actually, zwitterionic amino acids appear to be taken up in exchange either for cationic amino acids or for K + in voltage-clamped oocytes expressing rat BAT (Ahmed et al., 1995). The inhibition of phenylalanine uptake by higher extracellular K + concentrations (Fig. 6.49) could conceivably result from

A m i n o acid (50 p M )

IAA b

Charge flux c

Tracer flux c

Flux ratio

L-Phenylanine L-Arginine

6.9 -9.4

4.3 5.9

4.2 6.2

1.02 0.95

aAdapted from Ahmed et aL, 1995, with permission from American Society for Biochemistry & Molecular Biology, Inc. bin nA. Cln pmol oocyte -1 min -1.

reduction of the total chemical potential gradient of K + (Ahmed et al., 1995) or, more likely, from competition between phenylalanine and extracellular K § for transport. Interestingly, no such K § dependence of the current induced by zwitterionic amino acids is observed for the rabbit form of BAT (Busch et aL, 1994). The rabbit form of BAT may also catalyze amino acid homoexchange (e.g., Fig. 6.50) more readily than does the rat form (Ahmed et aL, 1995). Perhaps different members of the BAT family, or the transporters that they activate, vary in their ability to receive K + as a substrate, as is the case for transporters in the E A A T / A S C family. This parallel between BAT-associated and ASC-catalyzed transport extends also to a wide variation in the observed stoichiometries of their counter- and cotransport, respectively.

15-

<

10

E v

-

5

~

0.5

,

,

,

I

1

I

,

i

2

I

5

=

t

i

i

I

I

10

20

[K+] mM FIGURE 6.49 An increase in the K § concentration reduces the current associated with addition of L-phenylalanine to the medium apparently owing to their competition for uptake by Xenopus oocytes expressing rat BAT. Voltage-clamped ( - 6 0 mV) oocytes were exposed to 100/zM phenylamine at the indicated K § concentrations (adapted from Ahmed et al., 1995, with permission American Society for Biochemistry & Molecular Biology, Inc.).

ASC and Excitatory Amino Acid Transporters

L-arginine superfusion

40t

Before After

Before After

20 O

"E

0

"=-E~ ~, ..I

0 -20

-40

-o _=

-(30 -80L-amino acid: Arginine

Leucine

L-leucine superfusion Before After <

"~

Before After

20-

O"

~9 o

,~ -40 ~ -60 _.c -80

L-amino acid:

. Arginine

Leucine

FIGURE 6.50 Na+-Independent amino acid antiport in Xenopus oocytes expressing rabbit BAT appears to change from largely but not exclusivelyheteroexchange to principally homoexchange as intracellular amino acids are replaced by the extracellular substrate. The currents elicitedby 50 tzM L-arginine (upper panel) or L-leucine (lower panel) were greatly reduced in voltage-clamped (-50 mV) oocytes when oocytes had been exposed to the same amino acid for 3 hr prior to measuring current. In contrast, currents were greater when 50 IxM leucine (upper panel) or arginine (lower panel) was applied to oocytes that had been exposed to the other amino acid for 3 hr. We interpret these results to mean that the oocytes become loaded with the amino acid to which they are exposed for 3 hr and depleted of other amino acids. Subsequent exposure to the same amino acid elicits primarily homoexchange and little net current, whereas exposure to the other amino acid elicits primarily heteroexchange and even larger currents than before the 3-hr loading procedure (adapted from Chillar6n et al., 1996, with permission American Society for Biochemistry & Molecular Biology, Inc.). In contrast to the results presented here for rabbit BAT, the antiport associated with rat BAT expression appears to occur exclusively by heteroexchange at least in oocytes that have not been exposed to an amino acid for 3 hr prior to measuring the current they elicit (see text and Table 6.7).

2. The Stoichiometry Varies Widely for Both ASC-Catalyzed Na+/Amino Acid System and BAT-Associated (b~ Amino Acid Antiport As for Na+/amino acid cotransport via system ASC (e.g., Table 6.6), the stoichiometry of cationic and zwitterionic amino acid exchange that is associated with expression of the rabbit B A T protein varies widely depending on the zwitterionic amino acid substrate. Coady

231

and assodates (1996) observed one-to-one exchange of alanine for cationic acids via rabbit B A T expressed in X e n o p u s oocytes. Similarly, a one-to-one stoichiometry has been observed for exchange of other zwitterionic amino acids in association with expression of both the rabbit (Chillardn et al., 1996) and rat ( A h m e d et al., 1995) B A T proteins. In contrast, the stoichiometry of exchange of the alanine analog, 2-amino isobutyrate (AIB) is about 1 AIB molecule for every 30 cationic amino acid molecules (Coady et al., 1996). The authors explain this result with a model in which the cationic amino acid is much more likely than AIB (but not more likely than alanine) to dissociate from the system during the transport cycle. This model is similar to that proposed for system ASC (Fig. 6.34) in which the cosubstrates may have the same (case 3) or different (cases 1 and 2) probabilities of dissociating from the transport system (Koser and Christensen, 1971). It is currently uncertain how variable the stoichiometry of transport may be among other pertinent transport systems and proteins to be discussed in this section, although their close structural and functional similarities to systems ASC and b ~ (see below) is consistent with the possibility that their stoichiometries of transport also may be quite variable. Specifically, systems ASC, b ~ asc, and y+ each have a substrate receptor site composed of adjacent subsites for Na + and zwitterionic amino acids. Moreover, it is the variable association of different zwitterionic amino acids with their subsite that appears to produce variable stoichiometries of Na+/amino acid symport or cationic/zwitterionic amino acid antiport via systems ASC and b ~ respectively. These adjacent subsites for zwitterionic amino acids and Na + also function together as a single site to receive cationic amino acids in all four of these systems.

3. Na+-Independent Systems b ~ asc, and y§ Appear to Bind Na § and Other Cations at a Subsite That Resembles the Na + Receptor Subsite of EAAT/ASC Transport Proteins Almost 30 years ago, Thomas and Christensen (1970) noticed an interesting similarity in the substrate receptor sites of systems ASC and y+. They observed that cationic amino acids compete with zwitterionic amino acids plus Na + for the substrate receptor sites of these transport systems. In spite of this similarity, system y+ evolved mainly to transport cationic amino acids, whereas system ASC receives primarily zwitterionic amino acids under physiological conditions. Moreover, cationic amino acid transporter (CAT) proteins in various forms of system y+ fall into a family (2.3 in Table 8.2 of Chapter 8) apparently unrelated to the transport proteins in the A S C / E A A T family (2.27 in Table 8.2). In the case of

232

6. Transport Proteins That Propagate Solute Gradients

transporters in the E A A T subfamily, the anionic sidechain of their preferred substrates may nevertheless occupy a position near a Na § receptor subsite that has a design similar to the subsite that receives Na + in the ASC and CAT transport proteins. More recently, the Na+-independent systems b ~ (Van Winkle et al., 1990c) and asc (Young et al., 1988) were found also to contain the Na § subsite. While systems b~ asc do not require Na + in order to transport zwitterionic amino acids, Na + and other cations competitively inhibit transport of cationic amino acids by these systems. The reader may recall that transporters in the EAAT/ ASC family appear, according to some models, to bind several Na + ions for transport with each amino acid molecule. It was the relatively weak binding of the final of these Na § ions that was proposed in Section III,B,2 to trigger transport. It may also be this final Na § ion that competes with cationic amino acids for binding to the ASC transport proteins. Alternatively, Koser and Christensen (1971) proposed that only 1 Na § ion at a time migrates across the membrane with an amino acid molecule via system ASC (Fig. 6.34). Transport of more than one radiolabeled *Na + ion with each radiolabeled amino acid molecule(*S) was proposed to occur owing to displacement of the *Na + ion but not *S from the transporter by the unlabeled form of the solute (Koser and Christensen, 1971). For this reason, the binding of only a single Na + ion is shown in their model for the system ASC substrate receptor site (Fig. 6.51). It is either this single Na § binding subsite or the subsite for the binding of the last of several Na § ions by ASC that we propose here to resemble the subsite for Na + binding in systems y+, asc, and b We think it is unlikely that systems y+, asc, and b ~ have multiple subsites for binding Na + and other cations since they transport cationic amino acids apparently in a Na+-independent manner. The monovalent side-chain of a cationic amino acid molecule is, for the binding proposed here, equivalent to a single Na § ion (Fig. 6.51). Nevertheless, further studies are needed to explore the potentially important possibility that both these Na +independent systems and the Na+-dependent EAAT/ ASC transporters have multiple subsites to receive Na+. While it is interesting that amino acid transport proteins in different families appear to have evolved with the same general potential for binding and transport of inorganic cations and amino acids, this phenomenon would hold even more meaning if it included the capacity to bind precisely the same number of cations and amino acid molecules in about the same physical orientation to one another. In this regard, the binding of the Na § site (or subsite) inhibitor harmaline to systems asc and b ~247 is particularly strong (Young et al., 1988; Van WinO

,

+

,

FIGURE 6.51 Proposed positions of subsites for the recognition of a Na + ion and an L-hydroxyproline molecule for transport by system ASC. Similar subsites are proposed for Na + and the same or other zwitterionic and anionic amino acids on the E A A T / A S C proteins, the CAT (system y+) proteins, and the transport proteins in systems b ~ and asc (see text). It is also proposed that the two subsites together may serve as a receptor site for cationic amino acids. The diagrams are not intended to imply that Na + enters first. Rather, they were drawn to illustrate clearly the position proposed to be taken by Na + (adapted from Thomas and Christensen, 1970, with permission from Academic Press, Inc.).

kle et al., 1990c). Moreover, harmaline inhibition of cationic amino acid uptake by system b ~ is only partially competitive (Van Winkle et al., 1990c) perhaps because harmaline also binds to Na+-binding subsites not shared for binding by cationic amino acids. Hence, one wonders whether harmaline might interact with additional tight Na § binding subsites on systems b ~247and asc, such as those postulated in one case above for system ASC and the ASC transporters (Section III,B,2). Regardless of the precise number of Na § receptor sites on various systems and transporters, systems y+, asc, and b ~ appear to have at least one Na + subsite and one subsite for zwitterionic amino acids that resemble subsites for these solutes on E A A T / A S C transporters. These two subsites also constitute a receptor site for cationic amino acids in all of these cases except, perhaps, for the E A A T transporters to which weak binding of cationic amino acids is postulated but yet to be demonstrated. Hence, it appears that amino acid transport proteins in at least two evolutionarily unrelated families may have evolved to catalyze transport under the same constraints and consequently with similar outcomes. A1-

Additional AE and EAAT/ASC Protein Functions

ternatively, the proteins may have some essential components in their primary structures that have been conserved over time, but current procedures for analyzing primary structure appear insufficient for detecting the pertinent similarities. It will also be interesting to learn whether systems b ~ asc, and y+ serve as amino acidactivated C1- channels as is the case for the EAAT/ ASC transport proteins. Such a similarity would make the current parallels among the different amino acid transport systems and proteins even more striking. The observation that anion exchange (AE) proteins also serve as C1- channels supports the theory that such a property is inherent in the design of many different families of antiporters. The possible physiological significance of the alternative functions of AE and EAAT/ ASC proteins as C1- channels is discussed in the next section. IV. BOTH AE AND EAAT/ASC PROTEINS HAVE ADDITIONAL FUNCTIONS Transport proteins in the EAAT/ASC and AE families may perform functions in addition to amino acid transport and anion exchange. For example, the ability of AE1 to serve as a C1- channel and a taurine channel for volume regulation in the red blood cells of fishes appears to be at least as important as its ability to exchange anions (Perlman et al., 1996). Similarly, activation of EAAT4 and EAAT5 as C1- channels by glutamate may be the main physiological function of these proteins rather than their ability to transport this amino acid (Arriza et al., 1997). Hence, the apparently alternative transport functions of some members of these two families may actually be the primary functions of the proteins. We discuss in Sections IV,B and IV,C below the additional transport functions of the AE and EAAT/ ASC proteins. First, however, we consider the very interesting structural and metabolic functions of the Nterminal domains of the AE proteins. A. Structural and Metabolic Functions of AE Proteins The important structural functions of the AE1 protein were discussed above in regard to AEl-deficient mice and cattle (Peters et al., 1996; Inaba et al., 1996). AE1 tetramers have been proposed to serve as tethered adhesive particles in a model of erythrocyte membrane two-dimensional elasticity (Feng and MacDonald, 1996). Although AE1 is very important to erythrocyte structure and function, other structural and transport proteins that associate directly or indirectly with AE1 appear to function well enough in red cells to allow

233

some AEl-deficient animals to survive into adulthood. Presumably, other AE proteins also serve structural and metabolic functions in nonerythroid tissues, although these other proteins have not been studied as thoroughly as AE1. AE1 resides in the erythrocyte membrane in a megadalton complex with other transport proteins, glycolytic enzymes, the cytoskeleton, and hemoglobin (reviewed by Solomon, 1992). This complex also appears to include tyrosine kinase (Habib-Mohamed and Steck, 1986) and carbonic anhydrase (Kifor et al., 1993). Sequences within the first 200 amino acid residues of the N-terminal domain of AE1 have been shown to be needed for AE1binding to other proteins (Wang et aL, 1995; Ding et aL, 1996). Because allosteric effectors of some of these proteins influence the conformations of others, it has been proposed that they are arranged in a network capable of transmitting information at relatively long distances in and just beneath the red cell membrane (Solomon, 1992). Such a mechanism of amplification would also explain how cardiac glycoside binding to the approximately 250 Na+K+ATPase molecules in the red cell membrane can produce detectable effects on the kinetics of binding of stilbene compounds to the 6 x 105 AE1 dimers that are also present there (Solomon, 1992). In this model, some AE1 (band 3) tetramers are associated with Na+K+ATPase but most are not (Fig. 6.52). Nevertheless, since components of the putative megadalton complex are proposed to undergo more or less continuous disassembly and assembly, each relatively stable AE1 dimer may associate transiently with one or more Na+K+ATPase molecules during its lifetime. The proposed associations could conceivably help to coordinate several important metabolic processes in vivo.

Juxtaposition of Na+K+ATPase and glycolytic enzymes should supply the segregated pool of ATP apparently utilized by this transport protein (Parker and Hoffman, 1967; Proverbio and Hoffman, 1977; Mercer and Dunham, 1981). Similarly, apposition of glucose transport proteins and glycolytic enzymes (Fig. 6.52) would presumably help to organize and facilitate the glycolytic process (Solomon, 1992). Inclusion of tyrosine kinase in the megadalton complex (Habib-Mohamed and Steck, 1986) may help to regulate glycolysis through phosphorylation of the tyrosine residue at position 8 of AE1 (Schneider and Post, 1995). Phosphorylation of AE1 apparently disrupts its electrostatic and hydrogen bonding to aldolase (Schneider and Post, 1995) and possibly other proteins in the complex. The association of both hemoglobin and carbonic anhydrase with the complex probably helps to regulate transport of 02 and CO2 in the blood. Carbonic anhydrase converts the CO2 produced in peripheral tissues

234

6. Transport Proteins That Propagate Solute Gradients

FIGURE 6.52 Diagram of possible interactions among various transport proteins, enzymes, and the cytoskeleton in and just beneath the red blood cell membrane. The AE1 (Band 3 or 3) protein is shown as a tetramer that may, or more often, may not be associated with proteins, such as Na+K+ATPase, that are much less abundant in the membrane than is AE1. Also shown are interactions of stilbene compounds (e.g., DBDS), ouabain, cytochalasin B (CB), and cytochalasin E (CE) with AE1, Na+K+ATPase, the glucose transporter, and actin, respectively. The interactions between each of the substances and their target proteins are influenced by one another, thus indicating that the proteins are connected via a cytoskeletal and membrane network. Other abbreviation: 4.1, protein 4.1 (adapted from Solomon, 1992, with permission from Elsevier Science).

into H C O 3 - in red cells, and the resultant H C O 3 - is transported out of red cells by AE1 (Fig. 6.2). Moreover, hemoglobin binding to AE1 influences its binding of stilbene compounds and presumably other characteristics of the transport that it catalyzes (reviewed by Salhany, 1990). Hence, it is reasonable to suppose that conformational changes resulting from hemoglobin/O2 association/dissociation could influence H C O 3 - transport, although such an effect is yet to be demonstrated experimentally. In addition, the AE1 protein apparently has other transport functions, particularly in the red cells of fishes where it may be involved in cellular volume regulation. Association of AE1 with other transport proteins also involved in volume regulation could help to coordinate their activities. Moreover, some intracellular osmolytes may be produced or removed partly through glycolytic and other pathways, so association of the pertinent enzymes with AE proteins may contribute to coordination of volume regulation in some species. We now review briefly these other transport-related functions of AE proteins.

B. Additional Transport Functions of the AE Proteins Cellular volume regulation is now known to be tightly coupled to regulation of metabolism in mammals (e.g., Lang and Waldegger, 1997). For example, proteins and polysaccharides are hydrolyzed when nonerythroid cells shrink, whereas net synthesis of these macromolecules is favored when cells swell. These metabolic changes may have helped to counteract cellular shrinkage and swelling, respectively, in ancient organisms faced with an osmotically variable environment. While these ancient mechanisms may still operate for this purpose in some modern species, they appear to have been adapted to serve for metabolic regulation in mammals. Nevertheless, an association between the membrane transport proteins involved in volume regulation and the enzymes involved in intermediary metabolism could still help to coordinate these cellular processes in all species. The involvement of nonerythroid AE proteins both in volume regulation and in cellular structure and metabolism remains conjectural (see below). On the other hand, the importance of AE1 to cellular structure

Additional AE and EAAT/ASC Protein Functions

and metabolism is well established as discussed in the preceding section (IV,A). Moreover, AE1 in the red blood cells of fishes appears itself to help regulate the volume of these cells (e.g., Motais et al., 1997), and recent evidence indicates that mammalian AE1 may also serve such a function (Conejero, 1997). The contribution of mammalian AE1 to volume regulation would, however, be much smaller than the contribution of AE1 in fishes (see below). For volume regulation, the AE1 protein functions as a channel rather than as an obligatory anion exchanger. It has been known for some time that the mammalian AE1 also catalyzes C1- transport without exchange (reviewed by Passow, 1992). This channel-like activity may, however, be too small (i.e., only about 0.01% of total C1- transport) to be of much physiological significance in mammalian erythrocytes. In contrast, electrogenic C1- transport by AE1 in fish red blood cells may become a greater proportion of the total (e.g., Perlman et al., 1996). Particularly under hypotonic stress, the C1- channel activity of AE1 may increase in order to produce a regulatory volume decrease (Motais et al., 1997). In this case, however, the main function of AE1 is not C1transport. Instead, this transporter appears to catalyze release of organic osmolytes, such as taurine, sorbitol and urea, from fish erythrocytes. AE1 has for several years been suspected to be the volume-sensitive organic osmolyte/anion channel (VSOAC) in fish erythrocytes (Perlman et al., 1996; Motais et aL, 1997; Fi6vet et aL, 1998). Definitive proof of that identity was, however, not published until 1995 when Fi6vet and associates used a trout cDNA library to isolate and clone sequences encoding two closely related AE1 proteins. They then expressed the corresponding cRNAs in X e n o p u s oocytes. As summarized in Fig. 6.53, the trout AE1 proteins catalyze taurine transport and electrogenic C1- conductance as well as C1- exchange. In contrast, mouse AE1 catalyzes C1- exchange but no detectable C1- conductance or taurine transport when expressed in X e n o p u s oocytes (Fig. 6.53). The good correlation between the amounts of taurine transport and C1- conductance in oocytes expressing trout AE1 (Fig. 6.54) is consistent with the conclusion that they are catalyzed by the same transport process. These authors also showed that the relatively large extracellular Z-loop between M5 and M6 (Fig. 6.3) is needed for trout AE1 channel activity. When the longer Z-loop in the trout AE1 was replaced with the shorter mouse one (Fig. 6.53) or simply shortened by 24 residues (Fi6vet et al., 1995), anion exchange activity was retained, but channel activity was lost. In the reverse case, however, the longer trout Z-loop did not render the mouse AE1 protein more capable of catalyzing C1- conductance or taurine transport. Hence, the large Z-loop

235

of trout AE1 is necessary but by itself not sufficient for channel activity (Motais et aL, 1997). Full acceptance of the theory that AE1 serves as the VSOAC in trout erythrocytes awaits only further proof that this protein may lose its sensitivity to activation by hypotonic solutions when it is expressed in X e n o p u s oocytes. While AE1 is most likely the VSOAC in trout erythrocytes, VSOAC activity is also detected in numerous tissues in a variety of vertebrates including mammals (Strange et al., 1996). As just discussed, the mammalian AE1 is not a good candidate for a VSOAC, and most mammalian tissues do not express AE1. These other tissues do, however, express one or more of the numerous isoforms and subisoforms of AE2 and AE3, and it is conceivable that some AE2 and AE3 proteins catalyze volume-sensitive organic osmolyte/anion transport. As pointed out by Fi6vet et al. (1995), many of these proteins have a large Z-loop (e.g., compare mouse AE2 and trout AE1 to mouse AE1 in Fig. 6.3), and a large Zloop is needed by the trout AE1 for its channel activity. Interestingly, the relatively large Z-loop of mouse AE2 could have been produced during evolution in part by expansion of a trinucleotide now encoding seryl residues 861 to 865 (Fig. 6.3 and see Section II,A above). Since this large Z-loop is necessary but not sufficient for AE channel activity, however (Motais et aL, 1997), the publication of studies designed to determine whether one or more of the various isoforms and subisoforms of AE2 and AE3 can serve as channels is anticipated with much interest. 26 As for AE1, most proteins in the EAAT/ASC family also catalyze channel-like C1- transport. Moreover, the relative abilities of E A A T proteins to catalyze C1- transport varies greatly among them (e.g., Arriza et al., 1997) as is the case for AE proteins. Consequently, a major function of some of these proteins may be as channels. In the cases of EAAT4 and EAAT5, Amara and associates (Fairman et al., 1995; Sonders and Amara, 1996; Arizza et aL, 1997) found that glutamate transport activity may be relatively insignificant in relation to the ability of this amino acid to trigger the C1- channel activity of EAAT4 and EAAT5 in target neurons. C. EAAT/ASC Proteins also Function as Channels Most members of the EAAT/ASC amino acid transporter family also catalyze amino acid-activated C1- conductance. While this C1- channel activity usually accounts for less current than Na+/amino acid symport by 26 Another transporter that may be a VSOAC in mammals is the volume-regulated C1- channel C1C-3 (Duan et al., 1997), although it apparently has not been determined whether this protein also transports taurine.

236

6. Transport Proteins That Propagate Solute Gradients Wild types

"I-Z'I'*

MmM*

m

fl

Expressed proteins

M

tAE 1AN

NH

Trout versus mouse chimaeras MZM

TmT m

i

IM ' ~

M'

'i NH2

NH 2

2

* MmM = mAE1 Functional properties CI-/CI- exchange

Deletion mutant

* TZT : tAE I

+

4.

4.

4.

4.

CI- conductance Taurine transport Channel activity is independent of cytoplasmic tail Data are taken from Fi~vet et a !. (1995)

Z-loop is necessary for channel activity

Z-loop is not sufficient for channel activity

FIGURE 6.53 Summary of the effect of the Z-loop between transmembrane segments 5 and 6 of the trout (T) and mouse (M) AE1 proteins on their C1- conductance and taurine transport when these proteins are expressed in Xenopus oocytes. Both of these channel activities (but not exchange transport) are lost from trout AE1 (TZT) when its Z-loop (Z) is replaced with the mouse AE1 (MmM) Z-loop (m) as shown for the TmT chimera. In contrast, replacement of the mouse AE1 Z-loop (m) with the trout AE1 Z-loop (Z) has no effect on its transport function (compare the MZM chimera with the MmM wild-type AE1). Hence, the longer Z-loop of the trout AE1 is necessary but not sufficient for C1- conductance and taurine transport. Also shown are normal transport activities for a mutant trout AE1 protein (tAEIAN) that has the N-terminal domain deleted (adapted from Motais et aL, 1997, with permission from the Company of Biologists Ltd.).

E A A T 1 , E A A T 2 , a n d E A A T 3 ( W a d i c h e et al., 1995a), it a p p e a r s t o a c c o u n t f o r m o r e t h a n 95% of t h e c u r r e n t resulting from transport by EAAT4 and EAAT5 under p h y s i o l o g i c a l c o n d i t i o n s ( F a i r m a n et al., 1995; A r r i z a

130- Gm, p.S 120 110100-

~ g -o 8 0 e-

r

90 8070 605o -

_

4o 30-

J

20100

r,,,, 9

0

I 5

I

I 20 Jin Taurine, pmol/h.oocyte 10

ll5

i'

25

FIGURE 6.54 Correlation between taurine transport (abscissa) and C1- conductance (ordinate) in Xenopus oocytes expressing wild-type trout AE1. In general, both types of channel transport activity increase as oocytes injected with 3.5 ng of trout AE1 cRNA are incubated for from 1 to 6 days (adapted from Fi6vet et al., 1995, with permission from Oxford University Press).

et aL, 1997). H e n c e , it is r e a s o n a b l e t o c o n c l u d e t h a t a c t i v a t i o n of C1- t r a n s p o r t b y a n i o n i c a m i n o acid signaling m o l e c u l e s is t h e p r i n c i p a l f u n c t i o n of E A A T 4 a n d E A A T 5 . If this t h e o r y is c o r r e c t , t h e n s o m e E A A T proteins may have central rather than auxiliary roles in signal t r a n s d u c t i o n in n e r v e a n d o t h e r t i s s u e s (see S o n d e r s a n d A m a r a , 1996 f o r f u r t h e r d i s c u s s i o n ) . E l u c i d a t i o n of t h e p r e c i s e r o l e s of t h e E A A T p r o t e i n s in t h e f u n c t i o n i n g of n e r v e t i s s u e will n e e d t o i n c l u d e t h e c o n t i n u o u s ---10% a c t i v a t i o n of t h e i r C1- c o n d u c t a n c e t h a t s h o u l d o c c u r in t h e p r e s e n c e of t h e 3.25 m M e x t r a c e l l u l a r K + in b r a i n t i s s u e (e.g., L o u v e l et aL, 1994). 27 It will also b e i n t e r e s t i n g t o l e a r n w h e t h e r t h e C 1 - c h a n n e l 27 This estimate of about 10% activation of C1- channel activity is based on a Km value for K + of 17 mM as determined at +20 mV in oocytes expressing EAAT3. Hence, we need to know the more pertinent K + Km values for EAAT4 and EAAT5 if we want to know how extracellular K + may influence their function in brain. The K + Km values also should be determined at the membrane electrical potentials of cells in brain under various physiological conditions since it is expected that the Km value for extracellular K + will decrease as the membrane electrical potential is made more negative on the inside. Such a change in the K + Km value with membrane electrical potential is anticipated because the effective extracellular K + concentration should increase for a given actual extracellular K + concentration as the transmembrane electrical potential grows more negative on the inside of the membrane. (See Section II,B,5 of this Chapter and Section III,B of Chapter 7 for further discussion of the concept of effective concentration.)

Summary

activities of EAAT/ASC proteins may sometimes serve for organic osmolyte transport as is the case for AE1. Conversely, one wonders whether the C1- channel activities of AE proteins may also sometimes serve in signal transduction. V. SUMMARY In contrast to some assessments, we have found in this chapter that the kinetics of antiport and symport may be at least as complex as the kinetics of primary active transport by P- and F-/V-type ATPases (Chapter 5). Neither a ping-pong nor a two-site simultaneous model accounts well for the antiport catalyzed by anion exchange (AE) proteins. Moreover, this antiport may exhibit apparently hyperbolic kinetics as well as positive and negative cooperativity depending upon the conditions under which transport is measured. The function of AE proteins is further complicated in vivo owing to their also serving in some cases as channels for regulation of cellular volume. These transport functions may be regulated through interdependent associations of the proteins with other membrane proteins, the cytoskeleton, and cytosolic enzymes involved in intermediary metabolism. Similarly, the transport function of proteins in the E A A T / A S C family may vary among its members. While proteins in the ASC subfamily catalyze amino acid exchange, those in the E A A T family use Na § and K § gradients to catalyze transport of anionic acids against their total chemical potential gradients. In spite of their ability to concentrate anionic amino acids, some members of the E A A T subfamily probably function more

237

for anionic amino-acid-stimulated C1- conductance than for uptake of amino acid signaling molecules. Hence, some of these transporters may help to transmit signals, whereas others may help to terminate them. In the latter case, the transporters could conceivably progress from transporting 2 Na § ions with every amino acid molecule at high amino acid concentrations to 3 Na § ions with each molecule after the amino acid concentration has been lowered. We saw in this Chapter that the stoichiometry of Na+/amino acid cotransport may vary widely with the amino acid species, so the stoichiometry could conceivably be regulated independently for a given species. Such regulation might permit some E A A T transport proteins to take up glutamate more rapidly at higher extracellular concentrations and against larger gradients when its extracellular concentrations are lower. To our knowledge, the variety of alternative transport and metabolic functions, which have been observed for AE and EAAT/ASC proteins, have not been reported for other types of transport proteins such as the P- and F-/V-type ATPases (Chapter 5). As we have seen, however, alternative functions of transport proteins have been observed frequently enough for us to wonder whether multiple functions may actually be the rule for them rather than the exception. The service of mitochondrial aspartate amino transferase to transport fatty acids across the plasma membrane is perhaps the most surprising example of these multiple functions of transport proteins (discussed in Section VII of Chapter 4). Hence, although our discussion of K § channel proteins in the following Chapter will not extend beyond this well-defined transport activity, additional functions for them may remain to be discovered.

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Channel Proteins Usually Dissipate Solute Gradients

1. INTRODUCTION

functions, some channel-forming toxins, such as ahemolysin, may actually form pathways through biomembranes for the unregulated passage of small molecules (see summary in Table 8.3 in Chapter 8). These channels are, however, meant to be compatible with cellular death rather than life. They cause host cell lysis by catalyzing nonselective transport of water, inorganic ions, and low-molecular-weight organic solutes. The existence of such channels and the relatively large size of the pathways that they form in biomembranes provide evidence that when substrate selectivity or transport regulation are not required, intimate contact between transport proteins and their substrates or effectors may not be needed. Conversely, the substrate selectivity and regulation of transport discussed in this chapter require that the channel proteins make close contact with their substrates and effectors in order to sense their nature. While all of these channels may be classified together for convenience (Chapter 8), we suggest here that the principal characteristic that they have in common is speed. Consistent with the rate of the transport that they catalyze, the Q10 value for transport via channels is often considerably lower than for transport by other proteins (see also Section II,A of Chapter 4). Relatively low free energies of activation are, however, exactly what one might expect for proteins that catalyze more rapid processes. As a first approximation, biocatalysts function by lowering the free energy of activation to a value that can be achieved at temperatures compatible with life. The reader is reminded that even the fastest channel catalyzes transport of inorganic ions across biomembranes at a rate of only about 8% of that which could be achieved by free diffusion in the absence of the mem-

In contrast to transport proteins that form (Chapter 5) or propagate (Chapter 6) solute gradients by coupling transport to conspicuous sources of free energy, channel proteins appear to form pores through which ions pass by diffusion. The unusual characteristics of the proposed diffusion have been attributed to the narrowness of the channels and the effects on diffusion that such physical constraints produce. As the reader is by now aware, however, we view the transport catalyzed by channels as involving intimate contact between the transport protein and its substrate, as is the case for proteins that produce and propagate solute gradients. Moreover, it is likely to us that during their transport cycles, all of these proteins undergo at least small changes in their conformations that resemble such changes in enzymes and other transport proteins (e.g., Marban and Tomaselli, 1997). 1 This view also is supported by computer simulation of water migration across membranes via voltage-gated Na § channels. In this simulation, water and presumably Na § migration can occur when the channel protein is allowed to have the motion that it would have at 300~ but not when its motion is virtually frozen (Jakobsson, 1997). While the magnitudes of such protein motions that are needed for transport may vary widely among the proteins, we propose that virtually all transport proteins need to move to some extent in order to catalyze solute or solvent migration across biomembranes (see also Section III below). In contrast to the channel-catalyzed transport produced within an organism for its own physiological 1We define a conformational change as any change in the threedimensional structure of a protein biocatalyst that is needed for it to perform its transport or other activity or for it to be regulated.

239

240

7. Channel Proteins Usually Dissipate Solute Gradients

brane barrier (calculated by Stein, 1986). Consequently, it seems to us at least as useful to understand how these proteins slow solute migration relative to free diffusion as it is to understand how they speed migration across the barriers formed by phospholipid bilayers. As for the other types of transport proteins discussed in Chapters 5 and 6, such understanding should come in part from a detailed description of the structures of channel proteins and the ways in which they interact with their substrates.

ii. STRUCTURE, FUNCTION, AND EVOLUTION OF CHANNEL PROTEINS The scientific literature on substrate-selective transport channels is even more vast than such literature on symporters, antiporters, and primary active transporters. Moreover, complementary portions of this literature on channels are discussed in numerous excellent reviews and monographs (e.g., Hille, 1992; Pongs, 1992; Jan and Jan, 1992; Brown et al., 1993; Lang, 1993; Keynes, 1994; Dolly et al., 1994; Catterall, 1995; Christie, 1995; Chandy and Gutman, 1995; Herbert and Reeves, 1995; Guy and Durell, 1995, 1996; Verkman et al., 1996; Strange et al., 1996; Zagotta, 1996; Robertson, 1997; Marban and Tomaselli, 1997; Lehmann-Horn and Riidel, 1997; Benos et al., 1997; Nichols and Lopatin, 1997; Isomoto et aL, 1997; Scannevin and Trimmer, 1997; Yellen, 1997, Tester, 1997; Williams, 1997a,b; Maathuis et al., 1997; Roden and George, 1997; Okada, 1997; Barby and Hoffman, 1997; Jan and Jan, 1997; Durell et al., 1998; and Herbert, 1998 to name only a small proportion of the total). In fact, thorough discussion of the literature on the ionic channels of excitable tissues alone required the full contents of a book even before accumulation of much additional information over the past 15 years (e.g., Hille, 1984). For these reasons, we select for consideration here only a few types of channels in order to illustrate some of the characteristics of the transport catalyzed by this category of transport proteins. In particular, we limit our discussion to two of the three large families of voltage-sensitive cation channels (Fig. 7.1); those termed voltage-gated K + (Kv) channels and those more commonly known as inwardly rectifying K + (Kir) channels. We selected K + channels because of their wellstudied functions and the recent description of one of their structures at 3.2 A resolution (Doyle et al., 1998). These two families of K + channels perform numerous important physiological functions such as regulation of cellular volume, release of hormones and neurotransmitters, and myocyte and neuron excitability. We focus here, however, on our main purpose, which is to compare and contrast transport catalyzed by channels with that catalyzed by transport ATPases, symporters, and antiporters (Chapters 5 and 6). The reader is again re-

FIGURE 7.1 Putative evolutionary relationships among families in the voltage-gated channel superfamily. Each of the families shown may contain multiple subfamilies as discussed in the text for potassium voltage-gated (Kv) and inwardly rectifying (Kir) channels. Also shown are the putative secondary structures of the three major groups of channel proteins. The Kv and Kir channels have structures similar to voltage-gated sodium (Na +) and calcium (Ca 2+) channels because the K + channel proteins form tetramers in biomembranes. Other designations include: CNG, cyclic nucleotide-gated channels; Kvca2+, calciumdependent voltage-gated potassium channels; KATP, ATP-dependent Kir channels (e.g., Kir 6); and P2x, purinergic receptor (adapted from Christie, 1995, with permission from Blackwell Science Pty Ltd.).

ferred to the numerous reviews and monographs already available for more detailed discussions of the physiological functions of Kv, Kir, and other K + channels (e.g., Hille, 1992; Pongs, 1992; Jan and Jan, 1992; Brown et al., 1993; Dolly et al., 1994; Ruppersberg et al., 1993; Busch and Maylic, 1993; Keynes, 1994; Catterall, 1995; Christie, 1995; Chandy and Gutman, 1995; Herbert and Reeves, 1995; Robertson, 1997; Nichols and Lopatin, 1997; Isomoto et al., 1997; Yellen, 1997; Tester, 1997; Williams, 1997a,b; Jan and Jan, 1997; Durell et aL, 1998). A. K+ Channels in Eukaryotes and Prokaryotes Potassium channels are distributed widely among species in various kingdoms. For instance, a total of at least six subfamilies of Kv channels are known, and at least four of the subfamilies are expressed in all triploblastic metazoans (reviewed by Jan and Jan, 1997). The six subfamilies of a-subunits in mammals correspond to six Drosphilia channel genes termed Shaker, Shab, Shaw, Shal, ether-a-go-go (eag), and slowpoke (slo). Even diploblastic jellyfish have two genes in the Shaker subfamily, and plants and bacteria also express genes related to these subfamilies. For example, a plant Kir channel has a structure that resembles Kv channels in animals (Anderson et aL, 1992;

Structure, Function, and Evolution of Channel Proteins

Schactman et aL, 1992; Sentenac et aL, 1992; MtillerR6ber et aL, 1995). Since transport by members of both the Kv and Kir families in animals is sensitive to changes in membrane electrical potential, albeit usually in opposite ways, and because the families are distantly related (Fig. 7.1), the Kir-like functioning of the structurally Kv-like plant K + channel may reflect adaptation on a common theme. In support of this theory, Nichols and Lopatin (1997) point out that Kv channels may nevertheless show some mild deactivation as voltage is lowered (i.e., made less positive on the outside) as well as when it is raised. Furthermore, Kir channels show some tendency to close during hyperpolarization as well as during depolarization of the membrane (Nichols and Lopatin, 1997). 2 Hence, these two large families may have evolved mainly to open or close as a result of opposite changes in the membrane potential because these two types of transport activities are needed for the functioning of excitable animal cells. Nevertheless, each family retains vestiges of the functioning of the other. In fact, the transport characteristics of K + channel-related proteins of prokaryotes show that the transport functions of these proteins have great evolutionary plasticity. The reader may recall one such instance from Chapter 5 where we described a P-type ATPase in E. coli that is composed of several subunits. The KdpB subunit of the Kdp protein complex catalyzes ATP hydrolysis, but it has no inherent capacity to catalyze transport (e.g., Buurman et aL, 1995). Association of the KdpB subunit with the hydrophobic KdpA subunit (and a third subunit) leads, however, to transport of K + against its total chemical potential gradient. KdpA contains two motifs with limited sequence similarity to segments of Kir and Kv channels that are critical to their functioning (Jan and Jan, 1994). Because of the coupling of ATP hydrolysis to transport by the Kdp complex, it may be difficult for some readers to view the KdpA subunit as a channel rather than a carrier. Such difficulties disappear, however, if one adopts the position that all of these proteins catalyze transport by variations on the same fundamental mechanism (see Section I above and Section I of Chapter 4). B. Voltage-Gated K+ (Kv) Channels in Animals

Mammalian voltage-gated potassium channels are encoded by at least 18 genes in six subfamilies that correspond to the six Drosphilia K + channel genes listed 2 We USe the terms hyperpolarization and depolarization here to mean an increase or a decrease, respectively, in the magnitude of the inside negative membrane electrical potential. Consequently, even a highly polarized inside positive membrane is said here instead to be greatly depolarized relative to the physiologically normal inside negative resting membrane electrical potential.

241

above. The evolutionary relationships among some of the known members of the four more closely related of these subfamilies (Christie, 1995) are shown in Fig. 7.2. Current designations for the known mammalian isoforms (and their subfamilies) include Kvl.1 to Kvl.8 (Shaker-related, Kvl subfamily), Kv2.1 and Kv 2.2 (Shab-related, Kv2 subfamily), Kv3.1 to Kv3.4 (Shawrelated, Kv3 subfamily), Kv4.1 to Kv4.3 (Shal-related, Kv4 subfamily), H E R G (eag-related subfamily), and big K § (BK) or maxi K (slo-related subfamily) (reviewed by Christie, 1995 and Jan and Jan, 1997). The H E R G and maxi K or big K (BK) Kv channels (not shown in Fig. 7.2) differ from the other four subfamilies in that they also seem to interact with Ca 2§ as is the case for the corresponding Drosphila proteins. Nevertheless, only some of the properties of the mammalian Kv channels are equivalent to those of their counterparts in Drosophila (reviewed by Christie, 1995). All known Kv genes are expressed in brain, although not necessarily in the same region, and some of the genes are expressed in other tissues (reviewed by Chandy and Gutman, 1995). Interestingly, some of the isoforms also show specific patterns of subcellular localization in neurons. The molecular mechanisms regulating tissue and subcellular distribution of the Kv isoforms remain active areas of investigation (Chandy and Gutman, 1995) Since voltage-gated K § channels within the same subfamily form hetero- as well as homooligomers (Scannevin and Trimmer, 1997), the total variety of possible channels is much greater than the 18 or so known txsubunit monomers that are encoded by separate genes. Diversity of channel function also may be produced through RNA editing, at least in Kv2 in squid (Patton et al., 1997). In those subfamilies apparently containing only one gene (e.g., the slo-related BK channel), variety may nevertheless be produced through alternative splicing of the gene transcripts (e.g., Saito et al., 1997). Alternative splicing also occurs in the region of the gene transcript encoding the C-terminus of all known members of the Kv3 subfamily (Fig. 7.3). Interestingly, some of the splice variants of B K channels may contain clusters of up to 9 and 21 glycyl and seryl residues, respectively, near their N-termini (Saito et al., 1997), although no function has apparently been attributed to these clusters. Their apparent lack of function is consistent with our theory that such relatively innocuous clusters may form rapidly in exons of genes through trinucleotide expansion (see Section II,A,3 of Chapter 6). Trinucleotide expansion may, however, also have produced useful functions in some Kvl isoforms. Such expansions are particularly conspicuous in the Nterminal region of Kvl.4, as compared to other members of this subfamily where they appear to have produced conspicuous clusters of A,G,H, R, and E residues (Fig. 7.4). As a result, the N-terminal region of Kv 1.4 is

242

7. Channel Proteins Usually Dissipate Solute Gradients

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longer than other Kvl isoforms (Fig. 7.3). As pointed out by Chandy and Gutman (1995), Kv 1.4 is the only mammalian homolog of S h a k e r that shares its characteristic of rapid inactivation apparently owing to a longer N-terminal region (see below). S h a k e r itself (Fig. 7.4) as well as Shab (not shown) have relatively long Nterminal regions that may have been produced in part through expansion of trinucleotides encoding glutaminyl residues (reviewed by Chandy and Gutman 1995). These data further support the hypothesis presented in Chapter 6 in greater detail (Section II,A) that useful new functions may evolve in proteins in general and transport proteins in particular through expansions in

their genes of trinucleotides encoding clusters of amino acid residues. The functions of some voltage-gated K + channel c~subunits are also modulated through association with accessory/3-subunits. At least five /3-subunit proteins have been identified so far, and three of them are produced from the same gene through alternative splicing (England et al., 1995; McCormack et al., 1995). Interestingly, the/3-subunit of the BK channel renders the c~subunit sensitive (or more sensitive) to activation by Ca e§ (Knaus et al., 1994; Meera et al., 1996). In contrast, /3-subunits of the Kvl family enhance channel inactivation (Rettig et al., 1994; Yu et al., 1996). We provide

Structure, Function, and Evolution of Channel Proteins

243

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here an overview primarily of the structure and function of the c~-subunits of Kv channels and refer readers interested in the /3-subunit to other reviews (e.g., Dolly et al., 1994; Dolly and Parcej, 1996; Robertson, 1997). The c~-subunits of voltage-gated K + channels contain six putative transmembrane segments termed S1 to $6 (Fig. 7.5) (Jan and Jan, 1992; Pongs, 1992; Chandy and Gutman, 1995). Tetramers of the a-subunit appear to form a single central pathway across which K + ions migrate. The $5 and $6 membrane traverses of each subunit appear to interact to form the pathway, whereas the other membrane traverses form an outer shell in the putative three-dimensional structures of the channels (Fig. 7.5). Transport across the pathway is controlled by several components of the a-subunits. First, the N-terminus of each subunit forms an inactivating domain that blocks the inner entrance to the open channel (Fig. 7.6 and see below). The inactivating domain appears to interact with the intracellular loop between $4 and $5 (reviewed by Jan and Jan, 1997). Moreover, the extracellular loop between $5 and $6 extends no more than about one nanometer into the pathway (LO and Miller, 1995; MacKinnon, 1995; Miller, 1995; Ranganathan et aL, 1996), and it appears to form a selective ion coordination site (Fig. 7.7) or selectivity filter (Doyle et aL, 1998) (see

further discussion in Section III below). $6 also appears to contribute to ion selectivity, whereas transport is activated through the sensing of membrane depolarization primarily by $4 (Fig. 7.6). $4 in the Shaker channel appears to sense changes in membrane potential through positively charged cationic amino acid residues at every third position in the segment (Fig. 7.8), whereas steric interactions of noncharged residues in the four $4 segments of the tetramer appear to mediate cooperative interactions among the subunits (Smith-Maxwell et al., 1998). Membrane depolarization sufficient to activate the channel results in the net movement of about 10 positive charges of the channel protein tetramer from one side of the membrane to the other (Bezanilla and Stefani, 1994; Sigworth, 1994). In the closed state, no more than five residues in each $4 span the membrane (Larsson et al., 1996), which is particularly thin owing to the conformation of the channel protein tetramer in this state (Fig. 7.8). Only one of the cationic amino acid residues in each monomer lies within the membrane in the closed state (R365 in Shaker), and it appears to interact electrostatically with two anionic residues in $2 and $3 (Papazian et al., 1995). $4 moves outward during activation such that its cationic amino acid residue that is within the boundaries of the membrane in the closed state (Fig.

I I I 1150 I I I I I I I I I I mKvl I(MK1) M T V MSGENA I DEASTAPGHPQDGS YPRQA DHDDHE rKvl I(RBK1) ....A..... .____ ____._ ........ hKvl l(HUK(1)) -V ....A. ___._ ____._ xKvl l g r s h s l ) IA -M -.T.VL--.-.. HP -Q.. .. mKvlZ(MK2) AT-DPV ---AAL.-.-.. T DPErKvlZ(RCK5) AT-DPV .--AAL.---.. T -DPE. ...A A L hKvl2(HUK(IV)) AT-DPT -DPE_. dKvl2 AT--P---AAL.---.T DPE_. . bKvl2(BGK5) AT-DP-.-AAL T DPE._ xKvl20(5haZ) AT-DLT -GSVGFA-----DPEP mKvl J(MK3) VP DHL LEPEA-GGGG-D-Pa-GCGSGGGGGG CDR-EPLPP ALPAAGEQD rKvlJ(RGK5) VP-DHL L E P E A - G G G G - D P a - ~ c v s ~ GG CDR-EPLPP A L PAAGEQD hKvlJMPCN3)--VP-DHL LEPEV-DGGG- P ~ - G c GGGG CDR E P V P P S L PAAGEQD mKvl4 -E-AMV-A-SSGCNSHMPYGYAAQARARERERLAHSR-AA-AAVAAATAAVEGTGGSGGGPHHHHQTRGAYSSHDP-GSRGSRRRRRQRTEKKK LHHRQSSFPHCSDLMPSGSEEKILRELSEEEED.EEEEEEEEEGRFYYSEEDHGD rKvl4(RCK4) - E - A M V - A - S S G C N S H M P Y G Y A A Q A R A R A R E R E R L A H S R - A A - A A V A A A T A A V E G T G G S G G G P H H H H Q T R G A Y S S H D P - G S R G S R E E E A T R T E K K K K L H H R Q S S F P H C S D L M P S G S E E K I L R E L S E E E E D - E E E E E E E E E G R F Y Y S E E D H G D hKvl4(HPCN2)-E-AMV-A-SSGCNSHMPYGYAAQARARERERLAHSRRAA-RAVAAATAAVEGSGGSGGGSHHHHQSRGACTSHDP-SSRGSREEEATRSEKKKA HYRQSSFPHCSDLMPSGSEEKILRELSEEEED-EEEEEEEEEGRFYYSEDDHGD bKvl4(BAK4) - E - A M V - A - S S G C N S H M P Y G Y A A Q A R A R E R E R L A H S R ~ A A . A A V A A A T A A V E G G G G S G G S Q H H H H P S R G A C T S H D P - S G R G S R R R R R P H P E K K K V H H R Q S S F P H C S D L M P S G S E E K I L R D L S E E - D D E E - D D E E D E E E E G R F Y Y S E E D H G E rKvl J(Kv1) E I S LVPL GS A M T LRGGG EAG- -CVQTPRGECGCPPTSGLNNQSKETLLRGRTT LED-N QGGRPLPPMAQELPQPRRLSAEDEEGEGDPGLGTVEEDQAPQDAGS hKvl5(HPCNl) E I A L V P L - 0 0 AMTVRGGD EAR-GCGQATGGELQCPPTAGLSDGPKEPAPKGRGA Q-D-DSGVRPLPPLPDPGVRPLPPLPEELPRPRRPPPEDEEEEGDPGLGTVE-Q ALGTAS mKvl.B(MK6) RSEKSL TLAAPGEVRGPECEQQDAGEF rKvl.B(RCK2) RSEKSL TLAAPGEVRGPEGEQQDAGEF hKvl.B(HBK2) RSEKSL TLAAPGEVRGPEGEQQDAGDF APLK - E - A M A G I EGNGGPAGYRDSYHSS Q R P L L R S S N L P N SR S FPKLSEE-NANENGMGVPGS DYDCS Shaker - A AVAGLYGLGEDRQHRKKQQQQ QQHQKEQLEQKEEQKKIAERKLQLREQQLQR NSLDGYGS LPKLSSQ-EEGGAGHGFGGGPQHFEPIPHDHD

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-

..

I I r T Y - K I I I I I I C C E R V V I N I S G L R F E T Q L K T L A Q F P N L L G N P K K R M R Y F D P L R N E Y F F D R N R P S F D A I L Y Y Y Q S G GI R L R R P V N V P LI D M F S E E I K F YI E L G E E A M E K FI R E D E G F I K E mKvl.l(MK1) .......................... rKvl I(RBK1) .......................... . . . hKvl.l(HUK(I)) .. ......v............ s - - - T - - - - I...A .............. xKvl .10cShrl) mKvl.zMK2) ..................... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .........I rKvl.2(RCKS) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .........I hKvl.2 (HUK(IV)) . . . . .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ......... 1 .... dKvl.2 . . . . . .1. .E. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I.... bKvl.Z(BGK5) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 xKvl.2(XShaZ) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .........I .... -.G ................ ..C...E.... 0 . R - ................................ ....... 1 - 1 .... mKvI.XMK3) -.G . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . R. rKvl.J(RGK5) hKM.3WPCN3) ..G ...................C...E....D..R.......V......................... YSSVRYSD... . . . . V . . . . . . . . M . . . . . . . E . . . . D . E . . T Q - . . . . - - . . . . . . . . . . . . . . . . . . . . . . . . . . mKvl4 G C S Y T D L L PQ E D GGGGG rKvl.4(RCK4) G C S Y T D L L P Q D D G G G G G YSSVRYSD... . . . . V . . . . . . . . M . . . . . . . E . . . . D . E . . T Q . . . . . . . . . . . . . . . . . . . . . . . . . . . . ... hKvl.4(HPCN2)ECSYTDL L P Q D E G G G G YSSVRYSD... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .... bKd,4(wK4) E C S Y T D L L A Q D D G G G G G G G S G G G G Y S S V R Y S D . . ......V........ M. E....D.E..TQ.. ............................. . .L....... ....G D.A . - L H - - - - - - . . . . . . . . . . . . . . . G . . . . . . . . . . . rKvlS(KV1) LHHQ ........................... D A..LP. .............................. hKvl.5(HPCN1) LHHQ mKvl.B(MK8) QEAEGGGG ..S S .. L . . . . . . . . . . . . . R . . s L . . D . . . . D . G R . V . F . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . rKvl.B(kK2) QEAEGGGG hKvl.B(HBK2) PEAGGGGG APLK C Shaker FC

....

. . .

.....

I

~

....

...............

...

.......

.........

......... ....

................................

...... ...........

FIGURE 7.4 Amino acid residue sequence alignment among members of the Shaker-related Kvl subfamily of voltage-gated K' channels. Clusters of A,G,H,R, and E residues occur in the N-terminal segment of Kv1.4 proteins prior to the first membrane traverse (Sl), and a cluster of Q residues is present in Shaker beginning at residue 20. Dashes represent residues identical to those in the sequence at the top of the alignment (mKvl.l), conserved residues are marked by an asterisk at the bottom of the alignment, and periods indicate that the alignments vary owing only to conservative substitutions. Brackets above the alignment mark putative transmembrane segments (S1 to S6), the H5 or pore-forming region (P), and potential tyrosine kinase (TY-K) and protein kinase C (PKC) recognition sites. Asterisks above the alignment between S1 and S2 mark potential N-glycosylation sites that are underlined in the sequences themselves. Other underlined sequences in the rat Kv1.4 and Shaker mark areas known to mediate rapid N-terminal inactivation of the channels (see text for further discussion) (adapted from Chandy and Gutman, 1995, with permission from CRC Press).

W

EE RPLPEK

...................................................................................... ......................................... - - - - -- - --........................... ....

I I I S 1 mKvl.l(MK1) E Y Q R Q V W L L F E Y P E S S G P A R V I A I V S V M V I L = I rKvl.l(RBK11 hKvl l(HUK(i)) xKvl l(XSha1) F mKvl Z(MK2) F rKvl Z(RCK5) F hKv( 2 (HUK(1V)) F dKvl2 -F.-bKvl2(BGK5) F xKvl Z(XSha2) F K mKvl XMK3) D F rKvl3(RGK5) D F hKvl3WPCN3) D F mKvl4 -FKKrKvl4(RCK4) FK K hKvl WPCN2) FK K bKvl4(BAK4) FKKrKvl S(KV1) F hKvl WPCN1) F mKv1 a(MK6) P F rKvl qRCK2) P F hKvl WBKZ) P F APLK -F--R Shaker -K- -K

--- -- -- --- .- ----- --- .- ---. --- --------~

~

~

FIGURE 7.4 (Continued)

LPELK

I

I

FIGURE 7.4 (Continued)

I

148

247

Structure, Function, and Evolution of Channel Proteins

FIGURE 7.5 Proposed structures of voltage-gated (Kv) and inwardly rectifying (Kir) K + channels. The Kv channels appear to form tetramers with a central pathway for K +transport and inner and outer shells formed by the indicated membrane traverses (S1 to $6). Both the inner shell and the H5 loop (or pore-forming region) are retained in Kir channels, but the outer shell is not. $4 in the outer shell helps Kv channels sense voltage changes, and H5 contributes to K + selectivity and transport in both types of K + channels (see text) (adapted from Kubo et al., 1993, with permission from Macmillan Magazines Ltd.).

7.8) m o v e s to t h e o u t s i d e of t h e m e m b r a n e a l o n g with o n e o r t w o o t h e r c a t i o n i c a m i n o acid r e s i d u e s ( L a r s s o n et al., 1996). Similarly, t h r e e o r f o u r c a t i o n i c a m i n o acid r e s i d u e s in e a c h m o n o m e r t h a t lie o u t s i d e t h e s u r f a c e of t h e m e m b r a n e o n t h e i n s i d e of t h e cell in t h e c l o s e d s t a t e m o v e to p o s i t i o n s b e t w e e n t h e t w o s u r f a c e s of t h e m e m b r a n e in t h e o p e n s t a t e (Fig. 7.8). I n t h e o p e n state,

i

I

+

I +--+ m .

t h e s e r e s i d u e s a p p e a r to a s s o c i a t e with t h r e e a n i o n i c a m i n o acid r e s i d u e s in $2 a n d $3 t h r o u g h e l e c t r o s t a t i c i n t e r a c t i o n s ( P a p a z i a n et al., 1995). I n a d d i t i o n , t h e conf o r m a t i o n a l c h a n g e s a s s o c i a t e d w i t h a c t i v a t i o n fill t h e c r e v i c e t h a t p r o d u c e s t h e t h i n c o n f o r m a t i o n of t h e c h a n -

A

A

A

(]r

NJ

J~

+ ,--

9

B

~-'C

C FIGURE 7.6 Sketch showing functional components of the Kv channel a-subunit. Positively charged residues in $4 sense changes in voltage, and the N-terminal region (dashed line) causes N-type inactivation. This inactivation occurs in tetramers of the c~-subunit through interaction of an N-terminal segment with the intracellular loops between $4 and $5 as shown for two subunits on the right. The H5 loop and transmembrane segment $6 form much of the transport pathway, and amino acid residue substitutions in H5 result in loss of K + selectivity (adapted from Jan and Jan, 1997, with permission from Annual Reviews, Inc.).

FIGURE 7.7 The H5 loops (pore-forming regions) of the four asubunits of Kv (A) and Kir (B) potassium channels appear to form a K+-selective coordination site in the external portion of the transport pathway (C). Only three of the four subunits are shown in C (see Section III,A of the text for further details) (adapted from MacKinnon, 1995 with permission from Cell Press).

248

7. Channel Proteins Usually Dissipate Solute Gradients CLOSED

OPEN A359

OUTSIDE

.

.

.

.

R362

~

i.~i

:R371 :

INSIDE

FIGURE 7.8 Diagram of the migration of the Shaker (Kv) potassium channel transmembrane segment $4, in response to membrane depolarization. Only one of the four $4 segments in each tetramer is shown. In the closed state the channel appears to be only about half as thick as the hydrophobic core of the membrane phospholipid bilayer. Consequently, it forms a crevice in the membrane, which can be spanned by about five amino acid residues (363 to 367) with a structure intermediate in length to an a-helix (---8~,) and a/3 sheet (--~17,~). Only one of the seven cationic amino acid residues in $4 lies within the membrane in the closed state, and it appears to interact electrostatically with E293 in $2 and D316 in $3. Access to residues in the crevice from the external solution is restricted in the closed state. Upon membrane depolarization sufficient to open the channel, $4 migrates toward the outside so that about 12 residues now span the membrane (368 to 379) and the crevice is filled. In this conformation about four of the cationic amino acid residues lie within the membrane having entered it from the cytosolic side, and they appear to interact electrostatically with E293, D316, and a third anionic amino acid residue (E283) in $2. Since about two cationic amino acid residues leave the immediate vicinity of the membrane as four others enter it, a total of two to four of the positive charges in $4 appear to cross the membrane when the channel opens, yielding a total of about 10 positive charges for the tetramer. Repolarization reverses the process and closes the channel if it has not been inactivated (see text) (adapted from Larsson et aL, 1996 with permission from Cell Press).

nel in the closed state (Fig. 7.8). Nevertheless, 80% of transmembrane voltage difference appearsoto be imposed across a distance of only about --~12 A owing to the barrier formed by the selectivity filter of an open K + channel (Doyle et al., 1998). Consequently, a total of about 10 positive charges on the voltage-gated K + channel protein tetramer (2 or 3 charges per monomer) appear to move across the electrical field of the membrane during activation. Similarly, these charges move back to the other side of the membrane electric field when the channel is closed but susceptible to reactivation. Restoring the channel to its closed state is, however, sometimes more complex than simply reversing activation. Upon prolonged membrane depolarization, voltagegated K + channels may first be inactivated by a combination of two interdependent processes before the

channels return to a voltage-sensitive state. In N-type inactivation, each of the N-terminal domains (Fig. 7.6) of the four a-subunits (Fig. 7.5) independently may occlude the cytoplasmic end of the channel transport pathway (Hoshi et aL, 1990; Zagotta et al., 1990; MacKinnon et al., 1993). Since the N-terminal segment of S h a k e r also appears to occlude the BK calcium-activated potassium channel (Toro et al., 1994), all voltage-gated K + channels appear to utilize the same mechanism for N-type inactivation. Nevertheless, the rates of this inactivation may differ even among channels in the same subfamily (Chandy and Gutman, 1995). In contrast to N-type inactivation, C-type inactivation appears to involve the outer end of the channel transport pathway (Choi et aL, 1991; Hoshi et al., 1991; LopezBarneo et al., 1993). Differences in the sequence of the C-terminal region of the S h a k e r a-subunit produce very different kinetics of C-type inactivation (Hoshi et al., 1991). Normal C-type inactivation hinges critically on a threonyl residue near the outer end of the channel transport pathway at position 449 of the S h a k e r c~subunit (Hoshi et al., 1991). Conversion of this residue to a glutamyl or a lysyl one through site-directed mutagenesis leads to inactivation that is two orders of magnitude faster than in the wild-type. In contrast, replacement of T449 with a valyl residue greatly slows inactivation (Lopez-Barneo et aL, 1993). Although Ctype inactivation can occur in the absence of N-type inactivation, it is facilitated by the N-terminal type (Baukrowitz and Yellen, 1995). N-type inactivation appears to facilitate C-type inactivation by favoring dissociation of K + ions from their binding site at the outside of the transport pathway (Baukrowitz and Yellen, 1995 and 1996). By blocking migration of K + ions out of the cell, N-type inactivation appears to prevent these K + ions from occupying the K + binding site on the outside of the pathway. When K + does not occupy its outer binding site, the channel protein apparently can undergo the putative conformational change needed for C-type inactivation (Jan and Jan, 1997). It is currently unclear precisely how the channel returns to a state in which it is again susceptible to voltage gating following C-type inactivation. There appear however, to be components of the Ctype inactivation process in addition to those that are facilitated by N-type inactivation. For example, intracellular tetraethylammonium (TEA) blocks but does not inactivate the transport of K + out of cells. Consequently, intracellular TEA and the N-terminal domain appear to facilitate C-type inactivation by the same mechanism (Choi et al., 1991). Conversely, extracellular TEA slows C-type inactivation apparently because it occupies a site at the outside of the transport pathway and, thus, inhibits C-type inactivation like K + ions do (Choi et al., 1991; Jan and Jan, 1997). In addition, however, TEA on the

249

Structure, Function, and Evolution of Channel Proteins

inside of cells slows C-type inactivation even though one expects it only to increase the rate of this inactivation by blocking exodus of K + ions. This slowing of C-type inactivation by intracellular TEA is attributed to an as yet poorly defined allosteric effect of the drug (Baukrowitz and Yellen, 1996). Interestingly, the latter allosteric slowing of C-type inactivation is about equal in magnitude albeit opposite to the increase in the rate of C-type inactivation owing to the blocking of K + transport by intracellular TEA. Moreover, the ability of intracellular TEA to increase the rate of C-type inactivation in the presence of intracellular K + is greater than for related but larger and more hydrophobic compounds. These derivatives of TEA have been proposed to become more effective at facilitating C-type inactivation as they become larger and more hydrophobic because they also become better able to occupy the pertinent site long enough to permit the last K + ion to leave the outer K + binding site (Baukrowitz and YeUen, 1996; Sanchez and Blatz, 1995). This period during which the site is occupied is termed the dwell time, and it should be much shorter for TEA than for related compounds of similar abilities to increase the rate of C-type inactivation. For these reasons, the blocking of K + transport out of the cell may not account fully for the ability of all of these compounds or perhaps even the N-terminal domain to speed C-type inactivation. Once open, voltage-gated channels transport K + very rapidly (i.e., about 107 K + ions/channel/sec; Marban and Tomaselli, 1997). As for most other transport proteins, however, the details of the pathway across which ions migrate and the conformational changes that may be associated with transport remain an active field of investigation. Since the putative conformational changes associated with transport would be very rapid relative to those producing channel opening, closing, and inactivation, their detection and description will undoubtedly require more sophisticated techniques than those used to study slower conformational changes. First, however, we need to recognize that the mechanism of transport via channels may share fundamental characteristics with other transport proteins. As for all transport proteins that interact intimately with their substrates, both voltage-gated and inwardly rectifying K + channels are highly selective for their substrate. We consider the molecular basis for this K + selectivity below after we discuss the Kir family of mammalian K + channels. C. Inwardly Rectifying K+ (Kir) Channels The other broad category of voltage-gated K +channels in animals is described as inwardly rectifying because the inward current owing to K + transport is greater for a given driving force than the reverse current

resulting from an equal but opposite driving force. In contrast to Kv channels, which catalyze K + exodus when the membrane is depolarized, Kir channels catalyze K + uptake upon membrane hyperpolarization. Nevertheless, each of these two types of K + channels exhibit some of the characteristics of the other, perhaps owing in part to their distant phylogenetic relationship (e.g., Fig. 7.1). As for Kv channels, the Kir channel family is composed of six recognized subfamilies (Fig. 7.9). Although members of the same subfamily are more closely related to each other (50-70% amino acid residue sequence identity) than to members of other subfamilies, the precise relationships among the subfamilies are unclear. Consequently, various authors show the relationships among the subfamilies differently (e.g., compare Nichols and Lopatin, 1997 to Isomoto et al., 1997). Except for the apparently more distantly related Kir5 subfamily and the closer-than-usual relationship between Kirl and Kir4 (Fig. 7.9), the amino acid residue sequences of the Kir channel proteins in different subfamilies are all about 30 to 40% homologous (Isomoto et al., 1997). For these reasons, we may need to learn of additional structural characteristics on which to base evolutionary relationships beyond those that are known, before the relationships among the Kir subfamilies will be clear. Alternatively, our current inability to discern an obvious pattern of the evolution of the subfamilies may mean that most of them evolved more or less simultaneously from a common ancestor. The first Kir channel protein to be cloned, and the only known member of the Kirl subfamily (i.e., Kirl.1), is a weak inward rectifier that is expressed in kidney as well as brain (Boim et al., 1995; Ho et al., 1993; Kenna

3.2 3.4

Kir3

3.3 3.1 6.1 6.2

Kirl Kit4 I Kir2 Kir5

t

1.1 4.1

2.2 2.3 2.1

5.1

FIGURE 7.9 Proposed evolutionary relationships among members of the inwardly rectifying (Kir) family of potassium channels. With the exceptions of the more distantly related Kir5 subfamily, and the more closely related (to each other) Kirl and Kir4 subfamilies, the precise relationships among subfamilies are unclear and may be shown differently by various authors (see text for discussion) (adapted from Nichols and Lopatin, 1997, with permission from Annual Reviews, Inc.).

250

7. Channel Proteins Usually Dissipate Solute Gradients

et aL, 1994). At least six subisoforms of the protein are known to be produced through alternative splicing near the 5' end of the gene transcript (Shuck et al., 1994; Yano et aL, 1994; Zhou et al., 1994). In contrast, diversification is produced among strong inward rectifiers in the Kir2 and Kir3 subfamilies through expression of at least three isoforms in each of these subfamilies (Fig. 7.9). Members of the Kir2 and the Kir3 subfamilies are expressed in brain and heart as well as other tissues (reviewed by Nichols and Lopatin, 1997 and by Isomoto et al., 1997). Further diversification of the function of members of the Kir3 subfamily is produced through formation of hetero- as well as homotetramers (Krapivinsky et aL, 1995; Duprat et al., 1995; Ferrer et al., 1995; Isomoto et al., 1996; Kofuji et al., 1995; Spauschus et aL, 1996). In addition, Kirl and Kir2 subfamily members are regulated through phosphorylation (Xu et al., 1996; Fakler et al., 1994; Henry et al., 1996), whereas members of the Kir 3 subfamily are G protein-activated (Dascal et al., 1993; Kubo et al., 1993; Lesage et al., 1994). Interestingly, expression of Kir 6 channel activities appears to require coexpression of a high-affinity sulfonylurea receptor (Ammala et aL, 1996), perhaps indicating that regulatory proteins help to control the activities of Kir channel proteins (McNicholas et aL, 1996; Nichols and Lopatin, 1997). In this regard, the single known member of the Kir5 subfamily (Kir5.1) does not appear to form active (or inactive) channels when it is expressed alone (Bond et al., 1994). It can, however, form a channel with novel characteristics when it is coexpressed with Kir4.1 (Pessia et al., 1996). Interestingly, this novel channel is expressed in tetramers (Fig. 7.5) only when every other subunit is Kir5.1, whereas the tetramer formed by two adjacent Kir5.1 and two adjacent Kir4.1 subunits resembles the Kir4.1 homotetramer (Pessia et aL, 1996). Kir4.1 and Kir5.1 were discovered in brain (Bond et al., 1994; Takumi et aL, 1995), whereas both ubiquitously expressed (Kir6.1) and pancreas-specific (Kir6.2) isoforms of the Kir6 subfamily have been described (Inagaki et al., 1995 a,b). The tetrameric structure of Kir channels resembles the structure of Kv channels (Kubo et al., 1993) but without the outer shell formed by S1 to $4 (Fig. 7.5). Since the structure of the Kir channel monomer is homologous to the C-terminal portion of the Kv monomer and consequently does not contain the voltage-sensing $4, it is logical that Kir channels do not exhibit voltagegating. Nichols and Lopatin (1997) point out, however, that Kir channels show some tendency to be closed in hyperpolarized membranes (Koumi et aL, 1994; Lopatin et aL, 1995; Nichols et aL, 1994) and, thus, to open upon depolarization from the hyperpolarized state. Moreover, although Kir channels do not contain the Nterminal inactivation segment of Kv channels (Fig. 7.6),

they are apparently inactivated in an analogous way by intracellular cations (see below). Similarly, some Kv channels show a tendency to be inactivated by intracellular cations (Forsythe et al., 1992; Lopatin and Nichols, 1994; Rettig et al., 1992) and consequently exhibit mild inward rectification (Nichols and Lopatin, 1997). 1. Inward Rectification of Kir Channels Results Primarily from Inhibition of K + Exodus by Polyamines Both intracellular Mg 2+ and polyamines block outward current via Kir channels when the membrane is depolarized (reviewed by Nichols and Lopatin, 1997). Among these substances, the polyamine spermine appears to be primarily responsible for the block in vivo (Fakler et aL, 1995). In excised patches from X e n o p u s oocytes expressing a Kir channel, the spermine chelator ATP partly reverses spermine inhibition of outward current (Fig. 7.10). Hyperpolarization of the membrane leads to large inward currents via Kir channels owing to K + transport that is presumed to be no longer inhibited by intracellular spermine. We note, however, that for all pertinent voltages, the inward currents in the absence of spermine always exceed the currents in its

control /

10 ~M SPM 0 mM ATP

500

.

.

-50

.

.

.

0

50

100 mV

-500 pA

FIGURE7.10 Spermine(SPM) inhibits outward K+transport more strongly than it inhibits inward transport via inwardly rectifying K+ (Kir) channels. The K+ currents through excised patches from Xenopus oocytesexpressingKir2.1 (IRK1) were measured at variousvalues of electrical potential in the presence or absence of 10 tzM spermine on the cytosolicside of the membrane. Spermine stronglyinhibits net outward K+ transport (current) at depolarizing (in this case positive) values of membrane potential, but it only weakly inhibits net inward transport at hyperpolarizing (negative) values. In the absence of spermine, net flux approaches unidirectional flux as the value of the membrane potential is made to vary by a greater amount from the value producingno current. Consequently,spermine appears to inhibit unidirectional as well as net K+transport in each direction. The inhibition of outward current by spermine was largely reversed by the spermine chelator ATP (10 mM) (adapted from Fakler et al., 1995, with permission from Cell Press).

Structure, Function, and Evolution of Channel Proteins

presence (e.g., Fig. 7.10). Consequently, spermine appears to inhibit K § transport at all voltages regardless of the direction of transport. Spermine is a relatively long organic molecule with multiple positive charges at neutral pH (Fig. 7.11). Moreover, K § ions are proposed to bind at several sites within the transport pathway provided by channels (Hille and Schwarz, 1978; Hodgkin and Keynes, 1955 a,b). For these reasons, Lopatin and associates (1995) suggested that spermine and other polyamines may bind at the multiple K + binding sites within the pathway. Such a mechanism of inhibition could explain how polyamines might inhibit K § uptake, although they are present predominantly on the inside rather than the outside of cells. This possible mechanism of trans-inhibition of K § transport by polyamines is reminiscent of the proposed mechanism for inhibition of K + efflux by extracellular K + (Hodgkin and Keynes, 1955b). In this case, occupation of a channel transport pathway by K + ions from one side of the membrane appears to reduce the probability that K § ions from the other side will enter the pathway. Such transinhibition could result simply from the physical restraints on transport through a narrow pore. Alternatively or in addition, K § ions might occupy, say, an outward-facing conformation of the channel tetramer, thus reducing the proportion of tetramers available to receive K + on the inside. The kinetics of K § transport may be complex, however, since reduction of the K § concentration to zero on one side ofthe membrane surprisingly reduces K + flux through a plant K + channel to zero in both directions (Maathius et al., 1997). Apparently the channel protein must sense the presence of K § simultaneously on both sides of the membrane before it undergoes the conformational changes needed for it to open. 3 Although polyamines may inhibit Kir channel transport in both directions at all voltages, membrane hyperpolarization reduces the effect (Fig. 7.10) perhaps by reducing the effective intracellular polyamine concentration as well as the total polyamine chemical potential (see Section II,B,5 of Chapter 6 and Section III,B below regarding the concept of effective concentration). Reduction of the effective concentration of polyamines might reduce their ability to serve as cis- and transinhibitors of Kir-catalyzed K § transport. In this case, Kir channels probably should not be viewed as blocked 3 The effects on unidirectional flux of the substrate concentration on the other side of the membrane has also not been studied completely and systematically in most instances of electrically silent symport and antiport of anions and cations. In the latter cases the influence of membrane electrical potential on the effective ion concentrations on either side of the membrane has most frequently been overlooked. For example, the reader may recall from Chapter 6 that consideration of the effect of membrane electrical potential on the effective anion concentrations greatly influences interpretation of the kinetics of anion exchange.

25 |

or unblocked by cations such as polyamines. Rather, polyamine inhibition should be viewed as stronger as the membrane becomes less polarized owing to an increase in the effective polyamine concentration. Moreover, polyamines may inhibit K § exodus more strongly than K § uptake because polyamines usually are more concentrated on the inside than on the outside of cells and, hence, also compete with intracellular K § for reception by Kir channel proteins. These possibilities would be immediately amenable to our assessment here if transport by channels were more frequently studied by measuring unidirectional ion fluxes in addition to the net transport reflected by current. It is of course true that unidirectional flux becomes nearly identical to current at membrane electrical potentials that are sufficiently high or low. Nevertheless, it is still possible to measure transport in both directions, as is the case, for example, for channels apparently catalyzing the net exodus of taurine (e.g., Van Winkle et al., 1994). Even under conditions where unidirectional flux occurs primarily in one direction, a better understanding of the actual mechanisms by which channels catalyze transport might be gained if unidirectional flux were measured in both directions. For example, in the case of polyamine inhibition of Kir channel transport, what might be the results of a study in which the unidirectional K § fluxes as well as the K § current are measured? We predict that unidirectional K § exodus would be inhibited much more completely than uptake by intracellular polyamines at all membrane electrical potentials. Polyamine inhibition may, however, weaken for K § transport in both directions as the outside positive membrane electrical potential increases owing to reduction of the effective intracellular polyamine concentration. Nevertheless, cis-inhibition by polyamines should be nearly complete under physiological conditions in vivo. The intracellular concentration of free spermine ranges from about 8.2 to about 76/xM (the total intracellular spermine concentration is about 100-fold higher; Watanabe et al., 1991), whereas the spermine concentration producing 50% inhibition of strong inward rectifiers at a membrane potential of - 5 0 mV is no more than about 40 n M (Kubo et al., 1993; Bond et al., 1994). If these results were observed for Kir-catalyzed unidirectional K § efflux and influx, our view of the mechanisms by which polyamines inhibit current owing to K + transport would be altered significantly. For example, polyamines would not be thought to function through loss or gain of the ability to block Kir channels at high and low values, respectively, of the outside positive membrane electrical potential. Rather competitive inhibition of unidirectional K § exodus by polyamines would depend only on the effective concentrations of poly-

252.

7. Channel Proteins Usually Dissipate Solute Gradients

CH3

COO

COO-

!

I

+

SCH2CH2CHNH3

+ NH3CH2CH2CH2CIHNH3+ L-ORNITHINE

H2 N

~c

I

-HCO3

HC

N

L-METHIONINE +ATP -PP,Pi /N C" ~\ ig2+ II CH COO/ + I + C ~ N--CH (CHOH)2CHCH2SCH2CH2CHNH3 =..... o ~ I CH3

...........................................................S-ADENOSYLMETHIONINE / DFMO

MDL73811

-3

i...................................................... (~

...........................................................S-ADENOSYLMETHIONINE DECARBOXYLASE (SAMDC)

ORNITHINE DECARBOXYLASE (ODC)

N

~c

~ H2

-I-NH3CH2CH2CH2CH2NH3+ ~ PUTRESCINE

N

/

-HCO 3-

C II ~C~

CH / + , N--CH (CHOH)2CHCH2..r CH2CH2CH2NH3

LO

~H2

(.c

SPERMIDINE I SPERMINE SYNTHASE

7 HC

'~ N

I

CH3

DECARBOXYLATED ADENOSYLMETHIONINE

SPERMIDINE SYNTHASE

+ + + I NHzCH2CH2CH~NH2CH2CH2CH2CH.:,NH3

~

C II

fN

c~

/

CH

N -- CH (CHOH)2CHCH2SCH3 L_ O I METHYLTHIOADENOSINE

+ DECARBOXYLATEDI ADENOSYLMETHIONINE~ + " + + I NH3CH2CH2CHJNH2CH2CH2CH2CH2NH2CH2CH2CH2NH: CH2CH2CH2NHs+I SPERMINE

FIGURE 7.1 1 Structure and synthesis of the polyamines putrescine, spermidine, and spermine in mammals. The polycationic nature of the polyamines could allow them to bind to several K § binding sites along Kir channels. Apparently for this reason, the greater the number of positive charges associated with a given polyamine, the lower the concentration at which it produces 50% inhibition of K § exodus from cells via Kir channels (see text). Abbreviations: DFMO, a-difluoromethylornithine; MDL73811, 5'-(((z)-4-amino-2butenyl)methyl-amino)-5'-deoxyadenosine (adapted from Tabor and Tabor, 1976, with permission from Annual Reviews, Inc.).

I

253

Structure, Function, and Evolution of Channel Proteins

amines at different voltages, although such inhibition would be obscured when current is measured in hyperpolarized cells owing to rapid and less strongly inhibited K § uptake. What other characteristics of Kir catalyzed K § transport are anticipated from its asymmetric inhibition by polyamines? First, at voltages near the thermodynamic equilibrium for K + across the membrane, stronger inhibition of exodus than of uptake should lead to uptake of K § against its total chemical potential gradient (see Fig. 4.48 in Chapter 4 for a mechanism that could produce this K + total chemical potential gradient from a polyamine gradient). Such transport of K § ions against their total chemical potential gradient by Kir channels might have heretofore unappreciated physiological significance since the transport would oppose the tendency of the K § total chemical potential gradient to approach equilibrium in most cells (e.g., section VI,A of Chapter 3). This asymmetric function of Kir channels in association with p01yamines might also help to test our proposal that some transport proteins may produce total chemical potential gradients of their substrates without coupling to a conspicuous source of free energy. We proposed in Section IX,C,2 of Chapter 4 that some of the free energy needed to produce and maintain asymmetric membranes might be used by asymmetrically oriented proteins to catalyze solute migration against its total chemical potential gradient. Since much of the asymmetry of Kir channel function depends on the presence of intracellular polyamines, it might be possible to test this hypothesis by carefully examining K + transport in cells depleted of intracellular spermine and spermidine.

2. Surprisingly, Kir Channels May Normally Catalyze Net Transport against a K § Total Chemical Potential Gradient The major current in rat basophil leukemia (RBL1) cells results from K + transport by a strong inward rectifying channel (Lindau and Fernandez, 1986). When an inhibitor of polyamine synthesis is used to reduce the levels of spermine and spermidine in these cells, however (Bianchi et al., 1996), outward currents at slightly depolarized membrane potentials are increased substantially (Fig. 7.12). As a consequence of more rapid unidirectional K § exodus at reduced polyamine concentrations, the K § total chemical potential gradient more closely approaches zero, thus stabilizing a somewhat larger inside negative resting membrane electrical potential. Put another way, polyamines appear to affect Kir channel function by causing the channels to catalyze net K + uptake at some values of membrane electrical potential that would otherwise result in net K + exodus. For example, at about - 7 0 mV, net K + is transported

A

1~1

B 2

mV

mV / !

J

-1~50 -1

I

50

~

~

~

lu..

f

10

-1 -2

-10-

C

PUT

-20 -

,. fill

SPD -

t

t0.6 7< a ~

~ -,=o 0.2 0.0

-30-

C

0.2 0.1

-40 con

0.0 DMFO MDL 73.811 500 I.tM 50 l.tM

FIGURE 7.12 Effect of inhibitors of polyamine synthesis on K+ transport via the inwardly rectifying K+ channel (Kir2.1) in rat basophil leukemia (RBL-1) cells. Net K+ transport (current) via Kir2.1 channels was measured in RBL-1 cells with control (filled triangles partly obscured by open triangles) or reduced (filled circles) levels of spermine and spermidine (A and B). The levels of both spermine and spermidine were reduced using the inhibitor of polyamine synthesis, MDL73811 (filled circles), whereas another inhibitor (DFMO, open triangles) actually raised intracellular spermine levels (C). The mechanism of action of these inhibitors is shown in Fig. 7.11. The most effective polyamine, spermine (see text), greatly decreased outward currents at depolarizing values of membrane electrical potential. Measurements of current were too variable at large hyperpolarizingvalues of membrane potential to draw a conclusion about the effect of spermine in these experiments. A small inhibition of current by spermine is, however, generally observed for Kir channels in hyperpolarized membranes (e.g., Fig. 7.10). B is a magnification of a portion of the data shown in A (adapted from Bianchi et al., 1996, with permission from American Society for Biochemistry& Molecular Biology, Inc.).

into untreated RBL-1 cells (Fig 7.12), whereas the total chemical potential gradient of K + dictates that net K + transport should be out of the cells (i.e., the measured K + equilibrium potential is - 8 7 mV; Bianchi et al., 1996). Thus, polyamines cause the resting inside negative membrane electrical potential of RBL-1 cells to be somewhat less negative (e.g., -66.6 + 0.6 vs -69.3 _+ 0.3 mV in cells not depleted vs those depleted of spermine and spermidine, respectively; p < 0.01, A N O V A ) . In addition, these compounds cause membranes to be more sensitive to depolarizing stimuli (Bianchi et aL, 1996). In the absence of sufficient intracellular spermine

254

7. Channel Proteins Usually Dissipate Solute Gradients

and spermidine, Kir channels would catalyze more net K + exodus in resting cells, thus making their membranes more difficult to depolarize. Controlled studies are, of course, needed to test our proposal that Kir channels can catalyze net K + transport against a total chemical potential gradient when the channels are made more asymmetric by association with spermine and spermidine. The source of free energy needed to drive this otherwise endergonic K + transport may be derived primarily from that needed to produce and maintain the polyamine gradient. To be physiologically significant, such net transport of K + against a gradient would need to occur in cells at their resting membrane electrical potential. The resting membrane electrical potential is lower in magnitude than the potassium equilibrium electrical potential (EK) in most types of cells. Since many investigators assume that Kir channels cannot transport K + against its total chemical potential gradient, they expect slow net exodus rather than net uptake of K + to occur at the resting membrane potential. As discussed above, however, the EK in RBL-1 cells has been determined to be - 8 7 mV, whereas their normal resting membrane potential is -66.6 + 0.6 mV (Bianchi et aL, 1996). Since net transport via Kir channels in these cells appears to occur against the K + total chemical gradient at voltages between about - 8 7 and - 6 5 mV (Fig. 7.12), net K + transport does indeed appear to occur against the K + total potential gradient at the normal resting membrane potential. Even with reduced intracellular levels of polyamines, net K + transport is against its gradient at membrane electrical potentials between about - 8 7 and - 7 2 mV (Fig. 7.12), although net transport is with the gradient at the resting potential of -69.3 _+ 0.3 mV in this case. Nevertheless, it is conceivable that Kir channels may be able to function asymmetrically to produce and maintain a K + total chemical potential gradient under some conditions even without polyamines. In this regard, the voltageproducing K + current reversal by Kir channels is frequently observed to be about the same regardless of whether polyamines are present (e.g., Fig. 7.10). For these reasons, both the phenomenon of cis-inhibition by polyamines and the asymmetric function of Kir channels to catalyze net K + transport against its total chemical potential gradient may become significant particularly at or near physiologically normal values of the resting membrane electrical potential. These properties of Kir channels also depend on their being selective for K + ions, and the magnitude of the effects may depend on other kinetic characteristics such as that known as flux coupling. Both of these topics are considered in the following section.

III. KINETICS OF TRANSPORT VIA K+ AND OTHER CHANNELS The kinetics of voltage-gated ion channels extend well beyond consideration of the kinetics of transport alone and may include such characteristics as the kinetics of activation, opening, closing, inactivation, and recovery. It is, however, the fundamental function of channels in catalyzing transport that best links them to the functioning of other transport proteins. Moreover, students of channel operation have already produced many fine monographs and review papers on the other aspects of the kinetics of voltage-gated channels (e.g., Hille, 1992; Keynes, 1994) that should easily satisfy the interested reader. For these reasons, we focus in this section on the kinetics of transport per se rather than including all of the important physiological functions of channels. Before we consider the kinetics of K + transport via channels in Section III,B below, it is instructive to consider how K + channels select this ion over others for transport.

A. Molecular Basis of K+ Channel Substrate Selectivity The high selectivity of Kv and probably Kir channels for K + over other monovalent cations appears to result from the conserved amino acid residue sequence in the H5 (Figs. 7.5 and 7.7) or pore-forming (P) region. In particular, the GYG motif in this region (Fig. 7.13) is conserved among nearly all K+-selective channels (Chandy and Gutman, 1995), and the way in which these motifs in the four subunits of the channel interact with K + ions has been described in detail (Doyle et al., 1998). The eag Kv channel in Drosophila is slightly different from other K + channels in this part of its structure with a phenylalanine residue in place of the tyrosyl one. Interestingly, the eag channel is also permeable to Ca 2+ (Bruggemann et al., 1993), whereas other K + channels may interact with Ca 2+, but they are not known to transport it (Chandy and Gutman, 1995). Heginbotham et al. (1992) performed definitive studies to show that the GYG motif is required for K + selectivity. They pointed out that although cyclic nucleotide-gated (CNG) ion channels have a P-region that is homologous to that of K + channels, it lacks two adjacent residues in the GYG motif (Fig. 7.14). Moreover, they knew that CNG ion channels do not discriminate between K + and Na + (Yau et al., 1981; Zimmerman and Baylor, 1992; Kurahashi, 1989). For these reasons, they fostered expression of mutant forms of the GYG motif of the Shaker Kv channel in Xenopus oocytes

Kinetics of Transport Via K+ and Other Channels Shaker rKvl. 1 (RCK1) Shab rKv2.1 (DRK1)

Shaw

rKv3.1 (KV4)

Shal

mKv4.1 (mShal) rKv5.1 (11<8) rKv6.1 (K13) ECOKCH

slo

mSIo KAT1 AKT1 Kirl.la(ROMK1) Kir2.1(IRK1) Kir3.1(GIRK1)

eag

NSFFKS I PDAFWWAVVTMTTVGYGDMTPVGFWGKI V E-H-S Y--TIG D=K-V G-C-TTAL -D=K AS T-YKTLL HND-N LGL A-KT= I -MFH=H-N- IG Y-QT=S-M=A-K-T A =TV-KTIA F K=N-T A =TV-STIA F E=L QS --Y-KTTL-=N SPE-T ACV-RSTP-Q=F s I P fWwa v tMTTvGYGDm P G -PRI E-=MT .... S=E -V-SESA==F -AHRL-YWTCV==L-YCETVL-=TF -NQAL=YW=CV=LL-YAKTTL-==F A- L=NRYVTL=-S=T .... T FHAENPREM=F E- L=MRYVTSM=-S=T -H-NTKEM-F VEN I NG=TS- L=S=E-QV FRFVTEQCATA= V-EVNFTA- L=S=E-Q FRCVTDECP IAVANVYNF-S- L=F=E-EA YRYI TDKCPEG= P -RKSMYVTL==T=TC N=AAETDNE -=F t m t t vGyGd

255

Kvconsensus

all K § consensus

FIGURE 7.13 Conservation of the GYG motif in the pore-forming (P) region (H5 loop) of K+-selective channels. Dashes indicate the same amino acid residue as in Shaker, and double dashes represent conservative substitutions. Consensus sequences are given both for voltage-gated (Kv, top group) and for all of the K + channels shown (bottom). Uppercase letters in the consensus sequences indicate fully conserved residues, whereas lowercase letters represent the predominant amino acid residue. Only the eag Kv channel contains a phenylalanyl residue in place of the tyrosyl residue between the two conserved glycyl residues, and this difference may render it able to transport Ca 2+ (see text). Less familiar designations: ECOKCH, putative K + channel from E. coli (Milkman, 1994); KAT1 and AKT1, K + channels originally isolated using cDNA libraries prepared from Arabidopsis thaliana at all stages of development or at the seedling stage, respectively (Anderson et aL, 1992; Sentenac et al., 1992). (adapted from Chandy and Gutman, 1995, with permission from CRC Press).

to determine whether these amino acid residues are required for K § selectivity. Mutant proteins with the YG dipeptide deleted (Fig. 7.14) no longer select K + ions over other monovalent

N

$1

S2 $3

, Shaker CNGC Chimera Deletion

~ P

$4

$5 P region $5

C

region,

1234 56789012345678901234 DAFWWAVVTMTTVGYGDMTPVGFW YSLYWSTLTLTT IG ETPPPVRD DA FWWAVVTMTTVG ETPPPGFW DA FWWAVVTMTTVG DMTPVGFW

FIGURE 7.14 The P-regions of cyclic nucleotide-gated ion channels (CNGC) lack the last two (or the first two) of the three residues in the GYG motif of K+-selective channels. Although the P-regions are otherwise homologous in the two types of channel proteins, CNG channels do not discriminate between Na + and K + as K + channels do (see text). To begin to test the theory that the GYG motif is needed for K + selectivity, the Shaker K + channel mutants, labeled Chimera and Deletion, were produced and their channel activities were studied (see Fig. 7.15) (adapted from Heginbotham et aL, 1992, with permission from the American Association for the Advancement of Science).

alkali metal cations (Fig. 7.15). Moreover, the doubledeletion mutation renders the channel susceptible to inhibition by Ca 2+, and CNG channels are also inhibited by Ca 2+ (Heginbotham et al., 1992). Hence, deletion of these two amino acid residues renders the transport characteristics of the Shaker channel more like CNG channels than like Kv channels. As discussed above, the tyrosyl residue appears to influence interaction with Ca 2+, whereas the glycyl residues and the branched-chain residue that precedes the GYG motif (Fig. 7.14) appear to be more important for K + selectivity (Heginbothen et aL, 1994). Simple substitution of various residues for one of the latter three residues renders the Shaker channel nonselective. It is not yet known for sure whether other Kv channels or the Kir channels also require these residues for their K + selectivities. Conservation of the residues in all members of these two families as well as in plant Kir channels and even in a putative K + channel in E. coli (Fig. 7.13) (reviewed by Chandy and Gutman, 1995) is, however, consistent with the theory that they are needed for K + selectivity by all of these K + channels. In this regard, substitution of a threonyl residue for the conserved branched chain residue adjacent to the GYG motif in the Shaker channel protein does not disrupt selectivity

256

7. Channel Proteins Usually Dissipate Solute Gradients

00// 00//00,//

0.8

-0.8 '

Li*

,

-40

0

1.5

-1.5

40

f

,

-40

Cs +

<= o.o .( /f

-0.5

'

-40

0

mV

Na*

0

.

0.8

-0.8

40

K*

,

i -40

o y.

0

40

TMA +

o.o 40

-0.8

-80

I

-40

0

mV

FIGURE 7 . 1 5 Deletion of the last two (or the first two) of the three residues in the G Y G motif of the P-region of the Shaker channel renders it no longer selective for K + over other alkali metal cations. The mutant channel is, however, still impermeable to the large organic cation tetramethylammonium (TMA+). Consequently, outward K + currents are observed at the depolarizing membrane electrical potentials shown when the predominant cation in the bathing solution is 100 mM T M A + owing to migration of internal K + ions (100 m M ) into the bathing solution. No inward current of TMA + via mutant channels is observed, however, at hyperpolarizing membrane electrical potentials, in contrast to the inward currents of each of the akali metal cations tested. In the case of the wild-type Shaker channel (not shown), a current-voltage relationship similar to that shown for TMA + is observed for Na+, and the wild-type channel also transport Cs + much more poorly than does the mutant channel. (adapted from Heginbotham et al., 1992, with permission from the American Association for the Advancement of Science).

like other substitutions do (Heginbotham et al., 1994), and a threonyl residue occupies this position in one of the functional plant Kir channels (KAT1 in Fig. 7.13). The conserved aspartyl residue that follows the GYG motif (Fig. 7.13) appears also to be needed for K + selectivity as well as for K + transport activity in Kv channels. Its conversion to other than a glutamyl residue in all four monomers renders the tetrameric protein nonfunctional (Heginbotham et al., 1992, 1994). When, however, the equivalent of two of the four c~-subunits in the Kv2.1 tetramer have had this aspartyl residue converted to a threonyl one, transport function is retained and can be studied. Such mutation dosage reduction can be accomplished by producing cRNAs encoding covalently linked a-subunits in which only one of the two pertinent aspartyl residues is replaced by a threonyl residue. When these cRNAs are expressed in X e n o p u s oocytes, they produce active channels that appear to have the four domains of normal tetrameric Kv channels. Nevertheless, the mutant channels are less selective for K + over other monovalent alkali metal cations than are channels with all four aspartyl residues (Kirsch et al., 1995). Moreover, the mutant channels are strongly outwardly rectifying, whereas wild type Kv2.1 channels are not (Fig. 7.16). It is conceivable that the mutant channel has reduced affinity for extracellular but not for intracellular

K + (Kirsch et al., 1995). Interestingly, however, both exodus and uptake of K + are lost in channels in which all four aspartyl residues are converted to threonyl residues. Hence, these anionic residues appear to be needed for K + transport in both direction. Perhaps significantly, these aspartyl residues are not present in Kir channel proteins (Figs. 7.13 and 7.17). Adjacent to the residue replacing the aspartyl residues is, however, an arginyl residue that is conserved among Kir channels but not among Kv channels. Similarly, Kir channels have a conserved glutamyl residue that lies six residues in the N-terminal direction from the GYG motif (Fig. 7.17) that also is not present in Kv channels (Fig. 7.13). It will be interesting to learn whether replacement of the aspartyl residue adjacent to the GYG motif in Kv channels by nearby arginyl and glutamyl residues in the P-region of Kir channels shifts a K + binding site to a somewhat different location that facilitates inward rectification. For example, such a shift could conceivably influence the ability of a spermine molecule to interact with multiple K + binding sites along the Kir channel (see Nichols and Lopatin, 1997, for a more detailed discussion of how polyamines may bind to and thus inhibit Kir channels). In the case of both Kv and Kir channels, one also wonders whether charged residue side-chains near putative K + binding sites influence other characteristics of transport. For example, the Kv channel appears normally to accommodate up to four K + ions at a time (see discussion of flux coupling below), and accommodation of the four K + ions probably depends on the presence of K + binding sites along the transport pathway. Hence, conversion of two of the four pertinent aspartyl residues in the Kv channel tetramer to threonyl residues as discussed above could conceivably alter the number of K + ions accommodated in the channel. B. Flux and Flux Coupling Both the methods used to study transport by K + channels and the kinetic models applied to that transport are frequently quite different from the methods and models utilized for other transport proteins. Although Tester (1997) has emphasized that measurement of unidirectional fluxes via channels may best reflect physiological reality, few studies employ such measurements. Rather, most investigators measure current which reflects net flux in one direction or the other across the membrane. As discussed above, the measurement of current instead of unidirectional fluxes may obscure important aspects of K + channel function. Moreover, when unidirectional fluxes have been measured for channel transport, the resultant data have been used to support conclusions about the mechanism of transport through

257

Kinetics of Transport Via K§ and Other Channels

A

2o

B

30

_E I

I

-1

I

~

I

l

Em (mV)

150

I

FIGURE 7.16 Channels formed from Kv2.1 proteins expressed in Xenopus oocytes become strongly outwardly rectifying when two of the four aspartyl residues adjacent to the GYG motifs in the four P-regions are converted to threonyl residues. The K + current-voltage relationships are shown (A) for "wild-type" channels that have been modified so that each a-subunit is covalently linked to one adjacent a-subunit and (B) for similar but mutant channels in which alternate aspartyl residues adjacent to the GYG motifs in the four P-regions of the channel have been converted to threonyl residues. A nearly linear relationship obtains for wild-type channels in the presence of 120 mM K + on both sides of the membrane, whereas the inward currents owing to K + transport via mutant channels is greatly attenuated at inside negative membrane electrical potentials (adapted from Kirsch et aL, 1995, with permission from the Biophysical Society).

channels that are very different from conclusions drawn f r o m s i m i l a r d a t a f o r o t h e r t r a n s p o r t p r o t e i n s ( s e e Sect i o n I I I , B , 2 b e l o w c o n c e r n i n g flux c o u p l i n g ) . Justificat i o n f o r t h e a p p l i c a t i o n of t h e s e d i f f e r e n t m o d e l s app e a r s u s u a l l y n o t to e m a n a t e f r o m e x p e r i m e n t a l d a t a that distinguishes among the models but rather from our notions about how channels and transporters func-

mGIRKI(Kir3.1) mlRKl(Kir2.1) mBIR (Kit6,2) mKAs-2(Kir4,1 )

tion. A s w e s h a l l a g a i n see, t r a n s p o r t at a t u r n o v e r n u m b e r of 106 sec -1 o r h i g h e r is a s s u m e d t o b e t o o r a p i d t o r e s u l t f r o m c o n f o r m a t i o n a l c h a n g e s in c h a n n e l t r a n s p o r t p r o t e i n s . If t h e p e r t i n e n t c o n f o r m a t i o n a l changes are relatively small, however, then they become m o r e difficult t o r u l e o u t o n t h e b a s i s of t r a n s p o r t velocity.

NFPSIAFLFIFI IEITQTITI GIYFI~Gi

SMVWVVlI-AIYTR IGDLI-NKAHVG N Y T P I ~ ] A N V Y I-EITEAIT I GIYI'~Y CVFWL IIAILLH l i iID D T S K V S K - - A I C V l S E V N S F T A A F L F S I MVVWVLIIAIF A H D A P G E G T N- V PIC VIT s I H S F S S A F L F S l IEIVQVlT I VVWYLVL.AIVAH D L E L G P P A NHTPICVIVQVH T L T G A F L F S L IEISQ TIT I GIYIGIF H5

mlRK1 (Kir2.1) mBIR(Kir6.2) mKAB-2(Kir4.1 )

CVTDE MVTEE Y I SEE --

I A VFMVV LA I L I L I LA I V L L I

I V GCI I DAF I IlG IAVMAIKIMAKPK I V G L M I N A I M L I G ICIFMIKITAQAH VL TT I LE I F ! TIG ITFLAIKII ARPK M2

KR L RRAIETIL/IFSIK KR I

FIGURE 7.17 Sequence similarities among portions of the primary structures of representative members of various subfamilies of Kir channel proteins. Amino acid residues that are conserved among all members of the different subfamilies are enclosed in rectangles, and the two putative transmembrane regions (end of M1 and all of M2) are underlined. The H5 loop (P-region) contains several residues that are pertinent to K + selectivity and other characteristics of Kir transport, and the loop is underlined twice. The GY(F)G motif that is needed for K + selectivity is conserved among the proteins in the H5 loop, whereas an adjacent aspartyl residue that lies just C-terminal to the motif in other K + channels (Fig. 7.13) is not present in Kir channels. An arginyl residue that is two residues C-terminal to the GY(F)G motif is, however, conserved among Kir channels, whereas it is not present in other K + channels (Fig. 7.13). Finally, Kir channels have a conserved glutamyl residue lying six residues in the N-terminal direction from the GY(F)G motif that is not present in most other K + channels. It is proposed here that these differences between Kir and other K + channels may make the Kir channels particularly sensitive to inhibition by the polyamine, spermine (see text) (adapted from Isomoto et aL, 1997, with permission from the Center for Academic Publications, Japan).

148 147 135 133

197 185 183

258

7. Channel Proteins Usually Dissipate Solute Gradients

1. Unidirectional and Net Flux The unidirectional turnover numbers for transport via channels (i.e., the numbers of solute ions or molecules transported by each channel per second) have been found to exceed 10 6 sec -1, whereas these values range from 10 ~ to 105 sec -1 for other transport proteins (recall Section I of Chapter 4). Interestingly, the fastest known enzymes have turnover numbers that also exceed 106 sec -1, although most enzymes catalyze chemical changes at slower rates (summarized by Hille, 1992). It is, however, unclear why transport that exceeds a rate of 106 sec -1 is viewed as resulting from diffusion through a narrow pore, whereas transport that occurs at a rate of 105 sec -a or less is believed instead to result from protein conformational changes. No substrate-selective channel catalyzes transport at a rate that approaches the maximum achievable by free diffusion, and most catalyze transport at rates two or more orders of magnitude below this maximum. In our view, if transport by channels and other proteins had been studied primarily as a single discipline rather than as two or more, then no rate on the continuum from 10 ~ to about 108 sec -1 would stand out as distinguishing two fundamentally different transport mechanisms. Development of the notion that the mechanism of transport via channels differs fundamentally from the mechanism for other transport proteins may also stem from the principal method used historically to measure transport by these two categories of proteins. Transport by ion channels is most often studied by measuring electric current rather than unidirectional flux. Current reflects net flux, although this net flux approaches the unidirectional flux in one direction across the membrane as the total chemical potential gradient of the ion under investigation grows larger. The study of transport as the flow of current may, however, lead us to think of transport also as a more or less continuous flow of ions rather than as resulting from the repetitive catalytic events that are needed to overcome the free energy barrier to transport. The view of biomembrane transport as a flow may also have deterred us from designing experiments to determine whether substrate-selective channels undergo small conformational changes during transport. Channel proteins also probably receive the same species of substrate differently on opposite sides of the membrane, as appears to be the case for other transport proteins. 4 These alternative views of channels 4 It should also be noted that investigators who study channel transport might make the reverse argument that other transport proteins actually function as if they formed narrow pores through the membrane. While we hold the reverse position here, what seems to us to be most important is that the transport catalyzed by channels and transporters be studied under the assumption that a more or less unified mechanism of transport may eventually be found to account for both (see Lester and Dougherty, 1998 and the following section for a summary of recent efforts towards just such a unified mechanism).

and transporters also lead to fundamentally different views of the mechanisms by which they appear to catalyze the transport of more than one ion or molecule together across the membrane.

2. Is the Mechanism of Flux Coupling Different among Channels, Symporters, and Primary Active Transporters? In Chapter 4, we presented the equation l~i-- Vmax [s]n/([S] n + K0.5n)

(4.27)

to describe the flux coupling of transport of more than one of the same ion or molecule of a substrate(S) together in the same direction across a membrane where n is the number of such ions or molecules. In the absence of a propelling force (other than a substrate concentration gradient) that would favor transport of a given species of ion or molecule in one direction or the other (i.e., for functionally symmetric transport across a symmetric membrane) 5 one can divide the formula for unidirectional efflux by that for unidirectional influx to derive a relationship between the two fluxes termed the flux ratio (Vie/Vii). Furthermore, if one measures transport at substrate concentrations well below the K0.5 value, as is frequently the case for channels, then the expression for the flux ratio can be simplified as follows Vie/Vii = (Vmax [s]in/go.5n)/(Vmax [S]on/go.5n), which reduces to Vie/Vii-- ([S]i/[S]o) n,

(7.1)

where Vie is the unidirectional efflux, Vii is the unidirectional influx, [S]i is the intracellular substrate concentration, and [S]o is the extracellular substrate concentration. Equation (7.1) is applied to uniporters and channels where the value of n is the number of ions or molecules transported together in the same direction. When the value of n exceeds 1, flux coupling occurs. 6 Interestingly, however, the mechanism of flux coupling is assumed 5 The reader is reminded that such propelling forces are always potentially present, and some of them may be much less conspicuous than ATP hydrolysis or the membrane electrical potential (e.g., see Section IX,C,2 of Chapter 4). 6 The exponential relationship of flux coupling to the flux ratio could conceivably lead to large differences in the unidirectional fluxes of some substances even though their concentrations may be nearly equal on each side of the membrane. For example, water channels could conceivably transport several hundred water molecules together, as appears to be the case for water cotransport by the Na+dependent glucose transporter (Loo et aL, 1996). If such is the case, then relatively small differences in water concentration, as occur in osmotic imbalances, would result in exponentially more rapid transport via water channels to reverse the imbalance (see Section III of Chapter 4 for further discussion of the study of transport via water channels.)

259

Kinetics of Transport Via K+ and Other Channels

to differ for channels and transporters. In the case of channels, the value of n is taken to be the maximum number of ions or molecules that can occupy a channel at one time (e.g., Hodgkin and Keynes, 1955 a,b; Hille, 1992; Stampe and Begenisich, 1996). For transport catalyzed by channels, ions or molecules are viewed as migrating more or less single-file through a narrow pore. Transport via channels is believed to be too rapid to occur by a mechanism in which several ions or molecules first associate with and then are transported together by the transport protein. Such a simultaneous mechanism would likely be assumed, however, when the value of n exceeds 1 for the transport catalyzed by a uniporter. 7 These distinctions between transporters and channels begin to break down on consideration of bacterial ion transporters/channels (Jan and Jan, 1997). Bacterial transporters/channels are so named in part because it frequently is not obvious in which category they ought to be placed. For example, some of these transporters/ channels catalyze transport that is as rapid as that catalyzed by channels (Jan and Jan, 1997). It has also been concluded, however, that some of these proteins couple simultaneous transport of more than one ion to ATP hydrolysis (e.g., Fendler et al., 1996). We suggest again that a common mechanism underlies almost all of these transport processes. Most proteins classified as transporters (or carriers) catalyze slower transport than the proteins classified as channels, and members of the two categories may even use different propelling forces to produce net migration of their substrates in one direction or the other across the membrane. Similarly, the flux coupling that occurs in association with the transport catalyzed by some channels may result in part from trans-inhibition by the same species of substrate on the other side of the membrane (e.g., Hodgkin and Keynes, 1955b), whereas trans-stimulation (antiport) or no trans effect (uniport) are often expected for proteins classified as transporters. (But see also Section III,B,3 of Chapter 6 for a discussion of the trans-inhibition by some amino acids of the antiport catalyzed by systems ASC and the effect of such trans-inhibition on the stoichiometry of cosubstrate transport.) The use of such criteria to propose fundamentally different mechanisms for the flux coupling that occurs via channels and transporters has, however, not yet been supported experimentally. A simple and physiologically pertinent way to introduce an additional propelling force into the equation for the flux ratio of uniporters and channels is to assume that the membrane has an electrical potential and that 7 Interestingly, in the well-studied case of Na+/amino acid cotransport by system ASC, n may exceed one for Na + although Na + ions appear to migrate across the membrane one at a time (see Section III,B of Chapter 6).

the substrate is an anion or a cation. In this case, Eq. (7.1) becomes Vie/Vii = ( ( [ S ] i / [ S ] o ) e x p ( z s f ( X I t i -

Xrgo)/RT))n

'

(7.2)

where 5,, R, and T have their usual meanings (as defined in Chapter 3), z is the charge of a single substrate ion, and q~i - q~o is the membrane electrical potential. Equation (7.2) was derived originally by Ussing (1949) for the special case where n = 1, albeit from a different beginning than we use here. The Ussing equation was subsequently modified by Hodgkin and Keynes (1955 a,b), who found that n could exceed a value of 1 for some channels. The value of n has been found to exceed 1 for many ion channels (reviewed by Hille, 1992), including the voltage-gated Shaker K + channel (Stampe and Bengenisich, 1996). As discussed above, however, the conclusion that the value of n is the number of ions that can occupy the channel at the same time is based on the assumption that transport of ions through the channel is too rapid to occur by a mechanism other than diffusion of the ions through a narrow pore. In the case of voltage-gated K + channels, it has been concluded that up to 4 K + ions may occupy the channel pore at one time because n was found to equal 3.4 for the Shaker channel at a membrane electrical potential of - 3 0 mV (Stampe and Begenisich, 1996). Such a view apparently gained support recently by solution of the structure of the K + channel from Streptomyces lividans at 3.2 A resolution (Doyle et al., 1998). Nevertheless, such static structural studies scarcely rule out the possibility that small conformational changes may occur during transport. Moreover, occupation of the channel selectivity filter by only 2 K + ions in the static structure (Doyle et aL, 1998) is somewhat difficult to reconcile with observed higher values of n for the functioning of homologous K + channels. The selectivity filter appears to constitute the rate-limiting steps for transmembrane migration of K + ions (Doyle et aL, 1998), and the primary structures of the filter constituents are highly conserved among all K + channels. The filter is composed of the tvGyG motifs in the P-regions of the four channel monomers (Fig. 7.13), and the importance of this motif and other residues in the P region to K + selectivity were discussed in Section III,A above. In regard to the mechanism of K + transport, a summary of recent data supports the theory that cation channels may contain a single high-affinity substrate binding site flanked by sites of lower affinity (Lester and Dougherty, 1998). These findings fit the accepted view of transporters, but they are inconsistent with the conclusion that two or more nearly equivalent substratebinding sites span a pore through the membrane formed by cation-selective channels. In contrast, data for other transport proteins support the opposite view that anion channels, symporters, and even primary active trans-

260

7. Channel Proteins Usually Dissipate Solute Gradients

porters catalyze transport via pathways with multiple substrate binding sites, which is reminiscent of the accepted model for channels (Lester and Dougherty, 1998). While these data and models are still incomplete and may, at first, even seem mutually inconsistent, they support the notion that we may yet develop a unified theory of the mechanism by which all substrate-selective biomembrane transport proteins function. Now that the quaternary structure of a K + channel has been described at a high level of resolution (Doyle et aL, 1998), it will be interesting to apply to the structure the computer simulation that was mentioned in Section I above. According to this simulation, water and presumably Na + are unable to migrate across voltage-gated Na + channels unless the transport protein is able to move as it normally does at physiological temperatures (Singh et al., 1996; Jakobsson, 1997). The narrow part of the K + channel is lined with peptide backbone carbonyl oxygens (Doyle et al., 1998), much like the gramicidin channel (Chiu et aL, 1989; 1991, 1992, 1993), and the gramicidin peptides also must move according to the simulation in order to catalyze transport. While this movement may not be large, it seems to us to be a likely requirement for transport by nearly all such proteins. Hence, it appears that transport proteins should be placed on a continuum from those using only relatively small movements to accomplish transport to those that undergo relatively large conformational changes. The other common feature of most transport proteins is that they each form a pathway across which only certain solutes or the solvent may migrate owing to their physical and chemical properties. The close interaction between most of these proteins and their substrates apparently leads to slower migration of the substrate than would otherwise occur over the same distance by ordinary diffusion. It is also instructive to note again that the flux ratios defined by Eq. (7.2) do not necessarily hold for channels under allconditions in which they should apply. For example, instead of rapid net exodus no exodus of K + occurs via some plant K + channels when the extracellular K + concentration is zero (reviewed by Maathius et al., 1997). According to these authors, the channel proteins appear somehow to sense extracellular K +, which is thought to be a prerequisite for channel opening. While we do not intend to imply here that enough data exists for us to suggest that this sensing mechanism resembles obligatory exchange of the type discussed in the preceding chapter, the consequence of the finding is that exodus of K + can occur only when uptake also occurs, albeit frequently at greatly different rates. What, however, may be the result of the application of Eq. (7.2) to transport if we take this interdependence of uptake and exodus to its logical extreme of obligatory exchange?

Although this excursion may seem to be taking us far from the kinetics of transport via channels, it should be valuable for a number of reasons, including that it may seem like no more than an excursion. As just discussed, the mechanisms and consequently the kinetics of transport by channels and transporters may be viewed better as similar rather than as different. As we shall see, for example, when Eq. (7.2) is applied to obligatory exchange, the Nernst equation can be derived. The Nernst equation was derived initially in Chapter 3 for the Gibbs-Donnan equilibrium, and it is applied frequently to channel transport, although not usually to antiport. 3. How Does Obligatory Exchange of an Anion or a Cation Resemble a Gibbs-Donnan Equilibrium Established through Channel Transport?

The tendency of any ionic solute to migrate more rapidly down its total chemical potential gradient than against it across a membrane should not change with the means available for such migration as long as that means is not coupled to an additional source of free energy. Consequently, when the flux ratio must equal unity owing to obligatory exchange of a solute, Eq. (7.2) can be manipulated as follows n

Vie/Vii -- (([S]i/[S]o) exp

In(vie/Vii) = In (IS]i/IS]o) n + nZs 5( i- q%)/RT 0 =

(7.3)

In (Vie/Vii) - - I n ([S]i/[S]oY + nZs S ( q * i In ([Si]/[S]oY = - nZs S ( q * i -

(7.2)

q o)/RT.

o)/RT (7.4)

In its simplest form where n = 1, Eq. (7.4) is the Nernst equation, which was derived in Chapter 3 (Eqs. (3.23) and (3.24) and again here under the reasonable assumption that substrate concentrations are nearly equal to substrate activities in relatively dilute solutions. The Nernst equation describes an equilibrium in which the flux ratio is one for a charged solute although the concentrations of the solute are different on opposite sides of a semipermeable membrane. The concentration gradient results from the influence of the membrane electrical potential on the total chemical potential of the solute. The membrane electrical potential must, however, also be viewed as influencing the effective concentration of the charged solute on each side of the membrane. The effective solute concentrations must be influenced by membrane potential even for the equilibrium described by the Nernst equation because (a) the kinetics of unidirectional flux are expressed in terms of solute concentration rather than total chemical potential (e.g., Eq. (4.27) above and in Chapter 4) and (b) the inward-

Kinetics of Transport Via K§ and Other Channels

to-outward unidirectional flux ratio is 1 at equilibrium even though the absolute solute concentrations are usually different on opposite sides of the membrane. Similarly, when the unidirectional flux ratio in Eq. (7.2) is required to be 1 by obligatory exchange, the magnitude of the membrane electrical potential must increase as the concentration gradient of an anionic substrate is increased across the membrane. Such an increase in the magnitude of the membrane electrical potential would increase the effective substrate concentration on one side of the membrane and decrease it on the other. The concept of effective concentration was first introduced in chapter 6 to account for the transport kinetics observed for anion exchange (Section II,B,5). Experimental data presented in that chapter (Schnell and Besl, 1984) support the present notion that when the concentration of an ionic substrate is increased on one side of the membrane to increase unidirectional flux in one direction, compensatory changes in the membrane electrical potential (~i - ~o) also must occur to increase the rate of transport in the reverse direction. The compensatory changes in ~i - ~o increase the rate of transport in the reverse direction by increasing the effective substrate concentration on the other side of the membrane. In studies of anion homoexchange, Schnell and Besl (1984) observed that both the appropriate unidirectional flux and the value of XIt i - - XIt o changed when the substrate concentration was changed on one side or the other of the membrane of resealed red cell ghosts. The change in XIt i - - xi~to was s u c h that it changed the total chemical potential of substrate on the side of the membrane where there had been no change inactual concentration so that the velocity of transport could remain the same in both directions. Hence, we see that in this relatively simple system where the compositions of the solutions on both sides of the membrane are well controlled, it is indeed the case that the effective substrate concentrations can be viewed as remaining equal on both sides of the membranes as required for vie to equal vii in obligatory exchange. An interesting possible consequence of these constraints on obligatory exchange is that such exchange may occur only for charged solutes. For instances of exchange discussed in detail in this volume, it does indeed appear to be the case that obligatory or nearly obligatory exchange is observed only for charged solutes. For example, anion and cation exchange via the AE and EAAT/ASC proteins is obligatory (Chapter 6), whereas glucose exchange via the GLUT proteins is not (Chapter 4). 8 While the proportion of transport that 8 The effect of membrane electrical potential on Na§ threonine transport via system ASC in human fibroblasts (Bussolati et al., 1992) is also consistent with the theory that voltage influences the effective concentrations of charged substrates undergoing electrically silent exchange.

261

results from exchange may change for glucose in physiologically significant ways (see Section XI,F of Chapter 4) exchange may never become required for glucose transport. Obligatory exchange of glucose could conceivably occur if a change in its concentration on one side of the membrane were able to influence the values of Km and Vmax for transport of glucose on the other side. No such influence of substrate concentration on the values of the kinetic parameters is needed to produce obligatory exchange of a charged solute, however, as long as the membrane electrical potential is free to change in the appropriate way with changes in substrate concentration. 4. Changes in Membrane Electrical Potential Change the Effective Concentrations as Well as the Total Chemical Potentials of Charged Solutes on Both Sides of the Membrane

By now the reader has undoubtedly noted a failing of most existing kinetic equations to account fully for the unidirectional transport of charged solutes. This failing becomes particularly conspicuous under the constraints of obligatory exchange. The difficulty arises because most theoretical considerations of transport kinetics have been developed under the assumption that the velocity of transport varies with substrate concentration. In reality, however, the velocity of transport appears to depend on the total chemical potential of charged solutes of which their concentration is only a component. Since investigators of ion channels usually measure transport using electrical current, they remain acutely aware of the importance of membrane potential to the velocity of ion transport. This relationship becomes conspicuous at voltages far from the reversal electrical potential where currents approximate unidirectional fluxes. The relationship is obscured, however, and consequently perhaps almost forgotten in the case of the transporters when radiolabeled ionic solutes are used to study unidirectional fluxes and the membrane electrical potential is not measured. The fact that fluxes depend on voltage may be made obvious, however, by measuring the fluxes at different values of the membrane electrical potential. For example, in Fig. 7.10 the current produced by K + uptake appears to approach that which would be produced by its unidirectional flux at potentials approaching - 5 0 mV, and current varies about linearly with voltage near this value of the membrane electrical potential. 9 9 While we assert in several places that net flux approaches unidirectional flux as the membrane electrical potential grows larger in magnitude, data supporting this assertion are surprisingly scarce. Since unidirectional influx and efflux are rarely measured in concert with currents, it is conceivable that in some cases current constitutes only a small component to unidirectional flux owing to unanticipated increases in unidirectional influx and efflux with increases in the magnitude of the membrane electrical potential.

262

7. Channel Proteins Usually Dissipate Solute Gradients

Now it can be shown that

R T In (Vie~Vii) = A u tsi-o,

(7.5)

where Vie~Vii is the flux ratio for exodus and uptake and A Utsi_o is the total chemical potential gradient of the solute. The quantities on both sides of Eq. (7.5) are equal to each other because they are also equal to the same quantity

R T In

(Vie~Vii)

= A Utsi_o

= R T In ([S]i/[S]o) + Zs ~ - ( q t i - ~o) as shown in Eqs. (7.3) (n = 1) and (3.35) (see Chapter 3). In addition, we know from the preceding discussion that both Vie and Vii may vary with membrane electrical potential albeit in opposite directions for given intracellular and extracellular substrate concentrations (or activities). For these reasons, it might be better to derive transport kinetics in terms of solute total chemical potential rather than concentration. Such a derivation would require additional effort since the equations above were derived under the assumption that the substrate concentration is well below the Km (or K0.5) value. While many investigators might be willing to entertain such a change, most of us would probably prefer to continue to use present formulations. Moreover, unidirectional transport is frequently measured in large numbers of living cells where values of total chemical potential may be difficult to measure accurately and simultaneously with unidirectional flux. A reasonable approach may be simply to remember to think of effective substrate concentrations rather than actual ones, at least in qualitative assessments of the meaning of kinetic data obtained using radiolabeled tracers. Although precise values of the effective concentrations are unlikely to be available in most studies, their consideration may improve interpretations of the resultant kinetic data for transport of cations and anions. As we saw in Chapter 6 (Section II,B,5), consideration of effective substrate concentrations completely changed our conclusion about whether atlion exchange exhibits positive cooperativity under experimental conditions.

IV. SUMMARY While similarities in the function of channels and other transport proteins are emphasized at the end of this chapter, we describe novel aspects of the structures and functions of two distantly related families of K + channels near the beginning. These examples were chosen to represent transport and its regulation via channels that are selective for their substrates. Other channels, such as the a-hemolysin toxin family, differ from the

K + channels discussed here in that they are not selective and they evolved to lyse cells rather than to contribute to cellular function. Such toxins do indeed appear to function like nonselective pores, whereas substrateselective channels appear to interact intimately with their substrates in a manner analogous to enzymesubstrate interactions (Marban and Tomaselli, 1997). Both voltage-gated (Kv) and inwardly rectifying (Kir) K + channels function as tetramers in biomembranes. Their selectivity for K + over other monovalent alkali metal cations is attributed largely to the extracellular H5 loop or pore-forming (P) region between the putative transmembrane segments $5 and $6 or M1 and M2 in Kv and Kit channels, respectively (Fig. 7.5). That the P-region in each monomer contributes to K + selectivity and transport is supported by the observation that replacement of a conserved aspartyl residue in the loop of every other monomer in the Kv2.1 tetramer reduces the selectivity of the channel for K + over other monovalent cations (Kirsch et aL, 1995). Replacement of the same residue in all four subunits destroys Kv2.1 channel transport activity. While this aspartyl residue is not present in Kir channel proteins, a glutamyl residue at a different position in the P-region is present in Kir but not in Kv channel proteins. We suggest that this conserved glutamyl residue in Kir channel proteins serves a function similar to the conserved aspartyl reside in the P-region of Kv channel proteins. The first four putative transmembrane segments of each Kv channel monomer form an outer shell in the tetramer, which is not present in Kir channels (Fig. 7.5). Transmembrane segment $4 in the Shaker Kv channel senses changes in electrical potential across the membrane owing to its seven cationic residues at every third position. The segment moves toward the outside of the membrane upon depolarization thus permitting transport, and it can move back in again upon repolarization to close the channel. Prolonged depolarization leads, however, to an inactive state during which the channel is insensitive to changes in membrane electrical potential. Both rapid N-type and slower C-type components contribute to the inactivation process, and N-type inactivation facilitates C-type inactivation. While Kir channels have no such well-defined structural components that contribute to its opening, closing, and inactivation, they are regulated by intracellular polyamines. These polycations appear to block or inhibit K + exodus via the channel more strongly than they inhibit uptake probably owing to their more direct competition with intracellular than with extracellular K + for K + receptor sites. Moreover, hyperpolarization of the membrane reduces the effective concentration of intracellular polyamines, thus reducing their inhibition of rapid net K + influx. This effect of poly-

Summary

amines also appears to render the asymmetrically oriented Kir channels better able to catalyze net K + transport against its total chemical potential gradient. The surprising ability of Kir channels to catalyze uphill K + transport is coupled primarily to the free energy required to produce and maintain total chemical potential gradients of polyamines across the plasma membrane. Such uphill transport appears to occur especially at voltages at or near the resting membrane potential. Consequently, transport by polyamine-influenced Kir channels lowers the resting membrane potential somewhat and renders the membranes more sensitive to depolarizing stimuli. Polyamines are proposed to interact with multiple K + binding sites along the transport pathway formed by Kir channels. In this model, two spermine molecules are believed to span the pathway (Lopatin et aL, 1995), which may accommodate several K + ions at a time. In the case of the Shaker Kv channel, up to 4 K + ions are believed to be able to simultaneously occupy the channel (Stampe and Begenisich, 1996). Alternatively,

263

we propose that the channels may undergo small conformational changes to catalyze the migration of 3 or 4 four K § ions together. While different mechanisms have heretofore been proposed to account for the apparent migration together of more than one substrate ion or molecule via transporters and channels, we encourage investigators to search for a common explanation for the phenomenon. From such studies, common mechanisms for substrate-selective transport may also begin to emerge. Substrate-selective transport also usually is subject to regulation as discussed above for Kv and Kir channels. Regulation of transport is discussed more broadly in Chapter 9. The importance of transport regulation to interorgan nutrient flows is discussed in Chapter 10. In Chapter 8, the functions of all known transport proteins are summarized and a scheme for their systematic classification is presented. All of these findings and insights have been forthcoming owing to the many sophisticated procedures for studying transport that are now available, as summarized in Chapter 11.

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A Proposed System for the Classification of Transmembrane Transport Proteins in Living Organisms*

With the advent of genome sequencing, it became a tangible goal to recognize and understand the diversity of transmembrane molecular transport processes found in living organisms on this planet. To facilitate integration of all available relevant information into a limited number of conceptual frameworks, a system of transport protein classification had to be devised. Although tremendous effort had been devoted to the classification of enzymes, no comparable effort had been expended with transporters. Prior efforts to name and classify transport proteins have been restricted mainly to specific categories of transporters, e.g., amino acid porters (Christensen et aL, 1994) and ion channels (Alexander and Peters, 1997), although a broader but still incomplete effort was published recently (Griffith and Sansom, 1998). In this chapter, we describe our efforts to create a universally applicable system of classification for this functional class of proteins. For more detailed descriptions of the proposed system, the interested reader is referred to our World Wide Web site (http://www-biology.ucsd.edu/~msaier/transport/ titlepage.html). For our detailed genome analyses of transport proteins, see http://www-biology.ucsd.edu/ --~ipaulsen/transport/titlepage.html. Several recent publications from our laboratory deal with these topics (Pao et aL, 1998; Paulsen et aL, 1996, 1997, 1998; Saier, 1994, 1996, 1998).

I. INTRODUCTION Transport proteins mediate the uptake of essential nutrients as well as the exodus of end products of metabolism and toxic substances. They also maintain osmotic stability and control the ion contents of cells, thus being principal targets of homeostatic control mechanisms. Transporters export macromolecules that serve extracytoplasmic functions. They allow intercellular communication by facilitating release and uptake not only of macromolecules, but also of neurotransmitters, hormones, pheromones, alarmones, and a variety of other small signaling molecules. They allow organisms to use biological and chemical means to injure or destroy others by catalyzing export of deleterious toxins, virulence factors, and other substances. Finally, transporters provide the key constituents for the production of bioelectricity and the control of cellular electrical activities. All of these functions are universally required for the maintenance of life in its many distinct and varied forms. The importance of molecular transport to living organisms cannot be overestimated. Recent complete genome analyses have revealed that roughly 10% of all genes found in microorganisms encode transport proteins (Paulsen et al., 1998). This scope alone suggests the importance of transport to cellular function. We are only now coming to appreciate the tremendous variety of molecular transport mechanisms that have evolved in order to allow living organisms to cope with fluctuating environmental conditions while maintaining internal conditions compatible with life. A full appreciation of this scope will undoubtedly require decades of intense study.

II. WORK OF THE ENZYME COMMISSION AS A BASIS FOR THE SYSTEMATIC CLASSIFICATION OF TRANSPORT PROTEINS Enzymes have long been classified in accordance with the directives and recommendations of the E n z y m e Commission (EC) (Dixon and Webb, 1979). The Corn-

tMilton H. Saier, Jr., Department of Biology, University of California at San Diego, La Jolla, California 92093-0116.

265

266

8. Classification of Transport Proteins

mission developed its directives decades ago, before protein sequence data were available. Their system was based exclusively on function. It was assumed that proteins of similar catalytic function would be closely related and that they therefore should be grouped together. We now know, however, that two different enzymes catalyzing exactly the same reaction sometimes possess completely different amino acid sequences and three-dimensional structures, function by entirely different mechanisms, and apparently have evolved independent of each other, converging only with respect to the final reactions catalyzed. The EC classification system thus reflects only the reactions catalyzed by and the substrate specificities of the enzymes. It does not recognize the phylogenetic origins of these proteins. As has been extensively documented, molecular phylogeny is a reliable guide to protein structure and mechanism of action, but not necessarily to the specific process catalyzed or the substrate acted upon. Since the former characteristics are fundamental traits of a protein while the latter characteristics may be considered more superficial traits, sometimes merely reflective of single amino acyl residue changes in a protein, it would be reasonable to suggest that as more and more sequence and phylogenetic data become available, these data should provide a more reliable basis for protein classification. Since single-residue substitutions can give rise to different substrate binding properties, these characteristics are recommended for use in the final level of classification rather than at a primary level. We therefore suggest that the evolutionary process provides the most reliable indications of structure, mechanism, and function. If molecular phylogenetic studies can accurately retrace the evolutionary process, their use is encouraged as a basis for a rational system of protein classification. Some of the enzymes classified within the EC system are asymmetrically situated within an anisotropic, hydrophobic lipid membrane that separates two aqueous compartments. The resultant asymmetry allows these enzymes to catalyze vectorial as well as chemical modification reactions as clearly enunciated decades ago by Peter Mitchell (Mitchell, 1961, 1962; Mitchell and Moyle, 1959) and others (summarized by Christensen, 1962). Some of these integral membrane enzymes also catalyze transmembrane transport of small solutes without chemically destabilizing them. However, most currently recognized solute transporters do not catalyze a chemical reaction and consequently are not included within the EC classification system. The comprehensive system of classification proposed here has the potential to encompass all types of transporters, both those that are currently recognized and those that are yet to be discovered.

II1. PHYLOGENY AS A BASIS FOR PROTEIN CLASSIFICATION: CRITERIA FOR FAMILY ASSIGNMENT Homologous proteins are defined as those which share a common evolutionary origin (Doolittle, 1986). Such proteins are said to be members of a phylogenetic family. Homology can be recognized either by comparison of the primary structures (amino acid sequences) of proteins or by comparison of their tertiary (threedimensional) structures. Primary structure generally diverges more rapidly than does tertiary structure. Thus, when two homologous proteins have diverged in amino acid sequence to the extent that homology cannot be established on the basis of statistical analyses of their sequences, three-dimensional structural analyses can still be used to identify common protein folds (Bork et al., 1992; Doolittle, 1992, 1994). Unfortunately, very few transport proteins are understood in three dimensions. It is therefore necessary to depend on primary sequence analyses in order to provide reliable criteria for establishing common evolutionary descent. For the purposes of assigning a protein to a recognized family, we have followed previously enunciated criteria set up by Doolittle (1986, 1992, 1994). Thus, two sequences are optimally aligned and the percentage of identity is measured. The two sequences are then randomly shuffled and the percentage of identity of the randomized sequences is again determined. A comparison score is then computed which relates the probability that the percentage of identity observed for the two native protein sequences has occurred by chance. These comparison scores are expressed in standard deviations (SD). Because the native sequences are shuffled, the method corrects for potential compositional uniformity such as that observed with highly hydrophobic proteins. Several programs are available for comparison score computation. These include A L I G N (Dayhoff et al., 1983), Los Alamos (Kanehisa, 1982), RDF2 (Pearson and Lipman, 1988), and GAP (Devereux et aL, 1984). If the degree of similarity between the two native amino acid sequences is compared with a large number of random shuffles (i.e., ->500), reliable values are obtained. When two sequences give a comparison score within 3 SDs or less of the mean of the random shuffles, there is little or no evidence for homology. However, if they give a comparison score of 6 SDs above the mean, the probability ( p ) that the degree of similarity exhibited by these two sequences arose by chance is about 10 -9 . This result suggests that such sequences probably arose from a common ancestor by divergent evolution, but this degree of sequence similarity could conceivably have arisen by a convergent evolutionary process, particularly if the sequences compared are

Proposed ClassificationSystem short. Thus, homology is not established. On the other hand, when a comparison score is high, i.e., ->9 SDs above the mean of the random shuffles (p -< 10-19), the degree of similarity is considered to be too great to have arisen either by chance or by a convergent evolutionary process, so the two sequences are considered to be homologous (Doolittle, 1986). Methods for assessing the statistical significance and reliability of particular molecular sequence features have been presented (see, for example, Altschul et aL, 1990; Devereux et aL, 1984; Felsenstein, 1988; Karlin and Altschul, 1990). It should be noted that the term "convergence" has been used loosely by different investigators to refer to the independent evolution of a similar sequence (sequence convergence), a similar specificity or catalytic function (functional convergence), or a similar topology or structural scaffold (structural convergence) (Bork et aL, 1993). In the Transport Commission (TC) system proposed here, two proteins are assigned to the same family if they exhibit comparable regions of at least 60 residues that give a comparison score of at least 9 SD above the mean of the shuffles. If proteins A and B give a comparison score of at least 9 SD above the mean and proteins B and C do also, then proteins A and C are also homologous, even if they do not give a comparison score of 9 SD or greater above the mean. This presumption is called the "superfamily principle." However, if no pair of proteins between two established families of transporters give a comparison score of at least 9 SD above the mean for a stretch of at least 60 residues, we assign them to different families. Only when a "missing link" protein becomes available can the two families be joined into a single superfamily. A superfamily is thus rather arbitrarily defined as a collection of distantly related families (i.e., see discussion on the MF and ABC superfamilies in Section VII of this chapter).

IV. PROPOSED TRANSPORT PROTEIN CLASSIFICATION SYSTEM 1 According to the proposed classification system, transport proteins are grouped on the basis of four criteria, and each of these criteria corresponds to one of the four numbers within a suggested TC number for a particular type of transporter. The proposed four components are W.X.Y.Z, where W corresponds to the transporter type and free energy source used to drive 1The proposed system has been submitted to the Nomenclature Committee of the International Union of Biochemistryand Molecular Biologyfor its consideration. While it is anticipated that the proposed system will have an impact on any system that is finally adopted, the system that is eventually adopted could differ considerably from the one proposed here.

267

net transport (if any), X specifies the family or superfamily to which the transporter belongs, Y represents the subfamily (or family in a superfamily) in which the particular porter is found, and Z delineates the substrate(s) transported. Any two transport proteins in the same subfamily of a family that transport the same substrate(s) using the same mechanism are given the same TC number, regardless of whether they are orthologs (i.e., arose in distinct organisms by speciation) or paralogs (i.e., arose within a single organism by gene duplication). Sequenced homologs of unknown function are not normally assigned a TC number, and functionally characterized transport systems for which sequence data are not available are also not included. These deficiencies will be eliminated with time as the transport function of sequenced porters are characterized biochemically and as sequences become available for proteins in functionally characterized transport systems. The primary level of classification in the TC system proposed here is based on transporter type and free energy source. Thirteen primary categories are proposed as follows: 1. Channel-type facilitators. Proteins in this category have transmembrane channels that usually consist largely of a-helical spanners. Transporters of this type catalyze solute or solvent migration by a process independent of free energy other than that in the transmembrane total chemical potential gradient of the substrate. They allow passage through transmembrane channels or in some cases even aqueous pores without evidence of the relatively large conformational changes associated with transport via proteins historically termed carriers. Outer membrane porin-type channel proteins are excluded from this category and have been put into their own category (category 9). As over a dozen distinct channel-forming peptides and proteins have been elucidated in three dimensions (e.g., Table 8.1), the structural basis of channel formation is well understood for over 30% of the families shown in Table 8.2. 2. Carrier-type facilitators. Transport systems are included in this category if they utilize a relatively large conformational change to catalyze uniport (a single species is transported by a process not coupled to a change in chemical free energy), antiport (two or more species are transported in opposite directions in a tightly coupled process not directly linked to a form of free energy other than chemiosmotic free energy), and/or symport (two or more species are transported together in the same direction in a tightly coupled process not directly linked to a form of free energy other than chemiosmotic free energy). While the term "carrier" now appears unlikely to apply literally to any transport protein, it is retained here to help indicate that the confor-

268

8. Classification of Transport Proteins TABLE 8.1 TC number

Transport Proteins for Which Three-Dimensional Structural Data Have Been Reported a,b Protein

I. Channel-type peptides and proteins 1.6.1.1 Acetylcholine receptor 1.8.3.1 Melittin 1.8.4.1 Defensin 1 1.12.1.1 Colicin Ia 1.12.2.2 Colicin E1 1.13.1.1 Cry 1Aa 1.13.2.1 Cry 3Aa 1.14.1.1 a-Hemolysin 1.15.1.1 Aerolysin II. Porins 9.1.1.1 Porin (OmpC) 9.1.1.3 Porin (OmpF) 9.7.1.1 Porin (PorCa) III. Redox-driven proton pumps 6.3.2.1 Quinol:cytochrome c reductase 6.4.6.1 Cytochrome c oxidase 6.4.7.1 Cytochrome c oxidase IV. Light-driven proton pumps 7.1.1.1 Bacteriorhodopsin 7.2.1.1 Reaction Center

Family

Source

LIC CAP CAP Colicin Colicin ICP ICP aHL Aerolysin

Torpedo electric organ Bee venom Homo sapiens Escherichia coli Escherichia coli Bacillus thuringiensis Bacillus thuringiensis Staphylococcus aureus Aeromonas hydrophila

GBP GBP RPP

Escherichia coli Escherichia coli Rhodobacter capsulatus

QCR COX COX

Bos taurus Paracoccus denitrificans Bos taurus

BR RC

Halobacterium salinarium Rhodobacter spheroides

aThree-dimensional structural data for transporters included within TC categories 2-5, 8, 10, 11, and 99 are not yet available. However, X-ray crystallographic structures for soluble proteins or protein domains of several transporters in TC categories 3 and 4 are available (e.g., TC 3.1: extracytoplasmic receptors for ABC-type transporters. (Quiocho and Ledvina, 1996);TC 3.2:F1 components of F-type ATPases (Abrahams et al., 1994), and TC 4.1, 4.2, 4.3, and 4.6: the soluble IIA and IIB domains of PTS transporters (Liao et al., 1991; van Montfort et al., 1997)). Because the structures of the integral membrane constituents of these transport protein complexes are not known, these proteins are not tabulated here. bLiterature citations describing the structural data summarized in this table are available by reference to our web site (http://www-biology.ucsd.edu/-~msaier/transport/titlepage.html).

mational changes used by these proteins to catalyze transport are apparently much larger than the small movements required of channels (see Chapter 7). Three-dimensional structural data are not available for these porters or for those included in categories 3-5, 8, 10, 98, 99, or 100. 3. Active transporters driven by diphosphate (pyrophosphate) bond (usually in A T P ) hydrolysis. Transport proteins are included in this category if they hydrolyze the terminal diphosphate bond in ATP, in another nucleoside triphosphate or in diphosphate itself to drive the active uptake, and/or extrusion of a solute or solutes against their total chemical potential gradients. The transport protein may or may not be transiently phosphorylated, but the substrate is not phosphorylated. 4. PEP-dependent, phosphoryl transfer-driven group translocators. Proteins of the bacterial phosphoenolpyruvate:sugar phosphotransferase system are included in this category. The product of the reaction, derived from extracellular sugar, is a cytoplasmic sugarphosphate. Hence, these proteins are enzymes rather

than transporters because the latter do not chemically destabilize their substrates (Christensen, 1975). These proteins are included here for completeness because they do catalyze nutrient uptake into the cytosol although in a chemically altered form. 5. Decarboxylation-driven active transporters. Bacterial transport proteins that drive solute (e.g., sodium ion) uptake or extrusion by decarboxylation of a cytoplasmic carboxylic acid substrate are included in this category. 6. Electron flow-driven active transporters. Proteins that catalyze transport of a solute (e.g., an ion) energized by the flow of electrons from a reduced substrate to an oxidized substrate are included in this category. Threedimensional structural data, available for three of these enzyme complexes (Table 1), greatly help to conceptualize the mechanisms by which these ion pumps catalyze transport. 7. Light-driven active transporters. Transport proteins that utilize light energy to drive transport of a solute (e.g., an ion) are included within this category.

TABLE 8.2

Families of Transport Proteins (Proposed Abbreviations in Parentheses)

1. Channel-type transporters 1.1 The Major Intrinsic Protein (MIP) Family 1.2 The Epithelial Na + Channel (ENaC) Family 1.3 The Large Conductance Mechano-sensitive Ion Channel (MscL) Family 1.4 ATP-gated Cation Channel (ACC) Family 1.5 The Voltage-sensitive Ion Channel (VIC) Family 1.6 The Ligand-gated Ion Channel (LIC) Family of Neurotransmitter Receptors 1.7 The Glutamate-gated Ion Channel (GIC) Family of Neurotransmitter Receptors 1.8 The Channel-forming Amphipathic Peptide (CAP) Functional Superfamily 1.9 The Ryanodine-Inositol 1,4,5-triphosphate Receptor Ca2+ Channel (RIR-CaC) Family 1.10 The Chloride Channel (C1C) Family 1.11 The Holin Functional (Holin) Superfamily 1.12 The Channel-forming Colicin (Colicin) Family 1.13 The Channel-forming t~-Endotoxin Insecticidal Crystal Protein (ICP) Family 1.14 The c~-Hemolysin Channel-forming Toxin (c~HL) Family 1.15 The Aerolysin Channel-forming Toxin (Aerolysin) Family 1.16 Animal Inward Rectifier K + Channel (IRK-C) Family 1.17 The Organellar Chloride Channel (O-C1C) Family 1.18 The Channel-forming Colicin V (Colicin V) Family 1.19 The Channel-forming e-toxin (e-toxin) Family 1.20 The Transient Receptor Potential Ca 2+ Channel (TRPCC) Family 2. Carrier-type transporters (uni-, sym-, and antiporters) 2.1 The Major Facilitator Superfamily (MFS) 2.1.1 Sugar Porter (SP) Family 2.1.2 The Drug:H + Antiporter (14 Spanner) (DHA14) Drug Efflux Family 2.1.3 The Drug:H + Antiporter (12 Spanner) (DHA12) Drug Efflux Family 2.1.4 The Organophosphate:Pi Antiporter (OPA) Family 2.1.5 The Oligosaccharide:H + symporter (OHS) Family 2.1.6 The Metabolite:H + Symporter (MHS) Family 2.1.7 The Fucose:H + Symporter (FHS) Family 2.1.8 The Nitrate/Nitrite Porter (NNP) Family 2.1.9 The Phosphate:H + Symporter (PHS) Family 2.1.10 The Nucleoside:H + Symporter (NHS) Family 2.1.11 The Oxalate:Formate Antiporter (OFA) Family 2.1.12 The Sialate:H + Symporter (SHS) Family 2.1.13 The Monocarboxylate Porter (MCP) Family 2.1.14 The Anion:Cation Symporter (ACS) Family 2.1.15 The Unknown Major Facilitator (UMF) Family 2.1.16 The Aromatic Acid:H + Symporter (AAHS) Family 2.1.17 The Cyanate Transporter (CP) Family 2.1.18 The Polyol Transporter (PP) Family 2.2 The Glycoside-Pentose-Hexuronide (GPH):Cation Symporter Family 2.3 The Amino Acid-Polyamine-Choline (APC) Family 2.4 The Cation Facilitator (CF) Family 2.5 The Zinc (Zn 2+)-IrOn (Fe z+) Porter (ZIP) Family 2.6 The Resistance-Nodulation-Cell Division (RND) Family 2.7 The Small Multidrug Resistance (SMR) Family 2.8 The Gluconate:H + Symporter (GntP) Family 2.9 The L-Rhamnose Transporter (RhaT) Family 2.10 The 2-Keto-3-Deoxygluconate Transporter (KDGT) Family

2.11 2.12 2.13 2.14 2.15 2.16 2.17 2.18 2.19 2.20 2.21 2.22 2.23 2.24 2.25 2.26 2.27 2.28 2.29 2.30 2.31 2.32 2.33 2.34 2.35 2.36 2.37 2.38 2.39 2.40 2.41 2.42 2.43 2.44 2.45 2.46 2.47 2.48 2.49 2.50 2.51 2.52 2.53 2.54 2.55 2.56 2.57 2.58 2.59

The Citrate-Mg2+:H+ (CitM)-Citrate:H + (CitH) Symporter (CitMHS) Family The ATP:ADP Antiporter (AAA) Family The C4-Dicarboxylate Uptake (Dcu) Family The Lactate Transporter (LctT) Family The Betaine/Carnitine/Choline Transporter (BCCT) Family The Telurite-resistance/Dicarboxylate Transporter (TDT) Family The Proton-dependent Oligopeptide Transporter (POT) Family The Amino Acid/Auxin Transporter (AAAT) Family The CaZ+:Cation Antiporter (CaCA) Family The Inorganic Phosphate Transporter (Pit) Family The Solute:Sodium Symporter (SSS) Family The Neurotransmitter:Sodium Symporter (NSS) Family The Dicarboxylate:Cation (Na + or H +) Symporter (DCS) Family The Citrate:Na + Symporter (CSS) Family The Alanine or Glycine:Cation Symporter (AGCS) Family The Branched Chain Amino Acid:Cation Symporter (LIVCS) Family The Glutamate:Na + Symporter (GltS) Family The Bile Acid:Na + Symporter (BASS) Family The Mitochondrial Carrier (MC) Family The Cation-Chloride Cotransporter (CCC) Family The Anion Exchanger (AE) Family The Silicon Transporter (Sit) Family The NhaA Na:H + Antiporter (NhaA) Family The NhaB Na+:H+ Antiporter (NhaB) Family The NhaC Na+:H+ Antiporter (NhaC) Family The Monovalent Cation:Proton Antiporter-1 (CPA1) Family The Monovalent Cation:Proton Antiporter-2 (CPA2) Family The K + Transporter (Trk) Family The Nucleobase:Cation Symporter-1 (NCS1) Family The Nucleobase:Cation Symporter-2 (NCS2) Family The Nucleoside Uptake Porter (NUP) Family The Aromatic Amino Acid Porter (ArAAP) Family The Serine/Threonine Porter (STP) Family The Formate-Nitrite Transporter (FNT) Family The Metal Ion Transporter (MIT) Family The Benzoate:H + Symporter (BenE) Family The Divalent Anion:Na + Symporter (DASS) Family The Reduced Folate Carrier (RFC) Family The Ammonium transporter (Amt) Family The Triose Phosphate Translocator (TPT) Family The Nucleotide-Sugar Transporter (NST) Family The Ni2+-Co 2+ Transporter (NiCoT) Family The Sulfate Porter (SulP) Family The Mitochondrial Tricarboxylate Carrier (MTC) Family The Acetyl-Coenzyme A Transporter (AcCoAT) Family The Tripartite ATP-independent Periplasmic Transporter (TRAP-T) Family The Equilibrative Nucleoside Transporter (ENT) Family The Phosphate:Na + Symporter (PNaS) Family The Arsenical Resistance-3 (ACR3) Family

3. Pyrophosphate bond (ATP, GTP, P2) hydrolysis-driven active transporters 3.1 The ATP-binding Cassette (ABC) Superfamily

(continues)

TABLE 8.2 (Continued) 3.2 3.3 3.4 3.5 3.6 3.7 3.8 3.9 3.10 3.11

The H +- Na+-translocating F-type, V-type, and A-type ATPase (F-ATPase) Superfamily The Cation-translocating P-type ATPase (P-ATPase) Superfamily The Arsenical (Ars) Efflux Family The Type II (General) Secretory Pathway (IISP) Family The Type III (Virulence-related) Secretory Pathway (IIISP) Family The Type IV (Conjugal DNA-Protein Transfer or VirB) Secretory Pathway (IVSP) Family The Mitochondrial Protein Transporter (MPT) Family The Chloroplast Envelope Protein Transporter (CEPT) Family The H+-translocating Vacuolar Pyrophosphatase (H+-PPase) Family The Bacterial Competence-related DNA Transformation Transporter (DNA-T) Family

4. Phosphotransferases 4.1 The PTS Glucose-Glucoside (Glc) Family 4.2 The PTS Fructose-Mannitol (Fru) Family 4.3 The PTS Lactose-Cellobiose (Lac) Family 4.4 The PTS Glucitol (Gut) Family 4.5 The PTS Galactitol (Gat) Family 4.6 The PTS Mannose-Fructose-Sorbose (Man) Family 5. Decarboxylation-driven active transporters 5.1 The Na+-transporting Carboxylic Acid Decarboxylase (NaT-DC) Family 6. Oxidoreduction-driven active transporters 6.1 The Proton-translocating NADH Dehydrogenase (NDH) Family 6.2 The Proton-translocating Transhydrogenase (PTH) Family 6.3 The Proton-translocating Quinol:Cytochrome c Reductase (QCR) Superfamily 6.4 The Proton-translocating Cytochrome Oxidase (COX) Superfamily 6.5 The Na+-translocating NADH:Quinone Dehydrogenase (Na-NDH) Family 7. Light-driven active transporters 7.1 The Ion-translocating Bacteriorhodopsin (BR) Family 7.2 The Proton-translocating Reaction Center (RC) Family 8. Mechanically-driven active transporters 8.1 The H +- or Na+-translocating Bacterial Flagellar Motor (Mot) Family 9. Outer 9.1 9.2 9.3 9.4 9.5 9.6 9.7 9.8 9.9 9.10

membrane porins (fl-structure) The General Bacterial Porin (GBP) Family The Chlamydial Porin (CP) Family The Sugar Porin (SP) Family The Brucella-Rhizobium Porin (BRP) Family The Pseudomonas OprP Porin (POP) Family The OmpA-OprF Porin (OOP) Family The Rhodobacter PorCa Porin (RPP) Family The Mitochondrial and Plastid Porin (MPP) Family The FadL Outer Membrane Protein (FadL) Family The Nucleoside-specific Channel-forming Outer Membrane Porin (Tsx) Family 9.11 The Outer Membrane Fimbrial Usher Porin (FUP) Family

9.12 The Autotransporter (AT) Family 9.13 The Alginate Export Porin (AEP) Family 9.14 The Outer Membrane Receptor (OMR) Family 9.15 The Raffinose Porin (RafY) Family 9.16 The Short Chain Amide and Urea Porin (SAP) Family 9.17 The Outer Membrane Factor (OMF) Family 9.18 The Outer Membrane Auxiliary (OMA) Protein Family 9.19 The Glucose-selective OprB Porin (OprB) Family 10. Methyltransfer-driven active transporters 10.1 The Na+-transporting Methyltetrahydromethanopterin:Coenzyme M Methyltransferase (NaT-MMM) Family 98. Auxiliary transport-related proteins 98.1 The Membrane Fusion Protein (MFP) Family 98.3 The Cytoplasmic Membrane-Periplasmic Auxiliary-1 (MPA1) Protein with Cytoplasmic (C) Domain (MPA1-C or MPA1 + C) Family 98.4 The Cytoplasmic Membrane-Periplasmic Auxiliary-2 (MPA2) Family 98.6 The TonB-ExbB-ExbD/TolA-TolQ-TolR (TonB) Family of Auxiliary Proteins for Energization of Outer Membrane Receptor (OMR)-mediated Active Transport 98.7 The Phosphotransferase System Enzyme I (EI) Family 98.8 The Phosphotransferase System HPr (HPr) Family 98.9 The rBAT (rBAT) Family of Putative Transport Accessory Proteins 98.10 The Slow Voltage-gated K + Channel Accessory Protein (MinK) Family 99. Transporters of unknown classification 99.1 The Polysaccharide Transporter (PST) Family 99.2 The MerTP Mercuric ion (Hg2+) Porter (MerTP) Family 99.3 The MerC Mercuric Ion (Hg2§ Uptake (MerC) Family 99.4 The Nicotinamide Mononucleotide (NMN) Uptake Porter (PnuC) Family 99.5 The K + Uptake Porter (KUP) Family 99.6 The L-Lysine Exporter (LysE) Family 99.7 The Chromate Ion Transporter (CIT) Family 99.8 The Ferrous Iron Uptake (FeoB) Family 99.9 The Low Affinity Fe2+ Transporter (FeT) Family 99.10 The Oxidase-dependent Fe a§ Transporter (OFeT) Family 99.11 The Copper Transporter-1 (Ctrl) Family 99.12 The Copper Transporter-2 (Ctr2) Family 99.13 The Metal Ion (Mn2+ and iron) Transporter (Nramp) Family 99.14 The Cadmium resistance (CadD) Family 99.15 The Putative Amide Transporter (Ami) Family 99.16 The Canalicular Bile Acid transporter (C-BAT) Family 99.17 The Urate Transporter (UAT) Family 99.18 The Peptide Uptake Porter (PUP) Family 99.19 The Mg2+ Transporter-E (MgtE) Family 99.20 The Low-Affinity Cation Transporter (LCT) Family 99.21 The Membrane Targeting and Translocation (Mtt) Family

Proposed Classification System

Two such systems (Table 8.1) have been extensively studied from three-dimensional structural standpoints. 8. Mechanically driven active transporters. Transport proteins are included within this category if they directly drive the movement of a cell, organelle, or other physical structure by allowing the flow of ions (or other solutes) through the membrane down their electrochemical gradients. The Escherichia coli flagellar motor protein complex is considered to be an example of such a process, but F-type ATPases that couple ion movement to diphosphate bond formation or hydrolysis employing a mechanical device are not. These last mentioned systems utilize a mechanical process to couple transport to a chemical reaction ( Junge et al., 1997; Noji et al., 1997), and they are therefore placed in category 3. 9. Outer membrane channel-type facilitators (porins). The proteins of this category exhibit transmembrane/3-strands that form B-barrels through which solutes pass. They are found in the outer membranes of Gram-negative bacteria, mitochondria, and eukaryotic plastids. 10. Methyltransferase-driven active transporters. A single characterized protein, the Na+-transporting methyltetrahydromethanopterin:coenzyme M methyltransferase, currently falls into this category. 98. Auxiliary transport-related proteins. Proteins that function with or are complexed to known transport proteins are included in this category. An example would be a protein that facilitates solute transport across the two membranes of the Gram-negative bacterial cell envelope in a single step driven by the energy source (ATP or the pmf) utilized by a cytoplasmic membrane transporter. Energy coupling and regulatory proteins that do not actually participate in transport represent other examples. In some cases auxiliary proteins are considered to be part of a transport protein complex with which they function, and in such cases no distinct entry in category 98 is provided. Generally, then, a family of proteins is included in category 98 when its peripheral involvement in transport is established but it does not properly fit into a single family of transporters. 99. Transporters of unknown classification. Transport protein families of unknown classification are grouped under this number. These families include at least one member for which a transport function has been established, but either the mode of transport or the energy coupling mechanism is not known. They will be classified elsewhere when the transport process and energy coupling mechanism are characterized. 100. Putative transporters in which no family member is an established transporter. Putative transport protein families are grouped under this number and will either be classified elsewhere when the transport function of a member becomes established or will be elimi-

271

nated from the TC classification system if the proposed transport function is disproven. These families include a member or members for which a transport function has been suggested, but evidence for such a function is not yet compelling. The current index of transport protein families is presented in Table 8.2. There are 148 entries, each of which usually describes a single family or superfamily. Some of these families are large superfamilies with hundreds of currently sequenced members (e.g., the MFS; TC 2.1 and the ABC superfamily; TC 3.1; see Section VII of this chapter). Others are small families with only one or two currently sequenced members. Most families, however, are of intermediate sizes, with between 5 and 100 sequenced members. In a few instances (e.g., the channel-forming amphipathic peptide (CAP) functional superfamily (TC 1.8) and the holin (Holin) functional superfamily (TC 1.11)), the entry includes a number of functionally related families of peptides or proteins. In these cases, insufficient degrees of sequence similarity are observed between members of the different families included within the functional superfamily to establish homology. Table 2 therefore includes well over 150 families of transport systems. All of the families currently included in Table 8.2 will undoubtedly expand with time, and new families will be identified. The availability of new protein sequences will occasionally allow two or more currently recognized families to be placed together under a single TC number. In a few cases, two families are already known for which some evidence is available suggesting that these families are related, e.g., CPA1 and CPA2 families (TC 2.36 and 2.37), as well as the NCS1 and NCS2 families (TC 2.39 and 2.40). This evidence is usually based on (1) limited sequence similarities; (2) common function; and/or (3) similar protein size, topology, and structure. When "missing link" sequences or threedimensional structural data become available so that proteins of two families can be unequivocally grouped together within a single family, the lower TC number will be adopted for all of the family members, and the higher TC number will be abandoned. The complete index (Table 8.2) and representative tables describing some of the families to be discussed in this chapter will be presented below. The complete classification system is available on our web site (http://www-biology. ucsd.edu/--~msaier/transport/titlepage.html). It will be updated continuously as new information becomes available. Anyone noting errors or incomplete listings is encouraged to contact the chapter author providing the missing information and references by e-mail, fax, phone, or mail. In almost all cases, members of a transporter family utilize a single energy coupling mechanism, thus justify-

272

8. Classification of Transport Proteins

ing the use of transport mode and energy coupling mechanism as the primary basis for classification. However, a few exceptions have been noted. First, the arsenite (Ars) (TC 3.4) transporter of E. coli consists of two proteins, ArsA and ArsB. ArsB is an integral membrane protein which presumably provides the transport pathway for the extrusion of arsenite and antimonite (Silver et al., 1993). ArsA is an ATPase that energizes ArsBmediated transport. However, when ArsB alone is present, as in the case of the arsenical resistance pump of Staphylococcus aureus, transport is driven by the proton motive force (pmf) (Br6er et al., 1993). The presence or absence of the ArsA protein thus determines the mode of energy coupling. The use of such alternative sources of free energy to drive transport is not common and has been documented in only a few instances. When such an effect is reported, we shall classify the transporter in accordance with the more complicated energy coupling mechanism (in this case, as an ATP-driven primary active transporter (Class 3), rather than as a secondary carrier (Class 2)). This decision is based in part on the observation that the physiologically relevant form of energy coupling is usually the more complicated one. The potentially different energy coupling mechanisms will be described in the table characterizing that family (see our web site). Examples of secondary carrier families in which different transport modes have been reported include the mitochondrial carrier (MC) family (TC 2.29) and the triose phosphate translocator (TPT) family (TC 2.50). Proteins of both families are apparently restricted to eukaryotic organelles. Members of both families normally catalyze carrier-mediated substrate:substrate antiport and are therefore classified as secondary carriers. However, treatment of MC family members with chemical reagents, such as N-ethyl maleimide o r C a 2+ (Brustovetsky & Klingenberg, 1994, 1996; Dierks et al., 1990a,b; Jezek et al., 1994) or imposition of a large membrane potential (A~) across a membrane into which a TPT member has been incorporated (Schultz et al., 1993; Schwarz et aL, 1994; Wallmeier et al., 1992), has been reported to convert these antiport-catalyzing carriers into anion-selective channels capable of functioning by uniport. Another secondary carrier that may be capable of exhibiting channel-like properties is the KefC protein of E. coli (Booth et al., 1996). A few other examples have been documented in the scientific literature. The possibility of "tunneling" or "slippage" of ions and other solutes through carriers with little or no conformational change has been discussed (Fr61ich, 1988), but there is little evidence to show that conformational changes are not involved. Irrespective of these ambiguities, the more complicated carrier-type mechanism, which appears to be relevant under most physiological

conditions, provides the basis for classifying these proteins (i.e., as Class 2 carriers rather than Class 1 channels).

V. REPRESENTATIVE EXAMPLES OF CLASSIFIED FAMILIES All families are presented in our web site in tabular form with the format as shown in Tables 8.3-8.5. Other examples are published (Saier, 1998). Table 3 describes a family of bacterial toxins, the a-hemolysin channelforming toxin (aHL) family (TC 1.14). The description of this family is presented within the table, together with relevant references, and representative members of the family are tabulated with the accession numbers that render their sequences readily available. Table 8.4 presents a family of secondary carriers, the cation facilitator (CF) family (TC 2.4). Members of this family are specific for heavy metals (Co 2+, Cd 2+, and Zn2+). These proteins are derived from both bacteria and eukaryotes and can function with either inwardly or outwardly directed polarity. A third example is provided by the H +- or Na +translocating F-type, V-type, and A-type ATPase (FATPase) superfamilies (TC 3.2; Table 8.5). These transporters are composed of multiple components, and they use ATP hydrolysis to drive protons or Na + against their gradients (see Chapter 5). Some of them (but not others) can function reversibly. Some of these protein complexes (the F-type ATPases) are essential for oxidative and photosynthetic phosphorylation in bacteria, archaea, mitochondria, and chloroplasts. These proteins are believed to utilize a mechanical mechanism to couple ion transport to ATP hydrolysis or synthesis (Junge et al., 1997; Noji et al., 1997). Vl. CROSS-CLASSIFICATION OF TRANSPORT PROTEINS In addition to the primary classification system presented in Table 8.2, where classification is based on transporter type and family, we have cross-classified all transporters according to (1) substrate specificity and (2) database accession number. Cross-referencing allows one to readily identify a novel member of a family merely by associating its sequence with that of an established member of the database. All established members have recorded accession numbers in at least one of the three primary databases, GenBank (gb), SwissProt (sp), or protein information resource (pir). Thus, association of the novel protein sequence with an established transporter of known family, for example by screening the

273

Cross-Classification of Transport Proteins TABLE 8.3 TC number

The a-Hemolysin Channel-Forming Toxin (aHL) Family (TC Category 1.14) a

Name

Source

Example

a-Hemolysin of Staphylococcus aureus (spP09616)

1.14.1.1

a-Hemolysin

Gram-positive bacteria

1.14.2.1

Hemolysin II

Gram-positive bacteria

Hemolysin II of Bacillus cereus (gbU94743)

1.14.2.2

/3-Toxin

Gram-positive bacteria

/3-Toxin of Clostridium perfringens (pirI40856)

1.14.3.1

Leucocidin chain S

Gram-positive bacteria

Leucocidin chain S of Staphylococcus aureus (pirS32211)

1.14.4.1

Leucocidin chain F

Gram-positive bacteria

Leucocidin chain F of Staphylococcus aureus (pirS32212)

aThe a-hemolysin (aHL) of the human pathogen Staphylococcus aureus is secreted as a 33-kDa monomer. This monomeric species associates with animal cell membranes to form a 232-kDa homoheptameric transmembrane pore that promotes cell lysis by allowing transport of ions, water, and small solutes. The three-dimensional structure of aHL has been solved by X-ray crystallography to 1.9 ~, resolution (Song et al., 1997). The aHL forms a solvent-filled channel with a length of 100 ,~,, that runs along the sevenfold axis of the protein and ranges from 14 to 46 A in diameter. The transmembrane domain of the mushroom-shaped heptamer is the lower portion of the mushroom, consisting of a 14strand antiparallel/3-barrel to which each protomer contributes two B-strands, each 65 ,~ long. The interior of the B-barrel is primarily hydrophilic, and the exterior has a hydrophobic belt 28 ,~ wide. The aHL family consists of 13 currently sequenced proteins all of which are pore-forming toxins. Most are from Staphylococcal species, but one is from Bacillus cereus and one is from Clostridium perfringens. Thus, all are produced by low G + C Gram-positive bacteria. The S. aureus protein monomers are 308-326 residues in length while the B. cereus protein is of 412 residues and the C. perfringens monomer is of 336 residues. The phylogenetic tree for the aHL family reveals four clusters. The Staphylococcus a-hemolysin for which the three-dimensional structure is available comprises one branch, the B. cereus and C. perfringens proteins comprise a second, and all other members of the family fall into the remaining two clusters. The generalized transport catalyzed by these pore-forming toxins is: small molecules (in) ~ small molecules (out). References: Cooney, J., Kienle, Z., Foster, T. J., and O'Toole, P. W. (1993). The gamma-hemolysin locus of Staphylococcus aureus comprises three linked genes, two of which are identical to the genes for the F and S components of leukocidin. Infect. Immun. 61, 768-771. Supersac, G., Prevost, G., and Piemont, Y. (1993). Sequencing of leucocidin R from Staphylococcus aureus p83 suggests that staphylococcal leucocidins and gamma-hemolysin are members of a single, two-component family of toxins. Infect. Immun. 61, 580-587. Steinporsdottir, V., Frithriksdottir, V., Gunnarsson, E., and Andresson, O. S. (1995). Expression and purification of Clostridium perfringens betatoxin glutathione S-transferase fusion protein. FEMS Microbiol. Lett. 130, 273-278. Song, L., Hobaugh, M. R., Shustak, C., Cheley, S., Bayley, H., and Gouaux, J. E. (1996). Structure of staphylococcal a-hemolysin, a heptameric transmembrane pore. Science 274, 1859-1866.

databases using the various BLAST programs (Basic, Gapped, or PSI Blast) or the FASTA program (see Saier, 1994 for descriptions and primary references as

TABLE 8.4 TC number

well as Altschul et aL, 1997), allows rapid determination of its family identification, its probable mechanism of action, and often its substrate specificity.

The Cation Facilitator (CF) Family (TC Category 2.4) a

Name

Source

Example

2.4.1.1

Cd 2+, Z n 2+, C o 2+ efflux transporter

Bacteria

CzcD of Alcalgenes eutrophus (spP13512)

2.4.2.1

Mitochondrial Co 2+ uptake transporter

Yeast

Cotl of Saccharomyces cerevisiae (spP32798)

2.4.2.2

Mitochondrial Z n 2+, Cd 2+ uptake transporter

Yeast

ZnrP of Saccharomyces cerevisiae (spP20107)

2.4.2.3

Plasma membrane Z n 2+ efflux transporter

Animals

Zntl of Rattus norvegicus (gbU17133)

2.4.3.1

Vesicular Z n 2+ uptake transporter

Animals

Znt2 of Rattus norvegicus (gbU50927)

aThe CF family is a small but ubiquitous family, members of which are found in prokaryotes and eukaryotes. They transport heavy metals including cobalt, cadmium, and zinc. All members of the CF family possess six putative transmembrane spanners. These proteins exhibit an unusual degree of sequence divergence and size variation (300-750 residues). Eukaryotic proteins exhibit differences in cell localization and polarity. Thus, some catalyze heavy metal uptake while others catalyze efflux, and some are found in plasma membranes while others are in organellar membranes. Prokaryotic and eukaryotic proteins cluster separately. The mechanisms of free energy coupling are not well understood. The generalized transport for CF family members is: Me 2+ (in or out) ~ Me 2+ (out or in). References: Nies, D. H., and Silver, S. (1995). Ion efflux systems involved in bacterial metal resistances. J. Industr. Microbiol. 14, 186-199. Paulsen, I. T., and Saier, M. H., Jr. (1997). A novel family of ubiquitous heavy metal ion transport proteins. J. Membr. Biol. 156, 99-103.

274 TABLE 8.5

8. classification of Transport Proteins The H +- or Na+-Translocating F-Type, V-Type, and A-Type ATPase (F-ATPase) Superfamily (TC Category 3.2) a

TC number

Name

Source(s)

Example

3.2.1.1

H+-Translocating F-type ATPase

Bacteria; eukaryotic mitochondria and chloroplast

F-Type ATPase of E. coli (gbJ01594)

3.2.1.2

Na+-Translocating F-type ATPase

Bacteria

F-type ATPase of Propionigenium modestum (gbX58461)

3.2.2.1

H+-Translocating V-type ATPase

Bacteria; eukaryotes

V-Type ATPase of Thermus thermophilus (gbD63799)

3.2.2.2

Na+-Translocating V-type ATPase

Bacteria

V-Type ATPase of Enterococcus hirae NtpLMNOPQ (gbX76913)

3.2.3.1

H+-Translocating A-type ATPase

Archaea

A-Type ATPase of Methanosarcina mazeii AhaABCDEFG (gbU47274)

aF-Type ATPases are found in eukaryotic mitochondria and chloroplasts as well as in bacteria. V-Type ATPases are found in vacuoles of eukaryotes and in bacteria. A-Type ATPases are found in archaea. All such systems are multisubunit complexes with at least three dissimilar subunits embedded as a complex in the membrane (F0, a : b : c = 1:2:---12) and (usually) at least five dissimilar subunits attached to F0 (F1, c~:/3: y: 6: e = 3:3 : 1 : 1 : 1 for F-type ATPases). The y-subunit of the F1 component is believed to rotate relative to most of the other subunits in response to either ATP hydrolysis by F1 or proton transport through F0. Therefore H + transport and ATP synthesis may be coupled mechanically. The F1 portion of the bovine heart mitochondrial F-type ATPase has been solved to 2.8 A resolution. All eukaryotic F-type ATPases pump 3-4 H + out of mitochondria or into thylakoids of chloroplasts per ATP hydrolyzed. Bacterial F-type ATPases pump 3-4 H + and/or Na + (depending on the system) out of the cell per ATP hydrolyzed. These enzymes also operate in the opposite direction, synthesizing ATP when protons flow through the "ATP synthase" down the proton electrochemical gradient (the "proton motive force" or pmf). V-Type ATPases may pump 2-3 H + per ATP hydrolyzed. Phylogenetic clustering of the integral membrane constituents of F-type ATPases generally corresponds to the phylogenies of the organisms of origin, and consequently the systems in different organisms are probably orthologs. The a-subunit of F0 (1 copy per complex) spans the membrane six times. The b-subunits (2 copies per complex; heterodimeric in plant chloroplasts and blue-green bacteria) span the membrane once, and the c-subunits (called DCCD-binding lipoproteins; about 12 copies per complex) span the membrane two times. The c-subunits of Ftype ATPases are homologs to the c-subunits of V-type and A-type ATPases. Several of the integral membrane subunits in these protein complexes may be homologous to each other, but homology can be demonstrated only for the c-subunits based on sequence analyses alone. The generalized transport/reaction for F-type, V-type, and A-type ATPases is: nH+ (in) [or nNa + (in)] + ATP ~ nH+ (out) [or nNa + (out)] + ADP + Pi References: Abrahams, J. P., Leslie, A. G. W., Lutter, R., and Walker, J. E. (1994). Structure at 2.8 ,~ resolution of F1-ATPase from bovine heart mitochondria. Nature 370, 621-628. Solioz, M., and Davies, K. (1994). Operon of vacuolar-type Na+-ATPase of Enterococcus hirae. J. Biol. Chem. 269, 9453-9459. Takase, K., Kakinuma, S., Yamato, I., Konishi, K., Igarashi, K., and Kakinuma, Y. (1994). Sequencing and characterization of the ntp gene cluster for vacuolar-type Na+-translocating ATPase of Enterococcus hirae. J. Biol. Chem. 269, 11037-11044. Blair, A., Ngo, L., Park, J., Paulsen, I. T., and Saier, M. H., Jr. (1996). Phylogenetic analyses of the homologous transmembrane channel-forming proteins of the FoF1-ATPases of bacteria, chloroplasts and mitochondria. Microbiology 142, 17-32. Deckers-Hebestreit, G., and Altendorf, K. (1996). The FoFx-type ATP synthases of bacteria: Structure and function of the F0 complex. Annu. Rev. Microbiol. 50, 791-824. Goldsmith, E. J. (1996). Allosteric enzymes as models for chemomechanical energy transducing assemblies. FASEB J. 10, 702-708. Nakamoto, R. K. (1996). Mechanisms of active transport in the FoFx ATP synthase. J. Membr. Biol. 151, 101-111. Yamada, H., Moriyama, Y., Maeda, M., and Futai, M. (1996). Transmembrane topology of Escherichia coli H+-ATPase (ATP synthase) subunit a. FEBS Lett. 390, 34-38. Noji, H., Yasuda, R., Yoshida, M., and Kinosita, K., Jr. (1997). Direct observation of the rotation of F1-ATPase. Nature 386, 299-302. Rahlfs, S., and Mtiller, V. (1997). Sequence of subunit c of the Na+-translocating FIF0 ATPase of Acetobacterium woodii: Proposal for determinants of Na + specificity as revealed by sequence comparisions. FEBS Lett. 404, 269-271. Weber, J., and Senior, A. E. (1997). Catalytic mechanism of FI-ATPase. Biochim. Biophys. Acta 1319, 19-58.

T a b l e 8.6 p r o v i d e s i n f o r m a t i o n a b o u t t h e d i s t r i b u t i o n of t r a n s p o r t e r t y p e s b a s e d o n s u b s t r a t e specificity. C h a n n e l p r o t e i n s ( T C c a t e g o r y 1) a n d o u t e r m e m b r a n e p o r i n s ( T C c a t e g o r y 9) a r e s o m e t i m e s s e l e c t i v e w i t h r e s p e c t t o t h e s u b s t r a t e t r a n s p o r t e d , b u t t h e y a r e also s o m e t i m e s

n o n s e l e c t i v e o r o n l y m o d e r a t e l y specific for a class of i n o r g a n i c m o l e c u l e s . A few c h a n n e l p r o t e i n s a n d o u t e r m e m b r a n e p o r i n s e x h i b i t specificity for w a t e r o r s m a l l o r g a n i c m o l e c u l e s , a n d s e v e r a l f u n c t i o n in t h e t r a n s p o r t of p r o t e i n s .

275

Cross-Classification of Transport Proteins TABLE 8.6

TABLE 8.7

Distribution of Transporter Types Based on Substrate Specificity a

Class of compounds I. Inorganic molecules II. Carbon sources III. Amino acids and derivatives IV. Nucleic acid precursors V. Vitamins and cofactors VI. Drugs VII. Macromolecules

Channels & porins

1~

2*

c a r r ie r s

carriers

Compound transported

30

14

32

76

4 0

1 1

23 20

28 21

2

0

10

12

1

1

6

8

1 23

1 5

3 2

Substrate Specificities of ABC Transporters

5 30

aBold print indicates the major type(s) in each category of transporters.

Members of families of ATP-dependent primary carriers (TC category 3) usually exhibit a restricted range of substrate specificities, but members of one family, the ABC superfamily (TC 3.1; see below), can transport almost any molecule, large or small. Finally, members of families of secondary carriers (TC category 2) may function in the transport of inorganic molecules or proteins, but a large majority of them exhibit specificity for a particular class of organic molecules. Different types of transport proteins, included within different TC categories, thus in general exhibit specificities for different classes of molecules.

Vll. THE TWO LARGEST SUPERFAMILIES OF TRANSPORTERS. THE MF AND ABC SUPERFAMILIES Two superfamilies account for about 50% of all solute transporters known to be encoded within the genomes of several bacteria and archaea (Paulsen et al., 1998) as well as of the yeast Saccharomyces cerevisiae (unpublished results). One of them is the major facilitator superfamily (MFS; TC 2.1) of secondary carriers, and the other is the ATP-binding cassette (ABC; TC 3.1) superfamily of primary carriers. As the MFS has recently been described in detail (Pao et al., 1998), it will not be considered here further. The ABC superfamily is not only one of the two families with the largest representation in living organisms, it is also the most diverse with respect to substrate specificity and polarity. Table 7 summarizes the types of compounds known to be transported by ABC-type protein complexes and also relates the pumping polarities of these transporters. ABC protein complexes can

Polarity

Inorganic cations

In or out

Inorganic anions

In

Sugars

In

Organic anions

In

Amino acids and derivatives

In

Amines and polyamines

In

Peptides

In or out

Vitamin B12

In

Fea+-siderophores

In

Drugs

Out

Glutathione conjugates

Out

Heme

Out

Siderophores

Out

Steroids

Out

Pigments

Out

Polysaccharides

Out

Proteins

Out

Lipids

Out (flipping)

function with either inwardly directed polarity or outwardly directed polarity, but no one system has been described that couples energy to transport without exhibiting strict polarity. ABC protein complexes, unlike all other types, can transport almost any biological substance, regardless of size and nature. Thus, small molecules (inorganic ions; sugars; amino acids, etc.) as well as molecules of intermediate size (Vitamin B12, peptides, siderophores, etc.) and macromolecules (polysaccharides, proteins, phospholipids) are all substrates of various ABC protein complexes. We presume that this unusual flexibility reflects some basic and distinctive architectural feature of this transporter type (Paulsen et al., 1998; Saier, 1998). This situation contrasts with several other transport protein families, the members of which exhibit strict specificity for a single compound, e.g., the CaCA family (TC 2.19), all functionally characterized members of which are specific for Ca 2+, or the Amt family (TC 2.49), all characterized members of which are specific for N H 4 § VIII. MACROMOLECULAR TRANSPORT PROTEINS IN BACTERIA Macromolecules can be exported from cells and imported into cells by a variety of mechanisms (Table 8.8). Transport may occur by a uniport or channel-type

276

8. Classification of Transport Proteins TABLE 8.8

Macromolecular Transport Proteins in Bacteria

Category

Substrate

Family

VIIA

Complex carbohydrates

VIIB

Proteins

ATP-binding cassette Polysaccharidetransporter Polysaccharide transporter Resistance-nodulation-cell division Holin functional Diphtherin Toxin ATP-binding cassette Type II secretory pathway Type III secretory pathway Type IV secretory pathway Autotransporter Outer membrane receptor Type IV secretory pathway ATP-binding cassette

VIIC

Nucleicacids

VIID

Lipids

Abbreviation

ABC PST PST RND

TC number

3.1 99.1 99.1 2.6

Polarity

Membrane(s)

ATP ATP pmf pmf

Export Export Export Export

CM or both CM or both CM Both

Export Import Export Export Export

CM CM Both CM Both

Holin DT ABC IISP IIISP

3.1 3.5 3.6

None None ATP ATP and pmf ATP

IVSP

3.7

ATP

Export

Both

AT OMR (TonB) IVSP

9.12 9.14 3.7

None pmf ATP

Export Import Export

OM OM Both

ABC

3.1

ATP

Export (flipping)

CM

mechanism, by a proton motive force (pmf)-dependent mechanism, or by an A T P hydrolysis-dependent mechanism. While export systems for complex carbohydrates, nucleic acids, lipids, and proteins are known, only some of the protein export systems apparently function by channel-type mechanisms. Most functionally characterized macromolecular export systems depend upon A T P hydrolysis to drive transport. The mechanistic details of many of these processes have yet to be elucidated.

IX. CONCLUSIONS A N D PERSPECTIVES Molecular archeological studies of transport proteins have led us to propose a novel classification system based on both function and phylogeny. In this chapter, we have briefly described this system of classification and provided reference sources for its exploitation. Many of the benefits and novel information resulting from our studies have been summarized elsewhere (e.g., Pao et al., 1998; Paulsen et al., 1998; Saier, 1998; Saier et al., 1999). The analyses have led to a better understanding of fundamental principles in biology. For example, we have provided evidence that transporter families arose repeatedly and independently, at different times in evolutionary history, following different routes. In

1.11

Energy source

spite of similar apparent topological features, we believe that several transporter types must exhibit distinctive architectural features that confer differing capacities for functional diversification. G e n o m e analyses have helped to reveal the numbers of transport protein families and the breadth of their functionalities. These analyses have led us to devise a novel classification system based both on functionality and phylogeny. We believe that a functional-phylogenetic basis for transporter classification provides the most rational approach to protein classification available so far. It also provides the most information concerning the evolution, structures, and functions of any class of proteins. Note added in proof" Many new transporter families have been identified, and several of the described families have been redefined or expanded since this summary article went to press. The updated descriptions can be found in our web site. Acknowledgments I am grateful to Milda Simonaitis and Mary Beth Hiller for their assistance in the preparation of this manuscript. I also acknowledge useful discussions with Drs. Arnost Kotyk and Lon Van Winkle. Work in the laboratory of MHS was supported by USPHS Grants 5RO1 AI21702 from the National Institutes of Allergy and Infectious Diseases and 9RO1 GM55434 from the National Institute of General Medical Sciences, as well as by the M. H. Saier, Sr. memorial research fund.

C

H

A

P

T

[91

E

R

Regulation of Plasma Membrane Transport*

membrane vesicles where the membrane potential can be varied by the addition of membrane-permeable anions such as SCN- or by the addition of valinomycin in experiments where the intravesicular and extravesicular concentrations of K + are unequal. In some systems, e.g., the electrogenic H+-linked peptide transporter in kidney brush border membrane vescles (Daniel et aL, 1991) and the uptake of lysine into human placental brush border membrane vesicles (Eleno et aL, 1994), a detailed analysis of the effects of changes in membrane potential on the kinetics of substrate uptake into vesicles has been performed. Electrophysiological techniques have also been useful for the study of electrogenic transport in the case of transporters heterologously expressed in Xenopus oocytes (Kavanaugh, 1993). The short-term regulation of liver metabolism by glucagon represents one important example of a situation where membrane potential-related alterations in transport kinetics are of physiological importance. Unlike many other cells (Section X,C of Chapter 3), the liver cell membrane has a low permeability to K + ions, and the consensus membrane potential of - 3 5 mV is far removed from the K + diffusion potential of - 7 0 mV and is mainly set by the Na+/K + ATPase. One major effect of glucagon is to cause a rapid hyperpolarisation of the liver cell membrane to about - 5 0 mV (see Moule and McGivan, 1990a). This may be due in part to induction of K + permeability due to opening of specific K + channels and in part to activation of the Na+/H + exchanger by cAMP, presumably via a phosphorylation event (see Section III) with a consequent increase in intracellular Na + concentration and stimulation of the electrogenic Na+/K + ATPase (Moule and McGivan, 1990b). This membrane hyperpolarization stimulates the rate of transport into the liver cell of a number of important substrates. In isolated hepatocytes the initial rate of electrogenic Na+-alanine cotransport via System A (the known

1. INTRODUCTION In this chapter we offer a description of the major factors through which the activity of transporters can be modulated. General principles are described in terms of regulation of activity due to changes in transport driving force, modifications of endogenous transporters, or increase in the number of transporter molecules in the membrane. For each category selected examples have been considered in detail. As far as possible, the physiological significance of these changes is also discussed. Finally, the role of disorders of transport regulation in two pathological conditions of wide interest is reviewed.

II. REGULATION OF TRANSPORT BY CHANGES IN DRIVING FORCE: THE ROLE OF PLASMA MEMBRANE POTENTIAL If a substrate is transported across the cell membrane by an electrogenic transport process it is clear that the steady state accumulation ratio of that substrate is influenced by the magnitude of the cell membrane potential. However, the kinetic parameters of an electrogenic transport processes may also be affected by membrane potential. A theoretical analysis of the effect of membrane potential on electrogenic Na+-linked substrate cotransport (Geck and Heinz, 1976) showed that a change in membrane potential could affect either the Vmax of the transport or the Km for the substrate or both Km and Vmaxdepending on the assumptions made about the molecular mechanism involved. Many investigations demonstrating the electrogenic nature of Na +- or H +linked cotransport or uniport have been carried out on tOvidio Bussolati a, Gian Gazzola a, and John McGivan b. aIstituto di Patologia Generale, Universit~ degli Studi di Parma, Parma, Italy and bDepartment of Biochemistry, School of Medical Sciences, Bristol, United Kingdom.

277

278

9. Regulation of Plasma Membrane Transport

amino acid transport systems are summarized in Table 4.2 of Chapter 4 and Table 10.1 of Chapter 10) was stimulated by glucagon in a biphasic manner (Edmondson et al., 1985). The initial phase, which occurred within minutes, was independent of protein synthesis and was attributed to membrane hyperpolarization; this was followed by a slow protein synthesis-dependent induction of transport. Alanine is a major substrate for gluconeogenesis in liver, and perfusion experiments with isolated hepatocytes have shown that alanine transport at physiological alanine concentrations is a major control step for gluconeogenesis (Sips et al., 1980). Epidermal growth factor (EGF) also stimulated both alanine transport and gluconeogenesis in hepatocytes. In this case only a single rapid phase of stimulation was observed and this was independent of protein synthesis. Again this stimulation was attributed to membrane hyperpolarization via EGF stimulation of the Na+/H + exchanger and subsequent activation of the electrogenic Na+/K + ATPase (Moule and McGivan, 1987). The bile salt taurocholate is transported into hepatocytes by an electrogenic mechanism involving the uptake of 2 Na + ions/taurocholate. It was shown (Edmonson et al., 1985) that glucagon stimulates taurocholate uptake in hepatocytes via membrane hyperpolarization; this may be important in the physiological stimulation by glucagon of bile acid secretion. Recently, electrogenic Na+-uridine cotransport into hepatocytes has been shown to be rapidly stimulated by glucagon by a protein synthesis-independent mechanism, which is also very probably related to membrane hyperpolarization (Gomez-Angelats et al., 1996). It is not clear whether stimulation of transport by membrane hyperpolarization is of physiological importance in other situations. However, membrane depolarization may be important in pathological situations. For example, the rate of electrogenic L-arginine transport into pulmonary artery epithelial cells is decreased by hypoxia and this decrease can be correlated with hypoxia-induced membrane depolarisation (Zharikov et al., 1997).

I!i. REGULATION OF THE ACTIVITY OF EXISTING TRANSPORTERS THROUGH MODIFICATIONS OF TRANSPORTER MOLECULES A. Covalent Modifications of Channels and Carriers Almost all membrane proteins are modified after their synthesis. Carriers and channels do not elude this general rule. In many cases changes such as removal of leader sequences, glycosylation, and acylation are

integral parts of the synthetic pathway and represent irreversible steps of the processing mechanism that ensures correct folding and sorting of the protein. In other cases, chemical modifications represent convenient means to modulate protein function and constitute true short-term regulatory mechanisms. A common change of this type is protein phosphorylation; the formation of an ester bond between a phosphate group, donated by ATP, and the hydroxyl group of serine, threonine, or tyrosine side-chains. Convenience of phosphorylation, catalyzed by a vast array of protein kinases, derives from its easy reversibility through the activity of a number of protein phosphatases, several of which have been characterized in the past few years. Many regulatory pathways involve the activation of kinases and phosphatases; in some cases this may lead to modulation of the activity of transporter proteins. We have selected some examples in which the correlation between protein modification and functional change is particularly clear. However, other transporters are known to be substrates of phosphorylating/dephosphorylating enzymes.

1. Cystic Fibrosis Conductance Regulator Cystic fibrosis (CF) is the most common autosomic recessive disease in Caucasians and still provokes severe morbidity and premature mortality in most patients, notwithstanding the impressive therapeutic progresses obtained in the past few years. Cystic fibrosis is due to mutations of the CF gene, which encodes a single polypeptide chain of 1480 amino acids called CFTR (cystic fibrosis transmembrane regulator). The proposed structure of CFTR (Fig. 9.1) comprises two membrane-spanning domains (each formed by six transmembrane segments) and a highly complex cytoplasmic region, formed by a regulatory R-domain and by two highly homologous nucleotide-binding domains (NBD1 and NBD2). This structure allows CFTR to be ascribed to the superfamily of traffic ATPases (or ABC transporters, see Section VII of Chapter 8). These are prokaryotic and eukaryotic transport proteins with a homologous nucleotide-binding domain, such as many bacterial periplasmic permeases, the yeast STE6 gene product, the mammalian P-glycoprotein, and heterodimeric transporters TAP1/TAP2 (Higgins, 1992). Several studies had described the appearance of C1channels in cells transfected with CF cDNA. However, conclusive evidence that CFTR is itself a C1- channel was obtained by demonstrating that incorporation of purified CFTR into planar lipid bilayers produced the characteristic cAMP-activatable C1- conductance, which is deficient in CF cells (Bear et al., 1992; Tilly et al., 1992). The cAMP effect on C1- conductance was reproduced in artificial models, consisting of reconstitu-

Modifications of Transporter Molecules

FIGURE 9.1

ted CFTR and purified catalytic domain of PKA in the presence of ATP. From these and other studies a model was derived in which PKA phosphorylates the Rdomain of CFTR, whose open state is then gated by ATP binding to NBDs with subsequent hydrolysis of the nucleotide (Welsh et al., 1995). A direct interaction between PKA and CFTR (Berger et al., 1991; Tabcharani et aL, 1991) is consistent with the presence of a number of phosphorylation consensus sequences in the protein. Twelve of these sites (all serines except for one threonine) are located in the Rdomain, while one is close to NBD1 (Gadsby and Nairn, 1994). However, the analysis of tryptic phosphorylated peptides has demonstrated, in both in vitro and in vivo models, that only some of these sites, all constituted by serine residues, appear to be involved effectively in PKA stimulation of C1- fluxes (Cheng et aL, 1991; Picciotto et al., 1992). Site-directed mutagenesis studies have led to the identification of the five major phosphorylation sites $660, $700, $737, $795, and $813 (Cheng et al., 1991; Rich et aL, 1993; Chang et aL, 1993), whose overall removal produces a marked decrease in the open state probability of CFTR channel. However, even the all-mutated CFTR is still cAMP sensitive (although with a very low efficiency), thus suggesting the existence of nonclassic phosphorylation sites in CFTR. Addition of negative charge(s) to the R-domain through phosphorylation is thought to be an important mechanism in opening the channel by changing electrostatic interactions among the various domains of CFTR. Evidence for this mechanism has been obtained by replacing serine residues with aspartate (or glutamate). When more than

279

Structure of CFTR.

five serine residues are changed the CFTR channel opens even without cAMP addition (Rich et aL, 1993; Chang et aL, 1993). These results are compatible with the model of the "occluding ball" proposed for Shaker K + channels (Zagotta et aL, 1990; Hoshi et aL, 1990; and see Chapter 7) in which phosphorylation of the R-domain would change its conformation and relieve the occlusion of the CFTR channel exerted by the unphosphorylated form. Consistently, it has been demonstrated that PKAdependent phosphorylation induces a significant conformational change in the CFTR R-domain (Dulhanty and Riordan, 1994). However, a recent paper from Welsh's group claims that the addition of a phosphorylated Rdomain stimulates CFTR channel activity, while the unphosphorylated domain is without any inhibitory effect (Winter and Welsh, 1997), thus casting some doubts on the attractive "occluding ball" model. It should also be taken into account that not all the phosphorylation sites present on the R-domain have a stimulatory effect on CFTR (Wilkinson et aL, 1997) and that other kinases have been implicated in CFTR regulation (Picciotto et aL, 1992; Berger et aL, 1993; Dechecchi et aL, 1992, 1993; Sears et aL, 1995; Hwang et al., 1997; Tien et aL, 1994; French et al., 1995). It is also now generally accepted that CFTR dephosphorylation can be of great functional importance. Evidence for a regulatory effect of phosphatase activity on CFTR was obtained by studying the stimulating action of phosphatase inhibitors on CFTR-mediated C1- fluxes (Becq et aL, 1994, 1996). It is known that the CFTR channel can be rapidly inactivated upon membrane exci-

280

9. Regulation of Plasma Membrane Transport

sion for patch-clamp studies; this phenomenon has been attributed to the activity of membrane-associated phosphatases (Tabcharani et al., 1991; Hwang et al., 1993; Fischer and Machen, 1996). Direct intervention of the ubiquitous phoshatase PP2A in the dephosphorylation of CFTR has been demonstrated in excised membrane patches (Berger et al., 1993; Haws etal., 1994). In this case, since phosphatase is activated after cAMP stimulation, the effect of phosphatase activity is inhibitory and the dephosphorylated CFTR channel returns to basal activity. Also, more recently, protein phosphatase 2C has been involved in CFTR regulation (Travis et al., 1997). Thus, through the combined effects of phosphorylation and dephosphorylation of the various sites of Rdomain, an exquisitely fine tuning of CFTR channel activity appears possible. 2. Na+/H + Exchangers

Na+/H + exchangers (NHE) catalyze an electroneutral and strictly coupled antiport of Na + and H +. The various isoforms (Table 9.1) are involved in pH homeostasis, cell volume regulation, Na + absorption, transepithelial weak acid transport, and H + extrusion by epithelia. All the members of the family have two distinct parts (Figure 9.2; see Ts6 et al., 1993 and Yun et al., 1995, for reviews): an N-terminus region with 10-12 membrane-spanning domains, which on the extracellular side presents the site that binds amiloride and its analogs, and a large C-terminal cytoplasmic portion. The most important regulatory feature of NHEs consists in the marked sensitivity of the antiport to changes in the intracellular pH: at pHi values higher than 7.2 the antiport has no activity, while the more pHi shifts toward acid values, the more the activity of the antiport is stimulated via an increase in the affinity for H +. Using a mutant lacking the C-terminus (Wakabayashi et al., 1992), it was possible to demonstrate that the N-terminal region is sufficient to mediate the antiport. However, the dependence upon pHi was markedly shifted, al-

TABLE 9.1

though the allosteric activation by internal H + was still present. Therefore, it is generally accepted that NHEs have two functionally distinct domains. The N-terminal transmembrane domain performs the antiport and is endowed with an allosteric site for intracellular H + (the so called "pH sensor" or "H + modifier site"), while the C-terminus has a regulatory function. In agreement with this model, two regions (aa 636-656 and aa 567-635 of NHE1) of the cytoplasmic C-terminal domain have been identified (Wakabayashi et al., 1997), which are essential for pH sensitivity. However, this unifying structure hides marked differences in tissue distribution and physiological functions among the isoforms. From a structural point of view, isoform heterogeneity is due mainly to the poor conservation of the cytoplasmic domain (homology <30% among the various isoforms), while the N-terminus exhibits 50-60% homology. The activity of NHEs is sensitive to many hormones, growth factors, and tumor promoters (for review see Wakabayashi et al., 1997). For instance, the stimulation of antiport activity by serum, observed in many cell types, can be ascribed to an activation of one or more isoforms: the widely expressed, housekeeping NHE1 (Bianchini and Pouyssegur, 1996) and the specialized, tissue-specific isoforms NHE2 and NHE3. The mechanisms involved, however, appear to be distinct for the various isoforms. Studies by Pouyssegur and co-workers (Wakabayashi et al., 1992) have indeed demonstrated that serum increases the affinity for H + of NHE1. On the contrary, serum stimulation of NHE2 and NHE3 is mediated through changes in transport Vma x (Levine et al., 1993). Divergent kinetic effects are also seen in the case of treatment with phorbol esters, the classic nonphysiologic activators of PKC. These compounds stimulate NHE1 by increasing its affinity for H + (Paris and Pouyssegur, 1984), while they modulate the activity of NHE2 and NHE3 through changes in transport Vmax (Levine et al., 1993; Ts6 et aL, 1993b), increasing that of NHE2 and, conversely, decreasing that of NHE3.

The NHE (Na+/H + Exchanger) Family NHE4

NHE3

NHE2

NHE1

Amino acids a

820

813

831

717

Expression

Ubiquitous

Kidney, intestine, stomach

Kidney, intestine, stomach

Stomach

Activity increased by

Growth factors, phorbols, hypertonic incubation

Growth factors, phorbols

Growth factors

Hypertonic incubation

Activity decreased by

ATP depletion

ATP depletion

ATP depletion, hypertonic incubation

ATP depletion

Calmodulin binding site(s)

Present

Present

Present

Present

aData from rat NHEs.

Modifications of Transporter Molecules

FIGURE 9.2

281

Structure of NHE transporters.

The involvement of kinases in these effects has been suggested by several results: 1. All the isoforms of NHE have multiple consensus sites for kinases in the cytoplasmic C-terminus. 2. Upon stimulation with various growth factors NHE1 is phosphorylated on several serine residues in the C-terminal domain (Sardet et aL, 1990). 3. In NHE1 the major phosphorylation sites are located between amino acids 636 and 815. The deletion of this portion of the transporter markedly affects (but does not suppress completely) the serum stimulatory effect (Wakabayashi et al., 1993). What is the kinase involved in the regulation of NHEs? Structural features of NHE transporters and the rapid change in activity obtained under certain conditions (less than 1 min after serum exposure) would suggest a direct interaction of the carrier molecule with the kinase. The precise identification of the kinase involved, however, is still elusive. The problem is complicated

because several distinct kinases can influence in specific ways the activity of one or the other member of NHE family (Levine et aL, 1993; Kandasamy et aL, 1995). Due to the above-mentioned effect of phorbols, a candidate for NHE regulation is PKC. NHE has indeed a classic PKC consensus site, but this site appears not necessary for the phorbol effect (Wakabayashi et al., 1994). A good correlation between antiport activation and protein phosphorylation has been demonstrated for fibroblastic cells (Sardet et aL, 1990). However, phorbols increase NHE transport activity but lower phosphorylation of NHE1 in several other cell models (Rao et al., 1992; Sweeney et aL, 1995; Siczkowski and Ng, 1996). On the basis of these data, it is more probable that PKC activation modulates NHE activity by an indirect mechanism. Also PKA appears to influence the activity of NHE transporters, but this effect is only seen in some members of the family, because not all NHEs are endowed with PKA consensus sequences (Borgese et aL, 1992). In particular, PKA inhibits NHE3 activity and, in this

282

9. Regulation of Plasma Membrane Transport

case, the effect seems to be due, at least in part, to a direct phosphorylation of the antiport protein by PKA (Kurashima et al., 1997). Discrepancies between phosphorylation pattern and activity data have suggested that PKC and PKA, rather than interact directly with NHE1, would change the status of an intermediate kinase, which would be the direct NHE-regulator. MAP kinases, once proposed for the role (Sardet et aL, 1990), have now been implicated in growth factor activation of NHEs through indirect mechanisms rather than through direct phosphorylation of the antiporter (Aharanowitz and Granot, 1996; Bianchini et aL, 1997). The best candidates as NHE regulators are kinases structurally associated with the transporter, the so-called NHE1 kinases (Sardet et al., 1991), whose characterization is now in progress (Phan et al., 1997; Takahashi et aL, 1997). On the other hand, several data indicate that the change in the activity of NHE induced by growth factors is not mediated only by the phosphorylation of the transporter. The mutagenized transporter deprived of the major phosphorylation domain (Wakabayashi et al., 1992) still maintains the capability to respond, although with a reduced efficiency, to protein kinase activation, thus pointing to additional mechanisms for activity change. These data have led to the proposal that, a least in part, NHE response to kinase activation depends upon yet-unidentified ancillary regulatory proteins, which after phosphorylation would bind to critical regions of the NHE C-terminal domain (Wakabayashi et al., 1994; Wakabayashi et al., 1997). In this regard, it is well known that calmodulin, in its phosphorylated form, can interact with specific domains of the NHE molecule endowed with high regulatory activity (Wakabayashi et al., 1994; Bertrand et aL, 1994; see Section III,C below). 3. E A A T Transporters for Anionic Amino Acids

EAATs (excitatory amino acid transporters) are a family of high-affinity transporters for the anionic amino acids glutamate and aspartate (see for reviews Hediger et al., 1995; Malandro and Kilberg, 1996; and Chapter 6 of this volume). They have been cloned and characterized mainly in tissues and cell types of neural origin. In CNS EAATs are responsible for the termination of glutamatergic neurotransmission, a role of pivotal importance in CNS due to the neurotoxic effect of an overstimulation of NMDA glutamate receptors. The expression of human EAAT1 and EAAT2 (see Chapter 6 for the use of these denominations) is restricted to glial cells. Knockout mice for these glial transporters develop severe excitotoxicity, progressive paralysis, and neurodegeneration (Rothstein et aL, 1996). In contrast,

at least one member of the family, the neuronal type EAAT3, has been found expressed also in extraneural tissues of both mesenchymal and epithelial origin. Interestingly, knockout mice for EAAT3 develop only minor signs of neurotoxicity while presenting dicarboxylic aminoaciduria (Peghini et al., 1997). All EAATs operate a highly energized transport of their amino acid substrate, coupled with fluxes of sodium, potassium, and H + or OH- (discussed in detail in Section III of Chapter 6). Moreover, EAAT1 is endowed with a constitutive cation-selective pore (Vanderberg et aL, 1997), while two other members of the family, EAAT4 and EAAT5, combine transport activity with the properties of a ligand-gated C1- channel (Fairman et al., 1994; Arriza et al., 1997). The involvement of PKC activation in the regulation of transport activity of EAATs has been suggested for several years on the basis of data obtained both in cultured human fibroblasts (Franchi-Gazzola et al., 1991) and in rat glioma C6 cells (Casado et al., 1992) following three classical lines of evidence: 1. Stimulation of transport activity by phorbol esters and physiological activators of PKC, such as diacylglycerols. The effect is rapid, sustained, and independent of protein synthesis and membrane potential. 2. Prevention of the stimulation if PKC is downregulated by prolonged exposure to high concentrations of phorbols. 3. Suppression of the stimulation by relatively specific PKC inhibitors that poorly affect basal transport activity. A structural feature of E A A T carriers is the presence of one or more PKC consensus motifs. Moreover, the time course of stimulation by natural PKC activators can be very short (less than 1 min). Therefore, it is conceivable that phorbol stimulation of anionic amino acid transport could be attributed to a direct phosphorylation of the carrier by PKC. In agreement with this hypothesis, Casado and co-workers (Casado et al., 1993) have demonstrated an increased phosphorylation of the carrier in C6 glioma cells, predominantly at serine residues, after exposure to PKC. Moreover, the abundance of a specific PKC isozyme, PKCs, on the plasma membrane varies consistently with changes in aspartate transport (Franchi-Gazzola et al., 1996), thus further suggesting a direct interaction between the kinase and the carrier. While human fibroblasts seem to express an EAAT3 transporter, data about C6 cells are somewhat conflicting. Notwithstanding their glial origin, it has been indeed suggested that C6 cells express the neuronal-type EAAT3 (Palos et al., 1996; Dowd et aL, 1996). Other

283

Modifications of Transporter Molecules

authors have attributed glutamate transport in these cells also to an EAAT2-type carrier (Casado et al., 1993). Moreover, these authors have conclusively shown that this carrier is up-regulated by PKC activation when expressed heterotopically in HeLa cells and that substitution of a serine residue abolishes PKC stimulation without affecting basal transport activity. Another member of the family, EAAT1, exhibits a different behavior. The activity of this transporter, which has three consensus sequences for PKC, is indeed markedly lowered when cells are exposed to PKC activators (Conradt and Stoffel, 1997). Site-directed mutagenesis of all the three PKC consensus sites did not affect either transport inhibition or phosphorylation of EAAT1 (as judged by analysis of immunoprecipitated protein). Therefore, the effect is possibly due to either the involvement of nonconventional sites or the intervention of an intermediate kinase. It is likely that PKC sensitivity is a characteristic of the E A A T family, although the functional effects of phosphorylation vary depending upon the particular transporter (and, possibly, the particular PKC isoform) involved. Due to the extreme importance of E A A transporters for the control of neurotransmission, PKC modulation of their activity can have important functional consequences. Interestingly, PKC effects on the transport of neuromediators are by no means restricted to EAATs. Indeed, several changes in the activity of transporters of other neuromediators have been observed upon PKC activation, such as decrease in glycine transporter GLYTlb activity (Sato et al., 1995), increase of GABA influx through transporter redistribution (Corey et aL, 1994), and inhibition of the dopamine transporter (Kitayama et aL, 1994; Huff et aL, 1997; Zhu et al., 1997; Vaughan et al., 1997; Zhang et al., 1997). B. Oligomerization Many transporters and channels are composed of distinct subunits. This is the case of some vertebrate P-type ATPases (such as gastric H+K+ATPase and Na§247 which consist of a heterodimer composed of one large c~-subunit and of a small, heavily glycosylated/3-subunit. However, several experimental data point to more complex forms of subunit organization in these transporters. With radiation inactivation analysis, for example, the H+K+ATPase transport cycle has been suggested to involve targets of size roughly double the value predicted for a single heterodimer (Saccomani et aL, 1981; Rabon et al., 1988). Dimerization of the same transporter has been also suggested with chemical cross-linking studies (Rabon et al., 1990) and directly demonstrated after membrane solubilization (Shin and Sachs, 1996). Different levels of oligo-

meric organization can produce profound changes in the transport process. For instance, the structure needed for proton transport may change from dimeric to tetrameric, depending on the ATP concentration (Morri et aL, 1996) (see also Section II,C,4 of Chapter 5). Other transporters, such as NHEs, are single-chain polypeptides, but are hypothesized to work as oligomers. It is generally accepted that all the isoforms of the family consist of a single type of subunit (see above); moreover, transfection with a single cDNA ~s able to reconstitute Na§ + exchange activity in deficient cells (Levine et aL, 1993; Orlowski, 1993). However, multiple lines of evidence such as data on transitions in the apparent molecular size with thiol reagents (Fliegel et aL, 1993), results of radiation inactivation analysis (Beliveau et al., 1988), and kinetic evidence for positive cooperativity exhibited by pre-steady state kinetics in brush-border vesicles (Otsu et al., 1989, 1992, 1993), suggest that NHEs work as oligomeric complexes. In particular the studies by Otsu and co-workers demonstrated that the dependence of presteady state phase amplitude on the extravesicular Na § concentration exhibits a transition from sigmoidal to simple hyperbolic behavior upon the increase of the intravesicular pH from 5.7 to 7.7. This result suggests that the level of occupation of the H+-modifier site (that is, the intracellular pH), located in the N-terminal domain (see above), can modulate the interaction between distinct molecules of NHEs in the membrane. Although preliminary, these data indicate that the characteristics of the transport process can be changed through modifications of the oligomeric organization of carrier molecules. Functional oligomerization has been suggested for an increasing number of other transporters, mostly based on target size obtained by radiation inactivation analysis of transport processes compared with the molecular weight of the corresponding cloned transporters (see, for instance, Beliveau et aL, 1990; McCormick et al., 1991; Milner et al., 1994; Jette et al., 1996; Haugeto et al., 1996; Goldstein et al., 1996; Jette et aL, 1997). C. Allosteric Interactions In some instances allosteric regulation is of pivotal importance for the function of the transporter. For instance, through the allosteric interaction of protons with the "modifier site" located at the cytoplasmic region of the N-terminal portion, Na+/H + exchange through NHE transporters is directly linked to intracellular pH (see above). Moreover, this intrinsic property of NHE transporter is employed by other regulatory devices: the changes in the activity of NHE transporters caused by kinase activation are often produced by modulation of

284

9. Regulation of Plasma Membrane Transport

this modifier site, which is very close to the high-affinity binding site for calmodulin (Wakabayashi et aL, 1994; Bertrand et al., 1994). It is thought that, under normal conditions (low cytoplasmic Ca2+), the calmodulinbinding site interacts with the modifier site, lowering the affinity of the carrier for cytoplasmic H +. Conversely, upon cell stimulation, the increase in Ca 2+ activates kinases, thus leading to calmodulin phosphorylation and binding, with subsequent removal of modifier site inhibition, decrease in transporter Km for H +, and exchange stimulation (Wakabayashi et al., 1997). As far as CFTR is concerned, the presence of sufficient concentrations of ATP at the intracellular side of the membrane is indeed an absolute requirement for the already-discussed activation of CFTR channel by PKA (Anderson et al., 1991). It is, however, not yet clear if ATP interaction with one or both the nucleotide binding domains of the protein precedes the obligatory hydrolysis of the nucleotide. The view that emerges from these studies is that what we consider the basal level of activity of a transporter is often the combined result of the activity of subtle regulatory mechanisms. Moreover, it is also evident that covalent modifications, allosteric effects, and supramolecular interactions can effectively and intimately cooperate in setting the activity of a given transporter.

IV. REGULATION OF TRANSPORT BY CHANGES IN THE REPERTOIRE OF TRANSPORT PROTEINS IN THE PLASMA MEMBRANE In mammalian cells the membrane content of transporter proteins is not constant under all conditions. Transport activity can be stimulated by mechanisms which involve de n o v o protein synthesis and the putative insertion of additional transport proteins into the cell membrane. Such induction of transport activity may occur in response to hormones or to various types of cell stress such as substrate deprivation or exposure to a hypertonic medium. This has been particularly extensively studied in the case of amino acid transport system A, which serves as a good example for this type of transporter regulation. A. Hormonal Regulation of Transport The ubiquitous transporter system A is a Na +dependent, highly regulated mechanism for the uptake of small aliphatic amino acids in mammalian cells. System A activity in hepatocytes is induced by a number of hormones, in particular glucagon, catecholamines,

and glucocorticoids (see Kilberg and Haussinger, 1992 for a review). Stimulation by glucagon has been most studied since this is of the most obvious physiological significance. Protein synthesis-dependent induction of System A in hepatocytes by glucagon occurs after a lag period of at least 1 hr and is sensitive to cycloheximide and to tunicamycin, an inhibitor of protein glycosylation. Following glucagon treatment of rats, enhanced system A transport activity can be recovered in subsequently isolated liver plasma membrane vesicles (Schenerman and Kilberg, 1986) and also in Golgi vesicles (Caraiappa and Kilberg, 1990) and this is consistent with the postulate that glucagon-induced transport involves increased synthesis and insertion into the membrane of system A transporter molecules. It has been shown that in hepatocytes the decay of glucagon-enhanced transport after glucagon withdrawal is prevented by inhibitors of either RNA or protein synthesis, and it has been suggested that an additional newly synthesised protein is necessary for the degradation of induced activity (see e.g. Kilberg et al., 1985). Some properties of this putative protein have been deduced. However, since system A has not yet been cloned, the antibodies and cDNA probes necessary to demonstrate directly that induction of activity involves the synthesis of new transporter molecules are not available and the nature of any accessory proteins is still obscure. Like glucagon, insulin causes protein synthesis induction of system A activity after a lag period, but little is known of the molecular mechanisms involved. The same is true of induction by glucocorticoids and by catecholamines. Hormonal regulation of system A activity has been described also in extrahepatic tissues. B. Induction of Substrate Transport by Long-Term Substrate Deprivation Unlike induction of transport in bacterial and intestinal cells by nutrients (e.g., Van Winkle, 1999), a variety of mammalian cell types induce transport activity in response to substrate deprivation. This was originally thought to represent a defense mechanism whereby cells contrive to maintain adequate intracellular substrate concentrations until normal levels of substrate supply are resumed. However, the meaning of this response could be more complex and involve cell response to cell volume changes (see Section IV,C). In general, transport activity increases following substrate withdrawal after a lag period of some hours. Induction of transport activity depends on RNA and protein synthesis and is reversed over a period of time by readdition of substrate. The best-studied examples are the transport of small aliphatic amino acids via system A in a number of cell types and Na*-dependent gluta-

Transport Proteins in the Plasma Membrane

mate and phosphate transport in renal cells, but the phenomenon is widespread. In general it is not clear that protein synthesis-dependent induction of transport activity necessarily involves de n o v o synthesis of the transport proteins themselves, and more complex mechanisms may be involved.

1. Induction of System A Adaptive regulation of system A activity in response to amino acid deprivation was first characterized in fibroblasts, but was subsequently observed in many different cell types and is normally regarded as derepression of this transport system (see e.g., McGivan and PastorAnglada (1994) for a review). In hepatocytes, induction of transport is inhibited by cycloheximide and by tunicamycin and is prevented or reversed by the presence of single amino acids in the culture medium. Although single substrates of system A prevent the increase of activity, some compounds that are not substrates will also prevent the induction, and compounds that are not substrates of system A can effectively reverse system A induction. The increase in system A activity in hepatocytes is normally about two- to fourfold. In confluent cultures of renal epithelial cells, system A activity is undetectable in normal culture medium but reaches a considerable activity some hours after amino acid deprivation. That induction of system A activity may involve synthesis of a regulatory protein either in addition to or instead of induction of the system A protein itself has been suggested by experiments on cell mutants. The alar4 mutant of the CHO-K1 cell line exhibits an abnormally high constitutive activity that is not further increased on amino acid deprivation. In this case there is evidence that a mutation in a regulatory gene rather than in the system A gene is responsible for this behaviour (Moffett and Englesberg, 1984), but such an activator gene has not yet been characterized. A further complexity is added by the observation that repression of previously induced system A by added amino acids is dependent on protein synthesis in some cell types but not in others. The mechanism by which cells detect a deficiency of amino acids is also unclear. Asparagine synthase is induced by amino acid deprivation in FAO cells by a mechanism which exhibits a wide amino acid specificity. In this case, a regulatory element in the promoter region of the asparagine synthase gene has been identified as being involved in the induction process (Guerrini et al., 1991). However, this element is not present in the promoter of calreticulin, which is also induced by amino acid deprivation (Heal and McGivan, 1998), indicating that more than one mechanism may exist. Again, resolu-

285

tion of the mechanism of system A induction awaits the cloning of the system A protein

2. Glutamate Transport in Renal Epithelial Cells Renal epithelial cells express high activities of highaffinity Na+-dependent glutamate transport. In the bovine renal cell line NBL-1 this transport activity is catalyzed by an EAAT3-type transporter (see Section III,A,3 above); EAAT1 and EAAT2 transporters are absent in these cells. When NBL-1 cells are starved of amino acids, Na+-dependent glutamate transport activity increases after a lag period of some hours. As in the case of system A, this effect is blocked by cycloheximide and tunicamycin. Induction of transport is prevented by substrates of the glutamate transporter and by other substrates that can be metabolized to produce glutamate. Experimental conditions that specifically deplete the cells of glutamate cause induction of transport after a delay of some hours, suggesting that a fall in the intracellular glutamate level is responsible for the triggering of the process (Nicholson and McGivan, 1996). Induction of glutamate transport by substrate deprivation does not appear to involve the synthesis of new carriers since mRNA levels and the amount of immunoreactive protein do not increase. Since de n o v o synthesis of a glycoprotein is implicated, it has been suggested that glutamate deprivation induces the synthesis of a novel protein that activates the EAAT3 protein in the membrane (Nicholson and McGivan, 1996). However, induction of a so-far-uncharacterized isoform of the glutamate transport family is not ruled out.

3. Phosphate Transport in Renal Cells A physiologically important example of adaptive regulation of transport is that of phosphate transport in renal brush border membranes. Feeding animals a diet low in phosphate leads to an increase in renal phosphate reabsorption, and this can be attributed to an increase in the activity of renal Na+-phosphate cotransport (see Murer et al. (1996) for review). This phenomenon can be observed also in renal cell lines of proximal tubule origin cultured for prolonged periods in a phosphatedepleted medium. Two different types of renal Na § phosphate cotransporter have been cloned. These are known as type I and type II transporters and have widely different molecular weights. Only the type II transporter, when expressed in X e n o p u s oocytes, shows properties consistent with those of phosphate transport in renal membranes. Studies using specific antibodies and cDNA probes have shown that on feeding rats a lowphosphate diet there is a measurable increase in the rate of brush border membrane transport and immuno-

286

9. Regulation of Plasma Membrane Transport

reactive type II phosphate transport after 2 hr with nondetectable changes in mRNA levels, mRNA levels subsequently increased and there was a further increase in phosphate transport activity and transporter protein. After adaptation of rats to a low phosphate diet for several days, feeding of a normal-phosphate diet caused a marked decrease in both transport activity and transporter protein content within 2-4 hr, but the mRNA level remained high (Levi et al., 1994). It was concluded that the adaptive regulation of renal phosphate transport involved two components; first a rapid increase in membrane protein in the absence of an mRNA increase and, second, a slower response, which may represent changes in the rate of transcription or of mRNA stability. The detailed mechanisms underlying these events have not yet been elucidated. C. Induction of Transport by Exposure of Cells to Hyperosmotic M e d i u m

1. Transport of Compatible Osmolytes in Kidney Cells in the kidney medulla are exposed to hypertonic conditions during the production of concentrated urine. Such cells respond by increasing the Na+-linked uptake of compounds such as myo-inositol, betaine, and taurine. These compounds are known as compatible osmolytes since they are relatively inert, do not interfere with cell function, and may exert a protective effect on protein conformation; the accumulation of these solutes increases the intracellular osmotic pressure and the consequent influx of water helps to restore the cell volume. This situation is mimicked in a variey of cultured cells, particularly those derived from the kidney medulla such as MDCK cells but also in cells of glial, neuronal, and mesangial origin. Extensive studies particularly on MDCK cells have provided much information about the mechanisms involved in this increase in transport activity (see Kwon and Handler 1995, for a review). A number of transport proteins catalyzing the Na § dependent accumulation of osmolytes have been cloned from MDCK cells. These include a myoinositol transporter (SMIT), which is distantly related to the Na § glucose cotransporters, and a transporter for betaine (BGT-1), which belongs to a neurotransmitter gene family. Incubation of MDCK cells in hypertonic medium leads to a protein synthesis-dependent increase in transport, which is observed after a lag period of several hours and is preceded by a large increase in mRNA levels for SMIT and BGT-1. Increased protein synthesis has been shown to be due to increased gene transcription. In the case of the BGT-1 gene an element in the promoter region termed TonE has been shown to be involved in the osmotic response. Recent work has con-

centrated on the stress kinase pathways that may be activated by hypertonic exposure and may be involved in transporter induction. It has been reported that although the MAP kinase pathway is activated by hyperosmolarity, this pathway is not involved in transporter protein induction (Kwon et aL, 1995). A recent report has indicated that the activation of the p38 pathway is a necessary step in this process (Sheikh-Hamad et aL, 1998).

2. Transport of Amino Acids Amino acid transport activity in mammalian cells is induced in response to hypertonic exposure as a means of counteracting cell shrinkage and this has been extensively characterized in human fibroblasts (Dall'Asta et al., 1994). In particular, system A activity is induced in a wide variety of cell types including fibroblasts, CHO cells, renal epithelial and mesangial cells, thymocytes, and vascular smooth muscle cells (see McGivan and Pastor-Anglada 1994, for a review). Although amino acid deprivation and hyperosmotic stress both cause cell shrinkage, the induction of system A by these stress factors appears to involve two separate mechanisms, at least in some cases. For example, in the CHO mutant ala r which is insensitive to amino acid deprivation (see Section IV,B,1), system A activity is still induced by hypertonic stress. In NBL-1 cells induction by hypertonic stress does not occur unless system A has been previously derepressed by amino acid deprivation. Although the mechanism of system A induction by hypertonic stress has not been determined, there are indications that this may be more complex than an increase in the rate of transcription of the system A gene. In NBL-1 cells system A induction by hypertonic stress is blocked by the microtubule-disrupting reagent colcemide but not by tunicamycin, while system A induction, in response to amino acid deprivation, is blocked by tunicamycin but not by colcemide (Ruiz-Montasell et al., 1996). On the basis of these and other findings it has been suggested that in CHO and NBL-1 cells stimulation of system A activity by hypertonicity may be due to the synthesis of an osmotically regulated activator protein. In other cell types, such as cultured human fibroblasts, adaptive regulation and hypertonic stimulation of system A activity appear to be more closely related (Dall'Asta et al., 1996). In these cells changes in the osmolality of incubation medium interfere with the development of adaptive regulation and the increase in transport activity is completely blocked if medium osmolality is kept at values lower than 250 mosm/kg. The interpretation of all these results may be complicated by the absolute requirement to avoid secondary

287

Derangements in Transport Regulation

effects of the experimental conditions adopted on the gradient of sodium electrochemical potential, which constitutes the energy employed by the transport process mediated by system A. High-affinity glutamate transport via EAAT3 is also induced by hypertonic stress in renal epithelial cells but not in other cell types (such as CHO cells, hepatoma cells, fibroblasts, and endothelial cells). In the case of renal cells the increase in transport rate is accompanied by an increase in immunoreactive protein in the cell membrane and is preceded by a large increase in the abundance of EAAT3 mRNA, suggesting that a mechanism similar to the induction of myoinositol and betaine transport may be involved (Ferrer-Martinez et al., 1995).

V. COORDINATED REGULATION OF TRANSPORT SYSTEMS The regulation of cell volume is a function of vital importance for all living cells since the presence of impermeant polyions in the intracellular compartment poses a severe requirement for osmocompensatory devices. The classic "pump-and-leak" mechanism (Ussing, 1960; Tosteson and Hoffman, 1960) aimed to explain this basic situation. In addition, volume regulation contributes to cell survival under conditions of variable extracellular osmolality, such as those encountered by unicellular organisms and by certain cell types of higher organisms (e.g., kidney medulla). On the other hand, in multicellular organisms, where powerful specialized organ mechanisms (e.g., regulation of water intake and excretion) exist that maintain the extracellular osmolality fairly constant, potential dangers for cell volume homeostasis derive both from disorders of those mechanisms (e.g., hyperhydration and hypohydration states) and from changes in the cell content of osmolytes (e.g., under isotonic conditions such as in postabsorptive states for intestine and liver cells). Therefore, it is not surprising that different transport mechanisms are involved in cell volume regulation in different cell models. Several reviews have recently covered the field of cell volume regulation in several types of mammalian cells (see, for instance, Hoffmann, 1997; Brugnara, 1997; Hoffmann and Dunham, 1995). In particular, many channels and transporters co-operate in regulatory volume increase (RVI) and regulatory volume decrease (RVD), the integrated mechanisms that restore cell volume after, respectively, hypertonic shrinkage and hypotonic swelling (Table 9.2). The molecular basis for this coordination is still elusive. For example, the TonE element involved in transcriptional activation of BGT-1 expression promoted in MDCK cells by hypertonic treatment (see above) has not been

identified in the other genes of osmosensitive transporters for organic solutes. Membrane transport also undergoes significant modifications during the cell cycle. As far as mammalian cells are concerned, it has been known for several years that the progression of the G1 phase is associated with an increase in the activity of some transporters for ions and organic solutes, such as Na+K+ATPase, NHE transporters, Na§ - cotransport, and sodiumdependent transport of neutral amino acids, mostly attributable to the activity of system A (see Wakabayashi et al., 1997; Panet and Arian, 1991; McGivan and PastorAnglada, 1994, and the references cited therein). Since all those transporters are involved in cell volume regulation, an attempt to integrate these changes in terms of a cycle-associated increase in cell size has been made (Bussolati et aL, 1996). According to this model (Fig. 9.3) a marked cell swelling, due to an increased cell content of both organic and inorganic osmolytes, occurs during cell cycle progression. The increase in cell content of inorganic ions is mostly contributed by potassium chloride accumulation through Na+/K§ - cotransport. On the other hand, organic osmolytes are constituted by amino acids, with glutamine representing the largest fractional increase. Glutamine is accumulated through system A and can be then exchanged with other neutral amino acids through systems ASC and L, with a mechanism similar to that discussed in Chapter 4 (XI,E). Moreover, glutamine is also hydrolyzed to glutamate, which can also interact with distinct exchange routes. Na§247 activation would yield an enhanced capability to extrude sodium entering through the cotransport systems.

Vl. DERANGEMENTS IN TRANSPORT REGULATION A. Diabetes Mellitus A major metabolic effect of insulin is the stimulation of glucose transport in the so-called insulin-dependent tissues, such as skeletal muscle and adipose tissue. This dependency is characterized by the high expression of GLUT4 transporters (the members of the family are summarized in Table 9.3), whose translocation to the plasma membrane depends on the action of the hormone (James et al., 1988, 1989; Fukumoto et al., 1989; Charron et al., 1989; Birnbaum, 1989). In insulindependent diabetes mellitus (IDDM) pancreatic B-cells are destroyed by an autoimmune reaction and plasma insulin levels fall to very low values. Under these conditions glucose transport of insulin-dependent tissues is markedly inhibited and is rescued upon administration

288

9. Regulation of Plasma Membrane Transport TABLE 9.2

Examples of Membrane Transporters and Channels Activated in Cell Volume Regulatory Mechanisms

Transporter/channel

Representative cell models

Experimental condition a

Na+/K+ATPase

Various

RVI, RVD "Isotonic" swelling

Na+/K+/C1 cotransport

Endothelial cells Ehrlich c e l l s Erythrocytes Lymphocytes Lymphocytes Fibroblasts Muscle Kidneymedulla Kidneymedulla

RVI "Isotonic" swelling RVI RVI RVI RVI "Isotonic" swelling RVI RVI

Various

RVD

K+ channels (many types)

Various

Channels for organic osmolytes

Various

RVD RVI RVD

K+/C1 cotransport Na+/H+ antiport C1-/HCO3- antiport System A (cotransport of Na+ and neutral amino acids) SMIT (Na+/inositol cotransport) BGT-1 (cotransport of Na+ and betaine) C1- channels (many types)

Effects of transport activation

Loss of sodium and chloride (direct osmotic effect) Establishment of ion gradients necessary for channel and cotransport activity Increase in cell ion content Increase in cell ion content Increase in cell ion content Increase in cell ion content Increase in cell content of organic compatible osmolytes Increase in cell content of organic compatible osmolytes Increase in cell content of organic compatible osmolytes Loss of chloride Depolarization Loss of potassium Hyperpolarization Loss of organic, compatible osmolytes

aRVD, regulatory volume decrease (compensatory shrinkage after cell swelling in hypotonic solutions); RVI, Regulatory volume increase (compensatory swelling after cell shrinkage in hypertonic solutions); "isotonic" swelling, increase in cell volume under isotonic conditions (e.g., cell-cycle-associated increase in cell size).

of insulin (Berger et al., 1989). Since glucose influx is thought to be rate limiting for the rate of glucose utilization in peripheral tissues (Ziel et al., 1988), inhibition of glucose influx in I D D M significantly contributes to the decrease in peripheral glucose disposal, which is a hallmark of this form of diabetes. However, glucose consumption by insulin-dependent tissues is impaired also in the other, more common, form of diabetes, N I D D M (non-insulin-dependent diabetes mellitus). This is an effect of insulin resistance, which is a peculiar feature of the N I D D M syndrome, together with defective insulin secretion by pancreatic/~-cells. How much does glucose transport alteration contribute to insulin resistance of peripheral tissues in N I D D M ? G L U T 4 expression is significantly lower in the adipose tissue of N I D D M subjects (Garvey et aL, 1989, 1991; Sinha et aL, 1991). However, the presence of a similar defect in skeletal muscle cells is at best uncertain (Pedersen et aL, 1992; Eriksson et al., 1992). On the other hand, the G L U T 4 gene does not appear to be mutated in a significant portion of N I D D M patients (Kusari et al., 1991; Choi et aL, 1991). The role of glucose transport in the pathogenesis of diabetes has been reassessed in recent years employing transgenic mice. Several strains have been constructed that overexpress either GLUT1 (Marshall et aL, 1993) or G L U T 4 (Liu et al., 1993; Shepherd et aL, 1993). Both

types of transgenic animals exhibit increased peripheral glucose disposal and fasting hypoglycemia. However, GLUTl-overexpressing animals are insulin resistant, since they exhibit no increase in glucose consumption upon insulin treatment (Gulve et al., 1994; Buse et al., 1996). On the contrary, transgenic mice that overexpress the insulin-responsive transporter GLUT4, either in fat cells or in muscle, are insulin sensitive and exhibit an increased glucose disposal in response to the hormone (Deems et al., 1994; Hansen et al., 1995; Tsao et al., 1996; Charron et al., 1997). Moreover, restoration of G L U T 4 expression in muscle tissue of GLUT4-null mice, which are insulin resistant, rescues whole-body insulin action on glucose peripheral disposal (Tsao et al., 1997) and G L U T 4 overexpression improves glycemic control in a rodent model of N I D D M (Gibbs et al., 1995). Also, in a model of overt I D D M high-dose streptozotocininduced diabetes, overexpression of G L U T 4 is able to ameliorate glucose metabolism significantly (Tozzo et al., 1997). These data point to glucose transport as an important rate limiting step for the metabolism of the sugar in peripheral tissues and demonstrate the role of G L U T 4 as a mediator of the hypoglycemic effect of insulin. However, the situation in diabetic humans appears more complex. For example, the improvement of glucose disposal observed upon sulfonylurea therapy in N I D D M

FIGURE 9.3

A model for increase in cell size during the cell cycle. See text for details.

290

9. Regulation of Plasma Membrane Transport TABLE 9.3 The GLUT Familya GLUT1

Preferential substrates

Glucose

Affinity for substrates Typical expression

High (--~1 mM) Ubiquitous

Activity increased by

Glucose Hypoxia

GLUT2

Glucose Fructose Low (--~10 mM) Pancreas (/3 cells) Liver Glucose

GLUT3

GLUT4

GLUT5

GLUT7 b

Glucose

Glucose

Fructose

Glucose

High Neurons Blastocyst, Trophoblast Glucose

High Skeletal muscle Adiposetissue

Low Intestine Spermatozoa

? Liver

Insulin

Fructose

9

aGLUT6 is a pseudogene. b GLUT7 is localized in the endoplasmic reticulum and extrudes glucose produced by glucose-6-phosphatase into the cytoplasm.

obese subjects is not dependent on an increase in GLUT4 expression in skeletal muscle (Vestergaard et al., 1995). As discussed above, defective insulin secretion may also be present in NIDDM. Under physiological conditions, a major stimulus for triggering insulin secretion is a rise in blood glucose level. Since this mechanism is less efficient in NIDDM, many investigations have concerned the nature of glucose sensor of pancreatic/3cells and its efficiency in NIDDM. The results of these studies (reviewed by Effat et al., 1994) suggest that the primary glucose sensor is glucokinase, although a minor role is also played by GLUT2, the glucose transporter predominantly expressed by/~-cells. In contrast to the other GLUTs, GLUT2 has a high Km (15-20 mM) and, therefore, glucose influx in el-cells is proportional to blood glucose level over a wide range of hyperglycemic values. GLUT2 expression is sensitive to blood glucose levels, with prolonged hypoglycemia leading to the disappearance of the transporter and, conversely, hyperglycemia stimulating its expression (Chen et al., 1990). The expression of GLUT2 is severely down-regulated in animal models of N I D D M (Johnson et al., 1990; Orci et al., 1990; Thorens et al., 1990), although these changes appear to be secondary to the diabetic environment rather than primary defects (Thorens et al., 1992; Ohneda et al., 1993). More recently, it has been found that homozygous, but not heterozygous, GLUT2deficient mice have an abnormal glucose-stimulated insulin secretion (Guillam et al., 1997). Moreover, in a subpopulation of N I D D M subjects, affected by an earlyonset variant of the disease (MODY1), a nonsense mutation in the transcription factor HNF4a has been identified. This trancription factor controls the expression of GLUT2 and of several enzymes involved in glucose metabolism (Stoffel and Duncan, 1997). In summary, defective GLUT4 regulation contributes to the pathogenesis of NIDDM, although its rela-

tive importance remains to be assessed. In addition, it is possible that altered expression of GLUT2 plays a role in the pathogenesis of at least some variants of human NIDDM. B. Glucose Transport in Tumor Cells Dysregulated growth and parasitic behavior of tumor cells have been often attributed in the past to an enhanced uptake of nutrients. Evidence for this view came from tumor cells in vitro that exhibited an increased rate of amino acid and hexose uptake (Isselbacher, 1972). In particular, an enhanced transport of glucose has been correlated with the deviation toward aerobic glycolysis, a main metabolic change of malignant cells. However, this simple concept was developed from studies in highly malignant cells of transplantable tumors before the molecular biology of the GLUT family could be elucidated. More recent studies take into account the heterogeneity of G L U T transporters in different cells and the complex regulatory mechanisms that affect their expression. A somewhat complex picture has emerged from these studies. For instance, both the expression and the activity of glucose transporters were found to be sensitive to proliferative stimuli derived from cell transformation, oncogene expression, and growth factor signals (Flier et al., 1987; Rollins et al., 1988; Hiraki et al., 1989). It has been known for several years that the mechanisms underlying these changes may act at either the transporter m R N A (Birnbaum et al., 1987) or protein levels (Shawver et al., 1987). Data obtained in recent years suggest that an increased abundance of G L U T proteins is effectively detectable in human tumor cells and that, usually, overexpression of the transporters GLUT1 and GLUT3 is responsible for this change (Brown and Wahl, 1993; Nagamatsu et al., 1993; Nishioka et al., 1992). Systematic

Derangements in Transport Regulation

attempts to ascertain whether high glucose influx is a constant feature of human tumors have indeed demonstrated that the abundance of GLUT1 transporters is increased in many human carcinomas compared with normal epithelial tissues or even with benign epithelial tumors (Yamamoto et aL, 1990; Clavo et al., 1995; Younes et aL, 1996). What is the mechanism for GLUT transporter overexpression in human tumors? In some cases enhanced transport is effectively attributable to molecular changes in GLUT genes. This is the case in lung cancers, where a gene amplification of GLUT1 and GLUT3 is detectable and correlates with a bad prognosis (Ogawa et al., 1997). On the other hand, no amplification or rearrangement of GLUT genes was detected in many other instances (see Mellanen et al., 1994, for head and neck tumors). In these models transport overactivity probably derives from a regulatory change in transporter expression. This is well documented, at a molecular level, for hemangioblastoma, whose cells express an 80-kDa protein that reacts with a 10-nucleotide domain of the 3' untranslated region of the GLUT1 mRNA, stabilizing the messenger in a typical post trascriptional regulatory mechanism (Tsukamoto et al., 1996). Regulatory changes are also suggested when overexpression of GLUT transporters is associated with an enhanced expression of type II hexokinase (Shinohara et al., 1994), a result that points to the occurrence of a complex, coordinated mechanism that produces an overall acceleration of glucose metabolism. An important regulatory mechanism that influences glucose transport is the increase in sugar influx under conditions of hypoxia and metabolic inhibition, an effect conserved in tumor cells (Minn et al., 1996). Defects of perfusion and consequent local ischemia are quite common in solid tumors, and GLUT1 transporters are particularly abundant in ischemic areas (Clavo et al., 1995). Therefore, the possibility that an increased expression of GLUT1 derives from a response of cells to local conditions rather than from a genetic change linked to neoplastic transformation should be taken into consideration. However, in red blood cells stimulation of glucose transport by metabolic inhibition can also derive from modification of existing transporters (see Cloherty et al., 1996). This may also be the case in some tumor cells. Other regulatory mechanisms that influence hexose transport depend upon extracellular signals such as hormones or growth factors. In a well-documented example (Spieholz et al., 1995) it has been demonstrated that granulocyte-macrophage colony-stimulating factor stimulates glucose transport in melanoma cell lines, thus suggesting that autocrine or paracrine factors may be involved in transport alterations detected in tumor cells.

291

Similar findings were obtained also for EGF in astrocytic tumors (Murphy et al., 1992). In conclusion, although enhanced glucose transport appears to be a relatively common feature of tumor cells, it should be considered more an adaptation of the cells to a hostile environment than a tumor-specific metabolic feature. This complex array of changes has discouraged early attempts to employ GLUT transporters as reliable markers for cancer cells in vivo (Nelson et al., 1996). C. Amino Acid Transport in Tumor Cells As far as changes in amino acid transport are concerned, experimental results are contradictory and definite conclusions are very difficult to make. Early results from a number of models agree with the view that tumor cells are endowed with higher transport activity. For instance, the rate of uptake of amino acid substrates of the Na§ systems A and ASC markedly increases in virus- and chemically transformed cell lines (Borghetti et al., 1980; Boerner and Saier, 1982). However, all these studies did not always take into account possible disturbing effects of different proliferation rates of normal and neoplastic cell populations. More recent and controlled studies yield a different picture. No significant change is detected between normal and neoplastic models as far as the transport systems for neutral amino acids (Christensen's systems A, ASC, and L) are concerned (Nucci et al., 1988; Gazzola et al., 1990) when distortions from different proliferative activities, histotypes, and genetic heterogeneity are eliminated. However, detailed analyses of other neoplastic models indicated that peculiar changes in the activities of one or the other transport mechanisms can occur in a particular tumor cell. 3T3 mouse fibroblasts, transformed with activated ras or neu oncogenes, and C6 rat glioma cells are endowed with a very low activity of the Na§ high-affinity transport system for aspartate and glutamate, corresponding presumably to EAAT3 (Longo et al., 1988; Cho and Bannai, 1990; see above). Conversely, the activity of another transport mechanism for anionic amino acids, the Na § independent system X-c, is increased. The amino acid transport changes reported in these studies should be attributed neither to changes in cell growth nor to mutations of the carrier genes. Rather, they appear the results of a regulatory derangement. Employing a model of conditional expression of ras in 3T3 cells (Jaggi et al., 1986), it has been possible to demonstrate that suppression of EAAT3 activity is a reversible phenomenon. It is progressively detected upon oncogene expression

292

9. Regulation of Plasma Membrane Transport

(Uggeri et aL, 1995) and, conversely, released when ras expression is silenced (Uggeri et al., unpublished observations). Interestingly, suppression of anionic amino acid transport is only one aspect of a pleiotropic modulation of membrane functions, which also comprises an increased activity of Na+/H + antiport (Maly et al., 1989) and of furosemide-sensitive Na+/K+/CI - cotransport (Meyer et al., 1991). Somewhat comparable results are obtained with neoplastic cells derived from "spontaneous" human tumors. When chronic lymphocytic leukemia cells were compared with normal B lymphocytes, the activities of the Na+-dependent systems A and ASC for neutral amino acids have been found unchanged. On the contrary, tumor cells exhibited a marked decrease of the activity of another transport system for neutral amino acids, the Na+-independent system L (Segel and Lichtman, 1982; Segel et al., 1984), corresponding to a change in Vmax of more than one order of magnitude (Segel et al., 1989). Again, this modification is not apparently due to a genetic change involving the gene(s) of the transport system. Maturation of leukemia cells, triggered by a treatment with phorbol esters, led to the functional normalization of system L activity (Segel et al., 1988). Analysis of cell proteins expressed following phorbol treatment (Woodlock et al., 1993) revealed 14 newly synthesized distinct protein species, 4 of which exhibited photolabeling by O-diazoacetyl-L-serine (Segel et al., 1989), inhibitable by the L-type specific substrate 2-amino-2-carboxybicycloheptane (BCH). These results should soon be interpreted in relation to cloned system L transport and accessory proteins (e.g., Karai et al., 1998; Mastroberardino, 1998).

D. Expression of Ectopic or Novel Transporters in Tumor Cells A peculiar feature of malignant cells is abnormal or absent differentiation (anaplasia). Derangement of a differentiation program often causes tumor cells to produce inappropriate (ectopic) proteins, such as enzymes characteristic of fetal tissues or hormones typical of other tissues. Also, in selected cases, altered expression of transporters ensues as an aspect of the ectopic expression of proteins in tumors. For instance, breast cancer cells express the fructose-preferring transporter GLUT5, which is present in intestine and other tissues but not in normal breast epithelium (Zamora-Leon et al., 1996). Reversion to "fetal-type" transporters may also occur in tumor cells. This may be the case with system y+ for cationic amino acids, which exhibits a very low activity in "adult" hepatocytes but is clearly detectable in hepatoma cells (White and Christensen, 1982). Elucidation of the molecular basis for this change required

the characterization of the CAT (cationic amino acid transporter) family (see MacLeod et al., 1994, for review). The ubiquitous isoform CAT1 was identified with the previously cloned receptor for ecotropic murine retroviruses (Wang et al., 1991; Kim et al., 1991). Then two other members of the family, CAT2A and CAT2B, derived from alternative splicing of the same gene, were characterized in mouse liver and lymphocytes (Closs et al., 1993; Kavanaugh et al., 1994). The CAT family is expanding with the identification of a brain-specific transporter (Ito and Groudine, 1997; Hosokawa et al., 1997) and of human counterparts of rodent transporters (Closs et al., 1997). Although CAT2A and CAT2B are highly homologous, but for subtle differences in the active site, CAT2B is endowed with a 30- to 70-fold higher affinity for substrates than CAT2A. "Adult," differentiated hepatocytes express only the low-affinity CAT2A transporter, while no high-affinity transport system for cationic amino acid is present in these cells. Conversely, hepatoma cells express high affinity CAT2B and are endowed with high-affinity arginine transport. In other words, transformation may release differentiation-related suppression of CAT2B expression. The expression of different transporters due to anomalous differentiation may also underlie the discrepant properties of neutral amino acid transport observed for the first time in important early studies by Christensen and coworkers (Handlogten et al., 1981). It was later demonstrated (Chiles and Kilberg, 1986) that system A activity in H4 rat hepatoma cells was completely inhibited by the protein-modifying agent N-bromosuccinimide while system A in normal hepatocytes was unaffected; conversely system A activity in normal rat hepatocytes was sensitive to inhibition by the sulfydryl reagent N-ethylmaleimide while transport activity in hepatoma cells was not. Subsequent investigations showed that this differential inhibition of transport must be due to the presence of modified system A transport protein in the hepatoma cell line (Dudeck et al., 1988). In a study of glutamine transport in human hepatocytes and in the human hepatoma cell lines SK-Hep and Hep-G2 (Bode et al., 1995), it was shown that glutamine uptake was much faster in the hepatoma cells where it did not occur on the classic system N but, rather, on a system which had the characteristics of a modified system ASC. Glutamine transport in rat hepatoma cell lines was recently found to be catalyzed by two kinetically distinct transport systems (McGivan, 1998). One showed the characteristics of system N while the other was Na + dependent; did not accept substitution of Na + for Li+; was insensitive to external pH; and was inhibited by excess concentrations of leucine, histidine, and lysine. These properties differ from those of any

Summary

so-far characterized transport system in hepatocytes. The Na+-dependent interaction of the system with both cationic and zwilter ionic amino acids is consistent with expression of the rodent embryonal system B ~ (Van Winkle et aL, 1985 and 1990f) in rat hepatoma cells. It may be speculated that hepatoma cells express isoforms of the normal liver transporter proteins, although their relationship to cell transformation remains to be clarified.

VII. SUMMARY Starting from the earliest studies on mammalian plasma membrane transport it has become apparent that transport activities, like enzyme activities, are subject to both short-term and long-term regulation. In the past few years the cloning and sequencing of cDNA for various transport proteins has allowed the exploration of the mechanisms of transport regulation in greater molecular detail. This chapter covers the study of the regulation of transport from the original kinetic experiments to the most recent work using transgenic animals. Short-term mechanisms of regulation of transport activity include changes in the driving force, as exemplified particularly by the effects of membrane hyperpolarization by glucagon on liver cell substrate transport. Regulation of activity by phosphorylation has been characterized particularly in the cases of the CFTR protein and the Na+/H + exchanger family. In these cases phosphorylation of transport proteins on specific sites has been demonstrated directly and protein phosphorylation has been correlated with transport activity. Regulation of glutamate transport via the E A A T family is another good example of regulation of activity by phosphorylation. Other short-term mechanisms are possible but are less well characterized. For example, changes in subunit-subunit interactions may play a part in the regulation of some of transport ATPases that are heterodimers. There is increasing indirect evidence that a number of other transport proteins act functionally as oligomers and, again, changes in subunit interactions provide a potential mechanism of regulation. Finally, allosteric interactions of transport proteins with substrate molecules or with other proteins may play a part in the regulation of Na+/H + exchangers. It appears likely that as more information becomes available about the molecular structure of transport proteins, further such mechanisms will be recognized.

293

Long-term regulation of transport involves induction either of transport proteins or possibly in some cases of other proteins that may interact with and activate existing transporters. Hormonal induction of amino acid transport has been well characterized, but is still poorly understood at the molecular level. Another well-studied phenomenon is the induction of mammalian transport activities by long-term substrate deprivation; again, many of the molecular events involved are still obscure. Exposure of cells to hyperosmotic stress induces Na § dependent substrate transport in various cell types. The best-understood example of this is the transport of compatible osmolytes in renal medulla where a "tonicity sensitive element" in the promoter of the Na+-betaine cotransporter BGT-1 has been shown to be involved in the osmotic response. Evidence for the coordinated regulation of various transport activities in cell volume regulation and during the cell cycle are also discussed. An increasingly important area is the role of transport regulation in disease. Evidence for and against the views that defective GLUT4 regulation may be involved in non-insulin-dependent diabetes and that defective GLUT2 regulation may contribute to defective insulin secretion are reviewed. The molecular basis and role of altered glucose and amino acid transport activities in tumor cells are also discussed. Many problems in this area are at present unresolved. For example, how do cells recognize lack of substrate and how is this message translated into the synthesis of new transport molecules? Do "transport-activating" proteins exist, and, if so, how do they work? How do various hormones induce transport activity? Do closely similar isoforms of transporters with different activities exist in different tissues? Is defective transporter regulation a contributory factor in other pathophysiological conditions? Finally, what is the physiological importance of some of the phenomena seen in isolated cells? For example, the stimulation of alanine transport by glucagon in hepatocytes is clearly important in relation to the control of gluconeogenesis. On the other hand, the physiological significance of the complex regulation of the glutamate transporter family by phosphorylation is less apparent. Some of the answers to these questions await the cloning of both the promoter regions and coding regions of genes encoding further transport proteins. Others may come from further work using transgenic animals that do not express various transporters. Rapid advances in all these areas may now be expected.

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Biomembrane Transport and Interorgan Nutrient Flows: The Amino Adds t

port mechanisms situated at the cell-extracellular fluid interface (Waterlow et al., 1978; Biolo et al., 1992a). Amino acid (AA) movements between the major metabolic compartments of the body are summarized in Fig. 10.1. Amino acids may enter the free pool by uptake from the diet, release from protein breakdown, or by de n o v o synthesis. Amino acids are removed from the free pool either by reversible incorporation into proteins, by utilization in other biosynthetic processes (e.g., purine synthesis), by oxidation, or, to a negligible extent, under normal circumstances, by excretion. Over the long term these processes are usually highly regulated to maintain interorgan and whole-body nitrogen balance (and to avoid poisoning by excess of dietary aromatic and branched-chain amino acids). However the system is capable of rapid acute adaptation in response to changes in diet and physical activity. Physiological mechanisms such as growth and repair and pathophysiological mechanisms in disease and injury may result, respectively, in positive and negative whole-body nitrogen balance (Waterlow et al., 1978; Waterlow, 1995). Proteins are synthesized from a relatively small number of amino acids (20 translationally recognized, including the imino acid proline, plus a smaller number generated by posttranslational modifications). Compartmentation and metabolic specialization of tissues and organs are important factors both dictating and facilitating interorgan nutrition. Important elements here include the enzyme complement and the metabolite concentrations within specific compartments (e.g., cytosol, mitochondrion, and extracellular space). Both fluxes and concentrations of metabolites are important in complex systems and these show mutual dependence because of the net movement of metabolites between compartments in interorgan nutrition. Two ma-

I. INTERORGAN NUTRITION The concept of interorgan nutrition is encapsulated in the question posed by Christensen in a seminal review (Christensen, 1982): "How is each organ and each type of cell nourished by flows from other tissues of the same organism"? It is implicit that the nutritional requirements and processes of the various body compartments (i.e. tissues and their constituent cells) are not all the same. A cell may, of course, provide its own supply of a specific nutrient by de n o v o synthesis as well as, or instead of, receiving delivery through the blood supply. It should, however, be noted that the demonstrated ability of a cell to synthesize a particular nutrient does not necessarily mean that it can produce amounts sufficient for its own needs (this could only truly be demonstrated by blocking delivery from all external sources). The key concepts of interorgan nutrition are common to major nutrients including amino acids, sugars, and free fatty acids. In this article we focus mainly on the amino acids, which are of particular interest because of their structural diversity and their broad range of functions in cell biology.

II. INTERORGAN AMINO ACID NUTRITION: GENERAL PRINCIPLES AND KEY ISSUES A. Compartmentation within the Body The free amino acid pool of the body is subdivided into extracellular and intracellular "subpools" that exchange with one another by means of amino acid transt Peter M. Taylor, MichaelJ. Rennie, and SylviaY. Low,Department of Anatomy and Physiology, University of Dundee, Dundee DD1 4HN, Scotland, United Kingdom

295

296

10. Transport and Interorgan Nutrient Flows

dietary protein

extraceUular free AA pool

hydrolysis to

AA and peptides

J

cellular free AA pool

()

gut lumen

~

()

)

[

"

AA

other metabolites

3

FIGURE 10.1 Movements and exchanges of amino acids between different compartments of the body. (1) Protein synthesis; (2) protein breakdown; (3) transport mechanisms at cell membrane; (4) amino acid and peptide absorptive mechanisms in intestinal epithelium; (5) utilization in cell metabolism (e.g., purine biosynthesis, oxidation, excretory mechanisms); (6) de n o v o synthesis.

jor factors contributing to interorgan flows are first, the mass-action relationships established between "producer" and "user" compartments either locally or at a distance (the latter mediated through changes in composition of the intermediate "mobile" extracellular fluid compartment) and second, the work performed in active solute transport across cell membranes against the prevailing total chemical potential gradient. The relative contribution of these two mechanisms to interorgan nutrition continues to attract the attention of many investigators. Variables influencing interorgan amino acid fluxes therefore include: 9 The overall quantity of amino acids in the free pool (and hence their concentration in both extracellular and intracellular fluid compartments). 9 Rates of production and utilization within tissues. 9 Rate of delivery of amino acids to tissue in regional circulations. 9 The activity of the various transport mechanisms in cell membranes. This last variable (transport activity) determines the overall exchange and movement of amino acids between cells and extracellular fluid. A single amino acid may be transported into a specific cell type by several distinct transport mechanisms (Christensen, 1982, 1990; Malandro and Kilberg, 1996). The relatively weak specificity of transmembrane amino acid transport in series with the generally high specificity of intracellular enzymatic

catalysis, the specificities and capacities of these serial mechanisms, and their regulation by adaptive and endocrine mechanisms underpin interorgan amino acid nutrition. Much recent study in this area has been geared to answer the key question (as expounded by Christensen, 1990) of "to what degree does membrane transport, on the one hand, and the rates of metabolic reactions on the other hand, figure in the modulation of interorgan amino acid flows?" Another important issue is the extent to which these physiological processes match between capacities (i.e., Vmaxof transporters and enzymes) and loads (i.e., maximum expected flux under normal circumstances) so that "enough but not too much" (Diamond, 1993) transport or enzyme protein is produced and maintained. B. Methodological Issues Interorgan nutrition is de facto a scientific study of the whole body and the interactions between its constituent organs, tissues, and cells. Research at all levels of bodily organization is applied to this study and much useful information has been obtained at cellular and subcellular levels that can be readily extrapolated to higher levels of organization. The methods used to study the cellular and molecular biology of membrane transport and metabolism are discussed in other chapters (e.g., 4 and 11), but a brief update on current methodological approaches for investigation at whole-body and organ levels is included below.

lnterorgan Amino Acid Nutrition

1. Whole-Body Investigations Interorgan nutrition has traditionally been studied by sampling and analyzing the blood that supplies and drains the organ or tissue of interest. Quantitation of interorgan amino acid fluxes was pioneered in the late 1960s (e.g., Felig, 1975; Cahill, 1976; Christensen, 1982 for review) through the combination of measurements of blood flow and composition, where net flux flow • arteriovenous concentration difference of amino acid. The requirement of blood sampling in this type of work involves invasive methodologies but recent advances in the sensitivity of analytical instruments plus miniaturization of devices for measuring blood flow (at least in experimental animals) have facilitated advances of knowledge in the field of interorgan nutrition. The net nutrient fluxes obtained using these methods, although extremely useful, often mask larger unidirectional fluxes between cells and blood as well as intratissuc "cycling" of substrates. Unidirectional fluxes may be measured using tracers. The fate of an A A tracer introduced orally by bolus injection or by infusion into the plasma pool may be followed in one of two major ways (both protocols require sequential sampling of blood, preferably via a cannulated vein): 1. Time course of disappearance of bolus tracer load from plasma into exchangeable tissue pool(s) (Waterlow et al., 1978). Most AAs in the body are chemically combined into protein and the free A A pool therefore constitutes only a small proportion of the total, although it is this pool that turns over the most rapidly and is responsible for interorgan transfer of amino nitrogen and A A carbon. Linear kinetic modeling of tracer disappearance (e.g., Gastaldelli et al., 1997) is likely to reveal both rapidly and slowly exchanging free solute pools (compartments) for which size (distribution volume) and exchange rate constants can be estimated. Such multicompartmental analysis is useful for studying turnover but gives no information as to which tissues represent particular compartments. Radiotracers labeled with ~1C or 13N may be used to measure (or visualize by imaging) noninvasively the accumulation of tracers within tissues (e.g., Schmall et al., 1996). 2. Constant infusion (usually after a priming dose) of tracer into plasma to achieve a steady state (e.g., Carraro et al., 1994; Nurjhan et al., 1995; Hankard et al., 1995). This can be used to estimate whole-body appearance (Ra) of amino acids into the plasma (i.e., release of amino acids from cells to the extracellular fluid). Plasma appearance of essential AAs reflects their release by protein breakdown, whereas for nonessential

297

AAs both de n o v o synthesis and protein breakdown contribute to the overall appearance rate.

2. Regional and Tissue Studies The combination of arteriovenous concentration differences and blood flow measurements for tissues with accessible venous drainage in vivo provides useful information on net A A fluxes (calculated as flow • ([substrate]art - [substrate]ven)). Plasma flow should be used if the A A under study is effectively excluded from blood cells over the time course of the experiment or if plasma A A concentrations were obtained. Plasma and wholeblood concentrations of certain AAs (notably glutamate and aspartate) differ significantly from one another but the overall importance of blood cells in A A carriage is likely to be relatively minor under most experimental circumstances (Christensen, 1982; Felig et al., 1973). More detailed quantitative information on transport steps may be obtained if tracer is added at the arterial side of a tissue vascular bed and the venous blood is sampled during the first pass of tracer in the circulation. The paired tracer dilution method has been used in vivo (e.g., in the human forearm; Bonadonna et al., 1993) by examining tracer profiles in the effluent venous blood. The inclusion of tissue biopsies (or terminal tissue samples) increases the amount of information that can be obtained. Tissue-blood distribution ratios of AAs in vivo may be measured from chemical analysis of biopsy material or from steady state distribution of injected A A tracers (Christensen, 1982, 1990). Transportable tracer plus extracellular marker may be bolusinjected into an arterial circulation and tissue freezeclamped within 5 to 8 sec (i.e. less than the circulation time), in which case the tissue retention of tracer relative to extracellular marker can be used as an index of cellular extraction of tracer (e.g., Oldendorf, 1971). Amino acid flux may be calculated from this unidirectional extraction value if both tissue blood flow and plasma A A concentration are known. Alternatively, if constant infusion of tracer is used, a tissue biopsy at plasma steady state may be obtained from which tracer enrichment values may be measured. Using this information, a three-compartment model (Biolo et aL, 1992a, 1995) allows the quantitation of intracellular A A kinetics including transmembrane A A transport. The model is applicable in vivo (given access to the venous circulation draining the tissue of interest and a suitable method for measurement of tissue blood flow) and can be extended to include additional flux components representing appearance or removal of tissue A A through proteolysis and de n o v o A A synthesis or protein synthesis, respectively.

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10. Transport and Interorgan Nutrient Flows

C. Quantitative and Qualitative Importance of Amino Acid Flows b e t w e e n Tissues Interorgan nutrition is not merely a matter of random movement in the bloodstream of the amino acid products of protein digestion-absorption and their indiscriminate extraction by tissues. Some interorgan flows are directed because particular metabolic and membrane transport steps are restricted to certain sites in the body (e.g., the liver alone contains the full complement of urea cycle enzymes) and others because of particular structural relationships between tissues and t h e interconnecting vasculature. This is not always the case, however, and indeed many important substrate flows are of compounds that are produced or utilized by common metabolic pathways in a wide variety of cells (e.g., glutamine is synthesized by several tissues for release into the bloodstream). Equally, certain nutrients that are nonessential (or dispensable) as dietary components may be necessary as "essential" substrates for particular tissues, a requirement serviced by net synthesis and export by other tissues. This is the case for a number of nominally dispensable amino acids, notably glutamine (see Section IV,A). There are a variety of extremely important metabolic fluxes of major dispensable amino acids from one organ to another. For example, flows of alanine contribute to a glucose-alanine cycle where the amino groups released during muscle metabolism are directed to the liver in the form of alanine. The flows of alanine and glutamine between tissues are much greater than one would expect from protein hydrolysis alone and reflect large rates of de n o v o synthesis in various tissues. The amino acids with the largest role in intermediary metabolism are those for which component cells in the body are capable of appreciable rates of synthesis, i.e., they tend to be the dispensable amino acids. The sites of synthesis may well be very discrete and many tissues will require a supply of that amino acid from the sites of synthesis as well as from the diet. Equally the rate of synthesis within the particular cell may be inadequate for its requirements, calling for delivery by the dual methods of de n o v o synthesis and uptake. Amino groups may be transferred between the carbon skeletons of amino acids by a number of aminotransferases that usually operate at or near equilibrium. Glutamate occupies a central position in these transamination pathways, but it is largely sequestered within cells, reducing its importance as a vehicle of interorgan amino acid nutrition. The dispensable (or nonessential) amino acids glutamine and alanine play an especially important role in the overall transfer of amino nitrogen between tissues (Fig. 10.2). These amino acids are among the most abundant in the free pool, are intermediates in a wide variety

~CLE,

ADIP~

glutamine/

]alanine ~glutamine

urea

NH3

NH3 I

-~alanine

--1~ NH3

"~ ~ urea

I PERIVENOUS

1--73 LUNG glutamine

,~

FIGURE 10.2 Major pathways of interorgan amino acid metabolism in mammals. This schematic diagram shows the key sites of alanine and glutamine production and breakdown in intestine, liver, muscle, kidney, fat, and lung.

of metabolic pathways (both being readily interconverted with glutamate inside cells), and are carried by a broad range of transport mechanisms. Glutamine and, to a lesser extent, alanine are accumulated within certain cell types, notably muscle (the largest overall intracellular compartment of the body), by secondary active transport mechanisms. These accumulated amino acids represent a significant and important "store" of free amino nitrogen, which can be released into the blood circulation for transport to other tissues requiring nitrogen under particular circumstances. There are important links between the metabolism of amino acids and carbohydrate in terms of interorgan carbon flux (e.g., in provision of gluconeogenic substrates for the liver) and these tie in with movement of amino nitrogen groups (e.g., for urea synthesis), as considered in more detail in Section IV. Major pathways of interorgan metabolism of amino acids and related compounds that we consider in this chapter are : 9 Glutamine flows between peripheral (muscle, adipose), pulmonary, and splanchnic tissue. 9 Fluxes of alanine from muscle to viscera for urea and glucose synthesis. 9 Glutamate/glutamine and glycine/serine exchanges across the placenta.

interorgan Amino Acid Nutrition

9 Aromatic amino acid delivery to the central nervous system (for neurotransmitter synthesis). 9 Glutamine delivery to the kidney during disturbed acid-base balance. 9 Cystine and glutathione turnover. We also examine certain intercellular amino acid flows within a specific organ that may be quantitatively small (and/or highly localized) but nevertheless physiologically vital, e.g., Gln/Glu exchanges within the brain and the liver. A number of nominally dispensable amino acids (e.g., glutamine, arginine) show conditional essentiality, which is particularly apparent under conditions where body amino acid requirements are elevated, e.g., during growth or in certain disease states (see Section V). For example, a growing tumor may utilize glutamine at such a great rate that it successfully competes with other tissues (e.g., the intestine and immune system) and deprives them of their glutamine requirements, which are normally met by d e n o v o synthesis and export from tissues such as muscle, liver, adipose, and lung (Souba, 1992). Muscle contains a greater proportion of body protein than any other tissue and serves as the principal tissue determining the balance between whole-body amino acid catabolism and anabolism (Christensen 1990; Waterlow, 1995, Rooyackers and Nair, 1997). Muscle tissue functions in interorgan metabolism include the supply both of gluconeogenic and ureagenic substrates and of fuel to support immune, growth, and repair responses. The major potential source of nitrogen loss from the body is the renal ultrafiltrate, which contains smallmolecular-weight solutes such as amino acids and glucose at plasma concentrations. However, these solutes are actively and avidly reabsorbed by renal transport mechanisms, largely in the proximal tubule (Silbernagl, 1988 for review), and the overall renal clearance of amino acids from the plasma is virtually zero under normal circumstances (renal reabsorption of amino acids is considered in more detail in Sections IV,F and IV,G). D. Barrier Function of Cell M e m b r a n e s Cell membranes may be regarded as regulatory barriers controlling flows of metabolic intermediates between different cells, organs, and tissues. Numerous examples of this barrier function are described in the later sections of this chapter. Amino acid transport proteins in the cell membrane facilitate or direct flows of amino acids across the barrier as a consequence of their specific transport properties (outlined below in Section II,E).

299

The epithelial cell barriers of tissues including the gut, placenta, renal nephron, and lung have low intercellular permeability to solutes such as amino acids and these molecules must therefore traverse two membranes in series by a transcellular route if they are to be moved across the epithelium. Transcellular movement of amino acids may involve their intracellular metabolism such that the amino acids exiting one face of the cell are not necessarily those entering at the opposite face (examples of this are considered in Sections IV,F, V,A, and V,D). Amino acids moving between tissues through the bloodstream must also cross the capillary endothelium at both ends of their journey and in most tissues the intercellular junctions between endothelial cells have high solute permeability facilitating paracellular diffusion across the endothelial barrier, driven by local gradients of solute concentration. An important exception is in the central nervous system where blood capillaries in most regions have a low permeability even to small organic solutes and the capillary endothelium constitutes the "blood-brain barrier" (BB barrier). Because of the tight intercellular junctions between the endothelial cells lining capillaries of the brain, glucose for energy and amino acids for synthesis of proteins and many neurotransmitters must be transported across endothelial cell membranes of brain capillaries (see Sections IV,C and IV,F). This arrangement allows a large degree of control to be exerted over the composition of interstitial medium within the central nervous system (the cerebrospinal fluid), without which both chemical and electrical communication in the brain may be compromised by fluctuations in plasma solute composition (as occurs for example, during absorption of the constituents of a meal). This is particularly important for amino acid neurotransmitters such as glycine and glutamate, which are usually kept at very low concentrations in the brain interstitium by active transport processes that drive them into neural and glial cells (Smith and Cooper, 1992). Excesses of these particular amino acids are thus avoided, preventing excitotoxic actions that may produce lesions in active brain areas. Cerebrospinal fluid has relatively low concentrations of most amino acids and nutritional flows of amino acids to the brain must largely pass through this substrate-depleted medium. The uptake of amino acids into nerve and glial cells within the protection of the blood-brain barrier occurs by a variety of transport systems as for other tissues and active exchange of metabolites take place between different cell types in the brain (see Sections IV,C, IV,E, and IV,F). Specific transport systems for neurotransmitter amino acids are present within the brain (Smith and Cooper, 1992; Malandro and Kilberg, 1996; Kanai, 1997).

300

10. Transport and lnterorgan Nutrient Flows

E. Amino Acid Transport Systems and Their Functional Properties 1. Classification of Amino Acid Transport Systems The pioneering work of Christensen and colleagues in the early 1960s led to the identification of a wide variety of amino acid transport mechanisms or "systems" (see Table 10.1) based on criteria of function (Christensen, 1990; Van Winkle, 1992; Bertran et aL, 1994) and later of structure after the advent of molecular cloning (Bertran et al., 1994; Malandro and Kilberg, 1996). These transport systems differ in their substrate range, in their dependence on cosubstrates such as ions (particularly Na+), and in their regulatory properties such as sensitivity to hormones (e.g., insulin). Transporters for the translationally recognized amino acids (encompassing 19 amino acids plus the imino acid proline) are generally selective for a particular type of substrate.

TABLE 10.1

This selectivity is related to side-chain features such as charge (anionic, cationic, zwitterionic or "neutral"), size and structure (e.g., extra N in glutamine, asparagine, histidine). Amino acid transporters may be functionally subdivided as follows: 9 Na + dependent: Systems A, ASC (widespread), systems B ~ B ~ (epithelial brush borders, embryonic tissue), system N (liver, N m in muscle). 9 Na + and K + dependent: System X-AG widespread (especially neural and epithelial tissues). 9 Na + and C1- dependent: GABA, taurine, and glycine transporters (widespread, highly expressed in neural tissue). 9 Na + independent: Systems L, T, y+, b ~ Suffixes (+ or - ) denote carriage of appropriately charged amino acid substrates. The "system"-based nomenclature is largely derived from "model" or "para-

Summary of Some Known Amino Acid Transport Systems in Tissues and Cells of Higher Animals a

Ion dependence

System properties

Abbreviation

Analogous systems/special features

Widespread; Gly and Sar; variants known; N-methyl (N-Me) group tolerance

Na,C1

Imino, iminoglycine, and epithelial

Widespread, serves for most dipolar amino acids; often repressed; insulin-sensitive

Na

N-Me group tolerance

ASC

Widespread, some variation in scope; excludes N-Me amino acids, but often includes prolines; on protonation accepts analogous anionic amino acids; may accept Arg without Na +

Na (K)

Na+-independent asc system in erythrocytes has somewhat analogous scope

Bo,+

Wide-scope system, oocytes, blastocysts probably renal and intestinal brush borders; accepts cationic AAs limited by positions and bicyclic amino acids despite their bulk

Na

b ~ a Na+-independent analog more sharply limited by positions of branching, B/B ~ mouse blastocysts/ intestinal and renal epithelium

Gin, Asn, His; liver

Na

N m, insulin-sensitive Gln transporter in

/3-Ala, Tau, 4-aminobutyrate

Na, CI

System specific to 4-aminobutyrate.

Gly

Separate systems L1, etc., low Km analog develops on incubation of hepatocytes

Widespread, tolerates bulky and branched chains; high exchange property; bicyclic amino acids as model substrates T y+ y+L X-AG

System t in fibroblast lysosomes

Erythrocytes, hepatocytes, favours benzenoid amino acids Widespread, no discrimination between Arg, Lys and homologs, Homoserine

(Na for dipolar AA)

Epithelia, erythrocytes; similarities with b ~ substrates need Na +

(Na for dipolar AA)

but dipolar

Ubiquitous, includes neurons; similarly reactive with Aspand Glu-

X- G

Glu- and analogs, largely excluding Asp- and short analogs

X- C

Like X-G, except cystine competes/exchanges with Glu-, may be locked into exchange; explains some CNS Glu"binding"

muscle.

Na, K (H)

aAdapted from Christensen (1990) with permission from the American Physiological Society.

Variants, e.g., lysosomes, blastocysts, do not accept homoser + Na +

Interorgan Amino Acid Nutrition

digm" substrates, and kinetic discrimination between systems has been aided by design and synthesis of nonmetabolizable amino acid analogs that are at least nominally "system specific", e.g., N-methylaminoisobutyric acid (MeAIB) for system A and 2aminobicyclo[2,2,1]heptane-2-carboxylic acid (BCH) for system L. Cloned transporters (Malandro and Kilberg, 1996; Kanai, 1997; Dev6s and Boyd, 1998 for review) include (1) the CAT family (at least four isoforms) mediating system y+-like transport; (2) the EAAT superfamily including excitatory amino acid transporters for anionic amino acids (EAAT1-EAAT5; system X-AG, and structurally related ASC/ATB ~ carriers for zwitterionic amino acids); (3) the GABA transporter superfamily (including GABA, Gly, Tau, and Pro transporters); and (4) the BAT/4F2 proteins involved (probably as activating subunits) in mediation of system b~ transport. The major insulin-stimulated amino acid transport systems are A and N m, and they contain amino acid transport proteins yet to be cloned.

2. Transport Mechanisms and Kinetic Properties Amino acid transport systems effect saturable translocation of substrates across a cell membrane. The relationship between substrate concentration and rate of transport can be at least formally described by Michaelis-Menten kinetics and transport mechanisms are conventionally characterized in terms of maximal capacity (Vmax) and strength of interaction between transport protein and substrate (Km) (see Chapter 4). Physiological flux of an amino acid through a specific transport system is therefore dependent on its concentration at the cis-face of the membrane (and sometimes also at the trans-face) as well as the transport capacity of the system (Christensen, 1990; Guidotti and Gazzola, 1992). Together the amino acids represent "the largest group of mutually analogous nutrients" (Christensen, 1990). The zwitterionic amino acid template to which discrete side-chains are added is a common feature for molecular recognition within the grouping, although each amino acid has a side-chain differing in length, polarity, charge, and atomic complement. Evolutionary honing of molecular recognition in living organisms has progressed to the extent that individual amino acids are discriminated for binding to transfer RNAs and by many enzymes yet, in general, amino acid transport systems may discriminate only weakly among these analogs. The use of overlapping and common transport pathways for amino acids may therefore offer some advantage (e.g., in terms of improvements in metabolic efficiency), which in terms of evolutionary pressures must be weighted against the likelihood and potential consequences both of incorrect

301

substrate selections and of inappropriate competition between substrates (Christensen 1990). The relatively low specificity of amino acid transport for analogous structures means that changes in concentration of one amino acid may have considerable (and physiologically relevant) effects on transmembrane fluxes of other structurally related amino acids sharing the same transport system, either by c/s-competition or transacceleration (or in some cases trans-inhibition) (see Sections IV,F and V,D). Membranes of individual tissues or cell types are likely to express a variety of transport mechanisms with different ranges of substrate specificity and transport capacity (Table 10.1). Amino acid transporters with mechanisms generating a net charge (related to movement of ionic amino acids themselves or of inorganic cosubstrates such as Na +) produce measurable currents across the cell membrane and these rheogenic transport mechanisms may be of sufficient magnitude to significantly influence resting cell membrane potential (i.e., they may exert an electrogenic effect). Rheogenic transporters include EAAT and CAT isoforms (where charge is related to movement of the cationic amino acid). The glutamate transporter EAAT1 is also reported to cotransport hydrogen ions or countertransport hydroxyl ions in a manner capable of producing a fall in intracellular pH (this change is measurable in cells overexpressing EAAT1; Kanai et al., 1995). It is therefore conceivable that glutamate transport might increase cellular acidosis, a phenomenon with many effects such as accelerating glutamate utilization and enhancing ammoniagenesis (see Section V,D). Glutamate and aspartate transport may, therefore, result in delivery both of substrates and an acid load to cells extracting it from the plasma, hence multiplying the possible metabolic effects of the transport processes. Asymmetry of transport in terms of inward versus outward movement may well be inherent characteristics of transport systems of plasma membranes due to their asymmetric structure in the plasma membrane, imposed by their multiple transmembrane-spanning domain structure. The energization of the transport mechanism and the electrochemical gradient of the substrate will contribute to the overall net flow of solutes. Certain transport systems appear to prefer, or may even be locked into, either a hetero- or a homoexchange mechanism (e.g., Chillar6n et al., 1996; Zerangue and Kavanaugh, 1996b). Ion-coupled amino acid transporters may use favorable electrochemical gradients of cosubstrates (notably Na +) to move amino acids against a total chemical potential gradient by secondary active transport, utilizing the Na + gradient generated by primary active transport through the ATP-powered sodium pump. This mechanism enables several amino acids to be maintained in-

302

10. Transport and lnterorgan Nutrient Flows

side cells at a much higher total chemical potential than occurs in extracellular fluid. These transmembrane amino acid gradients may themselves be used to generate smaller gradients for other amino acids due to heteroexchange activity of amino acid transporters such as system L (Guidotti and Gazzola, 1992). For example, alanine accumulated in a cell by system A may passively exit down its concentration gradient through system L in exchange for a poor system A substrate such as leucine, enabling the latter substrate to be marginally accumulated inside the cell against its gradient (Chapter 4). Several amino acid transport systems (e.g., systems X-AG, B ~ b ~ and y+L) also show relatively low stereoselectivity for the bioactive L-forms. Certain D-amino acids (e.g., D-glutamate) are found in measurable quantities in plasma and are presumably "natural" substrates for these transporters, although under physiological circumstances they are unlikely to reach concentrations at which significant competition between L- and D-isomers will occur.

3. Regulation by Hormones and Other Exogenous Stimuli Amino acid transport activity may be modulated by the electrochemical gradient of substrate across the membrane and/or the number of operational transporters (see Chapter 9). Both of these factors may be influenced directly by the action of hormones or other stimuli. Insulin, glucagon, and corticosteroids are among hormones having effects on amino acid transport, alongside other exogenous stimuli such as nutrient supplementation and mechanical stimulation. Insulin and glucagon both affect activity of ion transport mechanisms in cell membranes, leading to changes in cell membrane potential (and hence electrochemical potential of charged substrates) that are capable of significantly altering energizing of rheogenic amino acid transport. In addition, second messengers produced within cells in response to hormones and other exogenous stimuli modulate a variety of intracellular signaling pathways that may lead to increased or decreased synthesis/breakdown of new transport proteins (i.e., altered transport capacity) or have more direct effects (e.g., through activation of signaling molecules such as protein kinase C or phosphatidylinositol 3-kinase), which, in certain cases, have very rapid effects on amino acid transport (e.g., Guidotti and Gazzola, 1992; McGivan and PastorAnglada, 1994; Low et al., 1997a). The most widespread hormone-sensitive amino acid transporter is system A, and insulin stimulates both system A and system N m in skeletal muscle (quantitatively the most important tissue for amino acid turnover). Insulin stimulation of system A in skeletal muscle cells is relatively rapid and is

not dependent on new protein synthesis (i.e., it is not blocked by inhibitors of transcription and translation). It is conceivable that the mechanism involved is analogous to that by which insulin stimulates glucose transport in muscle by recruitment of quiescent GLUT4 transporters from intracellular "stores" to the cell membrane (e.g., Tsakiridis et al., 1995). In contrast to its effects on system A, insulin stimulation of system N m in skeletal muscle requires new protein synthesis (Low et al., 1997a). Insulin also stimulates amino acid transport into other cell types (e.g., by increasing system A activity in fibroblasts) by mechanisms requiring new protein synthesis. Extremely rapid (within 1 min) responses of systems N, N m, and A to exogenous stimuli have been reported that cannot be explained simply by induced changes in cell membrane potential (Bode and Kilberg, 1991; Low et al., 1997a). Rapid swelling of liver or muscle cells results in rapid activation of system N/ N m and (at least for muscle) inactivation of system A. The opposite effects are seen after cell shrinkage and all responses (in muscle at least) are blocked by inhibition of phosphatidylinositol 3-kinase (using wortmannin) or tyrosine kinases (using genestein). The effects on system A may form part of a coordinated regulatory response to cell volume change (McGivan and PastorAnglada 1994; Low et al., 1997a), but those on system N/N m are likely to be counterregulatory and may form part of a feed-forward system for nutrient-induced modulation of cell metabolism (see Section II,F). Increased system A transport within 1 min of stimulation is also observed in lymphocytes treated with phorbol-ester, hence activating protein kinase C (Segel, 1992 for review). These rapid changes in transport activity are suggestive of a direct effect on the transport protein itself (e.g., changes in phosphorylation status) but this remains to be demonstrated directly. The activity of amino acid transport systems including A, N/N m, L, X-AG, and X-c may also be modulated during "adaptive regulation." This is a response to nutritional "stress" (amino acid starvation or replenishment) shown by many cell types, which consists of a derepression of transport activity during amino acid starvation and reversal after replenishment (Guidotti and Gazzola, 1992). These processes require de n o v o synthesis of protein (possibly of rapid-turnover repressor proteins which modulate transcription of transporter mRNA) and RNA and are generally most responsive to changes in concentration of amino acids that are substrates of the regulated transporter (Kilberg et al., 1994). Upregulation of system A transport in liver in vivo during fasting may be a physiologically important example of adaptive regulation (here in response to reduced hepatic alanine availability; see Sections IV,B, and V,B). Adaptive regulation in skeletal muscle cells in re-

Interorgan Amino Acid Nutrition sponse to glutamine deprivation or supplementation involves system N m (the principal glutamine transporter in this tissue) but also extends to system X-AG, the main supplier of the glutamine precursor glutamate (see Fig. 10.3); this process may involve coordinate regulation of glutamine transport, glutamate transport, and glutamine synthase to help maintain intramuscular glutamine concentration in the face of alterations in external supply (Low et aL, 1994). In contrast to this type of adaptive regulation, the small intestine (and other splanchnic tissues) adaptively up-regulates amino acid transport capacity in response to increased external amino acid concentration, as in, for example, a high-protein diet (Fafournoux et aL, 1983; Stevens, 1992). The physiological function of this mechanism is to maintain a "safety factor" (see Section III) between intestinal transport capacity and dietary intake of amino acids (Stevens, 1992; Diamond 1993; Van Winkle, 1999 for review). F. Amino Acids as Modulators of Metabolic Processes There are many reports of marked effects of amino acid supplementation on cellular metabolism (notably protein turnover) both in vitro (e.g. Bloomaart and Meijer, 1995; Kimball et al, 1996; Patti et aL, 1998) and in vivo (Casetellino et al, 1987; Patti et aL, 1998). Leucine, alanine, and glutamine appear to be particularly effective in modulation of protein turnover in vitro (Rennie et aL, 1989; Kimball et aL, 1996). The mechanisms underlying these effects remain poorly understood but they

~

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i~

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f = 0

.A

Gin

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T

I[~

' 5

' ' 101520

'

//

' 45

'50

' 55

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= 65

Time (h)

FIGURE 10.3 An example of adaptive regulation: Effect of Gin deprivation and Gin supplementation on glutamine and glutamate uptake in rat skeletal muscle cells. Glutamate (solid circles) and glutamine (solid triangles) uptake (0.05 mM) was measured in rat muscle cells in primary culture deprived of or resupplemented with 2.6 mM Gin at the times indicated. The figure represents the percentage change in Gln/Glu uptake relative to a control value (measured as Gln/Glu transport in Gin supplemented cells paired with Gindeprived cells over the same period). Representative values for a single preparartion (mean _ SEM for triplicate measurement) (adapted from Low et al., 1994,with permission from the Federation of American Societies for Experimental Biology).

303

include effects on gene transcription, on the abundance and activity of the protein synthetic machinery of ribosomes, and on the activity of intracellular signaling pathways. One contributing mechanism for those amino acids entering the cell by secondary active transport mechanisms (e.g., glutamine) may be linked to cell volume (H~iussinger, 1996; Low et al., 1997a). The net movement of solute into or out of a tissue represents the difference between unidirectional influx and unidirectional efflux of solutes across the cell membranes. This solute flux, unless it is associated with obligatory exchange of another solute, will also alter the osmotic activity of the cytosol. Large net solute fluxes may have sufficient effect on intracellular osmolarity to cause a significant change in intracellular volume due to osmotic reequilibration of water across the cell membrane. It is becoming increasingly apparent that such changes in cell volume are somehow sensed and transduced to intracellular signaling pathways, which modulate cell metabolism. Osmotic cell swelling (particularly in liver cells in the absorptive phase when large net, sodium-coupled uptakes of amino acids occur) generates an anabolic signal within the cell, whereas cell shrinkage produces the opposite catabolic effect (H~iussinger, 1996; Low et aL, 1997a,b). The mechanisms involved are probably analogous to those involved in the responses to cell stretch or in cell movement and allow a means by which nutrients (notably where they are concentrated into the cell by active transport processes) may influence cellular metabolism by a mechanism additional to any mass action effect. Certain Na+-coupled symport processes (e.g., the SGLT1 glucose transporter) may contribute additionally to cell volume change by acting as "water transporters" in conjunction with solute uptake (Meinild et al, 1998). The anabolic effect in both liver and skeletal muscle involves activation of the p70 s6 kinase signaling pathway via a wortmannin (i.e. PI 3-kinase)sensitive mechanism (Krause et aL, 1996; Low et aL, 1997a; see Fig. 10. 4). The initial sensor of cell volume change, at least in skeletal muscle, requires intact integrin-extracellular matrix interactions and may be initiated by integrin signaling mechanisms (Low et aL, 1997b; Fig. 10. 4). This pathway is distinct from, yet possibly associated with, the anabolic signaling pathway initiated by insulin receptors and may represent an important mechanism by which nutrients and hormones can interact for overall control of cell metabolism (Parsons, 1996; Ingber, 1997; Patti et aL, 1998). We have now established a direct link between a requirement for integrin-extracellular matrix interactions and signaling of cell volume increase to the anabolic process of glycogen synthesis (Low et al., 1997b) as well as to the rapid modulation of membrane amino acid transport activities (Low and Taylor, unpublished observations). Recent

304

10. Transport and Interorgan Nutrient Flows

uptake by pancreatic B-cells in response to elevated portal amino acid concentrations during absorption (by high Km transport processes) results in an increase in amino acid oxidation within the cells, increasing ATP production with concomitant effects on ATP-sensitive potassium channels, further resulting in cell depolarization, calcium influx, and exocytosis of insulin-containing vesicles (Docherty and Clark, 1994). G. Nutritional Status FIGURE 10.4 Cell signaling pathways involved in mediating metabolic and transport responses of skeletal muscle to cell volume change. Arrows represent activation unless indicated. The putative volume/ stretch sensor may incorporate sarcolemmal integrins linked to Gproteins and attached to both extracellular matrix and cytoskeleton. The G-protein-phosphatidylinositol 3-kinase "switch" is required to maintain activity of the sensor, which transduces signals through tyrosine kinase(s) to membrane transporters and downstream signaling cascades modulating cell metabolism. Activity of the putative "switch" may be modulated by receptor-mediated anabolic signals (e.g., insulin, growth factors), allowing interaction between parallel signaling mechanisms for overall control of metabolic processes. Alternative signaling pathways involving elements of the "switch" are shown by shaded arrows. Note the "positive feedback" loop linking solute transport, membrane stretch, and the sensor-transducer, allowing possible synergistic anabolic effects of nutrients and muscle stretch (adapted from Low et al., 1997a, with permission from the Federation of American Societies for Experimental Biology).

evidence has also shown that activation of the P70S6 kinase pathway modulates the activity of a number of proteins involved in modulation of protein synthesis (both translation initiation and elongation factors) (Proud and Denton, 1997). We are therefore now in a position to produce a verifiable model mechanism to explain the known anabolic effects of glutamine in promoting muscle protein synthesis, a phenomenon of proven clinical relevance (see Section V) but as yet of largely unknown origin. Transport of one amino acid may have effects on the metabolism of others. For example, uptake of Lglutamate in the kidney increases intracellular glutamate concentration and represses phosphate-dependent glutaminase, reducing glutamine breakdown at its first committed step (see Section V,D). Equally, competition between substrates for transport (e.g., at the blood-brain barrier) will affect delivery of individual substrates to a metabolic compartment (e.g., delivery of tryptophan to the brain for neurotransmitter synthesis; see Section IV,F). Other less-direct effects of amino acids on metabolism arise from their influence on insulin secretion. A number of amino acids (notably leucine) act as insulin secretagogs in the following manner: an increase in their

Interorgan fluxes are also dependent on the nutritional state of the animal and in many cases these fluxes are reversed in the fed and postabsorptive (typically overnight fasted) states. The metabolic changes associated with prolonged fasting are considered in Section V,B. The anabolic or fed state is characterized by an increase in plasma nutrient concentrations. Hepatic portal amino acid concentrations of amino acids may increase two- to three-fold, but rapid hepatic uptake and metabolism (largely oxidation) prevents these large increases from reaching the arterial circulation (Biolo et al., 1992b; Millward et al, 1996), although smaller systemic increases do occur (see Fig. 10.5). Increased cellular uptake of amino acids in the fed state results in transient increases in AA pool sizes in liver and skeletal muscle (see Fig. 10.5). Increased nutrient uptake into cells makes them swell and also tends to stimulate anabolic processes. Protein (especially in meat) is rich in branched-chain and aromatic amino acids, which in excess are toxic. A major function of interorgan nutrition is to provide cells with sufficient amounts of these largely nondispensable substrates for protein synthesis while preventing their build up, particularly as a result of the protein surplus in most Western diets (Waterlow, 1995; Millward et al., 1996). Protein is hydrolyzed to amino acids plus di- and tripeptides in the small intestine. The epithelial cells lining the small intestine are the first to see these products of macronutrient digestion in the gut lumen and are largely responsible for their absorption into the hepatic portal blood. A high proportion of hydrolyzed protein enters the intestinal enterocytes as small-molecularweight peptides, but most of these are hydrolyzed within the epithelial cells and are either utilized locally or exit across the basolateral membrane as their constituent amino acids (Stevens, 1992). The secondary active transport processes in the brush border membrane of the intestine include both sodium-coupled processes for amino acids and hydrogen ion coupled processes for small-molecular-weight peptides. The mammalian liver extracts a fair proportion of portal amino acid load at first pass in the fed state. Unidirectional fractional hepatic extractions (UFEs) of portal amino acids by rat

lnterorgan Amino Acid Nutrition

A Blood

~

Muscle

A

~ IW

A qW

t

I

--B-- Liver

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O = E

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Time after feeding (h) FIGURE 10.5 Effects of feeding on amino acid concentrations in rat tissues. Female Wistar rats were fasted for 60 hr then given a bolus protein-rich meal (0.5 g protein/100 g body weight). Tissues were isolated at timed intervals after feeding and samples were prepared for amino acid analysis by a Biotronik amino acid analyser. Results are the means _+ SEM for n - 3-8 rats (unpublished observations of P. M. Taylor and M. J. Rennie). (A) Blood, liver, and skeletal muscle amino acid concentrations. Values for liver and skeletal muscle refer to intracellular A A concentration obtained after correction for extracellular amino acids. Arterial blood values are given. (B) Hepatic portal and arterial blood amino acid concentrations.

liver vary from about 30 to 70%, depending on the properties of hepatic sinusoidal amino acid transporters (Table 10.2), but net extraction (NFE) is generally lower because intracellular metabolic activity is exceeded by uptake and amino acids rapidly exchange across the liver membrane. Amino acids are deaminated and oxidized as fuel for hepatic processes and the excess amino groups are excreted, largely in the form of urea. They may alternatively be interconverted into other amino acids or used as gluconeogenic substrates. The concentrations of organic nutrients in the hepatic portal vein

305

therefore exceed those entering the systemic circulation in the fed state. The lungs may act as a secondary tissue barrier to excess nutrients and potential xenobiotics entering the body through the digestive tract, because hepatic venous blood passes through the pulmonary circulation before entering the systemic circulation. Feeding elicits a postprandial suppression of muscle protein breakdown sufficient to minimize increases in blood amino acid concentrations, preventing excess amino acid oxidation (Millward et al., 1996). The liver plays a large and important role in determining the concentrations of blood amino acids, with the possible exception of the branched chain amino acids (Biolo et aL, 1992b; Millward et aL, 1996). Conversely, in the postabsorptive state, the liver may release amino acids and glucose produced from net breakdown of protein and glycogen, respectively, in which case hepatic venous solute concentrations will be greater than those entering via the portal vein and, to a lesser extent, the hepatic artery. In the postabsorptive state, release of alanine and glutamine from skeletal muscle accounts for over half the total net amino acid released from this tissue. This release is matched by net extraction of these amino acids in the splanchnic bed (mainly liver and small intestine) (Cahill, 1976, Biolo et aL, 1995b; Nurjhan et aL, 1995). Within the splanchnic circulation, glutamine is extracted by the small intestine but there is further release of alanine into the portal circulation so that the net uptake of alanine by the liver is larger than the observed arterial-hepatic vein difference (Windmueller and Spaeth, 1974; Buttrose et aL, 1987; see also Section IV). This intrasplanchnic pattern of amino acid movement is shown in man, rat, and dog. H. The Contribution of Protein Turnover Cells have an underlying requirement of amino acids for protein synthesis and they also generate free amino acids from protein breakdown. A fundamental function of amino acid transport systems in cell membranes is to provide routes of entry and exit for amino acids required or produced in cellular protein turnover. There are often differences between cell types in their requirement for individual amino acids for protein synthesis: for example, fibroblasts make collagen rich in glutaminederived prolyl residues and hepatocytes export proteins poor in BCAA residues (Waterlow et al., 1978; Millward et al., 1996). The human body contains about 11.5 kg of protein (typical value for a 70-kg man), which turns over at about 300 to 400 g/day (i.e. on the order of 3% per day; Rennie et al., 1994; Millward et al., 1996). Assuming a molecular weight of about 100 for amino acids in pro-

306

10. Transport and lnterorgan Nutrient Flows TABLE 10.2 Extraction of Amino Acids by Liver of Fasted and Refed Rats in Relation to Prevailing Amino Acid Concentrations in Hepatic Portal Vein (HPV)a NFE c

[Amino acid] HPV (mM) Amino acid

Fasted

Fed

UFE n

Fasted

Fed

Alanine Glutamine Glutamate Leucine Phenylalanine Lysine

0.41 ___0.03 0.40 ___0.01 0.19 ___0.01 0.12 ___0.01 0.06 ___0.01 0.53 +_ 0.02

1.17 ___0.10 0.66 +_ 0.07 0.27 ___0.04 0.56 ___0.06 0.22 _+ 0.02 1.13 ___0.12

0.39 ___0.03 0.70 ___0.08 0.37 +__0.06 0.48 ___0.03 0.50 _ 0.05 0.36 +__0.10

0.33 ___0.04 -0.27 ___0.04 -0.14 ___0.04 -0.10 ___0.06 -0.06 ___0.07 0.01 ___0.04

0.40 +__0.03 -0.05 ___0.10 0.16 ___0.05 0.36 ___0.03 0.23 +__0.04 0.25 ___0.04

aFasted and fed refer to rats treated as described in the legend to Fig 10.5. Table reproduced from Taylor and Rennie (1986) with permission from Portland Press Ltd. bUFE is unidirectional fraction extraction of tracer amino acid measured in peffused rat liver at physiological perfusate flow rates. CNFE is net fractional extraction of amino acid in vivo by liver of anesthetised rats, estimated as the difference between hepatic portal and hepatic venous [amino acid] divided by the portal concentration.

tein, we estimate that roughly 3.5 moles of amino acids per day exchange between protein and free pools due to whole-body protein turnover (rates for individual amino acids will reflect their relative abundance in body protein). The whole-body free amino acid pool (about 560 mmoles; see Table 10.3) therefore turns over about six times per day. Different proteins turn over at different rates and fractional rates of protein synthesis vary between different h u m a n tissues, being about 42% per day for kidney, 12% per day overall for splanchnic tissue, and 1.5% per day for muscle (Rennie et aL, 1994; Tessari et al., 1996). Certain rapidly dividing cell types, e.g., the intestinal epithelium, may turn over protein at a rate greater than 50% per day (Rennie et al., 1994; Millward et aL, 1996). Rapid cell division during growth and the mounting of the i m m u n e response involve increased rates of protein turnover in the dividing cells. The immediate source of amino acids for cellular protein synthesis is the free cytosolic pool, which receives amino acid by transport into the cell from the extracellular fluid and also from protein b r e a k d o w n (Waterlow et aL, 1978; Biolo et al., 1992a,b). Cellular supply of amino acids

TABLE 10.3

Intracellular fluid Extracellular fluid Total FAA pool

Estimated Whole Body Free Amino Acid Pools in a 70-kg-Man Total AA concentration (mmol/iiter)

Fluid volume (liter/70 kg man)

Whole body content (mmol)

18a 4 m

28 14 m

504 56 560

aEstimate based on average of total AA concentration in skeletal muscle, liver and intestine weighted for relative tissue volumes (calculated using information obtained from Waterlow et al., 1978).

from the plasma is d e p e n d e n t both on plasma flow and blood amino acid concentration in that particular region of the body (Table 10.4). The ultimate source of amino acids is dietary protein. The functional m i n i m u m requirement for protein in the diet is reported to be about 0.4g/kg/day (28 g per day for a 70-kg man), whereas Western diets typically include about 100 g/day protein (Waterlow, 1995; Millward et al., 1996). Protein oxidation represents about 12% of resting energy requirements but this decreases when total energy expenditure increases during exercise (Felig, 1975; Biolo et al., 1995a). One means of amino acid appearance into the cytosol (Table 10.5) is by protein breakdown. B r e a k d o w n of long-lived proteins occurs predominantly by autophagy in the lysosomes ( B l o m m a a r t et al., 1997), whereas extralysosomal proteolytic pathways (including the ubiq u i t i n - p r o t e a s o m e system) are responsible for the degradation of proteins with high turnover rates (Pickart, 1997). A n exception is found in skeletal muscle, where degradation of the slow-turnover myofibrillar proteins is normally initiated outside the lysosome because the large size of the myofibrillar apparatus precludes autophagy (Price and Mitch, 1998). M a c r o a u t o p h a g y is responsible for acceleration of proteolysis in many cell types when amino acid and insulin concentrations fall. M a c r o a u t o p h a g y in the absence of amino acids can reach values as high as 4 to 5% protein lost per hour in liver cells. A m i n o acids are effective inhibitors of the m a c r o a u t o p h a g y process, which is also inhibited by insulin and stimulated by glucagon ( B l o m m a a r t et al., 1997). This inhibition by amino acids primarily occurs at the initial ATPd e p e n d e n t sequestration step in which cytosolic material for breakdown becomes m e m b r a n e - b o u n d prior to fusion with existing lysosomes. Not all amino acids are

307

Interorgan Amino Acid Nutrition TABLE 10.4

A m i n o Acid Concentrations in Plasma, Skeletal Muscle, and Protein a

Plasma Amino acid

(pM)

Intracellular (ic) (pmoi.liter.ic water -1)

Distribution ratio

Protein (pmol.g -1)

Muscle protein (M %)

Normal fasting release (M%)

Alanine Glutamate Glutamine Valine Leucine Isoleucine Tyrosine Phenylalanine Methionine Glycine Proline Lysine Threonine Serine Arginine Histidine Aspartate Asparagine

419 40 761 271 104 67 97 69 29 303 231 212 172 153 99 104 13 58

3096 3800 21020 324 236 106 115 76 52 1131 995 1086 807 916 719 459 1746 405

7.4 95 27.6 0.8 2.3 1.6 1.2 1.1 1.8 3.7 4.3 5.1 4.7 6.0 7.3 4.4 134.3 7.0

120 189 incl. amide 84 116 59 29 45 25 134 73 112 75 70 60 34 125 incl. amide

8.7 13.7

31.5 9.6

6.1 8.4 4.3 2.1 3.3 1.8 9.7 5.3 8.1 5.4 5.0 4.3 2.5 9.0

4.9 3.4 1.5 2.6 2.5 1.5 9.6 6.9 8.7 6.1 0.4 2.9 3.7 3.9

aAdapted from Mackenzie et al. (1992) with permission from Plenum Publishing Corp.

equally effective as inhibitors of macroautophagy and in the liver it appears that leucine, phenylalanine, and tyrosine, in combination with alanine and glutamine, are the most important amino acids involved in control of autophagic proteolysis (Mortimore et al., 1988; Blommaart and Meijer, 1995). Leucine and glutamine are also implicated as regulators of proteolysis in the skeletal muscle although the extent to which this regulation is intra- and extralysosomal is, as yet, unclear. The contribution of extralysosomal proteolytic systems to protein degradation in muscle is more important than in liver

TABLE 10.5 Whole-Body Amino Acid Turnover in Postabsorptive Human Subjects

Amino acid

Appearance (pmoles.kg-l.hr -a)

Oxidation (pmoles.kg-l.hr -1)

Alanine

264-295

Arginine Glutamine

65 258-345

147-179

Glutamate Leucine

120 77-147

18-19

Phenylalanine

44-55

Subject a Adult (a,b) Adult (c) Adult (b,d,e) Adult (e) Adult (f,g,h) Adult (e,h)

aData obtained from (a) Carraro et al. (1994); (b) Nurjhan et aL (1995); (c) C a s t i l l o e t a l . (1994); (d) Hankard etal. (1995); (e) Matthews and Campbell (1992); (f) De-Feo et al. (1994); (g) Hankard et al. (1996); (h) Biolo et al. (1992b).

and accelerated breakdown of myofibrillar protein, e.g., during starvation, occurs via the ubiquitin-proteasome system (Price and Mitch, 1998). Swelling of liver cells has an inhibitory effect on autophagic protein degradation and also increases the sensitivity of the process to inhibition by amino acids (Blommaart et al., 1997). Alanine and glutamine may both exert their antiproteolytic effect (at least in the liver) via cell swelling, whereas leucine, tyrosine, and phenylalanine, which are not transported by sodiumcoupled mechanisms and do not accumulate inside cells, probably specifically exert their effects at the macroautophagic sequestration step by mechanisms that are independent of cell swelling. The dual mechanisms can account for the synergism between glutamine, alanine, and leucine in inhibition of hepatic proteolysis (Mortimore et al., 1988; Blommaart and Meijer, 1995). The rapid turnover of the jejunal mucosa is reflected by a fractional protein synthesis rate of greater than 50% per day (Rennie et aL, 1994; Waterlow, 1995). In the postabsorptive state, the small intestine relies mostly on amino acid supply from the blood stream and the release of indispensable amino acids from muscle (and also possibly from the liver) helps to maintain gut protein synthesis during this period (Felig, 1975; Waterlow et al., 1978). A recent study by Wolfe and colleagues on postabsorptive dogs (Biolo et al., 1995b) has revealed that intracellular amino acids released during protein breakdown in muscle appear to be preferentially released into the blood stream

308

10. Transport and lnterorgan Nutrient Flows

rather than reincorporated into protein. In contrast, in the intestine a higher proportion of amino acids were recycled into protein synthesis with hypothesized gains in overall efficiency. This study also indicates that differences in transmembrane transport kinetics between small intestine and muscle contribute to modulation of net amino acid transfer between the two tissues. Combined modulation of transport and metabolism is probably the usual situation. For example, rates of protein turnover and amino acid transport in skeletal muscle are both increased after exercise. This combination of effects results in a reduction of net muscle protein catabolism in dogs (Biolo et al., 1995b), although increased net synthesis of muscle protein also requires an increased supply of plasma amino acids. The importance of such increases in transport become obvious in their absence; when an increase in muscle protein turnover is unaccompanied by increased amino acid transport (e.g., during severe illness) and muscle protein synthesis is consequently not stimulated to the same extent as breakdown, net muscle catabolism is the result (Waterlow, 1995; Rooyackers and Nair, 1997; Price and Mitch, 1998). I. Modes of Carriage of Amino Acids in the Blood Amino acids are carried in the blood in free solution, bound reversibly to plasma proteins, or within blood cells (largely erythrocytes). In general, changes in the concentration of plasma amino acids are not followed by instantaneous net movement of amino acids between red cells and plasma. Nevertheless red cells will contribute to carriage of amino acids between tissues, although their relative importance will depend on the amino acid carried and the transit time across the tissue bed (Felig et al., 1973; Christensen, 1990). The anionic amino acids glutamate and aspartate show the highest erythrocyte/ plasma distribution ratio and they are most likely to have substantial interorgan carriage by blood cells. Blood cells may also have increased importance in interorgan AA exchanges involving tissues where significant extracellular metabolism of circulating amino acids occurs (e.g., in vasculature of the kidney) (Schwegler et al., 1992; see also Section V,D). The role of erythrocytes in amino acid carriage between tissues may be enhanced during starvation (Pico et al., 1994). The aromatic amino acids (especially tryptophan and its thyroid hormone derivatives) are the only amino acids that bind substantially to plasma proteins, and over 70% of plasma tryptophan may be bound to serum albumin (Smith and Cooper, 1992).

111. CONTROL OF INTERORGAN AMINO ACID METABOLISM: METABOLIC CONTROL THEORY AND SAFETY FACTORS Interorgan amino acid nutrition is, broadly speaking, a homeostatic phenomenon but the relative constancy of plasma concentrations of amino acids may be sacrificed under certain circumstances (see e.g., Sections V,B and V,D) to maintain the constancy of a flow between tissues rather than the reverse flow (the term homeorhysis has been applied to describe this effect; Christensen, 1990). Cycling of substrates is a common motif in metabolic pathways and such cycles may be intracellular (e.g., glycogen turnover in liver and triacylglycerol turnover in adipocytes; Frayn, 1996), intercellular (e.g., the glutamine "cycle" between periportal and perivenous hepatocytes; see Section IV,A), or interorgan (e.g., the glucose-alanine cycle; see Section IV,B). A physiological nutrient load imposed on a metabolic pathway meets a series of functional capacities (i.e., enzyme and transporter activities) and the ratios of these capacities provide some indication as to the importance of individual steps in control of the pathway (i.e., low capacities are likely to exert the greater degree of control). Within a cell or cell compartment the major control over metabolic pathways is exerted by the quantity or activity of an enzyme, and the first nonequilibrium reaction in a pathway (the committed step) is usually an important control element (Cornish-Bowden and Cardenas, 1990; Frayn, 1996). Transport across the plasma membrane has often been overlooked as a site for control of a metabolic pathway and there is an unjustified tacit assumption that transporters have equilibrium constants of unity and are therefore unlikely candidates as regulators. The integrated homeostatic behavior of metabolic processes in complex systems may be described in quantitative terms by Control Theory (Cornish-Bowden and Cardenas, 1990; Frayn, 1996, for review). This approach to understanding metabolic control in living systems (at least under well-defined steadystate conditions) involves study of the whole system as well as the properties of individual system parts (e.g., a specific enzyme) (see Fig. 10.6). For studies of the whole system, the effects of changing the activity of a single component enzyme or transporter on the total system flux may be examined. Here the influence of individual components is described by the flux control coefficient of that component (C• which is proportional to the overall change in system flux divided by the change in activity of a particular enzyme or transporter under study. The sum of these coefficients for all system components is unity. Studies of system parts consider each component of the metabolic pathway in isolation

309

Control of lnterorgan Amino Acid Metabolism Extracellular

Metabolic Control Coefficients (1) WHOLE SYSTEM For the system:-

A ~l

IS

~B ~-

His

-~ C ~$2

membrane

-~ D $3

j

s~ transporter S2 $3 enzymes

,~ Systemflux A ~

D

C x ot AJ / A activity of step Sx (NB: Z Cx = 1) C x = FLUX CONTROL COEFFICIENT OF Sx 9 For external effectors (eg hormones, inhibitors) which act on component reactions of a system:Reflector 0~ AJ [ A[effector] (Reflector = RESPONSE COEFFICIENT)

(2) SYSTEM PARTS 9Study each enzyme / transport step in isolation. Measure response in reaction rate (AVx) to change in substrate / effector concentration (A[effector]):Exeflect~ G~ AV x [ A[effector] (E,,eflect~ = ELASTICITY COEFFICIENT)

(3) INHIBITOR STUDIES C x = Reflector [ Exeffect~

FIGURE 10.6 Calculation and interpretation of metabolic control coefficients. See text for further details.

and, for each enzyme or transporter, the response in reaction rate to change in concentration of a substrate or inhibitor (the effector) can be defined in terms of an elasticity coefficient (Exeffect~ which is proportional to the change in reaction rate divided by the change in effector concentration. The influence of external effectors on the system flux can be described by a modified flux control coefficient termed the response coefficient (Reflector), which is proportional to the change in system flux divided by the change in concentration of the effector. The control coefficient of an individual step in a metabolic pathway can be estimated from the response coefficient of an effector and the elasticity coefficient of this effector for the step under investigation (e.g., Salter et aL, 1986) typically using as effectors either a step-specific inhibitor or a hormone known to induce the enzyme or transporter catalyzing a particular metabolic step. Using these types of approach the control coefficients for the various component steps in hepatic metabolism of aromatic amino acids and glutamine (Salter et al., 1986; Low et aL, 1993) have been established using isolated liver cells (Fig. 10.7). Care must be taken to ensure the selectivity of the inhibitors used under the experimental conditions (in these studies histidine, tryptophan, and

Cytosolic

/

protein

+ ',,_ / Na

/

9 ",, [ .. Na+ ,.,,,..L/_......~ 3.44

Gin

,,

I ~,~

/0.32 / v.

~/r

0.51Gls 2.96 mM

T - .... 1-2"54

rp

Trp

1_0.46 A ~,~......~ Trp 075 " i / \/ '~~ (0.25) 0.25

(0.75)

~

~ /

/

@/ \ \ ~

Mitochondrial 0.96

2-OG

~- Gin 1.22 ~ G1u / / i ;GUz

-~ kynurenine \ ~\ seriesofstepsto NADP coenzymes

A : Tryptophan2,3-deoxygenase

FIGURE 10.7 Pathways of glutamine and tryptophan catabolism in liver, showing experimentally derived fluxes in incubated rat liver cells. N and L represent systems N and L for amino acid transport across the plasma membrane. Glutaminase catalyses the conversion of Gln to Glu; 3H from [2-3H]GIn is lost as 3H20 during the conversion of Glu to 2-oxo-glutarate (2-OG) by transamination or action of glutamate dehydrogenase. GHz, glutamate-y-hydrazide. Numbers in normal type are the net rates of specific reactions, expressed as nmol/ min.mg dry weight of cells. Numbers in italics are the experimentally determined flux control coefficients for Gln and Trp metabolism. This diagram also shows the key steps in Trp metabolism (see lower portion of diagram). The italic numbers in parentheses show the change in flux control coefficients for Trp metabolism in liver cells from rats pretreated with dexamethasone.

glutamate y-hydrazide were used to inhibit system N, system L, and glutaminase respectively at concentrations which were without appreciable effect on other processes influencing glutamine metabolism). Certain general assumptions have been made regarding transport mechanisms when investigating these metabolic control structures, specifically (1) that forward and reverse transport rates are independent, (2) the values of kinetic parameters (Kin, Vmax)of the transport systems are the same on both sides of the plasma membrane, and (3) the equilibrium constant of a nonconcentrative transport process is unity. The calculated Cx values shown in Fig. 10.7 indicate that control of hepatic catabolism of both glutamine and tryptophan is shared largely between membrane transport and the first step of intracellular metabolism, establishing the importance of membrane transport as a control element for amino acid metabolism (at least in these specific circumstances), particularly under conditions in which there is induction of intracellular enzyme activities. The results relating to glutamine metabolism also highlight the contrasting effects of concentrative and dissipative transport mechanisms (here systems N and L respectively), which have control coefficients of oppo-

310

10. Transport and lnterorgan Nutrient Flows

site sign (the negative control coefficient for system L arises because the net glutamine flux through this transporter is out of the liver cell). The simultaneous activity of glutamine uptake and release mechanisms in the liver sinusoidal membrane may contribute to development of a translocation cycle between extracellular and intracellular glutamine pools (H~iussinger, 1990; Newsholme and Parry-Billings, 1990; Low et al., 1993). Such an arrangement facilitates fine regulation of cytoplasmic amino acid concentration with the retention of sensitivity to fluctuations in extracellular concentration to enable net hepatic glutamine flux to rapidly shift from release to uptake (e.g., as portal delivery increases after a meal; see Fig. 10.8) or vice versa (Taylor and Rennie, 1986; H~iussinger, 1990). Net hepatic extraction of several amino acids representative of the major types is less than unidirectional extraction (see Table 10.2), suggesting the existence of several translocation/exchange processes that could be responsible for the observed rapid increase in net extraction as portal amino acid

FIGURE 10.8 Hepatic glutamine and alanine fluxes and liver/ plasma amino acid concentrations during fasting and refeeding. Data refer to anesthetized Wistar rats and experimental protocols were as described in the legend to Fig. 10.5. Results are presented as the mean values for 5-8 rats (unpublished observations of P. M. Taylor and M. J. Rennie). Hepatic portal blood flow was measured using a 2-mm ultrasonic flow probe (Transonic Systems) and hepatic arterial blood flow was measured using radioactive microspheres (15/xm diameter). Net hepatic amino acid flux was estimated as the product of hepatic blood flow and hepatic influent-effluent blood [AA] difference. P, postabsorptive rats (overnight fast); F, 60-hr fasted rats; R, refed rats (sampled 0.75 hr after a bolus protein meal following 60 hr fast). A high liver:blood glutamine distribution ratio is maintained in all nutritional states, although liver glutamine exchange switches from net release to net uptake after refeeding. In contrast, the liver takes up Ala from blood in all nutritional states and the alanine distribution ratio collapses upon prolonged fasting as a result of increased hepatic Ala utilization (despite adaptive up-regulation of hepatic System A transport; note increase in Ala influx on proceeding from postabsorptive to fasted state). These observations are suggestive of a high degree of control for Ala transport in hepatic Ala utilization.

concentrations rise in the fed state. The notable exception to this generality is alanine, where unidirectional and net hepatic extraction are matched over a wide range of plasma concentrations, reflecting the major importance of catabolism of this amino acid to hepatic metabolism (see Table 10.2 and Fig. 10.8). Note that the range of transport mechanisms available to large zwitterionic (branched-chain and aromatic) or cationic amino acids is more restricted than for glutamine, such that net flux may be determined largely by the exchange properties of a single transport mechanism (e.g., system L for leucine) rather than by the difference in flux between two opposing mechanisms. Membrane transport may be an important step for control of interorgan and intercellular metabolic pathways as well as intracellular metabolism, although the situation is complicated by the fact that the transporter and enzyme complements in the different cell types are likely to be different. For example, alanine released across the basolateral membrane of the small intestine by the dissipative L and y+L-like systems is extracted from the portal plasma by hepatic system A, which is (at best) poorly expressed at the basolateral face of the intestine. In contrast, these same L and y+L-like systems in the intestine are likely to be the major routes of net glutamine uptake from blood because of the Na +dependence of system y+L for glutamine transport and the low intracellular glutamine concentration of intestinal epithelial cells (i.e. favorable mass-action ratio for glutamine uptake) arising from the high activity of intestinal glutaminase (see Sections IV,A and IV,B). Proposed transmembrane translocation processes (e.g., those related to glutamine-glutamate cycling between periportal and perivenous liver cells; H~iussinger, 1990) may be more complex than initially proposed in that the opposing transmembrane metabolite fluxes can occur in different cell types (e.g., glutamine uptake into periportal cells and release from perivenous cells); note that the control structure established for hepatic glutamine metabolism in Fig. 10.7 is related to isolated hepatocytes (predominantly periportal cells). The potential importance of particular membrane transport steps in modulation of intra- or interorgan metabolism may be assessed after establishing a number of factors including: 1. Km, Vmaxvalues of the transport system. 2. The concentrations of substrates at both faces of the membrane and hence both the extent of transporter saturation and the sizes and directions of substrate gradients. 3. The electrochemical potential of cosubstrates such as sodium ions and of anionic and cationic amino acids themselves. 4. The effect of changes in substrate gradients and concentrations on both unidirectional and net fluxes through a transport step.

Physiologically Important Flows of Amino Acids

5. Both short- and long-term effects of hormones and nutrients on transporter activity and expression. A transporter operating at somewhere near maximal capacity may provide a vital flux of substrate, but the ability to regulate that flux will depend on the extent to which the transport protein itself can be modulated in activity or abundance. In contrast, the flux through a transporter, which operates far from saturation (i.e., the Km for substrate is higher than the ambient substrate concentration), will be in direct proportion to the substrate concentration so that increasing substrate concentration will increase the flux and vice versa (see Fig. 10.9). These two distinct modes of operation of transporters may both be important under different circumstances. In the former case, the transporter itself appears to be the key element in the regulation, whereas in the latter situation the properties of the transporter are such that the processes of substrate delivery (e.g., from metabolism or blood circulation) largely dictate the magnitude of flux across the membrane. The question of the quantitative match between physiological capacities and loads is important in interorgan nutrition. For instance, how is the gma x of a transporter or enzyme related to the actual range of velocities under natural conditions? The physiological (and evolutionary) advantages of having "spare" capacity must be weighed against the associated maintenance costs (Diamond, 1993). Evidence from a variety of animal Feeding

>,, |

8F

Kmi 2 mM; Vmax 15

~6

"~ 5 ~#

,'~

" ~ < 1 2 ~ 0.

O0

0.50

!

_ _

organ systems (musculoskeletal, respiratory, alimentary) indicates that physiological systems maintain "safety factors" (i.e., the ratio, capacity/maximal load for the system), which are usually of the order of 2 (i.e., capacity is twice load). The total transport capacity for amino acids in the small intestine appears to be about twice that delivered by luminal digestion of dietary protein (similar results are obtained when considering carbohydrate/sugar). This relationship holds over a variety of species with different diets and can be shown experimentally to be maintained when dietary intake is increased or decreased, at least within certain limits (see Diamond, 1993, for review). A notable exception is for arginine transport in the cat intestine, where the safety factor approaches 10 (Buddington and Diamond, 1992). Arginine is a "hyperessential" nutrient for cats, which require dietary arginine to generate sufficient urea synthesis to cope with the high protein load in their carnivorous diet, and it is thought that this high safety factor helps to ensure that absorption of arginine is high relative to other amino acids (Diamond, 1993). There is major debate (see, e.g., Cornish-Bowden and Cardenas, 1990; Frayn, 1996) as to how sequential differences in series capacities may be used to modulate metabolic fluxes, but at least in the case of hydrolysis-absorption of sucrose in intact intestine it appears that the sequential steps (sucrose, glucose transport) are closely matched with a similar safety factor (Diamond, 1993). The evolutionary selection pressure on maintenance of relatively low safety factors is not always immediately obvious because the energetic costs of maintaining extra copies of enzymes and transporters is unlikely to represent a significant burden on the overall energy budget. One suggestion (Diamond, 1993) is that "competition" between nonstructural proteins simply for space within cells (and conceivably within membranes) prevents unnecessarily large build-up of individual protein species.

_+1insulin

KmiO.i mM----~,Vma-----x2.5 1.00

31 1

I

_

1.50

I

2.00

Amino acid concentration: (mM) FIGURE 10.9 Comparison of the relationship between plasma AA concentration and AA influx for transport systems with Km values above and below normal plasma A A concentrations. In this scheme, plasma A A concentration increases from 0.45 to 1.15 mM on going from postabsorptive to fed state (the change observed for Ala in experiments summarized in Fig. 10.5). For the transport system with Km of 2 mM, AA influx increases in direct proportion to plasma [AA] within this "physiological range," whereas for the system with Km of 0.1 mM there is little increase in AA influx over the same [AA] range because the system is close to saturation. Amino acid influx through this latter system may be increased physiologically by stimulation of transport activity (e.g., insulin stimulation of transport Vmax).

IV. PHYSIOLOGICALLY IMPORTANT FLOWS OF AMINO ACIDS AND RELATED COMPOUNDS The major net interorgan fluxes of alanine and of glutamine are summarized in Fig. 10.2. Certain intraorgan fluxes of amino acids are also very important for determining net metabolic flows; for example, within the liver and the brain (as outlined in Sections IV,A and IV,C). Skeletal muscle acts as the largest store of amino acids (both free in cytosol and bound in proteins) in the human body (see Table 10.4). Glutamine and alanine in particular are maintained with a high distribution ratio across the sarcolemmal membrane, yet are released into the bloodstream (Mackenzie et al., 1992;

312

10. Transport and Interorgan Nutrient Flows

Rennie et al., 1996). Other tissues such as lung (Welbourne, 1988; Plumley et al., 1990) and adipose (Frayn et al., 1991; Kowalski et al., 1997) are increasingly recognized to play a significant role in overall amino acid economy of the body, at least in terms of exchange of nonessential amino acids such as glutamine (Curthoys and Watford, 1995). We briefly consider physiologically important flows on the basis of individual amino acids. A. Glutamine Glutamine is the most abundant c~-amino acid in the body, representing over 60% of the total free amino acid pool (some 5 mmol glutamine/kg body weight). It is the main vehicle for the transfer of nitrogen between organs ultimately for excretion as urea or ammonia (Abumrad et al., 1989; Christensen, 1990; Neu et al., 1996) and it is an important intermediate in determining the distribution of nitrogen between urea, ammonia, and amino acids within the splanchnic bed (H~iussinger, 1990; Souba, 1991; Curthoys and Watford, 1995). Glutamine also serves as the major respiratory substrate in rapidly dividing cells (e.g., enterocytes, thymocytes, macrophages, lymphocytes) and it is a biosynthetic precursor for purine and pyrimidine synthesis (Newsholme and Parry-Billings, 1990; Souba, 1991; Neu et a/.,1996). In addition, glutamine has an important role in amino acid turnover within the central nervous system (see Section IV,C). The plasma glutamine pool turns over relatively rapidly at about 0.3 mmol/kg/hr in man (see Table 10.5), equivalent to 150% per day. A small portion of dietary glutamine is absorbed intact across the gut wall, but most of the free glutamine pool in the body results from de n o v o synthesis. Glutamine usually has the highest concentration of any amino acid in plasma (ranging between 0.5 and 0.8 mM) and reaches concentrations of up to 20 mM in skeletal muscle cells (see Table 10.4). System N and the muscle-specific variant system N m are high-capacity, Na+-dependent processes for glutamine accumulation in liver and muscle respectively (Christensen, 1990, for review). These systems select for glutamine, asparagine, and histidine (i.e., zwitterionic amino acids with a side-chain N (other than tryptophan)) substrates over other amino acids and they exhibit strong regulation by factors such as cell volume change (system N m is also stimulated by insulin) (Low et al., 1997a). These characteristics are important for modulating glutamine exchange across the liver and muscle membranes and hence for regulation of interorgan nitrogen flux. In the postabsorptive state, glutamine is released from muscle, thereby supplying fuel for the stomach, small intestine, and cells of the immune system. Glutamine and alanine together provide up to 80% of the

amino acid released from muscle owing largely to their de n o v o synthesis for removal of ammonium ion excess in the muscle (Cahill, 1976; Abumrad et al., 1989: Chris-

tensen, 1990, for review). Glutamine has an anabolic effect on whole-body and skeletal muscle protein turnover, increasing protein synthesis and (at least in muscle) inhibiting protein breakdown (Rennie et al., 1989, 1996). These effects appear to be relatively specific to glutamine and may be at least partly attributed to osmotic effects of glutamine on cell volume (see Sections II,F and V,D). The rate of muscle protein synthesis is closely correlated with the free glutamine concentration, and muscle glutamine concentration is in turn dependent on the activity of the system N m and glutamine synthetase. There are important interactions between interorgan flows and metabolism of glutamine and ammonia. Ammonia is produced in vivo by amino acid catabolism and bacterial degradation of urea in the large intestine (Wahren, 1988; H~iussinger, 1990). Under normal circumstances, hepatic urea synthesis is the primary mechanism by which the body accomplishes terminal disposal of ammonia, which is toxic to the central nervous system (H~iussinger, 1990; Watford, 1993). A large fraction of the ammonia entering the liver is produced through glutamine degradation in the intestine, but this is almost completely cleared from the plasma during first passage through the hepatic circulation (Buttrose et al., 1987; Souba, 1991). The liver synthesises both urea and glutamine from ammonia in periportal and perivenous hepatocytes respectively (Fig. 10.10). The functional organization of the liver acinus dictates that changes in flux through the urea cycle will have a major influence on substrate (ammonia, glutamate) supply for perivenous glutamine synthesis. Glutamine synthetase and glutaminase are simultaneously active in the liver in vivo (H~iussinger, 1990). Hepatic glutaminase is present in the mitochondria of the periportal cells and appears to act as an amplification system to help supply enough ammonia to activate carbamoylphosphate synthetase (H~iussinger, 1990; Curthoys and Watford, 1995). The activity of this latter enzyme is a major determinant of the rate of urea synthesis under most physiological circumstances (H~iussinger, 1990; Meijer et al., 1990). Hepatic (but not renal) glutaminase is also activated by ammonia, the end product, and indeed the enzyme activity may have an absolute requirement for ammonia. Ammonia resulting from hepatic glutaminase action is incorporated into urea (possibly by direct channeling to carbamoylphosphate synthetase) and much of the additional free ammonia used by this enzyme may derive from intestinal glutamine metabolism (Watford, 1993). The high affinity of glutamine synthetase for ammonium (Kin ~ 0.1 mM) in the distal population of

313

Physiologically Important Flows of Amino Acids

perivenous hepatocyte

periportal hepatocyte cytosol +

cytosol mitochondrion

.~~NH4+,N~ps Gin

~+ I

NH4+ ~

mitochondrion

1

Glu ~ C b m - P /-CPS

1

[

I

~ NH4

urea

T

Gln Glu

urea

Gln

N~

Gin

FIGURE 10.10 Glutamine and glutamate metabolism within a liver acinus. Abbreviations: G, glutaminase; GS, glutamine synthetase; CPS1, carbamoylphosphate synthetase 1; Cbm-P, carbamoylphosphate. See text for details (adapted from Haussinger, 1990, with permission from the Biochemical Society).

perivenous hepatocytes normally scavenges the ammonium that escapes urea synthesis (apparent Km = 2 to 5 m M ammonium for carbamoylphosphate synthetase) in periportal regions (H~iussinger, 1990). Intrahepatic compartmentation between periportal and perivenous hepatocytes also extends to transport mechanisms, notably the apparently restricted expression of sodiumcoupled glutamate transport to perivenous cells (Taylor and Rennie, 1987), where it appears to be important in providing glutamate for hepatic glutamine synthesis (see Fig. 10.10). The hepatic uptake of glutamine by the sodium-dependent transport system N shows an important element of control to subsequent glutamine catabolism in the liver (largely in the periportal region), as described in Section III. The liver therefore both receives and donates glutamine to the plasma as components of a glutamine/glutamate cycle. The overall result of this process is urea synthesis but the liver also becomes a net producer of glutamine in the fasted state (H~iussinger, 1990; Souba, 1991; Neu et al., 1996), due to a greater increase in activity of glutamine synthetase than of glutaminase in response to the extent of plasma ammonia concentration. Glutamine serves as a major fuel for the rapidly dividing cells of thesmall intestine and important end products of intestinal glutamine metabolism include both alanine and ammonia (Windmueller and Spaeth, 1974; Buttrose et aL, 1987; Souba, 1991). Alanine produced from intestinal glutamine metabolism is carried in the portal system to the liver where it acts as a substrate

for ureagenesis and gluconeogenesis. Ammonia acts as a synergistic activator of this overall metabolic process in the splanchnic bed and, since much of intestinal ammonia production results from glutamine metabolism, glutamine delivery to the splanchnic circulation is therefore an extremely important factor controlling ureagenesis (Buttrose et aL, 1987; Souba, 1991). Plasma glutamine concentration is a reasonable indicator of body nitrogen status (Abumrad et aL, 1989; Neu et aL, 1996) and increased arterial glutamine concentration leads to increased gut glutamine oxidation and ammonia production, which feeds forward through the portal vein to increase hepatic ureagenesis. The major functions of glutamine in renal metabolism are as a source of ammonia for ammonium excretion and as a metabolically generated base for wholebody pH regulation (Brosnan et aL, 1989; Curthoys and Watford, 1995). Catabolism of glutamine (and glutamate) in the kidney liberates ammonia, which is released into the plasma as well as the renal tubular lumen, and makes a significant contribution to total ammonia appearance in the systemic circulation (Welbourne, 1987; Welbourne and Dass, 1988). The major source of this glutamine appears to be muscle, although in chronic acidosis and prolonged fasting the liver may become an important source (Welbourne, 1987; H~iussinger, 1990). The kidney is capable of extracting large quantities of glutamine from renal plasma in severe acidosis (see Section V,D). The lungs may play a significant role in the fine tuning of glutamine homeostasis (Welbourne,

314

10. Transport and Interorgan Nutrient Flows

1988; Plumley et aL, 1990), facilitated by the high rate of blood flow through the tissue. The lungs have a high capacity for glutamine synthesis and it has been estimated that skeletal muscle and the lungs contribute approximately equally to the maintenance of blood glutamine concentration, at least in the rat (Plumley et al., 1990). Adipose tissue is also a site of glutamine synthesis and release in both man and rat, at rates which suggest that adipose tissue may play a substantial role in wholebody glutamine production (Frayn et al., 1991; Kowalski et al., 1997). There appears to be no significant net flux of ammonia into adipose tissue and the principal substrates for glutamine synthesis in this tissue appear to be derived from intracellular proteolysis (Kowalski et al., 1997). B. Alanine Alanine is quantitatively of similar importance to glutamine in interorgan nutrition but its lack of an amide nitrogen grouping results in alanine having a narrower range of functions than does glutamine (Felig, 1975; Abumrad et al., 1989). The combined additions of alanine from peripheral tissues (muscle, lung, adipose) and the intestine to portal plasma results in alanine being the main gluconeogenic amino acid flowing to the liver. Alanine is in consequence by far the largest contributor of the amino acids to hepatic gluconeogenesis, the glucose produced going largely to muscle where it is used in glycolysis or glycogen synthesis. These flows produce a glucose-alanine cycle (Felig, 1975; Christensen, 1990, for review) in which alanine synthesized in muscle by transamination from pyruvate is carried back to the liver for reconversion of the carbon skeleton to glucose. Physiologically this cycle moves amino groups from muscle to the liver but, although it contributes to the overall regulation of glucose economy, it does not move net carbon from amino acids to carbohydrate. Production and release of alanine by skeletal muscle increases markedly during exercise (see Section V,C). Systems ASC, A, and L participate in alanine transport into the liver cell, and system A may use the Na + gradient to transport amino acids against their gradients (Christensen, 1990). Alanine transport into the liver appears to increase with the portal alanine load (see Fig. 10.8), and transport may limit hepatic alanine utilization owing to the high intrahepatic capacity for alanine catabolism (e.g., for ureagenesis, gluconeogenesis) (Fafournoux et al., 1983). Processes contributing to modulation of hepatic system A transport activity, notably adaptive up-regulation during fasting or in response to corticosteroids, are extremely effective in controlling hepatic alanine catabolism (Fafournoux et al., 1983; Low et al., 1992).

C. Glutamate Glutamate is an abundant amino acid in cells (reflecting its importance in intermediary metabolism) but it is found at relatively low concentration in extracellular fluid. Glutamate cycles with glutamine in several important inter- and intraorgan metabolic pathways (e.g., see Sections IV,A, V,A, and V,D) and is also a major excitatory neurotransmitter in the central nervous system. Glutamate released by neurones during neurotransmission is largely taken up by astroglial cells which, in turn, release glutamine, which helps replenish the lost glutamate (Smith and Cooper, 1992; Kanai, 1997). A glutamate-glutamine cycle therefore provides neurones with a precursor for neurotransmitter biosynthesis (Brookes, 1993; Rothstein et al., 1994). This cycle is completed by intercellular movement of amino groups in branched-chain amino acids (see Fig. 10.11) (Smith and Cooper, 1992; Brookes, 1993). Glutamate concentration in the synaptic cleft is kept low by active uptake mechanisms in neuronal and astroglial cells which have high affinity and high "concentrating power" based on their stoichiometry. These excitatory amino acid transporters (EAAT1 to EAAT5) express low Km (values in low/~M range), sodium-, and potassium-dependent transport of L-glutamate and aspartate. E A A T isoforms have specific distributions within the central nervous system (see Fig. 10. 11) and in peripheral tissues. Coupling of glutamate transport by E A A T isoforms to movement of 2 or 3 Na +, plus reciprocal movement of K +, enables extremely high concentration gradients to be maintained across the plasma membranes of cells within the BB barrier. This process effectively excludes the neurotransmitter from the synaptic cleft and also clears it efficiently after excitatory release at the presynaptic terminal. Glutamate is also a precursor for synthesis of GABA (an important inhibitory neurotransmitter) in neuronal cells and a GABA-glutamineglutamate cycle between astrocytes and neurones has been proposed as a component of mechanisms replenishing GABA supplies in neurones (Smith and Cooper, 1992). D. Branched-Chain Amino Acids The liver removes excess amino acids delivered to it in the portal blood with the exception of the branchedchain amino acids (BCAA), which are removed by muscle and adipose tissue (Felig, 1975; Abumrad et al., 1989). Branched-chain amino acids obtained from the diet are generally deaminated in skeletal muscle with the amino groups produced used in de n o v o synthesis of alanine and glutamine. Branched-chain amino acid decarboxylation is the most important step for control

Physiologically Important Flowsof Amino Acids

BCKA ~ / MCT1 / ~ BCKA EAAT + 1/2 NH3 ~....Glu ~ )" Glu " A

Astrocyte

B C A A ~ - ~ BCAA Sys L1

Gln Sys L, ASC / ~ Gln tt

~-BCKA Neuron

BCKA ~- BCAA -------C--~BCAA + NH3 Glu ,,

- . . , ~ ..~

]~

~/Gin

EAAT3 -.~ Glu -9 Gin

Gln

BCAA AromaticAA

315

(lt Endothelium (blood-brain barrier)

I

BCAA AromaticAA

FIGURE 10.11 Glutamate/glutamineexchanges within the blood-brain barrier. This figure shows a diagrammatic representation of the exchanges of BCAA/BCKA, Gin, and Glu between neurons, astrocytes, and the blood-brain barrier. See Section II,E for more details of the denoted amino acid transport systems. Release of keto acids by astroglia cells is mediated by the monocarboxylate transporter MCT1.

of their catabolism in man (Harper et al., 1984; Rennie, 1996 for review). Ammonia accumulation in neurones due to the glutamate-glutamine cycle within the CNS is prevented by a branched-chain amino-keto acid shuttle (see Fig. 10.11). Here astroglial cells actively take up branchedchain amino acids, which are transaminated to keto acids and released into the interstitial space (BrOer et aL, 1997; Yudkoff, 1997). Keto acids are used by neurones for disposal of ammonia in reverse reactions. Much of the uptake of branched-chain amino acids in astroglial cells is by system L, where activity of this transport system appears to be related to the expression of the surface antigen for 4F2hc (Br6er et al., 1995). E. Arginine

Interorgan flows of the cationic amino acids arginine and ornithine (both substrates for system y+) have importance in regard to modulation of the urea cycle (Meijer et al., 1990; Closs, 1996 for review). The arginine economy of the body is effectively separated into hepatic and extrahepatic compartments because of the low level of arginine transport through the low-affinity isoform of the system y+ CAT transporter (CAT2a; Km 2 to 5 mM) at the hepatocyte sinusoidal membrane (Christensen, 1990; Closs, 1996). The liver normally appears to depend largely on arginine synthesized within

the urea cycle, but low Km isoforms of the CAT proteins are expressed upon liver cell transformation or regeneration when there is increased arginine demand for hepatic protein synthesis (Aulak et al., 1996; MacLeod et al., 1996). The kidneys are a major source of circulating arginine, having relatively low levels of arginase. Muscle and kidney both release arginine into the bloodstream but take up ornithine and citrulline which are urea cycle intermediates. Citrulline produced from metabolism of glutamine in the intestinal mucosa is believed to be a major substrate for renal arginine synthesis (Windmueller and Spaeth, 1974; Felig, 1975; Abumrad et aL, 1989). L-arginine is also the substrate of nitric oxide (NO) synthase and arginine uptake into NO-synthesizing cells (e.g., vascular endothelium, macrophages) is a potentially important step for modulation of NO synthesis in these cells (Closs, 1996; Dev6s and Boyd, 1998, for review). Nitric oxide has many crucial physiological functions and its release is modulated by factors including the activities of specific isoforms of CAT (cationic amino acid transporter) and NOS, for example: 1. NO released by endothelial cells (generated by eNOS isoform under the influence of intracellular signalling cascades) causes relaxation of arteriolar smooth muscle and is important for control of regional blood flow and mean arterial blood pressure. The "housekeep-

316

10. Transport and lnterorgan Nutrient Flows

ing" CAT1 isoform (Km ~ 0.15 mM) delivers arginine to these cells under normal conditions. 2. Cells of the immune system (notably macrophages) release NO upon activation (largely related to induction of iNOS isoform) as part of the immune response. Activation (e.g., by bacterial lipopolysaccharide) also induces synthesis of the CAT2 isoform, associated with increased arginine transport and NO synthesis. Both CAT1 and CAT2 express classical system y+ transport activity (Km ~ 0.15 mM for arginine) but macrophage/lymphocyte activation also appears to result in induction of system y+L transport (associated with upregulation of expression of 4F2hc protein), which makes an additional contribution to increased arginine uptake. Delivery of arginine to the intracellular compartment may become the most important factor for control of NO production by iNOS at high levels of cellular activation. 3. Within the nervous system, NO released from action of the nNOS isoform has important functions in control of neurotransmission and neural development. Both CAT1 and CAT2 contribute to arginine transport in cells of the central nervous system (Stevens et al., 1996; Dev6s and Boyd, 1998). F. Aromatic Amino Acids: Relationship to Thyroid Hormone Action The aromatic amino acids phenylalanine, tyrosine, and tryptophan are important substrates for metabolism, and both phenylalanine and tryptophan are essential dietary constituents (Waterlow et aL, 1978; Frayn, 1996). Both Tyr/Phe and tryptophan are precursors for synthesis of neurotransmitters (e.g., catecholamines and serotonin respectively) (Smith and Cooper, 1992 for review) and Tyr/Phe is, in addition, the substrate for generation of thyroid hormones in the thyroid gland (Schwartz et al., 1993). Entry of tryptophan into the brain across the blood-brain barrier has a direct effect on intracerebral tryptophan concentration and hence serotonin (5-hydroxytryptamine) synthesis, because the synthetic enzymes are not saturated at prevailing substrate concentrations within the brain (which are substantially less than plasma concentrations). Competition between tryptophan and other amino acids (including the BCAA) for entry into the brain across the bloodbrain barrier therefore indirectly modulates brain serotonin synthesis, with important physiological and pathophysiological consequences (Betz and Goldstein, 1978; Smith and Cooper, 1992, for review). For example, serotonin has been identified as a central "satiety" signal and relative changes in plasma tryptophan/BCAA ratios have been hypothesized to contribute to modulation of this signal. The high-affinity system L1 is the major

amino acid transporter of the BB barrier and facilitates movement of most zwitterionic amino acids into or out of the cerebrospinal fluid, whereas Na*-dependent amino acid transporters are poorly represented at this site (Betz and Goldstein, 1978; Smith and Cooper, 1992; Sanchez-del-Pino et al., 1995). System L1 also carries amino acid drugs such as L-DOPA into the brain. This transport system selects large, hydrophobic amino acids such as tryptophan and phenylalanine (with Km values of the order of 10/~M) over more polar substrates (e.g., Km for glutamine of 230-900/~M have been reported) and is predicted to be over 95% saturated at the bloodfacing surface of the BB barrier, because plasma amino acid concentrations greatly exceed transport Km values (Smith and Cooper, 1992; Sanchez-del-Pino et al., 1995). The total influx of amino acids into the brain through system L1 is therefore almost independent of their plasma concentrations, but competition between substrates is strongly influenced by changes in the relative abundance of particular amino acids (Christensen, 1990; Smith and Cooper, 1992 for review) as will occur during normal diurnal feeding cycles as well as in certain disease states (see Section V,D). Such competition also means that the apparent Km value for transport measured in vivo will be much higher than the actual value measured for a single substrate in isolation. Brain tryptophan uptake exceeds that predicted from free tryptophan concentration and it has been speculated that some tryptophan is "stripped off" albumin in the cerebral circulation. The transporter has high exchange properties and net fluxes for most substrates are relatively small, except for the BCAA (which are oxidized within the brain) and glutamine, which is released from the brain into the blood. A high proportion of this release is likely to be in exchange with large zwitterionic amino acids entering the brain through system L1 (Grill et al., 1992; Hilgier et al., 1992). The liver is the major site of aromatic amino acid metabolism in the body, extracting these amino acids from the plasma for protein synthesis in the fed state and slowly releasing amino acids generated from protein breakdown in the fasted state. These released amino acids may be used by other tissues during a prolonged fast (Cahill, 1976; Waterlow et al., 1978). Excess aromatic amino acids are oxidized by the liver (Salter et al., 1986), preventing their build-up in the plasma with consequent effects on amino acid exchanges at the BB barrier and therefore on cerebral metabolism. The aromatic amino acid transporters in the liver are the Na+-independent systems T (relatively specific for aromatic amino acids) and L (which has a broad scope for large zwitterionic amino acids) (Salter et al., 1986; Kemp and Taylor, 1997). The kinetic characteristics of systems L and T in hepatocytes appear to be such that at physio-

Physiologically Important Flows of Amino Acids

logical portal vein concentrations of amino acids the extent of competition for transport will be influenced by fluctuations of blood concentrations arising from changes in dietary intake or metabolic turnover (Salter et al., 1986). A metabolic control analysis of this pathway is described in Section III (see Fig. 10.7). Renal reabsorption of aromatic amino acids (see Fig. 10.12) may be used as a general example of transepithelial amino acid transfer (Silbernagl, 1988; Schwegler et al., 1992 for review). Aromatic amino acids are almost completely reabsorbed by the kidney under normal circumstances and only a minor portion of the absorbed amino acids are used for metabolism within the tubule. Renal reabsorption of amino acids is enabled by a battery of both Na+-dependent and Na+-independent transport mechanisms at both brush border and basolateral membranes (see Table 10.1). The Na§ systems B ~ and B ~247 cotransport amino acids with Na § into the epithelial cell down an electrochemical gradient of Na § ions (Christensen, 1990; Van Winkle, 1992, for review). Both transport systems will carry a range of zwitterionic amino acids and the B ~247 system will also accept cationic amino acids such as lysine and arginine as substrates. These Na§ transport processes are believed to be largely responsible for the bulk reabsorption of amino acids in the renal proximal tubule (Schwegler et al., 1992). The recently cloned ATB ~ transporter (Kekuda et al., 1997) is expressed in kidney cells and has functional characteristics resembling system B ~ although it has been suggested (e.g., Kanai, 1997) that the human ATB ~ and the mouse ASCT2 are equivalent

S 3 ~' (bO, *)

$1

AAA AAA Na § Cyst Na" AAA Na + (BO,*, B o)

CAA

AAA

(y*e) (e)

CAA

AAA

FIGURE 10.12 Aromatic amino acid and cystine reabsorption in the renal proximal tubule. Schematic diagram showing aromatic amino acid transport systems in the kidney. Note Na+-coupled reabsorption of aromatic amino acids in renal brush border. Abbreviations: A A A -aromatic amino acid, C A A = cationic amino acid; Z A A = zwitterionic amino acid; Cyst = Cystine. Cystine may be reduced intracellularly to the small Z A A cysteine for subsequent absorption back into the

bloodstream. The BAT-related cystine transport (putative system b~ is localized to the $3 (late) portion of the distal tubule.

317

to system ASC-like transporters in different species (see also Chapter 6). The Na+-independent transport of aromatic amino acids at the renal brush border appears to involve amino acid exchange mechanisms including system b ~ related to expression of BAT proteins (Schwegler et al., 1992; Palacfn, 1994; Chillardn et al., 1996). Basolateral aromatic amino acid exchanges between epithelial cells and the bloodstream appear largely to occur via Na+-independent mechanisms that may include the facilitative systems L and T plus exchange activities possibly including system y+L (Schwegler et al., 1992; Cheeseman, 1992; Palacfn 1994; Thwaites et al., 1996). The structurally related BAT and 4F2-hc glycoproteins are both expressed in renal epithelium (on brush border and basolateral membranes, respectively; Chillardn etal., 1996; Quackenbush etal., 1986). There is accumulating evidence that these two proteins act independently as subunits of heteromeric amino acid transporters (Palacfn, 1994; Wang and Tate, 1995; Peter et al., 1996). Both BAT and 4F2hc foster transport activities in X e n o p u s oocytes which appear to favor zwitterionic-cationic amino acid exchanges (resembling systems b ~ and y+L respectively; Palacin, 1994; Chillardn et al., 1996). Chillardn et al. (1996) propose a model for the physiological role of the b ~ transporter in renal reabsorption o f cationic amino acids (and cystine; see Section IV,G) dissipating the intracellular gradient of zwitterionic amino acids by heteroexchange at the brush border membrane. The inside-negative membrane potential of renal epithelial cells is likely to favor entry of cationic amino acids across the brush border, in which case the exchange activity of the BAT-induced amino acid transport would tend to transport zwitterionic substances such as aromatic amino acids into the tubule lumen. These released substrates are reabsorbed distally, presumably by Na+-coupled transporters such as B ~ and B ~ The observation that Type-1 cystinuria patients (defective in BAT) show increased urinary concentrations of cationic amino acids and cystine but not zwitterionic amino acids (Schwegler et al., 1992) indicates that additional processes for reabsorption of the latter exist and that BAT protein is not essential for this process (Chillardn et al., 1996). The kinetic properties of system y+L transport (as induced by 4F2hc in oocytes) appear to favor net movement of cationic amino acids out of the cell coupled to a Na+-dependent uptake of zwitterionic amino acids (Palacfn, 1994; Chillardn et al., 1996). Expression of this type of transporter mechanism in the renal basolateral membrane may be physiologically important for net transfer of cationic amino acids from kidney to blood (Chillardn et al., 1996), particularly arginine, given that the kidney is the major site of net arginine synthesis within the body. There is a surprisingly

318

10. Transport and Interorgan Nutrient Flows

low intrarenal accumulation of aromatic amino acids, considering the range of Na+-coupled processes favoring their entry into the cells (Silbernagl et al., 1996), and it is likely that the reabsorptive gradient is dissipated by the activity of BAT and 4F2-1inked exchanges. Heteroexchanges of zwitterionic amino acids also occur by these transport mechanisms (see Fig. 10.12). This may have particular relevance at the basolateral membrane, enabling metabolically important amino acids such as glutamine to be taken up into kidney cells in exchange for movement of essential amino acids back into the circulation, completing their reabsorption. The thyroid hormones T3 and T4 (triiodothyronine and thyroxine respectively) are large halogenated, aromatic amino acids generated by modification of iodinated tyrosine residues within follicles of the thyroid gland and released into the blood circulation (Schwartz et al., 1993). The thyroid gland predominantly releases T4, whereas T3, the most active of the two hormones, is largely generated by monodeiodination of T4 in liver and kidney. The liver is also the site of thyroid hormone degradation (Salter et al., 1986) and conjugated thyroid hormone catabolites are excreted in the bile (OudeElferink and Jansen, 1994). In addition, liver and kidney are both important target tissues for thyroid hormone action and their major source of nucleoreceptor-bound T3 is believed to be the circulating T3 pool (Hennemann et al., 1986; Schwartz et al., 1993). Perhaps surprisingly, the specific mechanisms by which thyroid hormones cross the blood-facing membranes of these tissues are not fully understood, although the mechanism is certainly not simple diffusion (see Chapter 4). Uptake into hepatocytes of T3 and T4 is known to be relatively stereospecific and saturable. It has been shown that there is a close relationship between transport of aromatic amino acids and thyroid hormones by system T in a number of tissues, involving a mechanism that may enable countertransport by heteroexchange of T3 with intracellular aromatic amino acids (Zhou et al., 1992). The binding of thyroid hormone to surface receptors may be an essential prerequisite for subsequent movement across the membrane by transport and/or receptor-mediated endocytosis and it has been suggested (Samson et al., 1992; Kemp and Taylor, 1997; Taylor et al., 1998) that interactions between receptors and transporters enable thyroid hormones to be "channeled" to transport mechanisms in the cell membrane. G. Cystine Cystine resulting from cysteine oxidation is released from peripheral tissue (largely skeletal muscle) at rates which are negatively correlated with plasma concentrations of amino acids but directly correlated with the

plasma cystine concentration (Dr6ge and Holm, 1997). It has been proposed that plasma cystine concentration is normally regulated by muscle protein catabolism and that a feedback mechanism exists whereby postabsorptive cystine release by muscle contributes to downregulation of hepatic urea synthesis with consequent amino acid retention (hepatic cystine catabolism generates sulfate and protons which would tend to reduce ureagenesis). Regulation of plasma cystine concentration appears to be disturbed in wasting catabolic conditions in tandem with a low plasma glutamine:cystine ratio (possibly related to impaired hepatic cystine and cysteine metabolism) and negative whole-body nitrogen balance (Dr6ge and Holm, 1997, for review). Cystine transported into cells is generally reduced to cysteine, so the intracellular pool of cystine is extremely small (Bannai and Tateishi, 1986; Christensen, 1990, for review). The transport system x-~ selects the tripolar form of cystine, as well as the similarly charged glutamate ion. Cystine and glutamate are exchanged by this transport system and the physiological flow is expected to be the uptake of cystine coupled to the efflux of glutamate. For example, astroglial cells in the CNS take up cystine, which is subsequently reduced to cysteine. Cysteine is then released by astrocytes and taken up by neurones (which have negligible cystine uptake). This shuttle mechanism may be important for maintenance of glutathione concentrations within neurones (Sagara et aL, 1993). The properties of the absorptive and reabsorptive transport systems for cystine appear to provide an illustration of mechanisms adapted to differences between physiological loads at the luminal surface of intestine and kidney. The initial renal filtrate has the same solute composition as plasma, whereas the intestinal luminal contents vary considerably with time and nutritional status and may be much lower than prevailing plasma concentrations. Bulk reabsorption of solutes from the kidney is, overall, less expensive energetically than intestinal absorption due to the lower transcellular solute gradients. In regard to cystine transport, both highaffinity (Na+-independent system b~ exchanger; see Fig. 10.12) and low-affinity (including Na+-depen dent) mechanisms are found in the kidney (Silbernagl, 1988; Chillar6n et al., 1996), whereas the intestinal epithelium may contain only the high-affinity system b ~ like mechanism (Ozegovic et aL, 1982; Palacin, 1994). The Na+-dependent transporter is found largely in the early part of the proximal tubule ($1 and $2 regions) where it is believed to be responsible for bulk reabsorption of cystine with the more distally located b~ mechanism ($3 proximal tubule) acting as a scavenger for remaining cystine (Silbernagl, 1988; Chillar6n et aL, 1996). Mutations in a gene (BAT) related to expression

Amino Acid Nutrition Under Special Circumstances

of system b~ transport are associated with the genetic disease cystinuria (see Section V,D). H. Glutathione and Conjugates Glutathione (GSH; y-glutamylcysteinylglycine) is a ubiquitous, low-molecular-weight thiol involved in a variety of metabolic processes including detoxification, reduction, and enzyme regulation. The importance of this tripeptide in protecting cells against reactive oxygen species, reactive drug intermediates, and free radicals is well documented (Meister, 1988; Christensen, 1990; Akerboom and Sies, 1992, for review). Most cells are capable of synthesizing glutathione from amino acid precursors and intracellular glutathione concentrations are usually in the millimolar range. Cellular glutathione is kept in the reduced form by glutathione reductase such that normally <5% of glutathione is in the disulfide form (GSSG; the product of GSH reductions). Glutathione and GSSG are pumped out of cells to be degraded extracellularly by peptidases located on the outer surface of cell membranes (Meister, 1988; Akerboom and Sies, 1992). High activities of these peptidases (including (y-glutamyl transpeptidase) are found in the apical membranes of epithelia including the kidney, intestine, and liver (bile canalicular membrane), enabling an active turnover of glutathione (export with reuptake of released constituent amino acids) within these organs. Membrane transport is therefore an important step in modulating glutathione turnover and function (Bannai and Takeishi, 1986; Akerboom and Sies, 1992). Plasma free-glutathione concentration is of the order of 15-30 ~M, related to an active interorgan glutathione turnover in which the liver is the major organ releasing glutathione and the kidney is the major site of extraction from the blood. Intraorgan turnover of GSH increases during fasting and exercise and in both situations increased hepatic GSH efflux is associated with elevated plasma GSH concentration (Akerboom and Sies, 1992; Dr6ge and Holm, 1997, for review). Plasma as well as cellular GSH has protective functions (e.g., against extracellular oxygen free-radicals at sites of inflammation) related to its reducing properties. The liver is a major site of glutathione synthesis for hepatic functions, detoxification, and export. Hepatic sinusoidal efflux of glutathione is about 10 times as great as that at the bile canalicular membrane. The liver supplies at least 90% of plasma glutathione, which is carried mainly bound to plasma protein such as albumin. Sinusoidal efflux of GSH is saturable, ATP dependent, and competitively inhibited at the intracellular face by organic anions (Bannai and Takeishi, 1986; Akerboom and Sies, 1992). The transport process normally operates at a nearmaximal rate. GSH transport at the canalicular mem-

319

brane is probably electrogenic but appears not to be ATP-dependent, i.e., it is distinct from the sinusoidal transport mechanism. GSH secreted into the bile may be subject to extensive metabolism and recycling of the amino acids via Na+-dependent amino acid transport systems localized in the canalicular membrane. GSSG and glutathione conjugates formed by liver metabolism are selectively transported into the bile canaliculae for biliary breakdown or excretion (MUller and Jansen, 1997; Keppler et aL, 1997, for review). This process is mediated by the unidirectional ATP-dependent export pumps MRP1 and MRP2 (members of the multidrug resistance protein family). MRP2 (also called the canalicular multispecific organic anion transporter or cMOAT) has been localized to the apical domain of polarized epithelia including the kidney proximal tubule and the hepatocyte canalicular membrane, whereas MRP1 is exclusively present in the basolateral membrane. Substrates of MRP1 and -2 include conjugated bilirubins and bile salts as well as glutathione Sconjugates. MRP1 and -2 appear to function in detoxification by hepatic/renal elimination of conjugated xenobiotics as well as in defence against oxidative stress (Keppler et aL, 1997). MRP1 may also be involved in hepatic sinusoidal GSH export. Organic anions including unconjugated bilirubin show potent inhibitory effects on sinusoidal GSH release, and interorgan turnover of GSH is altered in hyperbilirubinemia (Akerboom and Sies, 1992). The kidney is the major organ of glutathione removal and is responsible for at least 50% of plasma glutathione turnover reflected by an extraction of --~80% of both GSH and GSSG during a single passage. Glomerular filtration and peritubular absorptive mechanisms both contribute to this disappearance, which involves breakdown by peptidases at both surfaces of the renal epithelium followed by amino acid uptake.

V. AMINO ACID NUTRITION UNDER SPECIAL CIRCUMSTANCES A. Pregnancy and Lactation Amino acid uptake by uterine tissues (including placenta and fetus) during pregnancy is a vital aspect of maternal and fetal nutrition (Moe, 1995; Malandro et al., 1996). Uterine and fetal uptake of individual amino acids are not necessarily equal because amino acids can be produced or utilized by uteroplacental tissues including the myometrium and placenta (the latter being the primary site of metabolism). Major patterns of transplacental amino acid movement in vivo have been studied in the sheep by Battaglia and colleagues

320

10. Transport and Interorgan Nutrient Flows

(e.g., Vaughn et aL, 1995; Geddie et al., 1996; Chung et al., 1998). Indispensable amino acids are taken up across the placenta into the fetus for fetal growth although total uptake may significantly exceed the estimated normal protein accretion rate of the fetus, indicating significant fetal amino acid catabolism (Chung et aL, 1998). A large proportion of fetal glutamate taken up by the placenta is rapidly oxidized and the placenta excretes ammonia and also produces glutamine from placental glutamine synthetase (Vaughn et al., 1995). The placenta takes up glutamate and serine produced by the fetus (see Fig. 10.13). Some of the glutamine delivered to the fetus by the placenta is converted back to glutamate by the fetal liver, establishing a glutamate-glutamine shuttle promoting glutamate oxidation in the placenta and hepatic utilization of glutamine amide by the fetus (Vaughn et al., 1995). Hepatic type glutaminase is not found in the fetus, the fetal liver expressing renal-type glutaminase in line with the metabolic Gln-Glu cycling between fetus and placenta (Curthoys and Watford, 1995). The placenta may utilize branched-chain amino acids and release keto acids into both the fetal and maternal circulation (Chung et al., 1998). The deamination process will also result in the formation of glutamate via transamination and it has been suggested that this represents an additional source of glutamate to the placenta. A major metabolic function of the placenta (at least the ovine placenta) is glycine production from maternal serine (Geddie et aL, 1996). This conversion channels the/3carbon of serine into a variety of synthetic reactions that require activated single-carbon units, such as purine synthesis. Serine is also routed to the placenta from the liver of fetal lambs where it is produced from glycine supplied via the placenta. Under normal physiological conditions, transport by the maternal circulation is not a limiting factor for the uterine uptake of amino acids (Moe, 1995; Malandro et aL, 1996; Chung et aL, 1998).

FETUS

MOTHER

PLACENTA

oxidation

)

9

CO2

Glu

oxidation

B. Starvation Major changes in whole-body and interorgan metabolism occur if fasting is prolonged significantly beyond the normal postabsorptive period (Cahill, 1976; Waterlow, 1995). Bodily glucose requirements are met by hepatic (and renal) gluconeogenesis using precursors that include amino acids, largely in the form of alanine released from peripheral tissues. Adaptive up-regulation of hepatic alanine transport contributes to increased gluconeogenesis (Fafournoux et al., 1983; Low et al., 1992). This alanine is increasingly supplied from protein breakdown as starvation progresses. Renal glutamine utilization increases markedly due to metabolic acidosis associated with starvation (see Section V,D). Intestinal glutamine utilization continues to contribute to portal alanine load but the liver becomes an increasingly important source of this glutamine (Abumrad et al., 1989, for review) as it switches from synthesis of urea to glutamine to detoxify circulating ammonia (these changes reduce the extent of acidosis and conserve amino-N; see Section V,D). C. Exercise

~, +Nil3

Gin 9 >- Gly~,~Ser ~-~ Ser >-- BCAA 9

The surface membranes of the human placenta include Na+-dependent systems A, ASC, and B ~ and Na +independent systems L and y+L for zwitterionic amino acids and additional transporters for anionic and cationic amino acids, including EAAT1 and CAT1 respectively (Moe, 1995; Malandro et al., 1996; Novak and Beveridge, 1997; Kekuda et al., 1997; Dev6s and Boyd, 1998). Transport processes with high exchange activity such as systems ASC and L may be involved in the exchange activity at the fetal face of the trophoblast; for example, extracting serine from the fetus in exchange for delivery of other amino acids. After parturition, alanine and glutamine are used by the mammary glands of lactating animals, alanine at least appearing to serve as a lipid precursor. Milk proteins are relatively enriched in Gln+Glu residues and the mammary glands are a major site of glutamine utilization during lactation (Meijer et al., 1993).

Gln Ser BCAA 9 C02

FIGURE 10.13 Amino acid exchanges between fetus and mother across the placenta. This diagram shows the major exchanges of Gly, Ser, Glu, Gin, and B C A A between the fetus and mother across the placenta. Note Gin and Gly synthesis within the placenta. See text for further details.

The fuel requirements of muscle during aerobic exercise are largely met by oxidation of glucose and free fatty acids, whereas protein is spared from this fate (Felig, 1975; Rennie, 1996, for review). Nevertheless amino acid carbon skeletons may be used for de n o v o glucose synthesis during and after exercise and a major source of these amino acids is muscle protein (Carraro et al., 1994; Rennie, 1996). There are quantitative and qualitative changes in amino acid exchange across contracting muscles during exercise and a significant net

321

Amino Acid Nutrition Under Special Circumstances

output is observed only for alanine (Felig, 1975; Carraro et al., 1994). Alanine release increases in proportion

to exercise severity and is linked to the availability of pyruvate (Carraro et al., 1994; Rennie, 1996). Nitrogen sources for alanine production from pyruvate in exercising muscle are other amino acids as well as ammonia derived from the purine-nucleotide cycle. In short-term exercise, amino-N appears to be donated from endogenous amino acids (possibly including those produced from protein breakdown), notably B C A A . A s well as pyruvate-derived alanine, alanine derived from protein breakdown also increases (from 35 to 43% of total alanine release) during low-intensity exercise, due to accelerated whole-body protein catabolism (Carraro et aL, 1994). During prolonged exercise there is net uptake of BCAA into muscle where they are oxidized as fuel with continued alanine output (Felig, 1975; Biolo et aL, 1995a). The source of these BCAA is the splanchnic bed, at least partly from breakdown of gut proteins (Williams et al., 1996). Exercise appears to stimulate amino acid transport into muscle and the stimulatory effect of exogenous amino acids on muscle protein synthesis is enhanced by prior exercise (Biolo et al., 1995a; Rennie, 1996). The mechanism of this stimulation is poorly understood but includes contribution from increased transport activity as well as increased amino acid delivery to muscle secondary to the increased blood flow (Biolo et al., 1995a; Rennie, 1996; Williams et al., 1996). Alanine released from muscle is largely extracted from plasma by the liver for gluconeogenesis, although a transient increase in plasma alanine concentration is initially observed. There appears to be an increase in hepatic alanine transport capacity during prolonged exercise, associated with increased gluconeogenesis and hepatic glucose release (Felig, 1975). During recovery from exercise, hepatic alanine uptake exceeds muscle release, and hepatic gluconeogenesis from alanine is used to enhance repletion of liver glycogen stores. Plasma glutamine is also depleted in postexercise periods and these acute effects may be cumulative if inadequate recovery periods are taken between training bouts. Athletes suffering from overtraining syndrome maintain chronically low plasma glutamine, which may have adverse effects on the gastrointestinal and immune systems (Hack et al., 1997). D. Pathophysiological Conditions Body protein stores are at risk during disease, illness, and after injury (Felig, 1975; Rennie, 1985; Waterlow, 1995; Price and Mitch, 1998, for review). The protein stores are progressively depleted and the carbon skeletons of amino acids are used for gluconeogenesis, ketogenesis, and acid-base regulation. The pathways of in-

terorgan glutamine flux therefore show marked changes in various pathological circumstances (Rennie et al., 1989; Souba, 1992; Neu et al., 1996, for review). A reduction in muscle cell volume characteristic of many pathophysiological states (H~iussinger et aL, 1993) is associated with reduced rates of muscle glutamine uptake (see Fig. 10.14), lowered muscle glutamine concentration, and net muscle catabolism (Neu et aL, 1996; Rennie et al., 1996). We have evidence for a functional link between glutamine transport and cell volume in muscle (see Section II,F), which we propose makes an important contribution to the observed increase in glutamine release from muscle in these states (Rennie et al., 1996; Low et aL, 1997a). Acquired resistance to insulin and growth hormone in peripheral tissues (mainly skeletal

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FIGURE 10.14 (A) Correlation of muscle cell volume with nitrogen balance in humans. Skeletal muscle cell water was measured in needlebiopsy samples from quadriceps femoris by the chloride distribution method. A, healthy subjects (n = 17); B, liver tumors (n = 5); C and D, polytrauma, day 2 (C) and day 9 (D) after trauma (n = 11); E, acute necrotizing pancreatitis (n = 6)" and F, burns (n = 4). Results are expressed as means __+SEM (adapted from H~iussinger et al., 1993, with permission from Lancet Ltd.). (B) Correlation of the effects of Gin and insulin on Gin transport activity and cell volume of rat skeletal muscle cells. 3H-Labelled Gin (0.05 m M ) transport and cell volume (3H20 space) were measured over 1 or 2 min, respectively, in NaC1 medium after 30 min exposure to basic experimental medium without or with 2 m M Gin and without or with 66 nM insulin. A, control (nothing added); B, + insulin; C, + Gin; and D, insulin + Gin. Results are means ___ SEM for four preparations.

322

10. Transport and Interorgan Nutrient Flows

muscle) during severe catabolic states may also contribute to muscle wasting (Rooyackers and Nair, 1997). 1. Acidosis

Catabolic states (e.g., type I diabetes, prolonged fasting) are associated with a metabolic acidosis owing primarily to production of ketone bodies. In such situations ureagenesis is suppressed (conserving body bicarbonate pools) and glutamine becomes the major product of ammonia detoxification (see Welbourne, 1987; H~iussinger, 1990, for reviews). Regulation of the bodily hydrogen ion economy in chronic acidosis is highly dependent on flows of glutamine between organs capable of synthesizing and releasing this substrate and the kidneys, which are capable of extracting it from the plasma to produce NH+4 for direct excretion, thus conserving HCO-3 (Welbourne, 1987; Brosnan et al., 1989). Renal glutamine utilization increases markedly during chronic metabolic acidosis in association with reduced plasma glutamine concentration, an example of concentration being sacrificed to produce the necessary flow (Souba, 1992). Such changes in plasma composition (which also include decreased [HCO-3] and [Na3] and increased [NH+4]) affect substrate supply to liver and contribute to the reduction in urea synthesis and increase in glutamine synthesis by this tissue (Welbourne, 1987; H~iussinger, 1990; Meijer et al., 1990). Inhibition of amino acid uptake into liver cells during both acute and chronic metabolic acidosis also leads to a decrease in ureagenesis (Boon et al., 1996). A major advantage of glutaminogenesis over ureagenesis during acidosis is that no bicarbonate is consumed by the glutamine synthetase reaction and indeed subsequent glutamine oxidation generates bicarbonate (Welbourne, 1987; Brosnan et al., 1989; H~iussinger, 1990). Glutamine synthesis also conserves the amino nitrogen of the ammonia which otherwise would be excreted as urea. Furthermore, the diversion of hepatic ammonia detoxification from urea synthesis to the glutamine synthetase pathway in perivenous hepatocytes facilitates renal disposition of excess NH4 +. The drawback of glutamine synthesis is that the glutamate cosubstrate ultimately comes from body protein and as wasting of muscle or other lean tissue (e.g., gut) continues (Souba, 1992; Neu et al., 1996). The combined decrease in plasma pH and glutamine concentration during chronic metabolic acidosis also helps to redirect this amino acid from the intestine (which exhibits load- and pH-dependent glutamine uptake) to the kidneys (Welbourne, 1987). Renal (but not hepatic) glutaminase is inhibited competitively by glutamate over the concentration ranges seen in vivo (Curthoys and Watford, 1995). Renal glutamate concentration falls during chronic metabolic acidosis, thereby

reducing its inhibitory effect on glutaminase, allowing increased flux through the enzyme and higher rates of glutamine breakdown (Carter and Welbourne, 1997). These observations form the basis of a model of metabolic regulation, proposed by Welbourne and colleagues (Carter and Welbourne, 1997; Welbourne, personal communication 1998) to help explain how the kidney can increase ammonia production and excretion from glutamine during chronic metabolic acidosis. In this model, the regulatory mechanism involves a feedforward regulation of glutaminase by glutamate, related to extracellular glutamate availability and glutamate transport into kidney cells (see Fig. 10.15). There is evidence that glutaminase reacts with substrates and inhibitors from the cytosolic rather than the mitochondrial matrix compartments (Kvamme et al., 1991), opening up the possibility that renal extraction of plasma glutamate contributes to modulation of glutaminase flux by altering the size of the cytosolic glutamate pool. Basolateral y-glutamyl transferase (GGT) appears to be important for generating extracellular glutamate for renal uptake; approximately 40% of the glutamine extracted from plasma by the chronically acidotic rat kidney may enter renal cells as glutamate following extracellular glutamine hydrolysis by GGT at the basolateral membrane (Welbourne and Dass, 1988). This GGT activity is highly dependent upon the external concentrations of glutamine and bicarbonate (the latter as an activator: Mu and Welbourne, 1996) such that an approximately 50% fall in both of these concentrations, as occurs in metabolic acidosis, produces a marked limitation on the glutamate available for uptake into the cell with a concomitant reduction in the intracellular pool size.

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FIGURE 10.15 Glutamine (Gin) and glutamate (Glu) exchanges in the kidney. Gin reaches the tubule lumen by glomerular filtration. Extracellular lumenal and basolateral Gin hydrolysis is catalysed by y-glutamyl transferase (GGT) with subsequent glutamate transport into the cell. Renal glutaminase is normally suppressed by high cell [Glu]. In chronic metabolic acidosis, GGT is inhibited and cell [Glu] decreases with resultant activation of glutaminase and enhanced glutamine catabolism (adapted from Welbourne and Dass, 1988, with permission from Springer-Verlag).

Amino Acid Nutrition Under Special Circumstances

Glutamine transported into kidney cells (including filtered glutamine) is thus coupled to the activated glutaminase, yielding ammonia for tubular NH4 § excretion and bicarbonate, which is released into the plasma. Elevation of the plasma bicarbonate concentration increases GGT activity with resultant increase in cellular glutamate content to close the feedback loop by inhibiting glutaminase. In this scheme, both the extracellular and intracellular signals of metabolic acidosis (HCO3- and H § respectively) act to enhance glutamine utilization for bicarbonate regeneration to help restore normal acid-base balance. 2. Sepsis and Injury Muscle glutamine stores are released after injury or infection and are used as fuel, in repairing tissues and in mounting an immune response (Souba, 1991, 1992; Neu et aL, 1996 for review). Hepatic glutamine utilization is greatly increased during sepsis (supporting increased synthesis of nucleotides, glutathione, glucose, and urea) due to a combination of factors including increased hepatic glutamine delivery (Souba, 1991). Glutamine utilization by the immune system is also increased during sepsis (Newsholme and Parry-Billings, 1990; Neu et al., 1996). This overall increase in body glutamine utilization is supplied by increased synthesis in skeletal muscle (mostly at the expense of muscle protein) and the lungs (Rennie, 1985; Souba, 1992; Neu et aL, 1996). Lymphoid cells exhibit elevated rates of amino acid transport when they enter the proliferative cell cycle during the immune response (Segel, 1992 for review). Lymphoblastoid B cells show an increase in system L transport when passing from Go into the proliferative cycle and this persists through G1 and falls during the latter part of the S phase. Activated lymphocytes increase their amino acid transport rates reflecting increased Vmax values for systems A and ASC and, in some cases, systems y+, y+L, and L (Segel 1992; Dev6s and Boyd, 1998). The elevated amino acid transport in lectin-treated lymphocytes requires new protein synthesis and occurs in parallel with other processes of lymphocyte activation. 3. Cancer An increased rate of amino acid uptake is regarded as a general feature of tumor cells, and activation of system A is a consistent early event following cell transformation in vitro (e.g., Guidotti and Gazzola, 1992, for review). Rapidly dividing cells including those of malignant tumors use glutamine as a major fuel and a tumor may become the major organ of glutamine

323

utilization in advanced malignant disease (Souba, 1991, 1992). Here the plasma glutamine concentration becomes depleted and gut glutamine utilization is therefore reduced, whereas both hepatic and muscle glutamine output increase in a vain attempt to keep pace with glutamine utilization by the growing tumor (Neu et aL, 1996). 4. Endocrine Disorders a. Diabetes Mellitus

Insulin deficiency (Type I diabetes) or resistance (Type II diabetes) have a number of direct and indirect effects on amino acid transport and metabolism (Saltiel, 1996; Rooyackers and Nair, 1997) with an overall catabolic effect on protein turnover. The major protein anabolic effect of insulin on human skeletal muscle in vivo appears to be a decrease in protein breakdown (associated with hypoaminoacidemia) rather than an increase in protein synthesis (Rooyackers and Nair, 1997). Increased amino acid supply is therefore required for insulin to even marginally stimulate muscle protein synthesis in vivo. Insulin-stimulated transport processes (e.g., system A) are less responsive to changes in whole-body nutritional status of diabetics than healthy individuals, retarding any increase in amino acid uptake into tissues immediately after feeding (McGivan and PastorAnglada, 1994; Rooyackers and Nair, 1997). This is likely to reduce the anabolic potency of amino acids in diabetes, although there may be compensatory increases in basal transport activity during longer-term uncontrolled diabetes (e.g., Low et al., 1992). Systemic effects of poorly controlled diabetes (ketoacidosis, and elevated plasma corticosteroid concentrations) may have independent effects on amino acid transport and metabolism. b. Thyroid Disease

Thyroid hormones are essential for growth but have a profound catabolic effect on skeletal muscle during both hypo- and hyperthyroidism (Rooyackers and Nair, 1997). Thyroid hormones exert physiological (and pathophysiological) effects on tissues by altering gene transcription through interactions with regulatory elements in the cell nucleus (Schwartz et al., 1993). For example, the proportion of amino acids reabsorbed from the kidney is enhanced after T3 treatment in rat, suggestive of a direct effect of T3 on expression of amino acid transporters (Fleck, 1992). Changes in hepatic T3 turnover during altered thyroid status have been reported (Schwartz et al., 1993; De-Jong et aL, 1994). We have recently studied the possible role of hepatic T3 transport activity in mediating such changes, not least because pathophysiological modulation of this process

324

10. Transport and lnterorgan Nutrient Flows

could have important consequences for whole-body management of hyper- and hypothyroid states. We found that T3 and T4 inhibited tryptophan uptake by system T in rat liver membranes to an extent dependent upon the thyroid status of the donor rat, increasing in the order hypothyroid < euthyroid < hyperthyroid, although other kinetic parameters of tryptophan uptake and T3 binding were not influenced by thyroid status (Kemp and Taylor, 1997). We speculate that these results reflect changes in association between transporter and receptor in response to alterations in plasma thyroid hormone concentrations, enabling appropriate modulation of hepatic transport (uptake and/or release) of thyroid hormones to help limit pathophysiological changes in T3 and T4 abundance (Kemp and Taylor, 1997; Taylor et al., 1998). 5. Inborn Errors of Metabolism

The interdependence between fluxes of individual amino acids for whole-body amino acid economy is shown clearly when the concentration of a single amino acid (or a small group) is either decreased or increased in the circulation. Certain inborn errors of amino acid metabolism result in extremely high plasma levels of amino acids and their metabolites. For example, phenylketonuria (PKU; defective phenylalanine hydroxylase) results in elevated plasma concentrations of phenylalanine and its metabolites (Christensen, 1987, for review), whereas maple syrup disease (MSD; defective branched-chain keto acid dehydrogenase) results in increased plasma concentrations of BCAA (especially of leucine) and their keto acids (Chuang and Shih, 1995). These disturbances in plasma amino acid composition interfere with amino acid transport across the bloodbrain barrier and cause imbalances in neurotransmitter synthesis associated with impaired brain function. Plasma excess of aromatic or branched-chain amino acids may block amino acid efflux from tissues as well as influx by competing for exchange transport through systems having a broad-substrate range such as system L (Christensen, 1987a,b, 1990, for review). This leads to sequestration of certain amino acids (including glutamine, glycine, and alanine) within tissues such as muscle and liver with a consequent reduction in their plasma concentrations. Recent unpublished data by Dr. D. H. Morton and colleagues (Clinic for Special Children, Strasburg, Pennsylvania) demonstrates that therapy leading to correction of hyperleucinemia in a newborn MSD infant (over a 6-day period) is associated with reciprocal, restorative increases in plasma concentrations of amino acids including alanine, glutamine, lysine, glycine, and serine (e.g., Fig. 10.16). These observations may reflect a release of leucine inhibition on cellular

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FIGURE 10.16 Plasma leucine and alanine concentrations in a newborn child with maple syrup disease (MSD) during first 6 days of therapy to normalize hyperleucinemia. Unpublished observations of Drs. D. H. Morton and K. Strauss (Clinic for Special Children, Strasburg, Pennsylvania, USA). Plasma [leucine] at diagnosis was 2394 I~M, therapy was initiated and this value dropped steadily over 6 days to 100/~M (within the normal range). Plasma [alanine] shows a close inverse relationship with plasma [leucine], which may reflect a release of leucine inhibition on cellular release of alanine as tissue and plasma [leucine] decline.

efflux of these amino acids as the hyperleucinemia becomes normalized. Amino acid sequestration in the brain during untreated MSD may contribute to intracerebral water retention and characteristic brain edema (Morton, personal communication). In untreated PKU and MSD, the filtered load of certain amino acids and metabolites may exceed the reabsorptive capacity of the kidney and the excess spills over into the urine (Schwegler et al., 1992 for review). A small number of inborn errors of amino acid metabolism may be attributed directly to genetic abnormalities of amino acid transport processes. Type 1 cystinuria is characterized by mutations in BAT protein (Calonge et al., 1994). Failure to reabsorb cystine (which is poorly soluble in water) results in build-up of cystinecontaining renal calculi and consequently impaired renal function (Segal and Thier, 1995). Hartnup disorder (possibly a genetic defect in the system B ~ transporter; Scriver, 1988) is characterized by renal hyperexcretion and intestinal malabsorption of a variety of zwitterionic amino acids, notably tryptophan, and the major symptoms of the disorder are those of niacin deficiency apparently owing to tryptophan malabsorption (Scriver, 1988, Schwegler et aL, 1992 for review). E. Parenteral and Enteral Amino Acid Nutrition The importance of glutamine nutrition is now well recognized and various regimes of both enteral and parenteral glutamine supplementation in therapy for a wide range of diseases and injuries have been described (Neu et al., 1996, for review). Intestinal immune function can

Summary

be sustained by oral dietary glutamine supplementation, whereas for parenteral nutrition certain glutamine dipeptides are generally used because they afford increased stability to the compound. Glutamine supplementation is required to prevent gut mucosal atrophy during total parenteral nutrition. Glutamine supplementation (oral or intravenous) also appears to protect liver cells from oxidant injury by increasing intracellular glutathione content, whereas it decreases glutathione content of tumor cells (rat fibrosarcomas) (Rouse et al., 1995). This raises the possibility that oral glutamine supplementation could be used to enhance the selectivity of antitumour drugs by helping to protect normal tissue during chemotherapy.

Vl. SUMMARY This chapter provides an introduction to current concepts in interorgan nutrition as they relate to protein and amino acid metabolism in the whole body. Methodological issues and their limitations are discussed together with assessments of the scope and importance of amino acid flows between tissues and organs. We describe specific interorgan amino acid flows together with the influence upon them of a variety of physiological and pathophysiological conditions. We highlight the vital role of biomembrane transport of amino acids in the facilitation and control of flows between compartments. We also illustrate the use of metabolic control theory to determine the quantitative importance of transport processes to regulation of amino acid metabolism. The past 5 years have seen very rapid advances in our knowledge of the molecular structure of amino acid transporters. However, we still know remarkably little about how these transport proteins function, and we need to learn more about regulation of their expression.

325

Furthermore, cDNA encoding transport proteins in important amino acid transport systems, such as A and N, remain to be cloned. Nevertheless, there are already significant challenges and opportunities for the integration of the new molecular knowledge into a holistic view of interorgan amino acid nutrition. Many of the phenomenological descriptions of interorgan nutrition contained in this chapter should be capable of being explained on a mechanistic basis by combining the knowledge gained from classical flux measurements with the new molecular information. Particular opportunities are likely to arise from the use of antisense and transgenic animal technologies in order to establish the extent to which ontogenetic and physiological changes follow any reduction or enhancement of the expression of particular gene products (partial or total knockouts of EAAT1 to EAAT3 and CAT1 have been reported to date: Rothstein et al., 1996; Peghini et al., 1997; Perkins et al., 1997). The redundancy of cellular and organismal organization inevitably means that some of these approaches will lead to apparently null results, sometimes simply because we will be (at least initially) ignorant of the ways in which transgenic animals compensate for their deficiencies in amino acid transport. Nevertheless, such novel approaches will undoubtedly provide important new insights on interorgan nutrition and may yield tremendous surprises, such as the recognition of hitherto unrecognized functions associated with amino acid transport. Acknowledgements We are grateful to Drs. L. Van Winkle and H. N. Christensen for their constructive comments and Drs. S. Br/Ser and F. C. Battaglia for useful information provided during the writing of this chapter. We are particularly indebted to Drs. T. C. Welbourne and D. H. Morton for allowing us access to unpublished work. Thanks also to Mrs. D. M. Watt for excellent secretarial assistance. Authors' work presented here was supported by MRC, BBSRC, The Wellcome Trust, and the University of Dundee.

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Selected Techniques in Membrane Transport*

I. INTRODUCTION

Triton X-100, deoxycholate, cholate, urea, CHAPS (3-[- ( -cholamidopropyl)dimethylammonio]- 1-propanesulphonate), octaethylene glycol dodecyl ether, and decanoyl-N-methylglucamine (MEGA-10)--often in the presence of substrates or thiols as "protectants" against denaturation. Insoluble membrane components are removed from the membrane/detergent mix by centrifugation, after which the detergent must be removed from the solubilized protein fraction in the supernatant. One particular advantage of MEGA-10 (used in the solubilization of the hepatic zwitterionic amino acid transporter system A (Quesada and McGivan, 1988)) is that it can be easily removed by passing the protein fraction through Sephadex G-50, doing away with methods involving overnight dialysis to remove detergents. Zwitterionic amino acid transporter activities (including systems A, B, and N) are associated with very hydrophobic proteins (see Nakanishi et aL, 1994), making solubilization troublesome. Tamarappoo and Kilberg (1991) used a solution containing 2.5% cholate and 4 M urea to solubilize system N (the transporter of glutamine, asparagine, and histidine), and rapid reconstitution was made possible by precipitating the protein fraction with polyethylene glycol (PEG)-8000 and resuspending the protein pellet in a K +- and MgZ+-containing buffer (KMB). The protein was then reconstituted in artificial phospholipid membranes by mixing the PEGprecipitated protein fraction with asolectin (the protein:lipid ratio proving to be critical). After a freezethaw cycle and sonication, the resulting proteoliposomes were pelleted by centrifugation and resuspended in KMB. Protein content was determined as for vesicles, and the recovery of system N catalytic activity was assayed by measuring L-[3H]histidine uptake in the proteoliposomes. As for liver membrane vesicles, histidine

The study of membrane transport began over 80 years ago (e.g., Van Slyke and Meyer, 1913-1914) and, not surprisingly, investigators have adopted a broad range of techniques drawn from several different disciplines. Given this breadth, my intention is not that this chapter should be a complete manual, but rather a map to other resources. More detailed comments relate mainly to the most novel areas and to the author's bias toward the use of voltage-clamp techniques to study overexpression of transport proteins in Xenopus oocytes. II. PURIFICATION AND RECONSTITUTION OF TRANSPORT PROTEINS Purification of transport proteins from membrane preparations permits extensive biochemical (including Western) analysis of the transporter. Reconstitution of the protein fraction into liposomes allows investigators to perform functional assays of transport activity (adopting the same kind of radiotracer assays and protein determination methods used historically for membrane vesicles; see Section X,B) in order to evaluate the purification. Although the principles are the same, different transport systems have required that methods be adapted to suit individual properties such as hydrophobicity of the transport protein. The first step in purification of the transport activity involves solubilization of membrane preparations using detergentsmthese have included t Bryan Mackenzie, Brigham & Women's Hospital and Harvard Medical School, Boston, Massachusetts 02115

327

328

11. Techniques in Membrane Transport

uptake was Na+-dependent and inhibited by glutamine, but not by the system ASC substrates serine and threonine. Over 90% of the system N activity observed for membrane vesicles was recovered in the proteoliposomes, with a 20-fold enrichment. Several purification-reconstitution studies have offered insights into the structures of transport proteins. For example, the system A amino acid transporter appears to associate with the membrane-cytoskeletal components ankyrin and fodrin (Handlogten et al., 1996) and the a3-subunit of the integrin c~3/31(McCormick and Johnstone 1995). Purification-reconstitution techniques have also been adopted to study the Na+/glucose cotransporters from small intestine and kidney (reviewed in Semenza et al., 1984), a placental taurine transporter (Ramamoorthy et al., 1993), a synaptic vesicle glutamate transporter (Lewis and Ueda, 1998), vesicular monoamine transporters (Yelin and Schuldiner, 1998), and a plant H+/glutamine cotransporter (Weston et al., 1994). Why the zwitterionic amino acid transporters have been such frequent targets of purification-reconstitution studies may in part be due to the difficulties investigators have experienced in attempts at their expression cloning. However, purification-reconstitution studies will continue to be worthwhile even after cDNA cloning for at least two reasons. First, purification of transport proteins from overexpression systems may result in sufficiently high activity in proteoliposomes to facilitate biochemical or functional studies not presently feasible. For example, reconstitution of the cloned Na+/ H + antiporters NhaA and NhaB from E. coli permitted functional studies (e.g., determination of Na+/H + stoichiometry) that were not easy to perform in intact cells or vesicles (reviewed in Padan and Schuldiner, 1994). Second, in order to confidently ascribe a cDNA product to a particular transport system activity, the transport protein expressed in heterologous cells (e.g., cRNAinjected oocytes) will need to be reconstituted in proteoliposomes to prove finally that it is the transport catalytic agent per se. in. METHODS FOR ISOLATING cDNAs CODING FOR TRANSPORT PROTEINS Many membrane transport proteins have proved difficult to purify by conventional separation techniques due mainly to their hydrophobicity and often low abundance in membranes. Such circumstances impeded cDNA cloning either by amino acid sequencing of peptides to generate probes with which to screen a cDNA library or by generating antisera for the immune screening of a phage A cDNA expression library for the protein. Where no sequence information is available, trans-

port proteins may be cloned by relying on a functional assay; that is, "expression cloning." Expression cloning remains the only means by which we can isolate cDNAs encoding transporters in completely novel gene families, aside from more serendipitous discoveries that previously cloned cDNAs encode transporters. For example, it was found at the beginning of this decade that a cell-surface receptor, by which the murine leukemia retrovirus gains entry to the host cell, was a cationic amino acid transporter, CAT1 (Kim et al., 1991; Wang et al., 1991). A second isoform, CAT2 (Kakuda et al., 1993), had previously been identified as the T-cell early activation (Tea) gene product. Expression cloning typically involves the overexpression of cDNA libraries in mammalian host cell lines (Simonsen and Lodish, 1994) or RNA-injected amphibian oocytes (Sigel, 1990), coupled with a functional assay (e.g., radiotracer uptake) to identify the targeted transport activity. Sequential subdivision and screening of the cDNA library leads to the identification of a single clone, as illustrated for the recombinant vaccinia virus/ HeLa expression system (Fig. 11.1; see also Section IV,B). Detailed, up-to-date protocols for expression cloning using the X e n o p u s oocyte system appear elsewhere (Romero et al., 1998) and are summarized here. Because of the relative low abundance of many membrane transport proteins it has been necessary to construct plasmid libraries using cDNA synthesized from an enriched mRNA fraction. Poly(A) + RNA is sizeselected using sucrose density-gradient fractionation or preparative gel electrophoresis (Romero et al., 1998) and a positive fraction identified that stimulates radiotracer uptake in RNA-injected oocytes. Further enrichment may be possible by employing hormonal or dietary manipulations in an effort to specifically upregulate transporter mRNA. For example, system A-like neutral amino acid transport activity was at least doubled in oocytes injected with liver poly(A) + RNA isolated from glucagon-treated rats (Palac/n et al., 1990; Tarnuzzer et al., 1990) compared with RNA from normal rats. More recently, a rat intestinal H+-coupled metal-ion transporter was cloned from animals fed an iron-deficient diet for 3-4 weeks (Gunshin et aL, 1997). Since the success of expression cloning depends upon obtaining sufficient expression of exogenous transport activity over background (endogenous) levels, the latter may influence the choice of expression system. Unfortunately, endogenous systems have not always been characterized carefully, but knowledge of endogenous transport characteristics, as for amino acid transport in X e n o p u s oocytes (Campa and Kilberg, 1989; Mackenzie et al., 1994a; Taylor et al., 1989; Van Winkle, 1993), can significantly improve the design of the expression assay. For example, the predominant L-glutamine transporter

Heterologous Expression Systems for Transport Proteins

329

rat thyroid Na+/I - cotransporter (Dai et aL, 1996), and an adipocyte long-chain fatty acid transporter (FATP) (Schaffer and Lodish, 1994) to name only a few. Yeast mutants have been useful in the expression cloning of plant transporters (reviewed in Rentsch et aL, 1998), such as a peptide transporter from an Arabidopsis cDNA library. In this case, the cloned transporter functionally complemented a yeast peptide transport mutant (Steiner et al., 1994). Meanwhile, homology screening and PCR technology permitted the cloning of subtypes and isoforms related to transporters isolated by expression cloning. For example, the cDNA encoding SGLT1 (which displays high affinity for glucose) was used to screen a pig kidney LLC-PK1 cell line cDNA library under low-stringency conditions, leading to the isolation of a low affinity glucose transporter with 76% identity at the amino acid level (Kong et aL, 1993; Mackenzie et aL, 1994b).

IV. HETEROLOGOUS EXPRESSION SYSTEMS FOR TRANSPORT PROTEINS FIGURE 1 1.1 Use of vaccinia virus-encoded T7 RNA polymerase for cDNA library expression cloning of membrane transporters. cDNA libraries can be constructed in plasmid vectors downstream of the T7 RNA polymerase promoter. Meanwhile HeLa cells are infected with recombinant vaccinia virus containing the bacteriophage T7 RNA polymerase (under the control of the viral promoter P7.5) and, after 30 min, plated in 6- or 24-well plates. The viral-infected HeLa cells (in separate wells) are then transfected with individual pools of plasmid DNA. After 12-24 hr, virus and DNA are removed and the HeLa cells assayed for transport activity. This expression system is also frequently used to study the transport characteristics of individual clones (see Section III) (reproduced from Blakely et al. (1991) with permission from Academic Press).

in oocytes displays a strict requirement for Na § (Taylor et al., 1989), whereas the hepatic glutamine transporter system N tolerates Li+-for-Na § substitution. This distinction was exploited in the expression assay for system N (Taylor et al., 1992). The relative apparent Km (or K0.5) values for the test substrate may also be considered, as discussed in Section IV,D,3. Expression of heterologous RNA in oocytes was pioneered by Gurdon et al. (1971) and the first plasma membrane protein to be expressed in oocytes was an acetylcholine receptor (Sumikawa et al., 1981). Identification of the rabbit intestinal Na§ cotransporter SGLT1 by expression cloning (Hediger et al., 1987) led in the isolation of many cDNAs encoding transporters including the salamander renal electrogenic Na+/HCO~ cotransporter (Romero et al., 1997), the rabbit intestinal H+/peptide cotransporter (PEPT1) (Fei et al., 1994), the

A. General Comments The heterologous expression of transport proteins in prokaryotic or eukaryotic cells has been central in their cDNA cloning and structural and functional characterization. The criteria for a good expression system include the ease of use for transport experiments, the levels of expression (especially compared with background), and the amount of recombinant DNA work and cRNA synthesis involved (especially relevant in testing mutant or chimeric proteins). Each expression system has its strengths and limitations. When the importance of cellular processing of exogenous proteins is considered (e.g., protein folding, glycosylation, trafficking, and subunit assembly), eukaryotic and prokaryotic systems may be best applied to their own cell types. Several viral antigen/host cell systems are listed in Table 1 of Simonsen and Lodish (1994). The baculovirus/Sf9 insect cell expression system has been used in the expression of aquaporin-4 (Yang et al., 1997) and neurotransmitter transporters (Tate, 1998); the methylotrophic yeast Pichia pastoris expression system (Sreekrishna et al., 1997) has been used to express the mammalian peptide transporter PEPT1 (D6ring et al., 1997). Here we review two systems used for the transient expression of membrane transport proteins in established mammalian cell lines. We then revisit the X e n o pus oocyte expression system and, in the following sections, review biophysical approaches to studying cloned transporters expressed in X e n o p u s oocytes.

330

11. Techniques in Membrane Transport

B. Vaccinia Virus/HeLa Cell Expression System Fuerst et aL (1986) developed an infectious recombinant vaccinia virus that efficiently synthesizes T7 RNA polymerase. Bacteriophage T7 gene 1 was introduced into the genome of vaccinia virus under the control of the viral P7.5 promoter, which would ensure that T7 RNA polymerase was synthesized in high amounts in virus-infected monkey kidney CV-1 cells. When the virus-infected CV-1 cells were transfected with a plasmid containing the lacZ gene (downstream from the T7 promoter), Fuerst et al. obtained very high expression of/3-galactosidase. Among its several advantages, the vaccinia virus has a broad host range; it has a stable genome and possesses capping and methylating enzymes and poly(A) + RNA polymerase, which ensure mRNA stability in the host cell. Blakely and colleagues (Blakely et al., 1991) infected HeLa cells with recombinant vaccinia virus (VTF-7) expressing T7 RNA polymerase (Fig. 11.1). They constructed plasmids containing cDNA inserts encoding the Na+/glucose cotransporter SGLT1 or a Na+-coupled y-aminobutyric acid (GABA) transporter, each under the control of the T7 RNA polymerase promoter. Twelve hours after liposome-mediated transfection of viral-infected HeLa cells with a pBluescript SKII(-) plasmid containing the SGLT1 cDNA, the cells exhibited significantly elevated a-methyl-D-[14C]glucopyranoside (aMG) uptake. This was concomitant with a reduction in the endogenous D-aspartate transport activity. The aMG transport activity was Na + dependent, displayed a K0.5 value for aMG of 0.3 mM, and was inhibited by phlorizin, all of which are consistent with expression of the SGLT1 Na+/glucose cotransporter. Inserting the SGLT1 cDNA into a plasmid containing a 5' stem-loop structure and a 3' transcription terminator increased expression of SGLT1 even further. The expression of SGLT1 using the VTF-7/HeLa cell system was around 10-fold higher than had been obtained using the COS-7 cell system (Birnir et aL, 1990). For the GABA transporter (GAT1), Blakely et al. (1991) obtained transport rates 10a-fold higher than in nonspecific plasmid-transfected cells, in part because of a much smaller contribution of background due to endogenous transport (compared with glucose transport). The authors also described the potential usefulness of this expression system for screening cDNA libraries during the expression cloning of membrane transporters (see Section III and Fig. 11.1) and, more recently, used the VTF/HeLa system in voltage-clamp experiments with GAT1 (Risso et al., 1996). Whereas the VTF-7/HeLa expression system has been rewarding for studying many different membrane transporters, including those for glutamate (Trotti et al., 1996), the

levels of expression of other transporters have been disappointing. For example, the ASC1 transport activity expressed in HeLa cells was only 1- to 3-fold over the endogenous background system ASC activity (Shafqat et al., 1993; Tamarappoo et al., 1996). C. Epstein-Barr Virus Neutral Antigen/pDR2/ HEK 293 Cell Expression System Kilberg's group (Matthews et aL, 1997) set out to find a more efficient expression system that might suit transporters, such as ASC1, and chose human embryonic kidney HEK 293 cells made competent by stable transfection with the Epstein-Barr virus neutral antigen-1 (EBNA-1) cDNA (i.e., 293c18 cells). These 293c18 cells (which are available commercially) were transfected with the pDR2 plasmid containing cDNA inserts encoding ASC1 or the rat glutamate transporter EAAT1 downstream ofthe Rous sarcoma viral promoter, ensuring high rates of transcription of the cDNA insert sequence. The pDR2 plasmid also contains an Epstein-Barr virus origin of replication sequence so that it is stably maintained as an episome for several cell divisions (at 10-20 copies per cell), as well as the hygromycin resistance gene (hyg r) so that incubation with hygromycin B selects only those cells transfected with the pDR2 vector. Expression of ASC1 and EAAT1 in the pDR2/ 293c18 expression system was compared with that in the VTF-7/HeLa system by assaying amino acid transport activity (Table 11.1). The pDR2/293c18 system was roughly twice as effective for both transporters (by comparing increases over background activity). Unlike the VTF-7/HeLa system (Blakely et al., 1991), expression of endogenous cellular proteins does not appear to be affected by pDR2-transfection in 293c18 cells: pDR2transfection (without cDNA insert) did not reduce the endogenous activities of amino acid transport systems X-AG, A, and ASC, nor did ASC1 or EAAT1 expression affect systems X-AC or ASC activities respectively in the pDR2/293c18 system. Clearly, the pDR2/293c18 expression system is a viable alternative to the VTF-7/ HeLa model, at least for some transporters, and the hygromycin selection time is very flexible (even if 100% selection can take up to 10 days). D. X e n o p u s

Oocyte Expression System

1. Introduction

In recent years, the X e n o p u s laevis oocyte has become one of the most popular heterologous expression systems used in the study of membrane transport. Its size and resilience make handling and maintenance easy and facilitate micropipette techniques. Seasonal variability of frog oocytes continues to be a problem for

Heterologous Expression Systems for Transport Proteins

331

TABLE 11.1 Comparison of the Recombinant Vaccinia Virus/HeLa Cell and Epstein-Barr Virus Neutral Antigen/HEK293 Cell (293c18) Systems for the Expression of Amino Acid Transporters ASC 1 and Rat EAATIa Transfected cDNA

Expressionsystem

Vector only

Vector with cDNA i n s e r t

Expressionc.f. background

Na+-Dependent 5 tzM L-[3H]serine uptake/pmol.mg-l(protein).min-1 ASC1

VTF-7/HeLa pDR2/293c18

Rat EAAT1

VTF-7/HeLa pDR2/293c18

119 _ 2 305 ___7 697 + 32 4447 _+ 303 Na+-Dependent 50/zM L-[3H]aspartate uptake/pmol.mg-l(protein).min-1 50 _+_0.2 61 ___5

1106 __+76 2396 ___209

2.6x 6.4x

22x 39x

aCells were transfected with vector only or vector with transporter cDNA insert using pBluescript KS II (HeLa cells) or pDR2 (293c18 cells). Note. Na+-Dependent uptakes are mean _+SEM from three preparations in the case of VTF-7/HeLa cells, four for pDR2/293c18 cells. Serine uptake was measured in the presence of 5 mM 2-(methylamino)isobutyrate to minimize the contribution from the endogenous System A transporter. Data are from Matthews et al. (1997).

many groups. The use of X e n o p u s oocytes as an expression system has been described in detail by Colman (1984b) and I offer here only a brief description and update. Radiotracer techniques in oocytes and methods for the isolation of oocyte membrane proteins are also reviewed in this section, and biophysical approaches are discussed in Sections V and VI. 2. Preparation, Microinjection, and Maintenance of Oocytes Ovarian tissue is removed from the African clawed frog X e n o p u s laevis and placed in a modified Barths' medium (MBM) of composition 88 m M NaC1, 1 m M KC1, 2.4 m M NaHCO3, 0.82 m M MgSO4, 0.66 m M NaNO3, 0.75 m M CaC12, and 10 mM H E P E S and adjusted to pH 7.5 with Tris base. The ovarian tissue is cut or teased open and oocytes are disaggregated and defolliculated using collagenase. To do this, most investigators incubate the oocytes in calcium-flee M B M containing 2 mg.m1-1 collagenase A for 1 to 3 hr (with at least one solution change) with gentle rocking at room temperature followed by repeated rinsing with calcium-flee MBM. Alternatively, oocytes can be defolliculated manually, and this is sometimes done in a hypertonic K + solution. Healthy defolliculate stage V or stage VI (prophasearrested) oocytes have sharply contrasting animal (dark olive-brown) and vegetal (yellow) hemispheres, often with a narrow white band around the equator. Those oocytes with white patches on the animal hemisphere or with diffuse pigmentation should be discarded. Oocytes are maintained at 18~ in M B M with 10 mg liter -1 gentamicin. From anecdotal observations, penicillin, streptomycin, and fuel supplements such as pyruvate are usually unnecessary (but some modification of the medium may have to be made to accommodate specific

features of the cloned transporter being expressed). Oocytes can be injected with R N A (or D N A ) 2-24 hr after defolliculation. For microinjection of oocytes, I favor a positive-displacement system, the D r u m m o n d automated oocyte injector (a variable-volume model is available). Hydraulic systems are also commonly used but these require calibration, at least initially, as described (Colman, 1984b; Cupello et al., 1993). Ready-made micropipettes are also commercially available. R N A should be injected equatorially or at the vegetal pole to avoid damaging the nucleus. Most often 50 ng of c R N A (at 1/zg./z1-1 in sterile water) is injected, although maximal expression can normally be achieved with much less (and which in some cases may be preferable). The time courses for expression of different membrane transport proteins vary greatly, reaching a peak at 2-7 days. Beyond this point, prolonging the life of oocytes by storing them at 4~ for a day or two does not appear to introduce additional problems. Direct injection of D N A into the nucleus is also possible and is routinely used by many laboratories, saving on the time and expense of c R N A synthesis. Cytoplasmic coinjection of X e n o p u s oocytes with recombinant (T7) vaccinia virus and cDNA-containing plasmid vectors has been described for the expression of ion channels (Yang et al., 1991) but has not gained popularity for other types of transport proteins. 3. Radiotracer Assays in Oocytes In screening for the expression of transport proteins, radiotracer techniques normally offer the highest sensitivity and in some cases may be the only approach available to the investigator for characterizing a new clone. It is worth stating some fundamental points which similarly apply to radiotracer assays in other preparations.

332

11. Techniques in Membrane Transport

When maximum sensitivity is required (e.g., when screening a cDNA library), the radiolabeled substrate can be used alone (i.e., in the absence of any cold substrate); however, more useful information may be gained by using a substrate concentration close to (or just below) the expected K0.5 value for the transport activity of interest (especially if this differs from the K0.5 for any endogenous transporters). Increasing the concentration of nonradioactive substrate above the K0.5 is counterproductive: saturating the transport activity will certainly increase the total transport rate but the specific activity of radiolabeled substrate is reduced. Moreover, the initial transport velocities should be measured in these assays. The time course over which uptake increases linearly may be shorter in oocytes expressing high levels of exogenous transporters than in oocytes expressing only endogenous ones. Remarkably, the literature contains examples of assays performed at inappropriate substrate concentrations or in which initial rates may not have been measured. Such poorly designed assays could result in failure to detect expression of a particular transport activity.

4. Isolation of Oocyte Membranes Oocyte membrane preparations are useful for Western blotting (especially as part of the characterization of transporter mutants, see Section VIII) and for assessment of the binding of photoaffinity labels or radioactive ligands to membrane transport proteins. A relatively simple procedure, involving sucrose density-gradient fractionation of oocyte homogenates, allows separation of a crude membrane fraction from cytosolic and nuclear components. This crude membrane fraction comprises endoplasmic reticulum (ER), Golgi apparatus, and plasma membrane; contamination with nuclear membranes can be eliminated by prior enucleation of oocytes (Colman, 1984b). Corey et al. (1994) separated oocyte homogenates on discontinuous sucrose gradients and used marker enzymes to identify subcellular fractions. Using this method, they isolated ER, trans-Golgi, and plasma membranes and demonstrated that GAT1 undergoes translocation from trans-Golgi to plasma membrane following stimulation of protein kinase C in RNAinjected oocytes. V. VOLTAGE-CLAMP TECHNIQUES IN X E N O P U S OOCYTES A. Two-Microelectrode Voltage-Clamp Experiments

1. Introduction Voltage-clamp techniques in Xenopus oocytes offer several advantages over radiotracer techniques alone. While controlling the membrane potential in a single

oocyte, we can rapidly obtain extensive data including saturation kinetics, presteady-state kinetics, estimates of coupling stoichiometry and substrate selectivity, and we can begin to identify voltage-dependent steps in the transporter mechanism. 1 Since multiple measurements can be made using a single oocyte, direct comparisons among the data are possible. For example, we can compare the currents evoked by different substrates or the biophysical parameters obtained before and after treatment, say, with phosphorylation agents. Relatively few oocytes need to be injected, and at the end of voltageclamp experiments the oocytes can be saved for other tests (e.g., Western analysis). In addition to whole-cell and cut-open oocyte voltage-clamp experiments, singlechannel activities have been recorded from outside-out patches excised from oocytes expressing a serotonin transporter (Lin et aL, 1996).

2. Principles of the Conventional Two-Microelectrode Voltage Clamp Many systems transport a substrate or combination of substrates that carry net charge across the membrane. In such cases transport is rheogenic, generating currents that can be measured using microelectrodes. Under nonvoltage-clamped conditions, these transportermediated currents would result in changes in the membrane potential difference (Vm) and consequently influence other voltage-dependent activities (such as K + conductance) in the oocyte. Measuring substrate-evoked depolarizations does not therefore yield much useful information. By voltage-clamping we can measure the transportermediated currents. The electronics and theory of the two-microelectrode voltage clamp (Fig. 11.2) are discussed elsewhere (Finkel and Gage, 1985; ShermanGold, 1993). Briefly, microelectrodes (with DC resistances, 0.5-5 MI) ) are pulled from borosilicate glass, filled with electrolyte solution (typically 3 M KC1), and connected to the recording system through Ag-AgC1 electrodes. The voltage clamp is applied by an amplifier that compares the Vm signal (from the voltage-sensing electrode, ME1) with the chosen command potential (Vcommand). Its output voltage (which is proportional to e, the difference between Vm and Vcommand)forces current (I) to flow through the current-passing microelectrode (ME2) and into the oocyte. Membrane currents are measured with a virtual ground applied to the bath (via a Ag-AgC1 pellet or wire placed in the bath); for experiments that involve switching to C1--free solutions, the bath electrode must be isolated using a 3 M KC1agar bridge. The current recorded is equal in magnitude 1 Competition between substrates for transport must however be studied by other means.

Voltage-Clamp Techniques in Xenopus Oocytes

FIGURE 1 1.2 Two-microelectrode voltage-clamp experiment in Xenopus oocytes expressing cloned membrane transporters. ME1 and ME2 indicate microelectrodes 1 and 2; A1, unity-gain amplifier; A2, high-gain differential amplifier; ~, gain; Vm, membrane potential difference; Vc. . . . . d, command voltage (test potential); Glc, glucose. See Section V,A,2 for explanation./6~c refers to the additional current required to clamp the oocyte (at Vc. . . . . d ) during Na+/glucose cotransport.

333

at the new Vm. (2) Transporter-mediated presteady-state currents: these were more slowly decaying with time constants (r) of 3 - 7 ms; for other ion-coupled transporters, ~'ranges up to 50 ms. Interpretation of the presteadystate currents is discussed below. (3) Steady-state currents: comparing the currents (after reaching steady state) in the presence (Fig. 11.3A, right side) and absence (Fig. 11.3A, left side) of substrate at each Vm allows us to examine the current/voltage (I/V) relationships for the steady-state evoked currents (Fig 11.3C). Analyzing the I/V relationships and concentration dependence of the currents evoked by a range of sugar concentrations and over a range of extracellular Na + concentrations revealed that the sugar currents and Na + binding were voltage dependent and that SGLT2 operates according to an ordered, simultaneous transport

A

- Glc

800

but opposite in polarity to the sum of net charges migrating across the cell membrane.

0

+ Glc

j ....

I(nA) 3. Transporter-Mediated Currents

After impaling the oocyte with the microelectrodes, it is clamped at a chosen holding potential (Vh) normally close to the resting potential ( - 5 0 to - 6 0 mV). Current can be monitored continuously using a chart recorder, while superfusing test solutions. Such an experiment to measure glutamate saturation of E A A T 3 is shown in Fig. 6.37A of Chapter 6. However, without any additional solution changes, considerably more information can be obtained by applying step changes in Vm (typically for periods of 50-500 ms) before and after the addition of the test substrate; for example, by using the p C L A M P software package and Digidata-1200 interface (both Axon Instruments). p C L A M P version 7 also makes light work of measuring the input resistance (Rin) and capacitance (Cm) of the oocyte membrane. To study a pig kidney Na+/glucose cotransporter SGLT2 (Mackenzie et al., 1996b), step changes in Vm were applied from + 50 m V to - 1 5 0 m V each for 100 ms, first in the absence then in the presence of D-glucose (Fig. 11.3). The currents observed in oocytes expressing SGLT2 comprised three components. (1) Fast capacitive transient currents: since these were observed both in control and RNA-injected oocytes they are not related to the transporter; instead, these are rapidly decaying currents (with half-times of < 1 ms) that represent the current required to charge the m e m b r a n e capacitance

-800

-1600

B

C

Vm (mV)

-150 -100 -50 I

I

I

0

50

]

I

Zm(

+50 -50-150

100 ms

-1000 • (nA) FIGURE 1 1.3 Currents associated with SGLT2 expressed in Xenopus oocytes. (A) The oocyte was superfused in 100 mM NaC1 medium and held at Vh = --50 mV (the holding current is indicated by the dotted horizontal line) and Vmwas stepped to between +50 mV and -150 mV as shown by the protocol in B (for clarity, the currents at alternate voltage steps are omitted), first in the absence of sugar (-Glc, left panel) and then in the presence of 20 mM D-glucose (+Glc, right panel). (C) Subtraction of the steady-state currents (averaged over the 94-100 ms after each voltage step) in the absence of glucose from those in the presence of glucose (in A) yielded the steadystate current/voltage relationship for the glucose-evoked current in 100 mM Na + (evoked currents were absent in choline chloride) (redrawn from Mackenzie et al. (1996b) with permission from the American Society of Biochemistry and Molecular Biology).

334

11. Techniques in Membrane Transport

model in which Na + binds first (see Jauch and L~iuger, 1986). 2 Steady-state evoked currents of up to -1000 nA are typically obtained for ion-coupled transporters overexpressed in oocytes; those obtained with a plant H+/ amino acid cotransporter reached - 1 0 / z A (Boorer and Fischer, 1997). Although endogenous currents evoked by test substrates rarely exceed a few nanoamps, the proper control experiments should still be performed since they can provide important although more subtle information. For example, switching from an extracellular pH (pHo) of 7.5 to one of 5.5 resulted in a small inward current in control oocytes of - 6 +_ 3 nA (mean +_ SD), but four times that in oocytes expressing the H+-coupled metal-ion transporter DCT1 (Gunshin et al., 1997), suggesting that a H + "leak" pathway (uniport) could proceed in the absence of metal-ion substrate. Endogenous conductances in oocytes are documented in a number of useful publications (e.g., Barish, 1983; Miledi et al., 1989; Yao and Tsien, 1997). Inhibition of radiotracer uptakes cannot distinguish between transported substrates and nontransported ligands (blockers). Instead, this requires us to test for possible transport of a radiolabeled form of the inhibitor, which, even if available, is not the most economical way to identify a blocker. Blockers are, however, easily identified in voltage-clamp experiments as they (1) inhibit the substrate-evoked current, (2) block the leak current, and (3), at least competitive inhibitors abolish presteady-state currents, whereas they themselves evoke no inward current. It may, however, be difficult to distinguish a blocker from a substrate with a low velocity of transport using voltage-clamp experiments.

4. Presteady-State Currents in Ion-Coupled Transporters Ion-coupled transporters exhibit presteady-state (transient) currents following step changes in Vm (in the absence of driven substrate). Illustrated for SGLT2 (Fig 11.3), similar transient currents were observed for other Na+-coupled transporters including SGLT1 (Parent et aL, 1992, ID: 25), (Hazama et aL, 1997; Loo et al., 1993), the m y o - i n o s i t o l transporter SMIT (Hager et al., 1995), 2 These techniques and procedures are described under the assumption that the co- or countertransport is known to occur (e.g., that radiolabeled Na + and radiolabeled glucose have been shown to be cotransported, as described for SGLT1 in Section V,C below). In most instances such transport of all putative substrates has not yet been verified directly (e.g., see the discussion of the E A A T family of transport proteins in Chapter 6), in which case inferences such as those described here are provisional. The reader is reminded that the most information can be obtained about transport when both unidirectional and net fluxes are measured using radiotracer and voltage-clamp procedures, respectively, as discussed in Chapter 7 (e.g., see Sections II,C and III).

the GABA transporter GAT1 (Mager et aL, 1993), a serotonin transporter (Mager et al., 1994), the glutamate transporter EAAT2 (Wadiche et al., 1995b), and the thyroid iodide transporter NIS (Eskandari et aL, 1997). They are also common to the H+-coupled transporters, including the peptide transporter PEPT1 (Mackenzie et al., 1996a; Nussberger et al., 1997), the metal-ion transporter DCT1 (Gunshin et al., 1997), and a plant hexose transporter, STP1 (Boorer et al., 1994). These transient currents (charge movements) provide considerable information about the partial reactions within the transport cycle and reflect voltagedependent steps involved in the transport event. With the aid of computer simulation of transport models, transient currents have been attributed to (1) reorientation of the empty, charged transporter within the plane of the membrane and (2) movement of the driving ion (Na + or H +) into or out of the plane of the membrane during the binding/dissociation step (i.e., an "ion-well" effect) (Hazama et al., 1997; Loo et al., 1993; Mackenzie et al., 1996a,b). The models indicate, for example, that in the presence of Na + at Vm = - 5 0 mV, most of the carriers orient toward the extracellular side and bind Na +. Upon stepping Vm to positive potentials and/or reducing [Na+]o, the Na + dissociates and the unloaded charged carrier reorients toward the intracellular face. The transient currents are attenuated in the presence of the driven substrate (more evident by comparing the "off" responses in the absence and presence of glucose in Fig. 11.3) since the transporter is cycling, not transient. Transient currents are also abolished in the presence of blockers, since the transporter is "locked" into an immobile state. The transporter-mediated presteady-state current is isolated (1) by subtracting the capacitive transient (usually by fitting the first few points to an exponential decay) and the steady-state current from the total current record (the "fitted method") or (2) as the difference in total charge in the presence or absence of a specific blocker, such as phlorizin for SGLT1 (Hazama et al., 1997), kainate for EAAT2 (Wadiche et al., 1995b), or SKF89976A for GAT1 (Mager et al., 1993). The presteady-state currents for SGLT2 (Fig. 11.4) were integrated with time (to give charge, Q). The Q/Vm relationship was characteristically S-shaped and could be described by a single Boltzmann function (Eq. (11.1)), although it is more likely the aggregate of at least two Boltzmann functions (one for each voltagedependent step): Qmax = Qdep - Qhyp ( w h e r e Qdep and Qhyp represent the charge at depolarizing and hyperpolarizing limits); V0.5 is the Vm at the midpoint of charge transfer; z is the apparent valence; and F,, R, and T have their usual thermodynamic meanings.

Q - Qhyp_ Omax

1 1 + exp(z[Vm -- V0.5] ?-/R. T ) .

(11.1)

335

Voltage-Clamp Techniques in Xenopus Oocytes

A

+50

~~.~

B

mV - 5 0 -150

10 mM Na +

4 2

Vm(mY) I

"

-150

I

-1

I

I

-

50 -2

200 nA 50 ms

"

Q(nC)

-4

FIGURE 1 1.4 Presteady-state currents associated with SGLT2. (A) Presteady-state currents (recorded in 10 m M NaCI medium in the absence of sugar) were isolated using the fitted method (see text) following step changes in Vm (shown in the top panel) from 3 ms after the voltage step (and filtered at 100 Hz for display). (B) Charge/voltage relationship for 10 m M Na +. The data (mean _+ SEM of six determinations in a single oocyte) were fitted to the Boltzmann relation (solid line, Eq. (11.1)), which gave V0.5 = - 3 4 mV, Q m a x - - 11 nC, and z ~ 1. The -max/SUgarfor the sugar-evoked current was - 6 5 0 nA, so that the turnover rate (I~gar/Qmax) w a s ~60 see -1 (redrawn from Mackenzie et al. (1996b) with permission from the American Society of Biochemistry and Molecular Biology).

Since we have measured the total charge (Qmax) and obtained the apparent valence (z) of the movable charge from the data fit, we can calculate the number of SGLT2 functional units (CT) using the relation Qmax - C y ' z ' e for which e is the elemental charge. 3, 4 For SGLT2, CT was in the order 1011per oocyte, similar to that for other transporters expressed in oocytes, 101~ per oocyte. The validity of using Qmax t o determine the number of transporters in the membrane was confirmed by freezefracture electron microscopy. Zampighi et aL (1995) demonstrated that the density of particles appearing on the P face of the oocyte plasma membrane (up to 5000 p , m -2) matched the density estimated from Qmax measurements for SGLT1 and for a number of channels (whereas particle density in noninjected oocytes was around 200/zm-2). As Qmax is an index of the number of SGLT2 functional units in the oocyte membrane, the ratio of maximal sugar-evoked current (I~ga~, see Eq. 11.3) to Qmax gave a turnover rate of ~60 sec -1 (Fig. 11.4). 5 For other ion-coupled transporters, turnover rates ranged from 10a to 103 sec -a, in contrast to ~ 1 0 6 s e c -1 for most ion channels. The biophysical parameters have already proven useful in studying the characteristics of transporter mutants (see Section VIII,D) as for SGLT1 (Panayotova-Heier3 Apparent valence (z) is essentially identical to z6 described by some investigators, and is a reflection of the sum of movable charges multiplied by the fraction of the membrane field through which they move. 4e

=

--1.60

x

1 0 -19

coulomb.

5 *max/SUgarhas units nanoamps (nA), and 1 A = 1 coulomb.sec -1.

mann et al., 1994) and GAT1 (Bismuth et al., 1997), and the effects of second messengers upon transporter expression in oocytes. As we have seen, measuring Qmax and/max provides information on the number (C w), activity, and turnover rates of the transporter; in addition, membrane capacitance (Cm) is proportional to plasma membrane surface area. When injected into oocytes expressing GAT1, phorbol 12-myristate 13-acetate (PMA, a protein kinase C activator) rapidly caused a doubling of transporter activity (Quick et aL, 1997). This increase was associated with a doubling of the surface transporter number (CT) and a specific increase in Cm, suggesting the exocytic addition of transporters to the plasma membrane, a conclusion which was borne out by Western analysis of plasma membranes. Similar second messenger effects have been reported for SGLT1 (Hirsch et al., 1996), the human dopamine transporter hDAT (Zhu et al., 1997), and the mouse retinal taurine transporter (Loo et al., 1996a). Although heterologous expression of transporters in other cell types may be more like their expression in the native cell than is expression in oocytes, the latter expression system is not irrelevant since the second messenger effects just described were specific to the transporter--and even to the i s o f o r m ~ being expressed (Hirsch et aL, 1996). B. Cut-Open Oocyte Preparation Although somewhat less accessible than the conventional two-microelectrode voltage clamp, the cut-open oocyte preparation combines a number of useful lea-

336

11. Techniques in Membrane Transport

tures. First described in the analysis of K § channels (Taglialatela et al., 1992), the cut-open oocyte preparation involves the electrical isolation of a large "patch" of the oocyte membrane (about one-tenth of the total surface area) for current recording, while continuously perfusing the intracellular face. The principle of voltageclamping is the same as for the two-microelectrode voltage clamp except that membrane potential (Vm) is recorded using a low-access resistance (200 k~) electrode (Costa et al., 1994) at the intracellular face (without puncturing the membrane patch); the same electrode is used for internal perfusion. Also, the "intracellular" milieu is usually set to virtual ground, so under clamp conditions the bath is at 0 - Vmma technical but not functional distinction. As well as providing for very fast clamping of the oocyte membrane, with settling times as low as 50/zsec (Chen et al., 1996; Costa et al., 1994), this approach allows us to control intracellular concentrations. This control becomes important for building kinetic models and it proved valuable in investigating amino acid exchange associated with BAT expression (Coady et al., 1994) and determination of Na§ coupling stop chiometry from reversal potentials (Chen et al., 1995). In general, the cut-open oocyte preparation deteriorates much more rapidly than the whole-cell, two-microelectrode voltage clamp, limiting the amount of data that can be obtained from a single oocyte preparation. Although some improvements have been made (see Costa et al., 1994), it has not been possible to internally perfuse the cut-open oocyte at rates high enough to obtain rapid solution changes. C. Determining Coupling Stoichiometry for Cotransporters Expressed in Oocytes

1. Introduction For cotransporters, the transport of substrate (S) against a concentration gradient is achieved by coupling to the movement of a driving ion or cosubstrate (C) (such as Na § or H § down its electrochemical potential gradient. A knowledge of the C: S coupling stoichiometry may impact our understanding of the physiological roles of cotransporters and provide a focus for structure-function studies. Taking a theoretical system comprising a simple C/S cotransporter (with a neutral substrate and only one driving ion) and with no other pathways for the transport of S (i.e., no "external leaks"), the thermodynamic limit for the concentration gradient of S (i.e., [S]i/[S]o ) is given by the following modification (Eq. (11.2)) of Eq. (3.59) in which A is the transmembrane electrical potential gradient (see Turner, 1990):

[S]i/[S]o <- [([C]o/[C]i)exp(F- A ~ / R . T)]~c.

(11.2)

Taking the case of Na+-coupled systems, if ([C]o/[C]i ) e x p ( F A ~ / R . T ) is kept constant by the Na+/K+-ATPase and the stochastic action of the voltage-gated K + conductance (or additionally the Na§ + exchanger in the case of H+-coupled systems), the concentrative capacity of the transport system is limited by nc. Raising nc from 1 to 2 substantially increases the concentrative capacity of the transporter and makes it a more effective scavenger of extracellular substrate S. However, this is at an increased energy cost to the cell to maintain the electrochemical potential gradient for C. Several approaches exist for determining coupling stoichiometry, and these have been reviewed elsewhere for vesicle systems (Turner, 1990). The "activation" method, which has been used in vesicle preparations and in oocytes, involves measuring fluxes or evoked currents over a range of ligand (S) concentrations to find the least number of molecules of S required to bind to the transporter in each transport cycle, fitting data to Eq. (4.27) (Chapter 4). In our modified equation (Eq. (11.3)) as it applies to evoked currents (I),/max is the maximal evoked current, S is the concentration of S (either S or C above), K0S.5 is the concentration of S at which current is halfmaximal, and nH is the Hill coefficient for S: I =

/max"

SnH

.

(11.3)

(KS.5)~H + S~H The coupling stoichiometry has often been inferred from measurements of nil. For instance, if nH = 2 for Na § nH = 1 for alanine, an "apparent" coupling stoichiometry of 2 Na+:l alanine has been proposed. However, the Hill coefficient ( n i l ) iS no more than an empirical description of cooperativity in ligand binding. Whereas nH may in some cases be equivalent to or close to the coupling stoichiometry no, nH may underestimate nc if there is little or no cooperativity between multiple activator binding sites. There are also cases in which the actual stoichiometry may be less than that predicted by nH (e.g., see ASC-catalyzed transport in Chapter 6). Although nH remains a useful parameter for assessing cooperativity and classifying a particular transport system, the coupling stoichiometry should be confirmed by other methods. Here we review two direct methods for determining coupling stoichiometry under voltage-clamp conditions, each involving overexpression in oocytes which should minimize the contribution from external leaks (i.e., other pathways for transport of S within the cell).

2. Radiotracer Uptake Under Voltage Clamp Overexpression of the Na§ cotransporter SGLT1 in oocytes and measurement of 22Na or amethyl-D-[14C]glucopyranoside (aMG) accumulation in

Voltage-Clamp Techniques in XenopusOocytes

200 gM [14C]c~MG

50nA 3 FIGURE 1 1.5 Measuringthe Na+/glucose coupling coefficient in an oocyte expressing SGLT1. Continuous current recording (sampling at 10 Hz, filtered at 20 Hz) from an oocyte expressing SGLT1,voltageclamped at -70 mV, from the study described in Mackenzie et al. (1998). A stable baseline current was obtained in 100 mM Na+medium before superfusing 200 /zM a-methyl-i>[laC]glucopyranoside for 10 min (indicated by the solid bar) and washing out with Na+medium. The sugar-dependent charge (Q~MG),that is, the integral of the sugardependent current was -7.8 • 10-5 C, equivalent to 810 pmol of monovalent charge. In this oocyte 14C accumulation was 450 pmol, having subtracted the 14C accumulation in choline medium over 10 min in control oocytes (18 _ 1 pmol). We observed neither Na+dependent sugar uptake nor sugar-induced current in control oocytes. Since Q~MGwas shown to be equivalent to the Na+ flux, the Na+/ sugar couplingcoefficient under these conditionswas 1.8 in this oocyte.

voltage-clamped cells permitted direct comparison of simultaneous unidirectional uptakes and cotransporter currents in individual oocytes (Mackenzie et al., 1998). The first step was to confirm the identity of the sugarevoked current and the Na + flux. To do this, the oocyte was placed in a small-volume chamber (<100 ~1), voltage-clamped at Vh = --70 mV, and continuously superfused with a low-Na + (10 m M ) medium. Unlabeled a M G (10 m M ) was added along with 22Na for 10 min before washing out with unlabeled Na + medium. The sugar-evoked current was integrated with time (i.e., the area between baseline current and the current in the presence of sugar) to give the sugar-specific charge (Q~MG), converted to a molar equivalent using the Faraday, and compared with the 22Na accumulation over the same time course. 6 (Corrections were made for the minimal tracer uptakes observed in control oocytes, <3% of the fluxes for SGLT1.) The Q~MG and 22Na accumulation were identical, indicating that the sugarevoked current is carried only by Na + and validating the use of the sugar-evoked current as an index of Na + influx. By then comparing Q~MGwith the [14C]aMG accumulation in 100 mM Na + (as for the example in Fig. 11.5), a Na+/glucose coupling coefficient of 1.6 _+ 0.3 (mean 6Faraday (F) - 9.65 •

10 4

coulombs.mol-a.

337

_+ SD) was obtained at Vh -- --70 mV. The discrepancy from 2 can be explained by a sugar-uncoupled, internal leak pathway through the SGLT1 transporter; the leak (which could be inhibited by phlorizin) was about 5% of the sugar-coupled pathway at low Na+, but it can be up to 20% at 100 m M Na +. The Na + leak proceeds at "baseline" current, but it is abolished in the presence of sugar as coupled transport is the favored pathway (see kinetic models; Brown, 1995; Parent et al., 1992). Therefore, the sugar-dependent current is underestimated according to the magnitude of the leak pathway. In support of this conclusion, coupling coefficients close to 2 were obtained under saturating conditions (i.e., Vh = --110 mV) in which the leak current is much smaller relative to the sugar-dependent current, so the stoichiometry of the c o u p l e d pathway was 2 Na+:l sugar. This method has also been applied to the Na+/iodide cotransporter NIS (Eskandari et aL, 1997). Measuring radiotracer uptake under voltage-clamp in oocytes expressing the neurotransmitter transporters revealed that the currents were much larger than would be expected for coupled Na + transport. This observation helped to establish that these transporters also mediate thermodynamically uncoupled ion conductances (Mager et al., 1994; Sonders and Amara, 1996; Sonders et aL, 1997; Wadiche et al., 1995a).

3. Reversal Potentials

Chen et al. described determination of coupling stoichiometry based upon measuring reversal potentials in oocytes expressing SGLT1 (Chen et aL, 1995). Glucosedependent outward currents (within a reasonable Vm range) should not be expected for SGLT1 using the vsugar two-microelectrode voltage clamp, since ,~0.5 at the internal face (Chen et al., 1995) is nearly three orders of magnitude higher than the intracellular glucose concentration (< 50/xM; Umbach et al., 1990). Reversal potentials are therefore best measured using the cutopen oocyte preparation in which the intracellular ligand concentrations can be fixed. This approach relies upon our understanding that no current will flow through the transporter when the transmembrane electrochemical gradients for each ligand are balanced. For Na+/glucose cotransport, Chen et al. (1995) defined the reversal potential (Vr) according to Eq. (11.4), which can be derived from Eq. (11.2), with the introduction of the term Kc, the glucose concentration at which the Na + leak is equivalent to the glucose-coupled Na + transport (nc is the Na+/glucose coupling coefficient): V r - R . T I n ([sugar]o + Kc)[Na]o~C nc" F ([sugarli + Kc)[Na]~c "

(11.4)

338

11. Techniques in Membrane Transport

[Na]i and [Na]o were kept constant at 50 mM. With an extracellular [aMG] of 2 mM and [c~MG]i of 30 mM, Vm was stepped to between -160 mV and +40 mV, and the Vm at which current was zero (Vr) was determined. According to Eq. (11.4) at 22~ Vr predicted nc = 1.96, indicating a Na+/glucose coupling stoichiometry of 2:1. The coupling coefficient was independent of [aMG]o (in agreement with data for radiotracer uptake under voltage clamp, Mackenzie et aL, 1998) and [c~MG]i. As for the neuronal glutamate transporter EAAT3 (Zerangue and Kavanaugh, 1996a), which cotransports Na § and H § and countertransports K § in each glutamate transport cycle, additional terms can be added to Eq. (11.4), bearing in mind the polarity and direction of transport for each ion species (see Eq. (1) of Zerangue and Kavanaugh, 1996a), and the term Kc can be omitted for transporters for which it is known there is no significant leak pathway. In the case of EAAT3, it should also be remembered that transport of all of the putative ligands remains to be verified using radiotracers or ionselective microelectrodes.

Vl. PROBING TRANSPORT WITH ION-SELECrlVE MICROELECTRODES A valuable approach to probing membrane transport events is the use of ion-selective microelectrodes, which shows high resolution and rapid response times. Investigators commonly make use of ion-exchange resins introduced into silanized, single-barreled high-resistance microelectrodes pulled from borosilicate or aluminosilicate glass. 7 Bowling has described the construction of an intracellular pH ( p H i ) microelectrode in which the tip can be filled with proton cocktail (Bowling, 1989) or H + ionophore I-cocktail B (Fluka) and the microelectrode back-filled with phosphate buffer (pH 7.0) (Romero et aL, 1997). The microelectrode is then connected to an electrometer through a Ag-AgC1 pellet along with a conventional KCl-filled microelectrode as the reference electrode. Each pH-selective microelectrode must be calibrated using a range of buffer solutions of known pH. The slope of the calibration curve for a good pHi electrode will be - 5 5 to - 6 0 mV per pH unit over the useful range. Intracellular voltage and pH-sensing microelectrodes were used in the characterization of the tiger salamander renal electrogenic Na+/HCO~ cotransporter expressed in oocytes (Romero et aL, 1997). With a knowledge of both the intracellular volume and buffering capacity, 7 As for current, ion-selective microelectrodes measure net rather than unidirectional fluxes. Such net fluxes approach unidirectional fluxes at sufficiently steep transmembrane total chemical potential gradients.

reasonable estimates of the net H + influx can be calculated from the observed rate of change in pHi. The marked membrane potential dependence of the protoncoupled metal-ion transporter DCT1 necessitated measuring p H i under voltage-clamp conditions (Gunshin et al., 1997). By simultaneously following pHi and membrane current in oocytes expressing DCT1, we demonstrated both an uncoupled leak pathway carried by H + and intracellular acidification associated with the Fe 2+evoked current (Fig. 11.6). Additional processing of the p H i microelectrode signal was required in this experimental system to correct for the effects of current injection. The method involves signal subtraction and capacitance filtering (to eliminate noise arising from the 100-500 gigaohm resistances of the filled pHi microelectrodes) and will be described in detail elsewhere (M. F. Romero, M. A. Hediger, and W. F. Boron, manuscript in preparation). Intracellular Na + activity (a Na) c a n be measured in much the same way as pHi. Using a Na + resin (Fluka) in microelectrodes backfilled with 150 mM NaC1, Abdulnour-Nakhoul et al. demonstrated that norepinephrine stimulated Na+/K+ATPase activity in the perfused salamander proximal tubule (AbdulnourNakhoul et aL, 1994). Calibration with NaC1 and KC1 solutions indicated that the selectivity of the a Na i electrodes for Na § over K § was 35 to 1, with a slope of

pH o = 5.5 7.4 50 ~tM Fe 2§ 7.2 pHi lOm

7.0

Vh= --90 mV -100 -200

/

(nA)

~ ' -

FIGURE 1 1.6 Simultaneous recording of current and intracellular pH in an oocyte expressing DCT1. An oocyte expressing the metalion transporter DCT1 was impaled with three microelectrodes, ME1 and ME2 (see Section V,A,2 and Fig. 11.2) for voltage-clamping and a pH-sensing microelectrode (see section VI). The oocyte was clamped at -90 mV and superfused with pH 7.5 medium before switching to pHo = 5.5 and adding 50 tzM Fe 2+ for the period shown (redrawn from Gunshin et al. (1997) with permission from Macmillan Magazines Limited).

Structure-Function Studies of Transport Proteins

339

59 mV per 10-fold change in Na + concentration. More recently, this group has measured intracellular activities of Na§ K § and C1- in oocytes expressing the Na § channel ENaC (Nakhoul et al., 1998). Microelectrodes can even be made sensitive to specific charged substrates such as glutamate (Billups et al., 1998). In the next few years, we can probably expect to see an increase in the use of micropipette techniques in studying cloned transporters. The use of Na§ microelectrodes is all the more attractive given that the only real alternative is to use 22Na or 24Na.

pie, demonstrated that di- and tripeptide transport was associated with intracellular acidification (Thwaites et al., 1994; Wenzel et al., 1996). Fluorescence techniques can detect minute cell volume changes as a result of water transport in established cell lines (Alvarez-Leefmans et al., 1997). For oocytes, it may be somewhat easier to follow cell volume changes by measuring crosssectional area obtained from bright-field images. The latter approach was used to measure water transport associated with Na+/glucose cotransport mediated by SGLT1 (Loo et al., 1996b; Meinild et al., 1998).

Vll. OPTICAL METHODS FOR MEASURING MEMBRANE TRANSPORT

Vlll. STRUCTURE-FUNCTION STUDIES OF TRANSPORT PROTEINS

High spatial and temporal resolution are among several advantages that optical methods have over radiotracers in membrane transport studies. Critically reviewed by Verkman (1995), optical methods rely upon measuring the intensity of scattered light or the fluorescence of an entrapped indicator, sensitive to transportrelated changes (e.g., volume, pH, ion or substrate concentration, or membrane potential). In general, optical methods are "time-integrated" so that (at least in the past) voltage-clamp approaches have been preferable for measuring instantaneous changes associated with transport of ions or charged substrates. Application of fluorescence techniques in oocytes has been reviewed elsewhere (Cha et aL, 1998). The potential exists to identify very specific fluorescent probes that may serve as substrates for transporters of interest. For example, the optically active compound N-(2-hydroxymethyl)-6-methoxy-quinolinium (HEMQ +) is structurally similar to organic cations and has a single emission peak at 444 nm, unaffected by pH. Using a microscope-based epifluorescence system (AlvarezLeefmans et al., 1997), Bednarczyk and Wright (1998) observed that basolateral uptake of HEMQ + in superfused isolated renal proximal tubule segments was saturable and inhibited by organic cations. This inhibition and the strong inhibition of [3H]tetraethylammonium (TEA +) uptake by HEMQ + are consistent with the possibility that HEMQ § and organic cations share the same transport system, so HEMQ + may prove to be a useful tool in studying organic cation transport. A very popular membrane-impermeant fluorophore used in transport studies is 2',7'-bis(2-carboxyethyl)5(6)-carboxyfluorescein (BCECF), which can be used either as an indicator of cell volume (fluorescence is directly proportional to fluorophore concentration) or of intracellular pH (pHi) according to the choice of excitation/emission wavelengths. Measurement of pHi in BCECF-loaded oocytes and Caco-2 cells, for exam-

A. General C o m m e n t s A knowledge of the secondary and, if possible, tertiary structures of transport proteins is invaluable for designing experiments that may identify their key structures (e.g., substrate binding sites, coupling mechanisms, and translocation pathways). Three-dimensional crystal structures have been determined for several membrane proteins (Ostermeier and Michel, 1997), but carriertype transporters (i.e., type 2 proteins in Table 8.2 of Chapter 8) are not among them. Nevertheless, a great deal of information has come from noncrystallographic studies, and these have been elegantly described for the H+-coupled/3-galactoside transporter from Escherichia coli (Kaback and Wu, 1998). B. Secondary Structure and M e m b r a n e Topology Several resources are available for the prediction of transmembrane domains and membrane topology (sidedness) of transport proteins. Hydropathy analysis is part of the Genetics Computer Group (GCG) Wisconsin Package, but interpretation of hydrophobicity plots is rather subjective? The neural network PredictProtein boasts >95% accuracy in the prediction of transmembrane helices (Rost et al., 1995) based on multiple sequence alignments and >85% confidence in predicting topology (Rostet aL, 1996). 9 Secondary structure models should then be tested. For example, membrane topology modeling has been performed for proteins of the SGLT cotransporter family (Turk and Wright, 1997) and confirmed by biochemical tests in SGLT1. The SGLT1 primary sequence contains two N-glycosyslation consensus sites. When SGLT1 was translated in vitro in the presence of pancre8 http://www.gcg.com 9 http://www.embl-heidelberg.de/predictprotein/predictprotein.html

340

11. Techniques in Membrane Transport

atic microsomes, glycosylation was detected as a shift in the apparent size of the peptide in the presence or absence of the inhibitor endoglycosidase-H. Using partial transcripts and site-directed mutagenesis, it was revealed that only one (Asn-248) of the two N-glycosylation concensus sites in SGLT1 is glycosylated, confirming that the segment containing residue 248 is extracellular (Hediger et aL, 1991). Turk and colleagues (Turk et aL, 1996) introduced nonnative N-glycosylation sites throughout the peptide in order to identify additional extracellular loops. The N-glycosylation scanning mutants were expressed in X e n o p u s oocytes and glycosylation identified by gel shifts on Western blots. Evidence in support of the topological prediction for the /3-galactoside transporter included the findings from biophysical analyses and expression of a series of transporter-alkaline phosphatase (lacY-phoA) fusion proteins (reviewed in Kaback and Wu, 1998), in w h i c h phoA was used as a reporter enzyme. C. Noncrystallographic Determination of Helix Packing The three-dimensional arrangement of helical structures (helix packing) in the membrane may be critical to the translocation pathway, and interactions between residues from adjacent helices may be more important to molecular mechanisms than adjacent residues in the primary sequence. Modeling of the transmembrane helical organization was recently reviewed (Dieckmann and DeGrado, 1997) and three-dimensional structural modeling was performed for the GLUT1 facilitative glucose transporter (Zeng et aL, 1996). Using oligonucleotide-directed, site-specific mutagenesis in the fl-galactoside transporter, insertion of alanyl residues permitted identification of structurally or functionally important helix segments that were disrupted by introduction of the alanyl residue into the hydrophobic domain (Braun et al., 1997). Excimer fluorescence is one of several techniques (Kaback and Wu, 1998) that were used to determine helix packing in this transporter. Replacement of all eight cysteinyl (Cys) residues resulted in a "Cys-less" protein with 50% of wild-type activity (van Iwaarden et al., 1991). Insertion of a pair of Cys residues into the Cys-less/~-galactoside transporter permitted tagging with a thiol-specific fluorescent probe following solubilization and reconstitution. The fluorophore used was N-(1-pyrenyl)maleimide (PM), which can form an excited-state dimer (excimer) with unique emission if two conjugated ring systems are within 0.35 nm and correctly oriented. Excimer fluorescence identified four residues (Jung et al., 1993) that (1) form interhelical interactions, (2) are critical for H + coupling, and (3) are in close proximity to the putative

substrate translocation pathway. Based on these tertiary structural observations, Kaback has proposed a gratifyingly simple model for the mechanism of lactose coupling to H + translocation (Kaback and Wu, 1998). D. Identifying Key Residues and Structures in Transport Proteins Cysteine-scanning mutagenesis (in which nearly all 417 residues in the Cys-less/~-galactoside transporter have been systematically replaced with Cys) has revealed that replacement of very few residues disrupts transport activity (Kaback and Wu, 1998); those mutants that are affected can normally still bind substrate, and replacement of only four residues abolishes H + coupling. When designing mutagenesis experiments, short of systematically mutating every residue, there ought to be a clear rationale for producing each mutation. For example, histidyl (His) residues (undergoing protonation/deprotonation) may be involved in proton binding or the recognition of charged substrates. Mutation of His-57 in the human H+/peptide cotransporter PEPT1 abolished transport activity in RNA-injected oocytes. That this result was not due to decreased synthesis or impaired trafficking of mutant proteins was demonstrated by comparing confocal images of PEPTl-specific immunofluorescence in the membranes of oocytes expressing the mutant protein with those expressing the wild-type transporter (Fei et al., 1997). Of the SGLT1 mutations that do not result in normal expression of transport activity, most appear to cause trafficking defects (Lam et aL, 1998; Martin et aL, 1996) and may be of limited usefulness in determining transport mechanisms. Use of the broader approaches, such as constructing chimeric proteins, as for PEPT1-PEPT2 (D6ring et al., 1996) and SGLT1-SGLT2 (PanayotovaHeiermann et al., 1996), may therefore be prudent at this stage. The latter study suggested that the C-terminal half of SGLT1 contained the sugar translocation pathway, a hypothesis that was further tested by making a cDNA construct (C5) encoding only transmembrane helices 10-14 (Panayotova-Heiermann et al., 1997). Radiotracer and voltage-clamp methods (see Section V,A), together with electron microscopy and Western analysis, were used to monitor expression of C5 in oocytes and revealed that C5 behaved as a glucose uniporter. Further investigation of the glucose translocation pathway involved sulfhydryl-active MTS (methanethiosulfonate) reagents (Gallardo et al., 1998) with cysteinyl mutants of SGLT1. Accessibility to the MTS reagents was greatest under conditions in which the sugar binding site would be ready to accept sugar, suggesting that each of the mutations was associated with the sugar translocation pathway. Also, comparison of the effects of charged

341

Native Membrane Transport

MTS reagents suggested that, in the A468C mutant, residue 486 is located within the membrane electric field. The possibility now presents itself to use MTS reagents to probe dynamic, ligand-induced conformational changes in transport proteins, similar to studies performed on the/3-galactoside transporter using excimer fluoresence (Jung et aL, 1994; Kaback and Wu, 1998).

continued use ought to be important in evaluating gene knockouts, in pharmacological studies, in addressing clinical problems, and as an emphasis on physiology is restored. Most of the methods are not new and are not therefore reviewed in detail here. Regional and tissue studies and whole-body investigations are also discussed briefly in Sections II,B and III of Chapter 10.

IX. GENETIC APPROACHES TO UNDERSTANDING TRANSPORTER FUNCTION

B. Transport in Vesicles Isolated from Polarized and Nonpolarized Tissues

A broader appreciation of the physiological roles of specific transporters (especially in cases of apparent gene redundancy, such as within the EAAT and CAT transporter families) may come from genetic approaches such as gene ablation, knockout, and transgenic overexpression (see MacLeod, 1996). Gene targeting technology was reviewed by Porter and Dallman (1997) and has been employed in functional studies of the glutamate transporter family (Peghini et al., 1997; Tanaka et al., 1997). For example, mouse EAAT3 was knocked out by homologous recombination in which a neomycin resistance gene cassette was inserted into a fragment containing exon 1 of eaac-1 (which encodes mouse EAAT3), disrupting the eaac-1 gene and permitting positive selection (Peghini et al., 1997). Embryonic stem cells were transfected with the resulting fragment and injected into blastocysts, generating chimeric mice (eaac-1 + / - ) , which were then inbred to yield homozygous eaac-1 - / - mice. A battery of physiological and behavioral tests were used to evaluate the specific effects of knocking out EAAT3. In addition, mRNA hybridization in situ suggested that there was no appreciable compensatory increase in expression of other EAAT family transporters following knockout of mouse EAAT2 (GLT-1) (Tanaka et al., 1997), although such may, of course, not always be the case. Unfortunately, the effects of gene knockouts are seldom limited to a single gene since expression of other proteins may compensate for the deleted activity. For example, the first attempts at knocking out CFTR in mice failed to produce animals that developed lung disease, apparently because of a residual non-CFTR chloride channel activity. This was later remedied in an inbred congenic mouse strain (Kent et al., 1997). X. SUMMARY OF PREPARATIONS USED TO STUDY NATIVE MEMBRANE TRANSPORT

Studies in membrane vesicles have historically been central in the development of new concepts in understanding membrane transport and in characterizing transport systems. Vesicles can be prepared from native or reconstituted membranes, either by differential precipitation with Mg 2+ or by other fractionation methods (Hopfer, 1987; Johnson and Smith, 1988; McGivan, 1992). Marker enzyme assays are used to evaluate the membrane purification in terms of yield, enrichment, subcellular fractionation, and orientation (sidedness) of vesicles. In epithelia, disaccharidases (sucrase or maltase), aminopeptidase M, or alkaline phosphatase have been used as markers for the brush border membrane, and Na+/K+ATPase or hormone-stimulated adenylate cyclase may be present predominantly in the basolateral membrane (Hopfer, 1989; Murer et al., 1989). Such markers permit localization of transport systems to particular membranes. Essentially, the vesicle transport assay involves incubation of membrane vesicles with radiolabeled substrate for a period short enough to reflect the initial transport velocity followed by blocking of further transport (e.g., with an ice-cold, isoosmotic solution) and collection of the membranes by filtration at <0.6 txm. Methods have been reviewed extensively in the above references, and recent modifications include a fast-sampling, rapidfiltration technique (Oulianova and Berteloot, 1996). Notwithstanding the problems associated with vesicle heterogeneity and other limitations of vesicle preparations (Hopfer, 1984), a tremendous amount of information has come from vesicle transport studies. These investigations can be performed under a wide range of experimental conditions and have led, for example, to the identification and confirmation of driving forces such as Na + or proton electrochemical gradients (Ganapathy and Leibach, 1983; Hopfer, 1987; Toggenburger et aL, 1981).

A. Introduction

C. Transport in Freshly Isolated or Cultured Cells

Although molecular cloning of membrane transporters has shifted focus away from the more conventional preparations for studying membrane transport, their

Radiotracer measurements can be performed in freshly isolated cells in suspension or in cells cultured as a monolayer. Examples of their recent use includes

342

11. Techniques in Membrane Transport

salamander retinal glial cells (Barbour et al., 1991), human hepatocytes (Bode et al., 1995), intestinal epithelial cells (Tomita et al., 1995), MDCK cells (Brandsch et al., 1995), and Caco-2 cells (Thwaites et aL, 1994). Epithelial cells form monolayers when cultured on plastic surfaces, and these monolayers exhibit cell polarization characteristics upon reaching confluency. The apical membrane faces upward so that simply measuring radiotracer uptake represents predominantly apical transport. Exploiting this feature, it was demonstrated that polarization of the zwitterionic amino acid transport systems in human placental trophoblast is concomitant with differentiation and membrane specialization (Furesz et al., 1993). Cells are incubated in a physiological saline such as Krebs'-bicarbonate which requires equilibration with 95% 02:5% CO2 to prevent increases in pH. Replacement of bicarbonate with buffers such as HEPES or MOPS may facilitate pH control but will slightly depolarize the cell membrane, altering transport rates (McGivan, 1992). At the end of the incubation period radiolabeled substrate in the medium is separated from the cells by centrifugation of cells in suspension or by removing and washing the remaining medium from monolayers. The contribution of substrate metabolism should be considered in these preparations (McGivan, 1992), although it may not be a factor during initial rate measurements (see also Section X of Chapter 4). Problems of substrate metabolism can be bypassed by using inhibitors (e.g., aminooxyacetate) or by using nonmetabolizable substrates (e.g., 2-aminoisobutyric acid or glycyl-sarcosine), provided they are suitable model substrates. D. Transport in Perfused Tissues and Organs Extraction of substrates from the "blood" compartment in whole tissues or organs is best measured using the paired isotope dilution technique (Boyd and Yudilevich, 1987) in which the perfusate (e.g., Krebs'bicarbonate) contains radiolabeled substrate and a radiolabeled extracellular marker (e.g., mannitol). The extraction is calculated from the differences in radioisotope content before and after a single pass. Intracellular

metabolism of substrates is certainly a consideration in these preparations, which nevertheless seem appealing because they more closely reflect the physiological environment. That they contain mixed cell types (transporting and nontransporting) may reduce the sensitivity of the radiotracer assay; the contributions of intracellular metabolism and extracellular binding may render saturation kinetic studies inaccurate (McGivan, 1992). Still, much information has been gained from the bilaterally perfused placenta (Fisher and Karl, 1990; Mohammed et al., 1993; Page, 1991) and from the rat perfused hindlimb preparation (Hundal et al., 1987) in which practically the only cells involved in a quantitatively significant proportion of the transport are the skeletal muscle myocytes. Other tissues studied (Chapter 10) include perfused jejunal loops (Lister et al., 1995), gastric glands (Boron et al., 1994), and renal tubules (Nakhoul et al., 1990). The sensitivity of intact tissue experiments is limited by the need to perfuse the tissue or organ at high enough rates to ensure sufficient oxygenation, and the preparation will not normally tolerate much variation in the experimental conditions that can be applied (e.g., with respect to [Na+], pH, and temperature).

XI. COMMENTARY The explosion in molecular biological research has greatly impacted the field of membrane transport since the late 1980s to the extent that numerous cDNAs encoding transporters belonging to several different protein familes have been isolated. More transporters continue to be identified by expression cloning or homology screening and, with the genome projects, many other sequences will soon be forthcoming (see Chapter 8). Research has focused on overexpressing cloned transporters in mammalian cell lines or X e n o p u s oocytes and on the transport kinetics, substrate selectivity, tissue distribution, and regulation of cloned transporters. We are now witnessing the resurgence of classic micropipette techniques and the broadening of fluorescence approaches to studying membrane transport and structure-function relationships using the results of molecular biological research.

EPILOGUE

Hal Christensen, The Author Who Inspired This Volume

In 1972 Professor Arnost Kleinzeller, then a member of the Czech Academy of Science, characterized Hal Christensen as having had the courage to conclude very early that membrane transport could be given a molecular description. With that confidence, Christensen carved out with his colleagues a scholarly field so interdisciplinary that he was seen in congresses under the rubrics of physiology, biophysics, nutrition, pharmacology and even gastroenterology, as often as he was seen under his nominal identification with biochemistry. His pioneering work thus helped to establish the current models of interdisciplinary and multidisciplinary investigation in the biological sciences. Christensen's papers have appeared in some 48 journals in all and quite as often in journals at least partly identified with a field other than his primary discipline of biochemistry. Although his early training was highly connected to chemistry and his biological interests were largely hidden in his undergraduate years (between the ages of 15 and 20), his special physiological interests became clearly evident during his doctoral training at Harvard Medical School with Professor A. Baird Hastings. Hastings was himself as much a confessed physiologist as a physical biochemist, and no doubt these tendencies in Christensen were largely acquired from Hastings. It was while Christensen was completing his Masters degree at Purdue in April, 1935, that he received notice of an appointment to a University Fellowship at Harvard to work with Hastings and thus to become a truly interdisciplinary physical biologist. Christensen's early interdisciplinary tendencies are illustrated by his 8 years of membership on the nutrition study sections at the National Institutes of Health, on both research grant and training grant assignments,

in the 1960s. Furthermore, he was often called to serve on special study sections on NIH program projects in fields including gastroenterology, nephrology, neurology, and clinical chemistry. Similarly, he consulted on the assignments of postdoctoral fellowships in biology and agriculture with the NIH in the years 1957 to 1960, and with the Committee on Summer Institutes of the National Science Foundation from 1960 to 1963. His service as a part-time consultant for Burroughs Wellcome, both in London and in Raleigh, North Carolina from 1972 to 1974 continued the interdisciplinary emphasis. Moreover, he reviewed the more molecular biology-based literature for Nutrition Reviews for many years, continuing well into his retirement. As disciplinary barriers continued to shrink in the 1980s, he placed two of his major summaries in Physiological Reviews, and wrote twice for Trends in Pharmacologic Sciences. Such discipline-crossing interests had become the way of modern interdisciplinary scientists. True, during his 26 years in departmental chair positions at Tufts and the University of Michigan in the middle half of this century, Christensen found himself obliged to try to offer an unambiguously biochemical leadership. But even the first 6 years at Tufts included chairing the department of Nutrition along with that of Biochemistry. Activities during many of the Michigan years reinforced his claim to his initial identity as a biochemist. A term as chairman of the Michigan Section of the American Chemical Society, also an office he held in the Michigan College Chemistry Teachers Association, should have presented him unambiguously as a biochemist. For the latter association he did organize one of its annual sessions at Ann Arbor. While serving for some years

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on the Educational Affairs Committee of the American Society of Biological Chemists (ASBC) he also authored a plan for sessions devoted to educational innovations at national meetings, sessions that still continue. Christensen also, innocently, helped introduce to the ASBC (now the ASBMB) from the 1969 Federation of European Biochemical Societies meeting in Madrid, the system of poster presentations, which has come to dominate most such meetings. The disciplinary identities connected to Christensen's service on editorial boards range less widely than his other service, namely for The Journal of Biological Chemistry (1958-1963 and 1980-1985), for Biochimica et Biophysica Acta in Amsterdam (1972-1995), and also for Biomembrane Reviews of the same journal (to mid1990s). In an activity already mentioned, he served as a contributing editor for Nutrition Reviews most of the time from 1975 to 1995. His papers were at one time being cited as many as 600 times per year, a rate still averaging as often as once a day well after his retirement. Overall, however, these citations were distributed through a score of fields, many of them clinically oriented. He served many different journals as an occasional reviewer of submitted scientific papers and also in evaluating research proposals to national agencies of several countries for similarly diverse scientific fields. The title of a book Crossing Boundaries: Biological, Disciplinary, Human, the life story of Baird Hastings, may be relevant to the career of Christensen himself.

Christensen edited Hasting's autobiography and directed it to publication in 1989. These three boundaries were ones that Christensen unrelentingly crossed in fulfilling his lifelong interest in the transport of nutrient molecules across cellular membranes. Hence, it was in meeting the insatiable demands of his research interest that Christensen spread himself among so many disciplines. Christensen disregarded warnings, the first of these from Hastings in the late 1930s, that his record might well become so interdisciplinary that neither biochemistry, physiology, nor nutrition might regard him as "one of their own." Instead, however, these early experiences helped Christensen to reach out to people and their new ideas and to include everyone rather than excluding anyone. His integrity and creativity led him to make these choices early in his professional life. In my own personal experience, I was made to feel immediately welcome in his circle of colleagues despite my late arrival to that group in the last quarter of Christensen's long and illustrious scientific career. Those readers who might have questioned some of the more unusual ideas I have advanced in this book may now recognize that I was attempting in my own way to emulate this great man. Hal Christensen even at age 85, remains open to creative and consequently fragile new ideas. New ideas are of course the grist from which are selected conceptual advancements in all fields including biomembrane transport. This passion to explore is Christensen's legacy to interdisciplinary and multidisciplinary investigations in the biological sciences.

References

Alberts, B., Bray, D., Lewis, J., Raft, M., Roberts, K., and Watson, J. D. (1994). "Molecular Biology of the Cell," 3rd ed. Garland, New York. Alberty, R. A. (1969). Standard Gibbs free energy, enthalpy, and entropy changes as a function of pH and pMg for several reactions involving adenosine phosphates. J. Biol. Chem. 244, 3290-3302. Alexander, S. P. H., and Peters, J. A. (1997). "1997 Receptor and Ion Channel Nomenclature Supplement," 8th ed. Elsevier Trends Journals, Cambridge, UK. Alonso, M. A., Fan, L., and Alarc6n, B. (1997). Multiple sorting signals determine apical localization of a nonglycosylated integral membrane protein. J. Biol. Chem. 272, 30748-30752. Alper, S. L. (1994). The band 3-related AE anion exchanger gene family. Cell PhysioL Biochem. 4, 265-281. Alper, S. L., Kopito, R. R., Libresco, S. M., and Lodish, H. F. (1988). Cloning and characterization of a murine band 3-related cDNA from kidney and from a lymphoid cell line. J. BioL Chem. 263, 17092-17099. Alper, S. L., Stuart-Tilley, A., Simmons, C. F., Brown, D., and Drenckhahn, D. (1994). The fodri-ankyrin cytoskeleton of choroid plexus preferentially co-localizes with apical Na+K+-ATPase rather than with basolateral anion exchanger AE2. J. Clin. Invest. 93, 1430-1438. Altschul, S. F., Gish, W., Miller, W., Myers, E. W., and Lipman, D. J. (1990). Basic local alignment search tool. J. Mol. Biol. 215, 403-410. Altschul, S. F, Madden, T. L., Sch~iffer, A. A., Zhang, J., Zhang, Z., Miller, W., and Lipman, D. J. (1997). Gapped BLAST and PSI-BLAST: A new generation of protein database search programs. Nucleic Acids Res. 25, 33893402. Alvarez-Leefmans, F. J., Altamirano, J., and Crowe, W. E. (1997). Measurement of volume changes in single cells with fluorescent probes. AxoBits 20, 10-13. Amara, S. G. (1992). Neurotransmitter transporters: A tale of two families. Nature 360, 420-421. Ammala, C., Moorhouse, A., Gribble, F. Ashfield, R., Proks, P., Smith, P. A., Sakura, H., Coles, B., Ashcroft, S. J., and Ashcroft, F. M. (1996). Promiscuous coupling between the

Aaltonen, L. A., Peltomaki, P., Leach, F. S., Sistonen, P., Pylkkanen, L., Mecklin, J.-P., Jarvinen, H., Powell, S. M., Jen, J., Hamilton, S. R., Peterson, G. M., Kinzler, K. W., Vogelstein, B., and de la Chapelle, A. (1993). Clues to the pathogenesis of familial colorectal cancer. Science 260, 812-816. Abdulnour-Nakhoul, S., Khuri, R. N., and Nakhoul, N. L. (1994). Effect of norepinephrine on cellular sodium transport in A m b y s t o m a kidney proximal tubule. Am. J. Physiol. 267, F725-F736. Abrahams, J. P., Leslie, A. G. W., Lutter, R., and Walker, J. E. (1994). Structure at 2.8/k resolution of F1-ATPase from bovine heart mitochondria. Nature 370, 621-628. Abumrad, N. N., Williams, P., Frexes-Steed, M., Geer, R., Flakoll, P., Cersosima, E., Brown, L. L., Melki, I., Bulus, N., Hourani, H., Hubbard, M. and Ghishan, F. (1989). Interorgan metabolism of amino acids in vivo. Diabetes Metab. Rev. 5, 213-226. Aharonowitz, O., and Granot, Y. (1996). Stimulation of mitogen-activated protein kinase and Na+/H + exchanger in human platelets: Differential effect of phorbol ester and vasopressin. J. Biol. Chem. 271, 16494-16499. Ahmed, A., Peter, G. J., Taylor, P. M., Harper, A. A., and Rennie, M. J. (1995). Sodium-independent currents of opposite polarity evoked by neutral and cationic amino acids in neutral and basic amino acid transporter cRNA-injected oocytes. J. Biol. Chem. 270, 8482-8486. Ahn, J., Mundigl, O., Muth, T. R., Rudnick, G. and Caplan, M. J. (1996). Polarized expression of GABA transporters in Madin-Darby canine kidney cells and cultured hippocampal neurons. J. Biol. Chem. 271, 6917-6924. Akedo, H., and Christensen, H. N. (1962). Nature of insulin action on amino acid uptake by the isolated diaphragm. J. Biol. Chem. 237, 118-122. Akerboom, T. P. M., and Sies, H. (1992). Interorgan glutathione transport. In "Mammalian Amino Acid Transport: Mechanisms and Control" (M. S. Kilberg and D. H~iussinger, eds.), pp. 283-294. Plenum, New York/London. Albers, R. W. (1967). Biochemical aspects of active transport. Annu. Rev. Biochem. 36, 727-756. Alberts, B., Bray, D., Lewis, J., Raft, M., Roberts, K., and Watson, J. D. (1989). "Molecular Biology of the Cell," 2nd ed. Garland, New York.

345

346

References

sulphonylurea receptor and inwardly rectifying potassium channels. Nature 379, 545-548. Andersen, J. P., and SCrensen, T. (1996). Site-directed mutagenesis studies of energy coupling in the sarcoplasmic reticulum CaZ+-ATPase. Biochim. Biophys. Acta 1275,118-122. Andersen, J. P., and Vilsen, B. (1995). Structure-function relationships of cation translocation by Ca 2+- and Na+,K +ATPases studied by site-directed mutagenesis. FEBS Lett. 359, 101-106. Andersen, J. P., and Vilsen, B. (1998). Structure-function relationships of the calcium binding sites of the sarcoplasmic reticulum CaZ+-ATPase. Acta Physiol. Scand 163(Suppl. 643), 45-54. Anderson, M. P., and Welsh, M. J. (1992). Regulation by ATP and ADP of CFTR chloride channels that contain mutant nucleotide-binding domains. Science 257, 1701-1704. Anderson, M. P., Berger, H. A., Rich, D. P., Gregory, R. J., Smith, A. E., and Welsh, M. J. (1991). Generation of cAMP-activated chloride currents by expression of CFTR. Science 251, 679-682. Anderson, J. A., Huprikar, S. S., Kochian, L. V., Lucas, W. J., and Gaber, R. F. (1992). Functional expression of a probable Arabidopsis thaliana potassium channel in Saccharomyces cerevisiae. Proc. Natl. Acad. Sci. USA 89, 37363740. Andr6, B. (1995). An overview of membrane transport proteins in Saccharomyces cerevisiae. Yeast 11, 1575-1611. Aquila, H., Link, T. A., and Klingenberg, M. (1987). Solute carriers involved in energy transfer of mitochondria form a homologous protein family. FEBS Lett. 212, 1-9. Arriza, J. L., Eliasof, S., Kavanaugh, M. P., and Amara, S. G. (1997). Excitatory amino acid transporter 5, a retinal glutamate transporter coupled to a chloride conductance. Proc. Natl. Acad. Sci. USA 94, 4155-4160. Arriza, J. L., Kavanaugh, M. P., Fairman, W. A., Wu, Y.-N., Murdoch, G. H., North, R.A., and Amara, S.G. (1993). Cloning and expression of a human neutral amino acid transporter with structural similarity to the glutamate transporter gene family. J. BioL Chem. 268, 15329-15332. Asano, T., Katagiri, H., Takata, K., Lin, J. L., Ishihara, H,, Inukai, K., Tsukuda, K., Kikuchi, M., Hirano, H., Yazaki, Y., and Oka, Y. (1991). The role of N-glycosylation of GLUT1 for glucose transport activity. J. Biol. Chem. 266, 24632-24636. Atkins, G. L., and Nimmo, I. A. (1980). Current trends in the estimation of Michaelis-Menten parameters. A n a l Biochem. 104, 1-9. Aulak, K. S., Liu, J., Wu, J., Hyatt, S. L., Puppi, M., Hennings, S. J., and Hatzoglou, M. (1996). Molecular sites of regulation of expression of the rat cationic amino acid transporter gene. J. Biol. Chem. 271, 29799-29806. Auland, M. E., Roufogalis, B. D., Devaux, P. F., and Zachowski, A. (1994). Reconstitution of ATP-dependent amino phospholipid translocation in proteoliposomes. Proc. Natl. Acad. Sci. USA 91, 10938-10942. Bae, Y. M., Holmgren, E., and Crawford, I. P. (1989). Rhizobium meliloti anthranilate synthase gene: Cloning, sequence and expression in Escherichia coli. J. Bacteriol. 171, 3471-3478.

Baldwin, S. A. (1993). Mammalian passive glucose transporters: Members of an ubiquitous family of active and passive transport proteins. Biochim. Biophys. Acta 1154, 17-49. Banff, S., Servadio, A., Chung, M.-Y., McCall, A. E., Orr, H. T., and Zoghbi, H. Y. (1994). Characterization of the gene causing type 1 spinocerebellar ataxia and identification of the murine homolog. Am. J. Hum. Genet. 55, A17. Bannai, S. (1986). Exchange of cystine and glutamate across the plasma membrane of human fibroblasts. J. Biol. Chem. 261, 2256-2263. Bannai, S. and Tateishi, N. (1986). Role of membrane transport in metabolism and function of glutathione in mammals. J. Membr. Biol. 89, 1-8. Barbour, B., Brew, H., and Attwell, D. (1991). Electrogenic uptake of glutamate and aspartate into glial cells isolated from the salamander (Ambystoma) retina. J. Physiol. 436, 169-193. Barbry, P., and Hofman, P. (1997). Molecular biology of Na + absorption. Am. J. Physiol. 273, G571-G585. Barish, M. E. (1983). A transient calcium-dependent chloride current in the immature Xenopus oocyte. J. Physiol. 432, 309-325. Bartkiewicz, M., Hernando, N., Reddy, S. V., Roodman, G. D., and Baron, R. (1995). Characterization of the osteoclast vacuolar H+-ATPase B-subunit. Gene 160, 157-164. Bartl, F., Deckers-Hebestreit, G., Altendoff, K., and Zundel, G. (1995). The Fo complex of the ATP synthase of Escherichia coli contains a proton pathway with large proton polarizability caused by collective proton fluctuation. Biophys. J. 68, 104-110. Barzilay, M., and Cabantchik, Z. I. (1979). Anion transport in red blood cells. II. Kinetics of reversible inhibition by nitroaromatic sulfonic acids. Membr. Biochem. 2, 255-281. Bass6, F., Gaffet P., Rendu, F., and BienvenQe, A. (1993). Translocation of spin-labeled phospholipids through plasma membrane during thrombin- and ionophore A23187induced platelet activation. Biochemistry 32, 2337-2344. Baukrowitz, T., and Yellen, G. (1995). Modulation of K + current by frequency and external [K+]: A tale of two inactivation mechanisms. Neuron 15, 951-960. Baukrowitz, T., and Yellen, G. (1996). Use-dependent blockers and exit rate of the last ion from the multi-ion pore of a K § channel. Science 271, 653-656. Bear, C. E., Li, C., Kartner, N., Bridges, R. J., Jensen, T. J., Ramjeesingh, M., and Riordan, J. R. (1992). Purification and functional reconstitution of the cystic fibrosis transmembrane conductance regulator (CFTR). Cell 68, 809-818. Beckman, D. A., Brent, R. L., and Lloyd, J. B. (1994). Pinocytosis in the rat visceral yolk sac: Potential role in amino acid nutrition during the fetal period. Placenta 15,171-176. Beckman, D. A., Brent, R. L., and Lloyd, J. B. (1996). Sources of amino acids for protein synthesis during early organogenesis in the rat. 4. Mechanisms before envelopment of the embryo by the yolk sac. Placenta 17, 635-641. Beckman, D. A., Brent, R. L., and Lloyd, J. B. (1997). Leucine sources for the rat fetus. Placenta 18, 79-82. Beckman, D. A., Purgarelli, J. E., Jensen, M., Koszalka, T. R., Brant, R. L., and Lloyd, J. B. (1990). Sources of amino

References acids for protein synthesis during early organogenesis in the rat. 1. Relative contributions of free amino acids and of proteins. Placenta 11, 109-121. Beckman, D. A., Purgarelli, J. E., Koszalka, T. R., Brent, R. L., and Lloyd, J. B. (1991). Sources of amino acids for protein synthesis during early organogenesis in the rat. 2. Exchange with amino acid and protein pools in embryo and yolk sac. Placenta 12, 37-46. Beckmann, J. S., and Weber, J. L. (1992). Survey of human and rat microsatellites. Genomics 12, 627-631. Becq, F., Jensen, T. J., Chang, X. B., and Savoia, A. (1994). Phosphatase inhibitors activate normal and defective CFTR chloride channels. Proc. Natl. Acad. Sci. USA 91, 9160-9164. Becq, F., Verrier, B., Chang, X. B., Riordan, J. R., and Hanrahan, J. W. (1996). cAMP- and Ca2+-independent activation of cystic fibrosis transmembrane conductance regulator channels by phenylimidazothiazole drugs. J. Biol. Chem. 271, 16171-16179. Bednarczyk, D., and Wright, S. H. (1998). A novel, fluorescent compound for use in the study of renal organic cation (OC) transport. FASEB J. 12, A424-A424. [Abstract]. Beguin, P., Hasler, U., Beggah, A., Horisberger, J.-D., and Geering, K. (1998). Membrane integration of Na,K-ATPase a-subunits and /3-subunit assembly. J. Biol. Chem. 273, 24921-24931. Beguin, P., Wang, X., Firsov, D., Puoti, A., Claeys, D., Horisberger, J.-D., and Geering, K. (1997). The 3/subunit is a specific component of the Na, K-ATPase and modulates its transport function. E M B O J. 16, 4250-4260. Beliveau, R., Demeule, M., and Potier, M. (1988). Molecular size of the Na+/H § antiport in renal brush border membranes, as estimated by radiation inactivation. Biochem. Biophys. Res. Commun. 152, 484-489. Beliveau, R., Demeule, M., Jette, M., and Potier, M. (1990). Molecular sizes of amino acid transporters in the luminal membrane of kidney cortex, estimated by the radiation inactivation method. Biochem. J. 268, 195-200. Benos, D. J., Fuller, C. M., Shlyonsky, V. G., Berdiev, B. K., and Ismailov, I. I. (1997). Amiloride-sensitive Na § channels: Insights and outlooks. News Physiol. Sci. 12, 55-61. Berger, H. A., Anderson, M. P., Gregory, R. J., Thompson, S., Howard, P. W., Maurer, R. A., Mulligan, R. C., Smith, A. E., and Welsh, M. J. (1991). Identification and regulation of the cystic fibrosis transmembrane conductance regulator-generated chloride channel. J. Clin. Invest. 88, 1422-1431. Berger, H. A., Travis, S. M., and Welsh, M. J. (1993). Regulation of the cystic fibrosis transmembrane conductance regulator C1- channel by specific protein kinases and protein phosphatases. J. Biol. Chem. 268, 2037-2047. Berger, J., Biswas, C., Vicario, P. P., Strout, H. V., Saperstein, R., and Pilch, P. F. (1989). Decreased expression of the insulin-responsive glucose transporter. Nature 340, 70-72. Berk, P. D. (1996). How do long-chain free fatty acids cross cell membranes? Proc. Soc. Exp. Biol. Med. 212, 1-4. Berk, P. D., Zhou, S.-L., Kiang, C.-L., Stump, D., Bradbury, M., and Isola, L. M. (1997). Uptake of long chain free fatty

347

acids is selectively up-regulated in adipocytes of Zucker rats with genetic obesity and non-insulin-dependent diabetes mellitus. J. Biol. Chem. 272, 8830-8835. Berkelman, T., Houtchens, K. A., and DuPont, F. M. (1994). Two cDNA clones encoding isoforms of the B subunit of the vacuolar ATPase from barley roots. Plant Physiol. 104, 287-288. Bertran, J., Testar, X., Zorzano, A., and Palacfn, M. (1994). A new age for mammalian plasma membrane amino acid transporters. Cell. Physiol. Biochem. 4, 217-241. Bertrand, B., Wakabayashi, S., Ikeda, T., Pouyssegur, J., and Shigekawa, M. (1994). The Na+/H + exchanger isoform 1 (NHE1) is a novel member of the calmodulin-boding proteins: Identification and characterization of the calmodulinbinding sites. J. Biol. Chem. 269, 13703-13709. Betz, A. L., and Goldstein, G. W. (1978). Polarity of the blood-brain barrier: Neutral amino acid transport into isolated brain capillaries. Science 202, 225-227. Bezanilla, F., and Stefani, E. (1994). Voltage-dependent gating of ionic channels. Annu. Rev. Biophys. Biomol. Struct. 23, 819-846. Bianchi, L., Roy, M.-L., Taglialatela, M., Lundgren, D. W., Brown, A. M., and Ficker, E. (1996). Regulation by spermine of native inward rectifier K § channels in RBL-1 cells. J. Biol. Chem. 271, 6114-6121. Bianchini, L., and Pouyssegur, J. (1996). Regulation of the Na+/H + exchanger isoform NHEI: Role of phosphorylation. Kidney Int. 49, 1038-1044. Bianchini, L., L'Allemain, G., and Pouyssegur, J. (1997). The p42/p44 mitogen-activated protein kinase cascade is determinant in mediating activation of the Na+/H + exchanger (NHE1 isoform) in response to growth factors. J. Biol. Chem. 272, 271-279. Bienengraeber, M., Echtay, K. S., and Klingenberg, M. (1998). H + transport by uncoupling protein (UCP-1) is dependent on a histidine pair, absent in UCP-2 and UCP-3. Biochemistry 37, 3-8. Billups, B., Szatkowski, M., Rossi, D., and Attwell, D. (1998). Patch-clamp, ion-sensing and glutamate-sensing techniques to study glutamate transport in isolated retinal glial cells. In "Methods in Enzymology" (S. G. Amara, ed.), Vol. 296, pp. 617-632. Academic Press, San Diego. Biolo, G., Chinkes, D., Zheng, X. J., and Wolfe, R. R. (1992a). A new model to determine in vivo the relationship between amino acid transmembrane transport and protein kinetics in muscle. J. Parenterol. Enteral. Nutr. 16, 305-315. Biolo, G., Maggi, S. P., Williams, B. D., Tipton, K. D., and Wolfe, R. R. (1995a). Increased rates of muscle protein and amino acid transport after resistance exercise in humans. Am. J. Physiol. 268, E514-E520. Biolo, G., Tessari, P., Inchiostro, S., Bruttomesso, D., Fongher, C., Sabadin, L., Fratton, M. G., Valerio, A., and Tiengo, A. (1992b). Leucine and phenylalanine kinetics during mixed meal ingestion: A multiple tracer approach. Am. J. Physiol. 262, E455-E463. Biolo, G., Zhang, X-J., and Wolfe, R. R. (1995b). Role of membrane transport in interorgan amino acid flow between muscle and small intestine. Metabolism 44, 719-724.

348

References

Birkenh~iger, R., Hoppert, M., Deckers-Hebestreit, G., Mayer, F., and Altendorf, K. (1995). The Fo complex of the Escherichia coli ATP synthase: Investigation by electron spectroscopic imaging and immunoelectron microscopy. Eur. J. Biochem. 230, 58-67. Birnbaum, M. J. (1989). Identification of a novel gene encoding an insulin-responsive glucose transporter protein. Cell 57, 305-315. Birnbaum, M. J., Haspel, H. C., and Rosen, O. M. (1987). Transformation of rat fibroblasts by FSV rapidly increases glucose transporter gene transcription. Science 235, 14951498. Birnir, B., Lee, H. S., Hediger, M. A., and Wright, E. M. (1990). Expression and characterization of the intestinal Na+/glucose cotransporter in COS-7 cells. Biochim. Biophys. Acta 1048, 100-110. Bismuth, Y., Kavanaugh, M. P., and Kanner, B. I. (1997). Tyrosine 140 of the g-aminobutyric acid transporter GAT1 plays a critical role in neurotransmitter recognition. J. Biol. Chem. 272, 16096-16102. Bjcrgls, M., Gjesdal, O., Erickson, J. D., Torp, R., Levy, L. M., Ottersen, O. P., Degree, M., Storm-Mathisen, J., Seeberg, E., and Danbolt, N. C. (1996). Cloning and expression of a neuronal rat brain glutamate transporter. Mol. Brain Res. 36, 163-168. Black, P. N., and Di Russo, C. C. (1994). Molecular and biochemical analyses of fatty acid transport, metabolism and gene regulation in Escherichia coli. Biochim. Biophys. Acta 1210, 123-145. Blair, A., Ngo, L., Park, J., Paulsen, I. T., and Saier, M. H. (1996). Phylogenetic analyses of the homologous transmembrane channel-forming proteins of the FoFx-ATPases of bacteria, chloroplasts and mitochondria. Microbiology 142, 17-32. Blake, C. (t983). Exons present from the beginning? Nature (London) 3116, 535-537. Blakely, R. D., Clark, J. A., Rudnick, G., and Amara, S. G. (1991). Vaccinia-T7 RNA polymerase expression system: Evaluation for the expression cloning of plasma membrane transporters. A n a l Biochem. 194, 302-308. Bloch, R. (1974). Human erythrocyte sugar transport. J. Biol. Chem. 249, 3543-3550. Blommaart, E. F. C., and Meijer, A. J. (1995). Regulation of autophagy. In "Biochemistry of Cell Membranes" (S. Papa and J. M. Tager, eds.), pp. 119-135. Burkh~iuser-Verlag, Basel. Blommaart, E. F. C., Luiken, J. J., and Meijer, A. J. (1997). Autophagic proteolysis: control and specificity. Histochem. J. 29, 365-385. Blondeau, J.-P., Osty, J., and Francon, J. (1988). Characteristics of the thyroid hormone transport system of isolated hepatocytes. J. Biol. Chem. 263, 2685-2692. Bode, B. P., and Kilberg, M. S. (1991). Amino acid-dependent increase in hepatic system N activity is linked to cellswelling. J. BioL Chem. 266, 7376-7381. Bode, B. P., Kaminski, D. L., Souba, W. W., and Li, A. P. (1995). Glutamine transport in isolated human hepatocytes and transformed liver cells. Hepatology 21, 511-520.

Boerner, P., and Saier, M. H. (1982). Growth regulation and amino acid transport in epithelial cells: influence of culture conditions and transformation on A, ASC, and L transport activities. J. Cell PhysioL 113, 240-246 Boesze-Battaglia, K., and Schimmel, R. J. (1997). Cell membrane lipid composition and distribution: Implications for cell function and lessons learned from photoreceptors and platelets. J. Exp. Biol. 200, 2927-2936. Bogdanov, M., Sun, J., Kaback, H. R., and Dowhan, W. (1996). A phospholipid acts as a chaperone in assembly of a membrane transport protein. J. Biol. Chem. 271, 11615-11618. Boim, M. A., Ho, K., Shuck, M. E., Bienkowski, M. J., Block, J. H., Slightom, J. L., Yang, Y., Brenner, B. M., and Herbert, S. C. (1995). ROMK inwardly rectifying ATPsensitive K + channel. II. Cloning and distribution of alternative forms. Am. J. Physiol. 268, Fl132-F1140. Boldyrev, A. A., and Quinn, P. J. (1994). El/E2 Type cation transport ATPases: Evidence for transient associations between promoters. Int. J. Biochem. 26, 1323-1331. Bonadonna, R. C., Saccomani, M. P., Cobelli, C., and DeFronzo, R. A. (1993). Effect of insulin on system A amino acid transport in human skeletal muscle. J. Clin. Invest. 91, 514-521. Bond, C. T., Pessia, M., Xia, X. M., Lagrutta, A., Kavanaugh, M. P., and Adelman, J. P. (1994). Cloning and expression of a family of inward rectifier potassium channels. Recept. Channels 2, 183-191. Bonina, F., Lanza, M., Montenegro, L., Salerno, L., Smeriglio, P., Trombetta, D., and Saija, A. (1996). Transport of alphatocopherol and its derivatives through erythrocyte membranes. Pharmaceut. Res. 13, 1343-1347. Boon, L., Blommaart, P. J., Meijer, A. J., Lamers, W. H., and Schoolwerth, A. C. (1996). Response of hepatic amino acid consumption to chronic metabolic acidosis. Am. J. Physiol. 271, F198-F202. Boorer, K. J., and Fischer, W. N. (1997). Specificity and stoichiometry of the Arabidopsis H+/amino acid transporter AAP5. J. BioL Chem. 272, 13040-13046. Boorer, K. J., Loo, D. D. F., and Wright, E. M. (1994). Steadystate and presteady-state kinetics of the H+/hexose cotransporter (STP1) from Arabidopsis thaliana expressed in Xenopus oocytes. J. Biol. Chem. 269, 20417-20424. Boorer, K. J., Loo, D. D., Frommer, W. B., and Wright, E. M. (1996). Transport mechanism of the cloned potato H+/ sucrose cotransporter StSUT1. J. Biol. Chem. 271, 2513925144. Booth, I. R., Jones, M. A., McLaggan, D., Nikolaev, Y, Ness, L. S., Wood, C. M., Miller, S. T6temeyer, S., and Ferguson, G. P. (1996). Bacterial ion channels. In "Transport Processes in Eukaryotic and Prokaryotic Organisms"(W. N. Konings, H. R. Kaback, and J. K. Lolkema, eds.), Vol. 2, pp. 693-729. Elsevier, New York. Borgese, F., Sardet, C., Cappadoro, M., Pouyssegur, J., and Motais, R. (1992). Cloning and expressing a cAMPactivated Na+/H + exchanger: Evidence that the cytoplasmic domain mediates hormonal regulation. Proc. Natl. Acad. Sci. USA 89, 6768-6769. Borghetti, A. F., Piedimonte, G., Tramacere, M., Severini, A., Ghiringhelli, P., and Guidotti, G. G. (1980). Cell density

References and amino acid transport in 3T3, SV3T3, and SV3T3 revertant cells. J. Cell Physiol. 105, 39-49. Bork, P., Sander, C., and Valencia, A. (1992). An ATPase domain common to prokaryotic cell cycle proteins, sugar kinases, actin, and hsp70 heat shock proteins. Proc. NatL Acad. Sci. USA 89, 7290-7294. Bork, P., Sander, C., and Valencia, A. (1993). Convergent evolution of similar enzymatic function on different protein folds: The hexokinase, ribokinase, and galactokinase families of sugar kinases. Prot. Sci. 2, 31-40. Bormann, J., Hamill, O. P., and Sakmann, B. (1987). Mechanism of anion permeation through channels gated by glycine and y-aminobutyric acid in mouse cultured spinal neurones. J. Physiol. 385, 243-286. Boron, W. F., Waisbren, S. J., Modlin, I. M., and Geibel, J. P. (1994). Unique permeability barrier of the apical surface of parietal and chief cells in isolated perfused gastric glands. J. Exp. Biol. 196, 347-360. Boulter, J. M., Taylor, A. M., and Watts, A. (1996). Asymmetric and functional reconstitution of band 3 into pre-formed phosphatidylcholine vesicles. Biochim. Biophys. Acta 1280, 265-271. Bouvier, M., Szatkowski, M., Amato, A., and Attwell, D. (1992). The glial cell glutamate uptake carrier countertransports pH-changing anions. Nature 360, 471-474. Bowling, D. J. F. (1989). Intracellular and intercellular pH measurement with microelectrodes. In "Methods in Enzymology" (S. Fleischer and B. Fleischer, eds.), Vol. 174, pp. 331-338. Academic Press, San Diego. Bowman, E. J., O'Neill, F. J., and Bowman, B. J. (1997). Mutations of pma-1, the gene encoding the plasma membrane H+-ATPase of Neurospora crassa, suppress inhibition of growth by concanamycin A, a specific inhibitor of vacuolar ATPases. J. Biol. Chem. 272, 14776-14786. Boyd, C. A. R., and Yudilevich, D. (1987). Blood-tissue movement of amino acids in various organs: the basolateral membrane and its importance in the polarity of transepithelial transport. In "Amino Acid Transport in Animal Cells" (D. Yudilevich and C. A. R. Boyd, eds.), pp. 120-156. Manchester University Press, Manchester. Boyer, P. D. (1993). The binding change mechanism for ATP synthasemSome probabilities and possibilities. Biochim. Biophys. Acta 1140, 215-250. Brahm, J. (1983). Permeability of human red cells to a homologous series of aliphatic alcohols. J. Gen. Physiol. 81, 283-304. Brahm, J., Gasbjerg, P., and Funder, J. (1992). Kinetics of anion transport across the red human cell membrane. Progr. Cell Res. 2, 25-33. Brandl, C. J., and Deber, C.M. (1986). Hypothesis about the function of membrane-buried proline residues in transport proteins. Proc. Natl. Acad. Sci. USA 83, 917-921. Brandon, C., and Tooze, J. (1991). "Introduction to Protein Structure." Garland, New York. Brandsch, M., Ganapathy, V., and Leibach, F. H. (1995). H +peptide cotransport in Madin-Darby canine kidney cells: Expression and calmodulin-dependent regulation. Am. J. Physiol. 268, F391-F397.

349

Braun, P., Persson, B., Kaback, H. R., and von Heijne, G. (1997). Alanine insertion scanning mutagenesis of lactose permease transmembrane helices. J. Biol. Chem. 272, 29566-29571. Briggs, G. E., and Haldane, J. B. S. (1925). Note on the kinetics of enzyme action. Biochem. J. 19, 338-339. Br6er, S., Br6er, A., and Hamprecht, B. (1995). The 4F2-hc surface antigen is necessary for expression of system Llike neutral amino acid-transport activity in C6-BU-1 rat glioma cells: Evidence from expression studies in Xenopus laevis oocytes. Biochem. J. 312, 863-870. Br6er, S., Ji, G., Br6er, A., and Silver, S. (1993). Arsenic efflux governed by the arsenic resistance determinant of Staphylococcus aureus plasmid pi258. J. BacterioL 175, 3480-3485. Br6er, S., Rahman, B., Pellegri, G., Pellerin, J., Martin, J.-L., Verleysdonk S., Hamprecht, B., and Magistretti, P. J. (1997). Comparison of lactate transport in astroglial cells and monocarboxylate transporter 1 (MCT1) expressing Xenopus laevis oocytes. J. Biol. Chem. 272, 3009630102. Bronner, F. (1996). Cytoplasmic transport of calcium and other inorganic ions. Comp. Biochem. Physiol. B 115, 313-317. Brookes, N. (1993). Interaction between the glutamine cycle and the uptake of large neutral amino acids in astrocytes. J. Neurochem. 60, 1923-1928. Brosnan, J. T., Lowry, M., Vinay, P., and Halperin, M. L. (1989). Renal ammonium production- une vie canadienne. Can. J. Physiol. Pharmacol. 65, 489-495. Brown, A. M., Drews, J. A., and Hartmann, H. A. (1993). The potassium pore and its regulation. Ann. N.Y. Acad. Sci. 707, 74-80. Brown, D. A., and London, E. (1997). Structure of detergentresistant membrane domains: Does phase separation occur in biological membrane? Biochem. Biophys. Res. Commun. 240, 1-7. Brown, D., and Waneck, G. L. (1992). Glycosyl-phosphatidylinositol-anchored membrane proteins. J. Am. Soc. Nephrol. 3, 895-906. Brown, D., and Stow, J. L. (1996). Protein trafficking and polarity in kidney epithelium: From cell biology to physiology. PhysioL Rev. 76, 245-297. Brown, G. C. (1995). Rate control within the Na+/glucose cotransporter. Biophys. Chem. 54, 181-189. Brown, R. S., and Wahl, R. L. (1993). Overexpression of Glut-1 glucose transporter in human breast cancer: An immunohistochemical study. Cancer 72, 2979-2985. Brzezinski, B., Schroeder, G., Zundel, G., and Keil, T. (1992). Complex formation and proton transfer in the polarizable hydrogen bonds of two N,N'-dioxides + substituted phenol systems as a function of the pKa of the phenols: An FTIR study. J. Chem Soc.-Perkin Trans. 2, 819-823. Bruggemann, A., Pardo, L. A., Stuhmer, W., and Pongs, O. (1993). Ether-a-go-go encodes a voltage-gated channel permeable to K + and C a 2+ and modulated by cAMP. Nature 365, 445-448. Brutovetsky, N., and Klingenberg, M. (1994). The reconstituted ADP/ATP carrier can mediate H + transport by free

350

References

fatty acids, which is further stimulated by mersalyl. J. Biol. Chem. 269, 27329-27336. Brutovetsky, N., and Klingenberg, M. (1996). Mitochondrial ADP/ATP carrier can be reversibly converted into a large channel by Ca 2+. Biochemistry 35, 8483-8488. Buddington, R. K., and Diamond, J. M. (1992). Ontogenetic development of nutrient transporters in cat intestine. Am. J. Physiol. 263, G605-G616. Bullard, B., Leonard, K., Larkins, A., Butcher, G., Karlik, C., and Fyrberg, E. (1988). Troponin of asynchronous flight muscle. J. MoL Biol. 204, 621-637. Burke, J. R., Wingfield, M. S., Lewis, K. E., Roses, A. D., Lee, J. E., Hulette, C., Pericak-Vance, M. A., and Vance, J. M. (1994). The Haw River Syndrome: Dentatorubropallidoluysian atrophy (DRPLA) in an African-American family. Nat. Genet. 7, 521-524. Buttrose, M., McKellar, D., and Welbourne, T. C. (1987). Gutliver interaction in glutamine homeostasis: Portal ammonia role in uptake and metabolism. Am. J. Physiol. 252, E746E750. Btirki, E., Anjard, C., Scholder, J.-C., and Reymond, C. D. (1991). Isolation of two genes encoding putative protein kinases regulated during Dictyostelium discoideum development. Gene 102, 57-65. Busch, A. E., and Maylie, J. (1993). Mink channels: A minimal channel protein with a maximal impact. Cell Physiol. Biochem. 3, 270-276. Buse, M. G., Robinson, K. A., Marshall, B. A., and Mueckler, M. (1996). Differential effects of GLUT1 or GLUT4 overexpression on hexosamine biosynthesis by muscles of transgenic mice. J. Biol. Chem. 271, 23197-23202. Bussolati, O., Laris, P. C., Rotoli, B. M., Dall'Asta, V., and Gazzola, G. C. (1992). Transport system ASC for neutral amino acids. J. Biol. Chem. 267, 8330-8335. Bussolati, O., Uggeri, J., Belletti, S., Dall'Asta, V., and Gazzola, G. C. (1996). The stimulation of Na,K,C1 cotransport and of system A for neutral amino acid transport is a mechanism for cell volume increase during the cell cycle. F A S E B J. 10, 920-926. Buurman, E. T., Kim, K.-T., and Epstein, W. (1995). Genetic evidence for two sequentially occupied K + binding sites in the Kdp transport ATPase. J. Biol. Chem. 270, 6678-6685. Cahill, G. S. (1976). Starvation in man. Clin. Endocrinol. Metab. 5, 397-415. Cairns, J., and Foster, P. L. (1991). Adaptive reversion of a frameshift mutation in Escherichia coli. Genetics 128, 695-701. Calonge, M. J., Gasparini, P., Chillar6n, J., Chill6n, M., Galiucci, M., Rousaud, F., Zelante, L., Testar, X., Dallapiccola, B., Di Siverio, F., Barcel6, P., Estivill, X., Zorzano, A., Nunes, V., and Palacin, M. (1994). Cystinuria caused by mutations in rBAT, a gene involved in the transport of cystine. Nat. Genet. 6, 420-425. Campa, M. J., and Kilberg, M. S. (1989). Characterization of neutral and cationic amino acid transport in Xenopus oocytes. J. Cell Physiol. 141, 645-652. Capaldi, R. A., Aggeler, R., Turina, P., and Wilkens, S. (1994). Coupling between catalytic sites and the proton channel in F1 and Fo-type ATPases. Trends Biochem. Sci. 19, 284-289.

Caplan, M. J. (1997). Ion pumps in epithelial cells: Sorting, stabilization and polarity. Am. J. Physiol. 272, G1304G1313. Carafoli, E. (1992). The Ca 2+ pump of the plasma membrane. J. Biol. Chem. 267, 2115-2118. Carafoli, E., and Stauffer, T. (1994). The plasma membrane calcium pump: Functional domains, regulation of the activity, and tissue specificity of isoform expression. J. Neurobiol. 25, 312-324. Cariappa, R., and Kilberg, M. S. (1990). Hormone-induced system A amino acid transport activity in rat liver plasma membrane and Golgi vesicles: Evidence for a differential sensitivity to inactivation by N-ethylmaleimide during carrier maturation. J. Biol. Chem. 265, 1470-1475. Carraro, F., Naldini, A., Weber, J. M., and Wolfe, R. R. (1994). Alanine kinetics in humans during low-intensity exercise. Med. Sci. Sports Exer. 26, 348-353. Carter, P., and Welbourne, T. C. (1997). Glutamate transport regulation of renal glutaminase flux in vivo. Am. J. Physiol. 273, E521-527. Casado., M., Bendahan, A., Zafra, F., Danbolt, N. C., Aragon, C., Gimenez, C., and Kanner, B. I. (1993). Phosphorylation and modulation of brain glutamate transporters by protein kinase C. J. Biol. Chem. 268, 27313-27317. Casado, M., Zafra, F., Aragon, C., and Gimenez, C. (1991). Activation of high-affinity uptake of glutamate by phorbol esters in primary glial cell cultures. J. Neurochem. 57,11851190. Casanova, J. E., Breitfeld, P. P., Ross, S. A., and Mostov, K. E. (1990). Phosphorylation of the polymeric immunoglobulin receptor required for its efficient transcytosis. Science 248, 742-745. Casetellino, P., Luzi, L., Simonson, D. C., Haymond, M., and De Fronzo, R. A. (1987). Effect of insulin and plasma amino acid concentrations on leucine metabolism in man. J. Clin. Invest. 80, 1784-1793. Castillo, L., Ajami, A., Branch, S., Chapman, T. E., Yu, Y. M., Burke, J. F., and Young, V. R. (1994). Plasma arginine kinetics in adult man: Response to an arginine-free diet. Metabolism 43, 114-122. Catterall, W. A. (1995). Structure and function of voltagegated ion channels. Annu. Rev. Biochem. 64, 493-531. Cavieres, J. D. (1984). Calmodulin and the target size of the (Ca 2+ + Mg2+)-ATPase of human red-cell ghosts. Biochim. Biophys. Acta 771, 241-244. Celeste, A. J., Iannazzi, J. A., Taylor, R. C., Hewick, R. M., Rosen, V., Wang, E. A. and Wozney, J. M. (1990). Identification of transforming growth factor/3 family members present in bone-inductive protein purified from bovine bone. Proc. Natl. Acad. Sci. USA 87, 9843-9847. Cerneus, D. P., Strous, G. J., and van der Ende, A. (1993). Bidirectional transcytosis determines the steady state distribution of the transferrin receptor at opposite plasma membrane domains of BeWo cells. J. Cell Biol. 122, 1223-1230. Cha, A., Zerangue, N., Kavanaugh, M., and Bezanilla, F. (1998). Fluorescence techniques for studying cloned channels and transporters expressed in Xenopus oocytes. In "Methods in Enzymology" (S. G. Amara, ed.), Vol. 296, pp. 566-578. Academic Press, San Diego.

References

Cha, J.-H. J., and Dure, L. S. (1994). Trinucleotide repeats in neurologic diseases: An hypothesis concerning the pathogenesis of Huntington's disease, Kennedy's disease, and spinocerebellar ataxia type I. Life Sci. 54, 1459-1464. Chandy, K. G., and Gutman, G. A. (1995). Voltage-gated potassium channel genes. In "Handbook of Receptors and Channels: Ligand- and Voltage-Gated Ion Channels" (R. Alan North, ed.), pp. 1-71. CRC Press, Boca Raton, FL. Chang, X. B., Tabcharani, J. A., Hou, Y. X., Jensen, T. J., Kartner, N., Alon, N., Hanrahan, J. W., and Riordan, J. R. (1993). Protein kinase A (PKA) still activates CFTR chloride channel after mutagenesis of all 10 PKA consensus phosphorylation sites. J. Biol. Chem. 268, 11304-11311. Chaplin, D. D., Galbraith, L. J., Seidman, J. G., White, P. C., and Parker, K. L. (1986). Nucleotide sequence analysis of murine 21-hydroxylase genes: Mutations affecting gene expression. Proc. Natl. Acad. Sci. USA 83, 9601-9605. Chapman, D. (1993). Biocompatible surfaces based upon the phospholipid asymmetry of biomembranes. Biochem. Soc. Trans. 21, 258-262. Charron, M. J., Brosius, F. C., III, Alper, S. L., and Lodish, H. F. (1989). A glucose transport protein expressed predominantly in insulin-responsive tissues. Proc. Natl. Acad. Sci. USA 86, 2535-2539. Charron, M. J., Katz, E. B., and Zierath, J. R. (1997). Metabolic and molecular consequences of modifying GLUT4 expression in skeletal muscle. Biochem. Soc. Trans. 25, 963-968. Cheeseman, C. (1992). Role of intestinal basolateral membrane in absorption of nutrients. Am. J. of Physiol. 263, R482-R488. Che, A., Morrison, I. E. G., Pan, R.-J., and Cherry, R. J. (1997). Restriction by ankyrin of band 3 rotational mobility in human erythrocyte membranes and reconstituted lipid vesicles. Biochemistry 36, 9588-9595. Chen, C.-N., Denome, S., and Davis, R. L. (1986). Molecular analysis of cDNA clones and the corresponding genomic coding sequences of the Drosophila dunce + gene, the structural gene for cAMP phosphodiesterase. Proc. Natl. Acad. Sci. USA 83, 9313-9317. Chen, D. P., and Eisenberg, R. S. (1993). Flux, coupling, and selectivity in ionic channels of one conformation. Biophys. J. 65, 727-746. Chen, L., Alam, T., Johnson, J. H., Hughes, S., Newgard, C. B., and Unger, R. H. (1990). Regulation of b-cell glucose transporter gene expression. Proc. Natl. Acad. Sci. USA 87, 4088-4092 Chen, X.-Z., Coady, M. J., Jackson, F., Berteloot, A., and Lapointe, J.-Y. (1995). Thermodynamic determination of the Na+:Glucose coupling ratio for the human SGLT1 cotransporter. Biophys. J. 69, 2405-2414. Chen, X. Z., Coady, M. J., and Lapointe, J. Y. (1996). Fast voltage clamp discloses a new component of presteadystate currents from the Na+-glucose cotransporter. Biophys. J. 71, 2544-2552. Chen, X.-Z., Coady, M. J., Jalal, F., Wallendorff, B., and Lapointe J.-Y. (1997). Sodium leak pathway and substrate binding order in the Na+-glucose cotransporter. Biophys. J. 73, 2503-2510.

351

Cheng, S. H., Rich, D. P., Marshall, J., Gregory, R. J., Welsh, M. J., and Smith, A. E. (1991). Phosphorylation of the R domain by cAMP-dependent protein kinase regulates the CFTR chloride channel. Cell 66, 1027-1036. Chiles, T. C., and Kilberg, M. S. (1986). System A transport activity in normal rat hepatocytes and transformed liver cells: Substrate protection from inactivation by sulfhydrylmodifying reagents. J. Cell. Physiol. 129, 321-328. Chillar6n, J., Est6vez, R., Mora, C., Wagner, C. A., Suessbrich, H., Lang, F., Gelpi, J. L., Testar, X., Busch, A. E., Zorzano, A., and Palacin, M. (1996). Obligatory amino acid exchange via systems b~ and y+L-like. J. Biol. Chem. 271, 17761-17770. Chillar6n, J., Estevez, R., Mora, C., Wagner, C. A., Suessbrich, H., Lang, F., Gelpi, J. L., Testar, X., Busch, A. E., Zorzano, A., and Palacin, M. (1996). Amino acid exchange via systems b~ and y+L-like: A tertiary active transport mechanism for renal reabsorption of cystine and dibasic amino acids. J. Biol. Chem. 271, 17761-17770. Chiu, S.-W., Clark, M., Balaji, V., Subramaniam, S., Scott, H. L., and Jakobsson, E. (1995). Incorporation of surface tension into molecular dynamics simulation of an interface: A fluid phase lipid bilayer membrane. Biophys. J. 69,12301245. Chiu, S.-W., Gulukota, K., and Jakobsson, E. (1992). Computational approaches to understanding the ion channel-lipid system. In "Membrane Proteins: Structures, Interactions, and Models" (A. Pullman, B. Pullman, and J. Jortner, eds.). Kluwer, Amsterdam. Chiu, S.-W., Jakobsson, E., Subramaniam, S., and McCammon, J. A. (1991). Time-correlation analysis of simulated water motion in flexible and rigid gramicidin channels. Biophys. J. 60, 273-285. Chiu, S.-W., Novotny, J. A., and Jakobsson, E. (1993). The nature of ion and water barrier crossing in a simulated ion channel. Biophys. J. 64, 98-108. Chiu, S.-W., Subramaniam, S., Jakobsson, E., and McCammon, J. A. (1989). Water and polypeptide conformations in gramicidin channels--A molecular dynamics study. Biophys. J. 56, 253-261. Cho, Y., and Bannai, S. (1990). Uptake of glutamate and cysteine in C-6-glioma cells and in cultured astrocytes. J. Neurochem. 55, 2091-2097. Choi, K. L., Aldrich, R. W., and Yellen, G. (1991). Tetraethylammonium blockade distinguishes two inactivation mechanisms in voltage-gated K + channels. Proc. NatL Acad. Sci. USA 88, 5092-5095. Choi, W. H., O'Rahilly, S., Buse, J. B., Rees, A., Morgan, R., Flier, J. S., and Moller, D. E. (1991). Molecular scanning of insulin-responsive glucose transporter (GLUT4). gene in NIDDM subjects. Diabetes 40, 1712-1718. Chow, D. C., and Forte, J. G. (1995). Functional significance of the fl-subunit for heterodimeric P-type ATPases. J. Exp. Biol 198, 1-17. Christensen, H. N. (1962). "Biological Transport," 1st ed. W. A. Benjamin, Reading, MA. Christensen, H. N. (1975). "Biological Transport," 2nd ed. W. A. Benjamin, Reading, MA.

352

References

Christensen, H. N. (1982). Interorgan amino acid nutrition. Physiol. Rev. 62, 1193-1233. Christensen, H. N. (1984). Organic ion transport during seven decades: The amino acids. Biochim. Biophys. Acta 779, 255-269. Christensen, H. N. (1987a). Hypothesis: Where the depleted amino acids go in phenylketonuria and why. Perspect. Biol. Med. 30, 186-196. Christensen, H. N. (1987b). Role of membrane transport in interorgan amino acid flows: Where do the depleted amino acids go in phenylketonuria? In "Amino Acids in Health and Disease: New Perspectives" (S. Kaufman, ed.), pp. 1-17. A. R. Liss, New York. Christensen, H. N. (1990). Role of amino acid transport and countertransport in nutrition and metabolism. Physiol. Rev. 70, 43-77. Christensen, H. N. (1992). A retrovirus uses a cationic amino acid transporter as a cell surface receptor. Nutr. Rev. 50, 47-48. Christensen, H. N., and Antonioli, J. A. (1969). Cationic amino acid transport in the rabbit reticulocyte: Na+-dependent inhibition of Na+-independent transport. J. Biol. Chem. 244, 1497-1504. Christensen, H. N., and Handlogten, M. E. (1968). Modes of mediated exodus of amino acids from the Ehrlich ascites tumor cell. J. Biol. Chem. 243, 5428-5438. Christensen, H. N., and Handlogten, M. E. (1977). Does the non-saturable cell entry apply to the charge-free form of the amino acids? Biochim. Biophys. Acta 469, 216-220. Christensen, H. N., and Kilberg, M. S. (1995). Hepatic amino acid transport primary to the urea cycle in regulation of biologic neutrality. Nutr. Rev. 53, 74-76. Christensen, H. N., and Liang, M. (1966a). Transport of diamino acids into the Ehrlich cell. J. Biol. Chem. 241, 55425551. Christensen, H. N., and Liang, M. (1966b). Modes of uptake of benzylamine by the Ehrlich cell. J. Biol. Chem. 241, 5552-5556. Christensen, H. N., and Makowske, M. (1983). Recognition of anionic amino acids for hepatocyte transport and for neurotransmittory action compared. Life Sci 33, 22552267. Christensen, H. N., and Van Winkle, L. J. (1995). Ontogeny of transport systems for cationic amino acids in the placenta. Nutr. Rev. 53, 40-43. Christensen, H. N., Albritton, L. M., Kakuda, D. K., and MacLeod, C. L. (1994). Gene product designations for amino acid transporters. J. Exp. Biol. 196, 51-57. Christensen, H. N., Handlogten, M. E., and Thomas, E. L. (1969). Na+-facilitated reactions of neutral amino acids with a cationic amino acid transport system. Proc. Natl. Acad. Sci. USA 63, 948-955. Christensen, H. N., Liang, M., and Archer, E. G. (1967). A distinct Na+-requiring transport system for alanine, serine, cysteine, and similar amino acids. J. Biol. Chem. 242, 52375246. Christie, M. J. (1995). Molecular and functional diversity of K + channels. Clin. Exp. PharmacoL Physiol. 22, 944-951.

Chuang, D. T., and Shih, V. E. (1995). Disorders of branched chain amino acid and keto acid metabolism. In "Metabolic and Molecular Bases of Inherited Disease" (C. R. Scriver, A. L. Beaudet, W. S. Sly, and D., Valle, eds.), 7th ed., pp. 1239-1277. McGraw-Hill, New York. Chung, M., Teng, C., Timmerman, M., Meschia, G., and Battaglia, F. C. (1998). Production and utilization of amino acids by ovine placenta in vivo. Am. J. Physiol. 274, E13-E22. Clavo, A. C., Brown, R. S., and Wahl, R. L. (1995). Fluorodeoxyglucose uptake in human cancer cell lines is increased by hypoxia. J. Nucl. Med. 36, 1625-1632. Cloherty, E. K., Diamond, D. L., Heard, K. S., and Carruthers, A. (1996). Regulation of GLUTl-mediated sugar transport by an antiport/uniport switch mechanism. Biochemistry 35, 13231-13239. Closs, E. I., Albritton, L. M., Kim, J. W., and Cunningham, J. M. (1993). Identification of a low affinity, high capacity transporter of cationic amino acids in mouse liver. J. Biol. Chem. 268, 7538-7544. Closs, E. I., Graf, P., Habermeier, A., Cunningham, J. M., and Forstermann, U. (1997). Human cationic amino acid transporters hCAT-1, hCAT-2A, and hCAT-2B: Three related carriers with distinct transport properties. Biochemistry 26, 6462-6468. Coady, M. J., Chen, X.-Z., and Lapointe, J.-Y. (1996). rBAT is an amino acid exchanger with variable stoichiometry. J. Membr. Biol. 149, 1-8. Coady, M. J., Jalal, F., Chen, X., Lemay, G., Berteloot, A., and Lapointe, J.-Y. (1994). Electrogenic amino acid exchange via the rBAT transporter. FEBS Lett. 356,174-178. Colman, A. (1984a). Expression of exogenous DNA in Xenopus oocytes. In "Translation and Transcription" (B. D. Hames and S. J. Higgins, eds.), pp. 49-69. IRL Press, Oxford. Colman, A. (1984b). Translation of eukaryotic messenger RNA in Xenopus oocytes. In "Transcription and Translation" (B. D. Hames and S. J. Higgins, eds.), pp. 271-302. IRL Press, Oxford. Collander, R., and B~irlund, H. (1933). Permeabilit~itsstudieren an chara ceratophylla. Acta Botan. Fenn. 11, 1-14. Comfurius, P., Williamson, P., Smeets, E. F., Schlegel, R. A., Bevers, E. M., and Zwaal, R. F. A. (1996). Reconstitution of phospholipid scramblase activity from human blood platelets. Biochemistry 35, 7631-7634. Conejero, C. (1997). Anion exchanger AE1 as a candidate pathway for taurine transport in rat erythrocytes. Am. J. Physiol. 272, C1457-C1464. Conradt, M., and Stoffel, W. (1995). Functional analysis of the high affinity, Na+-dependent glutamate transporter GLAST-1 by site-directed mutagenesis. J. Biol. Chem. 270, 25207-25212. Conradt, M., and Stoffel, W. (1997). Inhibition of high-affinity brain glutamate transporter GLAST-1 via direct phosphorylation. J. Neurochem. 68, 1244-1251. Conradt, M., Storck, T., and Stoffel, W. (1995). Localization of N-glycosylation sites and functional role of the carbohydrate units of GLAST-1, a cloned rat brain L-glutamate/ L-aspartate transporter. Eur. J. Biochem. 229, 682-687.

References

Cooney, J., Kienle, Z., Foster, T. J., and O'Toole, P. W. (1993). The gamma-hemolysin locus of Staphylococcus aureus comprises three linked genes, two of which are identical to the genes for the F and S components of leukocidin. Infect. Immun. 61, 768-771. Corey, J. L., Davidson, N., Lester, H. A., Brecha, N., and Quick, M. W. (1994). Protein kinase C modulates the activity of a cloned y-aminobutyric acid transporter expressed in Xenopus oocytes via regulated subcellular redistribution of the transporter. J. Biol. Chem. 269, 14759-14767. Cornish-Bowden, A., and Cardenas, M. L. (1990). "Control of Metabolic Processes". Plenum, New York. Cornish-Bowden, A., and Eisenthal, R. (1978). Estimation of Michaelis constant and maximum velocity from the direct linear plot. Biochim. Biophys. Acta 523, 268-272. Coronado, R., Rosenberg, R. L., and Miller, C. (1980). Ionic selectivity, saturation, and block in a K+-selective channel from sarcoplasmic reticulum. J. Gen. PhysioL 76, 425-446. Costa, A. C. S., Patrick, J. A., and Dani, J. A. (1994). Improved technique for studying ion channels expressed in Xenopus oocytes, including fast superfusion. Biophys. J. 67, 395-401. Coupaye-Gerard, B., Zuckerman, J. B., Duncan, P., Bortnik, A., Avery, D. I., Ernst, S. A., and Kleyman, T. R. (1997). Delivery of newly synthesized Na+-K+-ATPase to the plasma membrane of A6 epithelia. Am. J. Physiol. 272, C1781-C1789. Cox, K. H., and Cox, J. V. (1995). Variant chicken AE1 anion exchangers possess divergent NHz-terminal cytoplasmic domains. Am. J. Physiol. 268, F503-F513. Cox, J. V., and Lazarides, E. (1988). Alternative primary structures in the transmembrane domain of the chicken erythroid anion transporter. Mol. Cell Biol. 8, 1327-1335. Cox, K. H., Adair-Kirk, T. L., and Cox, J. V. (1995). Four variant chicken erythroid AE1 anion exchangers. J. Biol. Chem 270, 19752-19760. Cox, K. H., Adair-Kirk, T. L., and Cox, J. V. (1996). Variant AE2 anion exchanger transcripts accumulate in multiple cell types in the chicken gastric epithelium. J. Biol. Chem. 271, 8895-8902. Craik, J. D., and Elliott, K. R. F. (1979). Kinetics of 3 - 0 methyl-D-glucose transport in isolated rat hepatocytes. Biochem. J. 182, 503-508. Cupello, A., Baldelli, P., Gheri, A., and Robello, M. (1993). A method for checking the volume of solutions microinjected into Xenopus oocytes. Anal. Biochem. 213, 172-174. Curthoys, N. P., and Watford, M. (1995). Regulation of glutaminase activity and glutamine metabolism. Ann. Rev. Nutr. 15, 133-159. Dai, G., Levy, O., and Carrasco N. (1996). Cloning and characterization of the thyroid iodide transporter. Nature 379, 458-460. Dall'Asta, V., Franchi-Gazzola, R., Bussolati, O., Sala, R., Rotoli, B. M., Rossi, P. A., Uggeri, J., Belletti, S., Visigalli, R., and Gazzola, G. C. (1996). Modulation of transport systems for neutral and anionic amino acids in mesenchymal cells. Biochem. Soc. Trans. 24, 864-869. Dall'Asta, V., Rossi, P. A., Bussolati, O., and Gazzola, G. C. (1994). Response of human fibroblasts to hypertonic stress:

353

Cell shrinkage is counteracted by an enhanced active transport of neutral amino acids. J. Biol. Chem. 269, 1048510491. Danbolt, N. C., Pines, G,. and Kanner, B. I. (1990). Purification and reconstitution of the sodium- and potassium-coupled glutamate transport glycoprotein from rat brain. Biochemistry 29, 6734- 6740. Dangott, L. J., Jordan, J. E., Bellet, R. A., and Garbers, D. L. (1989). Cloning of the mRNA for the protein that crosslinks to the egg peptide speract. Proc. Natl. Acad. Sci. USA 86, 2128-2132. Daniel, H., Morse, E. L., and Adibi, S. A. (1991). The high and low affinity transport systems for dipeptides in kidney brush border membrane respond differently to alterations in pH gradient and membrane potential. J. Biol. Chem. 266, 19917-19924. Dascal, N., Schreibmayer, W., Lim, N. F., Wang, W., Chavkin, C., DiMagno, L., Labarca, C., Kieffer, B. L., GaveriauxRuff, C., and Trollinger, D. (1993). Atrial G proteinactivated K + channel: expression cloning and molecular properties. Proc. Natl. Acad. Sci. USA 90, 10235-10239. Davies, J. M., Hunt, I., and Sanders, D. (1994). Vacuolar H +pumping ATPase variable transport coupling ratio controlled by pH. Proc. Natl. Acad. Sci. USA 91, 8547-8551. Dayhoff, M. O., Barker, W. C., and Hunt, L. T. (1983). Establishing homologies in protein sequences. Methods Enzymol. 91, 524-545. Dayhoff, M. O., Hunt, L. T., and Calderone, S. H. (1978). Composition of proteins. In "Atlas of Protein Sequence and Structure", Vol. 5, Suppl. 3, pp.363-373. National Biomedical Research Foundation, Washington, DC. Dean, A. M. (1998). The molecular anatomy of an ancient adaptive event. Am. Sci. 86, 26-37. de Cespedes, C., Thoene, J. G., Lowler, K., and Christensen, H. N. (1989). Evidence for inhibition of exodus of small neutral amino acids from non-brain tissues in hyperphenylalaninaemic rats. J. Inher. Metab. Dis. 12, 166-180. Dechecchi, M. C., Rolfini, R., Tamanini, A., Gamberi, C., Berton, G., and Cabrini, G. (1992). Effect of modulation of protein kinase C on the cAMP-dependent chloride conductance in T84 cells. FEBS Lett. 311, 25-28. Dechecchi, M. C., Tamanini, A., Berton, G., and Cabrini, G. (1993). Protein kinase C activates chloride conductance in C127 cells stably expressing the cystic fibrosis gene. J. Biol. Chem. 268, 11321-11325. Deckers-Hebestreit, G., and Altendorf, K. (1996). The FoF1type ATP synthases of bacteria: Structure and function of the F0 complex. Ann. Rev. MicrobioL 50, 791-824. De Feo, P., Volpi, E., Lucidi, P., Cruciani, G., Santeusanio, F., Bolli, G. B., and Brunetti, P. (1994). Chloroquine reduces whole body proteolysis in humans. Am. J. Physiol. 267, E183-186. De Jong, M., Docter, R., van der Hoek, H., Krenning, E. P., and Hennemann, G. (1994). Adaptive changes in transmembrane transport and metabolism of triiodothyronine in perfused livers of fed and fasted hypothyroid and hyperthyroid rats. Metabolism 43, 1355-1361. Demuth, D. R., Showe, L. C., Ballantine, M., Palumbo, A., Fraser, P. J., Cioe, L., Rovera, G., and Curtis, P. J. (1986).

354

References

Cloning and structural characterization of a human nonerythroid band 3-like protein. EMBO J. 5, 1205-1214. Deems, R. O., Evans, J. L., Deacon, R. W., Honer, C. M., Chu, D. T., Burki, K., Fillers, W. S., Cohen, D. K., and Young, D. A. (1994). Expression of human GLUT4 in mice results in increased insulin action. Diabetologia 37, 10971104. Devereux, J., Haeberli, P., and Smithies, N. O. (1984). A comprehensive set of sequence analyses for the VAX. Nucleic Acids Res. 12, 387-395. Dev6s, R., and Angelo, S. (1996). Changes in membrane and surface potential explain the opposite effects of low ionic strength on the two lysine transporters of human erythrocytes. J. Biol. Chem. 271, 32034-32039. Dev6s, R., and Boyd, C. A. R. (1998). Transporters for cationic amino acids in animal cells: Discovery, structure and function. Physiol. Rev. 78, 487-546. Dev6s, R., Rojas, A. M., and Angelo, S. (1997). Binding specificity and cation dependence of the two transporters for cationic amino acids in human erythrocytes: systems y+ and y+L. Amino Acids 13, 48-49. Diamond, J. M. (1993). Evolutionary Physiology. In "The Logic of Life" (C. A. R. Boyd and D. Noble, eds.), pp. 89-111. Oxford University Press, UK. Dieckmann, G. R., and DeGrado, W. F. (1997). Modeling transmembrane helical oligomers. Curr. Opin. Struct. Biol. 7, 486-494. Dierks, T., and Kr~imer, R. (1988). Asymmetric orientation of the reconstituted aspartate/glutamate carrier from mitochondria. Biochim. Biophys. Acta 937, 112-126. Dierks, T., Salentin, A., Heberger, C., and Kr~imer, R. (1990a). The mitochondrial aspartate/glutamate and A D P / A T P carrier switch from obligate counterexchange to unidirectional transport after modification by SH-reagents. Biochim. Biophys. Acta 1028, 268-280. Dierks, T., Salentin, A., Herberger, C., and Kr~imer, R. (1990b). Pore-like and carrier-like properties of the mitochondrial aspartate/glutamate carrier after modification by SH-reagents: Evidence for a preformed channel as a structural requirement of carrier-mediated transport. Biochim. Biophys. Acta 1028, 281-288. Ding, Y., Casey, J. R., and Kopito, R. R. (1994). The Major kidney AE1 isoform does not bind ankyrin (ANK1) in vitro. J. Biol. Chem. 269, 32201-32208. Ding, Y., Kobayashi, S., and Kopito, R. (1996). Mapping of ankyrin binding determinants on the erythroid anion exchanger, AE1. J. Biol. Chem. 271, 22494-22498. Dixon, M., and Webb, E. C. (1964). "Enzymes", 2nd ed. Academic Press, New York. Dixon, M., and Webb, E. C. (1979). "Enzymes", 3rd ed. Academic Press, New York. Docherty, K., and Clark, A. R. (1994). Nutrient regulation of insulin gene expression. FASEB J. 8, 20-27. Dolly, J. O., and Parcej, D. N. (1996). Molecular properties of voltage-gates K + channels. J. Bioenerg. Biomembr. 28, 231-253. Dolly, J. O., Rettig, J., Scott, V. E., Parcej, D. N., Wittkat, R., Sewing, S., and Pongs, O. (1994). Structure and regulation of cation channels. Biochem. Soc. Trans. 22, 473-478.

Doolittle, R. F. (1986). "Of Urfs and Orfs: A Primer on How to Analyze Derived Amino Acid Sequences." University Science Books, Mill Valley, CA. Doolittle, R. F. (1992). Reconstructing history with amino acid sequences. Prot. Sci. 1, 191-200. Doolittle, R. F. (1994). Convergent evolution: The need to be explicit. Trends Biochem. Sci. 19, 15-18. Dorer, D. R., Christensen, A. C., and Johnson, D. H. (1990). A novel RNA helicase gene tightly linked to the Triplolethal locus of Drosophila. Nucleic Acid Res. 18, 5489-5494. D6ring, F., Dorn, D., Bachfischer, U., Amasheh, S., Herget, M., and Daniel, H. (1996). Functional analysis of a chimeric mammalian peptide transporter derived from the intestinal and renal isoforms. J. Physiol. 497, 773-779. DOring, F., Theis, S., and Daniel, H. (1997). Expression and functional characterization of the mammalian intestinal peptide transporter PepT1 in the methylotropic yeast Pichia pastoris. Biochem. Biophys. Res. Commun. 232, 656-662. Dowd, L. A., and Robinson, M. B. (1996). Rapid stimulation of EAACl-mediated Na+-dependent L-glutamate transport activity in C6 glioma cells by phorbol ester. J. Neurochem. 67, 508-516. Dowd, L. A., Coyle, A. J., Rothstein, J. D., Pritchett, D. B., and Robinson, M. B. (1996). Comparison of Na+-dependent glutamate transport activity in synaptosomes, C6 glioma, and Xenopus oocytes expressing excitatory amino acid carrier 1 (EAAC1). Mol. Pharmacol. 49, 465-473. Dowhan, W. (1991). Phospholipid-transfer proteins. Curr. Opin. Cell Biol. 3, 621-625. Doyle, D. A., Cabral, J. M., Pfuetzner, R. A., Kuo, A., Gulbis, J. M., Cohen, S. L., Chait, B. T., and MacKinnon, R. (1998). The structure of the potassium channel: Molecular basis of K + conduction and selectivity. Science 280, 69-77. Dr6ge, W., and Holm, E. (1997). Role of cysteine and glutathione in HIV infection and other diseases associated with muscle wasting and immunological dysfunction. FASEB J. 11, 1077-1090. Drumm, M. L., Wilkinson, D. J., Smit, L. S., Worrell, L. T., Strong, T. V., Frizzell, R. A., Dawson, D. C., and Collins, F. S. (1991). Chloride conductance expressed by AF508 and other mutant CFTRs in Xenopus oocytes. Science 254, 1797-1799. Duan, D., Winter, C., Cowley, S., Hume, J. R., and Horowitz, B. (1997). Molecular identification of a volume-regulated chloride channel. Nature 390, 417-421. Ducarme, P., Rahman, M., Lins, L., and Brasseur, R. (1996). The erythrocyte/brain glucose transporter (GLUT1). may adopt a two-channel transmembrane a/fl structure. J. Mol. Model 2, 27-45. Dudeck, K. L., Dudenhausen, E. E., Chiles, T. C., Fafournoux, P., and Kilberg, M. S. (1987). Evidence for inherent differences in the system A carrier from normal and transformed liver tissue: Differential inactivation and substrate protection in membrane vesicles and reconstituted proteoliposomes. J. Biol. Chem. 262, 12565-12569. Dulhanty, A. M., and Riordan, J. R. (1994). Phosphorylation by cAMP-dependent protein kinase causes a conforma-

References tional change in the R domain of the cystic fibrosis transmembrane conductance regulator. Biochemistry 33, 40724079. Duprat, F., Lessage, F., Guillermare, E., Fink, M., Hugnot, J. P. et al. (1995). Heterologous multimeric assembly is essential for K + channel activity of neuronal and cardiac G-protein-activated inward rectifiers. Biochem. Biophys. Res. Commun. 212, 657-663. Durell, S. R., Hao, Y., and Guy, H. R. (1998). Structural models of the transmembrane region of voltage-gated and other K + channels in open, closed, and inactivated conformations. J. Struct. Biol. 121, 263-284. Edidin, M. (1996). Getting there is only half the fun. Curr. Topics Membr 43, 1-13. Edmondson, J. W., and Lumeng, L. (1980). Biphasic stimulation of amino acid uptake by glucagon in hepatocytes. Biophys. Biochem. Res. Commun. 96, 61-68. Edmondson, J. W., Miller, B. A., and Kumeng, L. (1985). Effect of glucagon on hepatic taurocholate uptake: relationship to membrane potential. Am. J. Physiol. 249, G427G433. Efrat S., Tal, M., and Lodish, H. F. (1994). The pancreatic/3cell glucose sensor. Trends Biochem. Sci. 19, 535-538 Ehrenfeld, E. (1998). Termination of the Physiology Study Section: The wrong way to make the right decision. The Physiologist 41, 1 and 15. Eisenthal, R., and Cornish-Bowden, A. (1974). The direct linear plot: A new graphical procedure for estimating enzyme kinetic parameters. Biochem. J. 139, 715-720. Eleno, N., Dev6s, R., and Boyd, C. A. (1994). Membrane potential dependence of the kinetics of cationic amino acid transport systems in human placenta. J. Physiol. 479, 291-300. E1-Hage Chahine, J.-M., Cribier, S., and Devaux, P. F. (1993). Phospholipid transmembrane domains and lateral diffusion in fibroblasts. Proc. Natl. Acad. Sci. USA 90, 447-451. E1-Mashak, E. M., and Tsong, T. Y. (1985). Ion selectivity of temperature-induced and electric field induced pores in dipalmitoylphosphatidylcholine vesicles. Biochemistry 24, 2884-2888. Elmoselhi, A. B., Samson, S. E., and Grover, A.K. (1996). SR Ca 2+ pump heterogeneity in coronary artery: Free radicals and IP3-sensitive and -insensitive pools. Am. J. Physiol. 271, C1652-1659. Endo, H., Matsuda, C., and Kagawa, Y. (1994). Exclusion of an alternatively spliced exon in human ATP synthase ysubunit pre-mRNA requires de novo protein synthesis. J. Biol. Chem. 269, 12488-12493. Engelke, T., Jording, D., Kapp, D., and Puhler, A. (1989). Identification and sequence analysis of the Rhizobium meliloti dctA gene encoding the C4-dicarboxylate carrier. J. Bacteriol. 171, 5551-5560. England, S. K., Uebele, V. N. Kodali, J., Bennett, P. B., and Tamkun, M. M. (1995). A novel K+channel /3-subunit (hKv/31.3). is produced via alternative mRNA splicing. J. Biol. Chem. 270, 28531-28534. Epand, E. M. (1998). "Lipid Polymorphism and Membrane Properties: Current Topics in Membranes", Vol. 44. Academic Press, San Diego.

355

Epplen, J. T. (1988). On simple repeated GATCA sequences in animal genomes: A critical reappraisal. J. Heredity 79, 409-417. Eriksson, J., Koranyi, L., Bourey, R., Schalin, J. C., Widen, E., Mueckler, M., Permutt, A. M., and Groop, L. C. (1992). Insulin resistance in type 2 (non-insulin-dependent) diabetic patients and their relatives is not associated with a defect in the expression of the insulin-responsive glucose transporter (GLUT-4). gene in human skeletal muscle. Diabetologia 35, 143-147. Eskandari, S., Loo, D. D. F., Dai, G., Levy, O., Wright, E. M., and Carrasco, N. (1997). Thyroid Na+/1 symporter: Mechanism, stoichiometry, and specificity. J. Biol. Chem. 272, 27230-27238. Espanol, M. J., and Saier, M. H. (1995). Topological and segmental phylogenetic analyses of the anion exchanger (band 3). family of transporters. Mol. Membr. Biol. 12, 193-200. Fafournoux, P., R6m6sy, C., and Demign6, C. (1983). Control of alanine metabolism in rat liver by transport processes or cellular metabolism. Biochem. J. 210, 645-652. Fairman, W. A., Vandenberg, R. J., Arriza, J. L., Kavanaugh, M. P., and Amara, S. G. (1995). An excitatory amino acid transporter with properties of a ligand-gated chloride channel. Nature 375, 599-603. Fajans, S. S., Christensen, H. N., Floyd, J. C., and Pek, S. (1974). Stimulation of insulin and glucagon release in the dog by a nonmetabolizable arginine analog. Endocrinology 94, 230-233. Fajans, S. S., Quibrera, R., Pek, S., Floyd, J. C., Christensen, H. N., and Conn, J. W. (1971). Stimulation of insulin release by a nonmetabolizable amino acid: Comparison with leucine and arginine. J. Clin. Endocrinol. Metab. 33, 35-41. Fakler, B., Br~indle, U., Glowatzki, E., Weidemann, S., Zenner, H. P. and Ruppersberg, J. P. (1995). Strong voltagedependent inward rectification of inward rectifier K + channels caused by intracellular spermine. Cell 80, 149-154. Fakler, B., Br~indle, U., Glowatzki, E., Zenner, H. P., and Ruppersberg, J.P. (1994). Kir2.1 inward rectifier K + channels are regulated independently by protein kinases and ATP hydrolysis. Neuron 13, 1413-1420. Falson, P., Menguy, T., Corre, F., Bouneau, L., Gomez de Gracia, A., Souli6, S., Centeno, F., MOiler, J. V., Champeil, P., and le Maire, M. (1997). The cytoplasmic loop between putative transmembrane segments 6 and 7 in sarcoplasmic reticulum Ca2+-ATPase binds Ca 2+ and is functionally important. J. Biol. Chem. 272, 17258-17262. Farge, E. (1995). Increased vesicle endocytosis due to an increase in the plasma membrane phosphatidylserine concentration. Biophys. J. 69, 2501-2506. Far/as, R. N., Chehin, R. N., Rintoul, M. R., and Morero, R. D. (1995). Differential effect of triiodothyronine and thyroxine on the liposomal membrane in liquid-crystalline and gel state. J. Mem. Biol. 143, 135-141. Fath, K. R., Mamajiwalla, S. N., and Burgess, D. R. (1993). The cytoskeleton in development of epithelial cell polarity. J. Cell Sci. 17, (Suppl.), 65-73. Fedosova, N. U., Cornelius, F., and Klodos, I. (1995). Fluorescent styryl dyes as probes for Na,K-ATPase reaction mech-

356

References

anism: Significance of the charge of the hydrophilic moiety of RH dyes. Biochemistry 34, 16806-16814. Fei, Y. J., Kanai, Y., Nussberger, S., Ganapathy, V., Leibach, F. H., Romero, M. F., Singh, S. K., Boron, W. G., and Hediger, M. A. (1994). Expression cloning of a mammalian proton-coupled oligopeptide transporter. Nature 368, 563-566. Fei, Y. J., Liu, W., Prasad, P. D., Kekuda, R., Oblak, T. G., Ganapathy, V., and Leibach, F. H. (1997). Identification of the histidyl residue obligatory for the catalytic activity of the human H+/peptide cotransporters PEPT1 and PEPT2. Biochemistry 36, 452-460. Felig, P. (1975). Amino acid metabolism in man. Annu. Rev. Biochem. 44, 933-953. Felig, P., Wahren, J., and R/if, L. (1973). Evidence of interorgan amino-acid transport by blood cells in humans. Proc. Natl. Acad. Sci. USA 70, 1775-1779. Felsenstein, J. (1988). Phylogenies from molecular sequences: Inference and reliability. Annu. Rev. Genet. 22, 521-565. Fendler, K., DrOse, S., Altendorf, K., and Bamberg, E. (1996). Electrogenic K + transport by the Kdp-ATPase of Escherichia coli. Biochemistry 35, 8009-8017. Feng, J., and Lingrel, J. B. (1995). Functional consequences of substitutions of the carboxyl residue glutamate 779 of the Na, K-ATPase Cell MoL BioL Res. 41, 29-37. Feng, L., Sun, W., Xia, Y., Tang, W. W., Chanmugam, P., Soyoola, E., Wilson, C. B., and Hwang, D. (1993). Cloning two isoforms of rat cyclooxygenase: Differential regulation of their expression. Arch. Biochem. Biophys. 307, 361-368. Feng, S., and MacDonald, R.C. (1996). A tethered adhesive particle model of two dimensional elasticity and its application to the erythrocyte membrane. Biophys. J. 70, 857-867. Ferrer, J., Nichols, C. G., Makhina, E. N., Salkoff, L., Bernstein, J., Gerhard, D., Wasson, J., Ramanadham, S., and Permutt, A. (1995). Pancreatic islet cells express a family of inwardly rectifying K + channel subunits which interact to form G-protein-activated channels. J. Biol. Chem. 270, 26086-26091. Ferrer-Martinez, A., Felipe, A., Nicholson, B., Casado, J., Pastor-Anglada, M., and McGivan, J. D. (1995). Induction of the high-affinity Na+-dependent glutamate transport system X-AO by hypertonic stress in the renal epithelial cell line NBL-1. Biochem. J. 310, 689-692. Ffrench-Constant, R. H., Mortlock, D. P., Shaffer, C. D., MacIntyre, R. J., and Roush, R. T. (1991). Molecular cloning and transformation of cyclodiene resistance in Drosophila: An invertebrate y-aminobutyric acid subtype A receptor locus. Proc. Natl. Acad. Sci. USA 88, 7209-7213. Fi6vet, B., Gabillat, N., Borgese, F., and Motais, R. (1995). Expression of band 3 anion exchanger induces chloride current and taurine transport: Structure-function analysis. E M B O J. 14, 5158-5169. Fi6vet, B., Perset, F., Gabillat, N., Guizouarn, H., Borgese, F., Ripoche, P., and Motais, R. (1998). Transport of uncharged organic solutes in Xenopus oocytes expressing red cell anion exchangers (AEls). Proc. Natl. Acad. Sci. USA 95, 10996-11001. Fillingame, R. H. (1997). Coupling H § transport and ATP synthesis in FIFo-ATP synthases: Glimpses of interacting

parts in a dynamic molecular machine. J. Exp. BioL 200, 217-224. Fincham, D. A., Mason, D. K., and Young, J. D. (1985). Characterization of a novel Na+-independent amino acid transporter in horse erythrocytes. Biochem. J. 227, 13-20. Finean, J. B., and Michell, R. H. (1981). Isolation, composition and general structure of membranes. In "Membrane Structure" (J. B. Finean and R. H. Michell, eds.), pp. 1-36. Elsevier/North-Holland Biochemical Press, Amsterdam. Finkel, A. S., and Gage, P. W. (1985). Conventional voltage clamping with two intracellular microelectrodes. In "Voltage and Patch Clamping with Microelectrodes" (T. G. Smith, H. Lecar, S. J. Redman, and P. W. Gage, eds.), pp. 47-94. Williams & Wilkins, Baltimore. Fischbarg, J., and Vera, J. C. (1995). Multifunctional transporter models: Lessons from the transport of water, sugars, and ring compounds by GLUTs. Am. J. Physiol. 268, C1077-C1089. Fisher, S. E., and Karl, P. I. (1990). Histidine transfer across the human placenta: Characteristics in the isolated perfused human placenta and the effect of ethanol. Placenta 11, 157-165. Fitscher, B. A., Elsing, C., Riedel, H.-D., Gorski, J., and Stremmel, W. (1996). Protein-mediated facilitated uptake processes for fatty acids, bilirubin, and other amphipathic compounds. Proc. Soc. Exp. Biol. Med. 212, 15-23. Fleck, C. (1992). Renal transport of endogenous amino acids. II. Influence of treatment with triiodothyronine or dexamethasone in immature and adult rats. Ren. Physiol. Biochem. 15, 266-276. Fliegel L., Haworth R. S., and Dyck J. R. B. (1993). Characterization of the placental brush border membrane Na+/H § exchanger: Identification of thio-dependent transitions in apparent molecular size. Biochem. J. 289, 101-107. Flier J. S., Mueckler M. M., Usher P., and Lodish H. F. (1987). Elevated levels of glucose transport and transporter messenger RNA are induced by ras or src oncogenes. Science 235, 1492-1495. Forsythe, I. D., Linsdell, P., and Stanfield, P. R. (1992). Unitary A-currents of rat locus coeruleus neuornes grown in cell culture: Rectification cause by internal Mg 2§ and Na § J. Physiol. 451, 553-583. Foster, D. L., and Fillingame, R. H. (1982). Stoichiometry of subunits in the H+-ATPase complex of Escherichia coli. J. Biol. Chem. 257, 2009-2015. Fraga, D., Hermolin, J., and Fillingame, R. H. (1994). Transmembrane helix-helix interactions in Fo suggested by suppressor mutations to Ala 24 ~ Asp/Asp 61 ~ Gly mutant of ATP synthase subunit c. J. Biol. Chem. 269, 2562-2567. Fragoso, S. P., and Goldenberg, S. (1992). Cloning and characterization of the gene encoding Trypanosoma cruzi DNA topoisomerase II. Mol. Biochem. Parasitol. 55, 127-134. Franchi-Gazzola, R., Visigalli, R., Bussolati, O., and Gazzola, G. C. (1996). Involvement of protein kinase Ce in the stimulation of anionic amino acid transport in cultured human fibroblasts. J. Biol. Chem. 271, 26124-26130. Franchi-Gazzola, R., Visigalli, R., Bussolati, O., Guidotti, G. G., and Gazzola, G. C. (1991). Phorbol esters stimulate

References the transport of anionic amino acids in cultured human fibroblasts. Biochem. Biophys. Res. Commun. 173, 13041310. Frankenhaeuser, B., and Moore, L. E. (1963). The effect of temperature on the sodium and potassium permeability changes in myelinated nerve fibres of Xenopus laevis. J. Physiol. 169, 431-437. Frayn, K. N. (1996). "Metabolic Regulation: A Human Perspective." Portland Press, London. Frayn, K. N., Khan, K., Coppack, S. W., and Elia, M. (1991). Amino acid metabolism in human subcutaneous adipose tissue in vivo. Clin. Sci. 80, 471-474. French, P. J., Bijman, J., Edixhoven, M., Vaandrager, A. B., Scholte, B. J., Lohmann, S. M., Nairn, A. C., and de Jonge, H. R. (1995). Isotype-specific activation of cystic fibrosis transmembrane conductance regulator-chloride channels by cGMP-dependent protein kinase II. J. Biol. Chem. 270, 26626-26631. Frillingos, S., Sahin-T6th, M., Wu, J., and Kaback, H. R. (1998). Cys-scanning mutagenesis: A novel approach to structure-function relationships in polytopic membrane proteins. FASEB J. 12, 1281-1299. Fr6hlich, O. (1988). The "tunneling" mode of biological carrier-mediated transport. J. Membr. Biol. 101, 189-198. Frye, L. D., and Edidin, M. (1970). The rapid intermixing of cell surface antigens after formation of mouse-human heterokaryons. J. Cell Sci. 7, 319-335. Fuerst, T. R., Niles, E. G., Studier, F. W., and Moss, B. (1986). Eukaryotic transient-expression system based on recombinant vaccinia virus that synthesizes bacteriophage T7 RNA polymerase. Proc. Natl. Acad. Sci. USA 83, 8122-8126. Fuhrmann, G. F., Haas, D., and Dittmar, T. (1992). Evidence for a high and low affinity C1-/HCO3- exchange system in human red cells. Progr Cell Res. 2, 99-101. Fukumoto, H., Kayano, T., Buse, J. B., Edwards, Y., Pich, P. F., Bel, G. I., and Seino, S. (1989). Cloning and characterization of the major insulin-responsive glucose transporter expressed in human skeletal muscle and other insulinresponsive tissues. J. Biol. Chem. 264, 7776-7779. Furesz, T. C., Smith, C. H., and Moe, A. J. (1993). ASC system activity is altered by development of cell polarity in trophoblast from human placenta. Am. J. Physiol. 265, C212-C217. Garcia-Sancho, J., Sanchez, A., and Christensen, H. N. (1977). Free carboxylate groups required for transport of neutral amino acids by the Erhlich ascites tumor cell. Biochim. Biophys. Acta 465, 426-428. Gallardo, E. M., Hirayama, B. A., Loo, D. D. F., Lam, J. T., Turk, E., and Wright, E. M. (1998). Identification of the sugar translocation pathway of the NA+/glucose cotransporter (SGLT1). FASEB J. 12, A1017. [Abstract] Ganapathy, V., and Leibach, F. H. (1983). Role of pH gradient and membrane potential in dipeptide transport in intestinal and renal brush-border membrane vesicles from the rabbit. J. Biol. Chem. 258, 14189-14192. Garlid, K. D. (1990). New insights into mechanisms of anion uniport through the uncoupling protein of brown adipose tissue mitochondria. Biochim. Biophys. Acta 1018, 151-154.

357

Garlid, K. D., Orosz, D. E., Modriansky, M., Vassanelli, S., and Je~ek, P. (1996). On the mechanism of fatty acidinduced proton transport by mitochondrial uncoupling protein. J. Biol. Chem. 271, 2615-2620. Gardner, M. L. G., and Atkins, G. L. (1982). Kinetic analysis of transport processes in the intestine and other tissues. Clin. Sci. 63, 405-414. Gartner, E. M., Liebold, K., Legrum, B., Fasold, H., and Passow, H. (1997). Three different actions of phenylglyoxal on band 3 protein-mediated anion transport across the red blood cell membrane. Biochim. Biophys. Acta 1323, 208-222. Garrick, R. A., Patel, B. C., and Chinard, F. P. (1980). Permeability of dog erythrocytes to lipophilic molecules: Solubility and volume effects. Am. J. Physiol. 238, C107-Cl13, Garrick, R. A., Patel, B. C., and Chinard, F. P. (1982). Erythrocyte permeability to lipophilic solutes changes with temperature. Am. J. Physiol. 242, C74-C80. Garvey, W. T., Huecksteadt, T. P., and Birnbaum, M. J. (1989). Pretranslational suppression of an insulin-responsive glucose transporter in rats with diabetes mellitus. Science 245, 60-63. Garvey, W. T., Maianu, L., Huecksteadt, T. P., Birnbam, M. J., Molina, J. M., and Ciaraldi, T. P. (1991). Pretranslational suppression of a glucose transporter protein causes insulin resistance in adipocytes from patients with noninsulin-dependent diabetes mellitus and obesity. J. Clin. Invest. 87, 1072-1081. Gastaldelli, A., Schwarz, J. M., Caveggion, E., Traber, L. D., Traber, D. L., Rosenblatt, J., Toffolo, G., Cobelli, C., and Wolfe, R. R. (1997). Glucose kinetics in interstitial fluid can be predicted by compartmental modeling. Am. J. Physiol. 272, E494-E505. Gazzola, G. C., Dall'Asta, V., Bussolati, O., Makowske, M., and Christensen, H. N. (1981). A stereoselective anomaly in dicarboxylic amino acid transport. J. Biol. Chem. 256, 6054-6O59. Gazzola, G. C., Dall'Asta, V., Franchi-Gazzola, R., Bussolati, O., Longo, N., and Guidotti, G. G. (1985). In "Cell Membranes and Cancer" (T. Galeotti, A. Cittadini, G. Neri, S. Papa, and L. A. Smets eds.), Elsevier, Amsterdam. Geck, P. (1971). Properties of an asymmetrical carrier model for the transport of sugars by human erythrocytes. Biochim. Biophys. Acta 241, 462-472. Geck, P., and Heinz, E. (1976). Coupling in secondary transport: Effect of electrical potentials on the kinetics of ion linked co-transport. Biochim. Biophys. Acta 443, 49-53 Gedde, M. M., Davis, D. K., and Huestis, W. H. (1997). Cytoplasmic pH and human erythrocyte shape. Biophys. J. 72, 1234-1246. Geddie, G., Moores, R., Meschia, G., Fennessey, P., Wilkening, R., and Battaglia, F. C. (1996). Comparison of leucine, serine and glycine transport across the ovine placenta. Placenta 17, 619-627. Gegelashvili, G., and Schousboe, A. (1997). High affinity glutamate transporters: regulation of expression and activity. MoL PharmacoL 52, 6-15. Gehrig, H., Mtiller, W., and Appelhans, H. (1992). Complete nucleotide sequence of band 3 related anion transport pro-

358

References

tein AE2 from human kidney. Biochim. Biophys. Acta 1130, 326-328. Gerasimova, T. I., Mizrokhi, L. J., and Georgiev, G. P. (1984). Transposition bursts in genetically unstable Drosphila melanogaster. Nature 309, 714-716. Gibbs, E. M., Stock, J. L., McCoid, S. C., Stukenbrok, H. A., Pessin, J. E., Stevenson, R. W., Milici, A. J., and McNeish, J. D. (1995). Glycemic improvement in diabetic db/db mice by overexpression of the human insulin-regulatable glucose transporter (GLUT4). J. Clin. Invest. 95, 1512-1518 Girvin, M. E., and Fillingame, R. H. (1995). Determination of local protein structure by spin label difference 2D NMR: The region neighboring Asp61 of subunit c of the F1Fo ATP synthase. Biochemistry 34, 1635-1645. Girvin, M. E., Rastogi, V. K., Abildgaard, F., Markley, J. L., and Fillingame, R. H. (1998). Solution structure of the transmembrane H+-transporting subunit c of the FIFo ATP synthase. Biochemistry 37, 8817-8824. Golan, D. E., Corbett, J. D., Korsgren, C., Thatte, H. S., Hayette, S., Yawata, Y., and Cohen, C. M. (1996). Control of band 3 lateral and rotational mobility by band 4.2 in intact erythrocytes: Release of band 3 oligomers from lowaffinity binding sites. Biophys. J. 70, 1534-1542. Goldsmith, E. J. (1996). Allosteric enzymes as models for chemomechanical energy transducing assemblies. FASEB J. 10, 702-708. Goldstein, L., Davis-Amaral, E. M., and Musch, M. W. (1996). Organic osmolyte channels: Transport characteristic and regulation. Kidney Int. 49, 1690-1694. Gomez-Angelats, M., del Santo, B., Mercader, J., FerrarMartinez, A., Felipe, A., Casado, J., and Pastor-Anglada, M. (1996). Hormonal regulation of concentrative nucleoside transport in liver parenchymal cells. Biochem. J. 313, 915-920 Gomperts, S. N. (1996). Clustering membrane proteins: Its all coming together with the PSD-95/SAP90 protein family. Cell. 84, 659-662. Gonz~ilez-Barroso, M. M., Fleury, C., Bouillaud, F., Nicholls, D. G., and Rial, E. (1998). The uncoupling protein UCP1 does not increase the proton conductance of the inner mitochondrial membrane by functioning as a fatty acid anion transporter. J. Biol. Chem. 273, 15528-15532. Goodson, H. V., Valetti, C., and Kreis, T. E. (1997). Motors and membrane traffic. Curr. Opin. Cell Biol. 9, 18-28. Gottardi, C. J., and Caplan, M. J. (1993). An ion-transporting ATPase encodes multiple apical localization signals. J. Cell Biol. 121, 283-293. Grabner, M., Friedrich, K., Knaus, H.-G., Striessnig, J., Scheffauer, F., Staudinger, R., Koch, W. J., Schwartz, A., and Glossmann, H. (1991). Calcium channels from Cyprinus carpio skeletal muscle. Proc. Natl. Acad. Sci. USA 88, 727-731. Graf, J., and H~iussinger, D. (1996). Ion transport in hepatocytes: Mechanisms and correlations to cell volume, hormone actions and metabolism. J. Hepatol. 24(Suppl. 1), 53-77. Green, H., and Wang, N. (1994). Codon reiteration and the evolution of proteins. Proc. Natl. Acad. Sci USA 91, 42984302.

Gregoriadis, G. (1993). "Liposome Technology", Vols. I-III, 2nd ed. CRC Press, Boca Raton, FL. Gregory, R. J., Kammermeyer, K. L., Vincent, W. S., and Wadsworth, S. G. (1987). Primary sequence and developmental expression of a novel Drosophila melanogaster src gene. Mol. Cell Biol. 7, 2119-2127. Griffith, J. K., and Sansom, C. E. (1998). "The Transporter Facts Book". Academic Press, San Diego. Grill, V., Bjorkman, O., Gutniak, M., and Lindqvist, M. (1992). Brain uptake and release of amino acids in nondiabetic and insulin-dependent diabetic subjects: Important role of glutamine release for nitrogen balance. Metabolism 41, 28-32. Groth, G., and Walker, J. E. (1997). Model of the c-subunit oligomer in the membrane domain of F-ATPases. FEBS Lett. 410, 117-123. Groth, G., Tilg, Y., and Schirwitz, K. (1998). Molecular architecture of the c-subunit oligomer in the membrane domain of F-ATPases probed by trytophan substitution mutagenesis. J. Mol. Biol. 281, 49-59. Grover, S., Woodward, S. R., and Odell, W. D. (1993). A bacterial protein has homology with human chorionic gonadotropin (hCG). Biochem. Biophys. Res. Commun. 193, 841-847. Groves, J. D., Wang, L., and Tanner M. J. A. (1998). Functional reassembly of the anion transport domain of human red cell band 3 (AE1). from multiple and non-complementary fragments. FEBS Lett. 433, 223-227. Grunewald, M., and Kanner, B. (1995). Conformational changes monitored on the glutamate transporter GLT-1 indicate the existence of two neurotransmitter-bound states. J. Biol. Chem. 270, 17017-17024. Grunewald, M., Bendahan, A., and Kanner, B. I. (1998). Biotinylation of single cysteine mutants of the glutamate transporter GLT-1 from rat brain reveals its unusual topology. Neuron 21, 623-632. Guerini, D., Foletti, D., Vellani, F., and Carafoli, E. (1996). Mutation of conserved residues in transmembrane domains 4,6 and 8 causes loss of Ca 2+ transport by the plasma membrane Ca 2+ pump. Biochemistry 35, 3290-3296. Guerrini, L., Gong, S. S., Mangasarian, K., and Basilico, C. (1993). Cis- and trans-acting elements involved in amino acid regulation of asparagine synthetase gene expression. Mol. Cell Biol. 13, 3202-3221. Guidotti, G. G., and Gazzola G. C. (1992). Amino acid transporters: Systematic approach and principles of control. In "Mammalian Amino Acid Transport: Mechanisms and Control" (M. S. Kilberg and D. H~iussinger, eds.), pp. 3-30. Plenum, New York/London. Guiltinan, M. J., Ma, D.-P., Barker, R. F., Bustos, M. M., Cyr, R. J., Yadegari, R., and Fosket, D. E. (1987). The isolation, characterization and sequence of two divergent/3-tubulin genes from soybean (Glycine max L.). Plant Mol. Biol. 10, 171-184. Gulve, E. A., Ren, J. M., Marshall, B. A., Gao, J., Hansen, P. A., Holloszy, J. O., and Mueckler, M. (1994). Glucose transport activity in skeletal muscles from transgenic mice overexpressing GLUT1: Increased basal transport is associ-

References ated with a defective response to diverse stimuli that activate GLUT4. J. Biol. Chem. 269, 18366-18370. Gunshin, H., Mackenzie, B., Berger, U. V., Gunshin, Y., Romero, M. F., Boron, W. F., Nussberger, S., Gollan, J. L., and Hediger, M. A. (1997). Cloning and characterization of a proton-coupled mammalian metal ion transporter. Nature 388, 482-488. Gurdon, J. B., Lane, C. D., Woodland, H. R., and Marbaix, G. (1971). Use of frog eggs and oocytes for the study of messenger RNA and its translation in living cells. Nature 233, 177-182. Guy, H. R., and Durell, S. R. (1995). Structural models of Na +, Ca 2+, and K + channels Soc. Gen. Physiol. Ser. 50,1-16. Guy, H. R., and Durell, S. R. (1996). Developing three-dimensional models of ion channel proteins. Ion Channels 4,1-40. Habib-Mohamed, A., and Steck, T. L. (1986). Band 3 tyrosine kinase: Association with the human erythrocyte membrane. J. Biol. Chem. 261, 2804-2809. Hack, V., Weiss, C., Friedmann, B., Suttner, S., Schykowski, M., Erbe, N., Benner, A., Brtsch, P., and DrOge, W. (1997). Decreased plasma glutamine level and CD4+T cell number in response to 8 wk of anaerobic training. Am. J. Physiol. 272, E788-E795. Hackstadt, T., Rockey, D. D., Heinzen, R. A., and Scidmore, M. A. (1996). Chlamydia trachomatis interrupts an exocytotic pathway to acquire endogenously synthesized sphingomyelin in transit from the Golgi apparatus to the plasma membrane. E M B O J. 15, 964-977. Hager, K., Hazama, A., Kwon, H. M., Loo, D. D. F., Handler, J. S., and Wright, E. M. (1995). Kinetics and specificity of the renal Na§ cotransporter expressed in Xenopus oocytes. J. Membr. Biol. 143, 103-113. Haghighat, N., and Van Winkle, L. J. (1990). A developmental change in follicular cell-enhanced amino acid uptake into mouse oocytes that depends on intact gap junctions and transport system Gly. J. Exp. Zool. 253, 71-82. Hagiwara, S., and Yoshii, M. (1979). Effects of internal potassium and sodium on the anomalous rectification of the starfish egg as examined by internal perfusion. J. Physiol. 292, 251-265. Haines, T. H. (1994). Water transport across biological membranes. FEBS Lett. 346, 115-122. Hajjar, D. P., and Nicholson, A. C. (1995). Atherosclerosis. Am. Sci. 83, 460-467. Hajela, K., Kayestha, R., and Sumati (1996). Carbohydrate induced modulation of cell membrane. II. Spin label study of fluidity changes in peripheral blood lymphocyte membrane. FEBS Lett. 380, 165-168. Hamasaki, N., and Okubo, K. (1996). Band 3 protein: Physiology, function and structure. Cell Mol. Biol. 42, 1025-1039. Hamasaki, N., Okubo, K., and Kang, D. (1992). Protein chemistry of the anion transport center of erythrocyte band 3. Progr. Cell Res. 2, 65-71. Hammerton, R., Krezeminski, K., Mays, R., Ryan, T., Wollner, D., and Nelson, W. (1991). Mechanism for regulating cell surface distribution of Na+, K+-ATPase in polarized epithelial cells. Science 254, 847-850. Handlogten, M. E., Garcia-Canero, R., Lancaster, K. T., and Christensen, H. N. (1981). Surprising differences in sub-

359

strate selectivity and other properties of systems A and ASC between rat hepatocytes and the hepatoma cell line HTC. J. Biol. Chem. 256, 7905-7909. Handlogten, M. E., Dudenhausen, E. E., Yang, W., and Kilberg, M. S. (1996). Association of hepatic system A amino acid transporter with the membrane-cytoskeletal proteins ankyrin and fodrin. Biochim. Biophys. Acta 1282, 107-114. Hankard, R. G., Darmaun, D., Sager, B. K., D'Amore, D., Parsons, W. R., and Haymond, M. (1995). Response of glutamine metabolism to exogenous glutamine in humans. Am. J. Physiol. 269, E663-E670. Hankard, R. G., Haymond, M. W. and Darmaun, D. (1996). Effect of glutamine on leucine metabolism in humans. Am. J. Physiol. 271, E748-E754. Hansen, P. A., Gulve, E. A., Marshall, B. A., Gao, J., Pessin, J. E., Holloszy, J. O., and Mueckler, M. (1995). Skeletal muscle glucose transport and metabolism are enhanced in transgenic mice overexpressing the Glut4 glucose transporter. J. Biol. Chem. 270, 1679-1684. Harper, A. E., Miller, R. H., and Block, K. P. (1984). Branched-chain amino acid metabolism. Annu. Rev. Nutr. 4, 409-454. Harper, M.-E. (1997). Obesity research continues to spring leaks. Clin. Invest. Med. 20, 239-244. Harris, S. L., Na, S., Zhu, X., Seto-Young, D., Perlin, D. S., Teem, J. H., and Haber, J. E. (1994). Dominant lethal mutations in the plasma membrane H+-ATPase gene of Saccharomyces cerevisiae. Proc. Natl. Acad. Sci. USA 91, 10531-10535. Haugeto, O., Ullensvang, K., Levy, L. M., Chaudhry, F. A., Honor6, T., Nielsen, M., Lehre, K. P., and Danbolt, N. C. (1996). Brain glutamate transporter proteins form homomultimers. J. Biol. Chem. 271, 27715-27722. H~iussinger, D. (1996). The role of cellular hydration in the regulation of cell function. Biochem. J. 313, 697-710. H~iussinger, D. (1990). Nitrogen metabolism in liver: Structural and functional organization and physiological relevance, Biochem. J. 267, 281-290. H~iussinger, D, Roth, E., Lang, F., and Gerok, W. (1993). Cellular hydration state: An important determinant of protein catabolism in health and disease. Lancet 341, 13301332. Haws, C., Finkbeiner, W. E., Widdicombe, J. H., and Wine, J. J. (1994). CFTR in Calu-3 human airway cells: Channel properties and role in cAMP-activated C1-conductance. Am. J. Physiol. 266, L502-L512. Hazama, A., Loo, D. D. F., and Wright, E. M. (1997). Presteady-state currents of the rabbit Na+/glucose cotransporter (SGLT1). J. Membr. Biol. 155, 175-186. Heal, R. D., and McGivan, J. D. (1998). Induction of calreticulin by amino acid deprivation in CHO cells. Biochem. J. 329, 389-394. Heale, S. M. ,and Petes, T. D. (1995). The stabilization of repetitive tracts of DNA by variant repeats. Cell 83, 539-545. Hebert, S. C. (1998). General principles of the structure of ion channels. Am. J. Med. 104, 87-98. Hebert, S. C., and Reeves, W. B. (1995). Introduction to a symposium on ion channels. Kidney Int. 48, 917-918.

360

References

Hediger, M. A., Coady, M. J., Ikeda, T. S., and Wright, E. M. (1987). Expression cloning and cDNA sequencing of the Na+/glucose cotransporter. Nature 330, 379-381. Hediger, M. A., Mendlein, J., Lee, H. S., and Wright, E. M. (1991). Biosynthesis of the cloned intestinal Na+/glucose cotransporter. Biochim. Biophys. Acta 1064, 360-364. Hediger, M. A., Kanai, Y., You, G., and Nussberger, S. (1995). Mammalian ion-coupled solute transporters. J. Physiol. (London) 482, 7S-17S. Heginbotham, L., Abramson, T., and 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., and MacKinnon, R. (1994). Mutations in the K + channel signature sequence. Biophys. J. 66, 1061-1067. Hegyvary, C. (1973). Effects of some organic solvents on the reactivity of sodium plus potassium ion-transport ATPase. Biochim. Biophys. Acta 311, 272-291. Hemken, P., Guo, X.-L., Wang, Z.-Q., Zhang, K., and Gluck, S. (1992). Immunologic evidence that vacuolar H + ATPases with heterogeneous forms of Mr = 31,000 subunit have different membrane distributions in mammalian kidney. J. Biol. Chem. 267, 9948-9957. Hennemann, G., Krenning, E. P., Polhuys, M., Mol, J. A., Bernard, B. F., Visser, T. J., and Docter, R. (1986). Carriermediated transport of thyroid hormone into rat hepatocytes is rate-limiting in total cellular uptake and metabolism. Endocrinology 119, 1870-1872. Henry, P., Pearson, W. L., and Nichols, C. G. (1996). Protein kinase C inhibition of cloned inward rectifier (HRK1/ Kit2.3). K + channels expressed in Xenopus oocytes. J. Physiol. 495, 681-688. Henthorn, P. S., Knoll, B. J., Raducha, M., Rothblum, K. N., Slaughter, C., Weiss, M., Lafferty, M. A., Fischer, T., and Harris, H. (1986). Products of two common alleles at the locus for human placental alkaline phosphatase differ by seven amino acids. Proc. Natl. Acad. Sci. USA 83, 55975601. Herbert, D. N., and Carruthers, A. (1992). Glucose transporter oligomeric structure determines transporter function. J. Biol. Chem. 267, 23839-23838. Hernfindez, J. A. (1998). Simple carrier kinetics in complex membrane transporters. J. Membr. Biol. 165, 235-242. Hernando, N., Bartkiewicz, M., Collin-Osdoby, P., Osdoby, P., and Baron, R. (1995). Alternative splicing generates a second isoform of the catalytic A subunit of the vacuolar H+-ATPase. Proc. Natl. Acad. Sci. USA 92, 6087-6091. Higgins, C. F. (1992). ABC transporters: From microorganisms to man. Annu. Rev. Cell Biol. 8, 67-113. Hilgier, W., Puka, M., and Albrecht, J. (1992). Characteristics of large neutral amino acid-induced release of preloaded I:glutamine from rat cerebral capillaries in vitro: Effects of ammonia, hepatic encephalopathy and gamma-glutamyl transpeptidase inhibitors. J. Neurosci. Res. 32, 221-226. Hill, A. E. (1995). Osmotic flow in membrane pores. Int. Rev. Cytol. 163, 1-42. Hill, M. W. (1974). The effect of anaesthetic-like molecules on the phase transition in smectic mesophases of dipalmi-

toyllecithin. I. The normal alcohol up to C = 9 and three inhalation anesthetics. Biochim. Biophys. Acta 356, 117-124. Hille, B. (1984). "Ionic Channels of Excitable Membranes", 1st ed. Sinauer Associates, Sunderland, MA. Hille, B. (1992). "Ionic Channels of Excitable Membranes", 2nd ed. Sinauer Associates, Sunderland, MA. Hille, B., and Schwarz, W. (1978). Potassium channels as multiion single-file pores. J. Gen. Physiol. 72, 409-442. Hiraki, Y., Garcia de Herreros, A., and Birnbaum, M. J. (1989). Transformation stimulates glucose transporter gene expression in the absence of protein kinase C. Proc. Natl. Acad. Sci. USA 86, 8252-8256. Hirata, R., and Anraku, Y. (1992). Mutations at the putative junction sites of the yeast VMA1 protein, the catalytic subunit of the vacuolar membrane H+-ATPase, inhibit its processing by protein splicing. Biochem. Biophys. Res. Commun. 188, 40-47. Hirayama, B. A., Loo, D. D. F., and Wright, E. M. (1997). Cation effects on protein conformation and transport in the Na+/glucose cotransporter. J. Biol. Chem. 272, 2110-2115. Hirayama, B. A., Lostao, M. P., Panayotova-Heiermann, M., Loo, D. D. F., Turk, E. and Wright, E. M. (1996). Kinetic and specificity differences between rat, human, and rabbit Na+-glucose cotransporters (SGLT-1).. Am. J. Physiol. 270, G919-G926. Hirsch, J. R., Loos, D. D. F., and Wright, E. M. (1996). Regulation of Na+/glucose cotransporter expression by protein kinases in Xenopus laevis oocytes. J. Biol. Chem. 271, 14740-14746. Ho, K., Nichols, C. G., Lederer, W. J., Lytton, J., Vassilev, P. M., Kanazirska, M. V., and Hebert, S. C. (1993). Cloning and expression of an inwardly rectifying ATP-regulated potassium channel. Nature 362, 31-38. Hodgkin, A. L., and Keynes, R. D. (1955a) Active transport of cations in giant axons from Sepia and Loligo. J. Physiol. 128, 28-60. Hodgkin, A. L., and Keynes, R. D. (1955b) The potassium permeability of a giant nerve fibre. J. Physiol. 128, 61-88. Hohl, D., Mehrel, T, Lichti, U., Turner, M. L., Roop, D. R., and Steinert, P. M. (1991). Characterization of human loricrin: Structure and function of a new class of epidermal cell envelope proteins. J. Biol. Chem. 266, 6626-6636. Hopfer, U. (1984). Criticisms and limitations of current transport studies with isolated membrane vesicles. Kroc Found. Ser. 17, 27-33. Hopfer, U. (1987). Membrane transport mechanisms for hexoses and amino acids in the small intestine. In "Physiology of the Gastrointestinal Tract" (L. R. Johnson, ed.), pp. 1499-1526. Raven, New York. Hopfer, U. (1989). Tracer studies with isolated membrane vesicles. "Methods in Enzymology" (S. Fleischer and B. Fleischer, eds.), Vol. 172, pp. 313-331. Academic Press, San Diego. Horn, R., and Patlak, J. (1980). Single channel currents from excised patches of muscle membrane. Proc. Natl. Acad. Sci. USA 77, 6930-6934. Hoshi, T., Zagotta, W. N., and Aldrich, R. W. (1990). Biophysical and molecular mechanisms of Shaker potassium channel inactivation. Science 250, 533-538.

References

Hoshi, T., Zagotta, W. N., and Aldrich, R. W. (1991). Two types of inactivation in Shaker K + channels: Effects of alterations in the carboxy-terminal region. Neuron 7, 547-556. Hosokawa, H., Sawamura, T., Kobayashi, S., Ninomiya, H., Miwa, S., and Masaki, T. (1997). Cloning and characterization of a brain-specific cationic amino acid transporter. J. Biol. Chem. 272, 8717-8722. Hsieh, T.-S., Brown, S. D., Huang, P., and Fostel, J. (1992). Isolation and characterization of a gene encoding DNA topoisomerase I in Drosophila melanogaster. Nucleic Acids Res. 20, 6177-6182. H0bner, S., Michel, F., Rudloff, V., and Appelhans, H. (1992). Amino acid sequence of band 3 protein from rainbow trout erythrocytes derived from cDNA. Biochem. J. 285, 17-23. Huff, R. A., Vaughan, R. A., Kuhar, M. J., and Uhl, G. R. (1997). Phorbol esters increase dopamine transporter phosphorylation and decrease transport Vmax. J. Neurochem. 68, 225-232. Hui, S. W. (1995). Geometry of domains and domain boundaries in monolayers and bilayers. Mol. Membr. Biol. 12, 45-50. Hundal, H. S., Rennie, M. J., and Watt, P. W. (1987). Characteristics of L-glutamine transport in perfused rat skeletal muscle. J. Physiol. 393, 283-305. Huntington's Disease Collaborative Research Group (1993). A novel gene containing a trinucleotide repeat that is expanded and unstable on Huntington's disease chromosomes. Cell 72, 971-983. Hwang, T.-C., Horie, M., and Gadsby, D. C. (1993). Functionally distinct phospho-forms underlie incremental activation of protein kinase-regulated C1- conductance in mammalian heart. J. Gen. Physiol. 101, 629-650. Hwang, T.-C., Wang, F., Yang, I. C., and Reenstra, W. W. (1997). Genistein potentiates wild-type and 6F508-CFTR channel activity. Am. J. PhysioL 273, C988-C998. Inaba, M., Yawata, A., Koshino, I., Sato, K., Takeuchi, M., Takakuwa, Y., Manno, S., Yawata, Y., Kanzaki, A., Sakai, J.-i., Ban, A., Ono, K.-i., and Maede, Y. (1996). Defective anion transport and marked spherocytosis with membrane instability caused by hereditary total deficiency of red cell band 3 in cattle due to a nonsense mutation. J. Clin. Invest. 977 1804-1817. Inagaki, N., Gonoi, T., Clement, J. P., Namba, N., Inazawa, J., Gonzalez, G., Aguilar-Bryan, L., Seino, S., and Bryan, J. (1995). Reconstitution of IKATP:An inward rectifier subunit plus the sulfonylurea receptor. Science 270, 1166-1170. Inagaki, N., Tsuura, Y., Namba, N., Masuda, K., Gonoi, T., Horie, M., Seino, Y., Mizuta, M., and Seino, S. (1995). Cloning and functional characterization of a novel ATPsensitive potassium channel ubiquitously expressed in rat tissues, including pancreatic islets, pituitary, skeletal muscle, and heart. J. Biol. Chem. 270, 5691-5694. Ingber, D. (1997). Tensegrity: The architectural basis of cellular mechanotransduction. Annu. Rev. Physiol. 59, 575-599. Inoue, T., Kanzaki, A., Yawata, A., Tsuji, A., Ata, K., Okamoto, N., Wada, H., Higo, I., Sugihara, T., Yamada, O., and Yawata, Y. (1994). Electron microscopic and physico-

361

biochemical studies on disorganization of the cytoskeletal network and integral protein (band 3). in red cells of band 4.2 deficiency with a mutation (codon 142:GCT ~ ACT). Int. J. Hematol. 59, 157-175. Ionov, Y., Peinado, M. A., Malkhosyan, S., Shibata, D., and Perucho, M. (1993). Ubiquitous somatic mutations in simple repeated sequences reveal a new mechanism for colonic carcinogenesis. Nature 363, 558-561. Inui, Y., and Christensen, H. N. (1966). Discrimination of single transport systems: The Na+-sensitive transport of neutral amino acids in the Ehrlich cell. J. Gen. Physiol. 50, 203-224. Iseki, K., Kaido, K.-I., Kobayashi, M., Sugawara, M., and Miyazaki, K. (1997). The effect of membrane surface potential on the permeability of anionic compounds across the apical membrane in human intestinal epithelial (Caco-2). cells. Biol. Pharmacol. Bull 20, 794-799. Ishii, T., Hata, F., Lemas, M. V., Fambrough, D. M., and Takeyasu, K. (1997). Carboxyl-terminal regions of the sarcoplasmic/endoplasmic reticulum Ca e+ and the Na§ Pases control their K § sensitivity. Biochemistry 36, 442-451. Isomoto, S., Kondo, C., Takahashi, N., Matsumoto, S., Yamada, M., Takumi, T., Horio, Y., and Kurachi, Y. (1996). A novel ubiquitously distributed isoform of GIRK2 (GIRK2B) enhances GIRK1 expression of the G-proteingated K § current in Xenopus oocytes. Biochem. Biophys. Res. Commun. 218, 286-291. Isomoto, S., Kondo, C., and Kurachi, Y. (1997). Inwardly rectifying potassium channels: their molecular heterogeneity and function. Jpn. J. Physiol. 47, 11-39. Isselbacher, K. J. (1972). Sugar and amino acid transport by cells in culture: Differences between normal and malignant cells. N. Engl. J. Med. 286, 929-933. Ito, K., and Groudine, M. (1997). A new member of the cationic amino acid transporter family is preferentially expressed in adult mouse brain. J. Biol. Chem. 272, 2678026786. Jacobson, K., Sheets, E. D., and Simson, R. (1995). Revisiting the fluid mosaic model of membranes. Science 268, 14411442. Jaggi, R., Salmons, B., Muellner, D., and Groner, B. (1986). The v-mos and H-ras oncogene expression represses glucocorticoid hormone-dependent transcription from the mouse mammary tumor virus LTR. EMB O J. 5, 2609-2616. Jakobsson, E. (1997). Computer simulation studies of biological membranes: Progress, promise and pitfalls. Trends Biochem. Sci. 22, 339-344. James, D. E., Brown, R., Navarro, J., and Pilch, P. F. (1988). Insulin-regulatable tissues express a unique insulin-sensitive glucose transport protein. Nature 333, 183-185. James, D. E., Strube, M., and Mueckler, M. (1989). Molecular cloning and characterization of an insulin-regulatable glucose transporter. Nature 338, 83-87. Jan, L. Y., and Jan, Y. N. (1992). Structural elements involved in specific K § channel functions. Annu. Rev. Physiol. 54, 537-555. Jan, L. Y., and Jan, Y. N. (1994). Potassium channels and their evolving gates. Nature 371, 119-122.

362

References

Jan, L. Y., and Jan, Y. N. (1997). Cloned potassium channels from eukaryotes and prokaryotes. Annu. Rev. Neurosci. 20, 91-123. Jauch, P., and Liauger, P. (1986). Electrogenic properties of the sodium-alanine cotransporter in pancreatic acinar cells. II. Comparison with transport models. J. Membr. Biol. 94, 117-127. Jencks, W. P. (1989). How does a calcium pump pump calcium? J. Biol. Chem. 264, 18855-18858. Jennings, M. L. (1982). Stoichiometry of a half-turnover of band 3, the chloride transport protein of human erythrocytes J. Gen. Physiol. 79, 169-185. Jette, M., Vachon, V., Potier, M., and Beliveau, R. (1996). The renal sodium/phosphate symporters: Evidence for different functional oligomeric states. Biochemistry 35,15209-15214. Jette, M., Vachon, V., Potier, M., and Beliveau, R. (1997). Radiation-inactivation analysis of the oligomeric structure of the renal sodium/D-glucose symporter. Biochim. Biophys. Acta 1327, 242-248. Je~ek, P., Orosz, D. E., Modriansky, M., and Garlid, K. D. (1994). Transport of anions and protons by the mitochondrial uncoupling protein and its regulation by nucleotides and fatty acids. J. Biol. Chem. 269, 26184-26190. Je~ek, P., Modriansky, M., and Garlid, K. D. (1997). A structure-activity study of fatty acid interaction with mitochondrial uncoupling protein. FEBS Lett. 408, 166-170. Jiang, J., Gu, B., Albright, L. M., and Nixon, B. T. (1989). Conservation between coding and regulatory elements of Rhizombium meliloti and Rhizobium leguminosarum dct genes. J. Bacteriol. 171, 5244-5253. Johansson, I., Karlsson, M., Shukla V. K., Chrispeels, M. J., Larsson, C., and Kjelbom, P. (1998). Water transport activity of the plasma membrane aquaporin PM28A is regulated by phosphorylation. The Plant Cell 10, 451-459. Johnson, J. H., Ogawa, A., Chen, L., Orci, L., Newgard, C. B., Alam, T., and Unger, R. H. (1990). Underexpression of /3 cell high Km glucose transporters in noninsulindependent diabetes. Science 250, 546-549. Johnson, L. W., and Smith, C. H. (1988). Neutral amino acid transport systems of microvillous membrane of human placenta. Am. J. Physiol. 254, C773-C780. Jones, P. C., Sivaprasadarao, A., Wray, D., and Findlay, J. B. C. (1996). A method for determining transmembrane protein structure. Mol. Membr. Biol 13, 53-60. JCrgensen, P. L. (1992). Na +, K+-ATPase, structure and transport mechanism. In "New Comprehensive Biochemistry: Molecular Aspects of Transport Proteins" (J. J. H. H. M. de Pont, ed.), Vol. 21, pp. 1-25. Elsevier, Amsterdam. Jung, K., Jung, H., and Kaback, H. R. (1994). Dynamics of lactose permease of Escherichia coli determined by sitedirected fluorescence labeling. Biochemistry 33, 3980-3985. Jung, K., Jung, H., Wu, J., Priv6, G. G., and Kaback, H. R. (1993). Use of site-directed fluorescence labeling to study proximity relationships in the lactose permease of Escherichia coli. Biochemistry 32, 12273-12278. Junge, W., Lill, H., and Engelbrecht, S. (1997). ATP synthase: An electrochemical transducer with rotatory mechanics. Trends Biochem. Sci. 22, 420-423.

Kaback, H. R., and Wu, J. (1998). From membrane to molecule to the third amino acid from the left with a membrane transport protein. Quart. Rev. Biophys. (in press). Kaback, H. R., Voss, J., and Wu, J. (1997). Helix packing in polytopic membrane proteins: The lactose permease of Escherichia coli. Curr. Opin. Struct. Biol. 7, 537-542. Kagawa, Y., and Hamamoto, T. (1997). Intramolecular rotation in ATP synthase: Dynamic and crystallographic studies on thermophilic F1. Biochem. Biophys. Res. Commun. 240, 247-256. Kakuda, D. K., Finley, K. D., Dionne, V. E., and MacLeod, C. L. (1993). Two distinct gene products mediate y+ type cationic amino acid transport in Xenopus oocytes and show different tissue expression patterns. Transgene 1, 91-101. Kallen, K.-J., Allan, D., Whatmore, J., and Quinn, P. (1994). Synthesis of surface sphingomyelin in the plasma membrane recycling pathway of BHK cells. Biochim. Biophys. Acta 1191, 52-58. Kamp, F., Zakim, D., Zhang, F., Noy, N. and Hamilton, J. A. (1995). Fatty acid flip-flop in phospholipid bilayers is extremely fast. Biochemistry 34, 11928-11937. Kanai, Y. (1997). Family of neutral and acidic amino acid transporters: Molecular biology, physiology and medical implications. Curr. Opin. Cell Biol. 9, 565-572. Kanai, Y., and Hediger, M. A. (1992). Primary structure and functional characterization of a high-affinity glutamate transporter. Nature 360, 467-471. Kanai, Y., Smith, C. P., and Hediger, M. A. (1994). A new family of neurotransmitter transporters: The high-affinity glutamate transporters FASEB J. 8, 1450-1459. Kanai, Y., Segawa, H., Miyamoto, K.-I., Uchino, H., Takeda, E., and Endou, H. (1998). Expression cloning and characterization of a transporter for large neutral amino acids activated by the heavy chain of 4F2 antigen (CD98). J. Biol. Chem. 273, 23629-23632. Kanai, T., Nussberger, S., Romero, M. F., Boron, W. F., Herbert, S. C., and Hediger, M. A. (1995). Electrogenic properties of the epithelial and neuronal high affinity transporter. J. Biol. Chem. 270, 16561-16568. Kanehisa, M. (1982). Los Alamos sequence analysis package for nucelic acids and proteins. Nucleic Acids Res. 10, 183-196. Kandasamy, R. A., Yu, F. H., Harris, R., Boucher, A., Hanrahan, J. W., and Orlowski, J. (1995). Plasma membrane Na+/ H + exchanger isoforms (NHE-1, -2, and -3) are differentially responsive to second messenger agonists of the protein kinase A and C pathways. J. Biol. Chem. 270, 2920929216. Kanner, B. I. (1996). Structure/function relationships in glutamate transporters. Biochem. Soc. Trans. 24, 843-846. Kanner, B. I., and Bendahan, A. (1982). Binding order of substrates to the sodium and potassium ion coupled L-glutamic acid transporter from rat brain. Biochemistry 21, 6327-6330. Karbach, D., Staub, M., Wood, P. G., and Passow, H. (1998). Effect of site-directed mutagenesis of the arginine residues 509 and 748 on mouse band 3 protein-mediated anion transport. Biochim. Biophys. Acta 1371, 114-122.

References Karlin, S., and Altschul, S. F. (1990). Methods for assessing the statistical significance of molecular sequence features by using general scoring schemes. Proc. Natl. Acad. Sci. USA 87, 2264-2268. Kavanaugh, M. P. (1993). Voltage dependence of facilitated arginine flux mediated by the system y+ basic amino acid transporter. Biochemistry 32, 5781-5785. Kavanaugh, M. P., Wang, H., Zhang, Z., Zhang, W., Wu, Y. N., Dechant, E., North, R. A., and Kabat, D. (1994). Control of cationic amino acid transport and retroviral receptor functions in a membrane protein family. J. BioL Chem. 269, 15445-15450. Kavanaugh, M. P., Bendahan, A., Zerangue, N., Zhang, Y., and Kanner, B. I. (1997). Mutation of an amino acid residue influencing potassium coupling in the glutamate transporter GLT-1 induces obligate exchange. J. Biol. Chem. 272, 1703-1708. Kawaguchi, Y., Okamoto, T., Taniwaki, M., Aizawa, M., Inoue, M., Katayama, S., Kawakami, H., Nakamura, S., Nishimura, M., Akiguchi, I., Kimura, J., Narumiya, S., and Kakizuka, A. (1994). CAG expansions in a novel gene for Machado-Joseph disease at chromosome 14q32.1. Nat. Genet. 8, 221-228. Kay, M. M. B., Cover, C., Schluter, S. F., Bernstein, R. M., and Marchalonis, J. J. (1995). Band 3, the anion transporter, is conserved during evolution: Implications for aging and vertebrate evolution. Cell Mol. Biol. 41, 833-842. Kekuda, R., Prasad, P. D., Fei, Y.-J., Torres-Zamorano, V., Sinha, S., Yang, T. L., Leibach, F. H., and Ganapathy, V. (1996). Cloning of the sodium-dependent, broad-scope, neutral amino acid transporter B ~ from a human placental choriocarcinoma cell line. J. Biol. Chem. 271,18657-18661. Kekuda, R., Torres-Zamorano, V., Fei, Y.-J., Prasad, P. D., Li, H. W., Mader, L. D., Leibach, F. H., and Ganapathy, V. (1997). Molecular and functional characterization of intestinal Na+-dependent neutral amino acid transporter B ~ Am. J. Physiol. 272, G1463-G1472. Kemp, H. F., and Taylor, P. M. (1997). Interactions between thyroid hormone and tryptophan transport in rat liver are modulated by thyroid status. Am. J. Physiol. 272, E809E816. Kenna, S., Roper, J., Ho, K., Hebert, S., Ashcroft, S. J., and Ashcroft, F. M. (1994). Differential expression of the inwardly-rectifying K-channel ROMK1 in rat brain. Mol. Brain Res. 24, 353-356. Kent, G., Iles, R., Bear, C. F., Huan, L. J., Griesenbach, U., McKerlie, C., Frndova, H., Ackerley, C., Gosselin, D., Radzioch, D., O'Brodovich, H., Tsui, L. C., Buchwald, M., and Tanswell, A. K. (1997). Lung disease in mice with cystic fibrosis. J. Clin. Invest. 100, 3060-3069. Keppler, D., Leier, I., and Ledlitschky, G. (1997). Transport of glutathione conjugates and glucuronides by the multidrug resistance proteins MRP1 and MRP2. J. BioL Chem. 278, 787-791. Keynes, R. D. (1994). The kinetics of voltage-gated ion channels. Quart. Rev. Biophys. 27, 339-434. Kifor, G., Toon, M. R., Janoshazi, A., and Solomon, A. K. (1993). Interaction between red cell membrane band 3 and cytosolic carbonic anhydrase. J. Memb r. B io I. 134,169-179.

363

Kilberg, M. S., Barber, E. F., and Handlogten, M. E. (1985). Characteristics and hormonal regulation of amino acid transport system A in isolated rat hepatocytes. Curr. Top. Cell Regul. 25, 133-163. Kilberg, M. S., Handlogten, M. E., and Christensen, H. N. (1980). Characteristics of an amino acid transport system in rat liver for glutamine, asparagine, histidine and closely related analogs. J. Biol. Chem. 255, 4011-4019. Kilberg, M. S., Handlogten, M. E., and Christensen, H. N. (1981). Characteristics of system ASC for transport of neutral amino acids in the isolated rat hepatocyte. J. BioL Chem. 256, 3304-3312. Kilberg, M. S., Hutson, R. G., and Laine, R. O. (1994). Amino acid-regulated gene expression in eukaryotic cells. F A S E B J. 8, 13-19. Kim, J. W., Closs, E. I., Albritton, L. M., and Cunningham, J. M. (1991). Transport of cationic amino acids by the mouse ecotropic retrovirus receptor. Nature 352, 725-728. Kim, H. R., Yew, N. S., Ansorge, W., Voss, H., Schwager, C., VennstrOm, B., Zenke, M., and Engel, J. D. (1988). Two different mRNAs are transcribed from a single genomic locus encoding the chicken erythrocyte anion transport proteins (band 3). Mol. Cell Biol. 8, 4416-4424. Kimball, S. R., Yancisin, M., Horetsky, R. L., and Jefferson, L. S. (1996). Translational and pretranslational regulation of protein synthesis by amino acid availability. Int. J. Biochem. Cell Biol. 28, 285-294. Kirsch, G. E., Pascual, J. M., and Shieh, C.-C. (1995). Functional role of a conserved aspartate in the external mouth of voltage-gated potassium channels. Biophys. J. 68, 18041813. Kirsch, J., and Betz, H. (1995). The postsynaptic localization of the glycine receptor-associated protein gephyrin is regulated by the cytoskeleton. J. Neurosci. 15, 4148-4156. Kitayama, S., Dohi, T., and Uhl, G. R. (1994). Phorbol esters alter functions of the expressed dopamine transporter. Eur. J. Pharmacol. 268, 115-119. Klapper, M. H.(1977). The independent distribution of amino acid near neighbor pairs into polypeptides. Biochem. Biophys. Res. Commun. 78, 1018-1024. Kleinfeld, A. M., Storms, S., and Watts, M. (1998). Transport of long-chain native fatty acids across human erythrocyte ghost membranes. Biochemistry 37, 8011-8019. Klingenberg, M. (1990). Mechanism and evolution of the uncoupling protein of brown adipose tissue. Trends Biochem. Sci. 15, 108-112. Klodos, I., Post, R. L., and Forbush, B., (1994). Kinetic heterogeneity of phosphoenzyme of Na,K-ATPase modeled by unmixed lipid phases. J. Biol. Chem. 269, 1734-1743. Knauf, P. A., Proverbio, F., and Hoffman, J. F. (1974). Electrophoretic separation of different phosphoproteins associated with Ca-ATPase and Na,K-ATPase in human red cell ghosts. J. Gen. Physiol. 63, 324-336. Knauf, P. A., Restrepo, D., Liu, S.-Q. J., Raha, N. M., Spinelli, L. J., Law, Y., Cronise, B., Snyder, R. B., and Romanow, L. (1992). Mechanisms of substrate binding, inhibitor binding, and ion translocation in band 3 and band 3-related proteins. Progr. Cell Res. 2, 35-44.

364

References

Knaus, H.-G., Folander, K., Garcia-Calvo, M., Garcia, M. L., Kaczorowski, G. J., Smith, M., and Swanson, R. (1994). Primary sequence and immunological characterization of /~-subunit of high conductance CaZ+-activated K + channel from smooth muscle. J. Biol. Chem. 269, 17274-17278. Kofuji, P., Davidson, N., and Lester, H. A. (1995). Evidence that neuronal G-protein-gated inwardly rectifying K + channels are activated by G beta gamma subunits and function as heteromultimers. Proc. Natl. Acad. Sci. USA 92, 65426546. Koide, R., Ikeuchi, T., Onodera, O., Tanaka, H., Igarashi, S., Endo, K., Takahashi, H., Kondo, R., Ishikawa, A., Hayashi, T., Saito, M., Tomoda, A., Miike, T., Naito, H., Ikuta, F., and Tsuji, S. (1994). Unstable expansion of CAG repeat in hereditary dentatorubral-pallidoluysian atrophy (DRPLA). Nat. Genet. 6, 9-13. Kong, C. T., Yet, S. F., and Lever, J. E. (1993). Cloning and expression of a mammalian Na+/amino acid cotransporter with sequence similarity to Na+/glucose cotransporters. J. Biol. Chem. 268, 1509-1512. Kopito, R. R., and Lodish, H. F. (1985). Primary structure and transmembrane orientation of the murine anion exchange protein. Nature 316, 234-238. Kopito, R. R., Lee, B. S., Simmons, D. S., Lindsey, A. E., Morgans, C. W., and Schneider, K. (1989). Regulation of intracellular pH by a neuronal homolog of the erythrocyte anion exchanger. Cell 59, 927-937. Kornberg, R. D., and McConnell H. M. (1971). Lateral diffusion of phospholipids in a vesicle membrane. Proc. Natl. Acad. Sci. USA 68, 2564-2568. Koser, B. H., and Christensen, H. N. (1971). Effect of substrate structure on coupling ratio for Na+-dependent transport of amino acids. Biochim. Biophys. Acta 241, 9-19. K6ster, A., Nagel, G., von Figura, K., and Pohlmann, R. (1991). Molecular cloning of the mouse 46-kDa mannose 6-phosphate receptor (MPR 46). Biol. Chem. Hoppe-Seyler 372, 297-300. Koumi, S., Sato, R., and Hayakawa, H. (1994). Modulation of voltage-dependent inactivation of the inwardly rectifying K + channel by chloramine-T. Eur. J. Pharmacol. 258, 281-284. Kowalski, T. J., Wu, G., and Watford, M. (1997). Rat adipose tissue amino acid metabolism in vivo is assessed by microdialysis and arteriovenous techniques. Am. J. Physiol. 273, E613-E622. Krapivinsky, G., Gordon, E. A., Wickman, K., Velimirovic, B., Krapivinsky, L., and Clapham, D.E. (1995). The Gprotein-gated atrial K + channel IKACh is a heteromultimer of two inwardly rectifying K+-channel proteins. Nature 374, 135-141. Krause, U., Rider, M. H., and Hue, L. (1996). Protein-kinase signalling pathway triggered by cell swelling and involved in the activation of glycogen-synthase and acetyl-CoA carboxylase in isolated rat hepatocytes. J. Biol. Cheml 271, 16668-16673. Kristensen, P., Eriksen, J., Blasi, F., and Dane, K. (1991). Two alternatively spliced mouse urokinase receptor mRNAs with different histological localization in the gastrointestinal tract. J. Cell BioL 115, 1763-1771.

Krolewski, J. J., and Dalla-Favera, R. (1991). The Itk gene encodes a novel receptor-type protein tyrosine kinase. E M B O J. 10, 2911-2919. Kubo, Y., Baldwin, T. J., Jan, Y. N., and Jan, L. Y. (1993). Primary structure and functional expression of a mouse inward rectifier potassium channel. Nature 362, 127-133. Kudrycki, K. E., and Shull, G. E. (1989). Primary structure of the rat kidney band 3 anion exchange protein deduced from a cDNA. J. BioL Chem. 264, 8185-8192. Kudrycki, K. E., Newman, P. R., and Shull, G. E. (1990). cDNA cloning and tissue distribution of mRNAs for two proteins that are related to the band 3 C1-/HCO-3 exchanger. J. Biol. Chem. 265, 462-471. Kurahashi, T. (1989). Activation by odorants of cation-selective conductance in the olfactory receptor cell isolated form the newt. J. Physiol. 419, 177-192. Kurashima, K., Yu, F. H., Cabado, A. G., Szabo, E. Z., Grinstein, S., and Orlowski, J. (1997). Identification of sites required for down-regulation of Na+/H + exchanger NHE3 activity by cAMP-dependent protein kinase: Phosphorylation-dependent and-independent mechanisms. J. Biol. Chem. 272, 28672-28679. Kurowski, B., and Ludwig, B. (1987). The genes of the Paracoccus denitrificans bcl: Nucleotide sequence and homologies between bacterial and mitochondrial subunits. J. Biol. Chem. 262, 13805-13811. Kusari, J., Verma, U. S., Buse, J. B., Henry, R. R., and Olefsky, J. M. (1991). Analysis of the gene sequences of the insulin receptor and the insulin-sensitive glucose transporter (GLUT-4) in patients with common-type non-insulindependent diabetes mellitus. J. Clin. Invest 88, 1323-1330. Kusumi, A., Sako, Y., and Yamamoto, M. (1993). Confined lateral diffusion of membrane receptors as studied by single particle tracking (nanovid microscopy): Effects of calciuminduced differentiation in cultured epithelial cells. Biophys. J. 65, 2021-2040. Kutchai, H., Chandler, L. H., and Geddis, L. M. (1980). Effects of anesthetic alcohols on membrane transport processes in human erythrocytes. Biochim. Biophys. Acta 600, 870-881. Kvamme, E., Torgner, I. A., and Roberg, B. (1991). Evidence indicating that pig renal phosphate-activated glutaminase has a functionally predominant external localization in the inner mitochondrial membrane. J. BioL Chem. 266,1218512192. Kwon H. M., and Handler J. S. (1995). Cell volume regulated transporters of compatible osmolytes. Curr. Opin. Cell Biol. 7, 465-471. Kwon, H. M., Itoh, T., Rim, J. S., and Handler, J. S. (1995). The MAP kinase cascade is not essential for transcriptional stimulation of osmolyte transporter genes. Biochem. Biophys. Res. Commun. 213, 975-979. Lai, S., Watson, J. C., Hansen, J. N., and Sze, H. (1991). Molecular cloning and sequencing of cDNAs encoding the proteolipid subunit of the vacuolar H+-ATPase from a higher plant. J. Biol. Chem. 266, 16078-16084. Lam, J. T., Martin, M. G., Turk, E., Steinmann, B., and Wright, E. M. (1998). Mutations of G318R and R468V in SGLT1 cause glucose-galatose malabsorption by trafficking defects. FASEB J. 12, A1017 [Abstract].

References Lang, F. (1993). Introduction to the molecules of transport: ion channels. Cell Physiol. Biochem. 3, 249. Lang, F., and Waldegger, S. (1997). Regulating cell volume. Am. Sci. 85, 456-463. Larsson, H. P., Baker, O. S., Dhillon, D. S. and Isacoff, E. Y. (1996). Transmembrane movement of the Shaker K + channel $4. Neuron 16,387-397. LaSpada, A. R., Wilson, E. M., Lubahn, D. B., Harding, A. E., and Fischbeck, K. H. (1991). Androgen receptor gene mutations in X-linked spinal and bulbar muscular atrophy. Nature (London) 352, 77-79. Latorre, R., and Miller, C. (1983). Conduction and selectivity in potassium channels. J. Membr. Biol. 71, 11-30. Lauffer, M. A. (1983). The significance of entropy-driven processes in biological systems. Comments Mol. Cell Biophys. 2, 99-109. L~iuger, P. (1980). Kinetic properties of ion carriers and channels. J. Membr. Biol. 57, 163-178. Laurent, B. C., Treitel, M. A., and Carlson, M. (1990). The SNF5 protein of Saccharomyces cerevisiae is a glutamineand proline-rich transcriptional activator that affects expression of a broad spectrum of genes. Mol. Cell Biol. 10, 5616-5625. Le Gall, A. H., Yeaman, C., Muesch, A., and RodriguezBoulan, E. (1995). Epithelial cell polarity: new perspectives. Semin. Nephrol. 15, 272-284. Lehmann-Horn, F., and Rtidel, R. (1997). Channelopathies: Their contribution to our knowledge about voltage-gated ion channels. News Physiolog. Sci. 12, 105-112. Lehninger, A. L., Nelson, D. L., and Cox, M. M. (1993). "Principles of Biochemistry," 2nd ed. Worth, New York. Leid, M., Kastner, P., Lyons, R., Nakshatri, H., Saunders, M., Zacharewski, T., Chen, J. Y., Staub, A., Garnier, J.-M., Mader, S., and Chambon, P. (1992). Purification, cloning and RXR identity of the HeLa cell factor with which RAR or TR heterodimerizes to bind target sequences efficiently. Cell 68, 377-395. Lesage, F., Duprat, F., Fink, M., Guillemare, E., Coppola, T., Lazdunski, M. and Hugnot, J. P. (1994). Cloning provides evidence for a family of inward rectifier and G-protein coupled K + channels in the brain. FEBS Lett. 353, 37-42. Lester, H. A., and Dougherty, D. A. (1998). New views of multi-ion channels. J. Gen. Physiol. 111, 181-183. Levi, M., Lotscher, M., Sorribas, V., Custer, M., Arar, M., Kaissling, B., Murer, H., and Biber, J. (1994). Cellular mechanisms of acute and chronic adaptation of rat renal P(i) transporter to alterations in dietary P(i). Am. J. Physiol. 267, F900-F980. Levine, S., Montrose, M., Tse, C. M., and Donowitz, M. (1993). Kinetic and regulation of three cloned mammalian Na+/ H § exchangers stably expressed in a fibroblast cell line. J. Biol. Chem. 268, 25527-25535. Levinson, G., and Gutman, G. A. (1987). Slipped-strand mispairing: A major mechanism for DNA sequence evolution. Mol. Biol. Evol. 4, 203-221. Lewis S. M., and Ueda, T. (1998). Solubilization and reconstitution of synpatic vesicle glutamate transport system. In "Methods in Enzymology" (S. G. Amara, eds.), Vol. 296, pp. 125-144. Academic Press, San Diego.

365

Liao, D.-I., Kapadia, G., Reddy, P., Saier, M. H., Reizer, J. and Herzberg, O. (1991). Structure of the IIA domain of the glucose permease of Bacillus subtilis at 2.2-A resolution. Biochemistry 30, 9583-9594. Lin, B., Nasir, J., MacDonald, H., Hutchinson, G., Graham, R. K., Rommens, J. M. and Hayden, M. R. (1994). Sequence of the murine Huntington disease gene: Evidence for conservation, alternate splicing and polymorphism in a triplet (CCG) repeat. Hum. Mol. Genet. 3, 85-92. Lin, F., Lester, H. A., and Mager, S. (1996). Single-channel currents produced by the serotonin transporter analysis of a mutation affecting ion permeation. Biophys J. 71, 31263135. Lindau, M. and Fernandez, J. M. (1986). A patch-clamp study of histamine-secreting cells. J. Gen. Physiol. 88, 349-368. Lindsey, A. E., Schneider, K., Simmons, D. M., Baron, R., Lee, B. S., and Kopito, R. R. (1990). Functional expression and subcellular localization of an anion exchanger cloned from choroid plexus. Proc. Natl. Acad. Sci. 87, 5278-5282. Linnertz, H., Urbanova, P., Obsil, T., Herman, P., Amler, E., and Schoner, W. (1998). Molecular distance measurements reveal an (c~/~)2 dimeric structure of Na+/K+-ATPase. J. Biol. Chem. 273, 28813-28821. Lister, N., Sykes, A. P., Bailey, P. D., Boyd, C. A. R., and Bronk, J. R. (1995). Dipeptide transport and hydrolysis in isolated loops of rat small intestine: Effects of stereospecificity. J. Physiol. 484, 173-182. Liu, D., Kennedy, S. D., and Knauf, P. A. (1995). 35C1nuclear magnetic resonance line broadening shows that eosin-5maleimide does not block the external anion access channel of band 3. Biophys. J. 69, 399-408. Liu, D., Kennedy, S. D., and Knauf, P. A. (1996). Source of transport site asymmetry in the band 3 anion exchange protein determined by NMR measurements of external C1affinity. Biochemistry 35, 15228-15235. Liu, M. L., Gibbs, E. M., McCoid, S. C., Milici, A. J., Stukenbrok, H. A., McPherson, R.K., Treadway, J.L., and Pessin, J.E. (1993) Transgenic mice expressing the human GLUT4/ muscle-fat facilitative glucose transporter protein exhibit efficient glycemic control. Proc. Natl. Acad. Sci. USA 90, 11346-11350. Loike, J. D., Hickman, S., Kuang, K., Xu, M., Cao, L., Vera, J. C., Silverstein, S. C. and Fischbarg, J. (1996). Sodiumglucose cotransporters display sodium- and phlorizindependent water permeability. Am.J. Physiol. 271, C1774C1779. Lolkema, J. S., Carrasco, N. and Kaback, H. R. (1991). Kinetic analysis of lactose exchange in proteoliposomes reconstituted with purified lacpPermease. Biochemistry 30, 12841290. Longo, N., Franchi-Gazzola, R., Bussolati, O., Dall'Asta, V., Nucci, F. A., and Gazzola, G. C. (1988). Amino acid and sugar transport in mouse 3T3 cells expressing activated ras and neu oncogenes. Ann. N.Y. Acad. Sci. 551, 374-377. Loo, D. D. F., Hazama, A., Supplisson, S., Turk, E., and Wright, E. M. (1993). Relaxation kinetics of the Na+/glu cose cotransporter. Proc. Natl. Acad. Sci. USA 90, 57675771.

366

References

Loo, D. D. F., Hirsch, J. R., Sarkar, H. K., and Wright, E. M. (1996a). Regulation of the mouse retinal taurine transporter (TAUT) by protein kinases in Xenopus oocytes. FEBS Lett. 392, 250-254. Loo, D. D. F., Zeuthen, T., Chandy, G., and Wright, E. M. (1996b). Cotransport of water by the Na+/glucose cotransporter. Proc. Natl. Acad. Sci. USA 93, 13367-13370. Lopatin, A. N., and Nichols, C. G. (1994). Internal Na + and Mg2+ blockade of DRK1 (Kv2.1) potassium channels expressed in Xenopus oocytes: Inward rectification of a delayed rectifier. J. Gen. Physiol. 103, 203-216. Lopatin, A. N., Makhina, E. N., and Nichols, C. G. (1995). The mechanism of inward rectification of potassium channels. J. Gen. Physiol. 106, 923-955. L6pez-Burillo, S., Garci~i-Sancho, J., and Herreros, B. (1985). Tryptophan transport through transport system T in the human erythrocyte, the Ehrlich cell and the rat intestine. Biochim. Biophys. Acta 820, 85-94. Lopez-Barneo, J., Hoshi, T., Heinemann, S.H., and Aldrich, R. W. (1993). Effects of external cations and mutations in the pore region on C-type inactivation of Shaker potassium channels. Recept. Channels 1, 66-71. Louvel, J., Avoli, M., Kurcewicz, I., and Pumain, R. (1994). Extracellular free potassium during synchronous activity induced by 4-aminopyridine in the juvenile rat hippocampus. Neurosci. Lett. 167, 97-100. Low, S. H., Chapin, S. J., Wimmer, C., Whiteheart, S. W., K~mtives, L. G., Mostov, K. E., and Weimbs, T. (1998). The SNARE machinery is inovlved in apical plasma membrane trafficking in MDCK cells. J. Cell Biol. 141, 1503-1513. Low, S. Y., Rennie, M. J., and Taylor, P. M. (1994). Na +dependent glutamate transport in cultured rat myotubes increases after glutamine deprivation. F A S E B J. 8, 127-131. Low, S. Y., Rennie, M. J., and Taylor, P. M. (1997a). Signalling elements involved in amino acid transport responses to altered muscle cell volume. F A S E B J. 11, 1111-1117. Low, S. Y., Rennie, M. J., and Taylor, P. M. (1997b). Involvement of integrins and the cytoskeleton in modulation of skeletal muscle glycogen synthesis by changes in cell volume. FEBS Lett. 417, 101-103. Low, S. Y., Salter, M., Knowles, R. G., Pogson, C. I., and Rennie, M. J. (1993). A quantitative analysis of the control of glutamine catabolism in rat liver cells: Use of selective inhibitors, Biochem. J. 295, 617-624. Low, S. Y., Taylor, P. M., Hundal, H. S., Pogson, C. I., and Rennie, M. J. (1992). Transport of L-glutamine and L-glutamate across sinusoidal membranes of rat liver: Effect of starvation, diabetes and corticosteroid treatment. Biochem. J. 284, 333-340. Lti, Q., and Miller, C. (1995). Silver as a probe of pore-forming residues in a potassium channel. Science 268, 304-307. Lucchesi, P. A., Cooney, R. A. Mangsen-Baker, C., Honeyman, T. W., and Scheid, C. R. (1988). Assessment of transport capacity of plasmalemmal Ca 2+ pump in smooth muscle. Am. J. Physiol. 255, C226-C236. Lukacs, G., Rotstein, O. D., and Grinstein, S. (1995). An overview of intracellular pH regulation: Role of vacuolar

H+-ATPases. In "Molecular Biology Intelligence Unitm Organellar Proton-ATPases" (N. Nelson, ed.), pp. 29-48. R. G. Landes Company, Austin, TX. Lutsenko, S., Anderko, R., and Kaplan, J. H. (1995). Membrane disposition of the M5-M6 hairpin of Na+,K+-ATPase alpha subunit is ligand dependent. Proc. Natl. Acad. Sci. USA 92, 7936-7940. Lux, S. E., John, K. M., Kopito, R. R., and Lodish, H. J. (1989). Cloning and characterization of band 3, the human erythrocyte anion exchange protein (ALl). Proc. Natl. Acad. Sci. 86, 9089-9093. Luxon, B. A. (1996). Inhibition of binding to fatty acid binding protein reduces the intracellular transport of fatty acids. Am. J. Physiol. 271, Gl13-G120. Maathuis, F. J. M., Ichida, A. M., Sanders, D., and Schroeder, J. I. (1997). Roles of higher plant K + channels. Plant Physiol. 114, 1141-1149. Mackenzie, B., Ahmed, A., and Rennie, M. J. (1992), Muscle amino acid metabolism and transport. In "Mammalian Amino Acid Transport: Mechanisms and Control" (M. S. Kilberg, and D. H~iussinger, eds.), pp. 195-231. Plenum Press, New York/London. Mackenzie, B., Harper, A. A., Taylor, P. M., and Rennie, M. J. (1994a). Na+/amino acid coupling stoichiometry of rheogenic System B ~ transport in Xenopus oocytes is variable. Pfliigers Arch. 426, 121-128. Mackenzie, B., Loo, D. D. F., Fei, Y.-J., Liu, W., Ganapathy, V., Leibach, F. H., and Wright, E. M. (1996a). Mechanisms of the human intestinal H+-coupled oligopeptide transporter hPEPT1. J. BioL Chem. 271, 5430-5437. Mackenzie, B., Loo, D. D. F., Panayotova-Heiermann, M., and Wright, E. M. (1996b). Biophysical characteristics of the pig kidney Na+/glucose cotransporter SGLT2 reveal a common mechanism for SGLT1 and SGLT2. J. BioL Chem. 217, 32678-32683. Mackenzie B., Loo, D. D. F., and Wright, E. M. (1998). Relationships between Na+/glucose cotransporter currents and fluxes. J. Memb. Biol. 162, 101-106. Mackenzie, B., Panayotova-Heiermann, M., Loo, D. D. F., Lever, J. E., and Wright, E. M. (1994b). SAAT1 is a low affinity Na+/glucose cotransporter and not an amino acid transporter. J. BioL Chem. 269, 22488-22491. MacKinnon, R. (1995). Pore loops: An emerging theme in ion channel structure. Neuron 14, 889-892. MacKinnon, R., Aldrich, R. W., and Lee, A. W. (1993). Functional stoichiometry of Shaker potassium channel inactivation. Science 262, 757-759. MacKrell, A. J., Blumberg, B., Haynes, S. R., and Fessler, J. H. (1988). The lethal myospheroid gene of Drosophila encodes a membrane protein homologous to vertebrate integrin/3 subunits. Proc. Natl. Acad. Sci. USA 85, 26332637. MacLennan, D. H., Rice, W. J., and Green, N. M. (1997). The mechanism of Ca 2+ transport by sarco(endo)plasmic reticulum Ca2+-ATPases. J. Biol. Chem. 272, 28815-28818. MacLeod, C. L. (1996). Regulation of cationic amino acid transporter (CAT) gene expression. Biochem. Soc. Trans. 24~ 846-852.

References MacLeod, C. L., and Kakuda, D. K. (1996). Regulation of CAT: Cationic amino acid transporter gene expression. Amino Acids 11, 171-191. MacLeod, C. L., Finley, K. D., and Kakuda, D. K. (1994). y+Type cationic amino acid transport: Expression and regulation of the m C A T genes. J. Exp. Biol. 196, 109-121. MacLeod, C. L., Finley, K., Kakuda, D., Kozak, C. A., and Wilkinson, M. F. (1990). Activated T cells express a novel gene on chromosome 8 that is closely related to the murine ecotropic retroviral receptor. Mol. Cell Biol. 10, 36633674. Maeda, M. (1994). Gastric proton pump (H+/K+-ATPase): Structure and gene regulation through GATA DNAbinding protein(s). J. Biochem. 115, 6-14. Mager, S., Min, C., Henry, D. J., Chavkin, C., Hoffman, B. J., Davidson, N., and Lester, H. A. (1994). Conducting states of a mammalian serotonin transporter. Neuron 12, 845-859. Mager, S., Naeve, J., Quick, M., Labarca, C., Davidson, N., and Lester, H. A. (1993). Steady states, charge movements, and rates for a coined G A B A transporter expressed in Xenopus oocytes. Neuron 10, 177-188. Majeau, N., and Coleman, J. R. (1991). Isolation and characterization of a cDNA coding for pea chloroplastic carbonic anhydrase. Plant Physiol. 95, 264-268. Makowske, M., and Christensen, H. N. (1982). Hepatic transport system interconverted by protonation from service for neutral to service for anionic amino acids. J. Biol. Chem. 257, 14635-14638. Malandro, M. S., and Kilberg, M. S. (1996). Molecular biology of mammalian amino acid transporters. Ann. Rev. Biochem. 65, 305-336. Malandro, M. S., Beveridge, M. J., Kilberg, M. S., and Novak, D. A. (1996). Effect of low-protein diet-induced intrauterine growth retardation on rat placental amino acid transport. Am. J. Physiol. 271, C295-C303. Maldonado, A. M., and Portillo, F. (1995). Genetic analysis of the fluorescein isothiocyanate binding site of the yeast plasma membrane H+ATPase. J. Biol. Chem. 270, 86558659. Maly, K., Uberall, F., Loferer, H., Doppler, W., Oberhuber, H., Groner, B., and Grunicke, H. (1989). Ha-ras activates the Na+/H § antiporter by a protein kinase C-independent mechanism. J. BioL Chem. 264, 11839-11842. Man, M. Z., Hui, T. Y., Schaffer, J. E., Lodish, H. F., and Bernlohr, D. A. (1996). Regulation of the murine adipocyte fatty acid transporter gene by insulin. Mol. Endocrinol. 10, 1021-1028. Manning, G. S. (1975). The relation between osmotic flow and tracer solvent diffusion for single-file transport. Biophys. Chem. 3, 147-152. Mantsch, H. H., and McElhaney, R. N. (1991). Phospholipid phase transitions in model and biological membranes as studied by infrared spectroscopy. Chem. Phys. Lipids 57, 213-226. Marban, E., and Tomaselli G. F. (1997). Ion channels as enzymes: Analogy or homology? Trends in Neurosci. 20, 144-147.

367

Marsh, D. (1995). Lipid-protein interactions and heterogenous lipid distribution in membranes. Mol. Membr. Biol. 12, 59-64. Marshall, B. A., Ren, J. M., Johnson, D. W., Gibbs, E. M., Lillquist, J. S., Soeller, W. C., Holloszy, J. O., and Mueckler, M. (1993). Germline manipulation of glucose homeostasis via alteration of glucose transporter level in skeletal muscle. J. Biol. Chem. 268, 18442-18445. Martin, M. G., Turk, E., Lostao, M. P., Kerner, C., and Wright, E. M. (1996). Defects in Na+glucose cotransporter (SGLT1) trafficking and function cause glucose-galactose malabsorption. Nat. Genet. 12, 216-220. Martonosi, A. N. (1996). Structure-function relationships in the CaZ+-ATPase of sarcoplasmic reticulum: Facts, speculations and questions for the future. Biochim. Biophys. Acta 1275, 111-117. Maruyama, K., and MacLennan, D. H. (1988). Mutation of aspartic acid-351, lysine-352 and lysine-515 alters the Ca + transport activity of the Ca+-ATPase expressed in COS-1 cells. Proc. Natl. Acad. Sci. USA 85, 3314-3318. Mastroberardino, L., Spindler, B., Pfeiffer, R., Skelly, P. J., Loffings, J., Shoemaker, C. B., and Verrey, F. (1998). Amino-acid transport by heterodimers of 4F2hc/CD98 and members of a permeable family. Nature 395, 288-291. Matsuda, C., Endo, H., Hirata, H., Morosawa, H., Nakanishi, M., and Kagawa, Y. (1993b). Tissue-specific isoforms of the bovine mitochondrial ATP synthase y-subunit. FEBS Lett. 325, 281-284. Matsuda, C., Endo, H., Ohta, S. and Kagawa, Y. (1993a). Gene structure of human mitochondrial ATP synthase ysubunit. J. Biol. Chem. 268, 24950-24958. Matthews, D. E., and Campbell, R. G. (1992). The effect of dietary protein intake on glutamine and glutamate nitrogen metabolism in humans. Am. J. Clin. Nutr. 55, 963-970. Matthews, J. C., Aslanian, A. M., McDonald, K. K. Yang, W., Malandro, M. S., Novak, D. A., and Kilberg, M. S. (1997). An expression system for mammalian amino acid transporters using a stably maintained episomal vector. Anal. Biochem. 254, 208-214. McClintock, B. (1984). The significance of responses to the genome challenge. Science 226: 792-801. McCormack, K., McCormack, T., Tanouye, M., Rudy, B., and Stuhmer, W. (1995). Alternative splicing of the human Shaker K § channel fll gene and functional expression of the B2 gene product. FEBS Lett. 370, 32-36. McCormick, J. I., and Johnstone, R. M. (1995). Identification of the integrin a3fla as a component of a partially purified A system amino acid transporter from Ehrlich cell plasma membrane. Biochem. J. 311, 743-751. McCormick, J. I., Jette, M., Potier, M., Beliveau, R., and Johnstone, R. M. (1991). Molecular size of a Na+-dependent amino acid transporter in Ehrlich ascites cell plasma membrane estimated by radiation inactivation analysis. Biochemistry 30, 3704-3709. McEver, R. P. (1992). Leukocyte-endothelial cell interactions. Curr. Opin. Cell Biol. 4, 840-849. McGivan, J. D. (1992). Techniques used in the study of plasma membrane amino acid transport. In "Mammalian Amino

368

References

Acid Transport: Mechanisms and Control" (M. S. Kilberg and D. H~iussinger, eds.), pp. 51-63. Plenum Press, New York. McGivan, J. D. (1998). Rat hepatoma cells express novel transport systems for glutamine and glutamate in addition to those present in normal rat hepatocytes. Biochem. J. 330, 255-260. McGivan, J.D., and Pastor-Anglada, M. (1994). Regulatory and molecular aspects of mammalian amino acid transport. Biochem. J. 299, 321-334. McNicholas, C. M., Guggino, W. B., Schwiebert, E. M., Hebert, S. C., Giebisch, G., and Egan, M. E. (1996). Sensitivity of renal K § channel (ROMK2) to the inhibitory sulfonylurea compound, glibenclamide, is enhanced by coexpression with the ATP-binding cassette transporter cystic fibrosis transmembrane regulator. Proc. Natl. Acad. Sci. 93, 8083-8088. Meera, P., Wallner, M., Jiang, Z., and Toro, L. (1996). A calcium switch for the functional coupling between a (hslo) and 13 subunits (Kv, Ca~) of maxi K channels. FEBS Lett. 385, 127-128. Meijer, A. J., Lamers, W. H., and Charmaleau, R. A. F. M. (1990). Nitrogen metabolism and ornithine cycle function. Physiol. Rev. 70, 701-748. Meijer, G. A., van-der-Meulen, J., and van-Vuuren, A.M. (1993). Glutamine is a potentially limiting amino acid for milk production in dairy cows: A hypothesis. Metabolism 42, 358-364. Meinild, A.-K., Klaerke, D. A., Loo, D. D. F., Wright, E. M., and Zeuthen, T. (1998). The human Na+-glucose cotransporter is a molecular water pump. J. Physiol. 508, 15-21. Meister, A. (1988). Glutathione metabolism and its selective modification. J. BioL Chem. 263, 17205-17208. Mellanen, P., Minn, H., Grenman, R., and Harkonen, P. (1994). Expression of glucose transporters in head-andneck tumors. Int. J. Cancer 56, 622-629. Mercer, R. W., Biemesderfer, D., Bliss, D. P., Jr.,. Collins, J. H., and Forbush, B. (1993). Molecular cloning and immunological characterization of the y polypeptide, a small protein associated with the Na, K-ATPase. J. Cell Biol. 121, 579-586. Meyer, M., Maly, K., Uberall, F., Hoflacher, J., and Grunicke, H. (1991). Stimulation of K + transport systems by Ha-ras. J. BioL Chem. 266, 8230-8235. Meyer, T., Mtinch, C., Knappenberger, B., Liebau, S., V61kel, H., and Ludolph, A.C. (1998). Alternative splicing of the glutamate transporter EAAT2 (GLT-1). Neurosci. Letts. 241, 68-70. Michaelis, L., and Menten, M. L. (1913). Die kinetik der invertinwirkung. Biochem. Z. 49, 333-369. Miledi, R., Parker, I., and Woodward, R. M. (1989). Membrane currents elicited by divalent cations in Xenopus oocytes. J. Physiol. 417, 173-195. Milkman, R. (1994). An Escherichia coli homologue of eukaryotic potassium channel proteins. Proc. Natl. Acad. Sci. USA 91, 3510-3514. Millar, I. D., Calvert, D. T., Lomax, M. A., and Shennan, D. B. (1997). Substrate specificity of the mammary tissue

anionic amino acid carrier operating in the cotransport and exchange modes. Biochim. Biophys. Acta 1326, 92-102. Miller, C. (1995). The charybdotoxin family of K + channelblocking peptides. Neuron 15, 5-10. Miller, M. J., Oldenburg, M., and Fillingame, R. H. (1990). The essential carboxyl group in subunit c of the F1Fo ATP synthase can be moved and H§ function retained. Proc. Natl. Acad. Sci. USA 87, 4900-4904. Milner, H. E., Beliveau, R., and Jarvis, S. M. (1994). The in situ size of the dopamine transporter is a tetramer as estimated by radiation inactivation analysis. Biochim. Biophys. Acta 1190, 185-197. Millward, D. J., Fereday, A., Gibson, N. R., and Pacy, P. J. (1996). Post-prandial protein metabolism. Baillieres Clin. Endcrinol. Metab. 10, 533-549. Minn, H., Clavo, A. C., and Wahl, R. L. (1996). Influence of hypoxia on tracer accumulation in squamous-cell carcinoma: in vitro evaluation for PET imaging. Nucl. Med. Biol. 23, 941-946. Mitchell, P. (1961). Coupling of phosphorylation to electron and hydrogen transfer by a chemiosmotic type of mechanism. Nature 191, 144-148. Mitchell, P. (1962). Metabolism, transport and morphogenesis: Which drives which? J. Gen. Microbiol. 29, 25-37. Mitchell, P., and Moyle, J. (1959). Coupling of metabolism and transport by enzymic translocation of substrates through membranes. Proc. R. Soc. Edinb. [Phys.] 28, 19-27. Mitra, A., van Hoek, A. N., Wiener, M. C., Verkman, A. S., and Yaeger, M. (1995). The CHIP28 water channel visualized in ice by electron cryo-crystallography. Nat. Struct. Biol. 2, 726-729. Modyanov, N., Lutsenko, S., Chertova, E., Effemov, R., and Gulyaev, D. (1992). Transmembrane organization of the Na+K+-ATPase molecule. Acta Physiol. Scand. Suppl. 607, 49-58. Moe, A. J. (1995). Placental amino acid transport. Am. J. Physiol. 268, C1321-1331. Moffett, J., and Englesberg, E. (1984). Recessive constitutive mutant Chinese hamster ovary cells (CHO-K1) with an altered A system for amino acid transport and the mechanism of gene regulation of the A system. Mol. Cell Biol. 4, 799-808 Mohammed, T., Stulc, J., Glazier, J. D., Boyd, R. D. H., and Sibley, C. P. (1993). Mechanisms of potassium transfer across the dually perfused rat placenta. Am. J. Physiol. 265, R341-R347. MOiler, J. V., Juul, B., and leMaire, M. (1996). Structural organization, ion transport, and energy transduction of P-type ATPases. Biochim. Biophys. Acta 1286, 1-51. Morii, M., Hayata, Y., Mizoguchi, K., and Takeguchi, N. (1996). Oligomeric regulation of gastric H+,K+-ATPase. J. Biol. Chem. 271, 4068-4072. Mortimore, G. E., Wert, J. J., and Adams, C. E. (1988). Modulation of the amino acid control of hepatic protein degradation by caloric deprivation. J. Biol. Chem. 263, 1954519951. Mosher, M. E., White, L. K., Hermolin, J., and Fillingame, R. H. (1985). H+-ATPase of Escherichia coli: An uncE

References mutation impairing coupling between F1 but not Fo-mediated H+translocation. J. Biol. Chem. 260, 4807-4814. Mostov, K. E., Apodaca, G., Aroeti, B., and Okamoto, C. (1992). Plasma membrane protein sorting in polarized epithelial cells. J. Cell Biol. 116, 577-583. Motais, R., Fi6vet, B., Borgese, F., and Garcia-Romeu, F. (1997). Association of the band 3 protein with a volumeactivated, anion and amino acid channel: A molecular approach. J. Exp. Biol. 200, 361-367. Moule, S. K., and McGivan, J. D. (1987). Epidermal growth factor, like glucagon, exerts a short-term stimulation of alanine transport in rat hepatocytes. Biochem. J. 247, 233-235. Moule, S. K., and McGivan, J. D. (1990). Regulation of the plasma membrane potential in hepatocytes. Mechanism and physiological significance. Biochim. Biophys. Acta 1031, 383-387. Moule, S. K., Bradford, N. M., and McGivan, J. D. (1987). Short-term stimulation of Na +- dependent amino acid transport by dibutyryl cyclic AMP in hepatocytes. Characteristics and partial mechanism. Biochem. J. 241, 737-743. Mu, X., and Welbourne, T. C. (1996). Response of LLC-PK1F + cells to metabolic acidosis. Am. J. Physiol. 270, C920C928. Mtiller, M., and Jansen, P. L. M. (1997). Molecular aspects of hepatobiliary transport proteins. Am. J. Physiol. 272, G1304-G1313. Mtiller-Berger, S., Karbach, D., K~Snig, J., Lepke, S., Wood, P. G., Appelhans, H., and Passow, H. (1995a). Inhibition of mouse erythroid band 3-mediated chloride transport by site-directed mutagenesis of histidine residues and its reversal by second site mutation of Lys 558, the locus of covalent HzDIDS binding. Biochemistry 34, 9315-9324. Mtiller-Berger, S., Karbach, D., Kang, D., Aranibar, N., Wood, P. G., Rtiterjans, H., and Passow, H. (1995b). Roles of histidine 752 and glutamate 699 in the pH dependence of mouse band 3 protein-mediated anion transport. Biochemistry 34, 9325-9332. Mtiller-ROber, B., Ellenberg, J., Provart, N., Willmitzer, L., Busch, H., Becker, D., Dietrich, P., Hoth, S., and Hedrich, R. (1995). Cloning and electrophysiological analysis of KST1, an inward rectifying K + channel expressed in potato guard cells. E M B O J. 14, 2409-2416. Munck, B. G. (1985). Transport of neutral and cationic amino acids across the brush-border membrane of the rabbit ileum. J. Membr. Biol. 83, 1-13. Munck, L. K., and Munck, B. G. (1994). Amino acid transport in the small intestine. PhysioL Res. 43,335-345. Munck, L. K., and Munck, B. G. (1995). Transport of glycine and lysine on the chloride-dependent ~-alanine (B ~ carrier in rabbit small intestine. Biochim. Biophys. Acta 1235, 93-99. Munck, B. G., and Munck, L. K. (1997). Na+-independent transport of bipolar and cationic amino acids across the luminal membrane of the small intestine. Am. J. Physiol. 272, R1060-R1068. Muragaki, Y., Mundlos, S., Upton, J. and Olsen, B.R. (1996) Altered growth and branching patterns in synpolydactyly caused by mutations in HOXD13. Science 272, 548-551.

369

Murer, H., Gmaj, P., Steiger, B., and Hagenbuch, B. (1989). Transport studies with renal proximal tubular and small intestinal brush border and basolateral membrane vesicles: Vesicle heterogeneity, coexistence of transport systems. In "Methods in Enzymology" (S. Fleischer and B. Fleischer, eds.)., Vol. 172, pp. 346-364. Academic Press, San Diego. Murer, H., Lotscher, M., Kaissling, B., Levi, M., Kempson, S.A., and Biber, J. (1996). Renal brush border membrane Na/Pi-cotransport: Molecular aspects in PTH-dependent and dietary regulation. Kidney Int. 49, 1769-1773. Murphy, S., Welk, G., and Kardos, S. (1992). Epidermal growth factor induces glucose transport in primary cell cultures derived from human astrocytic glioma biopsies. Neurosci. Lett. 138, 275-278. Muth, T. R., Gottardi, C. J., Roush, D. L., and Caplan, M. J. (1998). A basolateral sorting signal is encoded in the c~-subunit of Na-K-ATPase. Am. J. Physiol. 274, C688C696. Myher, J. J., Kuskis, A., and Pind, S. (1989). Molecular species of glycophospholipids and sphingomyleins of human erythrocytes: Improved method of analysis. Lipids 24, 396-407. Myung, J., and Jencks, W. P. (1995). There is only one phosphoenzyme intermediate with bound calcium on the reaction pathway of the sarcoplasmic reticulum calcium ATPase. Biochemistry 34, 3077-3083. Naftalin, R. J. (1984). The thermostatics and thermodynamics of cotransport. Biochim. Biophys. Acta 778, 155-175. Nagafuchi, S., Yanagisawa, H., Sato, K., Shirayama, T., Ohsaki, E., Bundo, M., Takeda, T., Tadokoro, K., Kondo, I., Murayama, N., Tanaka, Y., Kikushima, H., Umino, K., Kurosawa, H., Furukawa, T., Nihei, K., Inoue, T., Sano, A., Komure, O., Takahashi, M., Yoshizawa, T., Kanazawa, I., and Yamada, M. (1994). Dentatorubral and pallidoluysian expansion of an unstable CAG trinucleotide on chromosome 12p. Nat. Genet. 6, 14-18. Nagamatsu S., Sawa H., Wakizaka A., and Hoshino T. (1993). Expression of glucose transporter isoforms in human brain tumors. J. Neurochem. 61, 2048-2053. Nakamoto, R. K. (1996). Mechanisms of active transport in the FoF1 ATP synthase. J. Membrane Biol. 151, 101-111. Nakanishi, M., Kagawa, Y., Narita, Y., and Hirata, H. (1994). Purification and reconstitution of an intestinal Na+-dependent neutral L-a-amino acid transporter. J. Biol. Chem. 369, 9325-9329. Nakayama, K., Kim, W.-S., Torii, S., Hosaka, M., Nakagawa, T., Ikemizu, J., Baba, T., and Murakami, K. (1992). Identification of the Fourth Member of the Mammalian endoprotease family homologous to the yeast Kex2 protease. J. Biol. Chem. 267, 5897-5900. Nakhoul, N. L., Chen, L. K., and Boron, W. F. (1990). Intracellular pH regulation in rabbit $3 proximal tubule: Basolateral C1-HCO3 exchange and Na-HCO3 cotransport. Am. J. PhysioL 358, F371-F381. Nakhoul, N. L., Abdulnour-Nakhoul, S., Hering-Smith, K. S., and Hamm, L. L. (1998). Intracellular ionic activities in Xenopus oocytes expressing the epithelial sodium channel. F A S E B J. 12, A372. [Abstract]. Nelson, C. A., Wang, O. J., Bourque, J. P., and Crane, P. D. (1996). Targeting of glucose transport proteins for tumor imaging: Is it feasible? J. NucL Med. 37, 1038-1041.

370

References

Nelson, N. (1992a). Organellar proton-ATPases. Curr. Opin. Cell Biol. 4, 654-660. Nelson, N. (1992b). Evolution of organellar proton-ATPases. Biochim. Biophys. Acta 1100, 109-124. Nelson, N. (1995). Molecular and cellular biology of F- and V-ATPases. In: "Molecular Biology Intelligence Unit m Organellar Proton-ATPases" (N. Nelson, ed.), pp. 1-28. R. G. Landes, Austin, TX. Nelson, W. J. (1994). Regulation of cell adhesion and development of epithelial cell surface polarity. In: "Cell Biology and Membrane Transport Processes: Current Topics in Membranes" (M. J. Caplan, ed.), Vol. 41, pp. 123-142. Academic Press, San Diego. Nelson, W. J., and Veshnock, P. J. (1987). Ankyrin binding to (Na + + K +) ATPase and implications for the organization of membrane domains in polarized cells. Nature 328, 533-536. Neu, J., Shenoy, V., and Chakrabarti, R. (1996). Glutamine nutrition and metabolism: Where do we go from here? FASEB J. 10, 829-837. Newsholme. E. A., and Parry-Billings, M. (1990). Properties of glutamine release from muscle and its importance for the immune system. J. Parent. Enter. Nutr. 14, 63S-68S. Nichols, C. G., and Lopatin, A. N. (1997). Inward rectifier potassium channels. Annu. Rev. Physiol. 59, 171-191. Nichols, C. G., Ho, K., and Hebert, S. (1994). Mg2+-dependent inward rectification of ROMK1 potassium channels expressed in Xenopus oocytes. J. Physiol. 476, 399-409. Nicholson, B., and McGivan, J. D. (1996). Induction of high affinity glutamate transport activity by amino acid deprivation in renal epithelial cells does not involve an increase in the amount of transporter protein. J. Biol. Chem. 271, 12159-12164. Nies, D. H., and Silver, S. (1995). Ion efflux systems involved in bacterial metal resistances. J. Industr. Microbiol. 14, 186-199. Nikaido, H., and Saier, M. H. (1992). Transport proteins in bacteria: Common themes in their design. Science 258, 936-942. Nishigori, H., Yamada, S., Tomura, H., Fernald, A. A., Le Beau, M. M., Takeuchi, T., and Takeda, H. (1998). Identification and characterization of the gene encoding a second proteolipid subunit of human vacuolar H+-ATPase (ATP6F). Genomics 50, 222-228. Nishioka, T., Oda, Y., Seino, Y., Yamamoto, T., Inagaki, N., Yano, H., Imura, H., Shigemoto, R., and Kikuchi, H. (1992). Distribution of the glucose transporters in human brain tumors. Cancer Res. 52, 3972-3979. Noji, H., Yasuda, R., Yoshida, M., and Kinosita, K. (1997). Direct observation of the rotation of F1-ATPase. Nature 386, 299-302. North, R. A. (1995). "Handbook of Receptors and Channels: Ligand- and Voltage-Gated Ion Channels." CRC Press, Boca Raton, FL. NCrregaard, A., Vilsen, B., and Andersen, J. P. (1994). Transmembrane segment M3 is essential to thapsigargin sensitivity of the sarcoplasmic reticulum Ca2+-ATPase. J. Biol. Chem. 269, 26598-26601.

Novak, D. A., and Beveridge, M. J. (1997). Glutamine transport in human and rat placenta. Placenta 18, 379-386. Nucci, F. A., Bussolati, O., Dall'Asta, V., Gazzola, G. C., Giardini, R., and Guidotti, G. G. (1988). Derangements of cationic amino acid transport in fibroblasts from human desmoid tumor. In "Chemical Carcinogenesis" (F. Feo, P. Pani, A. Columbano, and R. Garcea, eds.), pp. 467-473 Plenum, New York. Nfifiez, E., and Arag6n, C. (1994). Structural analysis and functional role of the carbohydrate component of glycine transporter. J. Biol. Chem. 269, 16920-16924. Nurjhan, N., Bucci, A., Perriello, G., Stumvoll, M., Dailey, G., Bier, D. M., Toft, I., Jenssen, T. G., and Gerich, J. E. (1995). Glutamine: A major gluconeogenic precursor and vehicle for interorgan carbon transport in man. J. Clin. Invest. 95, 272-277. Nussberger, S., Steel, A., Trotti, D., Romero, M. F., Boron, W. F., and Hediger, M. A. (1997). Symmetry of H + binding to the intra- and extracellular side of the H+-coupled oligopeptide cotransporter PepT1. J. Biol. Chem. 272, 77777785. Ogawa, J., Inoue, H., and Koide, S. (1997). Glucose-transporter-type-l-gene amplification correlates with sialylLewis-X synthesis and proliferation in lung cancer. Int. J. Cancer 74, 189-192. Ohneda, M., Johnson, J. H., Inman, L. R., Chen, L., Suzuki, K., Goto, Y., Alam, T., Ravazzola, M., Orci, L., and Unger, R. H. (1993). GLUT2 expression and function in/3-cells of GK rats with NIDDM: Dissociation between reductions in glucose transport and glucose-stimulated insulin secretion. Diabetes 42, 1065-1072. Ohta, S., Goto, K., Arai, H., and Kagawa, Y. (1987). An extremely acidic amino-terminal presence of the precursor for the human mitochondrial hinge protein. FEBS Lett. 226, 171-175. Okada, Y. (1997). Volume expansion-sensing outwardrectifier C1~channel: Fresh start to the molecular identity and volume sensor. Am. J. Physiol. 273, C755-C789. Oldendorf, W. H. (1971). Brain uptake of radiolabelled amino acids, amines and hexoses after arterial injection. Am. J. Physiol. 221, 1629-1639. Ono, T., and Means, A. R. (1989). Calspermin is a testis specific calmodulin-binding protein closely related to Ca 2+/calmOdulin-dependent protein kinases. Adv Exp. Med. Biol. 255, 263-268. Opie, L. H. (1991). "The Heart: Physiology and Metabolism," 2nd ed. Raven, New York. Orci, L., Ravazzola, M., Baetens, D., Inman, L., Amherdt, M., Peterson, R. G., Newgard, C. B., Johnson, J. H., and Unger, R. H. (1990). Evidence that down-regulation of/3-cell glucose transporters in non-insulin-dependent diabetes may be the cause of diabetic hyperglycemia. Proc. Natl. Acad. Sci. USA 87, 9953-9957. Orlowsky, J. (1993). Heterologous expression and functional properties of amiloride high affinity (NHE1) and low affinity (NHE3) isoforms of the rat Na+/H § exchanger. J. Biol. Chem. 268, 16369-16377. Orr, H. T., Chung, M.-Y., Banff, S., Kwiatkowski, T. J., Jr., Servadio, A., Beaudet, A. L., McCall, A. E., Duvick,

References L. A., Ranum, L. P. W., and Zoghbi, H. Y. (1993). Expansion of an unstable trinucleotide CAG repeat in spinocerebellar ataxia type 1. Nat. Genet. 4, 221-226. Ostermeier, C., and Michel, H. (1997). Crystallization of membrane proteins. Curr. Opin. Struct. Biol. 7, 697-701. Oulianova, N., and Berteloot, A. (1996). Sugar transport heterogeneity in the kidney: Two independent transporters or different transport modes through an oligomeric protein? 1. Glucose transport studies. J. Membr. Biol. 153, 181-194. Otsu, K., Kinsella, J., Heller, P., and Froehlich, J. P. (1993a). Sodium dependence of the Na+/H + exchanger in the presteady state: Implications for the exchanger mechanism. J. Biol. Chem. 268, 3184-3193. Otsu, K., Kinsella, J., Koh, E., and Froehlich, J. P. (1993b). Proton dependence of the partial reactions of the sodiumproton exchanger in renal brush border membranes. J. Biol. Chem. 267, 8089-8096. Otsu, K., Kinsella, J., Sacktor, B., and Froehlich, J. P. (1989) Transient state kinetic evidence for an oligomer in the mechanism of Na+/H + exchange. Proc. Natl. Acad. Sci. USA 86, 4818-4822. Oude-Elferink, R. P., and Jansen, P. L. (1994). The role of the canalicular multispecific organic anion transporter in the disposal of endo- and xenobiotics. Pharmacol. Ther. 64, 77-97. Oxender, D. L., and Christensen, H. N. (1963). Distinct, mediating systems for the transport of neutral amino acids by the Ehrlich cell. J. Biol. Chem. 238, 3686-3699. Ozegovic, B., McNamara, P. D., and Segal, S. (1982). Cystine uptake by rat jejunal brush border membrane vesicles. Biosci. Rep. 2, 913-920. Padan, E., and Schuldiner, S. (1994). Molecular physiology of the Na+/H + antiporter in Escherichia coli. J. Exp. Biol. 196, 443-456. Page, K. R. (1991). Perfusion of isolated human placenta. Proc. Nutr. Soc. 51), 345-347. Palacfn, M. (1994). A new family of proteins (rBAT and 4F2hc) involved in cationic and zwitterionic amino acid transport: A tale of two proteins in search of a transport function. J. Exp. Biol. 196, 123-137. Palacfn, M., Werner, A., Dittmer, J., Murer, H., and Biber, J. (1990). Expression of rat liver Na+/L-alanine co-transport in Xenopus laevis oocytes. Biochem. J. 270, 189-195. Palos, T. P., Ramachandran, B., Boado, R., and Howard, B. D. (1996). Rat C6 and human astrocytic tumor cells express a neuronal type of glutamate transporter. Mol. Brain Res. 37, 297-303. Panayotova-Heiermann, M., Eskandari, S., Turk, E., Zampighi, G. A., and Wright, E. M. (1997). Five transmembrane behlices form the sugar pathway through the Na+/glucose cotransporter. J. Biol. Chem. 272, 20324-20327. Panayotova-Heiermann, M., Loo, D. D. F., Lostao, M. P., and Wright, E. M. (1994). Sodium/D-glucose cotransporter charge movements involve polar residues. J. Biol. Chem. 269, 21016-21020. Panayotova-Heiermann, M., Loo, D. D. F., Kong, C. T., Lever, J. E., and Wright, E. M. (1996). Sugar binding to Na+/ glucose cotransporters is determined by the carboxyl-terminal half of the protein. J. Biol. Chem. 271, 10029-10034.

3 71

Panet, R., and Atlan, H. (1991). Stimulation of bumetanidesensitive Na +/K +/C1- cotransport by different mitogens in synchronized human skin fibroblasts is essential for cell proliferation. J. Cell Biol. 114, 337-342. PaD, S. S., Paulsen, I. T., and Saier, M. H. (1998). The major facilitator superfamily. Microbiol. Mol. Biol. Rev. 62,1-34. Papazian, D. M., ShaD, X. M., Seoh, S.-A., Mock, A .F., Huang, Y., and Wainstock, D. H. (1995). Electrostatic interactions of $4 voltage sensor in Shaker K + channel. Neuron 14, 1293-1301. Parent, L., Supplisson, S., Loo, D. D. F., and Wright, E. M. (1992). Electrogenic properties of the cloned Na + glucose cotransporter: II. A transport model under nonrapid equilibrium conditions. J. Membr. Biol. 125, 63-79. Paris, S., and Pouyssegur, J. (1984). Growth factors activate the Na+/H + antiporter in quiescent fibroblasts by increasing its affinity for intracellular H +. J. Biol. Chem. 259, 1098910994. Parsons, J. T. (1996). Integrin-mediated signalling: Regulation by protein tyrosine kinases and small GTP-binding proteins. Curr. Opin. Cell Biol. 8, 146-152. Passow, H. (1992). The band 3 proteins: An introduction. Progr. Cell Res. 2, 1-6. Pathak, R. K., Yokode, M., Hammer, R. E., Hofmann, S. L., Brown, M. S., Goldstein, J. L., and Anderson, R. G. W. (1990). Tissue-specific sorting of the human LDL receptor in polarized epithelia of transgenic mice. J. Cell Biol. 111, 347-359. Patti, M-E., Brambilla, E., Luzi, L., Landaker, E. J., and Kahn, C. R. (1998). Bidirectional modulation of insulin action by amino acids. J. Clin. Invest. 101, 1519-1529. Patton, D. E., Silva, T., and Bezanil.la (1997). RNA editing generates a diverse array of transcripts encoding squid Kv2 K + channels with altered functional properties. Neuron 19, 711-722. Paulsen, I. T., and Saier, M.H., Jr. (1997). A novel family of ubiquitous heavy metal ion transport proteins. J. Membr. Biol. 156, 99-103. Paulsen, I. T., Beness, A. M., and Saier, M. H. (1997). Computer-based analyses of the protein constituents of transport systems catalyzing export complex carbohydrates in bacteria. Microbiol. 143, 2685-2699. Paulsen, I. T., Brown, M. H., and Skurray, R. A. (1996). Proton-dependent multidrug efflux systems. Microbiol. Rev. 60, 575-608. Paulsen, I. T., Sliwinski, M. K., and Saier, M. H. (1998). Microbiol genome analyses: Global comparisons of transport capabilities based on phylogenies, bioenergetics and substrate specificities. J. Mol. Biol. 277, 573-592. Pedersen, O., Kahn, C. R., and Kahn, B. B. (1992). Divergent regulation of the Glut1 and Glut4 glucose transporters in isolated adipocytes from Zucker rats. J. Clin. Invest. 89, 1964-1973. Pedersen, P. L. (1996). Frontiers in ATP synthase research: Understanding the relationship between subunit movements and ATP synthesis. J. Bioenerg. Biomembr. 28, 389-395. Peghini, P., Janzen, J., and Stoffel, W. (1997). Glutamate transporter EAAC-l-deficient mice develop dicarboxylic ami-

372

References

noaciduria and behavioral abnormalities but no neurodegeneration. E M B O J. 16, 3822-3832. Peng, S.-B., Crider, B. P., Xie, X.-S., and Stone, D. K. (1994). Alternative mRNA splicing generates tissue-specific isoforms of l l6kDa polypeptide of vacuolar proton pump. J. Biol. Chem. 269, 17262-17266. Penniston, J. T., and Enyedi, A. (1994). Plasma membrane Ca 2+ pump: Recent developments. Cell. Physiol. Biochem. 4: 148-159. Perkins, C. P., Mar, V., Shutter, J. R., Del Castillo, J., Danilenko, D. M., Medlock, E. S., Ponting, I. L., Graham, M., Stark, K. L., Zuo, Y., Cunningham, J. M., and Bosselman, R. A. (1997). Anemia and perinatal death result from loss of the murine ecotropic retrovirus receptor mCAT1. Gene Dev. 11, 914-925. Perlman, D. F., Musch, M. W., and Goldstein, L. (1996). Band 3 in cell volume regulation in fish erythrocytes. Cell. Mol. Biol. 42, 975-984. Persson, B., and Argos, P. (1996). Topology prediction of membrane proteins. Prot. Sci. 5, 363-371. Pessia, M., Tucker, S. J., Lee, K., Bond, C. T., and Adelman, J.P. (1996). Subunit positional effects revealed by novel heteromeric inwardly rectifying K + channels. E M B O J. 15, 2980-2987. Peter, G. J., Davidson, I. G., Ahmed, A., McIlroy, L., Forrester, A. R., and Taylor, P. M. (1996). Multiple components of arginine and phenylananine transport induced in neutral and basic amino acid transporter-cRNA-injected Xenopus oocytes. Biochem. J. 318, 915-922. Peters, L. L, Shivdasani, R. A., Liu, S.-C., Hanspal, M., John, K. M., Gonzalez, J. M., Brugnara, C., Gwyn, B., Mohandas, N., Alper, S. L., Orkin, S. H., and Lux, S. E. (1996). Anion exchanger 1 (band 3) is required to prevent erythrocyte membrane surface loss but not to form the membrane skeleton. Cell 86, 917-927. Peverali, F. A., D'Esposito, M., Acampora, D., Bunone, G., Negri, M., Faiella, A., Stornaiuolo, A., Pannese, M., Migliaccio, E., Simeone, A., Della Valle, G., and Boncinelli, E. (1990). Expression of HOX homeogenenes in human neuroblastoma cell culture lines. Differentiation 45, 61-69. Phan, V. N., Kusuhara, M., Lucchesi, P. A., and Berk, B. C. (1997). A 90-kD Na+-H + exchanger kinase has increased activity in spontaneously hypertensive rat vascular smooth muscle cells. Hypertension 29, 1265-1272. Piatkowski, D., Schneider, K., Salamini, F., and Bartels, D. (1990) Characterization of five abscisic acid-responsive cDNA clones isolated from the desiccation-tolerant plant Craterostigma plantagineum and their relationship to other water-stress genes. Plant Physiol. 94, 1682-1688. Picciotto, M. R., Cohn, J. A., Bertuzzi, G., Greengard, P., and Nairn, A. C. (1992). Phosphorylation of the cystic fibrosis transmembrane conductance regulator. J. Biol. Chem. 267, 12742-12752. Pickart, C. M. (1997). Targeting of substrates to the 26S proteasome. FASEB J. 11, 1055-1066. Pico, C., Liado, I., Pons, A., and Palou, A. (1994). Blood cell to plasma gradients of amino acids in arterial and venous blood in fed and fasted rats. Comp. Biochem. Physiol. (Comp. Physiol.) 107, 589-595.

Pines, G., and Kanner, B. I. (1990). Counterflow of Lglutamate in plasma membrane vesicles and reconstituted preparations from rat brain. Biochemistry 29,11209-11214. Pines, G., Danbolt, N. C., Bjoras, M., Zhang, Y., Bendahan, A., Eide, L., Koepsell, H., Storm-Mathisen, J., Seeberg, E., and Kanner, B.I. (1992). Cloning and expression of a rat brain L-glutamate transporter. Nature 360, 464-467. Pines, G., Zhang, Y., and Kanner, B. I. (1995). Glutamate 404 is involved in the substrate discrimination of GLT-1, a (Na + + K+ )-coupled glutamate transporter from rat brain. J. Biol. Chem. 270, 17093-17097. Pisoni, R. L., and Schneider, J. A. (1992). Amino acid transport by lysosomes. In: "Mammalian Amino Acid Transport" (M. S. Kilberg and D. H/iussinger, eds.), pp. 89-99. Plenum, New York. Pisoni, R. L., Thoene, J. G., and Christensen, H. N. (1985). Detection and characterization of carrier-mediated cationic amino acid transport in lysosomes of normal and cystinotic human fibroblasts. J. Biol. Chem. 260, 4791-4798. Plaza, A., Kono, D. H., and Theofilopoulos, A. N. (1991). New human V/3 genes and polymorphic variants. J. Immunol. 147, 4360-4365. Plumley, D. A., Austgen, T. R., Salloum, R. M., and Souba, W. W. (1990). Role of the lungs in maintaining amino acid homeostasis. J. Parenterol. Enteral. Nutr. 14, 569-573. Pohl, P., Saparov, S. M., and Antonenko, Y. N. (1997). The effect of a transmembrane osmotic flux on the ion concentration distribution in the immediate membrane vicinity measured by microelectrodes. Biophysical J. 72,1711-1718. Pongs, O. (1992). Molecular biology of voltage-dependent potassium channels. Physiol. Rev. 72, $69-$88. Popov, M., Tam, L. Y., Li, J., and Reithmeier, R. A. F. (1997). Mapping the ends of transmembrane segments in a polytopic membrane protein. Scanning N-glycosylation mutagenesis of extracytosololic loops in the anion exchanger, band 3. J. Biol. Chem. 272, 18325-18332. Porter, A. C. G., and Dallman, M. J. (1997). Gene targeting: Techniques and applications to transplantation. Transplantation 64, 1227-1235. Porter, W. R., and Trager, W. F. (1977). Improved nonparametric statistical methods for the estimation of Michaelis-Menten kinetic parameters by the direct linear plot. Biochem. J. 161, 293-302. Portillo, F., Eraso, P., and Serrano, R. (1995). The plasma membrane H+-ATPase of fungi and plants. In "Biomembranes" (A. G. Lee, ed.), Vol. 5, pp. 225-240. J.A.I. Press, Greenwich. Post, R. L., Kume, S., Tobin, T., Orcutt, B., and Sen, A. K. (1969). Flexibility of an active center in Na + K+adenosine triphosphatase. J. Gen. Physiol. 54, 306s-326s. Pratten, M. K., and Lloyd, J. B. (1997). Uptake of microparticles by rat visceral yolk sac. Placenta 18, 547-552. Prezioso, G., and Scalera, V. (1996). Sequential ordered mechanism for the sodium-glutamate transport in intestinal brush border membrane vesicles. Biochim. Biophys. Acta 1279, 144-148. Price, S. R., and Mitch, W. E. (1998). Mechanisms stimulating protein degradation to cause muscle atrophy. Curr. Opin. Clin. Nutr. Metab. Care 1, 79-83.

References Proud, C. G., and Denton, R. M. (1997). Molecular mechanisms for the control of translation by insulin. Biochem. J. 328, 329-341. Quackenbush, E., Gougos, A., Baumal, R., and Letarte, M. (1986). Differential localization within human kidney of five membrane proteins expressed on acute lymphoblastic leukemia cells. J. Immunol. 136, 118-124. Quesada, A. R., and McGivan, J. D. (1988). A rapid method for the functional reconstitution of amino acid transport systems from rat liver plasma membranes: Partial purification of System A. Biochem. J. 255, 963-969. Quick, M. W., Corey, J. L., Davidson, N., and Lester, H. A. (1997). Second messengers, trafficking-related proteins, and amino acid residues that contribute to the functional regulation of the rat brain GABA transporter GAT1. J. Neurosci. 17, 2967-2979. Quiocho, F. A., and Ledvina, P. S. (1996). Atomic structure and specificity of bacterial periplasmic receptors for active transport and chemotaxis: Variation of common themes. Mol. Microbiol. 20, 17-25. Rabon, E. C., Bassilian, S., and Jakobsen, L. J. (1990). Glutaraldehyde crosslinking analysis of the C12E8 solubilized H, K-ATPase. Biochim. Biophys. Acta 1039, 277-289. Rabon, E. C., Gunther, R. D., Bassilian, S., and Kempner, E. S. (1988). Radiation inactivation analysis of oligomeric structure of the H, K-ATPase. J. BioL Chem. 263, 1618916194. Rahlfs, S., and MUller, V. (1997). Sequence of subunit c of the Na+-translocating FaF0 ATPase of Acetobacterium woodii: Proposal for determinants of Na + specificity as revealed by sequence comparisons. FEBS Letts. 404, 269-271. Ramamoorthy, S., Kulanthaivel, P., Leibach, F. H., Mahesh, V. B., and Ganapathy, V. (1993). Solubilization and functional reconstitution of the human placental taurine transporter. Biochim. Biophys. Acta 1145, 250-256. Ranganathan, R., Lewis, J. H., and MacKinnon, R. (1996). Spatial localization of the K + channel selectivity filter by mutant cycle-based structure analysis. Neuron 16,131-139. Rao, R., and Slayman, C. W. (1993). Mutagenesis of conserved residues in the phosphorylation domain of the yeast plasma membrane H+-ATPase: Effects on structure and function. J. Biol. Chem. 268, 6708-6713. Rao, G. N., Sardet, C., Pouyssegur, J., and Berk, B. C. (1992). Phosphorylation of Na+/H + antiporter is not stimulated by phorbol ester and acidification in granulocytic HL-60 cells. Am. J. Physiol. 264, C1278-C1284. Raynard, R. S., and Cossins, A. R. (1991). Homeoviscous adaptation and thermal compensation of sodium pump of trout erythrocytes. Am. J. Physiol. 26tl, R916-R924. Regen, D. M., and Morgan, H. E. (1964). Studies of the glucose transport system in the rabbit erythrocyte. Biochim. Biophys. Acta 79, 151-166. Reithmeier, R. A. F., and Casey, J. R. (1992). Oligomeric structure of the human erythrocyte band 3 anion transport protein. Prog. Cell Res. 2, 181-190 Rennie, M. J. (1985). Muscle protein turnover and the wasting due to injury and disease. Br. Med. Bull 41, 257-264. Rennie, M. J. (1996). Influence of exercise on protein and amino acid metabolism. In "American Physiological Soci-

373

ety Handbook of Physiology on Exercise" R. L. Terjung, ed.), pp 995-1035. American Physiological Society, Bethesda, MD. Rennie, M. J., MacLennan, P. A., Hundal, H. S., Weryk, B., Smith, K., Taylor, P. M., Egan, C. J., and Watt, P. W. (1989). Skeletal muscle glutamine transport, intramuscular glutamine concentration and muscle protein turnover. Metabolism 38, 47-51. Rennie, M. J., Khogali, S. E. O., Low, S. Y., McDowell, H. E., Hundal, H. S., Ahmed, A., and Taylor, P. M. (1996). Amino acid transport in heart and skeletal muscle and the functional consequences, Biochem. Soc. Trans. 24, 869-873. Rennie, M. J, Smith, K., and Watt, P. W. (1994). Measurement of human tissue protein synthesis: An optimal approach. Am. J. PhysioL 266, E298-E307. Rentsch, D., Boorer, K. J., and Frommer, W. B. (1998). Structure and function of plasma membrane amino acid, oligopeptide and sucrose transporters from higher plants. J. Membr. Biol. 162, 177-190. Restrepo, D., Cronise, B. L., Snyder, R. B., Spinelli, L. J., and Knauf, P. A. (1991). Kinetics of DIDS inhibition of HL60 cell anion exchange rules out ping-pong model with slippage. Am. J. Physiol. 260, C535-C544. Restrepo, D., Cronise, B. L., Snyder, R. B., and Knauf, P. A. (1992). A novel method to differentiate between ping pong and simultaneous exchange kinetics and its application to the anion exchanger of the HL60 cell. J. Gen. Physiol. 100, 825-846. Rettig, J., Heinemann, S. H., Wunder, F., Lorra, C., Parcej, D. N., Dolly, J. O., and Pongs, O. (1994). Inactivation properties of voltage-gated K + channels altered by presence of/3-subunit. Nature 369, 289-294. Rettig, J., Wunder, F., Stocker, M., Lichtinghagen, R., Mastiaux, F., Beckh, S., Kues, W., Pedarzani, P., Schroter, K. H., and Ruppersberg, J. P. (1992). Characterization of a Shaw-related potassium channel family in rat brain. E M B O J. 11, 2473-2486. Reynafarje, B. D., and Pedersen, P. L. (1996). ATP synthase: Conditions under which all catalytic sites of the Fa moiety are kinetically equivalent in hydrolyzing ATP. J. Biol. Chem. 271, 32546-32550. Ricquier, D., Casteilla, L., and Bouillaud, F. (1991). Molecular studies of the uncoupling protein. FASEB J. 5, 2237-2242. Risso, S., De Felice, L. J., and Blakely, R. D. (1996). Sodiumdependent GAB A-induced currents in GATl-transfected HeLa cells. J. Physiol. 490, 691-702. Ritchie, R. J., and Prvan, T. (1996). Current statistical methods for estimating the Km and Vmaxof Michaelis-Menten kinetics. Biochem. Educ. 24, 196-206. Roberts, S. K., Yano, M., Ueno, Y., Pham, L., Alpini, G., Agre, P., and LaRusso, N.F. (1994). Cholangiocytes express the aquaporin CHIP and transport water via a channelmediated mechanism. Proc. Natl. Acad. Sci. USA 91, 13009-13013. Robertson, B. (1997). The real life of voltage-gated K + channels: More than model behavior. Trends Pharmacol. Sci. 18, 474-483.

374

References

Roden, D. M., and George, A. L. (1997). Structure and function of cardiac sodium and potassium channels. Am. J. PhysioL 273, H511-H525. Rodgers, A. J. W., Wilkens, S., Aggeler, R., Morris, M. B., Howitt, S. M., and Capaldi, R. A. (1997). The subunit 6subunit b domain of the Escherichia coli FIFo ATPase. J. Biol. Chem. 272, 31058-31064. Rolfe, D. F. S., and Brown, G. C. (1997). Cellular energy utilization and molecular origin of standard metabolic rate in mammals. Physiol. Rev. 77, 731-758. Rollins,B. J., Morrison, E. D., Usher, P., and Flier, J. S. (1988). Platelet-derived growth factor regulates glucose transporter expression. J. BioL Chem. 263, 16523-16526. Romero, M. F., Hediger, M. A., Boulpaep, E. L., and Boron, W. F. (1997). Expression cloning of the renal electrogenic Na+/HCO~ cotransporter. Nature 387, 409-413. Romero, M. F., Kanai, Y., Gunshin, H., and Hediger, M. A. (1998). Expression cloning using Xenopus laevis oocytes. In "Methods in Enzymology" (S. G. Amara ed.), Vol. 296, pp. 17-52. Academic Press, San Diego. Rooyackers, O. E., and Nair, K. S. (1997). Hormonal regulation of human muscle protein metabolism. Annu. Rev. Nutrition. 17, 457-485. Ross, J. C., Fennessey, P. V., Wilkening, R. B., Battaglia, F. C., and Meschia, G. (1996). Placental transport and fetal utilization of leucine in a model of fetal growth retardation. Am. J. Physiol. 270, E491-E503. Rost, B. Casadio, R., Fariselli, P., and Sander, C. (1995). Prediction of helical transmembrane segments at 95% accuracy. Prot. Sci. 4, 521-523. Rost, B., Fariselli, P., and Casadio, R. (1996). Topology prediction for helical transmembrane proteins at 86% accuracy. Prot. Sci. 5, 1704-1718. Rothman, J. E., and Wieland, F. T. (1996). Protein sorting by transport vesicles. Science 272, 227-234. Rothstein, J. D., Dykes-Hoberg, M., Pardo, C. A., Bristol, L.A., Jin, L., Kuncl, R.W., Kanai, Y., Hediger, M. A., Wang, Y., Schielke, J. P., and Welly, D. F. (1996). Neuron 16, 675-686. Rothstein, J. D., Martin, L., Levey, A. I., Dykeshoberg, M., Jin, L., Wu, D., Nash, N., and Kuncl, R. W. (1994). Localization of neuronal and glial glutamate transporters. Neuron 13, 713-725. Rouse, K., Nwokedi, E., Woodliff, J. E., Epstein, J., and Klimberg, V. S. (1995). Glutamine enhances selectivity of chemotherapy through changes in glutathione metabolism. Ann. Surg. 221, 420-426. Roush, D. L., Gottardi, C. J., Naim, H. Y., Roth, M. G., and Caplan, M. J. (1998). Tyrosine-based membrane protein sorting signals are differentially interpreted by polarized Madin-Darby canine kidney and LLC-PK1 epithelial cells. J. Biol. Chem. 273, 26862-26869. Ruiz-Montasell, B., G6mez-Angelats, M., Casado, F. J., Felipe, A., McGiven, J. D. and Pastor-Anglada, M. (1994). Evidence for a regulatory protein involved in the increased activity of system A for neutral amino acid transport in osmotically stressed mammalian cells. Proc. Natl. Acad. Sci. USA 91, 9569-9573.

Ruppersberg, J. P., Ermler, M., Knopf, M., Kues, W., Jonas, P., and Koenen, M. (1993). Properties of Shaker-homologous potassium channels expressed in the mammalian brain. Cell Physiol. Biochem. 3, 250-269. Sabbert, D., Engelbrecht, S., and Junge, W. (1996). Intersubunit rotation in active F-ATPase. Nature 381, 623-625. Sabbert, D., Engelbrecht, S., and Junge, W. (1997). Functional and idling rotatory motion within F1-ATPase. Proc. Natl. Acad. Sci. USA 94, 4401-4405. Saccomani, G., Sachs, G., Cuppolletti, J., and Jung, C. Y. (1981). Target molecular weight of the gastric (H + + K +)ATPase functional and structural molecular size. J. Biol. Chem. 256, 7727-7729. Sagara, J., Miura, K., and Bannai, S. (1993). Maintenance of neuronal glutathione by glial cells. J. Neurochem. 61,16721676. Saier, M. H. (1994). Computer-aided analyses of transport protein sequences: gleaning evidence concerning function, structure, biogenesis and evolution. Microbiol. Rev. 58, 71-93. Saier, M. H. (1996). Phylogenetic approaches to the identification and characterization of protein families and superfamilies. Microbiol. Comp. Genom. 1, 129-150. Saier, M. H. (1998). Molecular phylogeny as a basis for the classification of transport proteins from bacteria, archaea and eukarya. Adv. MicrobioL Phys. 40, 81-136. Saier, M. H., Eng, B. H., Fard, S., Garg, J., Haggerty, D. A., Hutchinson, W. J., Jack, D. L., Lai, E. C., Liu, H. J., Nusinew, D. P., Omar, A. M., Pao, S. S., Paulsen, I. T., Quan, J. A., Sliwinski, M., Tseng, T.-T., Wachi, S., and Young, G.B. (1998) Phylogenetic characterization of novel transport protein families revealed by genome analyses. Biochim. Biophys. Acta (in press). Saito, M., Nelson, C., Salkoff, L., and Lingle, C.J. (1997) A cysteine-rich domain defined by a novel exon in a Slo variant in rat adrenal chromaffin cells and PC12 cells. J. Biol. Chem. 272, 11710-11717. Sakura, H., Maekawa, T., Imamoto, F., Yasuda, K., and Ishii, S. (1988). Two Human genes isolated by a novel method encode DNA-binding proteins containing a common region of homology. Gene 73, 499-507. Salhany, J. M. (1990). "Erythrocyte Band 3 Protein." CRC Press, Boca Raton, FL. Salhany, J. M. (1992). Band 3 quaternary states and allosteric control of function. Progr. Cell Res. 2, 191-205. Salhany, J. M. (1996). Allosteric effects in stilbenedisulfonate binding to band 3 protein (AE1). Cell MoL Biol. 42, 10651096. Salter, M., Knowles, R. G., and Pogson, C. I. (1986). Quantification of the importance of individual steps in the control of aromatic amino acid metabolism. Biochem. J. 234, 635-647. Saltiel, A. R. (1996). Diverse signalling pathways in the cellular actions of insulin. Am. J. Physiol. 270, E375-E385. Samson, M., Osty, J., Francon, J., and Blondeau, J-P. (1992). Triiodothyronine binding sites in the rat erythrocyte membrane: Involvement in triiodothyronine transport and relation to the tryptophan transport System T. Biochim. Biophys. Acta 11118, 91-98.

References Sanchez, D. Y., and Blatz, A. L. (1995). Block of neuronal chloride channels by tetraethylammonium ion derivatives. J. Gen. Physiol. 106, 1031-1046. Sanchez-del-Pino, M. M., Peterson, D. R., and Hawkins, R. A. (1995). Neutral amino acid transport characterization of isolated luminal and abluminal membranes of the bloodbrain barrier. J. Biol. Chem. 270, 14913-14918. Sankaram, M. B., Marsh, D., and Thompson, T. E. (1992). Determination of fluid and gel domain sizes in twocomponent two-phase lipid bilayers. Biophys. J. 63, 340-349. Sansom, M. S. P. (1997). Structure of a molecular hole-punch. Nature 385, 390-391. Sardet, C., Counillon, L., Franchi, A., and Pouyssegur, J. (1990). Growth factors induce phosphorylation of the Na+/ H + antiporter, a glycoprotein of 110 kD. Science, 247, 723-726. Sardet, C., Fafournoux, P., and Pouyssegur, J. (1991). Thrombin, epidermal growth factor, and okadaic acid activate the Na+/H + exchanger, NHE1, by phosphorylating a set of common sites. J. Biol. Chem. 266, 19166-19171. Sato, K., Adams, R., Betz, H., and Schloss, P. (1995). Modulation of a recombinant glycine transporter (GLYTlb) by activation of protein kinase C. J. Neurochem. 65, 19671973. Scannevin, R. H., and Trimmer, J. S. (1997). Cytoplasmic domains of voltage-sensitive K + channels involved in mediating protein-protein interactions. Biochem. Biophys. Res. Commun. 232, 585-589. Schachtmann, D. P., Schroeder, J. I., Lucas, W. J., Anderson, J. A., and Gaber, R. F. (1992). Expression of an inwardrectifying potassium channel by the Arabidopsis KAT1 cDNA. Science 258, 1654-1658. Schaffer, J. E., and Lodish, H. F. (1994). Expression cloning and characterization of a novel adipocyte long chain fatty acid transport protein. Cell 79, 427-436. Schenerman, M. A., and Kilberg, M. S. (1986). Maintenance of glucagon-stimulated system A amino acid transport activity in rat liver plasma membrane vesicles. Biochim. Biophys. Acta 856, 428-436. Schlesser, A., Ulaszewski, S., Ghislain, M. and Goffeau, A. (1988). A second transport ATPase gene in Saccharomyces cerevisiae. J. Biol. Chem. 263, 19480-19487. Schmall, B., Conti, P. S., and Alauddin, M. M. (1996). Synthesis of [11C-methyl]-aminoisobutyric acid (AIB). Nucl. Med. Biol. 23, 263-266. Schmidt, C. E., Chen, T., and Lauffenburger, D. A. (1993). Simulation of integrin-cytoskeletal interactions in migrating fibroblasts. Biophys. J. 67, 461-474. Schneider, M. L., and Post, C. B. (1995). Solution structure of a band 3 peptide inhibitor bound to aldolase: A proposed mechanism for regulating binding by tyrosine phosphorylation. Biochemistry 34, 16574-16584. Schnell, K. F., and Besl, E. (1984). Concentration dependence of the unidirectional sulfate and phosphate flux in human red cell ghosts under selfexchange and under homoexchange conditions. Pfliigers Arch. 402, 197-206. Schroeder, J. I. (1995). Heterologous expression of higher plant transport proteins and repression of endogenous ion

375

currents in Xenopus oocytes. Methods Cell Biol. 50, 519-553. Schroeder, R. London, E., and Brown, D. (1994). Interactions between saturated acyl chains confer detergent resistance on lipids and glycosylphosphatidylinositol (GPI)-anchored proteins: GPI-anchored proteins in liposomes and cells show similar behavior. Proc. Natl. Acad. Sci. USA 91, 12130-12134. Schultz, B., Frommer, W. B., Fliagge, U.-I., Hummel, S., Fischer, K., and Willmitzer, L. (1993). Expression of the triose phosphate translocator gene from potato is light dependent and restricted to green tissues. Mol. Gen. Genet. 238, 357-361. Schultz, B., Frommer, W. B., Fltigge, U.-I., and Wagner (1994). Ion channel properties of the reconstituted chloroplast triose phosphate/phosphate translocation. J. Biol. Chem. 269, 29481-29489. Schwartz, H. L., Strait, K. A., and Oppenheimer, J. H. (1993). Molecular mechanisms of thyroid hormone action: A physiologic perspective. Clin. Lab. Med. 13, 543-561. Schwarz, F. P., and Inesi, G. (1997). Entropic drive in the sarcoplasmic reticulum ATPase interaction with Mg 2+ and Pi. Biophys. J. 73, 2179-2182. Schwarz, M. A., Gross, A., Steinkamp, T., Fltigge, U.-I., and Wagner, R. (1994). Ion channel properties of the reconstituted chloroplast triose phosphate/phosphate translocator. J. Biol. Chem. 269, 29481-29489. Schwegler, J. S., Sibernagl, S., Tamarappoo, B. K., and Welbourne, T. C. (1992). Amino acid transport in the kidney. In "Mammalian Amino Acid Transport: Mechanisms and Control" (M. S. Kilberg and D. H~iussinger, eds.), pp. 233260. Plenum, New York/London. Scriver, C. R. (1988). Nutrient-gene interactions: The gene is not the disease and vice versa. Am. J. Clin. Nutr. 48, 15051509. Segel, G. B. (1992). Amino acid transport in lymphocytes. In "Mammalian Amino Acid Transport: Mechanisms and Control" (M. S. Kilberg and D. H~iussinger, eds.), pp. 261274. Plenum, New York. Segel, G. B., Woodlock, T. J., and Lichtman, M. A. (1988). Trans-stimulation of L-system amino acid transport in normal and chronic leukemic human lymphocytes: Phorbol ester restores function in CLL. J. Cell PhysioL 134, 503-508. Segal, S., and Thier, S. O. (1995). Cystinuria. In "The Metabolic and Molecular Bases of Inherited Diseases" (C. H. Scriver, A. L. Beaudet, W. S. Sly, and D. Valle eds.), pp. 3581-3601. McGraw-Hill, New York. Sekler, I., Kobayashi, S., and Kopito, R. R. (1996). A cluster of cytoplasmic histidine residues specifies pH dependence of the AE2 plasma membrane anion exchanger. Cell 86, 929-935. Selski, D. J., Thomas, N. E., Coleman, P. D., and Rogers, K. E. (1993). The human brain homeogene, DLX-2: cDNA sequence and alignment with the murine homologue. Gene 132, 301-303. Semenza G. Kessler, M., Hosang, M., Weber, J., and Schmidt, U. (1984). Biochemistry of the Na +, D-glucose cotransporter of the small-intestinal brush-border membrane. Biochim. Biophys. Acta 779, 343-379.

376

References

Senior, A. E., Weber, J., and A1-Shawi, M. K. (1995). Catalytic mechanism of Escherichia coli F1-ATPase. Biochem. Soc. Trans. 23, 747-752. Sentenac, H., Bonneaud, N., Minet, M., Lacroute, F., Salmon, J. M., Gaynard, F., and Grignon, C. (1992). Cloning and expression in yeast of a plant potassium ion transport system. Science 256, 663-665. Sha, Q., Romano, C., Lopatin, A. N., and Nichols, C. G. (1996). Spermidine release from Xenopus oocytes: Electrodiffusion through a membrane channel. J. Biol. Chem. 271, 3392-3397. Shafqat, S., Tamarappoo, B. K., Kilberg, M. S., Puranam, R. S., McNamara, J. O., Guadano- Ferraz, A., and Fremeau, R. T., Jr. (1993). Cloning and expression of a novel Na +dependent neutral amino acid transporter structurally related to mammalian Na+/glutamate cotransporters. J. Biol. Chem. 268, 15351-15355. Shai, Y. (1995). Molecular recognition between membranespanning polypeptides. Trends Biol. Sci. 20, 460-464. Shainskaya, A., and Karlish, S. J. D. (1994). Evidence that the cation occlusion domain of Na/K-ATPase consists of a complex of membrane-spanning segments: Analysis of limit membrane-embedded tryptic fragments. J. BioL Chem. 269, 10780-10789. Shawver, L. K., Olson, S. A., White, M. K., and Weber, M. J. (1987). Degradation and biosynthesis of the glucose transporter protein in chicken embryo fibroblasts transformed by the src oncogene. Mol. Cell Biol. 7, 2112-2118. Sheikh-Hamad, D., Di Mari, J., Suki, W. N., Safirstein, A., Watts, B. A., and Rouse, D. (1998). p38 kinase activity is essential for osmotic induction of mRNAs for HSP70 and transporter for organic solute betaine in Madin-Darby canine kidney cells. J. Biol. Chem. 273, 1832-1837. Shepherd, P. R., Gnudi, L., Tozzo, E., Yang, H., Leach, F., and Kahn, B. B. (1993). Adipose cell hyperplasia and enhanced glucose disposal in transgenic mice overexpressing GLUT4 selectively in adipose tissue. J. Biol. Chem. 268, 2224322246. Sherman-Gold, R. E. (ed.) (1993). "The Axon Guide." Axon Instruments, Foster City, CA. Sigel, E. (1990). Use of Xenopus oocytes for the functional expression of plasma membrane proteins. J. Membr. Biol. 117, 201-221. Shin, J. M., and Sachs, G. (1996). Dimerization of the gastric H+,K+-ATPase. J. BioL Chem. 271, 1904-1908. Shinohara, Y., Yamamoto, K., Kogure, K., Ichihara, J., and Terada, H. (1994). Steady state transcript levels of the type II hexokinase and type 1 glucose transporter in human tumor cell lines. Cancer Lett. 82, 27-32. Shoemaker, D. P., and Garland, C. W. (1967). "Experiments in Physical Chemistry: Treatment of Experimental Data", pp. 22-23. McGraw-Hill, New York. Shuck, M. E., Bock, J. H., Benjamin, C. W., Tsai, T. D., Lee, K. S., Slightom, J. L., and Bienkowski, M. J. (1994). Cloning and characterization of multiple forms of the human kidney ROM-K potassium channel. J. Biol. Chem. 269, 2426124270. Siczkowski, M., and Ng, L. L. (1996). Phorbol ester activation of the rat vascular myocyte Na+-H + exchanger isoform 1. Hypertension 27, 859-866.

Sigworth, F. J. (1994). Voltage gating of ion channels. Quart. Rev. Biophys. 27, 1-40. Silbernagl, S. (1988). The renal handling of amino acids and oligopeptides. Physiol. Rev. 68, 911-1007. Silbernagl, S., Volker, K., and Dantzler, W. H. (1996). Compartmentation of amino acids in the rat kidney. Am. J. Physiol. 270, F154-F163. Silver, S., Ji., G., BrOer, S., Dey, S., Dou, D., and Rosen, B. P. (1993). Orphan enzyme or patriarch of a new tribe: The arsenic resistance ATPase of bacterial plasmids. Mol. Microbiol. 8, 637-642. Simons, K. (1995). Biogenesis of epithelial cell surface polarity. Harvey Lect. Ser. 89, 125-146. Simons, K., and Ikonen, E. (1997). Functional rafts in cell membranes. Nature 387, 569-572. Simonsen, H., and Lodish, H. F. (1994). Cloning by function: Expression cloning in mammalian cells. Trends Pharmacol. Sci. 15, 437-441. Simson, R., Yang, B., Doherty, P., Moore, S., Walsh, F., and Jacobson, K. (1995). The mosaic structure of cell membranes revealed by transient confinement of GPI-linked NCAM-125. Biophys. J. 68, A436. Singer, S. J., and Nicolson, G. L. (1972). The fluid mosaic model of the structure of cell membranes. Science 175, 720-731. Singh, C., Sankararamakrishnan, R., Subramaniam, S., and Jakobsson, E. (1996). Solvation, water permeation, and ionic selectivity of a putative model for the pore region of the voltage-gated sodium channel. Biophys. J. 71, 22762288. Sinha, M. K., Raineri-Maldonado, C., Buchanan, C., Pories, W. J., Carter, Su C., Pilch, P. F., and Caro, J. F. (1991). Adipose tissue glucose trm~sporters in NIDDM: Decreased levels of muscle/fat isoform. Diabetes 40, 472-477. Sips, H. J., Groen, H. J., and Tager, J. M. (1980). Plasmamembrane transport of alanine is rate-limiting for its metabolism in rat-liver parenchymal cells. FEBS Lett. 119, 271-274. Skinner, F. K., Ward, C. A., and Bardakjian, B. L. (1993). Pump and exchanger mechanisms in a model of smooth muscle. Biophys. Chem. 45, 253-272. Skulachev, V. P. (1991). Fatty acid circuit as a physiological mechanism of uncoupling of oxidative phosphorylation. FEBS Lett. 294, 158-162. Slade, M. B., Chang, A. C. M., and Williams, K. L. (1990). The sequence and organization of Ddp2, a high-copy-number nuclear plasmid of Dictyostelium discoideum. Plasmid 24, 195-207. Smit, L. S., Wilkinson, D. J., Mansoura, M. K., Collins, F. S., and Dawson, D. C. (1993). Functional roles of the nucleotide-binding folds in the activation of the cystic fibrosis transmembrane conductance regulator. Proc. Natl. Acad. Sci. USA 90, 9963-9967. Smith, Q. R., and Cooper, A. J. (1992). L-Amino acid transport in brain. In "Mammalian Amino Acid transport: Mechanisms and Control" (M. S. Kilberg and D. H~iussinger, eds.), pp. 165-194. Plenum, New York/London. Smith-Maxwell, C. J., Ledwell, J. L., and Aldrich, R. W. (1998). Role of the $4 cooperativity of voltage-dependent potassium channel activation. J. Gen. Physiol. 111, 399-420.

References Smythe, J. A., Peterson, M. G., Coppel, R. L., Saul, A. J., Kemp, D. J., and Anders, R. F. (1990). Structural diversity in the 45-kilodalton merozoite surface antigen of Plasmodium falciparum. Mol. Biochem. Parasitol. 39, 227-234. Solioz, M., and Davies, K. (1994). Operon of vacuolar-type Na+-ATPase of Enterococcus hirae. J. Biol. Chem. 269, 9453-9459. Solomon, A. K. (1992). Interactions between band 3 and other transport-related proteins. Progr. Cell Res. 2, 269-283. Sonders, M. S., and Amara, S. G. (1996). Channels in transporters. Curr. Opin. Neurobiol. 6, 294-302. Sonders, M. S., Zhu, S. J., Zahniser, N. R., Kavanaugh, M. P., and Amara, S. G. (1997). Multiple ionic conductances of the human dopamine transporter: The actions of dopamine and psychostimulants. J. Neurosci. 17, 960-974. Song, L., Hobaugh, M. R., Shustak, C., Cheley, S., Bayley, H., and Gouaux, J. E. (1996). Structure of Staphylococcal a-hemolysin, a heptameric transmembrane pore. Science 274, 1859-1866 SCrensen, T., Vilsen, B., and Andersen, J. P. (1997). Mutation Lys758 ~ Ile of the sarcoplasmic reticulum CaZ+-ATPase enhances dephosphorylation of EzP and inhibits the E2 to E1Ca2 transition. J. Biol. Chem. 272, 30244-30253. Sorrentino, D., Stump, D. D., Van Ness, K., Simard, A., Schwab, A. J., Zhou, S.-L., Goresky, C. A., and Berk, P. D. (1996). Oleate uptake by isolated hepatocytes and the perfused rat liver is competitively inhibited by palmitate. Am. J. Physiol. 270, G385-G392. Souba, W. W. (1991). Glutamine: A key substrate for the splanchnic bed. Annu. Rev. Nutr. 11, 285-308. Souba, W. W. (1992). "GlutaminemPhysiology, Biochemistry and Nutrition in Critical Illness" R. G. Landes, Austin, TX. Spauschus, A., Lentes, K. U., Wischmeyer, E., Dissman, E., Karschin, C., and Karschin, A. (1996). A G-proteinactivated inwardly rectifying K + channel (GIRK4) from human hippocampus associates with other GIRK channels. J. Neurosci. 16, 930-938. Spears, G., Sneyd, J. G. T., and Loten, E. G. (1971). A method for deriving kinetic constants for two enzymes acting on the same substrate. Biochem. J. 125, 1149-1151, Spieholz, C., Heaney, M. L., Morrison, M. E., Houghton, A. N., Vera, J. C., and Golde, D. W. (1995). Granulocytemacrophage colony-stimulating factor signals for increased glucose uptake in human melanoma cells. Blood 85, 973-980. Sreekrishna, K., Brankamp, R. G., Kropp, K. E., Blankenship, D. T., Tsay, J. T., Smith P. L., Wierschke, J. D., Subramaniam, A., and Birkenberger, L. A. (1997). Strategies for optimal synthesis and secretion of heterologous proteins in the methylotrophic yeast Pichia pastoris. Gene 190, 55-62. Stampe, P., and Begenisich, T. (1996). Unidirectional K +fluxes through recombinant Shaker potassium channels expressed in single Xenopus oocytes. J. Gen. Physiol. 107, 449-457. Stappen, R., and Kr~imer, R. (1994). Kinetic mechanism of phosphate/phosphate and phosphate/OH- antiports catalyzed by a reconstituted phosphate carrier from beef heart mitochondria. J. BioL Chem. 269, 11240-11246. Stanley, K. K. (1996). Regulation of targeting signals in membrane proteins. Mol. Membr. BioL 13, 19-27.

377

Stein, W. D. (1986). "Transport and Diffusion across Cell Membranes." Academic Press, Orlando, FL. Stein, W. D. (1990). Energetics and the design of the Na/KATPase J. Theor. Biol. 147, 145-159. Stein, W. D. (1990). "Channels, Carriers, and Pumps." Academic Press, San Diego, CA. Steiner, H. Y., Song, W., Zhang, L., Naider, F., Becker, J. M., and Stacey, G. (1994). An arabidopsis peptide transporter is a member of a new class of membrane transport proteins. Plant Cell. 6, 1289-1299. Steinporsdottir, V., Frithriksdottir, V., Gunnarsson, E. and Andresson, O.S. (1995) Expression and purification of Clostridium perfringens beta-toxin glutathione S-transferase fusion protein. FEMS Microbiol. Lett. 130, 273-278. Stevens, B. R. (1992). Amino acid transport in intestine. In "Mammalian Amino Acid Transport: Mechanisms and Control" (M. S. Kilberg and D. H~iussinger, eds.), pp. 149164. Plenum, New York/London. Stevens, B. R., Kakuda, D. K., Yu, K., Waters, M., Vo, C. B., and Raizada, M. K. (1996). Induced nitric oxide synthesis is dependent on induced alternatively spliced CAT-2 encoding L-arginine transport in brain astrocytes. J. Biol. Chem. 271, 24017-24022. Stoffel, M., and Duncan, S. A. (1997). The maturity-onset diabetes of the young (MODY1) transcription factor HNF4a regulates expression of genes required for glucose transport and metabolism. Proc. Natl. Acad. Sci. USA 94, 13209-13214. Stokes, D. L., Taylor, W. R., and Green, N. M. (1994). Structure, transmembrane topology and helix packing of P-type ion pumps. FEBS Lett. 346, 32-38. Stott, K., Blackburn, J. M., Butler, P. J. G., and Perutz, M. (1995). Incorporation of glutamine repeats makes protein oligomerize: Implications for neurodegenerative diseases. Proc. Natl. Acad. Sci. USA 92, 6509-6513. Strange, K., Emma, F., and Jackson, P. S. (1996). Cellular and molecular physiology of volume-sensitive anion channels. Am. J. Physiol. 270, C711-C730. Streisinger, G., Okada, Y., Emrich, J., Newton, J., Tsugita, A., Terzaghi, E., and Inoye, M. (1966). Frameshift mutations and the genetic code. Cold Spring Harb. Syrup. Quant. Biol. 31, 77-84. Stremmel, W., and Berk, P. D. (1986). Hepatocellular influx of [14C]oleate reflects membrane transport rather than intracellular metabolism or binding. Proc. Natl. Acad. Sci. USA 83, 3086-3090. Sttirmer, W., Apell, H.-J., Wuddel, I., and L~iuger, P. (1989). Conformational transitions and charge translocation by the Na,K pump: Comparison of optical and electrical transients elicited by ATP-concentration jumps. J. Membr. Biol. 110, 67-86. Suleymanian, M. A., and Baumgarten, C. M. (1996). Osmotic gradient-induced water permeation across the sarcolemma of rabbit ventricular myocytes. J. Gen. Physiol. 107, 503-514. Sumikawa, K., Houghton, M., Entage, J. S., Richards, B. M., and Barnard, E. A. (1981). Active multi-subunit ACh receptor assembled by translation of heterologous mRNA in Xenopus oocytes. Nature 292, 862-864.

378

References

Supersac, G., Prevost, G., and Piemont, Y. (1993). Sequencing of leucocidin R from Staphylococcus aureus p83 suggests that staphylococcal leucodidins and gamma-hemolysin are members of a single, two-component family of toxins. Infect. Immun. 61, 580-587. Sutherland, G. R., and Richards, R. I. (1995). Simple tandem DNA repeats and human genetic disease. Proc. Natl. Acad. Sci. USA 92, 3636-3641. Suzuki, Y., Nogi, Y., Abe, A., and Fukasawa, T. (1988). G A L l l protein, an auxiliary transcription activator for genes encoding galactose-metabolizing enzymes in Saccharomyces cerevisiae. MoL Cell Biol. 8, 4991-4999. Sweeney, F. P., Siczkowski, M., Davies, J. E., Quinn, P. A., McDonald, J., Krolewski, B., Krolewski, A. S., and Ng, L. L. (1995). Phosphorylation and activity of Na+/H + exchanger isoform 1 of immortalized lymphoblasts in diabetic nephropathy. Diabetes 44, 1180-1185. Tabcharani, J. A., Chang, X.-B., Riordan, J. R., and Hanrahan, J. W. (1991). Phosphorylation-regulated C1- channel in CHO cells stably expressing the cystic fibrosis gene. Nature 352, 628-631. Tabor, H., and Tabor, C. W. (1976). 1,4-Diaminobutane (putrescine), spermidine and spermine. Annu. Rev. Biochem. 45, 285-306. Taglialatela, M., Toro, L., and Stefani, E. (1992). Novel voltage clamp to record small, fast currents from ion channels expressed in Xenopus oocytes. Biophys. J. 61, 78-82. Takahashi, E., Abe, J., and Berk, B. C. (1997). Angiotensin II stimulates p90 rsk in vascular smooth muscle cells: A potential Na+-H + exchanger kinase. Circ. Res. 81, 268-273. Takase, K., Kakinuma, S., Yamato, I., Konishi, K., Igarashi, K., and Kakinuma, Y. (1994). Sequencing and characterization of the ntp gene cluster for vacuolar-type Na+-translocating ATPase of Enterococcus hirae. J. Biol. Chem. 269, 11037-11044. Takumi, T., Ishii, T., Horio, Y., Morishige, K., Takahashi, N., Yamada, M., Yamashita, T., Kiyama, H., Sohmiya, K., Nakanishi, S., and Kurachi, Y. (1995). A novel ATPdependent inward rectifier potassium channel expressed predominantly in glial cells. J. Biol. Chem. 2"/0, 1633916346. Tamarappoo, B. K., and Kilberg, M. S. (1991). Functional reconstitution of the hepatic System N amino acid transport activity. Biochem. J. 274, 97-101. Tamarappoo, B. K., McDonald, K. K., and Kilberg, M. S. (1996). Expressed human hippocampal ASCT1 amino acid transporter exhibits a pH-dependent change in substrate specificity. Biochim. Biophys. Acta 1279, 131-136. Tanaka, K., Miura, N., Satokata, I., Miyamoto, I., Yoshida, M. C., Satoh, Y., Kondo, S., Yasui, A., Okayama, H., and Okada, Y. (1990). Analysis of a human DNA excision repair gene involved in group A xeroderma pigmentosum and containing a zinc-finger domain. Nature 348, 73-76. Tanaka, K., Watase, K., Manabe, T., Yamada, K., Watanabe, M., Takahashi, K., Iwama, H., Nishikawa, T., Ichihara, N., Kikuchi, T., Okuyama, S., Kawashima, N., Hori, S., Takimoto, M., and Wada, K. (1997). Epilepsy and exacerbation of brain injury in mice lacking the glutamate transporter GLT-1. Science 276, 1699-1702.

Tanaka, M., Lai, J.-S., and Herr, W. (1992). Promoter-selective activation domains in Oct-1 and Oct-2 direct differential activation of an snRNA and mRNA promoter. Cell 68, 755-767. Tang, X.-B., Fujinaga, J., Kopito, R., and Casey, J. R. (1998). Topology of the region surrounding Glu 681 of human AE1 protein, the erythrocyte anion exchanger. J. Biol. Chem. 273, 22545-22553. Tanii, H. (1996). Anesthetic activity of monoketones in mice: Relationship to hydrophobicity and in vivo effects on Na+K+-ATPase activity and membrane fluidity. Toxicol. Lett. 85, 41-47. Tanner, M. J. A. (1997). The structure and function of band 3 (AE1): Recent developments Mol. Membr. Biol. 14, 155-165. Tanner, M. J. A., Martin, P. G., and High, S. (1988). The complete amino acid sequence of the human erythrocyte membrane anion-transport protein deduced from the cDNA sequence. Biochem. J. 256, 703-712. Tate, S. S., Yan, N., and Udenfriend, S. (1992). Expression cloning of a Na+-independent neutral amino acid transporter from rat kidney Proc. Natl. Acad. Sci. USA 89, 1-5. Tanner, M. J. A., Wainwright, S. D., and Martin, P. G. (1992). Structure, molecular genetics and topology of the human red cell anion transporter (AE1). Progr. Cell Res. 2, 17-22. Tarnuzzer, R. W., Campa, M. J., Qian, N. X., Englesberg, E., and Kilberg, M. S. (1990). Expression of the mammalian System A neutral amino acid transporter in Xenopus oocytes. J. Biol. Chem. 265, 13914-13917. Tate, C. G. (1998). Baculovirus-mediated expression of neurotransmitter transporters. In "Methods in Enzymology" (S. G. Amara, ed.), Vol. 296, pp. 443-445. Academic Press, San Diego. Tawa, N. E., and Goldberg, A. L. (1992). Suppression of muscle protein turnover and amino acid degradation by dietary protein deficiency. Am. J. Physiol. 263, E317-E325. Taylor, P. M., and Rennie, M. J. (1986). To what extent do kinetic properties of amino acid transporters in liver dictate the net exchange of amino acids in vivo? Biochem. Soc. Trans. 14, 1069-1070. Taylor, P. M., and Rennie, M. J. (1987). Perivenous localization of Na-dependent glutamate transport in perfused rat liver. FEBS Lett. 221, 370-374. Taylor, P. M., Hundal, H. S., and Rennie, M. J. (1989). Transport of glutamine in Xenopus laevis oocytes: Relationship with transport of other amino acids. J. Membr Biol. 112, 149-157. Taylor, P. M., Kemp, H. F., Davis, A., Birrell, J., and Peter, G. J. (1998). Aromatic amino acid transport in liver and kidney and possible links with transport of thyroid hormones. Nova Acta Leopoldina 78, 223-233. Taylor, P. M., Mackenzie, B., Low, S. Y., and Rennie, M. J. (1992). Expression of rat liver glutamine transporters in Xenopus laevis oocytes. J. Biol. Chem. 267, 3873-3877. Tessari, P., Garibotto, G., Inchiostro, S., Robaudo, C., Saffioti, S., Wettore, M., Zanetti, M., Russe, R., and Deferrari, G. (1996). Kidney, splanchnic and leg protein turnover in humans. Insights from leucine and phenylalanine kinetics. J. Clin. Invest. 98, 1481-1492.

References Tester, M. (1997). Techniques for studying ion channels: An introduction. J. Exp. Bot. 48, 353-359. Thibodeau, S. N., Bren, G., and Schaid, D. (1993). Microsatellite instability in cancer of the proximal colon. Science 60, 816-819. Thomas, E. L., and Christensen, H. N. (1970). Indications of spatial relations among structures recognizing amino acids and Na + at a transport receptor site. Biochem. Biophys. Res. Commun. 40, 277-283. Thomas, E. L., and Christensen, H. N. (1971). Nature of the cosubstrate action of Na + and neutral amino acids in a transport system J. Biol. Chem. 246, 1682-1688. Thorens B., Wu Y. J., Leahy J. L., and Weir G. C. (1992). The loss of GLUT2 expression by glucose-unresponsive/3 cells of db/db mice is reversible and is induced by the diabetic environment. J. Clin. Invest. 90, 77-85. Thureen, P. J., Narkewicz, M. R., Battaglia, F. C., Tjoa, S., and Fennessey, P. V. (1995). Pathways of serine and glycine metabolism in primary culture of ovine fetal hepatocytes. Pediatr. Res. 38, 775-782. Thwaites, D. T., Hirst, B. H., and Simmons N. L. (1994). Substrate specificity of the di/tripeptide transporter in human intestinal epithelia (Caco-2): Identification of substrates that undergo H+-coupled coupled absorption. Br J. Pharmacol. 113, 1050-1056. Thwaites, D. T., Markovich, D., Murer, H., and Simmons, N. L. (1996). Na+-independent lysine transport in human intestinal Caco-2 cells. J. Membr. Biol. 151, 215-224. Tien, X. Y., Brasitus, T. A., Kaetzel, M. A., Dedman, J. R., and Nelson, D. J. (1994). Activation of the cystic fibrosis transmembrane conductance regulator by cGMP in the human colonic cancer cell line, Caco-2. J. BioL Chem.

269, 51-54.

Tilly, B. C., Winter, M. C., Ostedgaard, L. S., O'Riordan, C., Smith, A. E., and Welsh, M. J. (1992). Cyclic AMPdependent protein kinase activation of cystic fibrosis transmembrane conductance regulator chloride channels in planar lipid bilayers. J. Biol. Chem. 267, 9470-9473. Toggenburger, G., H~iusermann, M., Mtitsch, B., Genoni, G., Kesssler, M., Weber, F., Hornig, D., O'Neill, B., and Semenza, G. (1981). Na+-dependent, potential-sensitive Lascorbate transport across brush border membrane vesicles from kidney cortex. Biochim. Biophys. Acta 646, 433-443. Tolner, B., Poolman, B., and Konings, W. N. (1992a). Characterization and functional expression in Escherichia coli of the sodium/proton/glutamate symport proteins of Bacillus stearothemophilus and Bacillus caldotenax. Mol. Microbiol. 6, 2845-2856. Tolner, B., Poolman, B., Wallace, B., and Konings, W. N. (1992b). Revised nucleotide sequence of the gltP gene, which encodes the proton-glutamate-aspartate transport protein of Escherichia coli K-12. J. Bacteriol. 174, 23912393. Tolner, B. Ubbink-Kok, T., Poolman, B., and Konings, W. N. (1995a). Characterization of the proton/glutamate symport protein of Bacillus subtilis and its functional expression in Escherichia coli. J. Bacteriol. 177, 2863-2869. Tolner, B., Ubbink-Kok, T., Poolman, B., and Konings, W. N. (1995b). Cation-selectivity of the L-glutamate trans-

379

porters of Escherichia coli, Bacillus stearothemophilus and Bacillus caldotenax: Dependence on the environment in which the proteins are expressed. Mol. Microbiol. 18, 123-133. Tomishige, M., Sako, Y., and Kusumi, A. (1998). Regulation mechanism of the lateral diffusion of Band 3 in erythrocyte membranes by the membrane skeleton. J. Cell Biol. 142, 989-1000. Tomita, Y., Takano, M., Yasuhara, M., Hori, R., and Inui, K. I. (1995). Transport of oral cephalosporins by the H+/ dipeptide cotransporter and distribution of the transport activity in isolated rabbit intestinal epithelial cells. J. Pharmacol. Exp. Therapeut. 272, 63-69. Toro, L., Ottolia, M., Stefani, E., and Latorre, R. (1994). Structural determinants in the interaction of Shaker inactivating peptide and a Ca2+-activated K + channel. Biochemistry 33, 7220-7228. Toyoshima, C., Sasabe, H., and Stokes, D. L. (1993). Threedimensional cryo-electron microscopy of the calcium ion pump in the sarcoplasmic reticulum membrane. Nature 362, 467-471. Toyofuku, T., Kurzydlowski, K., Narayanan, N., and MacLennan, D. H. (1994). Identification of Ser 38 as the site in cardiac sarcoplasmic reticulum Ca2+-ATPase that is phosphorylated by Ca2+/calmodulin-dependent protein kinase. J. Biol. Chem. 269, 26492-26496. Tozzo, E., Gnudi, L., and Kahn, B. B. (1997). Amelioration of insulin resistance in streptozotocin diabetic mice by transgenic overexpression of GLUT4 driven by an adiposespecific promoter. Endocrinology 138, 1604-1611. Travis, S. M., Berger, H. A., and Welsh, M. J. (1997). Protein phosphatase 2C dephosphorylates and inactivates cystic fibrosis transmembrane conductance regulator. Proc. Natl. Acad. Sci. USA 94, 11055-11060. Trotti, D., Rossi, D., Gjesdal, O., Levy, L. M., Racagni, G., Danbolt, N. C., and Volterra, A. (1996). Peroxynitrite inhibits glutamate transporter subtypes. J. Biol. Chem. 271, 5976-5979. Tsakiridis, T., McDowell, H. E., Walker, T., Downes, C. P., Hundal, H. S., Vranic, M., and Klip, A. (1995). Multiple roles of phosphatidylinositol 3-kinase in regulation of glucose-transport, amino-acid-transport, and glucose transporters in L6 skeletal-muscle cells. Endocrinology 136, 4315-4322. Tsao, T. S., Burcelin, R., Katz, E. B., Huang, L., and Charron, M. J. (1996). Enhanced insulin action due to targeted GLUT4 overexpression exclusively in muscle. Diabetes 45, 28-36. Tsao, T. S., Stenbit, A. E., Houseknecht, K. L., Zierath, J. R., Katz, E. B., and Charron, M. J. (1997). Muscle-specific transgenic complementation of GLUT4-deficient mice. Effects on glucose but not lipid metabolism. J. Clin. Invest. 100, 671-677. Ts6, C., Levine, S., Yun, C., Brant, S., Counillon, L., Pouyssegur, J., and Donowitz, M. (1993a). Structure/function studies of the epithelial isoforms of the mammalian Na+/ H + exchanger gene family. J. Membr. Biol. 135, 93-108. Ts6, C., Levine, S., Yun, C., Brant, S., Pouyssegur, J., Montrose, M. H., and Donowitz, M. (1993b). Functional charac-

380

References

teristics of a cloned epithelial Na+/H + exchanger (NHE3): Resistance to amiloride and inhibition by protein kinase C. Proc. Natl. Acad. Sci. USA 90, 9110-9114. Tseng-Crank, J. C. L., Tseng, G.-N., Schwartz, A., and Tanouye, M. A. (1990). Molecular cloning and functional expression of a potassium channel cDNA isolated from a rat cardiac library. FEBS Lett. 268, 63-68. Tsukaguchi, H., Shayakul, C., Berger, U. V., Mackenzie, B., Devidas, S., Guggino, W. B., van Hoek, A. N., and Hediger, M. A. (1998). Molecular characterization of a broad selectivity neutral solute channel. J. Biol. Chem. 273, 2473724743. Tsukamoto, H., Boado, R. J., and Pardridge, W. M. (1996). Differential expression in glioblastoma multiforme and cerebral hemangioblastoma of cytoplasmic proteins that bind two different domains within the 3'-untranslated region of the human glucose transporter 1 (GLUT1) messenger RNA. J. Clin. Invest. 97, 2823-2832. Turk, E., and Wright, E. M. (1997). Membrane topology motifs in the SGLT cotransporter family. J. Membr. Biol. 159, 1-20. Turk, E., Kerner, C. J., Lostao, M. P., and Wright, E. M. (1996). Membrane topology of the human Na+/glucose cotransporter SGLT1. J. Biol. Chem. 271, 1925-1934. Turner, R. J. (1990). Stoichiometry of coupled transport systems in vesicles. In "Methods in Enzymology," Vol. 191, pp. 479-494. Academic Press, San Diego. Uggeri, J., Belletti, S., Bussolati, O., Dall'Asta, V., and Gazzola, G. C. (1995). Suppression of anionic amino acid transport impairs the maintenance of intracellular glutamate in Ha-ras-expressing cells. Biochem. Biophys. Res. Commun. 211, 878-884. Ullrich, K. J. (1994). Specificity of transporters for 'organic anions' and 'organic cations' in the kidney. Biochim. Biophys. Acta 1197, 45-62. Ullrich, K. J., Rumrich, G., Neiteler, K., and Fritzsch, G. (1992). Contraluminal transport of organic cations in the proximal tubule of the rat kidney. Pfliigers Arch. 420, 29-38. Umbach, J. A., Coady, M. J., and Wright, E. M. (1990). Intestinal Na§ cotransporter expressed in Xenopus oocytes is electrogenic. Biophys. J. 57, 1217-1224. Umemoto, N., Ohya, Y., and Anraku, Y. (1991). V M A l l , a novel gene that encodes a putative proteolipid, is indispensable for expression of yeast vacuolar membrane H +ATPase activity. J. Biol. Chem. 266, 24526-24532. Unwin, N. (1995). Acetylcholine receptor channel imaged in the open state. Nature 373, 37-43. Ussing, H. H. (1949). The distinction by means of tracers between active transport and diffusion: The transfer of iodide across the isolated frog skin. Acta Physiol. Scand. 19, 43-56. Utsunomiya-Tate, N., Endou, H., and Kanai, Y. (1996). Cloning and functional characterization of system ASC-like Na+-dependent neutral amino acid transporter. J. Biol. Chem. 271, 14883-14890. Vadgama, J. V., and Christensen, H. N. (1984). Wide distribution of pH-dependent service of transport system ASC for

both anionic and zwitterionic amino acids. J. Biol. Chem. 259, 3648-3652. Vadgama, J. V., and Christensen, H. N. (1985). Discrimination of Na+-independent transport systems L, T and asc in erythrocytes: Na + independence of the latter a consequence of cell maturation. J. Biol. Chem. 260, 2912-2921. Vandenberg, R. J., Arriza, J. L., Amara, S. G., and Kavanaugh, M. P. (1995). Constitutive ion fluxes and substrate binding domains of human glutamate transporters. J. Biol. Chem. 270, 17668-17671. Vanderberg, R. J., Arriza, J. L., Amara, S. G., and Kavanaugh, M. P. (1997). Constitutive ion fluxes and substrate binding domains of human glutamate transporters. J. Biol. Chem. 270, 17668-17671. Van Dyke, R. W. (1996). Acidification of lysosomes and endosomes. In "Subcellular Biochemistry: Biology of the Lysosomes" (J. B. Lloyd and R. W. Mason, eds.), Vol. 27, pp. 331-360. Plenum, New York. van Helvoort, A., and van Meer, G. (1995). Intracellular lipid heterogeneity caused by topology of synthesis and specificity in transport. Example: sphingolipids. FEBS Lett. 369, 18-21. van Hille, B., Richener, H., Green, J. R., and Bilbe, G. (1995). The ubiquitous VA68 isoform of subunit A of the vacuolar H+-ATPase is highly expressed in human osteoclasts. Biochem. Biophys. Res. Commun. 214, 1108-1113. van Hille, B., Richener, H., Schmid, P., Puettner, I., Green, J. R., and Bilbe, G. (1994). Heterogeneity of vacuolar H § ATPase: Differential expression of two human subunit B isoforms. Biochem. J. 303, 191-198. van Iwaarden, P. R., Pastore, J. C., Konings, W. N., and Kaback, H. R. (1991). Construction of a functional lactose permease devoid of cysteine residues. Biochemistry 30, 9595-9600. van Monfort, R. L. M., Pijning, T., Kalk, K. H., Reizer, J., Saier, M. H., Jr., Thunnissen, M. M. G. M., Robillard, G. T., and Dijkstra, B. W. (1997). The structure of an energy-coupling protein from bacteria, IIB cell~176 reveals similarity to eukaryotic protein tyrosine phosphatases. Structure 5, 217-225. Van Slyke, D. D., and Meyer, G. M. (1913-1914). The fate of protein digestion products in the body. III. The absorption of amino acids from the blood by the tissues. J. Biol. Chem. 16, 197-212. Van Winkle, L. J. (1988). Amino acid transport in developing animal oocytes and early conceptuses. Biochim. Biophys. Acta 947, 173-208. Van Winkle, L. J. (1992). Amino acid transport during embryogenesis. In "Mammalian Amino Acid Transport: Mechanisms and Control" (M. S. Kilberg and D. H~iussinger, eds.), pp. 75-87. Plenum, New York. Van Winkle, L. J. (1993). Endogenous amino acid transport systems and expression of mammalian amino acid transport proteins in Xenopus oocytes. Biochim. Biophys. Acta 1154, 157-172. Van Winkle, L. J. (1999). Genetic regulation of expression of intestinal biomembrane transport proteins in response to dietary protein, carbohydrate and lipid. In: "Current Topics

References in Membranes: Gastrointestinal Transport." (K. E. Barret and M. Donowitz, eds.), Academic Press, San Diego (in press). Van Winkle, L. J., and Campione, A. L. (1987). Development of amino acid transport system B ~ in mouse blastocysts. Biochim. Biophys. Acta 925, 164-174. Van Winkle, L. J., and Campione, A. L. (1990). Functional changes in cation-preferring amino acid transport during development of preimplantation mouse conceptuses. Biochim. Biophys. Acta 1028, 165-173. Van Winkle, L. J., and Campione, A. L. (1991). Ouabain sensitive 86Rb+ uptake in mouse eggs and preimplantation conceptuses. Dev. Biol. 146, 158-166. Van Winkle, L. J., and Campione, A. L. (1992). Novel bumetanide-sensitive K § transport in preimplantation mouse conceptuses. Am. J. Physiol. 263, C773-779. Van Winkle, L. J., and Campione, A. L. (1996). Amino acid transport regulation in preimplantation mouse embryos: Effects on amino acid content and pre- and peri-implantation development. Theriogenology 45, 69-80. Van Winkle, L. J., Campione, A. L., Mann, D. F., and Wasserlauf, H. G. (1993). The cation receptor subsite of the choline transporter in preimplantation mouse conceptuses resembles a cation receptor subsite of several amino acid transporters. Biochim. Biophys. Acta 1146, 38-44. Van Winkle, L. J., Christensen, H. N., and Campione, A. L. (1985). Na+-dependent transport of basic, zwitterionic, and bicyclic amino acids by a broad-scope system in mouse blastocysts. J. Biol. Chem. 260, 12118-12123. Van Winkle, L. J., Campione, A. L., and Gorman, J. M. (1988a). Na+-independent transport of basic and zwitterionic amino acids in mouse blastocysts by a shared system and by processes which distinguish between these substrates. J. Biol. Chem. 263, 3150-3163. Van Winkle, L. J., Haghighat, N., Campione, A. L., and Gorman, J. M. (1988b). Glycine transport in mouse eggs and preimplantation conceptuses. Biochim. Biophys. Acta 941, 241-256. Van Winkle, L. J., Campione, A. L., Gorman, J. M., and Weimer, B. D. (1990a). Changes in the activities of amino acid transport system b ~247and L during development of preimplantation mouse conceptuses. Biochim. Biophys. Acta 1021, 77-84. Van Winkle, L. J., Mann, D. F., Campione, A. L., and Farrington, B. H. (1990b). Transport of benzenoid amino acids by system T and four broad scope systems in preimplantation mouse conceptuses. Biochim. Biophys. Acta 1027, 268-277. Van Winkle, L. J., Campione, A. L., and Gorman, J. M. (1990c). Inhibition of transport system b ~247in blastocysts by inorganic and organic cations yields insight into the structure of its amino acid receptor site. Biochim. Biophys. Acta 1025, 215-224. Van Winkle, L. J., Campione, A. L., and Farrington, B. H. (1990d). Development of system B ~ and a broad-scope Na+-dependent transporter of zwitterionic amino acids in preimplantation mouse conceptuses. Biochim. Biophys. Acta 1025, 225-233. Van Winkle, L. J., Haghighat, N., and Campione, A. L. (1990e). Glycine protects preimplantation mouse concep-

381

tuses from a detrimental effect on development of inorganic ions in oviductal fluid J. Exp. Zool. 253, 215-219. Van Winkle, L. J., Iannaccone, P. M., Campione, A. L., and Garton, R. L. (1990f). Transport of cationic and zwitterionic amino acids in preimplantation rat conceptuses. Dev. Biol. 142, 184-193. Van Winkle, L. J., Kakuda, D. K., and MacLeod, C. L. (1995). Multiple components of transport are associated with murine cationic amino acid transporter (mCAT) expression in Xenopus oocytes. Biochim. Biophys. Acta 1233, 213-216. Van Winkle, L. J., Mann, D. F., Weimer, B. D., and Campione, A. L. (1991). Na+-Dependent transport of anionic amino acids by preimplantation mouse blastocysts. Biochim. Biophys. Acta 1068, 231-236. Van Winkle, L. J., Mann, D. F., Wasserlauf, H. G., and Patel, M. (1992). Mediated Na+-independent transport of Lglutamate and L-cystine in 1- and 2-cell mouse conceptuses. Biochim. Biophys. Acta 1107, 299-304. Van Winkle, L. J., Patel, M., Wasserlauf, H. G., Dickinson, H. R., and Campione, A. L. (1994). Osmotic regulation of taurine transport via system beta and novel processes in mouse preimplantation conceptuses. Biochim. Biophys. Acta 1191, 244-255. Vasilets, L. A., and Schwarz, W. (1993). Structure-function relationships of cation binding in the Na+/K+-ATPase. Biochim. Biophys. Acta 1154, 201-222. Vaughn, P. R., Lobo, C., Battaglia, F. C., Fennessey, P. V., Wilkening, R. B,. and Meschia, G. (1995). Glutamineglutamate exchange between the placenta and fetal liver. Am. J. Physiol. 268, E705-E711. Vaughan, R. A., Huff, R. A., Uhl, G. R., and Kuhar, M. J. (1997). Protein kinase C-mediated phoshorylation and functional regulation of dopamine transporters in striatal synaptosomes. J. Biol. Chem. 272, 15541-15546. Verkman, A. S. (1995). Optical methods to measure membrane transport processes. J. Membr. Biol. 148, 99-110. Verkman, A. S., Van Hoek, A. N., Ma, T., Frigeri, A., Skach, W. R., Mitra, A., Tamarappoo, B. K., and Farinas, J. (1996). Water transport across mammalian cell membranes. Am. J. Physiol. 270, C12-C30. Verstergaard, H., Weinreb, J. E., Rosen, A. S., Bjorbaek, C., Hansen, L., Pedersern, O., and Kahn, B. B. (1995). Sulfonylurea therapy improves glucose disposal without changing skeletal muscle GLUT4 levels in noninsulindependent diabetes mellitus subjects: A longitudinal study. J. Clin. Endocrinol. Metab. 80, 270-275. Vilch6ze, C., McMullen, T. P. W., McElhaney, R. N., and Bittman, R. (1996). The effect of side-chain analogues of cholesterol on the thermotropic phase behavior of 1-stearoyl-2-oleoylphosphatidycholine bilayers: A differential scanning calorimetric study. Biochim. Biophys. Acta 1279, 235-242. Vilsen, B., Andersen, J. P., and MacLennan, D. H. (1991). Functional consequences of alterations to amino acids located in the hinge domain of the Ca2+-ATPase of sarcoplasmic reticulum. J. Biol. Chem. 266, 16157-16164. von Heijne, G. (1994). Membrane proteins: From sequence to structure. Annu. Rev. Biophys. Biomol. Struct. 23,167-192.

382

References

Wadiche, J. I., Amara S. G., and Kavanaugh, M. P. (1995a). Ion fluxes associated with excitatory amino acid transport. Neuron 15, 721-728. Wadiche, J. I., Arriza, J. L., Amara, S. G., and Kavanaugh, M. P. (1995b). Kinetics of a human glutamate transporter. Neuron 14, 1019-1027. Wahle, S., and Stoffel, W. (1996). Membrane topology of the high-affinity L-glutamate transporter (GLAST-1) of the central nervous system. J. Cell Biol. 135, 1867-1877. Wahren, J. (1988). Extra-hepatic ammonia metabolism. In "Advances in Ammonia Metabolism and Hepatic Encephalopathy" (P. B. Soeters, J. H. P. Wilson, A. J. Meijer, and E. Holm, eds), pp. 121-129. Elsevier, Amsterdam. Wakabayashi, S., Bertrand, B. Ikeda, T., Pouyssegur, J., and Shigekawa, J. (1994). Mutation of calmodulin-binding site renders the Na+/H + exchanger (NHE1) highly H+-sensitive and C a 2+ regulation defective. J. BioL Chem. 269, 1371013715. Wakabayashi, S., Fafournoux, P., Sardet, C., and Pouyssegur, J. (1992). The Na+/H + antiporter cytoplasmic domain mediates growth factor signals and controls H+-sensing. Proc. Natl. Acad. Sci. USA 88, 2424-2428. Wakabayashi, S., Shigekawa, M., and Pouyssegur, J. (1997). Molecular physiology of vertebrate Na+/H + exchangers. Physiol. Rev. 77, 51-74. Walbot, V., and Cullis, C. A. (1985). Rapid genomic change in higher plants. Ann. Rev. Plant Physiol. 36, 367-396. Wallmeier, H., Weber, A., Gross, A., and Fltigge, U.-I. (1992). Insights into the structure of the chloroplast phosphate translocator protein. In "Transport and Receptor Proteins of Plant Membranes" (D. T. Cooke and D. T. Clarkson, eds.), pp. 77-89. Plenum, New York. Wang, H., Kavanaugh, M. P., North, R. A., and Kabat, D. (1991). Cell-surface receptor for ecotropic murine retroviruses is a basic amino-acid transporter. Nature 352, 729-731. Wang, C. C., Moriyama, R., Lombardo, C. R., and Low, P. S. (1995). Partial characterization of the cytoplasmic domain of human kidney band 3.J. Biol. Chem. 270,1789217897. Wang, D. N. (1994). Band 3 protein: Structure, flexibility and function. FEBS Lett. 346, 26-31. Wang, D. N., Ktihlbrandt, W., Sarabia, V. E., and Reithmeier, R. A. F. (1993). Two-dimensional structure of the membrane domain of human band 3, the anion transport protein of the erythrocyte membrane. E M B O J. 12, 2233-2239. Wang, D. N., Sarabia, V. E., Reithmeier, R. A. F., and Ktihlbrandt, W. (1994). Three-dimensional map of the dimeric membrane domain of the human erythrocyte anion exchanger, band 3. E M B O J. 13, 3230-3235. Wang, Y., and Tate, S. S. (1995). Oligomeric structure of a renal cystine transporter: implications in cystinuria. FEBS Lett. 368, 389-392. Wang, Z., Schultheis, P. J., and Shull, G. E. (1996). Three Nterminal variants of the AE2 C1-/CHO3- exchanger are encoded by mRNAs transcribed from alternative promoters. J. Biol. Chem. 271, 7835-7843. Watanabe, S.-I., Kusama-Eguchi, K., Kobayashi, H., and Igarashi, K. (1991). Estimation of polyamine binding to macro-

molecules and ATP in bovine lymphocytes and rat liver. J. Biol. Chem. 266, 20803-20809.

Waterlow, J. C. (1995). Whole-body protein turnover in humansmPast, present and future. Annu. Rev. Nutr. 15, 57-92. Waterlow, J. C., Garlick, P. J., and Millward, D. J. (1978). "Protein Turnover in Mammalian Tissues and in the Whole Body." Elsevier/North-Holland, Amsterdam. Watford, M. (1993). Hepatic glutaminase expression: Relationship to kidney-type glutaminase and to the urea cycle. FASEB J. 7, 1469-1474. Weber, J., and Senior, A. E. (1997). Catalytic mechanism of F1-ATPase. Biochim. Biophys. Acta 1319, 19-58. Weimbs, T., Low, S. H., Chapin, S. J., and Mostov, K. E. (1997). Apical targeting in polarized epithelial cells: There's more afloat than rafts. Trends Cell Biol. 7, 393-399. Weisiger, R. A. (1985). Dissociation from albumin: A potentially rate-limiting step in the clearance of substances by the liver. Proc. Natl. Acad. Sci. USA 82, 1563-1567 Weisinger, R. A. (1996). When is a carrier not a membrane carrier? The cytoplasmic transport of amphipathic molecules. Hepatology 24, 1288-1295. Welbourne, T. C. (1987). Interorgan glutamine flow in metabolic acidosis. Am. J. PhysioL 253, F1069-F1076. Welbourne, T. C. (1988). Role of the lung in glutamine homeostasis. Contrib. Nephrol. 63, 178-182. Welbourne, T. C., and Dass, P. D. (1988). Gamma glutamyltransferase contribution to renal ammoniagenesis in vivo. Eur. J. Physiol. (Pflagers Arch.) 411, 573-578. Wells, R. D. (1996). Molecular basis of genetic instability of triplet repeats J. Biol. Chem. 271, 2875-2878. Welsh, M. J., Tsui, L.-C., Boat, T. F., and Beaudet, A. L. (1995). Cystic fibrosis. In "The Metabolic and Molecular Basis of Inherited Disease." ( C. R. Scriver, A. L. Beaudet, W. S. Sly, and D. Valle, eds.), pp. 3799-3876. McGraw-Hill, New York. Wenzel, U., Gebert, I., Weintraut, H., Weber, W. M., Clauss, W., and Daniel, H. (1996). Transport characteristics of differently charged cephalosporin antibiotics in oocytes expressing the cloned intestinal peptide transporter PepT1 and in human intestinal Caco-2 cells. J. PharmacoL Exp. Therapeut. 277, 831-839. Weston, K., Hall, J. L., and Williams, L. E. (1994). Characterization of a glutamine/proton cotransporter from Ricinus communis roots using isolated plasma membrane vesicles. Physiol. Plant. 91, 623-630. Wheeler, K. P., and Christensen, H. N. (1967). Interdependent fluxes of amino acids and sodium ion in the pigeon red blood cell. J. Biol. Chem. 242, 3782-3788. Wheeler, K. P., Inui, Y., Hollenberg, P. F., Eavenson, E., and Christensen, H .N. (1965). Relation of amino acid transport to sodium-ion concentration. Biochim. Biophys. Acta 109, 620-622. White, J. H., Kilbey, B. J., de Vries, E., Goman, M., Alano, P., Cheesman, S., McAleese, S., and Ridley, R. G. (1993). The gene encoding DNA polymerase a from Plasmodium falciparum. Nucleic Acids Res. 21, 3643-3646. White, M. F., and Christensen, H. N. (1982). The two-way flux of cationic amino acids across the plasma membrane

References of mammalian cells is largely explained by a single transport system. J. Biol. Chem. 257, 10069-10080. White, S.H. (1992). Amino acid preferences of small proteins: Implications for protein stability and evolution. J. Mol. Biol. 227, 991-995. White, S. H. (1994a). Global statistics of protein sequences: Implications for the origin, evolution, and prediction of structure. Ann. Rev. Biophys. Biomol. Struct. 23, 407-439. White, S. H. (1994b). The evolution of proteins from random amino acid sequences: II. Evidence from the statistical distributions of the lengths of modern protein sequences. J. Mol. Evol. 38, 383-394. White, S. H., and Jacobs, R.E. (1990). Statistical distribution of hydrophobic residues along the length of protein chains: Implications for protein folding and evolution. Biophys. J. 57, 911-921. White, S. H. and Jacobs, R. E. (1993). The evolution of proteins from random amino acid sequences. I. Evidence from the lengthwise distribution of amino acids in modern protein sequences. J. Mol. Evol. 36, 79-95. Whitesell, R. R., Tarpley, H. L., and Regen, D. M. (1977). Sugar-transport kinetics of the rat thymocyte. Arch. Biochem. Biophys. 181, 596-602. Wilbrandt, W. (1955). Secretion and transport of non-electrolytes. Symp. Soc. Exp. Biol. 8, 136-162. Wilkinson, D. J., Strong, T. V., Mansoura, M. K., Wood, D. L., Smith, S. S., Collins, F. S., and Dawson, D. C. (1997). CFTR activation: Additive effects of stimulatory and inhibitory phosphorylation sites in the R domain. Am. J. Physiol. 273, L127-L133. Williams, B. D., Wolfe, R. R., Bracy, D. P., and Wasserman, D. H. (1996). Gut proteolysis contributes essential amino acids during exercise, Am. J. Physiol. 270, E85-E90. Williams, K. (1997a). Modulation and block of ion channels: A new biology of polyamines. Cell Signal 9, 1-13. Williams, K. (1997b). Interactions of polyamines with ion channels. Biochem. J. 325, 289-297. Williamson, P., and Schlegel, R. A. (1994). Back and forth: The regulation and function of transbilayer phospholipid movement in eukaryotic cells. Mol. Membr. Biol. 11, 199-216. Wilson, D. B. (1978). Cellular transport mechanism. Annu. Rev. Biochem. 47, 933-965. Windmueller, H. G., and Spaeth, A. E. (1974). Uptake and metabolism of plasma glutamine by the small intestine. J. Biol. Chem. 249, 5070-5079. Winter, C. G., and Christensen, H. N. (1964). Migration of amino acids across the membrane of the human erythrocyte. J. Biol. Chem. 239, 872-878. Winter, C. G., and Christensen, H. N. (1965). Contrasts in neutral amino acid transport by rabbit erythrocytes and reticulocytes. J. Biol. Chem. 240, 3594-3600. Winter, M. C., and Welsh, M. J. (1997). Stimulation of CFTR activity by its phosphorylated R domain. Nature 389, 294-296. Wojtczak, L., Wieckowski, M. R., and Schtinfeld, P. (1998). Protonophoric activity of fatty acid analogs and derivatives in the inner mitochondrial membrane: A further argument

383

for the fatty acid cycling model. Arch. Biochem. Biophys. 357, 76-84. Wolf, G. (1997). A new uncoupling protein: A potential component of the human body weight regulation system. Nutr. Rev. 55, 178-179. Wood, P.G. (1992). The anion exchange proteins: Homology and secondary structure. Progr. Cell Res. 2, 325-352. Woodgett, J. R. (1990). Molecular cloning and expression of glycogen synthase kinase-3/factor A. E M B O J. 9, 24312438. Woodlock, T. J., Young, D. A., Boal, T. R., Lichtman, M. A., and Segel, G. B. (1993). Phorbol ester-induced membrane proteins in chronic leukemic B-lymphocytes. Candidate proteins for the L-system amino acid transporter. J. Biol. Chem. 268, 16020-16027. Woolford, J., McAuliffe, A., and Rohrschneider, L. R. (1988). Activation of the feline c-fms proto-oncogene: Multiple alterations are required to generate a fully transformed phenotype. Cell 55, 965-977. Woolf, C. M. (1968). "Principles of Biometry: Statistics for Biologists," 1st ed. Van Norstrand, London. Wright, J. K. (1989). Product inhibition during ion: Solute cotransport is an alternative to leaks as a cause of low accumulations. J. Membr. Biol. 109, 1-8. Xu, A., Hawkins, C., and Narayanan, N. (1993). Phosphorylation and activation of the Ca2+-pumping ATPase of cardiac sarcoplasmic reticulum by Ca2+/calmodulin-dependent protein kinase. J. Biol. Chem. 268, 8394-8397. Xu, Z. C., Yang, Y., and Hebert, S. C. (1996). Phosphorylation of the ATP-sensitive, inwardly rectifying K + channel, ROMK, by cyclic AMP-dependent protein kinase. J. Biol. Chem. 271, 9313-9319. Yamada, H., Moriyama, Y., Maeda, M., and Futai, M. (1996). Transmembrane topology of Escherichia coli H+-ATPase (ATP synthase) subunit a. FEBS Letts. 390, 34-38. Yamamoto T., Seino Y., Fukumoto H., Koh G., Yano H., Inagaki N., Yamada Y., Inoue K., Manabe T., and Imura H. (1990). Over-expression of facilitative glucose transporter genes in human cancers. Biochem. Biophys. Res. Commun. 170, 223-230. Yang, B., Van Hoek, A. N., and Verkman, A. S. (1997). Very high single channel water permeability of aquaporin-4 in baculovirus-infected insect cells and liposomes reconstituted with purified aquaporin-4. Biochemistry 36, 7625-7632. Yang, X. C., Karschin, A., Labarca, C., Elroy-Stein, O., Moss, B., Davidson, N., and Lester, H. A. (1991). Expression of ion channels and receptors in Xenopus oocytes using vaccinia virus. FASEB J. 5, 2209-2216. Yano, H., Philipson, L. H., Kugler, J. L., Tokuyama, Y., Davis, E. M., Le Beau, M. M., Nelson, D. J., Bell, G. I., and Takeda, J. (1994). Alternative splicing of human inwardly rectifying K + channel ROMK1 mRNA. Mol. Pharmacol. 45, 854-860. Yano, M., Marinelli, R. A., Roberts, S. K., Balan, V., Pham, L., Tarara, J. E., de Groen, P. C., and LaRusso, N. F. (1996). Rat hepatocytes transport water mainly via a nonchannel-mediated pathway. J. Biol. Chem. 271, 6702-6707. Yanbo, H., Philipson, L. H., Kugler, J. P., Tokuyama, Y., Davis, E. M. et al. (1994). Alternative splicing of human

384

References

inwardly rectifying K + channels ROMK1 mRNA. Mol. Pharmacol. 45, 854-860.

Yao, S. Y. M., Muzyka, W. R., Cass, C. E., Cheeseman, C. I., and Young, J. D. (1999). The BAT and 4F2hc amino acid transport proteins induce transport of uridine and pyruvate in Xenopus oocytes. Biochem. Cell Biol. (in press). Yao, Y., and Tsien, R. Y. (1997). Calcium current activated by depletion of calcium stores in Xenopus oocytes. J. Gen. Physiol. 109, 703-715. Yau, K. W., McNaughton, P. A., and Hodgkin, A. L. (1981). Effect of ions on the light-sensitive current in retinal rods. Nature 292, 502-505. Yelin, R., and Schuldiner, S. (1998). Purification of the vesicular monoamine transporters: From classical techniques to histidine tags. In "Methods of Enzymology" (S. G. Amara, ed.), Vol. 296, 64-72. Academic Press, San Diego. Yellen, G. (1997). Single channel seeks permeant ion for brief but intimate relationship. J. Gen. Physiol. 110, 83-85. Yi, S. J., Liu, S.-C., Derick, L. H., Murray, J., Barker, J. E., Cho, M. R., Palek, J., and Golan, D. E. (1997). Red cell membranes of ankyrin-deficient nb/nb Mice lack band 3 tetramers but contain normal membrane skeletons. Biochemistry 36, 9596-9604. Yoda, S., and Yoda, A. (1987). Phosphorylated Intermediates of Na,K-ATPase proteoliposomes controlled by bilayer cholesterol. J. Biol. Chem. 262, 103-109. Yokoyama, K., Muneyuki, E., Amano, T., Mizutani, S., Yoshida, M., Ishida, M., and Ohkuma, S. (1998). V-ATPase of Thermus thermophilus is inactivated during ATP hydrolysis but can synthesize ATP. J. Biol. Chem. 273, 2050420510. Younes M., Lechago L. V., Somoano J. R., Mosharaf M., and Lechago J. (1996). Wide expression of the human erythrocyte glucose transporter Glut1 in human cancers. Cancer Res. 56, 1164-1167. Young, J. D., Fincham, D. A., and Harvey, C. M. (1991). Cation and harmaline interactions with Na+-independent dibasic amino acid transport system y+ in human erythrocytes and in erythrocytes from a primitive vertebrate the pacific hagfish (Eptatretus stouti). Biochim. Biophys. Acta 1070, 111-118. Young, J. D., Mason, D. K., and Fincham, D. A. (1988). Topographical similarities between harmaline inhibition sites on Na+-dependent amino acid transport system A S C in human erythrocytes and Na+-independent system asc in horse erythrocytes. J. Biol. Chem. 263, 140-143. Yu, W., Xu, J., and Li, M. (1996). NAB domain is essential for the subunit assembly of both ~-~ and c~-/3 complexes of Shaker-like potassium channels. Neuron 16, 441-453. Yuan, G.-F., Fu, Y.-H., and Marzluf, G.A. (1991). nit-4, a pathway-specific regulatory gene of Neurospora crassa, encodes a protein with a putative binuclear zinc DNA-binding domain. Mol. Cell Biol. 11, 5735-5745. Yudkoff, M. (1997). Brain metabolism of branched-chain amino acids. Glia 21,92-98. Yun, C. H. C., Tse, C.-M., Nath, S. K., Levine, S. A., Brant, S. A., and Donowitz, M. (1995). Mammalian Na+/H + exchanger gene family. Am. J. Physiol. 269, G 1 - G l l .

Zabicipril, Y., Bendahan, A., Zarbiv, R., Kavanaugh, M. P., and Kanner, B. I. (1998). Molecular determinant of ion selectivity of a (Na + + K+ )-coupled rat brain glutamate transporter. Proc. Natl. Acad. Sci. USA 95, 751-755. Zabicipril, Y., Pines, G., and Kanner, B. I. (1994). Histidine 326 is critical for the function of GLT-1, a (Na + + K +)coupled glutamate transporter from rat brain. J. Biol. Chem. 269, 19573-19577. Zacharias, D. A., Dalrymple, S.J., and Strehler, E. E. (1995). Transcript distribution of plasma membrane C a 2+ pump isoforms and splice variants in human brain. Mol. Brain Res. 28, 263-272. Zachowski, A. (1993). Phospholipids in animal eukaryotic membranes: Transverse asymmetry and movement. Biochem. J. 294, 1-14. Zagotta, W. N., and Siegelbaum, S. A. (1996). Structure and function of cyclic nucleotide-gated channels. Annu. Rev. Neurosci. 19, 235-263. Zagotta, W. N., Hoshi, T., and Aldrich, R. W. (1990). Restoration of inactivation in mutants of Shaker potassium channels by a peptide derived from ShB. Science 250, 568-571. Zakim, D. (1996). Fatty acids enter cells by simple diffusion. Proc. Soc. Exp. Biol. Med. 212, 5-14. Zampighi, G. A., Kreman, M., Boorer, K. H., Loo, D. D. F., Bezanilla, F., Chandy, G., Hall, J. E., and Wright, E. M. (1995). A method for determining the unitary functional capacity of cloned channels and transporters expressed in Xenopus laevis oocytes. J. Membr. BioL 148, 65-78. Zavodnik, I. B., Lapshina, E. A., Palecz, D., and Bryszewska, M. (1996). The effects of palmitate on human erythrocyte membrane potential and osmotic stability. Scand. J. Clin. Lab Invest. 56, 401-407. Zavodnik, I. B., Piasecka, A., Szosland, K., and Bryszerwka, M. (1997). Human red blood cell membrane potential and fluidity in glucose solutions. Scand. J. Clin. Lab. Invest. 57, 59-63. Zeng, H., Parthasarathy, R., Rampal, A. L., and Jung, C. Y. (1996). Proposed structure of putative glucose channel in GLUT1 facilitative glucose transporter. Biophys. J. 70, 14-21. Zerangue, N., and Kavanaugh, M. P. (1996a). Flux coupling in a neuronal glutamate transporter. Nature 383, 634-637. Zerangue, N., and Kavanaugh, M. P. (1996b). ASCT-1 is a neutral amino acid exchanger with chloride channel activity. J. Biol. Chem. 271, 27991-27994. Zhang, L., Coffey, L. L., and Reith, M. E. (1997). Regulation of the functional activity of the human dopamine transporter by protein kinase C. Biochem. Pharmacol. 53, 677-688. Zhang, P., Toyoshima, C., Yonekura, K., Green, N. M., and Stokes, D. L. (1998). Structure of the calcium pump from sarcoplasmic reticulum at 8-A resolution. Nature 392, 835-839. Zhang, Y. and Fillingame, R. H. (1994). Essential aspartate in subunit c of FIFo ATP synthase: Effect of position 61 substitutions in helix-2 on function of Asp24 in helix-1. J. Biol. Chem. 269, 5473-5479. Zhang, Y., and Fillingame, R. H. (1995). Subunits coupling H + transport and ATP synthesis in the Escherichia coli ATP synthase. J. Biol. Chem. 270, 24609-24614.

References

Zhang, Y., Bendahan, A., Zarbiv, R., Kavanaugh, M. P., and Kanner, B. I. (1998). Molecular determinant of ion selectivity of a (Na + + K+ )-coupled rat brain glutamate transporter. Proc. Natl. Acad. Sci. USA 95, 751-755. Zhang, Y., Oldenburg, M., and Fillingame, R. H. (1994). Suppressor mutations in F1 subunit e recouple ATP-driven H+translocation in uncoupled Q42E subunit c mutant of the Escherichia coli F1Fo ATP synthase. J. Biol. Chem. 269, 10221-10224. Zharikov, S. I., Herrara, H., and Block, E. R. (1997). Role of membrane potential in hypoxic inhibition of L-arginine uptake by lung endothelial cells. Am. J. Physiol. 272, ,L78-L84. Zhou, H., Tate, S. S., and Palmer, L. G. (1994). Primary structure and functional properties of an epithelial K channel. Am. J. Physiol. 266, C809-C824. Zhou, Q., Zhao, J., Stout, J. G., Luhm, R. A., Wiedmer, T., and Sims, P. J. (1997). Molecular cloning of human plasma membrane phospholipid scramblase. J. Biol. Chem. 272, 18240-18244.

385

Zhou, Y., Samson, M., Francon, J., and Blondeau, J.-P. (1992). Thyroid hormone concentrative uptake in rat erythrocytes. Biochem. J. 281, 81-86. Zhou, Y., Samson, M., Osty, J., Francon, J., and Blondeau, J.-P. (1990). Evidence for a close link between the thyroid hormone transport system and the aromatic amino acid transport system T in erythrocytes. J. Biol. Chem. 265, 17000-17004. Zhu, S. J., Kavanaugh, M. P., Sonders, M. S., Amara, S. G., and Zahniser, N. R. (1997). Activation of protein kinase C inhibits uptake, currents and binding associated with the human dopamine transporter expressed in Xenopus oocytes. J. Pharmacol. Exp. Therapeut. 282, 1358-1365. Ziel, F., Venkatasen, N., and Davidson, M. B. (1988). Glucose transport is rate limiting for skeletal muscle glucose metabolism in normal and STZ-induced diabetic rats. Diabetes 37, 885-890. Zimmerman, A. L., and Baylor, D. A. (1992). Cation interactions within the cyclic GMP-activated channel of retinal rods from the tiger salamander. J. Physiol 449, 759-783.

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Index

A AB part of the qualitative ABC test, 126, 131 AB part of the quantitative ABC test, 130, 131 AB portion of the ABC test, 94, 95, 109, 110, 112 AB testing, 93-95, 109, 110 ABC testing, 126-131 ABC transporter superfamily, 275 Academic disciplines, 10 Accessory proteins, 111-114, Acetylcholine-gated ion channels, 19 Acholeplasma laidlawii cells, 19 Acidification of membrane-bound compartments, 58 Acidosis, 320, 322, 323 Actin filaments, 33, 35 Active transport, 49 Active transporters, driven by decarboxylation, 268, 270 diphosphate bond hydrolysis, 268, 269 electron flow, 268 light, 268, 270 methyltransfer, 270, 271 oxidoreduction, 270 mechanically, 270, 271 Activity coefficient, 40 Adaptive regulation, 285, 302-303 Adipose tissue, 298, 314 AE proteins, see also Anion exchanger as channels for organic osmolytes, 233-237 as C1- channels, 233-237 in regulation of metabolism, 233-234 structural function, 233-234 in volume regulation, 3-237 AE1 as a component of a megadalton, complex, 233 AE1 deficient mice and cattle, 171, 181,233 AE1 dimers, 233 AE1 extracellular Z-loop in channel function, 235-236 AE1 in fish red blood cells, 235

AE1 physiological function, 170-172, 233-234 AE1 tetramers, 231 AE2 evolution through trinucleotide expansions affecting Z-loop length, 235 Affinity constant, 90 Alanine, 298, 307, 310, 311,313, 314, 320, 321, 324 Albumin effects on fatty acid transport kinetics, 79 Aldolase, 233 Algal cell membranes, 74, 75, 77 Allosteric effectors, 233 Allosteric effects, 202, 218 Allosteric interactions, 283 Allosteric protein conformation free energy, 89 Allosteric regulation, 122 Allosteric transporter regulation, 170 Alpha subunit of Na § ATPase, 111 Alpha subunit subisoforms, 163 Alternative functions of transport proteins, 237 Alternative promoters in AE genes, 181-184 Alternative splicing and F-N-type ATPases, 163-166 of AE transcripts, 181-184 of P-type ATPase transcripts, 139, 141-144 of Type IIA SERCA ATPase, 142, 144 Amino acids, 295-325 clusters and protein evolution, 184-193 distribution ratio, 297, 307 exchange, 237 interorgan fluxes, 298 as metabolic signals, 303-304 sequestration, 124 structural diversity, 111 in study of transport heterogeneity, 103 transport, 115 Amino acid transport systems, 94, 103, 112, 300-302

387

Amino acid transporters, 103 as C1- channels, 233, 235-237 Amino acid transport-related proteins, 103 Amino acid whole body kinetics, 297, 306 Amino transferase reaction, 88 Ammonia, 298, 312, 313, 320, 322 Amphipathic helices, 137 Amphipathic stalk sequence, 137 Amphipathy, 13-15 Amphipatic phospholipids, 25 Analog inhibition analysis, 104, 107, 108, 111, 125 competing transport processes, 102, 103 studies, 126-131 Analysis of curve shape, 105 Anion exchange importance of effective anion concentrations, 205-207 multiple components of transport, 201-202 Anion exchanger, 27, 29, 170-207, 233-237; see also AE copies per cell, 170-171 allosteric effects, 201-207 allosteric function, 204 and alternative splicing and promoters, 181-184 anion pathway structure, 195-199 asymmetric function, 199, 204 conformational free energy difference, 201 conformations, 196-199 C-terminal associated domain, 172, 193-207 evolution, 172-184 evolution through trinucleotide expansions, 184 function in erythrocytes, 170-172 isoform and subisoform tissue distributions, 181-184 isoforms and subisoforms, 172-184 kinetics of transport, 199-207 models of function, 204-207 N-terminal cytosolic domain, 172-184 oligomerization, 199

388 Anion exchanger ( c o n t i n u e d ) positive and negative cooperativity, 201-207 sequence alignments, 172-180 structure/function, 194-207 Anion migration, 44 Anionic amino acid transporter, see Excitatory amino acid transporter; EAAT Anionic amino acid-stimulated C1conductance, 237 Ankyrin, 27, 145 Antiport, 4, 59-61, 83, 89, 101, 117, 119, 121-123, 145; see also Exchange simultaneous and ping-pong mechanisms, 170 Antiporters, 4, 83, 89, 119-121, 169-207 Apical membrane, 18, 30, 32-35, 120 Apical recycling compartment, 37 Apical sorting, 35, 37 Apical sorting signal, 35 Apical trafficking, 38 Apoptotic lymphocytes, 23 Aquaporin-1, 19, 21, 22 Aginine 299, 311,315-317 Arginine transport by system b ~ 81, 83 Arginine transport kinetics, 83 Aromatic amino acid, 316, 318 Arterial branch points, 14 Artificial bilayer, 29 Artificial proteoliposomes, 30 ASC and BAT-associated transport, widely variable stoichiometry, 231 ASC catalyzed Na + exchange, 80-fold increase owing to proline hydroxylation, 216 ASC catalyzed obligatory exchange, structural basis, 227-228, 230 ASC coupling ratio, 15-fold increase owing to proline hydroxylation, 216 ASC hydrophobicity, 211 ASC obligatory exchange, 212 ASC protein structure, 211-212 ASC substrate stoichiometry, 211-217 ASC systems and transporters, 211 ASC transport coupling ratios, 215-216 ASC transport kinetics, 212-217 ASC transport model, 215-217 ASC transport, asymmetric substrate treatment, 215 ASC transport, preferential unloading of one cosubstrate, 215 ASC transport, widely variable coupling ratio, 215-217 ASC transporter cycling, 80-fold increase owing to proline hydroxylation, 216 ASC/EAAT family, 231 ASC1 zwitterionic amino acid transport, 330, 331 Aspartate amino transferase, 79, 81 Asymmetric biochemical reaction, 88 Asymmetric biomembrane structure, 20 Asymmetric biomembranes, 89, 90 Asymmetric carbohydrate distribution, 23

Index Asymmetric conformations, 89, 90 Asymmetric fluidity, 25, 29, 38 Asymmetric K + transport via Kir channels, 253-254 Asymmetric lipid bilayer, 33 Asymmetric lipid distribution, 21, 23, 29 Asymmetric orientations of membrane proteins, 20, 21, 88, 89, 90, 169 Asymmetric phospholipid distribution, 38 Asymmetric reconstitution, 29 Asymmetric transport, 88 Asymmetric transporter function, 88 Atherosclerosis, 13, 14 ATPase orientation in biomembranes, 135 ATP-binding region, 139 ATP-binding site, 138 ATP hydrolysis, 49, 134, 136 ATP synthesis, 56, 57, 134 and electrically neutral phosphoric acid, 162-163 Auto phosphorylation of P-type ATPases, 146-152

B Band 3 protein see Anion exchanger Band 4.2 protein, 27 Basolateral endosomes, 35 Basolateral membrane, 18, 30, 32-35 Basolateral signals, 37 Basolateral sorting, 35, 37 Basolateral sorting signal, 36 BAT expression, 120, 121,230, 231 X e n o p u s oocytes, 222 BAT protein, 115 BAT subfamily, 229 BAT-associated transport system b ~ 120 BCECF cell volume measurement using, 339 intracellular pH measurement using, 339 Benzenoid amino acid transport, 112 Benzenoid amino acids, 110 Best fit of transport data to a hyperbola, 85 r subunit of Na+K+ATPase, 111, 112 B-barrel structure, 21, /3-galactoside transporter, 116 Bicyclic amino acid analogs, 109 Bilayers, 19, 29, 32 composition, 30 structure, 30 surface, 14, 16, 18 Binding change model, 158 for F-type ATPases, 159-163 of thermodynamic coupling, 160 Biocatalysts, 1 Bioenergetic coupling, 20, 50-52, 55, 59, 61, 64, 135 of ATP synthesis to proton transport, 158 Bioenergetically coupled processes, 49 Bioenergetically distinct conformations, 170 Bioenergetics of coupling, 134 of transport, 43

Biomembranes asymmetry, 1, 2, 19, 169 barriers, 1-3, 38, 42, 61, 86, 134, 239-240, 299 to lipophilic substances, 73-79 to simple diffusion, 66, carbohydrate composition, 23 component, 134, 135 composition 13, 30, 32 electrical potential, 221; see also Electrical membrane potential fluidity, 26, 53 leaflet, 15, 31 melting temperatures, 16 phospholipid composition, 23, 24 protein content, 19 reconstitution, 29 sector, 134, 137 skeleton, 33 skeleton fence model, 27 structure, 13, 30, 32 transport is not simple diffusion, 66, 67 of fatty acids, 79 of lipid-soluble substances, 69, 73-79 of water, 68, 69 process composition, 111 turnover, 13 Biomembrane-associated domains, 137, 138 of AE proteins, 194 Biomembrane-spanning helices, 137 Biomembrane-spanning segments, 19 Blastocyst, 8, 110, 112, 113 Blood, 305, 308, 310, 324 Blood-brain barrier, 299, 315, 316 b~ protein (BAT), 120 Brain, 314, 316, 324 Branched-chain amino acid, 314-316, 320, 321,324 Brown adipose tissue, 5 Brush border, 120 Brush border membrane vesicles, 222 Budding, 31

C C portion of the ABC test, 94, 95, 109, 130 C a 2+ ATPases, 59-61, 139, 141 molecule, 134 C a 2+ binding by SERCA ATPases, 146 C a 2+ containing compartments, 144 CaZ+-affinity mutants, 146 Calmodulin, 36 Calmodulin-binding domains, 144 of PMCA ATPases, 141-143 Calmodulin-binding domains, 144 Calmodulin-dependent protein kinase, 142 cAMP-dependent AE2b regulation, 183 Cancer, 323, 325 Carbohydrate synthesis, 23 Carbohydrate-rich zone, 24 Carbonic anhydrase, 170, 171,233 Cardiac glycoside, 233 Carrier, 65

Index Carrier kinetics, 91 Carrier-type facilitators, 267 Carrier-type transporters, 269 CAT subfamily, 229 CAT transport proteins, 232, 315, 316 Catalytic mechanisms, 135 Catalytic sites of channels, carriers, and enzymes, 65 Cation facilitator (CF) family ,272, 273 Cation inhibition of system b ~24796, 102 Cation migration, 44 Cation receptor subsite, 7 Cation selected to replace Na § 102 Cationic amino acid transport, 103 Cationic amino acid transporter (CAT), 231 Cationic amino acids, 121 Cell cycle, 287 Cell stretch, 303 Cell swelling and shrinking regulates metabolism, 72 Cell volume, 287, 303, 307, 321 Cell-cell interaction, 24 cell-matrix interaction, 24 Cellular volume regulation, 234, 237 Central dogma, 1 Ceramide, 24 Cerebroside, 18 CFTR, see Cystic fibrosis conductance regulator Channel and carrier proteins, possibly similar functional mechanisms, 258-262 Channel catalyzed transport, 65 Channel gating, 65 Channel genes in Drosophilia, 240-241 Channel inhibitors, 93 Channel protein conformational changes during transport, 239, 249, 257, 258, 260 Channel protein structure, function and evolution, 240-254 Channel proteins and dissipation of solute gradients, 239-263 Channel Q10 values, 239 Channel selectivity, 65 Channel substrate selectivity, 240 Channel transport flux coupling, 258-260 Channel transport is substrate-saturable, 81 Channel-catalyzed transport, 239-263 Channel-like transport of CI-, 63 Channels, 61, 65, 83, 90, 91, 119 Channels and diffusion, 239-240 Channels for inorganic anions, 98 Channel-type facilitators, 267 Channel-type transporters, 269 Chaperone, 32 Characteristics of inhibition, 106 of substrate metabolism, 99 of substrate transport, 99 of transport, 116 of transport activities, 103, 107-111 Chemical equilibrium, 41 Chemical potential, 40, 45, 64 change, 41, 42

differences, 44 gradient, 39, 43, 64 Chemical reaction, 43 Chemotherapy, 325 CHIP28, 19, 21, 22 Chlamydia trachomatis, 35, 36 Cholesterol, 15, 16, 18, 24 Choline inhibition of transport, 102 Choline substitution for Na+, 102 Choline transport, 115 Choline transport system, 114 Chorioallantoic placenta, 9, 10 cis-Golgi, 18, 30 cis-inhibition, 118-120, 123-125, 199 cis-stimulation, 119, 121 cis-trans isomerization, 140 C1- channels, 233, 235-237 C1C-3 in volume regulation, 235 transport kinetics, 82 conductance, 236 Classification of transport proteins, 265-276 system for, 267-272 Cleavage-stage conceptuses, 8, 110 Cloning, expression, 328-329 Closed system, 39 Coagulation, 23 Compartmentation, metabolic, 295, 296 Competing substrates, 81 interact in ten ways, 104-107 reveal transport heterogeneity, 104 Competition of solutes for transport, 93 Competitive inhibition, 86, 92-95, 97, 103, 104, 127, 130, 200 Complete genome analyses, 1,265 Complex inhibition, 96 Complex kinetics, 115 Confidence intervals, 86Conformation and free energy changes, 89 Conformation of transport-related proteins, 114-116 Conformational changes, 21, 43, 64, 89, 119, 134, 139, 152, 341 definition, 239 in Fo and F1 sectors, 160 of channel proteins, 239, 249, 257, 258, 260 Conformationally mobile kinks, 140 Consensus model, 137 Contamination of commercial preparations, 96 Cooperative functioning of P- and F-type ATPases, 166-167 Cooperative kinetics, unreliability for determination of transport stoichiometry, 217 Correction for nonsaturable uptake, 101 Cortical granule exocytosis, 22 Cotransport, 4, 57, 91,122 Countertransport, see Antiport; Exchange Coupled processes, 43, 61 Coupled transport, 4, 98, 121, 122 and chemical change, 134 uncoupling, 121 Cryoelectron crystallography, 19

389 Crystalline domain, 170 Crystalline lipid domains, 16, 17, 19, 20, 26, 28, 29, 37, 38, 53-55, 64, 148, 151, 167 Crystalline phase, 29 Crystalline state, 16 C-subunit isoforms, 165 C-terminal membrane-associated domain, 140 Current, 221 Current reversal, 121,221 Cut-open oocytes, 120 Cysteine, 318 Cystic fibrosis conductance regulator, 278-280, 284 Cystine, 317-319 Cystinuria, 317, 324 Cytokines, 14 Cytoskeletal interactions with AE proteins, 233-234 Cytoskeletal meshwork, 27, 28, 38 Cytoskeleton, 13, 15, 23, 24, 26-30, 33, 34, 37, 38, 145, 172, 233, 237 Cytosolic component of P-type ATPases, 139 Cytosolic domain, 137 Cytosolic enzymes, 237 Cytosolic protein domains, 34 Cytosolic regions and domains, 138 Cytosolic sectors, 137 Cytotoxic T lymphocytes, 25

D DCT1, see Metal-ion transport Deduction of nonsaturable transport, 100 Depolarization, definition, 241 Detection of multiple transport activities, 103, 104 Developmental regulation of expression of transport processes, 114 Diabetes mellitus, 287-290, 323 Diffusion, 2, 3, 47, 61,124 and channels, 239-240 halting by biomembranes, 64 kinetics, 67 of lipophilic substances, 73-79 not a mechanism of biomembrane transport, 66, 67 rate, exceeds that of transport, 67 of water, 68 Diffusion coefficient (D), 67, 68 in biomembranes, 70 not a mechanism of biomembrane transport, 66, 67 Diffusional permeability coefficients do not reflect simple diffusion, 73 Diffusional water permeability coefficients (Pd), 70, 71 Dimerization, 116 Dimers, 122 Dinucleotide expansions, 188, 189 Dinucleotide repeat sequences, 193 Diphosphadtylglycerol, 18 Dissociation constant (Kd), 90, 93, 125

390

Index

Dixon plot, 91, 92, 94, 95, 200 DNA polymerase slippage, 187 Dog red cell membrane, 75, 77 Domain C, 137-139 Donnan effect, 45 Drosophila channel genes, 240-241

E E1P-to-E2P transition, 150 EAAT (excitatory amino acid transporters) and ASC subfamily evolution, 208-211 co-substrate binding order, 221-223 EAAT1 glutamate transporter, 330, 331 EAAT2 subisoforms, 210 EAAT3 glutamate transporter, 338, 341 EAAT 4 and 5, as anionic amino acid activated C1- channels, possible effect of extracellular K + in brain, 236 EAAT-catalyzed homo- and heteroexchange, 224 evolution of K + countertransport, 208, 210 glycosylation scanning studies, 218 hydrophobicity plots, 212, 218 membrane topology, 218 mutant forms catalyze obligatory anionic amino acid exchange, 226-228 ping-pong model, 224 protein kinase, 218 protein structure, 217-218 proteins and some forms of system b ~ exchange amino acids for K +, 229 regulatory phosphorylation sites, 218 simultaneous two-site model, 224 substrate stoichiometry, 217-224 symporters, propagation of a Na + gradient into gradients of anionic amino acids, 217-225 transport kinetics, 218-225 model, 221-225 competition between K + and AA/H+/ Na +, 219-220, 223 K + or Na+/anionic amino acids are substrates, 217-225 EAAT/ASC and systems b ~ asc and y+, similarity of cation binding sites, 231-233 evolution of C1- channel transport, 210 protein sequence similarities, 208 proteins, 208-237 proteins as C1- channels, 233, 235-237 sequence alignments, 208 site directed mutagenesis, 225-228 substrate stoichiometry, 211-224 transport characteristics resemble system asc, b ~ and y+, 229 transport mechanism, 225-228 EAAT/ASC transporters, relationship to bacterial anionic amino acid and dicarboxylate transporters, 225-226, 228 Eadie-Hofstee plot, 86, 87

Effective anion concentrations, 206, 207 Effective extracellular K § concentration, 236 Effective intracellular polyamine concentration, 251 Effective ion concentrations, 207, 220, 222, 236 Effective ion concentrations and the membrane electrical potential, 260-262 EGF, see Epidermal growth factor Electric field, 134 Electrical current, 113 Electrical membrane potential, 21, 43, 45-47, 49-51, 59, 61, 64, 113, 121, 205-207, 221 effect on effective ion concentration, 205-207, 260-262 Electrical potential work, 62 Electrochemical potential, 302, 310 Electrogenic biomembrane transport, 113, 205 Electrogenic exchange, 222 Embryo, 8, 111 Embryo implantation, 8 Endergonic processes, 49 Endergonic transport processes 51 Endocytic vesicle, 58 Endocytosis, 13, 23, 31 Endogenous transport-related proteins, 113 Endolysosome, 30 Endoplasmic reticulum, 18, 30, 31 Endosome, 18, 30, 58 Endothelial cells, 13, 14 Energy conversion during transport, 217 Enthalpy change, 42, 43, 64 Enthalpy-driven freezing of lipid domains, 64 Enthalpy-driven processes, 39, 43, 44, 64 Entropy change, 42, 43, 64 Entropy of the membrane, 54 Entropy-driven melting of lipid domains, 64 Entropy-driven processes, 14, 39, 43, 44, 55, 64 Environmental effects on transport, 8 Enzyme inhibition, 93 Epidermal growth factor, 278 Epithelia, 120 Epithelial cell polarity biogenesis 32 Epstein-Barr virus neutral antigen, 330, 331 Equilibrium constant, 50 Erythrocytes, 23, 27 plasma membrane, 26, 74, 77 network, 233 two-dimensional elasticity, 233 structure and function, 233 volume, 76 Evolution and classification of F- and V-type ATPases, 152-155 during stress, 191 of transport proteins, 265-267 through trinucleotide expansions, 184-193

of voltage-gated channels, 240-246, 249-250 Exchange, 4, 118, 122, 125; see also Antiport nonobligatory, 118, 119, 125 obligatory, 118-121,123-125 Excitatory amino acid transporter, 63, 98, 208-211,217-237 282-283, 285, 287; see also EAAT Exercise, 320 Exergonic process, 49 Exocytosis, 13, 31 Expression cloning, 113 Expression of transport-related proteins in X e n o p u s oocytes, 115 Expression systems, heterologous, 329-332 Extracellular milieu, 58 Extramembrane components, 135, 138 Extramembrane sectors, 134, 137

F 4F2hc protein, 114, 115 F sector, 152 F-/V-type ATPases, 134-137 c~-,/3-, A-, B- and c-subunits, 153 evolution and classification, 152-155 isoforms, subisoforms and alternative splicing, 163-166 subunit stoichiometries, 152-156 superfamily, 272, 274 F1 sector conformational changes, 158 of F-type ATPases, 152 hydrophobic sleeve, 162-164 sector nucleotide binding sites, 158 subunit structure, 158 X-ray crystallographic studies, 158 Facilitated diffusion, 64, 83 Faraday constant, 44 Fatty acids binding to albumin, 79 flip-flop, 81 metabolism does not correlate with transport, 79 migration across biomembranes, 79 transport is not simple diffusion, 81 kinetics, 80 substrate saturability, 79 transporter, 79, 80 Fatty acid-binding protein, 79 Fatty acyl membrane composition, 26 Fatty streak, 14 Fetus, 9, 10, 320 First law of thermodynamics, 39 Fitting of experimental data to kinetic formulations, 85, 86 Flippases, 31, 33 Fluid mosaic model, 20, 24, 27, 28, 38 Fluid state, 16, 17, 19 Fluorescence techniques, 339 Flux and flux coupling via K + channels, 256-262

Index Flux coupling and flux ratio, 258-260 mechanism for channels, symporters and primary active transporters, 258-260 Flux ratio, 230 and flux coupling, 258-260 Fo c-subunit oligomer, 155 Fo sector of F-type ATPase, 152 Fo sector subunits, 155 FoF1-ATP synthases, s e e F-type ATPases Free energy, 5, 21 of activation, 239 alternate conformational states, 149 of ATP hydrolysis, 49-51, 55, 64, 117, 125 of ATPase conformational changes, 167 barrier, 25 biomembrane transport, 63, 64 change, 41-43, 64 for cotransport, 59 for coupled processes, 56 in a chemical reaction, 39 in migration of a solute, 39 of spontaneous processes, 40 catalyzed by Na § § 51 chemical sources, 133 conformational change, 133 conversions, 59 coupling, 133, 152 driven transport, 89 nucleotide binding 152 in phosphoric acid anhydride bond, 56 position of a solute, 64 of position of liquid and crystalline membrane lipid domains, 64 in proton gradients, 56 of proton transport, 158 of solute gradients, 125 in total chemical potential gradients, 63 transfer, 134, 148, 149, 166, 169 between solute gradients, 170 during the EIP-to-E2P transition, 150 of transition state, 170 of transport 49-51, 55, 117 of transporter conformations, 89, 169 Free fatty acid, 320 Freeze fracture, 20 F-type ATPase, 55-57, 133, 152, 159 F-type ATPase beta-subunit conformational changes, 161 binding change model, 159-163 bioenergetic coupling, 161 conformational changes, 152, 166-167 cytosolic sector (F1), 158 F1 sector structure/function, 155-158, 158-160 Fo, 152 y-subunit rotation, 161 subisoforms, 163-164 headpiece, basepiece and stalk region, 158 membrane sector (Fo), 155 nucleotide binding site, 159

pathway for proton transport, 155 proton pathway, 156-158 site-directed mutagenesis, 156, 159-161 structure, 152 structure/function, 166-167 subunits a and c, 156-158 Function, as a basis for protein classification, 266 Futile cycling, 118

G GABA transport, GAT1 transporter, 330, 334-335 Galactosyl ceramide, 18 Gel state, 16, 17, 19 Gene knockout, 341 Genome sequencing, 265 Gephyrin, 27 Gibbs-Donnan effect, 45, 46, 47, 49, 51, 57, 58, 61 Gibbs-Donnan, 51, 59, 62, 260 Glucagon, 278, 284 Gluconeogenesis, 314, 320, 321 Glucose, 314, 320 Glucose binding sites, 122 Glucose transporters (GLUTs), 19, 21, 93, 122, 233, 287-291 SGLT1 and SGLT2 Na+/glucose cotransporters, 329-330, 333-335, 334-341 Glucose-alanine cycle, 314 GLUT, s e e Glucose transporters Glutamate, 298, 313, 314, 320, 322; s e e a l s o EAAT Glutamine, 287, 298, 299, 307, 309, 310-314, 320-325 Glutamine-glutamate cycle, 310, 313, 314, 320 Glutaminyl clusters, 185 y-Glutamyltransferase, 322 Glutathione, 122, 319, 323, 325 Glycine, 320 Glycocalyx, 26, 76 Glycolipid rafts, 28, 38 Glycolipid synthesis, 25 Glycolytic enzymes, 233 Glycoprotein interactions, 28 Glycoprotein synthesis, 25 Glycosphingolipid rafts, 28, 37, 38 Glycosphingolipids, 33 Glycosylation, 24 Glycosyl-phosphatidyl-inositol-linked proteins, 28, 37, 38 Golgi, 30, 31, 58 Gradients of solute/solvent, 40 Gramicidin channels, 86 Growth factors, 14

H H § transport, 139 H+K+ATPase, 283 a- and/3-subunit isoforms, 136, 144, 145 trafficking, 145

391 Half-maximal transport velocity, 81, 82 Hanes plot, 86, 87 Harmaline, 97 Hartnup disorder, 324 Heat, 42 HEK 293 cell line, heterologous expression in, 330, 331 HeLa cells, 226, 227, 228 heterologous expression in, 329-330, 331 Helical wheel, 138 Helices, 147-148 Hemocompatible surfaces, 23 Hemoglobin, 233, 234 Hemolysin, 3, 4, 239 Hemolysin channel-forming toxin (HL) family, 272, 273 Hemolytic anemia, 171 Heteroexchange, 118 Heterogeneity of amino acid transport, 103 Heterogeneous biomembrane structure, 20 Heterologous transport related proteins, 113 Hexadecane/water partition coefficients, 74-79 Hexanucleotide expansions, 189 Hexanucleotide repeats, 192 Hill coefficient, 144 Hinge region, 137 Histidyl reactive agent (DEPC), 196, 198 Hofstee plot, 86, 87, 91, 92, 103,104, 116, 201,204-206, 212 Homoexchange, 118 Homologous proteins, 266 Hormonal regulation, 284 Human erythrocyte, 76 Human erythrocyte plasma membrane, 74, 77 Human neurodegenerative disorders, 187 Hydrogen-bonded chain, 157, 199 Hydrogen-bonded series of functional groups, 199 Hydrogen bonding in substrate transporter interactions, 88 Hydropathy analysis, 211,212 Hydrophilic amino acid residues, 137 Hydrophilic extrasystolic loop, 137 Hydrophilic side-chain, 138 Hydrophilic solutes, 125 Hydrophlic signaling molecules, 2 Hydrophobic binding site 57 Hydrophobic environment 134 Hydrophobic hydrocarbon chains, 16, Hydrophobic interactions between substrates and transport proteins, 15, 88 Hydrophobic region of a membrane, 69 Hydrophobic region of the bilayer, 70 Hyperbolas, 84, 87, 117 sum of 105, 106 Hyperbolic curve, 206 Hyperbolic kinetics, 202, 204, 212, 214, 221, 237 Hyperpolarization, definition, 241 Hyperpolarized membrane, 121

392

Index

Hyperpolarized oocytes, 121 Hypertension, 13 Hypotonic stress, 235

J

Jellyfish channel proteins, 240

transport and total chemical potential, 261-262 transport of lipid-soluble substances, 69 trans-stimulation, 119

K 1 Immobile proteins, 26 Impermeant anion, 45, 47 Infinite trans experiments, 84 Influenza virus hemagglutinin, 33 Inhibition analyses, 126-131 Inhibition heterogeneity, 105-107 Inhibition homogeneity, 105-106 Inhibition of alternate transport processes, 99 complete, 105-107, 131 incomplete, 105-108, 127, 128 partial, 108, 127, 128, 131 selective, 110 weak, 108, 127, 128, 131 Inhibitor concentrations, 96 Inhibitor protonation/deprotonation, 96 Initial velocity, 83, 99-102 Injury, 323 Inner leaflet, 23, 25, 26, 29, 31-33 Inorganic cation transport, 133 Inorganic ion transport, 134 Inorganic ion-dependent transport, 101 Inositol, 18 Insertion into lipid bilayers, 141 Insertion into membranes, 141 Insulin, 302, 321,323 Integral membrane protein insertion, 32 Integral membrane proteins, 19-21, 30 Integrin, 27, 29, 303 Interactions between substrates and transporters, 86 Interdisciplinary science, 10 Intermediary metabolism, 237 Interorgan nutrient flows and transport, 295-326 Intestinal epithelia, 119, 120 Intestine, 298, 312, 313, 320 Intracellular pH recording, 338-339 Intracellular substrate concentration, 99 Intracellular water, 76 Intramembrane ion binding sites, 139 Inwardly rectifying K + (Kir) channels, 80, 124, 249-254 Ion migration, 44 Ionic interactions between substrates and transport proteins, 88 Ion-selective microelectrodes, 338-339 Iron transport, see Metal-ion transport Isoforms, 141 Isolation of inorganic ion-dependent transport processes, 101-103 Isolation of inorganic ion-independent transport, 102 Isolation of transport activities, 103, 104, 107-111

K + channels, 61, 62, 86, 136, 240-263 flux and flux coupling, 256-262 pore-forming (P) regions (H5 loops), 247, 254-257 structure at 3.2,~ resolution, 240, 248, 259, 260 substrate selectivity, structural basis, 254-256 transport kinetics, 82, 254, 256-262 K + channel proteins, 237 site-directed mutagenesis, 254-256 K + channel-related proteins of prokaryotes, 241 K + flux and flux coupling via K + channels, 256-262 K + ion migration against its total chemical potential gradient, 90 K + transport against its gradient by Kir channels, 253-254 KdpK + ATPase, 136, 138 Ki values, 91-96, 106, 108, 109, 112, 123-125 Kidney, 298, 313, 317-320, 322 Kinesin like motor proteins, 35 Kinetic analyses, 98, 104 Kinetic constants, 119 Kinetic dependencies of amino acid transport on Na + concentration, 214 Kinetic differences among transport processes that form, propagate and dissipate solute gradients, 116 Kinetic formulations of transport, 83, 91 Kinetic mechanisms of transport, 170 Kinetic model of cotransport, 91 Kinetic modeling 117 Kinetic models, 116 Kinetic parameters, 84-86, 88-90, 99, 125 Kinetically distinct components of transport, 103, 104, 114 Kinetics anion exchange, 199-207 antiport, 117, 118 ASC-catalyzed symport and antiport, 212-217 cis-inhibition, 119 cis-stimulation, 119

diffusion, 67 EAAT transport, 218-225 fatty acid biomembrane transport, 79 K + channel transport, 254, 256-262 nonobligatory exchange, 118, 119 obligatory exchange, 118, 119 primary active transport, 117 saturable transport, 83 symport, 117 symport/antiport, 170 trans-inhibition, 119

transport, 65-131,207

uniport, 118, 119 voltage-gated ion channels, 254 Kir channels evolution, 249-250 functional dependence on polyamines, 250-254 homo- and hetero-tetramers, 250 mechanism of inhibition by polyamines, 251,256 structure and function, 247, 249-254 transport against the K + total chemical potential gradient, 253-254 Km values, 84-88, 90, 94-96, 108-110, 112, 115, 116, 123-125 for mediated transport, 100 Km/Vmaxvalues, 88 Kv channels alpha- and beta- subunits, 242-243 C-type inactivation, 248, 249 evolution, 240-246 evolution through trinucleotide expansion, 241-246 hetero- and homooligomers, 241 isoforms and subisoforms, 241-247 mechanism of voltage-gating, 243-248 N-terminus (N-type) inactivation, 243, 248 N-terminus and C-terminus lengths, 243 N-terminus and rapid inactivation, 241-242 RNA editing, 241 structure and function, 241-249 tetrameric structure and function, 243, 247-249 Kvl channel protein sequence alignments, 244-246

L Lactation, 320 Lactose transporter, 32 Lateral diffusion, 26 Laws of thermodynamics, 88 Leaflet, 15, 31 Leak-free condition of biomembranes, 61 Leaks, 59, 61, 62, 98 Leucine, 307, 324 Li + substitution for Na +, 102 Linear plot, 87 Linear transformations, 87, 103 Linear transformations of kinetic data, 86 Lineweaver-Burk plot, 80, 86, 87, 103 Lipid bilayer, 13, 15, 16, 20, 27-30, 33, 38; see also Phospholipid bilayer asymmetry, 33 barrier, 74, 79 Lipid composition, 29 Lipid domains, see Phospholipid domains Lipid molecules, 16, 18

Index Lipid motion, 24, 26, 27 Lipid phase transitions, 64 Lipid solubilities, 74, 75, 77, 78 Lipid sorting, 33, 37, 38 Lipid-soluble substances, 74, 78 Lipophile migration across biomembranes, 73-79 Lipophilic solutes, 124 Liposomes, 29, 116 Liquid lipid domains, 16, 17, 20, 26, 28, 29, 37, 38, 53-55, 64, 148, 151, 167, 170 Liquid phase, 29, 53 Liquid state, 16 Liquid-crystalline state, 16, 17, 19, 26 Liver, 305, 306, 309-314, 316, 318, 319, 321, 323 Liver periportal hepatocytes, 298, 310, 312, 313 Liver perivenous hepatocytes, 298, 312, 313 Lumenal leaflet, 33 Lumenal protein domains, 35 Lung, 298, 313, 314, 323 Lymphocyte plasma membrane, 26 Lymphocytes, 323 Lysosomal membrane, 30 Lysosomes, 9, 18, 58

M Macrophage, 14, 23 Major facilitator superfamily, 275 Mannitol substitution for Na+, 102 Maple syrup disease (MSD), 324 Mathematical expressions for simple diffusion, 67 Maximum velocity, s e e Vmax Maxwell's demon, 89 Medial-Golgi, 18, 30 Mediated transport, 83 Membrane, s e e Biomembrane Metabolic control theory, 308, 309 Metabolic effects on transport, 99 Metabolic functions of AE proteins, 233-234 Metabolic products of the substrate, 100 Metabolism-resistant substrate analogs, 100 Metal-ion transport, DCT1 H+/metal-ion cotransporter, 328, 334, 338 Mg2+ ATPase, 29, 31 Michaelis-Menten constant, s e e Km Michaelis-Menten equation, 84, 85, 87 Michaelis-Menten kinetics, 83, 117, 125, 160, 199, 200, 203 microsatellite DNAs, 188 Microtubule polymerization/ depolymerization, 43 Microtubules, 33-35 Migration of a solute against a total chemical potential gradient, 117 Mismatch repair, 187 Mitochondrial membrane, 30, 32 Mitochondrion, 5, 18 Mixed inhibition, 96

Mobile carrier model, 118 Molecular volume, 74 Monocytes, 13, 14, 23 Monomeric exchange, 30, 31 Monosialoganglioside, 18 Motor proteins, 27, 31, 33-35, 38 Multidisciplinary science, 10 Multimeric structure, 133 Multiple components of transport, 103, 104, 115, 116, 126-131 Multiple shared transport processes, 94 Multiple transport processes, 98 Muscle, 305, 307, 311-314, 320, 321,323 Mutagenesis studies of AE1,195-199 of K + channels, 254-256 Mutual inhibition, 131

393 Nonparametric statistical method, 86 Nonpermeant ions, 49 Nonsaturable component of transport, 108 Nonsaturable exodus, 101 Nonsaturable transport, 81, 101, 105, 126 Nonsaturable transport deduction, 100 Nonsaturable uptake, 101 Novel K + transporter, 114 N-terminal membrane-associated domain, 139 Nucleic acids, 1 Null phenotype cell line, 196 Nutrition, interorgan, 295-325 Nystatin, 200

O N Na + channel transport kinetics, 82 Na + inhibition of system b ~ 96, 102 Na + pump, 62 Na+//3-alanine symport, 212 Na+/amino acid coupling ratios, 215, 216 Na+/amino acid symporters, 208 Na+/Ca e+ exchange, 59-61 Na+/glucose symporter, 98 Na+/glutamate symport, 218 Na+/glycine symport, 212 Na+/H + exchangers, 280-283 Na+-dependent amino acid transport, 101 Na+-dependent component, 102 Na+-independent component, 102 Na+K+ATPase, 49, 59-62, 114, 233 a subunit, 111, 145 a and/3-subunit isoforms, 144, 145 c~ and/3-subunits, 111, 112, 136 (a/3)e tetrameres, 152 catalyzed processes, 52 conformations, 52-55, 62 tetrameres, 136 trafficking, 145 Negative cooperativity, 116, 160, 200-205, 237 Negative surface potential, 97 Nernst equation, 46, 260 Net flux, 4 N-ethylmaleimide (NEM), 111 Neural cell adhesion molecule, 28 Neutrophils, 24 N-glycosylation sites, 2 1 2 NHE s e e Na+/H + exchangers Nitric oxide, 315, 316 N-methyl-L-tryptophan, 110, 112, 113 Noncompetitive inhibition, 91, 92, 94, 97, 104, 127 Noncompetitive inhibitors, 200 Nonlamellar lipid structures, 13 Nonlinear plot, 87 Nonlinear regression analysis, 85, 103 Nonobligatory exchange, 118

Obligator exchange and the Nernst equation, 260-261 Obligatory exchange, 89, 118, 123 resembles a Gibbs-Donnan equilibrium 260-261 Obligatory exchanger, 199 Oligomeric structures, 134 Oligomerization, 136, 283 of P-type ATPases, 144, 145 of transport proteins, 116, 170 Oligomers, 122 Oligosaccharides assembly, 23 moieties, 24, 25, 29 interactions, 28 Olive oil/water partition coefficients, 74 One-cell embryo, 112 Organic osmolyte transport, 237 Orientation in proteoliposomes, 116 Osmolyte substitution, 102 Osmolytes, 286, 287 Osmotic gradients, 124 Osmotic pressure, 47-49 Osmotic regulation of water flux, 71, 72 Osmotic water permeability coefficient (Pf), 70, 71 Ouabain binding site, 144 Outer leaflet, 21, 23, 25, 26, 29, 31-34 Ovalocytosis, 27 Overtraining syndrome, 321

P p70s6 kinase, 304 Packing of transmembrane helices, 147 Parallel/3 sheet structure, 138 Parametric and nonparametric statistical tests, 85 Parenteral nutrition, 324 Partition coefficient (Kp), 69, 74-77 PEP-dependent phosphoryl transfer, 268 Perforin, 25 Peripheral proteins, 19, 20 Permeability, 2 upper limit, 78

394 Permeability coefficient (P), 68-70, 74-77 correction for size, 74-79 do not usually reflect permeation of the lipid bilayers, 73 not corrected for size, 77, 78 Permeable membrane, 46 Permeant ions, 48, 49 Permeation of biomembrane lipid layers is not ordinary diffusion, 66, 67 Peroxisomal membranes, 30, 32 Peroxisome, 18 pH gradient, 56 Phase boundary, 54 Phase transition, 16, 17, 19, 26, 29, 55, 64 Phase transition temperature, 53 Phenylketonuria (PKU), 324 Phorbols, 280-283 Phosphate transport, 285-286 Phosphatidylcholine, 18, 21-24, 26, 29, 33 Phosphatidylethanolamine, 18, 21, 23, 24, 29, 32, 33 Phosphatidylserine, 18, 21-26, 29, 33 Phosphocholine, 24 Phospholipids asymmetry, 31 bilayer, 13, 20, 29, 30, 76, 78, 134; s e e a l s o Lipid bilayer barrier, 74 distribution, 24 domain phase transition, 150, 152 domains, 16, 17, 19, 20, 26, 28, 29, 37, 38, 148, 170 exchange proteins, 31 head groups, 16 monomeric exchange, 30, 31 motion, 26 synthesis, 31 transbilayer transport, 31 Phosphoric acid anhydride bonds, 44, 49 Phosphoryl-accepting aspartyl residue, 138 Phosphorylation of SERCA ATPases, 138 sites, 138, 218 of P-type ATPases, 147 and transport regulation, 278-283 Phylogeny as a basis for protein classification, 266-267 Physiological substrate concentrations, 81, 82, 85 Ping-pong and two-site simultaneous model combined, 204-207 Ping-pong model, 118, 119, 125, 199, 200, 237 Ping-pong transport model of antiport, 170 PKA, s e e Protein kinase A PKC, s e e Protein kinase C Placenta, 9, 10, 319, 320 Plant Kir channel, 240-241 Plasma membrane, 13, 24, 30, 31 Plasma membrane fatty acid binding protein, 79, 80 Platelets, 22, 23 PMCA ATPases, 141 dimers, 136 subisoforms, 136

Index

Polyamines effect on asymmetric K § transport, 253-254 effective concentration, influence of membrane electrical potential, 250-254 inhibition of K § transport by Kir channels, 250-254 mechanism of inhibition of Kir channels, 251,256 structure and synthesis, 252 Porins, 270, 271 Positive cooperativity, 160, 200, 202, 203, 205, 237 Pregnancy, 319 Preimplantation conceptuses, 114 Preimplantation development, 113 Presteady-state currents, transporterassociated, 334-335 Primary active transport, 40, 42-44, 49, 57, 61, 83, 117, 121, 125 Primary structures, 266 Propelling forces, 3, 4 Protein, 304, 307, 321 Protein conformational changes, s e e Conformational changes Protein evolution through trinucleotide expansions, 184-193 Protein insertion, 20, 23 Protein kinase, 122 Protein kinase A, 279, 281,282 Protein kinase C, 281-283 Protein kinase-mediated phosphorylation sites, 212 Protein monomers, 116 Protein motion, 24, 25, 27 Protein sorting, 31, 33, 37, 38 Protein sorting signals, 37 Protein turnover, 304-307, 321-323 Protein-mediated transport is substrate saturable, 81 Proteoliposomes, 29, 30, 53, 115, 116, 226-228, 327 Proton-binding site of F-type ATPase, 156 Proton gradient, 5 Proton total chemical potential gradient, 56 Proton transport, 5, 133, 134 Protonophores, 5 P-type ATPase, 133-136, 138, 139, 140, 146, 147, 152 autophosphorylation, 146-152 cation pathway structure, 146-152 conformational changes, 148-152, 166-167 C-terminal membrane-associated domain, 137-141 cytosolic regions and domains, 137-139 dimerization, 167 domain C, 137-139 E1P-to-E2P transition, 148, 150 E2P mutants, 146 evolution and classification, 135-137 gene transcript splicing, 141-144 hinge region, 137-139

isoform and subisoform expression, 141 mechanism of ATP hydrolysis, 146-152 monomers, homodimers, heterodimers and heterotrimers, 144 N-terminal membrane-associated domain, 137-138 oligomerization, 151 region B, 137-139 site-directed mutagenesis, 146-52 sites of ATP hydrolysis and ion transport, 148-149 stalk, 137-139 structure, 134, 137-145 structure/function, 146-152, 166-167 Purified transport proteins, 115 Purity of commercial chemical preparations, 96 Putative transporters, 271 Putrecine structure, 252 Pyruvate transport, 115

Q Q10 values of biomembrane transport and permeation, 66, 78, 239 Quaternary structure, 19

R Radiotracer uptake assays, 331-332, 341-342 Reabsorption of cationic amino acids, 120 Reconstitution in proteoliposomes, 115, 116 Rectangular hyperbolas, s e e Hyperbolas Red blood cells of fishes, 235 Red cell ghosts, 203 Reductionism, 10 Region B 137, 139 Region J, 139 Regulation of metabolism through cell swelling and shrinking, 72 Regulation of transport, 65, 277-294 Regulatory site, 116 Regulatory volume decrease (RVD), 72 Regulatory volume increase (RVI), 72 Renal epithelia, 120 Reporter glycosylation sites, 218 Reproductive tract, 8 Resistance to diffusion (R), 67, 68 Reverse currents, 63 Reversibility of transport, 7

S Safety factor, 311 Saturable biomembrane transport, 84, 101, 125 Saturable fatty acid transport, 79, 81 Scramblase, 22 Scrambling of lipid asymmetries, 22, 31 Second law of thermodynamics, 39 Secondary active transporters, 4 Secretory vesicle, 58

Index Selectins, 24 Semiliki Forest virus, 33 spike glycoprotein, 35 Semipermeable membrane, 51, 57 Sepsis, 323 Sequence similarities among Kir channel proteins, 257 Sequence similarities in the pore-forming regions of K + channels, 255 SERCA ATPases, 139, 141, 142 dimers, 136 subisoforms, 136 Serotonin, 316 SGLT, see Glucose transporters S h a k e r channel, 243, 248, 254, 255, 259 Shared transport, 93 Shared transport processes, 94 Sheer stress, 13, 14 Sigmoidal curves, 117, 206 Sigmoidal kinetics, 214, 221,222 Signal peptides, 23 Signal sequences, 31, 32 Signaling through cell swelling or cell shrinking, 72 Simple diffusion, 83 Simultaneous mechanisms of symport/ antiport, 170 Simultaneous two-site mechanism, 200 Simultaneous two-site model, 199 Site-directed mutagenesis, 98, 139 AE1, 195-199 F-type ATPases, 159-161 K + channels, 254-256 P-type ATPases, 146-152 Size-corrected permeability coefficients, 74-79 Size-correction method for different solutes, 75 Slippage, 4, 59, 91, 98, 118, 121 Smooth muscle cells, 14 Sodium-proton antiport, see Na+/H + exchangers Solute gradients dissipation, 61, 116, 123, 125 via channel proteins, 239-263 formation, 116, 117, 125, 133 propagation, 116, 117, 123, 125 Solute gradients owing to asymmetric transporter function, 88 solute transport, 42, 43 Sorting signals, 34 Spectrin, 27 Spermidine structure, 252 Spermine structure, 252 Sphingomyelin, 18, 21, 22, 24, 29 Sphingomyelin synthesis, 24, 34 Spike glycoprotein, 33 Splice site A of PMCA ATPase gene transcripts, 141-143 Splice site C of PMCA ATPase genes transcripts, 141-143 Spontaneous processes, 39

Stalk region, 137, 138 Standard chemical potential, 40, 45 Standard free energy change, 41, 42, 50 Starvation, 320 Statistical comparisons of kinetic parameters, 85, 86 Statistical fit of transport data, 103 Statistical uncertainties in Km and Wmax values, 85 Steady state, 99 Steady-state solute gradients, 117 Steady-state approximation, 84 Stilbene compounds (DNDS, DIDS), 196, 198, 233 Stoichiometry of Na+/amino acid cotransport, 214, 237 of migration, 4 of transport, 4, 6, 7, 50, 59, 101,117 importance of direct measurement, 211-224 indirect cooperative kinetic method, 221-225 indirect measures are unreliable, 221-223 indirect thermodynamic method, 221-225 lack of correspondence between actual stoichiometry and that estimated thermodynamically, 222-223 lack of dependence on simultaneous entry, 215-217 measurement of, 336, 337 Stomatocytes, 23 Structure/function of P-type ATPase, 146-152 Subisoforms, 141 P-type ATPases, 141-144 Substitute osmolyte, 102 Substrate and transporter interactions, 86 binding order, 91 concentration effects of, 207 at half-maximal velocity, 84, 85 deprivation, 284-286 exodus, effect on uptake measurement, 99 inhibition, 202, 203 metabolism and biomembrane transport, 99 metabolites, 99 metabolized, 99 net uptake, 99 receptor sites, 103 saturable transport, 84, 101, 125 selectivity, 6 specificity, 103 Substrate/transporter interactions, 65, 86, 88, 125 Substrate-saturable fatty acid transport, 79, 81 Subunits of P-type ATPases, 144, 145 Sucrose substitution for Na +, 102

395 Sum of more than one rectangular hyperbola, 105, 106 Symmetric enzyme function, 88 Symmetric transporter function, 88 Symport, 59, 60, 91,101, 117, 122, 123 simultaneous and ping-pong mechanisms, 170 Symporter flux coupling, 258-260 Symporters, 4, 83, 89, 119, 121 169-170, 208-237 System A, 29, 94, 102, 112, 278, 284-287, 302, 310, 314, 320, 323, 327, 328 System ASC, 93, 94, 97, 98, 112, 212, 213, 229, 231,232, 314, 320, 323, 327; see also ASC System ASC transport model, 215-217 System ASC-like transporter (ASC), 208-217 System B, 94, 110, 112-114, 327 System B/b variants, 317, 318, 320, 324 System b+x, 94, 113, 114 System b+2, 94, 110, 113, 114 System/3, 8, 114, 212, 213 System B~ 94, 120 System B ~ 8, 93, 95, 102, 103, 107, 110-114, 120 System b ~ 96-98, 101-103, 106, 108-110, 112, 114, 120, 121,231,232 System b~ arginine transport, 83 System b ~ related amino acid transport protein (BAT), 104, 113, 229 System Gly, 94, 113, 114, 212, 213 System L, 94, 108-110, 112-114, 120, 121, 302, 310, 314, 316, 317, 320, 323 System N, 94, 302, 312, 327 System T, 93, 94, 110-114, 316-318 System X-/x-, 314, 318 System X-A, 114 System X-AO, 8, 94, 114 System xc-, 94, 114 System y+, 94, 97, 98, 229, 231,232, 315, 323 System y+L, 94, 97, 120, 121,229, 317, 320, 323

T T lymphocytes, 25 Tau/Gly channel1, 114 Tau/Gly channel2, 114 Taurine transport, 236 Taurocholate, 278 Techniques in biomembrane transport, 327-342 Temperature sensitivity of transport, 66 Tertiary (three dimensional) structures, 19, 116, 138, 266 Tetramer, 122 Tetramers, 231 Tetranucleotide expansions, 189 Tetranucleotide repeats, 188 Thermal energy, 5 Thermodynamic complexity, 117

396 Thermodynamic coupling of ATP hydrolysis and ion transport, 166 Thermodynamic expression ATP hydrolysis, 50 ATP synthesis, 56 bioenergetic coupling, 55 for a chemical reaction, 41 coupled processes, 56 migration of a solute, 42 solute gradient dissipation, 61 solute transport, 55 Thermodynamic principles and practical transport problems, 63 Thermodynamic principles of biomembrane transport, 63 Thermodynamic symmetry, 200 Thermodynamically asymmetric transporter function, 89 Thermodynamically symmetric transport, 20, 89 Thermodynamics and transport, 39-64 of coupling, 55, 57 of Na+/Ca § exchange, 59 Three dimensional model, 146 Thyroid hormone, 318, 323 Tight junctions, 32, 33 Tissue distribution of PMCA ATPase isoforms and subisoforms, 141-144 Total chemical potential, 4, 45-47, 64, 206 of Ca § 59 change, 44 gradient, 3, 4, 49, 58, 59, 61, 89, 120, 123-125, 134, 237 formation, 119 of water, 47, 48 Trafficking of Na§247 and H§247 145 Transepithelial transport, 9 Transferrin receptor, 37 Transgenic animals, 325 trans-Golgi, 18, 30 trans-Golgi network, 18, 30, 31 trans-4-hydroxy-L-proline, 110 trans-inhibition, 259

Translocators driven by proteins, 268 Transmembrane alpha-helices, 137 Transmembrane segments, 134 Transport activities for a single substrate, 63 Transport activities of a single protein, 63 Transport activity isolation and characterization, 98-110, 125-131 Transport and chemical change, 134 Transport and interorgan nutrient flows, 295-326 Transport ATPase, 133 Transport commission (TC) classification system, 267-272 Transport competition, 130 Transport components, 126-131 Transport coupling/uncoupling, 121, 122 Transport cycle, 117, 135 Transport experimental design, 95

Index

Transport heterogeneity, 103, 111, 131 Transport kinetics, 65 Transport kinetics and total chemical potential, 261,262 Transport mechanism, 311 Transport of fatty acids, 79 Transport of lipophilic substances, 69, 73-79 Transport of more than one of the same ion or molecule, 85 Transport of water, 68, 69 Transport pathway, 137 Transport process protein composition, 111 Transport proteins in bacteria, 275-276 catalysts, 64 classification, 265-276 conformational changes see Conformational changes cross classification, 272-275 expression in Xenopus oocytes, 113-116, 330-332 families, criteria for assignment, 266-267 function, 29 helix packing in, 340 multiple components of transport, 115, 116 purification of, 327 reconstitution of, 115, 116, 327 structure-function studies of, 339-341 three-dimensional structural data, 268 that form solute gradients, 133-167 that propagate solute gradients, 169-170, 208-237 Transport regulation, 124, 277-294 Transport related auxiliary proteins, 270 Transport stoichiometry see Stoichiometry of transport Transport systems (amino acid), 114, 300 Transport thermodynamics, 39-64 Transport, nonsaturable, 126 Transport, simultaneous, 119 Transporters asymmetry, 169 and solute gradients, 125 classification function-based, 265-267 phylogeny-based, 266-267 conformational change, 65, 119, 133, 151 conformational free energy, 89 definition, 65 function, 133 orientation, 88 reversal of, 90 structure, 133 symmetric operation, 123 topology, 20 transition state free energy, 170 of unknown classification, 270, 271 Transporter/substrate interactions, 65, 86, 88, 125 Transport-related auxiliary proteins, 271

Transport-related protein expression in Xenopus oocytes, 113-116 Transposable elements, 191 trans-stimulation, 84, 118-120, 123, 124, 199, 259 Triiodothyronine, 110 Trinucleotide expansion, 235 Trinucleotide expansion in Kv channel evolution, 241 Trinucleotide repeats, 187 trinucleotide residue expansion and protein evolution, 184-193 Trophoblast, 9 Trout AE1 proteins, 235 Trout cDNA library, 235 Tryptophan, 309, 316, 324 Tryptophan analogs, 108 Tryptophan transport, 108 Tryptophan, D-isomer, 110, 112 Tumor cells, 291-293 Tumorigenic cells, 23 Tunneling, see Slippage Turnover number of transport, 65 Turnover rate, measurement of, 335 Type I P-type ATPases, 135, 136 Type II ATPases, 136 Type II P-type ATPases, 135, 136 Type IIA ATPases, 135, 136, 142 Type IIA P-type ATPases, 141 Type IIB ATPases, 135, 142 Type IIB P-type ATPases, 141 Tyrosine kinase, 233

U Uncorrected permeability coefficients, 77, 78 Uncoupled transport, 4, 5 Uncoupling, 91 Uncoupling proteins, 5 Unidirectional efflux, 258 Unidirectional flux, 3, 83 Unidirectional influx, 258 Uniport, 118, 119, 121-123, 125 obligatory, 122 Uniporters, 83, 89-91, 120, 124 Unsaturated fatty acyl chains, 26 Unshared transport processes, 94 Unstable expansion, 187 Unstirred water layer width, 75, 76 Unstirred water layers, 73, 74, 78, 79, 121 Unstirred water volume, 76 Urea, 298, 312, 313, 315, 322 Uridine, 278 Uridine transport, 115 Uterus, 8

V Vaccinia virus, 329-331 Vacuolar H § ATPase, 58 Vascular endothelial cells, 23

Index Vesicle shedding, 22 Vesicles, transport in, 341 Vesicular budding, 22 Vesicular trafficking, 30, 31, 33-38 Vma x value, 83-87, 90, 91,108, 112, 115, 116 Voltage-sensitive cation channels, 240-254 Voltage-gated channel evolution, 240-246, 249-250 Voltage-gated K § (Kv) channels, 241-249 Voltage-gating mechanisms in Kv channels, 243-248 Volume regulation by AE proteins, 233-234 Volume-sensitive anion channel proteins, 141 Volume-sensitive organic osmolyte/anion channel (VSOAC), 235 VSOAC activity in mammals, 235 V-type ATPases, 55-57, 133, 134 c-subunit isoforms, 165 subunit isoforms and subisoforms, 164 variety among biomembranes, 164

W Water biomembrane transport, 68, 69, 339 Water channels, 82, 125 Water diffusional permeability coefficient (Pd), 70, 71 Water molecules inside water channels, 71 Water molecules that span water channels, 72 Water osmotic permeability coefficient (Pf), 70, 71 Water permeation of biomembranes, 68, 69 Water unstirred layers, 73, 74, 78, 79, 121 Widths of unstirred water layers, 76 Work performed owing to transport, 64 Wortmannin, 303

X X e n o p u s oocytes, 104, 114, 115, 120, 212,

213, 221,222, 226, 228-230, 235, 236, 254, 257, 328, 339 cut-open preparation, 335-336

397 heterologous expression in, 330-332 isolation of membrane fraction from, 332 microinjection of, 331 preparation of, 331 radiotracer assays using, 331-332, 336 voltage-clamp in, 332-338 X-ray crystallographic studies, 161,199 of F1 sector, 158 X-ray microanalysis, 17

Y Yolk sac placenta, 9, 10

Z Zero trans conditions, 91 Zero trans experiments, 84 Z-loop, in AE protein channel function, 236