Current Topics in Membranes and Transport, Volume 22: The Squid Axon

Current Topics in Membranes and Transport Volume 22 The Squid Axon

Advisory Board

M . P . Blaustein G. Blobel J . S . Cook P. A . Knuuf Sir H . L . Kornberg

P . Lauger C. A. Pasternak W . D . Stein W . Stoeckenius K . J . Ullrich

Contributors

D. A . Haydon Clay M . Armstrong P . F. Baker B . M . Hendry David Landowne Luis Beuuge' Raymond J . Lasek Ted Begenisich Walter F . Boron Rodolfo R . Llincis Leslie M . Loew A . Carruthers Lnwrence B . Cohen Donald R . Mutfeson H . Meves Franco Conti Toshio Narahashi R . DiPolo Hurish C. Pant Gerald Ehrenstein John M . Russell J . R . Elliott Brian M . Salzherg Harold Gainer Catherine Smith Paul E. Gallant Gloria M . Villegas Daniel L . Gilbert Jorge Villegas Robert Gould Raimundo Villegas

Current Topics in Membranes and Transport Edited by

Arnost Kleinzeller Department of Physioiogy University of Philudelphiu School of' Medicine Philadelphia, Pennsyluunia

VOLUME 22

THE SQUID AXON Gues I Editor

Peter F. Baker Department of Physiology University of London King's College London, England

1984

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Contents Contributors, x Preface, xiii Yale Membrane Transport Processes Volumes, xv

PART 1.

STRUCTURE

Squid Axon Ultrastructure

GLORIA M. VILLEGAS AND RAIMUNDO VILLEGAS 1.

11. 111. IV. V. VI.

Introduction, 3 Anatomical Organization. 4 Histological Organization, 6 Fine Structure, 7 Nerve Fractions, 29 Concluding Remarks, 32 References, 34

The Structure of Axoplasm

RAYMOND J. LASEK

I. Introduction, 39 11. Fine Structure of the Squid Giant Axon, 40 111. Axonal Transport in the Squid Giant Axon, 47 IV. Extruded Axoplasm from the Giant Axon, 49 V. Conclusions, 50 References, 5 I PART 11.

REGULATION OF THE AXOPLASMIC ENVIRONMENT

Biochemistry and Metabolism of the Squid Giant Axon

HAROLD GAINER, PAUL E. GALLANT, ROBERT GOULD. AND HARISH C . PANT

I. Introduction, 58 11. Molecular Composition of Axoplasm, 58

111. Functional Metabolic Studies Using the Giant Axon System, 68 V

vi

CONTENTS

IV. Concluding Remarks, 81 References, 81 Transport of Sugars and Amino Acids

P. F. RAKER AND A. CARRUTHERS I.

Introduction, 91

11. Concentration and State of Sugars and Amino Acids in Axoplasm, 92

Ill. Transport of Sugars by Squid Axons, 98 IV. Descriptive Models for Sugar Transport, I10 V. Transport of Amino Acids, 119 VI. General Conclusions, 128 References, 128 Sodium Pump in Squid Axons

LUIS BEAUGE I. Introduction, I3 I 11. Experimental Techniques, 132 111. Sodium Pump in Squid Axons, 134 References. 171 Chloride in the Squid Giant Axon

JOHN M. RUSSELL

I . Introduction, 177 11. [CI-] in Axoplasm and Hemolymph, 178 111. Transmembrane CI Movements, 180 IV. Summary and Discussion, 190 References, 192 Axonai Calcium and Magnesium Homeostasis

P. F. RAKER AND R. DIPOLO I. 11. Ill. IV.

Introduction. 195 Axonal Ca and Mg: The Problem of Bound versus Free, 196 Axonal Mg Homeostasis, 197 Axonal Ca Homeostasis, 201 References, 242

Regulation of Axonal pH

WALTER F. BORON I. Introduction, 249 11. Methodology, 250 111. Effect of Weak Bases and Acids on pH,, 2S2 IV. Effect of Metabolic Inhibitors o n pH,, 257 V . Ionic Mechanism of pH, Regulation, 258 References, 268

vi i

CONTENTS Hormone-Sensitive Cyclic Nucleotide Metabolism in Giant Axons of Loligo

P. F. BAKER AND A. CARRUTHERS I. Introduction, 271 11. Cyclic AMP Levels in Intact Axons, 272 111. CAMP Production and Breakdown in Perfused Axons, 273 1V. Conclusion, 276 References, 276 PART 111.

EXCITABILITY

Hodgkin-Huxley: Thirty Years After

H . MEVES

I. Introduction, 279 11. Action Potential, 280 111. Voltage-Clamp Currents, 2XX IV. Hodgkin-Huxley Analysis, 294 V. Reconstruction of the Electrical Behavior of the Squid Axon from the Hodgkin-Huxley Equations, 308 VI. Molecular Properties of the Na Channel: Permeability Ratios and Binding Constants, 31.5 VII. Concluding Remarks, 321 References, 322 Sequential Models of Sodium Channel Gating

CLAY M. ARMSTRONG AND DONALD R. MATTESON 1. Introduction, 332 11. Results, 334 111. Discussion, 350 References, 352 Multi-Ion Nature of Potassium Channels in Squid Axons

TED BEGENISICH AND CATHERINE SMITH I. Introduction, 353 11. Characteristics of Multi-Ion Pores, 355 111. Experimental Results, 358

IV. A Mathematical Model, 363 V. More on Energy Barriers, 366 VI. Summary and Conclusions, 368 References, 368 Noise Analysis and Single-Channel Recordings

FRANC0 CONTI I. Introduction, 371 11. Basic Concepts of Fluctuation Analysis, 372

viii

CONTENTS

111. The Physical Basis of Membrane Noise, 374

1V. Channel Noise in Nerve Membranes, 385 V . Single-Channel Recordings, 397 References, 401 Membrane Surface Charge

DANIEL L. GILBERT AND GERALD EHRENSTEIN

I. Introduction, 407 Channel Surface-Charge Density: Theory, 408 I l l. Neutralization of Channel Surface-Charge Density, 412 1V. Relation of Average Surface-Charge Density to Channel Surface-Charge Density, 414 V. Modifications of Surface Charge, 41.5 VI. Significance, 416 V11. Summary, 418 References, 418 11.

Optical Signals: Changes in Membrane Structure, Recording of Membrane Potential, and Measurement of Calcium

LAWRENCE B. COHEN, DAVID LANDOWNE. LESLIE M . I.OEW, AND BRIAN M . SALZBERC

1.

Introduction, 424 Intrinsic Signals. 426 Dye Measurements of Membrane Potential, 429 IV. Measurement of Changes in Internal Calcium Concentration, 439 V . Conclusions, 440 References, 441 11. 111.

Effects of Anesthetics on the Squid Giant Axon

D. A. HAYDON. J . R. ELLIOTT, AND B . M. HENDRY I . Introduction. 445 Effects of Anesthetics on the Action Potential and Sodium and Potassium Currents of the Squid Giant Axon, 451 111. Mechanisms of Sodium and Potassium Current Suppression, 464 Appendix: Effect of Membrane Thickness Changes on the Steady-State Inactivation of the Na Channel, 476 References. 478 11.

Pharmacology of Nerve Membrane Sodium Channels

TOSHlO NARAHASHI I. Introduction, 483 11. Ionic-Channel Block by Chemicals, 486 111. Ionic-Channel Modulation by Chemicals. 493 IV. Conclusions. 507 References. 51 1

ix

CONTENTS

PART 1V.

INTERACTION BETWEEN GIANT AXON AND NEIGHBORING CELLS

The Squid Giant Synapse

RODOLFO R. LLINAS

I. Introduction, 519 11. 111. IV. V. VI. Vil.

Anatomy, 521 Early Electrophysiology, 523 Synaptic Transmission without Action Potentials, 524 Electrophysiological Properties of the Postsynaptic Potential, 525 Voltage Clamp of the Presynaptic Element Using Square Pulses, 530 Voltage Clamp with Action Potential Waveform, 538 Keferences, 544

Axon-Schwann Cell Relationship

JORGE VILLEGAS I. Introduction, 547 1I. Axon-Schwann Cell Interface, 548 111. Axon-Schwann Cell Signaling, S54 IV. Schwann Cell Responses, 557 V. Cell-to-Cell Communication, 565 VI. Concluding Remarks, 567 References. 567

Index, 573 Contents of Previous Volumes, 579

Contributors Numbers in parentheses indicate the pages on which the authors’ contributions begin. Clay M. Armstrong, Department of Physiology, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania 19104 (331) P. F. Baker, Department of Physiology, University of London King’s College, London WC2R 2LS, England (91, 195, 271) Luis Beaugb, Divisi6n de Biofisica, lnstituto de Investigation Medica Mercedes y Martin Ferreyra, 5000 Cordoba, Argentina (131) Ted Begenisich, Department of Physiology, University of Rochester School of Medicine and Dentistry, Rochester, New York 14642 (353) Walter F. Boron, Department of Physiology, Yale University School of Medicine, New Haven, Connecticut 06510 (249) A. Carruthers, Department of Physiology, University of London King’s College, London WC2R 2LS. England’ (91, 271) Lawrence B. Cohen, Department of Physiology, Yale University School of Medicine, New Haven, Connecticut 06510 (423) Franco Conti, lstituto di Cibernetica e Biofisica, Consiglio Nazionale delle Ricerche. 1-16032 Camogli, Italy (371) R. DiPolo, Centro de Biofisica y I3ioquimic;r. InstitutoVene/.olano de lnvestigaciones Cientificas (IVIC), Caracas 1010-A, Venezuela (195) Gerald Ehrenstein, Laboratory of Biophysics, Nationdl Institute of Neurological and Communicative Disorders and Stroke, National Institutes of Health, Bethesda, Maryland 20205 (407) J. R. Elliott, Physiological Laboratory, University of Cambridge, Cambridge CB2 3EG, England (445) Harold Gainer, Laboratory of Neurochemistry and Neuroimmunology, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, Maryland 20205, and Marine Biological Laboratory, Woods Hole, Massachusetts 02543 (57) Paul E. Gallant, Laboratory of Neurobiology, National Institute of Mental Health, Alcohol, Drug Abuse, and Mental Health Administration, Bethesda, Maryland 20205, Laboratory of Preclinical Studies, National Institute on Alcohol Abuse and Alcoholism, Alcohol, Drug Abuse, and Mental Health Administration, Rockville, Maryland 20852, and Marine Biological Laboratory, Woods Hole, Massachusetts 02543 (57) Daniel L. Gilbert, Ldbordtory of Biophysics, National Institute of Neurological and Communicative Disorders and Stroke, National Institutes of Health, Bethesda, Maryland 20205 (407) Robert Gould, Institute for Basic Research in Developmental Disabilities, Staten Island. New York 10314, and Marine Biological Laboratory, Woods Hole, Massachusetts 02543 (57)

’

Present address: Department of Biochemistry, University of Massachusetts Medical Center, Worcester, Massachusetts 01605. X

CONTRIBUTORS

xi

D. A. Haydon, Physiological Laboratory, University of Cambridge, Cambridge CB2 3EG, England (445) B. M. Hendry, Physiological Laboratory, University of Cambridge, Cambridge CB2 3EG, England (445) David Landowne, Department of Physiology and Biophysics, University of Miami School of Medicine, Miami, Florida 33124 (423) Raymond J. Lasek, Neurobiology Center, Department of Developmental Genetics and Anatomy, Case Western Reserve University, Cleveland, Ohio 44106 (39) Rodolfo R. Llinas, Department of Physiology and Biophysics, New York University Medical Center, New York, New York 10016 (519). Leslie M. Loew, Department of Chemistry, State University of New York at Binghamton, Binghamton, New York 13901 (423) Donald R. Matteson, Department of Physiology, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania 19104 (331) H. Meves, I. Physiologisches Institut, Universitat des Saarlandes, 6650 Homburg-Saar, Federal Republic of Germany (279) Toshio Narahashi, Department of Pharmacology, Northwestern University Medical School, Chicago, Illinois 6061 1 (483) Harish C. Pant, Laboratory of Preclinical Studies, National Institute on Alcohol Abuse and Alcoholism, Alcohol, Drug Abuse, and Mental Health Administration, Rockville, Maryland 20852, and Marine Biological Laboratory, Woods Hole, Massachusetts 02543 (57) John M. Russell, Department of Physiology and Biophysics, University of Texas Medical Branch, Galveston, Texas 77550 (177) Brian M. Salzberg, Department of Physiology, University of Pennsylvania, Philadelphia, Pennsylvania 19104 (423) Catherine Smith, Department of Physiology, University of Rochester School of Medicine and Dentistry, Rochester. New York 14642 (353) Gloria M. Villegas, Centro de Biofisica y Bioquimica, lnstituto Venezolano de Investigaciones Cientificas (IVIC). and lnstituto lnternacional de Estudios Avanzados (IDEA), Caracas 1010-A, Venezuela (3) Jorge Villegas, Centro de Biofisica y Bioquimica, lnstituto Venezolano de Investigaciones Cientificas (IVIC), and lnstituto lnternacional de Estudios Avanzados (IDEA), Caracas 1010-A. Venezuela (S47) Raimundo Villegas, Centro de Biofisica y Bioquimica, lnstituto Venezolano de Investigaciones Cientificas (IVIC), and lnstituto lnternacional de Estudios Avanzados (IDEA), Caracas 1010-A, Venezuela ( 3 )

J. Z. YOUNG

A. L. HODGKIN

K . S. COLE

A. F. HUXLEY

Few preparations have contributed more to biology than the giant axon of the squid. The secret of its success is twofold: size and relevance to less tractable mammalian systems. This triumph of size over phylum merits examination. Despite its molluscan origin, most of t h e work on the squid axon finds remarkably close parallels in mammals, emphasizing the commonality of the basic building blocks from which biological systems are constructed. This is not to say that the squid axon has everything, but what it has is amenable to detailed experimental analysis. The great advantage of the squid axon is its large size, which makes it a relatively easy matter to perform experimental feats that are unthinkable on smaller cells; and even where manipulation is possible on a smaller cell, it is that much easier to do well with the larger squid axon. Although the squid axon played, and continues to play, a leading part in the elucidation of the mechanisms of nervous conduction, its usefulness extends far beyond this sphere. The recent discovery of particle movement in axoplasm promises, for instance, to put the squid axon in the vanguard of work on axoplasmic transport, and many other new developments are discussed in this book. We hope that readers will be encouraged to examine whether the squid axon can make a contribution to their particular field. A separate article on techniques is not included, but useful information can be found throughout the volume. The reader who needs more information is referred to the review by P. Rosenberg entitled “The Squid Giant Axon: Methods and Applications” (in Methods of’Nrrrrohiology 1, 1133, 1981) and to the excellent film “The Squid and Its Giant Nerve Fibre” [ J . Physiol. (London) 263, 99P. 19761, copies of which can be obtained from Dr. J. Cilpin Brown, The Laboratory, Citadel Hill, Plymouth, United Kingdom. M y task as editor has been simplified by the enthusiastic cooperation of many colleagues around the world. We wish to dedicate our efforts to all those who pioneered the use of the squid giant axon and hope this book will prove of value not only to those who regularly use squid axons, but to cell physiologists in general.

PETERF. BAKER xiii

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Yale Membrane Transport Processes Volumes Joseph F. Hoffman (ed.). (1978). “Membrane Transport Processes,” Vol. I . Raven, New York. Daniel C. Tosteson, Yu. A. Ovchinnikov, and Ramon Latorre (eds.). (1978). “Membrane Transport Processes,” Vol. 2. Raven, New York. Charles F. Stevens and Richard W. Tsien (eds.). (1979). “Membrane Transport Processes,” Vol. 3: Ion Permeation through Membrane Channels. Raven, New York. Emile L. Boulpaep (ed.). (1980). “Cellular Mechanisms of Renal Tubular Ion Transport”: Volume 13 of Current Topics in Membranes and Transport (F. Bronner and A. Kleinzeller, eds.). Academic Press, New York. William H. Miller (ed.). (1981). “Molecular Mechanisms of Photoreceptor Transduction”: Volume 15 of Current Topics in Membranes and Transport (F. Bronner and A. Kleinzeller, eds.). Academic Press, New York. Clifford L. Slayman (ed.). (1982). “Electrogenic Ion Pumps”: Volume 16 of Current Topics in Membranes and Transport (A. Kleinzeller and F. Bronner, eds.). Academic Press, New York. Joseph F. Hoffman and Bliss Forbush 111 (eds.). (1983). “Structure, Mechanism, and Function of the Na/K Pump”: Volume 19 of Current Topics in Membranes and Transport (F. Bronner and A. Kleinzeller, eds.). Academic Press, New York. James B. Wade and Simon A. Lewis (eds.). (1984). “Molecular Approaches to Epithelial Transport”: Volume 20 of Current Topics in Membranes and Transport (A. Kleinzeller and F. Bronner, eds.). Academic Press, New York.

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Part I

Structure

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CUKKENT TOPICS IN MEMBRANES AND TRANSPORT, VOLUME 22

Squid Axon Ultrastructure GLORIA M . VILLEGAS AND RAIMUNDO VILLEGAS Centro de Biofisica y Bioquimica Instituto Venezolano de Investigaciones Cient$cas (IVIC) and Instituto Internacional de Estudios Avanzados (IDEA) Caracas, Venezuela

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

I.

3

11. 111.

IV. Fine Structure. .......... A . Axon.. . . . . . . . . . . . . . B . Schwann Cell Layer . . . . V. VI.

References . . . . . . . . . . . . . .

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

29

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34

I. INTRODUCTION In nature, certain animals have developed special morphological features to support the mechanisms of rapid movements necessary for escape or attack. In the vertebrates, rapid nerve conduction is accomplished mainly by isolating the pathway along which the nerve impulse is transmitted with a special insulator, the myelin sheath. In some other animals, with largely unmyelinated nerve fibers, rapid conduction is achieved by a system of giant nerve fibers; the conduction velocity increasing with axon diameter. Among those animals having giant nerve fibers, the Mollusca, Cephalopoda, Decapoda (squids) are the ones that provide the best preparation for research in the neurosciences. As pointed out by Young (19359, the giant fiber system of the Cephalopoda was discovered by Williams at the turn of the century, but it was 3 Copyright 0 1984 by Academic Press, Inc All right\ of reproduction in any form reserved ISBN 0-12-153322-0

4

GLORIA M. VILLEGAS AND RAIMUNDO VILLEGAS

Young himself who called the attention of neuroscientists to this extraordinary preparation (1934, 1936). II. ANATOMICAL ORGANIZATION Giant nerve fibers exist in the decapodan nervous system at three levels (Young, 1939). Two first-order giant fibers emerge from the magnocellular lobes of the brain and then run caudally toward the palliovisceral lobes and there make synaptic contacts with the second-order giant axons. Two of the second-order giant fibers that emerge from the palliovisceral lobes join the pallial nerves, which run caudally toward the stellate ganglia and there synapse with the third-order giant axons that form part of the stellate nerves. Each stellate nerve carries a giant nerve fiber among numerous fibers of more usual size. The continuity and effectiveness of the system is accomplished by decussations (chiasmata) and synapsing of the giant fibers of the three different orders (Fig. 1) (Young, 1939; Martin, 1969). Besides the presence of giant axons, there is another unique feature of the squid nervous system. This feature, also noted by Young (1939), is that the first-order giant fibers not only decussate, but are fused at the point of decussation and there are neither septa nor membrane remnants separating the two axoplasms. This fusion, described by Martin (1969) as an H-shaped bicellular unit, apparently occurs during the larval or postlarval development in Loligo vulgaris, as Martin has observed the two decussating unfused axons (chiasm I) completely separated by two apposed intact membranes (Martin, 1969). In other types of cephalopods, such as Sepici and I l k x , also studied by Martin (1969), there is not a fusion of the first-order giant axons, but there exists either an electrical-type synaptic contact or a nonpolarized synapse. Both these arrangements achieve the bilateral spread of a unilaterally initiated impulse, thereby fulfilling the functional requirement of the system. Of the stellate nerves, the hindmost is the one used by most researchers because it is the largest in diameter and can be dissected for several centimeters of its length before shedding collaterals and penetrating into the muscles of the mantle. The hindmost stellar nerve is therefore universally known as the squid giant axon. Different species of squid are used over the world as sources of giant axons for research. The diameter of the giant axon in the most commonly used species ranges from about 200 pm in Sepia of$cinalis ; 200 to 500 pm in t h e squids Doryteuthis plei and Sepioreuthis sepiuideu, found in tropical waters; 400 to 900 p m in the squids Loligo pealei, Loligo forhesi, and

5

SQUID AXON ULTRASTRUCTURE

-

2"

SG

FIG.1. Schematic representation of the system of giant nerve fibers in the squid. The first-order giant fibers (1") emerge from the magnocellular lobes (MC) and run to the palliovisceral lobes (PV) to make synaptic contacts with the second-order giant fibers (2"). The first-order giant fibers are connected by a bridge ( B ) , and in such a way form a sort of Hshaped two-cell unit. One second-order giant fiber at each side integrates the pallial nerve and ends in the stellate ganglia (SG) after making synapse with the third-order giant fibers (3"). The stellate nerves, each having a third-order giant fiber, extend out from the stellate ganglia to innervate the mantle muscles. The hindmost and longest third-order giant fiber is the so-called squid giant axon. (Redrawn after Young, 1939, and Martin, 1969.)

Doryteuthis bleekeri, from cold waters; and up to 1500 p m in the giant squid Dosidicus gigas of the Humboldt current in the Pacific Ocean. According to Rosenberg (1981), the giant axon diameter seems to correlate better with the mantle circumference than with the mantle length, and this is also our observation in the tropical species S . sepioidea and D. plei. As described by Young (1936), the giant axon at its origin is a syncytial structure; this means that it originates from the fusion of several hundred small axons of the ganglionic cells. This fact constitutes one of the few exceptions to the neuron theory, which otherwise, according to Young (1939), contributes to prove the rule by showing the contrast between the function of two completely fused axons and that of two axons making contact through a synapse. Axons completely fused must work together

6

GLORIA M. VILLEGAS AND RAIMUNDO VILLEGAS

as a single cell, impulses spreading over the common axolemma. On the other hand, at the synapse, the cells are in contact but their cytoplasms are separated, and this organization prevents the spreading of a single impulse over the whole nervous system. I11.

HISTOLOGICAL ORGANIZATION

The so-called squid giant axon, as indicated above, is the giant nerve fiber unit present in the hindmost stellar nerve. Such a unit, like any other nerve fiber, is constituted by three different cellular elements: the neuronal element represented by the axon, which forms the most internal part or core; the glial element represented by the Schwann layer that surrounds the axon; and finally, the connective tissue element represented by the fibrocytes and collagen bundles that form the endoneurium and are situated most externally (Fig. 2). The fact that the giant axon has a very large diameter is perhaps responsible for the unique arrangement of glial cells found in the preparation. The small stellar nerve fibers resemble vertebrate nerves in that there are numerous Schwann cells along the length of the axon, but only one cell is needed to surround completely one or several axons in a single crosssection. In the case of the giant axon, several Schwann cells are organized in a single row, one cell thick, encircling the axon (Villegas and Villegas, 1968) (Fig. 2).

FIG.2. Schematic representation of the organization of the squid giant nerve fiber. The central cylinder (A) is the giant axon, which is surrounded by several Schwann cells arranged in a layer one cell thick (SC). The outer envelope corresponds to the endoneuriurn (E) and is integrated by fibrocytes and collagen.

SQUID AXON ULTRASTRUCTURE

IV.

7

FINE STRUCTURE

A. Axon

The axon, being a neuronal process, consists of a mass of cytoplasm (axoplasm) surrounded by a plasma membrane (axolemma), each in continuity with the neuron cytoplasm and the neuron plasma membrane, respectively. Axoplasm is a highly viscous material consisting of a ground substance in which at least three different types of filamentous structures contribute to form a cytoskeletal network. Interspersed in the network are numerous mitochondria and smooth endoplasmic reticulum profiles (Fig. 3). The three different types of threadlike elements have been identified as neurofilaments, microtubules (Davison and Huneeus, 1970), and actin microfilaments, the latter reacting specifically with heavy meromyosin to form typical arrowhead complexes (Witman and Rosenbaum, 1973). Neurofilaments and microtubules are the most prominent of the

FIG.3. Electron micrograph of the axoplasm of the giant axon in longitudinal section. A network of neurofilaments with a dotted appearance, intermingled with mitochondria (m), vesicular profiles of the endoplasmic reticulum (ER), and a few microtubules (arrows), is observed.

8

GLORIA M. VILLEGAS AND RAIMUNDO VILLEGAS

filamentous structures, although the microtubules have been difficult to preserve for electron microscopy because of their lability in solutions of high ionic strength. As reported by Davison and Huneeus (1970), Dr. Betty Geren Uzman, using glutaraldehyde-fixed preparations of Dosidicus gigas axons, was able to show only a few microtubules in the whole axon, but many more in similarly fixed freshly extruded axoplasm. Later, with improvements in the fixation methods by cannulation and irrigation of the axon interior with the fixative, it was possible to show numerous microtubules, especially in the periphery of the axoplasm, where they are oriented longitudinally (Endo et ul., 1979; Hodge and Adelman, 1980). The neurofilaments, on the other hand, are the most characteristic and commonly encountered fibrous element of the axoplasm. They are filaments 5-10 nm thick, grouped in bundles, which exhibit a dotted appearance and appear to run in all directions, but mainly longitudinally (Fig. 3) (G. M. Villegas, 1969). Metuzals (1969) and Metuzals and Izzard (1969) have studied the axoplasm of Loligo pealei with differential interference microscopy and electron microscopy and have been able to describe a spatial pattern of the neurofilaments characterized by a continuous threedimensional network, whose thread elements are more concentrated and closely packed in the peripheral ectoplasmic region, where they also are more parallel. The threads are formed by intercoiled unit filaments with cross-associations, forming ladder-like structures (Metuzals, 1969). This organization has been confirmed by Hodge and Adelman (1980) using the stereo electron microscope to observe thick sections of L . pealei giant axons. Hodge and Adelman described the presence of highly oriented 10nm neurofilaments, 5- to 6-nm actin microfilaments and microtubules, linked together by thin transverse bridges. At times, the microtubules appeared to be in close relationship with the axolemma, a fact already pointed out by Metuzals and Tasaki (1978) and by Endo et al. (1979), who observed some microtubules ending in or cross-bridged by filaments to the inner aspect of the axon plasma membrane. The existence of a continuous network in the axoplasm of the giant axon could serve as a morphological correlate for several important functions, as well as help explain other structural observations. The weakness of the birefringence of fresh axoplasm first noted by Bear et al. (1937b) could be explained by the wide range of angles of orientation of the filaments that form the network. A similar conclusion was reached by Richards of ul. (1943) in their pioneering ultrastructural studies of squid axoplasm. The early observation of Bear et (11. (1937a), who described a thin, membrane-like structure surrounding the extruded axoplasm, as well as the observation of Baker et d.(1962), who found a thin film of axoplasm remaining attached to the axolemma after perfusion of t h e axon, could both be explained by the existence of a peripheral, more dense

SQUID AXON ULTRASTRUCTURE

9

ectoplasmic region in the continuous network (Metuzals and Izzad, 1969). The folding and unfolding, the coiling and uncoiling, as well as the sliding of the filaments of the network could contribute to the mechanism of axonal transport (Metuzals, 1969). A further possible role for the network is in the control of excitability. Thus, there is a close correlation between conditions that maintain membrane excitability and conditions that keep microtubules intact (Chambers and Kao, 1952; Hodgkin and Keynes, 1956; Weisenberg, 1972), and this has led to the proposal that a probable function for the network of microtubules is to control the spatial interaction of proteins regulating ionic permeability (Matsumoto and Sakai, 1979), and specifically that microtubules and microtubule-associated proteins may be involved in the functioning of excitable membranes (Sakai and Matsumoto, 1978; Matsumoto et al., 1979). Direct evidence for interaction between excitation and axoplasmic structure is indicated by experiments in which the conduction of the nervous impulse is accompanied by changes in the light scattering of the axoplasm of the squid giant axon (Cohen et al., 1968). The slow reversal of the light-scattering changes suggests a long-lasting, impulse-associated structural change of radially oriented molecules associated with the membrane. As mentioned earlier, the other organelles present in axoplasm are mitochondria and agranular endoplasmic reticulum. Mitochondria have been observed in D.gigas axons in two different shapes. One is round, about 0.6 pm in diameter, mainly occupies the peripheral axoplasm (Fig. 4), and sometimes is observed very close to the axolemma. The other type is elongated, up to 3 pm long, and oriented longitudinally with respect to the axonal long axis (G. M. Villegas, 1969). In axons of the same species, mitochondria have been counted and appear to be slightly more concentrated in the periphery-40 mitochondria per 100 pm2 in the peripheral region and 30 units per 100 pm2 in a region situated 40 pm deeper in the axoplasm (Villegas and Villegas, 1968; G. M. Villegas, 1969). In the Doryteuthis and Sepioteuthis species studied by us, as well as in the pictures of L. pealei we have examined, the distribution of the mitochondria appears to be more homogeneous. In order to have an idea of this distribution, serial photographs were taken along two perpendicular lines extending from the borders of one cross-section of a D.plei giant axon. The pictures were arranged in a montage, and the mitochondria were counted in every 100 pm2 of area. A total of 456 mitochondria were counted in 1993 pm2,and their distribution varied between 13 and 38, units per 100 pm’, independent of the radial depth of the examined area. The endoplasmic reticulum appears to be formed by agranular membra-

FIG.4. Peripheral region of the giant axon of Dosidicus gigas showing several large, spherical mitochondria (m), as well a s some smaller profiles (arrows), corresponding to cross-sections of longitudinal mitochondria. SC, Schwann cell layer.

FIG.5. Axon-Schwann cell layer boundary of the giant fiber of S. sepinidea. Arrows point to several vesicular profiles lying under the axon membrane and very close to it. SC, Schwann cell layer; N , nucleus; A, axon.

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11

nous profiles 20-60 nm in diameter, filled with a moderately dense material. In longitudinal sections, long branching cisterns and tubules are observed, as well as rows of vesicles that might correspond to tubules artifactually disrupted (G. M. Villegas, 1969) (Fig. 3). Hodge and Adelman (1980) have described a close relationship between radially oriented cisterns of the agranular reticulum and the inner surface of the axolemma. Previously, Baker er al. (1962) had pointed out that the vesicles were more numerous within a few microns of the surface. We have made similar observations (Figs. 5 , 6, and 17) in thin sections of giant axons of D.plri and S . sepioidea and also in freeze-fracture replicas of the latter species (Villegas et al., 1984). In addition, Henkart er al. (1978), using the squid giant axon, have pointed out the close relationship of the peripheral endoplasmic reticulum cisterns and the sequestering of calcium. As demonstrated by those authors, the calcium ions penetrating the axon during stimulation are sequestered by the mitochondria and also by the peripheral vesicotubular structures of the endoplasmic reticulum. This modus operandi can be considered to be homologous with the system operating in skeletal muscle (Costantin et al., 1965). Another form of the axonal endoplasmic reticulum is a peculiar structure existing in the giant axon of D . gigas and occasionally seen in S . sepioideu. This structure consists of numerous grouped vesicles, 60-100 nm in diameter, and large masses of clustered dense granules, about 40 nm in size (Fig. 7), together forming a massive elongated body that has been followed up to 27 pm without discontinuity (G. M. Villegas, 1969). In S . sepioidea, however, the massive body is much smaller and consists mainly of clustered larger vesicles, 150-500 nm in diameter, with an interposed finely granular dense material (Fig. 8). Along the borders of the massive bodies several mitochondria are usually observed. This peculiar structure has been interpreted as a specialized form of the endoplasmic reticulum involved in glycogen metabolism, especially glycogenolysis, the liberated glucose being utilized by the adjacent mitochondria (G. M. Villegas, 1969). In conclusion, the axoplasm appears as a solid structure formed by disparate elements interconnected by bridges, not only between microtubules and neurofilaments, but also with terminations at the axolemma, on mitochondria, and on the cisternae of the endoplasmic reticulum (Hodge and Adelman, 1980). This organization and relationship of different structural elements appears to be arranged with a high degree of order, and this fact had led A. S. Hodge and W. S. Adelman (personal communication) to use the term “neuroplasmic lattice” to describe the axoplasm. By this term, they mean a quasi-crystalline arrangement comparable to some other molecular or macromolecular arrays existing in other tissues, and

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FIG. 6. Freeze-fracture replica of the giant fiber of S . sepioidea showing the axon (A) and the Schwann cell layer (SC). A large cistern of the endoplasmic reticulum (ER) is observed very close to the axon surface. Rf points a cytoplasmic face with several pits. E, endoneurium.

also comparable to the microtrabecular lattice forming the cytoskeleton described by Porter et al. (1979) and Wolosewick and Porter (1979). Surrounding the axoplasm is the axon plasma membrane or axolemma (Fig. 9a). It presents, in the electron microscope, the asymmetric threelayered pattern that characterizes cell membranes (Robertson, 1957). The thickness of the axolemma varies from 8 to 10.5 nm, the thickness increasing with the axon diameter (R. Villegas et al., 1971). The increase in thickness is due to variations in the thickness of the inner and outer leaflets of the membrane and appears to be related mainly to the deposi-

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FIG.7. Longitudinal section through the axoptasm of the giant fiber of Dosidicds gigns, showing part of a massive body formed by clustered vesicles (V) and a large mass of granules (C) resembling glycogen.

tion of mucopolysaccharides at the outer surface of the plasma membrane, a process that apparently occurs as the animal ages (G. M. Villegas and Villegas, 1968). Besides the typical three-laminar continuous pattern, other regions of the membrane seem to be formed by successive globules, 6-7 nm in diameter (G. M. Villegas and Villegas, 1968). In addition, spaced thickenings of the inner leaflet of the axolemma were also described (Fig. 9b) and later found to be part of structural complexes that involve both the axon and the Schwann cell plasma membranes. These complexes appear to be related to specialized sites for active transport, as well as possible sites for functional interaction between the axon and its satellite glial cell (G. M. Villegas and Villegas, 1976). The structural complex consists of a portion of the axolemma distinctly showing the threelaminar substructure undercoated by a dense material, 0.1 pm in length by 7-17 nm in thickness, and accompanied by a narrowing to disappearance of the axon-Schwann cell interspace (Fig. 9c). These complexes have an

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FIG.8. Massive bodylike structure found in the axoplasm of S. sepioidea giant axon, formed by vesicles, with a fine granular dense material placed among them. Mitochondria (m)are observed around the body.

incidence of 140 per 1000 pm of axon perimeter in the control samples, being appreciably diminished when Na+, K + , and Mg2+are absent from the incubating media, as well as when cardiac glycosides or high concentrations of Ca2+are present. When freeze-fracture replicas of the giant axon are examined, there appear some suggestive images that could be related to the structural complexes. These images consist of the tapering off of the axon-Schwann cell space, as observed in Fig. 16 below, and some sort of structure attached to the axolemma (see Fig. 17). This localization and spatial arrangement of the structural complexes also coincides with the sites of adenosinetriphosphatase (ATPase) (Sabatini et ul., 1968) and acetylcholinesterase (ChE) (G. M. Villegas and Villegas, 1974) activities of the axolemma, as demonstrated by histochemical methods with the electron microscope (Figs. 10 and 11). It is also worth mentioning that treatment of giant axons with detergents, such as

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sodium lauryl sulfate, showed that in the same single cross-section the axolernma was more affected in certain regions, not uniformly (G. M. Villegas and Villegas, 1968). This observation, together with the focal distribution of the enzymatic activities of the ATPase and ChE, the presence of the structural complexes, also distributed in foci, and the coexistence of both types of substructures (three-layered and globular) in the same membrane, are morphological correlates that support the local heterogeneity of the axon membrane. With improved methods of ultrathin sectioning and high-resolution electron microscopy, the axolemma was seen in the early 1950s, but its identification as the axon excitable membrane came later with the development of intracellular perfusion (Baker et al., 1962) and the use of a combined procedure consisting of diffusion of electron-dense markers and simultaneous electrophysiological recording of intracellular potentials from both the axon and the Schwann cell (R. Villegas et al., 1963). Before these findings, Schmitt and Geschwind (1957) had suggested that the excitable membrane could be a composite of the axolemma plus the Schwann cell; somewhat later, Sjostrand (1960) had proposed that the excitable membrane may be formed by the fusion of the axon and Schwann cell plasma membranes, though Frankenhaeuser and Hodgkin (1956) already had suggested the existence of a space interposed between the axon surface and the periaxonal layers. The fortunate existence in one of our tropical squids, S. sepioidea, of a Schwann cell large enough to be punctured with a micropipette inside its cytoplasm, allowed the recording of its membrane potential at rest and during the stimulation of the axon. In addition, the localization of the site where the tip of the microelectrode was located during the recording was established by releasing from the tip one drop of carmine-lithium solution, permitting further identification of the spot using histological methods and phase-contrast microscopy. It was found that, when the axon was depolarized, electrical potential difference across the Schwann cell membrane was not modified. In addition, action potentials were registered only from the axon (R. Villegas et al., 1963). These findings, together with the demonstration of the permeability of the periaxonal spaces to electron-dense markers, such as thorium dioxide (Fig. 12), led us to the identification of the axolemma as the excitable membrane (R. Villegas et al., 1963). The axolernma thus appears as the only effective barrier that discriminates ions and molecules by size and is interposed between the axon interior and the bulk of the extracellular space. Some studies have related the axolemma structurally and functionally

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FIG. 10. Histochemical demonstration of the ATPase activity in the giant axon of D. plei. Deposits of reaction product are observed distributed along the axolemma (arrows). A,

axon; SC, Schwann cell layer.

to the axoplasm and have proposed the existence of an axolemma-ectoplasm complex, which could be isolated by a refined technique for removal of the Schwann cell layer (Metuzals et al., 1981). The axolemma undercoating consists of a network of interwoven filaments that have been associated with maintenance of the normal structural characteristics of the membrane, as well as excitability (Inoue et al., 1974, 1976). Previous studies by Elfvin (1961) described an opaque axoplasmic material undercoating the axolemma at the level of the node of Ranvier in cat sympathetic fibers, and, later, a similar opaque layer was pointed out by Andres (1965) and Peters (1966) in axons of the central nervous system. Peracchia and Robertson (1971) also described an electron-dense material attached to the axonal membrane in crayfish giant nerve fibers and visible after electrical stimulation, asphyxia, or treatment with reducing agents. This alteration was attributed to conformational changes of some membrane proteins that produce unmasking of SH groups and consequently

FIG.9. Three electron micrographs (a, b, c) of the axon (A)-Schwann cell (SC) boundary of D. p k i giant axons, showing the substructure of the axolemma and the Schwann cell plasma membrane. The trilaminar pattern and the septated pattern are observed. In micrograph b, two structural complexes ( S ) appear spaced along the axolemma. Micrograph c, a higher magnification, shows in detail the components of the structural complex: the dense material lining the axolemrna, the continuous trilaminar pattern of the axon membrane, and the close approximation of the axon and the Schwann cell plasma membranes, the intercellular space almost disappearing.

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FIG. 1 I . Histochemical demonstration of the acetylcholinesterase activity in the giant axon of S. sepioidea. Positive reaction is observed distributed in foci along the axolemma (arrows). A . Axon; SC, Schwann cell layer.

the formation of osmium polymers, visualized as the dense undercoating. In S. sepioidea, the axolemma undercoating projects itself radially into the axoplasm (Fig. 13), and these projections are associated with vesicles and mitochondria.

FIG. 12. Part of the giant nerve fiber of D.plei incubated for 1 hr in artificial seawater containing thorium dioxide. The thorium micelles are observed at the endoneurium (E), and the smallest diffuse through the basal lamina (BL) and Schwann cell layer clefts to reach the periaxonal space (circles). A. Axon.

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FIG. 13. Axon (A)-Schwann cell (SC) boundary of the giant axon of S. sepioidea, exhibiting the dense material undercoating the axolemma. This material shows a feltwork texture and is intermingled with tubule-vesicular profiles (arrows) and mitochondria (m). Radial expansions of the material are observed protruding deep into the axoplasm.

B. Schwann Cell Layer This layer is formed by a single row of glial cells closely apposed to the axon surface (Fig. 2). A minute extracellular space separates the axolemma from the Schwann cell plasma membrane. This space measures 5-10 nm across (Fig. 9a), narrowing and often vanishing at the level of the structural complexes described above (Fig. 9b,c). At the outer surface, the Schwann cell layer is lined by a thick basal lamina, 0.2 p m across, which deeply indents the cells, sometimes coming all the way across and approaching the axonal surface (G. M. Villegas, 1969).The basal lamina is formed by a homogeneous material of medium electron density, very well limited and separated from the ground substance of the endoneurium (Fig. 14). In S. sepioidea and D.gigas giant axons, the basal lamina material is intermingled with collagen fibrils penetrating from the inner region of the endoneurium. Each Schwann cell is shaped like a rectangular sheet, measuring about 45 by 60 pm, its thickness varying in the different squid species: 0.20.8 p m in D. plei and 1.5-6 p m in D.gigas ( G . M. Villegas and Villegas, 1968), the thickest portion corresponding to the nuclear region. In S . sepioidea, the Schwann layer is I .5-5 p m thick (R. Villegas et al., l963), and in L. pealei Adelman et al. (1977) have reported a mean thickness of 0.95 p m in cross-sections and 0.91 p m in longitudinal sections. The surface of the Schwann cells is highly irregular, with foldings and invagina-

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FIG. 14. Electron micrograph showing the Schwann cell layer of a giant fiber of S . srpiuideu. This layer is lined at its outer surface by a thick basal lamina ( B L ) that presents deep invaginations (1) going across the layer and almost touching its axonal surface. Fibrils from the innermost region of the endoneurium (E) penetrate the basal lamina ground substance and also the invaginations. A, Axon.

tions, especially the lateral surfaces, which are deeply interdigitated with adjacent cells. By making repeated measurements, Adelman r f crl. (1977) have estimated that the cells are more extensively interdigitated in the longitudinal plane than in the cross-plane, the relationship being 8 or 10 to 4 or 6, respectively. These deep interdigitations of the lateral surfaces plus the pleats and infoldings of the same lateral membranes and also the outer and inner surface plasma membranes, confer on the Schwann cell layer its most striking and peculiar ultrastructural image, the presence of the so-called channels of the Schwann cell layer (Figs. 14 and 15). First visualized by Geren and Schmitt (1954) as double-contoured, osmiophilic, intracellular layers, sometimes arising from the outer membrane of the Schwann cell and reaching the axon-Schwann cell boundary, they were thought to represent the junction of two Schwann cells and were taken as a myelinlike structure. With improved methods, it was possible for us to clearly demonstrate that these tortuous osmiophilic double lines corresponded to pathways or channels, some of them continuous, connecting the extracellular with the axolemma-Schwann cell space (G. M. Villegas and Villegas, 1960). The channel walls were invaginations of the Schwann cell plasma membrane, and each one showed the trilaminar pattern of the unit membrane, as proposed by Robertson (1957). They were described as slitlike channels having a 6-nm lumen in the material embedded in meth-

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FIG. 15. Schwann cell layer (SC) of a Dosidicus gigas giant axon exhibiting the complicated pathway of the clefts. Arrows point to the sites where the clefts connect the axonSchwann cell interspace and the outer extracellular space at the level of the basal lamina (BL). A, Axon.

acrylate. Later, the permeability of the lumen was demonstrated by the penetration of electron-dense thorium dioxide micelles (G. M. Villegas and Villegas, 1964, 1968) (Fig. 12). The term “channel” that we used initially to describe these structures is somewhat confusing nowadays in view of the well-established use of the term “ionic channels” to refer to the molecule routes by which ions cross the excitable and other membranes. For this reason, the term “pathways” or “clefts,” as used by Adelman et al. (1977) to refer to the slitlike intra- and intercellular channels crossing the Schwann layer, seems to be explicit and more appropriate. The tortuous passage of the clefts, as well as the infoldings of the Schwann cell plasma membrane, appear very striking in images obtained after freeze-fracturing of clean giant axons of S . sepioidea species (G. M. Villegas et al., unpublished observations). When the fracture plane occurs more or less perpendicular to the interdigitations, the image observed corresponds well to the narrow clefts seen in thin sections (Fig. 16). When the fracture plane is oblique, the Schwann cell layer clefts appear as membrane faces in superposition (Fig. 17). In them, it is possible to observe pits or craters corresponding either to endocytosis or exocytosis. Another interesting feature revealed by freeze-fracturing is the cytoplasmic face (P face) of the Schwann cell plasma membrane, which reveals a mixed population of particles, as well as zones where exocytotic or endocytotic images are abundant (Fig. 18). These images suggest that

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FIG. 17. Freeze-fractured replica showing the axon (A), Schwann cell layer (SC), and endoneurium (E). The intercellular clefts appear mostly as undulating invaginations (N). At the upper left, part of a P face (Pf) with two pits and part of an E face (Ef) with a crater are observed. Note in the axoplasm a sort of flat structure apposed to the axolemma (arrow).

the cell is very active, a view consistent with observations made in thin sections where the Schwann cell cytoplasm appears rich in all types of membranous profiles, some corresponding to granular endoplasmic reticulum, some to Golgi complexes, and most appearing as smooth cisterns, vesicles, and microvesicles of different sizes (Fig. 19). In addition, Golgi complexes and lysosomes have been observed in the perinuclear region, and bundles of glial filaments have been reported, especially in the giant fiber of Dosidicus gigas (G. M. Villegas, 1969). Two FIG.16. Freeze-fracture replica of the Schwann cell layer of a S. sepioidea giant fiber. Axon (A) and Schwann cell are separated by an intercellular space, which appears to be almost obliterated at certain sites (0).The Schwann cell layer clefts are observed as superposed membrane fractured faces (Pf. Ef) or as narrow clefts in cross-fractures ( N ) . Craters, pits, and small mounds are seen at the E faces of the clefts (arrows), as well as scattered particles that appear to be somewhat larger than the ones covering the P face. Portions of Schwann cell cytoplasm (Cy) are seen among the fractured membranes.

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FIG.18. Freeze-fracture replica of the Schwann cell plasma membrane. The micrograph shows a region of the cytoplasmic face (Pf) where numerous pits occur. Part of the E face appears at the upper left, close to the endoneurium (E). A complementary image of the pit (mound) is observed in that face (arrow).

other rather peculiar, membranous structures have been seen in S. sepioidea and D. plei fibers. One corresponds to membranous profiles, interconnected in an orderly fashion and arranged in a lattice (Fig. 20), similar to the ones first described by Peterson and Pepe (1961) in cells surrounding the inhibitory synaptic endings of the crayfish stretch receptors, and later by Peracchia and Robertson (1971) and by Liebermann et al. (1981) in the giant nerve fibers of the same crayfish. In the lobster, Holtzman et al. (1970) reported the existence of the tubular lattice as patches of anastomosing tubules connected to the axon-Schwann cell space and also to the clefts of the Schwann cell layer. In the squid, this structure is not as abundant as in crayfish, but it has been seen opening into the lumina of the clefts. The other structure, found occasionally, appears as a pile of elongated flattened cisterns, about 2 pm long and formed by membranes 6 nm thick (Fig. 21). The appearance of the Schwann cell cytoplasm, as revealed by both thin-sectioning and freeze-fracture techniques, correlates well with the

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FIG.19. Thin section through the Schwann cell cytoplasm at the level of the paranuclear region. Different types of membrane vesicles, granules, and filaments, as well as tubular profiles and cisterns of the granular endoplasmic reticulum, fill all the space between the basal lamina (B1,) and the axon ( A ) boundary. In the axoplasm. clujlers of small. mediumdense vesicles are observed toward the surface.

high physiological activity attributed to this cell. Coupling of the axon and its satellite glia by a cholinergic system has been reported in the squid by J. Villegas (1972, 1973, 1974) and is described in detail in the article by J. Villegas in this volume. Schwann cells contain and synthesize acetylcholine (J. Villegas and Jenden, 1979; Heumann ef ul., 1981), and the axon houses acetylcholinesterase at the inner surface of the axolemma, as demonstrated by histochemical methods for electron microscopy ( G . M. Villegas and Villegas, 1974). The craters and pits corresponding to endocytosis or exocytosis of the Schwann cell could be homologs of similar profiles observed in synapses and attributed to the liberation of neurotransmitters (Pumplin and Reese, 1978). Thus, it can be speculated that some crater images may be related to sites of transference of the acetylcholine toward the axon. In a similar way, the craters could also be related to sites of passage toward the axon of proteins or other macromolecules synthesized in the Schwann cell cytoplasm, as described by Lasek et al. (1974).

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GLORIA M. VILLEGAS AND RAIMUNDO VILLEGAS

FIG.20. Part of the Schwann cell cytoplasm of a S. sepioideu giant axon showing the tubular lattice formed by interconnected membranous tubules. Some tubules seem to be connected to the Schwann cell clefts (arrows). BL, basal lamina.

The Schwann cell nucleus is also sheet shaped and contains a fine granular material with some larger granules scattered in the center and peripherally situated nucleoli (Fig. 22). The nucleus almost fills the cell space, the perikaryal cytoplasm being reduced to a thin rim. Nuclei as long as 24 pm have been seen in cross-sections of S. sepioideu giant axons. As many as eight nuclei have been counted in a single crosssection; however, the number of cells constituting the sheath at a single level is larger, since not all the nuclei are intercepted by the section (G. M . Villegas and Villegas, 1963). When the clefts that completely traverse the Schwann cell are counted, the number of cells can be doubled or tripled, as reportcd also for the crayfish medial giant axon (Lieberman et al., 1981). However, in the squid, such enumeration is more difficult owing to the presence of incomplete foldings from the external and internal aspects of the Schwann cells. Adelman et a / . (1977) have estimated the number of cells by counting the number of intercellular clefts abutting the periaxonal space in both cross and longitudinal sections, and taking into

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FIG.21. Part of the giant nerve fiber of S. srpioideci showing the axon (A), the Schwann cell layer (SC), and the endoneurium (E). In the Schwann cell cytoplasm, a pile of flattened, elongated cisterns formed by thin membranes occupies a space among the clefts.

consideration a characteristic invagination of the Schwann cell into the axon at each of the intercellular unions, as well as the average folding and branching of the interdigitation. They estimated, in L . peulsi fibers, 125 clefts entering the periaxonal space in a single cross-section of an axon 500 pm in diameter.

FIG.22. Part of a Schwann cell nucleus exhibiting a concentration of dense granules toward the central region and two peripheral nucleoli (NC).

GLORIA M. VILLEGAS AND RAIMUNDO VILLEGAS

C. Endoneurium The endoneurium consists of an almost regularly organized zone of connective tissue, averaging 6 pm thick in clean fibers, and formed by alternating layers of fibrocytes and collagen fibrils. This zone is interposed between the giant fiber and the bundles of small fibers and usually contains a blood vessel of very peculiar aspect (G. M. Villegas, 1969). Close to the giant fiber, the 2-pm-thick innermost region of the endoneurium is devoid of cells (Figs. 14 and 15) and appears to be populated by loosely or tightly packed fibrils, 20-30 nm thick, and with axial periodicity consistent with that of some type of embryonic collagen (Porter and Pappas, 1959). Adelman et al. (1977) have reported positive biochemical evidence for the existence of collagen by finding hydroxyproline in hydrolysates of squid axon sheaths. In certain squid species, such as L . ppalei, or D . plei, there is a distinct borderline between the basal lamina covering the Schwann cell layer and the closest endoneurial space. However, in Dosidicrrs gigcis and S. sepioidm, as mentioned above, the collagen fibrils penetrate the basal lamina and intermingle with its amorphous material; moreover, fibrils are seen in the invaginations of the basal lamina into the Schwann cytoplasm (G. M. Villegas, 1969) (Fig. 14). The rest of the endoneurial region exhibits a layered organization constituted by layers of fibrocytes separated by spaces filled with fibrils (Fig. 23). The fibrocytic cells are elongated units formed by a nucleus surrounded by a rim of cytoplasm that extends in long, delicate polar processes. Although orderly, and arranged in a laminated fashion, the endoneurium in the squid giant fiber does not appear to be a tight barrier for diffusion, since thorium dioxide micelles can penetrate all the way to the basal lamina of the Schwann cell layer (G. M. Villegas and Villegas, 1968). Investigations on the permeability of the giant axon to glycerol showed that a facilitated diffusion mechanism, evidenced from the efflux curve, could be attributed to the endoneurial cells, since their number and size (about 10 pm long by 2 pm thick, plus the long fine processes) agreed with the size calculated for a glycerol compartment of 4-16 p m (G. M. Villegas and Villegas, 1964). This calculated periaxonal compartment also could be correlated with the ionic X-phase of Shanes and Berman (1955) and the superficial ionic layer of Caldwell and Keynes (1960). A layer, similar in organization to the perineurium of vertebrate nerves, has not been observed in the squid around the giant fiber and the other nerve fibers of the bundle.

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FIG.23. Part of the endoneurial region of Dosidicus gigas fiber in which a fibrocyte (F) appears sectioned at the level of its nucleus (N). Note the alternation of cell processes (P) and collagen bundles (f).

V.

NERVE FRACTIONS

After its identification, attempts to isolate the axon-excitable membrane were directed to obtaining pure preparations. This was first accomplished by using a sequence of differential, discontinuous, and gradient centrifugations in sucrose solutions (R. Villegas and Camejo, 1968; Camejo e t a / . , 1969). Two membrane fractions were isolated from homogenates of D. gigas stellate nerves. One fraction, membrane fraction I , banded at a density of 1.090 g/ml, and the other, membrane fraction 11, banded at 1.140 giml, the difference in densities being produced by dissimilar protein to lipid ratios-29.5: 70.5 and 48.3: 51.7, respectively. The electron microscopic study of both fractions (Figs. 24 and 25) revealed that they consist exclusively of vesicles formed by a singlemembrane wall, measuring 9.5 to 11 nm thick in fraction I and 7 . 2 to 10 nm thick in fraction 11. Vesicles were greater in diameter and appeared cleaner in fraction I than in fraction 11. A moderately dense, nonhomogeneous material appeared attached to both membrane surfaces in fraction 11, and frequently it was interposed between the membranes of adjacent vesicles. On the other hand, the membranes forming the vesicles of fraction I showed a tendency to coalesce and form multilayered structures like myelin figures. The yield of the fractions, the ultrastructural features of the membrane fractions, and the distribution of the ATPase activity,

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FIG.24. Thin section of a pellet of membrane fraction I (axolemma-enriched preparation), prepared from giant nerve fibers of Dosidicus gigas squid. The pellet consists mainly of large vesicles formed by a single, clean membrane.

when compared with the findings in intact nerves, led Camejo et al. (1969) to identify fraction I tentatively as an axolemma-enriched preparation and fraction I1 as a preparation enriched in Schwann cell plasma membranes. This method of isolation was used later to obtain membrane fractions from lobster walking-leg nerves (Barnola et al., 1973) and squid optic nerves (Freund, 1976). In the garfish olfactory nerve (Chacko et al., 1974a)and in the squid optic nerve (Fischer et al., 1970), other procedures have also been applied to obtain axolemma-enriched preparations. The electron microscopic examination of thin sections of the pellets of the two fractions obtained from the optic nerves of S. sepioidea (G. M. Villegas, R. Villegas, and F. V. Barnola, unpublished results) revealed that fraction 1 is formed by a fairly uniform population of single-walled round vesicles, 160-340 nm in diameter (Fig. 26); membrane fraction I1 consists also of vesicles, less uniform in size and shape and less packed than those of fraction I, and usually intermingled with contaminants, such as mitochondria (Fig. 27). The yield of the fractions, as well as their

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FIG. 25. Thin section of a pellet of membrane fraction I1 (extra-axonal membranesenriched preparation), prepared from Dusidicus gigas giant nerve fibers. This fraction is mainly formed by vesicles smaller than those of fraction I and, usually, with a dense, amorphous material attached to the membranous walls.

ultrastructural aspect, also correlates well with the morphological features of the intact optic nerve. The isolation of squid nerve fiber membranes has opened the possibility of exploring the location and molecular counterparts of some membrane properties and functions. Nerve membranes isolated by the same procedure from walking-leg nerves of lobster have been used in our most recent work. The demonstration of tetrodotoxin receptors in vesicles spontaneously formed by the lobster membranes was considered an indication of the presence of sodium channels. The maximum binding of tritiated tetrodotoxin, in picomoles per milligram of protein, was 10.1 for the lobster fraction I vesicles (Barnola et al., 1974b) and 5.6 for the same fraction of the squid optic nerve (Freund, 1976). The presence of functional sodium channels in these membrane vesicles was later reported by Barnola and Villegas (1976). They showed that the sodium efflux from vesicles previously loaded with 22Nawas increased by veratrine and the increment was

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FIG.26. Thin section of a pellet of membrane fraction 1 isolated from optic nerves of the squid S. sepioideu. This fraction consists of nunierous round vesicles. each formed by a clean single membrane.

abolished by the addition of tetrodotoxin. A further step was achieved with the incorporation of the sodium channel-containing membrane fragments into artificial liposomes prepared with soybean phospholipids (R. Villegas et ul., 1977). Attempts to solubilize and purify the sodium-channel protein components from peripheral nerves have been reported (R. Villegas et al., 1980, 1981; R. Villegas and Villegas, 1981) and are the subject of our present investigations. VI.

CONCLUDING REMARKS

In the present work we have tried to make a somewhat broad review of the morphology of the squid giant axon, as revealed mainly by the electron microscope. It has also been our intention to try to correlate the structural details with the functional behavior of the giant fiber. It seems appropriate to summarize the most remarkable structural features of this preparation: ( I ) the large size of the giant axon, over 200 /*m

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FIG.27. Thin section of a pellet of membrane fraction I1 isolated from optic nerves of the squid S. sepioideu. As observed, this fraction is not homogeneous; it is composed of very irregular membranous vesicles, mitochondria, and some dense, amorphous material.

in diameter in most species; (2) the apparent polarization of the axoplasm constituents: neurofilaments, microtubules, endoplasmic reticulum, and possibly mitochondria, which appear to be more concentrated toward the periphery; (3) the characteristic Schwann layer, closely apposed to the axon surface and constituting a network of extracellular spaces; (4) the intimate contact, at certain specialized sites, between the axon and its satellite Schwann cell plasma membranes. In addition, the usefulness of the squid axon has been mentioned as a source for preparing membrane fractions. The latter have been important for biochemical and pharmacological studies in the neurosciences and have led to methods for isolation and purification of components of excitable membranes, such as the sodium channel. We can paraphrase Rosenberg’s words in his excellent and amusing review on methods and applications of the squid axon (1981)and say that, after collecting the material for the present article, we have gained a systematic insight into the organization and fine structure of the squid giant nerve fiber.

34

GLORIA M. VILLEGAS AND RAIMUNDO VILLEGAS

ACKNOWLEDGMENTS The authors express their thanks to Drs. Guillermo Whittembury and Jaime Requena for critical reading of the manuscript; to Mr. Freddy Sanchez for technical assistance; to Mr. Mardonio Diaz for photographic labor; to Messrs. JosC Luis Alvarez and Carlos Quintero for the drawings, and to Miss Isabel Otaegui and Mrs. Sol Van Praag de Rojas for secretarial help. REFERENCES Adelman, W. J., Moses, J., and Rice, R. V. (1977). An anatomical basis for the resistance in series with the excitable membrane of the squid giant axon. J. Neurocytol. 6 , 621-646. Andres, K. H. (1965). Uber die Feinstruktur besonderer Einrichtugen in markhaltigen Nervenfasern des Kleinhirns der Ratte. Z . Zellforsch. Mikroskop. Annt. 65, 701-712. Baker, P. F., Hodgkin, A. L., and Shaw, T. I. (1962). Replacement of the axoplasm of giant nerve fibers with artificial solutions. J . Physiol. (Londonj 164, 330-354. Barnola, F. V., and Villegas, R. (1976). Sodium flux through the sodium channels of axon membrane fragments isolated from lobster nerves. J. Gen. Physiol. 67, 81-90. Barnola, F. V., Villegas, R., and Camejo, G. (1973). Tetrodotoxin receptors in plasma membranes isolated from lobster nerve fibers. Eiochim. Biophys. Acta 298, 84-94. Bear, R. S . , Schmitt, F. O., and Young, J. Z. (1937a). The ultrastructure of nerve axoplasm. Proc. R . SOC. Ser. E 123, 505-519. Bear, R. S., Schmitt, F. O., and Young, J. Z. (1937b). Investigations on the protein constituents of nerve axoplasm. Proc. R . SOC. Ser. B 123, 520-529. Caldwell, P. C., and Keynes, R. D. (1960). The permeability of the squid giant axon to radioactive potassium and chloride ions. J. Physiol. (London) 154, 177-189. Camejo, G., Villegas, G. M., Barnola, F. V., and Villegas, R. (1969). Characterization of two different membrane fractions isolated from the first stellar nerve of the squid Dosidicus gigas. Biochim. Biophys. Acta 193, 247-259. Chacko, G . K., Goldman, D. E., Malhotra, H. C., and Dewey, M. M. (1974a). Isolation and characterization of plasma membrane fractions from garfish Lepisosteus osseus olfactory nerve. J. Cell B i d . 62, 831-843. Chacko, G . K . , Barnola, F. V., Villegas, R., and Goldman, 13. E. (1974b). The binding of tetrodotoxin to axonal membrane fractions isolated from garfish olfactory nerve. Biochim. Biophys. Acta 373, 308-3 12. Chambers, R., and Kao, C. Y. (1952). The effect of electrolytes on the physical state of the nerve axon of the squid and of Stentor. a protozoan. Exp. Cdl R c s . 3, S64-.573. Cohen, L. B . , Keynes, R. D., and Hille, B. (1968). Light scattering and birefringence changes during nerve activity. Niltitre (Lundon) 218, 438-441. Costantin, L. L., Franzini-Armstrong, C.. and Podolsky, R. J. (1965). Localization of calcium-accumulating structures in striated muscle fibers. Science 147, 158-160. Davison, P. F . , and Huneeus, F. C. (1970). Fibrillar proteins from squid axons. 11. Microtubule protein. J. Mu/. B i d . 52, 429-439. Elfvin, L.-G. (1961). The ultrastructure of the nodes of Ranvier in cat sympathetic nerve fibers. J . Ultrastruct. Res. 5 , 374-387. Endo, S., Sakai, H., and Matsumoto, G. (1979). Microtubules in squid giant axon. Ce// Struct. Funcr. 4, 285-293. Fischer, S.. Cellino, M., Zambrano, F., Zampighi. G., Tellez-Nagel, M., Marcus, D., and Canessa-Fischer, M. (1970). The molecular organization of nerve membranes. I . Isolation and characterization of plasma membranes from the retinal axons of the squid: An axolemma-rich preparation. Arch. Biochem. Biophys. 138, 1-15.

SQUID AXON ULTRASTRUCTURE

35

Frankenhaeuser, B., and Hodgkin, A. L. (1956). The after-effects of impulses in the giant nerve fibers of Loligo. J . Physiol. (London) 13, 341-376. Freund, P. (1976). Aislamiento y caracterizacibn de las membranas plasmaticas del nervio dptico del calamar Sepio/pu//tissepioideri. Master’s thesis, Sim6n Bolivar University, Caracas. Geren, B. B . , and Schmitt, F. 0. (1954). The structure of the Schwann cell and its relation to the axon in certain invertebrate nerve fibers. Proc. N u / / . A(,crd. Sci. U . S . A . 40, 863-870. Henkart, M. P., Reese, T. S . , and Brinley, Jr., F. J . (1978). Endoplasmic reticulum sequesters calcium in the squid giant axon. Science 202, 1300-1303. Heumann, R., Villegas, J., and Herzfeld, D. W. (1981). Acetylcholine synthesis in the 36, . Schwann cell and axon in the giant nerve fiber of the squid. J . N e u r o c ~ / i ~ m 765-768. Hodge, A. j.,and Adelman, W. J. (1980). The neuroplasmic network in Loligo and Hermissenda neurons. J . Ultrastruci. Res. 70, 220-241. Hodgkin, A. L., and Keynes, R. D. (1956). Experiments on the injection of substances into squid giant axons by means of a microsyringe. J . Physiol. (London) 131, 592-616. Holtzman, E., Freeman, A. R., and Kashner, L. A. (1970). A cytochemical and electron microscope study of channels in the Schwann cells surrounding lobster giant axons. J . Cell B i d . 44,438-445. Inoue, I., Tasaki, I., and Kobatake, Y. (1974). A study of the effects of externally applied sodium-ions and detection of spatial nonuniformity of the squid axon-membrane under internal perfusion. Eiophys. Chem. 2, 116-126. Inoue, I., Pant, H. C., Tasaki, I . , and Gainer, H. (1976). Release of proteins from the inner surface of squid axon membrane labeled with tritiated N-ethylmaleimide. J . Gen. Physiol. 68, 385-395. Lasek, R. J., Gainer, H., and Przybylski, R. J. (1974). Transfer of newly synthesized proteins from Schwann cells to the squid giant axon. Proc. Nut/. Acad. Sci. U . S . A .71, 1188-1 192. Lieberman, E. M., Villegas, J., and Villegas, G. M. (1981). The nature of the membrane potential of glial cells associated with the medial giant axon of the crayfish. N e ~ r o science 6 , 261-271. Martin, R. (1969). The structural organization of the intracerebral giant fiber system of cephalopods. The chiasma of the first order giant axons. Z . Zellforsch. 97,50-68. Matsumoto, G., and Sakai, H . (1979). Microtubules inside the plasma membrane of squid giant axons and their possible physiological function. J . Membr. B i d . 50, 1-14. Matsumoto, G . , Kobayashi, T., and Sakai, H. (1979). Restoration of the excitability of squid giant axon by tubulin-tyrosine ligase and microtubule proteins. J . Bio(Ac,m. 86, 1155-1 158.

Metuzals, J . (1969). Configuration of a filamentous network in the axoplasm of the squid (Loligo pealii L.) giant nerve fiber. J . Cell Biol. 43, 480-505. Metuzals, J., and Izzard, C. S. (1969). Spatial patterns of threadlike elements in the axoplasm of the giant nerve fiber of the squid (Loligo peulii L.) as disclosed by differential interference microscopy and by electron microscopy. J . Cell B i d . 43, 456-479. Metuzals, J . , and Tasaki, I. (1978). Subaxolemmal filamentous network in the giant nerve fiber of the squid (Loligopealei L.) and its possible role in excitability. J . Cell Eiol. 78, 597-62 1. Metuzals, J . , Tasaki, I., Terakawa, S., and Clapin, D. F. (1981). Removal of Schwann sheath from the giant nerve fiber of the squid: An electron-microscopic study of the axolemma and associated axoplasmic structures. Cell Tissue Res. 221, 1-15.

36

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M. VILLEGAS AND RAIMUNDO VILLEGAS

Peracchia, C., and Robertson, .I. D. (1971). Increase in osmiophilia of axonal membranes of crayfish as a result of electrical stimulation, asphyxia, or treatment with reducing agents. J . Cell B i d . 51, 223-239. Peters, A. (1966). The node of Ranvier in the central nervous system. Q. J . Exp. Physiol. 51, 229-236. Peterson, R. P., and Pepe, F. A. (1961). The fine structure of the inhibitory synapses in the crayfish. J . Biophys. Biochem. Cyrol. 11, 157-169. Porter, K . R., and Pappas, G. D. (1959). Collagen formation by fibroblasts of chick embryo dermis. J . Biophys. Biochem. Cyrol. 5, 153-166. Porter, K. R., Byers, H. R., and Ellisman, M. H. (1979). The cytoskeleton. I n “The Neurosciences, Fourth Study Program” (F. 0. Schmitt and F. G . Worden, eds.), pp. 703-722. MIT Press, Cambridge, Massachusetts. Pumplin, D. W., and Reese, T. S. (1978). Membrane ultrastructure of the giant synapse of the squid Loligo pealei. Neuroscience 3, 685-696. Richards, A. G . , Jr., Steinbach, H. B . , and Anderson, T. F. (1943). Electron microscope studies of squid giant nerve axoplasm. J. Cell. Cornp. Physiol. 21, 129-143. Robertson, J. D. (1957). New observations on the ultrastructure of the membranes of frog peripheral nerve fibers. J . Biophys. Biochem. Cyrol. 3, 1043-1048. Rosenberg, P. (1981). The squid giant axon: Methods and applications. In “Methods in Neurobiology” (R. Lahue, ed.), Vol. 1, pp. 1-133. Plenum, New York. Sabatini, M . R., DiPolo, R., and Villegas, R. (1968). Adenosin triphosphatase activity in the membranes of the squid nerve fiber. J . Cell B i d . 38, 176-183. Sakai, H . , and Matsumoto, G . (1978). ’Tubulin and other proteins from squid giant axon. J . Biochem. 83, 1413-1422. Schmitt, F. 0.. and Geschwind, N . (19571. The axon surface. Profi. Biopl7ys. C k m . 8, 165-2 15.

Shanes, A. M.. and Berman, J. (1955). Kinctics of ion movements in the squid giant axon. J . Gun. Physiol. 39, 279-300. Sjostrand, F. (1960). Electron microscopy of myelin and of nerve cells and tissue. In “Modern Scientific Aspects of Neurology” (J. N. Cumings, ed.), pp. 188-231. Arnold, London. Villegas, G . M. (1960). Electron microscopic study of the giant nerve fiber of the giant squid Dosidicus gigus. J . U1trasrricc.t. Res. 26, 501-514. Villegas, G . M., and Villegas, R. (1960). The ultrastructure of the giant nerve fiber of the squid: Axon-Schwann cell relationship. J. Ulfrastruct.Reg. 3, 362-373. Villegas, G. M., and Villegas, R. (1962). The endoneurium cells of the squid giant nerve and their permeability to 14C-glycerol.Biochim. Biophys. Acra 60, 202-204. Villegas, G. M . , and Villegas, R. (1963). Morphogenesis of the Schwann channels in the squid nerve. J . Ultrusrruct. Res. 8, 197-205. Villegas, G. M., and Villegas, R. (1964). Extracellular pathways in the peripheral nerve fibers: Schwann-cell-layer permeability to thorium dioxide. Biochim. Biophys. Acra 88, 231-233. Villegas. G . M.,and Villegas. R . (1968). Ullrastructurnl studies of the squid nerve fibers. J . G1.n. Physiol. 51, 44s-60s. Villegas, G. M.. and Villegas, J . (1974). Acetylcholinesterase localization in the giant nerve fiber of the squid. J. Ulfrustrucr. R e s . 46, 149-163. Villegas, G. M., and Villegas, J. (1976). Structural complexes in the squid axon membrane sensitive to ionic concentrations and cardiac glycosides. J . CeII R i d . 69, 19-28. Villegas, C. M . , Lane, N. J . , and Villegas, J . (1984). In preparation.

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Villegas, J. (1972). Axon-Schwann cell interaction in the squid nerve fiber. J . Physiol. (London) 225, 275-296. Villegas, J . (1973). Effects of tubocurarine and eserine on the axon-Schwann cell relationship in the squid nerve fiber. J . Physiol. (London) 232, 193-208. Villegas, J. (1974). Effects of acetylcholine and carbamylcholine on the axon and Schwann cell electrical potentials in the squid nerve fiber. J . Physiol. (London) 242, 647-659. Villegas, J., and Jenden, D. J. (1979). Acetylcholine content of the Schwann cell and axon in the giant nerve fiber of the squid. J . Neurochem. 32, 761-766. Villegas, R., and Carnejo, G. (1968). Tetrodotoxin interaction with squid nerve membrane lipids. Biochim. Biophys. Actu 163, 421-423. Villegas, R., and Villegas, G. M. (1981). Nerve sodium channel incorporation in vesicles. Annu. Rev. Biophys. Bioeng. 10, 387-419. Villegas, R., Villegas, L., Gimenez, M., and Villegas, G. M. (1963). Schwann cell and axon electrical potential differences. Squid nerve structure and excitable membrane location. J . Gen. Physiol. 46, 1047-1064. Villegas, R., Villegas, G . M., DiPolo, R., and Villegas, J. (1971). Nonelectrolyte permeability, sodium influx, electrical potentials, and axolemma ultrastructure in squid axons of various diameters. J . Gen. Physiol. 57, 623-637. Villegas, R., Villegas, G. M., Barnola, F. V., and Racker, E . (1977). Incorporation of the sodium channel of lobster nerve into artificial liposomes. Biochem. Biophys. Res. Cornmitn. 79, 210-217. Villegas, R., Villegas, G. M., Condrescu-Guidi, M., and Suarez-Mata, Z. (1980). Characterization of the nerve membrane sodium channel incorporated into soybean liposomes: A sodium channel active particle. Ann. N . Y. Acad. Sci. 358, 183-203. Villegas, R., Villegas, G . M., and Suarez-Mata, Z. (1981). Reconstitution of the sodium channel with partially solubilized lobster nerve membrane. J . Physiol. (Puris) 77, 1077-1 086. Weisenberg, R. C. (1972). Microtubule formation in vitro in solutions containing low calcium concentrations. Science 177, 1104-1 107. Witman, G., and Rosenbaum, J. (1973). Filamentous components of isolated squid axoplasm. B i d . Bull. 145, 460. Wolosewick, J. J., and Porter, K. R. (1979). Microtrabecular lattice of the cytoplasm ground substance. Artifact or reality. J . Cell B i d . 82, 114-139. Young, J . Z. (1934). Structure of nerve fibers in Sepiu. J . Physiol. (London) 83, 27P-28P. Young, J. Z. (1936). The structure of nerve fibers in Cephalopods and Crustacea. Proc. R . Soc. Ser. B 121, 319-337. Young, J. Z. (1939). Fused neurons and synaptic contacts in the giant nerve fibers of cephalopods. Phil. Trans. R. Soc. London Ser. B 229, 465-503.

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CURRENT TOPICS IN MEMBRANES A N D TRANSPORT, VOLUME 22

The Structure of Axoplasm RAYMOND J . LASEK Neurobiology Center Department of Developmental Genetics and Anutomy Case Western Reserve University Cleveland, Ohio

I. Introduction.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Fine Structure of the Squid Gian A . The Axoplasmic Matrix . . . B. Cross-Linking Elements in t C. Domains in the Axoplasmic D. Microfilaments in Axoplasm 111. Axonal Transport in the Squid Giant Axon 1V. Extruded Axoplasm from the Giant Axon ............... V. Conclusions . . . . . . . . . . . . . . . . . .

1.

49

INTRODUCTION

The squid giant axon is well known from numerous studies that have contributed significantly to our understanding of the properties of the action potential and the physical properties of excitable membranes. It is, of course, the large size of the giant axon in many species of squid that has made this preparation such a powerful biological model for membrane physiologists. The large size of the giant axon also makes it useful for studies of other aspects of axonal biology. It is possible to separate the axoplasm of the giant axon from the surrounding sheath so as to study the chemical composition of the axoplasm independent of the materials present in the nonneuronal cells that surround the axon (Bear et d., 1937; Roberts et al., 1958; Baker and Shaw, 1965; Lasek et al., 1973, 1979). Furthermore, it has been shown that fast axonal transport continues for 39 Copyright 0 1984 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN O-IZ-I53322-0

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RAYMOND J. LASEK

hours after the axoplasm is removed from the giant axon (Brady et al., 1983). This observation suggests that the isolated axoplasm retains most of its structural integrity and physiological capabilities in the absence of the plasma membrane. The purpose of this brief article is to evaluate some of the information that has been obtained by studying axoplasm isolated from the squid giant axon. One particularly important issue concerns the generality of this information to other axons. That is, to what extent is information obtained about the structure, chemistry, and physiology of axoplasm from the squid giant axon applicable to smaller axons in animals other than the squid? The possible similarities between the giant axon and vertebrate axons are of particular interest, because so much information is already available regarding the structure and function of vertebrate axons. This question of generality can be answered by evaluating the structure and chemistry of the giant axon and comparing it with axons from other animals.

II.

FINE STRUCTURE OF THE SQUID GIANT AXON

A. The Axoplasmic Matrix

Electron microscopic investigation of the giant axons of Loligo demonstrates that the structure of these axons is very similar to that of smaller molluscan axons (e.g., compare the studies of Hodge and Adelman, 1980, with those of Metuzals ~t d.. 19X3b). The primary structural component of any axon is the cytoplasmic matrix. The cytoplasmic matrix has been defined as all of the constituents of the cytoplasm exclusive of the membranous organelles such as the endoplasmic reticulum and mitochondria.' The cytoplasmic matrix includes the cytoskeleton, which consists of the three major cytoplasmic polymers (i.e., microtubules, intermediate filaments, and microfilaments). These polymers are crosslinked into a network that provides a structural framework for the cytoplasmic matrix.

'

The concept of the cytoplasmic matrix has evolved very rapidly during the last few years in order to encompass recent developments in electron microscopy. Dr. Keith Porter has played a particularly important role in developing the contemporary view of this subject. I queried him about a definition for the cytoplasmic matrix at a recent meeting entitled "The Cytoplasmic Matrix and the Integration of Cellular Function." which was held at the Fogarty International Center in Bethesda, Maryland. He proposed the definition that I have included here. A full report of this meeting will be published soon in the Journal of Cell Biology.

THE STRUCTURE OF AXOPLASM

41

Analyses of the cytoplasmic matrix of the squid giant axon, using conventional methods of transmission electron microscopy, indicate that neurofilaments and microtubules are the dominant structural components (Metuzals, 1969; Krishnan et al., 1979; Lasek et al., 1979). Structurally, the axoplasm of the giant axon is remarkably similar to that of large myelinated vertebrate axons (Metuzals et al., 1981a). In both cases, neurofilaments are more abundant than microtubules. This tends to be a general rule for large axons. That is, as axons increase in size during development, the number of neurofilaments that are added to the axon increases more rapidly than do the number of microtubules (Friede and Samorajski, 1970). Small axons tend to have more microtubules than neurofilaments but this ratio becomes reversed during development.

B. Cross-Linking Elements in the Axoplasmic Matrix

Another basic similarity between the cytoskeleton of the giant axon and vertebrate axons is the presence of a well-developed system of crossbridges that interconnect the neurofilaments and microtubules (Wuerker and Kirkpatrick, 1972; Ellisman and Porter, 1980; Metuzals et al., 1982). There appear to be at least two types of cross-bridges [i.e., those associated with neurofilaments and those associated with microtubules (Peters et al., 1976)l. The cross-bridges on the neurofilaments are probably formed by the extension of a region of the neurofilament proteins (Willard and Simon, 1981; Sharp et al., 1982). In particular, the high-molecularweight subunit of the triplet has been implicated in neurofilament crosslinking but evidence also indicates that all of the neurofilament subunit proteins may have a region that can be extended to form a projecting cross-bridge from the neurofilament (Geisler et al., 1983). The crossbridges that extend from microtubules apparently consist of the microtubule-associated proteins (MAPs) (Kirschner, 1978). In vertebrate axons, it has been shown that high-molecular-weight (HMW) MAPs can coat the surface of microtubules to form microtubules with cross-bridges that are similar in appearance to those observed in neurons (Kirschner, 1978). It is of interest that an HMW-MAP has been identified in squid brain ( G . Langford, personal communication) and that this HMW-MAP is present in squid axoplasm (Lasek and Morris, unpublished). The following evidence suggests that the squid HMW-MAP is the principal cross-bridge between the microtubules of the squid giant axon (Morris et al., 1981; Lasek, Morris, and Hodge, unpublished results). A cytoskeletal preparation consisting primarily of microtubules can be prepared from axoplasm by first stabilizing the microtubules in axoplasm

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RAYMOND J. LASEK

with taxol and then digesting the neurofilaments of the axoplasm using the intrinsic Ca2+-activatedprotease. This procedure produces a preparation from which most of the cytoplasmic matrix proteins have been extracted. The primary components that remain in these taxol-Ca2+-treated preparations are tubulin, the HMW-MAPS, and a fragment of the 60-kd neurofilament subunit, which is resistant to the intrinsic Ca2+-activatedprotease. Transmission electron micrographs of thin sections of the taxol-Ca2+ preparations reveal microtubules cross-linked by bridges. These bridges can be eliminated by adding trypsin to the preparation, and we have found that the HMW-MAPS are degraded simultaneously with the disappearance of the cross-bridges. The trypsin has no effect on the tubulin when analyzed by SDS-PAGE, and it has no effect on the integrity of the microtubules. When the cross-bridges disappear the taxol-Ca2+ preparation loses its stability and no longer behaves like a gel. That is, the taxolCa2+-treatedaxoplasm can be manipulated with forceps prior to the addition of trypsin; however, after trypsin treatment and the elimination of the cross-bridges, forceps rip right through the preparation and it no longer behaves like a solid piece of axoplasm. These observations suggest that HMW-MAPS are important structural components of the axoplasm because they cross-link the microtubules. C. Domains in the Axoplasmic Matrix

1. THESUBPLASMALEMMAL DOMAIN Recently, it has been noted that the cytoskeleton of large vertebrate axons can be divided into subdomains (Schnapp and Reese, 1982; Hirokawa, 1982). One domain is subjacent to the plasma membrane. This domain, which is approximately 100 nm thick, contains short, 4- to 5-nm filaments that appear to be associated with the plasma membrane (Metuzals and Tasaki, 1978; Schnapp and Reese, 1982; Hirokawa, 1982). Some of the 4- to 5-nm filaments are actin microfilaments (Hirokawa, 1982). A similar domain has been found near the plasma membrane of the squid giant axon and some of the microfilaments in this region can be decorated with subfragments of myosin, indicating that they are actin filaments (Metuzals and Tasaki, 1978). 2. MICROTUBULE A N D NEUROFILAMENT DOMAINS

The central region of the axoplasm can be divided into two additional domains: microtubule domains and neurofilament domains (Schnapp and Reese, 1982). That is, microtubules and neurofilaments tend to pref-

THE STRUCTURE OF AXOPLASM

43

erentially self-associate, forming bundles of either microtubules or neurofilaments. In the squid giant axon, the microtubules appear as bundles surrounded by neurofilaments because there are so many more neurofilaments than microtubules in the giant axon (Metuzals et al., 1983a,b). This segregation is illustrated in Fig. 1, which shows a lowpower electron micrograph of the squid giant axon. The microtubule domains have a very different substructure from that of the neurofilament domains. In Fig. I , the microtubule domains are much more electron dense than the neurofilament domains. This increased density apparently represents a higher concentration of protein components in the microtubule domains of the axoplasm. This higher density in the microtubule domains apparently represents cytoplasmic matrix proteins that preferentially associate with nondiffusable components organized by the microtubules. Schnapp and Reese (1982) have noted that the microtubule domains in turtle optic axons contain globular and filamentous structures that are much less abundant in the neurofilament domains. These globular and filamentous structures of the cytoplasmic matrix may correspond to the complex constellation of proteins that are transported slowly in the subcomponent of slow transport called slow component b (SCb). If this is the case, then the microtubule domains are apparently involved in the organization of a vast array of proteins such as actin, clathrin, calmodulin, and many metabolic enzymes because all of these proteins are present in SCb (Brady et al., 1981; Brady and Lasek, 1981; Brady and Lasek, 1982a,b; Garner and Lasek, 1982; Schnapp and Reese, 1982; Hirokawa, 1982). The neurofilament domains appear to be much simpler than the microtubule domains. The regions of the axoplasm occupied by neurofilaments are populated principally by the neurofilaments and their side bridges (Fig. 1). Apparently, the most abundant proteins of the cytoplasmic matrix tend to preferentially bind to constituents associated with the microtubule domains rather than to the neurofilament domains. Although it seems likely that some proteins will have a preferential tendency to bind to the neurofilaments rather than to the components of the microtubule domains, these proteins are far less abundant in the axoplasm than are those that are organized by the microtubule domain. The neurofilament domains also contain proteins that are in the aqueous phase of the axoplasm. These “free” proteins should be relatively uniformly distributed throughout the aqueous phase of the axoplasm, which permeates the microtubule and neurofilament domains (Lasek and Morris, 1982). Many of the proteins that are present in the aqueous phase also bind to components of the cytoplasmic matrix and therefore exist in an equilibrium between bound form and diffusable form. If these proteins

44

RAYMOND J. LASEK

45

THE STRUCTURE OF AXOPLASM

bind preferentially to particular components of the cytoplasmic matrix, these proteins will be present at higher concentrations in regions of the cytoplasm that contain nondiffusable binding sites. The microtubule domains apparently contain more binding sites for these proteins than do the neurofilament domains. These observations suggest that the basic structure of the cytoskeleton and the cytoplasmic matrix in the squid giant axon is similar to that of other axons such as myelinated vertebrate axons. The squid giant axon has already proven to be a useful model for studying axonal cytoskeletal proteins (Lasek et al., 1979). D. Microfilaments in Axoplasm

Studies of the protein composition of the squid giant axon suggest that the image of the cytoplasmic matrix provided by electron microscopy underestimates the important contribution of microfilaments to the structure of the axonal cytoskeleton. Table I lists the amount of the three TABLE I CYTOSKELETAL PROTEINS IN AXOPLASM FROM SQUIDGIANTAX ON^

Protein Tubulin Neurofilament protein Actin

THE

Fraction (70) of total axoplasmic protein

Concentration (mgW

22

5.6

13

3.3

6

I .4

From Morris and Lasek (1984).

FIG. 1. Transverse section of a Loligo giant axon fixed by immersion following removal of the connective sheath surrounding the axon by the method of Metuzals et al. (1981b). (A) This low-magnification image of the giant axon illustrates the segregation of the axoplasm into domains. The domains are distinguished by their position and their electron density. The subaxolemmal domain lies adjacent to the plasma membrane (a). The central axoplasm is subdivided into electron-lucent areas and electron-dense areas. The electron-lucent areas correspond to the neurofilament domains, and the electron-dense domains correspond to the microtubule domains. Note that the electron-dense microtubule domains are coextensive with the subaxolemmal domains. (B) is a higher magnification image of this region and illustrates the presence of large numbers of microtubules in the dense regions. Bar = 0.25 p m . This figure is similar to that in Metuzals et ul. (1983), which should be consulted for further details.

46

RAYMOND J. LASEK

principal cytoskeletal proteins of axoplasm from the squid giant axon and demonstrates that these proteins constitute roughly one-half of the proteins in the axon. Tubulin and the neurofilament proteins are the most abundant proteins in the axoplasm, which is consistent with the electron microscopic observation that microtubules and neurofilaments are the predominant components of the cytoskeleton (Lasek et al., 1979; Morris and Lasek, 1982). However, if we compare the amount of actin with that of tubulin or the neurofilament proteins, there is about three times as much tubulin as actin and about twice as much neurofilament protein as actin (Table I). A more useful comparison between these constituents is the amount of each type of polymer present in the axon. These values can be calculated from information that we have obtained on the proportion of each of these proteins that is polymerized in the axon. Table I1 lists the proportions of actin and microtubule and neurofilament proteins that are either in the form of monomer in the axoplasm as compared to polymer. Using these data and information about the basic structure of these polymers, we can compare the relative abundance of each type of polymer in the axon. That is, because we know approximately how many monomeric proteins are required to form a particular length of polymer, we can estimate the proportional length of the microfilaments, microtubules, and neurofilaments in the giant axon. The estimated proportion of the total length of neurofilaments : microfilaments : microtubules is, respectively, 27 : 3 : 1. (The estimated value for the neurofilaments was based on the assumption that neurofilament proteins have an average molecular weight of 100,000 and that there are 6 subunits in a cross-section of the neurofilaments. The actual structure of neurofilaments is not known and this estimate could be off by a factor of two or more.) These estimates of the proportion of cytoskeletal polymers in the axon are consistent with the observation that neurofilaments are the most abunTABLE I1 THEPROPORTION OF MONOMERIC AND POLYMERIC CYTOSKELETAL PROTEINSIN AXOPLASM~ Protein

Tubulin Neurofilament protein Actin

Monomer (%) 25 <5

45

From Moms and Lasek (1984).

Polymer (%) 75 >95

55

THE STRUCTURE OF AXOPLASM

47

dant structure in the axon. However, it is surprising that the microfilaments are three times more abundant than microtubules. Microtubules are easily seen in electron micrographs of the axon, but microfilaments have been very difficult to detect (LeBeaux and Willemot, 1975; Metuzals and Tasaki, 1978). This may be explained by the recent observation that microfilaments isolated from axoplasm of the giant axon are relatively short (Fath and Lasek, 1983). Very few microfilaments are more than 1 or 2 pm in length and most appear to be 0.5 pm or less in length. This fact, coupled with the small diameter of the microfilament (which measures 4-6 nm), may explain why microfilaments have been difficult to detect within the axon. Clearly, the role of the microfilaments in axonal structure and physiology must be reevaluated. In this regard, it should also be noted that studies of fast axonal transport clearly demonstrate that microfilaments are important in the translocation of particles in the axon (Brady, 1984). It remains to be determined whether the microfilaments participate directly in fast axonal transport (possibly as a substrate for myosin) or whether they play an important structural role like the microtubules. 111.

AXONAL TRANSPORT IN THE SQUID GIANT AXON

Comparisons of the physiological properties of the squid giant axon with other axons also indicate that it can be used as a model for understanding the physiology of axoplasm. One of the most important physiological properties of the axon concerns the mechanisms that transport materials between the cell body and the axon terminals. Axonal transport provides both a lifeline from the cell body to the axon for the metabolic support of the axon and as an information channel for chemical signals between the cell body and the periphery (Grafstein and Forman, 1980). Axonal transport can be divided into two basic components: (1) fast axonal transport, which is responsible for conveying membranous elements and their contents, and (2) slow axonal transport, which conveys the elements of the cytoplasmic matrix, including the cytoskeletal structures (Grafstein and Forman, 1980; Brady and Lasek, 1982b). No information exists regarding slow axonal transport in the giant axon. By contrast, the squid giant axon has become one of the most useful preparations for studying fast axonal transport (Allen et al., 1983; Brady et al., 1982). Fast axonal transport has been studied in the squid giant axon by directly visualizing the membranous particles in the giant axon. These studies have benefited greatly from the development of video-enhanced microscopy (Allen et al., 1982). In fact, studies on the squid giant axon with video-enhanced differential interference contrast microscopy have pro-

48

RAYMOND J. LASEK

vided detailed information about the movement of all the basic types of membranous particle transport. Three classes of particles have been recognized in the squid giant axon: ( 1 ) small particles (40-60 nm in diameter), which move principally in the orthograde direction, (2) medium-sized particles (100-150 nm in diameter), which move primarily in the retrograde direction, and (3) large particles (mitochondria), which move bidirectionally with a bias in the orthograde direction. Detailed analyses of the movements of these particles indicate that fast axonal transport is a highly conserved process, so that information obtained from the squid giant axon will apply to other axons. In fact, it is likely that fast axonal transport is a special example of intracellular particle transport more generally (Brady and Lasek, 1982b). Studies of fast axonal transport have been extended to axoplasm that is extruded from the squid giant axon (Brady P t al., 1983). Remarkably, fast transport continues for hours in the axoplasm after it is removed from the giant axon. The detailed properties of the rapid particle movements in isolated axoplasm are indistinguishable from those in the intact axon (Allen et al., 1983; Brady et al., 1983). These observations indicate that the plasma membrane is not required for normal translocation of membranous organelles by fast transport in the axon. Furthermore, they indicate that extrusion of the axoplasm does not disrupt the rather complex physiological processes that are known to be dependent upon the integrity of the cytoskeleton in the axon. That is, fast axonal transport is sensitive to disruption of microtubules or microfilaments and becomes very inefficient when the cytoskeleton is disordered and will fail if the cytoskeleton is substantially denatured (Grafstein and Forman, 1980). Thus, these observations on fast axonal transport in the giant axon demonstrate that the giant axon is a useful model for studying physiological processes and that the information gained from the giant axon will provide insight into the mechanisms that occur universally in axons. Furthermore, the discovery that fast transport continues in isolated axoplasm provides a permeable model for analysis of the mechanisms of fast axonal transport (Brady, 1984). Because the plasma membrane is separated from the axoplasm during extrusion, chemical probes can be introduced into the axoplasm by placing the axoplasm into a solution containing the particular probe (Brady et al., 1983). It is also possible to permeabilize the plasma membrane by other methods, such as the addition of a detergent, that produce holes in the lipid bilayer (Forman, 1982). However, this method suffers from the problem that detergents affect the properties of cellular components other than lipids. Thus, the isolated axoplasm from the squid giant axon can be considered a standard against which experiments on preparations per-

THE STRUCTURE OF AXOPLASM

49

meabilized by other methods can be judged (Brady, 1984). In fact, the results of various experiments on axons permeabilized by extrusion and from those permeabilized by other methods are in relatively good agreement (Forman, 1982; Adams, 1982). IV. EXTRUDED AXOPLASM FROM THE GIANT AXON

One of the most important advantages of the giant axon is that axoplasm can be removed from the axon for analysis without contamination from surrounding glial cells. The axoplasm can then be used for biochemical or physiological analyses (Roberts et al., 1958; Baker and Shaw, 1965; Baker et al., 1971; Lasek et al., 1973; Gilbert, 1975; Black and Lasek, 1977; Lasek et al., 1979; Pant and Gainer, 1980; Rubinson and Baker, 1979; Baumgold et al., 1981; Lasek, 1983). Recently, we have carefully evaluated the process of mechanical extrusion of axoplasm using video-enhanced microscopy (Brady and Lasek, unpublished results). A giant axon was placed under the light microscope (video-enhanced contrast) and then subjected to the process of extrusion, which involves cutting the axon at one end and placing pressure at the other end. Because the axoplasm is a gel, it yields only after sufficient pressure is applied to shear the axoplasmic network. By slowly increasing the pressure it was possible to reach a point at which extrusion of the axoplasm could be controlled. At pressures in excess of that required to extrude the axoplasm it was possible to observe the axoplasm moving through the axon. Our observations indicate, in agreement with those of Baker et al. (1962), that the axoplasm shears at a relatively sharp boundary that is located 10 ,urn beneath the plasma membrane. Although the axoplasm was disrupted at the boundaries of the shear plane, the disrupted region involved was relatively small. Particle movement continued in the remainder of the axoplasmic cylinder during the extrusion process, indicating that most of the axoplasm was unaffected by the extrusion process. In fact, if the pressure on the axon was released, so that the extrusion process was halted, the axoplasm in the region of the shear plane recovered and it was not possible to distinguish the sheared region from the unaffected axoplasm. These observations indicate that the extrusion process only effects a relatively limited region of the axoplasm located at the shear plane. Furthermore, extrusion produces a relatively uniform cylinder (Morris and Lasek, 1982) and leaves a IO-pm annulus of cortical axoplasm behind with the sheath. The clean separation between the cortical axoplasm and central axoplasm provides an opportunity for comparing the concentration of axo-

50

RAYMOND J. LASEK

plasmic proteins in the cortex and central axoplasm. We have carried out such a comparison for fodrin (brain spectrin) because our studies have indicated that significant amounts of fodrin are located in the extruded axoplasm (Fath and Lasek, unpublished results). Fodrin is generally considered to be principally localized subjacent to the plasma membrane (Levine and Willard, 1981). Quantitative analyses of the amount of fodrin in extruded axoplasm and the remaining sheath of the giant fiber indicate that more than 80% of the fodrin in the axon is located in the extruded axoplasm. The remainder of the fodrin is located in the sheath, which contains approximately 8% of the axoplasm. Although fodrin is more concentrated in the cortical region of the axoplasm, most of the fodrin is not associated with the cortex. These results differ from the observations made on other axons using immunofluorescence, in which the fodrin appeared to be localized exclusively beneath the plasma membrane. However, immunofluorescence is not quantitative and differences in concentration as little as threefold can produce sharp contrast in a photomicrograph. These observations on the squid giant axon suggest that the distribution of fodrin in neurons and probably in other cells must be reevaluated. V.

CONCLUSIONS

The squid giant axon provides a powerful model for understanding the structure and functions of axons generally. The most significant difference between the giant axon and other axons is its large size. However, the giant axon is really a collection of smaller axons. Each giant axon is a syncytium that can be traced back to a collection of cell bodies in the stellate ganglion. Apparently, the giant axon is formed from a fascicle of ordinary-sized axons that course together in the stellate nerves of the developing squid. These axons fuse with each other laterally to produce one common axoplasmic mass that is surrounded by a single plasma membrane. The process of fusion is documented in the case of the firstorder giant fibers (Martin, 1965, 1969). Only the axons fuse; the cell bodies retain their separate identities and each one contributes to the maintenance of the giant axon. Thus, the giant axon is the product of ordinary-sized molluscan neurons, each contributing axoplasm to a single giant axon. The development of the giant axon from ordinary-sized neurons working cooperatively explains how it is that the giant axon is so similar in its basic structure and physiology to other axons. Apparently, the evolution of giant axons in the squid did not involve any significant modification of

THE STRUCTURE OF AXOPLASM

51

basic neuronal processes. That is, except for the steps that are involved in the fusion of the embryonic precursors to the giant axon, the giant axon apparently retains all of the properties that characterize axons generally. Therefore, information obtained about the structure and function of the squid giant axon should be of great value in understanding the biology of axons whether they are small or large.

REFERENCES Adams, R. J. (1982). Organelle movement in axons depends on ATP. Nature (London)297, 327-329. Allen, R. D., Allen, N. S., and Travis, J . L. (1982). Video-enhanced contrast, differential contrast (AVEC-DIC): A new method capable of analyzing microtubule related motility in the reticulopodial network of Allogromiu luticolluris. Cell Motil. 1, 291-302. Allen, R. D., Metuzals, J., Tasaki, I., Brady, S. T., and Gilbert, S. (1983). Fast axonal transport in squid giant axon. Science 218, 1127-1129. Baker, P. F., and Shaw, T. I. (1965). A comparison of the phosphorous metabolism of intact squid nerve with that of the isolated axoplasm and sheath. J. Physiol. (London) 180, 424-438. Baker, P. F., Hodgkin, A. L., and Shaw, T. I. (1962). Replacement of the axoplasm ofgiant nerve fibers with artificial solutions. J . Physiol. (London) 164, 330-354. Baker, P. F., Hodgkin, A. L., and Ridgway, E. B. (1971). Depolarization and calcium entry in squid giant axons. J. Physiol. (London) 218, 709-755. Baumgold, J., Terakawa, S., Iwasa, K., and Gainer, H. (1981). Membrane associated cytoskeletal proteins in squid giant axons. J . Neurochem. 36, 759-764. Bear, R. S., Schmitt, R. O., and Young, J. Z. (1937). Investigations on the protein constituents of nerve axoplasm. Proc. R. Soc. London Ser. B 123,520-529. Black, M.M., and Lasek, R. J. (1977). The presence of transfer RNA in the axoplasm of the squid giant axon. J. Neurobiol. 8, 229-237. Brady, S. T. (1984). Basic properties of fast axonal transport and the role of fast axonal transport in axonal growth. Adu. Neurochem. (in press). Brady, S. T., and Lasek, R. J. (1981). Nerve-specific enolase and creatine phosphokinase in axonal transport: Soluble proteins and the axoplasmic matrix. Cell 23, 515-523. Brady, S. T., and Lasek, R. J . (1982a). The slow components of axonal transport: Movements, composition and organization. In “Axoplasmic Transport,” (D. G. Weiss, ed.), pp. 207-217. Springer-Verlag. Berlin and New York. Brady, S. T., and Lasek, R. J. (1982b). Axonal transport: A cell biological method for studying proteins that associate with the cytoskeleton. Methods CellBiol. 25,365-398. Brady, S . T., Tytell, M. M., Heriot, K., and Lasek, R. J . (1981). Axonal transport of calmodulin: A physiologic approach to identification of long term associations between proteins. J . Cell Biol. 89, 607-614. Brady, S. T., Lasek, R. J., and Allen, R. D. (1983). Fast axonal transport in extruded axoplasm from squid giant axon. Science 218, 1129-1 131. Ellisman, M. H., and Porter, K. R. (1980). Microtrabecular structure of the axoplasmic matrix: Visualization of crosslinking structures and their distribution. J . Cell Eiol. 87, 464-479. Fath, K . , and Lasek, R. J. (1983). Actin microfilaments are a major cytoskeletal component in squid axoplasm. E i o l . Bull. (Abstr.) 165, 489-490.

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RAYMOND J. LASEK

Forman, D. S. (1982). Vanadate inhibits saltatory organelle movement in a permeabilized cell model. Exp. Cell Res. 141, 139-147. Friede, R. L . , and Samorajski, T. (1970). Axon caliber related to neurofilaments and microtubules in sciatic nerve fibers of rat and mice. Anut. Rec. 167, 379-388. Garner, J., and Lasek, R. J. (1982). Cohesive axonal transport of the slow component b complex of polypeptides. J . Neurosci. 2, 1824-1835. Geisler, N., Kaufman, E., Fischer, S., Plessman, U., and Weber, K. (1983). Neurofilament architecture combines structural principles of intermediate filaments with carboxyterminal extensions increasing in size between triplet proteins. EMBO J . 2, 12951302. Gilbert, D. G. (1975). Axoplasm chemical composition in Myxicolu and solubility properties of its structural proteins. J . Physiol. (London) 253, 303-319. Grafstein. B . , and Forman, D. S . (1980).Intracellular transport in neurons. Phy.viol. Rev. 60, 1167-1282. Hirokawa, N. (1982). Cross-linker system between neurofilaments, microtubules, and membranous organelles in frog axons revealed by the quick freeze deep-etching method. J . Cell R i d . 94, 129-142. Hodge, A. J., and Adelman, W . J., Jr. (1980). The neuroplasmic network in Loligo arid Hermissenda neurons. J . Ultrastruct. Res. 70, 220-241. Kirschner, M. W. (1978). Microtubule assembly and nucleation. In?. Rev. Cyrol. 54, 1-69. Krishnan, N., Kaiserman-Abramof, I. R., and Lasek, R. J. (1979). Helical substructure of neurofilaments isolated from Myi-icolu and squid giant axons. J . CellBiol. 82,323-335. Lasek, R. J. (1983). Cytoskeletons reconstituted in vitro indicate that neurofilaments contribute to the helical structure of axons. In “Developing and Regenerating Nervous Systems” (P. W. Coates, R. R. Markwald, and A . I). Kenny, eds.). Alan R. Liss, New York. Lasek, R . J . , and Morris, J . R. (1982). The microtubule-neurofilament network: The balance between plasticity and stability in the nervous system. In “Biological Functions of Microtubules and Related Structures” (H. Sakai, H. Mohri, and G. G . Borisy, eds.), pp. 329-342. Academic Press, New York. Lasek, R. J., Dabrowski, C., and Nordlander, R. (1973). Analysis of axoplasmic RNA from invertebrate giant axons. Nature (London) New Biol. 244, 162-165. Lasek, R. J., Krishnan, N., and Kaiserman-Abramof, I. R. (1979). Identification of the subunit proteins of 10-nm neurofilaments isolated from axoplasm of squid and Myxicolu giant axons. J . Cell Bid. 82, 336-346. Lebeaux, Y . J., and Willemot, J. (1975). An ultrastructural study of the microfilaments in rat brain by heavy meromyosin labelling: The peridaryon, dendrites, and the axon. Cell Tiss. Res. 160, 1-36. Levine. J . . and Willard, M. (1981). Fodrin: Axonally transported polypeptides with the internal periphery of many cells. J . Cell B i d . 90, 631-643. Martin, R. (1965). On the structure and embryonic development of the giant fiber system of the squid Loligo uulgaris. Z. Zellforsch. Mikrosk. Anut. 67, 77-85. Martin, R. (1969). The structural organization of the intracerebral giant fiber system of cephalopods. The chisma of the first order giant axons. Z. Zellforsch. Mikrosk. Anut. 97, 50-68. Metuzals, J. (1969). Configuration of a filamentous network in the axoplasm of the squid ( L o l i g o pcwlei L.) giant nerve fiber. J . Cell B i d . 43, 480-505. Metuzals, J., and Tasaki, I. (1978). Subaxolemmal filamentous network in the giant nerve fiber of the squid (Loligo petrlc4 and its possible role in excitability. J . C‘rll B i d . 78, 597-62 1.

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53

Metuzals, J., Montepetit, V . , and Clapin, D. F. (1981a). Organization of the neurofilamentous network. Cell Tiss. Res. 214, 455-482. Metuzals, J., Tasaki, I., Terakawa, S., and Clapin, D. F. (1981b). Removal of the Schwann sheath from the giant nerve fiber of the squid: An electron microscopic study of the axolemma and associated axoplasmic structures. Cell Tiss. Res. 221, 1-15. Metuzals, J., Clapin, D. F., and Chapman, G. D. (1982). Axial and radial filamentous components of the neurofilamentous network. Cell Tiss. Res. 223, 507-518. Metuzak, J . , Clapin. D. F.. and Tasaki, 1. (1983a). The axolemma-ectoplasm complex of the squid giant axon. I n “Structure and Function in Exciiable Cells.” Plenum, New York. Metuzals, J.. Hodge. A., Lasek. R. J . , and Kaiserman-Abramof, I. R . (1983b). Neurofilamentous network and filamentous matrix preserved and isolated by different techniques from squid giant axon. Cc~llTiss. Res. 228, 415-432. Morris, J . R., and Lasek, R. J. (1982). Stable polymers of the axonal cytoskeleton: The axoplasmic ghost. J . Cell Biol. 92, 192-198. Morris, J. R., and Lasek, R. J. (1984). Monomer-polymer equilibria in the axon: Direct measurement of tubulin and actin as polymer and monomer in axoplasm. J . Cell Biol. (in press). Morris, J. R., Hodge, A. J., and Lasek, R. J . (1981). The microtubule network in the squid giant axon. Biol. Bull. 161, 308. Pant, H. C., and Gainer, H. (1980). Properties of a calcium-activated protease in squid axoplasm which selectively degrades neurofilament proteins. J . Neurobiol. 11, 1-12. Peters, A., Palay, S. L., and dewebster, H . L. (1976). “The Fine Structure of the Nervous System: The Neurons and Supporting Cells.” Saunders, Philadelphia. Roberts, N. R., Coelho, R. R., Lowry, 0. H., and Crawford, E. J. (1958). Enzyme activities of giant squid axoplasm and axon sheath. J . Neurochem. 3, 109-115. Rubinson, K . A , , Baker, P. F. (1979). The flow properties of axoplasm in a defined chemical environment: Influence of amines and calcium. Proc,. R. S o c . London S e r . B 205, 325-345. Schnapp, 8. J., and Reese, T. S. (1982). Cytoplasmic structure in rapid-frozen axons. J . Cell Biol. 94, 667-679. Sharp, G. A., Shaw, G., and Weber, K. (1982). lmmunoelectron microscopical localization of the three neurofilament triplet proteins along neurofilaments of cultured dorsal root ganglion neurons. Exp. Cell Res. 137, 403-413. Willard, M., and Simon, C. (1981). Antibody decoration of neurofilaments. J . Cell Biol. 89, 198-205. Wuerker, R. B., and Kirkpatrick, J . B. (1972). Neuronal microtubules, neurofilaments and microfilaments. I n t . Reu. Cytol. 33, 45-75.

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Part II

Regulation of the Axoplasmic Environment

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CUKRENI' . r o P i c s I N MEMBRANES A N D TKANSPOKT. VOLUME 22

Biochemistry and Metabolism of the Squid Giant Axon HAROLD GAINER,*.Il P A U L E . GALdLANT,'-."ll ROBERT GOULD,S.iiA N D HARISH C . PANT$.Il * Lriboratory of Ncrtror~hrrni.~tr?. tind N~~rrroitrrr~~rtnology Naiional Institute of Child Heultli and Human Deuelopmeni National Institutes of Health Beihesda, Maryland 'Laboratory of Neurobiology National Institute of Mental Heulth Alcohol, Drug Abuse, und Mental Health Adminisrration Bettiesda, Maryland flnstitrr~r~ ,for. Btrsic, Reserrrch in ~ i ~ v i ~ l o p t r i i ~Di.suhilities ntiil SrtrrrJn I.slond. New York ~Lubortrtoryof Predinic~trlSiiidie.s Ncttiontrl Institute o n Alr.ohol Ahrtsc~rind Alcoholism Alcohol Drug Abuse, and Mcntul Hatrlth Adtniwi.ctrtrrion Roc.kuillr~,Mrrrylnnd IIMrrrine Biologicwl Lahortrforv wood.^ H o l e , Mus.scrc,llrrsc~tts

1. 11.

Introduction ............................................................ Molecular Composition of Axoplasm. . . . . . . . . . . . . . . . . . . . . . . . . . . . . A . Ions and Other Small Mol B. Lipids and Their Metabolism.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

58

..........

66

111. Functional Metabolic Studies Using the Giant Axon System .......... A. Energy Metabolism and A B. Other Enzyme Systems . . . . . . . . . . . . . . .......... C. Protein Transfer from Schwann Cells to t xon . . . . . . . . . . . . . . . . D. Proteins and Axolemmal Properties.. ........................ 1V. Concluding Remarks . . . . . . . . . . . . . . . . ............................... References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

68

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

70 73 81

57 Copyright 0 1984 hy Academic Press. Inc. All rights of reproduction in any form reserved. ISBN 0-12-153322-0

58

HAROLD GAINER ET AL.

1.

INTRODUCTION

The authors of this chapter are a group of neurochemists from different laboratories, who gather together each summer at the Marine Biological Laboratory in Woods Hole with the common purpose of examining the biochemical properties of the giant axon of the squid. Our research interests are diverse and include lipid metabolism, protein biosynthesis and phosphorylation, immunology, proteases, and cytoskeletal proteinplasma membrane interactions. For each of us the squid giant axon represents a unique biological system that allows for the examination of specific and fundamental neurobiological mechanisms. Two features of the giant axon make it especially attractive to neurochemists who are interested in correlating biochemical and physiological processes. Both relate to the size of the axon. The first is the abundant and pure cytoplasm (axoplasm) available by extrusion methods. This allows us to study axoplasmic biochemistry, metabolism, and structure under tightly controlled experimental conditions. The second is that the large diameter of the axon permits intracellular perfusion. This is ideal for studies of membrane biochemistry that are to be correlated with one of the major functions of the axonal membrane, i.e., the production of action potentials. In addition, it is possible in this system to study glial-axonal interactions unequivocally. In short, the giant axon offers many opportunities to neurochemists that are not available from any other experimental preparations. In the following pages, we review some of the biochemical and metabolic information obtained as a result of research on the giant axon. As in the field of electrophysiology, biochemical studies on the giant axon are generally applicable to mammalian systems and have often provided the first insights into axonal properties, in general.

II.

MOLECULAR COMPOSITION OF AXOPLASM

Virtually all our knowledge about the molecular composition of axoplasm is derived from studies on axoplasm isolated from the giant axon of the squid. Approximately 73-88% of the total weight of the squid giant fiber is occupied by axoplasm (Bear and Schmitt, 1939). About 86-89% of the axoplasm (by weight) is water (Hodgkin, 1965; Koechlin, 1955). The intraaxonal (or axoplasmic) pH is reported to range between 7.0 (Caldwell, 1958; Bicher and Okhi, 1972) and 7.3 (Boron and DeWeer, 1976). The specific molecules found in squid axoplasm and some of their metabolic properties are discussed in the following sections.

BIOCHEMICAL PROPERTIES OF THE SQUID GIANT AXON

59

A. Ions and Other Small Molecules

The composition of squid (Loligo) giant fiber axoplasm with respect to the concentrations of major ionic, amino acid, carbohydrate, and highenergy phosphate substances is shown in Table I. The data are provided as ranges when more than one independent measurement was available, and the variation is probably due to differences in the species of Loligo (pealei versus forbesi) used, and differences in experimental conditions and methods of measurement. It should also be noted that the data in Table I represent total concentrations and that in many cases (e.g., for Ca2+, see chapter by Baker and DiPolo in this volume) a substantial fraction of the substance may be in a bound form. Although most of the cationic component of axoplasm osmotic pressure is due to inorganic ions (e.g., K + ) , most of the anionic component in the squid is composed of organic anions (e.g., isethionate, aspartate, and glutamate). The major storage form of high-energy phosphate in squid axoplasm, as in the case for most invertebrates, is arginine phosphate. The ionic composition of axoplasm has great bearing on the issue of which artificial axoplasmic solution to use for physiological and biochemical experimentation. It is well known that millimolar levels of Ca2+ (Hodgkin and Katz, 1949) and certain anions (Rubinson and Baker, 1979) can liquefy squid axoplasm and have deleterious effects on the excitability of perfused axons (Baker et al., 1962; Okawa et al., 1961; Tasaki and Takenaka, 1964; Tasaki, 1968). The effect of Ca2+ is due, in part, to activation of calcium-dependent neutral proteases, which are abundant in squid axoplasm (see Section III,B,3), and the effect of anions is probably related to their chaotropic properties [the order of chaotropic effectiveness is SCN>I>Br>NO3>C1>acetate>SO4>aspartate, glutamate> P04>F (see Tasaki, 1968; Rubinson and Baker, 1979; and Section III,D,2)]. We have found that many biochemical properties of the axons and their extruded axoplasm (e.g., phosphorylation of proteins, lipid metabolism) are modified by Ca2+and C1- in the artificial media. For this reason we show in Table I1 some artificial media described in the literature that we have found to be useful for biochemical studies on the squid giant axon (see Sections I1,B and 111,C). B. Lipids and Their Metabolism

McColl and Rossiter (1951; see also McMurray et al., 1964) were the first to analyze the lipids of the squid giant axon. In their study, they found that lipids extracted from different neural tissues, i.e., the central

60

HAROLD GAINER ET AL.

TABLE I CONCENTRATIONS OF IONSA N D OTHERMOLECULES I N Luligu AXOPLASM, BLOOD,A N D SEAWATER Substance Waterd Proteind Major ions' K+ Na+ C1-

Ca2+ Mg2+ Isethionate

so42+

po43

Major amino acidse Aspartate Glutamate Arginine Alanine Glycine Taurine Hornarine Betaine Major carbohydrates' Glycerol Glucose Mannose Fructose Succinate and fumarate High-energy phosphates' ATP Arginine phosphate

Axoplasm"

Bloodh

865 20

870 0. IS

Seawater' 966

-

323-400 44-65 40-151 0.4-7 6.4-20 164-250 7.5 2.S-17.8

20-22 440-456 560-578 10-1 1 54-55 1.6 8.1

8.3-10 423-460 506-580 9.3-10 48-53 28.2

33-100 6.2-8.4 0.36-2.2 1.7-16 4.5-18.4 81-106.7 20.4 73.7

-

-

-

-

-

-

4.35 0.24 0.92 0.24 17

0.7-1.7 1.8-5.7

-

-

-

3.5 3.4 4.4

-

-

-

-

-

-

-

-

-

-

-

-

-

-

Data from Deffner, 1961; Deffner and Hafter, 1960; Baker and Crawford, 1972; Koechlin, 1955; Hodgkin, 1965; Keynes, 1963; Steinbach, 1941 ; Steinbach and Spiegelman, 1943; Rosenberg and Khairallah, 1974. Data from Hodgkin, 1965; Deffner, 1961; Potts and Parry, 1964. Data from Hodgkin, 1965; Potts and Parry, 1964; Cavanaugh, 1975. Data are expressed in grams per kilogram. Data are expressed in millimoles per kilogram.

BIOCHEMICAL PROPERTIES OF THE SQUID GIANT AXON

ARTIFICIAL MEDIAUSEFUL

TABLE 11 BIOCHEMICAL STUDIESON EXTRUDED AXOPLASM

FOR

THE

61

SQUIDAXONA N D

Artificial axoplasm" Substance Na+ K' CaZ+ Mg2+

430 10

c1-

Tris PH Methane sulfonate Glutamate MOPS buffer EGTA Taurine ~

0

~ 3

External media," Gilbert (1974)

-

Isethionate Alanine Glycine Aspartate Arginine Betaine Glycerol Glucose Fructose ATP GTP PMSF

10 50 500 10 7.4 -

Rubinson and Baker ( 1979) 400 3 12.9 15.9 7.2 300b 100 206 10

150

Morris and Lasek (1982) 65 344 10 147.2 -

7.0 -

1.0 106.7 17.8 164.6 16.1 18.4 100.3 6.1 73.6 4.4 1.2 0.5 I .o 0.5

0. I

Data are expressed in millimoles per liter. We have substituted methyl sulfate for methane sulfonate, and imidazole for MOPS buffer, with no change in effectiveness of this medium for biochemical studies. a

ganglia, the pallial (or mantle connective nerve), the stellar nerve (or giant axon), and axoplasm extruded from the giant axon were similar in composition. Phospholipids were the principal components of all preparations, accounting for between 0.18 mg/100 mg of fresh tissue (extruded axoplasm) and 3.21 mg/100 mg of fresh tissue (central ganglia). The data are shown in Table 111. Levels of lipid in the giant axon and axoplasm are lower than in the pallial nerve and central ganglion, presumably because the axoplasm of the giant axon is composed predominantly of cytosolic and fibrous pro-

62

HAROLD GAINER ET AL.

LIPIDCOMPOSITION OF

SQUID

TABLE I11 AXOPLASM AND

OTHER

NEURALTISSUES"

Tissue

Central ganglion

Pallial nerve

Giant axon

Extruded axoplasm

Phospholipid (PL) Cholesterol (Chol) PL/Chol PC'IP SPH<'/PL

3.21 + 0.09 (4) 0.73 + 0.06* 4.4 0.42 0.14

0.98 (2) 0.3Ib 3.2 0.43 0.29

0.33 (2) 0.08 4. I 0.22

0.18 (2) 0.03 6.0 0.15

" Values are expressed as milligrams per 100 mg of fresh tissue. Numbers in parentheses represent the number of samples analyzed. Greater than 95% of the cholesterol is in the nonesterified form. Lecithin (PC) was determined by measuring the alkali-labile choline recovered as the reincke salt. Sphingomyelin (SPH) was measured as the difference between total and alkali-labile phosphorus. Data are from McColl and Rossiter (1951).

teins and fewer membranous organelles. McMurray et ul. (1964) noted the lack of cerebroside and the fact that cholesterol and sphingomyelin were lower than levels in the vertebrate nervous system. This is most likely due to the abundance of these lipids in the vertebrate myelin sheath. The phospholipid composition of the squid giant axon and extruded axoplasm was reexamined by Condrea and Rosenberg (1968), and also membrane fractions separated from the squid giant axon was examined by Camejo et al. (1969). Both studies used the chloroform : methanol cxtraction procedure developed by Folch et al. (1957). This method is still widely used because of its effectiveness in extracting phospholipids, neutral lipids, and glycolipids. The only lipids not extracted into the chloroform-rich phase are gangliosides [presumably absent from marine invertebrates (Vaskovsky et ul., 1970)] and the acidic polyphosphoinositides (Dawson and Eichberg, 1965), which are present in minor amounts in peripheral nervous tissues relative to PC,' PE, PI, etc. (Sheltawy and Dawson, 1966). Condrea and Rosenberg (1968) separated the extracted lipids on two-dimensional thin-layer chromatograms. The amounts of separated phospholipids were quantified by phosphorus estimation. The intact giant axon contained all the major phospholipids found in vertebrate nerves and other membrancs, i.e., PE, 31%; PC, 45%; PS, 8%; PI, 5%, and SPH, 11%. Extruded axoplasm contained much lower levels of lipid, but had I Abbreviations used: PA, phosphatidic acid; PC, phosphatidyl choline; PE, phosphatidyl ethanolamine; PI, phosphatidyl inositol; SPH, sphingomyelin.

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similar proportions of the major lipid species with the exception of SPH, which was not detected in axoplasm (however, its formation in axoplasm has been demonstrated using radioactive precursors, see below). As pointed out by Condrea and Rosenberg (1968), the corresponding plasmalogens and dialkyl ethers would not have been separated from the diacyl esters by these methods. Furthermore, the investigators did not look for phosphonolipids, which we have detected in lipid extracts of squid tissues (unpublished observations). As in the McColl and Rossiter study, it was found that the amounts of lipid present in axoplasm were nearly an order of magnitude less than those present in the sheath, i.e., 0.18 vs 1.03 (McColl and Rossiter, 1951), and 0.09 vs 0.5-2.0 (Condrea and Rosenberg, 1968). In the study of Camejo et al. (1969) the phospholipids present in two membrane fractions isolated from the giant axon of Dosidicus gigas were determined. The ratios of polar lipids (primarily phospholipids) to cholesterol were lower than were reported by McColl and Rossiter, i.e., 2. I and 2.6 for membrane fractions I and 11, compared to values of 3.2 to 6.0 (see Table HI). The amounts of the different phospholipids were similar to the values reported by Condrea and Rosenberg (1968) for whole axon, i.e., PC (44-46%), PE (32-35%), PS (10-15%), and SPH (9-10%). Steele et al. (1981) determined cholesterol-to-phospholipid molar ratios of 0.61 for giant axons from L . pealei, which on a weight basis would be around 3.2; this is close to the value reported by Zambrano et ul. (1971) for the membranes of squid retinal axons. Larrabee and Brinley (1 968) studied the incorporation of 32P-labeled inorganic phosphate (Pi) into lipids by intact giant axons and axoplasm separated from the giant axon by extrusion. They obtained the radioactive lipids by the Folch extraction procedure and separated the individual lipids by chromatography on paper impregnated with silicic acid. They found that PI, PE, and PA were formed by the giant axon and the ganglia when they were incubated in seawater containing 32P-labeledPi. Furthermore, labeled lipid was recovered both in extruded axoplasm and sheath (which were separated after the incubation), In addition, they demonstrated that 32P-labeledPi injected into the axon was incorporated into PI and PR, although the labeling of PE was equivocal. Finally, Larrabee and Brinley showed that extruded axoplasm incubated in seawater with 32Plabeled Pi synthesized phospholipids (mainly PI and PR) in a time-dependent manner, They were unable to detect the formation of PC, PS, or SPH from 32P-labeledPi precursor. Brunetti et al. (1979) reinvestigated the synthesis of PC by the squid giant axon using tritiated choline as a precursor. They found that phosphatidylcholine was synthesized in a time-dependent manner by both the squid giant axon and the stellate ganglia, the latter being roughly three to

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six times as active on a milligram of protein basis. The water-soluble components were chromatographically separated, and it was found that phosphatidylcholine and possibly trace amounts of CDP-choline were also formed. They took these results to indicate that synthesis of phosphatidylcholine was occurring by the de nouo pathway. Finally, these investigators showed that homogenates of the giant axon and stellate ganglion (fiber lobe) both catalyzed the incorporation of tritiated choline and ethanolamine into phospholipid under conditions designed to measure the Ca2+-dependentbase-exchange reaction. They did not, however, use extruded axoplasm to determine whether the incorporation of choline into PC was indigenous to the axonal cytoplasm. We have expanded these studies by monitoring the synthesis of lipids by intact giant axons and extruded axoplasm using a host of potential lipid precursors (Gould et ul., 1982a, 1983a,b). We found that the precursors choline, ethanolamine, serine, myo-inositol, phosphate, and glycerol were incorporated into phospholipids by extruded axoplasm. The radioactivity in individual lipid products was determined after their separation on thin-layer chromatograms. All of the major phospholipids were made, including phosphatidylcholine from choline, phosphatidylethanolamine from ethanolamine, phosphatidylserine and sphingomyelin from serine, phosphatidylinositol from myo-inositol, phosphatidylinositol phosphate from 3zP-labeledPi and y-labeled ATP. [2-3H]Glycerol was poorly incorporated into lipid. The major product comigrated with phosphatidylinositol, phosphatidic acid, and triacylglyceride. More recently, we examined the incorporation of galactose, sulfate, and glucosamine into lipid products (Gould rt ul,, 1983a,b; Gould, unpublished observations). We were unable to detect any incorporation with galactose or sulfate; i.e., less than 0.01% of the precursor was incorporated into lipid during a 90min incubation at room temperature. In contrast, glycerol, glucosamine, and choline were incorporated at 0.02-0.05%, whereas serine, myo-inositol, and phosphate were incorporated at 0.10-1 .O% of precursor during the 90-min incubations. The lack of incorporation of galactose and sulfate into cerebrosides and sulfatides agrees with the failure of McColI and Rossiter (1951), who were unable to detect these substances in the squid nervous system. The rate of incorporation of precursor into lipid was affected by the composition of the medium in which the axoplasm was suspended. Larrabee and Brinley used seawater, whereas we used media with a composition similar to that of axoplasm (Rubinson and Baker, 1979; Morris and Lasek, 1982; see Table 11). In our studies and those of Larrabee and Brinley, it was found that the pattern of labeling of 32P-labeledaxoplasmic lipids were different if the incorporation was carried out with intact axons or extruded axoplasm. It will be of interest to understand whether axo-

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plasmic lipid metabolism is influenced by the surrounding glial cells (and axolemma) that compose the sheath. I t is conceivable that the amounts, and even types, of lipids formed by axoplasm could be greatly altered by the inclusion of various agents and cofactors, e.g., CoA, fatty acids, CTP, ATP, divalent cations. In an effort to clarify the requirements for synthesis of some of the lipids in axoplasm, we have characterized several of the enzymes of lipid metabolism in extruded axoplasm (Gould et ul., 1983b). The following synthetic enzymes were studied: CDPdiglycerol-inositol 3phosphatidyltransferase (EC 2.7.8.1 I ) , ATP: diglyceride phosphotransferase (diglyceride kinase, EC 2.7.2.-), and phosphatidylinositol kinase (EC 2.7.1.67). In addition, we have demonstrated phospholipid-exchange protein activities capable of transferring PI and, to a lesser extent, PC between organelles and liposomes. These proteins may serve a role in the squid giant axon and in other axons of transferring phospholipids from sites of synthesis on internal membranes to the axolemma. The only other study of enzymes involved in lipid metabolism was that of Eyrich el ul. (1976), who demonstrated a phospholipase A2 activity in homogenates of squid (and frog) axons. However, these workers did not report any studies on extruded axoplasm. Several investigators have considered the relationships of lipids in nerve to impulse propagation. Larrabee and Brinley (1968) reported that repetitive electrical stimulation (at 75 or 100 Hz) of axons that had been injected with '?P-labeled phosphate caused an increase in the incorporation of precursor into PI relative to PA. We have reexamined this question (Gould et al., 1982b) and found (using acidified chloroform : methanol extraction) that there was a reproducible increase of label into phosphatidylinositol and both phosphatidylinositol phosphate and phosphatidylinositol bisphosphate compared with unstimulated controls. Furthermore, by extruding the axoplasm after the incubation, we demonstrated that the stimulation-dependent increase was confined to the axoplasm, but not the sheath, which would contain the electrically excitable axoplasmic membrane after extrusion. Dole and Simon (1973) used 32Pas a probe to look at the effect of levorphanol, a morphine-type narcotic compound that blocked the evoked action potential (Simon and Rosenberg, 1970), on the incorporation into phospholipids. They found that this drug caused an increase in the incorporation of the externally applied phosphate into their preparation (giant axon including adhering small fibers). Unlike the results of Larrabee and Brinley, they found that the incorporation of precursor into PA was increased relative to that of other lipids including PI. One difference between these two investigations was the method of application of precursor. Larrabee and Brinley injected it into the axoplasm while Dole and Simon added it to the bathing seawater solution. Brunetti et al. (1979) studied the effect of electrical stimulation

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on phospholipid synthesis using tritiated choline as precursor. They found that the incorporation of label into PC was depressed by the stimulation, whereas the incorporation into the water-soluble phosphorylcholine was enhanced. Steele et al. (1981) took an interesting approach to study the possible involvement of cholesterol in nerve excitation. By incubating the giant axon with liposomes, they were able to increase or deplete cholesterol in the giant axon preparations. By perfusing the giant axons with PC liposomes, they felt that they could specifically deplete cholesterol from the axolemmal membrane. Although they were able to influence membrane cholesterol, they were unable to alter the Na+ or K + currents in clamped axons. Because of the rapid flip-flop of cholesterol across the bilayer, and the possibility that cholesterol can be transferred between axonal and glial membranes as suggested in the study of Rawlins (1973), it might not be possible to deplete (or increase) the levels of cholesterol in the axolemma1 membrane sufficiently to see an effect on ionic currents, even though cholesterol may be important for excitability. Similarly, the studies of Rosenberg and his colleagues using phospholipases to study the importance of phospholipids in nerve excitation (see Rosenberg, 1973, 1981 for reviews) are difficult to interpret because the loss of excitability caused by phospholipase treatment may simply reflect leakiness of the excitable membranes. C. Proteins

The initial recognition that the squid giant axon was an extraordinarily valuable model for the analysis of fibrous proteins in relation to axoplasmic structure can be attributed to F. 0. Schmitt and colleagues (Bear et al., 1937; Schmitt, 1950). Early studies on the microtubule, intermediate filament, and microfilament associated proteins (Maxfield, 1953; Borisy and Taylor, 1967; Davison and Taylor, 1960; Huneeus and Davison, 1970; Gainer and Gainer, 1976) have evolved into more sophisticated studies of the proteins in axoplasm in relation to its cytoskeletal organization (Sakai and Matsumoto, 1980; Roslansky et al., 1980; Hodge and Adelman, 1980; Morris and Lasek, 1982; see also Lasek in this volume). In addition to these structural studies, there have been a variety of experiments on protein metabolism of the giant axon (see Section 111). Several workers have looked at the phosphorylation of proteins in squid axoplasm under a variety of experimental conditions (Pant et al., 1978a,b; 1979a-c), and it was first demonstrated in squid axoplasm that neurofilament proteins are major phosphoproteins in axons (Pant et al., 1978a,b).

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Subsequent work on Myxicola axoplasm and mammalian preparations (Eagles et al., 1981; Shecket and Lasek, 1982a) have demonstrated the generality of neurofilament phosphorylation and have extended these findings. Although axonal protein phosphorylation can be modified by electrical stimulation of the axon (Pant rt al., 1979a,b), the significance of these observations and of the neurofilament protein phosphorylation itself is still unclear. An early report of a specific calcium-binding protein in squid giant axons (Alema et al., 1973) remains undistinguished from the ubiquitous calmodulin that is present in axoplasm (see Lasek, this volume). Discussion of the proteins composing the subaxolemmal filamentous network in the giant axon (Metuzals and Tasaki, 1978; Metuzals et al., 1981) can be found in Section III,C,3. D. Nucleic Acids

Initial studies on ribonucleic acids (RNA) in the giant axons of the Chilean squid (Dosidicus gigas) suggested that there was no measurable RNA in axoplasm (Fischer et al., 1968). Lasek (1970), however, found significant levels of RNA in axoplasm from LoZigo giant axons. Whether this difference was due to a species or methodological difference remains unstudied. What is clear at present is that subsequent studies on Loligo axoplasm have confirmed the presence of RNA (Lasek er al., 1973; Giuditta et al., 1980). Squid axoplasm contains about 0.02% (wtv) of RNA, of which the predominant form is 4 S RNA (Lasek et al., 1973; Giuditta et al., 1980). Although there is a report of a small amount of ribosomal RNA in squid axoplasm (Giuditta et al., 1980), there is no convincing evidence that de nouo protein synthesis occurs in axoplasm (see Section 111,D). Definitive evidence exists to show that the 4 S RNA in axoplasm is transfer RNA (tRNA) and that axoplasm contains multiple species of tRNA (Black and Lasek, 1977; Giuditta et al., 1977; Ingoglia and Giuditta, 1982). The location of 4 S (or transfer) RNA in axons without appropriate levels of rRNA and mRNA (for protein synthesis) has also been indicated in vertebrate preparations (Ingoglia, 1978, 1982; Gunning e t al., 1979; Ingoglia and Zanakis, 1982). These findings have raised the question of what is the function of this tRNA in axoplasm. One possibility is that the tRNA in the presence of ATP and aminacyl-tRNA protein synthetase and transferase enzymes can add amino acids to the amino-terminus position of preformed proteins (for reviews of this posttranslational modification see Soffer, 1973, 1980). Such a mechanism has been postulated for axons (Droz and Koenig, 1971; Lasek et al., 1973; Black and Lasek, 1977; Ingoglia, 1978; Barra er al., 1973, 1980). Some evidence for the presence

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of an aminoacyl tRNA synthetase in squid axoplasm has been reported (Black and Lasek, 1977; Ingoglia and Giuditta, 1982). The tRNA and the synthetase and transferase enzymes are believed to be organized in a high molecular weight complex in situ (Bandyopahyay and Deutscher, 1971; Vadeboncoeur and La Pointe, 1980). No studies on the organization of this posttranslational modification system in squid axoplasm have been reported.

111. FUNCTIONAL METABOLIC STUDIES USING THE GIANT AXON SYSTEM

A. Energy Metabolism and ATPases It is well known that metabolic poisons such as cyanide and dinitropheno1 decrease the survival time of the squid giant axon in uitro (Segal, 1968). This is not surprising since there are several ATP-utilizing systems in the squid giant axon (for reviews see Matsumura and Clark, 1982; see also Beauge, Baker and DiPolo, and Baker and Carruthers in this volume). Adenosine triphosphate, which is maintained at a concentration of about I mM in axoplasm (see Table I), is the major energy source for active transport of ions and small molecules across the axolemma (Hodgkin and Keynes, 1955; Caldwell and Keynes, 1957; Caldwell, 1960; Baker and Shaw, 1965; Mullins and Brinley, 1967). Arginine phosphate, the other high-energy phosphate store in axoplasm, and phosphoenolpyruvate could restore ATP levels in cyanide-poisoned axons (Caldwell et d.,1964). Adenosine triphosphate can also be synthesized in perfused axons (Martin and Shaw, 1970) in the presence of AMP, and glutamate or aspartate (succinate and a-ketoglutarate were inadequate in this regard), indicating a subaxolemmal ATP-synthesizing system. The findings that CN--sensitive ATP production is present in both sheath and axoplasm is consistent with the observation that mitochondria are present both in the deep parts of the axoplasm and in the subaxolemma1 area (Villegas, 1969). Round (0.6 pm in diameter) mitochondria are found only near the axolemma, while long, thin (3 x 0.4 pm) mitochondria are found throughout the axoplasm. The density of the mitochondria is 40/100 pm2 in the periphery and 30/100 pmZdeeper into the axoplasm. The significance of the differences in mitochondrial density and shape are unknown. The mitochondria apparently utilize NADH in the process of oxidative phosphorylation, since CN-, amytal, and lack of O2all lead to a rapid accumulation of this substrate and depletion of NAD in axons, as measured by a fluorometric technique. This activity was largely confined to the mitochondrial fraction of isolated axoplasm (Doane, 1967).

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Various experiments on the carbohydrate metabolism of the squid axon indicate that there is little or no aerobic metabolism of glucose in axoplasm (Hoskin and Rosenberg, 1965; Hoskin, 1966). In contrast, anaerobic glycosis in axoplasm is greater than that of the sheath (Hoskin, 1966). The sheath cells appear to contain some pentose pathway activity and a significant citric acid cycle. This may account for a report of pyruvate conversion to C 0 2by intact axons (Hoskin er al., 1975). The absence of a citric acid pathway or aerobic metabolism in axoplasm is difficult to reconcile with the presence of abundant mitochondria in axoplasm and to the fact that cyanide, dinitrophenol, and oxygen depletion produce decreased ATP levels and NADH accumulation in axoplasm. One possible explanation for this paradox is that glucose may not be the major energy source for ATP generation in axoplasm. It is conceivable that amino acid catabolism is a more significant carbon source for energy production in squid axoplasm. In this regard, it is notable that glutamate and aspartate can support oxidative metabolism and Na+ transport in perfused axons (Martin and Shaw, 1970; Cannessa-Fischer e? al., 1968). An alternative possibility is that the sheath cells may provide an oxidative pathway for the axon in a type of metabolic cooperation. Some evidence for the movement of intact ATP across the squid axolemma has been reported (Pant er al., 1979b). In contrast, in squid muscle the carbohydrates are the primary sources of energy and aerobic glycolysis appears to be obligatory (Hochachka et al.. 1975). Lactate does not form, and there is no lactate dehydrogenase in the muscle. It is not known whether this enzyme exists in the giant axons. Work on glucose uptake in the giant axon (Baker and Carruthers, 1981a,b) shows that the uptake is a passive saturable process. The uptake rate is about 1 pmol cmP2sec-I, a rate more rapid than the rate of glucose metabolism of the axon. About 20% of the axon's rate of energy metabolism at rest is sensitive to ouabain (Baker and Shaw, 1965), and virtually all of the increased glucose uptake during stimulation is blocked by ouabain (Baker and Carruthers, 1981a). Thus, as in the case for mammals, a significant portion of the energy metabolism of the giant axon is devoted to Na+ and K + transport. The Na+,K+-ATPasein the squid axon is located in both axoplasm and sheath (Bonting and Caravaggio, 1962; Canessa, 1965; Cannessa-Fisher et al., 1968). In perfused axons part of the orthophosphate efflux is dependent upon internal Na+ and external K + and can be blocked by ouabain. In intact nerves poisoned by cyanide or ouabain, orthophosphate efflux is decreased by 20%, suggesting that the Na',K+-ATPase may account for a significant hydrolysis of ATP in the giant axon (Baker and Shaw, 1965). Most of the Na+,KS-ATPaseis concentrated in the sheath preparation (which includes glial cells and axolemma); this contains eight times greater ATPase activity than axoplasm on a weight basis (Canessa, 1965).

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The ATPase in axoplasm is relatively insensitive to ouabain, but is very sensitive to EDTA (Shecket and Lasek, 1982b). The principal ATPase activity in axoplasm requires Mg2+or Ca2+,has an optimum [K+] of 60 mM and a pH optimum of 7.5 (Shecket and Lasek, 1982b), and differs from myosin (ATPase) and Na+,K+-ATPase.This enzyme activity is similar in properties to ATPases found in vertebrate nerves (Khan and Ochs, 1974; Hammerschlag and Bobinski, 1981). Electron microscopic localization of ATPases in the giant axon has been studied (Sabatini et al., 1968). An ATP-dependent formation of lead phosphate (released phosphate from ATP was precipitated in the presence of PbN03) was found in axonal slices. These deposits were found on the cytoplasmic side of the axolemma, at Schwann cell membranes, and on mitochondria. Application of the cardiac glycosides, K strophanthodside or G strophanthin, reduced the deposits mostly in the axolemma and Schwann cell membranes, suggesting that these might be the sites of the Na+,K+-ATPases. B. Other Enzyme Systems

1 . DIISOPROPYL PHOSPHOFLUORIDATE (DFP) HYDROLYZING ENZYME While he was a member of the Nachmansohn group at Columbia University, F. C. G. Hoskin (now at the Illinois Institute of Technology) became interested in the high levels of DFPase activity unexpectedly found in squid axoplasm. Although this work was initiated in the context of a study of cholinergic systems in giant axons (see Section III,B,2), Hoskin later suggested that the coexistence of large amounts of isethionate (2-hydroxyethanesulfonate), taurine, and DFP-hydrolyzing activity in axoplasm might be related (Hoskin and Long, 1972; Hoskin and Brande, 1973; Hoskin et al., 1975; Hoskin, 1976). Subsequent work (Hoskin and Kordik, 1977) showed that isethionate does not arise from cysteine via taurine. However, cysteine (and H2S)can transfer its H-Sto a -C-Cacceptor intermediate, which in the presence of a thiosulfate sulfur transferase (rhodanase) or mercaptopyruvate sulfur transferase enzyme could form isethionate. The -C-Cacceptor in axoplasm is not known at present, and experimental measurements of such enzyme activities in axoplasm have not been performed. Although the relationship between isethionate metabolism and DFPhydrolyzing activity in axoplasm remains unclear, the squid DFP-hydrolyzing enzyme has been purified and coupled to agarose beads for the purpose of detoxifying unwanted nerve gas (Hoskin and Roush, 1982). There are two known DFPases. The type found in squid is a monomer of

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26,600molecular weight ( M , ) , relatively stable, inhibited by M Mn2+ and hydrolyzes DFP five times faster than Soman (a powerful inhibitor of cholinesterase, 1,2,2,-trimethylpropylmethylphosphonofluoridate, used as a chemical warfare agent). In contrast, the mammalian DFPase is a dimer (60,000M , monomers) and unstable; M Mn2+stimulates its activity and hydrolyzes Soman five times faster than DFP. Both enzymes act as acid anhydride hydrolases and do not have phosphatase- or ATPase-like activity. F. C. G. Hoskin (personal communication) speculates that the role of the squid DFPase in axoplasm may be connected to phospholipid or phosphonolipid metabolism in the axon.

2. CHOLINERGIC SYSTEMS Although it is now only of historical interest, the theoretical basis for the earliest studies of cholinergic systems in the squid giant axon was the Nachmansohn theory of nerve conduction (Nachmansohn and Neumann, 1975). This theory held that acetylcholine was essential for electrogenesis in the giant axon. These early studies are reviewed by Rosenberg (1973, 1981) and will not be described here. The idea that cholinergic systems may be involved in electrical events in the giant axon system, not on the axon itself but on the surrounding Schwann cells, has been revived by the Chilean group. Various pharmacological experiments have suggested that there may be a nicotine-type acetylcholine receptor on the Schwann cell (Villegas, 1973, 1974, 1975). Cholinesterase has been histochemically localized at Schwann cell-axon junctions (see Rosenberg, 1981), and acetylcholine synthesis has been reported in the giant nerve fiber of the squid (Heumann et al., 1981). 3. CALCIUM-DEPENDENT NEUTRAL PROTEASE Calcium-dependent neutral proteases (also referred to as calcium-activated proteases, calcium-activated neutral proteases, and calpain) are thiol proteases, with pH optima around neutrality, and they are found in the cytosol of a wide variety of tissues and cells (Ishiura, 1981; Murachi et al., 1981a,b). Their activation in situ often leads to the proteolysis of only one or few specific intracellular proteins, with selective physiological consequences depending on cell type. The presence of calcium-dependent neutral proteases (CANP) in nervous tissue was initially reported by Guroff (1964). The first indication, however, that CANP was of neuronal origin and had a specific endogenous protein substrate came from the work of Gilbert and his colleagues on axoplasm isolated from the giant axons of the fan worm Myxicdm and the squid Loligo (Gilbert, 1975; Gilbert et al., 1975; Anderson et al.,

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1976). This and subsequent work on both invertebrate and vertebrate axons showed that the CANP was a neutral, thiol protease specifically activated by Ca2+, which selectively degraded neurofilament proteins (Gilbert et al., 1975, Schlaepfer and Micko, 1978; Schlaepfer and Freeman, 1980; Pant et d . , 1979a; Pant and Gainer, 1980; Schlaepfer et al., 1981; Eagles et al., 1981; Tashiro and ishizaki, 1982; Malik et al., 1981). It is known that there are two types of CANPs extractable from various tissues, which have different elution positions in DEAE-cellulose chromatography, one of which has a high sensitivity (<40 p M ) and the other a low sensitivity (>500 p M ) to CaZ+(Mellgren, 1980; Dayton et al., 1981; Kubota et al., 1981; De Martino, 1981; Suzuki et al., 1981; Murachi et al., 1981a,b; Mellgren et al., 1982; and Hathaway et al., 1982). It appears that the form of CANP found in axons requires >500 p M of Ca2+(Pant and Gainer, 1980; Pant et al., 1982), whereas the form of CANP found acting on postsynaptic glutamate receptors (Baudry and Lynch, 1980; Baudry et al., 1981) requires lower levels of Ca2+(<40 pM).Murachi et al. (1981b) have shown that of 1 1 tissues studied that had CANP activity, nervous tissue (brain) was the least abundant in the high Ca2+ sensitivity form (referred to by those workers as Calpain I). Although some evidence is available for Calpain I in brain (see Baudry and L,ynch, 1980; Baudry et al., 1981), it is unlikely that this is the type of CANP associated with the neurofilaments in axons. In contrast, Calpain 11 (the low Ca2+sensitivity form of CANP) was quite abundant in brain tissue (Murachi et al., 1981b), and would appear to be the type of CANP found in axoplasrn (Pant and Gainer, 1980; Pant rt al., 1982). lshiura (1981) pointed out that although endogenous inhibitors of CANP have been found (Murachi et a l . , 1981a; Tashiro and Ishizaki, 1982), no endogenous factor has yet been found that could transform CANP I I to CANP I . Some that we have tried without success are AMP, ADP, ATP, trifluoroperazine, calmodulin, and tissue extracts (see also Pant et al., 1982). The only candidate at present is “unphysiological” levels of Ca”. There are three hypotheses with respect to the function of CANP in neurons. It is clear that although axons contain large amounts of CANP, it is in the CANP 11 form and is not activated in the axon under normal circumstances. This makes biological sense in the axon, where the neurofilaments are stable structural components and are degraded only after injury to the nerve when intracellular calcium concentrations could reach millimolar levels. Hence, one hypothesis is that the destruction of neurofilaments preceding nerve degeneration (after injury) is mediated by CANP (Schlaepfer, 1974; Schlaepfer and Micko, 1978). Another hypothesis suggested by Lasek and Hoffman (1976) was that the turnover of

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neurofilaments in nerve terminals may involve the CANP, and that one mechanism regulating the growth of axons might be via the action of CANP in the developing nerve terminals (Lasek and Black, 1977). A problem that remains for the latter proposal is that the axonal CANP directed at neurofilament degradation requires an unphysiological level (>500 wA4) of calcium ion for activation (see above). Even in the synaptic terminal, where during activity the Ca2+influx is relatively high, the internal calcium concentration rarely exceeds 10 p M (Llinas and Heuser, 1977). Studies on the distribution of CANP and endogenous protein substrates in the squid nervous system showed that there was no endogenous substrate or CANP in the nerve terminal, whereas the immediately contiguous area (i.e., the axon) contained both components abundantly (Pant et al., 1982). These studies were interpreted as support for the idea of a programmed turnover of neurofilament as they enter the nerve terminals, with a subsequent inactivation of the CANP (shown to occur with an appropriate time course for axonal CANP, see Pant et al., 1982). The third hypothesis involves the action of CANP I on glutamate receptors in the hippocampus. Baudry and Lynch (1979) first found that Ca2+ appeared to regulate glutamate receptor efficacy, and Vargas et al. (1980) showed that a cysteine proteinase was involved in the Ca2+induced increased binding of glutamate to hippocampal synaptic membranes. Subsequent work (Baudry and Lynch, 1980; Baudry et al., 1981) using the protease inhibitor leupeptin indicated that the protease involved was CANP I. No evidence for CANP I in the squid nervous system has been reported to date.

C. Protein Transfer from Schwann Cells to the Giant Axon

In Section II,D we pointed out that squid axoplasm contained tRNA but virtually no rRNA or mRNA. Despite this fact, when intact axons, isolated from their cell bodies, are incubated with radioactive amino acids, de nouo synthesized proteins are found within the axoplasm (Fischer and Litvak, 1967; Giudfitta et al., 1968; Lasek et al., 1973). Giuditta ef al. (1968) interpreted these data as evidence for de nouo protein synthesis within axoplasm, whereas Lasek et al. (1973) suggested that the labeled axoplasmic proteins were derived from protein synthesis in the Schwann cells with subsequent transfer of these proteins into the axon. The evidence in favor of the latter view (Lasek et al., 1977; Gainer et al., 1977) has been reviewed elsewhere (Gainer, 1978; Lasek and Tytell, 1981) and hence will be only briefly described here.

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The lines of evidence that support the glia-neuron protein transfer hypothesis (see Lasek et al., 1977; Gainer et al., 1977) are as follows: 1. Schwann cells surrounding the axon contain large numbers of polysomes and vigorously incorporate amino acids into proteins, whereas squid axoplasm contains no ribosomes and isolated axoplasm cannot synthesize protein in v i m . The failure of axoplasm to incorporate amino acids into protein is noteworthy, since other energy-dependent complex processes (e.g., lipid biosynthesis and axonal transport) do occur for many hours in isolated axoplasm. 2. Kinetic autoradiographic studies are consistent with this hypothesis (see Lasek et al., 1973, 1977). 3. Perfusion of the axon with artificial buffer for many hours, which would remove any soluble factors necessary for protein synthesis in the axon, does not affect the ability of the sheath cells to synthesize proteins, nor does it prevent the appearance of labeled proteins within the axonal space (i.e,, the perfusate, see Gainer et al., 1977). 4. Injection of RNase into intact axons, or intraaxonal perfusion of RNase, does not inhibit protein synthesis or its transfer to the axoplasm.

The proteins transferred from the Schwann cells to the giant axon are large in number and variety (Lasek et al., 1977; Tytell and Lasek, 1980, 1981; Lasek and Tytell, 1981). However, there are many proteins synthesized in the sheath that are not transferred to the axoplasm, implying a selective process. Efforts have been made to identify the transferred proteins by two-dimensional electrophoresis (Lasek and Tytell, 1981). Most of the transferred proteins do not appear to be structural (or high-abundance) proteins in axoplasm, although labeled proteins corresponding to actin and fodrin do appear to be transferred. No tubulin or neurofilament protein appears to be transferred. A 70,000 M , protein called “traversin” by the authors is the most heavily labeled protein in the axoplasm, but is a minor component in the sheath. Furthermore, the neuronal cell bodies in the stellate ganglion appear to synthesize little “traversin.” Hence, it is believed that the proteins supplied to axoplasm by the Schwann cells differ significantly from those proteins that are synthesized in the cell body and delivered by axonal transport. This indicates that the proteins of different origins may also have different functions in the axon. The mechanisms underlying the protein transfer remain speculative (see Gainer, 1978; Lasek and Tytell, 1981). Tytell and Lasek (1981) have some evidence that the transferred proteins in axoplasm behave as a macromolecular complex held together by ionic interactions.

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D. Proteins and Axolemmal Properties

Attempts to identify the macromolecules in the plasma membrane that are involved in membrane phenomena such as electrogenesis are best served by the availability of highly specific high-affinity ligands (e.g., tetrodotoxin for the Na+ channel; see Armstrong and Matteson; Narahashi, this volume). Given such radioactive ligands, it is possible to measure, and “fish out” by affinity chromatography, specific macromolecules that are either directly or indirectly involved in electrogenesis. The ideal experimental preparation for such biochemical work would be one with a high surface-to-volume ratio so as to increase the number of membrane molecules per mass of tissue. In this regard, the squid giant axon is an unfavorable preparation. However, chemical dissection studies on the cytoplasmic surface of the membrane under experimental conditions that allow for normal membrane excitability (e.g., perfusion) are particularly well suited to the squid giant axon. Such studies have been done extensively on the giant axon preparation using a type of biochemical pharmacology which assumes that the reagent being used is “specific” under the conditions used, and that the “assays” which are used measure various electrical and kinetic membrane parameters. The “reagents” fall into three classes: pH, group-specific reagents (e.g., sulfhydryl reagents), and enzymes (e.g., nucleases, proteases, and lipases). Many of these studies have been reviewed (Rosenberg, 1973, 1981; Tasaki, 1968; Brodwick and Eaton, 1982), and only a brief commentary on this subject will be presented below.

1 , CHEMICAL DISSECTION STUDIES The principal conclusion of the chemical dissection experiments in the squid axon is that proteins on the cytoplasmic side of the plasma membrane are involved in the generation of action potentials. This is because intraaxonally injected or perfused proteases can profoundly affect the action potential (Tasaki and Takenaka, 1964; Tasaki, 1968; Armstrong et al., 1973; Sevcik and Narahashi, 1975; Rojas and Rudy, 1976). Modification of action potentials can be produced by perfusions of either widespectrum proteases (e.g., Pronase) or more specific endopeptidases (e.g., trypsin, chymotrypsin, papain, and ficin). Carboxypeptidases have no effects, nor do nucleases. External applications of these proteases do not affect the action potential, presumably because the surrounding sheath cells and basement membranes prevent the access of the proteases to the axolemmal proteins. As might be expected, phospholipases are

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nonselectively destructive to the axonal membrane (Rosenberg, 1973, 1981). Little is known about the natures of the proteins involved. Whether they are intrinsic or extrinsic membrane proteins, or whether they are part of the macromolecules that constitute the sodium channel, is not known. One line of this type of work has led to a quasi-molecular model of sodium channel inactivation. Armstrong and Bezanilla ( 1977) proposed a model in which sodium-channel inactivation (the only process in the action potential that has been shown to be specifically modified by proteases) is mediated by a cytoplasmic-facing membrane protein that is flexible enough to plug the sodium channel. Rojas and Rudy (1976) showed that proteases that either cleaved specifically at basic amino acid residues (e.g., trypsin and alkaline protease b) or at hydrophobic amino acid residues (e.g., a-chymotrypsin) were equally capable of eliminating sodium inactivation, and that perfusion of various long-chain tetraethylammonium analogs could reverse the protease effect. They concluded that a hydrophobic amino acid residue (e.g., phenylalanine) was involved in stabilizing the protein on the membrane and that a basic amino acid residue on the protein served as the positive charge on the Armstrong-Bezanilla plug. Further experiments designed to test this model (Brodwick and Eaton, 1978, 1982) were generally confirmatory and suggested that the basic residue was arginine and the hydrophobic residue was tyrosine. The observation that prolonged action potentials in the squid axon are produced by alkaline perfusion media (Terakawa and Watanabe, 1976) was also interpreted as being consistent with this model. However, the pK of the decrease in sodium inactivation was found to be between 9.7 and 10.2, too low to be associated with the pK of arginine’s guanidinyl group (pK = 13), but which coincided with the titration range of tyrosine’s phenolic group (Brodwick and Eaton, 1978, 1982). Although this model is interesting, the inferences about protein structures and interactions are totally based on indircct data, and direct information about the protein structures is necessary before further progress can be made.

2 . MEMBRANE-ASSOCIATED PROTEINS In an attempt to gather information about the types of proteins that were located on the cytoplasmic side of the axolemma and their potential roles in maintaining excitability of the giant axon, several of us in collaboration with various members of I. Tasaki’s laboratory embarked on a program of covalent labeling of squid axon membrane proteins. Our initial effort (Gainer et al., 19741, which was to continue over several summers at the Marine Biological Laboratory, Woods Hole, was to use enzymatic

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radioiodination techniques to label proteins covalently on one or the other side of the plasma membrane. Such techniques had been successful in other experimental systems (see Carraway, 1975), and, since we perfused the reagents intraaxonally, only the inside surface of the axolemma was studied. During these perfusions, the size, duration, and conduction velocity of action potentials were routinely monitored. This was particularly important in the radioiodination studies, since the lactoperoxidase-Hz02 used at high concentrations was itself destructive to the excitable process (Gainer c f a / . , 1974; see also Brodwick and Eaton, 1978, 1982, who interpreted this as being due to the modification of the tyrosine residue discussed in Section lll,D, 1). Hence, in our studies concentrations of the reagents that were not deleterious to the action potential were used. This initial study suggested that a specific low molecular weight protein was labeled in the membrane and that this could be prevented by depolarization of the axon (Gainer et al., 1974). Because of technical limitations (i.e., the low levels of radioactivity, due to the above-described constraints on the conditions of in situ iodination), the proteins were poorly resolved on polyacrylamide gels. Takenaka et al. (1976) confirmed these results and further suggested that the low molecular weight protein was being released by the membrane into the perfusate during depolarization. Inoue et al. (1976) studied the release of labeled proteins in the perfused giant axon and reported that the depolarization-induced release required Ca2+in the external medium surrounding the axon. In these studies the proteins associated with the membrane were labeled by nondeleterious levels of N - [3H]ethylmaleimide, and the anion in the perfusion medium was F-. When the F- was replaced by chaotropic anions, various amounts of labeled proteins were released. The extent of protein release was correlated with the anion's position in the lyotropic series (i.e., SCN- >> Br > CI- >> F -), and the loss of the inward current in the axonal membrane was correlated with the extent of protein release. The conclusions of this study were that ( I ) the released proteins were not intrinsic membrane proteins, but subaxolemmal proteins associated with the membrane; and (2) that these subaxolemmal proteins may play some role in the regulation of the excitable process. The next stage of this work was to improve the resolution of the released proteins. This involved changes in the experimental paradigm as well as in the electrophoretic procedures. As a result of this effort (Yoshioka et al., 1978; Pant et af., 1978a,b), it became apparent that the released proteins were composed of both high and low molecular weight proteins and that these proteins might represent the subaxolemmal filamentous network visualized by electron microscopy in the perfused giant axon (Metuzals and Tasaki, 1978; Metuzals et al., 1981).

FIG.1 . Scanning electron micrographs of the internal surface of the axolemma in the perfusion zone in a giant axon. Left: The axon was internally perfused with a KF-containing perfusion solution for 45 min. Right: The axon was perfused with a KI-containing perfusion solution for 10 min. A I-pm scale line is shown beneath each figure. (From Baumgold er al.. 1981a.)

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In a combined morphological-biochemical study, Baumgold et al. (1981a) showed that the filamentous subaxolemmal network in the perfused axon was removed by perfusion with KI (Fig. 1). Analysis of the KI-released proteins by polyacrylamide slab gel electrophoresis and silver staining of the separated proteins provided the best resolution of the released proteins to date (see Fig. 2). The released proteins are composed of a variety of molecular species, many of which appear to be cytoskeletal

FIG.2. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis patterns of proteins from extruded axoplasm (WHOLE AXOPL.), the central core of axoplasm removed from the giant axon before perfusion (CORE), proteins found in the perfusate soon after initiation of perfusion with KF-containing solution (EARLY KF), proteins found in the K F perfusate 25 min after the onset of perfusion (LATE KF), proteins in the perfusate as a result of perfusion for 10 min of K1-containing solutions (KI). The differences in protein pattern between LATE K F and KI columns should represent the composition of the filamentous network shown in Fig. 1. Proteins on gels were stained by the silver method. Marker protein molecular weights (K = thousand) are shown at the left. (From Baumgold et a!., 1981a.)

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( e g , actin and tubulin). One high molecular weight protein that was rcleased and appeared to be concentrated in the subaxolemmal network (as opposed to axoplasm, Fig. 2 ) migrated on the gel above the N F 200 marker and at about the same molecular weight as fodrin (Levine and Willard, 1981). Whether this protein is fodrin or is related to it requires further study. Both proteins also share the property of being concentrated beneath the cell membrane. It has become clear that the proteins released from the subaxolemmal surface of perfused axons by depolarization or chaotropic anions are part of the filamentous network (Metuzals and Tasaki, 1978; Metuzals et ul., 1981) (Fig. 1). It has been shown that tetrodotoxin also can cause some protein release (Baumgold et al., 1981b),and that perfusion of pharmacological reagents that affect microtubules and microfilaments can influence the excitation process (Matsumoto et ul., 1979, 1980). However, Landowne et al. (1982) showed that p-lumicolchicine (which does not disrupt microtubules) was as effective as colchicine in blocking excitation in the giant axon. The roles of these membrane-associated proteins in the maintenance of excitability in the axon remain unclear. They could simply be protecting the membrane against mechanical injury (i.e., against pressure or bending), or they could be more directly involved in modulating the intrinsic membrane proteins (e.g., ion-channel proteins) that mediate the excitable process.

3. IMMUNOLOGICALPROBES Antibodies prepared against the axoplasm of the Chilean squid Dosidic'us gigus were internally perfused into giant axons and under certain

experimental conditions (after 3 hr of exposure) could produce conduction block (Huneeus-Cox, 1964; Huneeus-Cox and Fernandcz, 1967) with little effect on resting membrane potential. The specific experimental conditions that were necessary were ( I ) preperfusion of t h e axon by a 400 m M potassium cystcinate solution for about 10 min; and (2) inclusion of 30 mM potassium cysteinate in the standard perfusion medium (a potassium glutamate, potassium phosphate mixture) during the antibody perfusion. According to the authors, these conditions were absolutely essential for success, and control experiments using anti-bovine serum albumin and anti-neurofilament immunoglobulins under the same conditions did not cause conduction block. Thc authors suggested that these cysteine solutions were necessary to loosen the residual axoplasm so that the large antibodies could physically approach the plasma membrane in order to react with their antigens. The long perfusion times (3 hr) necessary to produce the effect were similarly explained. In view of the subaxolemmal

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filamentous network discussed above (Section III,D,Z), these explanations for the specific conditions used are not unreasonable. However, the experiments were essentially preliminary data, and very primitive (by today’s standards) electrophysiological parameters were measured. It is curious that no further work along these lines has been reported by these or other workers. A few reports have appeared showing that antibodies raised in other species cross-react with antigens present in the squid giant axon. Lasek (1977) has found that antibodies to Myxicola neurofilaments cross-react with an antigen in squid axoplasm, and Yoshioka et al. (1979) have shown that antibodies raised against tubulin isolated from the sea urchin stains regularly arranged fibers in the squid axon, presumably microtubules. This staining could be reduced by intracellular perfusion with KCl, colchicine, and cytochalasin B.

IV.

CONCLUDING REMARKS

In this article we have attempted to provide some information about the biochemistry and metabolism of the squid giant axon system. Because work on the giant axon is the province of a relatively small number of dedicated biochemists, restricted by the seasonal availability of squid, there are many gaps in the information. However, despite these limitations, a great deal of useful data has already been obtained from this model system. As we pointed out earlier, much of the information we have about the biochemistry and metabolism of axons in general is based on studies using the squid giant axon and its axoplasm. The giant axon will continue to serve as a “Rosetta stone” for neurobiologists, neurochemists, and cell biologists. Recent techniques of molecular biology and immunology applied to the giant axon will undoubtedly bring new and exciting insights into axonal function. REFERENCES Alema, S., Calissaano, P., Rusca, G., and Giuditta, A. (1973). Identification of a calcium binding brain specific protein in the axoplasm of squid giant axons. J. Neurochem. 20, 681-689. Anderton, B. H., Bell, C. W., Newby, B. J . , and Gilbert, D. S . (1976). Neurofilaments. Biochem. SOC. Truns. Meet. 53rd. London 4, 544-547. Armstrong, C. M., and Bezanilla, F. (1977). Inactivation of the sodium channel. 11. Gating current experiments. J . Gen. Physiol. 70, 567-590. Armstrong, C. M., Bezanilla, F., and Rojas, E. (1973). Destruction of sodium conductance inactivation in squid axons perfused with pronase. J. Gen. Physiol. 62, 375-391.

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Baker. P. F., and Carruthers, A. (1981a). Sugar transport in giant axons of Loligo. 1.Physiol. (London) 316, 48 1-502. Baker, P. F., and Carruthers, A. (1981 b). 3-0-methylglucose transport in internally dialysed giant axons of Loligo. J . Physiol. (London) 316, 503-525. Baker, P. F . , and Crawford, A. C. (1972). Mobility and transport of magnesium in squid giant axons. J . Physiol. (London) 227, 855. Baker, P. F., and Shaw, T. 1. (1965). A comparison of phosphorus metabolism of intact squid nerve with that of the isolated axoplasm and sheath. J . Physiol. (London) 180, 424-438. Baker, P. F., Hodgkin, A. L., and Shaw, T. I. (1962). Replacement of the axoplasm of giant nerve fibres with artificial solutions. J . Physiol. (London) 164, 330-350. Bandyopahyay, A. K., and Deutscher, M. P. (1971). Complex of aminoacyl-transfer R N A synthetase. J. Mol. Biol. 60, 113-122. Barra, H. S . , Rodriguez, J. A., Arce, C. A . , and Caputto, R. (1973). A soluble preparation from rat brain that incorporates into its own proteins I4C-arginine by a ribonucleasesensitive system and I4C-tyrosineby a ribonuclease insensitive system. J . Neurochem. 20, 97- 108. Barra, H. S . , Arce, C. A., and Caputto, R. (1980). Total tubulin and its aminoacylated and non-aminodcylated forms during the development of rat brain. Eur. J . Biochem. 1u9, 439-446. Baudry. M., and Lynch, G . (1979). Regulation of glutamate receptors by cations. Noturr (London) 282, 748-750. Baudry, M., and Lynch, G. (1980). Regulation of hippocampal glutamate receptors: Evidence for the involvement of a calcium activated protease. Proc. Nutl. Acad. Sci. U . S . A . 71, 2298-2302. Baudry, M., Bundman, M . C., Smith, E. R . , and Lynch, G . S . (1981). Micromolar calcium stimulates proteolysis and glutamate binding in rat brain synaptic membranes. Science 212, 937-938. Baumgold, J., Terakawa, S . , Iwasa, K., and Gainer, H. (1981a). Membrane-associated cytoskeletal proteins in squid giant axons. J . Neurochem. 36, 759-764. Baumgold, J., Gallant, P., Terakawa, S . , and Pant, H . (1981b). Tetrodotoxin affects submembranous cytoskeletal proteins in perfused squid giant axons. Biochem. Biophys. Res. Commun. 103, 653-658. Bear, R. S . , and Schmitt, F. 0. (1939). Electrolytes in the axoplasm of the giant nerve fibers of the squid. J . Cell. Comp. Physiol. 14, 205-215. Bear, K. S . , Schmitt, F. O., and Young, J. Z. (1937). Investigations on the protein constituents of nerve axoplasm. Proc. R . SOC. Ser. B 123, 520-529. Bicher, H., and Ohki, S. (1972). Intracellular pH electrode experiments on the giant squid axon. Biochim. Biophys. Acta 255, 900. Black, M. M., and Ldsek, R. J. (1977). The presence of transfer RNA in the axoplasm of the squid giant axon. J . Neurobiol. 8, 229-237. Bonting, S . L., and Caravaggio, L. L. (1962). Sodium-potassium activated adenosine triphosphatase in the squid giant axon. Nuture (London) 194, 1180-1181. Borisy, F. G . , and Taylor, E. W . (1967). The mechanical action of colchicine. J . Cell B i d . 34, 525-533. Boron, W. F., and DeWeer, P. (1976). Intrdcellular pH transients in squid giant axons caused by COz, NH,, and metabolic inhibitors. J . Gen. Physiol. 67, 91-112. Brodwick, M. S . , and Eaton, D. C. (1978). Sodium channel inactivation in squid axon is removed by high internal pH or tyrosine-specific reagents. Science 200, 1494-1496. Brodwick, M. S . , and Eaton, D. C. (1982). Chemical modification of excitable membranes.

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In “Proteins in the Nervous System: Structure and Function” (B. Haber, J. R. PerezPolo, and J. D. Coulter, eds.), pp. 51-72. Liss, New York. Brunetti. M., Guiditta, A , . and Porcellati. G. (1979). The synthesis of choline phosphoglycerides in the giant fibre of the squid. J . Nettrochem. 32, 3 19-324. Caldwell, P. C. (1958). Studies on the internal pH of large muscle and nerve fibers. J . Physiol. (London) 142, 22. Caldwell, P. C. (1960). The phosphorus metabolism of squid axons and its relationship to the active transport of sodium. J . Physiol. (London) 152, 545-560. Caldwell, P. C., and Keynes, R. D. (1957). The utilization of phosphate bond energy for sodium extrusion from giant axons. J . Physiol. (London) 137, 12p-13p. Caldwell, P. C., Hodgkin, A. L., Keynes, R. D., and Shaw, T. I. (1964). The rate of formation and turnover of phosphorus compounds in squid giant axons. J . Physiol. (London) 171, I 19-13 1. Camejo, G., Villegas, G. M., Barnola, F. V . , and Villegas, R. (1969). Characterization of two different membrane fractions isolated from the first stellar nerve of the squid, Dosidicus gigas. Biochim. Biophys. Acta 193, 247-259. Canessa, M. (1965). Properties of ATPase activities of membrane fractions from the sheath of squid giant axons. J . Cell. Comp. Physiol. 66, 165-167. Cannessa-Fisher, M., Zambrano, F., and Riveros-Moreno, V. (1968). Properties of the ATPase system from the sheath of squid giant axons. Arch. Biochem. Biophys. 122, 658-663. Carraway, K . L. (1975). Covalent labeling of membranes. Biochbn. Biophys. Actu 415, 379-40 I . Cavanaugh, G. M. (1975). “Formulae and Methods VI of the Marine Biological Laboratory Chemical Room,” pp. 67-68. Marine Biological Laboratory, Woods Hole, Massachusetts. Condrea, E., and Rosenberg, P. (1968). Demonstration of phospholipid splitting as the factor responsible for increased permeability and block of axonal conduction induced by snake venom. Biochim. Biophy.?. Actu 150, 271-284. Davison, P. F., and Huneeus, F. C. (1970). Fibrillar proteins from squid axons. Part 2: Microtubule protein. J . Mol. Biol. 53, 429-439. Davison, P. F., and Taylor, E. W. (1960). Physical-chemical studies of proteins of squid nerve axoplasm with special reference to the axon fibrous protein. J . Gen. Physiol. 43, 801-8 19. Dawson, R . M. C., and Eichberg, J. (1965). Diphosphoinositides and triphosphoinositides in animal tissue. Biochem. J . 96, 634-643. Dayton, W. R., Schollmeyer, J . V., Lepley, R. A., and Cortex, L. R. (1981). A calciumactivated protease possibly involved in myofibrillar protein turnover: Isolation of a low calcium requiring form of the protease. Biochim. Biophys. Acta 659, 48-61. Deffner, G. J. (1961). The dialysable free organic constituents of squid blood: A comparison with nerve axoplasm. Biochim. Biophys. Acta 47, 378-388. Deffner, G. J . , and Hafter, R. E. (1960). Chemical investigations of the giant nerve fiber of the squid. Biochim. Biophys. Actu 42, 200-205. DeMartino, G. N. (1981). Calcium-dependent proteolytic activity in the rat liver: Identification of two proteases with different calcium requirements. Arch. Biochem. Biophys. 211, 253-257. Doane, M. G. (1967). Fluorometric measurement of pyridine nucleotide reduction in the giant axon of the squid. J . Gen. Physiol. 50, 2603-2632. Dole, W. P., and Simon, E. J. (1974). Effects of levorphanol on phospholipid metabolism in the giant axon of the squid. J . Neurochem. 22, 183-185.

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Droz, B., and Koenig, H. L. (1971). Dynamic condition of protein in axons and axon terminals. Acru Neuropnrhol. (Berlin) 5, 109-1 18. Eagles, P. A. M., Gilbert, D. S . , and Maggs, A . (1981). The location of phosphorylation sites and calcium dependent proteolytic cleavage sites on the major neurofilament polypeptides from Myxicola infundibulum. Biochem. J . 199, 101-1 I I. Eyrich, T. L., Barrett, D., and Rock, P. A. (1976). Phospholipase activity in squid and frog axons. J. Neurochem. 26, 1079-1085. Fischer, S., and Litvak, S. (1967). The incorporation of microinjected 14C-aminoacids into TCA insoluble fractions of the giant axon of the squid. J . Cell. Physiol. 70, 69-74. Fischer. S . , Collins, M., Gariglio, A , . and Tellez-Nagel, 1. (196X). Protein and KNA metabolism of squid axons (Dosidicus gigcrs). J . Gcw. Phvsio/. ( S u p p l .J 51, 72s-XOb. Folch, J., Lees, M., and Sloane-Stanley, G. H. (1957). A simple method for the isolation and purification of total lipids from animal tissues. J . Biol. Chem. 226, 497-509. Gainer, H . (1978). Intercellular transfer of proteins from glial cells to axons. Trends Neurosci. 1, 93-97. Gainer, H., and Gainer, V. S. (1976). Proteins in the squid giant axon. In “Electrobiology of Nerve, Synapse and M u d e ” (J. P. Rueben c r ul., cds.), pp. IY-168. Raven. New York. Carbone, E., Singer, I . , Sisco, K., andTasaki, I . (1974). Depolarization induced Gainer, H., change in the enzymatic radio-iodination of a protein on the internal surface of the squid giant axon membrane. Comp. Biochem. Physiot. Ser. A . 47, 477-484. Gainer, H . , Tasaki, I., and Lasek, R. J. (1977). Evidence for the glia neuron protein transfer hypothesis from intracellular perfusion studies of squid giant axons. J. Cel/ Biol: 74, 524-530. Gilbert, D. L. (1974). Physiological uses of the squid with special emphasis on the use of the giant axon. In “A Guide to Laboratory Use of the Squid Loligo peulei” (.I,M. Arnold ei nl., eds.), pp. 45-54. Marine Biological Laboratory, Woods Hole, Massachusetts. Gilbert, D. S. (1975). Axoplasm chemical composition in Myxicola and solubility properties of its structural proteins. J . Physiol. (London) 253, 303-319. Gilbert, D. S . , Newby, B. J., and Anderton, B. H . (1975). Neurofilament disguise, destruction and discipline. Nature (London) 256, 586-589. Giuditta, A , , Dettbarn, W. D., and Brizin, M. (1968). Protein synthesis in the isolated giant axons of the squid. Proc. Nut/. Acad. Sci. U . S . A . 59, 1284-1287. Giuditta, A., Metafora, S . , Felsami, A., and Del Rio, A . (1977). Factors of protein synthesis in the axoplasm of squid giant axons. J . Neurochem. 28, 1393-1395. Giuditta, A., Cupello, A., and Lazzarini, G . (1980). Ribosomal RNA in the axoplasm of the squid giant axon. J . Neurochem. 34, 1757-1760. Gould, R. M. (1975). lnositol lipid synthesis localized in axons and unmyelinated fibers of peripheral nerve. Bruin Res. 117, 169-174. Gould, R. M., Pant, H., Gainer, H., and Tytell, M. (1982a). Studies of phospholipid synthesis in mammalian and invertebrate axons and terminals. In “Phospholipids of the Nervous System’’ (L. A. Horrocks, G. B. Ansell, and G. Porcellati, eds.), Vol. I , pp. 221-229. Raven, New York. Gould, R. M., Mancuso. C. A., Gallant, P., and Tasaki. 1. (1982b). Incorporation of 32P-phosphate into lipids and proteins by intact squid giant axons. Biol. Bull. 163, 400. Gould, R. M., Pant, H., Gainer, H . , and Tytell, M. (1983a). Phospholipid synthesis in the squid giant axon: Incorporation of lipid precursors. J . Neurochem. (in press).

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Gould, R. M., Spivack, W. D., Posnansky. M., and Robertson, D. (1983b). Phospholipid synthesis in the squid giant axon: Enzymes of phosphatidylinositol melabolism. J . Naitrochrtn. (in presb). Gunning, P. W., Por, S. B., Langford, C. J . , Scheffer, J., Austin, L., and Jeffrey, P. L. (1979). The direct measurement of the axoplasmic transport of individual RNA species: Transfer but not ribosomal RNA is transported. J . Neurochem. 32, 1737-1743. Guroff, G. (1964). A neutral, calcium-activated protease from the soluble fraction of the rat brain. J . Biol. Chem. 239, 149-155. Hammerschlag, R., and Bobinski, J. A. (1981). Ca" or Mg" stimulated ATPase activity in bullfrog spinal nerve. J . Neurochem. 36, 11 14-1 127. Hathaway, D. R., Werth, D. K., and Haeberle, J . R. (1982). Limited autolysis reduces the Ca" requirements of a smooth muscle Ca"-activated protease. J . Biol. Chem. 257, 9072-9077. Heumann, R., Villegas, J., and Herzfield, D. W. (1981). Acetylcholine synthesis in the Schwann cell and axon in the giant nerve fiber of the squid. J . N r t ~ r o c h e m .36, 765-768. Hochachka, P. W., Moon, T. W., Mustafa, T., and Storey, K . B. (1975). Metabolic sources of power for mantle muscle of a fast swimming squid. Comp. Biochem. Physiol. 52B, 15 1-1 58. Hodge, A. J., and Adelman, W. J., Jr. (1980). The neuroplasmic network in Loligo and Hermissenda neurons. J . Ultrustruct. Res. 70, 220-241. Hodgkin, A . L. (1965). "The Conduction of the Nervous Impulse." Thomas, Springfield, Illinois. Hodgkin, A. L . , and Katz, B. (1949). The effect of calcium on the axoplasm of giant nerve fibers. J . Exp. B i d . 26, 292-294. Hodgkin, A. L., and Keynes, R. D. (19.5s). Active transport ofcations in giant axons from Srpiu and Loligo. J . Physiol. (London) 128, 28-60. Hoskin, F. C. G. (1966). Anaerobic glycolysis in parts of the giant axon of squid. Nature (London) 210, 856-857. Hoskin, F. C. G. (1976). Distribution of DFP hydrolyzing enzyme EC-3.8.2.1 between sheath and axoplasm of squid giant axon. J . Neurochem. 26, 1043-1045. Hoskin, F. C. G . , and Brande, M. (1973). An improved sulphur assay applied to a problem of isethionate metabolism in squid axon and other nerves. J . Nrrrrorhrtn. 20, 13171327. Hoskin, F. C. G . , and Kordik, E. R. (1977). Hydrogen sulfide as a precursor for the synthesis of isethionatc in the squid giant axon. Arch. Bioc+em. Biophys. 180, 583-586. Hoskin, F. C. G . , and Long, R. (1972). Purification of a DFP-hydrolyzing enzyme from squid head ganglion. Arch. Biochrm. Biophy.s. 150, 548-555. Hoskin, F. C . G . , and Rosenberg, P. (1965). Penetration of sugars, steroids, amino acids, and other organic compounds into the interior of the squid giant axon. J . Gen. Physiol. 49,47-56. Hoskin, F. C. G., and Rousch, A. H. (1982). Hydrolysis of nerve gas by squid-type diisopropylphosphorofluoridate hydrolysing enzyme on agarose resin. Science 215, 1255-1257. Hoskin, F. C. G . , Pollock, M. L., and Prusch, R. D. (1975). An improved method for the measurement of I4C0?applied to a problem of cysteine metabolism in squid nerve. J . Nerrroc,/iern. 25, 445-449. Huneeus, F. C., and Davison, P. F. (1970). Fibrillar proteins from squid axons, Part 1: Neurofilament protein. J . Mol. B i d . 52,415-428.

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Huneeus-Cox, F. (1964). Electrophoretic and immunological studies of squid axoplasm proteins. Science 143, 1036-1037. Huneeus-Cox, F., and Fernandez, H. L. (1967). Effects of specific antibody on the excitability of internally perfused axons. J . Gen. Physiol. 50, 2407-2419. Ingoglia, N. (1978). The effect of intraocular injection of cordycepin on retinal RNA synthesis and on RNA axonally transported during regeneration of the optic nerves of goldfish. J . Neurochem. 30, 1029-1033. Ingoglia, N. (1982). 4 s RNA in regenerating oplic axons of goldfish. J. Neurosci. 2,331-338. Ingoglia, N., and Giuditta, A. (1982). Multiple species of transfer RNA are present in squid axoplasm. Soc. Neurosci. Annu. Meet., 13rh, Abstr. No. 313. Ingoglia, N . A,, and Tuliszewski, A. (1976). Transfer RNA may be axonally transported during regeneration of goldfish optic nerves. Brain Res. 112, 371-381. Ingoglia. N. A,, and Zanakis. M. F. (1982). Axonal transport of 4 s RNA. In “Axoplasmic Transport” (D. G. Weiss, ed.), pp. 161-169. Springer-Verlag, Berlin and New York. Inoue, I., Pant, H. C., Tasaki, I., and Gainer, H. (1976). Release of proteins from the inner surface of squid axon membrane labeled with tritiated N-ethyl maleimide. J . Gen. Physiol. 68, 385-395. Ishiura, S. (1981). Calcium dependent proteolysis in living cells. Life Sci. 29, 1079-1088. Keynes, R. D. (1963). Chloride in the squid giant axon. J . Physiol. (London) 169, 690. Khan, M. A., and Oehs, S. (1974). Magnesium or calcium activated ATPase in mammalian nerve. Bruin Res. 81, 413-426. Koechlin, B. A. (1955). On the chemical composition of the axoplasm of squid giant fibers with particular reference to its ion palterns. Biochim. Biophys. A c f a 1, 5 1 1-529. Kubota, S ., Suzuki, K., and Imahori, K. (1981). A new method for the preparation of a calcium-activated neutral protease highly sensitive to calcium ions. Biochem. Biopkys. Res. Commun. 100, 1189-1 194. Landowne, D., Larsen, J., and Taylor, K. (1982). Colchicine blocks nerve excitation: An optical study. Biol. Bull. 163, 386. Larrabee, M. G., and Brinley, F. J., Jr. (1968). Incorporation of labelled phosphate into phospholipids in squid giant axons. J . Neurochem. 15, 533-545. Lasek, R. J. (1970). The distribution of nucleic acids in the giant axon of the squid (Loligo pealii). J . Neurochem. 17, 103-109. Lasek, R. J. (1977). Neurofilaments: Structure, composition and axonal transport. Proc. I n t . Soc. Neurosci. 6, 75. Lasek, R. J., and Black, M. M. (1977). How do axons stop growing? Some clues from the metabolism of the proteins in the slow component of axonal transport. In “Mechanisms, Regulations, and Special Functions of Protein Synthesis in the Brain” (E. Roberts et a / . , eds.), pp. 161-169. Elsevier, Amsterdam. Lasek, R. J., and Hoffman, P. M. (1976). The neuronal cytoskeleton, axonal transport and axonal growth. I n “Cell Motility” (R. Goldman et a / . , eds.), pp. 1021-1049. Cold Springs Harbor Laboratory, Cold Springs Harbor, New York. Lasek, R. J . . and Tytell, M. A. (1981). Macromolecular transfer from glia to the axon. J . E x p . B i d . 95, 153-165. Lasek, R. J., Dabrowski, C., and Nordlander, R. (1973). Analysis of axoplasmic RNA from invertebrate giant axons. Nature (London) 244, 1188-1192. Lasek, R. J., Gainer, H . , and Przybylski, R. J. (1974). Transfer of newly synthesized proteins from Schwann cells to the squid giant axon. Proc. Natl. Acad. Sci. U . S . A .71, 1188-1 192.

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Lasek, R. J., Gainer, H., and Barker, J. L . (1977). Cell to cell transfer ofglial proteins to the squid giant axon: The glia neuron protein transfer hypothesis. J . Cell B i d . 74,501-523. Levine, J., and Willard, M. (1981). Fodrin: Axonally transported polypeptides associated with the internal periphery of many cells. J . Cell B i d . 90, 631-645. LlinBs, R. R., and Heuser, J. E. (1977). Depolarization-release coupling systems in neurons. Neurosci. Res. Prog. Bull. 15, 557-687. Malik, M. N., Meyers, L. A., Igbal, K., Sheikh, A. M., Scotto, L., and Wisniewski, H. M. (1981). Calcium-activated proteolysis of fibrous protein in central nervous system. Life Sci. 29, 795-802. Martin, K . , and Shaw, T. I. (1970). The production of adenosine triphosphate in perfused giant axons of Loligo. J . Physiol. (London) 208, 171-185. Matsumoto, G., Kobayashi, T., and Sakai, H. (1979). Restoration of the excitability of squid giant axon by tubulin-tyrosine ligase and microtubule proteins. J . Biochem. 86, 11551158. Matsumoto, G., Mirofushi, H . , and Sakai, H. (1980). The effects of reagents affecting microtubules and microfilaments on the excitation of the squid giant axon measured by the voltage-clamps method. Biomed. Res. 1, 355-358. Matsumura, F., and Clark, J . M. (1982). ATP-utilizing systems in the squid axon. Prog. Neurobiol. 18, 231-255. Maxfield, M. (1953). Axoplasmic proteins of the squid giant nerve fiber with particular reference to the fibrous protein. J . Gen. Physiol. 37, 201-216. McColl, J . D., and Rossiter, R. J. (1951). Lipids of the nervous system of the squid Loligo pealei. J . Exp. B i d . 28, 116-124. McMurray, W. C., McColl, J. D., and Rossiter, R. J. (1964). A comparative study of the lipids of invertebrate nervous system. In “Comparative Neurochemistry” (D. Richter, ed.), pp. 101-107. Macmillan, New York. Mellgren, R. L. (1980). Canine cardiac calcium-dependent proteases: Resolution of two forms with different requirements for calcium. FEBS L e f t . 109, 129-133. Mellgren, R. L., Repetti, A , , Muck, T. C., and Easly, J. (1982). Rabbit skeletal muscle calcium-dependent protease requiring millimolar Ca2+.Purification, subunit structure and Ca2+dependent autolysis. J . B i d . Chem. 257, 7203-7209. Metuzals, J., and Tasaki, I. (1978). Subaxolemmal filamentous network in the giant nerve fiber of the squid Loligo pealei and its possible role in excitability. J . Cell B i d . 78, 597-621. Metuzals, J., Tasaki, I., Terakawa, S . , and Clapin, D. F. (1981). Removal of the Schwann sheath from the giant nerve fiber of the squid: An electron-microscopic study of the axolemma and associated axoplasmic structures. Cell Tissue Res. 221, 1-15. Morris, J. R., and Lasek, R. J. (1982). Stable polymers of the axonal cytoskeleton: The axoplasmic ghost. J . Cell B i d . 92, 192-198. Mullins, L. J., and Brinley. F. J. (1967). Some factors influencing sodium extrusion by internally dialysed axons. J . Gen. Physiol. 50, 2333-2355. Murachi, T., Tanaka, K., Hatanaka, M . , and Murakami, T. (1981a). Intracellular Ca*+dependent protease (calpain) and its high molecular weight and endogenous inhibitor (calpastatin). Adu. Enzyme Regul. 19, 407-424. Murachi, T., Hatanaka, M., Yasumoto, Y., Nakayama, N., and Tamaka, K . (1981b). A quantitative distribution study on calpain and calpastatin in rat tissue and cells. Biochem. Int. 2, 651-656. Nachmansohn, D., and Neumann, E. (1975). “Chemical and Molecular Basis of Nerve Activity.” Academic Press, New York.

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Oikawa, T., Spyropoulos, C. S . , Tasaki, I . , and Teorell, T. (1961). Methods for perfusing the giant axon of Lolipo pealei. Acta Physiol. Scund. 52, 195-196. Pant, H. C., and Gainer, H. (1980). Properties of calcium-activated protease in squid axoplasm which selectively degrades neurofilament proteins. J. Neurohiol. 11, 1-12. Pant. H . C., Terakawa. S . , Yoshioka. T . , Tnsaki. I., and Gainer, H. (1976). Evidence for the utilization of extracellular gamma phosphorus-32 A‘I‘P for the phosphorylation of intracellular proteins in the squid giant axon. Biochim. Biop/?ys. Acltr 582. 107-1 14. Pant, H. C., Shecket, G.. Gainer, H., and Lasek, R. J. (1978a). Neurofilament protein is phosphorylated in the squid giant axon. J . Cell Biol. 78, R23-R27. Pant, H. C., Terakawa, S. , Baumgold, J., Tasaki, I., and Gainer, H. (1978b). Protein release from the internal surface of the squid giant axon membrane during excitation and potassium depolarization. Biochim. Biophys. Acta 513, 132-140. Pant, H. C., Terakawa, S . , and Gainer, H. (19794. A calcium activated protease in squid axoplasm. J . Neurochem. 32, 99-102. Pant, H. C., Yoshioka, T., Tasaka, I . , and Gainer, H. (1979b). Divalent cation dependent phosphorylation of proteins in squid giant axon. Bruin Rev. 162, 303-314. Pant, H. C., Pollard, H. B., Pappas, G. D., and Gainer, H. (1979~).Phosphorylation of specific, distinct proteins in synaptosomes and axons from squid nervous system. Proc. N u t / . Acud. Sci. U . S . A . 76, 6086-6090. Pant, H. C., Gallant, P. E., Could, R., and Gainer, H. (IY82). Distribution of calciumactivated protease activity and endogcnous substrates in the squid nervous system. J . Nc2urosc.i. 2, 1578-1587. Potts, W. T. W . , and Parry, G. (1964). “Osmotic and Ionic Regulation in Animals.” Pergamon, Oxford. Rawlins, F. A. (1973). A time-sequence autoradiographic study of the in vivo incorporation of [1,2-3H] cholesterol into peripheral nerve myelin. J. Cell Biol. 58, 42-53. Rojas, E., and Rudy, B. (1976). Destruction of the sodium conductance inactivation by a specific protease in perfiised nerve fibers from Loligo. J . Phy.tio/. ( L m d o r i ) 262, 501-531.

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CURRENT TOPICS IN MEMBRANES A N D TRANSPORT, VOLUME 22

Transport of Sugars and Amino Acids P . F . BAKER A N D A . CARRUTHERS' Lkpurtment o j Physiology Uniuersity of London King's College London, Englund

I. Introduction ..... . . . . . . . . . . . 91 11. Concentratio ugars . . . . . . . . . . . 92 A. Concentrations in Axoplasm Compared with Those in Squid Blood ...... 92 92 B. State of Sugars and Amino Acids in Axoplasm.. ....................... C. Metabolism of Sugars and Amino Acids in Axoplasm . . . . . . . . . . . 96 111. Transport of Sugars by ......................... 98 A. Kinetics of Transfer ...... B. Regulation of Sugar Transport in Squid Axons . . . . . . IV. Descriptive Models for Sugar Transport . . . . . . . . . . . . . . . . A. Hexose Transfer in the Giant Axon is Asymmetric . . B . Mechanisms for Hexose Transfer in the Giant Axon. C. Regulation of Transport by ATP.. .................................... I18 V. Transport of Amino Acids . . . . . . ................... A. Categories of Amino Acid Transport. ................................. I19 B. Glycine Transport ............................... C. Glutamate Transport. ............................ VI. General Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . ................................................. 128

1.

INTRODUCTION

Glucose and amino acids are present in squid blood and axoplasm in appreciable amounts (Deffner, 1961). They are taken up by the giant axon and contribute significantly to axonal function in a number of ways (Hoskin and Rosenberg, 1966; Baker and Potashner, 1973; Caldwell and Lea, 1978; Baker and Carruthers, 1981a). Thus, axoplasmic amino acids

' Present address: Department of Biochemistry, University of Massachusetts Medical Center, Worcester, Massachusetts. 91 Copyright 0 1984 by Academic Press. Inc. Ail rights of reproduction In any form reserved. ISBN 0-12-153322-0

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constitute at least one-third of the total intracellular osmotically active species, glucose and amino acids serve as intermediates in energy-rich phosphate synthesis, and synaptic transmission at the squid giant synapse is mediated by glutamate or aspartate-like species and uptake of these amino acids may influence transmission within the synapse. The detailed properties of sugar transport in the giant axon have been described in some detail by Caldwell and Lea (1978, 1979), and some information on glutamate transport has been provided by Baker and Potashner (1973). The magnitude of the observed fluxes covers a very wide range (see Table VIII). Highest is D-glucose, the maximum fluxes of which are of the same order as the ouabain-sensitive fluxes of Na and K . L-Glutamate and L-arginine are at the lower end of the range with fluxes almost three orders of magnitude lower than those of D-glucose. This article describes the basic properties of sugar and amino acid fluxes in the giant axon of Loligo forbesi and their regulation. Where appropriate data are available, we have attempted to model the transport systems to provide a rationale for experimental observations.

II.

CONCENTRATION AND STATE OF SUGARS AND AMINO ACIDS IN AXOPLASM

A. Concentrations in Axoplasm Compared with Those in Squid Blood

Table I lists the nonelectrolyte composition of squid blood and axoplasm. Most notable are the high concentrations in axoplasm of isethionate, aspartate, and taurine, although other amino acids, including glutamate, alanine, betaine, and homarine, are also present in appreciable amounts. D-Glucose levels in both blood and axoplasm are roughly 1 mM, which implies either a slow rate of hexose metabolism by the giant axon or a very high permeability to glucose. It is possible that some of the sugars and amino acids in squid blood are within cellular elements, but this possibility has not been investigated thoroughly. 6. State of Sugars and Amino Acids in Axoplasm

1 . SUGARS Sugars are essentially free to diffuse in axoplasm (Baker and Carruthers, 1981a; also see Table I1 and Fig. l ) . The longitudinal self-diffusion coefficient of D-ghCOse in axoplasm is 4 to 6 x cm2 sec-I. This

TABLE 1 NONELECTROLYTE COMPOSITION OF SQUID BLOOD A N D AXOPLASM~

(mM)"

Axoplasm (mMY

1.68 2 0. 3" 1.6 I .8 2.2 2.5 2.9 1.25 4.6 4.3 3.3 3.15 1.25 2.6 2. I 3.7 0.7 0.4 0.5 1.2

1.30 2 0. 7'' 182.9 87.8 23.6 12.9 9.5 4.4 118.5 81.8 22.6 3.2 3.2 2.7 1.2 0.8 0.7 0.6 0.5 2.7

42.6

556.9

3.6

49.8

Blood

Substance D-Glucose Isethionic acid Aspartic acid Glutarnic acid Glycine Alanine Serine Taurine Betaine Homarine Leucine Isoleucine Valine Proline Tyrosine Phen ylalanine Methionine Citrulline Threonine Total Total content (%) 'I

Data from Deffner (1961)

* Calculated assuming that

I g of blood contains I g of HZO. Calculated as millirnoles . kg H20 I. Data from Baker and Carruthers (1981a). I

TABLE 11 SELF-DIFFUSION OF SUGARS I N AXOPLASM A N D 0.4% AGARGEL A T 21°C" Diffusion coefficient (cm2 sec-') Axoplasm Sugar 3-O-Meth ylglucose D-Glucose a-Methyl-Dglucopyranoside 2-Deoxy-~-ghcose

Longitudinal

n

Agar gel

Radial

4.3 x

(2)

4.6

X

(1)

-

4.0 x lo-'

(2)

-

4.1 x

(I)

-

4.2 x

" Data are from Baker and Carruthers (1981a).

n

Longitudinal

n

(1)

4.3 x 10-6

(3)

4.2 x

(2)

-

-

P. F. BAKER AND A. CARRUTHERS

94

-

A

0.3

C

E

___,

I

t

0 -16

0

16-16

0

Distance from cenier of inlation imml

16

0

30 Time (mint

FIG.I . The self-diffusion of 3-0-methylglucose in agar gel and axoplasm. (A) Longitudinal self-diffusion of 3-0-methylglucose in 0.4% agar gel. Ordinate: activity in sample over total activity in the gel. Abscissa: distance from the center of injection in millimeters. The gel was injected with a 3-mm patch of I4C-labeledsugar. left at 20°C for 3 hr, frozen, and cut into 2-mm sections for counting. The bar marks the nominal site of injection, and the arrow the side of injection. The curve is drawn from the equation (Crank, 1956)

where Cis the concentration of tracer at time t and distance x from the middle of the injected patch, and C, is its concentration at I = 0. h is half the width of the injected patch. The smooth curve was drawn with D = 4.34 x cm2 sec-I. (B) Longitudinal self-diffusion of 14C-labeled3-0-methylglucose in axoplasm. The bar and arrow have the same meaning as in (A). Temperature, 21°C; t , 3 hr. The smooth curve was cm2 sec-' on the assumption of no loss of isotope. The loss of drawn with D = 4.41 x isotope during the experiment was 17%. Axon diameter was 63.5 p m . (C) Radial diffusion of 14C-labeled 3-0-methylglucose in axoplasm. At time zero I4Clabeled sugar was injected along the long axis of the fiber and samples of radioactivity emerging from the axon were taken every minute for the first 10 min, then at 2-min intervals. Temperature. IYC; axon diameter, 770 p m . The smooth curve is calculated from the equation

where y ( a , I ) is the concentration of tracer at time f at the edge of an insulated cylinder of radius a ; rx, is a positive root (>0) of J , ( a )= 0; JI and Jo are Bessel functions; and D is the diffusion coefficient, which was taken as 4.18 x 10P cmz sec-I. The application of this equation to 3-0-methylglucose efflux assumes that the efflux is directly proportional to 13-0methylg!ucose] inside and there is no lag in transporting the sugar out of the fiber. (From Baker and Carruthers, 1981a.)

95

TRANSPORT OF SUGARS AND AMINO ACIDS

Oe50

r

cathode

anode

u) L

C

a

0

0

-

0.25

L L

0

L

C

3 0

0

0

- 20

0

Distance from site of injection

+20

- mm

FIG.2. Mobility of L-( ''Clglutamate in an axon of Lo/i,yct.fitrhesi.The axon was injected over 3 m m with a mixture of ''Na and ['4Cjglutamate and subjected to a longitudinal electric field of I V/cm for 90 min. The injector entered the fiber from the end that was subsequently made the cathode. The axon was processed as described in the legend to Fig. 1A. Lines have been drawn by eye through the experimental points. Temperature, 21°C; axon diameter, 700 p m . (Unpublished data of Raker and Carruthers.)

compares well with the value 6.7 X lo-("an2sec-' for the diffusion coefficient of glucose in aqueous solutions at 25°C. Very similar values are found for the nonrnetabolizable sugars 3-0-methylglucose and a-methylD-glucopyranoside both in axoplasm and agar gel, indicating that the gel structure of axoplasm offers little resistance to the diffusion of sugars in the cytosol.

2. AMINOACIDS Only glutamate has been studied in any detail. The coefficient for longitudinal self-diffusion of glutamate in axoplasm is 6.5 X lo-(" cm2 sec-I. This compares with coefficients for diffusion in agar gel and aqueous solution of 6.6 x lop6 and 9 x lo-("cm2 sec-I, respectively. Figure 2 shows that not all glutamate injected into axoplasrn is free to diffuse. Here and an axon was injected axially with a small patch of ~-['~C]glutamate 24Na.A voltage ( 1 V/crn) was applied along the length of the fiber. 24Na moves to the cathodic (-uu) end of the fiber and a portion (65%) of the I4C to the anodic ( + u e ) end. However, the rest of the l4Cremains at the site of injection. This immobile I4C may arise from uptake of glutamate into

96

P. F. BAKER AND A. CARRUTHERS

organelles, from complexation of glutamate with axoplasmic constituents, or from metabolic conversion of glutamate to uncharged species. lnformation for other amino acids is lacking. C. Metabolism of Sugars and Amino Acids in Axoplasm 1. SUGARS

Table 111 lists the extent of metabolism of four different sugars by squid giant axons. Only glucose and 2-deoxy-~-glucose are metabolized significantly. 3-0-Methylglucose and a-methyl-D-glucopyranoside appear not to undergo metabolic transformation. A significant portion (45%) of the activity emerging from a-~-[U-'~C]glucose-injected fibers is acid labile and is probably 14C02.This implies that the giant axon is capable of oxidative metabolism of glucose.

2. AMINOACIDS Caldwell and Lea (1978) have reported that glycine is metabolized insignificantly after injection into axons. After several hours some 94% of the original material is still in the form of ['4C]glycine and very few of the emerging counts are acid labile. On the other hand, glutamate is metabolized significantly by the giant axon. One hour after injection of ['4C]glutaTABLE 111 METABOLISM OF SUGARS R Y GIANTA x o ~ OF s Loligo for.hesi*,h

96 Total activity as unchanged sugars % Total activity

Sugar

D-Glucose 2-Deoxy-~-glucose 3-0-Methylglucose a-Methyl-D-glucopyranoside

Axoplasm 58.6 19.2' 99.2 99

Effluent

55

as I4CO2 45

-

-

100

0 0

I00

From Baker and Carruthers (1981a). Temperature, 20°C. Exposure of axons to labeled sugar was for 3 hr with one exception. Exception where the duration of exposure to isotope was 20 min.

TRANSPORT

OF SUGARS AND AMINO ACIDS

97

mate, 89% of the injected radioactivity is recoverable as glutamate and the rest is distributed between at least 15 different metabolites of which aspartate, betaine, and alanine are most prominent. After exposure of injected axoplasm to a longitudinal electric field (Fig. 2), both mobile and immobile fractions of the radioactivity contain glutamate in approximately the same proportion. After injection of L-[ ''C]glutamate, roughly 0.02% of the injected radioactivity is lost each minute. Virtually all (9798%) of the 14C emerging from ~-[~~C]glutamate-injected fibers is acid labile and, presumably, 14C02.This is perhaps not surprising because glutamate and its derivatives may enter the citric acid cycle. Figure 3 shows that 14C02 exit from ~-['~C]glutamate-injectedfibers is strongly inhibited by cooling and by exposure to the respiratory inhibitor cyanide. The remaining 2-3% of the activity emerging from axons is almost entirely (90%) ~-[~~C]glutamate. The metabolism of other amino acids in squid axoplasm has not been investigated.

K

cholina

-

lOmM gluto

2 mM cyanide

l0C

0.0003

E

-t

0.0001

4 u

0

2

--

-

0

.0.

*'

0

?

0

0

ct,

~

c

o

w

I

I

I

0

1

2

o

o

o

3

w 00 4

TIME

-

. .. 0

0.0001

0

cg

0

0.

woo 00

I

I

5

6

I 7

00

I 8

Ihours)

FIG.3 . Efflux of I4Cfrom an axon of Loli,qo,forbrsi previously injected with ~-['~C]glutamate. Ordinate: fraction of total radioactivity lost per minute; abscissa: time. 0 , Total radioactivity; 0, radioactivity remaining after removal of the acid-labile (CO?)fraction. The bulk of non-acid-labile radioactivity is glutamate. Axon diameter, 924 fim; temperature, 20°C. (Unpublished data of Baker and Carruthers.)

98

P. F. BAKER AND A. CARRUTHERS

111.

TRANSPORT

OF SUGARS BY SQUID AXONS

A. Kinetics of Transfer

1 . FACILITATED TRANSFER Certain sugars penetrate the giant axons of the squid at rates several orders of magnitude greater than would be predicted for the free diffusion of sugars through a lipid bilayer. Thus, D-glucose, 2-deoxy-~-glucose, 3-O-methylglucose, and a-methyl-D-glucopyranoside are taken up rapidly by the giant axon, whereas mannitol, sucrose, and L-glucose penetrate the axon only very slowly (Hoskin and Rosenberg, 1965; Baker and Carruthers, 1981a). These findings demonstrate that squid axons possess a selective transfer process for sugars. The selectivity of this process is presumably a reflection of specific stereochemical requirements for transfer and, by analogy with other sugar transport systems, transfer is likely to be mediated by specific transport proteins that catalyze the rapid transbilayer flux of sugars (Jones and Nickson, 1981).

2. SUGAR TRANSPORT IS PASSIVE Two types of sugar transport have been described in biological tissues. In most cell types, hexose transfer is mediated by a passive mechanism. Here both unidirectional influx and efflux of sugar may be observed, but the direction of trans-membrane net sugar transport is always from high to low sugar concentration. This process proceeds until the concentration of sugar on both sides of the membrane is equal. Thus, transported sugar is not accumulated by cells in excess of the extracellular sugar concentration. The second type of hexose transfer is described as active. This is typical of transport in mucosal membranes of epithelial cells (Crane, 1977). Here net sugar transport may proceed uphill or from low to high sugar concentration and cells may accumulate sugar in excess of the extracellular sugar concentration. Such active hexose transfer is normally linked to or supported by the downhill influx of specific cations into the cell. Hexose transfer in the squid giant axon is mediated predominantly by a passive transport system (Baker and Carruthers, 1981a). Figure 4 shows that, at equilibrium, the 3-0-methylglucose space of an axon does not exceed the water content of the axoplasm and intracellular sugar does not exceed the extracellular sugar concentration. One possible exception to this is the transport of D-glucose at sugar concentrations in excess of physiological blood glucose levels. Here one component of glucose uptake requires the presence of extracellular Na+ ions (see Fig. 5 ) and as

TRANSPORT OF SUGARS AND AMINO ACIDS

99

& "'CI,

6

0

12

Time (hr)

FIG.4. Time course of 3-0-methylglucose uptake in axons of Loligoforbesi. Uptake of isotope from NaASW containing 1 mM 3-0-methylglucose was measured using the internal scintillator probe. The results of this analysis are shown by the filled circles (axon diameter, 775 pm; temperature, 23.5"C). Data from conventional influx experiments are also shown (0)(temperature, 21.5"C). Of the fiber wet weight, 90% is water; this is shown by the dashed line. (From Baker and Carruthers. 19Xla.)

A

t

t

0

50 [Glucose], (mM1

FIG. 5. (A) Comparison of the uptake of D-glucose in sodium artificial seawater (NaASW) and Na-free ASW. Axons were soaked for 20 min at 21°C in NaASW (0)or choline ASW (0)containing isotopic sugar. Four or more axons were used for each point. Glucose uptake in NaASW shows a clear discontinuity between 10 and 15 m M D-glucose. Glucose uptake in choline ASW can be fitted by a section of a rectangular hyperbola with apparent K , 3.6 mM and V,,, 11.4 pmol cm-? sec-I. ( B ) Dependence of glucose uptake on external Na concentration. External glucose concentration, 20 m M . Sodium was replaced isomotically by choline. Each point represents four fibers. The curve is drawn by eye. Temperature, 22.5"C. (From Baker and Carruthers, 1981a.)

P. F. BAKER AND A. CARRUTHERS

100

such, may be analogous to active, Na-dependent glucose transport (Baker and Carruthers, 1981a). Definite proof, the demonstration of accumulation of D-glucose in excess of extracellular glucose, however, is lacking. 3. F A C I L I T A TSUGAR ~D TRANSPORT Is A SATURABLE PROCESS

The concentration dependence of facilitated sugar transport in squid axons may, as a first approximation, be described by Michaelis-Menten kinetics. Figures 6 and 7 show that the rates of hexose influx and efflux increase monotonically at low sugar concentrations but begin to saturate as cxtra- and intracellular sugar concentrations are raised. Two types of kinetic constant may be derived from such studies: V,,,,, which is the maximum theoretical rate of transport; and the apparent K , , which is the concentration of sugar at which the rate of hexose transfer is V,,,/2. V,, is related to both the number of membrane transport sites and the catalytic rate constant for transport. The apparent K , is a measure of the

L

m m v)

1

I

I

0

I

1 20

[Sugar], (mM)

FIG.6. Uptake of analogs of D-glucose. Ordinate: influx in pmol cm-* sec-'. Abscissa: concentration in (mM). Axons were exposed to sodium artificial seawater containing sugar isotopes for 20 min at 21 ? 1°C. The number of fibers per point was four. All smooth curves are sections of rectangular hyperbolas calculated by Lineweaver-Burk analysis. The con, , 0.73 mM: V,,,, , 5.80 pmol cm ? sec '; 3 - 0 stants derived are 2-deoxy-D-glucose (01K,,, methylglucose ( 0 ) K . , , I .8 mM; V,,,;,,, 2.33 pmol crn ~2 sec I; a-methyl-D-glucopyr;inoside (u), Kn,,28.6 mM: v,,;,,, 3.15 pmol cm sec ~ I?'he . appropriate D-gkoSe influx points (m) are included for comparison. (From Baker and Carruthers, 1081a.)

101

TRANSPORT OF SUGARS AND AMINO ACIDS

affinity of the transport system for sugar. It is clear from Figs. 6 and 7 that 3-0-methylglucose transport in squid axons is mediated by an asymmetric transfer process where the V , and K , for zero-trans exit at 15°C (see Table 1V) are some fourfold greater than V , and K , for influx. This asymmetry is discussed in further detail in Section IV,A. 4. COMPETITION BETWEEN SUGARS FOR TRANSPORT

L-Glucose, P-methyl-D-glucopyranoside, glucuronic acid, fucose, mannitol, and sucrose are without effect on D-glucose uptake in squid axons. However, 3-0-methylglucose and a-methyl-D-glucopyranoside depress D-glucose uptake. 3-0-Methylglucose increases the K , for D-glucose uptake but is without effect on the V,,, for influx (Baker and Carruthers, 1981a). These findings are consistent with competitive inhibition of Dglucose influx and argue that D-glucose and 3-O-methylglucose compete for a common transport site at the exterior of the axon. Sugar efflux is also A

8

3

0

50

0

50

I3 OMGli ( m M )

FIG.7. Concentration dependence of 3-0-methylglucose exit from dialyzed axons of Loligoforbesi into sugar-free seawater. (A) Exit at 15°C. Ordinate: 3-0-methylglucose exit (pmol cm-2 sec-I). Abscissa: internal 3-0-methylglucose concentration ( m M ) .The smooth curve is a rectangular hyperbola with apparent K , 5.5 mM and V,,, 8.6 pmol cm-2 sec-'. The mean axon diameter is 694 pm; the mean E,,,, -53.2 mV. ( B ) Exit at 21°C. Ordinate: 30-methylglucose efflux. Abscissa: internal 3-0-methylglucose concentration. The curve is a rectangular hyperbola with apparent K , 15.6 mM and V,,, 20.2 pmol sec-'. The mean temperature is 21 2 1°C; the mean axon diameter, 755 pm; the mean E M ,-52.5 mV. The number of measurements is shown above the points. (From Baker and Carruthers, 198tb.)

102

P. F. BAKER AND A. CARRUTHERS

reduced by the presence of other sugar types at the interior of the axon. 30-Methylglucose efflux is inhibited reversibly by intracellular D-glucose, 2-deoxy-~-glucose,or a-methyl-D-glucopyranoside (Baker and Carruthers, 1981b). The apparent Ki for the inhibition of 3-0-methylglucose exit by 2-deoxy-~-glucoseand D-glucose are 7 and 12 mM, respectively. Comparison of these K ivalues with the apparent K , for influx of the sugars (0.7 and 3.6 mM) provides further evidence in support of the view that hexose transfer in the squid axon is asymmetric. 5 . INHIBITION OF SUGAR TRANSPORT BY CYTOCHALASIN B AND PHLORETI N

3-0-Methylglucose efflux from axons loaded with radiolabeled sugar by axial microinjection is inhibited by relatively low concentrations of phloretin or cytochalasin B. These agents are potent inhibitors of sugar transport in other cell types (Bloch, 1973; Clausen, 1975; Carruthers, 1983a,b), inhibiting sugar transport half-maximally at submicromolar concentrations. Hexose transfer in squid axons, however, is less sensitive to these inhibitors. The apparent Ki for inhibition of 3-0-methylglucose exit by cytochalasin B and phloretin are 0.1 and 0.2 mM, respectively. Furthermore, phloretin inhibits exit irreversibly in squid axons yet acts reversibly in mammalian cells (Widdas, 1980). 6. CONCLUSIONS The basic properties of hexose transport in squid axons are similar to those observed in most other animal cells (for review see Elbrink and Bihler, 1975; Carruthers, 1983b). Transport is facilitated, passive, selective, and saturable. These characteristics establish that, during the process of hexose transfer, sugars interact with a limited number of selective transport sites that are accessible at each surface of the membrane. 8. Regulation of Sugar Transport in Squid Axons

1. EFFECTOF ELECTRICAL ACTIVITY

A principal function of the giant axon is to conduct and transmit information to the mantle muscle. This may result in the axon alternating between periods of relative quiescence and periods of activity. During these active periods the giant axon gains Na+ ions and loses K. When these conditions are mimicked to an extreme degree by repetitive electrical stimulation of isolated axons, the uptake of D-glucose (and 2-deoxy-~glucose) is increased (Baker and Carruthers, 1981a); but uptake of the

TRANSPORT OF SUGARS AND AMINO ACIDS

103

nonmetabolizable sugar 3-O-methylglucose is unaffected. This suggests that the observed increase in uptake of D-glucose and 2-deoxy-~-glucose may be linked to the metabolism of hexose, not to a direct effect on the transport system. Ouabain, which inhibits Na-pump activity, greatly reduces the effect of electrical stimulation on glucose uptake, suggesting that the increased sugar uptake during stimulation is associated with enhanced Na-pump activity. Simultaneous determinations of sugar uptake and metabolism by stimulated axons show that there is indeed an increase in the metabolism of D-glucose and 2-deoxy-~-glucosein “active” axons (Baker and Carruthers, 1981a) and that this effect too is abolished by ouabain. It is possible that this increase in net sugar uptake is the result of a more powerful metabolic sink for axoplasmic sugar. A parallel may be drawn between these studies and experiments with mammalian neurons. Radiolabeled 2-deoxy-~-glucosehas become a popular tool as an intracellular marker of increased neuronal activity in the brain (Sokoloff er af., 1977). The labeled sugar is infused into a cranial artery, and later its intracranial position is located by gross or microautoradiography . As 2-deoxy-~-glucoseis metabolized to 2-deoxy-~glucose-6-phosphate after transport into the neuron it becomes essentially trapped within the cytosol. Those neurons in regions of the brain associated with increased activity appear to be more heavily labeled with 2deoxy-~-glucose-6-phosphatethan neurons in less active regions. It is not known, however, whether this is due to altered transport and subsequent metabolism of the sugar by active neurons or whether alterations in metabolism alone are responsible. The squid giant axon data support the view that increased metabolism rather than transport of sugar is responsible for this effect. 2. METABOLIC DEPLETION A N D SUGAR TRANSPORT Studies of sugar transport in synaptosomes from rat cerebral cortex and brain slices have shown that the metabolic poisons cyanide, 2,4-dinitrophenol, and iodoacetate inhibit 2-deoxy-~-glucoseuptake (Diamond and Fishman, 1973) and the hormone insulin is without effect on sugar uptake (Elbrink and Bihler, 1975). The mechanism by which metabolic depletion inhibits hexose uptake in synaptosomes is not known and stands in contrast with the stimulation of hexose transfer in muscle under comparable conditions and lack of effect of depletion on transfer in red cells, hepatocytes, and other cell types (Elbrink and Bihler, 1975; Clausen, 1975). Glucose uptake in squid axons is inhibited by the metabolic poison cyanide and is unaffected by insulin (Baker and Carruthers, 1981a). Cyanide also inhibits reversibly a-methyl-o-glucopyranoside and 3-O-methyl-

m

0

t

D Control

2 mMNaCN

L

1

0

20

[3OMGI, (mM) FIG.8. Effect of metabolic poisoning on 3-0-methylglucose efflux. (A) External 3-0-methylglucose concentrations are shown above the points. Cyanide (2 m M ) was applied halfway through the experiment. Temperature, 19°C; axon diameter, 770 prn. (B) Replot of the data from (A) showing the reduction in sugar exit as a function of external 3-0-methylglucose concentration before and during application of cyanide. Ordinate: reduction in sugar exit (5%); abscissa: external sugar concentration ( m M ) . The curves are rectangular hyperbolas with constants: control. Ki, 2.23 m M ; I,,, , 74.4%; after exposure to 2 m M cyanide for 2 hr, K i , 6.2 rnM, I,,,. 7%. (From Baker and Carruthers. 19Xla.)

TRANSPORT OF SUGARS AND AMINO ACIDS

105

glucose efflux in intact axons. This effect may arise from a reduction in the affinity of the transport system for sugar. One way to examine this possibility is to utilize the fact that efflux of 3-0-methylglucose from axons is reduced when sugar is present in the external medium. This inhibition of exit is dependent on the external sugar concentration and shows saturation kinetics (see Fig. 8). The concentration of sugar that reduces sugar exit half-maximally is similar to the apparent K , for sugar influx (see Section IV,A). Exposure of intact axons to cyanide increases the concentration of external sugar required to inhibit 3-0-methylglucose half-maximally (Baker and Carruthers, 1981a). Exposure to cyanide reduces the ATP content of the cytosol (Caldwell, 1960), increases the cytoplasmic ionized Ca (Baker et al., 1971), and very slowly brings about an increase in the Na and fall in the K content of axoplasm through inhibition of the Na pump. Any of these changes may account for the inhibition of hexose transfer in the poisoned axon. These possibilities may be tested directly in the squid giant axon, using the technique of intracellular dialysis (see Fig. 9, and Brinley and Mullins, 1967). With this method it is possible to control the solute composition of the cytosol while simultaneously measuring the permeability of the plasma membrane to sugar (Carruthers, 1979; Baker and Carruthers, 1981b). The results of such experiments show the following:

1. 3-0-Methylglucose exit is unaffected by altering cytosolic ionized Ca (with Ca-EGTA buffers) over the range 0.04-0.9 pM. 2. Increasing internal Na from 40 to 70 mM is without effect on transport. 3. Physiological cytosolic levels of glucose-6-phosphate, glucose-lphosphate, fructose-6-phosphate, and lactate are without effect on transport. 4. AMP, ADP, GDP, and physiological levels of cAMP are without effect on sugar transport. Very high concentrations of cAMP produce some inhibition (Fig. I I ) . 5. Depletion of cytosolic ATP decreases the rate of 3-0-methylglucose efflux, and the K , of the external site for 3-0-methylglucose is increased (i.e., affinity is reduced) from 0.72 to 3.2 mM (see Fig. 10, and Baker and Carruthers, 1981b). This effect of ATP depletion on 3-0-methylglucose exit from dialyzed axons is quantitatively similar to the effect of cyanide on exit from intact axons. Its effect can be reversed by including ATP in the dialysis solution, and the effect of ATP is mimicked by the hydrolyzable ATP analog cu,,B-methylene-5-ATP,but not by the nonhydrolyzable ATP analog P,y-methylene-5-ATP (Fig. 11, and Baker and Carruthers,

106

P.

A

Guard

V. elmrode

In

F. BAKER AND A. CARRUTHERS

Guard

-r,.-;--

_i

--

._

I

Dialysis capiilary

Flow

7 r - -Glass cannula

FIG. 9. Experimental arrangement for internal dialysis. (A) Efflux. A single dialysis capillary is inserted into the fiber via the right-hand cannula and an Em recording electrode via the left cannula. The porous section of the dialysis tubing is shown by the dashed lines. The fiber is superfused with seawater, and a guard system is operated to collect isotope entering the central compartment from the less well dialyzed end regions of the fiber. Fluid was withdrawn from the central compartment to maintain the fluid level constant. (B) Influx. Schematic view of the double-barreled dialysis capillary in position inside an axon (not to scale). Both capillaries are mounted in a Perspex block, and separate flow to each capillary is delivered by motor-driven syringes. The porous section of each capillary is shown by the dashed lines. When phenol red is added to the solution flowing through the collection capillary, a sharp cutoff in the coloration of axoplasm is seen on either side of the porous region of the collection capillary. When the guard capillary flow is halted, the coloration spreads beyond the region of axoplasm dialyzed by the collection capillary. (From Baker and Carruthers, 1981b.)

1981b). The inference is that hydrolysis of the y-phosphodiester bond of ATP leads, by an as yet undetermined mechanism, to an increase in the affinity of the hexose transfer system for its substrate, sugar. It is conceivable that this may occur through y-phosphoryl transfer from ATP to an allosteric site on the transport protein(s).

107

TRANSPORT OF SUGARS AND AMINO ACIDS

B

0

c

0

P -c

0

F -

1

0

20

( 3 OMGI, ImM)

FIG. 10. Effect of external 3-0-methylglucose on 3-U-methylglucose exit in a fiber dialyzed with medium containing 4 m M and zero ATP. (A) Ordinate: 3-0-methylglucose efflux in pmol cm-* sec - I . Abscissa: time in hours. The concentrations of 3-0-methylglucose and ATP in the dialysate and external 3-Omethylglucose are shown above the points. The fiber was dialyzed with zero-ATP medium for 1 hr before the radioactive sugar was added. Temperature, 14°C; axon diameter, 760 pm; EM at t = 0 hr, -53 mV; EMat t = 13 hr, -37 mV. (B) Redrawn from (A). Ordinate: percentage of reduction in 3-0-methylglucose exit by extracellular sugar. Abscissa: external 3-0-methylglucose concentration (mM). Both curves are rectangular hyperbolas with the following constants; 4 m M ATP in dialyzate (0):apparent K , for inhibition of exit by external sugars, 0.72 mM; maximum inhibition, 54.3%. Zero ATP in dialysate (@): apparent K , for inhibition of exit by external sugar, 3.22 mM; maximum inhibition, SS.S%. (From Baker and Carruthers. 19Rlb.)

108

P. F. BAKER AND A. CARRUTHERS

ATPi ( m M ) cAMPi ( m M ) a,P-MeATPi ImM) 0, y-MeATPi ( m M )

0 0 0

1

I

4

0 4

1 1 1

4

2

1

0 0

0

1

0

1

I

0

n o

L

*O O

I

1 6

1

Time Ihr)

FIG. 11. Effects of analogs of ATP on 3-O-methylglucose exit. Ordinate: 3-O-methylglucose efflux in pmol cm-* sec I . Abscissa: time in hours. The fiber was dialyzed with ATPfree medium containing 5 mM 3-O-methylglucose for 1 hr before the isotope was added. The various analogs of ATP and other nucleotides were added to the internal solution at different times during the experiment (see key above points). Temperature, 15°C; axon diameter, 690 wrn. (From Baker and Carruthers, 1981b.)

3 . EFFECTSOF pH

ON

SUGARTRANSPORT

Changing external pH from 5 to 10 is without effect on the K,, and V,,, for sugar uptake in squid axons (Baker and Carruthers, 1981a), demonstrating that the external site is relatively insensitive to H + ions. Reducing internal pH in intact axons by exposing fibers to seawater equilibrated with 5% C02 (Caldwell, 1958; Boron and de Weer, 1976) rapidly and reversibly reduces the rate of sugar exit (Baker and Carruthers, 1981a). Similarly, internal acidification of dialyzed fibers (pH 7 to 6.2) reduces sugar exit reversibly (Baker and Carruthers, 1981 b). These results provide further evidence for asymmetry of sugar transfer in the giant axon, the internal site being more sensitive to protons than the external site. This sensitivity to protons may, however, be rather more subtle than a simple interaction of the internal sugar binding site with H + ions. Protons may affect the interaction of the transporter with ATP or may interact at a second allosteric site. The internal pH dependence of the ATP sensitivity of hexose transfer must be determined to clarify these possibilities.

TRANSPORT OF SUGARS AND AMINO ACIDS

109

4. TEMPERATURE DEPENDENCE OF HEXOSE TRANSFER The kinetics of sugar transport in the giant axon are extremely sensitive to alterations in temperature. V,,, for D-glucose influx rises from 8.8 pmol sec-I at 0°C to approximately 16 pmol cmP2sec-I at 143°C. These results are consistent with an activation energy for D-glucose influx of 9.5 kcalhol. Beyond 15°C however, the V,,, for D-glucose influx falls, and at 21°C it is 11.2 pmol cm-2 sec-I. These findings are not dissimilar to the heat denaturation of enzyme systems. The apparent K , for D-glucose influx falls from 8.9 mM at 0°C to 3.6 mM at 21°C. 3-0-Methylglucose exit is rather more sensitive than influx to alterations in temperature. The activation energy for exit is 23 kcal/ mol. Furthermore, the exit rate continues to increase between 15 and 23°C. In contrast to the K , for influx, the K , for zero-trans exit of 3-0methylglucose increases from 5.5 mM at 15°C to 16 mM at 21°C (Baker and Carruthers, 1981a,b). These results imply that as temperature is raised the asymmetry of hexose transfer ( K , efflux/K, influx and V , efflux/V, influx) increases. A number of points arise from these studies. The lack of any obvious break in the Arrhenius plots of sugar exit suggests that, in keeping with most eukaryotic plasma membranes, there appears to be no gross bilayer phase transition. This is consistent with the high cholesterol content of the axolernma (45%), which acts to fluidize the bilayer at temperatures where the phospholipid acyl chains normally crystallize (Melchior, 1982).The apparent break in the Arrhenius plot of V,,, for uptake between 14.5 and 21°C is curious. A similar effect is not observed with exit, suggesting that more than one functional transport system may be present in the axolemma.

5 . CONCLUS~ONS

Sugar transport in the squid giant axon is regulated in a manner that may serve to maintain axoplasmic sugar levels. Compared to other tissues, D-glucose is metabolized only slowly by the giant axon (see Section 11). As such, under conditions where the metabolic requirements of the giant axon are increased, uptake of sugar will not be rate-limiting for metabolism. During increased neuronal activity the elevated rate of axoplasmic sugar phosphorylation ensures that an adequate net inflow of glucose may proceed. Similarly, under conditions where cytosolic ATP is reduced, the increase in the K , for sugar exit may serve to reduce the rate of loss of hexose from the axon to the exterior.

110

P. F. BAKER AND A. CARRUTHERS

IV. DESCRIPTIVE MODELS FOR SUGAR TRANSPORT A. Hexose Transfer in the Giant Axon is Asymmetric

1. GENERAL

A transport system may be termed asymmetric for a number of reasons. Here, the most fundamental use of this term is in its application to the basic kinetic properties of transport. Table IV opposite provides a rather complete kinetic analysis of 3-0-methylglucose transport in the giant axon of Loligo. The K , and V,, for zero-trans-3-0-methylglucose exit are some fourfold greater than the corresponding constants for influx. This is not unusual. Passive sugar transport in the human erythrocyte is also mediated by an asymmetric process in which the K , and V,, for sugar exit are some 10-fold greater than those for entry (Naftalin and Holman, 1977; Widdas, 1980). In other cell types [e.g., barnacle muscle, hepatocytes, avian erythrocytes, and adipocytes (for review see Carruthers, 1983a,b)] passive hexose transfer is symmetric. The asymmetry of transport does not imply that the giant axon will lose sugar at rates faster than the rate of uptake of sugar. Examination of Table IV shows that the kinetics of sugar exit are markedly affected by the presence of sugar in the extracellular solution. As discussed earlier, extracellular sugar inhibits the rate of sugar exit, but uptake is unaffected by intracellular sugar. As extracellular sugar is raised, K , is unaffected, but the V,,, for exit falls progressively until it is identical to V,, for influx. Equilibrium-exchange exit and uptake are, as is the case with all passive transport systems, identical and are not significantly different to zerotrans uptake. Here, transport in the giant axon differs significantly from hexose transfer in the human red cell. In the erythrocyte, transport is accelerated by the presence of sugar at the opposite, trans, side of the membrane. Both uptake and exit of sugar are affected in this way. The origin of hexose transfer asymmetry in the giant axon is discussed in Section 1V,B,2. 2.

ASYMMETRIC

RESPONSESTO pH A N D TEMPERATURE

As mentioned above, sugar transport in the giant axon is insensitive to external pH in the range 5 to 10, yet is inhibited by internal acidification. This provides direct evidence for the chemical asymmetry of sugar transport. The asymmetric behavior of hexose transfer is extremely sensitive to changes in temperature. If the temperature sensitivity of V,,, for glucose

TABLE IV: KINETICSOF

3-0-METHYLGLUCOSE

TRANSPORT IN

THE

DIALYZED SQUIDAXON ~~

Conditions Type of experiment

Blood (sugar)

Zero-trans fluxes Uptake

Exit Infinite-trans fluxes

4

Axoplasm (sugar)

J

4 I

varied

zero

Characterized by

Measured valuesa

K , uptake ( K R , ) V,,, uptake (V$!+i)

1.34 mM 1.99 pmol cm-* sec-l

c

zero

$+-q

5.5 mM 8.6 pmol c m - ? sec-'

I

Uptake

K, external ( K ; + , ) V,,, exchange Wee)

1.34 mM 1.99 pmol cm-' sec-'

Exit

K , internal (K7-J V,,, exchange ( V e e )

2.1 pmol

Uptake

K, internal (Kz',,) V,,, net entry (V;t,l)

6.0 mM 2.2 pmol c m 2 sec-l

Exit

K, external (Kf-J Vmx net exit (V;!+J

1.34 mM 8.6 pmol c m 2 sec-'

K, exchange (K")

1.39 mM 2.0 pmol cm-'sec-l

5.5 mM

sec-l

Infinite-cis fluxesb

Equilibrium exchange Internal (sugar) = external (sugar) uptake and exit

I I

a

V,,, exchange ( V e e )

Obtained by Lineweaver-Burk analysis of experimental data. Infinite-cis net fluxes were measured as the difference between unidirectional uptake and exit.

112

P. F. BAKER AND A. CARRUTHERS

uptake and 3-O-methylglucose exit in intact axons is compared, it is apparent that the Q15for glucose uptake between 0 and 15°C is 1.9 and that for 3-O-methylglucose exit over the same temperature range is 7.8 (Baker and Carruthers, 1981a). These data suggest strongly that the asymmetry between V,, for exit and uptake is reduced as temperature is lowered. Consistent with this view is the finding that the asymmetry between V,,, for exit and uptake at 21°C is 9 and at 15°C i s 4.4 (Baker and Carruthers, 1981b). Stated more plainly, sugar exit is more sensitive to temperature change than sugar entry. This is consistent with a greater energy barrier for the catalysis of exit than for influx. The observed increase in K , for glucose uptake and decrease in K , for 3-O-methylglucose exit as temperature is lowered from 20 to 0°C suggests that a reversal in the asymmetry of transfer may occur with reduced temperature. Proof of this thesis requires the full kinetic analysis of 3-O-methylglucose exit and entry over this range of temperatures. This sensitivity to temperature is rather different from that reported for hexose transfer in the human erythrocyte. Here the Qzofor influx between 0 and 20°C is 170 (Lacko et a / . , 1972), whereas the Qzofor exit between 7 and 17°C is 4 (Sen and Widdas, 1962).The high Qzofor influx is unlikely to arise from membrane crystallization at lower temperatures, for membrane cholesterol (40% of membrane lipid) suppresses bilayer phase transitions and behaves as a membrane fluidizer-plasticizer in erythrocyte membranes between - S and 50°C (Carruthers and Melchior, 1983b). These results suggest that the asymmetry between V,:,, for exit and entry in the erythrocyte increases as temperature is lowcred-quite the revzrse of the observations made with squid axons.

B. Mechanisms for Hexose Transfer in the Giant Axon

1. GENERAL When the plasma membrane shows a selective (and often very high) permeability to a particular substrate, this property is frequently attributed to the presence of specific carriers in the membrane with which the substrate combines to effect entry. Figure 12 illustrates two simple carrier models-the mobile carrier and the two-component carrier. The steadystate kinetic solutions for thesc carrier types have been reviewed in detail by Bowyer and Widdas (1958), Britton (1964), G. F. Baker and Widdas (1973), Lieb and Stein (1974), and, rather more concisely, by Baker and Carruthers (1981b).

TRANSPORT

OF SUGARS AND AMINO ACIDS A

113

Mobile carrier Membrane 0

Q

2

B

BiDolar carrier M

M

M

FIG.12. Models for the transport of sugars in the squid axon. (A) The mobile carrier. X is carrier, P is sugar on side 1. Q is sugar on side 2; a through h are rate constants for the appropriate reactions. (9)The two-component carrier. The carrier consists of two halves, each of which can bind sugar with a characteristic dissociation constant. When sugar binds, a conformation change occurs, permitting sugar to exchange between each half of the carrier in a central, intramembranous pool. The carrier then undergoes further conformational change releasing the sugar at the opposite side of the membrane. For simplicity, the carrier is shown operating in one direction only. (From Baker and Carruthers, 1981b.)

2. O R ~ G IOF N HEXOSETRANSFER ASYMMETRY Before any theoretical kinetic description of hexose transfer in squid axons may be formulated, it is necessary to determine whether the operational kinetics of transfer arise from factors intrinsic or extrinsic to the transport system. The most important data to consider are the high internal K,,, for exit and the inhibition of unidirectional exit by external sugar.

114

P.

F. BAKER AND A. CARRUTHERS

Here the following criticisms may be pertinent: (1) The high K , for exit and inhibition of exit by trans-sugar may result from an unstirred layer effect in the microenvironment of the inner surface of the bilayer; (2) the inhibition of exit may arise from a long-pore effect (Hodgkin and Keynes, 1955). The presence of an unstirred layer at the internal face of the membrane can be discounted for the following reasons.

1. The trans effect of external sugar persists in fibers dialyzed with internal sugar concentrations as high as 100 mM. 2. In Na seawater, external D-glucose promotes a sustained increase in the rate of 3-0-methylglucose exit, whereas in Na-free seawater glucose reduces exit (Baker and Carruthers, 1981b). As D-glucose uptake in Na seawater is greater than in choline (Na-free) seawater (see above), unstirred layer effects should be greater when the effects of trans-glucose are measured in Na seawater. 3 . The kinetics of inhibition of radiolabeled sugar exit by external unlabeled sugar should resemble competitive inhibition. The presence of an unstirred layer should lead to an increase in unlabeled sugar concentration below the membrane due to sugar influx. This cold sugar would then compete with labeled sugar for exit. As such, we would expect external sugar to increase the K , for exit but leave V,,, unchanged. This is not the case. V,,, for exit falls with increasing trans sugar but K , remains unchanged. These data are more consistent with noncompetitive inhibition. 4. The presence of unstirred layers should be detected by a plot of bulk solution sugar concentration/flux vs bulk solution sugar concentration (Lieb and Stein, 1974). Such transformations do not reveal the presence of unstirred layers. A long pore effect, if it existed, should be observed with both sugar influx and efflux; but entry is unaffected by intracellular sugar. A further pertinent observation is that inhibition of sugar exit by trans sugar is observed in dialyzed fibers where transmembrane solute asymmetry und symmetry are imposed (i.e., external and cytosolic solute concentrations are either nonidentical or identical) Baker and Carruthers, 1% 1 b’ These observations support the view that, unlike human erythrocyte hexose transfer (Carruthers and Melchior, 1983a), the observed opcrational kinetics of hexose transfer in the squid axon are a reflection of thc intrinsic activity of the transporters and are not the result of extrinsic modulation of transport.

115

TRANSPORT OF SUGARS AND AMINO ACIDS

3. THE MOBILECARRIER Tables V and VI summarize the steady-state solutions to the mobilecarrier flux equations. The kinetics (Lieb and Stein, 1974; Baker and Carruthers, 1981b) are formulated in relation to a number of resistance terms in a form used extensively by Geck (1971). The resistance terms may be calculated for 3-0-methylglucose transport in the giant axon (Table V) and applied to the derived flux equations. With the mobile carrier model we assume that each transfer protein may bind only one sugar molecule at a given time. Figure 12, which illustrates this model, implies that a single complex translocates the bilayer either loaded with sugar or empty. However, as with other enzymecatalyzed reactions, the kinetic equations ignore all physical transformations. In other words, no particular degree of translocational carrier movement through the membrane is assumed in a kinetic analysis of carrier-mediated transport, although illustrative models have sometimes given the impression that there is. With the mobile carrier there are eight rate constants to consider (see Fig. 12). If we assume that binding steps are rapid with respect to trans; membrane flux we may reduce the number of “significant” rate constants to four, namely, c , d , g , and h. If the rate of return of loaded carriers from outside to inside is slower than the rate of return of empty carrier to the TABLE V STEADY- STATE SOLUTI ONS FOR FHE

MOBILECARRIER

Re,

=

R12

f

Rzi

~

R,,,

n R I 2= lle + llh + lIc(u + d)/4 nR2, = lib + llg + Ild(h + <‘)lh wR, = Ilg + Ilh nRre = Ilb + I/(> t I/c(e t d)lc + lld(h + c)lh Kl? = hg/lr[l/b + I/(,((> + d)/e] Kzi = chlf” Ilr + Ild(h + (‘)ihl Constraint: aceh

=

bdfg

3-0-Methylglucose transport (15°C) RT2= 0.116 ern? sec pmol R21= 0.5 cm2 sec pmol R,, = 0.116 cmZsec pmol I Re, = 0.5 cm? sec pmol I K I z = 5.6 mM K 2 , = 5.78 mM

116

P. F. BAKER AND A. CARRUTHERS

TABLE V1 MOBILECARRIER F w x EQUATIONS"

a For fluxes in the direction 2-1 simply reverse the symbols I and 2. P is the concentration of sugar on side 1 of the membrane, and Q is the level of wgar at side 2 Table IV gives the meaning of symbols V f ' . , ,

etc.

inside (i.e., J , h ) , then, provided loaded and empty carrier move equally rapidly from inside to outside, but at the same rate as unloaded carrier returns to the inside (i.e., c = h = g = d ) , we may make the following predictions: ( I ) Extracellular sugar inhibits sugar exit. (2) Intracellular sugar is without effect on entry. (3) Equilibrium-exchange kinetics are identical to zero-trans influx. (4)External sugar is without effect on the K , for sugar exit, Furthermore, (5) external sugar will produce a redistribution of carrier between inside and outside. The first four predictions have been observed experimentally (Baker and Carruthers, 1981b). The results of the fifth prediction may be assessed using Eqs. (12), (13), (20), and (21) of Widdas (1952). The data presented here suggest that, in the absence of external sugar, carrier is equally distributed between both sides of the membrane. When sugar is present at saturating levels at both sides of the membrane, however, seven-eighths of the carrier must be at the outer face of the membrane and only one-eighth at the inner face. 4. REJECTIONCRITERIA

FOR T H E

ASYMMETRIC MOBILECARRIER

Hankin et al. (1972) have developed unambiguous rejection criteria for the mobile carrier. Specifically, these criteria were developed for simple

117

TRANSPORT OF SUGARS AND AMINO ACIDS

TABLE VII REJECTION CRITERIA FOR THE ASYMMETRIC MOBILECARRIER Asymmetry factor W

First rejection criterion W = lib llc

+

+

Ild Ile

;LZ -K=

+

+

llg Ilh

Ilb

+ clhd. ~K"

+ dlce' Kl',?

+

K"

llg

-

Ilc

+

Ild + lie + clbd + IIe + Ilh + dlre

+

Ilc Ilc

+

Ilh

+ Ilh + dlce' + dlce

Ile

Calculate W from this relationship. It must be similar to the asymmetry factor. W = 4.2. Second rejection criterion (W

+

1)K"

+ K;+?

=

0

Calculate W". If the asymmetry factor is significantly less than W", then the asymmetric carrier must be rejected. W"

=

3.1

application to transport systems that appear to be kinetically asymmetric, e.g., sugar transport in the squid giant axon. These criteria are listed in Table VII. Table VII shows that these criteria are satisfied. The data are, therefore, consistent with the asymmetric mobile-carrier model for transport.

5. THETWO-COMPONENT CARRIER

A model transport system with two sugar binding sites (existing simultaneously at opposite sides of the membrane) is also consistent with the data present here. Here, we assume that binding of sugar is rapid with respect to translocation. If K , and K2 are the apparent K,,, for exit and uptake, respectively, and V, and V2 velocity constants for exit and uptake, respectively, then in the absence of trans sugar uptake is given by &V2, where 1 +2

= 1

+ (K2")

P. F. BAKER AND A. CARRUTHERS

118

and exit by 4 ] V I , where

41 =

1 1

+ (K14Sli)

[S] is sugar concentration and i and o refer to inside and outside, respectively. When trans sugar is present, flux of sugar may occur through carrier with sugar bound at the cis site only or through carrier containing both cis and trans sugar. In the latter situation, the velocity constant may be altered to Vee.Here the fraction of total carrier available for uptake is given by $p#q or

Vee. Total unidirectional influx is, therefore, given by 42(1V2 + Net influx is given by influx - efflux and is 42(1V2 + ~ 4 ~V'" 4,lh(I- 42)V11+ 414?Vee or simply 42(1 - 41)V2 - 4dV1. If K I ,K 2 , V1, V 2 ,and Veeare given the values of K : L 2 , KY-,,, V : L 2 , V ? ! , and V e e , respectively, of Table VI, then the predictions of these equations are identical to those of the asymmetric mobile carrier. As such it is not possible to distinguish between either model for hexose transfer with currently available data. The methods suggested by Baker and Stone (1966) and Krupka and Deves (1981) could help to distinguish between these model types. Here fluxes are measured in the presence of two transport inhibitors, one acting at the outside and the other at the inside of the cell. The inhibition of transport mediated via the mobile-carrier model should be less than that calculated theoretically for the two-site transfer system. 1

C. Regulation of Transport by ATP

ATP appears to influence axolemmal hexose transfer by increasing the affinity of the transfer system for sugar. Baker and Carruthers (1981b) have proposed a scheme that can account for this effect. If K , , K 2 , and K3 are dissociation constants for interaction of the transporter with sugar in the absence of ATP, with ATP, and with sugar in the presence of excess ATP, respectively, then making the assumption that all reactions are rapid with respect to translocation, it can be shown that

where J is flux of sugar, n is a constant proportional to membrane trans-

119

TRANSPORT OF SUGARS AND AMINO ACIDS

porter density, V is a rate constant proportional to the rate of hexose transfer across the axolemma (nV = V,,,), and S is sugar. If K , > K 3 ,it is apparent that as ATP is increased the apparent K , for sugar uptake/exit approaches K3 but V,,, for transfer remains unchanged. Such interactions with ATP may be visualized as an ATP-induced conformational change in the transporter that increases the affinity for sugar while leaving translocation activity unaltered. V.

TRANSPORT OF AMINO ACIDS

A. Categories of Amino Acid Transport

Amino acid transport has been categorized on the basis of amino acid type (Heinz, 1972): the types include acidic, basic, and neutral amino acid transport systems. An analysis of the various categories of amino acid transport is lacking in the squid axon. Table VIII summarizes uptake rates for glycine and a number of other amino acids. TABLE VIlI AMINOACIDTRANSPORT I N THE GIANTAXON" ~~~

Amino acid Glycine Alanine Leucine Serine Proline Phen ylalanine Tyrosine Glutamateb Aspartate Arginine 6-Aminolevulinic acid' Glutamined Taurineb y-Aminobut yrated

~~~

Concentration (mM)

I 1 1

I 1 I 1 I 1 I 1 4.5 1 4.5

~

Uptake (fmol cm-* sec-') 500 440 500 400 400 I50 130 70 25 20 36 50 40 100

~~~~

N a dependence No

Yes Yes

Yes Yes

a Data are from Caldwell and Lea (1978) except as noted in footnotes b-d. For comparison, the maximum fluxes of D-glucose, 3-O-methylglucose, and 2-deoxy-D-glucose are 11.1, 2.33, and 5.8 pmol cm-? sec I , and at I m M . the fluxes of mannitol. L-glucose, and sucrose sec-', respectively. are 9, 7, and 10 fmol Unpublished observations of P. F. Baker, A . Carruthers, and S. J . Potashner. From Caldwell and Goldstuck (1979). " From Hoskin and Rosenberg (1966) and unpublished results of Baker and Carruthers. I

120

P. F. BAKER AND A. CARRUTHERS

Detailed knowledge of amino acid transfer in squid axons is limited to glycine and glutamate. Glycine transport is mediated by a facilitated, Naindependent system, whereas glutamate uptake is strongly Na dependent. Our discussion here will be limited to these two systems. 8. Glycine Transport

1. TRANSFER Is FACILITATED The permeability of the axolemma to glycine is approximately 0.7 x cm sec-I, which is some two to three orders of magnitude greater than the permeability of artificial bilayers to amino acids. Transport of glycine is, therefore, facilitated presumably by membrane proteins.

2. Is TRANSPORT PASSIVE? The axoplasm : blood ratio for glycine is 14.5 (Deffner, 1961). This constitutes a large net outwardly directed gradient for glycine. Net movement of glycine into the axon therefore requires energy input. However, it is not certain whether high axoplasmic glycine levels are maintained by glycine uptake or by intracellular glycine synthesis. Further, Rosenberg and Khairallan ( 1974) have argued that Deffner’s values for axoplasmic amino acid levels were too high owing to protein breakdown. Another complication is the inhibition of 70% of glycine uptake by the metabolic poison cyanide (Caldwell and Lea, 1978). These workers have argued that sensitivity to cyanide reflects a direct metabolic involvement in glycine uptake. However, Baker and Carruthers (1981a,b) have shown that the passive transfer of sugar in squid axons is affected by ATP depletion, but this inhibition does not indicate active transport. More work is required to clarify this point. A further factor that may be relevant is that glycine efflux is accelerated by a variety of extracellular amino acids, and it may be that glycine accumulation is brought about by exchange with other axoplasmic amino acids. 3. TRANSFER Is SATURABLE

Glycine uptake is mediated by a saturable process (Caldwell and Lea, 1978). However, there is good evidence to support the view that glycine uptake is mediated by more than one process. 1. The concentration dependence of uptake is not at all well approximated by simple Michaelis-Menten parameters (see Fig. 13). The data are more consistent with two parallel transfer systems each approximating to

121

TRANSPORT OF SUGARS AND AMINO ACIDS

c

I /

1

/

0

1

2

4

6

8

10

1

[Glycine]

(rnM-')

FIG. 13. (A) Glycine influx, measured by the paired axon method, plotted against external glycine concentration in artificial seawater. Each point is the mean from three or four axons; vertical lines are 2 SE of mean. Axon diameter is 470-813 pm. ( B ) A double reciprocal plot of the data in ( A ) . The dashed lines indicate only approximately the straight lines predicted for the two individual components of influx J , and J 2 . (From Caldwell and Lea, 1978.)

the Michaelis-Menten formalism. One process has high affinity (low K , ) and low V,,, for glycine transfer, and the other has low affinity (high K , ) and high V,,,. 2. Glycine uptake is inhibited (70%) by cyanide. 3. Glycine efflux is accelerated by extracellular glycine, but this trans acceleration of exit is unaffected by CN. These data suggest that CNsensitive and -insensitive uptake of glycine occur simultaneously. Definitive proof, however, requires the demonstration that trans acceleration of

122

P. F. BAKER AND A. CARRUTHERS

glycine exit is associated with a coupled influx of glycine in exchange for axoplasmic gl ycine. 4. COMPETITION FOR UPTAKE

Data concerning the competition between glycine and other amino acids for uptake are not available. 5 . EFFLUX

*

The fractional loss of l'4C]glycine from microinjected fibers is I . I 0.2 x lo-' min-I. Making the assumption that the cytosolic glycine concentration is 8.3 mM and that 87% of 14C efflux is in the form of [14C]glycine, Caldwell and Lea (1978) have calculated that this fractional loss is equivalent to an exit rate of 26 fmol cm-2 sec-I. This is considerably lower than entry rates. Glycine exit is accelerated by extracellular amino acids (Fig. 14). Table 1X summarizes the available data for this trans effect on exit. In addition to glycine, certain other neutral amino acids at 1 mM externally accelerate glycine exit to varying extents, but proline, taurine, acidic and basic amino acids have negligible effects. Glutamate, aspartate, and taurine uptakes into the giant axon are strongly Na-dependent 463 mM-Na+

463 m M ~ N a + 0-Na

z

I

"3

10

. E E

c

0 Ln

E

z

5

Time (min)

FIG.14. The effects of different external glycine concentrations on the glycine efflux in artificial seawater and zero Na+ (choline) seawater. An axon (diameter 800 p m ) was injected with ['4C]glycineat f = 0 min. Efflux of I4C label is plotted as a rate constant (fractional loss of counts per minute). Values for external glycine concentration were I 0.5 m M ; 2, I m M ; 3, 5 m M ; 4, 10 mM. (From Caldwell and Lea, 1978.)

123

TRANSPORT OF SUGARS AND AMINO ACIDS

and, on this basis, might not be expected to interact with Na-independent glycine transport. The acceleration of exit by trans-glycine is a saturable process. Acceleration is half-maximal at 0.5 mM external glycine, and the maximum acceleration of exit is 9.4-fold. The varying extent of acceleration of exit by other neutral amino acids at I mM is probably a reflection of the varying affinity of the exchange transfer system for these amino acids. A complete kinetic analysis of trans acceleration of glycine exit is required. 6. REGULATION

a . General. The effects of altered membrane potential, electrical activity, external and cytosolic pH, and temperature on glycine transport are not known.

TABLE IX STIMULATION OF G L Y C ~ N EFFLUX" E BY OTHERA M I N OA C ~ D S ~

Amino acid

Stimulated efflux a s percentage of glycine-stimulated efflux

Glycine L-C ysteine r-a-Alanine L-Serine L-Phenylalanine L-Leucine L-Isoleucine L-Tyrosine L-Threonine L-Valine L-Proline Taurine Glycylglycine GIyc ylglycylglycine L-Glutamic acid L-Aspartic acid r-Arginine

100.0 96.6 74.4 65.8 52.1 42.0 40.7 37.3 31.2 23.3 5.4 4.0 2.4 0.2 2.7 5.4 0

Stimulated efflux is the difference between the rate constant in artificial seawater (ASW) and that in ASW containing 1 m M amino acid. The values are single observations from six axons; each axon was tested with glycine first. From Caldwell and Lea (1978).

124

P. F. BAKER AND A. CARRUTHERS

b. Effect ofMetabolic Inhibitors. Cyanide inhibits glycine uptake after some delay (Caldwell and Lea, 1978). A kinetic analysis of this effect is lacking. Its delayed time-course suggests strongly that inhibition of uptake is the result of metabolic depletion; but whether this relates to the fall in ATP, rise in cytosolic ionized Ca, or something else is not known. c. Effects of Ouabain. The Na-pump inhibitor ouabain inhibits glycine uptake in the squid giant axon (Caldwell and Lea, 1979). This effect is not, however, specific for glycine transport because ouabain also inhibits orthophosphate, alanine, arginine, L-aspartate, L-glutamate (Caldwell and Lea, 1979), and D-glucose transport (Baker and Carruthers, 1981a). This effect appears not to be due to changes in axoplasmic levels of Na or K (Caldwell and Lea, 1979), nor is this phenomenon the result of accumulation of K in the periaxonal space. Caldwell and Lea (1979) have concluded that this effect might arise from activation of some cytosolic regulatory system initiated by Na-pump inhibition. Removal of K from the external milieu has only a small inhibitory effect on glycine uptake (Caldwell and Lea, 1979).As K removal also inhibits the Napump (Baker et al., 1969), it seems likely that the effect of ouabain on transport is mediated by a process rather more subtle than Na-pump inhibition per se. It is conceivable that ouabain may interact with the lipid matrix of the bilayer and influence transport systems through modifications in the bilayer state. Without further data, however, speculation along these lines is unwarranted.

C. Glutamate Transport I . CHARACTERIZATION OF FLUXES

Despite an adverse electrical gradient, the concentration of glutamate inside squid axons is maintained much higher than in squid blood. Some of this intracellular glutamate may be synthesized within the axons and some accumulated against the prevailing electrochemical gradient. Figure 15A shows that glutamate uptake into the giant axon has two components: one that increases linearly with external glutamate and a second that is a saturable function of external glutamate. Glutamate uptake into the sheath (Schwann cells plus residual small nerve fibers) can also be divided into two similar components. The saturable component of uptake into both axoplasm and sheath is Na-dependent whereas only the linear component is seen in seawaters in which Na has been replaced isosmotically by choline or lithium. Sodium-dependent uptake of glutamate into the giant axon exhibits an apparent K,,, of 100 pM and a V,,, of 32 fmol cm-2 sec-I at 20°C.

TRANSPORT OF SUGARS AND AMINO ACIDS 300 r

r

125

A

‘0

t

-

200

PI Y. (0

a a

2

;

--

100

a

0

0 0

1

2

3

[Glutamate]

4

5

mM 0

FIG. IS. Uptake of ~-/‘~C]glutamate into giant axons of Loligo ,firbesi. (A) Dependence of uptake on glutamate concentration in Na seawater ( 0 )and choline seawater (0). (B) Dependence of the saturable component of glutamate uptake on external Na at 20°C.

If all the glutamate in squid blood ( 2 . 2 mM) is free, and not bound within cellular elements, the glutamate influx under physiological conditions should amount to 120 fmol cm-2 sec-I, of which 32 fmol sec-’ is due to the Na-dependent, saturable component and 88 fmol cm-2 sec-’ to the Na-independent, linear component. After removal of C 0 2 , the rate conmin-I. Assuming stant for [14C]gIutamateefflux is in the region of 7 X an intracellular glutamate concentration of 25 mM, this is equivalent to a membrane flux of approximately 100 fmol cm-2 sec-I, which is quite close to the calculated glutamate influx under physiological conditions. Although this similarity in fluxes may seem satisfactory, the calculation of influx assumes that all the glutamate in squid blood is free, and this seems rather unlikely because in other invertebrates there is appreciable accumulation of glutamate in cellular components of the blood (see Evans, 1972). A lower free glutamate concentration in squid plasma would have the effect of increasing the importance of the Na-dependent uptake process relative to the linear component.

2. Na DEPENDENCE OF INFLUX The shape of the Na-activation curve suggests that more than one, and possibly two, sodium ions may be required to activate uptake (Fig. 15B). It is not known whether these Na ions are transported into the axon; but,

126

P. F. BAKER AND A. CARRUTHERS

by analogy with amino acid transport in other systems, this seems very likely (Schultz and Curran, 1970). It should be noted that the effects of Na on glutamate influx seem not to result from alterations in Ca influx or axoplasmic ionized Ca (Baker, 1972), because essentially similar results are obtained in Ca-free media. 3. PROPERTIES OF EFFLUX Glutamate efflux is remarkably insensitive to experimental manipulation. It is not obviously affected by external glutamate (10 mM), by replacement of external Na by choline, by K depolarization, by low temperatures or by the respiratory inhibitor cyanide (see Fig. 3). Efflux therefore behaves as though it is a passive leak of glutamate down the prevailing electrochemical gradient. 4. REGULATION

a . Eflects of Electrical Activity. In view of the fact that glutamate or some closely related molecule may be a transmitter at the squid giant synapse, the effects of electrical stimulation on glutamate influx and efflux are of particular interest. Neither the influx or efflux of glutamate is changed significantly during massive stimulation. Glutamate uptake and efflux are also unaffected by exposure of axons to external potassium concentrations that reduce the resting membrane potential close to zero. b. Metabolic Inhibitors. Glutamate uptake, but not efflux, is reduced to about half after exposure of axons to respiratory poisons. This effect is confined to the Na-dependent component of uptake and is somewhat more rapid in onset than inhibition of the sodium pump. The mechanism of this effect is unknown; but a rather similar sensitivity to metabolic inhibitors was described by Baker and Potashner (1973) in crab nerve, and they attributed their findings to metabolic regulation of the affinity of the uptake mechanism for glutamate. An analysis of glutamate fluxes in dialyzed or perfused axons should help clarify these and other questions. c. External p H . Glutamate uptake is rather constant between pH 7.5 and pH 9.5 but falls at acid pH, reaching approximately 30% of its control (pH 7.5) value at an external pH of 3.5. 5 . PHARMACOLOGY: ANTAGONISTS OF UPTAKE

In view of the role of glutamate as a transmitter molecule, glutamate uptake systems may be of importance in terminating transmitter action. We have examined a number of glutamate antagonists for their effects on glutamate uptake in squid axons. All measurements were carried out at a

127

TRANSPORT OF SLIGARS AND AMINO ACIDS

glutamate concentration of 100 kuM where uptake is largely Na dependent, but strictly speaking the antagonists should also be tested in choline seawater to establish whether they have any effect on the linear component of uptake (cf. Baker and Potashner, 1971). This was done in only a very few instances. Table X summarizes some of our main findings. Although a number of substances inhibit the uptake of glutam am ate, of those examined so far only cysteic acid and dihydrokainic acid have a higher affinity for the transport system than L-glutamate, and even in these cases the difference in affinity is not great. Both agents are known to inhibit glutamate uptake in other systems (see Baker and Potashner, 1971; Johnston e l ul., 1979). TABLE X INHIBITION OF GLUTAMATE UPTAKE

INTO

SQUID AXONS"

%

Amino acid L-Aspartic L- Aspartic-p-h ydroxamate

D-Aspartic o-Aspartic-p-h y droxamate N-Methyl-D-as partic

%

Inhibition 29 None 39 None 23

Amino acid

Inhibition

y-D-Ghtamylglycine Folic acid

None 42

Quinolinic acid

20 None

2,4-Pyridinedicarboxylic acid

Cysteic acid Taurin e L-2-Amino-4-phosphonobutyric ~-2-Amino-4-phosphonobulyric L-Glutamic

80 None 20 64 80

cis-2,3-Piperidinedicarboxylic trans-2,3-Piperidinedicarboxylic

D-Ghtamic

None

cis-2,4-Piperidinedicarboxylic

None

Glutaric

None

cis-2,S-Piperidinedicarbox ylic

None

N-Methylglutamic L-Glutamic-p-monohydroxamate D-aAminoadipic D-2-Aminophosphonovaleric DL-a- Aminosuberic

None None None 50 61

Kainic acid Dihydrokainic acid Quisqualic acid

None 85 None

Dipicolinic acid

None

70 17

a All measurements were made in sodium seawater containing 100 p M ~-['~C]glutamate and 500 p M antagonist under investigation at pH 7.6. Incubation was in general for 30 min at 20°C. Uptake was measured by counting extruded axoplasm. Inhibition is expressed relative to uptake in the presence of glutamate alone. All values are means of at least two paired determinations. Concentrations of cysteic acid, cis-2.3-piperidinedicarboxylicacid, and dihydrokainic acid required for half-maximal inhibition of uptake were approximately 30, 100, and 30 p M , respectively.

128

P. F. BAKER AND A. CARRUTHERS

VI. GENERAL CONCLUSIONS

The squid axon seems to be an excellent preparation in which to study transport of organic solutes, and it is surprising that it has been used so little. The giant axon has transport systems for sugars and amino acids, and where these have been studied in detail they reveal many features in common with the mammalian brain. A systematic analysis of the characteristics of the transport of organic molecules into squid axons may reveal a number of processes of widespread interest. Thus, Tasaki and Spyropoulos (1961) reported uptake of choline by squid axons, and subsequently Hodgkin and Martin (1965) described a saturable uptake system ( K , , 100 p M ; V,,, , 100 fmol cm-? sec - I ) that share many features in common with choline uptake into mammalian cells, and the axon also contains a sodium-sensitive uptake system for y-aminobutyric acid. In addition, the sheath takes up serotonin, and both sheath and giant axon have an active, hormone-sensitive, cyclic nucleotide metabolism (see article by Baker and Carruthers, pp. 271-276, this volume). The existence of transport systems and mechanisms by which they might be controlled in a cell that can be subjected to internal solute control by dialysis or perfusion offers many possibilities for experimental analysis of the mechanisms of permeation and their control. ACKNOWLEDGMENTS All our experimental work on squid axons was carried out at the Laboratory of the Marine Biological Station, Plymouth, U.K. We wish to thank the Director and Staff for the help they have given us over many years. Our work was supported by grants from the Medical Research Council. We are particularly indebted to Dr. J. C. Watkins for supplying many of the glutamate antagonists. REFERENCES Baker, G . F., and Widdas, W. F. (1973). The asymmetry of the facilitated transfer system for hexose in human red cells and the simple kinetics of a two component model. J . Physiol. (London)231, 143-165. Baker, P. F. (1972). Transport and metabolism of calcium in nerve. Prog. Biophys. Mol. B i d . 24, 177-223. Baker, P. F., and Carruthers, A. (19Xla). Sugar transport in giant axons of Loligo. J . P h y s i d . 316, 48 1-502. Baker, P. F., and Carruthers, A. (198lb). 3-0-Melhylglucose transport in internally dialysed giant axons of Loligo. J. Physiol. (London)316, 503-525. Baker, P. F., and Potashner, S. J. (1971). The dependence of glutamate uptake by crab nerve on external Na+ and K'. Biochim. Biophys. Actu 249, 616-622. Baker, P. F., and Potashner, S . J . (1973). The role of metabolic energy in the transport of glutamate by invertebrate nerve. Biochirn. Biophys. Acta 318, 123-139. Baker, P. F., and Stone, A. J. (1966). A kinetic method for investigating hypothetical models for the sodium pump. Biochim. Biophys. Acru 126, 321-329. Baker, P. F., Blaustein, M. P., Keynes, R. D.,Manil, J . , Shaw, T. I . , and Steinhardt, R . A.

TRANSPORT OF SUGARS AND AMINO ACIDS

129

(l96Y). The ouabain-sensitive fluxes of sodium and potassium in squid giant axons. J . Phy.rio1. (London) 200, 459-496. Baker, P. F., Hodgkin, A. L . , and Ridgway, E. B. (1971). Depolarisation and Ca entry in squid giant axons. J . Physiol. (London) 218, 709-755. Bloch, R. (1973). Inhibition of glucose transport in the human erythrocyte by cytochalasin B. Biochetnisiry 12, 4799-4801. Boron, W. F., and de Weer, P. (1976). Intracellular pH transients in squid axons caused by C 0 2 , NH, and metabolic inhibitors. J . Gen. Physiol. 67, 91-112. Bowyer, F., and Widdas, W. F. (1958). The action of inhibitors on the facilitated hexose transfer system in erythrocytes. J . Physiol. (London) 141, 219-232. Brinley, F. J., Jr., and Mullins, L. J. (1967). Sodium extrusion by internally dialysed squid axons. J . Gen. Physiol. 50, 2303-2331. Britton, H. G. (1964). The permeability of the human red cell to labelled glucose. J . Physiol. (London) 170, 1-20. Caldwell, P. C. (1958). Studies on the internal pH of large muscle and nerve fibres. J . Physiol. (London) 142, 22-62. Caldwell, P. C. (1960). The phosphorous metabolism of squid axons and its relationship to the active transport of sodium. J . Physiol. (London) 152, 545-560. Caldwell, P. C., and Goldstuck, N. D. (1979). The uptake of delta-aminolaevulinic acid into squid giant axons. J. Physiol. (London) 287, 2 2 ~ . Caldwell, P. C., and Lea, T. J. (1978). Glycine fluxes in squid giant axons. J . Physiol. (London) 278, 1-25. Caldwell, P. C., and Lea, T . J. (1979). The effect of ouabain on amino acid and orthophosphate influxes in squid axons. J. Physiol. (London) 289, 389-401. Carruthers, A. (1979). Internal dialysis as a method for studying sugar transport in large nerve and muscle fibres. J . Physiol. (London) 287, 5 - 6 ~ . Carruthers, A . (1983a). Sugar transport in giant barnacle muscle fibres. J . Physio/. (London) 336, 377-396. Carruthers, A. (1983b). Sugar transport in animal cells. The passive hexose transfer system. Prog. Biophys. M o l . Biol. (in press). Carruthers, A,, and Melchior, D. L. (W83a). Asymmetric or symmetric? Cytosolic modulation of human erythrocyte hexose transfer. Biochim. Biophys. Acru 728, 254-266. Carruthers, A,, and Melchior, D. L. (1983b). The relationship between bilayer physical state and bilayer water permeability. Biorhamisiff 22, 5797-5807. Clausen, T. (1975). The effect of insulin on glucose transport in muscle cells. Ctrrr. T o p . Mrrnbr. Trunsp. 5, 169-226. Crane, R. K . (1977). The sodium gradient hypothesis and other models of carrier-mediated active transport. Rev. Physiol. Biochrm. Phurmucol. 78, 99-159. Crank, J. ( 1956). “The Mathematics of Diffusion.” Clarendon Press, Oxford. Deffner, G . J . (1961). The dialysable free organic constitutents of squid blood: A comparison with nerve axoplasm. Biochim. Biophys. Actu 47, 378-388. Diamond, I . , and Fishman, R. A. (1973). High affinity transport and phosphorylation of 2deoxy-o-glucose in synaptosomes. J . Neurochem. 20, 1533-1542. Elbrink, J., and Bihler, I. (1975). Membrane transport: Its relation to cellular metabolic rates. Science 188, 1177-1 184. Evans, P. D. (1972). The free amino acid pool of the haemocytes of Curcinus tnuenm (L). J . EX^. B i d . 56, 501-507. Geck, P. (1971). Properties of an asymmetric carrier model for the transport of sugars by human erythrocytes, Biochitn. Biophvs. Actrr 241, 462-472. Hankin, B. L., Lieb, W. R., and Stein, W. D. (1972). Rejection criteria for the asymmetric carrier and their application to glucose transport in the human red blood cell. Biochitn. Biophys. Actti 288, 114-126.

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Heinz, E. (1972). In "Metabolic Transport" (L. E. Hokin, ed.). Vol. 6 . pp. 455-501. Academic Press, New York. Hodgkin, A. L., and Keynes, R. D. (1955). The potassium permeability of a giant nerve fibre. J . Physiol. (London) 128, 61-88. Hodgkin, A. L., and Martin, K. (1965). Choline uptake by giant axons of Loligo. J. Physiol. (London) 179, 2 6 ~ . Hoskin, F. C. G., and Brande, M. (1973). An improved sulphur assay applied to a problem of isethionate metabolism in squid axon and other nerves. J . Neurochem. 20, 13171327, Hoskin, F. C. G., and Rosenberg, P. (1966). Penetration of sugars, steroids, amino acids and other organic compounds into the interior of the squid giant axon. J . Gen. Physiol. 49, 47-56. Johnston, G. A. R., Kennedy, S. M. E., and Twitchen, B. (1979). Action of the neurotoxin kainic acid on high affinity uptake of L-glutamic acid in rat brain slices. J . Neurochem. 32, 121-127. Jones, M. N.. and Nickson. J. K. (1981). Monosaccharide transport proteins ofthe human erythrocyte membrane. Biochim. Biophys. Acrn 650, 1-20. Krupka, R. M., and Deves, Rosa (1981). An experimental test for cyclic versus linear transport models. The mechanism of glucose and choline transport in erythrocytes. J. Biol. Chum. 256, 5410-5416. I,acko, L.. Whittlee, B., and Kromphandt. H. (1972). Zur Kinetic der Glucose-Aufname in Erythrocyten. Eur. J. Biochfm. 25, 447-454. Lieb, W. R., and Stein, W. D. (1974). Testing and characterising the simple carrier. Riochim. Biophys. Acia 373, 178-196. Melchior, D. L. (1982). Lipid phase transitions and regulation of membrane fluidity in prokaryotes. Curr. Top. Membr. TronJp. 17,263-3 16. Naftalin, R. J., and Holman, G. D. (1977). Sugar transport in human red cells. In "Membrane Transport in Red Cells," (J. C. Ellory and V. L. Lew, eds.). Academic Press, New York. Kosenberg, P., and Khairallan, E. A. (1974). Effect of phospholipases A and C on free amino acid content of the squid axon. J . Ncrrrochern. 23, 55-64. Schultz, S.G., and Curran, P. F. (1970). Coupled transport of sodium and organic solutes. Physiol. Rev. 50, 637-718. Sen, A. K., and Widdas, W. F. (1962). Determination of temperature and pH dependence of glucose transfer across the human erythrocyte membrane measured by glucose exit. J. Physic)/. (London) 160, 392-401. Sokoloff, L., Reivich, M.. Kennedy, C., Des Rosiers, M. H., Patalak, C. S . , Pettigrew, K. D., Sakurada, 0.. and Shinohara. M. (1977). The ['4C]-deoxyglucosemethod for the measurement of local cerebral glucose utilization; theory, procedure and normal values in the conscious and unanesthetised albino rat. J. Neurochcm. 28, 897-915. Steele, J. A,, Pozawsky, M. J., Eaton, D. C., and Brodwick. M. S. (1981). Lipid vesiclemediated alterations of membrane cholesterol levels. Effects on Na' and K' currents in squid axon. J . Memb. Biol. 63, 191-198. pasaki, I., and Spyropoulos. C. s. (1961). Permeability of the squid axon membrane to several organic molecules. Am. J. Physiol. 201, 413-419. Widdas, W. F. (1952). Inability of diffusion to account for placental glucose transfer in the sheep and consideration of the kinetics of a possible carrier transfer. J. Physiol. (London) 118, 23-29. Widdas. W. F. (1980). The asymmetry of the hexose transfer system in the human red cell membrane. Ckrr. lop. Mrmbr. T r m s p . 14, 165-223.

CURRENT TOPICS IN MEMBRANES A N D TRANSPORT. VOLUME 22

Sodium Pump in Squid Axons LUIS BEAUGE Divisidn de Biofisica, Instituto de Investigacidn Medicu Mercedes y Martin Ferreyra Cdrdoba, Argentina

C. Internal Dialysis.. . . . . . . . . . . . . . . . D. Internal Perfusion. 111. Sodium Pump in Squid A. Active Na-K Exc ..................... B. Na-Na Exchange . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Uncoupled Na Efflux . . .

134

E. Na Fluxes in the Presence of Cardioactive Steroids . . . . . . . . . . . . . . 163 F. K-Free Effect ........................................ . . . . . . . . 168 G. Conclusions ........ 170 References.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 171

1.

INTRODUCTION

This is a critical comparative review of the work done on the Na pump in squid axons and other systems. During active transport of Na and K, and the associated biochemical occurrences, the responsible structure(s) react(s) with several ligands, suffering what is loosely called conformational changes. The views most accepted currently are illustrated in Fig. 1; they are given here to be used as a framework so that the reader may follow the discussion in the different sections of this article. For a detailed account, the reader is referred to specific reviews on the subject 131 Copyright 0 1984 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-153322-0

132

LUIS BEAUGE

FIG. 1. Modified Albers-Post scheme for the Na+,K+-ATPaseand the Na-K pump cycle and partial reactions. The solid lines represent the pathways for active Na,-KO exchange (Na+,K+-ATPase), Na-Na exchange (ATP-ADP exchange), K-K exchange (EH 2 0 P, exchange), and K,-Na, reversal (ATP synthesis). The dashed line is the possible pathway for the uncoupled Na efflux (Na,-ATPase). The dotted line is the hypothetical path for the K-like action of Na, (Na,,Na,,-ATPase); Na* denotes Na ions acting as K on K sites. The sites of inhibition by ouabain, oligomycin, NEM, and vanadate are indicated. For the original references see Albers, 1967; Skou, 1975; Glynn and Karlish, 1975; Robinson and F h h n e r , 1979; Jorgensen, 1980; De Weer, 1983.

(Albers, 1967; Skou, 1975; Glynn and Karlish, 1975; Robinson and Flashner, 1979; Jorgensen, 1980; De Weer, 1983). Other works directly concerned with the squid axon should also be consulted (Mullins, 1972, 1979; De Weer, 1975). II. EXPERIMENTAL TECHNIQUES A. Axoplasm Extrusion

Axoplasm extrusion has been used primarily for influx studies (Caldwell and Keynes, 1960). The axon is soaked in a radioactive solution for a certain time; after radioactivity is washed away from the extracellular space, the axoplasm is "rolled out" of the axon through one of its ends. Obviously this method does not allow the control of cytosol composition or biochemistry.

SODIUM PUMP IN SQUID AXONS

133

B. Microinjection

Microinjection was developed by Hodgkin and Keynes in 1956. It consists of the injection of substances into the axon by means of a microsyringe, and it finds its main uses in efflux studies. Although it does not permit control of the intracellular environment, it is extremely useful for the introduction into the cytosol of different solutes (ions, metabolites, substrates, enzymes, etc.) and has opened the door to the biochemistry of axons with their membrane and axoplasm as an integrated unit. C. Internal Dialysis

Internal dialysis was developed by Brinley and Mullins (1967). This indeed constitutes a major breakthrough in the methodology of transport studies in squid axons. A capillary tube (made of glass or plastic) with a selective “porous” region is longitudinally steered through the axon. The porous region allows the passage of solutes with a molecular weight of 1000 or less. Following a “dialysis” solution of known composition through the capillary gives the basis for effective control of small intracellular ions, metabolites, substrates, and nucleotides. At the same time it permits the measurement, with great accuracy, of the efflux and influx of substances tranported across the axon membrane.

D. Internal Perfusion Internal perfusion is aimed at controlling intracellular composition and metabolism. As proposed by Baker et al. (1962), most of the axoplasm was removed and replaced by a flowing solution. As such, it was not successful for use in studies on the Na pump owing to the large unspecific Na fluxes, but it was, and still is (with some modifications), the ideal tool for experiments on exitability. DiPolo and Beauge (1980) offered a modified version, where a small part of the axoplasm is removed by two intraaxonal cannulas of different diameters. Beginning with one overlapping inside the other, the cannulas are slowly separated, leaving the “perfused” region between their ends. Although successful in large axons, the percentage of failures increases sharply in axons with diameters of less than 500 pm. The survival time of the axons with this technique (up to I .5 and 2 hr) is much shorter than with internal dialysis (up to 12 hr in our experience). It is the feeling of this reviewer that, at the present time, internal dialy-

134

LUIS BEAUGE

sis is by far the best available method for studying the Na pump in squid axons. However, it has two major drawbacks: (1) the molecular weight of the solutes that can be “dialyzed” into the axons is too small; and (2) the equilibration time for nucleotides, especially in large axons, is rather long and the concentration at the inner membrane surface is uncertain. Complete control of the intracellular medium in squid axons will demand either a type of dialysis capillary with a much larger pore size or a more reliable perfusion technique.

111.

SODIUM PUMP IN SQUID AXONS

A. Active Na-K Exchange

1 . EFFLUXOF SODIUM a . Free Energy Source for Sodium Extrusion. The search for the free energy source for Na extrusion began with the experiments of Hodgkin and Keynes in 1955. Giant axons from Loligo forbesi and Sepia officialis (Hodgkin and Keynes, 1955) bathed with artificial seawater containing cyanide, 2,4-dinitrophenol, or azide presented a reduction in the efflux of Na and in the influx of K without changes in the fluxes of the same cations in the other directions. Cell metabolism was obviously related to active Na-K transport in nerves. Propelled by the introduction of the microinjection technique, Caldwell(l960) and Caldwell et al. (1960a,b) performed numerous experiments where Na efflux was followed in axons poisoned with cyanide and 2,4-dinitrophenol and injected with several “high energy” phosphate compounds. Figure 2, taken from Caldwell et al. (1960a), shows that total as well as KO-stimulatedNa efflux falls upon cyanide poisoning; in addition, the figure also shows that, when the exposure to cyanide has been long enough, the efflux of Na becomes larger in the absence of external K than in its presence (reversal of K-free effect). Injection of adenosine 5-triphosphate (ATP) restores a large fraction of Na efflux but not its KO sensitivity, which is regained somehow after removal of cyanide. On the other hand, if phosphoarginine is injected instead of ATP (not shown here) both Na efflux and its KOsensitivity are recovered. On this basis the authors proposed that although ATP could support Na efflux, arginine phosphate or the ratio ATP : adenosine 5‘diphosphate (ADP) was responsible for the coupling between Na and K transport (see also Section IILF). The development of the internal dialysis technique permitted a cleaner approach to the problem. In this way compounds were tested isolated

135

SODIUM PUMP IN SQUID AXONS

2mM

0

CN

1

5

::0.0010

L

z“

w ; 00005 C

s

+ \

7

L

L

O

1

0

5

2

ATP injected

6

I

I

7

Hours

FIG.2. Effect of injecting ATP into a CN-poisoned nerve fiber on Na efflux into K-free solution ( 0 )and 10 mM K (0).The length injected with ZZNawas 12 mm; that with ATP, I8 mm. Immediately after injection the mean concentration of ATP in the fiber was 4.8 mM. The ATP contained about 4% of the phosphorus as ADP. (Taken, with permission, from Caldwell et ul., 1960a.)

from those other compounds metabolically related to them and in the absence of the rather complicated biochemical changes likely to occur in poisoned cells. The accomplishments were indeed striking. By dialyzing the axons with ATP-free solutions, Brinley and Mullins (1968) were able to reduce the ATP content of the axoplasm to the micromolar range and with it to drop the efflux of Na to a small percentage of its original level (or of the levels found in similar axons with the injection technique). The residual Na efflux from axons nominally free of ATP is about 1 pmol cm-2 sec-’, and in many instances less than one-half that value. A normally KOstimulated Na efflux was restored when millimolar concentrations of ATP were added to the dialyzing media. Throughout their work the authors used cyanide to speed up ATP washout. In experiments performed some years later, BeaugC and DiPolo (1978, 1979a, 1981a) obtained essentially the same results in the absence of any metabolic inhibitor. Testing several phosphate compounds for their capability to sustain Na extrusion in ATP-free” axons, Brinley and Mullins gave convincing evidence that ATP is the unique free energy donor for the Na pump (see Table I). Their results deserve further consideration. On the one hand, the inability of phosphoarginine to promote any efflux of Na clearly indicates that the results of Caldwell et al. (1960b) were indeed due to the formation of ATP from the endogenous ADP and the exogenous phosphoarginine. On the other hand, the small stimulation found with uridine 5‘-triphosphate (UDP), cytidine 5’-triphosphate (CTP), and guanosine 5’-triphosphate (GTP) was originally attributed to a likely contamination with ATP; although this may have been the case, it is “

136

LUIS BEAUGE

TABLE I

ENERGYSOURCES FOR SODIUM EXTRUSION“ ~

~~

Number of experiments

Compound ATP*h PA PEP AcP G-3-P

AMP ADP* UTP* CTP* GTP* dATP*

Y

I

3 2 3 2

Physiological concentration

Test concentration

Na extrusion

(mM)

(mM)

(% of normal)

2-5 3

5 10 2.5

2-3 5 0.05-0. I

0.05-1

0.05

2

20 2 2 1.4

2

0.1-5

80- 100

<2 <2 <2 <5 4-10

1 1

<5 <5 4-10 -25-50

From Brinley and Mullins, 1968. Asterisk indicates Mg:’ set at 4 m M in excess of (-P,)

known that these nucleotides, especially CTP and inosine 5’-triphosphate (ITP) (ITP was not tested in axons), are also substrates of the ATPase enzyme; therefore, a direct effect on Na efflux cannot be ruled out (see Robinson and Flashner, 1979, for references). It is also interesting that there was no stimulation of Na efflux by acetyl phosphate (AcP), for it is known that AcP can phosphorylate the Na+,K+-ATPasein a way that is dependent on Na and competitively inhibited by ATP. The enzyme phosphorylated from AcP has the same spontaneous dephosphorylation rate as that phosphorylated from ATP, and there is indirect evidence for the existence of a similar K-promoted dephosphorylation (Bond et al., 1966, 1971; Bader and Sen, 1966) (for other views on the subject, see Dudding and Winters, 1971). On the basis of those facts alone, it was logical to expect some positive results with AcP. However, at that time the “regulatory” role of ATP in accelerating the E2(K)-EIK conformational change (disocclusion) (Post P I al., 1972) had not been established. The regulatory role of ATP is shared by other nucleotide polyphosphates including the nonhydrolyzable analogs. If AcP can phosphorylate the Na pump but cannot replace ATP as a disoccluding agent, turnover will stop, or be drastically reduced at the point of the E,(K) form; as a consequence the AcP-promoted Na-K exchange would be minimal or absent. It would be extremely important to test the effects of AcP on Na-K exchange with the addition of a nonhydrolyzable ATP analog; the idea is that the cycle

SODIUM PUMP IN SQUID AXONS

137

beginning with Na-dependent phosphorylation and Na translocation due to AcP will be completed by the ATP analog-facilitated K translocation. There is a problem in this proposal; a K-stimulated dephosphorylation of the AcP-produced phosphoenzyme has not been directly verified, and it is possible that it does not exist. If this is the case, it might still be feasible to detect an AcP-promoted Na extrusion in the absence of external as well as internal potassium, even without the addition of an ATP analog, owing to the absence of the E2K state under these conditions. These important experiments have yet to be done. Brinley and Mullins (1968) reported an ATP stimulation curve of Na efflux that had two components, one saturable and one linear. The saturable component had a K I l 2of about 30 p M ; that value was, and still is, completely unique. Considering the two roles of ATP, it means that the “regulatory” action is exerted in the squid axon with much higher apparent affinity than in any other kind of system investigated so far. In most of these experiments there was 5-10 mM phosphoarginine (PA) in the dialysis solutions. This was in order to prevent ATP losses in its transit from the dialysis capillary toward the cell membrane; any ATP hydrolyzed owing to axoplasmic ATPases would be resynthesized from PA and the released ADP. There is no direct evidence of how successful this “buffering system” is, and a failure in ATP resynthesis could account for the results; however, from what follows, this does not seem to be the reason. BeaugC and DiPolo (1978, 1979b, 1981a) found that the Na efflux with 50 pM ATP in the dialysate was not 50%, but only IS%, of that seen with ATP in the millimolar range. A study of the ATP dependence on total Na efflux showed a single saturable curve with a K1/2of about 200 ~ L M(Fig. 3 , top). These findings correlate very well with the ATP stimulation of a Na+,K+-ATPase activity present in membrane fragments isolated from optic nerves of the squid Sepiotheutis sepioidea (bottom of Fig. 3 ) . It could be argued that this correlation is just a coincidence, for the preparations come from different nerve tissues in different species. There are arguments against this view. First, this Na+,K+-ATPaseis activated by K and inhibited by ouabain in the same way as Na efflux in dialyzed axon from Loligo pealei. Second, these membrane fragments possess a Ca2’ ,Mg2+-ATPaseactivity that has kinetic characteristics almost identical to the ATP-dependent uncoupled Ca efflux in Loligo axons. These results have been confirmed in Na’,K+-ATPase isolated from Loligo optical nerves (Breitweiser, 1982). The problem is how to explain the observations of Brinley and Mullins (1968). There are various possibilities. I . The dialysis technique in both groups of experiments was essentially the same, but Beauge and DiPolo did not use cyanide; although not ex-

138

LUIS BEAUGE 1.0r

k

s E

0

-

0 c

Y

.-

-

g 0.5Y

L

X

L L

0

2

ATP concentration (mM1

ATP CONCENTRATION ( mM1

FIG.3. Top: ATP dependence of total Na efflux in squid axons dialyzed with 310 mM K, and 70 mM Na, and incubated in 10 m M K artificial seawater. The dialysis solutions contained variable ATP concentrations and 5 mM phosphoarginine. The data have been normalized to the levels of Na efflux with 3 mM ATP. Bottom: The ATP dependence of the ouabain-sensitive Na+,K+-ATPase activity in membrane fragments from squid optical nerves. The incubation mixture had the following composition (mM): NaCI, 460; KCI, 50; Tris-HC1, pH (25°C) 7.4,30; EGTA, 0.5; MgCI2, 4 mM in addition to the ATP concentration. The internal and external composition is given as rnillimolar. (Taken from Beaugk and DiPolo, 1981a.)

SODIUM PUMP IN SQUID AXONS

139

plored yet, it is possible that cyanide might directly or indirectly affect the Na pump. 2. Whereas Beauge and DiPolo used aspartate as the main anion in the dialysate, Brinley and Mullins used isethionate. It is now known that isethionate salts are quite impure and that among the contaminants are mono- and divalent cations, particularly calcium (DiPolo, 1977). Unfortunately the presence of Ca, through the induction of an ATP-dependent Na-Ca exchange, is not useful as an explanation because the K112 for ATP in that exchange reaction is also around 200 p M (DiPolo and Beauge, 1980). If an impurity in the isethionate is responsible, its nature remains unknown. 3. Brinley and Mullins used vanadate-containing ATP. It is conceivable that, as ATP was increased, vanadate reached its inhibitory concentration, thus reducing the stimulation of Na efflux (with a parallel reduction in the Kliz).There are two arguments against this possibility: first, it does not explain the existence of two components in the ATP curve; and second, the values of Na efflux with millimolar ATP concentrations were actually higher than those reported by Beauge and DiPolo. At present the reason for the discrepancy remains an open question.

b. External K Activation of Nu Efflux. In their classic paper, published in 1955, Hodgkin and Keynes formulated the basis for the need, and gave the experimental evidence for the existence, of a coupled Na-K active transport in nerve. This mechanism would reverse the net Na gain and K loss taking place during electrical activity, and it has been known since as the recovery process. Hodgkin and Keynes not only observed an Na efflux and a K influx depending on metabolism, but they showed that the removal of external K resulted in a large reduction (about a factor of 3) in the efflux of Na; i.e., external K ions were not only transported in but their presence was essential for the movements of Na in the other direction. From those observations, it seemed that active Na-K transport could be expressed either as (1) metabolically dependent Na efflux or K influx, or (2) KO-stimulatedefflux of Na. Time was to demonstrate that this view was an oversimplification. The efflux of Na as a function of the KOconcentration in Na-containing artificial seawater reaches saturation between 50 and 100 mM KOwith a of approximately 10 mM (Baker et al., 1969; De Weer, 1971; BeaugC and DiPolo, 1979b, 1981a). The KO-activation curve is sigmoid, but the sigmoidicity is more manifest when what is plotted is the ouabain-sensitive Na efflux (Baker et al., 1969) rather than the total KO-stimulatedflux (De Weer, 1971; BeaugC and DiPolo, 1979b, 1981a). When the external Na concentration is reduced, the for KOis also reduced without changes

140

LUIS BEAUGE

in the maximal rate of pumping, indicating a competitive inhibitory effect of Na, (Baker et al., 1969) (Fig. 4). On the other hand, other monovalent cations can replace K as external pump activators with varying efficiency; Sjodin and BeaugC (1967, 1968) and Baker et al. (1969) found that the order of effectiveness for equimolar concentrations followed the sequence K > Rb > Cs. The whole series of external activating cations follows the sequence T1 > K > Rb > NH4 > Cs > Li > Na; this has been shown for the Na+,K+-ATPaseactivity (Skou, 1960) and the active Na transport in several tissues (see BeaugC, 1975). It was a surprising finding that in axons Li not only did not have any measurable K-like effect but actually behaved like Na inhibiting the KOactivation of the Na pump; on the other hand, Li is unable to replace N a , in the Na-Na exchange flux (Baker et al., 1969). This peculiar behavior of squid axons may indicate that some properties of the Na pump, the external monovalent cation sites in this case, although similar, are not strictly identical in all species. c . Activation by Zntracellular N u . Over the years, experiments carried out on squid giant axons yielded a linear relationship between total Na

1

I

I

I

0

10

20

30

External K concentration ( m M )

FIG.4. Collected results of experiments on the K dependence of the ouabain-sensitive Na efflux in squid axons showing the effects of replacing external Na by choline or dextrose. l h e ordinate is the ouabain-sensitive Na efflux relative to that in 10 K (Na) artificial seawater (ASW), and the abscissa is the external K concentration (mM).0 , (460 Na)-ASW; U, (choline)-ASW; (dextrose)-ASW; (3, (230 Na-choline)-ASW; a, (230 Na-dextrose)ASW. Temperature 17-19°C. (Taken. with permission, from Baker et a / . , 1969.)

.,

SODIUM PUMP IN SQUID AXONS

141

efflux (into Na seawater) and Na, concentration. This behavior was observed in injected axons over a range of Na, between 16 and 220 mmol per kilogram of axoplasm (Hodgkin and Keynes, 1956; Sjodin and BeaugC, 1967, 1968; Baker, 1968; Baker et al., 1969) and in dialyzed axons with Na, between 5 and 230 mM (Brinley and Mullins, 1968). There are some explanations, none of them entirely satisfactory, that could be advanced to account for these results. One is that the density of pump units in axons is exceedingly high; Baker and Willis (1972) have estimated it to be between 1000 and 10,000 per square micrometer, which is within the range observed in frog skeletal muscle (Venosa, 1981), which shows a sigmoid relationship between rate of Na pumping and Nai concentration (Sjodin, 1971). Another is that the affinity of the squid pump for Na, is so low that Nq concentrations up to 200 mM are still within the approximately linear region of the activation curve; the data on Na activation of the Na+,K+ATPase in squid nerves (M. Campos, L. BeaugC, and R. DiPolo, unpublished; Breitweiser, 1982) are not consistent with this idea. A third possibility (L. J. Mullins, personal communication) is that the total Na efflux from these axons consisted of two components, one the Na pump and the other Na-Ca exchange. As the Na-Ca exchange increases with the square of Na, concentration (Baker, I968), a sigmoid relation for the Na pump could finally give a total linear response of Na efflux vs Na,. New experimental evidence, which appeared as an abstract (Kracke and De Weer, 1982), may indicate that perhaps external Na, via some type of Na-Na exchange flux, had something to do with the results described before. The total Na efflux from injected axons bathed in Na-free seawater and the ouabain-sensitive efflux of Na from axons dialyzed with 2 mM ethyleneglycol bis(3-aminoethyl ether-N,N’-tetraacetic acid (EGTA) in the presence of low Na, solutions followed a sigmoid relationship with Na, concentration. For the ouabain-sensitive flux, the authors give a Hill coefficient of 1.8 and a K , of 25 mM (without specifying what they mean by K , as they precisely stated a non-Michaelian relationship). Unfortunately, there is too little information to warrant discussing the matter further;.at any rate, a sigmoid relationship between the rate of Na pumping and Na, concentration no doubt accommodates itself very comfortably with current ideas about the Na pump. An expectation that should be borne out is an antagonism (of a competitive type) between internal Na and K ions. A relationship of this kind indeed exists, but it is complicated by the fact that ATP is also involved in this Na,-K, interaction (BeaugC and DiPolo, 1979b, 1981a). This aspect is discussed in Section III,A, I ,f. d . Effect of Znhibitors. The active Na-K transport is a complex process by which energy released from cell metabolism and stored in the

142

LUIS BEAUGE

form of ATP is transferred to a membrane-located system in order to perform osmotic work (cation translocation). In theory, such a mechanism can be inhibited in the following ways: (1) by slowing the release of free energy from metabolism; (2) by reducing the storage of that energy in the form of ATP (as happens with the so-called uncouplers in oxidative phosphorylation); (3) by interfering with the utilization or transfer of the free energy from ATP into the translocating system; and (4) by blocking cation translocation with or without interference with ATP hydrolysis; in the latter case we would be in the presence of a metabolic “uncoupler” of the pump. The use of inhibitors of types (I) and (2) allowed the demonstration that Na and K transport against a gradient indeed depended on cell metabolism (Hodgkin and Keynes, 1955; Caldwell, 1960; Caldwell et al., 1960a,b) and will not be dealt with further here. The present section will deal with those operating as described in (3) and (4). At this stage it is important to mention that so far all inhibitors acting as in (4) act also as in (3). In other words, there is no known compound that can uncouple ATP hydrolysis from Na-K translocation; in this regard Na+,K+-ATPase activity and Na-K translocation behave as a single entity. i . Cardiouctiue Steroids. Since Schatzman (1953) established that cardiac glycosides inhibit active Na and K transport in erythrocytes, these and other related steroids, basically strophanthin G (ouabain) and strophanthidin, have been widely used. On the assumption that these agents completely block Na,K-dependent ATP hydrolysis and active NaK exchange, without affecting “residual” enzymatic or transport activities, these steroids found their main application in the separation or isolation of those processes specifically related to the Na pump. (A possible complication derived from this assumption will be dealt with in Section 111,E.) In 1959, Caldwell and Keynes made the initial observation in squid axons M ouabain in seawater produced “at once” about 80% showing that M intracellularly injected solution reduction in Na efflux, whereas a had no effect; i.e., ouabain acted on the external membrane surface. Considering that the ligands favoring ouabain inhibition also favor the E2P state of the enzyme, it is assumed that this is the enzyme form to which cardioactive steroids bind more effectively (see Robinson and Flashner, 1979); nevertheless, the inhibition mechanism is by no means completely understood. In squid axons the dependence of ouabain inhibition on extracellular cation composition and cell metabolism has been worked out by Baker and Manil (1968) and by Baker and Willis (1972). Using a high-speed apparatus (collection times down to 2.5 sec), they showed that the rate of inhibition was concentration dependent, but even at saturating concentra-

143

SODIUM PUMP IN SQUID AXONS

tions (lop4M ) the rates were lower than the drop in Na efflux following removal of K, ; a puzzling observation was that in about 30% of the axons inhibition was preceded by a transient stimulation (see also De Weer, 1971). Removal of Na, reduced the rate of ouabain inhibition except when Li was used as a Na substitute; this is another indication that in squid axons external Li behaves mostly in an Na-like manner and has a very small K-like action, if any at all. In addition the rate of inhibition was faster in K-substituting than in choline- or dextrose-substituting seawater; this again is intriguing, for the K,-ouabain antagonism is a well-documented fact (Beauge and Adragna, 1971; Robinson and Flashner, 1979). Perhaps the existence of surface charges near the pump sites (Kracke and De Weer, 1980) is partially responsible for these differences. Finally, washout of ouabain already bound could be accomplished only in cyanide-poisoned axons, suggesting that the binding of the glycoside was influenced by the metabolic state of the cell. The K112for steady-state inhibition of Na efflux is about 5 to 7 x lo-* M , both for ouabain (Baker and Willis, 1972) and strophanthidin (De Weer, 1971). This value is about 10 times smaller than those obtained for the inhibition of Na+,K+-ATPase

TABLE 11 INHIBITION BY OUABAIN, STROPHANTHIDIN, VANADATE, AND OLICOMYCIN OF Na I N MEMBRANE FRAGMENTS EFFLUXI N SQUIDAXONSA N D Na+,K’-ATPase ACTIVITY FROM SQUID NERVESOR I N PURIFIEDNa+,K+-ATPaseFROM PIG KIDNEY Inhibition of Na+,K+-ATPaseactivity

Compound

Inhibition of Na efflux

Ouabain

Kllz= 7

Strophanthidin Vanadate

K1/: = 5 x lo-* M” Full inhibition of K,,-activated efflux

Oligomycin

N o effect at 10

(10

X

lo-* M a

Membrane fragments from squid nerves

Purified enzyme from pig kidney

Kllz = 9 x lo-’ M ” K I l 2= 3 X IO-’M‘ K I l 2= M“

-

K ~ , ?= 5 x 10-7 M‘

Kl12= 7 x lo-’ M”

K1/2 = 3

M)’

Baker ef a / . (I969a).

ME

” M. Campos and L. Beauge, unpublished data.

’ [

-

Breitweiser, 1982. De Weer, 1971. Beauge, 1979. Beauge and DiPolo, 1979a. De Weer et d.,1983.

X

lo-’ M ”

144

LUIS BEAUGE

activity present in membrane fragments from squid nerves (see Table 11). The discrepancy might reflect differences not only in cell metabolism, but also in ionic environments under the two circumstances. ii. Vanudlite. At the time the effects of vanadate on Na efflux in squid axons were investigated (Beauge and DiPolo, 1979a), the following facts about vanadate inhibition had been established (see Beauge et al., 1980 for references): (1) the inhibitor acted intracellularly; (2) inhibition was stimulated by Mg'+; and (3) rxtrurellr~larK acted as cofactor for inhibition. The experiments of Beauge and DiPolo were performed in dialyzed axons with the usual internal ionic composition (see legend to Fig. 5 ) , with 3 mM ATP and 5 mM PA. The external medium was Na seawater where K (or NHJ was varied from zero to 50 mM. The results reported in that paper confirmed that vanadate acts on the intracellular side. In addition, even for a 1 mM vanadate concentration, there was a time lag of about 1

10 K ASW a.

20.

70 No-310 K - 3 ATP- 5 PA 1 m M Vanadate [ O K 1 1 K I2.5K I 5 K 1 2 0 K 1 5 0 K

... . .. . 0

20

+

[KllrnMl

50

FIG.5 . Effect of different external K concentrations on the Na efflux in a vanadatepoisoned dialyzed squid axon. Note that the values of Na efflux in K-free Na artificial seawater (ASW) are the same in the presence and in the absence of vanadate. In the inset, the efflux of Na IS plotted as a function of the external K concentration: note the biphasic response to K O .All solute concentrations are given as millirnolar. (Data taken from BeaugC and DiPolo, 1979a.)

SODIUM PUMP IN SQUID AXONS

145

hr from the time the inhibitor was applied until a steady Na efflux was obtained; this is very important because ( 1 ) 1 hr is more than enough time to reach equilibrium distribution in the dialyzed region of the axons (Brinley and Mullins, 1967), and (2) the K l l zfor vanadate inhibition of the N a ’ .K+-ATPase activity in membrane fragments of squid nerves is about M (Table 11). A possible explanation for this behavior could be the existence of vanadate binding structures in the axoplasm, the presence of oxidation-reduction systems that can render vanadate ineffective (Grantham and Glynn, 1979), and/or the different ionic composition in the two assays. A discrepancy, although not as marked as that described here, can be found in the apparent affinity for vanadate in the inhibition of the Ca pump in dialyzed axons ( K I l ?= 7 x IWh M ) (DiPolo and BeaguC, 1981) and the Ca” ,Mg?+-ATPase activity in membrane fragments of squid nerves (M. Campos, R. DiPolo, and L. Bcague, unpublished data) ( K O 2= 3 x 10-7 M I . Figure 5 also shows that in the absence of external K there is no vanadate effect on Na efflux. The addition of KOto a vanadate-poisoned axon induces a biphasic response of Na efflux, an increase at low K, concentrations followed by an inhibition at high K, concentrations (the effect is almost saturated with 50 mM K,,); the magnitude of the inhibition is such that the actual levels of fluxes in the presence of sufficiently high K, are smaller than in its absence. This reverse K-free effect takes place under conditions completely different from those of cyanide poisoning (Caldwell et al., 1960a,b; De Weer, 1971) or apyrase injections (De Weer, 1971) owing to the lack of ADP and the presence of PA in the dialysis solutions. For these reasons, Beauge and DiPolo suggested that the reverse K-free effect in vanadate poisoning was the expression of the inability of ATP to accelerate the disocclusion of the E2(K)enzyme form (Fig. 1). This suggestion was confirmed by fluorescence studies with the unphosphorylated purified Na’,K’-ATPase enzyme (Karlish et al., 1979). Karlish and Pick (1981) suggested that all the effects of external K in the presence of vanadate were due to the formation of the E2(K) enzyme form after dephosphorylation. However, this explanation is not at all satisfactory; if this were the mechanism, each pump unit being dephosphorylated by KO and going through the E2(K) state should bind vanadate very tightly; the consequence for Na efflux would not be a biphasic K , response to K, but a monotonic decrease in flux as K, concentration is increased. By similar reasoning that explanation cannot account for biphasic KO effects on K influx either (Beauge st al., 1980). A more complex interaction of K ions with external pump sites remains as the more plausible explanation; this is further supported by recent findings (J. Sachs, unpublished: L. Beauge and G. Berberian, unpublished) than in vanadate-

146

LUIS BEAUGE

poisoned red cell there is no biphasic Rb, effect on Rb influx in the absence of external Na. iii. Oligomycin. Oligomycin blocks the conversion of an ADP-sensitive phosphoenzyme into a K-sensitive one (Fig. 1); in erythrocytes it inhibits Na-Na exchange, Na-K exchange, uncoupled Na efflux, and pump reversal, indicating not only that forward and backward reactions are sensitive to this agent, but also that the phosphoenzyme transformation is likely to be part of the cation translocating process. On the other hand, oligomycin increases the phosphoryl group exchange between ATP and ADP catalyzed by the Na+,K+-ATPaseenzyme (see Glynn and Karlish, 1976; Robinson and Flashner, 1979). Baker et al. (1969) reported that 12.5 pg of oligomycin per milliliter in the seawater (about 30 p M ) produced over a period of an hour a slow reduction in the efflux of Na into 10 mM K(Na)-ASW. The reason for this slow time course is not clear; one possibility is that the axon membrane is not very permeable to oligomycin; another is that the observed effect was indirect through a reduction in the levels of ATP. On the other hand, De Weer et al. (1982) found that at 10 p M concentration in the dialysis solution oligomycin failed to affect Na efflux. This contradicts the observation that the Na+,K+-ATPaseactivity present in membrane fragments from squid nerves has about the same sensitivity to oligomycin as the purified Na+,K+-ATPasefrom pig kidney (Table 11). Had De Weer et al. (1983) tried higher concentrations, they might have found inhibition of Na efflux in their experiments. The important points, and the reason why these data on oligomycin were included here, is that this compound and vanadate constitute two cases where the apparent affinity of an inhibitor acting intracellularly is much lower when perfused into the axon than when tried in a broken membrane preparation. Curiously, for inhibitors acting extracellularly (ouabain, strophanthidin) the reverse is the case (Table 11). With these observations in mind, it seems that caution should be exerted not to overestimate negative results nor to put much emphasis on the comparison of apparent affinities between inhibitors in in situ and in uitro experiments. e . Effects of Intracellular M g 2 + . In a series of elegant experiments performed mainly to determine the free-Mg concentration levels in squid nerves, De Weer (1976) obtained information on the influence of the Mg” concentration on the total Na efflux in injected axons. The optimal Mg2+ concentration was about 10 mM; as expected (Skou, 1957; see also Flatman and Lew. 1981), below and above those levels the efflux of Na was reduced. As De Weer pointed out, there are some uncertainties in these types of experiments, especially at high Mg2+ concentrations, not only

147

SODIUM PUMP IN SQUID AXONS

because of the possible damage due to several microinjections, but also because of changes in internal osmolarity upon those injections. Similar Mg2+ dependence curves have been found for the Na+,K+ATPase activity in membrane fragments from squid nerves (Breitweiser, 1982; M. Campos and L. BeaugC, unpublished); at high Mg2+ concentrations we have evidence indicating that the results are indeed a combination of Mg2+,osmolarity, and ionic strength effects (M. Campos and L. BeaugC, unpublished). The lowest Mg2+ levels obtained by De Weer were never below 400 p M . We have performed some experiments (L. BeaugC and R. DiPolo, unpublished) where the intracellular Mg2+concentration was taken practically to zero by dialyzing the axons with Mg-free ethylenediaminetet60 No-310 K - 150 C I - 1 EGTA 2 ATP - 5 PA 6 Mg - 0 EDTA 0 Mg - 2 EDTA

INTERNAL OATP-0 PA1 O M g - 2 EDTA EXTERNAL

I

I

.....*. . J

..”

VI Q

AXON

P Temp

100982 4OOurn 19°C

n

AlP I

HOURS

FIG.6. The effects of intracellular Mg on Na efflux in dialyzed squid axon in the absence and in the presence of strophanthidin. The levels of Na efflux obtained with 4 mM Mg, and no ATP are the same as those seen with no Mg, and 2 mM ATP; similar results were obtained when CDTA was used instead of EDTA. In the presence of 6 mM Mg, and 2 mM ATP the efflux of Na reaches its normal value, and it is inhibited as usual by strophanthidin [KO sensitivity is also restored (not shown here)]. During strophanthidin poisoning, removal of Mg, in the presence of ATP brings Na efflux to the values seen in the absence of digitalis and of Mg, and/or ATP. In summary, ATP is unable to promote any efflux of Na, under strophanthidin poisoned or unpoisoned conditions, if intracellular Mg is not simultaneously present. Unless otherwise stated, all solute concentrations are given as millimolar. (From an unpublished experiment of L. BeaugC, H. Rojas, and R. DiPolo.)

148

LUIS BEAUGE

raacetic acid (EDTA) [or trui?s-l,2-diaminocyclohexane N’,N,N’,N’-tetraacctic acid (CDTA)]-containing solutions. As Fig. 6 illustrates, the levels of Na efflux in the presence of 4 mM MgClz and no ATP are not different from those obtained with 3 mM ATP in the absence of internal Mg2+.After dialysis for 2 hours, the total efflux of Na regained its normal value and KOsensitivity when ATP and MgClz were both included in the dialysis solution. Experiments like that of Fig. 4 are to this reviewer as fundamental to the understanding of Na fluxes in axons as those of Brinley and Mullins (1968). These authors have shown that in the presence of intracellular Mg?’ almost all Na efflux is ATP dependent; in our case, we have shown that in the presence of internal ATP, almost all Na efflux depends on intracellular Mg’+. This point will be dealt with again when discussing ATP-dependent Na fluxes in the presence of strophanthidin. f. Interactions of the Nu Pump with Nu, K , and ATP. The kinetic consequences on the Na-K transport of the ATP-dependent E2(K)-EIK conformational change (Post et a/., 1972) (see Fig. I ) were first worked out by Beagi and DiPolo (1978, 1979b, 1981a, 1982) in dialyzed squid axons. The model predicted that, besides their ability to dephosphorylate the phosphoenzyme, the order of effectiveness of monovalent cations as external Na pump activators should depend on (1) the stability of the EZ(cation) complex formed after dephosphorylation (the degree of stability is TI > Rb > K > NH4 > Cs > Li), and (2) the actual levels of ATP. The dependence should be such that at low ATP concentrations those cations forming a loose complex with the enzyme were expected to be better activators of the pump than those forming a tight E-cation complex. These predictions were borne out; whereas at 3-5 mM ATP the sequence was K > NH4 > Rb, with 30-50 p M ATP it changed to NH4 >> K >> Rb. Removal of internal K (or changing the Nai : Ki concentration ratio) (Fig. 7) did not modify these ATP effects; the implication is that the E,(K) enzyme state is accessible to ATP but not to intracellular Na or K. The reduction of the ATP concentration, besides leading to a drop in the rate of Na pumping, increased the apparent affinity for KOas a pump activator (Beauge and DiPolo, 1979b) (Fig. 8 ) . This was observed in axons dialyzed with and without K; and cannot be attributed to a K accumulation in the extracellular space (BeaugC and DiPolo, 1981a). Moreover, this is predicted by the ATP-dependent disoccluding model. For a reaction with the following steps

(the irreversible steps are a simplification for axons dialyzed without Pi

I

OATP-OPA INTERNAL EXTERNAL

30

I

5 0 p M ATP - 5 mM PA

I

7 0 N o - 310K 1OK

-

15

-

30

x)

-

5

4e 0

0

*em

0

-

f -"

-

odb

Y

y

3 m M A T P - 5 m M PA

70No-OK

0

20 -

10-

OOO

.y

0

-

0

0 0 0 10

g

-

S-

0

0-0

*'*,

b

0

AXON

Qx=ooo

Temp

2

160578-A

8

8" 1

%.

0

0

saoo

0

0 .

O -0

o

0

+ - - e

O@" 0

17. 5.C

1

I

I

I

3

4

S

6

I 7

HOUT,

FIG.7. The effects of internal K at different internal ATP concentrations on the external cation activation of N a efflux in a dialyzed squid axon. The solute concentrations are given as millimolar. (Taken from BeaugC and DiPolo, 1979b.3

0

10

20 External

30 K

40

50

(mM)

FIG.8. Activation curve of Na efflux by external K in axons dialyzed with 5 mM ( 0 )or 50 y M ATP (0) together in both cases with 5 mM phosphoarginine. In the inset the effect of K is expressed as percentage increment in Na efflux over the levels of flux in K-free NaASW; the values of flux at 50 mM K were taken as 100%. The axons were dialyred with 310 mM K, and 60 mM N a , , but similar results were obtained in axons dialyzed without internal K. (Taken from Beauge and DiPolo, 1979b.)

150

LUIS BEAUGE

and Ki) the rate equation becomes

v =

(K,lK,+)’

+ 2KJK;

Et

+ [Ilk2 + (k-3 + k4)/k3k4 + l/kJ

Considering that k3 is the rate constant stimulated by ATP, it follows that a reduction in the concentration of the nucleotide will produce a decrease in the pump rate and an increase in the apparent affinity for KO.Similar behavior was found by Eisner and Richards (1981) in resealed human red cell ghosts. Another important kinetic consequence, not obvious at first sight, is that the reciprocal plot of v/Ko vs K, will yield straight lines only in the trivial case where the resulting sums and products of the rate constants between parentheses is close to 1 (Beauge and DiPolo, 1983). The reduction in the ATP concentration was followed by a decline in the K-free effect; this was not due to, or at least was not very much affected by, a KOaccumulation, for it appeared in axons dialyzed with and without K, (BeaugC and DiPolo, 1978, 1979b. 1981a). In low-ATP axons, when Rb was used as K substitute, in most instances the efflux of Na in 10 mM Rb was not different from that in K-free seawater (Fig. 7). However, reverse K-free effect, even with ATP concentrations as low as 20 p M , was never found. An ATP-K antagonism was also observed to take place at intracellular sites (BeaugC and DiPolo, 1979b, 1981a). However, in this case internal Na played an important role, to such an extent that, rather than speaking of an ATP-K, antagonism, one should refer to it as an (ATP + Na,)-K counteraction. This interrelationship is shown in Fig. 9 constructed with data taken from BeaugC and DiPolo (1981a); Ki has an inhibitory effect on Na efflux that is counteracted by both Na, and ATP. This can also be stated as: (1) K, reduces the apparent affinity of the Na pump for ATP, and this is antagonized by Na,; or ( 2 ) Ki diminishes the apparent affinity of the Na pump for Na,, and this effect is counteracted by ATP. These results are consistent with a kinetic scheme where the E,(K) occluding conformation can be reached also by K acting on intracellular pump sites before rephosphorylation (see Fig. 1); Na, pushes the equilibrium toward E,Na, whereas ATP brings the E2(K)state back into the EIK form. These results can also be explained on the basis of the ideas of Skou (1974), with ATP modulating the interconversion between E,Na and E,K. However, to follow the proposal of Skou demands an extra ATP site; although possible, this is less economical. 2 . INFLUXOF POTASSIUM

The uptake of potassium, the other half of the reaction cycle in the NaK active exchange mechanism, has not been as extensively studied as the

151

SODIUM PUMP IN SQUID AXONS

0

0.05 rnM ATP

3 mMATP

I

20 Internal

I

I

I

LO No concentration

I

60

A

(mM)

FIG.9. Interactions of ATP, Na,, and K, and their effect on Na efflux in dialyzed squid axons. The figure is a plot of the ATP-dependent Na efflux at 0 K, over the ATP-dependent Na efflux at 310 mM K, as a function of the Na, concentration at two concentrations of ATP. At 3 mM, internal K becomes inhibitory only at Na, concentrations below 30 mM. With 0.05 mM ATP, internal K is already inhibitory over 70 m M Na,. For a given Na concentration the sensitivity of Na efflux to K, increases when ATP is reduced. Each value is the mean 2 SEM. The number of axons is given in parentheses. (Data are taken from Beauge and DiPolo, 1981a.)

efflux of Na. The main reasons have been technical ones. Before the development of the internal dialysis technique there was no way to utilize the same axon as its own control in any test, something that was routine in efflux experiments; the usual approach, then, was to compare paired axons from the same squid. On the other hand, even with the dialysis method, a carefully controlled influx determination is not easy to perform, and additional sophistication in the technique is required (Brinley et al., 1975; DiPolo, 1979). a. Effect oflnhibitors. Hodgkin and Keynes (1955) found that in Sepia axons the influx of K from 10 mM K-Na ASW had a Qlo of about 3.3 (between 1.3"C and 20°C) and was 85% inhibited by 0.2mM dinitrophenol and 2 mM cyanide; 75% inhibition by 1 mM cyanide was reported in intact axons for the total K entry (Sjodin and BeaugC, 1967, 1968)and 53% inhibition by M ouabain (Baker et al., 1969). In axons subjected to internal dialysis, M strophanthidin or ouabain resulted in about 5060% reduction with an external K concentration of 10 mM (Brinley and Mullins, 1969) or 5 mM (De Weer et al., 1979). Neither of the inhibitors just mentioned had any effect on the passive movements.

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LUIS BEAUGE

6. EfJect i f A T P und Other Phosphorylntrd Compounds. In axons dialyzed with Na,, ATP stimulates the influx of K (Mullins and Brinley, 1969). The data are difficult to analyze, owing to the large dispersion from axon to axon and to undefined curves relating fluxes to ATP concentrations even in the same axon. Nevertheless, the authors claimed a KIl2for ATP of about 30 p M , that is, the same Kl,? reported for the ATP stimulation of Na efflux (Brinley and Mullins, 1968). An interesting observation made in the same paper is that all ATP-stimulated K uptake was abolished by strophanthidin. From the results in the preceding section, one wonders whether and why metabolic inhibitors yielded a larger inhibition of K entry than ouabain or strophanthidin; if that was indeed the case, the dialysis experiments with ATP-strophanthidin indicate only that ATP depletion was not the reason. Other phosphorylated compounds tested by Mullins and Brinley werc AP, AcP. phospho(eno1)pyruvate (PEP), and deoxy-ATP; as happened with Na efflux, only ATP and deoxy-ATP were able to stimulate a strophanthidin-sensitive uptake of K. c . Dependence on External K . There are two papers on this subject, and they are conflicting. On the one hand, utilizing the axoplasm extrusion technique in paired axons, Baker et al. (1969) found that the ouabainsensitive K influx as a function of K, concentration (measured at 3-10 and 30 mM KO) had a tendency toward saturation. On the other hand, in dialyzed axons Mullins and Brinley (1969), on the basis of four K concentrations tested, reported a linear relationship between the ATP-dependent K uptake and external K concentration up to 20 mM (the highest KO investigated). In one of his reviews, Mullins (1979) admits being prejudiced in favor of the dialysis data on the basis that measurements can be made using the same axon as a control. However, at the time his experiments were done the dialysis method had not yet been refined for influx measurements, and a close look at the data finds them to be rather disperse. The final answer will have to await further experiments utilizing the newly developed dialysis techniques and considering many more K concentration points. Mullins and Brinley (1969) also reported that, in the presence of 10 mM K,, replacing external Na with Li or choline did not appreciably modify the ATP-dependent K uptake. The results with Li are not surprising because Baker et d . (1969) have shown Li to be almost as powerful as Na in inhibiting the K , activation of the Na pump. On the other hand, the replacement of choline by Na, produced a noticeable increase in the apparent affinity of the external pump sites for K (Baker et al., 1969); with a K l i 2for K , of 10 mM in Na seawater one would expect choline to induce a large increase in K uptake. It is unfortunate that Mullins and Brinley did

SODIUM PUMP IN SQUID AXONS

153

not try the strophanthidin sensitivity in choline seawaters in order to rule out any possible change in the baseline of nonpumped K fluxes. d . Eflects of Internal Nu. In 1955 Hodgkin and Keynes observed that the total K influx had a transient increase following electrical stimulation of the axons, returning to its resting levels with a time constant of about 2-3 hr. They attributed these findings to changes in Na, but did not pursue the matter further. In 1968 Sjodin and BeaugC reported that an increase in Na, from 23 to 122 mrnol per kilogram of axoplasm brought about a doubling of the cyanide-sensitive K uptake. The actual Na, dependence on M influx was worked out by Mullins and Brinley (1969) in dialyzed axons. They found that Na, stimulated the ATP-dependent influx of K in a saturable fashion. The highest Na, concentration used was 80 mM; taking that concentration as saturating, the Kli2for Na, becomes about 20 mM.

3. N a : K COUPLING RATIO The coupling ratio has to do with the stoichiometry of the Na-K active exchange. At present there is no way of verifying the behavior of a single pump unit during a single cycle. This is a very important point, for it means that, although we can come up with integer numbers, the coupling ratio will always be a statistical average; the extrapolation to the operation of a unit pump during a single cycle is just by inference. Another important aspect derives from the fact that we are dealing with charged species. For a fully coupled system ( I : 1 ratio) the exchange is electroneutral and there is no need for extra ionic fluxes to preserve electroneutrality. On the other hand, for a coupling ratio different from 1 : 1 (electrogenic pump), electroneutrality requires that some ionic species, those with the largest mobilities, will have to move across the cell membrane to keep the balance of charges. They will move through a resistive pathway, and this ionic current will contribute to the electrical membrane potential. The “extra” fraction of the Emset by the pump will constitute an extra driving force for ion movements and will have to be taken into account. In this type of system, measurements of unidirectional fluxes of the actively transported species may not give the real coupling ratio. This is particularly true if the membrane has a high permeability for the lesser pumped species: and this happens to be the case for K ions in excitable cells. The magnitude and sign of the En, set u p by fully or partially uncoupled Na-K pump will be a function of ( I ) the degree of uncoupling, ( 2 ) the direction of the uncoupling (more ions moving outward or vice versa), (3) the magnitude of the fluxes, and (4) the membrane resistance (and also

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membrane capacitance in non-steady-state conditions). An elegant theoretical treatment of this problem has been developed by Frumento (1965); the reviews of Thomas (1972) and De Weer (1975) can also be consulted. a . From Flux Measurements. In Sepia axons Hodgkin and Keynes (1955) observed that the drop in Na efflux following removal of KOwas very close to the amount of K influx abolished by the same procedure or by the use of metabolic inhibitors. Although these authors favored a 1 : 1 Na : K coupling ratio, the door was left open for ratios larger than 1 : 1, and they even considered the possibility that the linkage between K uptake and Na efflux was not a rigid one. A ratio of 2 Na expelled per each K taken up was obtained by Baker et al. (1969) on the basis of the ouabainsensitive Na efflux and K influx at constant Nq and variable KOconcentrations; that 2 : 1 value was in close agreement with ouabain-sensitive Na: K stoichiometry obtained in red cells (Glynn and Karlish, 1975; Robinson and Flashner, 1979), a preparation particularly suitable for estimates of this kind. On the other hand, taking into account the cyanidesensitive Na and K fluxes, Sjodin and Beauge (1968) found a ratio that varied with the N q concentration from a value of 1 : 1 at 20 mmol of Nai per kilogram of axoplasm to 2 : 1 at 120 mmol per kilogram. Considering the influence of N& on the ATP-dependent Na efflux (linearly dependent on Nai) and K influx (following saturation kinetics), Mullins and Brinley (1969) calculated an exceedingly variable coupling ratio that went from 1 : I at low Nai up to 5 : 1 at high Na, concentrations. However, the finding of Kracke and De Weer (1982) that in injected and dialyzed axons Nai stimulates Na efflux not in a linear, but in a saturable, fashion (see Section III,A,l,c) puts the matter in a completely different perspective. The KIlZ of 2725 mM for the Nai-activated Na efflux is very close to the K112of 20 mM for the Nq-stimulated ATP-dependent influx of K (Mullins and Brinley, 1969), suggesting that both fluxes may indeed have identical dependence on internal Na; in other words, the Na- and K-active fluxes in squid axons would be coupled with a rigid stoichiometry. b . From Electrical Measurements. A Na : K coupling ratio larger than 1 : 1 requires that the balance of charges will have to be attained by the passive exit of anions, the passive entry of cations, or both. In all cases this implies an inwardly directed ionic current through the resistive membrane pathway; this in turn will lead to a potential drop that is a hyperpolarization. In their classic 1955 paper, Hodgkin and Keynes considered the possibility of a 3 : 2 Na: K coupling ratio and concluded that for the values of Na efflux and membrane resistance in Sepia axons the pump current should be about 1.8 ,uA/cm2with a corresponding hyperpolariza-

SODIUM PUMP IN SQUID AXONS

155

tion of 1.8 mV. The contribution of the Na pump to the resting membrane potential in Loligo axons was first established by De Weer and Guiduldig (1978) measuring the changes in membrane potential elicited by 10 p M strophanthidin; at 21”C, the strophanthidin-induced membrane depolarization was about 1.2 mV in fresh axons, increased with N q , and was abolished by cyanide; no changes in membrane conductance were detected upon treatment with the cardiotonic aglycon. The estimation of the “pump current” can be made in two ways: (1) from the values of membrane resistance and the Na pump-dependent membrane potential; and (2) under voltage-clamp conditions determining the changes in the holding current upon activation and/or inhibition of the Na pump. The first approach was used by Abercrombie and De Weer (1978), considering pump current and fluxes as those inhibited by digitalis. The values for current were 1.37k0.13 pA/cm2 at 2I0C, a result consistent with a 3 : 2 Na: K coupling. In axons with normal ADP content, that ratio remained constant upon changing the KOconcentration up to 20 mM; unfortunately the effects of Nai were not investigated. Another interesting observation was that in axons with elevated ADP content the efflux of Na in K-free ASW was higher than in axons with normal ADP and was not modified by increasing KOconcentrations. However, the pump current did increase with KO,but reaching values about 30% of those seen in normal axons. This behavior is to be expected if increasing KOproduces a shift from an electroneutral Na-Na into an electrogenic Na-K exchange. Using a lownoise voltage-clamp system, Rakowski and De Weer (1982) found that at 21°C and -58 mV membrane potential the ouabain-sensitive holding current averaged l .55 pA/cm2;compared with the ouabain-sensitive efflux of Na, this yielded a 3 : 2 Na : K coupling ratio. Again, no effects of changing internal Na concentrations were investigated. c . Membrane Potential and Na-K Pump. Pump current can be calculated from membrane resistance ( R , ) and pump-dependent potential (Emp)provided that under the conditions of the experiment the resistance through the pump pathway (R,) is much larger than the remaining resistance of the membrane (&); or, to put it in other words, the pump must be largely insensitive to Em over the range of the membrane-potential changes (Abercrombie and De Weer, 1978). Looking at the argument the other way around, an electrogenic pump is in theory expected to be influenced by the membrane potential; the question is, over what range of potentials? The answer to this comes from consideration of the pump electromotive force, Pemf(the difference between the free energy released by ATP hydrolysis and the work required to move the ions), and the

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actual values of Em (Hodgkin and Keynes, 1955; Abercrombie and De Weer, 1978). An oversimplified calculation of the free energy of ATP hydrolysis suggests that the difference between Pen,,.and Em is large enough to allow changes in Em without noticeable effects on pump rates (see above references). The actual experimental data seem to agree with that conclusion. No changes in the ATP-dependent Na efflux or K influx were observed in axons dialyzed with normal ATP content subjected to hyperpolarizations (passing electric current) up to 40 mV or to depolarizations (removing K i ) up to 60 mV (Brinley and Mullins, 1974). The lack of effect of membrane depolarizations on the ATP-dependent and the ouabain-sensitive Na efflux was observcd in axons dialyzcd with 3 mM and also with 30-50 p M ATP; in these cases dcpolarizations were attained by removing Ki or by “pulses” of 450 mM seawater (Beaugt and DiPolo, 1981a). It is possible then that over a wide range of En, the Na pump behaves as a constant current generator; however, other undetected changes (modification of the Na: ATP ratio, for instance) have not yet been excludcd.

B. Na-Na Exchange 1. AS

A

PARTIAL REACTIONOF

THE

PUMP CYCLE

u. Sodium-Sodiirm ExchangP Fluxes. The first evidence on the subject was provided by the work of Caldwell et al. (1960a,b). On the one hand, the efflux of Na from cyanide-poisoned axons not only progressively declined with time and lost its KO dependence, but external K became inhibitory, and a reverse K-free effect appeared; the injections of ATP restored the levels of flux but not their K, sensitivity, which was reinstated after injections of PA. On the other hand, 0.2 mM 2,4-dinitrophenol (DNP) at pH 8.0 caused a rise of Na efflux into K-free solutions but produced no change in K-containing seawaters, at the time that K influx was reduced to half. In 1969 Baker et ul. repeated these observations and showed in addition that (1) the Na efflux seen in cyanide- and dinitrophenol-poisoned axons in K-free solutions was abolished by ouabain, and (2) it was accompanied by ouabain-sensitive N a influx of the samc magnitudc; i.e., the Na pump was engagcd in a Na-Na exchange of 1 : 1 stoichiometry. The electroneutral properties of this exchange mode are further supported by the fact that injected axons with elevated ADP content and bathed in K-free sodium seawater have a large cardiotonic steroid-sensitive Na efflux that has no effect on membrane potential (Abercrombie and De Weer, 1978). Another very important observation was that, in the absence of Na,, partially poisoned axons responded to external K in a way similar to that in unpoisoned cells. This indicates that

SODIUM PUMP IN SQUID AXONS

157

the ability of K , to stimulate the Na pump is still present under these conditions (see also Abercrombie and De Weer, 1978; and De Weer et al., 1979). Baker et a / . (1969) considered that the Na-Na exchange was increased when the (ATP):(ADP) (P,) ratio was decreased (see also Caldwell, 1968). They proposed some possibilities to account for these Na movements, one of which involved a phosphoryl group transfer between ATP and ADP as had been found to be catalyzed by isolated N a + , K + ATPase enzyme (see Fig. I). By that time, a ouabain-sensitive I : I NaNa exchange requiring ATP and ADP, but presumably with no ATP hydrolysis, had also been described for human red cells incubated in K-free conditions (see Glynn and Karlish, 1975; Robinson and Flashner, 1979, for references). In 1971, utilizing injected axons, De Weer went through a long list of experiments designed to alter the cell biochemistry; the main finding was that all maneuvers utilized that brought the ATP : ADP ratio to the values reached by cyanide or dinitrophenol treatment produced alterations in the KOdependence of Na efflux similar to those seen with the inhibitors. In addition, inorganic phosphate, although it inhibited the Na pump, did not alter its response to external K. The conclusion that the ATP : ADP ratio was at least one of the key factors governing the shift between Na-K and Na-Na exchange modes of the Na pump seemed inescapable. The Na-Na exchange through the Na pump has been also studied in dialyzed axons. Figure 10 shows results of one of the experiments of De Weer et al. (1979) where Na-Na and Na-K exchanges were followed in the same nerve by simultaneous measurement of Na and K influx. In this particular experiment, the addition of 5 mrM ADP to the existing 5 mM ATP in the dialysis solution resulted in a large increase in Na influx (65% ouabain sensitive) and a very small reduction in the total K uptake. This was intcrpreted in terms of a Na-Na exchange arising "in uddition t o , rather than at the expenses of, a pre-existing Na-K exchange." The scheme of Fig. I , adapted with some constraints, could account for those data as well as for other features of the Na-Na and Na-K exchanges seen previously. Unfortunately, there is a lack of relevant information in the cxperimcnt of Fig. 10; for instance, there are no details of the dialysis solutions used, especially regarding MgClz concentration, and the rationale for the high nucleotide concentrations is not given. In addition, the author offers no explanation for the sizable ADP-stimulated ouabain-insensitive Na influx (about 35% of the total increase). Furthermore, it is an experimental fact that when ADP is added to the reaction mixture at the same concentration as ATP (both in the millimolar range) the Na+,K+ATPase activity is about 40% inhibited (M. Campos and L. Beauge, unpublished observation). It is intriguing that the K influx in Fig. 10 is so

Luis BEAUGE

158 INFLUX

pmol c I-2sec-'

I OUAEAIN A T P I A T P + ADP

3(

00 00 00

.

0°K

0 0

O " N ~ influx

0

0

influx

0

0 00 0

2c 0

0

0

0 00

00

0

000 0

1C

a 0"' ' 0

(

2

1 HOURS

FIG.10. Influx of 22Naand 42Kinto squid axons. The axon was dialyzed with a solution containing 5 mM ATP. At zero time, it was bathed in artificial seawater containing 22Na(425 mM) and "K (5 m M ) , and influx of each isotope into the internal perfusate was monitored with a two-channel gamma spectrometer. At 50 min the internal solution was changed to one containing, in addition to ATP, 5 mM ADP. At 90 min, lo-' M ouabain was added to the seawater bathing the axon. Temperature, 16°C. (Taken, with permission, from De Weer ei al., 1979.)

little affected when 5 mM ADP is included in the dialysis solution. There is also an important point regarding inorganic phosphate. In otherwise normal axons, De Weer (1971) found that injected Pi had no effect on the K sensitivity although there was a general inhibition of Na efflux. It is unlikely that, as suggested by Caldwell (1968), the available free energy from ATP hydrolysis determines the Na-Na exchange mode of the Na pump. However, the kinetic scheme of Fig. 1 implies that an increment in Pi will lead to accumulation of the E2PK andlor EzPNa intermediates (depending on the external ionic composition) and to an increase in Na influx. In this regard it is important to notice that Post et al. (1975) found that Pi stabilizes the phosphoenzyme formed from ATP, probably through the stabilization of an EPi-Na complex. This Pi effect was antagonized by ATP and by sodium. On the other hand, Garrahan and Glynn (1967) found that in human red cell ghosts the Na-Na exchange is lost if Ki is replaced by N q , an effect that could be reversed by high phosphate concentration. It would be interesting to follow the effects of Pi on Na-Na exchange in axons with low ATP :ADP ratio (low ATP and high ADP), or conversely, the effects of ADP in axons dialyzed with low ATP with and

SODIUM PUMP

159

IN SQUID AXONS

without Pi. Finally, despite the impressive evidence in favor of the ATP : ADP ratio in determining the Na-Na exchange (Baker et al., 1969a; De Weer, 1971), one still wonders whether this is the only element to be taken into account. In fact, there is still no experiment showing a large Na-Na exchange in axons dialyzed with ATP, ADP, and Mg concentrations similar to those seen in partially poisoned cells; the concentration of these three compounds used by De Weer et al. (1979) are far from those expected in cyanide- or DNP-treated nerves. b . ATP-ADP and Nu-Nu Exchange: The Stoichiometry Problem. The phosphoryl group exchange between ATP and ADP was investigated in axons dialyzed under conditions where a Na-Na exchange (taken as the ouabain-sensitive ADP-dependent Na influx) took place: among the usual components the axons were dialyzed with 100 mM Na, 5 mM ATP, 5 mM ADP, and I5 mM MgCI2. Against the original predictions the results were negative and no ouabain-sensitive ATP-ADP exchange was detected (De Weer et al.. 1983). This is in sharp contrast with the results of Cavieres and Glynn (1979) and Kaplan and Hollis (1980) in human red cells. De Weer et al. (1982) explained their results on the basis of an ordered binding (and release) of the internal ligands ATP, Mg, and Na, in such a way that the partial biochemical reaction would be ATP

+ El

ATP . El

+ATP #

Mg

El

. Mg

#

Mg . E,P . Na . ADP

Na Mg . E I P . Na + M g .

1

ADP

E 2 P . Na

This scheme not only accounts for the discrepancy between Na-Na and ATP-ADP exchanges observed by De Weer et a f . (1979, 1983), but suggests that the well-documented Mg inhibition of the ATP-ADP exchange reaction (see BeaugC and Glynn, 1979, for references) can occur at the same site at which Mg activates the reaction (slowing the release of ATP). However, a second site for Mg inhibition must exist, for it has been found that high Mg2+ levels also inhibit the Na-Na exchange (Flatman and Lew, 1981), and that observation cannot be explained by the scheme shown above. Actually, if one looks at the data of Flatman and Lew (1981) and extrapolates it to the experiments of De Weer et al. (1979), their Na-Na exchange should have been inhibited by the high Mg concentrations. One does not know how legitimate that kind of extrapolation is, but at any rate an Mg-dependence curve for Na-Na exchange in squid axons seems to be in order. Regarding the stoichiometry between Na-Na and phosphoryl group

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exchanges it must be remembered that not only is ATP-ADP exchange possible (or even enhanced) when the Na-Na exchange is blocked (as in the case of oligomycin treatment) (see Beauge and Glynn, 1979, for references), but that ATP-ADP exchange has been found in the absence of external Na (Kaplan and Hollis, 1980). Even without considering an ordered binding for the intracellular ligands, the evidence just presented proves that any attempt to find a stoichiometry between fluxes and biochemistry in this partial reaction would be futile. 2. Na-Na EXCHANGE AS A COMPLETE P U M PCYCLE

Some years ago (Beauge and Ortiz, 1971; Beauge and Del Campillo, 1976; Glynn and Karlish, 1976), it was suggested that external Na ions could stimulate the whole cycle of ATP hydrolysis by exerting a K-like action. The suggestion was made on the basis of ( I ) the poor selectivity of the external pump sites for monovalent cations; (2) the fact that ATPdepleted cells had a ouabain-sensitive Na,,-dependent Na efflux too high for the expected ADP levels; and ( 3 ) thc fact that even in the absence of internal ADP there was an Na,-stimulated ouabain-sensitive Na efflux. In 1980 Lee and Blostein provided convincing evidence that external Na can indeed exert a K-like effect both on Na efflux and ATPase activity. In squid axons the available data are confusing, and evaluation of the data is complicated by the fact that different dialysis solutions were used and different values for fluxes were reported. In axons dialyzed with Na + K and bathed in 10 mM K-NaASW, Brinley and Mullins (1968) found an ATP-stimulated Na influx of 28 pmol cm-? sec-l coexisting with an efflux of Na of 48 pmol cm-? sec-I; considering the relatively large errors the authors admit for their influx determinations, this could favor some Klike action of Na,. However, interpretation of the data is complicated by additional observations for which there is no satisfactory explanation: ( I ) Nai stimulated Na influx in the absence of ATP; ( 2 ) ATP stimulated Na influx in the absence of Nai; this latter flux was about 14 pmol ern-.? sec-l with no measurable Na efflux: and (3) in intact axons these authors found no effect of strophanthidin on Na influx. In axons dialyzed with 5 mM ATP, 100 mM Na (and presumably 15 mM MgC12 and bathed in 5 mM K-NaASW), De Weer et NI. (1979) found an almost nonexistent ouabain-sensitive Na influx over a total uptake of about 14 pmol cm-2 sec-'. Finally, Beauge and DiPolo (unpublished) observed that ATP had no effect on N a influx in the absence of Nai; in the presence of 60 mM Nai and in 10 mM K-NaASW with tetrodotoxin (TTX), 2 mM ATP stimulated an extra Na influx of about 18 pmol cm sec-' that was about 40% abolished by strophanthidin. However, the

S O D I U M P U M P IN SQUID AXONS

161

effects of external K (which should be inhibitory by competing with Na,) were not investigated. In summary, although there seems to be some evidence that in squid axons Na, can also exert a K-like action on the Na pump, the matter is by no means settled. If this mode operates, dialyzed axons appear as the ideal tool to study the stoichiometry of the fluxes.

C. Uncoupled Na Efflux

Squid axons bathed in solutions lacking both Na and K show a cyanideand ouabain-sensitive Na efflux that is actually larger than that seen in normal seawater. This Na efflux can represent (1) an effect of K leaking out of the cells and recaptured by the Na pump (which in the absence of Na,, has a very high affinity for KO),or (2) a truly uncoupled Na pump, that is an Na efflux not activated by, nor exchanged for, any external cation. We have performed a few experiments (Beaugk and DiPolo, unpublished) following changes in Na efflux upon Na, and K, removal in axons dialyzed with solutions containing ATP, PA, no ADP, and without K i . The response was not constant (sometimes there was an increase and other timcs a dccrease) nor was it reproducible within the same axon; in addition, when present, the magnitude of the increase never reached the values seen in axons dialyzed with K i . If these preliminary observations are correct, the implication is that most of the Na efflux from axons bathed in Na, ,K,-free solutions is due to K leaking out and being recaptured by the Na pump. There are other observations that support this view: if an uncoupled Na pump in axons behaves similarly to that present in red cells (Glynn and Karlish, 1976) or is affected by Na, in the same way as the dephosphorylation of the phosphoenzyme formed from ATP (Beauge and Glynn, 1979), one would expect Na, to be inhibitory at low concentrations and that Na efflux be restored, at least partially, at high Na, levels. The experiments of Baker et al. (1969) show that this is not the case; increasing Na, up to 450 mM (in choline or sucrose mixtures) resulted in a monotonic decline in the ouabain-sensitive efflux of Na (Fig. 11). A way to test the hypothesis of K recapture would be to determine the effects of ouabain on K efflux in axons immersed in Na,,K,-free solutions under voltage-clamp conditions (ouabain should increase K efflux in proportion to the efflux of Na it abolishes). To preserve electroneutrality, an anion efflux must accompany the exit of Na regardless of the mechanism of the Na efflux (K recapture or true uncoupled). There is no systematic study on anion efflux under these conditions in squid axons, but it has been done in another preparation. Baker (1964) found an amino acid efflux (a mixture of aspartic and glu-

162

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1

0.002

Na Dextrose

I

1

I

0 10

02

04

08

06

I

06 04

J

I

08 02

10 0

FIG.1 I . Reduction in ouabain-sensitive Na efflux into 0 K (dextrose) ASW on replacing the dextrose by Na in an injected squid axon. Axon diameter is 860 Fm;temperature, 18°C. The curve drawn through the points is a section of a rectangular hyperbola of the form M :MJ( I - C / k )where M is the Na efflux at any particular cation concentration (0.M , is the Na efflux in the absence of added cation, and k is a constant for any particular cation. M, was taken as 0.00185 and k as 0.15 (69 m M ) . (Taken, with permission, from Baker et al., 1969.)

tamic acids) from crab nerves immersed in Na, ,KO-freemedia; that efflux was increased with increasing Na,, was abolished by ouabain, and was reduced by the presence of a monovalent cation activator of the Na pump in the external solution. The reduction effectiveness of equimolar concentrations followed the sequence TI > K > Rb > Cs > Na > Li > choline

=

sucrose

that is, the same sequence as for the activation of the Na pump. The data on these monovalent cation effects could be fitted by rectangular hyperbofor K was 0.8 mM and for Na, 50 mM. On this basis one could las; the K112 expect about 10% of amino acid efflux remaining in K-free Na seawater (450 mM Na). However, the actual figures at that Na concentration have not been obtained. Although a direct extrapolation from crab to squid is not possible (especially from a quantitative viewpoint), it is interesting that the residual value of the Na,-inhibited ouabain-sensitive Na efflux in squid axons immersed in K-free solutions is also about 10% (Fig. 11). How much of that remaining Na efflux is uncoupled, K recapture, or a K-

SODIUM PUMP IN SQUID AXONS

163

like action of external Na is not known; if it were accompanied by an ouabain-sensitive amino acid (or anion) efflux, the last possibility (K-like action of Na) would be ruled out. D. K-K Exchange and Pump Reversal

There is no evidence in the literature concerning these two types of fluxes through the Na pump in squid nerves. In axons dialyzed without internal Na, Mullins and Brinley (1969) reported an ATP stimulation of K influx; this is unlikely to represent a K-K exchange because the axons were also dialyzed with PA, and it is not easy to conceive that enough Pi had accumulated to support any K-K exchange; on the other hand, no effects of inhibitors were tested. The proper experiment to test this would be to dialyze axons without N q and Na, and follow the effects of P, + ATP (or a nonhydrolyzable ATP analog) on the K fluxes in the absence and in the presence of ouabain. An ATP-stimulated Na influx in the absence of N q has been observed by Brinley and Mullins (1968). Again, there was also PA in the dialysis solutions; although some buildup of ADP and Pi near the inner membrane cannot be completely disregarded, other types of experiments are needed to study the matter. In this case the proper protocol would be to dialyze the axons without Nq and with full K, and to have them externally perfused in K-free Na seawater; the effects of ATP, ADP, and Pi, together and separately, on the ouabain-sensitive K efflux or Na influx should then be tested. E. Na Fluxes in the Presence of Cardioactive Steroids

The use of cardioactive steroids in the separation of pumped from nonpumped Na-K fluxes relies on the assumption that these agents fully inhibit the Na pump but do not modify or induce other mechanisms of ion movements. In 1968 Brinley and Mullins reported the puzzling observation that, in dialyzed axons, removal of ATP brought the Na efflux to very low values (approximately 1 pmol cm-2 sec-' ), whereas strophanthidin poisoning did not. In addition, in those axons with ATP already washed away, strophanthidin actually increased Na efflux and the steady levels of efflux attained with the digitalis were independent of the ATP concentration. As these results did not seem to be the consequence of an increased membrane permeability to Na, the authors concluded that strophanthidin induced an efflux of Na. The data of Brinley and Mullins (Fig. 8 in their 1968 paper) are straightforward and convincing; other experiments of a

164

LUIS BEAUGE

similar type (Beauge and Mullins, 1976) are not as clear owing to unstable fluxes and solution changes made many times before the steady-state value could be determined accurately. There are two other reports on increase in Na cftlux upparcntly mediated by cardiotonic steroids (Dc Weer, 1971; Baker and Willis, 1972), but in these cases there were transient responses that were not always reproducible. The matter was reinvestigated by Beauge and DiPolo (1981b). One of the observations of Brinley and Mullins (1968) was consistently reproduced; the other failed to appear with the same consistency: thc efflux of Na in strophanthidin-poisoned axons dialyzed with high concentrations of ATP was always larger than the efflux of Na in ATP-depleted axons; on the other hand, 10 p M strophanthidin was never able to stimulate any Na efflux in the absence of ATP (Fig. 12) over a whole variety of experimental conditions tested (with and without K i , KO,Mg,, Ca,, C q ) . A possible explanation for the discrepancy is the use of impure isethionate in the experiments of Brinley and Mullins (see also Section III,A, I ,a); if that is 60 N a - 310 K - 100 C I - 1 EGTA 0 ATP - 0 P A I 2 AMP-PCP I 2 PI 10 K ASW

INTERNAL EXTERNAL

2ATP - 5 PA OAMP-PCP 0 PI em

-"

0,

AXON 2 2 0 5 8 0 0 = LlOpm

N

-B

10pM stroph

0

"

1

HOURS

2

3

FIG. 12. The effect of ATP and the nonhydrolyrable ATP analog AMP-PCP, alon or in combination with P , . on the efflux or Na in dialyzed squid axon externally perfused with strophanthidin-containingseawater. Note the failure of 2 mM AMP-PCP, alone or in combination with 2 rnM P,. in promoting any efflux of N a above the base line. At the end of the experiment the usual sensitivity to ATP is observed. Unless otherwise stated, all concentrations are expressed a s millirnolar. 'The axon diameter is 410 ym: temperature, 17°C. (Taken from Beauge and DiPolo, 1981b.)

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the reason, it would be extremely important to find the compound that, acting from the inside and in combination with external strophanthidin, can stimulate Na efflux. The ATP-dependent efflux of Na remaining in strophanthidin occurs in exchange for external Na on a 1 : 1 basis. This Na-Na exchange has the following properties (BeaugC and DiPolo, 1981b): (1) it is stimulated by ATP with low apparent affinity ( K l l zabout 200 p M ) ; (2) adenylyl(P,ymethy1ene)diphosphonate (AMP-PCP), without and with P, cannot be used as an ATP substitute: ( 3 ) it is not modified by removing K,, , K , , Ca,,, Mg,, , or by increasing Cai : (4) it is not a part of the Cl- and Na-coupled transport, for it is observed at Cli concentrations, where that transport is fully inhibited (Russell, 1979); ( 5 ) it is not affected by M vanadate; (6) intracellular Mg is an absolute requirement (Fig. 6). There are two basic questions regarding the matter. One has to do with the reasons for the ATP requirements; the other, a crucial one, deals with the existence of this particular Na-Na exchange in unpoisoned axons. Let us begin with the second question: Is this Na-Na exchange induced by strophanthidin? The only argument in favor is shown in Fig. 5 in the work of BeaugC and DiPolo (1981b); in that figure, in the presence of 100 mM K O , the removal of Na, produced no change in Na efflux in the absence of strophanthidin but brought a drop in its presence. This behavior is as expected for steroid-induced flux. As has been pointed out, that experiment was not an ideal one because ( I ) the values of the Na,-dependent Na efflux were on the low side, and (2) the residual Na efflux in Nafree strophanthidin was rather large. Nevertheless, the difference in response before and after poisoning requires an explanation. That observation could be reconciled with an absence of induced fluxes provided that Na efflux is differently affected by K, in the Na and in Na-free (Tris) seawaters. This possibility was disregarded (Beauge and DiPolo, 1981b) because of the activation in Na efflux seen upon removal of Na, at low K concentrations. However, it is still possible that KO+ Tris (perhaps more noticeable at high K,) has some inhibitory action on the Na pump, which at low K concentrations is counteracted by the stimulation due to the removal of Na,. If 100 mM K, is not fully saturating in Na-ASW (and 50 mM certainly is not) when Na, is replaced with Tris, a small inhibition due to K, + Tris could be balanced out by a small activation duz to Nafree conditions resulting in no net change. The possibility of detection is further complicated by the high levels of Na efflux in these cases. In fact, if one looks carefully at Fig. 5 of BeaugC and DiPolo (1981b) (and this was also seen in the other experiment performed) there is a tendency for the total Na efflux in Tris to decline as K,, concentration increases. A11 this

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explanation will sound somewhat ad hoc; nevertheless, a complete KO activation curve of Na efflux in Na and Tris, with and without strophanthidin, should be worked out before the final word can be said. The arguments against cardiotonic steroids-induced Na fluxes are indirect, but by no means less important. One is related to vanadate inhibition. The efflux of Na into 10 mM K-Na seawater in axons dialyzed with 1 mM vanadate and 2 mM ATP (Fig. 13) is similar to that seen in strophanthidin- or ouabain-poisoned cells. Removal of Na, brings about a reduction of Na efflux to the values observed in the presence of strophanthidin (BeaugC and DiPolo, 1981b). There are two ways of accounting for these results: (1) that the removal of Na, increases the extent of vanadate inhibition and the similarity of the drop in fluxes is just fortuitous; or (2) that there is an ATP-dependent vanadate-insensitive Na-Na exchange. If INTERNAL EXTERNAL

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FIG. 13. Effect of Na, removal on Na efflux in a dialyzed squid axon poisoned with vanadate. All solute concentrations are expressed as millimolar. Tris was used as a Na substitute in seawater. Note that the Na efflux value in the presence of Na and vanadate is not different from values seen in strophanthidin- or ouabain-poisoned axons. (Unpublished experiment of L. Beaugk and R. DiPolo.)

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the second explanation is correct, it would be extremely unlikely that both vanadate and strophanthidin, which inhibit the Na pump through completely different mechanisms, could induce a similar Na-Na exchange. Another argument against induced fluxes refers to the ATP stimulation of Na efflux in the absence of external and internal K. From experiments like that of Fig. 7, once the leak fluxes have been subtracted (see Beauge and DiPolo, 1981a), the ATP-stimulated Na efflux becomes about 2-3 pmol cm-2 sec-' at 50 pM ATP and rises to about 7 pmol cm-? sec-I at 3 mM ATP. If these Na fluxes are an expression of an Na+-ATPase activity (uncoupled or Na,-stimulated), they should be activated by ATP acting with very high affinity reaching saturation at about 1-2 p M concentration (Glynn and Karlish, 1976); this is obviously not the case in these axons. There may be three possibilities for the discrepancy: ( 1 ) despite the large amount of PA present, the ATP concentrations reaching the inner membrane are much smaller than those present in the dialysis solution; (2) the uncoupled and Na,-stimulated Na pump in squid axons has much lower affinity for ATP than in other systems; and ( 3 ) we are in the presence of two different effluxes of Na; one is the uncoupled (or Na,-stimulated) ouabain-sensitive Na pump, which is activated by ATP with very high affinity, and the other is a preexisting (not cardioactive steroid-induced) Na-Na exchange stimulated by ATP with low affinity. This last possibility excludes the strophanthidin-induced Na fluxes. Strophanthidin induced or not, the reason for the ATP requirement in this Na-Na exchange remains unknown. If PA is not an excellent ATP buffer system, some ADP (and Pi) accumulation inside the axon might be possible, providing the basis for a steroid-resistant Na-dependent ATPADP (and Na-Na) exchange. On the other hand, a regulatory role for ATP similar to that observed for the ouabain-sensitive K-K exchange in red cells (Simons, 1975) would be conceivable only if the specificity of the binding site for ATP is much higher than in red cells, because, in axons, AMP-PCP could not substitute for ATP in sustaining the Na-Na exchange. It is exceedingly interesting, and intriguing that ATP alone cannot sustain these Na-exchange fluxes, but that intracellular Mg is needed as well (Fig. 6 ) . This is additional evidence against an ATP regulatory role of the type seen in the K-K exchange (Simons, 1975) for the nucleotidestimulated disoccluding step does not require Mg (Beaugt and Glynn, 1980). A regulatory effect where the agent is not ATP, but the MgATP complex, is not inconceivable; on the other hand, one is always tempted to associate ATP plus Mg with some kind of phosphorylating process. If the strophanthidin (or ouabain)-resistant ATP-dependent Na-Na is a transport process normally present in axons, the total Na efflux (leak

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subtracted) cannot be taken as equivalent to Na being pumped out (Section IILA, 1,a). Accordingly, the apparent affinity of the h a pump for ATP obtained previously will be valid only if both Na transport mechanisms, pumped and nonpumped, have a not very different ATP affinity. In addition, this sets a much lower value for two other modalities of pumped fluxes, uncoupled and Na, stimulated, that one can anticipate will be present under K-free conditions. F. K-Free Effect

The fractional drop in total Na efflux following removal of external K has proved to be a rather variable number in intact nerves (Hodgkin and Keynes, 1955) and injected axons (Caldwell et t i ) . , 1960a; Baker and Manil, 1968; Sjodin and Beauge, 1967, 1968; De Weer, 1971) as well as in axons being dialyzed (Mullins and Brinley, 1967; Beauge and DiPolo, 1981a). An explanation for this variability has not always been possible or satisfactory. Now we are aware of certain experimental conditions that, according to the accepted kinetic scheme for the Na pump (Fig. I ) , produce “predictable” alterations in the pump response to external K and Na; nevcrtheless, many unknown elements still remain in the picture. The cases where we think there is a reasonable explanation are listed below. 1. The ATP : ADP ratio. As proposed by Baker rt al. (1969) and De Weer (1971, 1975). a rcduction in the ATP: ADP ratio could account for an apparent loss of KOsensitivity by the N a pump. One can visualize that, with high levels of ADP, the ATP-ADP exchange reaction and its translocational counterpart, the Na-Na exchange fluxes, will be optimized; upon removal of K,, the system would lose its active Nai-K,, transport capability but would cngage in a Na-Na cxchange mode. Howcvcr, in axons dialyzed with high AT/‘ lcvr/,s (see Section IIl,B, I ,a), an ADP-stimulated Na-Na cxchange might occur not instead of, but in (rrlclitiori to, a preexisting, and not changing, Nai-K,, transport. De Weer r t t i l . (1979) have worked out a kinetic scheme that can account for their observations and in addition predicts a reversal K-free effect under certain conditions; but, as will be seen below, the reversal effect observed in poisoned axons may have a different explanation. 2. Changes in the ATP levels in the absence of ADP. In axons dialyzed with 5 mM PA and no ADP, a reduction in the ATP concentration goes together with a reduction, and sometimes a complete absence, of the Kfree effect (Beauge and DiPolo, 1978, 1979b, 19813). This cannot be ascribed to a K accumulation in the extracellular space because it is also

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observed in axons dialyzed without internal K (Fig. 7). The fact that this reduction is a function of the monovalent cation used to activate the pump (the cation-free effect is more sensitive to ATP with those cations forming a tight enzyme-cation complex after dephosphorylation) suggests that it is prcciscly the inhibition of the disoccluding step at low ATP that is responsible for these effects. At low ATP and in the absence of K (or its congeners) the Na pump can operate without any difficulty in its uncoupled o r Na,-stimulated mode, thus providing the basis for a lower relative activation by K (and its congeners) over Na at reduced levels of ATP. It is intriguing, though, that even with Rb a reversal cation-free effect was never found; perhaps this is just a coincidence and the ATP was at concentrations where the Rb-stimulated and -uncoupled plus Na,-stimulated rates of pumping balanced out. 3. Metabolic poisoning. Axons that have been poisoned with cyanide for a little more than an hour show about 75% inhibition of the total Na efflux (Caldwell et ul., 1960a; Sjodin and Beauge, 1967, 1968; De Weer, 1971). From Fig. 3 of BeaugC and DiPolo (1981a), this would correspond to an ATP concentration of approximately 80 p M (the ATP dependence of Na efflux was obtained in axons dialyzed without ADP and with PA, so a straightforward extrapolation of the type just made may very well be invalid). Following De Weer’s arguments (1971), the ADP concentration in axons with cyanide may have been around 0.5 mM. In other words, compared with normal cells, cyanide-poisoned nerves have the following changes in the nucleotide composition: ( I ) reduction in ATP, (2) increase in ADP, and ( 3 ) reduction in the ATP: ADP ratio. These biochemical changes are a combination of those seen in examples (1) and (2) above. What is the reason for the reversal K-free effect in these axons? One possibility is that it is just the consequence of the dramatic inversion in the ATP: ADP ratio (De Weer, 1971). On the other hand, very low ATP levels will largely affect the E?(K)-E,K disoccluding step (see Section llI,A, I , f ) being perhaps the major reason for the reduction of Na efflux in the presence of K,, . When external K is removed, two things will happen: ( I ) the E?(K) occluding form will no longer be formed from the outside, and (2) the high levels of ADP will stimulate ATP-ADP exchange and the N a fluxes going with that partial reaction; the final consequence will be a reversal K-free cffect. There is a complication in that explanation, and it concerns the observation (Beauge and Glynn, 1980) that ADP is almost as good a s ATP in its disoccluding capacity; at about 0.5 m M there is enough ADP in the axons to carry on the disoccluding step at 70% of its normal rate. Two possible ways out of this problem exist. One is related to the proposal that the low- and high-affinity ATP binding sites are actually the same site with alternating affinities (Moczydlowsky and Fortes,

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1981); it is conceivable that if ATP is the disoccluding nucleotide it will stay on the enzyme until phosphorylation; if ADP is the agent, it will have to be debound first and ATP allowed to get in for phosphorylation to be carried on; we will be in the presence of a type of substrate competitive inhibitor. The other way out is just to consider that for the ATP: ADP ratio of 1 : 10 in cyanide [in normal nerves it is 7 : I (De Weer, 197111, ADP should act both as a competitive substrate inhibitor and as a product inhibitor of the ATPase (Na-pumping) reaction, but stimulate the ATPADP (Na-Na) exchange. 4. Vanadate poisoning. The reversal K-free effect observed in axons poisoned with vanadate and dialyzed with no ADP is readily explained by the mechanism of inhibition, which involves (I) the blocking of the Ez(K)-EIK conformational change, and (2) the KO-promotedinhibition (see Section III,A,l,d). In the presence of vanadate, KOhas, then, two actions: it allows the inhibitor to bind, and it forms the E2(K) enzyme state after dephosphorylation; this results in a large inhibition of the NaK active transport (the uncoupled and Nao-stimulated are supposed to be inhibited, partially or totally, depending on the KOconcentration, even in the absence of vanadate). When KOis removed, the inhibition by vanadate is released and the uncoupled and Nh-stimulated Na pumping come into play; the end result is a higher Na efflux in the absence than in the presence of K , . The difference between vanadate poisoning and low ATP axons (both without ADP) simply may rely on the two actions of KO in the case of vanadate, whereas with low ATP the only factor is the slow disocclusion process. This may account for the lack of reversal in axons dialyzed with low ATP concentrations. Finally, the ATP-dependent strophanthidin-resistant Na-Na exchange should be kept in mind for some cases of variable K-free effect. If this exchange is a translocation mode normally present in axons, a modification in its rate will affect the total efflux of Na, and consequently the fractional drop in Na efflux when external K is taken away. G. Conclusions

The experimental evidence discussed in this work deals with a membrane-bound ATP-driven Na-K-coupled pump as the universal mechanism for the regulation of intracellular Na and K concentrations. The information gathered here has, on the one hand, helped to explain that mechanism and, on the other, added new elements requiring further clarification. In doing so it must be borne in mind that one is working with a particular preparation; no doubt it is extremely important to apply, and

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extract, general concepts, but room must be left for features belonging, perhaps, only to the squid. ACKNOWLEDGMENTS This work was supported by the Consejo Nacional de Investigaciones Cientificas y TCcnicas of Argentina (CONICET) and by Grant BNS 8025570 from the United States National Sciences Foundation. The author is an established investigator from CONICET. REFERENCES Abercrombie, R. F., and De Weer, P. (1978). Electric current generated by squid giant axon sodium pump: External K and ADP effects. Am. J . Physiol. 235, C63-C68. Albers, R. W. (1967). Biochemical aspects of active transport. Annu. Reo. Biochem. 36, 727-756. Bader, H., and Sen, A. K. (1966). K-dependent acyl phosphatase as part of the (Na + K) dependent ATPase of cell membranes. Biochim. Biophys. Acta 118, 116-123. Baker, P. F. (1964). An efflux of ninhydrin-positive material associated with the operation of the Na+ pump in intact crab nerve immersed in Na+-free solutions. Biochim. Biophys. Acta 88,458-460. Baker, P. F. (1968). Recent experiments on the properties of the sodium efflux from squid axons. J. Gen. Physiol. 51, 172s-179s. Baker, P. F., and Manil, J. (1968). The rates of action of K’ and ouabain on the sodium pump in squid axons. Biochim. Biophys. Acra 150, 328-330. Baker, P. F., and Willis, J . S. (1972). Inhibition of the sodium pump in squid giant axons by cardiac glycosides: Dependence on extracellular ions and metabolism. J . Physiol. (London) 224,463-475. Baker, P. F., Hodgkin, A. L., and Shaw, T. I. (1962). Replacement of the axoplasm of giant nerve fibers with artificial solutions. J. Physiol. (London) 164, 330-354. Baker, P. F., Blaustein, M. P., Keynes, R. D., Manil, J., Shaw, T. I., and Steinhardt, R. A. (1969). The ouabain sensitive fluxes of sodium and potassium in squid giant axons. J . Physiol. (London)200, 459-496. BeaugC, L. (1979). Vanadate-potassium interactions in the inhibition of Na,K-ATPase. In “Na,K-ATPase. Structure and Kinetics” (J. C. Skou and J. Norby, eds.), pp. 373387. Academic Press, New York. BeaugC, L., and Adragna, N. (1971). The kinetics of ouabain inhibition and the partition of rubidium influx in human red blood cells. J . Gen. Physiol. 57, 576-592. Beaugt, L., and Del Campillo, E. (1976). The ATP dependence of a ouabain sensitive Na efflux activated by external sodium, potassium and lithium in human red cells. Biochim. Biophys. Acta 433, 547-554. Beaugt, L., and DiPolo, R. (1978). ATP levels modify the activation of the Na+ pump by external cations in squid axons. Nature (London) 271, 277-278. Beaugt, L., and DiPolo, R. (1979a). In squid axons vanadate selectively inhibits the KOactivated Na efflux. Biochim. Biophys. Acra 551, 220-223. BeaugC, L.,and DiPolo, R. (1979b). Sidedness of the ATP-Na+-K+interactions with the Na pump in squid axons. Biochim. Biophys. Acta 553, 495-500. Beaugt, L.,and DiPolo, R. (1981a). The effects of ATP on the interactions between monovalent cations and the sodium pump in squid axons. J. Physiol. (London) 314, 457-480.

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HcaugC. L,and DiPolo. K. 119x1 b). An ATP-dependent odium-roditim exchange in strophanthidin poisoned dialyzed cquid giant iixons. ./. P l t ~ s i o l .(f,ondon) 315, 447-460. Heauge, L., and 1)iPolo. R. (1983). Sidedness of cations and ATP interactions with the sodium pump. C w r . 7 o p . M o n h r . h n s p . 19, 643-647. Bcauge. I,.. and Glynn. I . M . ( 1979). Sodium ions. acting at high-aftinity extra-ccllulur sites. inhihit sodium ATPase activity of the sodium pump by slowing dephosphory1;ition. J . P/ISS;O/. ( L o r r t l o ~289, ) 17-31. BeaugC, I-., and Glynn, I. M. (1980). The equilibrium between different conformations of the unphosphorylated sodium punip: Effects of ATP and of potassium ions, and their relevance to potassium transport. J . Physiol. (London) 299, 367-383. Beauge, L., and Mullins, L. J. (1976). Strophanthidin-induced sodium efflux. Proc. R. Soc. London Ser. B 194, 279-284. BcuugC, I , . . and Ortiz. 0. (1971). Sodium and rubidium tluxes in r;tt r'cd blood cclls. J . P/ly,si(l/.(London)218, 533-549. HcaugC. I > . . Cavierec. J . D., Glynn. I . M., and Gninlham, J. J . (1980). The ctfects of vanadate on thc lluxes of sodium and potahsium ions through the sodium pump. J . fhy.SiO/. IL~ltl~kltf) 301, 7-33. Bond, G . H., Bader, H., and Post, R. L. (1966). Acetyl phosphate as hubstrate for ( N a + K) ATPase. Fed. P m c . Frd. A m . Soc. Exp. B i d . 25, 567. Bond, G. H., Bader, H., and Post, R. L. (1971). Acetyl phosphate as a substitute for ATP in (Na + K)-dependent ATPase. Biochim. Biophys. Acrcr 241, 57-67. Breitweiser, G. E. (1982). Characterization of the (Na t K)-ATPase in a membranous preparation from the optic ganglion of the squid (Loligo p e a l r i ) . Biuchim. Biophys. A<'f
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on the sensitivity of sodium efflux from squid giant axons to external sodium and potassium. J . Gen. Physiol. 56, 583-620. De Weer, P. (1975). Aspects of the recovery processes in nerve. MTP Int. Reu. Sci. Physiol. S e r . 3, 231-255. De Weer, P. (1976). Axoplasmic free magnesium levels and magnesium extrusion from squid giant axons. J . Gen. Phvsiol. 68, 159-178. I)e Weer, P. (19x3). ( N a + K ) A‘1’I’:ise: Reaction mechanisms and ion translocating s t e p . Crtrr.. Top. Mcnrhr. Erft/l,\p. 19. De Weer, P., and Guiduldig. D. (1978). Contribution of sodium pump to resting potential of squid giant axons. A m . J . Physittl. 235, C55-C62. De Weer, P., Kennedy, B. G., and Abercrombie, R. F. (1979). Relationship between the Na:K exchanging and Na:Na exchanging modes of operation of the sodium pump. In “Na,K-ATPase. Structirrc and Kinetics” ( J . C. Skou and J . G . Norby. eds.). pp. 503515. Academic Press. New York. De Weer, I).. Breitweiser, G. E.. Kennedy, B. G., and Smith, G . (19x3). ADP-ATP cx. 19. change in intern;illy dialyLed squid giant axons. Cttu. 7’0pic.s M ~ t n h r Trctns. DiPolo, R. (1977). Characterization of the ATP-dependent calcium efflux in dialyzed squid giant axons. J . Gen. Physiol. 69, 798-813. DiPolo, R. (1979). Calcium influx in internally dialyzed squid giant axons. J . Gen. Physiol. 73, 91-113. DiPolo, R . , and Beauge, L. (1980). A new internal perfusion method for transport studies in squid giant axons. Biochiwz. Biophys. Actu 600, 853-859. DiPolo, R., and Beauge, L. (1981). The effects of vanadate o n calcium transport in dialyzed squid axons: Sidedness of vanadate-cation interactions. Biochirn. Biophys. Actu 645, 229-23 b. Dudding, W. F., and Winter, C. G. (1971). On the reaction sequence of the K-dependent acetyl phosphatase activity of the Na pump. Biochirn. Biophys. Actu 241, 650-660. Eisner, D. A , , and Richards, D. E. (1981). The interaction of potassium ions and A T P o n the sodium pump of resealed red cell ghosts. J . Pliysiol. (London)319, 403-418. Flatman, P. W., and Lew, V. L. (1981). The magnesium dependence of sodium pump medi;ited-sodium-patassium and sodium-sodium exchange in intact human red cells. J . P h y ~ i ~( l L. o ~ I ~315, o ~42 ) 1-446. Frumento, A . (1965). The elcctrical effects of an ionic pump. J . Tlrcrn.. Biol. 9, 253-262. Garrahan, P. J . , and Glynn. I . M . (1967). Factors affecting the relative magnitudes of the sodium-potassium and sodium-sodium exchangcs catalyzed h y thc sodium pump. J . Physiol.(London) 192, 189-216. Glynn, I. M., and Karlish, S. J. D. (1975). The sodium pump. Annu. Reu. Physiol. 37, 13-55. Glynn, I. M., and Karlish, S. J . D. (1976). ATP hydrolysis associated with an uncoupled Na efflux through the sodium pump: Evidence for allosteric effects of intracellular ATP and extracellular sodium. J . Physiol. (London)256, 465-496. Grantham, J. J., and Glynn. I . M. (1979). Renal Na,K-ATPase: Determinants of inhibition by vanadium. A m . J . Physiol. 236, F530-F535. Hodgkin, A. L . , and Keynes, R. D. (1955). Active transport of cations in giant axons from Sepin and Loligo. J . Physiol. (London) 128, 28-60, Hodgkin, A. L . , and Keynes, R. D. (1956). Experiments o n the injection of substances into squid giant axons by means of a microsyringe. J . Physiol. (London) 131, 592-616. Jorgensen, P. L. (1980). Sodium and potassium ion pump in kidney tubules. Physiol. Reu. 60, 864-917. Kaplan, J. H., and Hollis, R . J . (1980). External Na dependence of ouabain-sensitive ATP-

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ADP exchange initiated by photolysis of intracellularly caged-ATP in human red cell ghosts. Nature (London) 288, 587-589. Karlish, S. D. J., and Pick, U.(1981). Sidedness of the effects of sodium and potassium ions on the conformational state of the sodium-potassium pump. J . Physiol. (London) 312, 505-529. Karlish, S. D. J., Beaugt, L.. and Glynn, 1. M. (1979). Vanadate inhibits (Na + K) ATPase by blocking a conformational change of the unphosphorylated form. Nature (London) 282,333-335. Kracke, G. R., and De Weer, P. (1980). Are there membrane surface charges in the vicinity of the sodium pump? Biol. Bull. 159, 487. Kracke, G. R., and De Weer, P. (1982). The rate coefficient of the sodium efflux from squid giant axons. Biophys. J . 37, 220a. Lee, K. H., and Blostein, R. (1980). Red cell sodium fluxes catalyzed by the sodium pump in the absence of K ' and ADP. Nature (London) 285, 338-339. Moczydlowski, E. G., and Fortes, P. A. G. (1981). Characterization of 2',3'-0-(2,4,6-tnnitrocyclohexadienylidine) adenosine-5'-triphosphateas a fluorescent probe of the ATP site of sodium and potassium transport adenosine triphosphatase, J . Bio/. Chem. 256, 2346-2356. Mullins, L. J. (1972). Active transport of Na+ and K' across the squid axon membrane. I n "Role of Membranes in Secretory Processes," pp. 182-202. North Holland Publ., Amsterdam. Mullins, L. J. (1979). Transport across axon membranes. I n "Membrane Transport in Biology. 11. Transport Across Single Biological Membranes" ( G . Giebisch, D. C. Tosteson, and H. H. Ussing, eds.), pp. 161-210. Springer-Verlag, Berlin and New York. Mullins, L. J., and Brinley, F. J. (1969). Potassium fluxes in dialyzed squid axons. J . Gen. Physiol. 53, 704-740. Post, R. L., Hegyvary, C., and Kume, S. (1972). Activation by adenosine triphosphate in the phosphorylation kinetics of sodium and potassium ion transport adenosine-triphosphatase. J . Biol. Chem. 247, 6530-6540. Post, R. L., Toda, G., and Rogers, F. N. (1975). Phosphorylation by inorganic phosphate of sodium plus potassium ion transport adenosine triphosphatase. J . B i d . Chem. 250, 691-701. Rakowski, R. F., and De Weer, P. (1982). Direct measurement of electrogenic Na+/K+ pump current and resting inward Na' current in squid giant axons under voltage clamp control. Biophys. J. 37, 259a. Robinson, J. D., and Flashner, M. S. (1979). The (Na + K)-activated ATPase enzymatic and transport properties. Biochim. Biophys. Acta 549, 145-176. Russell, J. M. (1979). Chloride and sodium influx: A coupled uptake mechanism in the squid giant axon. J . Gen. Physiol. 73, 801-818. Schatzman, H. J. (1953). Herzglykoside als Hemmstoffe fur den aktiven Kalium und Natrium Transport durch die Erythrocytenmembran. Helu. Physiol. Aria 11, 346-354. Simons, T. J. B. (1975). The interaction of ATP analogues possessing a blocked gammaphosphate group with the sodium pump in human red cells. J . Physiol. (London) 244, 73 1-739. Sjodin. R. A. (1971). The kinetics of Na extrusion in striated muscle as functions of the external Na and K ions concentrations. J. Gm. Physicd. 57, 164-187. Sjodin, R. A. (1971). The kinetics of sodium extrusion in striated muscle as function of external sodium and potassium ion concentrations. J. Gen. Physiol. 57, 164-187. Sjodin, R. A., and BeaugC, L. (1967). The ion selectivity and concentration dependence of cation coupled active sodium transport in squid axons. Curr. M o d . Biol. 1, 105-115.

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Sjodin, R. A., and Beaugk, L. (1968). Coupling and selectivity of sodium and potassium transport in squid giant axons. J . Gen. Physiol. 51, 152s-161s. Sjodin, R. A . , and BeaugC, L. (1969). The influence of potassium-and sodium-free solutions on sodium efflux from squid giant axons. J . Gen. Physiol. 54, 664-674. Skou, J . C. (1957). The influence of some cations on an adenosine triphosphatase from peripheral nerves. Biochirn. Biophys. Acta 23, 394-401. Skou, J. C . (1974). Effect of ATP on Na:K affinity and catalytic activities of (Na + K)activated enzyme system. Ann. N . Y. Acad. Sci. 242, 168-184. Skou, J. C . (1975). The (Na+ + K') active enzyme system and its relationship to transport of sodium and potassium. Q.Rev. Biophys. 7 , 402-434. Thomas, R. C. (1972). Electrogenic sodium pump in nerve and muscle cells. Physiol. Rev. 52, 563-594. Venosa, A. (1981). Sitios de transporte activo de Na en mhsculo esquelktico de rana. Reunidn Cicntg. Soc. Argent. Siojh., 10th A, 23.

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CURRENT TOPICS I N M E M B R A N E S A N D TRANSPORT. V O L U M I ??

Chloride in the Squid Giant Axon JOHN M . RUSSELL Department of Phy.\iology cind Biophysics Univer.tiiy of Texas Medicul Brrinc h Galuesion. Texus

.................... I . Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . [CI-] in Axoplasm and Heniolymph.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .................... I l l . Transmembrane CI- Movements. . . . . . . . . . . . . . . . . A. Electrical or Conductive CI Transmembrane Movements. . . . . . . . . . . . . . . .................... B. Coupled C1- : Na+ : K' Uptake Process., . . . . . C. Role of CI- in pH, Regulation.. . . . . . .................... ..... ...... ion ....................... IV. Summary a References ................................................... 11.

I.

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INTRODUCTION

Although chloride is by far the most abundant inorganic anion in squid hernolymph and axoplasm, the study of this ion has lagged far behind that of the inorganic cations. There are undoubtedly several reasons for this apparent oversight on the part of axonologists. First, techniques for measuring CI- concentration were tedious and laborious until the advent of the coulometric chloridometer (Cotlove et ( I / . , I9SX). Sccond, the radioisotope of chloride, 36CI,has a very long half-life and consequently is available at much lower specific activities than are the isotopes for K', Na+, or Ca2+.However, probably the most important reason for the lack of detailed information about axonal handling of CI- is that, to the electrophysiologist, the study of C1F in the squid axon is singularly uninteresting. As will be discussed, C1- apparently plays a very minor role in axonal electrical phenomena. However, it has been known for a number of years that [Clt] in squid axoplasm is significantly higher than could be accounted for by a thermodynamically passive mechanism, thus suggesting the exis177

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JOHN M. RUSSELL

tence of specialized transport mechanism(s) for C1- in the squid axolemma. To date, two such transport processes have been identified and partially characterized. One process involves the coupled influx of CI-, Na+, and K +.This system is presumably responsible for the higher-thanequilibrium [Cl-Ii. The second identified C1- transport process involves the exchange of intracellular C1- (possibiy along with H + ) for extracellular HCO; and Na+. In effect, this latter process serves to extrude cellular acid equivalents and thus functions to maintain the normal, slightly alkaline, pH characteristic of squid axoplasm. The present article reviews the evidence concerning Cl- in the squid axon and the modes of its transmembrane movements.

II.

[CI-]IN AXOPLASM AND HEMOLYMPH

A description of C1- in the squid axon must begin with a discussion of its transmembrane distribution. In principle, the measurement of intracel-

lular ion concentrations in the squid giant axon is much less difficult than for most other cells. One simply extrudes a quantity of axoplasm from the cut end of an isolated axon and thereby dispenses with the potential errors inherent in methods requiring correction for extracellular fluid contamination. Beginning in 1939 (Bear and Schmitt, 1939) and continuing until recently, a large number of different investigators have measured axoplasmic chloride concentration utilizing a variety of techniques. Table I, Part A summarizes these findings. There is reasonably good agreement that axoplasmic [Cl-] is near 120 mM. Steinbach (1941) provided the single greatest exception to this agreement with his report that [Cl-Ii = 41 mM in freshly dissected axons (“within 20 min. of death”). Inasmuch as this atypically low value has sometimes been used by electrophysiologists as a representative figure, it is important to consider it further. Although Steinbach (1941) reported this low value for “freshly dissected” squid axons, he also noted that [Cl-Ii of dissected axons rapidly increased to a plateau value of about 86 mM. This rapid net uptake of CI- combined with other experiments demonstrating equally rapid net CI- efflux into low [CI-] external solutions implied that Steinbach’s axons were mediating net transaxolemmal CI- fluxes of around 280 pmol/cm2 sec. Such a large net flux is an order of magnitude greater than any reported unidirectional fluxes (see below), which suggests that the axons studied by Steinbach were very leaky. It was these implied large fluxes that prompted Keynes (1963) to perform the most extensive and careful investigation of axoplasmic CI- content conducted to date. Briefly, Keynes demonstrated that even in the freshest axoplasm available (obtained within 17 min of

CHLORIDE IN THE SQUID GIANT AXON

179

TABLE I CHLORIDE CONCENTRATION I N SQUID AXOPLASM A N D SQUID HEMOLYMPH ~~

[Cll (mM)"

Analytic technique

A. Squid axoplasm Volhard titration (ultrami130 croddaptation) Microdiffusion method of I09 Conway (1935) (analyzed whole axon, well cleaned) Wigglesworth technique 41 (1937) Ag-AgCI microelectrode 117" Microdiffusion method of 160 Conway (1935) 174 124 Electrometric titration (Sanderson, 1952) Coulometric titration 71 (Cotlove et ul., 1958) 111 I37 Coulometric titration Coulometric titration 130

B. Squid hemolymphd 474 Volhard titration 459 van Slyke (1923-1924) method 468 van Slyke (1923-1924) method 468 Coulometric titration

Species

Reference

Loligo peulei

Bear and Schmitt (1939)

Loligo forhesi

Webb and Young (1940)

Loligo pcwlei

Steinbach (1941)

Loligo peulei Loligo pealei

Mauro (1954) Koechlin (1955)

Loligo peulei Loligo forhesi

Deffner (1961) Keynes (1963)

Loligo peulei

Brinley and Mullins (1965a)" Russell (1976) Russell and Requena (unpublished)

Loligo prulei Loligo plei

Loligo pixilei Loligo p r d e i

Bear and Schmitt (1939) Manery (1939) Hayes and Pelluet (1947) Russell (1976)

All data are expressed as millimoles per liter of axoplasmic water. When the original data were not reported in these units, corrections have been made based on an axoplasmic water content of 87% (Manery, 1939; Koechlin, 1955). Assume that [Cl], = 470 mM; temperature = 20°C. These authors found a bimodal distribution of [Cl 1. The lower values came from animals caught early in the season from cold water, whereas the higher values came from animals caught later in season from somewhat warmer water. ,I All data are expressed as millimoles per liter of hemolymph water. Hemolymph is 87.6% water (Russell, 1976).

"

decapitation of the animal) the axoplasmic [Cl-] was high, averaging 123 mmol/kg axoplasmic water. In addition, by comparing C1- ionic activity measured with Ag-AgCI electrodes to CI- concentrations determined by electrornetric titration with Ag+, Keynes determined that axoplasmic C1was free in solution; no evidence for intracellular CI- binding could be

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JOHN M. RUSSELL

found. Furthermore, in contrast to Steinbach's findings (l94l), Keynes noted that axoplasmic [Cl-] values were stable after dissection. Storage of the axons in seawater for as long as 5.8 hr after their dissection had no effect on the [CI-I. We (J. M. Russell and J. Requena, unpublished observations) have confirmcd that [Cl-] of axons storcd in seawater is fairly stable for as long as 12 hr after dissection. We found that [CI-] decreased from 130 I-fr 3 mM (n = 7) in fresh axons (within 20 min of decapitation) to 123 k 6 mM ( n = 6) 12 hr after dissection. Thus, as noted in Table I , Part A, no other workers have been able to reproduce Steinbach's main observations, and it now seems certain that axoplasmic [Cl 1 is around 100-120 mM and stable with time after dissection. In order to determine whether the axoplasmic [Cl-] is in thermodynamic equilibrium with the extracellular [CI-1, it is obviously necessary to know the extracellular [CI-I. Table I, Part B shows that squid hernolymph, obtained by cardiac puncture, contains about 470 m.M C1-. If one assumes that this is a fair representation of the external fluid bathing the giant axon, then the equilibrium (Nernst) potential for CI- is between -33.8 and -38.5 (temperature = 20°C). Given that the resting membrane potential of the axon is -50 to -70 mV, it becomes clear that the axoplasm contains 2-4 times more CI-- than one would predict from passive considerations. This greater-than-equilibrium [Cl-1, raises two questions: ( 1 ) What is thc purpose of this high [Cl 1,; and (2) What is the mechanism by which it is established and maintained? 111.

TRANSMEMBRANE CI- MOVEMENTS

The results of studies of CI movements across the axolemma will furnish partial answers to the two questions posed above. A. Electrical or Conductive CI- Transmembrane Movements As mentioned in the Introduction, for the electrophysiologist, the study of CI- has not proved to be very interesting. There have been no reports of voltage-sensitive C1- conductances in the squid axon. The electrical conductance of the axolemma to C1- appears to be extremely low. In 1960, Caldwell and Keynes pointed out that the net ionic flux through the so-called "leak" conductance, i.e., all conductance that results from current carried by ions other than Na+ or K + , was 3-5 times larger than the unidirectional 36Clfluxes in squid axons. They also showed that the extra "CI influx associated with action potentials could be accounted for by the change in driving force, ix., the change in membrane resting potential. Thus, no change in membrane permeability or conductance to C1- was

CHLORIDE IN THE SQUID GIANT AXON

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apparent during an action potential. Adelman and Taylor (1961) specifically studied the contribution of C1F to the leakage current in the voltageclamped squid axon. They concluded that C1- conductance in a resting axon is negligible and that no more than 10% of the leakage current could be accounted for by inward movement of C1-. Similarly, Brinley and Mullins (196Sb) demonstrated that the transference number of CI-, i.e., that fraction of total transmembrane current carried by CI- , was only 0.04. These results may be taken to mean that the majority of CI- transmembrane movements do not occur via diffusion through so-called simple aqueous channels. Hence, C1- movements are not directly significant for electrophysiological properties of the axon. Keynes (1963) considered two other potential electrophysiological effects of the higher-than-equilibrium [Cl-I,. First, an increase of intracellular mobile anion levels would increase the specific conductance of the axoplasrn, thereby increasing the conduction velocity of action potentials. However, the increased [CI-1, (presumably at the expense of immobile anions) would ultimately result in only about a 10% increase in the conduction velocity. This does not seem to be a significant improvement in view of the energy expenditure necessary to achieve this [Cl-1,. The second potential effect would be that Ec, might act as a synaptically activated “battery” that could be important either at the cell bodies of origin of the axon or at its synapses. The problem with this suggestion is the lack of evidence for a depolarizing, CI--dependent synaptic potential at either of these locations. Hence, it appears that electrophysiological studies can teach us very little about either the purpose of the relatively high [Cl-Ii or how it is established and maintained. Thus, we must turn to the results of studies on what appear to be ion-coupled and electrically silent Cl-- transport processes. Two such processes have been partially characterized in the squid axon. One system mediates C1- influx and is tightly coupled to concurrent Na’ and K+ uptake. The second process mediates C1- efflux and is coupled to Na+ and HCO, influx. The latter process clearly functions to extrude intracellular acid loads, thereby permitting intracellular pH regulation (e.g., Boron and Russell, 1983). Although both processes require cellular ATP and extracellular Na+, current evidence suggests they are two separate processes, and they will be considered as such here. B. Coupled CI-:Na+ :K + Uptake Process

1. ROLEOF ATP

In his landmark study of CIF in the squid axon, Keynes (1963) also studied 36CIinflux in an effort to determine the basis of the relatively high

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JOHN M. RUSSELL

[Cl-Ii. In one series of experiments, axons were pretreated for 1.3-3 hr with inhibitors of ATP synthesis, 2,4-dinitrophenol (DNP) or cyanide (CN-). The axons were then soaked in seawater containing 36Clplus the inhibitor. After a fixed time, axoplasm was extruded and counted for radioactivity. Although DNP treatment clearly inhibited 36Clinflux, the effects of CN- were much less certain. Thus, convincing evidence for a direct role of ATP in CI- influx was still missing. The technique of internal dialysis, developed by Brinley and Mullins (e.g., 1967), permits the investigator to remove cellular ATP while maintaining constant the pHi, intracellular Na+ and K+ concentrations. Thus, by means of this technique a more direct test of an ATP requirement can be made. I have used the internal dialysis technique to examine the question of an ATP-dependent Cl- influx (Russell, 1976). Figure 1 represents data obtained from an internally dialyzed squid axon. The axon had been

Inside

4 ATP

Outside

0 CI+CN

,

0 ATP

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0 00 0 0 00

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FIG.1. Effect of ATP removal on unidirectional CI- influx. This squid axon was dialyzed with a fluid having the following composition ( m M ) : K+, 300; Na+, 58; Mg2+,7; CI-, 0; glutamate-, 36; taurine, 190; HEPES, 20; EGTA, 1.0; phenol red, 0.5; CN-, 2.0; and ATP, 4; pH = 7.35. The external fluid was squid seawater with the following composition (rnM): Na+, 431.4; K+.10; Ca*+,10; Mg2+.50; CI ,553; CN-. 2; HEPES, 10 total, 6.4 as the anion; EDTA, 0.1; pH = 7.8. At zero time, 36CIwas included in this external solution. At 48 rnin, the dialysis fluid was changed to one that contained no ATP and the [Mg2+]was reduced to 4 mM. CI- influx fell from about 21 pmol/cm2 sec to about 2 pmol/cm2 sec. Return of ATP to the dialysis fluid caused the CI- influx to increase to about 22 prnol/cm2 sec. Consult earlier publications for details of dialysis technique (Russell, 1976, 1979). Temperature, 17°C; axon diameter, 480 pm.

CHLORIDE IN THE SQUID GIANT AXON

183

dialyzed with a C1--free dialysis fluid (DF) for 45 min prior to the addition of 36Cl to the external fluid. We term this dialysis prior to addition of radioisotope the predialysis period. Experiments with C1--selective liquid ion-exchange microelectrodes show that such a predialysis regimen will reduce [Cl-Ii to 5 mM or lower. As discussed below, lowering [Cl-Ii stimulates the C1- : Na+ : K + uptake process. About 50 min after adding 36C1to the external fluid, the DF was changed to one that contained no ATP. Thereupon 36C1influx, which had stabilized at 21 pmol/cm2 sec, fell over a period of 60-80 min to a value of about 2 pmol/cm2 sec. Dialysis with the ATP-containing fluid restored 36CIinflux to a value of about 22 pmol/cm2 sec. Cyanide (2 m M ) was present throughout the experiment to prevent de nouo synthesis of ATP. Cyanide in the presence of ATP has no effect on C1- influx (Russell, 1976). Thus, these results clearly demonstrate a direct requirement for ATP by the C1- uptake process. The effect of ATP removal illustrated in Fig. 1 was quite large, i.e., removal of ATP resulted in a 10-fold decline of C1- influx. Such a large effect is because [Cl-Ii = 5 mM, and it has been shown that ATP-dependent C1- influx is inversely proportional to [Cl-Ii. For example, ATPdependent C1- influx is nearly zero when [Cl-li = 150 mM (Russell, 1976). Thus, in an undialyzed axon where [Cl-Ii = 120 mM (see above), ATPdependent C1- influx would be rather small. This fact probably helps to account for the uncertain effect of cyanide treatment in the axons studied by Keynes (1963).

2. ROLEOF Na’ When Keynes (1963) reduced “a+], to 25% of that in normal external fluid, he saw no effect on CI- influx into intact axons. This result is in marked contrast to what was later observed in internally dialyzed axons (Russell, 1979). The difference is partly attributable to the much lower [Cl-Ii achieved in the dialyzed axon. Figure 2 illustrates the external Na+ dependence of C1- influx. When external Na+ is completely replaced by MgZf, CI- influx becomes very low, 1-4 pmol/cm2 sec, just as it does when intracellular ATP is dialyzed out of the axon. Somewhat less inhibition of CI- influx is noted when lithium or choline are used as Na+ replacements (Russell, 1979). Removal of external Na+ from an ATP-free axon does not further reduce CIinflux, a result that suggests a common final pathway of action of ATP and external Na+. Ouabain (lo-’ M ) had no effect on CI- influx in agreement with the earlier results of Keynes (1963). Thus, the CI- uptake process requires not only ATP, but also external N a + .Such a result raises at least two questions. First, is Na+ cotransported with CI-? Second, is the CI-

184

JOHN M. RUSSELL 0 Na S S W

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FIG.2. Dependence of CI influx upon external Na’. This axon was internally dialyzed with a fluid having an identical composition to that used for the fiber illustrated in Fig. I except there was no cyanide. The external fluids (SSW) always contained 553 mM CI but “a’] was varied by replacement with Mg2’. The osmotic difference was made up with rnannitol. In this figure it is clear that CI- influx is directly proportional to the external “a’]. Qualitatively similar results are obtained when choline or lithium are used to replace Na+; however, quantitatively, these latter two cations appear to partially support CI- influx. Axon diametcr. 650 p i . (Adapted from Russell. IY7Y. by permission o f Jorrrr~ir/c!f’C;c.)?c,rtr/ ~

P/r!..cio/o,qy.)

influx a function of the Na+ chcmical or clcctrochcmical gradicnt across the axolemnia? Only the first of these questions has been answered. A series of experiments was performed to characterize Na‘ influx in order to identify similarities to CI- influx. All axons were treated with ouabain M ) and tetrodotoxin (TTX; lo-’ M ) to reduce background Na+ influx. Figure 3 shows an axon in which C1- and Nat influxes were measured simultaneously while [Cl-1, was varied between nominally 0 and 150 mM. It is clear that the influxes of both ions were stimulated by the reduction of intracellular CI , although it should bc noted here that CI- influx was stimulated more than Nat influx. The foregoing result strongly suggested that both CI- and Na’ influxes might be linked. Thus, it was reasoned that. if CI-Na cotransport existed, then Na+ influx ought to require extracellular C1- just as CI- influx required external Na’ (scc Fig. 2 ) . In six axons,. complete replacement of external CI with CH3S07 (methane sulfonate) resulted in a decrease of Na’ influx by about the same amount as observed when [CI-1, was increased to 150 mM. Further proof of a common pathway for C1- and Na+ influxes comes from the demonstration of ATP dependence of Na+ influx

185

CHLORIDE IN THE SQUID GIANT AXON Inside

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Outside

0 CI-4 ATP

SSW

x

-

-c c

+

Ouabain

I

+

150 CI-4 ATP

TTX

r

tf n

0

2

1

3

Time (Hr)

FIG.3. Effect of changing [Cl-1, on CI- and Na+ influxes into an ATP-fueled axon. The dialysis fluid used in this axon was identical to that presented in Fig. 1 with two exceptions: (1) it contained no Na+, therefore [K'] = 358 mM; (2) when [CI-] = 150 mM it was increased at the expense of glutamate. At zero time, 3bCland 24Nawere added to the ouabain and tetrodotoxin (TTX)-containing external fluid. After steady influxes were reached, the dialysis fluid was changed to one in which all the CI was replaced by glutamate-. This reduction of [Cl-1, clearly stimulated both C1- and Na' influxes, and the effect was reversible upon returning to the higher [CI-],. 24Nahas a half-life of 15 hr, so that the influx samples were counted right after the experiment and then again about 2 weeks later in order to separate the counts of the two isotopes. Temperature, 17°C; diameter, 500 p m .

and the fact that both C1- and Na+ influxes can be blocked by the diuretic agents furosemide or bumetanide (Russell, 1979).

3. ROLE OF K ' As noted above, removal of intracellular CI- stimulated CI- influx more than it did Na+ influx. A similar relationship between C1- and Na+ influxes was noted for ATP-dependent and furosemide- or bumetanideinhibited fluxes. Finally, it had been observed (Russell, 1980) that the external Na+-dependent C1- influx was greater than the external CI-dependent Na+ influx (Fig. 4). These results suggested that the CI- : Na+ cotransport process takes up more C1- than Na+. However, no systematic membrane potential changes were noted that could be attributed to a system that takes up anions in excess of cations. An alternative explanation is that another cation was cotransported. In view of the implication of K + in CI- : Na' transport processes in erythrocytes (e.g., Chipperfield,

186

JOHN M. RUSSELL

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FIG.4. Collated data illustrating that Na,-dependent CI- influx is greater than C1,-dependent Na’ influx. (A) Relation between the external sodium concentration and the CIinflux into the squid giant axons dialyzed with a fluid containing 4 mM ATP and SO mM sodium and nominally free of C1-. Nh-dependent C1- influx is defined as the difference between the CI- influx at a given external sodium concentration and the C1- influx when external sodium concentration = 0. External sodium was replaced by magnesium. The external chloride concentration was the same as normal seawater, the osmotic difference being made up with mannitol. These data are from 12 different axons. (B)Relation between sodium influx and the external chloride concentration. Axons were dialyzed with fluid containing 4 mM ATP and 58 rnM sodium and nominally free of chloride. CI,-dependent Na+ influx is defined as the difference between the Na+ influx at a given external chloride concentration and the Na+ influx when all the external chloride is absent. Methane sulfonate (CH2S0,) was used as the substitute anion for chloride. This figure represents data from 10 different axons. (From Russell. 1980. by permission of Annctls q f N ~ HYorA . Accttlrrny of Science. 1

1980; Dunham et al., 1980) and renal tubules (e.g., Greger and Schlatter, 1981), 1 examined the effects of removing external potassium on both CIand Na+ influxes. These experiments, performed under conditions already determined to yield maximal C1- : Na+ uptake (i.e., ATP present, intracellular C1- absent, normal extracellular C1- and Na+ concentrations), revealed that a significant portion of both CI- and Na’ influx requires the presence of extracellular K + . Furthermore, K +influx studied in the presence of ouabain and TTX shares the properties of CI : N a t cotransport. For example, K+ influx is inversely proportional to [ClIijust as are C1- and Na+ influx (Fig. 5).

187

CHLORIDE IN THE SQUID GIANT AXON lnslde

150 CI - 4 ATP

Outside

I

SSW

0 Cl-4 ATP

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160 CI-4 ATP

+ Ouabaln + TTX

Time (Hr)

FIG. 5. Effect of [CI-li upon both CI- and K+ influx measured simultaneously from the same axon. [Cl-1, was varied by substitution with glutamate. The dialysis fluid was similar to that presented in Fig. I except that it contained no Na', therefore [K+l, = 358 mM. The external fluid (SSW) contained M ouabain and lo-' M tetrodotoxin (TTX). The figure illustrates that both C1F influx (0)and K' influx (0) are inversely proportional to [CI-li. Temperature, 17°C; axon diameter, 475 pm.

Table I1 is a summary of the effects of various treatments upon the unidirectional fluxes of Cl-, Na+, and K + . The pattern that emerges from Table I1 is that C1- influx > Na' influx > K+ influx. In order to determine the stoichiometry of the purported Cl- : Na+ : K + cotransport process, experiments were performed in which the fluxes of two of the three ions were measured simultaneously, either as C1-/K+ or as Na+/K+. Two TABLE I1 SUMMARYOF CI-:Na+:K+ COTRANSPORT FLUXES~ Influx

Na, dependentb

CIO dependent'

K O

dependentd

Cli dependent'

Furosemide inhibited

23.6 14.3 7.6

24.8 18.3 8.5

~

c1 Na K a

16.5

-

9.3

11.0 9.5

-

16.9 10.9

-

ATP = 4 mM; [CI], = 0 except as noted below; data in pmollcm* sec. Na,+ totally replaced by Mg2+. C1; totally replaced by CH,SO7. K,+ totally replaced by Na'. [CI-1, changed from 150 to 0 and back to 150 mM by replacement of glutamate- with CI-.

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JOHN M. RUSSELL

treatments were used to quantitate the response of the unidirectional fluxes of the three ions. First, the effect of varying [Cl-Ii from 150 to 0 m M and back to 150 mM was studied. In this case the ratio of Cl;dependent CI- influx to K’ influx was 3.3, whereas the ratio of the Na+ influx to K’ influx was 1.8. Second, when axons were treated with 0.3 mM furosemide, the ratio of the furosemide-inhibited fluxes was CI-/K+ = 3.1 and Na+/Kt = 2.2. Thus, assuming a fixed stoichiometry, these data are consistent with a CI- : Na+ : K + uptake process that cotransports these ions in the approximate ratio of 3 C1F : 2 Na+ : I K + . Such a stoichiometry would be electroneutral, and as such the fluxes ought to be insensitive to changes of membrane resting potential. This was directly tested in the experiment illustrated in Fig. 6. Under conditions designed to favor CIF : Na+ : K+ uptake (i.e., [Cl-1; = 0 , 4 mM ATP, normal [CI-I,, “a+],, and [K’],) the C1 influx in this axon was almost 22 pmol/cmz sec. When the axon was treated with 75 p M veratridine, its membrane resting potential depolarized about 30 m V and CIF influx increased 7 pmol/cm2 sec. In order to determine whether any of the flux increase was via the coupled CI- : Na+ : K’ uptake process, the axon was next treated with 1 F M bumetanide to determine the magnitude of the coupled flux first in the depolarized axon and then in the repolarized axon after treatment with lo-’ M TTX. The bumetanide-sensitive CI- influx under both conditions of membrane potential was about 21 pmol/cm’ sec. If we assume that all the bumetanide-insensitive, voltage-dependent change in CI- influx (about 6-7 pmol/cm’ sec) was via a conductive pathway and that it represents a net flux (an assumption that could result in an overestimate of C1conductance), we calculate an axolemmal CI- conductance of 2.2 x mho/cm2. Hodgkin and Huxley (1952) have estimated “leak” conducmho/cm2, thus this result supports the contance to be about 26 x cept of a relatively small CI- movement via electrically sensitive pathways (see above). The foregoing studies on C1F uptake have raised some significant questions. Although it seems reasonable to assume that the CIF : Na+ : K + uptake process is the mechanism whereby the higher-than-equilibrium [Cl-Ii is maintained, we still do not know why [Cl-1, exceeds equilibrium values. In addition, we do not yet know exactly what energizes the uptake process; that is, what are t h e roles of ATP and the Na gradient? What is the role of K + ? It is tempting to speculate that this uptake process might function in cellular volume regulation. If so, the process as presently characterized is poised to inject osmotic particles into the axon, thereby causing it to swell. Hence, the transport process would be expected to be sensitive to, and stimulated by, some consequence of cell shrinkage. Clearly, that consequence cannot be a change of [Cl-li since osmotic shrinkage would be expected to raise [CIF]i whereas the coupled intake +

189

CHLORIDE IN THE SQUID GIANT AXON

I

T -30 -40

E

Veratridlne

.... ..@. 1-

Bumetanide

0.

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FIG.6. Effect of membrane potential depolarization on bumetanide-sensitive and -insensitive fractions of C1- influx. The axon was dialyzed with a CIF-free fluid containing 4 mM ATP. Membrane potential (V,) was measured by a 0.5 M KCI-filled glass micropipette inserted into middle portion of the axon alongside the dialysis tubing. A steady CI- influx of 22 pmol/cm2 sec was obtained. When 75 p M veratridine was added to the external fluid the membrane resting potential decreased from about -62 mV to about -33 m V and CIF influx increased to 29 pmol/cm2 sec. Thus, V , depolarizaion was accompanied by an increase of CI- influx of 7 pmol/cm2 sec (denoted as a in the figure). When 1 p M bumetanide was added to the external fluid to inhibit the CI- : Na+ : K' uptake mechanism, CI- influx declined to 7 pmol/cm'sec and V , hyperpolarired about I mV. The membrane potential was then repolarized by treating with lo-' tetrodotoxin (TTX), reaching a value of about -59.5 mV, whereupon CI- influx declined further to a value of about 1 pmol/cm2 sec, a decrease of 6 pmol/cm2 sec (denoted as b in the figure). By comparing a and b, we see that the fall of CI- influx after repolarization was essentially the same as the rise following depolarization despite the fact that repolarization was accomplished while the C1- : Na' : K+ transport mechanism was blocked. Thus, it appears that virtually all the voltage-sensitive C1- influx was via a bumetanide-insensitive mechanism. Axon diameter, 500 p m .

process is stimulated by a decrease of [Cl-Ii. It is obvious that further work is needed to understand this transport process and its physiological function. C. Role of CI- in pH, Regulation It has become increasingly evident that virtually all animal cells regulate their intracellular pH rather closely (Roos and Boron, 1981). At present, it appears that several transport processes can be used for pHi

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JOHN M. RUSSELL

regulation, depending upon the particular cell type. My colleague Dr. W. F. Boron and I have studied the pHi regulatory process of the squid axon in considerable detail. In this article I will concentrate upon the role of C1- in the pHi regulatory process. Dr. Boron covers the overall pH; regulatory process in more detail in another article of this volume (see pp. 249-269). The presence of intracellular CI- is an absolute requirement for the operation of the pHi regulatory process in the squid axon. When cellular C1- was dialyzed out by replacement with glutamate, axons were no longer capable of ejecting an intracellular acid load (Russell and Boron, 1976). Furthermore, we demonstrated that under conditions in which axons were extruding acid, there was an extra CI- efflux (Fig. 7) and this extra CI- efflux required the presence of ATP and HCO,. Both pHi regulation and the extra C1- efflux could be blocked by the anion transport inhibitor 4-acetamido-4'-isothiocyanostilbenedisulfonic acid (SITS). More recently we found that external Na+ was also a requisite for pHi regulation in the squid axon (Russell and Boron, 1979, 1982). A similar finding had been made earlier in the snail neuron by Thomas (1977).The extra C1- efflux associated with acid extrusion was also inhibited by removal of external Na+. Conversely, acid extrusion was found to be accompanied by an extra Na+ influx, and this extra Na+ influx required the presence of intracellular C1-. The interdependencies of the ion fluxes and acid extrusion provide strong evidence for a key role of CI- in the overall process. pHi regulation in the squid axon is believed to be accomplished by a tightly coupled ion-exchange transport process that exchanges internal C1- (and maybe H + ) for external HCO, and Na+ (or maybe NaC03) and requires ATP. Removal of intracellular CI- or any other of the required substances will completely stop acid extrusion. The rate of acid extrusion as a function of [Cl-Ii has been studied (Boron and Russell, 1983). We found that the dependence of the acid extrusion rate on [Cl-li followed simple Michaelis-Menten kinetics with an apparent K , of 84 mM ("a+], = 425 mM; [HCO,], = 12 mM; pH, = 8.0). This rather high apparent K , may partially explain the need for the higher-than-equilibrium [Cl-Ii found in axoplasm, since an equilibrium [Cl-Ii of -40-50 mM would seriously compromise the acid extrusion process. IV. SUMMARY AND DISCUSSION

The axoplasm of the squid giant axon contains significantly more free C1- than expected from thermodynamic equilibrium considerations, An

ATP-requiring, tightly coupled, cotransport process involving the uptake

191

CHLORIDE IN THE SQUID GIANT AXON

Internal ATP External

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FIG.7. These three panels represent data linking a component of Cl- efflux to intracellular pH regulation. We have shown that a squid axon can extrude an imposed intracellular acid load when ATP and CI- are present intracellularly and HCO, and Na+ are present externally. Removal of any one of these substances completely inhibits acid extrusion, as does treatment with SITS (Russell and Boron, 1976, 1982; Boron and Russell, 1983). In panel a are shown the effects of presenting an acid-loaded axon (dialyzed with a fluid containing 150 mM C1-, pH 6.7) with external HCOj (10 m M ) first in an ATP-depleted state, then after ATP replenishment via internal dialysis. In the right-hand part of panel a, all the requirements for acid extrusion have been met and there is a concomitant increase of CIefflux. Axon diameter is 625 pm. In panels b and c we see the effect on HCOF-stimulated CI- efflux of pretreating with 0.5 mM SITS. These two axons were a pair from the same animal. The axon whose data are represented in panel c was pretreated with SITS for 50 min before the experiment was begun. Both axons were dialyzed with a dialysis fluid containing 4 mM ATP, 150 mM CI- (pH 6.7). The experiments were begun with 10 mM HCO, in the external fluid. Upon removal of the HCOi, the efflux from the axon not pretreated with SITS fell by about 3 pmol/cm* sec, whereas no change was noted in the CI- efflux from the SITS-pretreated axon, indicating that SITS inhibits the C1- efflux associated with acid extrusion. Axon diameter: b, 500 pm; c, 510 p m . (Reprinted, by permission, from Russell and Boron, 1976).

of CI-, Na+, and K+ is apparently responsible for the higher-thanequilbrium [Cl-Ii. It is not yet known whether the Na+ gradient provides any energy for the uptake process or whether the required energy comes directly from ATP hydrolysis. Virtually all the unidirectional C1- influx can be accounted for by the coupled C1- : Na+ : K+ uptake process. Caldwell and Keynes (1960) estimated that the leakage conductance of the axolemma would correspond to an ionic flux of 67 pmol/cm2sec, whereas the unidirectional CI- influx remaining after inhibition of the voltageinsensitive C1- : Na+ : K + uptake process can be as low as 1 pmol/cm2 sec.

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Thus, CI- contributes very little to leakage conductance. It therefore seems clear that the most important transmembrane pathway for C1entry is via the tightly coupled C1- : Na+ : K + cotransport system. The extremely low electrical conductance of the axolemma to Cl- tends to reduce the backflux of C1- down its electrochemical gradient and thereby reduces the metabolic cost of maintaining the high [CIF],.One important means of net CI- efflux is via the ion exchange transport process, which serves to regulate pHi. This efflux is not large, being 2-4 pmol/cm2sec when pHi has been reduced to 6.7 (from normal pH, 7.35). The kinetic requirement of the pH,-regulatory process for intracellular C1F appears to mandate the relatively high [CI-1, since it has a K , of 84 mM. Thus a very important function of intracellular C1F is to serve as an exchange partner in the pHi regulatory transport process. ACKNOWLEDGMENTS The experiments of the author described herein were performed at the Marine Biological Laboratory (MBL) in Woods Hole, Massachusetts. The author wishes to thank the Director and personnel of the MBL for the facilities and services placed at his disposal. This work was supported by DHHS Grant NS- 11946 from the Institute of Neurological and Communicative Disorders and Stroke. REFERENCES Adelman, W. J., and Taylor, R. E. (1961). Leakage current rectification in the squid giant axon. Muture (London) 190,883-885. Bear, R. S ., and Schmitt, F. 0. (1939). Electrolytes in the axoplasm of the giant nerve fibers of the squid. J . Cell. Comp. Physiol. 14, 205-215. Boron, W. F., and Russell, J. M. (1983). Stoichiometry and ion dependence ofthe intracellular pH regulating niechanism in squid giant axons. J . G e r r . Physiol.82, 47-78. Brinley, F. J., and Mullins, L. J. (1965a). Variations in the chloride content of isolated squid axons. Physiologist 8, 121. Brinley, F. J., and Mullins, L. J. (196%). Ion fluxes and transference number in squid axons. J . Neurophysiol. 28, 526-544. Brinley, F. J., and Mullins, L. J . (1967). Sodium extrusion by internally dialyzed squid axons. J . G m . Physiol. SO, 2303-2332. Caldwell, P. C., and Keynes, R. D. (1960). The permeability of the squid axon to radioactive potassium and chloride ions. J . Physiol. (London)154, 177-184. Chipperfield, A. R. (1980). An effect of chloride on (Na + K) co-transport in human red blood cells. Nir/rtrtJ(Londorl) 286. 281. Conway, E. I. (1935). An absorption apparatus for the microdetermination of certain volatile substances. 111. The micro-determination of chloride with application to blood, urine and tissue. Biochein. J . 29, 2221-2235. Cotlove, E., l'rantham, H. V., and Bowman, R. L. (1958). An instrument and method for automatic, rapid. accurate and sensitive titration of chloride in biologic samples. J . Lab. C'lin. I I I V C ~ S51. / . 461-468.

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Deffner, G . G. J . (1961j. The dialyzable free organic constituents of squid blood. a comparison with nerve axoplasm. Bioc.him. Biophys. Actu 47, 378-388. Dunham, P. B., Stewart, G. W., and Ellory, J. C. (1980). Chloride-activated passive potassium transport in human erythrocytes. Proc. Natl. Acad. Sci. U . S . A . 77, 1711-1715. Greger, R., and Schlatter, E. (1981). Presence of luminal K', a prerequisite for active NaCl transport in cortical thick ascending limb of Heute's loop of rabbit kidney. PJluegers Arch. 392, 92-94. Hayes, F. R., and Pelluet, D. (1947). The inorganic constitution of molluscan blood and muscle. J . Mar. B i d . Assoc. ( U . K . )20, 580-585. Hodgkin, A. L., and Huxley, A. F. (1952). The components of membrane conductance in the giant axon of Loligo. J . Physiol. (London) 116, 473-496. Keynes, R. D. (1963). Chloride in the squid giant axon. J . Physiol. (London) 169, 69-705. Koechlin, B. A. (1955). On the chemical composition of the axoplasm of squid giant nerve fibers with particular reference to its ion pattern. J . Biophys. Biochem. Cytol. 1, 511-529. Manery, J. F. (1939). Electrolytes in squid blood and muscle. J . Cell. Comp. Physiol. 14, 365-369. Mauro, A. (1954). Electrochemical potential difference of chloride ion in the squid axonseawater systems. Fed. Proc. Fed. A m . Soc. Exp. Biol. 13, 96. Roos, A , , and Boron, W. F. (1981). Intracellular pH. Physiol. Rev. 61, 296-434. Russell, J. M. (1976). ATP-dependent chloride influx into internally dialyzed squid giant axons. J . Membr. Biol. 28, 335-349. Russell, J. M. (1979). Chloride and sodium influx: A coupled uptake mechanism in the squid giant axon. J . Gen. Physiol. 73, 801-818. Russell, J. M. (1980). Anion transport mechanisms in neurons. Ann. N . Y . Aiwd. S L . ~341, . 5 10-523. Russell, J. M., and Boron, W. F. (1976). Role of chloride transport in regulation of intracelM a r pH. Nature (London) 264, 73-74. Russell, J. M., and Boron, W. F. (1979). Intracellular pH regulation in squid giant axons. B i d . Bull. 157, 392. Russell, J . M., and Boron, W. F. (1982). Intracellular pH regulation in squid giant axons. In "Intracellular pH: Its Measurement, Regulation and Utilization in Cellular Functions" (R. Nuccitelli and D. W. Deamer, eds.), Kroc Foundation Series, Vol. 15, pp. 2212317. Liss, New York. Sanderson, P. H. (1952). Potentiometric determination of chlorides in biological fluids. Biochem. J . 53, 502-505. Steinbach, H. B. (1941). Chloride in the giant axons of the squid. J . Cell. Comp. Physiol. 17, 57-64. Thomas, R.C. (1977). The role of bicarbonate, chloride and sodium ions in the regulation of intracellular pH in snail neurones. J . Physiol. (London) 273, 317-338. van Slyke, D. D. (1923-24). The determination of chlorides in blood and tissues. J . Biol. Chem. 58, 523-529. Webb, D. A., and Young, J. Z. (1940). Electrolyte content and action potential of the giant nerve fiber of Loligo (forbesi). J . Physiol. (London) 98, 299-313. Wigglesworth, V. B. (1937). A simple method of volumetric analysis for small quantities of fluid: Estimation of chloride in 0.3 fi1 of tissue fluid. Biochem. J . 31, 1719-1722.

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CURRENT TOPICS IN MEMBRANES AND TRANSPORT, VOLUME 22

Axonal Calcium and Magnesium Homeostasis P. F. BAKER Department of Physiology University of London King’s College London, England

AND R . DIPOLO Centro de BioJisica y Bioquirnica Instituto Venezotano de Inuestigaciones Cientifcas (IVIC) Caracas, Venezuela

I. Introduction. ........................................................... 11. Axonal Ca and Mg: The Problem of Bound versus Free.. ................... 111. Axonal Mg Homeostasis.. ............................................... A. General.. . . . . . ..................... B. Major Features.. ................................................... C. Conclusions .......... IV. Axonal Ca Homeostasis .................................................

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B. Mechanisms of Intracellular C. Calcium Fluxes across the A

1.

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197 I97 I91 20 1 20 I 20 1 202 207 236 242

INTRODUCTION

The squid axon has proved to be a key preparation for unraveling the complexities of cellular Ca and Mg homeostasis. In this article we attempt to present a coherent account of the present state of knowledge; but 195 Copyright 0 1984 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-153322-0

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the reader is referred also to a number of earlier reviews that deal largely or in part with the squid axons (Baker, 1970, 1972, 1978; Blaustein, 1974; Brinley, 1978; Requena and Mullins, 1979; DiPolo and Beauge, 1980, 1983). II. AXONAL Ca AND Mg: THE PROBLEM OF BOUNDVERSUSFREE

The Ca and Mg contents of axoplasm differ by a factor of approximately 100. Thus, axoplasm extruded from a freshly dissected axon usually contains per kilogram between 50 and 200 pmol of Ca and 5 and 10 mmol of Mg. Both values tend to increase with time after dissection, especially if the axon is stored at low temperature. The Ca and Mg contents of squid blood are close to those of seawater (10 and 50 mM, respectively), but because of the presence of calciumbinding agents, most notably sulfate, the free Ca is probably nearer 4 mM (Blaustein, 1974). The free Mg has not been measured, but it seems likely to be a substantial proportion of the 50 mM total Mg. If divalent cations are distributed passively in accordance with the Nernst equation, with a resting potential close to 58 mV inside, negative axoplasmic Ca and Mg should be close to 0.4 and 5.0 M , respectively. As internal Ca and Mg never even remotely approach these values, yet 45Caand 2xMgcross thc axolemma quite readily, the axon must possess mechanisms for preventing Ca and Mg from reaching electrochemical equilibrium. Outwardly directed divalent cation pumps located in the axolemma are a major component of these mechanisms. A proper assessment of the energetic problems confronting these pumps requires a knowledge of the activity of Ca and Mg in axoplasm. For Mg, arguments from mobility and enzyme activity and direct measurements with Eriochrome Blue all indicate that between one-third and one-half of the total axoplasmic Mg is free (Baker and Crawford, 1972; Scarpa, 1974; Brinley and Scarpa, 1975; De Weer, 1976). The picture is quite different for calcium, where only a very small fraction, 0. I % or less, is free. Information on the state of Ca in axoplasm has been obtained from a variety of approaches. These include ( I ) self-diffusion and mobility of 45Ca (Hodgkin and Keynes, 1957; Baker and Crawford, 1792); (2) aequorin (Baker ef ul., 1971; DiPolo et al., 1976); (3) calcium-sensitive dyes (DiPolo et al., 1976), and (4) calcium electrodes (DiPolo et ul., 1983a; Baker and Umbach, 1983). The first technique shows that the bulk of the Ca in axoplasm is bound, and the last three are all in agreement that the free Ca in freshly dissected axons is 100 nM or less when calibrated

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against Ca-EGTA buffers in the presence of axoplasmic levels of HSand Mg2+.Axoplasmic free Ca tends to rise with storage of an axon, and there is also evidence that the distribution of free Ca is not always uniform across an axon, the free Ca immediately beneath the axolemma being higher than at the center under conditions of net Ca gain and the reverse pertaining under conditions of net loss. In conclusion, although chemical analysis of axoplasm shows it to contain 100 times as much Mg as Ca, the ratio of free Mg to free Ca is probably nearer 500,000 : 1. Axoplasm and squid blood also contain measurable amounts of other divalent cations including Sr, Mn, Zn, and Co. Despite the fact that each of these cations is strongly bound in axoplasm, in no case does the total axoplasmic concentration approach that expected for an ion distributed in accordance with the membrane potential. Thus, in Loligo forbesi, the respective axoplasmic contents of Sr, Mn, Zn, and Co are 2.9 pmollkg, 41 pmollkg, 63 pmollkg, and 28 nmol/kg, whereas the concentrations of Sr, Mn, and Zn in cell-free deproteinized samples of squid blood are 75 pmol/liter, 13 pmol/liter, and 16.5 pmoll liter, respectively. Apart from strontium, which seems to be handled rather like calcium (Baker and Singh, 1982), nothing is known of the mechanisms regulating these other trace metals.

111.

AXONAL Mg HOMEOSTASIS

A. General

In many ways the analysis of Mg homeostasis in the squid axon should be simpler than that of Ca because Mg is only weakly bound in axoplasm. Unfortunately, the short half-life and relative unavailability of 28Mghave severely limited work in this area. There is, nevertheless, good agreement in the literature on the main features of Mg homeostasis (Baker and Crawford, 1972; de Weer, 1976; Mullins et al., 1977; Mullins and Brinley, 1978; Caldwell-Violich and Requena, 1979). 6. Major Features

Four key observations are described below. 1. The presence of ATP and other cytoplasmic anions can account quantitatively for the bound fraction of axoplasmic Mg. Application of metabolic poisons brings about a small rise in ionized Mg roughly in parallel with the disappearance of ATP (DiPolo et al., 1976).

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2. Influx Mg from seawater averages 0.6 pmol c w 2 sec-' at 20°C. Influx is increased roughly 10-fold when external Na is replaced isosmotically by Li (Baker and Crawford, 1972). This effect is less marked in axons with a low internal Na content. Influx is also increased in choline seawater; but, although Caldwell-Violich and Requena (1979) have reported that axons immersed in Li seawater gain more Mg than axons immersed in choline, a direct comparison of 28Mg influx from Li and choline seawaters has not been made. Despite the clear evidence of a Naidependent Mg influx, no Mg,-dependent Na efflux has been reported. Although Mg influx from seawater is not altered in fully poisoned axons, the effects of poisoning on the increased influx from low-sodium solutions has not been examined. 3. Electrical stimulation in Na seawater containing 55 mM Mg and 10 mM Ca produces an extra Mg influx of 0.007 pmol cm-2 impulse. Inclusion of Mn in the seawater markedly reduces the extra Mg influx (Baker and Crawford, 1972). Stimulation also brings about a small rise in Mg efflux (de Weer, 1976). Depolarization with K at constant external Na does not obviously increase Mg uptake (Caldwell-Violich and Requena, 1979) or alter Mg efflux (Baker and Crawford, 1972; de Weer, 1976). 4. Efflux of Mg averages about 1 pmol cm-* sec - I . It is unaffected by ouabain or by removal of external K or Ca or by injection of arginine. It is reduced by injection of Na and is strongly inhibited by lowering the temperature, by replacement of external Na by choline, lithium, or potassium, by injection of apyrase, and by poisoning with cyanide (Baker and Crawford, 1972; de Weer, 1976). This last effect can be reversed by injection of ATP. Activation by external Na approximates to a section of a rectangular hyperbola (apparent K , for Na,, 40-100 mM), and external Mg seems to compete with Na such that lowering Mg, enhances Mg efflux. The effects of low Na, and cyanide are not secondary to altered axoplasmic ionized Ca (Baker and Crawford, 1972) (Fig. 1).

Efflux seems not to be affected by membrane potential, but it is inhibited by lanthanum (Ki approximately 0.5 mM), D600 (Ki 0.25 mM), and Mn (Ki - 50 mM) (de Weer, 1976; Baker and Crawford, 1972). Working with axons dialyzed with an ionized Mg in the physiological range, Mullins et al. (1977) have confirmed the dependence of Mg efflux on Na, and ATPi and its inhibition by raising internal Na. In addition they demonstrated that, in the nominal absence of ATP, Mg efflux can be increased up to and even beyond its value in the presence of ATP simply by raising Mg, (Fig. 2). This ATPi-insensitive Mg efflux is only partially reduced by removal of external Na. Had it been completely Na,-dependent, there would have been little doubt that the inhibitory effect of ATP

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AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS

I I

I I

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z u

00

t

I

0 1

I

I

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FIG.I . Effects of Na-free seawater and cyanide (2 mM) on the efflux of Mg (fraction of total Mg lost per minute) from an axon that had been preinjected with EGTA to give a final concentration of 2.5 mM. 0 ,Na seawater; 0,choline seawater. Loligoforbesi axon diameter is 875 pm; temperature. 21°C. (From Baker and Crawford. 1972.)

removal can be overcome by the simple expedient of raising internal Mg. As it is, more experiments are required to settle this very important point. Nevertheless, the findings of Mullins et al. (1977) are compatible with the observations of Caldwell-Violich and Requena (1979), who loaded axons with Mg to give a total axoplasmic Mg of 20 m M and shows that the axons Mg efflux pmol cm+ sec-1 ATPz3mM

FIG.2. Efflux of Mg as a function of Mgi in the absence (0)and in the presence (0)of ATP. Both sets of determinations were made on the same axon. Loligo pealei axon dialyzed with a K isethionate-glycine solution. (From Mullins ef a/., 1977.)

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can lose about 5 mM of this load in a Na-dependent manner even when fully poisoned (Fig. 3). This is approximately the range of internal Mg concentrations in which Mullins et al. observed a sizable ATP-independent Mg efflux.

I

I U

I

0

I

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2

I

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4

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Time (houd

Fic. 3. Time course of loss of a Mg load under different experimental conditions. Total axoplasmic Mg is plotted as a function of time for axons of Doryteufhis plri that had been loaded with Mg in Na-free Mg seawater and subsequently allowed to recover in the presence and in the absence of external Na. 100% of load refers to Mg, at zero time (15.2 mmolikg in control axons and 20.0 mmolikg in inhibited axons) and 0% of load refers to Mg, in freshly dissected axons (4.2 mmolikg). Filled symbols and the left axis refer to control axons; open symbols and the right axis refer to inhibited axons. Circles represent axons recovered in Na seawater, and triangles represent axons recovered in Na-free (Tris) seawater. Each point is the mean 5 SEM of 6-8 axons. (From Caldwell-Violich and Requena, 1979.)

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C. Conclusions Efflux of Mg is dependent on external Na and may reflect an exchange of intracellular Mg for external Na; but a Mg,-dependent Na influx has not been described. As Mg fluxes seem not to be sensitive to membrane potential, it is possible that cytosolic ionized Mg is determined largely by an electroneutral exchange of 2 Na' for 1 Mg2+.The existence of an MgNa exchange would be consistent with ( I ) the inhibition of Na,,-dependent Mg efflux by raising external Mg, and with (2) the finding that Mg influx is stimulated both by a rise in internal Na and a fall in external Na. The strongest argument against this interpretation remains the failure to observe an Mg,-dependent Na efflux on transferring an axon from Ca-free Mg-containing seawater to Ca- and Mg-free seawater. The possibility of some sort of cotransport of Mg into the axon with Ca deserves serious investigation. Nai-dependent Ca influx is strongly activated by internal Ca (see Section IV,C,2,c), and it is possible that Na,-dependent Mg influx has a similar requirement. I t follows that Na,-dependent Mg influx will be much reduced in Ca-free media and an Mg,-dependent Na efflux may be detectable only in the presence of elevated levels of internal Na and Ca. Provided the interpretation of Mg fluxes in terms of Na-Mg exchange is correct, the role of ATP would seem to be to increase the affinity of the exchange process for internal Mg. The mechanism by which this is achieved has not been examined. In addition to Na,-dependent Mg influx, there is also an entry of 28Mg associated with electrical activity. Although this may reflect augmented Na-Mg exchange, the experiments of Rojas and Taylor (1975) on axons perfused with nominally Na-free KF suggest that z*Mg may be entering through a voltage-dependent channel separate from the tetrodotoxin (TTX)-sensitive sodium channel. An Mg inward current has, however, not been detected. IV. AXONAL Ca

HOMEOSTASIS

A. General

The experimental a alysis of Ca homeostasis in squid a ons must contend with the powerful binding of Ca in axoplasm. Thus, alterations in binding tend to be reflected in altered fluxes and altered rates of binding are often accompanied by changes in cytosolic pH and nucleotide levels, which can in turn influence fluxes. In addition, changes in specific activity can pose serious problems.

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It follows that a detailed analysis of the various components that contribute to axonal Ca homeostasis is best performed under conditions where variables such as ionized Ca, pH, and ATP:ADP ratio can be subject to experimental control. This can best be achieved by techniques of intracellular perfusion or dialysis (Baker et al., 1961, 1962; Oikawa et al., 1961; Brinley and Mullins, 1967). For studies of axoplasmic binding in the absence of the surface membrane, it is often convenient to extrude axoplasm into a porous cellulose acetate tube of internal diameter roughly equal to that of the original axon (Baker et al., 1977; Rubinson and Baker, 1979). In this way the components of axoplasm can be subject to a controlled environment while still retaining their normal morphological relationships. Despite the greater possibilities for analysis provided by dialysis, it is always essential to relate findings with this technique to Ca homeostasis in intact axons to ensure that systems have not been modified or even lost when subject to a defined chemical environment. For these reasons, throughout this section we first summarize the main features of calcium homeostasis in intact fibers before considering in some detail their characterization under conditions of internal dialysis. 6. Mechanisms of lntracellular Bindlng

The Ca-binding capacity both of intact axons and isolated axoplasm is most impressive. Within a few seconds of injecting enough Ca to raise the axoplasmic Ca content from 100 pmol/kg to 200 pmol/kg, the free Ca measured with aequorin, arsenazo 111, or Ca electrodes has returned close to its initial resting value of 100 nM (Baker et ul., 1971; Baker and Schlaepfer, 1975, 1978; Brinley et ul., 1977; Brinley, 1978). Similarly, during electrical stimulation or immersion in low-Na media-both being conditions that produce a substantial net gain of Ca-only a very small fraction of this Ca uptake is detectable as free Ca. By comparing the Ca influx, determined in a separate set of experiments, with the observed rise in free Ca, Brinley et al. (1977) concluded that during conditions of net Ca gain only 111000 or less of the Ca entering axons of L. peulei is detectable as free calcium. One effect of axoplasmic binding is that a localized pulse of free Ca remains highly localized. This can be quantified in terms of a “space constant,” equal to (D/k)”*,where D is the diffusion coefficient for Ca in free solution and k is the uptake rate constant, for the spread of free Ca. Baker et al. (1971) calculated an effective space constant of 80 pm for axoplasm of L. forbesi at 20°C. Calcium-binding is altered, but not abolished, in fully poisoned axons.

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This change in Ca-binding seems to result in the main from inhibition of mitochondrial function because injection of apyrase, which lowers cytoplasmic ATP but leaves mitochondrial function intact, has no effect on free Ca, whereas exposure to the respiratory chain blockers cyanide or azide or the uncouplers DNP or FCCP results in a rise in ionized Ca. In the case of cyanide, the rise in free Ca occurs after a lag of 1-2 hr, whereas the uncoupler FCCP brings about a virtually immediate increase (see Fig. 4). In all cases the level to which the free Ca rises still represents only a small fraction of the total axoplasmic Ca. Exact quantification is difficult with aequorin because of the high rate of consumption of the photoprotein at micromolar concentrations of Ca; but, making allowance for this and the fact that the aequorin light is related to [Ca]’, the ionized Ca seems to reach the micromolar level. This has been confirmed both with arsenazo I11 and calcium-sensitive electrodes. The precise value depends on the past history of the axon and on the Na and Ca concentrations in the seawater in which the axon is poisoned; but the free Ca in poisoned axons is always higher than in unpoisoned axons. In the case of a freshly dissected axon immersed in Na seawater containing 4 mM Ca, the free Ca in cyanide usually stabilizes between 300 and 1000 mM. However, values of 10 p M or higher can be found in Ca-loaded axons poisoned

0

500

1000

1500

2000 )LM

FIG.4. Time course of rise in intracellular ionized calcium in squid axon stimulated at 100 pulses per second in artificial seawater that contains 112 mM Ca. Free-calcium concentration is measured with Arsenazo 111. Trace A: FCCP present from beginning of stimulation; trace B: FCCP added at the end of stimulation. Ordinate, ionized calcium; abscissa, duration of stimulation and estimated exogenous load introduced by stimulation. (From Brinley ef al., 1977.)

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either in Na-free seawater or Ca-rich Na seawaters. Poisoning with cyanide is largely reversible, the ionized Ca returning quickly toward the unpoisoned level after the cyanide is removed. By comparing the measured Ca uptake with the increment in free Ca, Brinley et a f . (1977) concluded that in a fully poisoned axon of L. pealei approximately 1-5% of the net Ca gain is detectable as free calcium. Thus, poisoning reduces the effectiveness of Ca buffering by a factor of at least 10, but appreciable buffering persists. The axoplasmic components contributing to buffering have been characterized in some detail by Baker and Schlaepfer (1975, 1978) on the basis of 45Ca binding by extruded axoplasm under conditions of equilibrium dialysis. As with all techniques where cytoplasmic constituents are exposed to a defined chemical environment, it is always possible that the medium used does not support the full physiological range of buffering and that endogenous Ca-binding molecules of low molecular mass may be lost during the dialysis procedure; but with these caveats in mind, their main findings are listed below. 1. Axoplasm binds Ca even in the nominal absence of ATP and in the presence of the mitochondria1 inhibitors cyanide and oligomycin. The extent of this binding depends on the ionized Ca to which the axoplasm is exposed. In the physiological range of ionized Ca, it is possible to distinguish a component of binding with an apparent K , for Ca of about 100 nM and a capacity of 10-60 pmol of Ca per kilogram of axoplasm and a component of much lower affinity that binds approximately 1000 nmol of Ca per kilogram of axoplasm per 1 p M increment in Ca over the range 0100 p M and shows no sign of saturation. This latter component probably includes the unknown Ca buffer designated X by Brinley (1978). The molecules contributing to energy-independent binding have not been identified positively although Alema et al. (1973) have isolated a Cabinding protein with a rather low affinity €or Ca that may contribute to the linear component described by Baker and Schlaepfer (1975, 1978) and by Head and Kaminer (1980) have shown that squid axoplasm contains roughly 200 mg of calrnodulin pcr kilogram of axoplasm. This is equivalent to 10 p M . On the assumption that cach calrnodulin molecule binds 4 Ca ions, the calmodulin content of squid axoplasm would provide 40 pmol of high-affinity Ca-binding capacity per kilogram of axoplam, i.c., very close to that determined by Baker and Schlaepfer. Morc work is clearly needed, especially to idcntify positivcly the components concerned and to determine their spatial distribution. It is also possible that other Ca-binding components exist because all the experimental work has utilized extruded axoplasm, and the technique of extrusion leaves behind a thin

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205

layer of peripheral axoplasm (Baker c f al., 1962), where it is possible that other important Ca-binding molecules may be located. 2. In the presence of the inhibitors cyanide and oligomycin, the addition of Mg-ATP does not promote further binding, but removal of the inhibitors results in very appreciable extra uptake of 45Ca.This uptake can be energized by either succinate or ATP, can be discharged by FCCP, and seems to reflect accumulation by mitochondria. It is half-maximal at a free Ca concentration of 20-60 ,uM and has a very large capacity. Experimental determination of the maximum 45Cabinding capacity at a given free Ca is likely to be an underestimate because, under the conditions of equilibrium dialysis used, energy-dependent binding (but not ATP-independent binding) probably does not reach a fully stable level within the useful life of the preparation-especially at free Ca concentrations close to physiological. This problem has been reinvestigated by direct microinjection of Ca into isolated axoplasm and measuring the resultant free Ca with a Casensitive electrode. These experiments, which avoid any changes in the chemical composition of axoplasm, suggest that the maximum Ca-binding capacity varies between 2 and 10 nmol of Ca per kilogram of axoplasm (Baker and Umbach, 1983) (Fig. 5). After loading axoplasm with enough Ca to raise the axoplasmic Ca content by 0.5-1.0 mmol/kg, the final steady level of ionized Ca remains close to 100 nM. If, however, Ca is delivered at a lower initial concentration, Brinley et al. (1978) found that squid mitochondria are not active in sequestering Ca at Ca2+concentrations much below 200-400 nM. The reason for this apparent hysteresis in presumed mitochondria1 function is not clear. As with ATP-independent buffers, it is possible that extruded axoplasm is not fully representative of the whole axon, and the peripheral axoplasm that remains behind when axoplasm is extruded may contain other energy-dependent buffer systems. There is no direct evidence for this suggestion, although a peripheral reticulum that seems to bind Ca and may reflect a Ca-binding system additional to those described above has been described. 3 . Axoplasmic Ca-binding is impaired by a variety of heavy metals and also by sodium (Baker and Schlaepfer, 1975, 1978; Baker and Umbach, 1983). This latter effect is of particular interest because alterations in axoplasmic Na could alter the Ca buffering capacity of axoplasm under physiological conditions-a rise in Na decreasing and a fall increasing the buffer capacity. The Na seems to act on the energy-dependent, mitochondrial, component of binding and, by analogy with other mitochondria, may promote Ca efflux from the mitochondria into the cytosol by some sort of Na-Ca exchange (see, e.g., Carafoli and Crompton, 1978; Nicholls and Akerman, 1982).

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4-

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FIG.5 . Titration of Ca-binding sites in a sample of axoplasm extruded from a Loligo forbesi axon that had been stored in seawater at 4°C for 5 hr. The upper part of the figure shows that immediately after injection the ionized Ca rose and then fell to a steady level, which is the value plotted in the lower half of the figure. Ionized Ca was measured with a calcium electrode. Temperature, 20°C. (From Baker and Umbach, 1983.)

AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS

207

In conclusion, the massive Ca-binding capacity of isolated axoplasm sems to be the result of energy-dependent and energy-independent processes. The only energy-dependent process that has been clearly identified is the mitochondrion, although there is some evidence for an endoplasmic reticulum type of system as well. Energy-independent binding probably involves a number of molecular species. Operationally, the nondialyzable fraction can be subdivided into components with high affinity and small capacity (possibly calmodulin) and with much lower affinity, but large capacity. In addition, components of low molecular mass that can be dialyzed out of axoplasm may also bind calcium. Although soluble anions such as ATP, aspartate, and citrate are present in axoplasm in millimolar concentrations, their affinity for Ca is rather low, and Brinley (1978) has estimated their aggregate Ca-binding capacity as no more than 200 nmol per kilogram of axoplasm under physiological conditions. It is always possible that axoplasm may contain dialyzable molecules with much higher affinity for Ca, but none has so far been identified. It follows that when axoplasm loaded with 45Cais poisoned, 4SCashould be transferred from energy-dependent Ca-binding sites to nondialyzable, energy-independent ones with a resultant small rise in ionized calcium. Although in agreement with the direct determination of free Ca by means of calcium electrodes, this conclusion seems to be at variance with the finding of Blaustein and Hodgkin (1969) that poisoning renders the bulk of the 45Cacontent of axoplasm dialyzable. This apparent discrepancy almost certainly results from the fact that Blaustein and Hodgkin followed the loss of 45Cainto a solution that, although nominally Ca-free, contained micromolar concentrations of Ca that could exchange with the bound Ca in axoplasm, thereby rendering a much larger fraction of axoplasmic Ca apparently dialyzable than is in fact the case.

C. Calcium Fluxes across the Axolemma

1. THEPROBLEM DEFINED Despite the prodigious Ca-binding capacity of axoplasm and the existence of a steady inward leak of Ca, the Ca content of axoplasm is maintained relatively stable with the intracellular binding systems far from saturation. This balance is achieved by the operation of outwardly directed Ca pumps in the axolemma that serve, on average, to achieve an exact match between Ca influx and Ca efflux. In this section we examine the various routes by which Ca enters and leaves the axon; before discussing these in detail, it is necessary to consider some of the technical

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problems that must be taken into account in the experimental analysis of these fluxes. Axoplasmic Ca binding poses a number of experimental problems because it severely restricts the diffusibility of Ca within the axon. Thus, injected 4’Ca takes a very long time to become uniformly distributed within the injected section of an axon (see Blaustein and Hodgkin, 1969), and, conversely, Ca entering across the axolemma penetrates only slowly to the center of the axon (see Baker ef at., 1971). From the standpoint of flux measurements, enough time (1-2 hr) must be left after injecting 45Ca for the isotope to become uniformly distributed in the axon, and the possibility must always be considered that measurements of Ca influx by extrusion of axoplasm could be in error because high levels of 45Camay remain bound in the peripheral axoplasm that is not normally extruded. Axoplasmic Ca binding is a particularly acute problem when measuring Ca fluxes by the technique of intracellular dialysis (Brinley and Mullins, 1967). Although monovalent cations (Na, K), monovalent anions (Cl, aspartate, etc.), nucleotides (ATP, ADP, etc.) and other substrates which are either free or very loosely bound to intracellular structures can be controlled effectively by this technique, for solutes such as Ca that are strongly bound to intracellular structures, one must overcome the binding in order to obtain reliable flux measurements (DiPolo, 1973; Beauge and DiPolo, 1981). This is particularly important for Ca influx experiments, since all the Ca that enters the cell in the dialyzed segment of the axon must be collected by the porous capillary. The energy-dependent systems can be turned off either by eliminating their natural substrates or by adding suitable inhibitors to the internal solution; but in order to neutralize the energy-independent systems an exogenous Ca-sequestering agent must be included in the dialysis fluid. This buffer should have affinity and permeability characteristics (high permeability through the porous capillary, poor permeability across the axon membrane) such that when introduced into the dialysis capillary it should reach an axoplasmic concentration sufficient to overcome all the endogenous buffer systems. Furthermore, it should be nontoxic to the cell. EGTA meets all these requirements. Thus, 1. Under dialysis conditions (1-2 mM total EGTA, more than 300 F M free EGTA), the Ca influx measured at rest and during electrical stimulation (DiPolo, 1979; DiPolo et af., 1982) is not different from that obtained in intact axons (Hodgkin and Keynes, 1957; Baker r t cd., 1969a; Brinley rt al., 1977). 2. In the absence of EGTA in the dialysis fluid, the measured Ca influx is 5-10 times smaller than the expected influx. Addition of sufficient

209

AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS

free EGTA (at constant Cap), increases the measured influx to the expected value. Further increments in the free internal EGTA cause no variations in the influx (DiPolo, 1979). 3. Experiments on isolated squid axoplasm show that the levels of EGTA normally used during dialysis (300 pM free) effectively remove all the bound Ca. This confirms theoretical calculations based on the known affinities and capacities of the axoplasmic Ca buffer systems (DiPolo and BeaugC, 1980). The use of EGTA and Ca-EGTA buffers brings with it another problem. At physiological and near physiological levels of ionized Ca, most of the total added Ca (cold + radioactive) is bound to the EGTA and only a very small fraction is free. If Ca-EGTA can escape across the axolemma, this “leak” flux could obscure the true Ca efflux (Brinley et af., 1975). Therefore the Ca-EGTA “leak” must be measured directly in order to establish the true value for the Ca efflux. The mean efflux of [I4Ca]EGTA per I mM EGTA is 4.8 k 1.2 fmol cm-2 sec-’ (f/CS) (DiPolo, 1978).Fortunately from the point of view of measuring Ca efflux the Ca-EGTA “leak” is not affected by removal of Nai, Ca, , and Na, or by changes in intracellular levels of ATP (see Fig. 6). Calcium-binding is not restricted to the interior of the axon. The experiments of Baker and McNaughton (1978) revealed the existence of extra-

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cellular calcium-binding sites that can severely distort the appearance of the Ca efflux from injected axons. Thus, when axons are transferred into nominally Ca-free media " T a efflux falls, only to creep slowly back to its initial level over the next 30 min (Fig. 7). Reintroduction of Ca, or exposure to low concentrations of EGTA, produces a sharp rise in efflux that quickly falls toward its original value. The simplest interpretation of these findings is that " T a leaving the axon must pass through an extracellular Ca-binding matrix. In the nominal absence of Ca, the matrix takes up an appreciable proportion of the radioactivity leaving the axon, and the radioactivity reaching the external solution only begins to reflect that leaving the axon once the extracellular high-affinity Ca-binding sites have been filled, Replacing cold Ca, which can occupy the Ca-binding sites, or EGTA, which acts as a mobile Ca chelator, permits the bound radioactivity to be released, resulting in a transient increase in the amount of radioactivity to be released, resulting in a transient increase in the amount of radioactivity leaving the preparation. Once recognized, these transients can be minimized either by avoiding large changes in extracellular Ca or by including small amounts of CaO.CQ15

.

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AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS

21 1

EGTA or Ca-EDTA in all the experimental solutions. In axons of L . forbesi, brief exposure to Pronase also greatly reduces Ca transients (Baker and McNaughton, 1978).

2. COMPONENTS OF Ca INFLOW

a . General Features of Ca Injiux. Influx of Ca into freshly dissected axons immersed in Na seawater containing 10 mM Ca is close to 0.15 pmol cm-* sec-' at 20°C (Hodgkin and Keynes, 1957; Baker and McNaughton, 1976). Uptake approximates to a linear function of Ca, over the range 0-30 mM, but shows signs of saturating as Ca, is increased further. The following are the most striking features of the resting Ca influx: (1) It is reduced to about one-fourth by external application of TTX. (2) It is increased dramatically, in a TTX-insensitive fashion, in Nafree media, especially if Nq is high. Partial replacement of Na, also increases Ca influx and the increase is greatest when K is used as substitute for Na,. (3) It is unaffected by concentrations of cardiac glycosides that completely inhibit the Na-K exchange pump. (4)It is elevated in fully poisoned axons. In addition, Ca influx is increased during periods of electrical stimulation. b. The Underlying Components. There is general agreement that the membrane processes underlying the resting Ca influx include Ca entry in exchange for internal Na and Ca entry through TTX-sensitive Na channels. In addition, the resting Ca influx may, under certain circumstances, include other, less well defined, components of net Ca influx and one of Ca-Ca exchange. Entry of Ca through TTX-sensitive Na channels is much increased during electrical stimulation, but can account for only part of the total Ca influx seen under these conditions. The remaining, TTX-insensitive, Ca inflow seems to reflect the operation of a distinct voltage-sensitive Ca channel. The evidence for, and major properties of, each of these membrane processes will be discussed in turn. c. Ca Inflow That Requires Internal Na (Ca,-Nai Exchange). Complete replacement of external Na by Li, choline, Tris, K, or sugar brings about a large increase in Ca inflow (Baker et al., 1967, 1969b). The magnitude of this increase is very dependent on the Na content of the axon, but is always larger in Li and K seawaters than in similar solutions based on choline, Tris, or sugar. Experiments on intact axons suggest that Ca influx increases at least as the square of internal Na and possibly as a higher power, Influx of Ca seems to be associated with a specific component of Na efflux (Baker et al., 1967, 1969a). Thus, in the presence of sufficient cardiac glycoside to give complete inhibition of the Na-K exchange

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P. F. BAKER AND R. DIPOLO

pump, replacement of external Na by Li stimulates Na efflux, and this stimulation is not seen in the absence of external calcium. The properties of this Ca,-dependent Na efflux closely parallel those of the Nai-dependent Ca influx (Fig. 8). The apparent affinity of both processes for Ca, is a few millimolar in Li seawater, but falls progressively when Li is replaced by Na. Both processes are stimulated by intracellular alkalinization and inhibited by intracellular acidification (Baker and McNaughton, 1977; Baker, 1978), and both processes are activated by certain monovalent cations including Li, K, and low concentrations of Na. They are also inhibited by external application of lanthanum (1 mM) and by inclusion of Mg, Mn, Co, or Ni in the seawater. Inhibition by these last three agents is half-maximal at about 5 mM and is fully reversible (T. J . A. Allen and P. F. Baker, unpublished results). D600 (0.1 mg/mol, 0.25 mM) causes only partial inhibition, and both TTX and ouabain are without effect. Injection of orthovanadate to give a final concentration of 0.1 mM also has no effect on Ca,-dependent Na efflux (Baker and Singh, 1981). B

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FIG.8. Comparison of the effects of external sodium ( 0 )and lithium (0) ions on the Cadependent Na efflux (A) and on the Ca influx (B) in squid giant axons. Isotonicity was maintained throughout the dextrose. Seawaters contained 460 mM of monovalent cation or an osmotically equivalent amount of dextrose, 55 rnM MgCl?, 11 mM CaClz, 2.5 mM NaHCO?, and lo-' M ouabain. Temperature, 18-20°C. (From Baker, 1970.)

213

AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS

The Ca,-dependent Na efflux is very sensitive to metabolic inhibitors and after application of cyanide (or FCCP) falls rather quickly: after about 0.5-1 hr no Ca,-dependent Na efflux can be detected. The Ca,-dependent Na efflux recovers rapidly on removing the cyanide. As mentioned in Section IV,C) metabolic inhibitors bring about both a fall in ATP and a delayed rise in ionized calcium. The rapid fall in Ca,-dependent Na efflux correlates rather well with the disappearance of ATP, and this is supported by the finding that injection of apyrase, which lowers cytoplasmic ATP without altering ionized Ca, also brings about a rapid fall in Ca,dependent Na efflux. The other possibility, that Ca,-dependent Na efflux is inhibited secondarily to a rise in cytosolic ionized Ca is difficult to examine experimentally in intact axons because injection of Ca-chelating agents such as EGTA, EDTA, CDTA, and NTA all inhibit Ca,-dependent Na efflux (and the associated Ca influx) from unpoisoned axons (Baker, 1970, 1972; Baker and McNaughton, 1976a) (Fig. 9). Appreciable inhibition is still seen even when the chelating agent is injected in its Mg or Ca form. This is a puzzling observation because one might have expected that steepening the inward gradient for Ca would, if anything, increase the rate of Ca,-dependent Na efflux. A very plausible explanation for these findings, however, has emerged from studies on dialyzed axons (DiPolo, 1979; DiPolo e l a l. , 1982). When squid axons are dialyzed with solutions containing Na, Ca, and ATP, an Nai-dependent Ca influx can be demonstrated. An interesting M ouabain

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P. F. BAKER AND R. DIPOLO

property of the Nq-dependent Ca influx in squid axons is its dependence on the level of internal ionized Ca as is shown in Fig. 10. In the complete absence of Caf' (2 mM EGTA and no added Ca), there is no measurable Nai-dependent Ca influx. Raising the ionized Ca increases this component until it saturates at micromolar Ca concentrations. Most of the activation occurs in the range of Caf' from 0.1 to 1 .O p M and is absolutely dependent in the presence of Nai and ATP. The Nq and ATP requirements argue against a simple Ca-Ca exchange mechanism being responsible for this phenomenon. Whether the Ca, that activates Ca influx is also translocated to the outer face of the axolemma remains an open question; however the magnitude of the Nq-dependent Ca influx activated by Cq (about 200 fmol cm -2 sec-l at K l i 2 )is much greater than that of the Ca,-dependent Ca efflux (about 10 fmol cm-* sec-I) under similar experimental conditions, which argues strongly against this possibility. These experiments provide strong support for the view that Ca,-Nq exchange is activated by intracellular calcium. This form of activation must provide a system of positive feedback, the physiological implications of which have not been explored. In the dialyzed axon, internal Na activates the Ca influx along a sigmoidal curve, suggesting, in agreement with studies on intact axons, that more than one Na is exchanged for each Ca ion. The half-saturation constant for activation of the Ca influx by Nai decreases from about

-

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215

AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS

60 mM at 0.06 pM C q to 38 mM at 0.8 p M Cai (Fig. 11). At relatively high Cq (0.8 p M ) , the relation between Ca influx and Nai can be best fitted on the assumption that two Na ions exchange with each Ca ion, but this relationship does not hold at low levels of ionized Ca, where the data are more consistent with exchange of one Ca for more than two Na ions. This finding might imply that the stoichiometry of Nai-Ca, exchange may vary with the level of ionized Cai (see Mullins and Brinley, 1973, but a clear answer requires simultaneous measurement of both Na,-dependent Ca influx and Ca,-dependent Na efflux, and this has not so far been possible. The only direct data on the stoichiometry of Ch-Nq exchange come from the experiments of Baker et al. (1969b) on intact axons. They concluded that in Li seawater roughly 3-5 Nq exchanged with each Ca,, whereas in choline seawater the value was closer to 3 Na: 1 Ca. It should be stressed, however, that these values were not determined simultaneously on a single axon. Nevertheless, the data strongly suggest that more positive charge leaves the fiber associated with the Na arm of the exchanger than enters with calcium. It is not clear whether the external monovalent cations that activate Nq-dependent Ca influx (see Fig. 8) are also transported; but even in sugar solutions the Ca,-dependent Na efflux

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216

P. F. BAKER AND R. DIPOLO

is greater than the associated Ca influx, suggesting that the exchanger is inherently electrogenic. The possibility has been examined directly by measuring Ca,-dependent Na efflux under conditions where the membrane potential is varied electrically (Baker and McNaughton, 1976b; Allen and Baker, 1983, and unpublished observations). The finding is quite clear. Electrical depolarization stimulates Ca,-dependent Na efflux and electrical hyperpolarization inhibits. For a given depolarization, the stimulation brought about by a rise in external K is much greater than that brought about electrically in the absence of external K (Fig. 12). In other words, a large fraction of the stimulation of &,-dependent Na efflux brought about by external K is due to K ions per se, and only a small part is due to depolarization. K ions

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FIG.12. Comparison of the effects on Ca,-dependent Na efflux of altering the membrane potential either by KOor current. Note that electrical hyperpolarization fails to remove all the activation brought about by 50 m M K, and electrical depolarization fails to mimic the effects of applying K externally. The experiment was carried out in choline seawater conM ) . The axon was preinjected with TEA, taining tetrodotoxin (0.6 p M ) and ouabain HEPES, and MOPS to give a final concentration of each close to 20 mM, pH 7.3. Temperature, 18°C. (Unpublished data of T. J. A. Allen and P. F. Baker.)

AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS

217

are, therefore, acting very like Li ions (which do not alter E m )to stimulate Ca,-dependent Na efflux; but in both instances depolarization can bring about further activation. A full voltage sensitivity of Ca-dependent Na efflux is not available, but the flux is still increasing at +40 mV (T. J. A. Allen and P. F. Baker, unpublished results). Experiments with dialyzed axons support the view that external K activates Ca,-Na, exchange. Thus, prolonged exposure to external K causes a maintained increase in Ca influx. This increment is observed only in the presence of internal Na, and its magnitude depends on the level of Ca?', an increment in ionized Ca from 0.1 to 0.6 pM increasing Ca entry 10-fold. The effects of K and potential have not been compared directly in the dialyzed axon. In summary, the Na,-dependent Ca influx requires ATP and intracellular Na and is activated by internal Ca and inhibited by external Na and a variety of externally applied di- and trivalent cations including Mn, Co, and La. It appears to involve the exchange of at least 3 Na for each Ca, and, consistent with this stoichiometry, the rate of exchange is increased by depolarization and decreased by hyperpolarization. d . Ca Influx through Tetrodotoxin (TTX)-Sensitive N u Channels. The resting Ca influx into freshly dissected axons immersed in Na seawater containing 10 mM Ca is about 0.16 pmol cm-* sec-'. Addition of TTX to the seawater reduces this flux to about one-quarter (unpublished observations of P. F. Baker), suggesting that Ca can enter the axon through TTXsensitive Na channels that are open at the resting potential. The finding of Baker et al. (1969a) that TTX reduces the resting Na influx from 27.6 to 11.2 pmol cm-* sec-' is fully consistent with this view. The actions of TTX on resting Ca influx have been investigated in some detail in dialyzed axons. Under conditions chosen to resemble physiological (Na, = 40 mM; Ca, = 0.1 pM; ATP = 2 mM) the resting Ca influx varies from 0.1 to 0.14 pmol cm-2 sec-' when Na, and Ca, are 440 and 10 mM, respectively (DiPolo, 1979; DiPolo et al., 1982), which is in the range reported for intact axons (Hodgkin and Keynes, 1957; Baker et al., 1969b; Blaustein and Hodgkin, 1969; DiPolo and Beauge, 1979, 1982). Under these conditions, 60-70% of the total Ca influx is sensitive to external TTX, 20% is dependent on internal Na (Na,-dependent Ca influx), and the remaining 10% has the characteristics of an unspecific leak (insensitive to Na,, Cai, ATP, and D600). Inhibition by TTX is half-maximal at 5 nM, which is very similar to the concentration required to block half the Na channels. The existence of open Na channels in the dialyzed axon can be demonstrated in axons depleted of ATP, where the passive Na influx is about I8 pmol cm-* sec-' in the absence of TTX and only I .3 pmol cm-* sec-l in the presence of 1 p M TTX (Id. Rojas and R. DiPolo, unpublished

21 8

P. F. BAKER AND R. DIPOLO

information). DiPolo e f al. (1982) have estimated the ratio pca:pNafor the TTX-sensitive channel to be close to 1 : 10. In view of the fact that Ca enters through TTX-inhibitable channels in the resting axon, it would be surprising if electrical stimulation did not enhance Ca entry by this route. The extra Ca influx associated with electrical activity averages 0.006 pmol cm-* per impulse at 11 mM Ca,. It increases roughly linearly over the range 0-112 mM Ca, and a flux of similar magnitude can still be detected in fully poisoned axons. Voltageclamp experiments on axons containing aequorin showed that the voltagesensitive extra entry of Ca can be divided into two roughly equal components: one that is inhibited by TTX and a second that is not (Baker et uf., 1971). The time course and other properties of the TTX-inhibitable component are identical with those of the Na channel, leaving little doubt that Ca entry through the Na channel is an important route by which Ca enters axons. The data suggest that pca:p~~ for the Na channel is about 1 : 100, which implies greater selectivity during activity than at rest. Further support for entry of Ca through Na channels was obtained in experiments on perfused axons (Tasaki et al., 1966, 1967; Watanabe et al., 1967; Meves and Vogel, 1973), where, under appropriate conditions, Ca action potentials can be observed. Ca action potentials are usually seen only in axons that have been immersed in Na-free seawaters containing 100-200 mM CaCI2 after perfusion with low ionic strength media based on glycerol or sucrose and containing 25 mM CsF. Under these conditions it is possible to detect a slowly inactivating, TTX-sensitive, Ca inward current. The reversal potential of this current is close to +55 mV, and it would seem that Cs ions can move outward through the TTXsensitive channel. The data of Meves and Vogel(l973) are consistent with the existence in perfused axons of a TTX-sensitive Na channel that also permits the entry of calcium. The ratio pca:pNafor this channel is close to 1 : 10; this, when compared with the aequorin data mentioned above, suggests that perfusion with low ionic strength media may alter the selectivity of the N a channel. e . Ca Influx through TTX-Znsrnsitive Cu Chunnels. The main evidence for the existence of specific Ca channels in the squid axon (as opposed to the squid giant synapse) comes from experiments in which intracellular free Ca was monitored with aequorin (Baker et al., 1971). Voltage-clamp analysis revealed TTX-sensitive and TTX-insensitive components of Ca influx. After a step depolarization, the TTX-insensitive component increased with much the same time course as the outward K current, but was unaffected by TEA concentrations that blocked the outward K current. In addition, it exhibited a bell-shaped voltage dependence with a peak close to zero potential (Fig. 13A), was rather insensitive to external

219

AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS

A

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FIG. 13. Voltage dependence of (A) tetrodotoxin (TTX)-insensitive Ca entry measured with aequorin in an intact axon of Loligo forbesi and (B) Ca-inward current in a dialyzed axon of L . pealei. The curves in (A) relate to Na seawater containing TTX and 112 mM Ca, 0 Mg (curve I), 20 mM Ca, 0 Mg (curve 2), and 20 mM Ca, 90 m M Mg (curve 3). Abscissa: Depolarization from the resting potential of 60 mV. Data in (B) were obtained from an axon dialyzed with a solution containing 20 mM fluoride, 180 mM glutamate, and 200 mM N methylglucamine together with sucrose to give a final osmolarity of 980 mOsm/liter, pH 7.4. The external medium was based on trismethane sulfonate and contained 300 nM TTX. HP, potential at which the axon was maintained between pulses. (Panel A from Baker et a/., 1971; panel B from DiPolo et ul., 1983.)

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Na, was reduced by external Mg and external application of Mn, Co, La, and D 600, and was not maintained but declined (inactivated) slowly over a time course of many seconds (Baker ef al., 1971, 1973a,b). Although in the experiments using aequorin no clearly defined inward Ca current was detected, Baker et (11. (1971) and Baker et al. (1973a,b) ascribed the voltage-sensitive and TTX-insensitive phasic component of Ca entry to a Ca channel. This conclusion seemed to receive strong support from the discovery of an inward Ca current in the presynaptic terminal of the squid giant synapse (Katz and Miledi, 1969; LlinAs et d.,1981). the properties of which bore a striking resemblance to those of the putative axonal Ca channels. If, instead of an electrical depolarization, axons are depolarized by rapid exposure to high potassium, the aequorin response consists of phasic and tonic components (Baker et al., I973b). Baker et ul. (1973b) and Baker and Glitsch (1975) interpreted the phasic response as reflecting a transient opening of a Ca channel and the maintained, tonic, response as the manifestation of another voltage-sensitive process-perhaps Na,-Ca, exchange-which, as discussed earlier, is sensitive to membrane potential. Experimental evidence indicates that the tonic component of the aequorin signal is due to a Na-Ca exchange process, since it is greatly reduced in axons containing low Na, (Mullins and Requena, 1981) and is absent in axons dialyzed without internal Na. On the other hand, the interpretation of the phasic component has been questioned by Mullins (1981), Mullins and Requena (1981). and Mullins et al. (1977), who suggest that both phasic and tonic components of Ca entry may be manifestations of Na,Ca, exchange. Their new piece of evidence is that the Ca transients evoked by exposure to high external K are much reduced in size, although still very definitely transient, in axons that have been depleted of Na by repetitive stimulation in Li seawater. At first glance this interesting observation seems to give strong support to an explanation in terms of Na,-Ca, exchange, although it is still necessary to explain why the exchange exhibits phasic and tonic components. Here, Mullins invokes the fact that accumulation of Ca tends to bring about a fall in cytosolic pH (Baker, 1972; Meech and Thomas, 1977; Mullins and Requena, 1979) and Na,-Ca, exchange is known to be sensitive to cytosolic pH (see Section IV,C), a fall in pH, inhibiting, and a rise accelerating, Na,-Ca, exchange (Baker and McNaughton, 1977; Baker and Honerjager, 1978; Baker, 1978). The main experimental objection to this explanation is that small, transient light responses can be seen when K is applied in media containing rather little Ca, whereas exposure of these same axons to Ca-rich seawaters based on lithium brings about a much larger, but maintained, increase in

AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS

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aequorin light output. This suggests that the transient response is a feature of K depolarization, not solely a function of Ca entry. Other observations also argue against Mullins’ interpretation. Thus, (1) the phasic component of entry is strongly reduced by D 600, which has rather little effect on Nq-Ca, exchange; (2) the phasic component of entry begins to decline at potentials positive of zero, whereas Nq-Ca, exchange does not fall and, if anything, increases at potentials positive of zero (Allen and Baker, 1983, and unpublished observations), and (3) a phasic component of Ca entry persists in apyrase-injected axons where Na,-Ca,, exchange is completely inhibited. In addition, DiPolo et al. (1983a) have described an inward Ca current in dialyzed squid axons (Fig. 8). The voltage dependence, inactivation characteristics, and pharmacological properties of these Ca currents are remarkably similar to those described by Baker e t al. (1971, 1973b) on the basis of aequorin experiments and also closely resemble the Ca currents characterized in some detail by Lliniis et uf. (1981) at the squid giant synapse. Because these inward Ca currents are not maintained, but inactivate over a period of seconds, they are unlikely to contribute significantly to the net Ca entry during prolonged depolarization where the main factor determining Ca entry is likely to be Nai-Ca, exchange. If the TTX-insensitive phasic component of Ca entry does indeed reflect Ca entry through a Ca channel, we still need an explanation for the finding of Mullins and Requena (1981) that the phasic response is much reduced in Na-depleted axons and increased in Na-loaded ones. Two possible explanations merit consideration: (1) It may be a property of the aequorin light response, which is known to be proportional to [CaiI2,and the light output in response to an increment in Ca entry (ACa) will be (Y (CaR + ACa)*, where CaRis the resting ionized Ca. Even if ACa remains constant, reducing CaR,as may happen in Na-depleted axons, will lead to an apparent reduction in the size of the Ca transient. (2) Alterations in Nai may change the buffering capacity of axoplasm such that as Nai is reduced the buffering capacity increases. This is in line with the experimental findings discussed in Section IV,B. More experiments are needed to settle this very important question. In addition to Ca entering through TTX-sensitive Na channels and TTX-insensitive Ca channels, other experiments have been interpreted in terms of Ca entry through the voltage-sensitive K channel (Keynes et al., 1979; Inoue, 1980); however, more experiments are needed to establish this possibility. f. Ca InJEux by Other Routes. Although the pathways that have been discussed so far probably constitute the major routes by which Ca enters

222

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R. DIPOLO

axons under physiological conditions, additional pathways become evident in fully poisoned axons, and it is quite possible that these may contribute, in small measure, to the resting Ca influx in unpoisoned axons (see Baker and McNaughton, 1978). Ca influx is increased in fully poisoned axons. At least part of this increased Ca influx probably reflects the operation of an exchange of external Ca for internal Ca. In support of this view is the finding that a component of the Ca efflux from poisoned axons is dependent on the presence of external Ca. However, no one has demonstrated that injection of enough EGTA to lower internal free Ca in fully poisoned axons inhibits Ca influx. The properties of Ca influx and Ca,-dependent Ca efflux in poisoned axons resemble those of Ca,-dependent Na efflux in unpoisoned axons in being activated by certain monovalent cations in the external solution. Thus the flux is much larger in Li and K seawaters than in choline or Tris seawaters. As Ca,-dependent Na efflux is completely inhibited in poisoned axons (see Section IV,C), it is possible that under these conditions the external Ca-binding site now interacts with an internal Ca-binding site instead of one that binds sodium, There is general agreement that axons dialyzed with high levels of Ca exhibit Ca,-dependent Ca efflux, and Blaustein (1977) has shown that this flux is activated by both external and internal monovalent cations, but a full analysis of the properties of the Ca influx into ATP-depleted axons is still needed. A characteristic of Ca influx into unpoisoned axons is its stimulation following replacement of external Na by a whole host of Na substitutes, such as Li, choline, and dextrose. Such stimulation is still seen in poisoned axons, where Ca, no longer exchanges for Nai. After injection of apyrase to lower ATP and inhibit Nai-Ca, exchange without increasing Cai, removal of Na, still brings about a marked rise in cytosolic Ca (measured with aequorin), suggesting that Ca may now be entering the axon in an uncoupled fashion. More experiments are clearly needed, especially under conditions of internal dialysis. 3. COMPONENTS OF Ca OUTFLOW

a . General Features of Ca Efflux: Intact Axons. As mentioned in Section IV,C, the study of Ca efflux can be complicated by the presence of an extracellular Ca-binding matrix of high affinity; however, when the properties of this matrix are taken into account, the bulk of the Ca efflux from unpoisoned axons comprises two apparently independent components: one that requires extracellular Na and a second that is not obviously coupled to the movement of any other ion (see Fig. 7). The Na-dependent component normally comprises 10-30% of the total Ca efflux. Poisoning the axon increases the total Ca efflux, which is now composed largely of

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Na- and Ca-dependent components and the uncoupled flux is no longer detectable (Baker et ul., 1967; Rojas and Hidalgo, 1968; Blaustein and Hodgkin, 1969; Baker and McNaughton, 1976a, 1978). The loss of the uncoupled efflux seems to result rather directly from the fall in ATP, whereas the increase in Na,-dependent and Ca,-dependent fluxes reflects the increase in free Ca. Thus the total Ca efflux falls dramatically if intact axons are depleted of ATP under conditions where the rise in free Ca does not take place, for instance after injecting apyrase or exposure to cyanide after injection of enough EGTA to buffer the free Ca at its resting level (Baker and Glitsch, 1973; Baker and McNaughton, 1976a). Although the uncoupled flux seems to be absolutely dependent on ATP or something derived from it and the Na-dependent flux persists in the nominal absence of ATP, the Na-dependent flux is not unaffected by the metabolic state of the axon. Thus, the curve relating Na-dependent flux to external Na is sensitive to the metabolic state of the axon, the apparent affinity for external Na being markedly reduced, in fully poisoned axons (Baker and Glitsch, 1973; Baker and McNaughton, 1976b). b. General Features of the Ca Efflux: Diulyzed Axons. Many of the problems outlined above should be greatly simplified in dialyzed axons where ionized Ca and ATP can both be controlled. However, many of the early experiments with dialyzed axons failed to detect the uncoupled Ca efflux, and this has caused some confusion in the literature. The reason for this is not altogether clear; but may be related to the failure in many early experiments to maintain a low, physiological, ionized Ca and high ATP: ADP ratio in the dialysis fluid. The easiest way to ensure a high ATP: ADP ratio is to include arginine phosphate as well as ATP in the dialysis solution. In dialyzed axons in which the concentration of free Ca is kept constant with EGTA, four components of Ca efflux can be distinguished: one dependent on external Na (Na,-dependent Ca efflux) (DiPolo, 1973; Blaustein, 1974; Mullins and Brinley, 1975), one activated by external Ca (Ca,-activated Ca efflux) (DiPolo, 1973; Blaustein, 1974,1977), the uncoupled efflux (Ca efflux observed in the absence of Na,, Ca,, and Mg,) (DiPolo, 1978; DiPolo and Beauge, 1979), and the “leak” of Ca-EGTA (Brinley et al., 1975; DiPolo, 1978). The relative contributions of these components to the total Ca efflux is markedly influenced by the experimental conditions. There is still some controversy concerning the nature of the uncoupled or “residual” Ca efflux. It has been proposed that it may be part of the Na-Ca exchange system (Requena and Mullins, 1979; Blaustein and Nelson, 1982). Two specific arguments have been advanced: (1) that the residual uncoupled flux may actually reflect exchange of Ca for the Tris or

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choline used to replace external Na; and (2) that the residual uncoupled flux may in fact be Na,Ca, exchange maintained by Na pumped or leaking into the very restricted mesaxonal space between the axon and surrounding Schwann cells (Frankenhaeuser and Hodgkin, 1956). The first possibility can be ruled out by the finding that the uncoupled Ca efflux in both intact and dialyzed axons is not altered when Na, and Ca, are replaced isosmotically by nonelectrolytes such as glucose, mannitol, or sucrose. Rather clear evidence against the second (Na leak) possibility is provided by Fig. 14, where it is shown that, in the absence of Na,,, removal of internal Na has rather little effect on the magnitude of the ATP-dependent uncoupled efflux (see also DiPolo and Beauge, 1979). These experimental findings, taken together with the fact that changes in Na,, Na,, Ca,, and ATP do not seem to affect the ‘‘leak’’ of Ca-EGTA, strongly support the existence of a genuine ATP-dependent, apparently uncoupled, Ca efflux in squid axons. The importance of the level of ionized Ca in determining the relative sizes of the various components of the Ca efflux is illustrated in Fig. 15,

I OATP-OPAI I KT

60 No

.

3lOK

2 rnMATP-5rnMPA IONa

I

OK

I

60No

310K

TIME (hr) FIG. 14. The effect of external and internal potassium ions on the ATP-dependent uncoupled Ca efflux in an axon dialyzed with 0.06 p M Ca2+and 2 mM ATP. Tris ions were used to substitute for K+.Note that 100 mM KO has no effect (O), that the uncoupled component is inhibited by the removal of K,, and that there is further inhibition upon removing Na, in the absence of K,. Temperature, 18°C; axon diameter, 500 pm. (Unpublished data of R. DiPolo and L. A. Beaugk.)

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which compares the behavior of the Ca efflux from two axons dialyzed with similar internal solutions, except that in one ionized Ca is 80 nM (i.e., close to physiological) and in the other it is 200 p M , approximately 250 times higher than the resting level. The most relevant conclusions derived from these experiments can be summarized as follows: 1. At low internal ionized Ca, and in the absence of ATP (Fig. 15A), the Ca efflux stabilizes at a value not significantly different from the expected Ca-EGTA "leak" (2-3 fmol cm-2 sec-I) (see also DiPolo, 1977).

4 30 0[**

0

*-0

v 0

0

0

ATP 1-

1

-0

I

3 TIME (hr)

ooOoo,

4

FIG. 5. Calcium efflux and internal free Ca. Free Ca was 80 nM in (A) and 200 p in (B). The dialysis fluid contained ( m M ) : K, 310; Na, 60; Mg, 4 in excess of the ATP concentration; Tris, 20; chloride, 99; aspartate, 310; total EGTA, 1; glycine, 330; oligomycin, 10 pg/ml; cyanide, I ; osmolarity, 980 rnOsm; pH, 7.3. The external solution contained (rnM): Na, 440; K , 10; Mg, 50; Ca, 10; Tris, 20; C1, 590; EDTA, 0.1; osmolarity, 1000 rnOsm; pH, 7.6. Temperature, 18°C. Vanadate ( V ) was added internally at a concentration of 2 m M . (Unpublished data of R. DiPolo and L. A. Beauge.)

226

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2. At low internal ionized Ca, internal ATP promotes a sizable increase in the Ca efflux above that of the Ca-EGTA “leak.” Removal of Na, and Ca, causes a small drop in the efflux (10-15%), confirming the observation that under nearly physiological conditions the Ca efflux is largely ATPdependent (85-90%) and Na,, and Ca, insensitive (DiPolo and Beauge, 1979, 1980). 3. At high internal ionized Ca and in the absence of ATP (Fig. 15B), Ca efflux is much greater than the Ca-EGTA “leak” and completely dependent on external Na and Ca, although dependence on Ca, is not shown in Fig. 15. This observation, originally reported in dialyzed axons by DiPolo (1973), shows that activation of the Ca efflux by Na can occur in the nominal absence of ATP. 4. At high internal ionized Ca, ATP activates the Ca efflux in the absence of Na, and Ca, (uncoupled component) and induces a further activation of the Na,-dependent component (ATP-stimulated, Na,-dependent Ca efflux) (DiPolo, 1974; Mullins and Brinley, 1975; Blaustein, 1977). It should be noted that at high internal ionized Ca and in the presence of ATP, the Na,-dependent component of Ca efflux is almost 10 times greater than the uncoupled component. This result contrasts markedly with that obtained at low physiological levels of ionized Ca, where the uncoupled component of Ca efflux predominates. To summarize, experiments with dialyzed axons give clear evidence for an ATP-dependent component of Ca efflux that has a high affinity for internal Ca, but relatively low capacity, and a Na-dependent component that persists in the nominal absence of ATP and has a rather low affinity for internal Ca, but a large capacity (Fig. 16). The properties of these two components of the Ca efflux are summarized in Table I and discussed in more detail below. c‘. ATP-Dependent, Apparently Uncoupled C a Efflux. As its name implies, this component of Ca efflux has a very specific requirement for ATP. Of a variety of other nucleotides examined, only 2-deoxy-ATP and t h e hydrolyzable analog AMpCpp can replace ATP; other naturally occurring high-energy phosphate compounds (CTP, UTP, UDP, ADP, AMP, and acetyl phosphate) do not promote Ca efflux (DiPolo, 1977). The nonhydrolyzable anaIog AMppCp inhibits the ATP-stimulated Ca efflux, probably by competing with ATP at the active site. The dialysis technique has made possible a detailed analysis of the kinetics of activation of the uncoupled flux by ATP. Activation proceeds along a hyperbolic curve with half-maximal activation requiring 20 p M ATP (Fig. 17). Internal Mg is an absolute requirement for the activating effect of ATP, and the flux is fully activated by 0.2 mM Mg. All these data strongly suggest that ATP is

AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS

P

0

uncoupled

A A No, -dependen1

0 01

01

I

10

100

FIG. 16. The activation of both the ATP-dependent uncoupled and the Na,-dependent Ca efflux by Ca,2+. 0, Data obtained in the absence of external Na and Ca; A , A, data obtained in the presence of external Na. The ATP concentration was 2 mM and the "a], was 45 mM. Open symbols are read on the right-hand scale; filled symbols are read on the left-hand scale. The points represent the mean of two or more determinations. Error bars are shown for more than four determinations. Ionized calcium in the dialysis medium was controlled by varying the ratio Ca-EGTA :free EGTA at a constant total EGTA of 1 mM. The apparent dissociation constant for the Ca-EGTA complex was chosen as 0.15 p M (DiPolo er a / . , 1976). The horizontal dashed line corresponds to a resting Ca influx of about 44 f/CS (from 4 mM Ca,). (From DiPolo and BeaugC, 1979, 1980, and unpublished data of DiPolo and Beauge.)

hydrolyzed in association with the uncoupled Ca efflux; but the small size of the flux has so far precluded a direct demonstration of this important point. The activation of the uncoupled efflux by Ca, is best described by a section of a rectangular hyperbola: half-maximal activation requires about 200 nM Cai and the flux saturates at about 200 fmol cm-2 sec-I. Of particular interest is the finding that the size of the ATP-dependent Ca efflux is unaffected by alterations in internal sodium but is rather sensitive to changes in internal potassium, a rise in Ki activating the flux. In addition, the flux is rather insensitive to changes in internal pH over the range pH 6.5 to 8.5, but is inhibited as pHi is lowered below 6.5 (see Fig. 20). The sensitivity to the presence of internal K seems to be an effect of the K ions per se rather than the result of alterations in membrane potential, because changing Em by varying external K has no effect on the uncoupled Ca

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COMPARISON

OF UNCOUPI-ED AND

Component

TABLE I Na-DEPENDENT COMPONENTS OF Ca EFFLUX Uncoupled

Na-dependent

[Cali giving half-maximal activation ATP

0.2 pM

10 p M

Essential

[ATP] giving half-maximal effect Na , Na,

20 pM

Persists in nominal absence, but ATP alters Na-activation kinetics 300 pM

K,

Activates No effect Inhibits ( K , = 7 p M ) Inhibits Inhibits

Essential Inhibits, especially in the absence of ATP Activates at constant En, Inhibits No effect at 100 p M Inhibits Small activation

N o effect

Activates

De polarization Internal vanadate External lanthanum ( I mM) Increase in external pH from pH 8.0 to pH 9.0 Increase in internal pH from pH 7.3 to pH 8 3

No effect No effect

efflux. This asymmetrical action of K ions is indicative of a role for internal K as an activator of the ATP-dependent Ca efflux. A similar role for K ions has been reported for other preparations, such as sarcoplasmic reticulum and red blood cells (Duggan, 1977; Rega rt al., 1973). Interestingly, in these cases Na ions can partially substitute for K in the activation of the Ca pump. This might explain the further small inhibition of the uncoupled Ca efflux induced by the removal of Na, in the absence of Ki (see Fig. 14). The fact that the K-like effect of Na ions is not seen in the presence of a normal internal K concentration (DiPolo and Beauge, 1979) supports this suggestion. It is not known whether the activating monovalent cation is also transported. Recent experiments with voltage-clamped dialyzed axons indicate that Ki also activates Na,,-dependent Ca efflux (R. DiPolo and H . Rojas, unpublished results.) A question of some importance is whether the ATP-dependent Ca efflux is really uncoupled in the sense that the outward movement of positively charged Ca is not compensated by the inward movement of some other positive charge or the outward movement of negative charge. No evidence for such coupling has been found, and the uncoupled Ca efflux is not obviously sensitive to changes in Em.The only data that might be interpreted in these terms is the finding that the uncoupled Ca efflux is strongly inhibited at alkaline external pH (Fig. 18) (DiPolo and Beauge,

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AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS

02

06

08

10

FIG.17. Activation of the components of the Ca efflux by ATP. The graph shows the components of the Ca efflux activated by ATP only in relative terms. The fraction of the Ca efflux that is Na, dependent and persists in the absence of ATP is not shown. Ordinate: Ca efflux relative to that in the presence of 2 mM ATP. Abscissa: ATP concentration in the dialysis fluid. Phosphoarginine (PA) ( S mM) was always present in the internal medium when testing different [ATP]. The usual procedure was to deplete the axons of ATP by dialyzing them with ATP-free solutions and then to follow the Ca efflux at different levels of ATP. The ATP-dependent Na,-dependent component represents the difference between the total Ca efflux measured in Na-ASW and that measured in the absence of Na, and Ca, (ATPdependent uncoupled). Note a 10-fold difference in the apparent affinity for ATP between the two components. The points represent the mean SEM. The number of experiments is given in parentheses. (From DiPolo and Beauge, 1979, and unpublished data of DiPolo and Beauge.)

*

1982), which suggests that the Ca pump has a requirement for external H ions. This might arise either from the presence of an ionizable group or because the uncoupled pump actually exchanges internal Ca for external H. Niggli et al. (1982) have provided some evidence for ATP-dependent Cai-H, exchange by the erythrocyte Ca pump. The finding that alkaline external pH inhibits the ATP-dependent Ca efflux reversibly has been a useful experimental tool to separate Na-dependent and “uncoupled” components of pumping without recourse to alterations in external sodium. In addition to alkaline external pH, the “uncoupled” Ca efflux is inhib-

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P. F. BAKER AND R. DIPOLO

3

.

0.0015-

A

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E m

k

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‘&

0.

0.0010-

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s

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.-c0

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i

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1

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1

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P-

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0, 3

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7.0

8.0

9.0

10.0

PHO

FIG.18. Influence of external pH on uncoupled Ca efflux from an intact axon of Loligo fouhcsi. (A) Raw data; (B) collected data from this and a second experiment. pH, was buffered with a mixture of HEPES and PIPES. Temperature, 20°C. Na was replaced isosmotically by choline. (Unpublished data of P. F. Baker.)

ited by external Ca (DiPolo and Beauge, 1982), by external application of lanthanides, and by internal application of vanadate. Lanthanum has proved to be very nonspecific, since it blocks both the “uncoupled” and the Na,-dependent components of the Ca efflux (Baker et al., 1969a; van Breemen and de Weer, 1970; Baker and McNaughton, 1976a, 1978); but vanadate is much more specific (DiPolo et al., 1979; DiPolo and Beauge, 1981; Baker and Singh, 1981). Orthovanadate has been shown to be a

AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS

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potent inhibitor of the Na and Ca transport systems in a variety of tissues (see BeaugC and Glynn, 1978; Bond and Hodgkins, 1980; Barrabin et al., 1980), and the main effects of internally applied orthovanadate on the squid axon are listed below.

1. Internal vanadate inhibits the ATP-dependent uncoupled Ca efflux with high affinity (K112= 7 p M ) . The inhibition is fully reversible. 2. In the absence of ATP, vanadate has no effect on the Na,-dependent component of Ca efflux. Vanadate is also without effect on the Ca,dependent Na efflux. 3. The site for vanadate inhibition of the uncoupled Ca efflux shows positive interaction with K ions: internal K increases the affinity of the intracellular vanadate binding site. External K has no effect on the inhibition. 4. External Ca antagonizes the inhibition by internal vanadate. In axons dialyzed with 1 mM vanadate, raising Ca, from 0 to 3 mM brings about 50% reversal of the inhibition. This effect can be overcome by further increasing the internal vanadate concentration. The remarkable similarity in the cationic requirements and sidedness of vanadate inhibition between squid axons and other preparations that are known to exhibit ATP-driven Ca pumps strongly indicates that this active mechanism is responsible also for the uncoupled Ca efflux in squid axons. An ATP-driven pump of this type must manifest itself as a membranebound, Ca-dependent ATPase. Membrane fragments isolated from the optic nerves of the squid exhibit Ca*+,Mg2+-ATPaseactivity (Beauge et al., 1981). This preparation produced a high yield of axolemma with negligible contamination from Schwann cells and intracellular organelles (Condrescu et al., 1984). The mernbrane-bound ATPase is half-maximally activated by 0.5 p M Ca and 20 p M ATP, is insensitive to ouabain and oligornycin, and is inhibited by nanomolar concentrations of vanadate. In addition, like many other Ca2+-ATPases, it is activated by calmodulin (Condrescu et al., 1984), and membrane vesicles from this preparation show ATP-dependent Ca uptake (L. Osses, unpublished results). The biochemical characteristics of this Ca2+-ATPaseclosely resemble the behavior of the uncoupled Ca pump and also share many features in common with Ca*+-ATPases from other membrane preparations (Schatzmann, 1981). d . Nu,-Dependent Ca Efflux. The Na,-dependent Ca efflux is, by definition, the difference in Ca efflux between Na-containing and Na-free solutions. This method of defining the flux carries with it a number of possible pitfalls. For instance: (1) Are all Na-free solutions equally without effect as Na substitutes? and (2) Is the flux that remains in zero Na, a

232

P. F. BAKER AND R. DlPOLO

genuine flux c,omponent that coexists with the Na-dependent flux in solutions containing Na, or is it generated from the Na-dependent flux under Na-free conditions? As mentioned in Section IV,C, Na,, can be replaced by a number of substitutes, both ionic and nonionic, without materially altering the size of the Na,,-dependent component of Ca efflux, suggesting that none of these agents exerts a Na-like action. The Na substitute used most commonly is choline. Obtaining a clear-cut answer to the second point is more difficult; but internal vanadate and external alkalinity both inhibit the uncoupled Ca efflux in a fairly specific manner, and the reduction in Ca efflux is much the same in the absence and in the presence of external sodium, suggesting that in the presence of external sodium the uncoupled Ca efflux does, indeed, coexist with the Na-dependent Ca efflux. In unpoisoned axons and axons dialyzed in the presence of ATP, halfmaximal activation of the Na,-dependent Ca efflux requires 40-60 mM Na,, and the curve relating the extent of activation to Na, usually approximates to a section of a rectangular hyperbola (Baker and Glitsch, 1973; DiPolo, 1974), although close inspection sometimes reveals it to be slightly sigmoidal (Fig. 19) (Blaustein, 1977). This sigmoidicity becomes very apparent in poisoned axons or in axons dialyzed free of ATP, where half-maximal activation may require as much as 300 mM Na, (Fig. 19) (Baker and Glitsch, 1973; Baker and McNaughton, 1976a; DiPolo, 1974; Blaustein, 1977). In order to return the kinetics of activation by Na, to the unpoisoned state, high levels of nucleotide (Kllz= 300-400 p M ) are required, and, of a variety of nucleotides examined, only ATP, 2-deoxyATP, and the hydrolyzable analog AMpCpp are effective (DiPolo, 1976). This suggests that the alteration in kinetics may involve ATP hydrolysis. In the presence of ATP. the apparent affinity for Na, is not obviously affected by varying the intracellular free Ca (Fig. 19). The internal Ca concentration required for half-maximal activation of the Na,-dependent Ca efflux also depends on whether ATP is present. Data from several laboratories (DiPolo, 1973; Brinley et al., 1975; Blaustein, 1977) show rather variable results, probably reflecting the different experimental conditions used. In an extensive study on this subject, Blaustein (1977) found that in axons dialyzed with 4 mM ATP and a low Na, (5 mM), the curve relating Na,-dependent Ca efflux to Cq was fitted best by a rectangular hyperbola with an apparent half-saturation constant of 0.73 pM. In the absence of ATP, the curve shifted to higher Cai concentrations, half-maximal activation now requiring about 8 pM. Somewhat different results have been obtained in axons of L. pealei and D. plei dialyzed with a solution containing 2 mM ATP and a mean Nai of 45 mM. Here the Na,-dependent Ca efflux is poorly activated at low Ca,, but

_-__----a

, / -

/

X

4

0

100

2 00

I

1

300

4 00

I

5 00

[NO], I m M ) FIG. 19. Dependence on Na, of Na,-dependent Ca efflux in an axon of Loligo pealei dialyzed with different concentrations of free Ca. (A) Raw data; (B) normalized data. Open symbols: in the presence of ATP (4 m M ) ;filled symbols: in the nominal absence of ATP. Internal free Ca 2.5 p M (A), 0.5 pLM (O), 0.3 p M (a), and 100 pM ( 0 ) .The dialysis fluids also contained FCCP (10 p M ) and oligomycin (31 pg/ml). Temperature, 15°C. The curves have been drawn on the assumption of an exchange of 3 Na for 1 Ca. (From Bbdustein, 1977.)

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P. F. BAKER AND R. DIPOLO

increases dramatically in the micromolar range to a value of 2000 fmol cm-2 sec-I. Half-maximal activation requires approximately 10 p M Ca. The difference between this value and that obtained by Blaustein might result from the much lower Nai used in Blaustein’s experiments (5 mM as opposed to 45 mM). Internal Na under physiological conditions normally lies between 30 and 50 mM. Raising internal Na can inhibit Na,-dependent Ca efflux, especially in axons depleted of ATP. Thus, in axons in which the ATP has been removed by dialysis, the Nh-dependent Ca efflux is reduced when Nai is elevated. The kinetics suggest that two Na ions may compete with one Ca ion for an identical binding site (Blaustein et al., 1974; Blaustein, 1977). Although Blaustein (1977) did not report changes in the N q concentration giving half-maximal inhibition (30 mM) in the presence or in the absence of ATP, more recent experiments by Requena (1978), which confirm previous observations of DiPolo (1977), show that ATP increases the internal Na concentration required for half-maximal inhibition by almost a factor of 10. This striking observation suggests that ATP can, in some way, relieve the inhibitory effect of internal Na. The conditions under which this takes place are approximately those where Ca,-dependent Na efflux is activated (see Section IV,C,2,c). It is important to note that while Na, is intimately involved as an inhibitory substrate of the carrier-mediated Na,dependent Ca efflux, this ion has no effect on the magnitude of the ATPdependent uncoupled Ca efflux (see Fig. 15) (DiPolo and Beauge, 1979) or on its affinity for ATP (DiPolo and Beauge, unpublished data), thus supporting the existence of two, functionally different, Ca transport mechanisms. The only direct evidence that external Na actually exchanges with internal Ca comes from experiments of Blaustein and Russell (1975), who were forced to use grossly unphysiological conditions in order to obtain measurable fluxes. Working with an internal ionized Ca of 160 p M , an external Na of 200 mM, and in the presence of TTX, they found an exchange of approximately 1 Cq for 3 Na,. A nonelectroneutral exchange is also suggested by the sensitivity of Na,-dependent Ca efflux to membrane potential. It is generally agreed that both electrical and potassium depolarization inhibit Na,-dependent Ca efflux, both in the presence and in the absence of ATP (Blaustein et al., 1974; Mullins and Brinley, 1975; Baker and McNaughton, 1976). The inhibitory effect of external K can be completely mimicked by electrical depolarization, and electrical hyperpolarization strongly activates Na,-dependent Ca efflux (Allen and Baker, 1983) (see Section IV,C,2,c). No really specific inhibitor of Na,-dependent Ca efflux has yet been

235

AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS

found; but the range of substances that do not affect Na,-dependent Ca efflux is almost as interesting as the range that does. Thus, external ouabain and internal orthovanadate, which block the Na-K exchange pump and the uncoupled Ca pump, respectively, are without effect on the Na,dependent Ca efflux, and internal alkalinity, which is without effect on the uncoupled pump, strongly activates the Na,-dependent Ca efflux (Fig. 20). These same conditions also strongly activate Ca,-dependent Na efflux, providing some support for the view that these two fluxes may reflect forward and reverse modes of the same system. External lanthanum inhibits Na,-dependent Ca efflux (van Breemen and de Weer, 1970) as it does Ca,-dependent Na efflux (Baker et al., 1969b) and the uncoupled Ca pump (Baker and McNaughton, 1978). Less complete inhibition is brought about by external Mn2+(Ki 50 mM), cadmium (Ki - 5 mM), and cobaltous ions. The organic Ca antagonist D 600 (0.1 mgiml) causes a very small fall in the Na,-dependent Ca efflux, which is only a little larger than the 1% ethanol in which it is normally dissolved. The Na,-dependent Ca efflux is strongly inhibited by tetracaine (0.5 mg/ml).

-

.

lo(

I

1.0 "nccup*d

60

A

70 13

INTERNAL pH

80

90

74

00 EXTERNALoH

135

I

99

FIG.20. The effects of pH on both ATP-dependent uncoupled (open symbols) and Naodependent (filled symbols) components. (A) Internal pH. Ordinate: steady-state Ca efflux at different pH, values relative to that at pH 7.3. (B) External pH. Ordinate: steady-state Ca efflux at different pH, relative to that at pH 7.5. Note the inhibition of the uncoupled at alkaline pH, and in the presence of SO m M Ca,. The symbols refer to different axons. Mean temperature, 18°C. (From DiPolo and BeaugC, 1982.)

236

P. F. BAKER AND R. DIPOLO

D. Maintenance of the Steady State 1. GENERAL The long-term maintenance of the resting Caft in an axon requires that the passive resting Ca influx together with the extra Ca influx during nervous activity be balanced by Ca efflux. The experimental evidence from both intact and dialyzed axons demonstrates that squid axons are endowed with two well-defined mechanisms for extruding calcium: an ATP-driven Ca pump and a carrier-mediated Na,-dependent Ca efflux. In these circumstances it is important to determine the relative contribution of the two mechanisms to Ca homeostasis in a living nerve cell. Any transport mechanism responsible for maintaining the resting ionized Ca should be capable of operating effectively at Ca;+ similar to those found in a living axon and should be capable of generating a transmembrane Ca gradient close to 10'. For Na,-dependent Ca efflux to be responsible for this Ca distribution, sufficient energy must be stored in the Na electrochemical gradient. The voltage dependence of this component of Ca efflux strongly points to an electrogenic exchange, and from activation kinetics Blaustein (1977) suggested that three or more Na may be required to activate the efflux of one Ca. A similar conclusion had been reached earlier by Baker et al. (1969a), who found that the influx of one Ca is associated with the exit of 3-5 Na ions. Provided a 4 Na : 1 Ca stoichiometry pertains, the carrier-mediated mechanism is indeed able to account for the observed 1Ca:+] : [Ca;'] ratio (Baker, 1972; Blaustein, 1977; Mullins, 1977). However, in a real axon most of the Ca entry under resting conditions is not carrier mediated and the Na,-dependent Ca efflux mechanism will have to balance all the observed resting Ca influx. The large body of evidence presented in this article shows this not to be the case. In fact, an Na,,-dependent Ca efflux of the magnitude needed to balance the physiological Ca influx (40-60 fmol cm-2 sec-.') is never attained at the normal Ca't, but only at a value almost 10 times higher (see Fig. 16). It is therefore highly unlikely that the Na-Ca exchange system with its relatively low affinity for CaZt (apparent Kcai about 10-50 times higher than the resting [Ca;']) is the principal mechanism responsible for maintaining the low [Ca2+lifound in fresh axons. The kinetic properties of the Na-Ca exchange system described here (low affinity, high capacity) are in no way peculiar to squid axons. They have been described in other preparations (Reeves and Sutko, 1978; Caroni and Carafoli, 1980; Schellenberg and Swanson, 19XI), suggesting that this behavior may be a generalized characteristic of the Na-Ca antiporter. The high capacity for Ca transport exhibited by the Na-Ca exchange system may play an important role in removing large amounts of

AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS

237

Ca from the cytosol after transient increases in internal calcium. This could be of particular significance during depolarization-repolarization cycles such as occur at nerve terminals and in muscle, or in any other situation in which the internal ionized Ca near the membrane reaches relatively high values. The other obvious candidate that must be considered for the primary role in regulating Caf+in squid axons is the “uncoupled”’ ATP-driven Ca pump. A primary role for this pump is strongly supported by the following observations.

1. The uncoupled Ca efflux is able to operate effectively at Caf+ concentrations normally found in fresh axons. The apparent affinity for Caf+ is only a factor of two higher than the resting free Cai. The data shown in Fig. 16 demonstrate that in the physiological range of ionized Ca the magnitude of the ATP-dependent Ca efflux is sufficient to balance the “leak” influx of Ca. Even though the maximal transport capacity of the Ca pump is only one-tenth that of the Na-Ca exchanger, the contribution of the two processes to net Ca extrusion should become equal only when Ca’+ rises to 1 p M (see Fig. 16). 2. An uncoupled Ca efflux is present in axons (intact and dialyzed) bathed in seawater, proving that this modality of Ca transport operates under physiological conditions. Similarly, this mechanism has been shown to effect net extrusion of Ca in dialyzed axons in the absence of ionic gradients (Fig. 21) and in intact axons under nearly normal conditions. 3. For the ATP-driven Ca pump to be responsible for the resting Caz+, the ratio of the apparent affinities of the external and internal transport sites for Ca must be greater than or equal to the observed ratio of [Ca?’]: [Ca’’]. An estimate of the affinity of the transport site for Ca when it faces the external medium can be deduced from experiments of DiPolo and Beauge (1982) in which they examined the inhibition of the ‘‘uncoupled’’ Ca efflux by external Ca. At physiological external pH (7.8), the Ca concentration required to achieve half-maximal inhibition is greater than 4 x M . As the apparent affinity constant for internal Ca under similar experimental conditions is close to 2 x lo7 M (DiPolo, 1979; DiPolo and BeaugC, 198O), the ratio of affinities of the external and internal Ca transporting sites must be greater than lo5, which is adequate to maintain the observed Ca gradient. 2. NET MOVEMENTS OF CALCIUM Rather clear evidence for the relative importance of the two components of Ca efflux under physiological conditions can be obtained from an

238

P. F. BAKER AND R. DIPOLO

I NTI IOK-%No-390TRIS-IOMg-0 35 p MCo2+-2ATP EXTl IOK-%NO-390TRIS-IOMg-0 35 pM Co2+

0

50

I00 I50 TIME (min)

200

250

FIG.21. The effect of ATP on unidirectional Ca fluxes in a single dialyzed squid axon. Zero ionic gradients were obtained using the same medium on either side of the membrane (see upper portion of the graph); 1 rnM ATP, 1 mM cyanide, and 10 pg of oligomycin per milliliter were included in the dialysis solution. For Ca influx measurement 45Cawas added to the external fluid, and the perfusate was collected at 5-min intervals. The Ca influx was measured, then the radioactive external medium was removed and the chamber washed carefully. During this period (30 min), the axon was dialyzed to remove the isotope accumulated during the influx measurement. The efflux of Ca was measured by dialyzing the axon with a 45Ca-containingmedium. The [Ca2’li was set in both internal and external solutions at 0.35 p M (CalnIal= 1.4 m M , EGTA,,I,I = 2 m M . The level of the Ca-EGTA “leak” is indicated by the horizontal dashed line. Temperature, 19°C; axon diameter, 400 p m . (Unpublished data of R. DiPolo.)

analysis of recovery from an imposed Ca load. Isolated axons tend to maintain a fairly constant Ca content and, if loaded with Ca by electrical stimulation, immersion in low sodium or microinjection will reestablish their normal resting Ca content during a subsequent recovery period of 1-2 hr in seawater. The rate and extent of recovery is much the same in unpoisoned and poisoned axons (Fig. 22) (Requena and Mullins, 1981). This observation has been thought to indicate that ATP is not essential for the reestablishment of the resting Caf’ . However, this interpretation must be viewed with caution because, although total Ca falls in fully poisoned axons, ionized Ca remains in the micromolar range (Baker et al., 1971; Brinley et al., 1978; Baker and Umbach, 1983; DiPolo et al., 1983b), and so the recovery seen in poisoned axons is quantitatively not the same as that seen in unpoisoned ones. It is precisely what might be expected from the operation of the Na-Ca exchange system alone. Another way to examine this problem is to impose a defined Ca load at a specified ionized Ca and to examine the requirements for extrusion of the load. This can be achieved by injecting a Ca-EGTA buffer; using this

239

AXONAL CALCIUM AND MAGNESIUM HOMEOSTASIS 200 I80

I60

410 Choline, Control

140

120 h

E

-B4

100

80

\

60

Na, Control

40

Na,, + FCCP + IAA 2c

a

I

I

I

I

I

I

20

40

60

80

100

120

Time (min)

FIG.22. The time course for unloading calcium from Ca-loaded axons of D. plei under different experimental conditions. (From Requens and Mullins, 1979.)

approach, Baker and Singh (1981) found, in contrast to Requena and Mullins (1981), that, at an ionized Ca close to physiological, net extrusion of Ca was inhibited by the mitochondria1 uncoupler FCCP and by internal vanadate (Fig. 23). Net extrusion was also impaired in Na-free media, presumably because Ca influx is also stimulated under these conditions. These findings suggest that an ATP-dependent and vanadate-sensitive system is important in the physiological range of ionized calcium. Overall, the most likely interpretation of these experiments is that, in the absence of ATP and at high Ca’+, Na,-dependent Ca efflux can extrude a significant fraction of the imposed load; once Caz+ reaches about 1 p M this process is no longer able to effect net extrusion of Ca, and a steady state is reached. In unpoisoned axons much the same state of affairs pertains when Caf+ is high; but instead of Ca!+ remaining in the

240

[

P. F. BAKER AND R. DIPOLO

A

E

0

t

/t+V=nAd'l*

C I

12

-

0.6

-

2 L

01

a

EE

9: V

0

0

t

I I I 0 60

I I

i

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0

60

FIG.23. Influence of vanadate, FCCP, and Na+-free (choline) seawater on net movements of Ca" in axons of Loligu forbesi measured by atomic absorption spectrometry. Axons immersed in seawater containing ( m M ) :4 0 2 3 Na+ (for choline C): I00 Mg?+;3 C d + , 10 K + , 2.5 HCO;, 61hCI- (pH 7.8). Comparisons wcrc made between paired axons from the same animal; a different symbol is used for each pair. At zero time, axons were injected (arrow) over their full length with a solution containing 24 mM Ca?' and 400 mM EGTA (pH 7.2) and injected a second time with vanadate (4 mM) or FCCP (80 pg ml-I) as indicated. At the time indicated, axoplasm was extruded and a sample from the middle section of the fiber was taken for analysis. All axons were excitable. Temperature, 21°C. (From Baker and Singh. 1981.)

micromolar range it is taken down below 100 nM by operation of the ATPdependent Ca-extrusion system. These data strongly suggest therefore that the two Ca efflux systems work in parallel to pump Ca out of the squid axon. The concerted action of two parallel Ca transport systems does not seem to be a unique property of squid axons. It has been proposed for several other preparations (Caroni and Carafoli, 1980; Kurzinger et d., 1980; Sulakhe and St Louis. 1981; Akerman and Nicholls, 1981).

3 . MODULATION OF Ca HOMEOSTASIS The various processes that have been described in this article all serve to maintain a fairly constant level of ionized calcium in axoplasm. They are, however, themselves subject to modulation, and such modulation may lead to either transient or maintained changes in ionized Ca that may be of profound physiological significance. Table I1 summarizes some of the main effects of variables, such as ATP levels, Em,pH,, and Na,. Others, such as cyclic nucleotides, methylation, and products of PI metabolism, have yet to be examined in the squid axon. The effect of each

EFFECTS OF

SOME

PHYSIOLOGICALLY

TABLE 11 VARIABLES ON AXONAL CALCIUM HOMEOSTASIS

IMPORTANT

Calcium inflow Calcium binding

ATP depletion Depolarization Hyperpolarization Decrease in pH, Increase in pH, Increase in Na,

Calcium outflow

ATP dependent

ATP independent

Voltage sensitive channels

.1

-

?

-

.1 ?

.1

-

t 1

? ?

? ?

-

-

-

Na,-Ca, exchange

1 T .1 .1 t

t

Ca pump

Na,-Ca, exchange

Ionized Ca

.1

Altered

t

-

.1

-

1 t

T

1

N o change or J, Variable Variable

1

t

t

242

P. F. BAKER AND R. DIPOLO

perturbation is complex, and the magnitude and duration of any change in ionized Ca may depend on the past history of the axon and on the relative importance of intracellular binding systems and membrane transport in the region of axoplasm under consideration. Modulation of the basic mechanisms is probably an important means of “fine-tuning” cellular Ca homeostasis, and, although many such modulatory mechanisms may not be represented in the squid axon, it seems likely that the squid axon will continue to serve as an excellent preparation in which to unravel at least some of the complex interactions of cellular calcium homeostasis. ACKNOWLEDGMENTS This work was supported by grants from the Medical Research Council of Great Britain (to P. F. B.) and the Consejo Venezolano de lnvestigaciones Cientificas y Tecnologicas (CONICIT), PNUD-UNESCO, RLA, and NSF-BNS-USA (to R. D.). REFERENCES Akerman, K. E. O., and Nicholls, D. S. (1981). Ca2+transport by intact synaptosomes the voltage dependent Ca2+ channel and a re-evaluation of the role of sodium/calcium exchange. Eur. J. Biochem. 117, 491-497. Alema, S.,Calissano, P., Rusca, G., and Guiditta, A. (1973). Identification of a calciumbinding brain specific protein in the axoplasm of squid giant axons. J . Neurochern. 20, 68 1-689. Allen, T. J. A., and Baker, P. F. (1983). Comparison of the effects of potassium and electrical depolarizations in Na-Ca exchange in squid axons. J . Physiol. (London) 345,80P. Almeida, S., and de Meis, L. (1977). pH-induced changes in the reactions controlled by the low- and high-affinity Ca2+binding sites in sarcoplasmic reticulum. Biochemistry 16, 329-334. Baker, P.F. (1970). Sodium-calcium exchange across the nerve cell membrane. In “Calcium and Cellular Function” (A. W. Cuthbert, ed.). Macmillan, New York. Baker, P. F. (1972). Transport and metabolism of calcium ions in nerve. Prog. Biophys. Mol. Biol. 24, 177-223. Baker, P. F. (1978). The regulation of intracellular calcium in giant axons of Loligo and Myxicola. Ann. N. Y. Acad. Sci. 307, 250-268. Baker, P. F., and Crawford, A. C. (1972). Mobility and transport of magnesium in squid giant axons. J . Physiol. (London) 227, 855-874. Baker, P. F., and Glitsch, H. G. (1973). Does metabolic energy participate directly in the Na-dependent extrusion of Ca ions from squid axons? J . Physiol. (London) 233, 44-4hP. Baker, P. F., and Glitsch, H. G. (1975). Voltage-dependent changes in the permeability of nerve membranes to calcium and other divalent cations. Philos. Trans. R . Soc. London Ser. B 270, 389-409. Baker, P. F., and Honerjiiger, P. (1978). Influence of carbon dioxide on level of ionized calcium in squid axons. Narirre (London) 273, 160-161. Baker, P. F., and McNaughton, P. A. (1976a). Kinetics and energetics of calcium efflux from intact squid axons. J . Physiol. (London) 259, 103-144.

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Baker, P. F., and McNaughton, P. A. (1976b). The effect of membrane potential on the calcium transport systems in squid axons. J . Physiol. (London) 260, 24P. Baker, P. F., and McNaughton, P. A. (1977). Selective inhibition of the Ca-dependent Na efflux from intact squid axons by a fall in intracellular pH. J . Physiol. (London) 269, 78-79P. Baker, P. F., and McNaughton, P. A. (1978). The influence of extracellular calcium binding on the calcium efflux from squid axons. J . Physiol. (London) 276, 127-150. Baker, P. F., and Schlaepfer, W. W. (1975). Calcium uptake by axoplasm extruded from giant axons of Loligo. J . Physiol. (London) 249, 37P. Baker, P. F., and Schlaepfer, W. W. (1978). Uptake and binding of calcium by axoplasm isolated from giant axons of Loligo and Myxicola. J . Physiol. (London) 276, 103-125. Baker, P. F., and Singh, R. (1981). Influence of vanadate on calcium fluxes and net movement of calcium in intact squid axons. Biochirn. Biophys. Acta 646, 450-456. Baker, P. F., and Singh, R. (1982). Metabolism and transport of strontium in giant axons of Loligo. J . Physiol. (London) 330, 373-392. Baker, P. F., and Umbach, J . (1983). Calcium electrode determination of ionized Ca and Cabuffering capacity of squid axoplasm. J . Physiol. (London)341, 61 P. Baker, P. F., Hodgkin, A. L., and Shaw, T. I. (1961). Replacement of the protoplasm of a giant nerve fibre with artificial solutions. Nature (London) 190, 885-887. Baker, P. F., Hodgkin, A. L., and Shaw, T. I. (1962). Replacement of the axoplasm of giant nerve fibres with artificial solutions. J . Physiol. (London) 164, 330-354. Baker, P. F., Blaustein, M. P., Manil, J . , and Steinhardt, R. A. (1967a). A ouabain-insensitive, calcium-sensitive sodium efflux from giant axons of Loligo. J . Physiol. (London) 191, IOOP. Baker, P. F., Blaustein, M. P., Hodgkin, A. L., and Steinhardt, R. A. (1967b). The effect of sodium concentration on calcium movements in giant axons of Loligo forhesi. J . Physiol. (London) 192, 43P. Baker, P. F., Blaustein, M. P., Hodgkin. A. L . . and Steinhardt. R . A. (1969a). The influence of calcium on sodium efflux in squid axons. J . Physiol. (London) 200, 431-458. Baker, P. F., Blaustein, M. P . , Keynes, R. D., Manil, J . , Shaw. T. I., and Steinhardt, R. A . (1969b). The ouabain-sensitive fluxes of sodium and potassium in squid giant axons. 1. Physiol. (London) 200, 459-496. Baker, P. F., Hodgkin, A. L., and Ridgway, E . B. (1971). Depolarization and Ca entry in squid giant axons. J . Physiol. (London) 218, 709-755. Baker, P. F., Meves, H., and Ridgway, E. B. (1973a). Effects of manganese and other agents on the calcium uptake that follows depolarization of squid axons. J . Physiol. (London) 231, 511-526. Baker, P. F., Meves, H., and Ridgway, E. B. (1973b). Calcium entry in response to maintained depolarization of squid axons. J . Physiol. (London) 231, 527-548. Baker, P. F., Knight, D. E., and Pattni, R. D. D. (1977). Porous cellulose acetate tubing provides a suitable support for isolated protoplasm during studies under controlled conditions. J . Physiol. (London) 266, 6P. Barrabin, H., Garrahan, P. F., and Rega, A. F. (1980). Vanadate inhibition of the CaZ'ATPase from human red cell membranes. Biochim. Biophys. Acra 600, 796-804. Beauge, L. A., and DiPolo, R. (1981). An ATP-dependent sodium-sodium exchange in strophanthidin poisoned dialyzed squid giant axons. J . Physiol. (London) 315, 447460. Beauge, L. A., and Glynn, I. M. (1978). Commercial ATP containing traces of vanadate alters the response of (Na+ + K + ) ATPase to external potassium. Nature (London) 272, 551-552.

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Bers, D. M. (1982). A simple method for the accurate determination of free (Ca) in CaEGTA solutions. Am. J . Physiol. 242, C404-C408. Blaustein, M. P. (1974). The interrelationship between sodium and calcium fluxes across cell membranes. Reu. Physiol. Biochem. Exp. Phormucol. 70, 33-82. Blaustein, M. P. (1977). Effects of internal and external cations and of ATP on sodiumcalcium and calcium-calcium exchange in squid axons. Biophys. J . 20, 79-1 10. Blaustein, M. P., and Hodgkin, A. L. (1969). The effect of cyanide on the efflux of calcium from squid axons. J . Physiol. (London) 200, 497-527. Blaustein, M. P., and Nelson, M. T. (1982). Sodium calcium exchange: Its role in the regulation of cell calcium. I n “Membrane Transport of Calcium” (E. Carafoli, ed.), pp. 217-236. Academic Press, New York. Blaustein, M. P., and Russell, J. M. (1975). Sodium-calcium exchange and calcium-calcium exchange in internally dialyzed squid giant axons. J . Menrbr. Biol. 22, 285-312. Blaustein, M. P., Russell, J . M., and De Weer, P. (1974). Calcium efflux from internally dialyzed squid axons: The influence of external and internal cations. J . Supramol. Struct. 2, 558-581. Bond, G. H., and Hugdkins, P. M. (1980). Inhibition of red cell Ca”-ATPase by vanadate. Biochim. Biophys. Acfa 600, 781-790. Brinley, F. J . (1978). Calcium buffering in squid axons. Annu. Reu. Biophys. Bioeng. 7, 363392. Brinley, F. J . , and Mullins, L. J. (1967). Sodium extrusion by internally dialysed squid axons. J . G e n . Physiol. 50, 2303-2331. Brinley, F. J . , and Scarpa, A. (1975). Ionized magnesium concentration in axoplasm of dialysed squid axons. FEES L e t t . 50, 82-85. Brinley, F. J . , Spangler, S. G . , and Mullins, L. J. (1975). Calcium and EGTA fluxes in dialysed squid axons. J . Gen. Physiol. 66, 223-250. Brinley, F. J . , Tiffert, T., Scarpa, A., and Mullins, L. J. (1977). Intracellular calcium buffering capacity in isolated squid axons. J . C e n . Physiol. 70, 355-384. Brinley, F. J . , Tiffert, T., and Scarpa, A. (1978). Mitochondria and other calcium buffers of squid axon studied in sifu. J . Gen. Physiol. 72, 101-127. Caldwell, P. C. (1960). The phosphorus metabolism of squid axons and its relation to the active transport of sodium. J . Physiol. (London) 152, 561-590. Caldwell-Violich. M . . and Requena, I . (1979). Magnesium content and net fluxes in squid giant axons. J . Gen. Physiol. 74, 739-752. Carafoli, E . , and Crompton, M. (1978). The regulation of intracellular calcium. Curr. Top. Membr. Transp. 10, 151-216. Caroni, P., and Carafoli, E. (1980). An ATP-dependent Ca2+pumping system in dog heart sarcolemma. Nuture (London) 283, 765-767. Condre.;cu. M . , Osses. I > , , and DiPolo. R. (1984). Partial purification and characterization of the (Ca” + Mg?+)-ATPasefrom squid optic nerve membrane. Biochirn. B i o p h y s . Actu 769, 26 I.

De Weer, P. (1976). Axoplasmic free magnesium levels and magnesium extrusion from squid giant axons. J . Gen. Physiol. 68, 159-178. De Weer, P. (1980). Effects of intracellular adenosine-5’-diphosphateand orthophosphate on the sensitivity of sodium efflux from squid axons to external sodium and potassium. J . Gen. Physiol. 56, 583-620. De Weer, P., and Lowe, A. G. (1973). Myokinase equilibrium. An enzymatic method for the determination of stability constant of magnesium complexes with adenosine triphosphate, adenosine diphosphate, and adenosine monophosphate in media of high ionic strength. J . Biol. C h e m . 248, 2829-2835.

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DiPolo, R. (1973). Calcium efflux from internally dialyzed squid axons. 1. Gen. Physiol. 62, 575-589. DiPolo, R. (1974). The effect of ATP on Ca efflux in dialysed squid giant axons. J . Gen. Physiol. 64, 503-517. DiPolo, R. (1977). Characterization of the ATP-dependent calcium efflux in dialysed squid axons. J. Gen. Physiol. 69, 795-814. DiPolo, R. (1978). Ca pump driven by ATP in squid axons. Nature (London) 274, 390-392. DiPolo, R. (1979). Calcium influx in internally dialysed squid giant axons. J . Gen. Physiol. 73,YI-113. DiPolo, R . , and Beauge, L. A. (1979). Physiological role of ATP-driven calcium pump in squid axons. Nature (London) 278, 271-273. DiPolo, R., and Beauge, L . A. (1980). Mechanisms of Ca transport in the giant axon of the squid and their physiological role. Cell Calcium 1, 147-169. DiPolo, R., and Beauge, L. A. (1981). The effects of vanadate on calcium transport in dialyzed squid axons. Sidedness of vanadate-cation interactions. Biochim. Biophys. Act0 645, 229-236. DiPolo, R., and Beauge, L. A. (1982). The effect of pH on Ca2+extrusion mechanisms in dialyzed squid axons. Biochim. Biophys. Acta 688, 237-245. DiPolo, R . , and Beauge, L. (1983). The calcium pump and sodium-calcium exchange in squid axons. Annu. Rev.Physiol. 45, 313-324. DiPolo, R., Requena, J . , Brinley, F. J., Mullins, L. J., Scarpa, A., and Tiffert, T. (1976). Ionized calcium concentration in squid axons. J . Gen. Physiol. 67, 433-467. DiPolo, R., Rojas, H.. and BeaugC, L. A. (1979). In squid vanadate inhibits uncoupled calcium efflux but does not inhibit the NdCa exchange. Nature (London) 281, 228229. DiPolo, R., Rojas, H., and Beauge, L. A. (1982). Ca entry at rest and during prolonged depolarization in dialysed squid axons. Cell Culcium 3, 19-41. DiPolo, R., Caputo, C . , and Bezanilla, F. (1983a). A voltage-dependent calcium channel in the squid axon. DiPolo, R., Rojas, H., Vergara, J., Lopez, R., and Caputo, C. (1983b). Measurement of intracellular ionized calcium in squid axons using calcium-selective electrodes. Biochim. Biophys. Acta 728, 311-318. Duggan, P. F. (1977). Calcium uptake and associated adenosine triphosphate activity in fragmented sarcoplasmic reticulum. J . Biol. C h e m . 252, 1620-1627. Frankenhaeuser, B., and Hodgkin, A. L. (1956). The after-effects of impulses in the giant nerve fibres of Loligo J . Physiol. (London) 131, 341-376. Gill, 1. L . , Grollman, E. F., and Kohn, L. D. (1981). Calcium transport mechanisms in membrane vesicles from guinea pig brain synaptosomes. J . B i d . Chem. 256. 184- 192. Head, J . F., and Kaminer. B . (1980). Calmodulin from the axoplasm of the squid. Biol. Bu11. Woods H o l e 159, 485. Henkart, M., Reese, T. S . , and Brinley, F. J. (1978). Endoplasmic reticulum sequesters calcium in the squid giant axon. Science 202, 1300-1303. Hodgkin, A. L., and Keynes, R . D. (1957). Movements of labelled calcium in squid axons. J . Physiol. (London) 128, 28-60. Inoue, I. (1980). Separation of the action potential into a Na-channel spike by tetrodotoxin and by tetraethylammonium ion in squid giant axons internally perfused with dilute Na-salt solutions. J . Gen. Physiol. 76, 337-354. Katz, B., and Miledi, R. (1969). Tetrodotoxin-resistant electric activity in presynaptic terminals. J . Physiol. (London) 203, 459-487.

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Keynes, R. D., Malachowski, 0.C., and van Helden, D. F. (1979). The effect of calcium on the sodium gating current in the squid giant axon. J . Physiol. (London) 295, 54P. Kurzinger, K., Stadtkus, C., and Humprecht, B. (1980). Uptake and energy-dependent extrusion of calcium in neural cells in culture. Eur. J . Biochem. 103, 597-61 I . LlinBs, R . , Steinberg, I. Z., and Walton, I . C . (1981). Presynaptic calcium currents in squid giant synapse. Biophys. J . 33, 289-322. Luxoro, M., and Yanez, E. (1968). Permeability of the giant axon of Dosidicus gigas to calcium ions. J . Gen. Physiol. 51, 115s. Meech, R.‘W., and Thomas, R. C. (1977). The effect of calcium injection on the intracellular sodium and pH of snail neurones. J . Physiol. (London) 265, 867-879. Meves, H . , and Vogel, W. (1973). Calcium inward currents in internally perfused giant axons. J . Physiol. (London) 235, 225-265. Mullins, L. J. (1977). A mechanism of Na/Ca transport. J . Gen. Physiol. 70, 681-696. Mullins, L. J . (1981). Calcium entry upon depolarization of nerve. J . Physiol. (Paris) 77, 1139-1 144. Mullins, L. J., and Brinley, F. J. (1967). Some faftors influencing sodium extrusion by internally dialysed squid axons. J . Gen. Physio/. 50, 2333-235s. Mullins, L. J . , and Brinley, F. J . (1975). The sensitivity of calcium efflux from squid axons to changes in membrane potential. J . Cen. Physiol. 50, 2333-235s. Mullins, I,. J . . and Brinley, F. J . (1978). Magnesium influx in dialysed squid axons. J . Membr. Biol. 43, 243-250. Mullins, L. J., and Requena, J . (1979). Calcium measurement in the periphery of an axon. J . Gen. Physiol. 74, 393-413. Mullins, L. J . , and Requena, J. (1981). The ‘‘late’’ Ca channel in squid axons. J . Gen. P h y s i d . 78, 683-700. Mullins, L. J . , Brinley, F. J., Spangler, S. G . , and Abercrombie, R. F. (1977). Magnesium efflux in dialysed squid axons. J . Gen. Physiol. 69, 389-400. Nicholls, D . , and Akerman, K. (1982). Mitochondria1 calcium transport. Biochim. Biophys. AC~U 683, 57-88. Niggli, V . , Sigel, E., and Carafoli, E. (1982). The purified Ca2+pump of human erythrocyte membranes catalyzes an electroneutral Ca2+ - H + exchange in reconstituted liposoma1 systems. J . Biol. Chem. 257, 2350-2356. Oikawa, T., Spyropoulos, C. S., Tasaki, I . , and Teorell, T. (1961). Methods for perfusing the giant axon of Loligo pealei. Acra Physiol. Scand. 52, 195-196. Reeves, J . P., and Sutko, J. L. (1979). Sodium-calcium ion exchange in cardiac membranes vesicles. Proc. Natl. Acad. Sci. U.S.A. 76, 590-594. Rega, A . F., Richards, D. E., and Garrahan, P. J . (1973). Calcium-ion dependent p nitrophenyl phosphatase activity and calcium ion dependent adenosine triphosphatase. Activity from human erythrocyte membranes. Biochem. J . 136, 185-194. Requena, J. (1978). Ca efflux from squid axons under constant sodium electrochemical gradient. 1. Gen. Physiol. 72,443-470. Requena, J . , and Mullins, L. J . (1979). Calcium movement in nerve fibres. Q.Reu. Biophys. 12, 371-460. Rojas, E., and Hidalgo, C. (1968). Effect of temperature and metabolic inhibitors on 45Ca outflow from squid giant axons. Biochim. Biophys. Acta 163, 550-556. Rojas, E., and Taylor, R. E. (1975). Simultaneous measurement of magnesium, calcium and sodium influxes in perfused squid giant axons under membrane potential control J . Physiol. (London) 252, 1-27. Rubinson, K. A., and Baker, P. F. (1979). The flow properties of axoplasm in a defined

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chemical environment: Influence of anions and calcium. Proc. R . Soc. London Ser. B 205, 323-345. Scarpa, A. (1974). Indicators of free magnesium in biological systems. Biochemistry 13, 2789-2794. Schatzmann, H . J. (1981). The plasma membrane Ca pump of erythrocyte and other animal cells. In “Membrane Transport of Calcium” (E. Carafoli, ed.), pp. 41-108. Academic Press, New York. Schellenberg, G . , and Swanson, P. (1981). Sodium-dependent and calcium-dependent calcium transport by rat brain microsomes. Biochim.Biophys. Acta 648, 13-27. Singh, R . (1980). Influence of vanadate on calcium fluxes in squid axons. J . Physio/. (London) 305, 32P. Sulakhe, P. V., and St. Louis, P. J. (1980). Passive and active calcium fluxes across plasma membranes. Prog. Biophys. Mol. Biol. 35, 135-195. Tasaki, I., Watanabe, A., and Singer, I. (1966). Excitability of squid giant axons in the absence of univalent cations in the external medium. Proc. Nat. Acad. Sci. U . S . A .56, 1 1 16-1 122. Tasaki, I., Watanabe, A . , and Lerman, L. (1967). Role of divalent cations in excitation of squid giant axons. A m . J . Physiol. 213, 1465-1474. Van Breemen, C., and De Weer, P. (1970). Lanthanum inhibition of T a efflux from squid giant axon. Nuiurc (London) 226, 260-261. Watanabe, A., Tasaki, I., Singer, I., and Lerman, L. (1967). Effects of tetrodotoxin on excitability of squid giant axons in sodium-free media. Science 155, 95-97.

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CUXRENT TOPICS IN MEMBRANES AND TRANSPORT, VOLUME 22

Regulation of Axonal pH WALTER F . BORON Department of Phy5iolog.y Yule University School of Medicine New Huuen. Connecticut

I. Introduction.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Methodology . . . . . . . . . . . . . . . 111. Effect of Weak Bases and Aci , ................................... A . NH3 and NH:

E. Possible Mechanisms of Transport. ........................... References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

I.

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INTRODUCTION

The intracellular pH (pH,) of squid giant axons is of interest for two reasons. In the first place, the squid axon serves as a valuable model of pH, regulation in other systems. Second, because numerous physiological processes are sensitive to changes in pH,, the acid-base physiology of squid axons is of particular interest to those working on this preparation. The intracellular pH of squid axons was first studied by Caldwell (1954, 1958). and later by Spyropoulos (1960), Bicher and Ohki (1972), and Boron and De Weer (l976a). These workers, all of whom used pH-sensitive microelectrodes to measure pH,, established that the normal pH, of squid axons (i.e., -7.3 at about 20°C) is far too high for H+ and/or HCO, to be in electrochemical equilibrium across the axon’s cell membrane. These 249 Copyright 0 1984 by Acddemlc Presa, Inc All righta of reproduction in any form rererved ISBN 0-12-153322-0

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data corroborated similar conclusions being drawn at about the same time from experiments on other excitable cells (see Caldwell, 1956; Waddell and Bates, 1968; Roos and Boron, 1981). Specifically, the squid axon’s membrane potential (V,) of --60 mV and external pH (pH,) of -7.8 would predict an equilibrium pHi of -6.8, a half pH unit lower than the actual pHi . Clearly, if pHi is to be kept at the relatively alkaline value of 7.3, an active-transport mechanism must continuously extrude acid from the axon as rapidly as the acid accumulates in the cell as the result of cellular metabolism and the passive fluxes of ions. In this article I will first review some of the factors that modulate pHi and then summarize our understanding of the active-transport mechanism responsible for pHi regulation in squid axons.

II. METHODOLOGY

The experiments described in this article were performed in Woods Hole, Massachusetts, on giant axons of the squid Lofigo pealri. The axons were dissected and cleaned according to standard practices and cannulated at both ends in a dialysis chamber (see Fig. 1). The experiments were of four types. In the first, the only additional manipulation was the insertion of two microelectrodes through opposite end-cannulas (Fig. 1, top, but without dialysis tube). The first microelectrode, an exposed-tip pH-sensitive electrode (Hinke, 1967), is illustrated schematically in Fig. 2. This electrode is capable of exceptionally stable and reproducible recordings, although its large size precludes its use in cells smaller than the squid axon. The second microelectrode was an open-tipped reference electrode (tip size, <5 pm), filled with KCI or potassium glutamate (1 M). In a second group of experiments, a dialysis tube as well as the microelectrodes was inserted into the axon (Fig. 1, top). The internal dialysis technique, developed by Brinley and Mullins (1967), permits the control of intraaxonal concentrations of low-molecular-weight ( M , < 1000) solutes. In a third and fourth group of experiments, we measured isotopic fluxes of Nat and CIF. These axons were internally dialyzed, but not impaled with microelectrodes. Effluxes (Fig. 1, middle) were measured by presenting an isotope to the axoplasm via the dialysis fluid (DF) and assaying the radioactivity of the artificial seawater (ASW), which continuously superfused the axon. Influxes were measured by placing an isotope in the ASW and measuring the radioactivity of the collected DF (Fig. I , bottom). The standard ASW was composed of (mM): 425 Na+, 12 K ’ , 10 Ca2+,50 Mg, 542 C1-, and 30 mM buffer 4-12-hydroxyethyl]-I-piperazinepropanesulfonic acid (EPPS) titrated to pH 8.0 (-970

YDROLYZED

TUBING

DIALYSIS TUBING

--L-y---l

END W E L L

DIALYZED REGION

GUARD

&---

-

END WELL

GUARD

S U C T l O N y

------

FIG. I . Internal dialysis apparatus. T o p : pH, experiments. The squid giant axon (400600 bm in diameter) is tied onto a glass cannula at both ends. A length of dialysis tubing (- 140 p n in diameter) is introduced through one cannula, traverses the length of the axon, and exits through the opposite end cannula. In its hydrolyzed region (dashed lines), the tubing is permeable to low-molecular-weight solutes (i.e., M,< 1000). A pH-sensitive microelectrode (see Fig. 2) is inserted through one cannula, and an open-tipped reference (V,) electrode, through the other. Their tips are fewer than one axon diameter apart. The axon's central region, which is continuously circumfused with artificial seawater (ASW), is isolated from the cut ends by Vaseline petroleum jelly seals. In some pH, experiments, no dialysis tubing was present. Middle: Isotopic efflux experiments. **Na or "CI is presented to the axon's interior via the dialysis fluid (DF) and is collected in the ASW. Isotope exiting into the guard region is discarded; it represents a flux from a part of the axon whose composition is not well controlled by dialysis. Bottom: Isotopic influx experiments. The isotope is presented via the ASW and collected in the DF. The guard regions are completely obliterated by Vaseline petroleum jelly seals. This prevents the entry of isotope through membrane surrounding axoplasm not optimally controlled by dialysis. To maximize the collection of isotope that may diffuse laterally toward the end wells, the hydrolyzed region of the dialysis tube is somewhat wider than the central region over which the isotope is presented. (From Boron and Russell, 1983; reproduced by permission of the Rockefeller University Press.)

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PH -sensitive

\

Frc. 2. Protruding tip pH microelectrode. Designed by Hinke (1967). this electrode consists of an insulating lead glass shank (Corning 0120), through which protrudes a cone of pH-sensitive glass (Corning 0150). The two pieces of glass are welded together on a microforge. The outer diameter of the insulating glass is s 125 pm for the terminal 3-4 cni. The pH glass has a diameter of 40-60 pm at the glass seal, and an exposed length of 100-300 pm.

mosmlkg). The standard DF contained (mM): 350 K + , 50 Na+, 7 Mg'+, 150 CIF, 264 glutamate, 210 taurine, 10 N-2-hydroxyethylpiperazine-N'-2ethanesulfonic acid (HEPES), I .0 ethylene glycol bis(p-aminoethyl ether)-N,N'-tetraacetic acid (EGTA), 0.5 phenol red, and 4.0 ATP (950960 mOsm/kg). For further details, consult Boron and De Weer (197ha) and Boron and Russell (1983). 111.

EFFECT OF WEAK BASES AND ACIDS ON pHi

A. NH3 and NH:

When an axon is exposed to a solution containing NH, and NH: (NH; = NH3 + H + ; p K ; 9 . 9 , pHi at first rises rapidly (see Fig. 3A, segment ah) as the highly permeant NH3 enters the cell and then com-

253

REGULATION OF AXONAL pH

6 9L

IS mm

C

FIG. 3. Effect of NHJNH: on pH,. (A) Short-term exposure. The axon is exposed to artificial seawater (ASW) (pH 7.70 throughout) in which 10 mM NaCl has been replaced with NH4CI. The influx of NH3produces an alkalinization as the weak base combines with H' to form NH; (segment ab). When the external NH3/NH; is removed, pH, falls to a value slightly below its initial one (compare c and a). (B) Long-term exposure. When the treatment with NH3/NH; is more prolonged, the initial alkalinization ( c d ) is clearly followed by a slower acidification during a period termed the plateau phase ( d e ) .The subsequent washout of NHJNH; leads to a pH, undershoot of exaggerated size (comparef'and c ) . (C) Proposed mechanism for segment di,plateau-phase acidification. The passive entry of NH: is followed by the intracellular dissociation of a small amount of this weak acid to H + and NH3. The resultant increase of [NH,], leads to a net efflux of NH1. Thus, the NH3/NH: couple shuttles H + into the axon during the exposure to NH,/NH;. (From Boron and De Weer, 1976a; reproduced by permission of the Rockefeller University Press.)

bines with H' to form its conjugate weak acid, NH;. This entry and protonation of NH3 continues until [NH,], reaches [NH,],, at which point the increase in pH, is expected to halt. When the external NH3 and NH; are removed, these processes reverse, and pHi falls ( h c ) ;note, however, that pHi undershoots its initial value by a small amount (compare a and c ) , The magnitude of the NH,-induced intracellular alkalinization is determined by [NH,],, the initial pH, (the law of mass action predicts that the higher the starting pH,, the smaller the subsequent pHi increase), and the

J69

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WALTER F. BORON

intracellular buffering power (p). Indeed, the change in pHi produced by application or withdrawal of extracellular NH3 is used to calculate /3 (see Boron, 1977); in squid axons (pHi = 7.3-7.8) p = 9 mM (Boron and De Weer, 1976a). Experiments in which the axon is exposed to NH3 and NH: for more prolonged periods indicate that the cell membrane is permeable to NH: as well as to NH,. In the example of Fig. 3B, exposing the axon to NH3 and NH: causes an initial rise in pHi ( c d ) ,followed by a slower decline during a period termed the "plateau phase" (de).Throughout this plateau phase, the NH,f electrochemical gradient favors the influx of this ion.' As NH: passively enters the axon, most2 of it remains NHd and has no immediate effect on pHi. A small fraction, however, dissociates into H+ and NH, and produces a plateau-phase acidification. Because [NH3], rises slightly above [NH,],, NH3 passively leaves the cell. Thus, the oppositely directed fluxes of NH3 and NH: acidify the cell by shuttling protons inward (Fig. 3C). When the NHJNH: is removed from the external solution (point e ) , virtually all intracellular NH: exits the cell as the highly permeant NH3, after having first given up its H+ and lowered pHi (segment ef). This dissociating intracellular NHd includes not only the NH4f derived from the NH3that entered during segment cd, but also the NH: that directly entered the cell during segment de. Note that the magnitude of the pHi undershoot (compare points c andf) is greater than that of the plateau-phase acidification (compare d and e ) , even though both are caused by the influx of NH:. This apparent discrepancy arises because only a small fraction of the entering NH: dissociates during the plateau phase; the rest dissociates only after removal of external NH3/NH:. We have found that the magnitude of the pHi undershoot upon removal of the external NH,/NH,f is directly related to the amount of NH; that had previously entered. Thus, the small pHi undershoot in Fig. 3A reflects the relatively short exposure to NHJNH:. The importance of the NH: electrochemical gradient is evidenced by an experiment in which this gradient is inverted by depolarizing the axon with high [K+],. Not only does pHi rise during the plateau phase (owing to NH: efflux), but removal of external NH:/NH3 produces a pHi shortfall rather than an undershoot (W. F. Boron and P. De Weer, unpublished observation). I At point d in Fig. 3B, [ N H J ] is ~ approximately equal to [NH,],. If the dissociation constant ( K : ) of the reaction NH; = NH3 + H+is the same inside and outside the cell, then it follows that [H'li INH:li = tG lNH31i = K: [NH31, = [H+I,, [NH:],. In other words, the At point d, EHis -0 m V , equilibrium potential for NH: is the same as that for H + (EH). whereas the membrane potential is ---58 mV. Thus, the electrochemical gradients strongly favor the influx of both H +and NH; . * The fraction of the entering NH: which remains NH: is determined by the relationship between the pK: (-9.5) of the reaction NH: = NH3 + H', and the current pH,. At a pHi of 7.7, the ratio of entering NH; which remains NH: to that which dissociates is about 50 : 1.

REGULATION OF AXONAL pH

255

NH3 is by no means the only weak base that produces sizable increases of intraaxonal pH. Methylamine, triethanolamine, and the K+-channel blocker 4-aminopyridine all cause comparable intracellular alkalinizations (W. F. Boron and P. De Weer, unpublished observation).

B. COOand HCO;

When an axon is exposed to a C02-containing solution, pHi falls rapidly (see Fig. 4A, segment ab) as C02 enters the cell and combines with H 2 0 to form H2CO3, which then dissociates into H+ and HCO;. CO2

+ H20 = H2CO3 = H' + HCO;

The entry of C02 and the resultant intraaxonal acidification continue until [C0,li = [CO,],. The extent of the intracellular acidification depends on [CO,],, p, and the initial pHi (because C02 + H 2 0 = H+ + HCO,, the higher [H+],the less the tendency for the reaction to proceed to the right). When the C02/HCO; is removed from the external solution, these processes reverse and pHi returns toward its initial level ( c d ) .However, if, during the C 0 2 exposure, pH, is sufficiently high (e.g., 7 . 9 , pHi returns to a value slightly higher than the initial one (compare points a and d ) . This overshoot is caused by the axon's pHi-regulating system (see below). In the experiment of Fig. 4A, the fall of pHi brought about by the application of C02 appears to be sustained (segment bc) only because the C02 exposure is relatively brief. If the exposure is more prolonged (see Fig. 4B), the initial intracellular acidification (ab) is followed by a slow rise in pHi (bc). This alkalinization cannot be accounted for by passive transport processes, because the predicted passive fluxes of H+ (an influx) or HCO, (an efflux) would each tend to further acidify the axon. The slow segment -bc pH, recovery must therefore be caused by an activetransport mechanism that extrudes acid from the cell (Fig. 4C). When the external C02/HC0; is removed, pHi overshoots its initial value (compare a and d ) by an exaggerated amount. The magnitude of this overshoot is a direct reflection of the amount of acid extruded from the axon during the preceding CO, exposure. Thus the overshoot after the 25-min C02 exposure of Fig. 4A is only about 20% as large as the overshoot following the 125-min C 0 2 exposure of Fig. 4B. Although snail neurons (Thomas, 1976b) and barnacle muscle fibers (Boron et al., 1979) can return pHi to normal (provided pH, and [HCO;], are sufficiently high) after even very large intracellular acid loads (i.e., sufficient to lower pHi by 0.5-1.0), squid axons cannot return pHi to normal during an in vitro experiment unless the acid load is quite small

WALTER F. BORON

5 % co2

B 7.4 75[

4

'

7-

I

>

-

/

50

FIG.4. Effect of Cot on pH,. (A) Brief exposure. An intact axon is exposed to ASW containing 5% c o 2 / 5 0mM HCO;; extracellular pH (pH,) is 7.5 throughout. The influx of C 0 2causes pH, to fall (segment ah) and apparently to level off ( h c ) . The subsequent return to a HCO1-free ASW (buffered with 5 mM Tris-HEPES) produces a slight overshoot of the initial pH, (compare d and a ) . (B) Prolonged exposure. In a more lengthy experiment, it is clear that the initial C02-induced fall in pHi (ah)is followed by a slower recovery ( h c ) .When the COz is now washed out, pH, overshoots its initial value by an exaggerated amount (compare d and ( I ) . (C) Proposed mechanism of segment -6c alkalinization. During the plateau phase of the above C 0 2 exposure, the electrochemical gradients for H' and HCO; favor a net influx and efflux, respectively, of these two ions. Thus, the observed increase in pH, must be due to an active transport process. Although depicted here as a proton extruder, this process could involve the uptake of alkali and/or the efflux of acid. The transport process has been generically described as "acid extrusion." (From Boron and De Weer, 1976a; reproduced by permission of the Rockefeller University Press.)

REGULATION OF AXONAL pH

257

(i.e., lowering pHi by -0. I ) . At a pH, of 7.5, a [HCO.;], of 50 mM, and a pHi of 7.0, the axon’s acid-extrusion rate is exceeding low, being able to raise pHi by only -0.1 pH unit per hour (Boron and De Weer, 1976a). After a few hours, the pH, recovery halts even though pHi is far below its normal value. We suspect that long-term metabolic changes either decrease the acid extrusion rate and/or increase the acid-loading rate, thereby preventing a complete pHi recovery. IV.

EFFECT OF METABOLIC INHIBITORS Off pH1

Cyanide (CN-), 2,4-dinitrophenol (DNP), and azide (Ny) all produce a slow and prolonged intracellular acidification that begins after a short delay. In addition, the latter two substances cause a more rapid but brief phase of acidification that begins very soon after they are applied. As illustrated in Fig. 5A, cyanide has little effect on pHi for the first 5 min, after which it causes pH, to decline by -0.3 over the next -40 min (Boron and De Weer, 1976a). The lack of an immediate acidification implies that the influx of the neutral weak acid (HCN = H+ + CN-) is not followed by a substantial dissociation into H + and CN-. Indeed, the dissociation is expected to be very small because HCN has such a high pK; (-10). The slow fall in pHi is likely the result of a buildup of acidic metabolites,

FIG.5 . (A) Effect of cyanide on pH, and V , . Exposing the axon to ASW containing 2 mM NaCN (pH, = 7.70) causes, after a 5-rnin delay, a slow fall of pH, of -0.3. Simultaneously, there is a transient depolarization, followed by a sustained hyperpolarization. The effect of CN is only partially reversible in the first few minutes. (B) Effect of azide on pH, and V,. Application of 3 mM NaN, (pH, = 7.70) causes a rapid internal acidification followed, after a brief delay, by a slower pH, decline. The membrane voltage undergoes a transient depolarization. followed by a sustained hyperpolarization. The changes are only partially reversible in the first several minutes. (From Boron and De Weer, 1976a; reproduced by permission of the Rockefeller University Press.)

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WALTER F. BORON

following inhibition of oxidative phosphorylation. It is unlikely to be due to the shuttling of protons across the cell membrane by HCN/CN-, inasmuch as [CN-Ii is exceedingly low (-2 p M ) . Removal of cyanide produces only a small pHi recovery over the first several minutes. The application of azide (3 mM, pH, = 7.70) causes an abrupt fall in pHi of -0.1 (see Fig. SB), followed by a period of stability. If the azide is removed during this period, pHi rapidly returns to its initial value (not shown). Thus, the rapid azide-induced acidification is probably the result of the influx and dissociation of the neutral weak acid (HN3 = Ht + N;; pKA = 4.7). During lengthier exposures to azide, pHi eventually begins a slow decline, similar to the slow phase of acidification produced by cyanide. This slow phase of acidification is likely caused by the build-up of acidic metabolites, the result of azide's uncoupling of oxidative phosphorylation. The inward shuttling of protons across the cell membrane by the HN3/N: couple is probably not a major contributor to this acidification, inasmuch as the slow pHi fall begins only after a lengthy delay, and because reversing the N; electrochemical gradient has only a small effect on pHi (Boron and De Weer, 1976a). Removal of the azide causes a rapid pHi recovery, but only to the same extent as the initial, rapid phase of azide-induced acidification. 2,4-Dinitrophenol (1 mM, pH, = 7.70) has an effect similar to that of azide, except that the slow phase of acidification follows the rapid, initial one without an appreciable delay. V.

IONIC MECHANISM OF pH1 REGULATION

The existence of an axonal pHi-regulating mechanism is indicated by the relatively high value for steady-state pHi (i.e., -7.3 vs the equilibrium pHi of -6.8) and by the plateau-phase recovery of pHi from a C02-induced acid load (see Fig. 4B). These results suggest that an axonal pHi regulator responds to intracellular acid loads by extruding acid from the cell. Detailed mechanistic information can be gained by studying the recovery of pHi from acute, intracellular acid loads. A. Dependence on External HCO;

In the experiment of Fig. 4B, the recovery of pHi from the COJnduced acid load necessarily took place in the presence of external HCO,. The experiment of Fig. 6 was designed to test whether HCO; is required for the recovery of pHi from an acute acid load (Boron and De Weer, 1976b). The acid load is applied by pretreating the axon with an ASW containing 50 mM NH: (pH, = 7.7). As described above, the application of the NH3/

259

REGULATION OF AXONAL pH

F FIG.6 . Effect of external HCO; on recovery of pH, from an acid load. The axon is acid loaded by pretreating it with artificial seawater (ASW) containing 50 mM NH: (pH, = 7.7), and then washing out the NHJNH:. As shown in Fig. 3B, the washout causes pH, to undershoot its initial value; i.e., the cell is acid loaded. In the nominal absence of HCO; (segment AB), pH, fails to recover. During the application of ASW containing 10 mM HCO; at the same pH,, however, pH, recovers more rapidly (segment C D ) . After washout of the CO,/HCO;, pH, no longer recovers (point E ) . (From Boron and De Weer, 1976b; reproduced by permission of Macmillan Journals, Ltd.)

NH: causes an abrupt increase in pHi (due to NH3 influx) followed by a slower fall (due to NH; influx). When the external NH3/NH: is washed out, pHi falls to a value substantially lower than its initial one (i.e., the cell is acid loaded). In the nominal absence of HCO, (segment AB), pHi recovers only very slowly, even though pH, is 8. The application of ASW containing 0.4% C 0 2 and 10 mM HCO; at constant pH, causes a transient acidification (BC) due to the influx of C 0 2 .This is followed, however, by a marked acceleration of the pHi recovery (compare CD and AB). This suggests that external HCO; is required for acid extrusion in the squid axon, The rate of the segment -CD recovery can be used to calculate the rate of acid extrusion (i.e., the equivalent net HCO; influx, JFbo3). This is simply the product of the recovery rate (dpHJdt), intracellular buffering power (p),and axonal volume-to-surface ratio ( p ) . A more extensive series of experiments (Boron and Russell, 1983), similar in design to the experiment of Fig. 6 (i.e., in nondialyzed axons),

260

WALTER

KINETICS OF

F. BORON

TABLE I AXON'SpH,-RECUt.ATING

THE SQUID

SYSTEMS''

External HCO, External Na' Internal CIP

30 21 38

2.3 +- 0.2 77 -t 13 84 f 15

10.6 2 0.4 10.3 t 0.6 19.6 1.2

*

Kinetic parameters obtained from nonlinear, leastsquares curve fit. n is number of data points (measurements of net, equivalent HCO, flux); V:,,, is apparent maximal velocity; KL, is substrate concentration at half VLdx. Standard conditions: [Na'], = 425-437 mM, [HCO;], = 12 mM, pH, = 8.0, [el-], = 100 m M , pH, = 6.7.

has shown that the dependence of acid-extrusion rate on [HCO,], follows Michaelis-Menten kinetics (see Table I). The apparent K , for external HCOT is -2.3 mM (pH, = 8.0, pHi = 6.7, T = 22°C) and the apparent VmaXis 10.6 pmol ~ r n sec-' - ~ (at a physiological [CI -Ii of 70-100 mM). The magnitude of the acid-extrusion rate has been carefully documented (Boron and Russell, 1983) in a series of experiments on internally dialyzed axons. Isotopically measured fluxes of Na+ and C1- (see below) were determined under identical conditions (pHi = 6.7, [ClFIi= 150 mM, pH, = 8.00, [HCO;], = 10 mM, (Na'l, = 425 mM, T = 16°C) for the purpose of determining the stoichiometry of acid extrusion. The approach in the acid-extrusion rate experiments was to dialyze the axon with a lowpH fluid until pHi fell to -6.7, at which time dialysis was halted, thereby returning control of pHi to the axon. After dialysis was halted, pHi failed to recover (i.e., increase) until HCOC was added to the external solution. From thc rate of HCOi-stimulated pHi recovcry, axon diameter, and buffering power,' the acid-extrusion rate (J$io,) was computed. In 15 axons, the mean JnHeEOjwas 7.5 pmol cm12 sec-' (see Table 11). This acid extrusion was completely blocked by the anion-flux inhibitor 4-isothiocyano-4'acetamido -2,2'-disulfonate (SITS), as had previously been demonstrated The non-C02 or intrinsic buffering power (PI)was determined by halting dialysis, blocking the pHi,-regulating systems with SITS, and then exposing the axon to ASW containing 0.2 mM NH; at pH 7.75 (see Boron, 1977; Boron and Russell, 1983). PI, the quotient of the calculated change in [NH;], and the measured pHi change, came to 11.2 mM. The total buffering power (&) is the sum of PI and the COz buffering power (PcOz).The latter is calculated from the relation pco, = 2.3 [HCO, Ii,and comes to I .7 mM under the conditions of these experiments. Thus, PT = 11.2 + 1.7 = 12.9 mM.

REGULATION

OF AXONAL

261

pH

TABLE I1

AXON'S ~H,-RECUI.ATING SYSTEM"

STOICHIOMETRY OF THE SQUID

Ion

Na' CIHCO;

Influx

Efflux

3.4 t 0.4 (13) 0.1 t 0.2 (6) -

0.0 t 0.1 (6) 3.9 t 0.2

Net flux (pmol cm sec-I) +3.4

-3.8

(15)

-

1.5

%

0.6

(15)

Mean kSE, with number of experiments given in parentheses. The HCOj flux was calculated from the rate of pH, recovery and I S the net, equivalent HC03 flux referred to in the text. Standard conditions: temperature = 16°C. [Nail, = 425 mM. [HCO,] = 12 mM, pH, = 8.0, [CI 1, = 150 mM, pH, = 6.5.

for this preparation (Russell and Boron, 1976), as well as the snail neuron (Thomas, 1976a) and barnacle muscle (Boron, 1977). 6. Dependence on External Nai

In 1977, Thomas showed that acid extrusion in snail neurons requires external Na+, an observation confirmed on giant barnacle muscle fibers (Boron et ul., 1979). The experiment of Fig. 7 demonstrates how the recovery of pH, from an NHJNH4f-induced acid load depends on "a+], in the squid axon (Boron and Russell, 1983).After removal of external NHl/ NH;, pH, is allowed to fall and level off (segment ab) in a nominally HC0;-free ASW. Application of 10 mM HCO; ASW containing 425 mM Na+ (bc)causes a rapid recovery of pH, (calculated JFbo3 = 9.7 pmol cm-2 sec-I). The pH, recovery is slowed (cd) by reducing "a+], to 15 mM (JE& = 1.1 pmol cm-2 sec-I), and then accelerated (de) by raising "a+], to 100 mM (J$& = 6.4 pmol cm-2 sec-I). The results of a series of such experiments showed that the dependence of acid-extrusion rate on "a+], follows Michaelis-Menten kinetics, with an apparent K , of 77 mM (pH, = 8.00, pH, = 6.7, temperature = 22°C). Not only does acid extrusion require external Na+, but it is accompanied by a net influx of Na+. Unidirectional Na+ effluxes and influxes (measured with 22Na)were determined in experiments in which axons were dialyzed to a pH, of 6.7 and a [CI-1, of 1.50 m M .The pH,-regulating system-linked Na+ flux was defined as the increase in the Na+ flux pro-

262

WALTER F. BORON

klOO N$ PHO 7 7 0

,

12 HCOi

[Notlo:

425

pHo 8.00 IS I00

Fie. 7 . Dependence of acid-extrusion rate on [Na+l0.The axon was acid loaded by pretreating it with artificial seawater (ASW) containing 100 mM NH: (pH, = 7.70). After washout of the NH1/NH:, pH, failed to recover from the acid load (segment ub) until HCO; was added to the ASW. The subsequent pH, recovery was rapid when "a'], was 425 mM ( h c ) , but it slowed considerably when "a+], was reduced to 15 mM ( c d ) . Raising "a+], to 100 mM (de) brought the pH, recovery rate to a value intermediate between the first two values. (From Boron and Russell, 1983; reproduced by permission of the Rockefeller University Press.)

duced by the application of 10 mM HCO, (pH, = 8.0) ASW. We found that, under standard conditions, the unidirectional Na+ efflux mediated by the axon's pHi-regulating system is approximately zero, and that the influx is about 3.4 pmol cm-2 sec-' (Boron and Russell, 1983). Thus, the pHi-regulating system mediates a net Na' influx (see Table 11). This net Na+ influx shares all the properties of the HCO: flux: (1) it is blocked by SITS; (2) it requires internal C1-; and (3) it requires internal ATP. The

263

REGULATION OF AXONAL pH pH 6.6 k 2 0 0 CI-

Inside:

'

pH, 8.00

'

Outside:

12 HCOj

I

5:"l

7.4

7.2

350 CI-

pHi 7.0

6.8

6.6

30 rnin

FIG.8. Dependence of acid-extrusion rate on [CI-1, . The axon was loaded by predialyzing (segment ab) it with a fluid at pH 6.6, containing 200 mM C1-. When dialysis was halted, returning control of pH, to the axon ( b c ) ,there was no pH, recovery until 12 m M HCO, was added to the artificial seawater (cd). The inset shows the results of three similar experiments (during segments bc and c d ) in which the axons were dialyzed to [CI-1, levels of 0, 100, and 350 mM CI-. (From Boron and Russell, 1983; reproduced by permission of the Rockefeller University Press.)

magnitude of the net Na+ influx is about 0.45 as large as the J;;CO3obtained under identical conditions. This is consistent with a HCO, : Na+ stoichiometry of 2 : 1. C. Dependence on Internal CI-

When internal CI- is removed by dialyzing the axon with a C1-free solution, acid extrusion is blocked (Russell and Boron, 1976). This dependence on internal CI- is demonstrated by the experiment of Fig. 8 . The axon is dialyzed with a low-pH solution containing 200 mM CIF (segment ab). After dialysis is halted, returning control of pHi to the axons, there is no pHi recovery (bc) until HCOj is added to the external solution. The HCOy-stimulated rise in pHi (at the beginning of the segment cd pHi recovery) corresponds to an acid-extrusion rate of 15.0 pmol cm-2 sec-I. The inset shows three additional experiments (each on a different axon) at [Cl-Ii values of 0, 100, and 350 mM. It is clear that the pHi-recovery rate gradually increases as [ClFIi is raised. The results of 38 similar experi-

264

WALTER

F. BORON

ments show that the dependence of acid-extrusion rate on [C1-Ii follows Michaelis-Menten kinetics, with an apparent K , for internal C1- of 84 mM (see Table I). This value is in the range of reported normal values of [CIVliin squid axons (Brinley and Mullins, 1965) and indicates that the axon's pHi-regulating system is normally only about half saturated with respect to intracellular C1- . This is in marked contrast to the other two substrates (i.e., external Na+ and HCO,), for which the transporter is nearly saturated under physiological conditions. The dependence of acid extrusion on internal C1- has also been demonstrated for snail neurons by Thomas (1977). Acid extrusion not only has an absolute dependence on internal C1-, it is accompanied by a net CIV efflux. When the pHi of dialyzed axon is lowered to 6.7 under the aforementioned standard conditions (pHi = 6.7, [Cl-li = 150 mM, pH, = 8.00, [HCO,], = 10 mM, "a+], = 425 m M , temperature = 16°C) the addition of 10 mM HCO, to the ASW has minimal effect (stimulation of -0. I pmol ern--* sec - I ) on the unidirectional CIinflirx (see Table II), measured with "CI. The CIV c f f h x , however, increases by about 3.9 pmol cm-2 sec-'. The net CI- efflux, 3.8 pmol cm-* sec-I, is 51% as large as the net equivalent HCOY flux, but is approximately the same as the net N a + influx. The external HCOy-stimulated C1efflux has all the properties expected of a flux tightly linked to the pHiregulating system (Boron and Russell, 1983): it ( I ) is blocked by SITS, (2) requires external Na+, (3) requires internal ATP, and (4) is stimulated by IOW pHi. D. Dependence on Internal ATP

If squid axons are treated with either cyanide (2 mM) or DNP (1 mM), pHi fails to recover from a C02-induced acid load. This inhibition is reversible in both cases (Boron and De Weer, 1976b). Although Caldwell (1960) had previously shown that treatment with cyanide or DNP reversibly lowers [ATPI, substantially, the above experiments left open the possibility that cyanide and DNP inhibit the pHi regulator directly, not through their effect on ATP levels. The experiment of Fig. 9 was designed to determine whether, in the continuous presence of cyanide, the addition of ATP would restore the axon's ability to recover from an acid load. In the first part of the experiment, the axon is dialyzed with an ATP-free solution for 1 hr; the dialysate's pH is lowered to 6.6 during the final 30 min to acid load the axon. When dialysis is halted (point a ) , returning control of pH, to the axon, there is no pHi recovery (segment u b ) , even

265

REGULATION OF AXONAL pH INTERNAL FLUID

-

ATP fro. pH Z3

'

pH 6.6

EXTERNAl FLUID

~

8mMATP pH 7.3' pH 6.6

, HC03

FIG.9. Dependence of acid extrusion on ATP. The axon was exposed throughout to artificial seawater containing 2 mM CN. Predialysis with an ATP-free fluid at pH 7.3 was followed with a similar fluid at pH 6.6. After pHi fell to -6.8, dialysis was halted, returning control of pH, to the axon. There was no pH, recovery, however, either in the absence (segments ah and c d ) or in the presence of HCO;. After ATP was reintroduced to the axon (de), in the continued presence of ATP, the application of HCO, elicited a pH, recovery (fg) which was blocked by SITS ( g h ) .(From Russell and Boron, 1976; reproduced by permission of Macmillan Journals. Ltd.)

during stimulation with HCO.7 (bc). After the axon is dialyzed with an ATP-containing solution ( d e ) ,however, application of HCO, does elicit a recovery of pHi from the acid load ( f g ) , and this recovery is blocked by SITS ( g h ) . Thus, ATP or a related substance is required for acid extrusion. As noted above, ATP is also required for the net Na+ influx and C1efflux that are coupled to acid extrusion. Although ATP is necessary for transport, it is not clear whether hydrolysis of ATP is stoichiometrically linked to transport, or whether ATP merely fills a catalytic role. E. Possible Mechanisms of Transport

The above data indicate that the squid axon possesses a SITS-sensitive ion-transport mechanism that responds to intracellular acid loads by extruding acid from the cell. The transport not only requires external HCO? and Na+ and internal C1-, but is accompanied by a net Na+ influx and a net C1- efflux. The data are consistent with a stoichiometry of one Na+ entering the axon for each C1- exiting, and for every two acid equivalents

266

WALTER F. BORON

neutralized inside the axon (see Table 11). The implication of the stoichiometry data that transport is electrically neutral is corroborated by the observation that changes in membrane potential are without significant effect on the acid extrusion rate. These data are accounted for by each of the models of Fig. 10. The first model (Thomas, 1977) has Nat and HCOj entering in exchange for C1- and H+. In the second, the exit of H+ is replaced by the entry of a second HCO,. In the third model, the entry of a single C0:- replaces that of two HCOT. Finally, according to the last model (Becker and Duhrn, 1978; Boron, 1980), Na+ and COi- enter only after forming the ion pair NaCOT (Garrels et al., 1961). These models are equivalent thermodynamically. The free-energy change associated with the overall transport process (AGnec)is the sum of the individual freeenergy changes of the transported ions. In the following example, these free-energy changes are computed for the relatively low pHi of 6.7, assuming normal values for [Cl-]; and “a+];. The calculations are based on the first model of Fig. 10. OUT

IN

FIG.10. Models of acid extrusion. The models are equivalent thermodynamically. They all predict electroneutral transport with a stoichiometry equivalent to one NaCtaken up for each C1- lost, and for each two protons neutralized inside the cell tie., by the uptake of HCO; or a related species and/or the efflux of H+). (From Boron and Russell, 1983; reproduced by permission of the Rockefeller University Press.)

REGULATION OF AXONAL pH

AGNa = RT In(INa+],/INa+l,) RT In([HCOi],/[HCOj],) ACcl = RT In([CI-l,/[CI 1,) AGH = RT In([H+],/[H+],)

AGHCo, =

267 RT ln(50/425) RT ln(0.5/10) = RT ln(550/100) = RT ln(10'8/10-6 7, =

=

=

= = =

-2.14RT -3.00RT +1.70RT -3.00 RT

Thus, the overall free-energy change of the acid-extrusion reaction (pHi = 6.7) is -6.44 RT, the negative value indicating that the reaction ought to proceed spontaneously as written (i.e., extruding acid from the cell). At higher pHi values the calculated AG,,, is less negative (i.e., [H+Iifalls and [HCOjli rises). However, even at the normal pHi 7.3, where the observed is greatly reduced and the accompanying Na+ and C1- fluxes are undetectably low, the AG,,, is still -3.66 RT, and thus still favors the net forward reaction by a wide margin. A similar relationship among AGnet,acid extrusion, and pHi exists in barnacle muscle. In this cell, pHi is regulated by a Na/HCO,-Cl/H exchanger similar to that of the squid axon. The acid-extrusion rate in barnacle muscle falls to near zero at the pHi of 7.3-7.4, even though there is sufficient energy in the ion gradients to drive pHi to -8. The barnacle's pHi-regulating system also mediates bidirectional C1- fluxes and bidirectional Na+ fluxes. Interestingly, all these fluxes are blocked at pHi values greater than -7.3 (Boron et al., 1979; Russell et al., 1983). Thus, all known modes of ion transport by the pHi regulator are blocked at pHi values exceeding 7.4. It is unlikely that the inhibition of all modes of transport at pHi > 7.4 is due solely to unavailability of substrate (i.e., intracellular H+). Indeed, if the apparent transporter-mediated C1- influx and Na+ efflux were due to the microscopic reversibility of the pHi regulator, then one might expect low [H+Iito stimulate rather than inhibit these fluxes. It would thus appear that there is an allosteric site for H+ on the internal surface of the barnacle muscle's pHi regulator. We would predict that this site must be protonated in order for the transporter to operate in any mode. Inasmuch as the squid axon's pHi regulator does not appear to mediate an appreciable Cl- influx or Na+ efflux during acid extrusion (see Sections V,B and V,C), it will be difficult to establish the presence or the absence of a similar allosteric H+ site in this preparation. It is interesting to note that the Na-H exchanger, which regulates pHi in several vertebrate cell types, has a pHi dependence that is similar to that of the Na/ HCO,-Cl/H system (see Aickin and Thomas, 1977; Boron and Boulpaep, 1982a), which suggests that it may have an internal allosteric site for H+. This suspicion has been confirmed by the vesicle studies of Aronson et al. (1982).

268

WALTER F. BORON

Concerning the ATP dependence of the squid axon’s pHi regulator, the preceding thermodynamic calculations point out that there is more than sufficient energy in the gradients of the transported ions to account for the activity of the pH, regulator. If the ion gradients were the only source of energy for the transporter, then one might expect transport to bc reversed by appropriately inverting the gradient of one or more of the transported ions. When such gradient inversions are imposed on the barnacle muscle’s Na/HCO,-CI/H exchanger, for example, transport clearly reverses (Russell et ul., 1983). Reversal of the pH, regulator, however, has yet to be demonstrated in squid axons, despite considerable effort (Boron, unpublished). The squid axon pHi regulator’s dependence on ATP and its resistance to reversal could be explained by a tight coupling of transport to ATP hydrolysis. However, we cannot yet rule out the possibility that ATP is required merely for catalytic purposes, and that reversal of acid extrusion is difficult to observe only for kinetic reasons. Although the four models of Fig. 10 are equivalent thermodynamically, they do not necessarily make the same kinetic predictions. In particular, the ion-pair model predicts that the acid-extrusion rate should not depend on “a’],, or [HCO;], per se, but on [NaCOj] at a given pH. Indeed, the Na+ and HC03- data of Table I indicate that whether [Na ‘I, is varied at a constant [HCOj], of 12 mM or [HCOI], is varied at a constant “a+] of 425 mM, the acid-extrusion rates fall on the same velocity vs [NaCO;], curve. The apparent K , for NaCOr is 74 W M (Boron and Russell, 1983). More recently, these studies have been expanded to include the [Na’], dependence of acid extrusion at three [HCOI] levels (12, 6 , and 3 mM), and the [HCOj], dependence at three “a+] levels (425, 212, and 106 rnM) (Boron, unpublished). In all cases, the data fall on the same velocity vs [NaCOy1, curve, in agreement with the ion-pair model. REFERENCES Aickin, C . C., and Thomas, R. C. (1977). An investigation of the ionic mechanism of intracellular pH regulation in mouse soleus muscle fibers. J . Physiol. (London) 273, 295-3 16. Aronson, P . S . , Nee, J . , and Suhm, M. A. (1982). Modifier role of internal H’ in activating the Na-H exchanger in renal microvillus vesicles. Nature (London) 299, 161-163. Recker, B . F., and Duhm, J. (1978). Evidence for anionic cation transport of lithium, sodium and potassium across the human erythrocyte membrane induced by divalent anions. J . Physio/. (London) 282, 149- 168. Bicher, H . , and Ohki, S. (1972). Intracellular pH electrode experiments on the giant squid axon. Biochirn. Biophys. Aria 255, 900-904. Boron, W. F. (1977). lntracellular pH lransients in giant barnacle muscle fibers. A m . J . Physiol. 233, C61-C73. Boron, W. F. (1980). Intracellular pH regulation. Curr. T ( J ~ Membr. . Trcinsp. 13, 3-22.

REGULATION OF AXONAL pH

269

Boron, W. F., and Boulpaep, E. L. (1982a). Intracellular pH regulation in the renal proximal tubule of the salamander: Na-H exchange. J . Gen. Physiol. 81, 29-52. Boron, W. F., and Boulpaep, E. L. (1982b). lntracellular pH regulation in the renal proximal tubule of the salamander: Basolateral HCO, transport. J . Gen. Physiol. 81, 53-94. Boron, W. F., and De Weer, P. (1976a). Intracellular pH transients in squid giant axons caused by C 0 2 , NH,, and metabolic inhibitors. J . Gen. Physiol. 67, 91-112. Boron, W. F.. and Ile Weer. P. (1976b3. Active proton transport stimulated by CO?/HCO:r. blocked by cyanide. Nature (London) 259, 240-241. Boron, W. F., and Russell, J . M. (1983). Stoichiometry and ion dependencies of the intracelMar-pH-regulating mechanism in squid giant axons. J . Gen. Physiol. 81, 373-399. Boron, W. F., McCormick, W. C., and Roos, A. (1979). pH regulation in barnacle muscle fibers: Dependence on intracellular and extracellular pH. A m . J . Physiol. 237, C185C193. Brinley, F. J., and Mullins, L. J. (1965). Variations in the chloride content of isolated squid axons. Physiologisl 8, 121. Brinley, F. J . , and Mullins, L. J. (1967). Sodium extrusion by internally dialyzed squid axons. J . Gen. Physiol. 50, 2303-233 I. Caldwell, P. C. (1954). An investigation of the intracellular pH of crab muscle fibers by means of microglass and micro-tungsten electrodes. J . Physiol. (London) 126, 169-180. Caldwell, P. C. (1956). lntracellular pH. Int. Reu. Cyrol. 5, 229-277. Caldwell, P. C. (1958). Studies on the internal pH of large muscle and nerve fibers. J . Physiol. (London) 142, 22-62. Caldwell, P. C. (1960). The phosphorus metabolism of squid axons and its relationship to the active transport of sodium. J . Physiol. (London) 152, 545-560. Garrels, R. M., Thompson, M. E., and Siever, R. (1961). Control of carbonate solubility of carbonate complexes. A m . J . Sci. 269, 24-45. Hinke, J . A. M. (1967). Cation-selective microelectrodes for intracellular use. In “Glass Electrodes for Hydrogen and Other Cations” (G. Eisenman, ed.), 464-477. Dekker, New York. Roos, A., and Boron, W. F. (1981). lntracellular pH. Physiol. Rev. 61, 296-434. Russell, J. M . , and Boron, W. F. (1976). Role of chloride transport in regulation of intracellular pH. Nature (London) 264, 73-74. Russell, J . M., Boron, W. F., and Brodwick. M. S. (1983). lntracellular pH and Na fluxes in barnacle muscle with evidence for reversal of the ionic mechanism of intracellular pH regulation. J . Gen. Physiol. 82, 47-78. Spyropoulos, C. S. (1960). Cytoplasmic pH of nerve fibers. J . Neurochem. 5 , 185-194. Thomas, R. C. (1976a). Ionic mechanism of the H + pump in a snail neurone. Nature (London) 262, 54-55. Thomas, R. C. (1976b). The effect of carbon dioxide on the intracellular pH and buffering power of snail neurones. J . Physiol. (London) 255, 715-735. Thomas, R . C. (1977). The role of bicarbonate, chloride and sodium ions in the regulation of intracellular pH in snail neurons. J . Physiol. (London) 273, 317-338. Waddell, W . J . , and Bates, R. G. (1968). Intracellular pH. Physiol. Reu. 49, 285-329.

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CURRENT TOPICS IN MEMBRANES AND TRANSPORT, VOLUME 22

Hormone-Sensitive Cyclic Nucleotide Metabolism in Giant Axons of Loligo P . F . BAKER AND A . CARRUTHERS Department of Physiology King’s College University of London London, Englund

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Cyclic AMP Levels in Intact Axons.. ..................................... 111. cAMP Production and Breakdown in Perfused Axons.. ..................... IV. Conclusion. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References .............................................................

271 272 273 276 276

I. INTRODUCTION

Although the squid axon has served as an excellent model system for the study of many aspects of neuronal function, very little attention has been given to the possibility that it may also be useful for investigating the control of cyclic nucleotide metabolism. We have begun to examine cyclic nucleotide metabolism in giant axons of Loligo forbesi (Baker and Carruthers, 1982). Measurements on extruded axoplasm reveal concentrations of cAMP and cGMP within the range found in many other cell types, and, at least for CAMP,the concentration in axoplasm is subject to hormonal control. This last finding was unexpected and suggests that the squid axon may prove to be an excellent preparation in which to probe the regulation of hormone action: for instance, its dependence on the intracellular and extracellular environments and its sensitivity to membrane potential. 271 Copyright 0 1984 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-153322-0

272

P. F. BAKER AND A. CARRUTHERS

II. CYCLIC AMP LEVELS IN INTACT AXONS The CAMP content of axoplasm is usually in the range 25-150 n M and that of cGMP 100-500 nM. Table I summarizes our findings on the cAMP contents of axoplasm extruded from axons that have been subjected to various treatments. Particularly noteworthy are the findings that the phosphodiesterase inhibitor caffeine elevates the level of cAMP in axoplasm and electrical stimulation lowers CAMP. This latter effect is reduced in a nominally Ca-free medium. The most dramatic changes, however, are found with certain agents that, in other systems, arc known to act a h neurotransmitter substances or hormones. At a concentration of 500 pM octopamine, tyramine, and substance P have no significant effect on the cAMP content of axoplasm; but at this same concentration, the level of cAMP is increased three- to fourfold by 5-hydroxytryptamine (serotonin), doubled by secretin, and halved by antidiuretic hormone. 1.-Glutamate,

TABLE 1 cAMP CONTENT OF S Q U I D AXOPI ASM A r 1 1 R E X I ~ O S U OFK ~ I N I AC I AXONSTO VARIOUSTREATMENTS”

Solution 10 Ca, 400 Na SW 10 Ca, 400 Na SW Ca-free Na SW Ca-free Na SW 10 Ca, 400 Na SW containing: Serotonin (500 p M ) Serotonin (SO0 pLM) methysergide (1 m M ) ADH (500 p M ) Caffeine ( 5 mM) Cyanide (2.5 m M )

Conditions of exposure -

cAMP content relitive to control in ASW

I .o 0.5

Number of determinations -

0.70

5 3 3

15 rnin

3.3

7

15 min 15 min 15 rnin 5 hr

I .25 0.37 2.0 0.09

Stirn. 200 Hz, 20 min 15 rnin Stim. 200 Hz, 20 min

1 .o

I? Axons of Lo/igo,forbesiwere exposed to the treatment specified for IS min at 20-22°C before the axoplasm was extruded. weighed, and homogenized in ice-cold ethanol. The cAMP content of the ethanolic extract was measured by radioimmunoassay. The cAMP content varied appreciably between animals and tended, on average, to tall during the squid season (September-December). For these reasons, cAMP levels are expressed relative to control axons immersed in artificial seawater (SW) containing (mM): NaCI, 400; KCI, 10; MgCI2, 100; CaCI2, 10; NaHCO,, 2.5; pH 7.8.

CYCLIC NUCLEOTIDE METABOLISM IN SQUID GIANT AXONS

273

carbamylcholine, and dopamine have not been examined at SO0 pM, but are without significant effect at 100 p M . We have examined only the action of 5-HT in any detail. Figure I shows the relation between 5-HT concentration and cAMP levels in axoplasm. The increase in cAMP content is half-maximal at a 5-HT concentration of approximately 1 pM. This observation together with the finding that the effect of 5-HT is abolished by inclusion in the external medium of the 5-HT antagonist methysergide (Table I) suggests that the squid axon contains conventional 5-HT receptors. The response to 5-HT is half complete in about 2 min and is fully reversible. Exposure of highly cleaned axons to 3H-labeled 5-HT reveals very considerable binding to the axon sheath. This binding is a saturable function of the 5-HT concentration and seems to reflect a large uptake of 5-HT into the Schwann cell sheath. Uptake into the interior of the giant axon is a linear function of 5-HT concentration over the range 0 to 50 p M . 111.

cAMP PRODUCTION AND BREAKDOWN IN PERFUSED AXONS

The great advantage of giant axons is that their internal environment can be controlled by perfusion or dialysis, and we were curious to discover whether the action of 5-HT persists in perfused axons. Figure 2

FIG.1 . Cyclic AMP content of axoplasm extruded from axons of Lo/i,yo,forbe.si after 15 min of exposure to different concentrations of 5-HT. The data were obtained late in the squid season when, as noted in the footnote to Table 1, the resting cAMP content of axoplasm was quite low. The best fit to'the data gives an apparent affinity for 5-HT of I .44 rt 0.45 p M . Temperature. 20-22°C.

P. F. BAKER AND A. CARRUTHERS

274

0

0 tlme

IATPI,

mM

I,

FIG.2 . Cyclic AMP content of perfusate emerging from an axon of Loligoforbesi. The perfusion medium was (rnM): KzS04.200; taurine, 400; MgCL, 10; NaCI, 20; CaCI:, 1.S; ECTA, 5 ; Tris, 5 ; pH 7.3; FCCP, 8 y M . Axon diameter, 821 prn; temperalure, 12°C. 5-HT ( 5 0 0 p M ) was applied during thc period indicated by the solid bar.

shows this to be the case. Cleaned axons were doubly cannulated and perfused by the method of Oikawa et al. (1961), in which a fine glass capillary is used to remove a central core of axoplasm before perfusing fluid through the space created. When the perfusion solution contained ATP, the perfusate emerging from the axon contained easily measurable concentrations of CAMP. Production of cAMP was half-maximal at an ATP concentration of 0.12 mM, and at 12°C the maximum rate of cAMP production was 4 fmol c w Z sec I . Figure 2 shows that the amount of cAMP emerging from the axon was increased by external application of SHT and also by inclusion of GTP in the perfusion medium. A systematic analysis of the ionic and nucleotide requirements of these effects has not been made. As the level of CAMP in axoplasm and in the perfusate emerging from a perfused axon presumably reflects the relative rates of cAMP synthesis by adenylyl cyclasc and breakdown by phosphodiesterase, we have attempted to measure these enzyme activities both separately and simultaneously in a single perfused axon. We have made use of CAMP-binding protein to sequester cAMP emerging from axons perfused either separately or with a mixture of 14C-labeledATP and 'H-labeled CAMP.The I4C associated with the binding protein is assumed to reflect synthesis of CAMP, and the difference in 'H associated with the binding protein in samples of the solution entering and leaving the axon provides a measure of cAMP disappearance, presumably by breakdown. When axons are perfused at 4.S pllmin with 0.1 ,uM CAMP, approximately 3-10% of the CAMP disappears during a single passage through the axon. The disappearance of cAMP is almost completely prevented by 10 mM caffeine,

CYCLIC NUCLEOTIDE METABOLISM IN SQUID GIANT AXONS

WM nM cAMPi

100 nMATP,

275

I

FIG.3. Simultaneous measurement of cAMP synthesis and breakdown in a single axon of Loligo forbesi. Artificial axoplasm (see legend to Fig. 2 ) containing 100 nM [I4C]ATPand 100 nM ['HICAMP was perfused through the axon at 4.5 pl/min. Temperature, 12°C; axon diameter, 820 pm.

which rules out the possibility that cAMP is being lost from the axon and supports the view that it is being broken down by a caffeine-sensitive process, presumably phosphodiesterase. We have not as yet made a systematic analysis of the factors that affect cyclase and phosphodiesterase activities in perfused axons, but Fig. 3 gives some idea of the potential of this approach. This experiment, in which, it should be stressed, conditions for optimal cyclase and diesterase activity may not have been achieved, reveals a number of interesting features of cAMP metabolism in squid axons. These include the findings that (1) the activity of cyclase but not diesterase is increased by raised external K ; (2) externally applied 5-HT inhibits diesterase and only slightly stimulates cyclase; (3) raising ionized Ca inhibits diesterase cyclase; and (4)caffeine inhibits diesterase with no effect on cyclase. These alterations in cyclase and diesterase activity are in the right direction to explain the effects of 5-HT on cAMP levels in intact axons (Table I), but do not help account for the effects of electrical stimulation. The dramatic inhibition of cAMP breakdown by 5-HT suggests that much of the phosphodiesterase in squid axons is closely associated with the axolemma.

276

P. F. BAKER AND A. CARRUTHERS

IV. CONCLUSION

These few experiments suggest that the perfused squid axon will prove to be an excellent preparation in which to study hormonal control of CAMP metabolism. It will be of particular interest to extend these observations to cGMP, the level of which seems to be decreased by 5-HT, and to investigate the effects of membrane potential on cyclic nucleotide metabolism in perfused axons. A further question that must be addressed concerns the function of hormone-sensitive cyclic nucleotide metabolism in squid axons. In view of the modulatory effects of cyclic nucleotides on the electrical properties of other molluscan neurons (see Siegelbaum rt al., 1982), it may be rewarding to examine the effects of 5-HT and cyclic nucleotides on the electrical properties of squid axons; to date, however, no experiments of this kind have been reported. ACKNOWLEDGMENTS The experiments described above were supported by the Medical Research Council of Great Britain and carried out at the Laboratory of the Marine Biological Association, Plymouth. We wish to thank the Director and staff of the Marine Riological Association for supplying material and facilities for this work. REFERENCES Baker, P. F., and Carruthers, A. (1982). Cyclic AMP metabolism in squid giant axons and giant barnacle muscle fibres. J . Physial. (London)326, 13P. Oikawa, T., Spyropoulos, C. S . , Tasaki, I., and Teorell, T. (1961). Methods for perfusing the giant axon of Loligo pecrlii. Actu Physial. Scund. 52, 195-196. Siegelbaum, S. A,. Camardo, J. S., and Kandel, E. R. (1982). Serotonin and cyclic AMP close single channels in Aplysin sensory neurones. N ~ f r t r (Landan) e 299, 413-417.

Part Ill

Excitability

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CURKENT TOPICS IN MEMBRANES AND TRANSPORT. VOLUME 22

Hodgkin-Huxley: Thirty Years After H . MEVES I . Physiologisches lnstitut Universitar des Saarlandes Homburg-Saar, Federal Republic of Germany

I.

...

....................................... 11. 111. ................................... .......................... IV. V. Reconstruction of the Electrical Behavior of the Squid Axon from the Hodgkin-Huxley Equations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Molecular Properties of the Na Channel: Permeability Ratios and Binding Constants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V l f . Concluding Remarks ........................ References .................................

279 280 288 294 308

VI.

315

32 I 322

I. INTRODUCTION

The biological function of the squid axon is to conduct action potentials. This article summarizes the present knowledge about the action potential and the ionic currents that generate the action potential. Most of our knowledge about the origin of the action potential comes from the work of Hodgkin and Huxley (1952a-d). The main aim of this article is to describe (1) the analysis of the ionic currents by Hodgkin and Huxley, (2) recent measurements of the Hodgkin-Huxley parameters and of a few additional parameters not contained in the original analysis, and (3) the reconstruction of the action potential and of various excitation phenomena from the Hodgkin-Huxley equations. As will be seen, the HodgkinHuxley analysis-with a few modifications and additions-even after 30 years provides a suitable basis for describing the macroscopic ionic currents in the giant nerve fiber of the squid. As pointed out by Hodgkin ( 1 9 5 the ~ main objective of the HodgkinHuxley equations was “to obtain an empirical description of the way in 279 Copyright 0 1984 hy Academic Press, Inc. All right> of reproduction in any form re\erved. ISBN 0-12-153322-0

280

H. MEVES

which the permeability depends on time and membrane potential.” However, in formulating the equations Hodgkin and Huxley had in mind a simple physical picture that attributes the changes in ionic permeability to the movement of charged particles (gating particles) within the membrane (see Section V l l ) . In recent years, it has become possible to measure the charge movement associated with the gating system. The results of these gating current measurements are not consistent with the simple “Hodgkin-Huxley model” (see Section VII). The problems arising from this discrepancy are discussed in a review (French and Horn, 1983) and in other article5 in this book: the present article is limited to a description of the action potential and the macroscopic ionic currents. II. ACTION POTENTIAL

Action potentials of the squid giant axon, recorded under different experimental conditions, are illustrated in Figs. 1-3 and 5. An axon that is taken out of the animal produces an action potential with an after-hyperpolarization or underswing following the spike (Fig. IS). When the axon is left in its natural position in the animal, the resting potential is more negative and the after-hyperpolarization is missing (Fig. 1A; see also Moore and Cole, 1960). Presumably, in the latter case the leakage conductance is smaller and the resting potential closer to the K equilibrium potential. The action potential of a squid axon of 500 p m diameter propagates at

mV

A

04

mV

rnsec

B

0.4 msec

FIG. I . Action potentials recorded with a capillary microelectrode from an axon in its natural position in the animal (A) and from an isolated axon (B). Temperature. 8.S”C in A , 12.5”C in B. (From Hodgkin and Keynes. published in Hodgkin, 19.58).

HODGKIN-HUXLEY: THIRTY YEARS AFTER

281

room temperature at a velocity of about 20 m/sec (Fig. 2A). Between 5 and 20°C the Qloof velocity is 1.7-1.8 (Chapman, 1967; Easton and Swenberg, 1975). Larger Qlovalues are observed at temperatures between - 19 and -5°C (achieved by lowering the freezing point by means of glycerol) (Kukita, 1982). The conduction velocity is nearly proportional to the for axons between 32 and 520 pm in square root of the diameter ( u diameter according to Burrows et al., 1965). The square-root dependence of conduction velocity on diameter is predicted by cable theory, provided that (1) the extracellular resistance is small compared with the intracellular resistance and (2) axon diameter is the only factor that varies with fiber size (Hodgkin, 1954; Jack rt ul., 1975). Lowering the intracellular resis-

-

FIG.2. Effect of a low-impedance axial wire on the conduction velocity of the action potential. (A) Action potentials recorded by two capillary microelectrodes inserted at points 16 mm apart; conduction velocity, determined from the time between the peaks, is 21.5 m/sec. (B) Introducing a low impedance wire longitudinally into the axon increases the conduction velocity several hundred times, so that the two action potentials coincide; the wire is a platinized Pt wire, about 20 mm long, and the tips of the microelectrodes are 2 mm from each end of the wire. Axon diameter is about 500 p m : temperature, 20°C. (From del Castillo and Moore, 1959.)

282

H. MEVES

tance by introducing a low-impedance metal wire into the axoplasm leads to a dramatic increase in conduction velocity (Fig. 2B). The propagated action potential enters the region with the wire at one end and traverses it almost without delay: the region with the wire is "space clamped"; i.e., at any instant the potential is the same at every point. The conspicuous foot (slow initial rise time) of the action potential in Fig. 2B is due to the fact that the current generated by the action potential approaching the region with the wire discharges simultaneously the whole membrane over the 20-mm-longwire. Figure 3 compares the action potential outside the wire region (uppermost record) and the action potential in the wire region recorded 650 pm from the end of the wire (lowermost record). The latter shows again a pronounced foot and reaches its peak with a considerable delay. It propagates back toward and beyond the end of the wire, resulting in a double peaked action potential in the region near the tip of the axial wire. The second peak is the reflected action potential. Lowering the internal resistance by means of an axial wire is equivalent to an increase in axon diameter; the experiment of Fig. 3 may help to explain the changes in the shape of the action potential that occur during antidromic invasion of a spinal motoneuron (Ramon el al., 1976). The potential at the peak of the action potential (active membrane potential) is dependent on the external Na concentration or more strictly Na activity (Hodgkin and Katz, 1949a). As shown in Fig. 4A, the changes in active membrane potential that result from small increases or decreases of external Na agree approximately with those expected for an Na electrode. This supports the idea that the membrane (which is mainly permeable to K ions in the resting state) becomes selectively permeable to Na ions during the rising phase of the action potential. From the Goldman (1943) equation

Hodgkin and Katz calculated a permeability ratio PK : PNa: PCIof 1:0.04:0.45 for the resting membrane and a permeability ratio P K : P N: Pa ~ of 1 : 20 : 0.45 for the active membrane. The action potential remains unchanged when the bulk of the axoplasm is removed and the interior of the fiber is perfused with an artificial salt solution (Fig. 5). Two different perfusion techniques are in use: the roller technique (Baker ef d., 1961, 1962a) and the Tasaki technique (Oikawa et al., 1961). In the former, the axoplasm is squeezed out with a small roller. In the latter, the bulk of the axoplasm is removed by inserting one or two glass pipettes longitudinally into the axon and forcing perfusion fluid through the pipettes (for details see Tasaki et al., 1962; Adelman and Gilbert, 1964; Fishman. 1970; Bezanilla and Armstrong, 1972).

HODGKIN-HUXLEY: THIRTY YEARS AFTER

1

283

i

. .... ..

FIG.3. Changes in shape of an action potential propagating (from top to bottom) into the region with the internal wire. The internal wire, introduced from the bottom end of the axon, is shown as a thick black bar, and the vertical line corresponds to the axon membrane. The action potential was recorded by an intracellular electrode, introduced from the top end, at various points indicated by arrows. The uppermost record was obtained about 15 mm from the axial wire. (From Ramdn et nl., 1976.)

T

2

m

T.

1 Y 0 )

+

a

1

1 0 m

+

1

1

z

+

d

1

o

1

o

1

1

1

1

c

rn 0

Y 0)

I

I

I

( A U ) piuaiod

lOUJOlU(

1

1

a

h

I

1

1

a

HODGKIN-HUXLEY: THIRTY YEARS AFTER

285

The perfused axon technique is in wide use because it allows changing the intraaxonal fluid in a simple way and measuring the resulting changes in resting potential, action potential, and membrane currents. The first experiments were concerned with the effects of varying the internal Na and K activities on the resting and active membrane potential (Baker et al., 1962b). Different internal Na and K activities were obtained by mixing isotonic solutions of Na2S04and K?S04in appropriate ratios. As shown in Fig. 4B, with isotonic K2S04as perfusion fluid a resting potential (0)of -65 mV and an overshoot ( 0 )of 40 mV were observed. A 1 : 1 mixture of isotonic K2S04 and Na2S04 ([K], = [NaIi = 0.22 M ) produced a slight decrease of the resting potential and a clear reduction of the overshoot. Further lowering of [KIi definitely reduced the resting potential and rendered the fiber inexcitable. The measurements could be fitted by Eq. (1) with PNa:P K = 7 for the active membrane, PNa: PK = 0.08 for high values of the resting potential, and PNa: P K = 0.03 for low values of the resting potential and for the underswing (neglecting the contribution of CI ions). Numerous studies on perfused axons showed that action potentials can be obtained with Na-free external media containing certain univalent organic cations or certain divalent inorganic cations. Non-sodium action potentials are more easily observed in perfused axons than in intact axons because, by choosing an appropriate perfusion fluid, the outward currents (which tend to cancel the relatively small inward current carried by the Na substitute) can be greatly reduced. Action potentials of perfused axons in Na-free seawater with 300-430 mM guanidinium, hydrazinium, hydroxylammonium, or ammonium have been described by Tasaki and Singer (1966), Watanabe et al. (1967), and Binstock and Lecar (1969). These action potentials are blocked by small concentrations of tetrodoFIG.4. Effect of external and internal Na on the action potential. (A) Average change in active membrane potential plotted against l~g([Na],,,,/[Na],,,~~,~~). Dashed line was calculated from

where [Nalleilwater is the Na concentration of the normal seawater (455 m M ) and [Na],,,, is the Na concentration of the test solution (made by diluting seawater with isotonic dextrose or by adding solid NaCl to seawater). Figures next to the open circles give the number of experiments. (From Hodgkin and Katz, 1949a.) (B) Effect of replacing K by Na in perfusion fluid on active membrane potential ( 0 ) .resting potential (0). and potential at bottom of underswing ( X ). Abscissa: activity (moles/liter) of K and Na in perfusion fluid; K and Na activities were altered by mixing isotonic K2S04and Na2S04.External solution: artificial seawater with "a], = 0.32 and [K], = 0.0068 mol/liter. The curves were drawn from Eq. (I), neglecting the contribution of CI ions, with PNa/PK = 7 for upper curve, PNa/PK = 0.08 for middle curve, and PNaIPK= 0.03 for lower curve. (From Baker et al., 1962b.)

286

H. MEVES

0

1

2

3

4

5

6

7

8

mec

FIG. 5 . Action potentials recorded with an internal electrode from a perfused axon (internal solution 500 mM K2S04)(A) and from an intact axon (B). Temperature, 16°C in A , 18°C in B. (From Baker er a / . , 1961.)

toxin (TTX), as is the normal Na action potential, suggesting that the univalent organic cations move through the channels normally used by Na ions. Calcium action potentials were studied by Tasaki et af. (1966, 1967) and Watanabe et al. (1967) and later by Meves and Vogel (1973) and Inoue (1980). According to Tasaki and co-workers, Sr and Ba (but not Mg) are able to replace Ca. Figure 6 shows examples of Ca action potentials on different time bases. Calcium action potentials are characterized by their long duration and slow rising phase; the maximum rate of rise in record B of Fig. 6 is 2 V/sec, i.e., much smaller than the maximum rate of 400 V/sec

287

HODGKIN-HUXLEY: THIRTY YEARS AFTER C

20 sec

D

OUT :

7 +40 0 mV

U 100 msec

U 100 msec

-40

E 100 mM CaCI2

100rnMCaCI2

+ 1 )rM TTX

LOmV

FIG. 6. Calcium action potentials of perfused axons, elicited by current pulses through an internal wire. (A-C) External solution was 100 mM CaCI,; internal solution, 25 mM CsF; isotonicity was maintained with sucrose; records on slow and fast time base show responses to sub- and suprathreshold pulses. Temperature was 16°C. (From Meves and Vogel, 1973). (D and E) External solution is indicated above records; internal solution was 20 m M N a F + 10 mM sodium phosphate; isotonicity was maintained with glycerol; records show responses to sub- and suprathreshold pulses. Note that the action potential persists in the presence of 1 p M tetrodotoxin (TTX). Temperature, 19°C. (From Inoue, 1980.)

for a normal Na action potential. Calcium ions can move through Na channels and through K channels, at least under certain experimental conditions. In the experiment of Fig. 6, A-C, the axon was perfused with 25 m M CsF and the K channels were blocked by the internal Cs (cf. Chandler and Meves, 1965; Bezanilla and Armstrong, 1972). The action potential was solely generated by Ca ions moving through the Na channels and was completely abolished by external TTX. In the experiment of Fig. 6, D and E, by contrast, the internal electrolyte was 20 m M NaF and 10 m M sodium phosphate. Part of the Ca inward flux occurred through the K channels, and the action potential persisted in the presence of external TTX; the TTX-insensitive Ca action potential, however, was

288

H. MEVES

blocked by internal tetraethylammonium. This interpretation of the Ca action potentials is supported by voltage-clamp experiments, which will be described in Section V1. 111.

VOLTAGE-CLAMP CURRENTS

The first records of voltage-clamp currents of the squid axon were published by Cole (1949) and Hodgkin et al. (1949). A detailed description and a mathematical analysis of the currents were presented by Hodgkin et al. (1952) and Hodgkin and Huxley (1952a-d). Further important details are found in the papers by Cole and Moore (1960a) and Moore and Cole (1 963). Examples of voltage-clamp currents recorded by Moore and Cole (1963) and Hodgkin r t al. (1952) are shown in Figs. 7 and 8. In Fig. 7A we see an early transient current that is inward for small depolarizations and reverses sign around 50 mV and a delayed current that is outward. Current voltage curves for the early current and the steady-state current are illustrated in Fig. 7B. The curve for the early current shows a region of steep negative slope; the positive slopes of the two curves are both equal to a conductance of about 100 mS/cm’ and are constant over a large potential range. The reversal potential, V,,,, of the early current is more accurately determined from records on fast time base (Fig. 8). The value of V,,, depends on the external Na concentration and changes according to the Nernst relation for the Na equilibrium potential (Hodgkin and Huxley, 1952a; Moore and Adelman, 1961). The early current measured at a potential V decreases with decreasing external Na concentration according to Eq. (la). I‘

-=

I

(LNal~ilNaI,)exp[(V,,, - V)F/RTl - 1 cxp[(V,,, - V)F/RT] - 1

where I and I ’ are the currents at the concentrations “a], and “a];, respectively (Hodgkin and Huxley, 1952a; Adelman and Taylor, 1964). This equation was derived by Hodgkin and Huxley (1952a) on the assumption that “the chance that any individual ion will cross the membrane in a specified interval of time is independent of the other ions” and is, therefore, called the independence principle (see also Section VI). Using the voltage-clamp technique on perfused axons in seawater with radioactive 22Na, Atwater et al. (1969) applied depolarizing pulses of varying amplitude and measured (1) the influx of ZZNaby determining the radioactivity in samples of the perfusion fluid and (2) the ionic flux during

A

B

10 1 mA/cm*

5

0

-5

0

I

2

3

4

msec

FIG.7. Voltage-clamp currents from a squid axon. (A) Five records superimposed. Ordinate: membrane current density, outward current upward. Abscissa: time after change of membrane potential. Membrane potential during voltage-clamp pulse is given on the right of each curve. The membrane was held at -70 mV between pulses. Temperature. 10°C. (From Moore and Cole, 1963.) ( B ) Current voltage curves from an experiment similar to that in (A). Peak of early current (Iha)and steady-state current (IK)are plotted against the pulse potential; the resting potential is indicated by an arrow. (From Cole and Moore, 1960a.)

290

H. MEVES 4kc/;ec

143 mV I30

I17 I04

91

-

I mA/cmZ

FIG.8. Reversal of the early current. Five voltage-clamp currents associated with pulses of different amplitude (see figures on the right) on fast time base. Resting potential was about -62 mV; temperature, 3.5"C. Early current is inward for pulses 91 and 104 rnV and outward for pulses 130 and 143 m V ; reversal of early current is at 117 mV, i.e., at a potential of 55 mV. (From Hodgkin et ul., 1952.)

the early permeability change by integrating the early current; the ratio tracer flux : electrical flux was close to unity (average 0.92 in 17 measurements on six different axons). These findings provide conclusive evidence that the early current is a Na current. The delayed outward current can be identified as K current for three reasons:

I . In tracer experiments on Sepia axons the quantity of K leaving the axon in a given time is equivalent to the electric charge carried by the outward current in the same time (Hodgkin and Huxley, 1953). 2. The reversal potential of the instantaneous current following a long depolarizing pulse varies with [K], although not as steeply as the theoretical K potential (Hodgkin and Huxley, 1952b). The discrepancy is probably due to accumulation of K ions in a space outside the membrane (Frankenhaeuser and Hodgkin, 1956; Adelman et al., 1973). No such discrepancy is found in Myxicola axons, where periaxonal K accumulation is less pronounced (Binstock and Goldman, 1971). 3. The delayed outward current is seen in perfused axons with K as the only internal cation and becomes smaller when the internal KCI is partially replaced by NaCI, RbCI, CsC1, choline chloride, OF sucrose (Chandler and Meves, 1965; Chandler et al., 1965). [Partial substitution of internal K by other cations, in particular by Na or Cs, reduces the delayed outward current below the values expected from the independence principle; this inhibitory action of internal Na and Cs has been studied in detail by Bezanilla and Armstrong (1972), who showed it to be strongly voltage dependent .]

291

HODGKIN-HUXLEY: THIRTY YEARS AFTER A

mV

.m

-10 -40

~525mA/cm2

I msec

I 0

I I

1

1

I

3

3

4

FIG.9. Examples for pure Na currents recorded from a perfused axon (A) and from an intact axon (B). (A) Seawater with 325 m M Na and 50 mM Ca; perfusion fluid 380 m M K , 10 mM Na, and 20 m M TEA; holding potential -70 mV; temperature 2°C. (From Bezanilla and Armstrong, 1977.) (B) Seawater with 470 mM Na, 1 1 mM Ca, and 10 mM 4-AP; holding potential, -70 mV; 50 msec prepulse to -90 mV; temperature, 8. IT. (Kimura and Meves, unpublished observation; from Meveb, 1978.) In (A) and (B) TTX-insensitive currents are subtracted. The numbers indicate the potential during the clamp pulse.

For a quantitative analysis, the early current (INa)and the delayed have to be separated. To separate the two currents, outward current (IK) Hodgkin and Huxley (1952a) took voltage-clamp records in seawater with full Na and in seawater with reduced Na. Simpler methods for separating the two currents became available through the discovery that certain substances selectively block either ZNa or IK.The pufferfish poison tetrodotoxin (TTX) in nanornolar concentrations blocks INawithout affecting IK (Moore and Narahashi, 1967). Intracellular tetraethylammonium (TEA) in millirnolar concentrations blocks the K outward current (but not the K inward current recorded in high-K seawater) (Armstrong and Binstock, 1965); the early current remains normal although often reduced in size (see Armstrong and Binstock, 1965; Lipicky et af., 1978). Blockage of the K channels can also be achieved by internal Cs (see above) or by 4arninopyridine (4-AP) (Yeh et af., 1976; Meves and Pichon, 1977), which acts both externally and internally. Examples of pure Na currents recorded from perfused or intact axons are shown in Figs. 9 and 10. Blockage of the K channels was achieved with 20 mM TEA internally (Fig. 9A), 10 mM 4-AP externally (Fig. 9B), or 300 mM Cs internally (Fig. 10). In Fig. 9A and B the records are corrected for the TTX-insensitive currents; for this purpose the pulse program was repeated after adding a high concentration of TTX to the seawater, and the currents remaining in TTX (capacitative current, leak-

292

H. MEVES

u 0

0.5 msec

0

0 5 msec

FIG.10. Sodium currents associated with depolarizing pulses of different duration at tw o different holding potentials. Conditions: Perfused axon in seawater with 107.5 mM Na and 10 mM Ca; perfusion fluid 300 mM Cs; holding potential Vh is indicated above records; pulse potential 20 mV; temperature, 7°C. The turning-on of INilcould be fitted by Eq. ( I1) with Sr = 0, T, = 202 psec for Vh = -60 m v , and 6r = 30 psec, T , = 210 psec for Vh = - 100 m v . The time constants for the tail currents are 101 psec at Vh = -60 mV and 36 psec at Vh = - 100 mV. (From Keynes and Rojas, 1976.)

age current, unblocked rcst of K current) were subtracted from the original records by means of a computer. Before describing details of the records in Figs. 9 and 10, an important technical point, the series resistance artifact. must be discussed. Between the internal and external potential electrode is the membrane and, in addition, a small series resistance R , made up of two components-an external component representing the resistance of the dense layer of connective tissue and Schwann cells around the axon, and an internal component arising in the axoplasm or perfusion fluid between the internal electrode and the membrane. The potential that is seen and held constant by the clamp amplifier is the membrane potential plus the voltage drop of the membrane current across R,. Hodgkin et al. (1952), Moore and Cole (1963), Binstock et al. (19751, and Keynes and Rojas (1976) have described methods for measuring R,. R, is usually between 5 and 10 R cm? but rises to much higher values in axons perfused with low-salt solutions (e.g., 27 R cm2 for an axon perfused with a 60 mM salt solution; see Keynes and Rojas, 1976). With R, = 5 s1 cm2 and peak ZNa = -5 mA/cm*, the voltage would be in error by -25 mV. Consequently, the negative resistance branch of the INa(V)curve in Fig. 7B and all voltage-dependent rate constants of the Na system would be considerably shifted to the left, resulting in a distortion of the time course of ZNa (Taylor et al., 1960; Chandler and Meves, 1970c; Ramon et al., 1975). By contrast, the voltage error produced by an outward current is positive; i.e., the series resistance makes the I,(V) curve in Fig. 7B less steep (Taylor et al., 1960; Ram6n et al., 1975). To compensate for the voltage error, or at least for part of it, a potential proportional to the membrane current (obtained from

HODGKIN-HUXLEY: THIRTY YEARS AFTER

293

5 msec

FIG.I I .

Example for pure K currents recorded from an intact axon in seawater with 10

M tetrodotoxin (TTX). Holding potential is -60 mV; pulse potential as indicated; temperature, 9°C. (From Armstrong, 1971.)

a potentiometer whose setting depends on the size of R,) is added to the command voltage pulse (compensated feedback of Hodgkin et al., 1952). This method was used for obtaining the records shown in Figs. 9 and 10. An additional precaution was taken in Fig. 10: was drastically decreased in order to reduce ZNa and thereby the voltage drop ZN,R,. Successful series resistance compensation is indicated by the flat time course of INaat -40 mV in Fig. 9 A and B and by the very fast tail currents in Fig. 10. Figures 9 and 10 demonstrate three impotant properties of the Na current: 1. The current through the Na channel is inward for small and mediumsized depolarizations ( V < VreV) but becomes outward for large pulses ( V > Vrev) (Fig. 9). 2. The Na current is transient: it increases (or activates) fast, reaches a peak and then decreases (or inactivates) more slowly (Fig. 9); on fast sweep speed, the time course of activation is seen to be sigmoidal (Fig. 10). 3. Open Na channels close rapidly at the end of short depolarizing pulses (Fig. lo), a process much faster than the inactivation during a maintained depolarization (Fig. 9).

An example of pure K currents recorded from an intact axon is shown in Fig. 1 1 . The Na currents are blocked by lo-’ M TTX in the seawater. The K outward currents do not inactivate during the 13-msec pulses used in Fig. I I , but often exhibit a slow droop for longer pulse durations. The

H. MEVES

294

depolarizing pulses are followed by slowly decaying inward current tails carried by K ions that have accumulated in the periaxonal space during the depolarizing pulse (Frankenhaeuser and Hodgkin, 1956; Adelman et al., 1973). The size of the inward current tails increases with increasing pulse duration. Slow inactivation of the K outward currents (time constant 10-33 sec at 9 T ) occurs during depolarization produced by high-K seawater (Ehrenstein and Gilbert, 1966). IV. HODGKIN-HUXLEY ANALYSIS

Hodgkin and Huxley (1952d) described the voltage-clamp currents by a set of mathematical equations. The basic ideas underlying the HodgkinHuxley analysis were that (1) the membrane currents are due to ions moving down their electrochemical gradients given by the driving forces (V - VNa),( V - V K ) , ( V - Vkeak); (2) INaand ZK are independent from each other and have a different voltage and time dependence, suggesting that Na and K ions move through different channels (dual-channel hypothesis); (3) activation (m)and inactivation ( h ) of the Na conductance are parallel processes that start at the beginning of a depolarizing pulse and follow exponential time courses; and (4) the voltage dependence of g N a and gKmay be due to electrically charged molecules inside the membrane that alter their position according to the electric field, thereby opening or closing a pathway for the ions. Point 1 above is supported by the finding that ZNa and ZK reverse sign at potentials close to the Na potential VNa or K potential VK, respectively (see Section 111). The dual-channel hypothesis (point 2) provides a simple explanation for the different kinetics of ZNa and IK and their selective blockage by drugs; it received strong support from the Pronase experiment of Armstrong et al. (1973), which shows that, after complete destruction of Na inactivation by Pronase, the K conductance still increases in a normal fashion. Points 3 and 4 will be discussed below and in the article by Armstrong and Matteson in this volume on sodium channels and gating. The ionic currents are given by the equations INa [K

[leak

(v - V N a ) g N a = (v - v N a ) g N a m 3 k = (v - V K ) g K = (v - v K ) g K n 4 =

=

(v -

Vleak)gleak

(2) (3) (4)

where rn, h , and n are variables describing Na activation, Na inactivation, and K activation, respectively. The three variables are potential depen-

295

HODGKIN-HUXLEY: THIRTY YEARS AFTER

dent and change exponentially with time. The third power of rn and the fourth power of n were chosen to fit the sigmoidal turning-on of ZNa and the delayed rise of ZK (see Figs. 10 and 11). The values of the three variables are given by the relations dmldt

=

a,(l - rn) - &rn

dhldt

=

trhfl

h ) - Phh

(6)

dnldt

=

an(l - n ) - Pnn

(7)

-

(5)

where a and p are rate constants that depend on membrane potential V and temperature. The steady-state values are

nee

+ Pn)

= an/(an

(10)

and the corresponding time constants are T = I l ( a + p). The steady-state values of n , rn, and h and their time constants T , , T,, and Th are plotted against membrane potential in Fig. 12. The values are from Hodgkin and Huxley (1952d), but they are replotted on an absolute membrane potential scale. Both n, and rn, reach unity at small positive potentials while h, becomes zero. The slope of the n,(V) curve is smaller rn, msec I nm

L -50

0

T,, msec

I

f h , msec

I

,h

50

V, millivolts

aK = 36 mmho/cm2 V, = -77 m V

V, millivolts

V, millivolts cNa

VN,

120 mmho/cm2 50 m V

0.3 mmho/cm2

gt

V,

=

-54.4 m V

FIG. 12. Steady-state values of the variables n , rn, and h and their time constants T , , T,, and T~ as a function of membrane potential. Values from Hodgkin and Huxley (1952d) are replotted on an absolute membrane potential scale, taking the resting potential (rp) in the experiments of Hodgkin and Huxley as -65 mV. Numerical values of gK, gNa,g, and the equilibrium potentials are at the bottom of the figure. (From Cole, 1968.)

296

H. MEVES

than that of the m,( V ) curve. The curves for the three time constants are bell-shaped with a maximum near the resting potential. The HodgkinHuxley equations for the ratc constants a! and /3, rewritten for absolute membrane potential, can be found in Table 6 of Adrian d.(1970). From Eqs. (2) and (3) the Na current and the K current during a depolarizing clamp pulse are obtained as (If

INa = Iha[I - exp(-t/T,)13[h, - ( h , - I) exp(-t/q,)] I K

=

Ik,[l - e x p ( - t / ~ , ) ] ~

(I I) (12)

where

IK1.

=

(V -

VK),QK~

(14)

Equations ( I 1) and (12) arc valid fora holding potential sufficicntly negative so that m, and h,, the activation and inactivation variables at zero time, are zero and unity, respectively. If the depolarizing pulse leads to complete inactivation (h, = O), the third term in Eq. (1 1) is reduced to exp(-tl Th). By fitting Eqs. ( I I ) and (12) to records of f N a and IK,numerical values for Iha, T,, h,, and Th or for I K Z and T , are obtained. The fitting procedure has been described by Hodgkin and Huxley (1952d); the same procedure has been used by Keynes and Rojas (1976). Nowadays, the fit is best done by computer (e.g., Kimura and Meves, 1979). In nodes of Ranvier a proper fit of Na currents recorded at different pulse potentials requires different exponents of m (Neumcke et a l . , 1976). In squid axons, by contrast, a perfect fit with a constant exponent 3 can be obtained (Keynes and Kimura, 1983). However, at strongly negative holding potentials it is necessary to introduce an initial delay a t , i.e., to use ( t - st) instead o f t in Eq. (11). The initial delay 6t is negligible at a holding potential Vh = -60 mV,but it is noticeable at Vh = - 100 mV (see Fig. 10) (Keynes and Rojas, 1976; Keynes and Kimura, 1983; Taylor and Bezanilla, 1983). There is uncertainty whether it is correct to use gNafor perfused axons. The use of gNais justified for intact axons in normal seawater because the instantaneous f N i , ( V ) curve is linear (Hodgkin and Huxley, 1952b).This is not the case for axons perfused with 300 m M KCI or 275 mM K F (Chandler and Meves, 1965; Begenisich and Cahalan, 1980b). Therefore, instead of gNa,the permeability coefficient P N adefined by the constant-field equation (Goldman, 1943) has been used by some authors (e.g., Kimura and Meves, 1979). Sometimes m, calculated by using gNadecreases at positive potentials; as shown in Fig. 8 of Keynes and Rojas (1976), the decrease disappears when PNais used instead of gNa.

HODGKIN-HUXLEY: THIRTY YEARS AFTER

297

Figures 13-15 show measurements of m, and 7 ,as a function of membrane potential [see also m i and peak g N a curve in Fig. 5 of Oxford (1981) and in Fig. 2 of Keynes et al. (1982)l. The new measurements are, in general, consistent with the original measurements of Hodgkin and Huxley. Values for mcc[as obtained from in Eqs. ( 1 1) and (13)] are plotted against potential V in Fig. 13. The points have been fitted with Eq. (15).

where Vmidis the potential at which m, = 0.5 and k is a reciprocal measure of the maximum slope in terms of how many millivolts are required to change m, e-fold. The dashed curve (Vmid= -32 mV, k = 12.5 mV) and the continuous curve (Vmid= -34 mV, k = 18 mV) are fits of all points and of points between -70 and -20 mV, respectively. Both values for Vmidare close t o Vmid = -60 + 25 = -35 mV in Fig. 8 of Hodgkin and Huxley

mV

FIG. 13. Voltage dependence of m, for perfused axonb in seawater with 107.5 m M Na. Different symbols denote different axons. The perfusion fluid contains 300 or 350 mM Cs. Holding potential is between -50 and -100 mV; temperature, 5-7°C. Continuous and dashed curves were calculated from Eq. (15) with Vmld = -34 mV, k = 18 mV for the continuous curve, and Vmld= -32 mV, k = 12.5 mV for the dashed curve. (From Keynes and Ro-jas, 1976.)

298

H. MEVES

/

299

HODGKIN-HUXLEY: THIRTY YEARS AFTER

600

400 u Ln m a

200

I

-60

I -40

I

-20

I

I

I

0

20

40

mV

FIG.15. Time constant 7, at different potentials determined by fitting the rising phase of

ZNa(0,T,,, on) or from Na current tails at the end of depolarizing pulses (0, T, on). The graph

also shows values of the time constant T,, I for the fast relaxation of the gating current measured on the same axon (0).Internal solution: 350 mM CsF + 400 mM sucrose; external solution: artificial seawater with 86 mM NaCl for the measurement of Na currents, Na-free Tris seawater with 2 p M tetrodotoxin for the measurement of gating currents. Holding potential was -100 mV; temperature, -0.1"C. (From Keynes and Kimura, 1983.)

pulses (see Fig. 10) by means of the equation 7, = 3 T N a l a l . Keynes and Kimura (1983) have measured T , at the same potentials with both methods and found no significant difference (Fig. 15). The difference seen in Fig. 15 at positive pulse potentials was not observed in other experiments. Measurements of T N by ~Oxford ~ (1981) ~ ~ confirm ~ Fig. 14 in showing that 7, values calculated from the tail current time constants are appreciably larger than the Hodgkin-Huxley values. In contrast to Fig. 14, however, Oxford was unable to obtain a continuous 7, curve over the whole potential range; in order to draw a continuous curve, T N tail ~ (rather than T, = 37Natal~) had to be plotted. In contrast to Fig. 15, 7, values derived were clearly from tail currents by means of the equation T,,, = 37Natall larger than those derived from the turning-on of fNa at the same voltage. The reason for the discrepancy between the results of Keynes and coworkers (see Figs. 14 and 15) and those of Oxford (1981) is not clear. A situation opposite to that described by Oxford seems to exist in molluscan neurons where T , derived from Ca inward current tails is much smaller than T, obtained from the turning-on of fc,,at the same potential (Byerly and Hagiwara, 1982). An inequality 7, > T, on (Oxford, 1981) or T , off < 7, on (Byerly and Hagiwara, 1982) would suggest that the pathway of the reaction resting e conducting is different for the forward and the backward direction.

300

H. MEVES

In recent years, the question whether N a activation ( m ) and Na inactivation ( h ) are independent or coupled has received much attention (for a review, see French and Horn, 1983). Hodgkin and Huxley (1952d) discussed the two different mathematical approaches corresponding to these two alternatives: “First, we might assume that the sodium conductance is determined by a variable which obeys a second order differential equation. Secondly, we might suppose that it is determined by two variables, each of which obeys a first-order equation.” Hodgkin and Huxley chose the second alternative (see point 3 at the beginning of this section), “since it was simpler to apply to the experimental results.” Hoyt (1963, 1968) solved the second-order differcntial equation corresponding to the first alternative and presented an analysis of the data of axon 17 of Hodgkin and Huxley (1952d) in terms of a coupled activation-inactivation model. Goldman and Schauf (1972) (see also Goldman, 1975, 1976) obtained experimental evidence for coupling in Myxicoltr giant axons. Gating current experiments demonstrated that the asymmetrical charge displacement in the squid axon membrane can be immobilized by conditioning depolarization (Bezanilla and Armstrong, 1974; Armstrong and Bezanilla, 1977; Meves and Vogel, 1977), an observation usually taken as evidence in favor of activation-inactivation coupling. On the other hand, an analysis of the gating statistics of single Na channels of rat myotubes showed that N a channels need not open before they inactivate (Horn et al., 1981). Even if a strictly sequential reaction scheme (in which the active state must be passed through before the inactive state can be reached) is ruled out, there still remains the possibility of partial coupling between m and h ; an example would be a scheme in which the reaction resting + active consists of several steps and the first few of these steps are necessary for inactivation to start. In the discussion as to whether rn and h are independent (Hodgkin and Huxley) or coupled (Hoyt, 1963, 1968; Goldman, 1976; Armstrong and Bezanilla, 1977),the question of the exact time course of the development of Na inactivation has played an important role. Chandler et al. (1965) showed on perfused squid axons that, upon depolarization, inactivation develops along an exponential time course without an obvious initial delay. However, a sigmoid time course with a pronounced initial delay (suggestive of activation-inactivation coupling) was found by Goldman and Schauf (1972) on Myxicola axons and by Bezanilla and Armstrong (1977) on perfused squid axons. The problem was reinvestigated by Gillespie and Meves (1980a) on squid axons, Bean (1981) on crayfish giant axons, Kniffki ut al. (1981) on nodes of Ranvier, and Goldman and Keny o n (1982) on Myxiculu axons. An example is shown in Fig. 16. The pulse program (see inset) consists of a conditioning pulse to -43 mV of

301

HODGKIN-HUXLEY: THIRTY YEARS AFTER

-400

-100

0

1

I

I

A r (msec) 2

3 1

I

r

'

. :

N

5

-40

\

-

\

-8

-10

.-

-63-

-

-63 mV

At

0

10

20

30

Ar (msec)

FIG. 16. Time course of development of Na inactivation at V = -43 mV in a perfused axon (external solution: seawater with 157 mM NaCI; internal solution: 218 mM K F + 54 m M TEA). Pulse program (see inset) consists of a conditioning pulse to -43 mV of variable duration Ar and a constant test pulse to -8 mV. The difference INapear - I , (the latter determined with Ar = 45 msec) is plotted on a logarithmic scale against At (lower scale) and fitted by a straight line with 7h= 10.55msec. Points for A? = 0-3 msec replotted on expanded scale (upper scale) and again fitted by a straight line with T,,= 10.55 msec. Temperature, 3.8"C. (From Gillespie and Meves, 1980a.)

variabIe duration Al, immediately followed by a test pulse to -8 mV [similar to the pulse program in Fig. 1 of Hodgkin and Huxley (1952c)l. Test pulse current ZNa peak (minus ZNa peak measured with At = 45 msec) is plotted on a logarithmic scale against At. The points are well fitted by a straight line with Th = 10.55 msec. Points for At = 0-3 msec replotted on an expanded scale (upper scale) can again be fitted by a straight line of the same slope. Figure 16 shows no obvious initial delay in the development of inactiva-

302

H. MEVES

tion; however, the small slope of the lines makes it difficult to exclude with certainty a deviation of the earliest points from the straight lines. When the pulse program of Fig. 16 is used with stronger conditioning pulses, there is an (apparent) initial delay in the development of inactivation. As shown by Gillespie and Meves (1980a), the delay disappears when the conditioning pulse and the test pulse are separated by a 3-8 msec interval, thus ensuring that m (which rises during thc strong conditioning pulse) has decreased again before the beginning of the test pulse (see also Kniffki et al., 1981). The upper limit for a true initial delay in the development of inactivation was found to be 50-100 psec (at temperatures between 0 and 13°C and at membrane potentials between -40 and 15 mV). Similarly, KniMti ct al. (1981) concluded for the node of Ranvier that a delay, if it exists at all, is less than 100 psec. A clear delay in the development of inactivation, however, is seen in crayfish axons (Bean, 1981) and in Myxicola axons (Goldman and Kenyon, 1982); it amounts to 0.1-0.2 msec at 3.S"C and to 0.1-0.8 msec at S'C, respectively. Possiblc reasons for the discrepancy of the experimental findings have been discussed by Bean (1981) and Goldman and Kenyon (1982). Numerical values for T h , the time constant of Na inactivation, in intact axons at 10°C can be obtained from the equations Th = ( a h

+ Ph)-'

These equations arc derived from Eqs. (23) and (24) of Hodgkin and Huxley (1952d), assuming a resting potential of -62 mV in their experiments and a Qloof 3 for both a h and P h . An example of measurements of Th is shown in Fig. 17A. The data are from 21 axons perfused with 270 mM KF. The time constant q, was determined either from the decay of Zwa during a depolarizing pulse or by means of a double-pulse program with variable duration of the first pulse or variable duration of the interval between the two pulses; in squid axons Th values determined from the decay of I,, are identical to those measured with the double-pulse method (Bezanilla and Armstrong, 1977; Gillespie and Meves, 1980a). The Th values in Fig. 17A can be compared with the Hodgkin-Huxley values for intact axons: at -90 mV the values are sirnilar (1.92 msec from Fig. 17A for 8.1-10.4"C, 2.32 msec from Hodgkin and Huxley for lO"C), whereas at -50 and 0 mV the values in Fig. 17A (9.02 and 1.37 msec) are considerably larger than the Hodgkin-Huxley values

303

HODGKIN-HUXLEY: THIRTY YEARS AFTER

A msec

5.0 4.0 3.0 2.0 1.0 0-

.. .. I

L

1

-100

-50

0

50

rnV

B

0.9 0.8 0.7 0.6 0.5 0.4 0.3 0.2 0.1 0

-

-

-100

0

-50

50 rnV

FIG. 17. Collected values of T~ and h, vs the membrane potential for perfused axons (external solution: seawater with 118-157 mM NaCl; internal solution: 270 mM K F + 60 mM TEA) at 8.1-10.4"C. (A) Th values from 21 axons. (B) 17, values for I0 of the 21 axons. For pulse programs, see text. Curves for Th = ( a h t ph) I and 11, = (Yh/((Yh + ph) are calculated from ah = 0.11 exp(-

Ph = O.S/[exp((From Gillespie and Meves, 1981.)

-)V t18 62

F) $)I t 0.66 exp( -

304

H. MEVES

(3.70 and 0.69 msec) (see also Fig. 7 of Meves, 1978). Recent measurements of Th on intact axons (Fig. 7 of Meves, 1978; Fig. 8 of Gillespie and Meves, 1981) are in reasonable agreement with the Hodgkin-Huxley values. It seems that inactivation develops more slowly in perfused axons than in intact axons. Replacing the K in the perfusion fluid by Na further increases 'rh (Chandler and Meves, 1970~;Gillespie and Meves, 1981). On 10 ofthe 21 axons of Fig. 17A, the k ( V ) curve was measured, using 30-msec prepulses to different potentials, immediately followed by a constant test pulse to about 0 mV, Figure 17B shows normalized test pulse current plotted against prcpulse potential. The experimentally determined relation between h, = & h / ( a h + ,&) and membrane potential V can be fitted by the same equations for a h ( V )and &(V) that were used to fit the Th values in Fig. 17A. The potential at which h, = 0.5 is Vh = -50 mV in Fig. 17B; i.e., at the normal resting potential of a squid axon (-60 or -65 mV) more than 50% of the Na channels can be activated. As shown by Wang et al. (1972) and Kimura and Meves (1979), lowering the temperature shifts the A,( V ) curve to more negative potentials (V, = -47 mV at 15°C v h = -55 mV at OT), a phenomcnon responsible for the stronger blocking effect of certain drugs and insecticides on the action potential at low temperatures (Wang et a l . , 1972). For V > -50 mV, most of the points in Fig. 17B are above the computed curve, indicating that h, is larger than calculated from the equations for (Yh(v) and P h ( V ) , That h, does not reach zero for strong depolarizations is shown by records labeled +80 mV and +70 mV in Figs. 9A and 9B, respectively, from a perfused and an intact axon. Incomplete Na inactivation in intact axons was also demonstrated by Shoukimas and French (1980). The phenorncnon is most pronounced in axons perfused with NaF solution. As shown in Fig. 18, the Na currents inactivate only partially during depolarizing pulscs of 3Y-msec duration. The ratio INi,at 39 mscc to peak INaincreases with increasing pulse potential, indicating that inactivation becomes less and less complete at positive potentials; i.e., 17, not only fails to reach 7,ero at small negative potentials, but actually increases again in the positive potential range. Chandler and Meves (1970a,b) explained this finding by a second open state of the Na channel as indicated by the sequence

where h , represents the normal open state originally described by Hodgkin and Huxley, x is the inactivated state, and hz is a second open state, whose population increases with increasing potential. As pointed out by Chandler and Meves (1970b), an alternative assumption would be

305

HODGKIN-HUXLEY: THIRTY YEARS AFTER

B

A

4-

I

n

mV

3 -

/

+I

/d

84 74 64 54

2U 2 \

3-

44

34 24 10.5

0-

-1

c

0.5 -22.5

-

! "

I

( 1 1 1 1

0

10

20 30 msec

40

FIG.18. Voltage-clamp currents from an axon perfused with 300 mM NaF and immersed in seawater with 470 m M NaCl. (A) Superimposed membrane currents associated with pulses from the holding potential (-76 mV) to the potentials indicated. (B) peak current (0) and current at 39 msec (8)as a function of pulse potential; the dashed line indicates leakage current. Temperature, 0°C. (From Chandler and Meves, 1970a.)

that peak conductance gNam3h,and maintained conductance gNam3h2reflect two sepurute populations of Na channels that differ in their kinetics. The second hypothesis is favored by Matteson and Armstrong (1982), who found the size of the maintained currents to be little affected by cooling, whereas peak Na currents decrease substantially as the temperature is lowered. Two measurements of the steady-state Na inactivation curve of the squid axon are shown in Fig. 19 (see also Figs. 7 and 8 of Bezanilla and Armstrong, 1977). In Fig. 19A the axon was perfused with NaF solutions of different pH; for pHi = 5.5-7.2 the h,(V) curves have flat minima at about 0 mV and rise as the potential is made more positive; for pHi = 9.810.5, the minima continue increasing but the rises are steeper. As Fig. 19B shows, the noninactivating component h2 is smaller in a CsF-perfused axon than in NaF-perfused axons (see also Oxford and Yeh, 1979). It can be completely and reversibly abolished by 1 mM octanol in seawater (A in Fig. 19B). Externally applied scorpion venom, on the other hand, increases the noninactivating component h2 (see Fig. 6 of Gillespie and Meves, 1980b). In addition to fast Na inactivation described by the h parameter of Hodgkin and Huxley, a voltage-dependent attenuation of Na conductance

306

H. MEVES

10-

- 9 0 mV

_r" tr'On'r' 0 Dmsrr

08.

-Po=104-

3 Q2

,(L-

-80 , -60 O L

-40

-20

O

20

Marr&ane Potential (mVI

FIG.19. Steady-state Na inactivation curve of the perfused squid axon under different experimental conditions. (A) Axon perfused with NaF solutions (containing 200 mM Na) of different pH,; for pulse program, see inset; ordinate: normalized test pulse current; abscissa: membrane potential during 42 msec conditioning pulse. (From Carbone er ul., 1981.) (B) Axon perfused with 275 m M CsF, pH 7.3; pulse program similar to that in A; ordinate and abscissa as in A: measurement before (e),during (A),and after (wash; 0) external application of I mM octanol; tempenittire, I0"C. (Prom Oxford and Swenson. 1979.)

has been reported that exhibits much slower kinetics ("slow inactivation''). That restoration of excitability after a strong and long-lasting depolarimtion is much slower than expected from h kinetics as first pointed out by Narahashi (1964), then by Adelman and Palti (1969). Quantitative measurements of the slow inactivation parameter s and its time constant 7, were pcrformed by Chandler and Mcves (1970d) and Rudy (1978). As shown in Fig. 20, the voltage dependence of the steady-state value s, and of the time constant 7, is similar to the voltage dependence of h, and 7 h (Fig. 17). but the s l ( V )curve is less steep than the h , ( V ) curve and T , is three orders of magnitude larger than Th. Internal perfusion with Pronase or N-bromoacetamide removes fast inactivation, but slow inactivation remains (Rudy, 1978; Oxford rt al., 1978), suggesting that fast and slow inactivation are different processes. Whereas Na activation and Na inactivation have been studied by a great number of authors, relatively few quantitative investigations of the K current exist. Recent values of T,,, determined by fitting the rising phase of ZK with Eq. (12), are shown in Fig. 21 and can be seen to agree with the original Hodgkin-Huxley values. The quantitative analysis of fK is more difficult than that ofZNa.Figure 3 of Hodgkin and Huxley (1952d) shows a small but consistent deviation between computed gK(t)curves and experimental points; better agreement might have been obtained by raising n to a higher power than 4. A hyperpolarking prepulse further

307

HODGKIN-HUXLEY: THIRTY YEARS AFTER

A

FIG.20. Voltage dependence of the steady-state value s, (A) and of the time constant T , (B) of slow Na inactivation. Measurements on axons perfused with CsF (Rudy, 1978) (A)or with NaF (Chandler and Meves, 1970d) (A).In B, points at V < -50 mV represent time constants of recovery from a strong and long-lasting depolarization, whereas points at V > -50 mV are time constants of development of slow inactivation. Curves were drawn from the equations s, = 1/{1 7, = I/('&

a,

=

+ exp[(V -

Vh)/kl}

+ ps)

0.000016 exp(-0.097 V) (sec) (sec)

ps = 0.125 exp(0.035 V)

Measurements were at 5°C (A) or at 16-16.5"C, but were adapted to S"C assuming a Qloof 3

(A).(From Rudy, 1978.)

delays the onset of ZK (Frankenhaeuser and Hodgkin, 1957; Cole and Moore, 1960b; Keynes and Kimura, 1980). To fit the pronounced delay seen with strong hyperpolarizing prepulses the exponent of n has to be increased to 25 or, alternatively, a long initial delay has to be introduced (Cole and Moore, 1960b). In squid axons, the hyperpolarization-induced delay corresponds to a parallel shift of I K ( t )along the time axis; i.e., the currents are superimposable by translocation along the time axis (Moore and Young, 1981; see, however, Clay and Shlesinger, 1982). A further complication in the analysis of IKarises from the accumulation of K ions in the periaxonal space (Frankenhaeuser and Hodgkin, 1956; Adelman et al., 1973). The resulting continual decrease of the driving force (V - VK)leads, if not taken into account, to an underestimate of

308

H. MEVES

FIG.21. Voltagc dependence of 7, for a perfused axon in seawater with 215 mM NaCl and 3 X I0 M tetrodotoxin. Perfusion fluid consists 0 1 150 mM KF and 780 mM sucrose: Holding potential, -70 mV; temperature, 7°C. Symbols 0 and x denote two series of measurements on the same axon. Curve for T, = (a, + fin) calculated from Eqs. (12) and (13) of Hodgkin and Huxley (1952d), taking the resting potential in their experiments as -60 mV. (From Haydon and Kimurd, 1981.)

gK and, in addition, distorts the time course of gK(see Fig. 8 of Adclman cf d., 1973). Adelman ct nl. (1973) have recalculated a , ( V ) and p,,(V)

taking into account K accumulation in the periaxonal space; their values for a,, and Pnare smaller than the original values of Hodgkin and Huxley (1952d) (see also Adclman and FitzHugh, 1975). V.

RECONSTRUCTION OF THE ELECTRICAL BEHAVIOR OF THE SQUID AXON FROM THE HODGKIN-HUXLEY EQUATIONS

From the equations for the ionic currents, the action potential of the squid axon can be reconstructed. Mathematically, the situation is simplest for a membrane action potential obtained under space clamp (see Section 11). In this situation there is no flow of longitudinal current because at any instant the potential is the same at every point. The equation for a membrane action potential is Z, = C,JV/dt

+ Z N ~+ ZK + Ilenk

(16)

with C , taken as 1 pF/cm*. For a propagated action potential, Eq. (16) is

309

HODGKIN-HUXLEY: THIRTY YEARS AFTER

replaced by a partial differential equation with distance along the axon (x) and time ( t ) as independent variables:

(a12R)62V16x2= C,dV/dt

+ IN,+ I K + IIeak

(17)

where a is the radius of the axon and R the specific resistance of the axoplasm. In the case of an action potential propagating at constant velocity 6, Eq. (17) is simplified into the ordinary differential equation

(a12RO2)d2Vldt2= C,dVIdt -I-I N a

4-

+

(18)

Equations (16)-( 18) have all been used to compute action potentials and to reproduce some of the classical excitation phenomena. Figure 22 (lower part) shows propagated action potentials, computed from Eq. (18), at three different temperatures. Amplitude, form, duration, and velocity agree remarkably well with the experimental records in Fig. 22 (upper part). Cooley and Dodge (1966) and Moore et al. (1975) have calculated propagating action potentials from the partial differential Eq. (17). Joyner et al. (1978) extended the method to simulate propagation in excitable cells with a wide range and complexity of morphologies; they calculated the effect of (abrupt or gradual) changes in diameter or of axon branching on the size, rate of rise, and conduction velocity of an action potential (see also Joyner et al., 1980). The equations allow calculation of the conduction velocity as a function of temperature (Huxley, 1959) and channel density (Hodgkin, 1975; Adrian, 1975). As shown by Chapman (1967) and Easton and Swenberg (1979, the experimentally observed effect of temperature on conduction velocity is in reasonable agreement with the effect computed by Huxley (1959). With increasing density of Na channels, N , the maximum Na conductance per unit area, gNa,becomes bigger and the conduction velocity, 8, grows correspondingly (Fig. 23A). However, as pointed out by Hodgkin (1975), increasing N leads to the displacement of more gating charges and thus to an increase of the membrane capacity CM.The latter is given by CM=

co + NCC,

where Co is the capacity of the membrane without channels and C:, is the capacity associated with each gate. In Fig. 23B, 6 has been calculated from Hodgkin-Huxley equations modified by the addition of a term I g a representing the Na gating current; the latter has been assumed to be proportional to dmldt (an assumption no longer considered correct; see, for instance, Keynes and Kimura, 1983) and given by the equation ICa= 3Nz’e(dmldt) where N is the number of channels per unit area of membrane, z’ is the apparent valence of the gating particles, and e is the charge on an electron. The increase of CMdue to the presence of gating current

310

H. MEVES

L

80 -

100 rnV

6040-

20-

-

.L

0 1

I

l

l

l

l

l

r

r

l

~

l

i

,

l

l

l

l

,

r

J

FIG.22. Propagated action potential at three different temperatures. Top: Experimental records with time marks I msec apart (Fig. 3 of Hodgkin and Katz, 194Yb). Bottom: compuled from Eq. (18); the rate constants CY and /3 have been assumed to rise with temperature with a Qloof 3 whereas the temperature dependence of gNa,&, and R has been neglected. (From Huxley, 1959.)

reduces H for RNil = 120 mS cm-? and 18.5"C from 18.5 rn/sec (Fig. 23A) to 14.7 m/sec (Fig. 238). Since increasing gNnincreases 8 and increasing CM decreases 8, the curve relating 8 to N or 2~~has a flat maximum for gNa= 100-300 mS cm-2 (Fig. 23B). These values of K N correspond ~ to N = 417-1250 pm-' (with a single-channel conductance yNil= 2.4 pS as

HODGKIN-HUXLEY: THIRTY YEARS AFTER

31 1

FIG.23. Calculated conduction velocity, 0, plotted against Na conductance, gNa;with single-channel conductance yNa = 2.4 pS the value ENa = 120 mS c m Zcorresponds to a channel density of 500 p m 2 .(A) On the assumption that no gating currents are associated with the opening of the Na channels. (B) On the assumption that before each channel opens, three gating particles with an effective valence z' = 2.5 move in the membrane. 0, 18.5"C; 0 , 6.3"C. Temperature is assumed to change only the rate constants for the conductance variables (Qlo = 3). The axon radius is 238 p m , and the internal resistivity is 35.4 Rcm. (From Adrian, 1975.)

assumed by Adrian, 1975). Using yNa = 4 pS obtained from the noise measurements of Conti et af. (1973, the formulation of Adrian (1975) would give N = 250-750 pm-*. The density of Na channels determined from the binding of tritiated tetrodotoxin to squid giant axons is N = 553 t 119 pm-* (Levinson and Meves, 1975). The value of N obtained from noise measurements is 330 pm-2 (Conti et a/., 1975). Both values of N lie in the range where 6, has its maximum. The strength-duration curve was calculated from Eq. (17) by Cooley and Dodge (1966) and can be compared with experimental results (see, for instance, Guttman, 1966). Calculation and experiment show that the height of the membrane action potential is a continuous graded function of the stimulus strength, not an all-or-none phenomenon; the effect of stimulus strength is most pronounced at 35°C but also visible at 15°C (Cole et al., 1970). Equations (16)-( 18) predict repetitive firing in response to a constant current of sufficient intensity [see Jack et al. (1975), pp. 331-342, for summary of literature]. Except on rare occasions, squid axons in normal seawater do not exhibit maintained repetitive firing in response to a steady current. However, they fire repetitively with an infinite train when bathed in low Ca or low Mg seawater. In the calculated as well as in the

312

H. MEVES

experimental axon the frequency of firing depends on the strength of the steady stimulating current [see Fig. 11.14 of Jack et al. (197s) and Fig. S of Fohlmeister et al. (1980)l. With the original Hodgkin-Huxley parameters, the calculation predicts repetitive firing at frequencies between SO and 125 Hz, but slight modifications of the parameters (e.g., reducing the leakage conductance or shifting the voltage dependence of Na inactivation or K activation) extend thc firing rate to values as low as 10 H r . Guttman et al. (1980b) reported that repetitive firing of a squid axon can be annihilated by a brief depolarizing or hyperpolarizing pulse ofthc proper magnitude applied at the proper phase (Fig. 24A). As shown by Best (1979), the same result is obtained from the Hodgkin-Huxley cquations (Fig. 24B). For other nonannihilating perturbations, repetitive firing resumes with unaltered frequency but with phase resetting; this is seen both A

FIG.24. Annihilation of repetitive firing of squid axon by a brief depolarizing perturbation. (A) Space-clamped squid axon in Mg-free seawater with 10 mM Ca at 22.3"C. Axon was stimulated with a barely suprathreshold step of current of 30-msec duration; stimulus (above) and response (below). In the lower record a brief depolarizing pulse of 0.15 msec duration was superimposed upon the 30-msec step of current and completely annihilated the repetitive firing. Calibration: 2 pA/div, 100 mV/div, 2 mseddiv. (From Guttman er u / . , 1980h.) (€3) Calculated repetitive action potentials. Ordinate scale givcs deviation from resting potential. A constant current of 8.75 pA/cm' was present continuously. After an instantaneous voltage perturbation of - 5 mV (applied at *), repetitive activity is replaced by subthreshold oscillations. (From Best, 1979.)

HODGKIN-HUXLEY: THIRTY YEARS AFTER

31 3

in the experiments (Guttman e f al., 1980b) and in the calculations (Best, 1979). Computations on the Hodgkin-Huxley axon for sinusoidally varying stimulating parameters were done by Nemoto et al. (1975) and Holden (1976). The Hodgkin-Huxley model behaves in the same way as the real axon (Hirsch, 1965; Guttman et al., 1980a): the model and the axon respond to cyclic inputs of certain frequencies by a series of action potentials that, in the steady state, are phase locked to the driving function, i.e., are generated at fixed phases with respect to the driving cycle. Fohlmeister et al. (1980) compared Bode plots (i.e., amplitude and phase of impulse density plotted against frequency of the perturbing sine wave) obtained from squid axons with Bode plots calculated from the original Hodgkin-Huxley model and from the model modified to include the effects of K accumulation in the periaxonal space (Adelman and FitzHugh, 1975). Their results suggest the existence of an as yet uncharacterized slow process in the axon, since they see a low-frequency peak in the Bode plots of the experimental axons that is not present in the computer simulations. Sinusoidal waves of small amplitude and varying frequency are used to determine the membrane admittance Y = W 8 V as a function of frequency. To measure the admittance under voltage clamp, DeFelice et al. (1981) superimposed a small (1 mV) sinusoidal wave 6V on the holding potential, varied the frequency of this wave continuously between 20 and 200 Hz over a period of 30 sec, and recorded the small resulting change in membrane current, 61. The admittance goes through a minimum at each holding potential at some particular frequency (see Fig. 2 of DeFelice et al., 1981). Poussart et al. (1977) and Fishman et al. (1979) used a broadband signal (pseudorandom noise) source and discrete Fourier transform techniques; this enabled them to measure the admittance over the whole frequency range from 10 to 1000 Hz within a quarter of a second or less. Figure 25A shows as an example the magnitude and phase of the admittance of a squid axon at 10 different membrane potentials. The last curve (at -54 mV) is similar to the first curve (at -59 mV), indicating that the preparation has suffered little if any deterioration. The measurements were done in the presence of tetrodotoxin (TTX), and the admittance is, therefore, solely determined by g K , g L , and the membrane capacitance. For all potentials the admittance reaches a minimum at a certain frequency (“antiresonance”). With increasing depolarization and increasing K conductance, g K , the admittance curve is shifted to larger absolute values and the “antiresonance” becomes less pronounced. The increase of the admittance at frequencies larger than the “antiresonance” frequency reflects the decrease of the capacitative resistance with increasing

31 4

H. MEVES

A

B Y, II

0. 809 0. 4 0"

0"

I

10

I

1

..I

100 HZ

T TX / I

,

, , I

1000

30" ,

I

,

,

, , )

,

-4c

~

A

f

1

10

U

100

1000

U

1

Hz

FIG. 25. Comparison of the complex admittance measured in a squid axon (A) with a calculation from the Hodgkin-Huxley model (B) for g,, = 0 [artificial seawater with 1 p M tetrodotoxin (TTX)].(A) Measurements on an unperfused axon; the magnitude plots (above) and phase plots (below) were obtained at the indicated membrane potentials (sequence -59, -54, -49, -44, etc., to + 16 mV and back to -54 mV); acquisition time was 250 msec for lower curves and 100 msec for upper three curves at large depolarizations. (B) Calculations from the linearized Hodgkin-Huxley model. Temperature was 10°C in (A) and 63°C in (B). (From Poussart el id., 1977.)

frequency (see Fig. 10 of Fishman et nl., 1981). The phase function ( LY) first goes negative with increasing frequency and then makes a sharp transition very near the antiresonance frequency to a positive phase. The experimental admittance is compared with the theoretical admittance predicted from the Hodgkin-Huxley equations (see Fig. 25B). Since 6V and 61 are small, the equations can be used in their linearized form. The details of the linearization are given by Hodgkin and Huxley (1952d), Chandler er al. (1962), Mauro ~t al. (1970), and DeFelice (1981). As shown in Fig. 2SB, an "antiresonance" is also predicted from the linearized Hodgkin-Huxley equations for V > -70 mV. The phase functions also behave similarly, including the range of membrane potential over which the phase curves cross one another. When the Na channels are not blocked, the antiresonance is sharper and the experimental results are

HODGKIN-HUXLEY: THIRTY YEARS AFTER

31 5

more complex, but again the linearized Hodgkin-Huxley equations are capable of producing and accounting for most of the significant features that have been observed (Poussart et af., 1977; Fishman et al., 1979). VI. MOLECULAR PROPERTIES OF THE Na CHANNEL: PERMEABILITY RATIOS AND BINDING CONSTANTS

More information about the properties of the Na channels has been obtained by studying the ionic selectivity of the channels and the permeation of ions through the open channels. In axons perfused with a Na-free K salt solution, an early outward current, clearly separated from the delayed outward current, is observed at large depolarizations (Chandler and Meves, 1965; Rojas and Atwater, 1967; Cahalan and Begenisich, 1976) (see Fig. 26A). The early K outward current is blocked by 10 nM tetrodotoxin. Its time course corresponds closely to the time course of the Na permeability measured by repolarizing the membrane to the resting potential at different times after the beginning of the large depolarizing pulse. The steady-state inactivation of the early outward current resembles the steady-state inactivation of the Na inward current. These experiments provide fairly conclusive evidence that the K ions that flow through the membrane during the early outward current are moving through the same channels that are normally used by Na ions. A quantitative estimate of the relative permeability of the Na channel for Na and K can be obtained from the equation

where V , is the equilibrium potential at which the early current changes its sign, [Na], and [KIiare the activities of external Na and internal K , and PNaand PKare the Na and K permeabilities of the active membrane. Chandler and Meves (1965) found PNa/PK = 12. Similar ratios were obtained by Atwater et af. (1969), Binstock and Lecar (1969), and Cahalan and Begenisich (1976). Thus, the Na channel is about 12 times more permeable to Na than to K. Closer investigation showed, however, that the ratio PNa/PK depends markedly on the internal K concentration. Chandler and Meves (1965) observed that diluting the internal K solution with isotonic sucrose appeared to reduce the selectivity of the channel. The phenomenon was studied in detail by Cahalan and Begenisich (1976). Figure 26 illustrates voltage-clamp records and current-voltage curves before and after dilu-

31 6

H. MEVES

A

B

/O

/"

0.5 msec FIG.26. Thc effect of internal K on the selectivity of the Na channels in a perfused axon. (A) Currents through the Na channels with artificial seawater outside and 275 mM KF inside (top) or SO mM KF inside (bottom). lsotonicity of the internal solution is maintained with sucrose; delayed outward currents are blocked by internal tetraethylammonium. Depolarizations in steps of 10 mV from -70 to 110 mV (top) and from -70 to 120 m V (bottom). Temperature, 5°C. (B) Peak current-voltage relations for the traces in (A); 0 , 2 7 5 mM K F inside; U, 50 m M K F inside; the potassium activities, reversal potentials, and permeahility ratios PNaIPK (abbreviated by PIP) calculated from Eq. (19) are shown in the inset. 'Ihe arrow at 97 mV indicates the predicted reversal potential for 50 mM K F on the assumption that there was no change in selectivity upon diluting. (From Cahalan and Begenisich, 1976.)

tion of the internal KF solution by isotonic sucrose. The reversal potential is 61 mV with 275 mM KF and 75 mV with 50 m M KF, corresponding to PNaIPK= 8.3 and 3.2, respectively. Without a change in YNaIPK the reversal potential in 50 mM KF should have been 97 mV. As shown by Cahalan and Begenisich (1976), the decrease in PNaIPKis due to the decrease in [K],, not to the decrease in ionic strength or [CI],. In Fig. 27 average values of the permeability ratio PNaIPK are plotted against the internal K activity [K];. The ratio PNa/PK decreases linearly with decreasing [K],; possible explanations for this phenomenon are discussed below. To determine the relative permeability of the Na channel for other inorganic cations (Li, Rb, Cs) or for organic cations (NH4, guanidinium), the reversal potential of the early current was measured after replacing the external Na or the internal K by the respective ion. From these measurements the results shown in Table I were obtained. The finding

317

HODGKIN-HUXLEY: THIRTY YEARS AFTER

plotted against internal potassium activity [K],. Filled FIG. 27. Permeability ratio PNa/PK circles are averages from several experiments. The solid line is the linear regression line given by the equation P,,/PK = 0.029[K],+ 2.21

with [K], as millimolar. (From Cahalan and Begenisich, 1976.)

that PLiis nearly equal to PNa(see also Moore et al., 1966) is consistent with the old observation that Li ions can fully replace Na ions in the action potential mechanism. For the inorganic cations the permeabilities follow the sequence of the ionic radii; the organic cation NH4,however, is more permeant than expected from its ionic radius. The relative permeaTABLE I RELATIVEPERMEABILITY OF THE Na CHANNEL FOR VARIOUS INORGANIC A N D ORGANIC CATIONSO Permeability Reference

Li

Na

Guanidinium

Chandler and Meves (1965) Binstock and Lecar (1969) Cahalan and Begenisich (1976) Ionic radius (A)b

1.1 -

1

-

1

-

1

112.1

-

0.60 0.95

-

NH4

K

Rb

Cs

- 1/12 1/40 1/61 - 113.7 113.3 li8.l 1122 1159 1.48 1.33 1.48 1.69

a All values (except that for Li) were measured with infernal application of the ions (concentrations 200-550 mM); the value for Li was obtained with external application; the value 1/3.7for NH, was measured with infernal or external application. Crystal radii are quoted from Robinson and Stokes (1959).

31 8

H. MEVES

bility of the Na channel for Cs ions is very small, but not zero; a tetrodotoxin-sensitive Cs outward current in CsF-perfused fibers has been demonstrated by Meves and Vogel (1973), Armstrong and Bezanilla (1974), and Keynes and Rojas (1974). Interestingly, the permeability for guanidinium ions is higher than that for NH4 ions, while the reverse is true for the node of Ranvier (Hille, 1971). As the relative permeability for K depends on [K],(see above), the relative permeabilities for guanidinium, NH4, Rb, and Cs depend on the internal concentration of these ions (Cahalan and Begenisich, 1976; Begenisich and Cahalan, 1980a). The values given in Table 1 were determ i n d with internal concentrations between 200 and 550 mM. With smaller internal concentrations, larger values are obtained. For instance, guanidinium at an internal concentration of 225 mM is about one-half as permeant as Na (P = PNa/2.1 in Table I), whereas at 50 mM it becomes as permeant as Na (P = pN,/1.01). By contrast, experiments with external NH4 and internal Na show that PNdIf"H4 does not vary greatly with [NH41, or "a];; neither does PNnIPK or P N a / P ~vary ~ 4 with "a],. The observations of Cahalan and Begenisich (1976) and Begenisich and Cahalan (1980a) could be explained by saying that ion permeability ratios are functions of ionic concentration for some ions on the internal side of the membrane. Lowering the internal concentration of these ions may alter the structure of the Na channel so that the selectivity is altered. Alternatively, the observations can be accounted for by a three-barrier, two-site energy profile model. As shown by Begenisich and Cahalan (1980a), such a model produces the observed effects without explicitly making permeabilities functions of concentration. The model is attractive because it also predicts the experimentally observed saturation of outward current through N a channels that occurs at high concentrations of the internal permeant ion (Begenisich and Cahalan, 1980b). Figure 28A shows the peak outward current as a function of internal ion activity for perfused axons with Na, NH4, or K solutions of varying concentration inside and Na-free Tris seawater outside. The outward currents were measured at a pulse potential of 50 mV. The currents seem to saturate as the ionic activity is increased. The continuous lines in Fig. 28A are fits of rectangular hyperbolas to the data representing the binding of ions to a single saturable binding site. The equilibrium constants, K,, obtained by this fitting procedure are 623, 268, and 161 mM for Na, NH4, and K , respectively. Similar results were obtained at other pulse potentials; with increasing pulse potential, the K , values for Na and K decreased. The data of Fig. 28A are shown again in Fig. 28B. The continuous lines here represent the predictions of the three-barrier, twosite model used by Begenisich and Cahalan (1980a) for describing Na-

31 9

HODGKIN-HUXLEY: THIRTY YEARS AFTER A

7-

, ,

8 7-

I

6-

Independence

Na

:

5I

0

ts

3-

3-

Ql (r

V m = 50 mV Rectangular hvperbola

300 Internal activity (mM)

100

200

2-site model

400

100

1

1

I

200

300

400

Internal activity ImMI

FIG.28. Peak outward current through Na channels as a function of internal ion activity at a membrane potential of 50 mV. Perfused axon in Na-free Tris seawater. A,internal Na; H, internal NH,; 0 , internal K. For activities below 200 mM the ionic strength was kept constant by means of tetrarnethylammonium. Current has been normalized to unity at an activity of about 35 m M (+). Mean values TSE of mean are plotted except where the symbols are larger than these limits. (A) Continuous lines are fits of rectangular hyperbolas to the data according to the formula y = ymax[x/(x+ K,)] with K , = 623, 268, 161 m M for Na, NH4, and K , respectively. The dashed line is the expectation from the independence principle. (B) Same data as in A but the continuous lines are computations from a threebarrier, two-site model. (From Begenisich and Cahalan, 1980b.)

channel reversal potentials. The model provides a good fit to the experimental data. It seems likely that permeant ions at the inner side of the membrane bind to a saturable binding site, the innermost well in the model. Saturation of the ionic current is not consistent with the independence principle (see Section 111). As indicated by the dashed line in Fig. 28A, a linear relation between current and ion activity would be expected if ions moved independently of each other. Deviations from the independence principle in perfused axons have also been described by Chandler and Meves (1965) and Binstock and Lecar (1969): after replacing the internal K by Na or NH4, the current through the Na channel was smaller than predicted by the independence principle. In intact axons injected with radioactive 22Na, the efflux of sodium associated with voltage-clamp pulses is reduced to 50% by Na-free seawater; i.e., it does not obey the independence principle (Landowne, 1977). By contrast, the effect of changes in external Na on the peak inward current of squid axons is correctly described by the independence principle (see Section 111).

320

H. MEVES

The Na channel is also to some extent permeable to Ca. In intact axons injected with aequorin, the early phase of the light response to depolarization is blocked by TTX and seems to reflect Ca entering through the Na was estichannels (Baker Pt a / , , 1971); from these experiments PCa/PNa mated as 1/100; i.e., the Na channel is even less permeable to Ca than to Cs. A larger permeability for Ca was found in axons perf‘used with diluted salt solutions. Meves and Vogel (1973) (see also Meves, 1975) measured the small TTX-sensitive Ca inward and Cs outward currents in perfused axons with 100 mM CaC12 + sucrose outside and 25 mM CsF + sucrose inside. From the reversal potential and the ion activities, the permeability ratio PCa/Pc,was obtained as 1.6111 by means of a modified constant-field equation; i.e., the permeability for Ca is about I .6 times larger than for Cs. From the changes in reversal potential produced by small amounts of Na in the internal or external solution the ratio PCJPN, was found to be l/lO-l/7, indicating a substantial loss of selectivity under perfusion with diluted salt solutions. As shown by Inoue (1980) on perfused axons, Ca ions can move even through the K channels, provided they are not blocked by internal Cs. Figure 29 shows current voltage curves from two axons with 100 mM A

B

T T X treated

LI A”,

I N 3ONa OUT 1 0 0 ca

(pA lcm21

-100

Pic;. 29. Effect ofexternal tetrodotoxin (TTX) and internal tetraethylammonium (‘I‘EA)on the current voltage curve of perfused axons with 100 mM CaCI2 + glycerol outside and 20 m M NaF, 10 m M sodium phosphate + glycerol inside. (A) Current voltage curve before (0) and after ( 0 )addition or 3 p M TTX to the external solution. (B)Current voltage curves in the presence of 1 p M T T X in the external solution; 0, without TEA; A. with 20 niM’IEA in internal solution; 0 , with 20 mM TEA in external solution. Temperature, 19°C. (From Inoue, 1980.)

HODGKIN-HUXLEY: THIRTY YEARS AFTER

321

CaClz + glycerol outside and 20 mM NaF + 10 mM sodium phosphate + glycerol inside. External TTX reduces (but does not block) the Ca inward current and, in addition, shifts its reversal potential to more positive internal potential (Fig. 29A). The Ca inward current remaining in the presence of TTX is totally blocked by internal (but not by external) TEA (Fig. 29B). The Ca inward current through the K channels can give rise to a TTX-insensitive Ca spike (see Fig. 6E). It is clear that a situation where the reversal potential of the K channel is 44 or 27 mV positive (see Fig. 29A and B) is not a normal situation. In addition to the Ca currents through Na and K channels, a Ca current through separate voltage-dependent Ca channels has been described (DiPolo et al., 1983); it may contribute to the late TTX-insensitive Ca entry observed in aequorin-injected squid axons (Baker et al., 1971). The cation selectivity of the resting membrane of the squid axon was determined by observing the relationship between resting potential and the external concentration of T1, K, Rb, Cs, Na, and Li (Hagiwara et al., 1972). The relative permeabilities are TI, 1.8; K, 1.0; Rb, 0.72; Cs, 0.16; Na < 0.08; Li < 0.08. The same sequence (except for TI) has been found by Baker et al. (1962b). The finding that TI is twice as permeant as K agrees with the observations of Hille (1973) on nodes of Ranvier. Presumably, the diameter of the K channels in squid axons is similar to that estimated for the node of Ranvier (3.0-3.3 A according to Hille, 1973). VII.

CONCLUDING REMARKS

The essential conclusion is that even after 30 years the Hodgkin-Huxley equations with a few relatively minor modifications provide a correct quantitative description of the Na and K currents of the squid giant axon. Modifications became necessary to include slow inactivation, incomplete inactivation, and delayed rise of the ionic currents after a hyperpolarizing prepulse. The equations allow us to reconstruct the membrane action potential, the propagated action potential, and the strength-duration curve, to predict repetitive firing and account for most of the significant features observed in impedance measurements. The equations are widely and successfully used to investigate and characterize the effects of toxins and drugs on the ionic currents. Hodgkin and Huxley (1952d) had a physical picture or model in mind when they formulated their equations (see also Hodgkin, 1958). “For the sodium channel we assumed that three simultaneous events, each of probability m, opened the channel to Na and that a single event of probability (I-h) blocked it. These events . . . may be thought of as the movement of three activating particles and of

322

H. MEVES

one blocking particle to a certain region of the membrane. The probability that there will be three activating particles and no blocking particle is therefore m’h. To account for the change in potassium conductance we assumed that a path for potassium was formed when four charged particles had moved to a certain region of the membrane under the influence of the electric field.”

Measurements of gating currents suggest that the simple “HodgkinHuxley model” does not give a correct description of the physical events. It was originally hoped that the mobile charges that generate the gating current could be identified with the three activating particles of the “Hodgkin-Huxley model” (Keynes and Rojas, 1974, 1976). Closer investigation showed, however, that this identification is not possible (Armstrong and Bezanilla, 1974; Meves, 1974; Keynes and Kimura, 1983); neither the voltage dependence of the charge movement nor the time constants of the on and off gating currents directly reflect the voltage and time dependence of the activation variable m. The evidence is summarized in a review by French and Horn (1983) and in subsequent articles of this volume. The task of the future will then be twofold: (1) to develop a physical model that accounts for the known properties of gating currents and ionic currents; (2) to reconstruct the time and voltage dependence of the ionic currents from the time and voltage dependence of the gating currents as Hodgkin and Huxley (1952d) have reconstructed the action potential from the ionic currents. ACKNOWLEDGMENTS The author is indebted to Dr. J. M. Simard for reading the manuscript and to the Deutsche Forschungsgemeinschaft SFB 38 for financial support. KEFERENCES Adelman, W. J., Jr., and FitzHugh, R. (197.5). Solutions of the Hodgkin-Huxley equations modified for potassium accumulation in a periaxonal space. Fed. Proc. Fed. A m . Soc. Exp. Biol. M, 1322-1329. Adelman, W. J., Jr., and Gilbert, D. L. (1964). Internally perfused squid axons studied under voltage clamp conditions. I. Method. J . Cell. Comp. Pliysiol. 64, 423-428. Adelman, W. J . , Jr., and Palti, Y. (1969). The effects of external potassium and long duration voltage conditioning on the amplitude of sodium currents in the giant axon of the squid, Loligo pealei. J . Gen. Physiol. 54, 589-606. Adelman, W. J . , Jr., and Taylor, R. E. (1964). Effects of replacement of external sodium chloride with sucrose on membrane currents of the squid giant axon. Biophys. J . 4, 451-463. Adelman, W. J., Jr., Palti, Y.,and Senft, J. P. (1973). Potassium ion accumulation in a periaxonal space and iis effect on the measurement of membrane potassium ion conductance. J. Membr. B i d . 13, 387-410. Adrian, R. H. (197.5). Conduction velocity and gating current in the squid giant axon. Proc.. R. SOC. London Ser. B 189, 81-86.

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Cahalan, M. D., and Begenisich, T. (1976). Sodium channel selectivity: Dependence on internal permeant ion concentration. J . Gen. Physiol. 68, I 11-125. Carbone, E., Testa, P. L., and Wanke, E. (19x1). Intracellular pH and ionic channels in the Loligo uulgoris giant axon. Biophys. J. 35, 393-413. Chandler, W. K., and Meves, H. (1965). Voltage clamp experiments on internally perfused giant axuns. 1.Physial. (Lundon) 180, 788-820. Chandler, W. K., and Meves, H. (1970a). Sodium and potassium currents in squid axons perfused with fluoride solutions. J. Physiol. (London) 211, 623-652. Chandler, W . K., and Meves, H. (1970b). Evidence for two types of sodium conductance in axons perfused with sodium fluoride solution. 1.Physiol. (London) 211, 653-678. Chandler, W. K., and Meves, H. (1970~).Kate constants associated with changes in sodium conductance in axuns perfused with sodium fluoride. J. Physiol. (London) 211, 679-705. Chandler, W. K., and Meves, H. (1970d). Slow changes in membrane permeability and longlasting action potentials in axons perfused with fluoride solutions. J . Physiol. (Lundon) 211, 707-728. Chandler, W. K., FitzHugh, R., and Cole, K. S . (1962). Theoretical stability properties of a space-clamped axon. Biophys. J . 2, 105-127. Chandler, W. K., Hodgkin, A. L., and Meves, H. (1965). The effect of changing the internal solution on sodium inactivation and related phenomena in giant axons. J . Physiol. (London) 180, 821-836. Chapman, R. A. (1967). Dependence on temperature of the conduction velocity of the action potential of the squid giant axon. Nnture ([,ondon) 213, 1143-1 144. Clay, J. R.,and Shlesinger, M. F. (1982). Delayed kinetics of squid axon potassium channels do not always superpose after time translation. Biophys. J . 37, 677-680. Cole, K. S. (1949). Dynamic electrical characteristics of the squid axon membrane. Arch. Sci. Physiul. 3, 253-258. Cole, K. S. (1968). “Membranes, Ions and Impulses.” Univ. of California Press, Berkeley. Cole, K. S . , and Moore, J. W. (1960a). Ionic current measurement in the squid giant axon membrane. 1. Grn. Physiol. 44, 123-167. Cole, K. S., and Moore, J. W. (1960b). Potassium ion current in the squid giant axon: Dynamic characteristic. Biophys. J . 1, 1-14. Cole, K . S.,Guttman, R., and Bezanilla, F. (1970). Nerve membrane excitation without threshold. Proc. N d . Acnd. Sci. U.S.A. 65, 884-891. Conti, F., DeFelice, L. J . , and Wanke, E.(197.5). Potassium and sodium ion current noise in the membrane of the squid giant axon. J . Physial. (London) 248, 45-82. Cooley, J. W., and Dodge, F. A , , Jr. (1966). Digital computer solutions for excitation and propagation of the nerve impulse. Biophys. J . 6, 583-SYY. DeFelice, L. J. (1981). “Introduction to Membrane Noise.” Plenum, New York, DeFelice, L. J., Adelman, W. J., Jr., Clapham, D. E., and Mauro, A. (1981). Second order admittance in squid axon. I n “The Biophysical Approach to Excitable Systems’’ (W. J. Adelman and D. E. Goldman, eds.), pp. 37-63. Plenum, New York. del Castillo, J . . and Moorc. J . W. (19.59). On increasing the velocity of a nerve impulse. J . Physiol. (Lnndon) 148, 665-670. DiPolo, R., Caputo, C., and Bezanilla, F. (1983). Voltage-dependent calcium channel in the squid axon. Proc. Nail. Acud. Sci. U.S.A. 80, 1743-1745. Easton, D.M., and Swenberg, C. E. (1975). Temperature and impulse velocity in giant axon of squid Loligo pecrlei. A m . J . Physiul, 229, 1249-1253. Ehrenstein, G., ilnd Gilbert, D. L. (1966). Slow changes in potassium permeability in squid giant axon. Biophys. J. 6, 553-566.

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Fishman, H. M. (1970). Direct and rapid description of the individual ionic currents of squid axon membrane by ramp potential control. Biophys. J. 10, 799-817. Fishman, H. M., Poussart, D., and Moore, L. E. (1979). Complex admittance of Na’ conduction in squid axon. J. Membr. Biol. 50, 43-63. Fishman, H. M., Moore, L. E., and Poussart, D. (1981). Squid axon K conduction: Admittance and noise during short- versus long-duration step clamps. In “The Biophysical Approach to Excitable Systems” (W. J. Adelman and D. E. Goldman, eds.), pp. 6595. Plenum, New York. Fohlmeister, J. F., Adelman, W. J., Jr., and Poppele, R. E. (1980). Excitation properties of the squid axon membrane and model systems with current stimulation. Statistical evaluation and comparison. Biophys. J. 30, 79-98. Frankenhaeuser, B., and Hodgkin, A. L. (1956). The after-effects of impulses in the giant nerve fibres of Loligo. J . Physiol. (London) 131, 341-376. Frankenhaeuser, B., and Hodgkin, A. L. (1957). The action of calcium on the electrical properties of squid axons. 3. Physiol. (London) 137, 218-244. French, R. J., and Horn, R. (1983). Sodium channel gating: Models, mimics, and modifiers. Annu. Rev. Biophys. Bioeng. 12, 319-356. Gillespie, J. I., and Meves, H. (1980a). The time course of sodium inactivation in squid giant axons. J . Physiol. (London)299, 289-307. Gillespie, J. I., and Meves, H. (1980b). The effect of scorpion venoms on the sodium currents of the squid giant axon. J . Physiol. (London) 308, 479-499. Gillespie, J. I., and Meves, H. (1981). The effect of external potassium on the removal of sodium inactivation in squid giant axons. J. Physiol. (London)315, 493-514. Goldman, D. E. (1943). Potential, impedance, and rectification in membranes. J . Gen. Physiol. 21, 37-60. Goldman, L. (1975). Quantitative description of the sodium conductance of the giant axon of Myxicola in terms of a generalized second-order variable. Biophys. J. 15, 119-136. Goldman, L. (1976). Kinetics of channel gating in excitable membranes. Q . Reu. Biophys. 9, 49 1-526. Goldman, L., and Kenyon, J. L. (1982). Delays in inactivation development and activation kinetics in Myxicola giant axons. 1. Gen. Physiol. 80, 83-102. Goldman, L . , and Schauf, C. L. (1972). Inactivation of the sodium current in Myxicola giant axons; evidence for coupling to the activation process. 1. Gen. Physiol. 59, 659-675. Guttman, R. (1966). Temperature characteristics of excitation in space-clamped squid axons. J. Gen. Physiol. 49, 1007-1018. Guttman, R., Feldman, L., and Jakobsson, E. (1980a). Frequency entrainment of squid axon membrane. J. Memhr. Biol. 56, 9-18. Guttman, R., Lewis, S . , and Rinzel, J. (1980b). Control of repetitive firing in squid axon membrane as a model for a neurone oscillator. J. Physiol. (London) 305, 377-395. Hagiwara, S . , Eaton, D. C., Stuart, A. E., and Rosenthal, N. P. (1972). Cation selectivity of the resting membrane of squid axon. J. Memhr. B i d . 9, 373-384. Haydon, D. A., and Kimura, J. E. (1981). Some effects of n-pentane on the sodium and potassium currents of the squid giant axon. J. Physiol. (London) 312, 57-70. Hille, B. (1971). The permeability of the sodium channel to organic cations in myelinated nerve. J. Gen. Physiol. 58, 599-619. Hille, B. (1973). Potassium channels in myelinated nerve. Selective permeability to small cations. J . Gen. Physiol. 61, 669-686. Hirsch, H. R. (1965). Squid giant axon: Repetitive responses to alternating current stimulation. Nature (London) 208, 1218-1219.

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Hodgkin, A. L. (1954). A note on conduction velocity. J . Physiol. (London) 125, 221-224. Hodgkin, A. L. (1958). Ionic movements and electrical activity in giant nerve fibres. Proc. R . Sac. London Ser. B 148, 1-37. Hodgkin, A. L. (1975). The optimum density of sodium channels in an unmyelinated nerve. Pliilos. Trans. R . Soc. London S p y . B 270, 297-300. Hodgkin, A. L., and Huxley, A. F. (1952a). Currents carried by sodium and potassium through the membrane of the giant axon of Loligo. J. Physiol. (London) 116,449-472. Hodgkin, A. L., and Huxley, A. F. (1952b). The components of membrane conductance in the giant axon of Loligo. J. Physiol. (London) 116, 473-496. Hodgkin, A. L., and Huxley, A. F. (1952~).The dual ell’tct of membrane potential on sodium conductance in the giant axon of Loliyo. J. Physiol. (London) 116, 497-506. Hodgkin, A. L., and Huxley, A. F. (1952d). A quantitative description of membrane current and its application to conduction and excitation in nerve. J . Physiol. (Lundon) 117, 500-544. Hodgkin, A. L., and Huxley, A. F. ( 1953). Movement of radioactive potassium and membrane current in a giant axon. J. Physiot. (London) 121, 403-414. Hodgkin, A. L., and Katz, B. (1949a). The effect of sodium ions on the electrical activity of the giant axon of the squid. J. Physiol. (London) 108, 37-77. Hodgkin, A. L., and Katz, B. (1949b). The effect of temperature on the electrical activity of the giant axon of the squid. J. Physiol. (London) 109, 240-249. Hodgkin, A. L., Huxley, A . F., and Katz, B. (1949). Ionic currents underlying activity in the giant axon of the squid. Arch. Sri. Physiol. 3, 129-150. Hodgkin, A. L., Huxley, A. F., and Katz, B. (1952). Measurement of current-voltage relations in the membrane of the giant axon of Luligo. J . Physiol. (London) 116, 424-448. Holden, A. V. (1976). The response of excitable membrane models to a cyclic input. Biol. Cybernet. 21, 1-7. Horn, R., Patlak, J., and Stevens, C . F. (1981). Sodium channels need not open before they inactivate. Nature (London) 291, 426-427. Hoyt, R. C. (1963). The squid giant axon. Mathematical models. Biophys. J . 3, 399-43 I . Hoyt, R. C. (1968). Sodium inactivation in nerve fibers. Biophys. J . 8, 1074-1097. Huxley, A. F. (1959). Ion movements during nerve activity. Ann. N . Y . Arad. Sci. 81, 22 1-246. I n w e , I. (1980). Separation of the action potential into a Na-channel spike and a K-channel spike by tetrodotoxin and by tetraethylammonium ion in squid giant axons internally perfused with dilute Na-salt solutions. J. Gen. Physiol. 76, 337-354. Jack, J. J . B., Noble, D., and Tsien, R. W. (1975). “Electric Current Flow in Excitable Cells.” Clarendon, Oxford. Joyner, R. W., Westerfield, M., Moore, J . W., and Stockbridge, N. (1978). A numerical method to model excitable cells. Biophys. J . 22, 155-170. Joyner, R. W . , Westerfield, M., and Moore, J. W. (1980). Effects of cellular geometry on current How during a propagated action potential. Biophy.~.J . 31, 183-194. Keynes, R. D., and Kimurd, J. E. (1980). The effect of starting potential on activation of the ionic conductances in the squid giant axon. J . Physiol. (London) 308, 17P. Keynes, R. D., and Kirnura, J. E. (1983). Kinetics of activation of the sodium conductance in the squid giant axon. J . Physiol. (London) 336, 621-634. Kcynes, R. D., and Rojas, E. (1974). Kinetics and steady-state properties of the charged system controlling sodium conductance in the squid giant axon. J . Physiol. (London) 239. 393-434.

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Keynes, R. D., and Rojas, E. (1976). The temporal and steady-state relationships between activation of the sodium conductance and movement of the gating particles in the squid giant axon. J . Physiol. (London) 255, 157-189. Keynes, R. D., Greeff, N. G . , and Van Helden, D. F. (1982). The relationship between the inactivating fraction of the asymmetry current and gating of the sodium channel in the squid giant axon. Proc. R . Soc. London Ser. B 215, 391-404. Kimura, J. E., and Meves, H. (1979). The effect of temperature on the asymmetrical charge movement in squid giant axons. J . Physiol. (London) 289, 479-500. Kniffki, K.-D., Siemen, D., and Vogel, W. (1981). Development of sodium permeability inactivation in nodal membranes. J . Physiol. (London) 313, 37-48. Kukita, F. (1982). Properties of sodium and potassium channels of the squid giant axon far below 0°C. J . Membr. B i d . 68, 151-160. Landowne, D. (1977). Sodium efflux from voltage clamped squid giant axons. J . Physiol. (London) 266,43-68. Levinson, S. R., and Meves, H. (1975). The binding of tritiated tetrodotoxin to squid giant axons. Philos. Trans. R . Soc. London Ser. B 270, 349-352. Lipicky, R. J., Gilbert, D. L., and Ehrenstein, G . (1978). Effects of yohimbine on squid axons. Biophys. J . 24, 405-422. Matteson, D. R . , and Armstrong, C. M. (1982). Evidence for a population of sleepy sodium channels in squid axon at low temperature. J . Gen. Physiol. 79, 739-758. Mauro, A,, Conti, F., Dodge, F., and Schor, R. (1970). Subthreshold behavior and phenomenological impedance of squid giant axon. J . Gen. Physiol. 55,496-523. Meves, H. (1974). The effect of holding potential on the asymmetry currents in squid giant axons. J . Physiol. (London) 243, 847-867. Meves, H. (1975). Calcium currents in squid giant axon. Phil. Trans. R . Soc. London Ser. B 270, 377-387. Meves, H. (1978). Inactivation of the sodium permeability in squid giant nerve fibres. Prog. Biophys. Mol. Biol. 33, 207-230. Meves, H., and Pichon, Y. (1977). The effect of internal and external 4-aminopyridine on the potassium currents in intracellularly perfused squid giant axons. J . Physiol. (London) 268, 51 1-532. Meves, H., and Vogel, W. (1973). Calcium inward currents in internally perfused giant axons. J . Physiol. (London) 235, 225-265. Meves, H., and Vogel, W . (1977). Inactivation of the asymmetrical displacement current in giant axons of Loligo forbesi. J . Physiol. (London) 267, 377-393. Moore, J. W., and Adelman, W. J., Jr. (1961). Electronic measurement of the intracellular concentration and net flux of sodium in the squid giant axon. J . Gen. Physiol. 45, 77-92. Moore, J . W., and Cole, K. S. (1960). Resting and action potentials of the squid giant axon in vivo. J . Gen. Physiol. 43, 961-970. Moore, J. W., and Cole, K. S. (1963). Voltage clamp techniques. In “Physical Techniques in Biological Research” (W. L. Nastuk, ed.), Vol. VIB, pp. 263-321. Academic Press, New York. Moore, J. W., and Narahashi, T. (1967). Tetrodotoxin’s highly selective blockage of an ionic channel. Fed. Proc. Fed. A m . Soc. Exp. Bid. 26, 1655-1663. Moore, J . W., and Young, S . H. (1981). Dynamics of potassium ion currents in squid axon membrane. A re-examination. Biophys. J . 36, 715-722. Moore, J. W., Anderson, N. C., Blaustein, M. P., Takata, M., Lettvin, J. Y . , Prickard, W. F., Bernstein, T., and Pooler, J . (1966). Alkali cation specificity of squid axon membrane. Ann. N . Y . Acad. Sci. 137, 818-829.

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Moore, J. W., Ramon, F., and Joyner, R. W. (1975). Axon voltage-clamp simulations. I. Methods and tests. Biophys. J. 15, 11-24. Narahashi, T. (1964). Restoration of action potential by anodal polarization in lobster giant axons. 1.Cell. Cutnp. fhysiol. 64, 73-96. Nemoto, I., Miyazaki, S . , Saim, M., and Utsunomiya, T. (1975). Behavior of solutions of the Hodgkin-Huxley equations and its relation to properties of mechanoreceptors. Biophys. 1. 15, 469-479. Neumcke, B.,Nonner, W . , and Stampfli, K. (1976). Asymmetrical displacement current and its relation with the activation of sodium current in the iiiernbrane of frog myelinatcd nerve. tf//iiyers Arch. 363, 193-203. Oikawa, T., Spyropoulos, C. S . , Tasaki, l., and 'reorell, T. (1961). Methods for perfusing the giant axon of Loligo peulii. Act0 Physiol. Scnnd. 52, 195-196. Oxford, G. S . (1981). Some kinetic and steady-state properties of' sodium channels after removal of inactivation. J . Grn. Physiol. 77, 1-22. Oxford, G.S . . and Swenson, R. P. (1979). n-Alkanols potentiate sodium channel inactivation in squid giant axons. Biophys. J . 26, 585-590. Oxford, G. S . , and Yeh, J. Z. (1979). Interference with sodium inactivation gating in squid axons by internal monovalent cations. Biophys. J . 25, 195a. Oxford. G . S . , Wu, C . H., and Narahashi, T. (1978). Removal of sodium channel inactivation in squid giant axons by N-bromoacetamide. .I. Gen. Physiol. 71, 227-247. Poussart, D.,Moore, L.. E., and Fishman. H. M. (1977). Ion moveinents and kinetics in squid axons. I. Complex admittance. Ann. N . Y. Acad. Sci. 303, 354-379. Ramon. F., Anderson, N . , Joyner, R. W . , and Moore, 1. W. (1975). Axon voltage-clamp simulations. IV. A multicellular preparation. Eiopliys. 1. 15, 55-69. Ramcin, F., Moore, J. W., Joyner, R. W . , and Westerfield, M. (1976). Squid giant axons. A model for the neuron soma? Biuphys. J . 16, 953-963. Robinson, R. A., and Stokes, R. H. (1959). "Electrolyte Solutions," 2nd ed. Butterworths, London. Rojas, E., and Atwater, I. (1967). Effect of tetrodotoxin on the early outward currents in perfused giant axoiis. Proc. Nut!. Acud. Sci. U . S . A . 57, 1350-1355. Rudy, B. (1978). Slow inactivation of the sodium conductance in squid giant axons. Pronase resistance. 1.Physiol. (London) 283, 1-21. Shoukimas, J. J., and French, R . J . (1980). Incomplete inactivation of sodium currents in nonperfused squid axon. Biophys. J . 32, 8.57-862. Tasaki, l., and Singcr, 1. (1966). Membrane macromolecules and nerve excitability: A physico-chemical interpretation of excitation in squid giant axons. Ann. N . Y. Actid. Sci. 137, 792-806. Tasaki, I . , Watanabe, A , and Takenaka, .J. (1962). Resting and action potential of intracellularly perfused squid giant axon. f v o c . Nrctl. Acad. Sci. U.S.A. 48, 11771184. Tasaki, I., Watanabe, A., and Singer, I. (1966). Excitability of squid giant axons in the absence of univalent cations in the external medium. Proc. N u t / . Actid. Sri. U.S.A. 56, I 1 16-1 122. Tasaki, I., Watanabe, A . , and Lerman, L. (1967). Role of divalent cations in excitation of squid giant axons. A m . J . Physiul. 213, 1465-1474. Taylor, R. E., and Bezanilla, F. (1983). Sodium and gating current time shifts resulting from changes in initial conditions. .I. Gen. Physiol. 81, 773-784. Taylor, R. E., Moore, J . W., and Cole, K . S.(1960). Analysis ofcertain errors in squid axon voltage clamp measurements. Biophys. J . 1, 161-202.

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Wang, C. M., Narahashi, T., and Scuka, M. (1972). Mechanism of negative temperature coefficient of nerve blocking action of allethrin. J . Pharmrrcol. Exp. Ther. 182, 442-453. Watanabe, A., Tasaki, I . , Singer, I., and Lerman, L. (1967). Effects of tetrodotoxin on excitability of squid giant axons in sodium-free media. Science 155, 95-97. Yeh, J. Z., Oxford, G. S., Wu, C. H., and Narahashi, T. (1976). Dynamics of arninopyridine block of potassium channels in squid axon membrane. J . Gen. Physiol. 68, 519-535.

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CURRENT TOPICS IN MEMBRANES A N D TRANSPORT, VOLUME 22

Sequential Models of Sodium Channel Gating CLAY M . ARMSTRONG AND DONALD R . MATTESON Depurlment of Physiology University of Pennsylvania School of Medicine Philadelphia, Pennsylvania

I. Introduction.. . . . . . .

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

332

11. Results ...................................... A. How Many Closed Conformations Does a So

B. The Models.. .................................... C. Energy Diagrams for the Seq D. Calculation of Is and Conduc E. Conductance and I , Kinetics ......................................... F. Steady-State g-V and Q-V Relations. .............. G . Single-Channel Simulations . H . A Voltage-Independent Polymerization Step. .......................... 111. Discussion .............................................................

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

339

347 350 352

It is argued that state diagrams are the natural language for describing the behavior of ionic channel proteins. Several means are available for determining the number of states and the rates of the transitions among them: measurement of macroscopic and single channel ionic currents, and measurement of gating current. The relative advantages of the three techniques are examined by analyzing variants of a four-state model of the sodium channel activation process. The four-state model is the simplest that is compatible with macroscopic current kinetics in the squid giant axon. The mathematical description of the histograms that summarize single-channel kinetics is very similar to that for the time course of macroscopic current. With the exception of the closed time histogram, macro331 Copyright 0 1984 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-153322-0

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scopic currents and single-channel histograms give little informal ion about the transition rates among closed states. Gating current gives information about all steps in the activation sequence that involve charge movement. At very negative voltages, the steepness of the conductancevoltage relation gives a good measure of total gating charge movement. At more positive voltages, the steepness of the curve is dependent on the probability of the states intermediate between fully closed and fully open. The kinetics of any steps with voltage-independent rate constants must be very rapid. 1.

INTRODUCTION

‘This article has two major purposes: to cxamine the advantages of state diagrams in the description of ionic-channel behavior, and to discuss the means available for determining the number of states and the rate constants of the transitions among them. Ionic channels are sensitive to membrane voltage because they have mobile “gating charges” as part of their structure (Hodgkin and Huxley, 1952; Armstrong and Bezanilla, 1974; Keynes and Rojas. 1974). These charges redistribute after a change of the membrane field, imparting a twist to the channel structure that may open or close it. Movement of these charges generates “gating current,” which is associated with the gating process, and only indirectly with ionic current flow through the channels. Gating current gives direct information about the molecular rearrangements of the channel protein as it opens and closes. Complementary information can be obtained by measuring ionic current flow through the open channels, either macroscopically or as single-channel events (Sigworth and Neher, 1980; Patlak and Horn, 1982). Macroscopic currents are valuable among other things for determining average behavior of the channels, for measuring the gating charge per channel, and for indicating the number of closed states. Single-channel measurements give a relatively direct measure of some of the rate constants. Ionic channels are now known to be composed of large proteins. The natural language for describing the behavior of channel proteins kinetically is the state diagram (Fitzhugh, 1965). Those familiar with the conciseness of the Hodgkin and Huxley (1952) expression for the number of open sodium channels Fraction open = m3h may be persuaded only with difficulty that there is virtue in expressing

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333

this relation in the much more cumbersome form of its equivalent state diagram (Diagram 1) (Fitzhugh, 1965; Armstrong, 1969; Hoyt, 1971). activation gute:

closed 3

inactivation gate: closed

activation gate:

closed 3

inactivation gute: open

2closed 2 P

closed

2closed 2 P

open

- 2~

closed 1 closed

2P

za

2P

closed I open

u

a

v

3P

open closed

3P

open open

DIAGRAM 1

In Diagram 1 the upper entry of each row refers to the activation process, which, in Hodgkin and Huxley's formulation, can be in any of four states: closed by 3, 2, I , or none (i.e., the gate is open) of their hypothetical gating particles. The lower entry refers to the inactivation gate, which can be closed or open. The channel conducts only when both gates are open. What are the advantages of the state diagram? With a structurally unknown protein, the rates of the transitions among conformational states must be determined empirically and cannot be derived from first principles. There are assumptions implicit in the Hodgkin and Huxley formulation that contribute to its conciseness but make it not purely empirical. Specifically, the forward rate constants in the diagram above have the ratio 3 : 2 : I going from left to right, and the reverse rate constants have the same ratio going from right to left. Further, the rate constants for closing and opening of the inactivation gate are assumed to be the same regardless of the state of the activation gate. One can easily imagine, instead, that inactivation might be more rapid from one state than from another. The Hodgkin and Huxley equations thus can be viewed as a special case of the state diagram, simplified by the two assumptions just noted. They are perfectly appropriate to a mode1 with independent gates and gating particles, but inappropriate to the transitions among conformational states of a protein, where the transition rates follow no simple rules. Although cumbersome, a state diagram is better suited to this problem, and, fortunately, is easy to handle computationally. To illustrate the information derivable from the available experimental approaches, and the utility of state diagrams, we have chosen to examine several four-state models of the activation gating process for the sodium channel. Four states, three closed and one open, is the smallest number that is consistent with the sigmoid rise of sodium conductance following a

334

CLAY M. ARMSTRONG AND DONALD R. MATTESON

depolarization, and its exponential decay on repolarization, as explained in Section 11. We have further simplified by not including inactivation. 11.

RESULTS

A. How Many Closed Conformations Does a Sodium Channel Have? As noted above, the sodium conductance increases along a sigmoid curve after a depolarization. The foot of the sigmoid gives a good indication of the minimum number of closed states or conformations, as illustrated in Fig. I . The curves show the theoretical time course of channel activation for channel models with one, three, and five closed states:

closed,

- - - - closed

open

closed3

closed2

closed,

open

closed,

closed,

closedz

closed,

-

open

(For economy of space, the states of the middle scheme will be referred to as X4, X 3 , X2,and X f , where Xf is the open state.) The calculations assume that all channels are in the leftmost state at the beginning of the trace, and that all are open at trace end. Rate constants are in the figure caption. The lag in the activation of the channels is progressively larger as the number of closed states increases, because more closed states must be

0

time

FIG. I . Conductance time course calculated from sequential models with 2, 4, and 6 states (1, 3, and 5 closed states) illustrates that the delay in the rise of conductance is clearly larger as the number of closed states increases. Within each model the forward rate constants were equal and had values of 2.S, 5.0. and 6.3 rnsec-' for the 2-, 4-, and 6-state models, respectively. It was assumed that all channels start in the leftrnost closed state and that all reverse rate constants were zero, so that all channels end in the open state.

SEQUENTIAL MODELS OF SODIUM CHANNEL GATING

335

traversed before the conductance begins to rise. To produce a lag it is necessary that the states be traversed sequentially. In the four-state scheme (Diagram 2), some of the channels would begin to open without a

x3

-x4

DIAGRAM 2

lag, because there is a direct path to the conducting state from all the other states, and the scheme is thus not compatible with the experimental observations. The calculations in Fig. 1 assume that the forward rate constants within a given scheme are equal (see figure caption). Subsequent calculations show that the lag is not highly sensitive to the relative values of the rate constants. If, however, some are much faster than others, the lag (as a fraction of the half-time) is reduced, and, for example, five closed states with highly unequal rates can give the same lag as three closed states with equal rates. The minimum number of closed states compatible with the rise of the sodium conductance is three. The lag in activation, and the complexity it introduces into modeling, are so striking that one is compelled to ask what advantages there are to a nerve fiber in having several closed states? A reasonable answer is that the closed states serve as a filter, and thus give the fiber some noise immunity. Were there only one closed state, the immediate opening of the channels on depolarization would make the fiber much more susceptible to triggering by brief random thermal voltage changes, produced by chance accumulations of negative or positive ions. Closing kinetics of sodium channels at negative voltages are described by a single exponential. In the four-state models, a single step (X: + X,) is required to close the channel, and this has exponential kinetics.

B. The Models The preceding section gives the rationale for considering four-state models of sodium channel activation, of the generic form

x481

Ul

--

x,

Pz

x:

a1

PI

x:

where X4. X 3 , and X2 are closed states and X: is the conducting state.

336

CLAY M. ARMSTRONG AND DONALD R. MATTESON

The specific embodiments to be considered are

The first sequence is equivalent to the Hodgkin and Huxley rn' scheme, as described in the Introduction. The behavior of these schemes, as manifested in macroscopic and single-channel currents and in gating current, are examined in the next sections. C. Energy Diagrams for the Sequential Models Associated with each step in the sequences given above is a movement of gating charge that makes the step voltage dependent. The charge movement is not nccessarily the same for all steps. It is rigidly linked to the transition, which cannot occur without the charge movement. The steps and their associated charge movement can be convcniently summarized with Eyring-type diagrams like the one in Fig. 2 (top left), which pertains to the m3 sequence. The ordinate is free energy, and the abscissa is the quantity of gating charge that has moved from resting toward active configuration, explained momentarily. The four stable conformational states are the free-energy minima distributed along the abscissa, with transitional states in between. The minima correspond to closings of the activation gate by three, two, one, or zero of the m particles. The horizontal distance between two neighboring states is proportional to the charge movement associated with the transition between them. Equal spacing of the states, as in Fig. 2, means that all transitions involve the same charge movement. Charge movement and the free-energy difference between, for example, states X3and X2 are related by Eq. (1). QxI-x,

= A(Gx~ ~

Gx,)/AV

(1)

Qx7+xzis the charge movement that occurs when a single channel undergoes the X3to X2 transition. G (in meV) is the free energy of a channel in the state specified by the subscript, and A V is a change of mcmbrnne voltage in millivolts. Gating charge movement equals the change in free energy per change in membrane voltage. The solid curve in Fig. 2 (top left) shows the energy profile for an m3

SEQUENTIAL MODELS OF SODIUM CHANNEL GATING

337

m3

Fic,. 2. Free-energy diagram\ for the models at V , , the voltage where X4 and X: are equally probable. Free energy is given on the ordinate; the abscissa represents gating charge. The dashed curve at the upper left illustrates how the energy profile would be slanted at a membrane voltage more negative than V , .

channel at the membrane potential where X4 and Xf are equally probable. At this potential, which will be called V,, half of the m particles are in their “gate open” position (all three must be in this position actually to open the gate). In squid axons this voltage is -30 to -35 mV (Hodgkin and Huxley, 1952). At V,, states X3 and XI are three times more probable than states X4 and XT purely for statistical (or entropic) reasons: there are three ways of arranging the indistinguishable rn particles to get states X1 and X!, and only one arrangement each for Xq and X?. (If the particles were distinguishable, both states X3 and X2 would be replaced by three states.) The factor three corresponds to a free-energy difference of I . I kT ( k is Boltzmann’s constant; T is the absolute temperature), or about 27.5 meV per particle at room temperature. At a more negative voltage (Vi, - V,,,,) the diagram would be slanted as shown by the dashed line (Fig. 2, top left): the relative energies and probabilities of all the states have changed in favor of the ones to the left in the diagram. The other parts of Fig. 2 show energy diagrams for the 1 : I : 1 and 1 : 2 : 3 sequences at the voltage (VJ for which states X4 and X r are equally probable. The four states of the 1 : 1 : 1 sequence have identical free energies and probabilities at V,. The 1 : 2 : 3 sequence is in a sense the comple-

338

CLAY M. ARMSTRONG AND DONALD R. MATTESON

ment of the m 1 sequence: states X4 and XT are three times more probable than XZ and X3.The last diagram ( z = 6 ) represents a channel with only two possible states, closed and open. The four diagrams form a progrcssion with regard to the probability of occupancy of the intermediate states. which are relatively probable in the m’ sequence and steadily less so in the I : I : I and 1 : 2 : 3 sequences. Finally, for z = 6, the limiting case, there are no intermediate steps, or, stated another way, the probability of intermediate states is zero.

D. Calculation of I, and Conductance

Membrane potential affects the relative energies of the states because there is charge movement through the membrane field during the conformational change from, say, X3 to X2. Suppose n charged groups on a gating molecule move in the transition, that qbis the charge on a given group, and that d, is the fraction of the membrane field through which group i moves. Conformation and charge distribution are rigidly linked, so all the groups must move simultaneously when the transition occurs. Total charge movement for this step (in electronic charges per channel) is given by Eq. (2).

(2) QxJ+x? = (qidi + q2d2 + ... + q d t t ) From electrical measurements there is no way to distinguish between two electronic charges moving halfway through the field, and one charge moving all the way. Q here is the charge movement referred to the complete membrane field. Gating current associated with the one-way step X3+. X2 is the product of the charge per channel and the number of channels per unit time that undergo transition [Eq. (3)]. N~x3QxI-x~ (3) N here is the total number of channels, and X3 is the fraction of the channels that are in state X3. For the same step in reverse, the equation is lg

=

(4) Qxrx3 is cqual to Q x ~ +and ~ ~will be called Q2,3.For each of the other steps there is a similar pair of equations, and total gating current is the sum of that for the three forward and the three reverse steps [Eq. ( S ) ] . 1, = -NPzXZQX~+X, ~

I,

=

N(rulX.4 - PlXdQ1.4

+ (aJ3

-

P.XZ)O21 + (aiX2

-

PiXT)Q1,2 ( 5 )

SEQUENTIAL MODELS OF SODIUM CHANNEL GATING

339

In the following section, gating current is calculated from Eq. ( 5 ) with the assumption that the charge movement is equal for all the steps. That is, (21.2 = Q2.3

= Q3,4

(6)

This is necessarily so for the m3 sequence, since each of the steps represents the movement of one m particle through the membrane field. To simulate the conductance, each of the sequential schemes was described by four first-order differential equations of the form

dX3ldt = a3X4

- (a2

+ p3)X3 + ,&X2

(7)

These equations were solved numerically for XT. It was assumed that before the voltage change all the a’s are zero and all the channels are in state X4. E. Conductance and I, Kinetics

The calculations in Fig. 3 simulate currents generated by three depolarizations that produce steady-state Xr values of 0.99, 0.5, and 0.1. The time course of conductance (XT) is quite insensitive to the ordering of the rate constants, and the XT curves for the three sequences are almost superimposable except at the intermediate depolarization. Further calculations, which are not illustrated, show that, as the rate constants are made steadily more unequal, the sigmoid shape of the curve tends to be lost, and the time course comes to resemble an exponential with a time constant that is the reciprocal of the smallest rate constant. The time course of Zg is much more sensitive to the order of the rate constants than is conductance. For the largest depolarization the m3 sequence predicts an exponentially declining I,, while the 1 : 2 : 3 sequence has an initial rising phase, and the I : 1 : I sequence an initial plateau. The reason is that charge movement slows down as channels move to the right in the m3 sequence, where the rate constants are steadily smaller, and current is thus largest initially. Exactly the opposite is true for the 1 : 2 : 3 sequence; charge movement is faster as channels move to the right, so Z, increases in amplitude. For the I : 1 : 1 scheme, charge movement neither accelerates nor decelerates on moving right in the diagram, resulting in an initial plateau. At less positive voltages, I , calculated from the I : 2 : 3 sequence has an initial fast component, whereas that calculated from the other sequences is more nearly exponential. This results from the large value of p3 in the 1 : 2 : 3 sequence, which recycles channels from X3 back to X4, generating

340

CLAY M. ARMSTRONG AND DONALD R. MATTESON A

C.

iii

I

I \

B

FIG.3. Simulated gating current (Ig;top traces in each panel) and the fraction of open channels ( X : ; lower 1r;ices in each panel) for three depolari7ations that yield sle;idy-si;itc values of ( A ) 0.1, ( B ) 0.5, and (C) 0.99 for thc open lniclion. The time coursc of I , is very scnsitive lo values of the rate constants, while Xr is less sensitive. The rate constants (in msec I) were as follows: a1

a?

83

Pz

( A ) m' I: I: 1 1:2:3

2.80

1.86

0.93 1.07

2.14

3.21

1.86

1.86 2.69

1.86 3.09

4.04 7.76

3.09 5.17

3.09

1.35

(B) m3 I :I :I 1:2:3

6.29 4.40

4.19 4.40

2.10 0.55

1.09

4.40 2.39

2.39

1.64 2.39

3.38

6.76 10.1

5.42

2.71

(C) m3

I:I:I 1:2:3

21.5

14.2

11.9

11.9

7.16 14.3

8.14

PI

2.59

7.16 0.024 0.048 0.072 0.12 0.12 0.12 21.5 0.63 0.42 0.21 11.9

current of opposite sign and leading to the rapid decline in the initial phase of zg. For the rn3 sequence much charge movement occurs before conductance (XT) rises to, e.g., 10% of its final value: the area under the I,,rn3 curve to the left of the point where XT is 0.1 is larger than for the other sequences. This is a consequence of the faster rate constants to the left in the r n 3 sequence: many channels move from X4to X3 and XZbefore some of them take the slower step to X f . Simulations of conductance (X:) and 1, during the closing of the chan-

SEQUENTIAL MODELS OF SODIUM CHANNEL GATING

341

FIG.4. The predicted time course of gating current and conductance as the channels close. It is again clear that the time course of gating current is quite sensitive to the rates of transition between states. The rate constants (in msec-’) were

P3 rn’ 1: I : 1 1 :2 : 3 cr3, u 2 ,a ,

P? PI

1.0 2.0 3.0 3.0 3.0 3.0 9.0 6.0 3.0

were zero in all cases, and Xf was I .O at the beginning of the calculation.

nels following repolarization are shown in Fig. 4. The calculations assume that Xr = 1.0 when the step is applied, and that all the constants aiare zero after the step, causing all the channels to end in state X4. Rate PI has been given the same value for the three sequences ( 1 .O in arbitrary units). The conductance curve for the three sequences (XT) is a single exponential with time constant 1 .O in arbitrary units under these conditions, for the closing of the channels is a one-step process with first-order kinetics (Xr -+ X2). The predicted Igcurves (Fig. 4) are the same as those in Fig. 3 , but inverted and with different relative amplitudes. Zg for the m 3 sequence is exponential, with a time constant three times slower than that for the conductance curve. Thus, any of the sequences can be fitted to the decay of conductance by appropriately setting a single rate constant. Gating current time course, on the other hand, depends on all the rate constants. This section can be summarized by saying that gating current time course is strongly affected by the values and ordering of the rate constants, and macroscopic current much less so. For the three schemes considered, macroscopic currents were similar or identical, while the predicted gating current traces were usually quite distinct. F. Steady-State g-V and 9-V Relations

Each of the sequences makes predictions about gating charge distribution and steady-state conductance. It is assumed in the following calculations that the charge movement associated with each step is the same, and

342

CLAY M. ARMSTRONG AND DONALD R. MATTESON

equal to 2e (electronic charges) per step moving through the entire field, or a total of six electronic charges per channel. The steady-state distribution among the various states was determined over a range of voltagcs, and Qti(normalized charge movement) was calculated from the equation Q,, = X3/3

+ 2x213 + X:

(8)

Q, is zero when all the channels are in state X4 (X, = l.O), and 1.0 when all are in statc X:. In terms of the m 3sequence, these assumptions correspond to having three m particles per channel, each with a charge of 2e. Each particle has two possible positions, and in passing from one position to the other it traverses the entire membrane field. It should be noted that the behavior of Hodgkin and Huxley's m particles is similar but somewhat more complex. Calculations of conductance and gating charge as a function of voltage are shown in Fig. 5 , where X r - V and QII-Vcurves are plotted for the four

-604--20

0

2 0 4 0

V. rnV

FIG.5 . Steady-state conductance (X:) and charge distribution (Q)as a function ofincmhrane volt;ige, for the four models of Fig. 2. Ry definition, Q is zero when all ch;tnnels :ire in state Xq.and 1.0 when all arc in statc XT. V , was assumed to he -30 m V .

SEQUENTIAL MODELS

OF SODIUM CHANNEL GATING

343

models represented in Fig. 2. The curves change in an orderly way as one progresses from the m3 scheme, which has intermediate states (X, and X2) of relatively high probability, to the single-particle, two-state scheme ( z = 6), for which there are no intermediate states. As X2 and X3 become less probable, both the XT and the Qn curves become steeper near their midpoints, and they draw closer together until they coincide for the singleparticle model. It is interesting to note that for a given sequence the XT and Qn curves at their respective midpoints are not much different in steepness, even for the m’ sequence, where the discrepancy is greatest. For all sequences, however, the steepness of the two curves is quite different at very negative voltages, where, in the limit, conductance changes e-fold for 4 mV (kt/6e) while the Q,, curves change e-fold for 12.5 mV (kt/2e).

G. Single-Channel Simulations Single-channel recordings can be used to construct three useful histograms: open time, closed time, and latency to first opening. To evaluate the kinetic information available from these histograms, we have simulated single-channel records for the three sequences and constructed histograms from them. We have also derived equations for the histograms of four-state models, following the procedure of Colquhoun and Hawkes (1981). An interesting point arising from the derivations is that the equations describing single-channel behavior are very similar to those for macroscopic currents, as discussed below. Single-channel simulations were performed in the following way. The probability that a channel undergoes a transition from one state to another (e.g., from X3 to X,) in time interval At is approximately equal to the rate constant governing the transition ( a z )multiplied by A t , if At is sufficiently short. A channel in state X3 at the beginning of an interval can be in X4, Xz, or X3 at the end of the interval. The respective probabilities are p 3 A t , a2At, and 1 - (p3+ az)Ar. To decide among these possible fates, the range 0 to 1 was divided into three intervals, each equal to the probability of one of the possible outcomes; and a random number between 0 and 1 was drawn that necessarily fell into one of the intervals, thus determining the outcome. Simulated single-channel records generated by this procedure are shown in Fig. 6 for each of the three sequential models. The rate constants used here, and in all remaining calculations in this section, were the same as those in Fig. 3B, corresponding to a steady-state value for X; of 0.5. The most obvious feature in the simulations is that the frequency of short closings is highest in the 1 : 2 : 3 sequence and lowest in the 3 : 2 : 1

344

CLAY M. ARMSTRONG AND DONALD R. MATTESON

N L n r u w mm

u

In3

A

Jlnnm-Lm I u u L 1 l-Lmm3m r r w llnLmrr rnrNlJ I . M M ~.

__

7

Bwm r rnrumr~ I U. - i l l u r n J L l L J N

LJ-KmNm

m m.m

c h . p c L

.-rwuLr-

1

m n n nN L ~ mmI r m m Lm-. SmULlrl

~

T

FIG. 6 . Simulations of single-channel behavior for depolariLations to the membrane vol~ageresulting in a steady-statc value o f X: of0.S.Each trace simulates the response to a single depoliirization. Brief closings arc niore frequent for thc I : 2 : 3 sequence (C) than for and least frequent for the m' sequence (A). At the beginning of each the 1 : 1 : I sequence (8) trace the channel was in state X q . Upward deflections indicate channel opening. Rate constants were the same as those used for Fig. 3B.

sequence. This is because the rate constant that cuts short the opening, 1 : 2 : 3 sequence and smallest in the 3 : 2 : 1 sequence. The channel records generated from the 3 : 2 : 1 sequence were used to construct the open-time histogram shown in Fig. 7. A total of 148 openings were measured and plotted with a bin width of 50 p e c . The smooth curve in the figure is the probability density function of open times predicted for the same model. This function is a single exponential and has the form

P I , is largest in the

fO> = PI exP(-Plr)

(9)

Thus, if the channel has only a single way to close, the open-time distribution gives a meabure of the closing rate constant. More generally, if there is a single open state that can close in several ways, the open-time distribution is a single exponential with rate constant equal to the sum of all closing rate constants.

SEQUENTIAL MODELS

OF SODIUM CHANNEL GATING

345

Time, msec

FIG.7 . Histogram of open times from simulations of the mi sequence and the predicted distribution of this parameter. The smooth curve is Eq. (9) (with PI equal to 1.64 msec-I), scaled by the total number of openings (148). If the channel has only one way to close, a single exponential fit to the open time histogram gives a measure of the closing rate constant.

The distribution of latency to first opening, and the distribution of closed intervals, were derived as described by Colquhoun and Hawkes (1981). The probability density functions of these parameters are the sum of three exponential terms. Assuming that the channel starts in the resting state X4, the distribution of latencies to first opening is given by

wheref(t) is the probability density that the first channel opening following depolarization occurs at time t ; and L I , L 2 ,and L3 are the three roots of the equation s3

+ s2(ff,+ ff? +

ffj

+ pz + pj)

+ s(ffla2 + a I f f 3 + f f 2 f f 3 + alp3 + a3p2 + p2p3)

+ ( ~ 1 ( ~ 2 ( =~ 3 0

(1 I )

Plots of this equation for all three models are shown in Fig. 8A. The three

346

CLAY M. ARMSTRONG AND DONALD

A

R. MATTESON

B

i

i

3

2

2

3

T l m e , msec

FIG.8. Predicted distribution of latency to first opening for the three sequential models. The probability density functions [Eq. (lo)] are shown in (A), where the ordinate is in units of probability per millisecond, and the cumulative distributions of first opening latencies arc shown in (B). These predicted distributions for the three models arc quite similar, indicating that this parameter is insensitive to the values and ordering of the rate constants. thc values of the rate constants were the same as those used in Fig. 3B (SIX Fig. 3 caption).

curves are very similar. In view of the random noise associated with histograms constructed from a small number of events, as in Fig. 7 , latency to first opening may not be very useful for estimating rate constants. The cumulative distribution of this parameter, which is simply the integral (from 0 to t ) of the probability density function, is shown in Fig. 8B of the figure. These curves resemble, but are not exactly like, the sigmoid curves in Fig. 3B, which plot the time course of conductancc using the same rate constants. As discussed later, the macroscopic current predictions yield curves identical to those shown in Fig. 8B if PI is assumed to be zero. The equation describing the probability density function of closed intervals following openings is f(t) = ~ [ Cexp(LIt) I - C2 exp(L2t) + C3 exp(l3r)l

where CI =

(%a3 +

L,(w +

'y3

+

P3)

+ m J 5 2 - LI)(L3 - L , )

c2 = (a2u3 + Lz('y2 + 'y3 + p3) + L C3

= ('y2a3

W * - L,)(L3 - L,)

+ L3(a2 + a3 + 03) + L:)/(L3- LI)(L3 - L2)

(12)

347

SEQUENTIAL MODELS OF SODIUM CHANNEL GATING

1 .o

0.5

1lO

1.5

Time, msec

FIG.9. Predicted distributions of closed intervals. The probability density functions [Eq. (12)] are shown in (A) (ordinate in units of probability per millisecond), and the cumulative

distributions are shown in (B). The closed time distribution is clearly quite sensitive to the values of the rate constants. The rate constants were the same as those used in Fig. 3B (see Fig. 3 caption).

Plots of Eq. (12) and its integral are shown in Fig. 9A and 9B, respectively. The closed-time distribution is clearly quite sensitive to the rates of transitions between states. With respect to the simulations presented above, it was noted that the 1 : 2 : 3 sequence yielded the highest frequency of very brief openings, and this point is made again, in a more quantitative way, by the curves shown in Fig. 9A. This type of analysis may have limited usefulness, since experimentally a patch recording must contain only one channel for the closed times to be interpreted in this simple way (see Section 111).

H. A Voltage-Independent Polymerization Step The calculations presented so far have assumed equal voltage dependence or charge movement for all steps. It is perhaps more likely that some steps have greater voltage dependence than others, and an extreme example is a sequence with one voltage-independent step that could represent, for example, a polymerization of the type postulated to occur in

348

CLAY M. ARMSTRONG AND DONALD

R. MATTESON

the formation of an alamethicin channel (Baumann and Mueller, 1974). In the sequence

X,*is the conducting state, and the rate constants a and b of the step X I+ X,* are independent of voltage. The first three steps are identical to the m3 sequence, except that Xi is nonconducting. X:) on steady-state conductance The effect of the fourth step (Xi (X;) and Qn is determined by the ratio of rate constants a and h and is independent of their absolute values. X,* and Qn curves for two values of this ratio are shown in Fig. 10A, and for comparison the predictions of the m 3scheme are also given. If a = b, a maximum of one-half of the channels can be in state X,* and the saturation level is shifted to the left relative to the Xr, m’ curve. The Q curve for a = b (Q, a = b) is slightly steeper than the Q,m3 curve and lies to the left of it. For a = 100b, the Q and the X: curve are markedly steepened and are in close proximity to each othcr. They lie far to the left of the curves for the m 3 scheme. Their proximity can be viewed as a consequence of the low probability of occupancy of the intermediate states, as mentioned above. The kinetic consequences of a polymerization step depend on the amplitude of LI and b relative to the a’s and p ’ s . At one extreme, if the time constant of X , + X; [ l l ( n + h)] is relatively quick, the time course of X: is largely determined by the three slower steps. At the other extreme, if XI + X: is very slow, X: rises almost exponentially. This is illustrated in Fig. lOB, which shows calculations of these extreme cases together with predictions of the m 3 sequence. Rate constants are given in the figure caption. The gating currcnt predictions are identical in all three cases. In general, the fourth step would have some effect on the time course of Z,, but it has none at all in the cases illustrated because p is zero. The kinetics of conductance are the same whether the slow voltageindependent step is first or last, as illustrated in Fig. 1OC. The calculations are based on the scheme --f

-xz-x,

x, r‘x, P

3u

2u

2P

a-

30

xi

That is, the first rather than the last step is slow (rate constants are in the figure caption) and voltage independent. Unlike conductance, gating current time course is drastically altered by this transposition. A maximum estimate for the time constant of a voltage-independent step can be obtained experimentally at very large positive potentials, where a is large and the voltage-independent step might become rate

C

A

V, mV FIG. 10. Calculations for sequences with a voltage-independent step. ( A ) Conductance (Xg) and charge ((2)are plotted as a function of membrane voltage for two ratios of the rate constants 11 and h of a voltage-independent last step. The ordinate is fraction of total charge moved for the Q curves, and fraction of channels open for the X,* curves. The predictions of the m' sequence ( X t , m3and Q, m3)are shown for comparison. The overall effect of the voltage-independent last step is to draw the Q and XR curves together and shift them to the left. (B) The time course of gating current (1,) and open fraction (X:) for two different values of the rate constant u. The rate constants were (I = 90 msec-' ((1 = 30a) or (I = 0.1 msec I (u = a/30);with a = 3 msec-', = 0. and h = 0 in all cases. At the beginning of the calculation X, was 1 .O. For comparison, I, and conductance (X:) for the m3 sequence have been plotted, the latter at half the scale of the X,* curves, since X: rises to a final value of I .O and X i to a final value of 0.5. I, is completely independent of the final step. (C) The time course of gating current ( I , ) and conductance for a sequence with a voltage-independent first step. The rate constants were the same as those used in part (B) of this figure, except that (I and h pertain to the first step (see text). Unlike conductance, I , is greatly slowed by placing the voltage-independent step first.

350

CLAY M. ARMSTRONG AND DONALD R. MATTESON

limiting. In squid axons, the rise of conductance becomes steadily faster with V , with no indication of a saturation value. If there is a voltageindependent step, it must have very rapid kinetics. 111.

DISCUSSION

Channel behavior will inevitably be summarized in the future in terms of state diagrams. We have considered the methods available for getting information on the number of states required in such a diagram, and the rates of the transitions among them. In the following we summarize the best means of studying several aspects of channel gating, and then discuss the rather surprising fact that the mathematics of macroscopic currents and single-channel histograms are almost identical. 1. The number of closed states can be determined most directly from the lag in the activation of channels following a voltage step, or, almost equivalent, the histogram of latency to first opening of single channels. The effect on latency of the number of open states was illustrated in Fig. 1 . Very rapid steps contribute less to the latency, which thus gives a minimum estimate of the number of closed states. In simultaneously fitting gating and Na currents, it was found necessary to use five closed states, because fits of gating current required a number of rapid steps (Armstrong and Gilly, 1979). There is also considerable evidence for the existence of more than one open state, but this has been set aside here for simplicity. 2 . Charge per channel can best be determined from the steepness of the foot of the conductance-voltage curve, a method first employed by Hodgkin and Huxley (1952). At very negative voltages, where few channels are open and almost all of the gating charge is in its rcsting position, the g-V curve gives a good estimate of total gating charge per channel. At more positive voltages the steepness of the curve is strongly affected by the probability of the intermediate states, being steeper if they are improbable, as illustrated in Fig. 5. 3. Gating current is the most sensitive indicator of the transition rates among closed states, as illustrated in Fig. 3. The closed-time histogram for single channels also gives much information on these rates, provided the recording is from a single channel. As pointed out by Colquhoun and Hawkes (1981), however, this histogram is difficult to interpret if there is more than one channel. With a single channel, and a single closing path, one knows the state of the entire “system,” i.e., the one channel, immediately after it closes: it is in state X2. If there are several channels, the state of only one is known immediately after a closing, and the uncertainty regarding the others makes interpretation very complicated.

SEQUENTIAL MODELS OF SODIUM CHANNEL GATING

-

351

4. The closing rate constant P I , for the step XT X2, is best determined from the open time histogram or from the decay of macroscopic current at negative voltage. Mean channel lifetime is the time constant of the closing step, provided that there is only one path out of the conducting state. If there are several paths, the histogram reflects the sum of their rate constants. For the sodium channel there are at least two paths out of the conducting state: deactivation (i.e., closing of the activation gate), and inactivation. Patlak and Horn (1982) removed inactivation by the use of N-bromoacetamide and then measured directly the closing rate constant, p, in present terms, in rat myotubes. 5. Variation of initial conditions. In studying kinetics it is important to know the initial state of the channels. In the schemes considered here, all the channels can be put in state X4 by making V very negative, or in state Xf by making it positive. An advantage unique to single-channel measurements from a patch with a single channel is the possibility of knowing precisely when the channel enters state Xz, as discussed under 3 above: if there is a single closing path, a channel is in state X2 immediately after closing. 6. Single-channel recordings give direct information about “flickering,” rapid transitions from open to closed, that cannot readily be studied using the other methods. 7. Single-channel conductance can be most directly determined from single-channel records. There is always the possibility that single-channel conductance is much higher than one would guess from the current steps, because limited frequency response of the measuring system filters out high-frequency flickering. To our surprise, the mathematics for macroscopic currents and singlechannel histograms turned out to be very similar. The open-time histogram is described by Eq. (9) in Section II,G. Exactly the same operation describes the time course of macroscopic current for the generic scheme with a I equal to zero:

=x,P2====== x*-xt a?

x4

P?

a2

PI

It is necessary to assume that some of the channels start in state XT. The equation for the histogram of latency to first opening [Eq. (lo)] also describes the macroscopic current for the generic scheme if PI is zero:

and if the channels start in state X4.

352

CLAY M. ARMSTRONG AND DONALD R. MATTESON

Finally, the equation for the closed-time histogram (Section II,G) describes the scheme when p, is zero

if one assumes that all the channels start in state X2. The close mathematical relation between,the histogram equations and the macroscopic current equations suggests that the two measurements yield similar information. In general this seems to be true, with the important exception of the closed-time histogram. which is very senbitive to the transition rates among closed states. It may be that the method of maximum likelihood estimators explored by Horn and Lange (1983) will prove to be a better way of extracting kinetics information from single-channel events. Interest in channel states and kinetics may grow as structural data on thc channel proteins become available. REFERENCES Armstrong, C. M , (1969). Inactivation of the potassium conductance and related phenomena caused by quaternary ammonium ion injected in squid axon. J . Gen. Physiol. 54, 553. Armstrong, C.M., and Bezanilla, F. (1974). Charge movement associated with the opening and closing of the activation gates of the Na channels. J. Gen. Physiol. 63, 533. Armstrong, C. M., and Ciilly. Wm. F. (1979). Fast and slow steps in the activation of sodium channels. J . G p r i . Physiol. 74, 691. Haumann, G . , and Mueller, P. (1974). A molecular model of membrane excitability. 1. Suprumnl. Strucr. 2, 538. Colquhoun, D.. and Hawkes, A. G . (19x1). On the stochastic properties dsingle ion chiinnels. Prof,. R. Soc. Londorr S r r . B 211, 205. Fifzhugh, R . (196.5). A kinetic model of the conduct:incc changes in nerve menibriine. J . Cell. C'omp. Pl1ysiol. Mi (Suppl. 21, 11 1. Hodgkin, A . I.., and Huxley. A . F. (1952). A quiintit;itive description of membranc current and its applicalion t o conduction and excitation in nerve. J. Physiol. (I.oridorc) 117,

SO. Horn, K.. and Lange, K. (1983). Estimating kinetic constants liom single channel data. niop/iy.s. J . 43, 201. Hoyt, R. (1971). Independence of the bodium and potassium conductance channels, a kinetic argument. Biophys. J . I t , 110. Keynes, R., and Rojas, E. (1974). Kinetics and stexdy state properties of the charged system controlling sodium conducldnce in the squid giant axon. J. Pliysiol. (Loridort)233,28P. Patlak, J., and Horn, R. (1982). The effect of N-bromoacetamide on single sodium channel currents in excised membrane patches. J. Gen. Yhysid. 79, 333. Sigworth, F. .I..and Neher, E. (1980). Single Na+ channel currents observed in cultured rat muscle cells. Nutitre (London) 287, 447.

CURRENT TOPICS IN MEMBRANES A N D TKANSPOKI. VOLUME 22

Multi-Ion Nature of Potassium Channels in Squid Axons TED BEGENISICH A N D CATHERINE SMITH Depurtment of Physiology Universiry o j Roc,hester School of Medicine and Dentistry Rochester, New York

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Characteristics of Multi-Ion Pores . . . . ...................... A. Concentration-Dependent Charact ...................... B. Pore Characteristics in the Presence of Blocking Ions. ............. C. Unidirectional Fluxes through Pores . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

.................... B. Blocking Ions . . . . . . . . . . . .

3.55 355 357 357 358

. . . . . . . . . . . . . 359

A Mathematical Model .............................. A. Current vs Concentration,. ......................................... B . Block by Cesium Ions.. ............................. C. Unidirection Flux Ratio . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. More on Energy Barriers . . . . ........................

IV.

References . . .

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

I.

353

363 364 365 365 366

368

INTRODUCTION

The squid axon has played a pivotal role not only in the study of electrical excitability, as described in several articles in this volume, but also in the study of ionic permeation. Hodgkin and Katz (1949) applied the theory of Goidman (1943) to the squid axon and demonstrated changes in the relative sodium and potassium permeabilities during the action potential. One of the results of this theory is Eq. ( l ) , which describes how the 353 Copyright Y 1984 by Academic Press. Inc. All rights of reproductlon in any form reserved. ISBN o - i ? - i c m ~ - n

354

TED BEGENlSlCH AND CATHERINE SMITH

current through an ionic pathway carried by a cation X ( I , ) depends on membrane potential, V,.

In this equation P , is the permeability of ion X ; [XI,, and [Xli are the external and internal concentrations (activities) of ion X. R , T , and F have their usual thermodynamic meanings. If t h e pathway is permeable to two cations, the potential at which the net current is zero is given by Eq. (2).

The derivation of these equations requires several assumptions, the most fundamental of which is that the ion movement obeys the independence principle. Hodgkin and Huxley (1952) gave a concisc definition of independence: “The chance that any individual ion will cross the membrane in a specified interval of time is independent of the other ions which are prescnt.” Since this definition applies to the diffusion of ions in aqueous solutions, ion permeation models based on this principle can be called ‘‘free-diffusion’’ models . While Eq. (1) and (2) are concerned with net ionic currents, there is another equation [Eq. (3)] involving unidirectional fluxes that must be obeyed by ion pathways that allow independent ion movement. MJM, = exp(V, - Vx)FIRT

(3)

In Eq. (3) M e is the efflux of the ion kinder study and M ,is the corresponding influx; Vx is the equilibrium or Ncrnst potential for this ion. l h i s equation has been derived by Ussing (1949) and Hodgkin and Huxley (1952) and is a special case of the more general treatment of Tcorell ( 1949).

These three equations have, until recently, formed the theoretical basis for our understanding of ionic permeation in excitable cells. That we have relied so heavily on these equations is, in retrospect, wrprising, since almost 30 years ago the potassium channel of Sclpiu (first cousin to squid) was shown not to pass the flux ratio test represented by Eq. ( 3 ) (Hodgkin and Keynes, 1955). Hodgkin and Keynes found that their data could be described by this equation only if the right-hand side was raised to a power near 2.5-a clear violation of independence. This parameter is usually called the flux ratio exponent. These authors were also the first to explain the deviation from independence by considering that K + ions moved through a very narrow pore in “single file.” They constructed an

POTASSIUM CHANNELS IN SQUID AXONS

355

ingenious mechanical model to simulate their results, and they derived a mathematical model as well. In the last 30 years much progress has been made (especially by Heckmann, 1965a,b, 1972) in the theoretical understanding of ion permeation in narrow pores. This effort culminated in what is already a classical report by Hille and Schwarz (1978). These authors described several types of experiments that could be used to distinguish among various models of ion permeation. In particular they showed the special behavior to be expected from multi-ion, singlefile pores. This article summarizes these theoretical predictions and discusses the related experimental data on squid axon potassium channels responsible for the repolarization of the nerve action potential. Often called the “delayed rectifier,” these channels can be distinguished from other conductances by their strong sensitivity to membrane potential, by high ionic selectivity, and by their reaction to various pharmacological agents. These potassium-selective channels (or pores) exhibit many of the permeation characteristics of multi-ion pores. In addition to a review of the data, we present a simple model of this potassium pore that accounts, at least qualitatively, for the wide range of experimental results. II. CHARACTERISTICS OF MULTI-ION PORES

Table I [adapted from Table I of Hille and Schwarz (1978)l summarizes some of the various permeation properties expected of pores obeying independence (i.e., free-diffusion pores), one-ion pores, and pores simultaneously containing as many as m ions. Some of these predictions apply only to special circumstances, usually symmetric ion concentrations. It will be important to realize such limitations when comparing these predictions to experimental results. A. Concentration-Dependent Characteristics

The zero voltage slope conductance of a pore that can simultaneously contain up to m permeant ions in a membrane bathed by equal concentrations of the permeant ion has rather remarkable properties. The conductance will increase at first linearly with permeant ion concentration. There may then be up to rn - 1 maxima in the conductance as the concentration is raised further. Finally, at very large concentrations the conductance may actually decrease as all sites become filled and the permeant ions block their own passage through the pore. The relative permeability of two different ions in a multi-ion pore also has rather special properties. The usual method of obtaining the relative

SOhlE

TABLE I PERhiEATlON PROPERTIES OF CERTAIN TYPES OF

PORES"

Pore type Permeation property

Free diffusion

One-ion

rii-ion

Pot ass i u rn Two components

Armstrong (personal communication)

sort of

Oxford and A d a m (personal communication) Adelman and French (1978)

Conductance v5 concentration

Linear

Saturating

Concentration-dependent permeability ratio Power dependence of blocker concentration Effective valence of monovalent blocker Change of current from side with blocker when permeant ions are added to opposite side

No

No

Up to m-1 maxm a and finally self-block Yes

First

First

Up to m'th

Second (Csl

05Z'5I

0CZ'S I

OSZ'

0.6

None

Decrease

Possible increase

Increase

Flux vs trans concentration

None

Decrease

Decrease

Decrease

Flux ratio exponent

n'

n' = 1

I s n ' 5 m

ti' =

(' Adapted from Table

=

1

I of Hille and Schwarz (1978).

5 m

5

Z'

2.5 1.6 c n'

5

5

1.3

3.3

Keferences

Adelman and French (1978) Adelman and French (19781. Bezanilla and Armctrong (1972). Armstrong and Taylor (1980) Hodgkin and Keynes (1955). Begenibich and De Weer (1977) Hodgkin and Keynes (1955) Begenisich and De Weer (1980)

POTASSIUM CHANNELS IN SQUID AXONS

357

permeabilities for two cations is to bathe the membrane with ions of one type on one side and ions of another type on the opposite side. Then the zero-current potential is measured and the ratio of the permeabilities is determined from Eq. (2). This ratio is independent of ion concentration in both free-diffusion pores and one-ion pores (although it may be voltage dependent). In multi-ion pores the permeability ratio is, in general, a function of the concentration of the two ionic species. B. Pore Characteristics in the Presence of Blocking Ions

Another class of experiments useful in studying ion permeation properties makes use of blocking ions: ions that are impermeant (or sparingly permeant) and block the passage through the pore of permeant ions. In multi-ion pores several blocking or permeant ions may have to move to new locations within the pore before the blocking particle can reach its blocking location. This gives the block some of the properties expected if the block were produced by a multivalent multimer of the blocking particle. Specifically, the block may depend on the blocking ion concentration raised to a power greater than one and up to a power of m. Additionally, the voltage dependence of the block produced by, for example, a monovalent ion may be more steeply dependent on voltage than the maximum of an e-fold change for 25 mV. This effect can be described by the effective valence of the blocker, which can attain a value of up to m ,the number of ions allowed in the multi-ion pore. Another distinguishing feature of multi-ion pores occurs when the concentration of permeant ions on the side of the membrane opposite to that of the blocker is raised. In free-diffusion pores this would have no effect on the ionic current. In one-ion pores the current from the side with blocker is expected to decrease as the pore is occupied by the permanent ions. In multi-ion pores the current may increase as the increased occupancy of the pore by the permeant ion also clears the pore of blocking ions. C. Unidirectional Fluxes through Pores

Other interesting properties of multi-ion pores are revealed when unidirectional fluxes, rather than net flux or current, are considered. For example, in free-diffusion pores the efflux of an ion will be independent of the external concentration of that ion. For both one-ion and multi-ion pores, the efflux is expected to decrease with increasing external concentration. Perhaps the most revealing feature of multi-ion pores can be seen in the flux ratio exponent, n’. This parameter has a value of one for both free-

358

TED BEGENISICH AND CATHERINE SMITH

diffusion and one-ion pores, but can be as large as the maximum number of ions allowed in a multi-ion pore. The particular value of n’ for a muitiion pore depends upon, but is not directly related to, the number of ions in the pore. Since occupancy is both concentration and voltage dependent, a’ will also, in general, be a function of these parameters. 111.

EXPERIMENTAL RESULTS

A. Concentration-Dependent Effects

In contrast to the careful analysis in myelinated nerve (Hille, 1973) relatively little work has been done on K channel selectivity in squid axons. It has been known for some time that the potassium channels of squid are permeable to Rbf ions (Pickard ~t al., 1964). In fact, PRhIPKis near I .0, as determined from reversal potential measurements with exter1966). At about this nal ion replacement in intact axons (Moore rt d., same time Binstock and Lccar (1969) showed that NHd ions were also permeant, but less so than K + ions. From this period until the present there was no systematic quantitative investigation of potassium channel selectivity in squid. In 1981 Oxford and Adams reported the sequence of relative permeabilities as TI+ > K + > Rb+ > NH: > Lit > Na+. I n additional unpublished experimcnts these authors obtained a value of PNH41PK of 0.14 from the change in zero current potential when 300 mM internal potassium was replaced by 300 mM NH4 with 220 m M external potassium. The reciprocal experiment of replacing external potassium whilc keeping ammonium with internal potassium constant yielded a permeability ratio of about 0.5. Similar results were obtained with rubidium. This type of behavior does not occur in pores obeying independence. It is certainly the kind of result expected of multi-ion pores. However, since the ionic conditions used on these experiments produce substantial changes in zero current potential, it is possible that the change in the permeability ratio is an indirect effect of voltage-dependent permeability ratios in a one-ion pore. More experiments need to be done to eliminate this latter possibility. There has been even less work on how potassium channel current or conductance depends upon the K + ion concentration. C. M . Armstrong (personal communication) has remedied this situation, with data of the type shown in Fig. 1A. Plotted in this figure is potassium channel current as a function of internal K + concentration. The current increases at first steeply with K + concentration and then gradually approaches an approxi-

POTASSIUM CHANNELS IN SQUID AXONS 1.6

1.8

a

Y

u

5 Y

B I

c

FIG. 1. Potassium channel current as a function of the concentration of internal potassium. ( A ) Data points are from axon JLO82P of C. M. Armstrong (personal communication). The outward potassium channel currents at V , = 120 mV are plotted versus internal K concentration. The external solution contained no potassium. The values have all been normalized to the current with 100 m M K. The solid lines represent “best” nonlinear leastsquares fits to saturating hyperbolas with (curve 2) and without (curve I ) a linear cornponent. (B) Same as (A), but the solid line was computed from the four-barrier, three-site model.

mate linear phase. The solid lines in this figure represent nonlinear leastsquares fitted to a rectangular hyperbola and to the sum of a rectangular hyperbola and a linear function. The two-component fit is clearly better than a single saturating function. Figure 1B shows the same data as Fig. IA, but the solid line in this figure is from the four-barrier, three-site model described below.

B. Blocking Ions

Several ions can block the current through squid axon potassium channels. Internal Rb+ ions, even though fairly permeant as deduced from reversal potential measurements, reduce potassium-channel currents (Chandler and Meves, 1965). This result explains the older finding of Baker et al. (1962) that internal Rb+ ions prolong squid axon action potentials. Rubidium is an example of a relatively permeant blocker. Internal sodium ions are also blocking ions that can permeate the potassium channel,

360

TED BEGENISICH AND CATHERINE SMITH

but only at rather large potentials (Bezanilla and Armstrong, 1972; Chandler and Meves, 1965; French and Wells, 1977). The most carefully studied potassium-channel blocking ion is cesium. This ion can block from either side ofthe membrane (Adelman and Senft, 1966; Adelman and French, 1978) and is probably slightly permeant, but even less so than sodium (R. French, personal communication). In their exccllcnt study using external Cs' ions as potassium channel blockers, Adelman and French (1978) found that the block could not be described at all potentials by the binding of one cesium ion per channel. Rather, a two-cesium : one-channel stoichiometry was required at negative potentials and a 1 : 1 stoichiometry at more positive voltages. A second power of the Cs concentration inherent in the 2 : 1 stoichiometry would be expected if the potassium pore allowed occupancy by at least two ions. A change in the power-dependence is expected as voltage and concentration alters the degree of occupancy. At positive potentials and low cesium concentrations, there may be little occupancy, so a I : 1 stoichiometry is expected. At more negative potentials and higher cesium concentrations, the multi-ion nature is revealed by higher Cs : channel stoichiometries. Dose-response data for internal Cs+ block at four values of membrane voltage as determined by Adelman and French (1978) are plotted in Fig. 2A. Shown along with the data points are t h e predictions of the 2 : 1 Cs : K pore stoichiometry at -106 and -86 mV, and of the I : 1 stoichiometry at -46 and -26 mV. Shown in Fig. 2B are the computations from the model described later. One of the striking results obtained by Adelman and French (1978) was that the voltage dependence of the block produced by 200 mM Cs' was equivalent to this ion moving 130% across the membrane voltage. 'They also showed that the effective valence of Cs+ was a function of C s ' Concentration, ranging from a value of 0.6 at 5 mM Cs+ to 1 .O at 20 niM and finally 1.3 at 200 m M C s ' . These remarkablc results are simply the consequences expected from a pore that can simultaneously be occupied by several ions. In a study using internal tetra-N-alkylammonium ions as blockers, French and Shoukimas (1981) showed a deviation from a simple I : I stoichiometry at high concentrations of blocking ions. While they did not analyze this point in detail, the explanation may be the same as that for external C s + :the multi-ion nature of the K' pore. The block of K channels by external Cs ' is rcduced by increasing the internal K' concentration as shown in Fig. 3A. This figure replots data from Adelman and French (1978) for 200 mM external Cs+ and two concentrations of internal potassium of 265 mM (filled squares) and 150 m M +

FIG.2. Concentration dependence of blockage of potassium channel current by external cesium. (A) Data are from Adelman and French (1978). The ordinate represents current in the presence of various concentrations of cesium relative to current in the absence of cesium. The abscissa values are external cesium concentrations on a logarithmic scale. Four values of membrane potential are represented by the four symbols: V , = - 106 mV (A), -86 (H), -46 (V),-26 ( 0 ) Standard . error limits are plotted except where these are smaller than the size of the symbol. The solid lines for - 106 and -86 mV were computed from a 2 : I Cs: K pore stoichiometry, the lines at -46 and -26 mV from a 1 : I stoichiometry. ( B ) Same as (A), but the solid lines are from the four-barrier, three-site permeation model. The external and internal K concentrations were 240 and 350 mM, respectively. A

WKI 1=15(mM B:Kl I = ? 6 h M

-1

-

FIG.3. Potassium-channel current in the presence of 200 mM external cesium at two different concentrations of internal potassium. (A) Data redrawn from Adelman and French (1978). The solid lines in panel A have no theoretical meaning. (B) Calculations from the four-barrier, three-site model simulating the experiment shown in panel A. The external concentration is 240 mM and internal K concentrations were 340 and 350 mM, respectively.

362

TED BEGENISICH AND CATHERINE SMITH

(filled circles). Raising the concentration of internal potassium clearly reduces the amount of block produced by cesium. Shown in panel B of Fig. 3 is the simulation of this experiment with the four-barrier, three-site model described below. This ability of internal K+ to increase net inward current through the potassium channel in the presence of external cesium demonstrates that the block occurs within the K‘ permeation pathway. The observed increase is not consistent with free diffusion pores. It is, however, one of the properties of multi-ion pores in the presence of blocking ions. Armstrong and Taylor (1980) showed that this effect can occur with blockers on either side of the membrane. In their experiments Ba’+ ions were used as blocking ions, and it was found that block by internal Ba2+ was reduced by external K+ and that internal K + inhibited the block produced by external Ba2+. C. Flux Studies

In their historic study of unidirectional fluxes through the potassium channel of S q i u axons, Hodgkin and Keynes (1955) found that the efflux of potassium ions was inhibited by external K+ ions, More recently, Begenisich and De Weer (1977) used internal perfusion and voltage-clamp techniques and showed that this inhibition was a function of membrane potential. This type of behavior cannot occur through free-diffusion pores where ions move independent of each other, but is consistent with both one-ion and multi-ion pores. The multi-ion nature of a pore is probably most clearly revealed by a measurement of the ratio of the unidirectional fluxes. A determination of a flux ratio exponent greater than one identifies the pore under question as a multi-ion pore. In their ingenious experiments on Sepia, Hodgkin and Keynes (1955) found n’ values near 2.5. Begenisich and De Weer (1980) showed that for squid axon potassium channels n’ was a function of membrane potential and internal K+ concentration. As described above, n’ will be influenced by the degree of occupancy of the pore, which in turn depends upon concentration and potential. Begenisich and De Weer (1980) found n’ (near zero mV) to be approximately 2.3 for high internal K+ and about 1.5 for low internal K +.These values increase as the potential is made more negative and can reach values as large as 3.3 near -40 mV with a high internal K+ concentration. However, because of the technical difficulties introduced by ion accumulation in the periaxonal space, the precise values of these numbers must not be taken too literally. Nevertheless, it is clear that n‘ for the potassium pore

POTASSIUM CHANNELS IN SQUID AXONS

363

in squid exhibits values much larger than unity and these values are probably functions of concentration and membrane potential.

IV. A MATHEMATICAL MODEL The results described above are clearly consistent with the general expectations of a multi-ion pore, not with those of one-ion or free-diffusion pores as indicated by the entries in Table I for potassium channels. Nevertheless, it remains to be seen whether or not all the various results can be duplicated by a specific multi-ion pore model. This section describes a four-barrier three-site model for potassium ion permeation and for block produced by cesium ions. Hille (1975), Hille and Schwarz (1978), and Begenisich and Cahalan (1980) have discussed the mathematical techniques used to construct models of ionic pores. In these models ions “jump,” according to a Poisson process, from site to site over an intervening energy barrier. The jump frequency decreases exponentially with the free-energy difference be1949). tween the site and the top of the barrier (Eyring et d., It is clear from the results summarized in Table I that the potassium channel in squid axon membranes is a multi-ion pore. The cesium data might be described by a two-site model but such a model would provide a maximum flux ratio exponent of two. A three-site model would allow n’ values up to three which, given the experimental uncertainties in this parameter, may be sufficient. A three-site model that allows all three sites to be occupied by any combination of two different ions present on both sides of the membrane has 27 states. These states are connected by 96 forward and reverse rate constants. However, most of the rate constants are not independent; rather, they are linked through the values of the free energy barriers and wells. In a three-site model there are for each type of ion four energy barriers and three wells, and thus seven parameters placing the barriers and wells along the electric field. To reduce the number of adjustable parameters, we have assumed symmetrically shaped, equally spaced barriers and wells. We additionally constrained the potassium barriers to be symmetric (at zero applied potential) about the middle of the pore. This proved to be too much a constraint for Cs, so the two external Cs barriers were made equal (see Fig. 4). We found it necessary to include also one other adjustable parameter representing ionic repulsion within the pore. There are several ways in which this can be done within the constraints of microscopic reversibility (Hille and Schwarz, 1978). We have chosen to assume that the repulsion

364

TED BEGENlSlCH AND CATHERINE SMITH

10

K+ 10

-9

-11.5

FIG.4. Free-energy profiles for K and Cs ions, The values for the peaks and valleys are energies in units of RT. The abscissa represents fractional distance along the membrane potential gradient.

occurs only between ions in adjacent sites. This repulsion is represented by a factor t.' similar to that used by Hille and Schwarz (1978). These authors also discussed possible values for F ranging from I to 100. We found a rather modest value of 4 to be sufficient (and necessary) for a reasonable description of the experimental data. Figure 4 shows the free-energy diagrams (at zero applied membrane potential) for K + and Cs+ used to simulate the experimental data. We have not attempted a quantitative fit owing both to experimental uncertainties and computational tcdium. Nevertheless, the calculations using this model demonstrate that a single set of barriers can indeed duplicate, at least qualitatively, a wide variety of experimental data. A. Current vs Concentration

Figure IB illustrates the relationship between current and K + concentration for this model pore along with the experimental data (C. M. Armstrong, personal communication). It was mainly the attempts to fit these

POTASSIUM CHANNELS IN SQUJD AXONS

365

data that required the presence of ionic repulsion within the pore. With no repulsion, the early increase in current with concentration could be duplicated but the current saturated at quite low concentrations. The presence of ionic repulsion in the model spreads out the concentration range over which the various sites become filled with ions. The result is analogous to a sum of saturable binding curves with different values of dissociation constant. Increasing the amount of repulsion by using larger F values improves the fit in the high concentration region. 8. Block by Cesium Ions

Figure 2B shows a plot of the amount of potassium pore current blocked as a function of Cs' concentration. The solid curves are the prediction from the three-site model at four values of membrane potential. The fits are reasonable but not as good as those shown by the solid lines in panel A. However, in panel A the stoichiometry was allowed to change as a function of voltage. A fixed stoichiometry cannot describe the data at all potentials. Figure 3B represents the model current-voltage relation of the potassium channel pore in the presence of external cesium. Shown are computations for two values of internal potassium, simulating the experimental results of Fig. 3A. The model is able to reproduce the increase in inward potassium channel current (at negative voltages) produced by elevated internal potassium levels. Figure 5 illustrates the expected block of potassium channel current by internal Cs+ ions. The inset plots the data on an expanded current scale and demonstrates a slight negative resistance region and then an increase in current as Cs+ ions are cleared from the channel by the large potential gradient. This result is similar to results with internal Cs+ reported by Adelman (1971) and Bezanilla and Armstrong (1972) and to data with internal Na+ ions obtained by French and Wells (1977). Although not shown, the model predicts the known ability of external Kf ions to reduce the block. C. Unidirection Flux Ratio

The four-barrier, three-site permeation model has so far been able to reproduce qualitatively the significant features of potassium channel permeability and block by Cs+ ions. The computed values of n' for this model with 35 mM external and 350 mM internal potassium are 2.57 and 2.6 at potentials of zero and -40 mV, respectively. The values are certainly in

366

TED BEGENISICH AND CATHERINE SMITH

0.4 x lo5,

-50

-0.5

x 105

1

V,(mV)

100

1 ccs31= OmM 2 CCsIi=SCmhf

FIG. 5 . Potassium channel current and internal cesium. Calculations from the fourbarrier, three-site model are shown with and without 50 m M internal cesium. The inset shows the same calculations on an expanded current scale illustrating a small negative resistance region and a further increase in potassium channel current at larger potentials. The external and internal K concentrations were 100 and 275 m M , respectively.

the range reported by Hodgkin and Keynes (1 955) and Begenisich and De Weer (1980),but the voltage dependence is less steep than the experimental values. The model does predict slightly decreasing values of a’ with lower internal K concentrations. The n’ value at zero mV is reduced to 2.49 as internal Kf is lowered from 350 to 200 mM. +

V.

MORE ON ENERGY BARRIERS

The free-energy profile (in the absence of an external electric field) of ions in pores arises from at least three sources: the so-called “image” potential, hydration energies, and chemical binding energies. Figure 6 shows one way in which these three types of energies can combine to produce the total free energy diagram for K+ ions that was used here. An “image” potential results when an ion is near the interface between

367

POTASSIUM CHANNELS IN SQUID AXONS HYDRATION

v

-21.1

FIG.6 . Image, binding, and hydration energies. Similar to Fig. 4.

two regions of different dielectric constant, in this case water and membrane. The ion induces an electric field in the surrounding media such that the ion is in a lower energy state near the interface than in the middle of the pore. This energy is illustrated in Fig. 6 by the curve labeled IMAGE. Since potassium channels have both high selectivity and high ion permeation rates, ions probably go through the pore neither fully hydrated nor fully dehydrated. Rather, some waters of hydration are probably lost upon entering the pore perhaps owing to physical constrictions at either end. The middle of the pore may be wide enough to allow more association between the ion and water than the narrow regions. The barriers labeled HYDRATION in Fig. 6 represent this simplified situation. Ions no doubt interact with some chemical constituents of the pore. Indeed, external and internal hydrogen ions bind to potassium channels with pK values of 4.5 and 6.5, respectively. These chemical groups may also bind K + , Cs+, and other cations. These binding energies are labeled CHEMICAL in Fig. 6. The sum of the three types of energies described above is shown in Fig. 6 as the profile marked TOTAL. This is exactly the same profile as

368

TED BEGENISICH AND CATHERINE SMITH

that shown for K+ ions in Fig. 4. There may be other energies involved in ion translocation including entropic energies. Neverthelcss, the simple ideas sketched here show how the energy barrier profiles that describe reasonably well the available data can be constructed from the various underlying physical processes.

VI. SUMMARY AND CONCLUSIONS Column 5 of ‘Table I summarizes the known permeation properties of the potassium channels in squid giant axons. A picture of this channel as a multi-ion pore clearly emerges. The various figures showing the computations of a particular four-barrier, three-site model illustrate that this rather simple pore model can readily reproduce the experimental data in a qualitative and, in many cases, a quantitative manner. ACKNOWLEDGMENTS We thank Clay M. Armstrong. Robert French. and Gerry Oxford for providing unpublished data without which this article would have been incomplete. We also thank Karen Vogt, Wendy Keck, and Marlene Bauer for their processing of words. This work was in part supported by USPHS Grants NS-14138 and NS-00322 to T. Begenisich.

REFERENCES Adelman, W . I . , JI-. (1971). Electricid studies of inlcrnally pelfused q u i d axuns. f i r “Biuphykics and Physiology o f Excitable Memhi-anes” ( W . J . Adelman, Jr. , Ed.). pp. 274319. Van Nobtimd Rcinhold, New York. Adelman, W. J . , Jr., and French, H. J . (1978). Blocking of the squid axon potassium channel by external caesium ions. J . Physicil. (1,onclon)276, 13-25. Adelman, W. J., Jr., and Senft, J. P. (1966). Voltage clamp studies on the effect of internal cesium ion on sodium and potassium currents in the squid giant axon. 1.Gen. Physiol. SO, 279-293. Armstrong, C. M., and Taylor, S . R. (1980). Interaction of barium ions with potassium channels in squid giant axons. Biophys. J. 30, 473-488. Baker, P. F., Hodgkin, A . L., and Shaw, T. I . (1962). The effects of changes in internal ionic concentrations on the electrical properties of perfused axons. J . f h y s i o l . (London) 164, 355-314. Begenisich, T., and Cahalan, M. D. (1980). Sodium channel permeation in squid axons. I . Reversal potential experiments. J . Physiol. (London) 307, 217-242. ch, T., and De Weer, P. (1977). Ionic interactions in the potassium channel of squid nt axons. Nutitre (London) 269, 710-71 I . Begenisich, T., and De Weer, P. (1980). Potassium tlux ratio in voltage-clamped squid giant axons. J . Gen. Physiol. 76, 83-98. Rezanilla. F., and Armstrong, C. M. (1972). Negative conductance caused by the entry of sodium and cesium ions into the potassium channels of squid axons. J . Gen. Physioi. 60, 588-608. Binstock, L.. and Lecar, H. (1969). Aminonium ion currents in the squid giant axon. .I Gen. . Physiol. S3, 342-361.

POTASSIUM CHANNELS IN SQUID AXONS

369

Chandler, W. K., and Meves, H. (1965). Voltage clamp experiments on internally perfused giant axons. J . Physiol. (London) 180, 788-820. Eyring, H., Lumry, R., and Woodbury, J . W. (1949). Some applications of modern rate theory to physiological systems. Rec. Chem. f r o g . 100, 100-114. French, R., and Shoukimas, J . J. (1981). Blockage of squid axon potassium conductance by internal tetra-N-alkylammonium ions of various sizes. Biophys. J. 34, 271-291, French, R., and Wells, J. B. (1977). Sodium ions as blocking agents and charge carriers in the potassium channel of the squid giant axon. J . Gen. Physiol. 70, 707-724. Goldman, D. E. (1943). Potential, impedance, and rectification in membranes. J . Gen. Physiol. 27, 37-60. Heckmann, K. (196Sa). Zur Theorie der “single-file”-diffusion. Z . Plivs. Chem. 44, 184203. Heckmann, K. (1965b). Zur Theorie der “single-file”-diffusion. Z . Phys. Chem. 46, 1-25. Heckmann, K. (1972). Single file diffusion. I n “Biomembranes” (F. Kreuzer and J. F. G . Slejers, eds.), Vol. 3, pp. 127-153. Plenum, New York. Hille, B. (1973). Potassium channels in myelinated nerve. Selective permeability to small cations. J . Gen. Physiol. 61, 669-686. Hille, B. (1975). Ionic selectivity, saturation and block in sodium channels. A four barrier model. J . Gen. Physiol. 66, 535-560. Hille, B., and Schwarz, W. (1978). Potassium channels as multi-ion single-file pores. J . Gen. Physiol. 72, 409-442. Hodgkin, A. L., and Huxley, A. F. (1952). Currents carried by sodium and potassium ions through the membranes of the giant axon of Loligo. J . Physiol. (London) 116,449-472. Hodgkin, A. L., and Katz, B. (1949). The effect of sodium ions on the electrical activity of the giant axon of the squid. J . Physiol. (London) 108, 37-77. Hodgkin, A . L., and Keynes, R. D. (1955). The potassium permeability of a giant nerve fibre. J . Physiol. (London) 128, 61-88. Moore, J . W . . Anderson, N.. Blaustein. M . . Takata. M., Lcttvin. J . Y . . I’ickurd. W . F.. Bernstein. T.. and Pooler, J . ( 1966). Alkali cation selectivity of squid axon membrane. Ann. N . Y . A c d . S I . ~ 137, . 818-829. Oxford, G., and Adams, D. J . (1981). Permeant cations alter K channel kinetics and permeability. Biophvs. J . 33, 70a (Abstr.). Pickard, W. F., Lettvin, J . Y.. Moore, J . W . . Takiita. M., Pooler, J . , and Bernstein. T. (1964). Cesium ions do not pass the membrane of the giant axon. Pro(.. Ntrrl. Ac,trtl. Sci. U.S.A. 52, 1177-1183. Teorell, T. (1949). Membrane electrophoresis in relation to bio-electrical polarization effects. Arch. Sci. Physiol. 3, 205-218. Ussing, H. H . (1949). The distinction hy means of tracers between active tranhport and diffusion. Ac,f(i Physiol. Sccind. 19, 43-56.

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CURRENT TOPICS IN MEMBRANES AND TRANSPORT, VOLUME 22

Noise Analysis and Single-Channel Recordings FRANC0 CONTI lstiruto di Cibernerica e Biojisicu Consiglio Nuzioncile delle Ricwche Camogli, Italy

I.

Introduction . . . . . . .

11. Basic Concepts of FI 111. The Physical Basis of Membrane Noise A. Equilibrium N o i s e . . . .

B. Transport Noise.. . . . . C. Channel N o i s e . , . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Extrinsic Channel Noise . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1V. Channel Noise in Nerve Membranes.. .................................... A. K Channels. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9. Na Channels.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Single-Channel Recordings .......................................... References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1.

3x0 382 3x5 385 392 397 40 I

INTRODUCTION

There is now abundant evidence that the permeability changes underlying nerve excitation involve a relatively low density of physically distinct and noninteracting ionic pathways, generated by specific proteins embedded in a common lipid matrix. This notion has been quantified in the last 15 years through estimates of ionic-channel densities and single-channel currents, obtained with four different techniques. In addition to binding studies (Moore er af., 1967; Ritchie and Rogart, 1977; Strichartz er al., 1979) and to gating-current measurements (Armstrong and Bezanilla, 1973; Almers, 1978; Meves, 1978), fluctuation analysis and single-channel recordings have provided some of the most convincing data. Fluctuations have long been understood in physics as the macroscopic 371 Copyright D 1984 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-153322-0

372

FRANC0 CONTl

manifestation of the discreteness of matter (Lax, 1960). The first unequivocal correlation between electrical fluctuations and quantal membrane conductance changes was demonstrated in the postsynaptic membrane of the frog neuromuscular junction (Katz and Miledi, 1970; Anderson and Stevens, 1973), where ionic channels are primarily gated by their interaction with a chemical agonist. The measurements, yielding an estimate of about 20 pS for the quantal conductance step, provided the strongest evidence that synaptic channels are aqueous pores. Since then, noise analysis has become a standard method of investigation of agonist-receptor interactions in postsynaptic membranes. The theory and application of fluctuation studies in this field have been reviewed elsewhere (Neher and Stcvens, 1977; Colquhoun and Hawkes, 1977). I n this article we consider the application of noise measurements to the study of thc ionic channels which are responsible for the voltage modulation of the conductance of axonal membranes. Recordings of single-channel events provide the most direct evidence of the discrete nature of membrane permeability changes. Such recordings were first obtained from artificial lipid bilayers containing very low concentrations of extrinsic ionophores (Bean et d., 1969; Ehrenstein and Lecar, 1977). The direct observation of single-channel events in natural membranes (Neher and Sackmann, 1976) is one of the greatest achievements in electrophysiology after the classical work of Hodgkin and Huxley (1952). The application of patch recording techniques to the study of ionic channels in the squid axon membrane (Conti and Neher, 1980) is discussed in Section V of this article. II. BASIC CONCEPTS OF FLUCTUATION ANALYSIS

The full appreciation of noise analysis requires some knowledge of the theory of stochastic variables, as treated by several textbooks (e.g., Papoulis, 1965; Bendat and Piersol, 1971). Outlined below are the minimal basic concepts needed for understanding the general results. Fluctuations of a physical observable originate from the random nature of the microscopic contributions to its overall value. Their analysis involves primarily measurements of mean square deviations and time correlations, which provide information about the size and the temporal characteristics of the microscopic events. This information is easily derived from simple properties of random variables, if the elementary contributions are statistically independent. With reference to the particular system that concerns us, let I be the

NOISE ANALYSIS AND SINGLE-CHANNEL RECORDINGS

373

electric current flowing through a membrane as a result of the contributions, i G ) ( j = 1 , ..., N ) , from N ionic channels: N

Let us denote expected values with angle brackets, ( ). If the channels are all identical and statistically independent, the mean of I , P I , and its uariance, af,are given by Eqs. (2) and (3). N

N

af = ( ( I - pI)*)=

,C = ((i(J)- p l ) * )= NUT 1

(3)

where pl and af are the mean and the variance of single-channel currents. Dividing Eq. ( 3 ) by Eq. (2) term by term yields vflp,= a:/p]

(4)

Equations (1)-(4) are valid at any time, as long as the means are identified with overall averages over a large number of observations repeated under identical macroscopic conditions. However, for stationary conditions the means are time-independent , and they can be approximated by appropriate time averages. The information provided by vT/p, depends on the particular structure of single-channel current fluctuations, although it can be stated quite generally that this quantity is proportional to the size of single-channel currents. Indeed, scaling the latter by a factor produces the same scaling of v.?/p,. The correlation between fluctuations at any two instants in time, tl and f 2 , is characterized by the autocovariance function, TI(tl, t 2 ) , defined by

rdrl rl(tl,f 2 )

7

h ) = (“(tl) - pdt1)ll~(f2)- Pl(f2)l)

(5)

Note that = rl(f2, t l ) . Furthermore, in stationary conditions rI(tl,t 2 )depends only on the absolute value of ( f 2 - tl). The time correlation of the fluctuations is then characterized by an even function of time,

W).

w)= wl,tl + t )

(6)

An important concept of fluctuation analysis is the recognition of a strict analogy between the time correlation of spontaneous fluctuations of any observable and its macroscopic relaxation following an initial displacement of the system from the stationary state (Onsager, 1931; Lax,

374

FRANC0 CONTI

1960; Kubo, 1975). For most physical systems, both these processes are described by a s u m of linear relaxation components and C , ( t )is the sum of several exponentially decaying functions with different time constants. In the analysis of stationary fluctuations, C,(t)can be estimated from time averages over samples of random signals having much longer duration than the characteristic correlation times of the fluctuations. An equivalent characterization is obtained through the power spectrum, S , ( f ) , measured by the square modulus of the Fourier spectrum of the signal at any frequency, f.The latter quantity is directly related to C,(r) by

The power spectra most frequently encountered in fluctuation analysis are those associated with linear relaxation processes and described by the Lorentzian function, L ( f ; 7). L(,f,7) = 47/11 i- ( 2 7 r ~ . f ) ~ ]

(8)

which can be obtained from Eq. (7) when C , ( t ) = Oi7.

111.

THE PHYSICAL BASIS OF MEMBRANE NOISE

The application of fluctuation analysis to the study of ion transport in biological membranes has been treated in several reviews (Verveen and DeFelice, 1974; Conti and Wanke, 1975; Neher and Stevens, 1977; Colquhoun and Hawkes, 1977; DeFelice, 1977). This section summarizes the basic physical principles underlying the interpretation of the membrane noise measurements discussed later. Membrane noise will in general contain several components that are not simply related to the discrete nature of ionic channels. An understanding of all possible noise sources is therefore essential for the unambiguous interpretation of membrane noise in terms of channel fluctuations. The statistical nature of all microscopic phenomena that contribute to the macroscopic properties of a biological membrane is always governed by the general laws of thermal equilibrium between any elementary cornponent and its local environment. However, for the sake of discussion, we can distinguish three major categories of membrane fluctuations, arising from distinct physical mechanisms: equilibrium noise, transport noise, and channel noise. The only reservation about this classification is that, in general, it may be impossible actually to dissect membrane noise into corresponding independent components. Only membrane current fluctuations under ideal voltage-clamp conditions are considered. Noise mea-

NOISE ANALYSIS AND SINGLE-CHANNEL RECORDINGS

375

surements in other conditions yield in principle the same kind of information (Wanke et al., 1974). A. Equilibrium Noise

In a membrane that is part of a system in thermodynamic equilibrium at the absolute temperature, T , the elementary events contributing to current fluctuations are determined by the general law of equipartition of energy, These fluctuations, often simply referred to as thermal noise, are well characterized according to thermodynamic principles. Their amplitude is determined by Boltzmann's constant, k , and their power spectrum is given by the Johnson-Nyquist formula (Johnson, 1928; Nyquist, 1928) [Eq. (913. S',"4'(f)= 4kTRe{ Y p ' ( f ) }

(9)

where Re{ } denotes the real part of a complex quantity, and Y$"(f) is the alternating current membrane admittance that one would measure in equilibrium conditions. It is important to emphasize that Y E ' is a purely theoretical quantity that would be measurable only if the driving forces acting on all membrane permeant ions were balanced by means of some external constraints. This is illustrated in Fig. 1 , showing the equivalent scheme of YEq' for a squid axon membrane with voltage-dependent ionic conduc-

c,,

FIG.1. Equivalent scheme of the equilibrium admittance of a simple HH model of squid axon membrane. From left to right, the parallel branches represent: the loss-free component of membrane capacitance; the equivalent loss capacitances arising from the charge redistributions associated with the state transitions of the K' and Na' channels; the leak, the potassium. and the sodium conductances. CM,T , , , T,,,, ,&. gK, and gNnhave the same meaning as in the HH equations. The voltage-dependent capacitors, C, , C , , and Ch,can be calculated from Eqs. (10) and ( I I ) using standard HH parameters and assuming particular values for the densities of K + and Na+ channels.

376

FRANC0 GONTI

tances modeled according to the simplest molecular mechanism compatiblc with the Hodgkin and Huxley (HH) equations (Hodgkin and Huxley,

1952). Two distinct features of Fig. 1 are worth stressing. First, YL“’ differs from the actual membrane admittance because it does not contain the purely phenomenological components arising from the voltage dependencc of ionic conductances (Mauro ct d., 1970). Indeed, these components vanish when the driving forces are set to zero and, contrary to what was suggested by Fishman (1975a), they do not per se generatc thermal noise as real electrical pathways. On the other hand, Yp’contains the rncmbrane admittance arising from the charge displacements associated with the gating of ionic channels (Almers, 1978). The scheme of Fig. 1 assumes. for the sake of illustration, that the n , M . and h processes yield independent gating-current contributions, characterized by effective charge movements “per gate” of z,,.el,,, and z h electronic charges, respectively. The first-order kinetics of the HH parameters imply that the x gating process (x = n , m , h ) is a linear relaxation with time constant T , , thereby producing an equivalent RCseries admittance with capacitancc, C , , given by (Lhuger, 1978) (ZxeCJ2 x(l c, = M , kT

-

where e,>is the electronic charge, and M , is the density of x gates. Denoting by NKand NNathe density of K + and Na+ channels, and accepting a literal interpretation of the HH equations, we must set Mn = 4 N K , Mni = 3 N N , , ,and M h = NNd.Furthermore, for self-consistcncy, zx must be related to the voltage dependence of the x gate through

(z,e,)lkT

=

d/d& In ( p x l a x )

(1 I )

where a, and p, are the forward and backward ratcs of the x process and EM is the membrane potential. According to Eq. (9) the power spectrum of the x gating current fluctuations is given by

Gating current noise spectra calculated according to the model of Fig. 1 are shown in Fig. 2. The numerical values for the various parameters used in the calculations were obtaincd from standard HH data (Colc, 1968) and assuming NK= 50 pm-2 and NNa= 300 pm-* (Conti et ai., 1975). Membrane noise power spectra of this type actually have been demonstrated in

377

NOISE ANALYSIS AND SINGLE-CHANNEL RECORDINGS

B A -30 m V . 5 0 mV

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

-

..............".-.........-...... .._..........-....... STol ............................ ....-...-.........",. "..".._.-...Sih. ....-............. ............,.L:_..... Sah. Sgm ...... I

__

...-..... ....-._..._.. -....... _........_........

lozz-

-PIIPIIII

..

"I.."

~

-.........-...-.......-........- "... .........,.*-...."._............ -"."... .. . . %" .:. . .: . . ... ... ..... ..... ... . .. .''

d6.

Slh . S s h

......

. . . . . 1 . . .

-

STo~

............_A...."

I

'gh

1

I

1

......

Sgm

-

.. .,,,.- ....-....-........_. ".... :. ...,+' ,I.*

s&+;rr.

.* ..

....'.::'sg ... 10ZB

I

I

lipid bilayers in association with the random translocation of hydrophobic ions or ion carrier complexes (Kolb and Lauger, 1977, 1978). The membrane admittances associated with the reversible ion transport through open channels are schematized in Fig. 1 by pure resistors. As such, they are expected to yield white noise with constant spectral density, drawn in Fig. 2 as Sth.Indeed, the roll-off of these admittances is expected only at high frequencies ( > I MHz), comparable with the rates at which ions can cross an open channel (Lauger, 1978). B. Transport Noise

A biological membrane does not normally operate at near thermodynamic equilibrium, and one measures in actual experiments a large "excess noise." We shall call "transport noise" those excess fluctuations

378

FRANC0 CONTI

that are merely associated with the presence of a net ionic transport, independently of possible fluctuations in the macroscopic structure of the pathway through which the transport occurs. This noise will depend on the nature of the transported ions, on their net flux, and on their interactions with the membrane structure that provides the pathway. The best characterized excess current fluctuations are called shot noise (Van der Ziel, 1970). They arise from the discrete nature of the electric charge and, in the simplest case, they are estimated by assuming that the electric current results from a large number of statistically independent needle pulses (shots), each carrying the same charge, ze,. For a unidirectional charge flux producing a mean current, f, the power spectrum of shot noise at low frequencies is given by Schottky’s formula [Eq. (13)]

s,,(f’)= 2

~ l ~ i l

(13)

whereas at high frequencies it decays to zero according to the time characteristics of the single-shot events (Van der Ziel, 1970). When the mean net current, 1, results from the imbalance of fluxes of the same quanta1 nature occurring in two directions, Schottky’s formula yields an upper limit for the excess noise produced by such unbalance. The flow of ions across a membrane is likely to occur through discrete steps corresponding to jumps over activation barriers. This mechanism will give rise to a more general type of shot noise, in which the shot current pulses occurring at any ion jump cannot be treated as statistically independent events (Lauger, 1975; Frehland, 1978, 1980; Frehland and Stephan, 1979; Kolb and Frehland, 1980). An instructive simple example treated by Lauger (1975) is the transport noise across a channel having a single-ion binding site. In this case there are four kinds of elementary shot events, corresponding to the jumps of an ion in and out of the channel in either direction. For monovalent ions the single-shot charge is the fraction of e, given by the fraction of membrane thickness traveled by the ion in the corresponding jump. At low frequencies and for large driving forces, the combination of two successive jumps in the same direction appears as a unitary event, and the noise spectrum is given by Eq. (13) with z = 1 . However, at high frequencies the individual shots appear as statistically independent and the excess noise obeys Eq. (13) with z < 1. This occurs above the characteristic frequency, f = 1 / ( 2 T T j ) , where T~ is the correlation time between the two possible states of channel occupancy. The general treatment of shot noise in discrete transport systems (Frehland, 1978, 1980; Frehland and Stephan, 1979) leads to the same qualitative conclusions: the excess noise produced by a net ion flow has a power spectrum with a low plateau at very high frequencies, and a low frequency amplitude determined by the clustering of shot events on a

NOISE ANALYSIS AND SINGLE-CHANNEL RECORDINGS

379

large time scale. The change in spectral amplitude occurs in a broad frequency range characterized by the kinetics of the transitions between different states of ion occupancy of the transport channel. For ionic channels in nerve membranes, which can allow the flow of at least lo6ions per second, these transitions must have very short relaxation times, yielding roll-off frequencies above 100 kHz. Thus, for all practical purposes, the shot noise produced by these ionic fluxes has a white (constant) power spectrum. In addition to shot noise, nonequilibrium systems usually exhibit more complicated fluctuations associated with long-range interactions between their components. Empirically, transport noise in excess of thermal and shot noise is often found to be described, within wide frequency intervals, by a spectral density proportional tof-E, where E is close to unity. Such llfnoise has been measured in a variety of electrical transport systems, such as carbon resistors (De Felice, 1979, glass microelectrodes (De Felice and Firth, 1970),and thick synthetic membranes (De Felice and Michalides, 1971; Michalides et al., 1973; Dorset and Fishman, 1975). llf noise is a common feature of current flow in semiconductors and metals (Hooge, 1976) as well as in ionic solutions (Hooge and Gaal, 1970). There is no satisfactory theory of llfnoise, although particular models have been proposed for specific systems (Neumcke, 1978). In the ion flow through membrane pores, llfnoise must involve longrange correlations between each pore and its surroundings. Diffusion polarization near the pore ends may give rise to such correlations (Neumcke, 1975). However, since these processes have characteristic times of less than sec (Neumcke, 1975) the buildup of their effects is expected to be reset whenever the pores stay closed for much longer periods. This argument is strongly supported by the observation that permanent gramicidin A dimers in lipid bilayers produce a large llfnoise (SauvC and Bamberg, 1978), whereas transient dimers produce only Lorentzian noise (Neher and Zingsheim, 1974; Kolb el al., 1975) with a relaxation time equal to their mean lifetime. A mechanism that could give rise to long correlation noise is suggested by the observation that K + channels stay open much longer at high extracellular potassium concentrations (Stuhmer and Conti, 1979; Swenson and Armstrong, 1981). Because of potassium accumulation, long-lived K+ channels have a greater chance of staying open even longer. It would be interesting to theorize on whether this effect could produce significant llf noise. Other highly speculative models of Ilfmembrane noise, involving longrange interactions of the ionic channels with the lipid matrix of the membrane (Lundstrom and McQueen, 1974; Clay and Schlesinger, 1976) or correlations between the states of neighboring channels (Holden and

380

FRANC0 CONTI

Rubio, 1976, 1978), cannot easily be ruled out. It must be stressed, however, that the most recent and most accurate data on nerve membrane noise do not show much of Ilfnoise components, so that interest as to theories of Ilfnoise is greatly dccrcased in this field. C. Channel Noise

The most important component of nonequilibrium noise in excitable membranes is that produced by fluctuations in the macroscopic state of ionic channels. This noise is clearly separable from thermal and shot noise because it occurs on a much longer time scale (Conti and Wanke, 1975; Frehland, 1979). The full theoretical characterization of general and specific channel noise models has been given in several papers (Stevens, 1972; Hill and Chen, 1972; Chen and Hill, 1973) and reviews (Conti and Wanke. 1975; Neher and Stevens, 1977; Colquhoun and Hawkes, 1977). The essential results are summarized below. Let the nerve membrane contain N channels, each undergoing independent fluctuations between n possible conformational states. At any fixed membrane potential let p; be the probability for a channel to be in state (j), passing the current i,. The mean and the variance of the total membrane current are then given by

Furthermore, the autocovariance of the stationary current fluctuations is expressed by the general formula

where the time constants, q , ..., rn-l, are solely determined by the rate constants of channel-state transitions, while t h e rclative amplitudes r I , ..., r, depend also on the ii's. Correspondingly. the power spectrum of channel noise is given by

,

NOISE ANALYSIS AND SINGLE-CHANNEL RECORDINGS

381

Finally, the relaxation of the mean membrane current during a voltageclamp step to EMis expressed by

where the amplitudes A I , ..., A , - , depend on the membrane voltage prior to the voltage step. Particular channel-noise models are derived by specifying the number of channel states, the currents in each state, and the matrix of state transition rates. A general simplification is obtained by assuming that single-channel currents have only one non-zero value, i (Begenisich and Stevens, 1975; Sigworth, 1977). In this case Eqs. (14) and (15) become

where p is the probability of finding a single channel in any one of its open states. Equation (20) shows that estimates of single-channel currents and of channel densities may be obtained from measurements of noise variance without the need of particular assumptions about the detailed kinetics of channel-state transitions. On the other hand, the spectral distribution of channel noise needs to be compared with the expectations of specific kinetic schemes. HH schemes, derived from a straightforward statistical interpretation of the HH equations (Stevens, 1972; Hill and Chen, 1972), are the simplest frame of reference for discussing channel-noise measurements in nerve membranes. The HH scheme of K+ channels assumes only one open-channel state, having probability n4, and passing the current iK. Equations (14), (15), and (17) become in this case ZK = N K ~ K ~ ~ 2 (TIK

= I K i K ( 1 - n4)

(21)

(22)

4

s~,(f)= l K i K I=

I

(;)(I

-

n)'n'4-')t(f,Tn/l)

(23)

where ZK is the mean potassium current, and n and 7, are the usual HH parameters describing potassium activation. For Na channels the HH scheme assumes only one open state, having

382

FRANC0 CONTI

probability m3h, and passing the current iNa.Denoting the mean sodium current by IN&,Eqs. (14) and (15) become

The power spectrum of Na channel noise can bc split into two components, s I ~ ; , ( . f )= sm(f’)

+ Sh(f)

(26)

whcre, in terms of HH parameters, m , n , T,, and 7 h r Sh(f) =

INaiNam3(1

-

h)L(.f;Th)

(27)

and, for 711 >> 7, (Conti er ul., 1975),

Examples of channel-noise spectra, calculated according to Eqs. (23), (27), and (28) for the standard HH axon of Fig. 1, are shown in Fig. 3. The same channel densities of Fig. 2 were used for the calculations. The NK estimate (50 pm-2) is taken from squid axon noise data (Conti et ul., 1975); the NNavalue (300 pm-*) is an average of estimates, ranging between 160 and 540 pm-2, from noise data (Conti et al., 1975) and from toxin-binding studies (Levinson and Meves, 1975; Keynes ef al., 1975; Strichartz et al., 1979). For the standard HH axon these values imply ohmic open-channel conductances, Y N ~and Y K , of 4 pS and 7.2 pS. The main point illustrated by Fig. 3 is that the expected H H channel noise is much higher than the sum of the other noise cornponcnts of Fig. 2 up to frequencies of the order of 10 kHz. Notice also that for small depolarizations the HH model calculations predict a trivial relative contribution of the inactivation component of Na channel noise, Sh.

D. Extrlnsic Channel Noise Membrane conductance fluctuations similar to channel noise can be induced artificially by chemicals that block reversibly the conductance of open ionic channels. Consider the simplest case of a first-order blocking reaction: ki

C+B=C* k-i

where C is the open channel, B is the blocking substance, and C* is the

NOISE ANALYSIS AND SINGLE-CHANNEL RECORDINGS

lo

''

r

383

A

FIG.3. Theoretical channel-noise spectra according to the HH kinetic scheme of K + and Na+ channel fluctuations. The spectra drawn in the figure refer to 1 cm2 of squid axon membrane containing 50 K+ channels and 300 Na+ channels per square micrometer. The calculations were made according to Eqs. (231, (271, and (28) using standard values of the HH parameters at two different membrane potentials, EM = -50 mV (A), and EM= -30 mV (B). The various spectra drawn in the figure are the K + channel noise, S,; the activation component of Na' channel noise, S,; and its inactivation component, Sh. S,,, is the total channel noise expected from an intact axon.

blocked channel. This case is equivalent to that of a two-state channel. Denoting with [B] the concentration of the blocker, the transition rates from C to C* and vice versa, are given by kl [B] and k l .The mean and the variance of current fluctuations are then given by Eqs. (19) and (20), where the open-channel probability, p , is in this case

Furthermore, the noise spectral distribution is a simple Lorentzian with time constant, T, given by: T =

(k-,

+ kl[B])-'

= p/k-l

(30)

When the blocking reaction affects channels that are also fluctuating independently between open and closed configurations, the two noise contributions are separable, provided that T is much shorter than the

384

FRANC0 CONTI

mean lifetime of the open-channel state. However, the extrinsic noise variance is reduced according to the mean fraction of time during which the channels are closed, whereas the intrinsic channel noise is calculated according to an effective single-channel current reduced by the mean fraction of time during which the open channel is blocked. Figure 4 shows the theoretical expectations for the extrinsic noise produced in squid giant axons by TEA ions, known to block reversibly the K+ channels (Armstrong, 1975; Fishman et ul., 197%; Moore et ul., 1979). The noise power spectra were calculated for the same standard axon of Figs. 1-3, assuming a first-order TEA blocking reaction with k , = 0.5 mM-' msec-' and k - , = 0.2s msec-I. These values are consistent with a half-blocking TEA concentration of 0.5 mM (Armstrong, 1975) and a cutoff frequency of the Lorentzian noise, induced by 10 mM TEA at -30 mV, of 9 0 Hz (Moore et d.,1979). Figure 4 shows the power spectrum of the intrinsic K+ channel noise (no TEA) and thosc of the total noise expected in the presence of 5 mM and SO mM TEA. Notice that the latter spectra contain only a small contribution from the intrinsic slow-channel fluctuations. Notice also that for [TEA] 2 5 mM the low-frequency plateau of both the intrinsic and the induced noisc decrease proportionately

-30 mV

I

0 1

10'

1

Hz

1o'$

Fic. 4. Calculated spectra of the extrinsic current noise induced by the reversible block of K + channels by tetraethylammonium (TEA) on a square centimcier of squid axon niembrane at EM = -30 mV. The spectrum for 0 m M TEA is identical to S, in Fig. 3 R . The spectra for 5 mM TEA and 50 mM TEA were obtained by assuming a half-block TEA concentration of 0.5 mM and an unblocking rate constant of 0.25 msec-'.

NOISE ANALYSIS AND SINGLE-CHANNEL RECORDINGS

385

to [TEAI2, while the total noise variance decreases only proportionately to [TEA]. This is because the cutoff frequency of the TEA-induced noise increases linearly with [TEA]. IV. CHANNEL NOISE IN NERVE MEMBRANES

Since the first direct measurements, the voltage fluctuations of a nerve membrane were found to be much larger than expected from thermal noise, to be correlated with the voltage-dependent potassium currents, and to be unaffected by the block of active transport (Verveen and Derksen, 1965; Derksen, 1964). However, the spectral distribution of these early data, obtained from node of Ranvier preparations of the frog, was mainly described as llfnoise. The same description applied to current noise measurements from voltage-clamped preparations of the lobster giant axon (Poussart, 1971). Positive identifications of channel noise in nerve membranes were obtained only after the successful application of fluctuation analysis to the study of chemically activated channels of postsynaptic membranes (Katz and Miledi, 1970; Anderson and Stevens, 1973). Only these more recent results of noise analysis that have contributed to the characterization of voltage-gated channels in nerve membranes are reviewed in this section. A. K Channels

Relaxation noise components associated with the open-close kinetics of K+ channels were first observed in frog node preparations (Siebenga and Verveen, 1971, 1972; Siebenga et al., 1973, 1974). The correlation between the noise spectral distribution and the kinetics of potassium currents was verified only qualitatively, but the identification of K+ channel noise was well supported on pharmacological grounds. On top of a l/f background, the voltage fluctuations showed a pseudo-Lorentzian component that was influenced by TEA, but insensitive to tetrodotoxin (TTX) (Siebenga et al., 1974). Assuming that the K+ channels have only two states with equal probability, the conductance of an open K + channel, YK, was estimated to range between 10 and 37 pS. These values are much higher than later estimates, obtained also in the same laboratory (Van den Berg et al., 1977). Considering that the model used for the analysis should lead to underestimate YK, it seems obvious that in these early measurements the K+ channel noise was largely overestimated. One of the difficulties of noise measurements in nerve membranes is meeting the requirement of steady-state conditions while avoiding the

386

FRANC0 CONTl

problems that arise from the slow inactivation of the ionic channels (Ehrenstein and Gilbert, 1966; Rudy, 1978; Fox, 1976; Brismar, 1977) and from electrode reactions. This limits the applicability of a true stationary noise analysis only to relatively small membrane depolarizations. Most of these difficulties were overcome by Begenisich and Stevens (1975) by measuring simultaneously mean potassium currents and current fluctuations during repeated voltage-clamp depolarizations of relatively short duration. This analysis was applied to TTX-treated frog nodes in the membrane potential range of -48 to + 16 mV. The data were reported to be well fitted by Eq. (20) assuming a simple ohmic behavior of the open K ' channel with yK= 4 pS. This result, confirmed by more recent data of Neumcke et ul. (l98Oa),provided the first direct evidence that K + channels of frog nodes have only one significant non-zero conductance value. An interesting physiological finding of the latter work is a significant difference between the YK values of motor fibers (-2.7 pS) and those of sensory fibers (-4.6 pS). The applicability of Eq. (20) makes the estimate of YK from noise data fairly independent of the kinetics of the state transitions of K+ channels. Which kinetic model fits most adequately the spectral distribution of K+ channel noise is more open to discussion. Stevens (1977) reported a power spectrum of K channel fluctuations, from the experiments of Begenisich and Stevens (1975), which is well fitted by two Lorentzians with quite different cutoff frequencies, one in the range expected from the HH model and the other about one order of magnitude higher. Van den Berg et al. (1977) reported that the power spectra of current fluctuations attributed to K + channel noise are fitted by a single Lorentzian both during relatively short-step depolarizations (phase I ) and during prolonged depolarizations (phase 2). However, while the voltage dependence of the phase 1 noise follows roughly the expectations of the HH model, the phase 2 noise had completely different characteristics. Fishman et ul. (1981) have investigated the same problem in squid giant axons, but they found that in this preparation the noise spectra and the membrane admittance do not change between phase I and phase 2. The K+-channel noise spectra reported by Neumcke ct nl. (1980a) were fitted with a curve that declines beyond the half-power frequency,f,, as (.flfc)1.5. It is argued that this description is consistent with a model of multistate channel with state transitions occurring over a large number of identical energy barriers. K+-channel noise has been studied in squid giant axons using two different techniques. The first observations were reported by Fishman (1973, 1975a), who measured voltage and current noise from small patches ( to 10-5cm2)of membrane isolated electrically by a peripheral continuous flow of sucrose solution (Fishman, 1975b). The total membrane noise was

NOISE ANALYSIS AND SINGLE-CHANNEL RECORDINGS

387

fitted by the sum of a llfcomponent and a relaxation component that was abolished by intracellular TEA. The half-power frequency of the relaxation noise was in the range expected from the kinetics of potassium currents, but it was found to be in systematic quantitative disagreement with the expectation of a simple HH model. The same conclusion was reached in later measurements using the same technique (Fishman et al., 1975a,b). The latter works emphasized also the presence of sharp corners in the noise power spectra that were inconsistent with the superposition of linear relaxation components. Since patch recordings do not allow a reliable measurement of mean currents (Fishman, 1975b),these data did not yield estimates of YK. The measurements from sucrose-isolated small membrane patches have been subject to criticism (Wanke et al., 1974; Conti and Wanke, 1975) based mainly on the fact that in this preparation the power spectra of voltage and current noise are very similar in shape, contrary to what was expected from the strong frequency dependence of the squid axon membrane impedance (Cole, 1941; Mauro et al., 1970; Conti, 1970). Several arguments have been offered against these criticisms (Fishman et al., 1975a). However, it is a fact that some features of the noise recorded from membrane patches isolated with sucrose flow have no obvious interpretation. Indeed, the interpretation of “peaking” noise spectra in terms of cyclic transitions between channel states (Fishman et al., 1977) is hardly reconcilable with the fact that nerve excitation has no immediate need of metabolic energy (Baker er al., 1962). The other mechanism proposed by Fishman et al. (1977), involving a major role of gating currents, is even less likely, considering the trivial noise contribution expected from this source (see Fig. 2). A further puzzling feature of patch-noise measurements, which is peculiar to this preparation, is the observation of a large Kf-channel noise at very negative membrane potentials [see Fig. 5 of Fishman et al. (1975b), and Fig. I of Moore et al. (1979)1, for which IK should be practically zero. Quite different results were obtained by measuring the electrical fluctuations of large areas of squid axon membrane in standard space-clamp conditions (Wanke et al., 1974; De Felice et al., 1975; Conti et al., 1975). As illustrated in Fig. 5 , this preparation showed large differences between spectra of voltage noise and current noise, the latter lacking completely the peaking features of the former, which merely reflect the frequency dependence of the axonal membrane impedance. An extensive analysis of the current fluctuations measured during small steady depolarizations (Conti et al., 1975)revealed the presence of relaxation components, sensitive to TTX and TEA, that were analyzed as a function of temperature and membrane potential. The block of Na cur-

388

FRANC0 CONTI

J

1 1

1

I

I

I

1 I l l 1

I

1

11llllL

10

FIG.5. Current noise power spectrum, 0; voltage noise power spectrum, e; and the square modulus of the impedance, x , in a large area (0.36 cm2)of squid axon membrane in normal physiological conditions at resting potential (between -55 and 60 mV). From Wanke et t r l . (1974).

NOISE ANALYSIS AND SINGLE-CHANNEL RECORDINGS

389

rents by addition of TTX was shown to produce a significant decrease of membrane noise, mainly in the high frequency region of the spectrum, and this reduction was attributed to removal of Na+ channel noise, as discussed later. On the other hand, the noise spectra from axons injected with large concentrations of TEA (-70 mM) were strongly depressed at low frequencies, but comparable to those of normal axons at high frequencies. The TTX-insensitive noise, mainly attributed to K+ channels, was described as the sum of a l/Jand a Lorentzian component, and the latter was compared with the theoretical expectation according to Eq. (23). The temperature and voltage dependence of the amplitude and cutoff frequency of the Lorentzian spectrum was found to be in fair quantitative agreement with the HH model. The data from several axons were fitted with Eqs. (21)-(23) using mean I , , n, and 7, values obtained from step voltage-clamp measurements, allowing one to estimate NK as about 60 Fm-2. This yielded a yKestimate of about 12 pS, calculated according to a measured maximum potassium conductance, gK,of 70 msec cmP2. The analysis of channel-noise data using NKas a fit parameter is subject to several sources of error. First, channel-density estimates depend on separate measurements of membrane area. Second, NKmay vary significantly from one preparation to another. Finally, the relationship between noise variance of channel density [obtained by dividing, term by term, Eq. (22) by Eq. (21)] is strongly dependent on a separate evaluation of the open-channel probability, which may be quite inaccurate at low depolarizations. A better strategy of channel noise analysis avoids the use of Eq. (21) and uses the single-channel current as the fit parameter (Conti et a/., 1976a,b). This method is applied in the analysis of the noise spectra shown in Fig. 6 . The data (F. Conti and E. Wanke, unpublished) were obtained from a squid giant axon preparation using the same technique described by Conti et al. (1973, with two improvements: (1) The noise of the voltage-clamp amplifier was further reduced by using four parallel low-noise JFET’s (C413N Teledyne Crystalonics, Massachusetts) for each differential input; (2) the manual compensation of slow voltage drifts was substituted by a slow additional feedback system, thereby eliminating a possible source of stray low-frequency noise. Figure 6A shows current noise spectra at three different levels of steady membrane potential, measured when the axon was immersed in artificial seawater (ASW) at 6°C. Figure 6B shows similar data measured from the same axon after addition of 300 mM TTX to the extracellular medium. A clear reduction of membrane noise, most pronounced at high frequencies, is observed in the latter measurements. The dashed lines in Fig. 6B are theoretical spectra given by the sum of thermal and Schottky noise estimated from the measured mean currents, plus the K+ channel noise calcu-

390

FRANC0 CONTI

A

ASW

1

1

I

lo2

1

1

I

H Z

10'

ld

B ASW- TTX

102

1B

ld I

1

I

1

I

I

10

*

H Z

to4

NOISE ANALYSIS AND SINGLE-CHANNEL RECORDINGS

391

lated from Eq. (23), using standard HH values for n (Hodgkin and Huxley, 1952; Cole, 1968) and selecting T, and iK values that provided a reasonable fit by eye of the low-frequency data. Although not shown in the figure, a fair fit of all data of Fig. 6B (perhaps with the exception of that at -48 mV) could have been obtained also at high frequencies by adding to the theoretical spectra the expected contribution from the amplifier's noise. It is obvious, however, that such contribution, as well as that of thermal and shot noise, is insignificant below 100 Hz. The values of iKyielding the theoretical curves of Fig. 6B correspond to single-channel chord conductances ranging from 5.3 to 7.2 pS with a mean of 6.3 pS which is about one-half of the y K estimate given by Conti et al. (1975). The discrepancy is only apparent, most likely ascribable to the rectification of potassium currents (French and Wells, 1977). Indeed, the estimate of Conti er al. (1975) based upon gKmeasurements, is applicable to the chord conductance for large depolarizations, which is consistently expected to be much larger than the chord conductance near the resting potential. The values of T, yielding the theoretical spectra of Fig. 6B were 50 to 100% larger than expected from the HH fit of voltage-clamp currents. A better agreement could have been obtained by adding a llfcomponent to the theoretical spectra as done by Conti er al. (1975). This would have shifted the cutoff of the Lorentzian spectra toward higher frequencies, without changing significantly iK. However, such I/fcomponent would in any case have been much smaller than in Conti er al. (1975), suggesting that the origin of the llfnoise in the latter work was mostly artifactual. Furthermore, in view of our discussion of llfnoise, and in view of the K + FIG.6. Power spectra of current fluctuations in a large area (0.36 cmZ)of squid giant axon membrane voltage-clamped at various steady potentials, indicated in the figure. Temperature, 6°C. The spectra in (A) were obtained with the axon immersed in artificial seawater (ASW); those in B were measured after switching the extracellular solution to ASW containing 300 m M TTX. The mean membrane currents in ASW, IASW, were -0.7 p A at -62 mV; 0.44 pA at -56 mV; 1.84 p A at -53 mV. Mean currents in ASW-TTX, l m x :0.1 /.LAat -62 mV; 1.64 p A at -56 mV; 3.84 pA at -53 mV; 7.6 p A at -48 mV. Leak currents, IL, were calculated by extrapolating the linear I-V membrane characteristics measured at membrane potentials negative of -80 mV. The mean potassium current, I K .was measured as: / K = I T ~ X- II ; the mean sodium current in ASW was calculated as: IN;,= IAsW- !mx. The dashed lines in ( B ) are the sum of the thermal and shot noise of IKand 11, plus the K' channel noise calculated according to Eq. (23) using standard HH values for n and using the following values of 7,and iK: 20 msec and 0.06 pA at -62 mV; 20 msec and 0.09 pA at -56 mV; 18 msec and 0.09 pA at -53 mV; 17 msec and 0.11 pA at -48 mV. The iK values correspond to chord conductances ranging between 5.3 and 7.2 psec. The solid lines in (A) are the sum of the theoretical spectra in (B) and those of Fig. 7, obtained from the fit of Na+ channel noise. Data from F. Conti and E. Wanke (unpublished).

392

FRANC0 CONTI

channel noise data from frog nodes already discussed (Neumcke et al., 1980a; Stevens, 1977), it seems more reasonable to attribute the apparent discrepancy between the time constants that fit relaxation and noise data, to some inadequacy of the HH description of the K t channel kinetics. Indeed a much better fit of the data of Fig. 6B, particularly at -48 mV, could be obtained by adding to the theoretical spectra a fast relaxation component as done by Stevens (1977). The reversible block of K t channels by TEA, as discussed in Section III,D, provides a typical example of extrinsic channel noise, which can be exploited to study the properties of K+ channels. The current noise induced by TEA and by the TEA analog triethyldecylammonium (TEDA) has been observed in squid giant axons with the patch recording technique (Fishman et al., 1975b; Moore et al., 1979). The noise spectra were found to be consistent with the simple first-order blocking reaction proposed by Armstrong (1973, and tentative estimates of y K .from the application of Eq. (20) with p << 1, were reported to range from 1.2 to 2.1 pS. Unfortunately, these values cannot be taken too seriously because of the general inaccuracy of steady-state current measurements from membrane patches (Fishman, 1975b). In addition, these estimates were obtained from axons perfused with high TEA concentrations (>S mM), which reduce the potassium currents by more than 10-fold. In those conditions it seems that the value of ZK could at best only be guessed. 6. Na Channels

The first positive identification of Na+ channel noise was obtained by Conti et al. (1975) from measurements of the kind shown in Fig. 6 . The membrane noise that is abolished by blocking the Na currents with TTX was found to have all the general features expected for the Na' channel noise according to thc simple HH scheme discussed in Section III,C. These features included (1) temperature dcpendence; (2) correlation between tNaand noise amplitude; (3) fair quantitative agreement between the high-frequency cutoff of the noise and the kinetics of sodium activation. It was also found that the TTX-sensitive noise persisted after intracellular injection of 70 mM TEA, which reduced the potassium currents by more than 100-fold. Without placing much confidence in the data below 100 Hz, which they considered to be affected by too large errors, Conti et al. (1975) analyzed the differences between ASW and TTX spectra according to Eqs. (24)-(28), estimating NNato be about 330 pm-' and thereby deducing a yNa value of about 4 pS (to be consistent with R N a = 120 msec cm-2).

393

NOISE ANALYSIS AND SINGLE-CHANNEL RECORDINGS

Figure 7 illustrates a similar analysis of the data of Fig. 6 using iNaas the main fit parameter. The spectral data points of Fig. 7 were obtained from the difference of sampled data from Fig. 6A and B . It should be noticed that the difference spectra do not show any significant amplifier’s noise contamination, in agreement with the fact that TTX produces insignificant changes of the membrane impedance at the frequencies at which this noise becomes important. The dashed lines drawn in Fig. 7 are theoretical spectra calculated according to Eqs. (26)-(28) using standard HH values for m , h , and ‘rh (Hodgkin and Huxley, 1952; Cole, 1968), using the ZNa values measured by the change in membrane current upon TTX addition, and selecting the iNa and 7, values that yielded reasonable fit by eye of the high-frequency data. The estimated thermal and shot noise associated to 1~~ was disregardable. Gating noise M’US not added to the theoretical fit, consistent with the fact that gating currents are not affected by TTX (Armstrong and Bezanilla, 1974). The fit of the three difference spectra of Fig. 7 was obtained for 7, values about 50% higher than those of a standard HH axon, and for iNavalues corresponding to an open Nachannel conductance, yNa,in the range of 5 to 6.3 ps. This is fairly close

-81

mv

0

I

10

I

I

10”

nz

1

1

I

1

I

I

FIG. 7 . Sampled difference spectra of the Na+ channel noise which is suppressed by tetrodotoxin in the experiment of Fig. 6 . The dashed lines are the theoretical spectra predicted by the HH model, Eqs. (26)-(28), calculated according to standard HH values of rn, h , and ~ h using , the ZNa values measured as described in Fig. 6, and using the following fitted values of T , and iNa:0.35 msec and -0.61 pA at -62 mV; 0.45 msec and -0.72 pA at -56 mV; 0.5 msec and -0.72 pA at -53 mV. The iNs values correspond to Na’ channel chord conductances of 5.0, 6.2, and 6.3 psec. The same values of all parameters were used to calculate the spectra drawn as solid lines, except that the inactivation component, Sh,was arbitrarily amplified to obtain a better fit of the low-frequency data. The Sh amplification factors were: 5000 at -62 mV, 700 at -56 mV, and 100 at -53 mV.

394

FRANC0 CONTI

to the mean yNa estimate of Conti et aI. (1975). obtained from less accurate data. The dashed lines in Fig. 7 are a poor fit of the data below 100 Hz, which lie almost all above the curves. A similar discrepancy was also observed by Conti et al. (1975) and tentatively attributed to llfnoise. An alternative interpretation, suggested by an accurate analysis of Na+ channel noise in frog nodes (Conti et al., 1980) is that the slow relaxation component of is much higher than expected according to Eq. Na-channel noise, Sh(f), (27). The solid lines in Fig. 7, yielding a fair fit of the data in the whole measured frequency range, were calculated for the same parameters of the dashed curves, allowing the &(f) component to have an amplitude two to three orders of magnitude higher than predicted by Eq. (27). The sum of the theoretical fits of K+ channel noise (Fig. 6B) and of Na+ channel noise (Fig. 7) is shown as solid lines in Fig. 6A. The Na+ channel noise measurements of Conti et af. (1975) have been repetitively judged to be artifacts by Fishman P t ul. (1975b, 1977), who stated that the intracellular injection of TEA should produce extrinsic K+channel noise, rather than allowing to detect intrinsic Na+-channel noise. The model calculations of Figs. 3 and 4 show clearly that the extrinsic noise produced by the 70 mM TEA is expected to be trivially small in comparison with Na+-channel noise. Furthermore, had the TEA noise contribution been significant in the experiments of Conti et al. (1975), the apparent Na+-channel noise measured in TEA-injected axons would have been much larger than that measured from TTX experiments. Strangely enough, however, the latter argument has been reversed to conclude that the Naf-channel noise obtained from TTX experiments of the kind illustrated in Figs. 6 and 7 must also be artifactual (Fishman rt al.. 1977). Noise measurements from small patches of squid axon membranes have allegedly failed to reveal Na+ noise under normal ionic conditions (Fishman et nl., 1975b, 1977). Na+-channel noise was reportedly observed in this preparation only after the replacement of the intracellar potassium with impermeant solutes or with cesium (Fishman et al., 1977). The Na+ noise spectra were defined as difference between the spectra measured at various depolarizations and a reference spectrum measured at -90 mV. An attempt to correlate the Nat noise variance with mean sodium currents according to Eq. (20) yielded yNa values increasing monotonically from I to 40 pS in the membrane potential range of -70 to +20 mV. I t is very difficult to assess the relevance of these results to the understanding of the normal functioning of Na+ channels. Quite apart from the questionable method of analysis, the properties of the preparation used in these experiments seem quite different from those of normal axons. For example, the apparent steady-state sodium conductance measured by Fishman

NOISE ANALYSIS AND SINGLE-CHANNEL RECORDINGS

395

et al. (1977) increases monotonically with voltage, in the range of -60 to 20 mV, with no obvious sign of sodium inactivation. This casts serious doubts about the correct identification of sodium currents, especially considering that the patch recording technique does not allow to add TTX extracellularly during a single experiment. Na+-channel noise in frog nodes was first detected by measuring stationary voltage fluctuations in the absence of potassium currents, blocked by intracellular cesium and TEA (Van den Berg et al., 1975). The TTXsensitive noise was reported to be in fair agreement with the expectations of the HH model, predicting a predominance of the Sh component at the relatively large depolarizations (EM= -30 and -40 mV) applied in these studies. Assuming standard HH parameters for frog nodes (Hille, 1967), yNawas estimated to be in the range of 2-5 pS. Frog node sodium current fluctuations in the wider range of membrane potentials (-60 to -20 mV) were extensively studied by Conti et al. (1976a) using the pulse technique (Begenisich and Stevens, 1975). Na+ noise was measured from difference spectra, before and after application of 150 nM TTX, in preparations having 95% of the potassium currents blocked by 10 mM TEA. Apart from a small llfcomponent, the data were well fitted by Eqs. (24)-(28), using approximately the same values of rn, h , T,, and T h , which yielded a good HH fit of the macroscopic sodium currents. Recognizing that large errors could have been introduced by the pulsing technique in the measurement of low-frequency fluctuations, the authors could not reach any trustworthy conclusion about the Shcomponent of the Na+ channel noise predicted by the HH scheme, or about the real presence of a llfNa noise in the nodal membrane. However, it was argued that the iNavalues estimated from the fit of the high-frequency fluctuations were fairly independent of the detailed fitting scheme. These values yielded a mean y N a estimate in normal fibers of 7.9 pS. This estimate was also found to be little affected by various agents that block or delay sodium inactivation (Conti et al., 1976b), a result that argued in favor of the all-or-nothing nature of the inactivation process. By analyzing differences of successive noise samples, Conti et al. (1980) were able to eliminate most of the errors due to systematic drifts and slow nonstationarities, extending the reliable range of Na+-channel noise measurements i n frog nodes down to frequencies of 3 Hz. The mean yNaestimate obtained in the latter work was 8.85 pS, in excellent agreement with the previous results. The noise spectra were fitted by the mere superposition of Lorentzian components, but the spectral contribution associated with the inactivation of sodium currents was found to be much larger than expected from Eq. (27). Furthermore, significant quantitative discrepancies were found between the T, values estimated from the fit of

396

FRANC0 CONTI

the high-frequency noise according to Eq. (28) and those yielding accurate HH fits of the voltage-clamp currents. The major inadequacy of the HH model was recognized to arise from the assumption of independence between activation and inactivation of Na+ channels, an assumption that is also independently challenged by the results of gating-current measurements (Armstrong and Bezanilla, 1977; Nonner, 1980). In qualitative agreement with the analysis of squid axon gating currents (Armstrong and Gilly, 1979), it was concluded that the HH scheme is also inadequate to describe in detail the kinetics of the Na+-channel state transitions associated with sodium activation. The low-frequency fluctuations measured by Conti r t al. (1980) could be better fitted by the sum of two Lorcntzians, consistent with the observation that sodium inactivation in frog nodes is diphasic (Chiu, 1977; Nonner, 1980). This feature was analyzed in detail by Neumcke et ul. (1980b) in a study of the modifications of sodium inactivation produced by Anornonia toxin I1 or by intracellular iodate ions. I t was reported that the modifying agents primarily increase the time constant of the slower phase of sodium inactivation, changing also its voltage dependence. In agreement with previous results (Conti rf d., 1976b), the estimated YN,, was rather insensitive to these modifications. The most extensive test of the all-or-nothing character of Na+ channel conductance was obtained by Sigworth (1977, 1980a,b) from measurements of nonstationary noise in frog nodes. In Sigworth’s analysis, the variance and the mean of sodium currents were evaluated as ensemble averages over many repeated step depolarizations. At any time from the onset of the voltage steps these quantities were found to be related by Eq. (201, yielding estimates of the number of Naf channels per node that are independent of the voltage step. In addition, the voltage dependence of the estimated single-channel current was found to match very well the instantaneous Z-V characteristics of the macroscopic sodium currents. The chord conductance of the open Na+ channel near EM = 0 was estimated to be about 7 pS (Sigworth, 1980a). This value was not changed when the sodium currents were reduced by TTX, by saxitoxin, or by depolarizing prepulses (Sigworth, 1980b),but a 60% reduction of Y N was ~ observed upon lowering the extracellular pH to 5.0 Nonstationary fluctuations allow the study of membrane states in which the Na channels have a large probability of being open. a situation that never occurs in normal fibers in stationary conditions. At the time at which the sodium currents reach their peak value, this probability was estimated by Sigworth (19804 to be 0.6 for EM= -5 mV,and 0.9 for EM= 125 mV, in fair agreement with the predictions ofthe H H model. Another important application of nonstationary noise analysis was obtained by

NOISE ANALYSIS AND SINGLE-CHANNEL RECORDINGS

397

Sigworth (1981)from measurements of the autocovariance defined by Eq. ( 5 ) . The data were taken from frog nodes during the inactivation phase of sodium currents, following nearly maximal activation. For any fixed t ~ , the decay of r,(t,,t z ) with ( t 2- t l ) was described by two exponentials in agreement with the diphasic time course of sodium inactivation (Chiu, 1977; Nonner, 1980). However, it was observed that the relative amplitude of the two exponentials changed markedly as a function of t l , a finding that argues against models that postulate several inactivated states and only one open state for the Na+ channel (Chiu, 1977; Nonner, 1980). V. SINGLE-CHANNEL RECORDINGS

The analysis of channel noise discussed in Sections I11 and IV is based upon the fundamental assumption that ionic channels have a relatively small number of states that are kinetically distinguishable within the time resolution of the measuring apparatus. In other words, it is assumed that the set of all possible microscopic states of any ionic channel can be divided into a small number of subsets with (1) random state transitions within a single subset occurring at much faster rates than the transitions between different subsets, and (2) a mean lifetime of any subset well within the range of the recording instruments. It is then legitimate to identify each subset with a macroscopic channel state and to describe the state transitions of a channel as a simple Markovian process (Lax, 1960). The ultimate proof of the validity of the above picture can be obtained only from direct observations of single-channel events, because other interpretations of channel noise data are in principle possible. For example, by assuming that only the triggering of the channel opening is a random event, whereas the closing follows a deterministic exponential time course, one would predict the same channel-noise spectra that are expected for ionic channels with two-state Markovian transitions, and the former model was in fact used for the original analysis of the noise from chemically activated channels in the postsynaptic membrane (Katz and Miledi, 1970). This ambiguity was definitely resolved by the first successful recordings of single-channel currents from small membrane patches of denervated muscle fibers (Neher and Sackmann, 1976), which showed an all-or-nothing behavior with open-time distributions consistent with a Markovian description. The patch-clamp technique has been applied to squid giant axons by Conti and Neher (1980) to record discrete current events attributed to the opening and closing of single K + channels. Two major modifications of the original method had to be introduced for this particular preparation.

398

FRANC0 CONTI

1. In order to allow a close enough approach to the axon membrane by

the patch-recording pipette, the latter was introduced intracellularly after extensive perfusion of the axon with Pronase. 2. In order to increase the size of the single-channel currents for negative membrane potentials, when the opening of a K+ channel is a rare event, the equilibrium potential for potassium ions was made positive by reversing the normal potassium gradient. Condition 2 had the two additional advantages that (1) the low intracelMar ionic strength improved the electrical seal around the patch area, and (2) the high extracellular potassium concentration prolonged the opening time of K+channels (Stuhmer and Conti, 1979; Swenson and Armstrong, 1981). A major disadvantage, however, was that the large intracellular resistivity caused a strong spillover of nonstationary currents, from the whole axon clamp into the patch-clamp system. The major features of the observations made by Conti and Neher (1980) in the above experimental conditions are illustrated in Fig. 8. The fluctuations of the current flowing through the patch area (- 1 pm2) showed large changes when the membrane voltage was varied within the range over which the activation of potassium currents occurs. A sharp maximum in

+34 417

- 8

-19

m 35

i

-25

-31

I

5 PA

125 MSEC

I

3 PA

5 0_N.S E C

FIG.8. Recording of K+ currents through a small patch of squid axon membrane, voltage-clamped through an L-shaped glass pipette pressed against its intracellular face. Temperature, 5.5"C. ( A ) Several traces were recorded at the membrane potentials indicated. (B) Four records from the same membrane patch at higher resolution, showing the burstlike character of activity of single K+ channels. From Conti and Ncher (1980).

NOISE ANALYSIS AND SINGLE-CHANNEL RECORDINGS

399

the noise variance near the half-activation voltage subsided, for more negative voltages, into low-noise records containing discrete fluctuations in the form of inwardly directed square current pulses (last two traces of Fig. 8A). These fluctuations were interpreted as contributions from individual K+ channels, which cannot be seen at more depolarized voltages because several channels in the patch area and under the rim of the pipette are open simultaneously, have very short closing times, and produce smaller current jumps according to a smaller driving force. The amplitude distributions of the unitary currents at negative voltages was found to be strongly skewed because of the reduced size of the currents from openings having short durations or occurring under the rim of the pipette. The edge of the distributions on the higher side was taken as a measure of well resolved full-size single-channel currents, yielding an estimate of YK of about 17 pS in fair agreement with that obtained from noise analysis (Conti et al., 1975). Figure 8B shows examples of single-channel events, at a better revolution than in Fig. 8A, which illustrate a most interesting feature of these waveforms. It is seen that a seemingly unitary event contains several short interruptions, whose average number was estimated as 1.8 per event. This burstlike appearance of single-channel currents is inconsistent with the HH scheme of K+ channel kinetics, which predicts in the same conditions that only one out of four openings should be followed by a reopening after a brief closing interval. The frequent interruptions suggest the presence of a closed state of the Kf channel, which is in relatively fast equilibrium with its open state. It is obviously tempting to correlate this observation from patch recordings to the high-frequency components of K+ channel noise, which are not fitted by the HH scheme, as discussed in Section IV,A. The patch-recording experimental setup used by Conti and Neher (1980) was very elaborate; it required the manufacturing of very complicated whole-cell recording electrodes and of special perfusion and recording pipettes. Furthermore, the bandwidth of the recordings was seriously limited by the large capacitance of the long patch-recording pipette, inserted longitudinally inside the axon. An easier preparation that might allow to improve considerably the recording of single-channel events from squid axons has been developed by Llano and Bezanilla (1980) using small segments of cut-open axons positioned between two compartments. Unfortunately, the reported measurements of voltage-clamp currents from this preparation so far have not included recordings of single-channel events. The possibility of achieving seal resistances larger than lo9 ohm (giga seals), by a mere suction through the patch pipette, has improved the

400

FRANC0 CONTI

resolution of single-channel recordings by more than one order of magni, Unfortunately, the development tude (Neher, 1980; Hamill er ~ d . 1981). of giga seals requires a perfectly clean membrane surface, a condition that is easily achieved only in tissue-cultured cells. Thus, in the experiments on squid axons described above, gentle suction often produced moderate improvements of the seal, but never dramatic ones, presumably because a lot of cytoskeletal material remains attached to the axolemma even after extensive enzymatic treatmcnt. It appears, therefore, that the best patch recordings of voltage-activated channels will likely be obtained from cultured cells. Indeed, currcnts from single Na' channels have alrcady been observed in cell-attached (Sigworth and Neher, 1980) and in excised patches (Horn et al., 1981) of the membrane of cultured muscle cells. Two main conclusions were drawn from the observations of Sigworth and Neher (1980). The single Na+ channel conductance was estimated to be about 18 pS, in fair agreement with estimates of yNain frog nodes after taking into proper account differences in temperature and in sodium concentration. On the other hand, the relationship between the mean lifetime of an open channel and the time constant of sodium activation did not conform to the predictions of the HH schcme, being more consistent with a kinetic model in which the final opening step is rate limiting in the channel activation process (Armstrong and Gilly, 1979). The single-channel recordings of Horn et al. (1981) confirmeJ the order of magnitude of the yNaestimates. Furthermore, it was reported that the statistics of single-channel currents argued against a strictly sequential scheme for the activation-inactivation kinetics of Na+ channels, in which the inactivation step occurs only from the open state. In summary of the results discussed in this section, it can be stated that patch recordings have provided a most convincing definite proof of the notion that guided the interpretation of noise data: the basic phenomenon underlying the voltage-dependent conductance of electrically excitable cells is the all-or-nothing behavior of independent ionic channels, sparsely distributed over the cell surface. The large number of independent estimates of the unitary conductances of voltage-gated channels lie in the rather narrow rangc of 3 to 18 pS, providing thc best proof that in the open configuration these channels have a porelike structure spanning the membrane. Concerning the detailed kinetics of the conformational transitions of ionic channels, it is easy to foresee that more exhaustive information will be soon obtained, particularly from high-resolution patch recordings. The inadequacy of the single HH kinetic schemes is not news any more. Unfortunately, it seems that any more faithful picture will lack the beautiful mathematical and physical simplicity that was determinant for the choice operated by Hodgkin and Huxley.

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ACKNOWLEDGMENTS

I thank Professor E. Wanke for permission to report unpublished data and Dr. R. Fioravanti for help in the calculations of theoretical noise spectra. REFERENCES Almers, W. (1978). Gating currents and charge movements in excitable membranes. Rev. Physiol. Biochem. Pharmacol. 82, 97-182. Anderson, C. R., and Stevens, C. F. (1973). Voltage clamp analysis of acetylcholine produced end-plate current fluctuations at frog neuromuscular junction. J . Physiol. (London) 235, 655-691. Armstrong, C. M. (1975). K pores of nerve and muscle membranes. In “Membranes: A Series of Advances” (G. Eisenman, ed.), Vol. 3, pp. 325-328. Dekker, New York. Armstrong, C. M., and Bezanilla, F. (1973). Currents related to movement of gating particles of the sodium channels. Nature (London) 242, 459-461. Armstrong, C. M . , and Bezanilla, F. (1974). Charge movement associated with the opening and closing of the activation gates of the Na channels. J . Gen. Physiol. 63, 533-552. Armstrong, C. M., and Bezanilla, F. (1977). Inactivation of the sodium channel, 11. Gating current experiments. J . Gen. Physiol. 70, 567-590. Armstrong, C. M., and Gilly, W. F. (1979). Fast and slow steps in the activation of Na channels. J Gen. Physiol. 74, 691-711. Baker, P. F., Hodgkin, A. L., and Shaw, T. I . (1962). Replacement of the axoplasm of giant nerve fibres with artificial solutions. J . Physiol. (London) 164, 330-354. Bean, R. C., Shepherd, W. C., Chan, H., and Eichner, J. T. (1969). Discrete conductance fluctuations in lipid bilayer protein membranes. J . Gen. Physiol. 53, 741-755. Begenisich, T., and Stevens, C. F. (1975). How many conductance states do potassium channels have? Biophys. J . 15, 843-846. Bendat, J. S . , and Piersol, A. G. (1971). “Random Data: Analysis and Measurement Procedures.” Wiley, New York. Brismar, T. (1977). Slow mechanism for sodium permeability inactivation in myelinated nerve fibre of Xenopus laevis. J . Physiol. (London) 270, 283-297. Chen, Y.,and Hill, T. L. (1973). Fluctuations and noise in kinetic system: Application to K’ channels in the squid axon. Biophys. J . 13, 1276-1295. Chiu, S. Y . (1977). Inactivation of sodium channels: Second order kinetics in myelinated nerve. J . Physiol. (London) 273, 573-596. Clay, J. R., and Schlesinger, M. F. (1976). Theoretical model of the ionic mechanism of l/f noise in nerve membrane. Biophys. J . 16, 121-136. Cole, K . S. (1941). Rectification and inductance in the squidgiant axon. J . Physiol. (Landon) 25, 29-5 I . Cole, K. S. (1968). “Membranes, Ions and Impulses.” Univ. of California Press, Berkeley. Colquhoun, D., and Hawkes, A . G . (1977). Relaxation and fluctuations of membrane currents that flow through drug-operated channels. /‘roc.. R . Soc. London Sor. B 199, 231-262. Conti, F. (1970). Nerve membrane electrical characteristics near the resting state. Biophysik 5, 71-81. Conti, F., and Neher, E. (1980). Single channel recordings of K’ currents in squid axons. Nuture (London) 285, 140-143. Conli, F., and Wanke. E. (1975). Channel noise in nerve membranes and lipid hilayers. Q. Reu. Bioplr.v.s. 8, 45 1-506.

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Conti, F., DeFelice, L. J . , and Wanke, E. (1975). Potassium and sodium ion current noise in the membrane of the squid giant axon. J . Physiol. (London) 248, 45-82. Conti, F., Hille, B., Neumcke, B., Nonner, W.. and StPmpfli, R. (1976a). Measurement of the conductance of the sodium channel from current fluctuations at the node of Ranvier. J . Phy.siol. (London) 262, 699-727. Conti, F., Hille, B . , Neumcke, B . , Nonner, W., and Stampfli, R. (1976b). Conductance of the sodium channel in myelinated nerve fibres with modified sodium inactivation. J . Phv.siol. (London) 262, 729-742. Conti, F., Neunicke, B . , Nonner, W., and Stampfli, R. (1980). Conductance fluctuations from the inactivation process sodium channels in myelinated nerve fibres. J . Physiol. (London) 308, 217-238. DeFelice. L. J. (1975). I/f resistor noise. J. Appl. P h y s . 47, 350-352. DcFelicc, L. J , (1977). Fluctuation analysis in neurobiology. / n / . Kuu. Ntwrohio/. 20, 169-208. DeFelice, L. J., and Firth, D. R. (1970). Spontaneous voltage fluctuations in glass microelectrodes. IEEE Truns. Biomed. Eng. 18, 339-351. DeFelice, L. J., and Michalides, J . P. L. M. (1971). Electrical noise from synthetic membranes. J . Mernhr. B i d . 9, 261-290. DeFelice, L. J., Wanke, E., and Conti, F. (1975). Potassium and sodium current noise from squid axon membranes. Fed. Proc. Fed. A m . Soc. Exp. Bid. 34, 1338-1342. Derksen, H. E. (1965). Axon membrane voltage fluctuations. Acta Physiol. Pharmucol. N e ~ r l 13, . 373-466. Dorset, D. L., and Fishman, H . M. (1975). Excess electrical noise during current flow through P O I - O U ~ nicnibranes scparating ionic solutions. J . Mernbr. B i d . 21, 29 I - 309. Ehrenstein, G., and Gilbert, D. L. (1966). Slow changes in potassium permeability in the squid giant axon. Biophys. J . 6, 553-566. Ehrenstein, G . . and Lecar. H . (1977). Electrically galed ionic channels in lipid bilayers. Q.~ e u Biop/lv.s. . 10, 1-34. Fishman. H . M . (1973). Relaxation spectra of potassium channel noise from squid axon membrane. Pror. N o d . Arod. Sri. U . S . A . 70, 876-879. Fishman, H . M. (1975a). Noise measurements in axon membranes. Fed. Proc. Fed. A m . SOC. Exp. B i d . 34, 1330-1337. Fishman, H . M. (1975b). Patch voltage clamp of squid axon membrane. J . Memhr. Biol. 24, 265-277. Fishman, H. M., Poussart, D. J . M., and Moore, L. E. (1975a). Noise measurements in squid axon membrane. J . Mernhr. Biol. 24, 281-304. Fishman, H. M., Moore, L. E., and Poussart, D. J . M. (197%). Potassium-ion conduction noise in squid axon membrane. J. Mernhr. Biol. 24, 305-328. Fishman, H. M., Moore, L. E., and Poussart, D. J. M. (1977). Ion movements and kinetics in squid axon. I I . Spontaneous electrical fluctuations. A n n . N . Y . Acrrd. S c i . 303, 3W-423. Fishman, H. M., Moore, L. E., and Poussart, D. (1981). Squid axon K conduction: Adrnittance and noise during short versus long duration step clamps. I n "The Biophysical Approach to Excitable Systems" ( W . J . Adelman, Jr. and D.E. Goldman, eds.), pp. 65-95. Plenum, New York. Fox, J . M. (1976). Ultra-slow inactivation of the ionic currents through the membrane of myelinated nerve. Biochirn. B i o p h y s . Actu 426, 232-244. Frehland, E. (1978). Current noise around steady states in discrete transport systems. Biophys. Chem. 8, 255-265.

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Frehland, E. (1979). Theory of transport noise in membrane channels with open-closed kinetics. Biophys. Struct. Mech. 5, 91-106. Frehland, E. (1980). Current fluctuations in discrete transport systems far from equilibrium. Breakdown of the fluctuation dissipation theorem. Biophys. Chem. 12, 63-71, Frehland, E.,and Stephan, W. (1979). Theory of single-file noise. Biochim. Biophys. Acta 553, 326-341. French, R. J., and Wells, J. B. (1977). Sodium ions as blocking agents and charge carriers in the potassium channel of the squid giant axon. J . Gen. Physid. 70, 707-724. Hamill, 0.P.. Marly, A.. Neher. E., Sackmann, B.. and Sigworth, F. J. (1981). Improved patch-clamp techniques for high resolution current recordings from cells and cell-free membrane patches. Pf/iigcr.s Arch. Gesumte Physiol. 391, 85-100. Hill, T. L., and Chen, Y. (1972). On the theory of ion transport across the nerve membrane. IV. Noise from the open-close kinetics of K' channels. Biophys. J . 12, 948-959. Hille. B. (1967). A pharmacological analysis of the ionic channels of nerve. Thesis, Rockefeller University; University Microfilms, Ann Arbor, Michigan (No. 689584). Hodgkin, A. L., and Huxley, A . F. (1952). A quantitative description of membrane current and its application to conduction and excitation in nerve. J. Physiol. (London) 117, 500-523. Holden, A . V.,and Rubio, J. E. (1976). A model for flicker noise in nerve membranes. Biol. Cybern. 24, 227-236. Holden, A. V . , and Rubio, J. E. (1978). Retardation currents in excitable membranes and models of flicker noise. Biol. Cybern. 30, 45-54. Hooge, F. N . (1976). I/f noise. Physica 83B, 14-23. Hooge, F. N., and Gaal, J. L. M. (1970). Fluctuations with a l/f spectrum in the conductance of ionic solutions and in the voltage of concentration cells. Philips Res. Rep. 26,77-90. Horn, R., Patlak, J., and Stevens, C. F. (1981). Sodium channels need not open before they inactivate. Nature (London) 291, 426-427. Johnson, J. B. (1928). Thermal agitation of electricity in conductors. Phys. Rev. 32,97-109. Katz, B . , and Miledi, R. (1970). Membrane noise produced by acetylcholine. Nature (London) 225, 962-963. Keynes, R. D., Bezanilla, F., Rojas, E., and Taylor, R. E. (1975). The rate of action of tetrodotoxin on sodium conductance in the squid giant axon. Philos. Trans. Soc. London Ser. B 270, 365-375. Kolb, H . A., and Frehland, E. (1980). Noise-current generated by carrier-mediated ion transport at non-equilibrium. Biophys. Chem. 12, 21 -34. Kolb, H. A , , and Lauger, P. (1977). Electrical noise from lipid bilayer membranes in the presence of hydrophobic ions. J . Membr. Biol. 37, 321-345. Kolb, H. A,, and Lauger, P. (1978). Spectral analysis of current noise generated by carriermediated ion transport. J. Membr. B i d . 41, 167-187. Kolb, H. A., Lauger, P., and Bamberg, E. (1975). Correlation analysis of electrical noise in lipid bilayer membranes: Kinetics of gramicidin A channels. J . Membr. Biol. 20, 133-154. Kubo, R. (1975). Statistical mechanical theory of irreversible processes. General theory and simple applications to magnetic and conduction problems. J . Physiol. Soc. Jpn. 12, 570-586. Lauger, P. (1975). Shot noise in ion channels, Biochim. Biophys. Acta 413, 1-10. Lauger, P. (1978). Transport noise in membranes. Current and voltage fluctuations at equilibrium. Biochim. Biophys. Acta 507, 337-349. Lax, M. (1960). Fluctuations from the nonequilibrium steady state. Rev. Mod. Phys. 32, 25-64.

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Levinson, S. R., and Meves, H. (1975). The binding of tritiated tetrodotoxin to squid giant axon. Philos. Trans. R . Soc. London Sera B 270, 349-352. Llano, I., and Bezanilla, F. (1980). Current recorded from a cut-open giant axon under voltage clamp. Proc. N u t / . Acud. Sri. U . S . A . 77, 7784-7486. Lundstrom, I., and McQueen, D. (1974). A proposed l/f noise mechanism in nerve cell membranes. J . Theor. B i d . 45, 405-409. Mauro, A,, Conti, F., Dodge, P., and Schor, K.(1970). Subthreshold behavior and phenomcnologic;il impedance of the squid giant axon. J . Gen. Phy.sio/. 55, 497-523, Meves, H. (1978). lntramembrane charge movement in squid giant nerve fibres. In “lons in Macromolecular and Biological Systems” (E. H. Everett and B. Vincent. eds.), pp. 284-302. Scientechnica, Bristol. Michalides, J . P. L. M . , Wallaart, R. A. M., and DeFelice, L. J. (1973). Electrical noise from PVC-membranes. PJugers Arch. G e s a m f e Physiol. 341, 97-104. Moore, J . W., Narahashi, T., and Shaw, T. I. (1967). An upper limit to the number of sodium channels in nerve membrane‘? J . Physiol. (London) 188, 99-105. Moore, L. E., Fishman, H. M., and Poussart, D. J. M. (1979). Chemically induced K’ conduction noise in squid axon. J . Memhr. B i d . 47, 9Y-112. Neher, E. (1980). Unit conductance studies in biological membranes. In “Techniques in Cellular Physiology” (P.F. Baker, ed.), pp. 4-19. Elsevier, Amsterdam. Neher, E., and Sackmann, B. (1976). Single-channel currents recorded from Inembrdne of denervated frog muscle fibres. Nature (London) 260, 799-802. Neher, E., and Stevens, C. F. (1977). Conductance fluctuations and ionic pores in membranes. Annu. R e v . Biophys. Bioeng. 6 , 345-381. Neher, E., and Zingsheim, H. P. (1974). The properties of ionic channels measured by noise analysis in thin lipid membranes. Pjiigers Arch. Gesamie Physiol. 351, 61-67. Neumcke, B . (1975). l/f membrane noise generated by diffusion process in unstirred solution layers. Biophys. Struci. Mech. 1, 295-309. Neumcke, B. (1978). l/f noise in membranes. Biophys. Struci. Merh. 4, 379-399. Neumcke, B., Schwarz, W., and Starnpfli, R. (1980a). Differences between K channels in motor and sensory nerve fibres of the frog as revealed by fluctuation analy Arrh. Crsumte Physiol. 387, 9-16. Neumcke, B . , Schwarz, W., and Stampfli, H. (1980b). Modification of sodium inactivation in myelinated nerve by Anernonia Toxin I1 and iodate. Analysis of current fluctuations and current relaxations. Biochim. Biophys. Acia 600, 456-466. Nonner, W. (1980). Relations between the inactivation of Na channels und the immobilization of gating charge in frog myelinated nerve. J . Physiol. (London) 299, 573-603. Nyquist. H . (IY2X). Thermal agitation of electric charge in conductors. Phys. Reu. 32, 110-1 13.

Onsager, L. (1931). Reciprocal relations in irreversible processes. 11. Phys. Rev. 38, 22652279. Papoulis, A. (1965). “Probability, Random Variables, and Stochastic Processes.” McGrawHill, New York. Poussart. D. J . M. (1971). Membrane current noise in lobster axon under voltage clamp. Biophys. J . 11, 21 1-234. Ritchie, J. M., and Rogart. R. B. (1977). The binding of saxitoxin and tetrodotaxin to excitable tissue. Rev. Physio/. Biochem. Pharrnucol. 79, 1-45. Rudy, B. (1978). Slow inactivation of the sodium conductance in squid giant axons. Pronase resistance. J . Physiot. (London) 283, 1-21. Sauvk, K., and Bamberg, E. (1978). l/f noise in black lipid membranes induced by ionic channels formed by chemically dimerized gramicidin A. J . Membr. B i d . 43,317-333.

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Siebenga, E., and Verveen, A. A. (1971). The dependence of the l/f noise intensity of the node of Ranvier on membrane potential. Proc. Eur. Biophys. Congr., Is(, pp. 219-223. Siebenga, E., and Verveen, A. (1972). Membrane noise and ion transport in the node of Ranvier. Biomembranes 3, 473-482. Siebenga, E., Meyer, A., and Verveen, A. A. (1973). Membrane shot noise in electrically depolarized nodes of Ranvier. Pjugers Arch. Gesarnte Physiol. 341, 97-104. Siebenga, E.,De Goede, J.. and Verveen. A . A . (1974). The influence of T r X , DNP, and TEA on membrane flicker noise and shot effect noise of the frog node of Ranvier. PJiig. Arch. Gesarnte Physiol. 351, 25-34. Sigworth, F. J. (1977). Sodium channels in nerve apparently have two conductance states. Nutiue (London) 270, 265-267. Sigworth, F. J. (1980a). The variance of sodium current fluctuations at the node of Ranvier. J . Physiol. (London) 307, 97-129. Sigworth, F. J . (1980b). The conductance of sodium channels under conditions of reduced current at the node of Ranvier. J . Physiol. (London) 307, 131-142. Sigworth, F. J . (1981). Covariance of nonstationary sodium current fluctuations at the node of Ranvier. Biophvs. J . 34, 1 11-133. Sigworth, F. J . , and Neher, E. (1980). Single Na+ channel current observed in cultured rat muscle cells. N o t w e (London) 287, 447-449. Stevens, F. (1972). Inferences about membrane properties from electrical noise measurements. Biophys. J . 12, 1028-1047. Stevens, C. F. (1977). Study of membrane permeability changes by fluctuation analysis. Nutrire (London) 270, 39 1-396. Strichartz, G . K.. Rogart, R. B . , Ritchie, J. M. (1979). Binding of radioactively labeled saxitoxin to the squid giant axon. J . Mambr. Biol. 48, 357-364. Stiihmer, W., and Conti, F. (1979). The effect of high extracellular potassium on the kinetics of potassium conductance of the squid axon membrane. Annu. Meet. Dtsch. Ges. Biophys.. p. 84. Swenson, R. P., and Armstrong, C. M. (1981). K + channels close more slowly in the presence of external Kt and Rb’. Nafure (London) 291, 427-429. Van den Berg, R. J., De Goede, J., and Verveen, A. A. (1975). Conductance fluctuations in Ranvier nodes. PJlugers Arch. Gesamte Phy.sio/. 360, 17-23. Van den Berg, R. J., Siebenga, E., and DeBruin, G. (1977). Potassium ion noise currents and inactivation in voltage-clamped node of Ranvier. Nature (London) 265, 177-179. Van der Ziel, A. (1970). “Noise: Sources, Characterization, Measurements.” Prentice-Hall, New York. Verveen, A. A., and DeFelice, L. J. (1974). Membrane noise. Prog. Biophys. Mol. B i d . 28, 189-265. Verveen, A. A , , and Derksen, H. H. (1965). Fluctuations in membrane potential of axons and the problem of coding. Kybernetik 2, 152-160. Wanke, E., DeFelice, L. J., and Conti, F. (1974). Voltage noise and current noise in space clamped squid giant axon. Pfliiprs Arch. Gesamle PhVsio/. 347, 63-74.

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CURRENT TOPICS IN MEMBRANES AND TRANSPORT, VOLUME 22

Membrane Surface Charge

Introduction.. . . ............................ Channel Surface............................ Neutralization of Channel Surface-Charge Density. ........................ Relation of Average Surface-Charge Density to Channel Surface-Charge Density . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Modifications of S VI. Significance . . . . . VII. Summary. . . . . . . . References . . . . . . I. 11. 111. IV.

407 408 412

I. INTRODUCTION

Since both lipids and proteins are present in the membrane and each contain groups that are charged at the appropriate pH, it is not surprising that many biological membranes possess surface charges. The membrane of the squid giant axon is no exception, and the surface charge for this membrane has been previously reviewed (Gilbert, 1971 ; Lakshminarayanaiah, 1976, 1977; Lakshminarayanaiah and Bianchi, 1977). There is experimental evidence for a surface charge on both the outer and inner surfaces of the squid giant axon membrane. In this article we summarize this evidence, with emphasis on more recent work. We also describe how this surface charge relates to membrane channel proteins. One method of determining membrane surface charge is to measure the electrophoretic mobility of cells. This approach provides an estimate of the average charge density over the entire cell membrane. Another method-the one that we emphasize in this article-utilizes the voltage407 ISBN 0-12-153322-0

408

DANIEL L. GILBERT AND GERALD EHRENSTEIN

dependent properties of membrane ionic channels to provide estimates of surface charge density near the channels. The voltage-dependent properties actually depend on the electric field in the membrane, not on the voltage measured with electrodes. Therefore, when agents are added to change the effect of the surface charge on the membrane electric field, graphs describing these properties as a function of membrane voltage translate along the voltage axis. The membrane surface contains ionic groupings that may contribute to the membrane surface charge. There has been much speculation on thc nature, location, and significance of these ionic sites. From electrophoretic studies, it is known that many cells contain a negative outer surface charge of about 1 electronic charge per 2000 A* (EM, 1967), which corresponds to an average distance of about 45 A between the negative fixed charges. It has not yet been determined which membrane constituents are responsible for these charges. Since electrophoretic measurements relate to average properties over the entire cell membrane, this surface charge docs not necessarily havc any relation to the surface charge seen by membrane ionic channels. II. CHANNEL SURFACE-CHARGE DENSITY: THEORY

The manner in which surface charge influences the membrane electric field, and thereby shifts graphs of properties that depend on electric field along the voltage axis, can be seen in Fig. 1. In this figure, the external potential is assumed to be zero and the internal potential measured with an electrode is -65 mV. In the absence of surface charge, thcsc would also be the potentials at the membrane surfaces. When surface charges are present, they give rise to a component of electric field at each surface and to a redistribution of ions to form diffuse double layers. The double layers are usually confined to a few angstroms at the membrane surfaces. Figure 1 shows the potential profiles of the double layers in a typical axon membrane and also shows that the potential at each surface is reduced by the amount ofthe potential across the double layer at that surface. Thus, expected shifts along the voltage axis can be determined by calculating the double-layer potentials. The equation relating voltage shift, surface-charge density, and ionic concentration (Gilbert and Ehrenstein, 1969; Gilbert, 1971) is df [ K M exp

(&Vt) + 13 [i [exp (gVt) =

G

ci

-

1]}-’’*

(1)

i= 1

where d, is the average spacing between surface charges; K , equilibrium

409

MEMBRANE SURFACE CHARGE

C

EXTERNAL PHASE

INTERNAL PHASE

>

-r W 0

z

* -65

W

[L

W

LL

LL

E -46

~

~

-

~

~~

~~~

-J

5 $ -65 II-

?

I

-7e

FIG. 1. Potential profile across squid axon membrane. The distance is not drawn to scale. From Gilbert (197 I ).

binding constant; M , concentration of binding ion: z , valence of binding ion; F , Faraday constant (96,500 C/mol); R , gas constant (8.314 J/kmol); T, absolute temperature; G, 41,800 (DT)-n.5(A2/electroniccharge) (mole K/liter)o,s.D is the dielectric constant. For water at IWC, the value of G = 270 (A*/electronic charge) (mole/liter)o.s;n , number of types of ions in solution; V , , voltage shift; ci , concentration of ith ion; zi , valence of ith ion. We consider three mechanisms for changing the electrical double layer at a membrane surface. These are binding, screening, and modifying the surface charge. With reference to Eq. (11, binding corresponds to increasing the concentration M of a specific ion that binds to surface charge, screening corresponds to increasing the concentrations ci of any ions in solution, and modifying the surface charge corresponds to changing the average spacing between surface charges dt . In this section we consider binding and screening. In Section V we consider modification of surface charge. If the voltage shift is plotted as a function of the logarithm of the binding ion concentration, a titration curve is obtained, as illustrated in Fig. 2. If the binding constant is increased, this curve is shifted to the left with no limits. If the binding constant is decreased, the relative importance of screening compared to binding increases. The curve marked K = 0.0001 mM-' is a curve in which weak binding and screening both occur. The limit as to how much the titration curve can be shifted to the right corresponds to K = 0, where only screening occurs. This limit is shown in Fig.

41 0

DANIEL L. GILBERT AND GERALD EHRENSTEIN

+NoE l l eIncreased Ionic Slrenqlh cl To Added CQ Due

-501

-60

7

- I O - ~ IO-1

I

10’

lo2

10’

lo4

lo5

lo6

lo7

CALCIUM CONCENTRATION ( m M )

FIG. 2. ‘Theoretical binding and screening effects of adding calcium chloride to the external solution.

2 as a slight shift to the right. Experimentally, increased binding caused by increased concentration of a neutralizing ion is always accompanied by increased screening. For pedagogic purposes, however, we have drawn a dashed curve in Fig. 2 marked “no ionic strength effect” to correspond to the hypothetical case of increased binding ( K = 0.0001 mM-I) without increased screening. The three curves in Fig. 2 are very similar, especially at low calcium concentrations. Thus, binding and screening, which are different ways of neutralizing surface charge, have rather similar effects on the voltage across the double layer. In actual practice, the distinction between these two influences can be made using cation substitution. Since all cations of the same concentration and valence should have the same screening effect, any difference must be peculiar to the binding or other biological property of the cation. If the hydrogen cation is used, then the screening effect is negligible, since the lowest pH that can be reached is about 3, corresponding to a hydrogen ion concentration of 1 mM. Thus, Hille (1968) and Mozhayeva and Naumov (1970, 1972a-c), using this cation substitution procedure, have

MEMBRANE SURFACE CHARGE

41 1

shown for the frog node that binding is the mechanism by which cations influence the voltage across the double layer. With the assumption that the surface charge is temporally and spatially uniform, the voltage shift described by Eq. (1) should occur with no change in shape or amplitude for parameters that depend upon the electric field across the membrane. As we will indicate subsequently, however, this requirement does not apply to all surface-charge models. By contrast, all surface-charge models predict shifts along the voltage axis. Two examples of voltage-dependent parameters that are commonly used to measure shifts along the voltage axis, and hence to determine membrane surface charge, are relative conductance and the time constant for the conductance to reach steady state. Voltage clamp experiments do not provide values of relative conductances, but rather provide values of membrane currents. In order to obtain membrane conductance, it is necessary to divide the current by the appropriate driving force. Hodgkin and Huxley (1952) used a linear driving force in their squid axon model. An improvement of accuracy can be obtained by direct determination of driving force by measurement of instantaneous membrane currents (Fohlmeister and Adelman, 1982). The membrane surface-charge density determined from measurements of shifts along the voltage axis is applicable only to the region of the membrane near the ionic channel whose properties are being considered. More specifically, the measurements relate to the region near the voltage sensor of the ionic channel. The particular location of discrete surface charges relative to the voltage sensor influences the membrane electric field. Thus, Eq. (l), which was derived for a uniform surface charge, may be called into question. Since Cole (1969) first considered the implications of the discreteness of surface charge for biological membranes, a number of papers (Brown, 1974; Nelson and McQuarrie, 1975; Sauve and Ohki, 1979; Enos and McQuarrie, 1981) have addressed this issue. As clearly summarized by Brown (1974), both uniform surface-charge models and discrete surfacecharge models fit experimental titration curves about equally well, but give average spacing between surface charges that differ by as much as a factor of two. The magnitude and direction of this difference depends on the exact position of the discrete charges relative to the charge sensor. In fact, the assumption of uniform surface charge results in estimates of charge spacing that lie between the estimates based on two examples of discrete charge distribution considered by Brown (1974). Thus, in the absence of detailed information about the specific location of discrete surface charges, it seems appropriate to use the uniform sur-

412

DANIEL L. GILBERT AND GERALD EHRENSTEIN

face-charge model and Eq. ( I ) to determine approximate surface-charge densities. If the surface charge in the vicinity of a channel consists of a very small number of discrete charges, then, as shown by Attwell and Eisner (IY78), a change in shape can accompany the shift of a graph of a voltage-dependent parameter along the voltage axis. The reason for this can best be seen for the case of a single surface charge. In this case, when a binding ion is added, there are two classes of channels-those whose surface charge is neutralized and those whose surface charge is not neutralized. The overall response to a voltage pulse, therefore, has components corresponding to two different electric fields. This, in turn, leads to a shape change. There is another reason for a voltage-dependent parameter to exhibit a change in shape as surface charge is neutralized. The surface charge of an open state of a channel may be different from the surface charge for a closed state. Gilly and Armstrong (1982) proposed such a difference to explain why addition of zinc causes a much larger shift along the voltage axis of sodium-channel opening kinetics than of sodium-channel closing kinetics. This type of effect could result in a change of shape or amplitude for a time-constant parameter that depends upon both opening and closing rates. In view of the possibility that there are small numbers of discrete channels or that there are differences in surface charge between different states of a channel, shifts that are accompanied by changes in shape and/ or amplitude and differences in shifts for different parameters do not, in themselves, argue against the presence of membrane surface charge near a channel. 111. NEUTRALIZATION OF CHANNEL SURFACE-CHARGE DENSITY In accordance with Eq. (I), the surface charge of a given channel can bc determined by titration with an ion that tends to neutralize the surface charge by binding or by screening or both. An example of such a titration curve is shown in Fig. 2. In practice, it may be difficult to obtain data over a large range of concentrations, and sometimes only a few points have been obtained. Table I shows the average spacing between surface charges found in the squid giant axon and lists values for both sodium and potassium channels for both the internal and external membrane surfaces. For each case, there is good agreement among all measurements. The surface-charge densities at the external surfaces are very similar for the sodium channel

41 3

MEMBRANE SURFACE CHARGE

TABLE I AVERAGESPACING BETWEEN SURFACE CHARGES I N SQUID GIANTAXONS dt

(A)

Reference

internal Internal

27 26

Chandler et a / . (1965) Carbone ei a / . (1981)

Na

External

11

Na

External

8

Begenisich (1975) based on data of Frankenhaeuser and Hodgkin ( 1957) Carbone ei a / . (1978)

K

Internal

>30

K

Internal

>30

K

External

I1

K K K

External External External

II 8 13

Channel

Membrane surface

Na Na

Based on data of Begenisich and Lynch (1974) Based on data of Wanke et a / . ( 1979) Begenisich (1975) based on data of Frankenhaeuser and Hodgkin (1957) Gilbert and Ehrenstein (1969) Carbone et al. (1978) Fohlmeister and Adelman (1982)

and the potassium channel. Also, for both channels, the surface-charge density is much larger at the external surface than at the internal surface. The actual value of the surface-charge density at the internal surface of the potassium channel has not yet been measured, but it must be small, since neither the addition of calcium ions (Begenisich and Lynch, 1974) nor the addition of protons (Wanke et al., 1979) caused significant shifts along the voltage axis. The surface-charge densities referred to in Table I are only for binding or screening experiments on squid giant axons. if the charge density of a channel depends primarily on the structure of the channel, itself, and if all channels of a certain type, such as sodium channels, are rather similar, it would be expected that the surface-charge densities for other preparations would be similar to those listed in Table I. Table I1 shows the average spacing between surface charges found for preparations other than the squid giant axon. It includes relatively recent data on tunicate eggs and snail neurons in addition to the data previously tabulated by Lakshminarayanaiah (1977). In view of the variety of methods used, there is generally good agreement among the various preparations listed in Table I1 and there is also generally good agreement between the average spacing between surface charges in Table I1 and the comparable values for the squid giant axon listed in Table I.

414

DANIEL L. GILBERT AND GERALD EHRENSTEIN

TABLE I1 AVFRAGE SPACING BETWEEN SURFACE CHARGES ON Ex I ~ R N A I MEMBRANE SURFACE F O R VARIOUS PREPARATIONS

Channel

Preparation

4 (A)

Na

Frog node

15

Na Na Na

Frog node Frog node Toad

20 9

Na Na Na Na Na

Toad Crayfish Myxicolu Myxicola Tunicate egg

R 7 9 II 9

K

K

Frog node Toad

20 17

K K K

Toad Myxicola Myxicola

14 18 9

Ca Ca

Tunicate egg Snail neuron

13

9 9

Reference Gilbert and Ehrenstein (1970) based on data of Hille (1968) Drouin and Neumcke (1974) Hille er a / . (1975) Begenisich (197s) based on data of Brismar (1973) Vogel (1974) d’Arrigo (1973) Schauf ( 1975) Begenisich (1975) Ohrnori and Yoshii (1977) Mozhayeva and Naumov (1970) Begenisich (1975) based on data of Brismar (1973) Vogel (1974) Begenisich (1975) Schauf (1975) Ohrnori and Yoshii (1977) Wilson et a / . (1983)

IV. RELATION OF AVERAGE SURFACE-CHARGE DENSITY TO CHANNEL SURFACE-CHARGE DENSITY

As mentioned before, electrophoretic measurements indicate that the external surfaces of biological membranes have an average surfacecharge density of about 1 charge per 2000 A2. This relatively low density is consistent with measurements using alamethicin as a probe to explore the inner and outer surface potential of the frog node of Ranvier (Cahalan and Hall, 1982). Alamethicin, a peptide antibiotic, penetrates the lipid phase of the membrane and produces an ionic channel. The observed voltage shifts with calcium indicated that the outer surface of the lipid phase has very little negative charge. The external surface-charge densities at ionic channels shown in Table I are about 10-20 times larger than the overall external membrane surface-charge density. This indicates that most of the external surface charge in the vicinity of a channel comes from the channel structure itself. Since the channel structure has a relatively large surface charge, the

MEMBRANE SURFACE CHARGE

415

question can be raised whether this channel surface charge can account for the overall surface charge of the entire axonal membrane. If we assume that the surface charge on the channel is fairly uniform, the channel contribution per unit area to the overall membrane surface charge would be about 10-20 times larger than an average area of membrane. However, only a small fraction of the membrane area is taken up by channels. Even for an extremely high channel density of 1000 channels per square micrometer and an area per channel of about 1000 AZ,the fraction of the total membrane area taken up by channels is only about 1%. Thus, even though the protein channel contributes most of the surface charge seen by the potential sensor of the channel, the lipids and nonchannel proteins contribute most of the surface charge over the entire membrane surface. V.

MODIFICATIONS OF SURFACE CHARGE

We have previously considered neutralization of membrane surface charge by binding o r screening. In this section, we consider modifications of membrane surface charge corresponding to changes in d, of Eq. ( I ) . These modifications are brought about by chemical agents and may result in either an increase or a decrease in surface charge. Since not much of this work has been performed on squid axons, we will discuss other membrane surfaces where such modifications have been made. It will be interesting to note in the future whether the results of this section also apply to the squid axon membrane. Batrachotoxin (BTX) produces a voltage shift in the hyperpolarizing direction for both the sodium conductance and the sodium activation time constant in the frog node (Khodorov and Revenko, 1979) and in neuroblastoma (Huang et al., 1982). The magnitude of the time constant is also decreased. Aconitine shows similar effects in the myelinated frog nerve (Schmidt and Schmitt, 1974). Cahalan and Pappone (1981) noted that trinitrobenzenesulfonic acid (TNBS) increased the negative surface charge of frog skeletal muscle. These authors assumed that TNBS reacted with amino groups to produce this increase in negativity. They observed a shift in the sodium inactivation parameter and a similar shift for its time constant. The change in the magnitude of the time constant was within twofold. Fluorescent dyes have been used to determine the membrane potentials of many cells (Conti, 1975; Waggoner, 1976; Cohen and Salzberg, 1978). The binding of these dyes to the membrane may modify the surface charge (Krasne, 1980). For example, bromosulfophthalein, an anion dye, increases the negative surface charge of mitochondria1 membranes (Sch-

41 6

DANIEL L. GILBERT AND GERALD EHRENSTEIN

wenk rt ul., 1977). Also, a fluorescent dye may be used to probe membrane surface charge. The fluorescence of the cationic 9-aminoacridine is quenched by the chloroplast thylakoid membrane, indicating a negative surface charge of this membrane (Searle and Barber, 1978). Chiu et ul. (1980) used the binding of the fluorescent dye ANS- as an assay to determine the surface potential of the sarcoplasmic reticulum in skeletal rnuscle. Their measurement corresponded to a surface potential of - 10 to - 15 mV. It has been suggested that salicylate increases the negative surface charge in sheep cardiac muscle. This would produce an increase in the potassium ion concentration on the surface, and this increase would stimulate the sodium-potassium pump (Cohen er ul., 1979). Cyanate increases the electrophoretic mobility of human erythrocytes, presumably by blocking amino groups and consequently producing an increase in the surface charge (Durocher et al., 1973). On the other hand, chlorprornazine (Tenforde rt uf., 1978) reduces t h e electrophoretic mobility of erythrocytes, presumably by decreasing the surface charge. Other chemicals that appear to decrease the surface charge are guanides, which decrease the negative charge on mitochondria (Schafer, 1976). Small doses of X-irradiation increase the surface charge in polymorphonuclear granulocytes and FL cells as measured by an increase in the electrophoretic mobility of these cells, whereas largc doses cause a decrease (Redmann and Reichel, 1977). The decrease in surface chargc due to X-irradiation is dependent upon the presence of calcium (Sato et ul., 1979). Vitamin E deficiency produces a slight increase of the inner mitochondrial membrane negative surface charge. This conclusion was reached by using a spin-label partitioning technique for measuring surface charge (Quintanilha et ul., 1982). The partitioning of a cationic electron paramagnetic resonance spin label detergent has also been used to determine membrane surface potentials (Quintanilha and Packer, 1977).

VI.

SIGNIFICANCE

Surfacc-charge effects can lead to repetitive firing. Calculations made by Huxley (1959) using the Hodgkin-Huxley equations and assuming that surface-charge affccts only the gating parameters indicate that a change in surface potential causes a change in resting potential whose magnitude is about one-half that of the surface potential change. For example, if the surface potential were to become more positive by 10 mV, the resting

41 7

MEMBRANE SURFACE CHARGE

membrane potential as measured in the bulk phases would become depolarized by 5 mV. Guttman and Barnhill (1970) pointed out that if the surface potential were to become 8 mV more negative, repetitive firing would occur. The repetitive firing often seen in the squid axon when the external calcium concentration is reduced may be the result of a more negative surface potential caused by a reduction in external calcium concentration (Guttman and Barnhill, 1970). Surface-charge effects can modify drug action. A negative surface charge makes the cation activity larger at the surface than in the bulk phase, as indicated by a,

=

ab exp(zFV,lRT)

(2)

where as is the activity of the species at the surface; a b , activity of the species in the bulk solution; z , valence of the species; V , , surface potential. However, the electrochemical potential of the species is the same in the bulk phase as that at the surface. The difference in the activities between the surface and bulk phases is counterbalanced by the surface potential. If the effect of a drug is dependent upon its electrochemical potential, then altering the surface potential of the membrane should not influence the action of the drug. However, if the effect of a drug depends upon its activity, then the activity at the membrane surface is a function of the surface potential. Electrochemical potentials determine ionic movements through membrane channels, whereas activities usually determine the kinetics of active transport processes. An example of the influence of surface charge on drugs is the action of tetrodotoxin and saxitoxin in myelinated nerve (Hille et al., 1975). Surface charge on the surface of the T tubules in skeletal muscle may play a role in controlling electromechanical coupling (Dorrscheidt-Kafer, 1976, 1979). It has also been proposed that surface charges can regulate membrane protein (Ostroumov and Vorobiev, 1978) and membranebound enzymes, possibly by altering substrate concentrations at the surface (Wojtczak and Nalecz, 1979). During energization of the mitochondria1 inner membrane, the outer surface of this membrane becomes more negative (Kamo et al., 1976; Quintanilha and Packer, 1977). It thus appears, at least for this membrane, that the negative surface charge is not a constant, but is changing as a function of its activity. These changes can provide a means for regulation of cellular function. It has also been suggested that changes in surface charge might cause pinocytosis and formation of microvilli (Anderson, 1979) and that cellcell interaction due to electrostatic forces can be influenced by the surface charge (Ninham and Parsegian, 1971; Dolowy and Holly, 1978).

418

DANIEL L. GILBERT AND GERALD EHRENSTEIN

VII.

SUMMARY

The surface-charge density at ionic channels is considerably larger than the average surface-charge density over the membrane as a whole. The surface-charge density at ionic channels is also considerably larger on the external surface than on the internal surface. These generalizations apply not only to squid axon membranes, but also to a wide variety of other membranes. There arc a number of potentially important effects of the surfwe charge\. These include modulation of the function ofcclls by regulation of the surface charge and influences on the binding of drugs. REFERENCES Andersson, G . ( 1979). Self-electrophoresis of membrane molecules and chromosomes: Unifying hypothesis of cell cycle controls. J. Theor. Biol. 77, 1-18. Attwell, D., and Eisner, D. (1978). Discrete membrane surface charge distributions. Effect of fluctuations near individual channels. Biophys. J. 24, 869-1175, Begenisich, T. (1975). Magnitude and location of suiface charges on Myxicokc giant axons. J . G m . Physiol. 66, 47-65. Begenisich, T., and Lynch, C. (1974). Effects of internal divalent cations on voltageclamped squid axons. J . Gen. Physiol. 63, 675-689. Brismar, T. (1Y73). Effects of ionic concentration on permeability properties of nodal membrane in myelinated nerve fibres of Xenopus lueuis. Potential clamp experiments. Acid Physiol. Scand. 87, 474-484. Brown, R. H., Jr. (1974). Membrane surface charge: Discrete and uniform modelling. Prog. Biophys. M o l . B i d . 28,341-370. Cahalan. M . I)., aiid Hall, A. (19x2). Aliimethicin channels incorpoi-ated i n t o frog nndc of Kanvier: Ciilcium-induced inactivation and membrane s u i f i ~ echaiges. 1. Gcn. Phv.viol. 79, 4 I 1-430. Cahalan, M. D., and Pappone, P. A. (1981). Chemical modification of sodium channel surface charges in frog skeletal muscle by trinitrobenzene sulphonic acid. J . Physiol. (I.ondon) 321, 127- 139. Carbone, E., Fiordvanti, R., Prestipino, G., and Wanke, E. (1978). Action of extracellular pH on Na' and K' membrane current5 in the giant axon of L o l i p ) [ ~ i d g ~ Ji .. Meriihr. ~. nioi. 43,295-31s. Carbone, E., lesta, P. L., and Wanke, E. (1981). Intracellular pH and ionic channels in the Loligo uulgctris giant axon. Biophys. J . 35, 393-413. Chandler, W. K., Hodgkin, A. I.., and Meves, H. (1965). The effect of changing the internal solution on sodium inactivation and related phenomena in giant axons. J . f'hysiol. (London) 180, 821-836. Chiu. V . C. K . , Mouring, D., Watson, B. D., and Haynes, D. H. (1980). Measurement of surface potential and surface charge densities of sarcoplasmic reticulum membranes. J . Mernhr. B i d . 56, 121-132. Cohen, I., Noble, D.,Ohha, M., and Ojeda, C . (1979). The interaction of ouabain and salicylate on sheep cardiac muscle. J . Physiol. (London) 297, 187-205. Cohen, I , . > and Salzberg. B. (1978). Optical measurement of membrane potential. Rev. Ptiysinl. Biochem. Pharmacd. 83, 35-88.

MEMBRANE SURFACE CHARGE

41 9

Cole, K. S. (1969). Zeta potential and discrete vs. uniform surface charges. Biophys. J. 9, 465-469. Conti, F. (1975). Fluorescent probes in nerve membranes. Annu. Reu. Biophys. Bioeng. 4, 287-3 10. D’Arrigo, J . S . (1973). Possible screening of the surface charges on crayfish axons by polyvalent metal ions. J. Physiol. (London) 231, 117-128. Dolowy, K., and Holly, F. J. (1978). Contribution of interfacial tension changes during cellular interaction to the energy balance. J . Theor. Biol. 75, 373-380. Dorrscheidt-Kafer, M. (1976). The action of Ca2+,Mg2+and H+ on the contraction threshold of frog skeletal muscle. Evidence for surface charges controlling electro-mechanical coupling. f‘fliigers Arch. 362, 33-41. Dorrscheidt-Kafer, M. (1979). The interaction of ruthenium red with surface charges controlling excitation-contraction coupling in frog sartorius. Pjiigers Arch. 380, 181-187. Drouin, H.,and Neumcke, B. (1974). Specific and unspecific charges at the sodium channels of the nerve membrane. PJliigers Arch. 351, 207-229. Durocher, J. R., Glader, B. E., and Conrad, M. E. (1973). Effect of cyanate on erythrocyte membrane surface charge. Proc. Soc. Exp. Biol. Med. 144, 249-251. E M , R. (1967). Fixed charge in the cell membrane. J. Physiol. (London) 189, 351-365. Enos, B. E., and McQuarrie, D. A. (1981). The effect of discrete charges on the electrical properties of membranes. 11. J . Theor. Biol. 93,499-522. Fohlmeister, J. F., and Adelman, W. J. (3982). Periaxonal surface calcium binding and distribution of charge on the faces of squid axon potassium channel molecules. J. Mernhrcrnr Biol. 70, 115-123. Frankenhaeuser, B., and Hodgkin, A. L. (1957). The action of calcium on the electrical properties of squid axons. J. Physiol. (London) 137, 218-244. Gilbert, D.L. (1971). Fixed surface charges. In “Biophysics and Physiology of Excitable Membranes” (W. J. Adelman, Jr., ed.), pp. 359-378. Van Nostrand-Reinhold, Princeton, New Jersey. Gilbert, D. L., and Ehrenstein, G. (1969). Effect of divalent cations on potassium conductance of squid axons: Determination of surface charge. Biophys. J. 9, 447-463. Gilbert. D.L., and Ehrenstein, G. (1970). Use of a fixed charge model to determine the pK of the negative sites on the external membrane surface. J . Gen. Physiol. 55, 822-825. Gilly, W. F., and Armstrong, C. M. (1982). Slowing of sodium channel opening kinetics in squid axon by extracellular zinc. J . Gen. Physiol. 79, 935-964. Guttman, R., and Barnhill, R. (1970). Oscillation and repetitive firing in squid axons. J . Gen. Physiol. 55, 104-1 18. Hille, B. (1968). Charges and potentials at the nerve surface: Divalent ions and pH. J . Gen. Physiol. 51, 221-236. Hille, B . , Ritchie, J . M., and Strichartz, G. R. (1975). The effect of surface charge on the nerve membrane on the action of tetrodotoxin and saxitoxin in frog myelinated nerve. J . Physiol. (London) 250, 34P-35P. Hodgkin, A. L., and Huxley, A. F. (1952). A quantitative description of membrane current and its application to conduction and excitation in nerve. J. Physiol. 117, 500-544. Huang, L. M., Moran, N . , and Ehrenstein, G . (1982). Batrachotoxin modifies the gating kinetics of sodium channels in internally perfused neuroblastoma cells. Proc. N d . A c a d . Sci. U . S . A . 79, 2082-2085. Huxley, A. F. (1959). Ion movements during nerve activity. Ann. N . Y . Acrid. Sci. 81, 22 1-246. Kamo, N . , Muratsugu, M.. Kurihara, K., and Kobatake. Y. (1976). Change in surface charge density and membrane potential of intact mitochondria during energization. FEBS Lett. 72, 247-250.

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Khodorov, B . I., and Revenko, S . V. (1979). Further analysis ofthe mechanism of batrachotoxin on the membrane of myelinated nerve. Neuroscience 4, 1315-1330. Krasne, S . (1980). lnteractions of voltage-sensing dyes with membranes. 11. Spectrophotometric and electrical correlates of cyanine-d ye adsorption to membranes. Biophys. J . 30, 441-462. I.akshminarayanai;ih. N. ( 1976). Surface charges on menihranes. J . Metnhr. B i d . 29, 243-253. l,akshminarayanaiah, N . (1977). Evaluation of membrane surface charge density: A discussion of some models. Bull. Murh. B i d . 39, 643-662. Lakshminarayanaiah, N., and Bianchi, C. P. (1977). Membranes, ions, and drugs. Adu. G e n . Cell. Pharmacol. 11, 1-70. Mozhayeva, G . N., and Naumov, A. P. (1970). Effect of surface charge on the steady-state potassium conductance of nodal membrane. N u t w e (London) 228, 164- 165. Mozhaeva, G. N., and Naumov, A. P. (1972a). Effect of surface charge on stationary potassium conductivity of the Ranvier node membrane. I. Change in pH of external solution. (Biofizika 17,412-420); Neurosci. Behau. Phvsiol. 6 , 166-174. Mozhaeva, G. N . . and Naurnov, A. P. (1972b). Effect of surface charge on stationary potassium conductance of the Ranvier node membrane. 11. Changes in ionic strength of external solution. (Biofzika 17, 618-622); Neurosci. Rehau. Physiol. 6 , 228-233. Mozhaeva, G . N . , and Naumov, A. P. (1972~).Effect of surface charge on stationary potassium conductmce of Ranvier node membrane. 111. Effect of divalent cations. Biofizika 17, 794-808 (in KUSSbdn). Nelson, A. P., and McQuarrie, D. A. (1975). The effect of discrete charges on the electrical properties of a membrane. 1. J . Theor. B i d . 55, 13-27. Ninham, B. W . , and Parsegian, V. A . (1971). Electrostatic potential between surfaces bearing ionizable groups in ionic equilibrium with physiologic saline solution. J . Theor. Biol. 31, 405-428. Ohmori, H . , and Yoshii, M . (1977).Surfice potential reflected in both gating and permeation mechanisms of sodium and calcium channels of the tunicate egg cell membrane. J . Phvsiol. (I.onc/on)267, 429-463. Ostroumov, S. A.. and Vorobiev, L. N. (1978). Membrane potential and surface charge densitities as possible generalized regulators of membrane protein activities. J . Theor. B i d . 75, 289-297. Quintanilha, A. 'T.,and Packer, I,. (1977). Surface charge of the inner mitochondria1 rnembrane monitored by impermeable spin labels. Riophys 1, 17, 257a. Quintanilha, A. T., Packer. Id.+Davics. 1. M. S . , Kacanelli. T. L., and Davies, K. J. A . (19x2). Membrane effects of vitamin E deficiency: Bioenergetic and surface charge density studies of skeletal muscle and liver mitochondria. Ann. N . Y. Actid. Sci. 393, 32-47. Kedmann, K . . :ind Keichel, G . (1977). Die Wirkung v o n Kimtgenstrahlen auf clas Transmembranpotenlial und die elektrophoretische Reweglichkeit polyrnorphkerniger Granulozyten und FL-Zeilen. Rad. Environ. Biuphys. 14, 21-30. Sato, C.,Nishizawa, K., and Kojirna, K. (1979). Calcium-dependent process in reduction of cell surface charge after X-irradialion. Int. 1. Radiar. B i d . 35, 221-228. Sauve, R., and Ohki, S. (1979). Interactions of divalent cations with negatively charged membrane surfaces. I. Discrete charge potential. J . Theor. B i d . 81, 157-179. Schafer, G. (1976). Commentary. On the mechanism of action of hypoglycemia-producing biguanides. A reevaluation and a molecular theory. Biorhem. Pharmacol. 25, 20052014.

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Schauf, C. L. (1975). The interactions of calcium with Myxicola giant axons and a description in terms of a simple surface charge model. J. Physiol. (London) 248, 613-624. Schmidt, H., and Schmitt, 0. (1974). Effect of aconitine on the sodium permeability of the node of Ranvier. Pfliigers Arch. 349, 133-148. Schwenk, M . , Burr, R., Baur, H., and Pfaff, E. (1977). Interaction of bromosulfophthalein with mitochondria1 membranes: Effect on ion movements. Biochem. Pharmacol. 26, 825-832. Searle, G. F. W., and Barber, J. (1978). The involvement of the electrical double layer in the quenching of 9-aminocridine fluorescence by negatively charged surfaces. Biochirn. Biophys. Acra 502, 309-320. Tenforde, T. S., Yee, J. P . , and Mel, H. C . (1978). Electrophoretic detection of reversible chlorpromazine-HCI binding at the human erythrocyte surface. Biochim. Biophys. Acta 511, 152-162. Vogel, W. (1974). Calcium and lanthanum effects at the nodal membrane. Pftiigers Arch. 350, 25-39. Waggoner, A. (1976). Optical probes of membrane potential. J. Membr. Biof. 27, 317-334. Wanke, E., Carbone, E., and Testa, P. L. (1979). K+ conductance modified by a titratable group accessible to protons from the intracellular side of the squid axon membrane. Biophys. J . 26, 319-324. Wilson, D. L., Morimoto, K., Tsuda, Y . , and Brown, A. M. (1983). Interaction between calcium ions and surface charge as it relates to calcium currents. J. Membr. B i d . 72, 117-1 30. Wojtczak, L., and Nalecz, M. J. (1979). Surface charge of biological membranes as a possible regulator of membrane-bound enzymes. Eur. J. Biochem. 94, 99-107.

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CURRENT TOPICS IN MEMBRANES A N D TRANSPORT. VOLUME 22

Optical Signals: Changes in Membrane Structure, Recording of Membrane Potential, and Measurement of Calcium LAWRENCE B. COHEN Department of Physiology Yale University School of Medicine N e w Haven, Connecticut

DAVID LANDOWNE Department of Physiology and Biophysics University of Miami School of Medicine Miami, Florida

LESLIE M . LOEW Department of Chemistry State University of N e w York at Binghamton Binghamton, N e w York

BRIAN M . SALZBERG Department of Physiology University of Pennsylvania Philadelphia, Pennsylvania

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Light Scattering

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B. Series Resistance.. . . . . . . . . . . C. Mechanisms for Potential-Depe D. The Squid Axon as a Dye-Scre IV. Measurement of Changes in Interna

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423 Copyright 0 1984 by Academlc Press. Inc. All righlr of reproduction in any form reserved. ISBN 0-12-153322-n

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I. INTRODUCTION

The squid giant axon has had a powerful influence on optical measurements from excitable cells. On one hand, the attractiveness of this preparation probably delayed the discovery of optical correlates of individual action potentials by 25 years. Because the surface membrane comprises only a small fraction of the giant axon preparation, optical signals from this membrane are relatively small. When Schmitt and Schmitt (1940) tried to measure birefringence changes in axons, they chose the giant axon from the squid, and were unable to detect a signal. From an estimate of the sensitivity of their apparatus (Schmitt, 1950; Schmitt and Schmitt, 1940), it appears that they would have detected the signal if they had chosen a preparation with a larger proportion of axon membrane, such as a crustacean leg nerve. Hill and Keynes (1949) first observed the changes in light qcattering in a crab nerve that are associated with 5-sec trains of stimuli at SO pcr second. It was not until 20 years later that the use of signal-averaging techniques allowed routine measurcments of the scattering (Cohen and Keyncs, 1968) and bircfringence changes (Cohen et GI., 1968) resulting

1

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5 msec BIREFRINGENCE

FIG.I . Results of the first attempt to measure birefringence changes in a squid giant axon. There was a transient decrease in intensity following the stimulus. The light passing through perpendicular polarizers with the nerve between the polarizers at 45" to the plane of polarization of the incident light was measured. The arrow labeled S indicates the time ofthe stimulus. For rnuasurcnicnts of intrinsic signals thc lcngth of the vertical arrow to the right of the trace represents the stated value of the change in intensity divided by the resting intensity, per sweep. AH records are traced from the original. Temperature, 13°C. (L. B. Cohen and R. D. Keynes, unpublished results.)

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from single stimulations in a crab nerve. As expected, similar experiments carried out on squid giant axons resulted in signals that were much smaller. Figure 1 shows the results of the first measurement of birefringence changes in giant axons. Even after averaging 27,500 trials, the record is quite noisy, although there is a clear transient decrease in intensity that occurs just after the stimulus. Even though the giant axon was not the best preparation for detecting optical signals, improvements in apparatus and the discovery of larger effects have made these measurements easier. A relatively large optical signal, an absorption change measured in a single trial from an axon stained with a merocyanine dye (XVII, see Fig. 11) is illustrated in the top trace of Fig. 2. The action potential is shown on the bottom trace. In this situation, where the signals are large enough to measure accurately, then the possibilities for analysis that the giant axon offers can make it the preparation of choice.

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2 msec

FIG.2. Absorption change during an action potential of an axon stained with dye XVII. A single sweep was photographed after digitization; no averaging was used. For this and other figures where absorption changes are illustrated, the length of the vertical arrow to the right represents the stated value of the change in absorption in a single sweep divided by the resting absorption. The incident light filter was centered at 750 nm. The response time constant of the light-measuring system was 170 psec. The axon was stained in a 0.1 mgiml solution of the dye. (From Ross e / a / . , 1977.)

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Four different kinds of optical signals have been examined on squid axons.

I . Changcs in intrinsic optical properties of axons have bccn measured in an attempt to study the alterations in axon structure that gcncrate or accompany the action potential. This kind of cxpcriment is described in Section I I . 2 . Absorption and fluoresccncc changes from axons stained with a variety of dyes (extrinsic signals) were first measured with the goal of learning about membrane structure. However, almost all these dye signals proved to depend exclusively and linearly on membrane potential. Therefore they could not provide information about the structural basis of the dramatic increases in membrane permeability that give rise to the action potential. At present the main interest in these signals is that they allow optical measurements of membrane potential when electrode measurements of potential are difficult or impossible. Section 111 is devoted to these extrinsic signals. 3 . Both intrinsic and extrinsic signals have been found whose time course far outlasts the time course of the action potential. These include light scattering (Hill and Keynes, 1949; Cohen and Keynes, 1971), birefringence (Watanabe and Terakawa, 1976), and absorption of stained axons (Warashina, 1979). Axon swelling or changes in axoplasmic proteins have been proposed as mechanisms that would account for these very slow signals (Hill, 1950; Cohen and Keynes, 1971; Watanabe and Terakawa, 1976). Slow signals are not discussed further in this article. 4. The first biological use of metallochromic indicator dyes to measure changes in concentration of intracellular free calcium was in a squid giant axon, This kind of experiment is described in Section IV. II. INTRINSIC SIGNALS Two intrinsic optical properties of axons have been shown to change during the action potential-birefringence and light scattering. A. Birefringence

The early measurements of the birefringence of squid axons during voltage-clamp steps suggested that the birefringence changes were potential dependent (Cohen el d., 1968, 1971). The results in Fig. 8 of Cohen et al. (1971) indicated that birefringence depended on the square of mem-

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brane potential. A more recent measurement by Landowne et al. (1983) is illustrated in the left panel of Fig. 3. Consistent with the nonlinear voltage dependence, larger optical signals (top traces) were found during hyperpolarizations than during equivalent depolarizations. In addition, Landowne et al. (1983) found that the birefringence change occurred more slowly in the depolarizing direction than in the hyperpolarizing direction. Further, when 30 m M colchicine was added (Fig. 3B) to the internal perfusion solution, both the inward currents (bottom trace) and the birefringence signal during the depolarization were dramatically reduced. A similar reduction was obtained with P-lumicolchicine, a drug that does not act on microtubules. This correlation between reduction in inward current and alteration of the birefringence response is not universal, since tetrodotoxin blocks the current without affecting the birefringence change. In the presence of colchicine the birefringence signal reverses direction at smaller depolarizations than in the absence of colchicine. Thus one possible interpretation of the result in Fig. 3B is that colchicine binding leads to a large shift in the voltage-birefringence relationship as well as a large shift in the current-voltage relationship.

0 mV

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FIG.3. The effect of colchicine on the birefringence response of an internally perfused squid axon. The upper traces represent the potential impressed across the axon membrane. The middle traces are the corresponding birefringence responses, and the lower traces are the membrane currents. The arrows beside the optical records represent a change in light intensity of 3 x 10 >, and increase in light intensity is plotted downward. (A) control; (B) after 7 min of internal perfusion with 30 mM colchicine. In this and subsequent figures where voltage-clamp steps are used, hyperpolarizing was downward, inward current was downward. Axon from Lolig(:op d e i , diameter 0.39 mm; temperature, 0°C; 100 Na/400 K ; optical trace filtered at 30 kHz, 1024 sweeps. (From Landowne c t ( I / . , 1983.)

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6. Light Scattering

Some components of the light-scattering changes that occur during action potentials are potential dependent (Cohen et a)., 1972a). Howevcr, measurement of light scattering during voltage-clamp steps (Fig. 4) quickly showed that some of the scattering signal was not directly potential dependent, since the scattering changes were very much larger during depolarizations than during hyperpolarizations and the time courses of the scattering changes were qualitatively different from the time course of the change in potential. Figure 4 illustrates scattering measured at two angles, 90" and at more forward angles. In neither case is the scattering Forward angles

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FIG. 4. Light-scattering changes (heavy lines) at forward angles and at right angles during voltage-clamp steps. At both angles, there were large changes during the depolarizing step and only small changes during the hyperpolarizing step. Top two traces, scattering; middle trace, potential; bottom trace, current density. The holding potential was the resting potential. The response time constant of the light-measuring apparatus was 70 psec; 900 sweeps were averaged.

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change during the depolarizing step potential dependent; furthermore the signals at the two different angles differ in time course, suggesting that their origins cannot be the same. To determine whether the large changes found during the depolarizing step were dependent on current or permeability, we compared the 90" scattering resulting from brief 50- and 92-mV depolarizing steps. The 50mV step caused an increase in sodium permeability, which resulted in a large inward current. The 92-mV step also caused a large increase in sodium permeability, but the current was small because the potential reached was near the equilibrium potential for sodium ions. So there were large permeability increases for both steps, but only the 50-mV step resulted in a large current. Figure 5 illustrates the results of such an experiment for the 90" scattering. Only the 50-mV step, with its large current, gave rise to the scattering increase; no scattering change resulted from the 92-mV step, with its small current, and we concluded that the scattering change was dependent on current rather than permeability. It was then necessary to determine the current dependence of the scattering change. Since the increase in Fig. 5 reached a peak some 5-10 msec after the end of the current, it clearly did not depend directly on the instantaneous value of the current. Further experiments (Cohen et al., 1972b) showed that the size of the scattering change depended on the time integral of the current, not on the ion carrying the current. However, the result in Fig. 5 shows clearly that the signals occur more slowly than the time integral. The forward-angle scattering change also depended on the time integral of the current, but its time course was more rapid; its time course could be exactly matched by the time course of the integral of the current (Cohen et al., 1972b). It was concluded that the slower, rightangle scattering change resulted from a volume change in the periaxonal space that was dependent on a transport number effect. The origins of the faster forward-angle scattering changes have not been identified. 111.

DYE MEASUREMENTS OF MEMBRANE POTENTIAL

A. Potential Dependence

Shortly after the discovery of the intrinsic optical signals that accompany excitation, Tasaki el al. (1968) measured changes in fluorescence of stained axons during action potentials. These fluorescence (and later absorption) changes were shown to be potential dependent (Cohen et al., 1970, 1974; Conti and Tasaki, 1970; Patrick et al., 1971; Davila et al., 1974; Conti, 1975; Ross et al., 1977; Gupta et al., 1981; Grinvald et al.,

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ET AL.

50 mV

92 mV

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5x10-s

U 2 msec FIG.5 . Changes in 90"light scattering (heavy lines), resulting from two different depolarizing steps (bottom trace). A 50-mV step (continuous curve, bottom trace) led to a large increase in conductance and a large inward current (continuous curve, middle trace). A large scattering increase resulted. The large, 92-mV, depolarizing step (dashed curve, bottom trace) was near the equilibrium potential, and thus the current (dashed curve, middle trace) was much smaller, even though the conductance increase was still large. When the current was reduced, there was no clearly demonstrable scattering change. The holding potential was the resting potential. This axon had been microinjected with tetraethylammonium bromide, final concentration 24 mM, to block delayed outward currents. The responsetime constant of the light-measuring system was 610 psec; 256 sweeps were averaged. (From Cohen el al., 1972b.)

1982) although there was some early disagreement about this conclusion (Conti et al., 1971; Tasaki ut at.. 1972). The result illustrated in Fig. 2 shows that relatively large signal-to-noise ratios can be obtained in single trial measurements from squid axons when the bandwidth of the light-measuring system was restricted. When a much shorter response time constant (5 psec) was used, the result illustrated in Fig. 6 was obtained. The average of 32 light-measurement trials is shown as the dots, and the simultaneous electrode measurement is shown as the smooth line. When scaled in the manner shown, it is clear

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I miec

FIG. 6. Changes in absorption (dots) of a giant axon stained with dye XVII during a membrane action potential (smooth trace) recorded simultaneously. The change in absorption and the action potential had the same time course. The incident light filter had a peak transmission of 750 nm;32 sweeps were averaged. The response time constant of the lightmeasuring system was 5 psec. (From Ross ef al., 1977.)

that the absorption change had a time course very similar to that of the potential change, suggesting that absorption was sensitive to changes in membrane potential rather than the ionic currents or the membrane conductance increases that occur during the action potential. This suggestion was tested by measuring absorption during voltage-clamp steps (Fig. 7). During both the hyperpolarizing and the depolarizing steps the absorption signal had the same shape as the change in membrane potential. When four potential steps were imposed and the size of the absorption change was plotted against the size of the potential step, a straight line was obtained. The absorption signal of this dye was linearly related to membrane potential over the range 2200 mV from the resting potential.

B. Series Resistance There was, however, an arbitrary adjustment of compensation for the series resistance in the experiment illustrated in Fig. 7. Results from earlier voltage-clamp experiments (Fig. 8 of Davila P I a/., 1974) showcd that without series-resistance compensation the optical signal during a depolarization had a component with a time course similar to that of the

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3 msec

PIG.7. Changes in absorption of an axon stained with dye XVll (top trace) during hyperpolarizing and depolarizing potential steps (middle trace). The bottom trace is currcnt density. The absorption changes had the same shape as the potential changes and were insensitive to the large currents and conductance changes that occurred during the depolarizing step. The holding potential was the resting potential. The incident light filter had a peak transmission at 750 nm; 128 sweeps were averaged; the time constant ofthe light-measuring system was 20 Fsec. The axon was stained in aO.O1 mdml solution of the dye. (From Ross et a / . . 1977.)

ionic currcnt. This component was substantially diminished whcn a somewhat arbitrary umount of Compensation was used. Howcvcr, in the experimcnt illustratcd in Pig. 7, the compensation was completely arbitrary, t h c amount of compensation chosen to yield an absorption change during the depolarizing and hyperpolarizing steps that was most symmetrical. Salzberg and Bezanilla (1983) have repeated this kind of experiment, and, instead of adjusting the compensation to make the optical recording appear sanitary, they have used the optical measurement to determine the size of the series resistancc. This measurement is illustrated in Fig. 8. The lower trace shows the depolarizing and hyperpolarizing potential steps of magnitude V, and Vh, respectively. The upper trace shows the resulting changes in extrinsic absorption, expressed as transmitted intensities, Td and Ti,. Then, I ; , : T h : :(V,,t l,,,R\e,-):V h . Since I,,, is mcasured independently, together with the voltages and transmitted intensities,

RT

(VhTdITh - Vd)IZm

yields the value of the total series resistance directly (in ohm-cm2), with-

OPTICAL SIGNALS

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FIG. 8. Illustration of the method used to determine the value of the series resistance optically. The lower trace shows the depolarizing and hyperpolarizing potential steps of magnitude V , and Vh, respectively. The upper trace shows the resulting changes in extrinsic absorption, recorded simultaneously. and expressed as tranbmitted intensities, Td and Th. R,,, = (VhTd/Th- Vd)/Im. Series resistance was uncompensated. Diameter = 511 pn. (LP 307). The axon was stained with dye XVll (see Fig. 11). (From Salzberg and Bezanilla, 1983.)

out introducing any compensation. The results of this kind of measurement were in agreement with estimates made from electrical measurements on the same axons.

C. Mechanisms for Potential-Dependent Signals

Once it is established that the absorption and fluorescence changes are potential dependent, and thus that they occur in the axon membrane, what further can be said about their origin? The possible origins of these extrinsic signals fall into two classes, nonstructural and structural. The nonstructural class includes those mechanisms that could give rise to extrinsic signals without any change in membrane structure, and the structural class includes those mechanisms by which membrane structure changes cause extrinsic signals. When extrinsic signals were first discovered there was hope that they might provide structural information. This hope has not been fulfilled (Cohen, 1973). All recent attempts to study the mechanism(s) responsible for the extrinsic optical signals have attempted

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to understand the signals without postulating changcs in membrane structure. The spectra of the potential-dependent signals may provide clues about their origins. If the change in absorption in a squid axon stained with a merocyanine dye (dye I of Cohen et ul., 1974) is measured as a function of wavelength, the result shown in Fig. 9 is obtained. This spectrum has a relatively complicated shape, but it can be synthesized rather accurately from the difference spectra of monomers and dimers of the dye derived from spectral studies of dye bound to lipid vesicles (Waggoner and Grinvald, 1977). Although the extraction of monomer and dimer spectra involves some assumptions, the ability to resynthesize the spectrum in Fig. 9 supported the idea that a potential-dependent , monomer-dimer equilibrium would explain the signals found with this dye. However, it has been difficult to explain the results of three subsequent experiments (L. B. Cohen, A. Grinvald, K. Kamino, and B. M . Salzberg, unpublished results) with this simple hypothesis. First, the spectrum was found to depend on the concentration of the dye in the staining solution. If a lower concentration was used, the crossover point (near 550 mm in Fig. 9) was shifted toward longer wavelengths. (In fact. this result argues against any two-state mechanism, not only the specific monomer-dimer mechanism that was originally proposed.) Second, the emission spectrum in fluorescence measurements was strikingly dependent on the concentration of dye in the solution used to stain the axon. Third, the shape of the excitation spectrum depended on the emission wavelength that was used for the

' h (nm) AA

PIG.9. The wavelength dependence of the changes in absorption of unpolarized light (A), measured during 50-mV depolarizing steps on axons stained with dye I. The peak tranmihsion of Ihc inteiference filters ( 10-nm width at half-height) i s shown on the abscissa. The ordinate is the intensity changes in absorption measurement corrected for the efficiency of thc apparatus. The direction of the absorption change was wavelength dependent, reversing sign twice. There were absorption increases at wavelengths between 550 and 600 nm, and absorption decreases between 450 and 530 nm and between 610 and 640 nm. The solid curves connecting the experimental points were drawn arbitrarily. (From Ross et al., 1977.)

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measurements. None of these findings could be explained by a simple potential-dependent shift in a monomer-dimer equilibrium, where the monomers were highly fluorescent and the dimers weakly or nonfluorescent. Two additional mechanisms, dye rotation and energy transfer from monomer to dimer, have been suggested to explain these and other features of signals found with this dye (Dragsten and Webb, 1978; Ross et al., 1977). The complicated shape of the difference spectrum for dye I, in particular the long-wavelength reversal seen at 600 nm, was consistent with a monomer-dimer mechanism and is inconsistent with any mechanism that led to a simple wavelength shift in the absorption spectrum. One mechanism that does predict a simple wavelength shift is electrochromism, a direct effect of the large changes in membrane electric field on the dipole structure of the dye molecules. Loew et af. (1979) and Loew and Simpson (1981) have in fact found that dyes such as dye XXVI (see Fig. 11) do have absorption changes whose spectra are consistent with a simple shift in absorption spectrum. A simple shift is also observed in the fluorescence excitation spectrum. Furthermore, these signals occur rapidly (<5 psec) (Loew et al., 1984), also consistent with the possibility of an electrochromic effect. However, recent measurements of the fluorescence emission changes, both on squid axons and on bilayers, suggest that electrochromic mechanisms probably cannot account for all of the fluorescence signals seen with these dyes (Loew et al., 1984). In Fig. 10A the emission spectrum component of the fluorescence change on bilayers of dye XXVI is shown. As expected for an electrochromic mechanism, this signal does reverse sign, but the reversal occurs at 545 nm instead of 565 nm, where the fluorescence is maximum. In addition the fluorescence change is not symmetrical about the reversal. When a similar experiment was carried out on squid axons, the result of Fig. 1OB was obtained. In this case, the sign reversal is further blue-shifted (to 520 nm) from the wavelength of maximum emission and the fluorescence change is even less symmetrical about the reversal wavelength. Thus, the fluorescence signal for this dye cannot be totally understood in terms of an electrochromic mechanism. Even more difficult to interpret in terms of spectral shifts is the fact that the absorption changes of some membrane-bound dyes were in the same direction at all wavelengths (for example, dye XVII,Ross et al., 1977), a result that is inconsistent with any simple wavelength shift in an absorption spectrum, as required by either electrochromism or a monomerdimer mechanism. When the spectra of these dyes were measured using polarized light, the signals were highly dichroic and very complex indeed (Ross et af., 1977; Warashina, 1979, 1980; Gupta et al., 1981). Warashina

A 11

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3pM Di -6. ASPPS

2 0 0 H z . 100mV ex = 462 nm

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FIG. 10. (A) Emission response of dye XXVI to a 100-mV potential step across the hemispherical bilayer. A standard tungsten lamp was used to correct the results for the wavclength dependence of the photomultiplier tube. (B) Relative and absolute (uncorrected) emission response specfrii of squid axon stained with dye XXVI. Voltage-clamp steps of 50 mV and excitation at 470 nm. (From Loew e / d.,1984.)

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(1979, 1980) suggested that these spectra may be explained by a potentialdependent movement of dye between two binding sites. At one site the dye would be oriented perpendicular to the plane of the membrane and in the electric field. At the second site the dye would be randomly oriented and in a region of smaller field strength. The finding of a marked dichroism implies that there is a dye-related birefringence signal. In the squid giant axon this extrinsic birefringence signal is large (Gupta et al., 1981). However, in spherically symmetric preparations, signals in opposite quadrants can have opposite signs, and the signal-to-noise ratios are thereby reduced (Boyle and Cohen, 1980). The extrinsic absorption and fluorescence signals have been used to measure membrane potential in a variety of physiological and neurobiological preparations. Some of these uses have been reviewed (Cohen and Salzberg, 1978; Waggoner, 1979; Freedman and Laris, 1981). D. The Squid Axon as a Dye-Screening Preparation

More than 1000 dyes have been tested on squid axons in an effort to find probes with relatively large signals and a minimum of pharmacological and photodynamic effects (Cohen et al., 1974; Ross et al., 1977; Gupta et al., 1981). Figure 11 illustrates the structures, wavelengths, fractional changes, and signal-to-noise ratios obtained with seven dyes that seemed relatively promising as potentiometric probes on squid axons. The fractional changes are uniformly small. Even though the signal-to-noise ratios in squid axon experiments are large (Fig. 2), the membrane area is much smaller in most applications, and the signal-to-noise ratios are reduced. Because of the small size of the signals, efforts to find better dyes and to improve the apparatus have continued. The relative signal sizes obtained on squid axons and four other preparations of excitable cells have been compared. Barnacle ganglia (Grinvald et al., 1981), neuroblastoma cells (Grinvald et al., 1982), salamander olfactory bulb (Orbach and Cohen, 1983), and rat cortex (Orbach et al., 1982) were each screened with 15-50 dyes that had been tested on squid axons. These comparisons suggested that signals could be obtained with each of the dye types illustrated in Fig. 1 1 on all five preparations. However, individual dyes in the five classes often exhibited dramatically different relative signal sizes from preparation to preparation. For example, one styryl dye, an analog of XXVI, had a very small signal in experiments on squid axons but was one of the best available dyes on rat cortex. Thus, it may be necessary to test several dyes of each class with each new preparation. The results obtained on squid axons are applicable only in a general way.

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MEASUREMENT OF CHANGES IN INTERNAL CALClUM CONCENTRATION

Dr. John Cooper first suggested to Dr. Larry Pinto that metallochromic indicator dyes might be useful for detecting changes in internal free-calcium concentration. The squid axon was an obvious test preparation, since it would be easy to inject the dye internally and easy to control calcium influx with the use of depolarizing voltage-clamp steps. In an axon injected with arsenazo I11 and bathed in artificial seawater containing a high concentration (1 12 mM) of calcium, a decrease in transmitted light intensity at 660 nm in response to individual depolarizing voltage-clamp steps was detected by averaging over many trials (Fig. 12C) (Brown et al., 1975). This signal was reversibly attenuated by replacing most of the external calcium, [Ca],, with magnesium (Fig. 12D). The difference between the signals recorded in high and low [Ca], was not attributable to a change in light-scattering or other optical properties of the axon itself. N o difference was detectable at X = 750 nm, where the absorption of the dye is negligible. Furthermore, at a wavelength shorter than the isosbestic point for Ca-dye complexation (530 nm), an absorption change of opposite sign was elicited by voltage-clamp steps. If it is assumed that all the calcium enters the axon uniformly during the first 300 psec of the depolarizing step, that there are no diffusion delays, that the complexation reaction had no delay, and given that the lightmeasuring system had a time constant of 170 psec, then the absorption increase should have the time course shown as the faint dotted curve in Fig. 12C. The measured absorption increase lags the calculated curve by 400 psec, suggesting that the Ca-dye complexation reaction must be faster than 400 psec. Baylor el al. (1982) discussed the apparent discrepancy between this result and the results obtained using temperature jump (Scarpa et al., 1978) or stopped flow (Ogawa et al., 1980), which would suggest a substantially slower response time. FIG. 11. Dyes that have relatively large changes in absorptions (abs.), birefringence (biref.), and/or fluorescence (fluor.) when added externally to squid giant axons. The source of dye, the incident wavelength for the largest signal in absorption or fluorescence, the time required for the inward current to be reduced by 50% in oxygenated seawater, the fractional changes, and the signal-to-noise ratios (SIN) are given. Only relatively large signals are indicated. Birefringence measurements were not made for dyes XXlII and XXVI.Sources: H may be obtained from Dr. A. S. Waggoner, Center for Fluorescence Studies, Carnegie Mellon University, 4400 Fifth Ave., Pittsburgh, PA 15213; NK, from Nippon KankohShikiso Kenkyusho Co. Ltd., 2-3 Shimoishii 1 Chome, Okayama-shi, Okayama, Japan; LL, from Dr. L. Loew, Department of Chemistry, SUNY Binghamton, Binghamton, NY. (From Gupta et al., 1981.)

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A

-u-~---

B. COHEN

ET AL.

165 mV

I

FIG. 12. Changes in ahsorption during hyperpolarizing voltngc-clamp steps of a n ai-senazo Ill-injected axon in high-calcium ((3 and low-calcium (D) artificial seawaters. The two superimposed records in trace C were recorded before and after that shown in D. Potential steps are shown in trnce A , and current density recorded in low-calcium seawater is shown in trace B. When the external calcium was reduced, the absorption change resulting from the depolarizing step was also reduced. The holding potential was the resting potential. The dotted curve in trace C is explained in the text. The pcak transmission of the interference filter was 660 nm. (From Brown PI d.,1975.)

Since the report of Brown el ul. in 1975, arscnazo 111 and its analog have been used by other investigators to measure internal calcium first in squid axons and subsequently in many other cells. One group has used both metallochromic dyes to measure calcium and voltage-sensitive dyes to measure potential changes in an effort to study excitation-contraction coupling in striated muscle (Coward-Baylor et al., 1981). V.

CONCLUSIONS

The squid giant axon has had a critical role in the introduction of optical methods for monitoring membrane potential and for monitoring intracellular free calcium using metallochromic indicator dyes. The axon still provides a convenient screening preparation for testing new dyes, but the results obtained in axons cannot be assumed to apply in detail to other preparations. The giant axon will probably continue to be an important preparation in the attempt to study structural events correlated with excitability and in the analysis of the mechanisms responsible for potential dependent dye signals.

44 1

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ACKNOWLEDGMENTS Supported by Grants NS08437, NS137809, GM25190, and NS16824 from the National Institutes of Health. REFERENCES Baylor, S. M., Chandler, W. K., and Marshall, M. W. (1982). Use of metallochromic dyes to measure changes in myoplasmic calcium during activity in frog skeletal muscle fibres. J . Physiol. (London) 331, 139-177. Boyle, M. B., and Cohen, L. B. (1980).Birefringence signals that monitor membrane potential in cell bodies of molluscan neurons. Fed. Proc. Fed. Am. Soc. Exp. B i d . 39,2130. Brown, J. E., Cohen, L. B., De Weer, P., Pinto, L. H., Ross, W. N., and Salzberg, B. M. (1975). Rapid changes of intracellular free calcium concentration: Detection by metallochromic indicator dyes in squid giant axon. Biophys. J . 15, 1155-1160. Cohen, L. B. (1973). Changes in neuron structure during action potential propagation and synaptic transmission. Physiol. Reu. 53, 373-418. Cohen, L. B., and Keynes, R. D. (1968). Evidence for structural changes during the action potential in nerves from the walking legs of Maia squinado. J. Physiol. (London) 194, 85-86P. Cohen, L. B., and Keynes, R. D. (1971). Changes in light scattering associated with the action potential in crab nerves. J . Physiol. (London) 212, 259-275. Cohen, L. B., and Salzberg, B. M. (1978). Optical measurement of membrane potential. R e v . Physiol. Biochrin. Phortncic-ol. 83, 35-88. Cohen, L. B . , Keynes, R. D., and Hille, B. (1968). Light scattering and birefringence changes during nerve activity. Nature (London) 218, 438-441. Cohen, L. B., Landowne, D., Shrivastav, B. B . , and Ritchie, J . M. (1970). Changes in fluorescence of squid axons during activity. B i d . Bull. Mar. B i d . L a b . Woods Hole 139,418-419. Cohen, L. B., Hille, B., Keynes, R. D., Landowne, D., and Rojas, E. (1971). Analysis of the potential-dependent changes in optical retardation in the squid giant axon. J . Physiol. (London) 218, 205-237. Cohen, L. B., Keynes, R. D., and Landowne, D. (1972a). Changes in light scattering that accompany the action potential in squid giant axons: Potential-dependent components. J . Physiol. (London) 224, 701-725. Cohen, L. B., Keynes, R. D., and Landowne, D. (1972b). Changes in axon light scattering that accompany the action potential: Current dependent components. 1.Physiol. (London) 224, 727-752. Cohen, L. B., Salzberg, B. M., Davila. H . V . , Ross, W. N., Landowne, D., Waggoner, A. S . , and Wang, C . H. (1974). Changes in axon fluorescence during activity: Molecular probes of membrane potential. J . Mernbr. Biol. 19, 1-36. Conti, F. (1975). Fluorescent probes in nerve membranes. Annu. Rev. Biophys. Bioeng. 4, 287-310. Conti, F.,and Tasaki, I. (1970). Changes in extrinsic fluorescence in squid axons during voltage-clamp. Science 169, 1322-1324. Conti, F.,Tdsaki, I., and Wanke, E. (1971). Fluorescence signals in ANS-stained squid axons during voltage clamp. Biophysik 8, 58-70. Coward-Baylor, S.. Chandler. W. K., and Marshall, M. W . (1981). Studies in skeletal muscle using optical probes of membrane potential. I n "The Regulation 0 1 Muscle Contraction: Excitation-Contraction Coupling" (A. D. Grinnell and M. A. B . Brazier, eds.). pp. 97-130. Acadcmic Press, New York.

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Davila, H. V., Cohen, L. B., Salzberg, B. M., and Shrivastav, B.B. (1974). Changes in ANS and TNS fluorescence in giant axons from Loligo. J. Membr. B i d . 15, 29-46. Dragsten, P. R., and Webb, W. W. (1978). Mechanism of membrane potential sensitivity of the fluorescent membrane probe merocyanine 540. Biochemistry 17, 5228-5240. Freedman, J. C., and Laris, P. C. (1981). Electrophysiology of cells and organelles: Studies with optical potentiometric indicators. Int. Rev. C y d . Suppl. 12, 177-246. Grinvald, A., Cohen, L. 8..Lesher, S., and Boyle, M. B. (1981). Simultaneous optical monitoring of activity of many neurons in invertebrate ganglia using a 124 element photodiode array. J. Neurophysiol. 45, 829-840. Grinvald, A , , Hildesheim, R., Farber, 1. C., and Anglister, L. (1982). Improved fluorescent probes for the measurement of rapid changes in membrane potential. Biophys. J. 39, 301-308. Gupta, R. K., Salzberg, B. M., Grinvald. A., Cohen, L. B.,Kamino, K., Lesher, S., Boyle, M. B., Waggoner, A. S. , and Wang, C. H. (1981). Improvements in optical methods for measuring rapid changes in membrane potential. 1.Mernhr. Eiol. 58, 123-127. Hill. D. K. (1950). The volume change resulting from stimulation of a giant nerve fibre. J . Phy.sinl. (London) 111, 304-327. Hill, D. K., and Keynes, R. D. (1949). Opacity changes in stimulated nerve. J. Physiol. (London) 108, 278-28 I . Landowne, D., Larsen, J. B.,and Taylor, K.T. (1983). Colchicine alters the nerve birefringence response. Science 220, 953-954. Loew, 1,. M.. and Simpson, L. 1.. (1981). Charge shift probes of membrane potential. A probable electrochromic mechanism for p-aminostyrylpyridinium probes on a hemispherical lipid bilayer. Biophys. J . 34, 3.53-365. Loew, L. M., Scully, S . , Simpson, L., and Waggoner, A. S. (1979). Evidence for a chargeshift electrochromic mechanism in a probe of membrane potential. Narure (London) 281, 497-499. Loew. L. M . , Cohcn, L.B., Salzberg, B . M.. Obaid, A . I,.. and Rezanilla. F. (19x4). Charge shill probes of membrane potential. Cheracterizatiun of aniinostyrylpyridinitim dyes on the squid giant axon (in preparation). Ogawa, Y.,Harafugi, H., and Kurebayashi, N. (1980). Comparison of the characteristics of four metallochromic dycs as potential calcium indicators for biological experiments. J . Lliochem. 87, 1293-1303. Orbach, H. S . , and Cohen, L. B. (1983). Optical monitoring of activity from many areas of the in uitro and in uiuu salamander olfactory bulb: A new method for studying funclional organization in the vertebrate CNS. J. Nenrosci. 3, 22.5 1-32h2. Orbach, H. S., Cohen, L. B.,and Grinvald, A. (1982). Optical monitoring of evoked activity in the visual cortex of the marine rat. Bid. Bull. 163, 389. Patrick, J., Valeur, B., Monnerie, L., and Changeaux, J . P. (1971). Changes in extrinsic fluorescence intensity of the electroplax membrane during elcctrical excitation. J. Mcmhr. Biol. 5, 102-120. Ross, W. N., Salzberg, B. M., Cohen, L. B., Grinvald, A., Davila, H. V., Waggoner, A. S., and Wang, C. H. (1977). Changes in absorption, fluorescence, dichroism and birefringence in stained giant axons: Optical measurement of membrane potential. J . Mernhr. Biol. 33, 141-183. Salzberg, B. M., and Bezanilla, F. (1983). An optical determination of the series resistance in Lolino. J. C k n . Physiol. 82, 807417. Scarpa, A,, Brinley, F. J., and Dubyak, G . (1978). Antipyrylazo 11, a middle range Ca2+ metallochromic indicator. Biochemistry 17, 1378-1 386.

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Schmitt, F. 0. (1950). Morphology in muscle and nerve physiology. Biochim. Biophys. Acra 4, 68-77. Schmitt, F. O., and Schrnitt, 0. H . (1940). Partial excitation and variable conduction in the squid giant axon. J . Physiol. (London) 98, 26-46. Tasaki, I . , Watanabe, A,, Sandlin, R., and Carnay, L. (1968). Changes in fluorescence, turbidity, and birefringence associated with nerve excitation. Proc. N d . Acad. Sci. U . S . A . 61, 883-888. Tasaki, I., Watanabe, A., and Hallett, M. (1972). Fluorescence of squid axon membrane labelled with hydrophobic probes. J . Membr. Biol. 8, 109-132. Waggoner, A. S . (1979). Dye indicators of membrane potential. Annu. Rev. Biophys. Bioeng. 8, 47-68. Waggoner, A. S . , and Grinvald, A. (1977). Mechanisms of rapid optical changes of potential sensitive dyes. Ann. N . Y. Acad. Sci. 303, 217-241. Warashina, A. (1979). Spectral analyses of absorption changes associated with nerve excitation in dye-stained crab nerve. Biochim. Biophys. Acra 554, 51-61. Warashina, A. (1980). Further studies on absorption changes arising in dye-stained nerves during excitation. J . Membr. Biol. 53, 207-213. Watanabe, A., and Terakawa, S . (1976). Alteration of birefringence signals from squid giant axons by intracellular perfusion with protease solutions. Biochim. Biophys. Acra 436, 83 3-842.

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CURRENT TOPICS IN MEMBRANES AND TRANSPORT, VOLUME 22

Effects of Anesthetics on the Squid Giant Axon D . A . HAYDON, J . R . ELLIOTT, AND B . M . HENDRY Physiological Laboratory Uniuersity of Cambridge Cambridge, England

I. Introduction . . . . . . . . . ............................................. A. The Squid Axon a ode1 for Studies of Anesthesia.. . . . . . . . . . . . . . . . . B. Lipid versus Protein Sites.. ......................................... C. Early Studies of Anesthetics on Squid Nerve.. ........................ 11. Effects of Anesthetics on the Action Potential and Sodium and Potassium Currents of the Squid Giant Axon. ....................................... A. Action Potentials.. ................................................. B. Voltage-Clamped Axons. ............................................ 111. Mechanisms of Sodium and Potassium Current Suppression. ................ A. Shifts in the Steady-State Inactivation Curve .......................... B. Shifts in the Steady-State Activation Curve ........................... C. Reduction of the Maximum Conductances gNaand gK . . . . . . . . . . . . . . . . . . D. Time Constants. ................................................... E. Concluding Remarks. . . . . . Appendix: Effect of Membrane Changes on the Steady-State .................................. Inactivation of the Na Channel. References. ............................................................

1.

445 445 447 449 451 451 454 464 464 468 471 473 476 478

INTRODUCTION

A. The Squid Axon as a Model for Studies of Anesthesia

Physiological studies of anesthetic effects on a whole organism are very difficult to interpret. It is particularly hard to investigate the mechanisms of anesthesia at the molecular level using a tissue preparation as complex as the mammalian central nervous system (CNS). For this reason preparations have been sought, both artificial and biological, on which the 445 Copyright 0 1984 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-153322-0

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origins of anesthetic effects can be examined more satisfactorily. In view

of current thinking about the basis of CNS activity and function, the models most often employed are in vitro isolated nerve cell preparations and artificial lipid bilayers. The axonal membrane of the squid is arguably the best understood excitable tissue of all and constitutes an obvious model on which anesthetic molecules can be examined. It is unlikely that general anesthesia results from precisely the same mechanism as impulse blockade in peripheral nerve, but a number of parallels do exist. The strong correlation between hydrophobicity and anesthetic potency holds for both general and local anesthesia, whether expressed as the Meyer-Overton rule or as the free energy of transfer of the CH2 group to an anesthetic site from an aqueous environment (Schneider, 1968; Seeman, 1972; Haydon and Urban, 1983b). The wide rangc of substances known to be gencral anesthetics also cause a reversible blockade of peripheral nerve impulses as well as affecting synaptic transmission. It is widely held that the concentration of a given anesthetic required to block a peripheral nerve is some 3 to 10 times that required to produce general anesthesia (Seeman, 1972). While the existence of this discrepancy is not surprising, its origin is not clear. Several studies have shown that ether and chloroform, in particular, depress synaptic transmission at concentrations lower than those required to block transmission in a peripheral nerve (Sherrington, 1947; Larrabee and Posternak, 1952; Austin and Pask, 1952; Somjen, 1963), but for methanol, cthanol, propanol, and urethane the reverse is true and the nerve is more susceptible than the synapse (Larrabee and Postcrnak, 1952). Such comparisons of anesthetic concentrations required to affect nerve, synapse, and CNS function are further complicated by the observation that small axons are more sensitive to anesthetics. But the extent to which this is an equilibrium phenomenon is not clear (Franz and Perry, 1974; Staiman and Seeman, 1977; Haydon and Hendry, 1982). The anatomical site of general anesthetic action in the CNS is not known, but suggested areas include the reticular activating system (RAS) of thc brainstem and the cerebral cortex (Richards, 1980). Each of these contain numerous very small axons that might be highly sensitive to anesthetics. In peripheral nerve, anesthetics affect impulse conduction velocity and threshold behavior at concentrations below those required to block conduction. Perturbations of this nature may be significant in the CNS, where complex integration of nerve cell activity is required to maintain normal function. In short it is by no means clear whether general anesthetics have their main effects on synapses or nerve axons, and it appears likely that anesthetic effects on both need further study before these ideas can be transferred to the CNS. The sodium and potassium channels of the squid giant axon membrane

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show remarkable similarities to the equivalent channels in amphibian and mammalian nerve. This is seen, for example, in the high-affinity binding of tetrodotoxin (TTX) to all these sodium channels with similar blocking effect. Ionic currents through mammalian sodium and potassium channels can be fitted by equations almost identical to those originally proposed by Hodgkin and Huxley (1952) for squid (Neumcke and Stampfli, 1982). It is such parallels that justify experiments on the squid axon as relevant to mammalian nerve conduction. The sodium channel is not confined to the axonal membrane but is present in the membranes of nerve cell bodies, in cardiac muscle membranes, in both external and internal skeletal muscle membranes, and possibly in the membranes of the fine dendritic processes of central nerve cells (Jack et al., 1975; Noble, 1979). For this reason the distinction between anesthetic effects on axonal conduction and on synaptic function is not clear-cut. Synaptic function is intimately dependent upon sodium-channel activity, and at certain sites in the CNS small anesthetic-induced perturbations of sodium-channel function may be vital. Such sites include the fine dendritic cylinders conveying information from axodendritic synapses to the nerve cell body, and the axon hillock of the cell body where action potentials are initiated and where small changes in threshold may alter the cell firing rate significantly. Aside from these general reasons for interest in anesthetic modulation of ion channels in squid nerve, there is a specific objective for certain workers. The molecular mechanisms by which the sodium and potassium channels conduct ions and are turned on and off by membrane voltage changes are poorly understood. Anesthetic effects on the ionic currents and on the so-called “gating currents” may help to constrain the possible models for normal channel function. This idea has led to the investigation of similarities between anesthetic effects and normal sodium-channel inactivation, but the results are as yet difficult to interpret (Oxford and Swenson, 1979; Swenson and Oxford, 1980; Oxford and Yeh, 1982; Fernandez et al., 1982). B. Lipid versus Protein Sites

There is no general agreement as to whether neutral lipophilic general anesthetic molecules act via a lipid or protein site, and this disagreement extends to their actions on the squid axon. For certain clinical local anesthetics and related molecules (such as lignocaine and procaine), the position is clearer in that good evidence indicates a direct blockade of sodium channels by binding to a site within the pore itself (Hille, 1977; Cahalan, 1978). Local anesthetics acting in this manner, usually with an active charged form, are not the major concern of this article, which will instead

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consider the mechanisms by which neutral lipophilic anesthetics, such as halothane, chloroform, methoxyflurane, the alkanes, and the n-alkanols, inhibit impulse propagation, Some authors hold the view that even this group of substances act by direct binding to membrane proteins (see, e.g., Richards et ul., 1978; Franks and Lieb, 1982). This is a view for which we do not feel there is strong evidence, and one purpose here is to examine the extent to which the observed effects of anesthetics on the squid axon can be explained in terms of certain simple concepts of lipid bilayer structure and lipid-protein interactions. A number of significant differences exist between the effects on the nerve axon of “specific” local anesthetics such as procaine and of general anesthetics such as halothane and n-hexane. Procaine and lignocaine show a high selectivity for blockade of sodium channels compared with potassium channels (Taylor, 1959; Hille, 1977), whereas halothane and the n-alkanes appear to inhibit both channels to an approximately equal degree (Shrivastav et ul., 1972; Haydon and Kimura, 1981; Haydon and Urban, 1983a-c). Lignocaine appears to act in its cationic form from the inside of the axon and has a voltage dependence of block indicative of a site within the channel. Lignocaine and related molecules also show a use-dependent or frequency-dependent rate of block consistent with a model in which access of the cationic local anesthetics to their site of action is possible only when the sodium channel gates are open. This frequency dependence appears to be characteristic of the direct channelblocking molecules, and it is not seen with halothane, methoxyflurane, or diethyl ether in amphibian nerve (Kendig et al., 1979) nor with n-hexane in blockade of the impulse in squid axons (B. M. Hendry, unpublished observations). This lack of observed use dependence in the actions of neutral anesthetics on nerve implies a difference in their mode of action. On the other hand, it would be wrong to argue that any molecule that produces a usedependent block must therefore be acting by direct protein binding. For example, interactions between an anesthetic and the lipid bilayer can produce changes in bilayer structure likely to affect the voltage dependence of the gated ion channels, Such changes include alteration of dipole moments (Haydon, 1975; Reyes and Latorre, 1979) and bilayer thickncss (Haydon et al., 1977). n-Pentane, which thickens artificial bilayers, is known to shift the voltage dependence of the steady-state sodium-channel inactivation curve (h,) in the hyperpolarizing direction (Haydon and Kimura, 1981). This shift will increase the value of the time constant for recovery from inactivation Th at membrane voltages close to the resting potential (Hodgkin and Huxley, 1952; Frankenhaeuser and Hodgkin, 1957). This in turn could lead to an increase in the relative refractory

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period and a differential block of high-frequenc y trains of action potentials that would mimic a use-dependent inhibition. The best reason for scepticism about a direct channel-blocking mechanism for small neutral lipophilic molecules can be discerned by returning to the correlation between potency and hydrophobicity. For small hydrophobic molecules such as cyclopropane, halothane, and carbon tetrachloride, it is difficult to reconcile the success of this correlation for local anesthesia with a site of action inside the pore. In the case of a large amphipathic local anesthetic, it is possible to imagine that it can insert a hydrophobic portion into a nearby hydrocarbon-like phase at the same time as leaving a tertiary or quaternary nitrogen in the channel itself. This explanation is unlikely in the case of a small hydrophobic anesthetic. However, the possibility that these neutral anesthetics act via a hydrophobic site on the sodium-channel protein cannot be ruled out by such arguments.

C. Early Studies of Anesthetics on Squid Nerve

Until the end of the 1970s there had been few studies of general anesthetics on the squid axon and fewer still of anesthetic effects on the voltage-clamped axon. Since about 1978 or so there has been a considerable expansion in this work, and here we shall summarize the position prior to this expansion. The substances that had been examined included ethanol (Moore et al., 1964; Moore, 1966; Spyropoulos, 1957), a series of alkanols (Armstrong and Binstock, 1964), barbiturates (Narahashi et al., 1969), trichloroethylene (Shrivastav et al., 1976), and ketamine (Shrivastav, 1977). All these substances were found to produce a reversible inhibition of action potential propagation, and their effects before complete inhibition included decreased rate of rise of the action potential, decreased conduction velocity, and increased threshold. Spyropoulos (1957) demonstrated that 3-7% ethanol inhibited the membrane action potential in squid and that this effect could be reversed by low temperature or high pressure. Moore et al. (1964) applied ethanol under voltage-clamp conditions and analyzed the resulting data in terms of the variables of the Hodgkin-Huxley equations. They reported that 6% ethanol reversibly reduced both sodium and potassium currents approximately equally, with gNaand gK falling to 66 and 68% of control values, respectively. There was also a slight hyperpolarizing shift of the h, curve. Armstrong and Binstock investigated alkanols between ethanol and pentanol and n-octanol. They reported that the alcohols reduced gNa,and

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their data siiggcst that there is probably a small (-5 mV) shift in the depolarizing direction of the g,,-V relationship. A similar effect appears to have been present in the gK-V relationship with the shortchain alcohols. Thcse two studies did not test accurately for effects , 7,) of sodium and potassium channel on the time constants ( T ~ ?-hr gating. Shrivastav rr ul. (1976) studied the effects of trichloroethylene on voltage-clamped squid axons at concentrations similar to those found during general anesthesia in man. The sodium and potassium currents were reduced by the anesthetic. Sodium currents appeared to be reduced by a combination of a reduction in gNawith a reduction in EN^, the reversal from +50 to +30 mV. The h, curve potential for sodium current (INa), showed a hyperpolarizing shift of about 6 mV and a reduction in slope. The finding of an alteration in ENa is most important, as it implies that the selectivity of the sodium channel is altered by the anesthetic. Such a change in selectivity would lend support to the idea of direct anestheticprotein binding. However, it is likely that the apparent effects of trichloroethylene on E N a were artifactual. These sodium current measurements were performed without series resistance compensation, without subtraction of leakage currents, and in the presence of full potassium currents. In these circumstances accurate records of the current passing through the sodium channel are impossible to obtain. The major source of error in their estimate of ENaappears to be the uncorrected leakage currents, which [as Fig. 3 of Shrivastav ef ul. (1976) shows] dominates any true component of ZNaat membrane potentials greater than +20 mV. In those studies in which ZNa has been accurately obtained by subtraction of TTX traces in the absence of K+ currents, it is interesting that no significant effect on ENa of anesthetics such as ether, halothane, chloroform, methoxyflurane, or n-pentane has been seen (Bean et al., 1981; Haydon and Kimura, 1981; Haydon and Urban, 1981). Prior to recent investigations of anesthetic effects on squid nerves, a relatively small number of substances had been tested. Certain common features had been uncovered, including the tendency to inhibit both sodium and potassium currents in approximately equal measure. However, there seemed to be no clear pattern to explain the relative contribution of changes in total available conductance (gNaand &) and changes in the voltage dependence of the steady-state gating parameters (rn,&, h,, and nm), The effects of anesthetics on the time constants of channel gating were not well quantified. Section I1 of this article describes the data obtained from recent work on the effects of n-alkanes, n-alkanols. ethers, esters, and halogenated hydrocarbons on the squid axon.

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II. EFFECTS OF ANESTHETICS ON THE ACTION POTENTIAL AND SODIUM AND POTASSIUM CURRENTS OF THE SQUID GIANT AXON

This section is concerned with the details of how a range of simple organic substances affects the action potential and currents of the squid giant axon. Much of this information has come from voltage-clamp experiments. The results of such experiments are conveniently discussed in the terms used by Hodgkin and Huxley (1952). However, while this procedure provides a valuable phenomenological framework for the analysis of the suppression of excitability, it reveals nothing directly concerning the molecular origins of the effects. This latter topic will be examined in Section Ill when the available facts have been presented. A. Action Potentials

Surprisingly little has been reported concerning the effects of simple organic molecules on action potentials in single nerve axons. Armstrong and Binstock (1964) examined the membrane (i.e., space-clamped) action potential in the squid giant axon when exposed to various alcohols and commented only that the peak of the spike was suppressed much more than the following hyperpolarization. The effects of n-pentane on propagated action potentials in giant axons of the tropical squid Doryteuthis plei were studied by Haydon et al. (1977). As the height of the action potential decreased, the propagation rate also decreased. Hendry (unpublished results) noticed that the suppression and recovery of propagated action potentials in Loligo forbesi usually involved some hysteresis in the movement of the peak. Examples for n-hexane and halothane are shown in Fig. 1. During the experiments on Doryteuthis plei mentioned above, it was found that in n-pentane solutions the axons often exhibited spontaneous action potentials or gave several spikes from a single stimulus (Fig. 2). These phenomena occurred soon after the axons were exposed to a nerveblocking concentration of the n-pentane and disappeared before the steady-state suppression of the action potential was reached. Furthermore, it was only during the application of the hydrocarbon, never during the reversion to artificial seawater, that the spontaneous activity was observed. Subsequently it was found that many of the test substances mentioned in this article produce similar effects. The concentrations re-

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H

H

H

FIG.1. Effects of n-hexane and halothane on the squid action potential. Photographs on left-hand side show the inhibition produced by 64 f i M n-hexane (upper) and subsequent recovery in anesthetic-free artificial seawater (lower). The axon was stimulated at I-min intervals. Photographs on right-hand side show inhibition by and recovery from S m M halothane. This axon was stimulated at 15-sec intervals. The calibration bars are 20 mV and 1 msec. The icrnpcrature was 6 2 1°C. (Unpublished results of B . M. Hcndry.)

quired are often well below those for impulse blockage. Apart from the suggestion made originally (Haydon ef al., 1977) that an asymmetric change in the surface dipole potentials of the membrane might be responsible, no mechanism for the spontaneous excitability has been proposed. To conclude this brief discussion of the non-voltage-clamped intact axon, it should be mentioned that all the anesthetics examined produced some depolarization. At impulse-blocking concentrations this depolarization varied considerably with the substance and to some extent with the axon; some results are shown in Table I. At lower concentrations (even by x 10) depolarizations of 1-2 mV are still observed.

ANESTHETIC EFFECTS ON THE

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SQUID GIANT AXON

U

5 rnsec

FIG. 2. Train of propagated impulses arising from a single stimulus in the squid giant axon (Doryteuthis plei) during exposure to 0.7 saturated n-pentane in artificial seawater. The record was taken approximately 2 min after contact with the alkane solution. The resting and action potentials in the control state were -60 and 108 mV, respectively. Temperature, 18°C. [From Haydon el ul. (1977); reproduced by courtesy of ElseviedNorth-Holland Biomedical Press.]

TABLE I DEPOLARIZATION OF INTACT AXONSOF Loligo forbesi VARIOUS ANESTHETICS"

Substance n-Butane n-Pentane C yclopropane Cyclopentane Cyclohexane Carbon tetrachloride Chloroform Halothane Diethyl ether Dichloromethane Methoxyflurane Methoxyflurane n-Pentanol n-Octanol

Concentration (mM) 0.675 0.306 7.0 0.76 0.216 1 .o 3.75 3.75 100

10 3 1 14.8 0.29

BY

Depolariza tion imV) 3.3 2.4 3.6 5.8 2.0 6.7 6.5 3.5 3.0 4.6 13.6 8.3 9 1.7

a The concentrations are close to those that reduce the peak Na current by 50%. The depolarizations are averages for the numbers of axons shown in parentheses. Temperature = 6 * I T .

454

D. A. HAYDON ET AL.

B. Voltage-Clamped Axons With a few exceptions, the hydrocarbons, halogenated hydrocarbons, ethers, alcohols, and other surface-active substances all suppress the peak sodium current rather more than the maximum potassium current. The exceptions are found among the halogenated ethers (D. A. Haydon and B. W. Urban, unpublished results). It is not surprising, therefore, that more attention has been paid to the sodium than to the potassium currents, and, inevitably, this will be reflected in the present discussion. 1. SODIUM CURRENTS

The main topics for consideration in this article are the mechanisms by which the sodium current is suppressed by the hydrocarbons, alcohols, etc. It is obviously convenient to work with concentrations of these substances that, in the steady state, produce a reduction in the sodium current of about 50%. A collection of these concentrations for intact giant axons of L . forbesi is presented in Table 11. The concentrations are quite close to those given by Seeman (1972) for the blockage of impulses in frog sciatic nerve. The corresponding concentrations determined during perfu-

TABLE I1 CONCENTRATIONS THATPRODUCE APPROXIMATELY 50% SUPPRESSION OF T H E MAXIMUM Na I N VOLTAGE-CLAMPED INTACT SQUIDAXONSO CURRENT ~

Concentration* (mmol liter I )

Substance n-Butane n-Pentane n-Hexane Cyclopropane Cyclopentane C yclohexane Dichloromethdne Chloroform Carbon tetrachloride Halothane Diethyl ether Methox y flurane Benzyl alcohol

0.68 ( I atrn) 0.28 (0.9 saturated) 0.06 (0.95 saturated) 7.0 ( I atm) 0.7 (0.37 saturated) 0.19 (0.35 saturated) 8 3 1 2 75 1

x

Substance

Concentration' (rnmol liter I )

n-Heptanol n-Oc tanol n-Nonanol n-Decdnol n-Octyl(oxyethylene)3 alcohol n-Dec yl(oxyethylene), alcohol n-Decyl(oxyethylene)4 alcohol Methyl n-octonoate Glycerol 1-monooctanoate Dioctanoylphobphatidylcholine

14.8 3.52 0.93 0.2Y 0.068 0.022 0.44 0.035 0.05 0.35 1.3 0. I5

n-Pentanol 11- Hexanol

Artificial seawater; 6 2 1°C.

* Interpolated from unpublished results of D. A. Haydon and B. W. Haydon and Urban (1983b).

Urban.

455

ANESTHETIC EFFECTS ON THE SQUID GIANT AXON

sion experiments (CsF + sucrose solutions inside; 75% of Na replaced by choline outside) are usually a little higher, though not by more than a factor of two. The reason for this has not been elucidated. Different holding potentials, prepulses, and Na concentrations do not have sufficient effect to account for the discrepancies. A remaining possibility stems from the fact that the Na currents (especially their inactivation) are affected by CsF perfusion (Meves, 1978) and might therefore have different susceptibilities to anesthetics under these conditions. The reversibility of the effects of the anesthetics on the currents in a healthy axon is usually good (290%) though rarely complete. It appears that the natural deterioration of an isolated axon, especially under voltage clamp, is accelerated in various ways by exposure to anesthetics. The nature of the deterioration depends on the type of anesthetic. More detailed comments on this question can be found in Haydon and Urban (I983a-c). One of the clearest conclusions so far reached is that, at the concentrations for 50% suppression of the peak Na current, most if not all aspects of the current are to some extent affected by each of the substances examined. These effects will be described in terms of the Hodgkin-Huxley parameters obtained by analysis of the voltage-clamp current records for axons perfused with CsF. An example of these records (for diethyl ether) can be seen in Fig. 3, and the plots of the peak inward currents versus membrane potential are shown in Fig. 4. These two figures serve to illustrate several of the features found for many of the other test substances. Thus, both the time to the peak and the time for inactivation for the currents is shortened (Fig. 3), the reversal potential is affected slightly or not at all, and the maximum in the current-voltage curve shifts, in this instance to less negative membrane potentials (Fig. 4). The poor reversal at low depolarizations was also a fairly common observation. The analysis of the current records has been described often, but it may A

B

C

FIG.3. Inward currents before (A), during (B), and after (C) exposure of a CsF-perfused axon to 100 mM diethyl ether in artificial seawater. The depolarizations from the holding potential of -70 mV were 20-50 mV in 5-mV steps, SO-I00 mV in 10-mV steps, and 100-160 mV in 20-mV steps. A prepulse to -90 mV of SO-msec duration was applied immediately before each depolariLation. (Data of Haydon and Urban, 1983c.)

456

D. A. HAYDON ET AL.

- 20

- 40

40

20 r

1

I

1

- 8 ' 9 el

V (mV)

1

I

o x *

6o

'

-

0'

80 I

x

a

. 0

X . X

8

-

-400 .

0 0

X

*

a

e

X

0

-800X

0 0

X

X

x

- 1200 0

-

0

Q

xoaFIG.4. Relationship between the peak current ( I , ) and the membrane potential (V) for an axon before (O),during (O), and after ( X ) exposure to 100 m M diethyl ether. The plots were calculated from the records of Fig. 3.

be helpful to give a brief resume of the procedure and of the relevant equations of Hodgkin and Huxley that define the parameters to be discussed. The Na current, Z N ~ for , both the control and test solutions may be fitted by means of an equation derived from those of Hodgkin and Huxley (1952); i.e., INa

=

fkJ1 - exp(-t/~,)l~Lh,(l - exp(-hh)) + exp(-thh)l

(1)

where t is the time from the step depolarization, and T,, and 7 h are, respectively, the time constants for the activation and inactivation of the current. fha is the current that would be reached if no inactivation were to occur. h, is the steady-state inactivation parameter (see below). The fitting procedure yields I h A , T,, and h, for each depolarization. The Na conductance, gNa,can be expressed in terms of the maximum Na conductance gNaby the expression 7h1

g N a = g,,m3h

(2)

where, under the conditions in the experiments to be discussed, m

2

m,(I - e-'"m)

(3)

ANESTHETIC EFFECTS ON THE SQUID GIANT AXON

457

and h

2

h,(] - e-"7h) +

e-'lTh

(4)

Thus m, is the steady-state activation parameter. a. The Maximum Na Conductance (gNa).In Table I11 is summarized a set of results showing the effects of various organic substances on the Na currents of L . forbesi. In column 3 are given the suppressions of the maximum Na current for each substance, and in column 4, the suppression of the maximum conductance. Except in two instances, where the reduction of the currents is relatively small (n-butane and cyclopropane), the maximum conductance is suppressed. This implies that either the number of operational Na channels per unit area of membrane is reduced or the conductance per channel decreases. It is not at present known which of these possibilities actually occurs. In addition, it is apparent that the suppression of the maximum conductance is less in every instance than the suppression of the maximum current. Since the current at any given potential is directly proportional to gNa[through Eq. (2)] it follows that the test substances must affect the currents to some extent through either or both of the m or h factors. Finally, if normalized to Ik/Ip = 0.5, gka/gN,is smaller for the nonpolar, non-surface-active substances (e.g., the hydrocarbons and carbon tetrachloride) than it is for the strongly surface-active substances such as the alcohols. The two ratios are 0.60 and 0.88, respectively.

b. Steady State Inactivation (h=). Several examples of the steadystate inactivation versus membrane potential curves are shown in Fig. 5 . The curves are usually shifted to some extent by the test substances, and the movement AVh of the point at which h, = 0.5 is given in Table 111. The nonpolar molecules, i.e., the hydrocarbons and carbon tetrachloride, all produce substantial shifts in the hyperpolarizing (negative) direction. Surface-active substances such as the alcohols, oxyethylene alcohol, the monoglyceride, and diethyl ether have relatively little effect on the curves, and dioctanoylphosphatidylcholine causes a significant displacement in the positive direction. The halogenated hydrocarbons other than carbon tetrachloride (dichloromethane and chloroform) lie in intermediate positions, whereas halothane seems to belong to the surface-active group. By contrast, methoxyflurane and methyl octanoate, both of which are polar and amphipathic, produce substantial negative shifts akin to the hydrocarbons. It is now known that other esters and aliphatic ketones behave similarly (J. R. Elliott, D. A. Haydon, and B. M. Hendry, unpub-

458

D. A. HAYDON ET AL.

TABLE 111 EFFECTS OF VARIOUS

TYPES OF

ORGANIC' S U B S T A N C F ON T H E PARAMETERS OF T H E

HODGKIN-HUXI E Y EQUATIONS FOR T H E Na CUKRI5NT"

n-Butane n-Pentane n-Hexane Cyclopropane C yclopentane C yclohexane Dichloromethane Chloroform Carbon tetrac hloride Halothane Diethyl ether Methoxyflurane n-Pentanol n-Octanol n-Octyl (oxyethylene)J alcohol Methyl n-octanoate Glycerol 1-rnonooctanoate Dioctdnoyl phosphatidylcholine

0.675 0.275 0.06 7.0 1.33 0.379 15.0 5.0 1 .o 4.3 I00 3.0 14.8 0.33

0.97 0.78 0.74 0.81 0.41 0.60 0.52 0.49 0.47 0.56 0.61 0.51 0.46 0.48

1.0 0.75 0.73 1.07 0.64 0.77 0.80 0.77

-8,s - 14.0

- 13.0 -7.8

- 16.5 - 13.7 -4.3 -8.5

0.72

-7.0

0.85

-1.8

0.90 0.85 0.96 0.86

-3.6 -10.0 2.8

-1.9

2 2

-5.5 -7.3 -6.3 4.4 -4.6 -5.5 9.5 6.3

0.75 0.67 0.58 0.79" 0.85 1.01 0.74 0.78

0.98 0.87 0.90 0.68" 0.73 0.89 0.55 0.67

0.0 8.5 2.0 6.7 16.0 9.8

0.78 0.67 0.55 0.62 0.71 0.79

0.77 0.70 0.62 0.68 0.57 0.79

3 3 4

2 4 4 3

2 2

3 3 4

0.405 0.35

0.58 0.60

0.81 0.76

-2.3 -10.0

8.3 7.0

0.71 0.76

0.83 0.61

2 2

1.46

0.47

0.86

-3.5

14.5

0.74

0.50d

2

0.1

0.71

0.83

7.5

0.74

0.R3d

2

5.3

" Ibllpgives the reduction in the peak inward current, and fiha/g~a is thc corresponding reduction in the maximum Na conductance. AV, and AV, indicate, respectively, the shifts in the midpoints of the steady-state inactivation (h,) and steady-state activation (m,) curves. [ ? # r h l p and [ T ~ / T , ] pare the ratios of the time constants for inactivation and activation in test conditions to those for the control both at the peak of the T curve (see Figs. 7 and 8). The results are averages for the numbers of axons given in the last column and have been taken from Haydon and Urban (1983a-c). Superscript t, test; no superscript, control. Two axons. Three axons. One axon.

lished results). A prominent feature of all the h, curves that undergo appreciable negative shifts is that their slopes decrease. c . Strudy-State Activation ( m m ) .As for steady-state inactivation. effects on this parameter have been described in terms of the movement AV,.,, of the midpoint of the curve of m, against membrane potential.

459

ANESTHETIC EFFECTS ON THE SQUID GIANT AXON 10

p

Q Q

0%

t

n-pentane

"0 0-

Q

I h,

0.5

00

h,

I

. O

dioctanoyl phorphotidylcholine

FIG.5. Effects of lipophilic and amphipathic substances on the steady-state inactivation (h,) curve of the CsF-perfused squid giant axon. The open and the filled circles are for the control and test records, respectively. The latter were taken under quasi-steady-state conditions, i.e., ca. 20 min after exposure of the axon to the test substance. V is the membrane potential. The concentrations were n-pentane, 275 p M ; cyclopropane, 7 mM; chloroform, 7 mM; methyl octanoate, 0.4 mM; methoxyflurane, 3 mM; diethyl ether, 100 mM; halothane, 5 mM; n-octanol, 0.29 mM; dioctanoylphosphatidylcholine,0.1 mM. Temperature, 6 t 1°C. [Results of Haydon and Urban (1983a-c), where other experimental details are given.]

Results for some systems are shown in Fig. 6, and numerical values of AV, for all the substances examined are given in Table 111. Some negative shifts of the rn, curve are observed, but the majority are positive. The negative values are all for hydrocarbons (though the cyclopropane result is positive) and, numerically, the AV, is always less than the AVh. No clear changes occur in the slopes of the m , curves, but the poor accuracy of the data for rn, 5 0.5 may conceal small effects. d . The Time Constants for Inactivation and Activation ( T ~ ,T , ) . Curves of time constants versus membrane potential are shown in Figs. 7 and 8. The peaks of the curves shift in the same direction and approximately to the same extent as do the corresponding steady-state curves. Apart from the shifts, the peaks of the curves are suppressed in almost every instance. The straight-chain hydrocarbons reduce Th more than T,, whereas for the cyclic hydrocarbons, the reverse is true. In

460

D. A. HAYDON ET AL.

FIG.6 . Effects of lipophilic and amphipathic substances on the steady-state activation for the control and test records, respectively. V is the membrane potential. The experimental details were as for Fig. 5 . The concentrations were n-pentane, 306 p M ; cyclopropane, 7 m M ; chloroform, 5 mM; methoxyflurane, 3 mM; diethyl ether, 100 m M ; n-octanol, 0.29 m M . (Results of Haydon and Urban, 1983a-c.) (m,) curve of the CsF-perfused squid giant axon. The open and filled circles are

fact, cyclohexane has no effect on the peak height of Th although it does lower the curve at larger depolarizations. For the other substances at larger dcpolarizations, the Th is always reduced. The 7, sometimes increases slightly, but this is evidently the result of a depression combined with a shiCt in the positive direction, where the shift predominates. The results of Table 111 confirm in some instances the findings of earlier workers. In particular the reduction by alcohols of gNaand the concomitant positive shift of the Na steady-state activation were observed by Armstrong and Binstock (1964). Also, Oxford and Swenson (1979) and Swenson (1979) have noted that Th is reduced by n-alkanols. Although for crayfish rather than squid giant axons, the results of Bean et al. (1981) for halothane and diethyl ether are of special interest, As in the experiments of Haydon and Urban (1983c), it was found that gNaand the h, curve were scarcely affected, while q,was suppressed. Fernandez et al. (1982) examined the effect of chloroform on the charge movement in the squid giant axon. A reduction in the magnitude of the “on” gating current was found, but, in agreement with the results of Haydon and Urban (1983c), no influence on the activation kinetics at relatively high depolarizations. No

46 1

ANESTHETIC EFFECTS ON THE SQUID GIANT AXON

0

0 0

0 cyclopropane

0

methoxyflurane

n-octanol

halothone

0

h‘

h‘ Irnsec)

(rnrec)

0 0

4 -

0

8

0 0

h-

000 0

.

0

0 0

p

0 . 0

0

08 0

. Q o g

0

.

.,O,.oo

0

0.

FIG. 7. Effects of lipophilic and amphipathic substances on the time constant (7J for inactivation of the Na current in the CsF-perfused squid giant axon. The open and filled circles are for the control and test records, respectively. V is the membrane potential. The experimental details were as for Fig. 5. The concentrations were cyclopropane, 7 mM; methoxyflurdne, 3 rnM; halothane, 5 mM; n-octanol, 0.29 mM. (Results of Haydon and Urban, 1983a-c.)

data were reported for lower depolarizations, where the rn-system shift and T, peak suppression become clearer. 2. POTASSIUM CURRENTS

All the substances listed in Table I11 suppress the potassium current to some extent, but in many instances the concentration required to produce 50% suppression is larger than for the peak Na current. For the hydrocarbons and carbon tetrachloride, a comparison of the reductions in peak inward and steady-state outward currents in intact axons has been given (Haydon and Urban, 1983a). On balance, the Na current is the more sensitive, but exceptions are listed and in no case is the difference very great. For the alcohols and several other substances, on the other hand, concentrations some threefold higher are required to suppress the K current to the same extent as for the Na current (Armstrong and Binstock, 1964;

462

-

D. A. HAYDON ET AL.

01

0

-50

0

V (mV)

50

' -50

I

0

V ImVl

54

FIG.8. Effects of lipophilic and amphipathic substances on the time constant (T,,,) for activation of the Na current in the squid giant axon. The open and filled circles are for the control and test records, respectively. V is the membrane potential. Experimental details were as for Fig. 5 . The concentrations were n-pentane, 275 y M ; cyclopropane, 7 m M ; diethyl ether, 100 m M ;n-octanol, 0.29 mM. (Results of Haydon and Urban, 1983a-c.)

D. A. Haydon and B. W. Urban, unpublished results). The reversibility of the suppression of the K current is subject to the same remarks as for the Na current but, unlike the Na currents, certain substances, such as the nalkanols, often produce an over-recovery. This effect can be seen in the results of Moore et al. (1964) for ethanol and has been found for other alcohols by Haydon and Urban. A further feature of the K currents is that some nonionic as well as ionic anesthetics induce a form of inactivation. n-Decanol is a particularly good example of this (Swenson, 1981) (Fig. 9), although lower homologs exhibit the same phenomenon to a lesser extent (Paternostre and Pichon, 1982; D. A. Haydon and B. W. Urban, unpublished results). The quantitative analysis of the effects of anesthetics on the K currents, to give the Hodgkin-Huxley parameters is, unfortunately, fraught with difficulties. Apart from irreversibility (e.g., over-recovery) and the need to introduce a further parameter (for "inactivation"), there are also the

ANESTHETIC EFFECTS ON THE SQUID GIANT AXON

463

5 msec

FIG. 9. Effect of n-decanol (-0.2 mM) on the K current of a voltage-clamped KFperfused squid giant axon in choline artificial seawater containing 0.3 p M tetrodotoxin. The upper record is the control; the lower was taken 45 min after introduction of the n-decanol. The holding potential was -60 mV, and the depolarizations were 20-130 mV in IO-mV steps. Temperature, 6 f 1°C. (Unpublished results of D. A. Haydon and B. W . Urban.)

usual problems associated with the accumulation of K+ in the periaxonal space and hence the difficulty of ascertaining the driving force for the currents. For these reasons, no values of gK,T,, and n, of comparable reliability to those for EN;,, T,, and m, can be given. Some qualitative or semiquantitative results are, however, available. Aliphatic hydrocarbons suppress the maximum K current (Haydon and Urban, 1983a) and for npentane gKand gNaare reduced by comparable amounts by comparable concentrations (Haydon and Kimura, 1981). A similar statement can be made for the n-alkanols from the work of Armstrong and Binstock (1964) and Haydon and Urban (1983b, and unpublished data). n-Pentane reduces T, (by approximately a factor of two at 306 p M ) for membrane potentials less negative than -40 mV (Haydon and Kimura, 1981), and diethyl ether reduces T, for crayfish giant axons (Bean et ul., 1981). Shifts in n,, analogous to those seen for m,, seem to occur for several of the substances listed in Table 111 (D. A. Haydon and B. W. Urban, unpublished data). In summary, the hydrocarbons and alcohols, together with some other substances of Table 111 seem, in the main, to have rather similar effects on Na and K channels.

464

D. A. HAYDON ET AL.

111.

MECHANISMS OF SODIUM AND POTASSIUM CURRENT SUPPRESSION

It is quite clear from the results described in Section 11 that each of the test substances perturbs the currents in several different ways. Further, it is usual for both steady-state and kinetic parameters to be affected. There is no possibility of accounting for the observations in terms of a single physical concept, such as fluidity, and thus, as far as nerve impulse suppression is concerned, a unitary hypothesis is out of the question. In this section it is suggested that the main effects could originate from four distinct phenomena: membrane thickening, surface potential, surface tension, and fluidity changes. It is convenient to discuss first the steady-state inactivation. A. Shifts in the Steady-State Inactivation Curve

Although not alone in producing shifts in the h, curve, the nonpolar molecules have been found to produce much the largest effects, and always in the negative sense. It has been established beyond reasonable doubt that changes in the ionic double layer can cause the h, curve to move on the voltage axis (Chandler et a / . , 1965), but it does not seem likely that the hydrocarbons could act in this way. Not only are they nonionic substances, but also they are known not to have a large influence on the area per molecule of lipids in bilayers (Fettiplace er al., 1971). In fact, since they are not surface active, it is difficult to see how the hydrocarbons could change the membrane surface charge density anything like enough to account for the observed shifts. It is also important to remember that the hydrocarbons reduce the slope of the h, curve, and this is not in accordance with an explanation in terms of ionic double layers. Henzocaine has been shown to shift the h, curve in the negative direction in frog nodal membranes, and this has been accounted for by postulating adsorption at a site in the Na channel such that closure of the inactivation gate occurs (Hille, 1977). This proposal will be considered later when the results for methyl octanoate and methoxyflurane (Table 111) are discussed. As far as the hydrocarbons are concerned, however, there are awkward counterarguments. For example, why should n-pentane act in this way when n-pentanol, even at 50 times the concentration. has almost no effect on h,? A mechanism by which the hydrocarbons shift the h, curve has been proposed by Haydon and Kimura (1981). The essence of the mechanism is that the membrane is thickened by the adsorption of hydrocarbon into its

ANESTHETIC EFFECTS ON THE SQUID GIANT AXON

465

interior and that this affects the electric field experienced by the inactivation sensor. Certain assumptions are involved, and, in order to show these clearly, a rather more detailed description than was given originally is presented in the Appendix to this article. It is shown there that for a membrane with equal surface potentials on its two sides the shift AVh of the midpoint of the h, curve should be

AV,

=

Vh[(d'/d) - 11

(5)

where d and d' are the effective hydrophobic layer thicknesses of the membrane under control and test conditions, respectively, and Vh is the membrane potential at which h, = 0.5, as in the original (approximate) relationship [see Appendix, Eq. (Al)] of Hodgkin and Huxley (1952). In addition, the ratio of the slopes at the midpoint of test and control curves is [see Appendix, Eq. (All)]

Unlike Eq. ( 5 ) for the shift, this result does not depend on any assumption concerning surface potentials. The equation for h, consistent with Eqs. ( 5 ) and (6) is obtained from Eqs. (A6) and (A7) (see Appendix) by putting the sum of the surface potentials to zero and is

as in Haydon and Kimura (1981). In Fig. 10 is shown a test of the above expressions. The constants v h and k were obtained by fitting the control h, curve with Eq. (7), where did' = 1. The value of d/d*for the pentane-treated axon was then calculated from the observed shift (AVh) using Eq. (9,and from Eq. (7) the whole curve of hk versus V was plotted. The result (continuous line) fits the points well. In particular, the value of did' (= 0.769) calculated from the shift is consistent with the change in slope at the midpoint. This exercise has been carried out for 6 hydrocarbons and for carbon tetrachloride (28 axons in all) without finding any serious discrepancies (Haydon and Kirnura, 1981; Haydon and Urban, 1983a). The model proposed to account for the h, results for nonpolar molecules is, from the above considerations, internally consistent. In addition there is independent evidence that the adsorption of nonpolar molecules thickens the axon membrane. It has been known for some time that, at the concentrations that produce nerve blockage, the hydrocarbons can thicken lipid bilayers by as much as 90%. This conclusion has been

D. A. HAYDON ET AL.

466

- 100

- 60

.20

V (mVl

FIG.10. The steady-state inactivation parameter (h,) as a function of membrane potential hefore (O), during (O), and after ( x ) exposure of an axon to 306 pM n-pentane. The right-hand continuous curve was calculatcd from Eq. (7) with did' = I V,, = -S2.6 mV, and k = 6.41 m V . The left-hand curve was also calculated from Eq. (7) with Vh and k unchanged but with dld' = 0.769. (From Ilaydon and Kimura, 1981; reproduced by courtesy of the Jourrrt~lof P h y s i o l o g y . )

reached from capacity and optical reflectance measurements (Dilger PI a / . , 1982, and references therein) and from X-ray diffraction studies (Lea, 1979; Mclntosh rt ul., 1980). More direct evidence comes from Xray diffraction measurements on the myelin membranes of sciatic nerve (Padron et ul,, 1979) and from capacity measurements at relatively high frequencies on the squid giant axon membrane (Haydon rt ul., 1980; Haydon and Urban, 1983a). Some capacity results for n-pentane at 306 p,M are shown in Fig. 11. The interpretation of impedance data for the squid axon membrane is not straightforward, but, apart from the frequency dependence of the capacity, the effects of hydrocarbons are very similar to those for lipid bilayers. Thus a thickening is at present the most obvious and satisfactory explanation. In contrast to the hydrocarbons, the alcohols do not produce appreciable shifts in the h, curve (Table 111). Consistent with this are findings that the alcohols have little or no influence on the capacity of the axon membrane (Armstrong and Binstock, 1964; Haydon and Urban, 1983b). The capacity of lipid bilayers is also not significantly affected by alcohols (Ebihara et a/., 1979; Elliott and Haydon, 1979; Elliott, 1982; Reyes and Latorre, 1979), and from X-ray diffraction and nuclear magnetic resonance experiments it has been concluded that the bilayer thickness is not greatly changed (Ebihara et al., 1979; Turner and Oldfield, 1979). Other substances listed in Table 111 cannot be discussed in detail com-

467

ANESTHETIC EFFECTS ON THE SQUID GIANT AXON

(

W

C

y

O 10

':-" ,

50 100 frequency (kHz1

500

FIG. 11. Mean capacity per unit area ( C )for nine squid giant axons in artificial seawater (upper curve) and in 306 F M n-pentane solution (lower curve). (From Haydon et 01.. 1980; reproduced by courtesy of the Joitmui of Physiology.)

parable to that for the hydrocarbons and alcohols, since less is known about them. Effects on the membrane capacity have been examined in most instances (Haydon and Urban, 1983b,c), but the interpretation of the results in terms of membrane thickness is hampered by ignorance as to how these substances are distributed and by the fact that their dielectric constants are different from that of hydrocarbon chains. There is also little knowledge of how bilayer thickness is affected. It can be seen, however, that, broadly speaking, the less surface-active substances give the largest hyperpolarizing shifts in the h, curve. In other words, those substances that could be expected to absorb into the nonpolar interior of the membrane, so causing thickening, have the largest effect. Three substances, however, deserve special mention. Dioctanoylphosphatidylcholine shifts the h, curve in the positive (depolarizing) direction. According to the above arguments, this could result from a thinning of the membrane. There is no direct evidence that this actually occurs. However, the adsorption of short-chain phospholipids into a membrane must presumably bring about a thinning. The other two substances are methoxyflurane and methyl octanoate. Both shift the h, curve appreciably in the negative direction and both reduce the slope of the curve. On this evidence it would seem that these molecules must absorb into the middle of the membrane. The fact that both contain polar groups would, on the other hand, make this seem rather unlikely. It is known that esters and ketones generally behave rather like methyl octanoate in producing a negative h, shift. The explanation may therefore be that the presence of a sufficiently electronegative oxygen atom leads to a specific interaction, with perhaps the channel protein, and that this in-

468

D. A. HAYDON ET AL.

creases locally the partitioning into the membrane interior (Haydon and Urban, 1983b). It is of interest that benzocaine which, as mentioned earlier, shifts the h, curve for nodal membranes in the negative direction, has an ester group. The possibility arises, therefore, that the mechanism of action of benzocaine is similar to that of methyl octanoate and other esters. Haydon and Urban (1983b) suggested that some kind of specific interaction with the channel protein occurs, but the main effect of this is to produce a local membrane thickening. A small pocket of lipid near the interior of the channel, perhaps between protein helices, might account for these effects. While the thickening model accounts reasonably satisfactorily for the observations, some questions remain. For example, quantitative tests are clearly desirable. In particular it would be valuable to compare estimates of the thickening obtained from the h, shift with independent determinations of thickness change. Such a comparison, using the membrane capacity at 100 kHz, has been attempted (Haydon and Urban, 1983a). It is shown that, although there is an obvious correlation between the two estimates, they are not equal. The discrepancy may originate from an inadequate interpretation of the membrane capacity. It must also be remembered that the membrane capacity would, at best, reflect an average membrane thickness, whereas the h, voltage sensor would respond to local changes. The latter could well differ, owing to geometrical factors and variations in lipid compositions, from the average value. Another interesting question arises from the fact that, according to the unsimplified model, the surface dipole potentials should influence the h, shift. However, molecules such as the alcohols, which should alter these dipole potentials, have very little effect. It is possible that the adsorption of the alcohols is similar on both sides of the membrane, in which case no field change would occur. But the likelihood that the membrane is asymmetric and the fact that dipole potentials offer an attractive explanation for m, shifts (see below) are against this suggestion. Another possibility will be mentioned in the next section. B. Shifts in the Steady-State Activation Curve

The nonpolar molecules, with the exception of cyclopropane and carbon tetrachloride, shifted the rn, curve in the hyperpolarizing (negative) direction. These effects are similar to, though not as large as, those of the nonpolar molecules on the h, curve, and membrane thickening is an obvious explanation. Dioctanoylphosphatidylcholine moved the m, curve in the depolarizing direction. A similar effect was found for the h, curve and

ANESTHETIC EFFECTS ON THE SQUID GIANT AXON

469

a membrane thinning could account for both phenomena. Explanations based on thickness changes are not adequate, however, for many of the results. Thus, all the polar molecules shifted the m, curve in the depolarizing direction without appearing to change membrane thickness. In addition, cyclopropane and carbon tetrachloride produced depolarizing shifts even though membrane thickening occurred in these systems. Finally, the hyperpolarizing shifts in the m, curves for other nonpolar molecules were always less than the shifts in the h, curves. A tendency to shift the m, curve in the depolarizing direction seems therefore to be present in all the systems. Two possibilities have been suggested for the positive m, shifts, surface tension or surface free-energy variations and asymmetric surface dipole potential changes (Haydon and Urban, 1983b). The adsorption of anything into the membrane must affect its surface tension in some respect, and, since the surface tension contributes to the chemical potential of species in a surface (Defay et al., 1966; Aveyard and Haydon, 1973), proteins in a membrane could be perturbed by this effect (Haydon and Urban, 1983b). The main objection to this possibility is that not all the substances examined would be likely to cause similar changes in the surface tension. The hydrocarbons and n-alkanols have been shown to raise the tension of lipid bilayers (Needham and Haydon, 1983; Elliott ad Haydon, 1979), but it is unlikely that the oxyethylene alcohols and dioctanoylphosphatidylcholine would do so. The role of asymmetric dipole potentials in excitable membranes was speculated upon some years ago (Haydon, 1975). It is necessary to assume that the voltage sensor for whatever process is involved is a dipole that responds to the internal field of the membrane. It is also assumed that the normal form of the potential variation across the resting membrane is as shown in Fig. 12, where only the membrane potential ( V ) as conventionally measured and the surface dipole potentials ( xo and xi) have been depicted. Diffuse double-layer potentials and membrane thickening may also be present (as in Fig. A l ; see Appendix), but these features are not essential for the mechanism to be discussed. The large positive values of the surface dipole ( xo,Jpotentials are assumed from measurements of the surface potentials of lipids at air-water and oil-water interfaces. For phosphatidylcholine and phosphatidylethanolamine, values of 420-450 mV have been reported (Papahadjopoulos, 1968; Hladky and Haydon, 1973). In order that the m, curve should move to more positive potentials, it is necessary for the internal field to increase on adsorption of the anesthetic, as indicated by the dashed line in Fig. 12. Obviously, such an increase in the field could occur either by an increase of the dipole potential on the

470

D. A. HAYDON ET AL. dipole (headqrwDI

outside

inside

sensor

0 hydro-

FIG. 12. A schematic illustration of the axon membrane and the activation voltage sensor (top) with the supposed electrical potential variation across it (bottom). V is the membrane potential a5 normally measured; xu and x, are surface dipole potentials. The dashed line indicates the change in the potential profile that would result from the adsorption of an anesthetic on the inner surface of the membrane.

outside of the membrane or by a decrease on the inside. However, it seems from the available data (see, c.g., Gaines, 1966) that the phospholipids have larger dipole potentials than any of the substances in Table 11 1. Thus the adsorption of the anesthetic would probably lower the dipole potential. Hence, in order to change the field appropriately, the adsorption would have to be predominantly on the inside of the membrane. Two questions immediately arise: Why should the adsorption be larger on the inside? Would the adsorption be sufficient to produce the observed shifts? The axon membrane is known to be asymmetric in the sense that the channel proteins respond differently to various substances according to the side on which they are presented. Nevertheless, no direct evidence of asymmetry in the lipid has been reported. It is known, however, that the myelin membrane of frog sciatic nerve is asymmetric. Caspar and Kirschner (1971) have suggested from spectroscopic data that the outside of the myelin membrane may be richer in cholesterol than the inside. Cholcsterol inhibits the adsorption of a number of anesthetics into lipid bilayers (Miller et al., 1977; Haydon et al., 1977; Simon et al., 1979a,b; Smith et al., 1981), so there is clearly some reason to think that the field may change as indicated in Fig. 12. The magnitude of the change required to account for the m, shift is small compared with the absolute values of the dipole potential, i.e., 5 2 0 mV in -400 mV. Thus an increase of 5 5 % in

ANESTHETIC EFFECTS ON THE SQUID GIANT AXON

471

the area per molecule of the phospholipids could reduce the dipole potential sufficiently although, if the expansion was accomplished by the adsorption of an anesthetic with an appreciable dipole potential itself, the increase would have to be larger. Nevertheless, the adsorptions required are not out of line with those expected or observed. For a monoglyceride with a dipole potential of -320 mV (Hladky and Haydon, 1973), an adsorption of approximately 1 monoglyceride per 10 phospholipids should induce a shift of -10 mV. More direct evidence has been furnished by Reyes and Latorre (1979). These authors concluded that -4 mM chloroform and 8 mM benzyl alcohol reduced the dipole potential of phosphatidylethanolamine bilayers by -16 mV and -10 mV, respectively. The concentrations are both very similar to those for blockage of the nerve impulse. The m, shift for 12 mM benzyl alcohol is 10 mV and the shift for chloroform (Table 111) also corresponds closely with the surface potential decrease. It is obviously premature to conclude that the shifts in the m, curves originate from surface dipole potential changes. However, it is worth noting that this type of mechanism is attractive if only because very small perturbations in the membrane surface can produce large variations in the internal field. It is also of interest that the Na- and K-channel activation seem to behave similarly. In view of the foregoing discussion, it is slightly surprising that there is little or no indication that the surface dipole potentials affect the h, curve. This may be related to the generally accepted conclusion that parts of the Na-channel protein involved in inactivation are exposed to the inside of the axon. If this is so, then changes in the lipid on the inside could be unimportant, simply because of a lateral separation between its dipoles and the inactivation voltage sensor.

C. Reduction of the Maximum Conductances gNaand gK

The nonpolar substances not only shift the h, curve substantially, but they also tend to suppress gNaand gKmore readily than do the polar or surface-active substances. It appears likely, therefore, that the suppression in g is related to a thickening of the membrane. The possibility that membrane thickness (combined with membrane tension) could affect ionchannel stability arose from studies in artificial bilayers of the polypeptide gramicidin (Haydon et al., 1977). The gramicidin channel is formed from two helical molecules joined head to head by hydrogen bonds, and it has a length of -28 A. This structure has a high stability in membranes of thickness comparable to its own length, but, in membranes of greater thickness, most of the dimers dissociate. Thus, in bilayers with a hydro-

472

D. A. HAYDON ET AL.

carbon thickness of 48 A, the stability can be - x lo4 lower. The mechanism proposed has been described in detail by Hendry et ul. (1978). Briefly, it is supposed that when the membrane thickness is larger than the length of the channel, a deformation or dimpling of the lipid is necessary in order that the two gramicidin monomers may meet in the middle of the membrane (Fig. 13). This deformation requires that work be done against surface forces. A quantitative test of the model was carried out and found to account for the conductance results to within experimental error (Hendry et uf., 1978). Although there is no suggestion that the Na and K channel proteins resemble gramicidin, the general ideas outlined above could well apply to such structures. These channel structures are presumably tailored to fit the lipophilic regions of the bilayer, and their conformation is very probably maintained by hydrogen bonds. Any perturbation in the surface thermodynamic properties of the lipid is thus liable to affect the intramolecular bonding and hence the conformation. If the adsorption of n-pentane can affect so much the stability of the hydrogen-bonded gramicidin structure, it seems quite feasible that it would disrupt the channels of the nerve axon. Two other observations are consistent with this type of mechanism, First, the effects of the hydrocarbons on the Na and K channels are very similar, as might be expected for such a nonspecific process. Second, the n-alkanols and various other amphipathic substances (at nerve block con-

aqueous solution

FIG. 13. A grarnicidin dirneric ion channel spanning a lipid bilayer. The dashed lines indicate the normal thickness of the bilayer (i.e., comparable to the length of the channel) whereas the continuous lines depict a bilayer thickened through the adsorption of a nonpolar anesthetic. The hydrogen bonds that link the two polypeptide molecules are represented by the vertical dashed lines. Other hydrogen bonds in the helices are not shown. If the surface tension of the bilayer is finite, the polypeptide channel is stressed by the thickening.

ANESTHETIC EFFECTS ON THE SQUID GIANT AXON

473

centrations) that do not appreciably affect membrane thickness have a relatively small effect on gNaand gK.The fact that some of these surfaceactive substances (e .g., the n-alkanols) increase the membrane tension (Elliott and Haydon, 1979; D. Needham, personal communication) is not necessarily relevant unless there were, in normal membranes, a thickness mismatch between protein and lipid. The importance of membrane thickness for the functioning of a transport protein has been demonstrated in at least two other systems, the Na+,K+-ATPase(Johannsson et al., 1981a) and the Ca2+,Mg2+-ATPase(Johannsson et al., 1981b). In both instances it was found that whether the thickness was varied by changing the chain length of the lipid or by the adsorption of hydrocarbon, the ATPase activity was optimal only for a limited range of thickness. As for the Na and K channels of the nerve axon, however, no strong evidence as to the mechanism was obtained. Armstrong and Binstock (1964) showed that at concentrations higher than those mentioned above, n-alkanols reduced gNasignificantly. There is apparently, therefore, a binding site for the n-alkanols that is somewhat weaker than that involved in the initial nerve blockage. A binding site of somewhat similar strength has been described for long-chain quaternary ammonium ions (Rojas and Rudy, 1976). From the fact that the blockage of the Na conductance by these ions is voltage dependent it is probable that the site in question is on the interior surface of the channel. D. Time Constants

The time constants T, and t-h for activation and inactivation of the Na channel appear to be influenced by anesthetics in at least two ways. First, the curves of time constant versus membrane potential are shifted on the voltage axis, as are the corresponding steady-state parameters m, and h, . These shifts, as estimated from the positions of the maxima in the time constants, are similar in magnitude to those of m, and h, curves and apparently originate from the same physical effects. Second, the peak value of the time constant is, with one exception, reduced by the anesthetics. Furthermore, if the voltage shift is allowed for, it is found that the time constants decrease at all depolarizations. The results for the K channel, though limited as yet, are similar to those for the Na channel. There is no direct evidence as to the origin of the decrease in the time constants, but it is remarkable that practically all the substances examined produced qualitatively the same effects. The substances in question have molecular dimensions that are somewhat smaller than those of the usual membrane lipids, and there is abundant evidence that the adsorp-

474

D. A. HAYDON ET AL.

tion of such substances into lipid bilayers increases the fluidity of the structure. Thus, there is circumstantial evidence that the reduction in time constants originates from an increase in bilayer fluidity. But, even if this mechanism is correct, there remains a great deal to be accounted for at the quantitative level. For example, it is not clear why the normal alkanes reduce Th more than T, whereas, for the cyclic alkanes, the reverse occurs. It is notable that the smaller hydrocarbons and alcohols are more effective (at concentrations for -50% suppression of fNa) than larger molecules, but surprising that cyclohcxane produces no reduction in q,whereas n-hexane is very potent. The concentration dependence of the changes in time constant has not been studied. But the rates of change of T , , Th. and h, as anesthetic is introduced have been compared for n-pentane (Haydon and Urban, 1983a). The results showed that the time constants decreased slightly more rapidly than the h, curve shifted, but the effect was not sufficient to suggest that fluidity changes might be especially important at low anesthetic concentrations. Similar studies for halothane and chloroform have reinforced this conclusion.

E. Concluding Remarks 1 . LIPID SITESVERSUS PROTEINSITES

Evidence and arguments have been presented in the foregoing sections to show that, for most of the substances examined, the major effects on the voltage-clamp currents could have arisen from perturbations of the membrane lipid. These perturbations provide reasonable and self-consistent explanations for effects on the sodium current steady-state activation and inactivation curves, effects on the maximum conductances gNHand g K ,and effects on the time constants of channel gating. An additional piece of evidence is that, in the effects of the homologous series of IZalkanols, the apparent standard free energy of adsorption per methylene group is very similar in the nerve axon to that for adsorption in a phospholipid bilayer (Haydon and Urban, 1983b). The success of such lipid-based explanations should not be taken to rule out the possibility of direct anesthetic-protein interactions. For certain substances (e.g., methoxyflurane) circumstantial evidence has been described that direct protein interaction may contribute significantly to their anesthetic effects. It seems likely that attempts to provide a complete distinction between protein and lipid as the site of action oversimplify the problem. For example, the form of the channel proteins is not known and it is quite possible

ANESTHETIC EFFECTS ON THE SQUID GIANT AXON

475

that lipid is present in small aggregates or pockets between protein helices or subunits. If this is so, then interactions with the lipid could scarcely occur without simultaneous interaction with the protein. The great advantage at present of hypotheses based on interactions with lipids is that they suggest further perfectly feasible experiments on well-defined systems. For example the suggestion of asymmetrical dipole potential changes to explain anesthetic-induced shifts of m, is amenable to quantitative testing in monolayer and artificial bilayer systems.

2. THE SQUIDAXONA N D GENERAL ANESTHESIA The experiments and hypotheses discussed in this article have no immediate relevance to the clinical phenomenon of general anesthesia. They have been primarily concerned with the mechanisms by which anesthetics affect the function of the sodium channel. Nevertheless a number of insights have resulted that may prove to be of more general importance. It is striking to find that a chemically simple anesthetic substance can affect sodium-channel function in a variety of ways. These effects are usually additive and may even be synergistic in reducing the physiological inward sodium current. For example a depolarization of the resting axon, a negative h, shift, a positive m m shift, a reduction in gNa,and a reduction in 7 h will all tend to reduce inward sodium current in the intact cell. The complexity of effects on a whole animal containing many different types of ion channels in different functional roles is potentially enormous. Nevertheless if ion channels can be observed at the molecular level, the possibilities for explaining anesthetic effects in simple physicochemical terms appear to be very encouraging. One final point can be made concerning the relevance of the sodium channel itself to general anesthesia. The anesthetic concentrations required to inhibit the maximum inward Na current by 50% are 3 to 10 times those associated with general anesthesia in man (Steward et al., 1973). However, the most important cause of Na-current inhibition by a number of anesthetics (including halothane) is a depolarizing shift in m,. It follows that for these anesthetics the inward current produced by small depolarizations must be reduced by much more than 50%. In terms of axonal function this distinction may not matter. It does, however, mean that the threshold for impulse initiation is particularly sensitive to anesthetics. Consequently, the firing rate of neurons in the CNS may be significantly altered by anesthetic concentrations that do not affect peripheral axonal conduction. A mechanism of action of this nature, involving sodium-channel perturbation, for clinical anesthetics such as halothane cannot be ruled out.

476

D. A. HAYDON ET AL.

APPENDIX: EFFECT OF MEMBRANE THICKNESS CHANGES ON THE STEADY-STATE INACTIVATION OF THE Na CHANNEL

The main assumption of the following theoretical model is that the steady-state Na-channel inactivation is determined primarily by the internal electric field of the membrane. In ordcr to develop this supposition, the membrane interior will be considered to be an isotropic dielectric structure. In view of the fact that membranes are apparently highly nonisotropic, this is not an encouraging beginning. However, for lipid bilayers, at least, the electrical capacity can be predicted remarkably accurately on this basis and constant field equations are known to account reasonably well for numerous observations on the ion permeability of membranes. The variation of the electric potential across the membrane is assumed to be broadly as shown in Fig. AI. The membrane potential, V , is that between two points far from the surfaces. At the membrane surfaces are electrical double layers arising from the segregation of ions (ionic double a

I

k

d’

Y

b

FIG.A l , (a) A schematic illustration of the membrane and the dipolar inactivation voltage sensor. The thickness, d, increases to d‘ in the presence of the test substance. (b) The variation of the electrical potential assumed to exist for the normal membrane (continuous line) and for the membrane under test conditions (dashed line). I$ denotes the ionic doublelayer potential, and x the dipole potential; &, and xu are referred to a point far from the surface on the outside as zero, and, correspondingly. & and xi are referred to a point far from the surface on the inside. Thus, in the diagram, +o and 4, are negative while xo and xi are positive. V , is the difference of potential between the two inner surfaces of the membrane. As for V . the sign convention for V , is “inside-outside,” and in the diagram both are therefore negative,

477

ANESTHETIC EFFECTS ON THE SQUID GIANT AXON

layer) and from the orientation of the dipoles of the polar groups of the membrane lipids and other components. Hodgkin and Huxley (1952) proposed an empirical expression for the dependence of h, upon the membrane potential. In its approximate form and for the present conventions, this is

1

=

hm

1

+ exp[( V - Vh)/kl

where V,, the potential at which h, = 0.5, and k are constants. The slope of the relationship at h, = 0.5 is 1 4k

Equation (Al) may be rewritten in terms of the internal field of the membrane by introducing the membrane thickness d and the potential difference V , between the inner surfaces (Fig. Al), i.e., 1

hm = 1

f

exp[( V,ld

-

VA/d)lk/dl

where V; is a constant equal to the value of V, when h, Fig. Al, Vm = V

- (xo +

$0)

+ (Xi +

=

0.5. From

4i)

(A4)

From Eqs. (A3) and (A4),

h,

=

1

1

+ exp{[(V - xo - +o + xi + 4i)/d - VA/d]/k/d)

(A5)

When h, = 0.5, V = V, [Eq. (Al)] and hence, from Eq. (A5),

v;

40) - (xi -k 4i)

(A61 The slope of the h, curve at the midpoint remains as given by Eq. (A2). When exposed to a test solution, the membrane thickness changes to d', and the surface potentials may also change. Both effects give rise to a change in the internal field, and the dependence of h, on V now becomes vh =

( x o -k

1

hl= 1

+ exp{[(V - xb - +:,+ xi + +:)/dt - VL/d]/k/d}

047)

The terms containing V,!, and k do not change because, as in Eq. (Al), these are empirical constants obtained by fitting the control h, curve. The membrane potential Vh at which hk = 0.5 is now

v', = (d'id) VA + (xb + (#&)- (xi + 4;)

(A8)

478

D. A. HAYDON ET AL.

The shift AVh in the midpoint of the h, curve is AVh = Vb

~

Vh = Vb[d'/d

1J

~

+ (x:, - x,,) + (4'0 - 4'))

- (xi - xi) - (4 - 4i) The gradient of Eq. (A7) at h' = 0.5 is

and the ratio of the slopes for test and control conditions is thus

The change in slope does not therefore depend on the surface potentials of the membrane or on the influence of the test substance of these potentials. The shift AVh does, however, depend upon both these factors. But if Eq. (A9) reduccs to

and if, further, from Eq. (A6) Vh

=

v;,

and Avh = Vh (d'/d - 1) which is the relationship given in Haydon and Urban (1983a) and (except for a sign change) in Haydon and Kimura (1981).

REFERENCES Armstrong, C. M . , and Binstock, L. (1964). The effects of several alcohols on the properties of the squid giant axon. J . Gen. P h y s i d . 48, 265-277. Austin, G . M . , and Pdsk, E. A. (1952). Effect of ether inhalation upon spinal cord and root action potentials. J . Phy.siol. (London) 118, 405-41 1. Aveyard, R . . and Haydon, D. A . (1973). "An Introduction to the Principles of Surface Chemistry." Cambridge Univ. Press, London and New York. Bean, B. P., Shrager, P., and Goldstein, D. A. (1981). Modification of sodium and potassium channel gating kinetics by ether and halothane. J . Gen. Pliysiol. 77, 233-253.

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Cahalan, M. D. (1978). Local anaesthetic block of sodium channels in normal and pronasetreated squid giant axons. Biophys. J . 23, 285-31 1. Caspar, D. L. D., and Kirschner, D. A. (1971). Myelin membrane structure at I0 A resolution. Nutiire (London) New Biol. 231, 46-52. Chandler, W. K., Hodgkin, A. L., and Meves, H. (1965). The effect of changing the internal solution on sodium inactivation and related phenomena in giant axons. J . Physiol. (London) 180, 821-836. Defay, R., Prigogine, I . , Bellemans, A,, and Everett, D. H. (1966). “Surface Tension and Adsorption.” Longmans, Green, New York. Dilger, J. P., Fisher, L . R., and Haydon, D. A. (1982). A critical comparison of electrical and optical methods for bilayer thickness detemination. Chem. Phys. Lipids 30, 159- 176. Ebihara, L., Hall, J. E., MacDonald, R. C . , Mclntosh, T. J., and Simon, S. A. (1979). Effect of benzyl alcohol on lipid bilayers. A comparison of bilayer systems. Biophys. J . 28, I 85- 196. Elliott, J. R. (1981). Anaesthetic mechanisms: Effects of alcohols on lipid membranes. Ph.D. dissertation, University of Cambridge. Elliott, J . R., and Haydon, D. A. (1979). The interaction of n-octanol with black lipid bilayer membranes. Biochim. Biophys. Acta 557,259-263. . Bezanilla, F., and Taylor, R. E. (1982). Effect of chloroform on charge Fernandez, .IM., movement in the nerve membrane. Nature (London) 297, 150-152. Fettiplace, R., Andrews, D. M., and Haydon, D. A. (1971). The thickness, composition and structure of some lipid bilayers and natural membranes. J . Membr. B i d . 5, 277-296. Frankenhaeuser, B., and Hodgkin, A. L. (1957). The action of calcium on the electrical properties of squid axons. J . Physiol. (London) 137, 218-244. Franks, N. P., and Lieb, W. R. (1982). Molecular mechanisms of general anaesthesia. Nature (London) 300, 487-493. Franz, D. W., and Perry, R. S. (1974). Mechanisms for differential block among single myelinated and non-myelinated axons by procaine. J . Plzysiol. (London) 236, 193-220. Gaines, G . L. (1966). “Insoluble Monolayers at Liquid-Gas Interfaces.” Wiley. New York. Haydon, D. A. (1975). Functions of the lipid in bilayer ion permeability. Ann. N . Y . Acud. Sci. 264, 2-16. Haydon, D. A., and Hendry, B. M. (1982). Nerve impulse blockage in squid axons by nalkanes: The effect of axon diameter. J . Physiol. (London)333, 393-403. Haydon, D. A . , and Kimura. J. E . (1981). Some effects of n-pentane on the sodium and potassium currents of the squid giant axon. 1.Physio!. (London) 312, 57-70. Haydon, D. A,, and Urban, B. W. (1981). Effects of different types of general anaesthetic on the sodium current of the squid giant axon. J . Physiol. (London) 319, 27-28P. Haydon, D. A,, and Urban. B. W. (1983a). The action of hydrocarbons and carbon tetrachloride on the sodium current of the squid giant axon. J . Physiol. (London) 338, 435-450. Haydon, D. A., and Urban, B . W. (1983b). The action of alcohols and other non-ionic surface active substances on the sodium current of the squid giant axon. J . Physiol. (London) 341, 41 1-427. Haydon, D. A.. and Urban, B. W. (1983~).The effects of some inhalational anaesthetics on the sodium current of the squid giant axon. J . Physiol. (London) 341, 429-439. Haydon, D. A., Hendry, B. M., Levinson, S . R., and Requena, .I(1977). . Anaesthesia by the n-alkanes. A comparative study of nerve impulse blockage and the properties of black lipid bilayer membranes. Biochim. Biophys. Acta 470, 17-34.

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Haydon, D. A., Kequena, J., and Urban, B. W. (1980). Some effects of aliphatic hydrocarbons on the electrical capacity and ionic currents of the squid giant axon membrane. J. Physiol. (London) 309, 229-245. Hendry, B. M., Urban, B. W., and Haydon, D. A. (1978). The blockage of the electrical conductance in a pore-containing membrane by the n-alkanes. Biochim. Biophys. Acta 513, 106-1 16. Hille, B. (1977). Local anaesthetics: Hydrophilic and hydrophobic pathways for the drugreceptor interaction. J. Gen. Physiol. 69, 497-515. Hladky, S. B.. and Haydon, D. A. (1973). Membrane conductance and surface potential. Biochirn. Biophys. Acta 318, 464-468. Hodgkin, A. L., and Huxley, A. F. (1952). A quantitative description of membrane current and its application to conduction and excitation in nerve. J . Physiol. (I,ondorr) 117, 500-544. Jack, J. J. B., Noble, D., and Tsien, R. W. (1975). “Electric Current Flow in Excitable Cells.” Clarendon, Oxford. Johannsson, A., Smith. G . A., and Metcalfe, J. C. (1981a). The effect of bilayer thickness on the activity of (Na+ + K+)-ATPase. Biochim. Biophys. Act0 641,416-421. Johannsson, A., Keightley, C. A,, Smith, G . A , . Richards, C. D., Hesketh, T. R., and Metcalfe, J . C. (1981b). The effect of bilayer thickness and n-alkanes on the activity of the (Ca2+ + Mg*’)-dependent ATPase of sarcoplasmic reticulum. J. Biol. Chem. 256, 1643-1650. Kendig, J. J., Courtney, K. R., and Cohen, E. N. (1979). Anesthetics: Molecular correlates of voltage- and frequency-dependent sodium channel block in nerve. J . Pharmacol. Exp. Ther. 210, 446-452. Larrabee, M. G . , and Posternak, J. M. (1952). Selective action of anesthetics on synapses and axons in mammalian sympathetic ganglion. J, Neurophysiol. 15, 91-1 14. Lea, E. J . A. (1979). Effect of n-alkanes on membranes in lipid-water systems. I n f . J. Biol. Macromol. 1, 185-187. Mclntosh, T. J . . Si-mon, S. A., and MacDonald, R. C. (1980). The organisation of n-alkanes in lipid bilayers. Biochim. Biophys. Acfa 597, 445-463. Meves, H. (1978). Inactivation of the sodium permeability in squid giant nerve fibres. Prog. Biophys. Mu/. Biol. 33, 207-230. Miller, K.W . , Hammond, L., and Porter, E. G. (1977). The solubility of hydrocarbon gases in lipid bilayers. Chem. Phys. Lipids 20, 229-241. Moore, J. W. (1966). Effects of ethanol on ionic conductances in the squid axon membrane. Psychosom. Med. 28,450-457. Moore, J. W., Ulbricht, W., and Takata, M. (1964). Effect of ethanol on the sodium and potassium conductances of the squid axon membrane. J . Cen. Physiol. 48, 279-295. Narahashi, T., Moore, J. W., and Poston, R. N. (1969). Anesthetic blocking of nerve membrane conductances by internal and external applications. J . Ncurobiot. 1, 3-22. Needham, D., and Haydon, D. A. (1983). Tensions and free energies of formation of “solventless” lipid bilayers: ’The measurement of high contact angles. Bio~phys.J. 41, 251-257. Neumcke, 8.. and Stampfli, R. (1982). Sodium currents and sodium current fluctuations in rat myelinated nerve fibres. J. Physiol. (London) 329, 163-184. Noble, D. (1979). “The Initiation of the Heart-beat,” 2nd ed. Clarendon, Oxford. Oxford, G. S., and Swenson, R. P. (1979). n-Alkanols potentiate sodium channel inactivation in squid giant axons. Biophys. J. 26, 585-590. Oxford, G.S., and Yeh. J. Z. (1982). Molecular mechanisms of sodium channel inactivation: Discriminations using temperature, octanol and ion competition. Biophys. J. 37, 104a.

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48 1

Padron, R., Mateu, L., and Requena, J. (1979). A dynamic X-ray diffraction study of anesthesia action. Thickening of the myelin membrane by n-pentane. Biochim. Biophys. Acta 552, 535-539. Papahadjopoulos, D. (1968). Surface properties of acidic phospholipids: Interaction of monolayers and hydrated liquid crystals with uni- and bi-valent metal ions. Biochim. Biophys. Acta 163, 240-254. Paternostre, M., and Pichon, Y. (1982). Effects of propanol and heptanol on the potassium conductance in giant axons of Loligo. J . Physiol. (London) 328, 31-32P. Reyes. J., and Latorre, R. (1979). Effect of the anesthetics benzyl alcohol and chloroform on bilayers made from monolayers. Biophys. J . 28, 259-280. Richards, C. D. (1980). The mechanisms of general anaesthesia. In “Topical Reviews in Anaesthesia” (J. Norman and J. Whitwam, eds.), pp. 1-84. Wright, Bristol. Richards, C. D., Martin, K., Gregory, S . , Keightley, C. A., Hesketh, T. R., Smith, G. A., Warren, G . B., and Metcalfe, J. C. (1978). Degenerate perturbations of protein structure as the mechanism of anaesthetic action. Nature (London) 276, 775-779. Rojas, E., and Rudy, B. (1976). Destruction of the sodium conductance inactivation by a specific protease in perfused nerve fibres from Lo/igo. J . Physiol. (London) 262, 501-53 I . Schneider, H. (1968). The intramembrane location of alcohol anesthetics. Biochim. Biophys. Acts 163,451-458. Seeman, P. (1972). The membrane action of anesthetics and tranquilizers. Pharmacol. Reu. 24, 583-655. Sherrington, C. S . (1947). “The Integrative Actions of the Nervous System,” 2nd ed. Cambridge Univ. Press, London and New York. Shrivastav, B. B. (1977). Mechanism of ketamine block of nerve conduction. J . Pharmacol. Exp. Ther. 201, 162-170. Shrivastav, B. B., Roberts, J. T., and Kitz, R. J. (1972). The effects of general anesthetics on the sodium and potassium conductances of the squid giant axon. Fed. Proc. Fed. A m . SOC.Exp. Biol. 31, 550 ab. Shrivastav, B. B., Narahashi, T., Kitz, R. J . , and Roberts, J . D. (1976). Mode of action of trichloroethylene on squid axon membranes. J . Pharmacol. Exp. Ther. 199, 179-188. Simon, S . A . , Stone, W. L., and Bennett, P. B. (1979a). Can regular solution theory be applied to lipid bilayer membranes? Biochim. Biophys. Acra 550, 38-47. Simon, S . A., McIntosh, T. J., Bennett, P. B., and Shrivastav, B. B. (1979b). Interaction of halothane with lipid bilayers. Mol. Pharmacol. 16, 163-170. Smith, R. A., Porter, E. G., and Miller, K. W. (1981). The solubility of anesthetic gases in lipid bilayers. Biochim. Biophys. Acta 645, 327-338. Somjen, G. G. (1963). Effects of ether and thiopental on spinal presynaptic terminals. J . Pharmacol. Exp. Ther. 140, 396402. Spyropoulos, C. S. (1957). The effects of hydrostatic pressure upon the normal and narcotinized nerve fiber. J . Gen. Physiol. 40, 849-857. Staiman, A,, and Seeman, P. (1977). Conduction-blocking concentrations of anaesthetic increase with nerve axon diameter: Studies with alcohol, lidocaine and tetrodotoxin on single myelinated fibres. J . Pharmacol. Exp. Ther. 201, 340-349. Steward, A., Allott, P. R.,Cowles, A. L., and Mapleson, W. W. (1973). Solubility coefficients for inhaled anaesthetics for water, oil and biological media. Br. J . Anuesth. 45, 282-293. Swenson, R. P. (1979). Like a local anesthetic? Octanol immobilises gating charge. Biophys. J . 25, 136a.

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Swenson, R. P. (1981). Inactivation of potassium current in squid axon by a variety of quaternary ammonium ions. J . Gen. Physiol. 77, 255-271. Swenson, R. P., and Oxford, 0 . S. (1980). Modification of sodium channel gating by longchain alcohols: Ionic and gating current measurements. In “Molecular Mechanisms of Anaesthesia” (B. R. Fink, ed.), Vol. 2, pp. 7-16. Raven, New York. Taylor, R. E. (195Y). Effect of procaine on electrical properties of squid axon membrane. Am. J . Physiol. 196, 1071-1078. Turner, G. L., and Oldfield, E. (1979). Effect of a local anesthetic on hydrocarbon chain order in membranes. Nature ( l o d o n ) 277, 669-670.

CURRENT TOPICS IN MEMBRANES AND TRANSPORT, VOLUME 22

Pharmacology of Nerve Membrane Sodium Channels TOSHIO NARAHASHI Department of Pharmacology Northwestern University Medical School Chicago, Illinois

I . Introduction.. . . . . . . . . . . . . . . ....................................... Ionic-Channel Block by Chem .............. . . . . . . . . . . . . . . . . . . . . . . . . . A. Technological Developments in the Study of Channel Block.. . . . . . . . . . . . B. Voltage- and Current-Dependent Block ............................... C. Kinetic Schemes of Sodium-Channel Block .....................

11.

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

References .............................................................

483 486 486 487 489 493 493 495 500 506 507 51 1

I. INTRODUCTION

A variety of approaches and techniques has been used for the study of the mechanisms whereby excitable membrane ionic channels function. Since the channels open and close rapidly, leading to ionic fluxes and membrane potential changes, electrophysiological techniques such as intracellular microelectrode and voltage clamp have been used most extensively. There are, however, limits to this approach. Because the ionic channels are composed of macromolecules, presumably those of proteins and phospholipids, our eventual goal should be to identify both chemical and functional aspects of the channels. Attempts are being made to clarify the chemical nature of the channels with the aid of various biochemical techniques such as isolation, purification, and identification of channel 483 Copyright 0 1984 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-153322.0

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TOSHIO NARAHASHI

macromolecules. One of the powerful approaches is to use chemical probes, often called chemical or pharmacological dissection (or characterization) of ionic channels, which has been gaining ever increasing popularity during the past 15 years or so. A variety of natural toxins, chemicals, and even therapeutic drugs that possess certain specific actions on ionic channels is being used as tools to characterize the function and topography of the channels (see reviews by Narahashi, 1974; Khodorov, 1979; Catterall, 1980; Hille, 1976; Ritchie, 1979; Lazdunski et ul., 1980; Yeh, 1982). The history of pharmacological probes for excitable membrane ionic channels began in 1960 when tetrodotoxin (TTX), the puffer fish poison, was discovered to possess a highly specific blocking action on the sodium channel of skeletal muscle membrane (Narahashi et al., 1960). This hypothesis was more definitely demonstrated using voltage-clamped lobster giant axons (Narahashi et al., 1964). Saxitoxin (STX), the toxic component isolated from the dinoflagellate Gonyaulax catanella, was found to exert the same sodium channel-blocking action as TTX (Narahashi et al., 1967). Both TTX and STX have since become very useful and powerful probes for the study of ionic channels, including the estimation of sodium channel density, the study of potassium and calcium channels, the isolation, purification, and identification of sodium channels, and the study of synaptic and neuromuscular transmission (see reviews by Narahashi, 1972, 1974; Ritchie, 1979). Another important contribution of TTX and STX is that their studies aroused interest in other chemicals and toxins as probes for excitable membrane ionic channels that were largely ignored before. Squid giant axons have been used extensively for the study of action of various neuroactive agents. There are at least three reasons that make this preparation almost ideal for the analysis of drug-ionic channel interactions. First, the squid axon has been the prototype of excitable cells as the theory of the ionic mechanism was developed primarily with this preparation (Hodgkin and Huxley, 1952a-d; Hodgkin et ul., 1952). Second, this preparation permits performing near perfect voltage-clamp experiments. Third, it is relatively easy to perfuse the squid axon internally, a condition mandatory for the study of drug action under voltage-clamp conditions. Since topics related to sodium-channel gating and potassium channels are covered in other articles of this volume, the present article is primarily concerned with the sodium channels of squid axons as related to the action of a variety of chemicals. Two basic types of drug action are recognized. One is a drug-channel interaction that leads to a block of the channel, and the other is the one that causes a modulation of the channel

485

PHARMACOLOGY OF NERVE MEMBRANE SODIUM CHANNELS

including changes in gating kinetics. This article deals with these two types of drug action. First, general features of the sodium-channel block caused by drugs will be reviewed without going into much detail. Second, the mechanisms whereby the sodium-channel kinetics are modulated by certain chemicals will be discussed. Chemicals acting on sodium channels of nerve membranes may be classified into three large groups (Fig. 1): ( I ) blockers, such as TTX, STX, local anesthetics, and pancuronium; (2) modulators, such as batrachotoxin (BTX), grayanotoxin (GTX), veratridine, aconitine, scorpion toxins, and pyrethroids; and (3) inactivation inhibitors, such as Pronase, N-bromoacetamide, high and low internal pH, deoxycholate, sea anemone toxins, and goniopora toxin. Modulators are those agents that modify the kinetics of opening and/or closing of the sodium-channel gating by various mechanisms. Inactivation inhibitors are those chemicals that prolong the falling phase of sodium current primarily through inhibition of sodium inactivation. Thus modulators and inactivation inhibitors have

Tet rodotoxi n Pancuronium - on

OAC

OH OH

Botrachotoxi n

(+)-trans-Tetromethrin

FIG. I .

Groyanotoxin I

(+)-trans-Allethrin

Structures of tetrodotoxin, pancuroniurn, batrachotoxin, grayanotoxin I ,

(+)-trans-tetrarnethrin, and (+)-trans-allethrin.

486

TOSHIO NARAHASHI

some features in common, and therefore this classification should not be regarded as reflecting the precise mechanism of action. II. IONIC-CHANNEL BLOCK BY CHEMICALS

Channel block by chemicals may be divided into two categories with respect to the gating status of the channel that exhibits a high affinity for the blocker. One is “closed-channel block” as represented by polyarginine (Wu and Yeh, 1982) and TTX. The closed acetylcholine (ACh)channel block has been demonstrated for histrionicotoxin and tetraethylammonium (Masukawa and Albuquerque, 1978; Adlcr clt d., 1979) and N-octylguanidine (Farley r t al., 1981). However, some chemicals exhibit both closed- and open-channel block (sce below). The other type is “open-channel block.” The drug molecule can bind to a channel only whcn it is open, thereby causing a block. A number of chemicals have been demonstrated to cause this type of block in various kinds of ionic channels, as will be described later (see review by Yeh, 1982).

A. Technological Developments in the Study of Channel Block

The modern era of the study of drug-channel interactions began when voltage-clamp techniques were used to demonstrate the block of sodium and potassium channels of squid axons caused by procaine and cocaine (Taylor, 1959; Shanes et al., 1959). The discovery of the highly selective and potent sodium-channel blocking action of tetrodotoxin (Narahashi et al., 1960, 1964) ignited a widespread interest in using specific chemicals as probes for the study of ionic channels (see Narahashi, 1974; Yeh, 1982). A quanta1 jump in technological developments for the study of ionic channels was made when Neher and Sakmann (1976) successfully recorded opening and closing of individual ionic channels associated with the extrajunctional ACh receptors from the denervated skeletal muscle using a glass capillary applied onto the membrane surface. More recently, a gigaohm-seal patch-clamp technique has been developed whereby the activity of a limited number of ionic channels confined in a very small area of the membrane could be clearly observcd (Sigworth and Neher, 1980; Neher, 1981; Hamill r t ul., 1981). During the past few years, this technique has been applied to a variety of ionic channels including those associated with the ACh receptor (Ogden et ul., 1981; Jackson and Lecar, 1979; Steinbach, 1977; Neher and Steinbach, 1978; Sakmann et al., 1980;

PHARMACOLOGY OF NERVE MEMBRANE SODIUM CHANNELS

487

Sakmann, 1978), those associated with the glutamate receptor (CullCandy et al., 1980; Patlak et al., 1979; Gration et al., 1981; Colquhoun and Sakmann, 1981; Hamill and Sakmann, 1981), sodium channels (Fukushima, 1981b; Patlak and Horn, 1982; Sigworth and Neher, 1980; Horn and Patlak, 1980; Horn et af., 1981a-c; Quandt and Narahashi, 1981, 1982a; Quandt et al., 1982; Yamamoto et al., 1982), potassium channels (Fukushima, 1981a; Conti and Neher, 1980; Marty, 1981; Lux et al., 1981; Pallotta et af., 1981; Quandt and Narahashi, 1982b; Ohmori et al., 1981; Coronado and Miller, 1979, 1980, 1982), and calcium channels (Neher et a/., 1981; Lux and Nagy, 1981; Fenwick et al., 1982; Hagiwara and Ohmori, 1983). Unlike the conventional “macroscopic” current-recording techniques, the patch-clamp techniques permit observation and analysis of single-channel behavior at a level very close to the molecular events. Single-channel analysis has proved to be extremely powerful in elucidating the mechanisms whereby a chemical or ion blocks the channel. This is not only because the opening and closing of individual channels can be measured directly in the absence and in the presence of a test chemical, but also because the step from the channel opening to the drugbound block can be directly measured (for open-channel blockers) without complication caused by the channel gating mechanism. B. Voltage- and Current-Dependent Block

The drug-induced block of ionic channels is often affected by the membrane potential. The voltage-dependent block has been studied extensively for various channels and for various chemicals: e.g., the sodiumchannel block by local anesthetics (Strichartz, 1973; Courtney, 1975), 9-aminoacridine (Yeh, 1979), pancuronium (Yeh and Narahashi, 1977), strychnine (Shapiro, 1977), ACh-channel block by local anesthetics (Deguchi and Narahashi, 1971), histrionicotoxin (Masukawa and Albuquerque, 1978), amantadine (Tsai et a/., 1978), and certain alkylguanidines (Farley et al., 1981). The voltage dependence of block is an important parameter, as the site of drug binding within the membrane electric field can be calculated (Woodhull, 1973). In some cases, however, the apparent dependence of drug-induced block of channels on voltage is due actually to a current dependence, the degree of block changing with the direction of membrane current. This has been shown for the potassium-channel block by tetraethylammonium derivatives (Armstrong, 1969, 1971), the sodium-channel block by paragracine (Seyama

488

TOSHIO NARAHASHI

P t a / . , 1980), and the ACh-channel block by methyl- and ethylguanidine (Vogel et al., 1984). Figure 2 illustrates the current-dependent block of sodium channel by paragracine (Seyama ~t a / . , 1980).The sodium current gradually decreased during repetitive stimulations in the presence of the toxin inside the squid axon only when it flowed in the outward direction.

B O

30 rnV

30 rnV

E 0.4

-0.25

0.2

0

-a5

30

10

F

0.5

0.5 10

20

30

10

M

30

FIG.2. Changes in the amplitude of peak sodium currents elicited by repetitive pulses to 0, +30, and +70 m V during the internal application of 0.23 mM paragracine. Panels A, B, and C were obtained with an axon in bathing solutions containing 445 mM [Najoand 50 mM INal,; panels D. t,and F with II I m M [Nal,,and SO m M [Na],. The sodium currents do no1 decrease during repetitive stimulations while flowing inward at t 30 mV (B), whereas the currents do decrease gradually while flowing outward at the same potential (E). (From Seyama rt d . , 1980.)

PHARMACOLOGY OF NERVE MEMBRANE SODIUM CHANNELS

489

C. Kinetic Schemes of Sodium-Channel Block The open and closed sodium-channel block caused by a drug can be illustrated by simplified kinetic schemes. 611

H-H

Open channel block

C

2 O eI

41

kinetics

ffh

O*

SCHEME 1 Closed channel block

C

-

H-H

kinetics

C*[

-

blIf 0

o

Ph

e

1

ah

*

e '*I

SCHEME 2

where C, 0, and I represent the closed, open, and inactivated states of the sodium channel, respectively, and C* and O* represent the drug-bound closed state and drug-bound open but nonconducting state, respectively.

1 . CLOSED-CHANNEL BLOCK The closed sodium-channel block caused by polyarginine is represented by Scheme 2 without the steps in the bracket. Polyarginine bound to the sodium channel of the squid axon in its closed configuration, thereby blocking the ion conduction and gating mechanism (Wu and Yeh, 1982). The inability of the drug-bound sodium channel to open was demonstrated by a decrease in gating current. The closed sodium-channel block caused by TTX is represented by Scheme 2, including the steps in the bracket. In lobster and squid giant axons, no change in sodium-current kinetics was observed in the course of TTX block (Takata et al., 1966). In single-channel recording experiments with cultured neuroblastoma cells, it was found that external application of 3 nM TTX decreased the number of conducting channels observed for a given number of trials (applications of step depolarizations), but the channel open time and current amplitude remained unchanged (Quandt et al., 1982). Analyses of the dose-response curve indicated that TTX blocked the individual sodium channel on a one-to-one stoichiometric basis with an apparent dissociation constant of 2 nM. This value for

490

TOSHIO NARAHASHI

the apparent dissociation constant agrees well with that obtained in squid giant axons (3 nM; Cuervo and Adelrnan, 1970). The gating current was not affected by TTX (Armstrong, 1975; Almers, 1978). These results suggest that TTX blocks the sodium channel regardless of its gating state. 2. OPEN-CHANNEL BLOCK

Open-channel block has been demonstrated for the potassium-channel block caused by tetraethylammonium derivatives (Armstrong, 1969, 1971), the sodium-channel block by pancuronium (Yeh and Narahashi, 1977), local anesthetics (Strichartz, 1973; Courtney, 1975; 1980; Courtney et ul,, 1978; Hille, 1977; Khodorov et nl., 1976; Yeh, 1978, 1980). 9aminoacridine (Yeh, 1979), N-alkylguanidines (Kirsch et al., 1980), strychnine (Shapiro, 1977), and N-methylstrychnine (Cahalan and Alrners, 1979); and the ACh-channel block by local anesthetics (Steinbach, 1968a,b; Neher and Steinbach, 1978; Adams, 1977), amantadine (Tsai er al., 1978), histrionicotoxin (Masukawa and Albuquerque, 1978), tetraethylammonium (Adler et al., 1979), decamethonium (Adams and Sakmann, 1978), and N-octylguanidine (Farley et al., 1981). Depending on the rate of drug binding to the open channel relative to the channel-opening kinetics, the time course of membrane ionic current could undergo a drastic change. For example, pancuronium accelerated the falling phase of sodium current (Fig. 3). This was interpreted as being A.

CONTROL

c. 1x10-3nr

pc

FIG.3. Effect of internal application of 1 X 10 M pancuronium (Pc) on sodium currents in an axon internally perfused with 20 mM tetraethylammonium. (A, C) Families of sodium currents associated with 20-mV step depolarizing pulses (40-160 mV) from the holding potential of -80 mV. (B) The upper traces are superimposed sodium currents associated with depolarizing pulses to f 8 0 mV before and during application of PC, and the lower traces are those with pulses to 0 mV. Note that the rising phase of sodium current is not affected, whereas its falling phase is accelerated. Temperature, 6.3"C. (Prom Yeh and Narahashi, 1977.)

PHARMACOLOGY OF NERVE MEMBRANE SODIUM CHANNELS

49 1

due to an open-channel block, not to a change in inactivation, because a similar acceleration by pancuronium was observed in the axon treated with Pronase, which destroyed the inactivation mechanism (Fig. 4). The block of sodium channel at its open state also results in a frequency- or use-dependent block by local anesthetics (Strichartz, 1973; Courtney, 1975; Schwarz et al., 1977). Single-channel recording experiments provide us with more detailed insight into the open-channel block. Two cases can be predicted in the behavior of single sodium channel in the presence of an open-channel blocker depending on the rates of drug binding to and unbinding from the open channel. If the rates are slow enough to be resolved by the recording system, bursts or flickerings will be observed while a channel is open. However, if the rates are too fast to be resolved, then the observed current represents an average level of the closed and open states. a . Open-Channel Block with Bursts. The open-channel block exhibiting bursts can be schematically illustrated (Scheme 3). Duration of pulse group (t,) A

T

\

Blocked time (t,)

I

t

open time (to) SCHEME 3

A. i x I O - ~M PC CONTROL

CONTROL

8. Ix n

M PC CONTROL

2 rnsec

FIG. 4. Time course of pancuronium (PC)-induced decay of sodium current in axons internally treated with 20 mM tetraethylammonium after internal Pronase treatment. (A) M PC. (From Yeh and Narahashi, M PC; (B) control and 1 X Control and I x 1977.)

492

TOSHIO NARAHASHI

For the purpose of analysis of these parameters, Scheme 1 can be simplified to Scheme 4.

The mean open time (r,,), blocked time ( t b ) , and burst period (t,) are given by Eqs. (1)-(3) (Neher and Steinbach, 1978).

where V is the membrane potential and Q is the drug concentration. Equations (1)-(3) predict that: (1) the values for t o , t b , and rg are random variables with exponential distribution; (2) the open time ( t o ) decreases with an increase in drug concentration (Q) [Eq. (l)]; (3) the burst period ( t , ) increases with an increase in Q [Eq. (31; and (4) the blocked time ( t b ) is independent of Q [Eq. (2)]. The value for p can be estimated from the mean open time at various membrane potentials in the absence of drug [Eq. (l)]. Using the value of p thus obtained, the rate constant for drug binding, f(V), can be estimated from measurements of to in the presence of drug [Eq. (l)]. The rate constant for drug unbinding, &( V), can be obtained from the blocked time in the presence of drug [Eq. (2)]. Thus the dissociation constant Kd can be calculated from Eq. (4). Kd = h( Wf( V)

(4)

If a monovalent cationic blocking agent acts from the internal membrane surface (e.g., local anesthetics), the voltage dependence of the dissociation constant is given by Eq. ( 5 ) . &(V) = Kd(0)exp(-6VF/RT)

(5)

where Kd(0) is the dissociation constant at 0 mV, 6 is the fraction of electrical distance of the binding site, and F, R, and T are the Faraday constant, gas constant, and absolute temperature, respectively. h. Open-Channel Block without Bursts. If the drug binding and unbinding are too fast to record, no flickering will be observed. Instead the average amplitude of current is expected to decrease, taking a value

PHARMACOLOGY OF NERVE MEMBRANE SODIUM CHANNELS

493

somewhere between the closed and open levels. In such a case, the voltage-dependent block can be given by Eq. (6) (Coronado and Miller, 1979) where Z, and Z represent single-channel current amplitudes in the absence and in the presence of drug, respectively. In the above analyses for both types of open-channel block, it is important to realize that the dissociation constant Kd(V ) thus measured repre0).This sents the value uncontaminated with the channel gating (C value cannot be obtained by measurements of macroscopic peak sodium current. Instantaneous sodium current must be measured if the conventional macroscopic voltage-clamp experiments are performed. 111.

IONIC-CHANNEL MODULATION BY CHEMICALS

Sodium-channel modulators are characterized by their ability to slow the sodium-channel kinetics. Very often the modified sodium channels exhibit voltage dependence greatly shifted in the direction of hyperpolarization with little or no inactivation. This leads to a large depolarization of the membrane. There is evidence in support of the notion that a population of sodium channels is modified in an all-or-none manner depending on the concentration of the modulator. Because of unique actions of the modulators in changing the kinetics of sodium channels, and also because of their membrane depolarizing action, these chemicals have been used extensively as tools for the study of sodium channels. The modulator-induced depolarization is a convenient means by which neurotransmitters can be released under experimental conditions. A. Batrachotoxin

Batrachotoxin is one of the toxic components contained in skin secretion of the Colombian poison arrow frog Phyllobates aurotaeniu. They include batrachotoxin, homobatrachotoxin, batrachotoxin A, and pseudobatrachotoxin. Batrachotoxin (BTX) has a potent and irreversible depolarizing action on membranes of nerves, skeletal muscles, and cardiac muscles. The chemistry and pharmacology of BTX have been reviewed (Albuquerque et al., 1971; Albuquerque and Daly, 1976). In the squid giant axon, BTX caused a large depolarization that exceeded the zero potential level at high concentrations (Narahashi et al., 1971 ; Albuquerque et al., 1973). The depolarization was restored by lowering the exter-

494

TOSHIO NARAHASHI

nal sodium concentration to 1 mM or by adding TTX to the external medium. No depolarization was induced by BTX in the absence of sodium ions in both external and internal media. These observations indicate that the BTX-induced depolarization is due primarily to an increase in sodium permeability at resting conditions (Fig. 5). Khodorov and his associates have discovered interesting changes in sodium-channel kinetics caused by BTX using the node of Ranvier of the frog (Khodorov, 1978; Khodorov et al., 1975; Khodorov and Revenko, 1979). In the presence of BTX, the peak sodium current was followed by a slow, steady-state sodium current that was blocked by TTX. With advancement of BTX intoxication either by increasing the concentration or by prolonging the treatment time, the slow current increased while the peak current decreased. Furthermore, the kinetics of the slow current were greatly shifted in the direction of hyperpolarization, enabling the modified sodium channel to open at large negative membrane potential with little or no inactivation. This caused a depolarization. The BTXmodified sodium channels were less selective to sodium, becoming significantly or measurably permeable to thallium, ammonium, guanidine, potassium, rubidium, cesium, calcium, and even methylamine. We have used squid giant axons for extensive analyses of the BTX action on the sodium-channel kinetics (J. Tanguy, V . M. Scruggs, and T.

I

T T X TTX ' ~ E ~ l ~ ~ A S ' #

A SW

TTX ImM

I mMNo

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I

I mMNo

BTX

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-

-

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2

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c D

._ *

-75

0

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I

20

40

I

1

1

I

60

80

I00

I20

Time i m i n 1

Effects on the resting membrane potential of an internally perfused squid giant batrachotoxin (B'IX)applied internally, 1000 nM tetrodotoxin (TTX) externally, or 1 mM Na externally. Standard internal solution (SIS) contains 50 m M Na. ASW, artifical seawater. (From Narahashi ti of.,1971.) FIG.5 .

axon of 550 nM

PHARMACOLOGY OF NERVE MEMBRANE SODIUM CHANNELS

495

Narahashi, unpublished observation). In the squid axon perfused externally and internally with K-free media, the peak transient sodium current was followed by a steady-state sodium current during a step depolarization of the membrane. The steady-state sodium current was markedly increased by BTX, and the sodium current during a step depolarization resembled the sodium current devoid of inactivation. The curve relating the slow sodium current to the membrane potential was shifted in the direction of hyperpolarization. These two changes in sodium-channel kinetics caused by BTX can account for the observed depolarization. Thus, the squid axon responds to the action of BTX basically in the same manner as the node of Ranvier of the frog. B. Grayanotoxins

Grayanotoxins are the toxic principles isolated from the leaves of plants belonging to the family Ericaceae (Leucothoe, Rhododendron, Andromeda, Kalmia). In squid axons, grayanotoxin 1 (GTX I) and a-dihydrograyanotoxin 11 (a-H2-GTX11) caused a large depolarization due to a specific increase in resting sodium permeability (Seyama and Narahashi, 1973; Narahashi and Seyama, 1974; Hironaka and Narahashi, 1977). Voltageclamp measurements showed that the resting sodium permeability was increased by GTX by a factor of 10-100 depending on the concentration. Tetrodotoxin antagonized the action of GTX in a noncompetitive manner with an apparent dissociation constant of 40 nM, a concentration approximately 10 times higher than the apparent dissociation constant to block peak sodium current.

1. EFFECTSON SODIUM-CHANNEL KINETICS Despite such a large increase in sodium permeability at rest, the nerve membrane was still capable of generating sodium currents if depolarized from a large negative holding potential. However, the sodium currentmembrane potential relationship revealed three changes caused by GTX I (Seyama and Narahashi, 1981): (1) the peak current was reduced; (2) the reversal potential was shifted in the hyperpolarizing direction; and (3) the peak sodium current could be produced at large negative potentials where no current was generated under normal conditions. Analyses of sodium conductance kinetics indicated that both sodium activation and inactivation parameters, including steady-state values and time constants (m,, T, , h,, T,,),were greatly shifted in the direction of hyperpolarization. Experiments with prolonged depolarizing steps (100-200 msec) have revealed a secondary, slowly rising current component (Seyama and

496

TOSHIO NARAHASHI

Narahashi, 1981). The slow current was blocked by TTX. However, the concentration of TTX required completely to block the slow current was higher than that required to block peak current; TTX at 30 nM blocked the slow current only partially, and concentrations of 300 nM or higher were required for complete block. Furthermore, several remarkable features were noted in the GTX-induced slow current (Fig. 6): (I) With increasing concentration of GTX I, the amplitude of slow current increased while that of peak current decreased; (2) the slow current was generated at large negative potentials (e.g., by a depolarizing step from the holding potential of -150 to -100 mV) where no peak current was produced; (3) the reversal potential for slow current was shifted in the hyperpolarizing direction as compared with that for normal peak current; (4) the slow current hardly inactivated during prolonged depolarizing steps; and ( 5 ) the tail current associated with a step repolarization during the slow current was expressed by two exponential functions with time constants of 3.2 and 42.4 msec (for repolarization from -70 to - 110 mV). Observation (5) suggests that there are two processes or pathways involved in the return of the slowly opened sodium channel to its closed state(s). The observed shifts of the reversal potentials for peak and slow currents in the hyperpolarizing direction in the GTX-treated axon suggest that the cation selectivity of the sodium channel is decreased. The reversal potentials for peak currents were measured in external media containing various concentrations of potassium but no sodium and in an internal medium containing 300 mM K+ and 100 mM Na'. The permeability ratio PK/PNa before application of GTX I changed from 0.098 to 0.99 as the reversal potential changed from - 18 to -50 mV as a result of decrease in external potassium concentration. GTX I increased the permeability ratio

(mv)-lO -15*b/ f mAlc m')

-7

-1

/'

-- --- - -- - --< -< -r

0 1 C h S U )

FIG.6. Membrane currents associated with step depolarizations from the holding potenI50 to -70 mV (upper tracings) and to - 10 m V (lower tracings) before and during internal perfusion of various concentrations of grayanotoxin 1 (GTX I) in the presence of 20 mM tetraethylammonium inside a squid giant axon. (From Seyama and Narahashi, 1981.) tial of

-

PHARMACOLOGY OF NERVE MEMBRANE SODIUM CHANNELS

497

to 0.13 at -18 mV and to 1.57 at -50 mV. Thus the selectivity of sodium channel in generating peak current is decreased by GTX I, and the selectivity in both control and GTX-treated membranes decreases with hyperpolarization and/or with the decrease in external potassium concentration. Permeability ratios Px/PNa for other test cations were also measured at various membrane potentials and in the absence and the presence of GTX I. For guanidine, methylguanidine, choline, and cesium, the ratio Px/PNa increased with hyperpolarization and with application of GTX I. However, the increase by GTX I was less than that for PK/PNa. In contrast to relatively small increases in Px/PNa during the peak current caused by GTX I, the increases in the permeability ratio during the slow current were remarkable. In this experiment, the possible contribution of potassium channels to the observed slow current was eliminated for hydroxylamine, formamidine, by the use of TTX. The ratios Px/PNa and ammonium were estimated to be 1.58, 0.71, and 1.10, respectively. Changes in internal pH had dramatic effects on the GTX-induced slow current. An increase in the pH from 7.3 to 9.0 caused the slow current to increase and the peak current to decrease. The titration curve for the amplitude of the slow current in the internal pH range from 5.6 to 9.7 showed two dissociation constants, one at pH 6.1 and another at pH over 9.0 (Fig. 7). In summary, this study has revealed several striking features of the action of GTX I on squid axon membranes. First of all, the peak sodium conductance parameters are shifted in the direction of hyperpolarization. Second, a slow, noninactivating current appears. This current occurs at

lntracellular pH

FIG.7 . Titration curve for the slow sodium currents in 2 (GTX 1). (From Seyama and Narahashi, 1981.)

X

1O-j

M grayanotoxin

1

498

TOSHIO NARAHASHI

large negative membrane potential and is therefore responsible for the GTX-induced depolarization. With increasing GTX concentration, the slow current increases while the peak current decreases. Third, TTX blocks the slow current, but requires higher concentrations than those to block the peak current. Thus, the slow current appears to flow through a sodium channel modified by GTX I. Fourth, the modified channel is less selective to sodium than the normal sodium channel that generates peak current. Fifth, the modified channel exhibits two dissociation constants, one at internal pH 6.1 and the other at pH over 9.0. Sixth, the tail current associated with a step repolarization during the slow current in GTX I decays with a dual exponential time course, suggesting that two steps or pathways are involved for the modified open sodium channel to close. These observations are compatible with kinetic scheme 5 .

c

*

eo

*

e I*

SCHEME 5

C , 0, and I refer to the closed, open, and inactivated states of normal sodium channel, respectively, and C*, O*, and I* are the corresponding states of the GTX-induced modified channel. The reaction O* +-I* may be extremely slow or nonexistent. 2. ACTIVE MOIETIES IN

THE

GKAYANOTOXIN MOLECULE

The modulation of sodium channels caused by GTX I has many features in common with that caused by other modulators such as BTX, veratridine, and aconitine. Thus it has become crucially important to identify the moieties in the GTX molecule that are responsible for the effect and to identify the common groups in the molecules of the other modulators. In order to accomplish this goal, the structure-activity relationship for CiTX has been studied (Masutani ct al., 1981). This was the first step in this series of experiments, and in order to facilitate measurements of a large number of derivatives, a simple preparation and a simple technique were employed. It was more important to select the test compounds carefully, so that GTX molecules were modified in a highly systematic manner. The preparation chosen for this study was the frog sartorius muscle. When exposed to an effective concentration of GTX, the muscle fiber generated an action potential without electrical stimulation, which was

PHARMACOLOGY OF NERVE MEMBRANE SODIUM CHANNELS

499

followed by a sustained depolarization lasting as long as 10-20 min at the potential level of -40 to -30 mV. This was a remarkable effect and could easily be detected by the conventional intracellular microelectrode technique. The potencies of GTX derivatives were estimated by measurements of critical concentrations that were defined as the concentrations at which an action potential was generated spontaneously and followed by a sustained depolarization. The concentration of a derivative was cumulatively increased until the response was obtained. Thirty-four GTX derivatives were examined. a-Dihydrograyanotoxin I1 (a-H2-GTX 11) was one of the most potent GTX derivatives and was used as the reference or parent compound. The results clearly indicate that 3P-OH or 2p,3p-oxide, 5@-OH,6P-OH, and lop-methyl groups are essential for GTX molecules to exert the physiological action. The former three groups are distributed on a 3./ plane forming a triangle, and the 10P-methyl group is located adjacent to the SPOH group. Batrachotoxin, veratridine, and aconitine appear to possess the equivalent groups (Table I). From a limited number of the derivatives so far examined, the following hypothesis has been put forward. Batrachotoxin has three reactive oxygen groups 3P-OH, 3a,9a-oxide, and 1laOH aligned on an a plane forming a triangle, and a 2'-methyl group on the pyrrole ring or methyl group on the 2'-ethyl moiety is in the proximity of the three reactive oxygen groups, thus resembling the situation of GTX. TABLE I COMPONENTS OF ALIGNMENT OF THREE REACTIVEOXYGEN GROUPSI N THE GRAYANOTOXIN (GTX), BATRACHOTOXIN (BTX), A N D VERATRIDINE MOLECULES ESSENTIAL FOR OPENING SODIUM CHANNELS" B and B': proton acceptors. AH or B": proton donor or acceptor

Interatomic distance (A)

Reactive oxygen groups

Compound

B

B'

BB'

B'A(B")

A(B)-

AH or B" 6P-OH

3.1

2.8

4.9

1 la-OH 14a-OH

2.3 3.1

2.6 2.8

3.9 4.4

B

~~~

GTX

BTX Veratridine

3P-OH or 2P,3/3-oxide 3a-OH 17a-OH

From Masutani et al. (1981).

SP-OH

3a,Ya-Oxide 12a-OH

500

TOSHIO NARAHASHI

In the veratridine molecule, the 3-methoxymethyl group on benzoyl moiety can take a conformation in the proximity of the triangle formed on an a plane by 17a-OH, 12a-OH, and 14a-OH groups. In the aconitine molecule, l6P-OCH3, 15a-OH, and 8a-OCOCH) (or 14a-OCOC6H5) are aligned on a triangle plane, and the methyl portion of acetyl group may be able to approach 15a-OH. It then appears that three reactive oxygen groups aligned on a triangle plane with a methyl group in the proximity play a crucial role for GTX, BTX, veratridine, and aconitine to modulate the sodium channel. Two of the reactive hydroxyl or oxide groups can be assumed to be proton acceptors (B and B‘), because the OH group can be replaced by an oxide group that can function only as a proton acceptor. The third reactive OH group could act as either proton donor (AH) or proton acceptor (B”). The distances among the three reactive groups have been estimated. The B-B’ distances are 3.1, 2.3, and 3.1 for GTX, BTX, and veratridine, respectively; the B’-B” distances are 2.8, 2.6, and 2.8 A; the B”-B distances are 4.9, 3.9, and 4.4 A. This study has opened a road toward the molecular mechanism of interactions of these modulators with the sodium channel. C. Pyrethroids

Pyrethroids are the synthetic derivatives of pyrethrins, which are the ~ h e ~ ~ ~ toxic components contained in the flowers of ~ h ~ s a n species. These compounds have been widely used as insecticides. From the neurophysiological point of view, they constitute an extremely interesting group of chemicals, which exhibit highly specific and very potent effects on nerve membrane sodium channels. They share some features with other sodium channel modulators, such as BTX, GTX, and veratridine. 1. INTERACTION OF PYRETHROIDS WITH THE SODIUM CHANNEI

Allethrin, a synthetic pyrethroid, causes repetitive after-discharges to be induced by a single stimulus in nerve fibers as a result of an increase in depolarizing (negative) after-potential (Narahashi, 1962a,b). This change in the after-potential is due to an inhibition of sodium inactivation and potassium activation (Narahashi and Anderson, 1967; Wang et ul., 1972; Murayama et ul., 1972). The mechanism most sensitive to allethrin is sodium inactivation, and the sodium current is greatly prolonged before any other change in membrane current is observed after application of allethrin. We have now undertaken detailed voltage-clamp analyses to determine the nature of interaction of pyrethroids with the sodium channel (Lund and Narahashi, 1981a,b). In order to observe the sodium current uncontaminated by potassium

501

PHARMACOLOGY OF NERVE MEMBRANE SODIUM CHANNELS

component, cesium-substituted potassium-free internal solution was used. The potassium-channel blocking agents, 4-aminopyridine and 3,4diaminopyridine, did not give satisfactory results owing to time-dependent recovery from the block. Tetramethrin markedly slowed the falling phase of sodium current without any effect on its rising phase and peak amplitude (Fig. 8). A high concentration of tetramethrin (3 X M ) was used in these experiments in order to see the effect very clearly. The tail current associated with step repolarization was also large (Fig. 8). With prolonged depolarizing pulses, it became clear that the peak sodium current was followed by a secondary slow current and that only the slow current was seen at small depolarizations. With further prolongation of pulses, the current developed slowly and then decayed very slowly, and the tail current also decayed slowly. The slow current as well as the peak current in the presence of tetramethrin was blocked by 3 X M TTX, indicating that both currents were carried through the sodium channel. Despite the drastic prolongation of sodium current by tetramethrin, the peak component of sodium current was not affected. The time constants of activation (7,) and inactivation (q,)were totally unchanged by tetramethrin. The steady-state sodium inactivation curve as measured with 30msec prepulses was not significantly shifted, although a foot appeared at large depolarizations. The tail current in tetramethrin was expressed by two exponential functions, and the initial phase was identical to the control. Since the tail current also represents the m kinetics, this result agrees with that for 7,. These results strongly suggest that, in the tetramethrinpoisoned axon, there is a population of sodium channels that open and close with normal kinetics and voltage dependence. It then follows that the remaining population of sodium channels may be modified by tetramethrin to give rise to slow kinetics of opening and closing. Therefore, the nature of the slowly developing current was analyzed next. The amplitude of the slowly decaying tail conductance is a measure of A.

Control

B. Treated

Control Treated,

FIG.8. (A) Sodium currents associated with step depolarizations from the holding potential of - 100 mV to -50, -30, - 10, 0, 10, 30, 50, and 70 mV in a control axon. (B) Same as (A), but after internal treatment with 3 X M tetramethrin. (C) Sodium currents recorded at - 10 mV before and 6 min after treatment with tetramethrin. Scale = 1 mA/cm*and 1 msec for (A) and (B); 0.25 mA/cm2and 2 msec for (C). (From Lund and Narahashi, 1981b.)

TOSHI 0 NARAHASHI

502

the population of the tetramethrin-modified channels, and the time course of the development of this slow tail conductance represents the kinetics of arrival of channels in this modified open state. Detailed measurements with squid axons showed that the time course of the arrival of channels in the modified open state was clearly second order, having a quickly developing phase with a time constant of about 1-2 msec and a slowly developing one with a time constant of about 15-30 msec (Fig. 9). Thus there appear to be at least two kinetic steps or pathways that lead to the modi-

8

Time

(mscc)

FIG.9. (A) Amplitude of the slowly decaying tail current after conditioning pulses to X lo-' M tetrarnethrin. (B) Tail currents recorded at -80 m V after conditioning the membrane at +40 mV for various +40 mV for various durations in an axon trealed with 8

durations. Record represents multiple sweeps. Scale = 0.25 mAlcm2, 10 msec. (C) The rising phase of A ( 0 )and the kinetic analysis of the rising phase of A (open symbols) revealing two time constants. (From Lund and Narahashi, 1981b.)

PHARMACOLOGY OF NERVE MEMBRANE SODIUM CHANNELS

503

fied open state of channels. It seems that some channels are modified in the resting state and then open and close in response to changes in membrane potential, whereas others are modified only after they have opened by the normal process. This second modification pathway is necessarily fast, since it must compete for the normal state with inactivation. This model predicts that any treatment that interferes with the h process should potentiate the tetramethrin modification of sodium channels. This has been shown to be the case (Lund and Narahashi, 1982). After pretreatment with Pronase, deoxycholate, or N-bromoacetamide, the doseresponse relation for the tail-current amplitude was shifted in the direction of low dose and the maximum response was more than doubled (Fig. 10).

2. STUDYOF ISOMERS-CHANNEL BINDINGSITES Pyrethroid can exist in two optical and two geometrical isomers. It turned out that (-)-trans and (-)-cis forms of tetramethrin are almost completely inactive on the nerve, whereas (+)-trans and (+)-cis forms are highly active. Such a differential action provided us with an excellent opportunity to study the sites of action of tetramethrin on the sodium

(+)-Trans -Tetramethrin Concentration (MI

FIG.10. Dose-response relationships for (+)-trans-tetramethrin after pretreatment with Pronase (0),N-bromoacetamide (NBA) (A), deoxycholate (DOC) (El),and without pretreatment (0).The initial amplitude of the slowly decaying tail current (I,,,,) following a ISmsec conditioning pulse was normalized to the peak current recorded at -20 mV [[,(-20)] before application of tetramethrin and is plotted as a function of the concentration of tetramethrin. (From Lund and Narahashi, 1982.)

504

TOSHIO NARAHASHI

channel. The experiments were performed with the internally perfused and voltage-clamped squid giant axons (Lund and Narahashi, 1982). The membrane was step depolarized (e.g., to +40 mV) for 15 msec and then repolarized to the holding potential of -80 mV. The tail current associated with the repolarization decayed with second-order kinetics. Since the amplitude of the slowly decaying phase was a measure of the conductance system that was in the modified open state, the tail-current amplitudes were measured 2 msec after repolarizing the membrane to -80 mV. The tail-current amplitudes were normalized to the peak current recorded at -20 mV before application of pyrethroid, so that data from different axons could be pooled together and compared. The tail-current response to the active (+ )-trans-tetramethrin was partially blocked by 3 or 6 nM TTX. The maximum responses were decreased by 29 and 57%, respectively, but the apparent dissociation constant was little affected, suggesting a noncompetitive interaction (Fig. 11). A higher concentration (300 nM) of TTX completely abolished the tailcurrent response. The (-)-trans and (-)-cis isomers were virtually inactive on the sodium M. However, when applied 15 min channel at a concentration of 3 X prior to an active (+) isomer, either form of the (-) isomers was effective in suppressing the action of the subsequently applied (+) isomer. The antagonism was either competitive or noncompetitive in nature depending 07

O6I

(t)- trans-Tetramethrin Concentration ( M )

FIG. 11. Dose-response relationships for (+)-trans-tetramethrin obtained as in Fig. 9 M (0) and 6 x after pretreatment with 3 x M (0)tetrodotoxin (TTX), and without pretreatment (0).(From Lund and Narahashi, 1982.)

PHARMACOLOGY OF NERVE MEMBRANE SODIUM CHANNELS

505

on the combination of (+) and (-1 isomers. The noncompetitive antagonism was observed for the (-)-trans isomer against the (+)-trans, the (-)trans against the (+)-cis, and the (-)-cis against the (+)-trans, whereas the competitive antagonism was seen for the (-)-cis isomer against the (+)-cis isomer (Fig. 12). These results can be accounted for by a scheme of binding sites in the sodium channel (Fig. 13). There are a trans site and a cis site in the channel to which the (+)-trans and (+)-cis isomers bind, respectively, with a high affinity. The (-)-cis isomer binds to the cis site also with a high affinity, causing a competitive antagonism against the (+)-cis isomer. In addition, there is a negative allosteric site to which the (-)-trans isomer binds with a high affinity, causing a noncompetitive

A

I"'

COnird

/-

B

100

/

80/

(tJ-c~s-Tetmmrhr~n Concentram (MI

FIG.12. Dose-response relationships obtained as in Fig. 10 and plotted as percentage of M and 1 x the maximum response. (A) Antagonism of (+)-trans-tetramethrin by 3 x M (-)-trans-tetramethrin. (B) Antagonism of (+)-cis-tetramethrin by 1 x M (-)M (-)-cis-tetratrans-tetramethrin. (C) Antagonism of (+)-trans-tetramethrin by 3 x M (-)-cis-tetramethrin. Values methrin. (D) Antagonism of (+)-cis-tetramethrin by 1 X shown are mean 2 SEM, where n = 3. (From Lund and Narahashi, 1982.)

506

TOSHIO NARAHASHI (+)-trans

(- k trans

(high affinity)

(low affinity)

(-)-Irons (high affinity1

TTX

J,

negative allosteric site

channel

.r

(-)

CIS

‘74

(low affinity)

b)-cis (high affinity)

7(

1 CIS

(hlgh affinity)

FIG.13. Hypothetical model for the interactions of tetramethrin with the sodium channel. TTX, tetrodotoxin. (From Lund and Narahashi, 1982.)

antagonism against the (+)-trans and (+)-cis isomers. Tetrodotoxin binds to another site, thereby causing a noncompetitive antagonism against the (+)-trans and (+)-cis isomers. In summary, these optical and geometrical isomers of tetramethrin provide us with excellent probes for the study of drug binding sites in the sodium channel. They can also be used to identify the binding sites of othcr typcs of sodium-channel modulators, and experiments are in progress in our laboratory along this line. D. Slngle-Channel Studles

The drastic modifications of the sodium-channel kinetics caused by various modulators predict significant changes in behavior of single channels. It should be emphasized that both “macroscopic” and “microscopic” analyses must be performed in order to interpret the kinetics of sodium channels and their modification caused by chemicals. In our laboratory, batrachotoxin, tetramethrin, and veratridine have so far been studied for their effects on single sodium channels (Quandt and Narahashi, 1982; Yamamoto et al., 1982; Yoshii et al., 1983). Although these experiments were conducted with cultured neuroblastoma cells, not squid axons, the results are crucial for interpreting the macroscopic current data obtained with squid axons. 1. BATRACHOTOXIN

Individual sodium channels opened during a step depolarization of the membrane from a holding potential of -90 to -50 mV (Quandt and Narahashi, 1982). The amplitude histogram showed a mean value of 1.1 PA.

PHARMACOLOGY OF NERVE MEMBRANE SODIUM CHANNELS

507

The open time could be described by a Poisson distribution with a mean of 2.2 msec. The probability of opening during a depolarizing step first increased and then decreased slowly, forming the time course of the “macroscopic” sodium current. An example of such an experiment with a normal cell is shown in Figs. 14A-D. Batrachotoxin caused several drastic changes (Figs. 14E-H). Channels opened longer, and the open time could be described by two exponential functions. The slower components had a mean duration of 62.5 msec. The mean amplitude decreased to 0.79 PA. The probability of opening formed a curve mimicking a macroscopic sodium current without inactivation. Openings of channels could be observed at large negative membrane potentials (e.g., -80 mV) where no opening was seen before treatment with BTX. This explains the large depolarization of membrane caused by BTX (Narahashi et al., 1971). Another important observation with single-channel recording is that two populations of sodium channels, one normal and the other modified by BTX, are clearly discernible. When the amplitude of each opening is plotted against the open time, the population of sodium channels in the presence of BTX can be separated into two groups, one having the normal amplitude and open time and the other having the reduced amplitude and prolonged open time (Fig. 15). This strongly suggests that BTX modifies individual sodium channels in an all-or-none manner.

2. TETRAMETHRIN Since tetramethrin induces a slow sodium current in squid and crayfish giant axons as described above, it is also expected to change the single sodium-channel current. This was actually the case (Yamamoto et al., 1983). The open time was greatly prolonged by tetramethrin, exhibiting two populations-one with the normal value and the other with a prolonged time (Fig. 16). However, the single-channel current amplitude was not affected by tetramethrin. IV. CONCLUSIONS

Much progress has been made on the squid axon pharmacology during the past two decades. Classical voltage-clamp analyses with internally perfused axons have revealed a number of important features of drugchannel interactions. These include the mechanisms of sodium-channel block caused by tetrodotoxin and local anesthetics, and the mechanisms

B

C

351

'=1

D

0.0241

E

, Amplihrde (PA)

F

d "1

M"1 h

G

25

SO8

Open Time (msec)

A

2.01 *

.

. : .

n

a

. *

20

40

ao

6.0

10.0

OPEN TIME (msec)

J

2o

40

80 loo OPEN TIME (msec)

60

120

140

160

FIG. 15. Two populations of open states for sodium channels observed in membrane patches exposed to 10 p M batrachotoxin (BTX). Scattergrams of the amplitude vs the duration (open time) of each conducting state are shown. (A) All channel openings from the experiments shown in Fig. 14A-D are represented. No clear correlation between these parameters exists. Results from excised membranes exposed to BTX (Fig. 14E-H) are shown in panel B. Note the longer time scale for the abscissa compared to panel A. The larger-amplitude open states have short open times, and the events with long open times all have the smallest current amplitudes. (From Quandt and Narahashi, 1982a.) FIG. 14. Properties of single sodium channels in neuroblastoma cells NIE-115 before and after exposure to batrachotoxin. (A-D) An outside-out, cell-free membrane patch was hyperpolarized to a holding potential of -90 mV; pulses then were applied to depolarize the membrane to -50 mV. Membrane current recordings during 14 of these pulses are shown in A. Inward current is downward. The time course of the depolarization ( V , ) is shown by the lowermost trace in A ( I 1°C). (B) The distribution of current amplitudes. Xis the mean of the distribution, and IT is the standard deviation. The continuous line plots a normal distribution with the parameters as given. ( C )The number of channel openings that had durations greater than the times on the abscissa are plotted. The line shows a best fit to a single exponential curve with the parameters as given. 2 is the mean duration of the open state. The probability that a channel was open as a function of time from the beginning of the depolarizing pulse is plotted in D. The termination of the pulse is marked by the arrow. (E-H) The above properties were measured for a second patch of membrane in an experiment similar to that shown in A-D, except 10 pM batrachotoxin (BTX) was added to the internal solution in the patch pipette. Holding potential, -90 mV. Pulses to depolarize the membrane to -54 mV were given. Fourteen representative examples of evoked membrane currents are shown in E. The vertical lines in each record are residual, uncompensated capacitative currents at the onset and termination of the depolarization. Note that the time scale is different from that in A (10°C). F, G, and H are the distribution of the amplitudes of the open states, the distribution of durations of the open states, and the probability of channels being open, respectively. for the membrane exposed to BTX. The solid line in G is that for the equation given and has two exponential terms. (From Quandt and Narahashi, 1982a.)

510

TOSHIO NARAHASHI

A

CONTROL

OPEN TIME (rnsec)

OPEN TIME (rnsec)

FIG. 16. (A) Control. Distribution of the open time of single sodium channels in an inside-out membrane patch excised from the neuroblastoma cell NIE-I IS. The number of events having an open time longer than that indicated on the abscissa is plotted. The distribution can be fitted by a single exponential function with a decay rate constant of 0.59 msec-I. (9)Tetramethrin. Same membrane patch as that in the control, but after exposure to an internal solution containing 60 pM (+)-trans-tetramethrin. The distribution of the open time longer than 5 msec is fitted by a single exponential function with a decay rate constant of 0.06 msec-I. The distribution of the open time shorter than 5 msec can be divided into two component>. The component obtained by subtraction of the long time constant component from the total population is plotted in the inset using an expanded time scale and can be fitted by a single exponential function with a decay rate constant of 0.53 msec I , This value is very close to the rate constant obtained before application of tetramethrin, and this component probably represents the unmodified population of sodium channels in the presence of tetramethrin. (From Yamamoto et d.,1983.)

whereby the channel kinetics are drastically modified by chemicals, to mention a few. These results are now being integrated into those recently obtained by patch-clamp single-channel recording techniques, albeit different materials have been used. In the light of the extremely powerful approach provided by single-channel recording techniques, the classical voltage-clamp data using squid giant axons are becoming very useful, as they provide us with the basic picture with a high degree of accuracy of measurements. ACKNOWLEDGMENTS

Our studies cited in this article were supported by NIH Grants NS14143 and NS14144.1 thank Janet Henderson for secretarial assistance.

PHARMACOLOGY OF NERVE MEMBRANE SODIUM CHANNELS

511

REFERENCES Adams, P. R. (1977). Voltage jump analysis of procaine action on frog end-plate. J . Physiol. (London) 268,291-318. Adams, P. R . , and Sakmann, B. (1978). Decamethonium both opens and blocks endplate channels. Proc. Null. Acad. Sci. U . S . A . 75, 2994-2998. Adler, M., Oliveira, A. C., Albuquerque, E. X., Mansour, N. A., and Eldefrawi, A. T. (1979). Reaction of tetraethylammonium with the open and closed conformations of the acetylcholine receptor ionic channel complex. J . Gen. Physiol. 74, 129-152. Albuquerque, E. X., and Daly, J. W. (1976). Batrachotoxin, a selective probe for channels modulating sodium conductances in electrogenic membranes. In “The Specificity and Action of Animal, Bacterial and Plant Toxins. Receptors and Recognition” (P. Cuatrecasas, ed.), Ser. B, Vol. I , pp. 297-338. Chapman & Hall, London. Albuquerque, E. X . , Daly, J., and Witkop, B. (1971). Batrachotoxin: Chemistry and pharmacology. Science 172, 995-1002. Albuquerque, E. X., Seyama, I., and Narahashi, T. (1973). Characterization of batrachotoxin-induced depolarization of the squid giant axons. J . Pharmacol. Exp. Ther. 184, 308-314. Almers, W. (1978). Gating currents and charge movements in excitable membranes. Rev. Physiol. Biochem. Pharmacol. 82,96-190. Armstrong, C. M. (1969). Inactivation of the potassium conductance and related phenomena caused by quaternary ammonium ion injection in squid axons. J . Gen. Physiol. 54, 553-575. Armstrong, C. M. (1971). Interaction of tetraethylammonium ion derivatives with the potassium channels of giant axons. J . Gen. Physiof. 58, 413-437. Armstrong, C. M. (1975). Ionic pores, gates, and gating currents. Q . Rev. Biaphys. 7, 179-2 10. Cahalan, M. D., and Almers, W. (1979). Interactions between quaternary lidocaine, the sodium gates, and tetrodotoxin. Biophys. J . 27, 39-56. Catterall, W. A. (1980). Neurotoxins that act on voltage-sensitive sodium channels in excitable membranes. Annu. Reu. Pharmacol. Toxicol. 20, 15-43. Colquhoun, D., and Sakmann, B. (1981). Fluctuations in the microsecond time range of the current through single acetylcholine receptor ion channels. Nufirre (London) 294, 464-466. Conti, F., and Neher, E. (1980). Single channel recordings of K+ currents in squid axons. Nature (London) 285, 140-143. Coronado, R., and Miller, C. (1979). Voltage-dependent caesium blockade of a cation channel from fragmented sarcoplasmic reticulum. Nafure (London) 280, 807-810. Coronado, R . , and Miller, C. (1980). Decamethonium and hexamethonium block K+ channels of sarcoplasmic reticulum. Nature (London) 288, 495-497. Coronado, R., and Miller, C. (1982). Conduction and block by organic cations in a K+selective channel from sarcoplasmic reticulum incorporated into planar phospholipid bilayers. J . Gen. Physiol. 79, 529-547. Courtney, K . R. (1975). Mechanism of frequency-dependent inhibition of sodium currents in frog myelinated nerve by the lidocaine derivative GEA 968. J . Pharmacol. Exp. Ther. 195, 225-236. Courtney, K. R. (1980). Structure-activity relations for frequency-dependent sodium channel block in nerve by local anesthetics. J . Pharrnacol. Exp. Ther. 2l3, 114-1 19. Courtney, K. R., Kendig, J. J., and Cohen, E. N. (1978). The rates of interaction of local anesthetics with sodium channels in nerve. J . Pharmacol. Exp. Ther. 207, 594-604.

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Horn, R., Patlak, J., and Stevens, C. (1981~).The effect of tetramethylammonium on single sodium channel currents. Biophys. J. 36, 321-327. Jackson, M. B., and Lecar, H. (1979). Single postsynaptic channel currents in tissue cultured muscle. Nature (London) 282, 863-864. Khodorov, B. I. (1978). Chemicals as tools to study nerve fiber sodium channels; effects of batrachotoxin and some local anesthetics. I n “Membrane Transport Processes” (D. C. Tosteson, A. 0. Yu, and R. Latorre, eds.), Vol. 2, pp. 153-174. Raven, New York. Khodorov, B. I. (1979). Some aspects of the pharmacology of sodium channels in nerve membrane. Process of inactivation. Biochem. Pharmacol. 28, 1451-1459. Khodorov, B. I., and Revenko, S. V. (1979). Further analysis of the mechanisms of action of batrachotoxin on the membrane of myelinated nerve. Neuroscience 4, 1315-1330. Khodorov, B. I., Peganov, E. M., Revenko, S. V., and Shishkova, L. D. (1975). Sodium currents in voltage clamped nerve fiber of frog under the combined action of batrachotoxin and procaine. Brain Res. 84, 541-546. Khodorov, B. I., Shishkova, L., Peganov, E., andRevenko, S. (1976). Inhibition of sodium currents in frog Ranvier node treated with local anesthetics. Role of slow sodium inactivation. Biochim. Biophys. A c f a 433, 409-435. Kirsch, G. E., Yeh, J. Z., Farley, J. M., and Narahashi, T. (1980). Interaction of n-alkylguanidines with the sodium channels of squid axon membrane. J . Gen. Physiol. 76, 3 15-335. Lazdunski, M., Balerna, M., Barhanin, J., Chicheportiche, R., Fosset, M., Frelin, C., Jacques, Y., Lombet, A., Pouyssegur, J., Renaud, J. F., Romey, G., Schweitz, H., and Vincent, J. P. (1980). The voltage-dependent sodium channel as a drug receptor. I n “Neurotransmitters and Their Receptors” (U. Z. Littauer, Y. Dudai, I. Silman, V. I. Teichberg, and Z. Vogel, eds.), pp. 51 1-530. Wiley, New York. Lund, A. E., and Narahashi, T. (1981a). Modification of sodium channel kinetics by the insecticide tetramethrin in crayfish giant axons. Neuroroxicology 2, 213-229. Lund, A. E., and Narahashi, T. (1981b). Kinetics of sodium channel modification by the insecticide tetramethrin in squid axon membranes. J. Phurmucol. Exp. Ther. 219, 464-473. Lund, A. E., and Narahashi, T. (1982). Dose-dependent interaction of the pyrethroid isomers with sodium channels of squid axon membranes. Neuroroxicology 3, 11-24, Lux, H. D., and Nagy, K. (1981). Single channel Caz’ currents in Helix pomatia neurons. PJugers Arch. 391, 252-254. Lux, H. D., Neher, E., and Marty, A. (1981). Single channel activity associated with the calcium dependent outward current in Helix pomatia. PJugers Arch. 389, 293-295. Marty, A. (1981). Ca-dependent K channels with large unitary conductance in chromaffin cell membranes. Nature (London) 291,497-500. Masukawa, L. M., and Albuquerque, E. X. (1978). Voltage- and time-dependent action of histrionicotoxin on the endplate current of the frog muscle. J. Gen. Physiol. 72, 351-367. Masutani, T., Seyama, I., Narahashi, T., and Iwasa, J. (1981). Structure-activity relationship for grayanotoxin derivatives in frog skeletal muscle. J. Pharmacol. Exp. Ther. 217, 812-819. Murayama, K., Abbott, N. J., Narahashi, T., and Shapiro, B. I. (1972). Effects of allethrin and Condylacris toxin on the kinetics of sodium conductance of crayfish axon membranes. Comp. Gen. Pharmacol. 3, 391-400. Narahashi, T. (1962a). Effect of the insecticide allethrin on membrane potentials of cockroach giant axons. J . Cell. Comp. Physiol. 59, 61-65. Narahashi, T. (1962b). Nature of the negative after-potential increased by the insecticide allethrin in cockroach giant axons. J . Cell. Cornp. Physiol. 59, 67-76.

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Narahashi, T. (1972). Mechanism of action of tetrodotoxin and saxitoxin on excitable membranes. Frd. Proc. Fed. Am. Soc. Exp. Biol. 31, 1124-1132. Narahashi, T. (1974). Chemicals as tools in the study of excitable membranes. Physiol. Reu. 54, 813-889. Narahashi, T., and Anderson, N. C. (1967). Mechanism of excitation block by the insecticide allethrin applied externally and internally to squid giant axons. Toxicol. Appl. Pharmacol. 10, 529-547. Narahashi, T., and Seyama, I. (1974). Mechanism of nerve membrane depolarization caused by grayanotoxin I. J . Physiol. (London) 242, 471-487. Narahashi, T., Deguchi. T., Urakawa, N., and Ohkubo, Y. (1960). Stabilization and rectification of muscle fiber membrane by tetrodotoxin. A m . J . Physiol. 198,934-938. Narahashi, T.,Moore, J . W., and Scott, W. R. (1964). Tetrodotoxin blockage of sodium conductance increase in lobster giant axons. 1. Gen. Physiol. 47, 965-974. Narahashi, T., Haas, H. G . , and Therrien, E. F. (1967). Saxitoxin and tetrodotoxin: Companson of nerve blocking mechanism. Science 157, 1441-1442. Narahashi, T., Albuquerque, E. X., and Deguchi, T. (1971). Effects of batrachotoxin on membrane potential and conductance of squid giant axons. J . Gen. Physiol. 58,54-70. Neher, E. (1981). Unit conductance studies in biological membranes. In “Techniques in Cellular Physiology” (P. F. Baker, ed.). Elsevier, Amsterdam. Neher, E., and Sakmann, B. (1976). Single-channel currents recorded from membrane of denervated frog muscle fibres. Nature (London) 260, 779-802. Neher, E., and Steinbach, J . H. (1978). Local anaesthetics transiently block currents through single acetylcholine-receptor channels. J . Physiol. (London) 277, 153- 176. Neher, E., Marty, A., and Sigworth, F. J . (1981). Currents through individual ionic channels in chemically and electrically excitable cells. I n t . Biophys. Congr., 7[h, Pun-Am. Biochutn. Congr., 3rd, p. 297 (Abstr.). Ogden, I). C., Sicgclbaum, S. A., and Colquhoun, D. (1981). Block of acetylcholineactivated ion channels by an uncharged local anaesthetic. Narore ( Z ~ r i d ~ ~289, nl 596-59X.

Ohmori, H., Yoshida, S., and Hagiwara, S . (1981). Single K ’ channel currents of anomalous rectification in cultured rat myotubes. Proc. Natl. Acad. Sci. U . S . A . 78, 4960-4964. Pallotta, B. S . , Magleby, K . L., and Barrett, J. N. (1981). Single channel recordings of CaZ+activated K+ currents in rat muscle cell culture. Nature (London) 293, 471-474. Pdtlak, J., and Horn, R . (1982). Effect of N-bromoacetamide on single sodium channel currents in excised membrane patches. J . Gen. Physiol. 79, 333-351. Patlak, J. B . , Gration, K. A. F., and Usherwood, P. N. R. (1979). Single glutamate-activated channels in locust muscle. Nature (London) 278, 643-645. Uuandt, F. N., and Narahashi, T. (1981). Characteristics of single sodium channel currents and their modification by batrachotoxin in neuroblastoma cells. Annu. M e e f . Soc. Neurosci., 11th. 7, 902 (Abstr.). Quandt, F. N., and Narahashi. T. (1982a). Modification of single Na’ channels by batrachotoxin. Proc. Natl. Acad. Sci. U.S.A.79, 6732-6736. Quandt, P. N., and Narahashi, T. (1982b). Properties of delayed rectified K channels in neuroblastoma cells. Annu. M c c i . Soc. Ncurosci., I Z i h , 8, 124 (Abstr.). Quandt, F. N., Yeh, J. Z., and Narahashi, T. (1982). Contrast between open and closed block of single Na channel currents. Eiophys. J . 37, 319a. Kitchie, J. M. (1979). A pharmacological approach to the structure of sodium channels in myelinated axons. Annu. Reu. Nerrrusci. 2, 341-362. Sakmann, 8.(1978). Acetylcholine-induced ionic channels in rat skeletal muscle. Fed. Proc. Fed. A m . Soc. Exp. Biol. 37, 2654-2659.

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Sakmann, B., Patlak, J., and Neher, E. (1980). Single acetylcholine-activated channels show burst-kinetics in presence of desensitizing concentrations of agonist. Nature (London) 286,71-73. Schwarz, W., Palade, P. T., and Hille, B. (1977). Local anesthetics. Effect of pH on usedependent block of sodium channels in frog muscle. Biophys. J . 20, 343-368. Seyama, I., and Narahashi, T. (1973). Increase in sodium permeability of squid axon membranes by a-dihydrograyanotoxin 11. J . Pharmacol. Exp. Ther. 184, 299-307. Seyama, I . , and Narahashi, T. (1981). Modulation of sodium channels of squid nerve membranes by grayanotoxin I. J . Pharmacol. Exp. Ther. 219,614-624. Seyama, I., Wu, C. H., and Narahashi, T. (1980). Current-dependent block of nerve membrane sodium channels by paragracine. Biophys. J . 29, 531-537. Shanes, A. M., Freygang, W. H., Grundfest, H., and Amatniek, E. (1959). Anesthetic and calcium action in the voltage clamped squid giant axon. J . Gen. Physiol. 42,793-802. Shapiro, B. I. (1977). Effects of strychnine on the sodium conductance of the frog node of Ranvier. J . Gen. Physiol. 69, 915-926. Sigworth, F. J., and Neher, E. (1980). Single Na+ channel currents observed in cultured rat muscle cells. Nature (London) 287, 447-449. Steinbach, A. B. (1968a). Alteration by Xylocaine (lidocaine) and its derivatives of the time course of the endplate potential. J . Gen. Physiol. 52, 144-161. Steinbach, A. B. (1968b). A kinetic model for the action of Xylocaine on receptors for acetylcholine. J . Gen. Physiol. 52, 162-180. Steinbach, J. H. (1977). Local anesthetic molecules transiently block currents through individual open acetylcholine receptor channels. Biophys. J . 18, 357-358. Strichartz, G. (1973). The inhibition of sodium currents in myelinated nerve by quaternary derivatives of lidocaine. J . Gen. Physiol. 62, 37-57. Takata, M., Moore, J. W., Kao, C. Y., and Fuhrman, F. A. (1966). Blockage of sodium conductance increase in lobster giant axon by tarichatoxin (tetrodotoxin). J . Gen. Physiol. 49, 977-988. Taylor, R. E. (1959). Effect of procaine on electrical properties of squid axon membranes. Am. J . Physiol. 196, 1071-1078. Tsai, M.-C., Mansour, N. A , , Eldefrawi, A. T., Eldefrawi, M. E., and Albuquerque, E. X. (1978). Mechanism of action of amantadine on neuromuscular transmission. Mol. Pharmacol. 14,787-803. Vogel, S. M., Watanabe, S. , Yeh, J. Z . , Farley, J . M., and Narahashi, T. (1984). Currentdependent block of end-plate channels by guanidine derivatives. J . Gen. Physiol. In press. Wang, C. M., Narahashi, T., and Scuka, M. (1972). Mechanism of negative temperature coefficient of nerve blocking action of allethrin. J . Phormoco/. Exp. Ther. 182, 442-453. Woodhull, A. M. (1973). Ionic blockage of sodium channels in nerve. J . Gen. Physiol. 61, 687-708. Wu, C. H . , and Yeh, J. Z. (1982). Effects of polyarginine on gating current of squid sodium channels. Biophys. J . 37, 314a. Yamamoto, D., Quandt, F. N., and Narahashi, T. (1982). Modification of single sodium channels by the insecticide tetramethrin. Annrr. M w t . Soc. Nertrosci., I21h. 8, 25 I (Abstr.). Yamamoto, D., Quandt, F. N., and Narahashi, T. (1983). Modification of single sodium channels by the insecticide tetrdmethrin. Brain Res. 274, 344-349. Yeh, J. Z. (1978). Sodium inactivation mechanism modulates QX-314 block of sodium channels in squid axons. Biophys. J . 24, 569-574.

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Yeh, J. Z. (1979). Dynamics of 9-aminoacridine block of sodium channels in squid axon. 1. Gen. Physid. 73, 1-21. Yeh, J. Z. (1980). Blockage of sodium channels by stereoisomers of local anesthetics. In “Molecular Mechanisms of Anesthesia” ( 8 . R. Find, ed.), pp. 35-44. Raven, New York. Yeh, J. Z. (1982). A pharmacological approach to the structure of the Na channel in squid axon. 111 “Proteins in the Nervous System: Structure and Function” (B. Haber, J. Prez-Polo, and J. Coulter, eds.), pp. 17-49. Liss, New York. Yeh, J. 2.. and Narahashi. T. (1977). Kinetic analysis of pancuronium interaction with sodium channels in squid axon membranes. J. Cen. Physid. 69, 293-323. Yoshii, M . , Luke, V. S., and Narahashi, T. (1983). Effect of veratridine on single sodium channel currents. Annrc. Meet. SUC.Newusci.. Ijrh. 9, 674.

Part IV

Interaction between Giant Axon and Neighboring Cells

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CURRENT TOPICS IN MEMBRANES AND TRANSPORT, VOLUME 22

The Squid Giant Synapse RODOLFO R . LLINAS Department of Physiology and Siophysics N e w York Uniuc,rsity Mrdic,ul Center. New York, New York

I. Introduction..

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

11. Anatomy ................................

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111. Early Electrophysiology . . IV. Synaptic Transmis ction Potentials .......................... V. Electrophysiological Properties of the Postsynaptic Potential. ............... A. Quanta1 N a t u r e . . ...................... ............. B. Nature of the Transmitter Substance.. ... ............. C. Equilibrium Potential. ................. ........................ D. Voltage Dependence of the Synaptic Cur ............. VI. Voltage Clamp of the Presynaptic Element Using Square Pulses.. ........... A. Presynaptic Calcium Current.. ...................................... B. Postsynaptic Potential as a Function of Presynaptic Calcium Current.. .. C. Synaptic Delay .................................................... D. A Model for Synaptic Transmission.. ................................ VII. Voltage Clamp with Action Potential Waveform.. ......................... A. Relationship between Amplitude of Presynaptic Spikelike Depolarizdtion and lc,,.............................................. 540 B. Relationship between Ca Current, Transmitter Release, and EPSP Latency for an Action Potential . . . . . . . . . . . . . . . . 542 References ..... ....................................... 544

1.

INTRODUCTION

Among the experimental preparations utilized for the study of the socalled depolarization release coupling in chemical synaptic transmission, none has been as useful as the squid giant synapse. In particular, the rather large but otherwise morphologically simple presynaptic terminal has allowed direct inquiry into the electrophysiology that accompanies transmitter release. Indeed, most of what is currently known about the 519 Copyright 6 1984 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-153322-0

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electrical events leading to synaptic transmission, from a presynaptic vantage point, has been obtained in this structure. The “giant synapse” is the last junction in a chain of giant neurons that form the escape system in squid. This is a bilaterally symmetrical system that originates with the first-order giant neurons located one on each side of the ventral magnocellular lobe of the brain. They receive-among others-visual, vestibular, and tactile inputs (Young, 1938). Each of the two first-order neurons sends its axon toward the midline, where it fuses with its counterpart to form a cytoplasmic bridge. After forming this bridge the two axons course caudally to make axo-axonic synapses with the second-order giant cells in the dorsal magnocellular lobe. The axons of the second-order neurons in turn leave the brain in the pallial nerves and terminate at the stellate ganglia (found at each side of the dorsomedial region of the mantle). In each stellate ganglion the second-order axon enlarges to form a “palm” that branches to generate 8-10 presynaptic terminal digits (Fig. IA). Each one of these terminals contacts a thirdorder axon, the last link in the chain. This contact takes place via spinal protrusions emerging from the third-order axon (Fig. IB), which demarcate the extent of the junction (arrows). The third-order axons are the postsynaptic elements of the giant synapses. They arise from the fusion of the axons of many small neurons located in the stellate ganglion (Fig. 1A) and spread radially to innervate the mantle musculature. Thus, the giant synapses are axo-axonic junctions between the axon of the second-order giant neuron in the brain and the giant axons of the stellate ganglion. The synapse that has received the most attention is the largest and most distal of these junctions and is commonly referred to as “the giant synapse” (Fig. 1A). FIG. 1. Light microscopic photograph of the stellate ganglion illustrating the giant synapse. In panel A the presynaptic fiber was stained with methyl blue and showed seven preterminals arising from the second-order giant fiber. The postsynaptic element, stained with procion red, is the largest of the tertiary giant axons. These axons are generated by the fusion of a large number of smaller axons originating in the neurons of the stellate ganglion. Retrograde staining of the fibers of origin of the most distal giant axon is observed in this picture (from R. LlinBs, M. Sugimori, and J. M. Bower, unpublished results). The actual synapse is made between dendrite-like protrusions arising from the postsynaptic axon and the presynaptic terminal. These protrusions are visible in the upper edge of the tertiary axon (panel B) following methyl blue staining. The arrows indicate the extent of the synaptic contact. In panel C, aequorin light emission from the preterminal digit during depolarization is large enough to generate transmitter release. The electrode to the left measured voltage while that to the right injected current. The pre- and postsynaptic fibers are outlined in the picture. (LlinBs, Sugimori, and Bower, 1983, and unpublished observations.) Panel A, x 16; panel B, x57; panel C, ~ 8 3 .

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It. ANATOMY The first-order giant cell was first described by Williams in 1910, as were the pre- and postsynaptic fibers in the stellate ganglion. The synapse itself was missed by this author, who believed that the presynaptic axons, after entering the ganglion, divided into several branches, “one of which enters each of the larger nerves which arise from that ganglion, and thus the fiber branches pass to various parts of the mantle” (Williams, 1910, p. 74). The actual synaptic junction was described over 20 years later by J . Z. Young (1936, 1938), who was also the first to describe accurately all the components of the giant neuronal system (Young, 1939). Today it is agreed that the actual contact in the giant synapse is made between the smooth surface of the presynaptic terminal and spinelike processes that protrude from the postsynaptic axon (Fig. IB) (Young, 1939; Robertson, 1963) and envelop the terminal (Young, 1973; Pumplin and Reese, 1978). These dendrite-like processes branch to form the junctions, as shown in Fig. 2, the postsynaptic bulbous heads having a diameter of approximately I pm (Hama, 1962; Pumplin and Reese, 1978). The number of contacts between a presynaptic terminal and postsynaptic element has been estimated to be 10,000 (Pumplin and Reese, 1978). The site of synaptic contact is limited to the “head” of each spinous process by the curvature of the postsynaptic elements and the presence of connective tissue surrounding the necks of the postsynaptic spines. At the ultrastructural level the presynaptic terminal demonstrates the usual synaptic vesicles (Fig. 2), which have been observed with both transmission electron microscopy (Hama, 1962; Castejon and Villegas, 1964; Martin and Miledi, 1975; Pumplin and Reese, 1978) and freeze-fracture techniques (Pumplin and Reese, 1978). The vesicles are located immediately opposite the postsynaptic spines (Hama, 1962; Pumplin and Reese, 1978), suggesting that vesicle fusion (the active zone) takes place at that point. As will be seen later, because transmitter can be released within a very short time after presynaptic calcium entry, the site of ionic influx current and that for transmitter release must be very close to each other (Llinas, 1977). Furthermore, based on those measurements, it has been postulated that the active-zone particles observed by freeze fracture at the presynaptic membrane could represent the actual site of calcium entry (Pumplin et al., 1981). Preliminary quantification of particle number on freeze-fracture replicas is consistent with the view that these particles are the calcium channels (Pumplin et al., 1981), the average number of active zone particles per synapse being 1 . 1 x 10’.

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EARLY ELECTROPHYSIOLOGY

Electrophysiologically, the squid giant synapse was first studied by J. Z. Young, who demonstrated that activation of the secondary giant axon could produce contraction of the mantle (1939). The first recordings from the synapse were made by T. H. Bullock (1946, 1948); his extracellular recordings of the synaptic potential produced by activation of the presynaptic terminal led him to suggest that transmission in this synapse, although rectifying, was electrical. In short order, Bullock and S. Hagiwara (1957) demonstrated by intracellular recordings from the postfiber that transmission in the junction was probably chemical and that the size of the presynaptic action potential affected the EPSP amplitude. The following year, Hagiwara and Tasaki (1958) were the first to demonstrate directly that transmission was chemical. They recorded the effects of postsynaptic polarization on EPSP size and measured synaptic current by voltage clamping the postsynaptic fiber, using a technique similar to that developed for the isolated giant axon (Hodgkin et al., 1952). In the same year Bryant (1958) carried out the first detailed pharmacological study of this synapse and demonstrated that oxygen was necessary to maintain synaptic transmission. He also noted that synaptic transmission was optimal when the magnesium in the saline was lowered and the calcium raised. Also belonging to this early period are the studies of Takeuchi and Takeuchi (1962) and Miledi and Slater (1966), who investigated the relationship between presynaptic spike height and postsynaptic response in some detail. They demonstrated that transmitter release is related to the action potential height with respect to the resting level, but not to its absolute amplitude. Another puzzling finding at the time was that, although calcium (Takeuchi and Takeuchi, 1962; Miledi and Slater, 1966) and magnesium (Takeuchi and Takeuchi, 1962) had a marked effect on the amplitude of the postsynaptic current, these ions did not change the height of the presynaptic action potential. Takeuchi and Takeuchi (1962) concluded that release was not triggered directly by the action potential but through some intermediate process; Miledi and Slater showed that in a calcium-free saline, extracellular iontophoresis of calcium at the synaptic junction restored transmitter release (1966). The latter authors postuFIG.2. Electron micrograph of the junctional sites in the giant synapse. The presynaptic apposition to the postsynaptic protrusions (post) indicates the surface of the preterminal. The active zone is indicated by arrows to the right. The pre- and postsynaptic axons are separated from each other by a glial interphase ( G ) seen here filling the space between these two axons. (Modified from Pumplin and Reese, 1978.)

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lated that extracellular calcium was required for transmission and that reactions leading to transmitter release were triggered by the combination of calcium with a “Ca-receptor” on the external surface of the membrane. IV. SYNAPTIC TRANSMISSION WITHOUT ACTION POTENTIALS

Up to this time research in the squid giant synapse had been important in demonstrating the relation between amplitude of the presynaptic action potential and transmitter release as well as in elucidating the properties of the postsynaptic response, but the next steps in the study were probably the more significant to our understanding of the mechanism of transmitter release. In 1966 it was shown, after pharmacological blockage of the action potential by tetrodotoxin (TTX), that release of transmitter from the presynaptic terminal, as determined by the postsynaptic response, could be obtained by direct depolarization of the preterminal (Bloedel et al., 1966; Katz and Miledi, 1966; Kusano et al., 1967). These experiments indicated that transmitter release was independent of the mechanisms generating the presynaptic action potential. Furthermore, blockage not only of sodium but also of voltage-dependent potassium conductance (Katz and Miledi, 1967; Kusano et al., 1967) could lead to prolonged presynaptic release. In addition, presynaptic depolarization beyond a certain level suppressed transmitter release during the period of depolarization. However, transmission did occur at the break of the polarizing pulse (Katz and Miledi, 1967; Kusano et al., 1967). The voltage at which no transmitter was released was called the “suppression potential,” and the synaptic response seen at the break of the pulse was termed the “off” excitatory postsynaptic potential (Katz and Miledi, 1967). In short, at this time it was agreed that the presynaptic action potential itself was not required for transmitter release (Bloedel et al., 1966; Katz and Miledi, 1967; Kusano rr al., 1967). It was recognized that extracellular calcium was required for transmission (Katz and Miledi, 1967; Kusano, 1968) and that possibly an inward calcium current across the preterminal membrane was necessary for transmitter release (Katz and Miledi, 1967). Historically. the next significant steps in our understanding of synaptic transmission at this junction were the demonstration that (1) the presynaptic terminal is capable of generating calcium-dependent action potentials when the sodium and potassium conductances are blocked (Katz and Miledi, 1969a); (2) during synaptic transmission there is a measurable presynaptic calcium entry, as determined by the use of intracellular aequorin (Llinas er al., 1972) and of Arsenazo I11 (Charlton, Smith, and

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Zucker, 1982). An example of the aequorin light emission is illustrated in Fig. 1C for a prolonged depolarizing pulse to the presynaptic terminal following intracellular aequorin injection at the current injection point (right electrode). The aequorin light is restricted to the area which corresponds to the release site, although aequorin was shown to have diffused into the “palm” of the second-order giant axon (Llinas, Sugimori, and Bower, 1983, and unpublished observations). This light emission was observed using an I.T.T. T-4144 double microchannel plate with an overall light gain of 5 X lo5;(3) the suppression potential is accompanied by a complete abolition of the inward calcium current, as determined by the disappearance of aequorin light response (Llinas and Nicholson, 1975); and finally, (4) transmission may be triggered by direct injection of calcium into the presynaptic terminal (Miledi, 1973; Charlton, Smith, and‘ Zucker, 1982; Llinas, Sugimori, and Bower, 1983). These findings indicated that the presynaptic fiber has a large voltage-dependent calcium conductance such that an inward flow of calcium follows depolarization of the presynaptic terminal, and that the increased level of intracellular calcium triggers transmitter release. Taken together, the findings regarding synaptic transmission by direct presynaptic depolarization prompted us to apply the well-known voltageclamp technique (Cole, 1949) to the presynaptic terminal in order to attempt to measure the calcium current allegedly responsible for transmitter release. However, before reviewing this latter set of results and the model of synaptic transmission derived therefrom, the properties of the postsynaptic response will be considered.

V. ELECTROPHYSIOLOGICALPROPERTIES OF THE POSTSYNAPTIC POTENTIAL

Another aspect of transmission at the squid junction concerns the ionic conductances generating the excitatory postsynaptic potential (EPSP). A. Quanta1 Nature

Synaptic potentials are generated at the giant synapse by the spike activation of the presynaptic fiber. This synaptic potential is brought about by a quanta1 release of synaptic transmitter from the presynaptic terminal due to a mechanism similar to that operant at other synapses, in particular the neuromuscular junction (cf. .Katz, 1969). Miniature potentials (with an approximate amplitude of 5-10 p V ) have been described at the stellate junction by Miledi (1967) and by Mann and Joyner (1978). For

526

RODOLFO R. LLINAS

a postsynaptic component of 50 mV, a quantum content of lo4 has been estimated (Miledi, 1967). The time course of the EPSP under normal conditions is dependent on its amplitude due to the direct activation of gK by the postsynaptic potential (Westerfield and Joyner, 1982). This was demonstrated by voltage clamping the synaptic potential at rest membrane potential. Under such conditions the synaptic current does not show the undershoot that follows the normal EPSP due to delayed rectification. Similar results were obtained following intra-axonal injection of tetraethyl ammonium. B. Nature of the Transmitter Substance

The identity of the excitatory transmitter in this synapse is not actually known. The most likely candidate, glutamic acid, while depolarizing the postfiber following iontophoretic injection, was rejected as a possibility by Miledi (1969), who determined that the synaptic potential generated by glutamic acid does not have the same reversal potential as that of the naturally evoked EPSP. However, as will be discussed below, for geometric reasons it may be quite difficult to mimic the exact timing and distribution of the natural transmitter release with a point source iontophoretic injection, and thus this point may still be open to further research. C. Equilibrium Potential

The equilibrium potential for this excitatory response was initially studied by Takeuchi and Takeuchi (1962) and Miledi (1969). These investigators concluded that the ionic basis for this potential was similar to that underlying the endplate potential at the neuromuscular junction. The EPSP was proposed to be generated by a sodium and potassium conductance change, the ratio of these two conductances being close to unity. On the other hand, Gage and Moore (1969) suggested that the EPSP was sufficiently close to EN^ for it to be generated by a sodium permeability change exclusively. This controversy has stemmed from difficulties in the measurement of reversal potentials (cf. Werman, 1972), especially since this junction is distributed over a considerable fraction of a length constant and demonstrates geometric irregularities in the postsynaptic element (Young, 1973). To achieve a preparation in which the postsynaptic terminal region of the fiber can be maintained at a close to isopotential voltage, without the use of an axial wire, a double intracellular oil injection method was developed

THE SQUID GIANT SYNAPSE

527

(Llinas et af.,1974). Mineral oil, tinted with Sudan black for visualization purposes, was pressure-injected into the postsynaptic fiber flanking the site of functional contact between the pre- and postfibers as shown in Fig. 3A. The oil, which expanded in a spherelike manner from the point of impalement, filled the lumen of the axon to appose the internal surface of the plasma membrane. Because of the double oil block, the current injected was limited to this isolated pocket. After achieving the double oil gap, in order to obtain a membrane with nearly linear current voltage properties, tetraethylammonium (TEA) was injected into the presynaptic terminal to block gK and tetrodotoxin (TTX) was locally applied onto the postsynaptic fiber. Under these conditions, the reversal potential for this EPSP could be accurately determined. The results from a current-clamp experiment are shown in Fig. 3B. The reversal of the EPSP occurred at approximately +20 mV, and the relationship between peak synaptic potential and the membrane potential ( V , ) was close to linear. The nonlinearity seen at the beginning of the potential record represents residual gNa, since local application of TTX did not totally cover the surface of the postsynaptic pocket in its entirety. A similar set of records was obtained in five other preparations, as shown in the plot in Fig. 3C. The range of reversals in this set of results was from +I8 to +25 mV with a rather small scatter. Since in all these cases a good double internal gap was obtained, it was concluded that the reversal potential for the EPSP occurs at a positive membrane potential. A significant question at this point was whether the EEpspcorresponded to ENa.The measurements described above argued against this conclusion, since ENais generally +40 mV. However, given the multiple penetration and the time required for the establishment of the experimental paradigm, it was possible that an increase in [Na+Iicould have affected ENa (Moore and Adelman, 1961). To resolve this question, a set of voltage-clamp experiments was undertaken to determine EN^ directly and, at the same time, to compare it with EEpsp. In these experiments the time course and reversal potential of the synaptic current as well as the value of EN^ were determined (Llinas et af., 1974). V , was held at a constant hyperpolarizing level (- 100 mV), and at 10-sec intervals the potential was driven for 12 msec to a pulse potential between - 150 and + 100 mV in 10-mV steps. The stimulus to the presynaptic fiber was timed so that the synaptic conductance change occurred after the small residual f K .As the postsynaptic membrane potential was successively pulsed to higher positive values, the synaptic current (PSC) reversed in sign (Fig. 4A). In Fig. 4B, PSC is plotted as a function of membrane potential. This experiment also demonstrated that the reversal

Giant Axon

-

Oil Injection System

Current

B

J

.

FIG.3. (A) Diagram of experimental technique. The oil drop was injected by pressure from a micrometer syringe (left). The membrane potential level was regulated by a control amplifier (AD170) via feedback. The control amplifier was fed the membrane potential level through the preamplifier (Preamp) by an RC circuit (Rs-Cs). Control pulses were inlroduced from a computer though the resistance (Rcj. The current was injected into the postsynaptic fiber through a second microelectrode and measured by amplifier (AD1 18). The presynaptic terminal was activated by means of an intracellular electrode or with an external bipolar electrode. (B) Reversal of the excitatory postsynaptic potential (EPSP) after application of tetrodotoxin and tetraethylammonium. (C) Plot of the synaptic potential amplitude against membrane potential (MP) level. Reversal potentials in these five examples occurred between the levels of IS and 25 mV positive. (Modified from Llinas ef a / . , 1974.)

c

A

O-

-

L" PSt MAX

-I

-

-21 I

'

mnc

1

I

2

3

t

-21

I

I

I

rnsec

2

1

3

FIG.4. Time course and reversal of synaptic currents. (A) Plot of postsynaptic current (PSC) versus time for a family of step clamps from - 100 mV to 0, 10, ..., 90 mV. (B) Plot of postsynaptic current (PSC) versus the control potential for a family of step clamps shown in A. (C) Comparison of time course for synaptic current at two membrane potential levels, - 100 and +90 mV. Note the lack of change in the time course for synaptic currents evoked at these two levels. (Modified from Llintis et a / . , 1974.)

530

RODOLFO R. LLlNAS

potential for the postsynaptic current is negative to ENa.Indeed, in five experiments ENawas measured at 42 5 6 mV (mean t SD) and EIlc at 19 2 2 mV (mean SD). Thus, current and voltage-clamp experiments on this isolated compartment indicate that EEpspis close to ENa,but that other ions, probably K + , may contribute to the EPSP since EEpspis 20-25 mV more negative (+ 18+25 mV) than ENa(+40-+45 mV).

*

D. Voltage Dependence of the Synaptic Current

In addition to studying EEpsp,another aspect investigated with the above technique was the time course of the PSC (Llinhs et nl., 1974) to determine whether the rate of decline of the falling phase changed with membrane potential. For several levels of membrane potential in each experiment, the PSCs were isolated from the data records, and each PSC was converted to a value between 0 and 1 by scaling each point by the maximum PSC. The natural logarithms of this function for PSCs at membrane potentials of -100 and +90 mV are shown in Fig. 4C. Although the curve for V = +90 mV is noisier than that for V = - 100 mV, it is clear that the time courses are essentially identical. The first 3 msec of the PSC was plotted in each case, and the time constant was determined as the negative reciprocal of the slope of the linear decreasing phase on the logarithmic scale. The average values for the time constant of decay of the PSC at - 100 mV and at +90 m V was 0.85 r 0.21 msec and 0.94 k 0.18 msec, respectively (mean k SD, n = 5). It was concluded, therefore, that, in contrast to the findings at other junctions, the time course of the PSP in the squid giant synapse does not appear to be substantially modified by the potential across the postsynaptic-subsynaptic membrane, as confirmed by Miledi (cf. Cull-Candy and Miledi, 1980). Nevertheless, the voltage sensitivity requires further study, particularly at single-channel level, since the voltage sensitivity of postsynaptic channels may be rather subtle and may vary in magnitude and even in the direction of the electric field sensitivity. VI. VOLTAGE CLAMP OF THE PRESYNAPTIC ELEMENT USING SQUARE PULSES

The next step in studying the properties of the presynaptic elements that govern transmitter release was the development of a presynaptic voltage-clamp paradigm. The methodology employed in this voltageclamp study is described in detail in the original papers (LlinBs, Steinberg

THE SQUID GIANT SYNAPSE

531

and Walton, 1976, 1980, 1981a,b). After double or triple microelectrode impalement of the presynaptic terminal and blockage of g N a and g K with TTX and TEA + 4-AP, respectively, the membrane potential of the terminal was depolarized in square voltage-step fashion using a voltage-clamp circuit. The membrane potential was usually held at -70 mV. (Throughout this article, polarizations are given relative to holding potential; -70 mV.) A. Presynaptic Calcium Current

A rapid presynaptic depolarization (usually 4-10 msec) triggers the onset of a rather slow inward calcium current that does not inactivate and reaches a peak whose amplitude varies with the level of depolarization (Fig. 5A,B). At a depolarization step of 30 mV (from -70 to -40 mV) (Fig. 5A) and small inward current (lowest record) is seen that begins very slowly and shows a close-to-linear increase with time. At the end of the pulse, a fast tail current is observed (see Fig. 5C). These transient currents and the postsynaptic potentials that accompany them are blocked by manganese (I0 mM) or cadmium chloride (1 mM) (LlinBs, 1977; Llinas et al., 1980) and are absent if calcium is removed from the extracellular medium. The postsynaptic response generated by this current is illustrated in the middle trace. Note that the postsynaptic response during the pulse also has a close-to-linear rate of rise, and that the EPSP that follows the break of the voltage-clamp pulse (“off” response) is associated with the inward tail current. The calcium current (Zca) increases in amplitude with increasing levels of depolarization, reaching a maximum in most synapses at 60 mV from rest (-10 mV absolute value), a value quite close to that determined by Baker and co-workers for the giant axon using aequorin (Baker et al., 1971). At this level, the sigmoidal character of the current onset is evident (Fig. 5B, bottom trace), after which the current reaches a plateau, indicating the absence of inactivation, at least for pukes of up to 100 msec. At the end of the pulse, a tail current is again apparent. Accompanying this increase in Zca, a change is seen in the shape, rate of rise, and ampiitude of the EPSP. At 60 mV presynaptic depolarization the EPSP reaches a peak, begins to decline during the pulse, and is followed by a secondary transmitter release, the “off’ response (Fig. SB,arrow in middle trace). As the presynaptic voltage step is increased beyond 60 mV, the level of the “on” presynaptic current decreases in amplitude, while the tail current increases to reach a maximum near I10 mV depolarization (+40 mV

532

RODOLFO R. LLlNAS

A

D 30 mV

B

:;,-

(10%

E

60 mV

F

C 130mV

r

FIG.5. Synaptic transmission during voltage clamp of presynaptic terminal. (A-C) Experimental data. Top trace, presynaptic voltage; middle trace, postsynaptic response; lower trace, calcium current. The S shape of the current onset can be seen at 60 mV depolariLation from a holding potential of -70 mV. Note the fast tail current and the “on” and “off’ excitatory postsynaptic potential (EPSP). (D-F) Numerical solution to mathematical model. Top trace, vesicle depletion; other traces as in A-C. 0, Recorded EPSP. [Ca”],, = 10 mM. (Modified from Llinas er al., 1981b.)

533

THE SQUID GIANT SYNAPSE

absolute value). At the same time, the rate of rise and amplitude of the “on” EPSP decreases, while the “off’ response increases. The “on” current continues to decrease beyond 110 mV until the “suppression potential” (Katz and Miledi, 1967) is reached at close to 130 mV depolarization (+60 mV) (Fig. SC, lowest trace). At this point, no inward current is seen during the voltage pulse, but a large tail current is observed following the pulse break. Similarly, no synaptic transmission is observed for the duration of the pulse, and a sharp, short-latency “off’ postsynaptic response is generated by the tail current. A clear correlation can thus be seen between the characteristics of presynaptic ZCa and the postsynaptic response, the “on” EPSP being related to the Zca during the pulse and the “off’ EPSP to the tail Ica. The relationship between the amplitude of the presynaptic voltage clamp depolarization and the amplitude of the inward steady-state caland tail current is shown in Fig. 6A. As seen in that cium current (Ica) figure, ZCa follows a bell-shaped curve with a rather rapid rise, reaching a peak near 60 mV corresponding to an absolute membrane potential value of - 10 mV. The decrease in current amplitude at higher levels of depolarization is asymptotic, making the estimation of a reversal potential by extrapolation very difficult. The tail currents increase sharply for increasing levels of depolarization up to about 80 mV (+ I0 mV), at which point they begin to level off reaching a maximum near 110 mV (+40 mV).

A Pre I

Post

v

nA

60

600-

400

.

200

1

0 0

mV

40

80

Pre V

120 mV

I

0

40

80

120 mV

Pre V

FIG.6. Dependence of calcium current (A) and EPSP (B) on presynaptic depolarization. (A) 0,Steady state fca;0, tail current. (B) 0, “On” EPSP; X , “off” EPSP. The continuous line in panels A and B show that solution of mathematical model [Ca*+], = 10 mM. In the Pre V axis, 0 corresponds to a holding potential of -70 mV. (Modified from Llinfis et al., 1981a,b.)

534

RODOLFO R. LLlNAS

The above description of calcium currents in the presynaptic terminal, and the more recent description by Charlton, Smith, and Zucker (1982), which basically confirms our own results, assumes that the inward current faithfully reproduces Zca; that is, it assumes that no other current flows simultaneously with ICHto either increase, decrease, or otherwise change its time course. Because a possible contamination of our inward current with an outward K current could not be ruled out in a preterminal, the study of DiPolo et al. (1983) is very relevant to this point. These authors have demonstrated the presence in the squid giant axon of a “late” calcium current measured under conditions where calcium is the only permeant cation. In this study (DiPolo ef al., 1983), sodium and potassium were replaced by large organic ions and the sodium and potassium channels were blocked with TTX and TEA, respectively, to exclude possible flow of calcium via those channels. Since a clear inward current could be obtained with 10 mM [Ca],. these results demonstrate the presence of calcium channels in this axon. In addition this calcium current. which is a thousand times smaller than its presynaptic counterpart, has the same voltage-dependent characteristics described above and shows halfinactivation in about 45 sec. B. Postsynaptic Potentlal as a Function of Presynaptic

Calcium Current

The quantitative relationship between the amplitude of the presynaptic voltage-clamp depolarization and the postsynaptic potential is shown in Fig. 6B, where the filled circles represent the “on” and the crosses the “off” postsynaptic response. The peak amplitude for the postsynaptic potential occurs, as does the peak Ica,with a voltage step of 60 mV from rest level, and the suppression potential with depolarization pulses in the vicinity of 120-130 mV amplitude. The amplitude of the “on” EPSP was measured as the maximum level of postsynaptic membrane potential prior to the termination of the voltape-clamp step. The amplitude of the “off” response was taken as the difference between the “on” EPSP and the maximum postsynaptic potential after the termination of the clamp. As can be seen in Fig. 6, the voltage dependence of the calcium current and of the postsynaptic response is very similar, suggesting a close-tolinear relationship between these two variables. In fact, when Ica is plotted against the “on” EPSP amplitude using double-logarithmic coordinates, a mean slope of 1.36 (20.13, n = 8) is obtained. Similarly, a mean slope of 1.11 (?0.22, n = 4) is obtained when tail current amplitude and “off” EPSP amplitudcs arc plotted using double-logarithmic coordinates

535

THE SQUID GIANT SYNAPSE

(Llinas et al., 1981b). These findings indicate that the S-shaped curve relating presynaptic depolarization to postsynaptic response (Katz and Miledi, 1967; Kusano et ul., 1967; Llinas and Nicholson, 1975) reflects mainly the nonlinear dependence of gcaon voltage and that, at least at the levels of ZCa studied, transmitter release demonstrates a first- to secondorder dependence on gca. C. Synaptic Delay

Other results obtained from these voltage-clamp studies relate to the nature of synaptic delay. As discussed in previous papers (LlinAs et ul., 1976; Llinas, 1977) the temporal relationship between depolarization and transmitter release may be divided into two parts, a and b (Llinas, 1977). The first component of this delay, a , is related to the time required for opening of the calcium channels, and the b portion relates to the time between calcium entry and the onset of the postsynaptic potential. Examples of these two components of synaptic delay can be obtained by comparing the “on” and “off” release as shown in Fig. 7A and B. Here, in record A, a 60-mV presynaptic voltage-clamp step demonstrates a characteristic I-msec delay for the “on” response (composed of components (I and b). The record in Fig. 7B was taken at the suppression potential and illustrates the tail calcium current and the latency (about 200 psec; comprising the b component alone) for the “off” synaptic potential generated by this short current injection. The actual synaptic delay may, in fact, be slightly shorter since the falling phase of the pulse is not instantaneous. These two components were measured directly (Llinas, 1977) by comparing the time of onset of the voltage-clamp pulse with the first sign of

A

B

7

1 mrec

FIG.7. Synaptic delay for the “on” and “off’ EPSP. (A) The “on” response is shown for a 60-mV presynaptic voltage clamp from a -70 mV holding potential; latency 773 psec. (B) The “off” response is seen following a voltage clamp to the suppression potential (I30 mV from hold); 200-psec latency. (Modified from LlinAs cr (I/.. 1981b.)

536

RODOLFO R. LLINAS

inward calcium current (giving a value for a) and then determining the delay between the current onset and the initiation of the postsynaptic potential (giving a value for b). D. A Model for Synaptic Transmission

Based on the above findings, a kinetic model was developed for the relation between presynaptic depolarization and calcium current and between calcium current and transmitter release (Llinas er d.,1976, 1981a,b). The model for the calcium current indicates that the calcium conductance is best simulated assuming fifth-order Michaelis-Menten kinetics and assuming that the fifth-order kinetics correspond to five noncooperative conformational changes occurring prior to the channel opening. These conformational changes, which are voltage- and timedependent, are thought to be governed, as in the case of the Hodgkin and Huxley model (1952), by forward and backward rate constants between the active and inactive states of the subunits in the channels. According to the model, each Ca channel is composed of n subunits, each of which exists in either of two active states, (S) and ( S f ) . The transition between these states is governed by the rate constants, k , and X z . Thus, ki se S'

k2

A gate or channel ( G ) is open when all n subunits are in the S' state. The number of "open" channels [GIis proportional to ( S f ) " ,and the Ca current is given by ([GI) times the current flow per gate per unit time ( j ) . Thus,

P I N Cex~(-80V) ~ - GI 1

+ Kr,,exp(-80V)

where [GI,is the total number of channels, open and closed; k l and kz are the rate constants for the opening and closing, respectively, of a channel subunit; t is the time; V is membrane voltage relative to absolute zero; c, is [C'a''],; C,, i5 I C n ? ' ] , , K ; i5 the equilibrium constant for the transfer of calcium from outside the cell to the calcium site in the channel, at zero membrane potential; PI is a proportionality factor; and 80 is the solution to 2e/kT, where e is the elementary electric charge, k is Boltzmann's constant, and T is the absolute temperature (291°K = I S T ) . The solution of this equation for prcsynaptic clamp voltages of 30, 60,

537

THE SQUID GIANT SYNAPSE

and 130 mV is given in Fig. 5D-F (lowest traces). An expression for the steady-state current Zca is obtained by simplifying the equation to eliminate the time-dependent component. The solution to the equation for fc, for various values of V is given by the continuous lines in Fig. 6A. Several factors must be considered when modeling the remaining steps in synaptic transmission. (1) The high specificity of calcium ions in promoting vesicle fusion and transmitter release (cf. Katz, 1969) strongly suggests that a specific binding entity for calcium, probably a protein (Baker and Schaepfer, 1978), is involved in the process. Such a factor is included in this model and will be referred to as the fusion-promoting factor (fpf). (2) As fusion is initiated, we assume that vesicles are depleted from the immediate vicinity of the plasma membrane (the immediately available store) and that this may become a factor in the transmission process. (3) The time required for the diffusion of the transmitter across the synaptic 8ap is expected to be very short (less than 1 psec for a gap width of 200 A and a diffusion coefficient of cm2sec-I) and thus is not a significant consideration. (4) The opening and closing time constants for the gating of the postsynaptic receptor transmitter complex have been assumed to be proportional to the rate of transmitter release, and it is also assumed that the postsynaptic receptors are far from saturation. ( 5 ) The electrical constants of the postsynaptic terminal membrane are included. In accordance with the above considerations, a three-compartment scheme was developed to model transmitter release and postsynaptic current, each compartment having forward and backward first-order rate constants (Llinas et al., 1981b). These will be considered in turn. The first compartment consists of a calcium protein reaction involving fpf. The behavior of fpf is assumed to be as follows: If a pulse of calcium ions enters the presynaptic digit, fpf binds these ions, forming a complex fpf * Ca2+directly determined by the amplitude of Icaand the time of exposure to the calcium ions. This complex then becomes activated by firstorder kinetics into a species fpf" * Ca?+that facilitates fusion of vesicles with the plasma membrane at a rate proportional to its amount. The species fpf* . CaZ+can then revert to an inactive form, again by first-order kinetics. Similarly, the complex fpf . Ca2' can be inactivated directly without being converted into fpf * Ca2+.The chain of events may be summarized by the following reactions. fpf Ca2+

kb

I

k

-inactive form

fpP Ca2+

k

inactive form

where k, is the rate constant for activation, and kb and ki are the rate constants for the inactivation of fpf . Ca2+and fpr" . Ca2+,respectively.

538

RODOLFO R. LLlNAS

The second compartment relates to a membrane fusion factor-synaptic vesicle reaction leading to transmitter release ( T R ) . Thus, TR = kf(fpf* . Ca2+)Q

where kf is the rate constant for vesicle fusion and Q is the fractional number of vesicles still available from the immediate store. (The rate constants k,, kb , k, , and kf are voltage-dependent.) The third component concerns synaptic transmitter triggering the postsynaptic conductance that generates the EPSP. The computation of EPSP amplitude and time course for three levels of presynaptic voltage clamp are illustrated in Fig. 5 D-F (solid lines). Values obtained experimentally are given by open circles; those calculated using the model for the peak postsynaptic “on” and “off” EPSP are given by the continuous lines. The relationship between Zca and postsynaptic response as determined by the model is given by the solid lines in Fig. 6B. As can be seen in Figs. 5 and 6 , the present model of presynaptic calcium current, transmitter release, and postsynaptic response agrees reasonably well with the voltage-clamp data. The description of the relation between calcium current and postsynaptic potential predicted by the model is of further interest if it can predict the events triggered by an actual presynaptic action potential and if it can give a quantitative description of the time course and magnitude of these predicted events. Thus, as illustrated in Fig. 8. an experimentally obtained action potential was utilized to obtain [via Eqs. 14 and 18 from Llinas ~t al. (1981a) and Figs. 10 and 11 from Llinis et (11. (1981b)l the time courses of (a) the number of open channels, (b) the calcium current, (c) the postsynaptic current, and (d) the postsynaptic potential.

VII. VOLTAGE CLAMP WITH ACTION POTENTIAL WAVEFORM In order to assess directly the validity of the model and to test further the relationship between presynaptic Ca entry and transmitter release, voltage-clamp experiments were designed in which the command pulses simulated presynaptic action potentials (Llinis et al., 1982). A technique similar to that introduced to dctcrmine the ionic conductances during action potentials (Starzak and Starzak, 1978) was utilized. The action potential rccordcd intracellularly from a presynaptic terminal was stored digitally. After pharmacological blockage of gNn and g K , the presynaptic action potential previously stored in the digital buffer was fed into the command amplificr of the voltage-clamp circuit.

539

THE SQUID GIANT SYNAPSE

mV

""F O

L

-

n

a =G

b = Ic

a

c = Ipor,

FIG.8. Theoretical solution for propagating action potential in the presynaptic terminal and related steps in synaptic transmission. Reconstruction of events during synaptic transmission are based on Eqs. 10, 1 1 , 14, and 18 of Llinas et ul. (1981b). a, Calcium channel formation; b, calcium current: c , postsynaptic current; d, postsynaptic potential. (Modified from Llinas ef a / . , 1981b.)

The waveform of the action potential obtained through the voltageclamp circuit was quite close to that originally recorded. This is illustrated in Fig. 9, where the pre- and postsynaptic action potentials recorded prior to g N a and gK blockage are shown in A. The artificial presynaptic action potential using the voltage-clamp circuit is shown in Fig. 9B. A comparison of Fig. 9A and B is illustrated in C and shows that the action potential-like voltage transient is very similar to the original prespike. This artificial spike produces a synaptic potential (Post in B) having the same initial rate of rise and latency as the original postsynaptic response. The Zca generated by the artificial spike is shown as an up-going transient in Fig. 9B. The method for determining Zc, is illustrated in Fig. 9D-F. The artificial spike and the total current (Itotal) are shown in Fig. 9D. Following addition of CdClz to the external bath there is a reduction in the latter part of Jtotal(only the leakage current remains) and the postsynaptic potential is abolished (E). Superposition of the previous two current records (F) shows the difference between the total current (which includes Ica) and the leakage current. The computed difference ( 1 ~at~twice ) the gain is shown as an upward deflection in Fig. 9F.

RODOLFO R. LLINAS A

B

C

FIG.9. Determination of the ,/ generated by a presynaptic action potential. (A) Recordings of the presynaptic (Pre) and the postsynaptic (Post) action potentials prior to blockage of gNuand gK. (B) The presynaptic action potential in A is used as a command voltage, generating a postsynaptic potential (Post). The,/ triggering the release of transmitter is also shown. (C) Superposition of A and B show the similarity between original and simulated presynaptic action potentials and between delay and rate of rise of the postsynaptic responses. Arrow, the original action potential. (D-F) Determination of ,/ (D) From another synapse, the simulated presynaptic spike (Pre), total current (reduced leakage and lCa), and postsynaptic response. (E) A similar experiment after addition of 0.5 mM CdCI2to the bath, showing the disappearance of the postsynaptic response. (F) Superposition of D and E, showing the difference (lower arrow) in inward current between D and E. Upper arrow, time course and amplitude of Ica. Note that the current calibration relates to lCu only. (From Llinfis pf a!., 1982.)

.

A. Relationship between Amplitude of Presynaptlc Spikelike Depolarization and Ica The latency and time course for the Zca evoked by an artificial action potential is shown in detail in Fig. IOA-C for the synapse illustrated in Fig. 9D-b'. Typically this current has a late onset (during the falling phase of the action potential), a fast rate of rise, and a slightly slower rate of fall. This waveform is quite close to that predicted on the basis of our model (LlinAs et al., 1981b). However, while the latency and amplitude of Ica matched quite well, the falling time was slightly faster than predicted, as was the time constant of the tail current for this particular set of results. Indeed, the mean time constant of the tail current following a step voltage-clamp pulse was found to be 331 psec (SD 14 psec, n = 56) rather than that reported previously (540 psec, SD k 83 psec, n = 68). This change reflects an increase in voltage-clamp speed brought about by faster amplifiers. Because the time course for the fall of I,-, and for the tail current was

*

54 1

THE SQUID GIANT SYNAPSE

D

L

A

40 5 mV

II

H

j : jl f f l f

D-

B

20

I

’

4 *. 1001 60

I

J;

50

70 90 Rr-voltage. mV

110

70 90 Re-voltage.mV

110

20

J

0

msec

50

1\

FIG. 10. Amplitude of simulated presynaptic spike, I,,, and postsynaptic potential. (A-C) Time course and amplitude of ICaevoked by a 87.5-mV artificial presynaptic spike (same as that in G). (A) Experimental results showing the truncated (due to high gain) piebynaptic potential, the Ca current (noisy trace), and the postsynaptic response. (B) Result generated by the model, with the presynaptic spike in A superimposed on experimental results. (C) Same records as B at increased gain to demonstrate synaptic delay (onset of presynaptic spike measured by linear extrapolation of rising phase to baseline). (D-G) Relationship of presynaptic spike height to I,, and postpotential. The experimental and modeling results are superpositioned as in B and C. The relationship between presynaptic voltage (pre-voltage) and postsynaptic potential (post-potential) (H), presynaptic voltage and Zca (I), and I,, and postsynaptic potential ( J ) are shown for five experiments. Because the absolute values of these variables differ with the size of the terminal, the results were normalized to give a clearer view of variability among preparations. The results from the theoretical model are shown as a continuous line in plots H-J. (From Llinhs cz al., 1982.)

quite reproducible in our new results (Llinas et al., 1982), we modified the voltage dependence of the forward and backward rate constants, k, and k 2 , in our model (LlinBs et al., 1981a), to achieve a better fit with the new data. The expressions for k, and k2 are kl

=

ky exp(ez,V/kT)

k2 = k: exp(ez2VIkT)

where kp and k; are independent of membrane potential; e is the elemen-

542

RODOLFO R. LLlNAS

tary electric charge; k is Boltzmann’s constant; T is the temperature; and zI and z2 designate the number of charges that move across the membrane by the transformation of S and S’ into the activated transition state. These values reflect the dipole moment changes (normal to the membrane) between open and transition state ( z , )and closed and transition state (z2). (For a full explanation, see LlinAs et al., 1981a, 1982.) The modifications consisted in shifting the orientation of these dipoles such that in the closed state they sense 6% less and in the open state 6% more of the electric energy field than previously. This brought the modeled tail ZCa (solid line in Fig. IOB and C) close to the currents obtained in most experiments without substantially altering the onset or amplitude of the modeled “on” lea. [Modification of the value of n (cf. Llinfis et al., 1981a) failed to achieve the small correction in the tail-current time constant.] The relation between the height and duration of the action potential and the presynaptic current (and postsynaptic response) could be determined directly by modifying the artificial action potentials. In Fig. IOD-G the Icaand the postsynaptic response elicited by a I-msec artificial spike are illustrated at four amplitudes covering the steepest portion of the depolarization-release curve (40-90 mV). The peak Zca amplitudes are given for each trace. An 80-mV spike elicits a mean lc,of 309 nA (k 77; n = 13). This variability in ICaamplitude among synapses is closely related to the size of the pre-terminal. The records in D to G represent a superposition of experimental and model results (as in Fig. IOB and C). Note that the amplitude, time course, and duration of Ic, for each record is very close to that of the model. (To facilitate comparison, the current amplitude was normalized at 75 mV depolarization and expressed as a percentage of the highest value.) A similar agreement may be seen for the postsynaptic potentials following the parameters of the original model. The relationship between the amplitude of the artificial action potential and Icacurrent (measured as described above) for five experiments is illustrated in Fig. 101. This plot shows a typical S shape that approaches a plateau near 110 mV the minimum action potential required to generate a Ca current (Fig. 101) or a synaptic potential (Fig. 10H) being, for this synapse, 45 mV from holding. The S-shaped relationship is also very similar to that predicted by the model (solid lines). B. Relationship between Ca Current, Transmitter Release, and EPSP Latency for an Action Potentlal

Using the above technique, the relationship between ZCa and transmitter release was studied (Llinas et al., 1982). The EPSP amplitude expressed as percentage of the value for a 75-mV spike is plotted as a function of Zca

543

THE SQUID GIANT SYNAPSE

for five synapses in Fig. IOJ. The relationship between total Ca charge and EPSP amplitude is similar to that shown in Fig. lOJ, having an average slope of 1.47 (range 1.01 to 2.08, n = 13). The latency and amplitude for the postsynaptic response following TTX was quite close to that obtained prior to pharmacological manipulation. This is significant since, even when [Ca2+]owas increased to 100 mM (in the absence of [Mg2+],), addition of TTX did not increase synaptic delay, indicating that the Ca entering the terminal through the sodium channels (Baker et al., 1971) does not contribute to transmitter release. The latency between the onset of Ca entry and transmitter release was found to be 375 65 psec ( n = 5 ) . This latency is, as expected from our model (Llinas et al., 1981a, i.e., assuming first-order kinetics for the Ca fusion promoting factor (fpf) activation), slightly longer than that of the experimentally observed 200 @seefor the “off potential generated by the tail Zca. The slightly longer delay is also present in the results from the model (Fig. IOC). A best fit for the modeled EPSP was obtained where the minimum latency observed for the “off” EPSP (approximately 200 psec) was added as a constant to the model, as indicated in previous publications (Llinas et al., 1981b; Meves and Pichon, 1977). In summary, the results of these experiments gave us the first measurements of the Ca currents generated by an action potential in the presynaptic terminal of a chemical synapse. Furthermore, these results indicate that artificial action potentials are as effective in releasing transmitter as the original action potential itself. Also, as concluded from modeling based on previous experimental results, the action potential releases transmitter via a Ca current that begins during the falling phase of the action potential; that is to say, normal ZCa is largely a tail current (Llinas et al., 1976, 1981b). Indeed, this current is larger than the “on” current but slightly smaller than the tail Zca generated by a square pulse of the same amplitude. The advantages of a tail Ca current in evoking synaptic transmission are clear. First, if the increase in gcacomes late during the action potential, the onset of this current will be well matched to the increased driving force for Ca as the membrane potential returns to resting level. Thus, at the peak of the conductance the driving force for Ca will be maximal. Because I,, is both voltage and time dependent, the amount of transmitter released will be related not only to the amplitude of the action potential, but also to its duration. The larger the amplitude (up to 110 mV) and duration (up to the asymptote of the Zca “on” kinetics) and the faster the rate of fall of the action potential, the larger the amount of Zca. In short, because the amplitude of the tail current depends on the magnitude and duration of the transmembrane electric field, especially in the normal range of action potential amplitude and duration, modulation of these parameters will regulate de facto the amount of transmitter released.

*

”

544

RODOLFO R. LLINAS

Since this technique allows direct control of the time and voltage parameters of the presynaptic depolarization, this approach is deemed useful in studying chemical synaptic transmission. In particular, transmission changes following specific modifications in the waveform and amplitude of this depolarization (Miledi and Slater, 1966; Takeuchi and Takeuchi, 1962; Shapiro et al., 1980) or modifications of the ionic and pharmacological character of the external milieu may be investigated independently of the possible action of such variables on gNaand g K . In addition, the effect of modulator of synaptic release may also be addressed directly (Dunlap and Fishbach, 1978; Kupfermann, 1979). On the other hand, it is also quite clear that our present understanding of the depolarization-release coupling is very far from complete. That voltage-dcpcndent Ca entry is the trigger for transmitter release is firmly established; however, the biophysical, biochemical, and cell biological steps that are actually triggered by such Ca influx remain some of the most fascinating unknowns in this as well as other synaptic junctions. Much remains to be done. ACKNOWLEDGMENTS Research was supported by USPHS Research Grant NS14014 from the National Institute of Neurological and Communicative Disorders and Stroke. REFERENCES Baker, P. F., and Schaepfer, W. W. (1978). Uptake and binding of calcium by axoplasm isolated from giant axons of Loligo and Myxicola. J . Physiol. (London) 276, 103-125. Baker, P. F., Hodgkin, A. L., and Ridgway, E. B. (1971). Depolarization and calcium entry in squid giant axon. J . Physiol. (London) 218, 709-755. Bloedel, J. R., Gage, P. W., LlinCis, R., and Quastel, D. M. J. (1966). Transmitter release at the squid giant synapse in the presence of tetrodotoxin. Nature (London) 212,49-50. Bryant, S . H. (1958). Transmission in squid giant synapses. The importance of oxygen supply and the effects of drugs. J . C e n . Physiol. 41, 473-484. Bullock, T. H. (1946). A preparation for the physiological study of the unit synapse. Nature {London) 158, 555-556. Bullock, T. H. (1948). Properties of a single synapsc in the stellate ganglion of squid. J. Neitrophysiol. 11, 343-364. Bullock, T. H., and Hagiwara, S. (1957). Intracellular recording from the giant synapse of the squid. J. Gen. Physiol. 40, 565-577. Castejon, 0. J.. imd Villegas, G . M. (1964). Fine structure of the synaptic contacts in the stellate ganglion of the squid. J . Ultrasrruct. Res. 10, 585-598. Charlton, M. P., Smith, S . J., and Zucker, R. S.(1982). Role ofpresynaptic calcium ions and channels in synaptic facilitation and depression at the squid giant synapse. J . Physiol. (London) 323, 173-193. Cole, K. S . (1949). Dynamic electrical characteristics of the squid axon membrane. Arch. Sci. Physiol. 3 , 253-258. Cull-Candy, S. G., and Miledi, R. (1980). Factors affecting the channel kinetics of glutamate receptors in locust muscle fibres. In “Receptors for Neurotransmitters, Hormones and

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Pheromones in Insects” (D. B. Satelle. L. M. Hall, and J . G . Hildebrand, eds.). pp. 161-173. Elsevier, Amsterdam. DiPolo, R., Caputo, C., and Bezanilla, F. (1983). Electrical recording of a voltage-dependent Ca conductance in the squid axon. Proc,. Nail. Acad. Sci. U . S . A . 80, 1743-1745. Dunlap. K . , and Fischbach, G. D. (1978). Neurotransmitters decrease the calcium component of sensory neuron action potentials. Nuiure (London) 276, 837438. Gage, P. W., and Moore, J . W. (1969). Synaptic currents at the squid giant synapse. Science 166, 510-526. Hagiwara, S . , and Tdsaki, 1. (1958). A study on the mechanism of impulse transmission across the giant synapse of the squid. J . Physiol. (London) 143, 114-137. Hama, K . (1962). Some observations on the fine structure of the giant synapse in the stellate ganglion of the squid. Doryteuphis bleekeri. Z . Zellforsch. 56, 437-444. Hodgkin, A. L., and Huxley, A. F. (1952). The components of membrane conductance in the giant axon of Loligo. J . Physiol. (London) 116,473-496. Hodgkin, A. L., Huxley, A . F., and Katz, B. (1952). Measurement of current-voltage relations in the membrane of the giant axon of Loligo. J . Physiol. (London) 116, 424-448. Katz, B. (1969). “The Release of Neural Transmitter Substances” (Sherrington Lecture X). Thomas, Springfield, Illinois. Katz, B., and Miledi, R. (1966). Input-output relation of a single synapse. Nature (London) 212, 1242-124s. Katz, B . , and Miledi, R. (1967). A study of synaptic transmission in the absence of nerve impulses. J . Physiol. (London) 192, 407-436. Katz, B., and Miledi, R. (1969a). Tetrodotoxin-resistant electric activity in presynaptic terminals. J . Physiol. (London) 203, 459-487. Kupfermann, I. (1979). Modulatory action of neurotransmitters. Annu. Rev. Neurosci. 2, 447-46s. Kusano, K. (1968). Further study of the relationship between pre- and postsynaptic potentials in the squid giant synapse. J . Gen. Physiol. 52, 326-345. Kusano, K., Livengood, D. R.. and Werman, R. (1967). Correlation of transmitter release with membrane properties of the presynaptic fiber of the squid giant synapse. J . G e n . Physiol. 50, 2579-2601. Llinis, R . (1977). Calcium and transmitter release in squid synapse. Soc. Neurosci. Symp. 2, 139-1 60. Llinas, R.,and Nicholson, C. (1975). Calcium role in depolarization-secretioncoupling: An aequorin study in squid giant synapse. Proc. Natl. Acad. Sci. U . S . A . 7 , 187-190. Llinris, R . , Blinks, J. R., and Nicholson, C. (1972). Calcium transient in presynaptic terminal of squid giant synapse: Detection with aequorin. Science 176, 1127-1129. Llinas, R., Joyner, R. W., and Nicholson, C. (1974). Equilibrium potential for the postsynaptic response in the squid giant synapse. J . Gen. Physiol. 64, 519-535. Llinas, R., Steinberg, I. Z., and Walton, K. (1976). Presynaptic calcium currents and their relation to synaptic transmission. Voltage clamp study in squid giant synapse and theoretical model for the calcium gate. Proc. Natl. Acad. Sci. U . S . A .73, 2918-2922. LlinAs, R., Steinberg, I. Z., and Walton, K. (1980).Transmission in the squid giant synapse: A model based on voltage clamp studies. J . Physiol. (Paris) 76, 413-418. Llinas, R., Steinberg, I. Z., and Walton, K . (1981a). Presynaptic calcium currents in squid giant synapse. Biophys. J . 33, 289-321. LlinBs, R., Steinberg, 1. Z., and Walton, K. (1981b). Relationship between presynaptic calcium current and postsynaptic potential in squid giant synapse. Biophys. J . 33, 323-3s I.

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Lliniis, R., Sugimori, M., and Simon, S. M. (1982). Transmission by presynaptic spike-like depolarization in the squid giant synapse. Proc. N u t / . Acud. Sci. U . S . A . 79, 24152419. Llinas, R., Sugimori, M., and Bower, J. M. (1983). Visualization of depolarization-evoked presynaptic calcium entry and voltage dependence of transmitter release in the squid giant synapse. Biol. Bull. 165, 529. Mann, D. M., and Joyner, R. W. (1978). Miniature synaptic potentials at the squid giant synapse. J. Newobio!. 9, 329-335. Martin, R.. and Miledi, K. (1975). Presynaptic complex in giant synapse of squid. J. Neurucyrol. 4, 121-129. Meves, H., and Pichon, Y.(1977). The effect of internal and external 4-aminopyridine on the potassium currents in intracellularly perfused squid giant axons. J. Physiol. (London) 268, 511-532. Miledi, R. (1967). Spontaneous synaptic potentials and quanta1 release of transmitter in the stellate ganglion of the squid. J. Physiol. (LondonJ192, 379-406. Miledi, R. (1969). Transmitter action in the giant synapse of the squid. Nature (London) 223, 1284- 1286. Miledi. R. (1973). Transmitter release induced by injection of calcium ions into nerve terminals. Proc. R. Soc. London Ser. B 183,421-425. Miledi, R., and Slater, C. R. (1966). The action of calcium on neuronal synapses in the squid. J. Physiol. (London) 184, 473-498. Moore, J. W., and Adelman, W. J. (1961). Electronic measurement of the intracellular concentration and net flux of sodium in the squid axon. J. Gen. Pkysiol. 45, 77-92. Pumplin. D. W., and Reese, T. S. (1978). Membrane ultrastructure of the giant synapse of the squid Loligo peuleii. Neuroscience 3, 685-696. Pumplin, D. W., Reese, T. S., and Lliniis, R. (1981). Are the presynaptic membrane particles the calcium channels? Pruc. N u t / . Acud. Sci. U.S.A. 78, 7210-7213. Robertson, J. D. (1963). The occurrence o f a subunit pattern in the unit membranes of club endings in Mauthner cell synapses in goldfish brains. J. Cell B i d . 19, 201 -221. Shapiro, E., Castellucci, V. F., and Kandel, E. R. (1980). Presynaptic membrane potential affects transmitter release in an identified neuron in A p l y . ~ by i ~ modulating the Ca” and K’ currents. Proc. Nutl. Acud. Sci. U.S.A.77, 629-633. Slarzak, M. E., and Starzak, R. J. (1978). An action potential clamp to probe the cffcctivenew of space clamp in axons. lEEE Truns. Eiomed. Eng. 25, 201-204. Takeuchi, A , , and Takeuchi, N. (1962). Electrical changes in pre- and postsynaptic axons of the giant synapse of Luligo. J. Gen. Physiol. 45, 1181-1 193. Werman, R. (1972). CNS cellular level: Membranes. Annu. Keo. Physiol. 34, 337. Westerfield, M., and Joyner, K. W. (1982). Postsynaptic factors controlling the shape of potentials at the squid giant synapse. Neuroscience 7 , 1367-1375. Williams, L . W. (1910). “The Anatomy of the Common Squid, Loligo peulii.” Brill, Leiden. Young, J. Z. (1936). Structure of nerve fibers and synapses in some invertebrates. Cold S p r i n ~Harbor Symp. Quiint. B i d . 4, 1-6. Young, J. Z. (1938). ‘The functioning of the giant nerve fibres of the squid. J. Exp. Biol. 15, 170- 185. Young, J . Z. (1939). Fused neurons and synaptic contacts in the giant nerve fibres of cephalopods. Philos. Truns. R. Soc. London S e r . B 229, 465-503. Young, J. Z.(1973). The giant fibre synapse of Loligo. Emin Rrs. 57, 457-460.

CURRENT TOPICS IN MEMBRANES AND TRANSPORT, VOLUME 22

Axon-Schwan n Cell Relationship JORGE VILLEGAS' Centro de Biofsica y Bioquirnica In.stituto Venezoluno de I n v ~ , . s t i ~ ( i ~ ~ i CicntiJic,us one.\ ( I V I C ) rind Insrituto intemucionul de E.strtdio.s Avuzudos (IDEA 1 Caracus, Vunezuela

1.

Introduction. .

.........

D. The Cell of Schwann. 111. Axon-Schwann Cell Signal

550

. . . . . . . . . 554

..........

554

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

VI. Concluding Remarks . . . . . . . . . . . . . .

I. INTRODUCTION

Most nerve cells in the central and the peripheral nervous systems are surrounded by satellite cells. This close proximity of neuronal and glial elements in the tissue suggests the existence of a similar intimate functional relationships between them. Such a possibility has been the subject of intensive research and debate (Glees, 1955; Schmitt and Geschwind, i Present address: Centro de Neurociencias, Instituto Internacional de Estudios Avanzados, Apartado 17606, Parque Central, Caracas, 1015-A, Venezuela.

547 Copyright 0 1984 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-153322-0

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1957; Causey, 1960; Kuffler and Nicholls, 1966; Lasansky, 1971; Watson, 1974; Somjen, 1975; Orkand. 1977; Varon and Somjcn, 1979; Treherne, 1982). Nevertheless, the physiological function of satellite cells is largely unknown except for their role in the formation of myelin, which helps speed conduction of nerve impulses. Thus, it remains to discover what role, if any, the Schwann cell plays in the normal functioning of unmyelinated nerve fibers. The fast-conducting unmyelinated fibers of the squid offer exceptional opportunities for the study of the relationships between the axon and its satellite Schwann cells. As pointed out by G. M. Villegas and R. Villegas in this volume, the large size of the axon dictates a special axon-glia relationship, which consists in a multiplicity of Schwann cells necessary to cover up the perimeter of the axon. This structural anangement has facilitated the detection of effects of axonal excitation on the Schwann cell membrane potential that otherwise would have escaped detection.

11.

AXON-SCHWANN CELL INTERFACE

The boundary region between the giant axon and the Schwann cell layer is characterized by the presence of a narrow intercellular cleft delimited by the axolemma (axon plasma membrane) and the Schwann cell plasma membrane (G. M. Villegas and Villegas, 1960). The existence of this periaxonal space was proposed by Frankenhaeuser and Hodgkin ( I 965) after their demonstration of a transient accumulation of potassium ions in the vicinity of the surface-excitable membrane during conduction of nerve impulses. Thus, by reinterpreting the electron micrographs of the surface of the giant axon of Loligo previously published by Geren and Schmitt (1954), they suggested that the sharply defined osmiophilic layer or membrane observed at the surface of the axoplasm represented the excitable membrane, and that the space between this layer and a similar one seen at the inner border of the Schwann cell was the region in which potassium ions accumulate. Furthermore, since thcir experiments indicated that potassium ions escape from that limited space through a lowimpedance unselective layer, they suggest the possibility that ions pass through the space between the osmiophilic double-edged intracytoplasmic membranes observed by Geren and Schmitt (Frankenhaeuser and Hodgkin, 1956). This hypothesis was criticized by Schmitt and Geschwind (l957), in the light of evidence advanced earlier on the similarities between the intracytoplasmic system of double-edged membranes and myelin, considering that the light areas between the membranes were

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filled with a dense material rather than with an aqueous solution. Thus, in discussing the site of interaction between neuronal and extraneuronal phases, Schmitt and Geschwind (1957) mentioned the need to find a preparation in which a microelectrode could be inserted into the Schwann cell to measure the potentials across each of the effective boundaries. Such an experiment was carried out by R . Villegas et al. (1963) in the giant nerve fiber of the tropical squid Sepioteuthis sepioidea.

A. Inner Boundary The axon-excitable membrane is well characterized in other articles of this volume concerned with the different processes that take place at this membrane. However, it seems relevant for understanding the relationships between the axon and its satellite cells to call the attention of the reader to the presence in the axolemma of specialized sites that form part of structural complexes, where the axolemma and the plasma membrane of the Schwann cell come into close apposition. Such complexes have been related to specialized sites for active transport, as well as to sites for functional interaction between the axon and its satellite cell (G. M. Villegas and Villegas, 1976; see also G . M. Villegas and R. Villegas in this volume).

B. Outer Boundary The outer limit of the intercellular space is formed by a single row of glial cells arranged in a mosaic disposition around the perimeter of the axon and along the entire nerve fiber ( G . M. Villegas and Villegas, 1'968). The surface of the Schwann cells is highly irregular with foldings and invaginations. Furthermore, in images of the Schwann layer obtained after freeze-fracturing of giant nerve fibers of S. sepioidea there appeared a mixed population of particles of different sizes in the cytoplasmic face (P face) of the plasma membrane, and zones where exocytotic or endocytotic craters are abundant (see G . M. Villegas and R. Villegas, in this volume). The pleats and infoldings of the plasma membrane at the inner surface of the Schwann cell are also accompanied by openings into the periaxonal space of the intercellular clefts, separating adjacent Schwann cells, which in L . pealei fibers have been correlated with a characteristic invagination of the Schwann cell into the axon (Adelman et al., 1977). It has been possible to see in S. sepioidea Schwann cells membranous profiles, interconnected in an orderly fashion and arranged like a tubular lattice opening also into the clefts (a.M . Villegas, personal communica-

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tion). Thus, unlike the continuous inncr limit formed by the axolemrna, the outer limit of the pcriaxonal space appears to be interrupted by relativcly large (10-nm) slitlike intercellular pathways. C. The Perlaxonal Space

The axolemma-Schwann cell space represents the actual extracellular environment of the axon. This space measures about 5-10 nm across, and its volume is so exceedingly small that potassium ions leaving the axon during conduction of nerve impulses are able to increase the concentration of this ion in the immediate vicinity of the axolemma. Thus, the propagation of a single impulse, which is known to release about 4 x mol of potassium per square centimeter of membrane surface (Shanes, 1954; Keynes and Lewis, 1951), produces an apparent rise of about 1.6 mM in the concentration of this ion in the periaxonal space, whereas much larger concentrations are built up during the conduction of nerveimpulse trains and following axonal depolarizations under voltage-clamp conditions (Frankenhaeuser and Hodgkin, 1956; Adelman et al., 1973). On the other hand, after activity, the excess potassium accumulated in the periaxonal space disappears along an exponential curve (time constant of 30- 100 msec) by diffusing away through low-resistance unselective pathways and by reentering the cells (Frankenhaeuser and Hodgkin, 1956). The resistance of the periaxonal space and extracellular pathways to potassium diffusion in Lofig0 nerve fibcrs is about 4-7 a cmz. This value is not different from that found for the ratio between the fraction of the total surface area available for diffusion (A) and the length of the diffusion path (Ax) followed by water molecules through the axonal sheaths, in the giant fiber of Doryteuthis plei species (R. Villegas and Villegas, 1960; R. Villegas et ul., 1962). The demonstration of the permeability of the periaxonal space to electrodense markers such as thorium dioxide (G. M. Villegas and Villegas, 1964), together with the finding of a much higher resistance (lo3 bl cm2) to current flow across the plasma membrane of the Schwann cell (J. Villegas, 1972), supports the conclusion that the intercellular clefts are the main pathways for the rapid diffusion of ions between the external solution and the axonal surface. D. The Cell of Schwann

The term Schwann cell is used to designate any cell that enfolds a nerve fiber within its cytoplasm, whether this cell is in the somatic or the visceral part of the peripheral nervous system (Causey, 1960). The most

AXON-SCHWANN CELL RELATIONSHIP

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remarkable structural feature of the cell of Schwann in the giant nerve fiber of the squid is its high surface-to-volume ratio. This has been interpreted as a sign of high energy expenditure by the cell (Geren and Schmitt, 1954; Schmitt and Geschwind, 1957; Coelho etal., 1960). The surface of the cell is largely increased by its many foldings and invaginations, especially at the lateral surfaces, which are deeply interdigitated with the neighboring cells (G. M. Villegas and Villegas, 1963, 1968; Adelman et al., 1977). These deep interdigitations, pleats, and infoldings of the lateral surfaces of the cell determine the tortuous passage of the extracellular clefts formed by them. Thus, although the thickness of the Schwann cell varies among different species from 0.9 to 6.0 pm, the extracellular pathways are about one order of magnitude longer than that (G. M. Villegas and Villegas, 1960; Adelman et al., 1977). The study of the physiological properties of the satellite cells of the giant axon has been greatly aided by two preparations in which the contribution of the Schwann cell to the parameters being measured can be easily distinguished from that of the axon. One is the axoplasm-free nerve fiber sheath preparation obtained by Coelho et al. (1960) from Loligo nerve fibers by slitting the giant axon open longitudinally. In this preparation the Schwann cells continue to respire and maintain a high intracellular potassium concentration for many hours. The other is the giant nerve fiber of the tropical squid S . sepioidea, in which the Schwann cells are 1-5 pm thick and can be impaled with glass capillary microelectrodes and their membrane properties studied (R. Villegas et al., 1963; J. Villegas et al., 1968). Thus, the combination of the slitting technique with the use of S . sepioidea giant nerve fibers has permitted some of the physiological properties of the Schwann cell to be explored. 1. MEMBRANE POTENTIAL

In the intact nerve fiber under resting conditions, the Schwann cell maintains an electrical potential difference of about 40 mV, inside negative, across its plasma membrane. The Schwann cell potential is less negative than the resting membrane potential of the neighboring axon (-60 mV), but higher than the electrical potentials of the endoneurial cells (R. Villegas et al., 1963). The values found for the membrane potential in different Schwann cells impaled along the same nerve fiber vary between -33 and -46 mV. However, the average level of -40 mV has been found to represent a rather constant reference potential level, present both in the intact nerve fibers and in the slit fibers under resting conditions (J. Villegas, 1972). Similar Schwann cell resting-membrane potential values are recorded in experiments carried out at sea level and at 1600 m above it.

552 2.

JORGE VILLEGAS

IONIC PERMEABILITY

The membrane potential of the Schwann cell is determined by t h e ionic concentration gradients and the permeability of its surface membrane to potassium and, to a lesser extent, other ions. Thus at high external potassium concentration the Schwann cell electrical potential is related to the logarithm of [K+],, the external potassium concentration, in a quasilinear fashion (Fig. 1). However, the linear portion of the curve is characterized by a slope of 45 mV per 10-fold change in [K+],. This slope is smaller than that of 52 m V per 10-fold change in concentration of the equivalent regions of the curve obtained with axons of the same nerve fibers (J. Villegas ~t n f . , 1968).Thus, under resting conditions thc behav-

J

I

I

I

0.1

1.0

10

100

lK+lo

mM

Fic. 1. Relationship between Schwann cell membrane potential and [K+l,, the external potassium concentrution. Values are mean * SEM of the potentials measiired in the abbence (0) and in the presence ( 0 )of 10 M carbamoylcholine in the external scawater medium. (After J . Villeges, 1974.)

AXON-SCHWANN CELL RELATIONSHIP

553

ior of the Schwann cell membrane in the squid nerve fiber departs from that found by Kuffler et al. (1966) for the glial membrane, which closely follows the theoretical slope of 59 mV, even at concentrations lower than the physiological range (Kuffler et al., 1966; Kuffler and Nicholls, 1966). This apparent discrepancy, together with the fact that the measured resting potentials in the squid Schwann cells are lower than those found in the above cited glial cells, has been interpreted either as (I) the result of leakage current during electrode penetration of the Schwann cells (Kuffler and Nicholls, 1976), or (2) the result of the contribution of different ionic species and transport mechanisms to the membrane potential of the Schwann cell ( J . Villegas et id., 1968; J . Villegas, 1968). In the unstimulated giant nerve fiber of S. sepioidea, the permeability of the Schwann cell membrane to sodium ions appears to be low. Thus, large decreases of the external sodium concentration produce only small variations in the membrane potential of the cell (J. Villegas et al., 1968). The existence of sodium pathways in the Schwann cell membrane has been revealed in S. sepioidea nerve fibers treated under resting conditions with veratrine and grayanotoxin (J. Villegas et al., 1976). The effects of these compounds on the membrane potential of the Schwann cell were similar to those they produce on the resting membrane potential of the giant axon. Thus, the depolarizations induced by veratrine and grayanotoxin in the Schwann cells were reversed by decreasing the external sodium concentration or by external application of tetrodotoxin (J. Villegas et al., 1976). However, the Schwann cell sodium pathways are different from those of the axolemma in that they are not voltage dependent. The electrical, behavior of the Schwann cell plasma membrane to the injection of depolarizing and hyperpolarizing currents into the cell is ohmic, and no regenerative responses have been observed in the Schwann cell even with displacements of 30 mV in membrane potential produced by depolarizing current pulses (J. Villegas, 1972). The sensitivity of the Schwann cell resting membrane potential to other ionic species normally present in the external medium is also small. Large decreases in the external concentrations of chloride, calcium, and magnesium produce only small variations in the membrane potential of the cell; whereas a 10-fold increase in the external concentration of calcium is accompanied by a 10-mV hyperpolarization of the Schwann cell (J. Villegas et al., 1968). It has been found that exposure of the cells to high calcium concentrations is accompanied by a significant decrease in the Schwann cell intracellular sodium concentration (J. Villegas and D. W. Herzfeld, unpublished results). These experimental findings confirm that, in addition to potassium, the Schwann cell plasma membrane is also permeable to other ionic species.

554

JORGE VILLEGAS

The experimental results illustrated in Fig. 1 also provide a good estimate of the internal concentration of freely diffusible potassium. The Nernst equation, which describes the behavior of an ideal potassium electrode, predicts that, when the internal and the external potassium concentrations are the same, the membrane potential is zero. Thus, by extrapolating to zero membrane potential, a value of 210 mM can be obtained ( J . Villegas et al., 1968), which is in good agreement with that of 220 mM found in the same nerve fibers by direct flame photometric measurements of total Schwann cell potassium concentrations (J. Villegas et al., 1965, 1968), as well as with the values reported by Coelho et al. (1960) for the periaxonal cells in Loligo giant fibers. From these values it can be estimated that the equilibrium potential for the distribution of this ion across the Schwann cell membrane is of the order of -78 mV as referred to the external solution (J. Villegas, 1968). Thus, under steady-state conditions the intracellular potassium in excess of the membrane potential is maintained constant by an inward movement of this ion through the cell membrane against its electrochemical potential difference. In agreement with this, prolonged immersion of the tissue in seawater solutions containing cardiac glycosides (strophanthoside K, strophanttin G ) produces a significant decrease of the Schwann cell intracellular potassium concentration (J. Villegas et al., 1968). A similar estimate of the equilibrium potential for the distribution of sodium across the cell membrane gave a +9 mV value, as referred to the external seawater medium of the nerve fibers analyzed by flame photometry (J. Villegas, 1968). Thus, although the Schwann cell intracellular sodium concentration found in these nerve fibers is rather high (240-400 mM), in the steady-state sodium should be extruded from the cell against its electrochemical potential difference in order to balance its passive entry (J. Villegas, 1968). On the contrary, chloride appears to be distributed almost in equilibrium across the Schwann cell membrane, since the equilibrium potential for this ion is about -32 mV and the measured membrane potential is -40 mV (J. Villegas, 1968). 111.

AXON-SCHWA"

CELL SIGNALING

A. Electrical Signals

Simultaneous intracellular recordings of the axon and Schwann cell electrical potentials obtained by R. Villegas et al. (1963) in the giant nerve fiber of S. sepioidea revealed that during the propagation of a nerve impulse the action potential is recorded only from the axon. It was also

AXON-SCHWANN CELL RELATIONSHIP

555

found that the Schwann cell resting membrane potential remains unchanged during the injection of depolarizing current into the axon through a third impaling microelectrode (R. Villegas et al., 1963). Similar results were obtained by Kuffler and Potter (1964) in leech ganglia. Also, synchronized conduction of nerve impulses had no direct effect on the neighboring glial membrane in the optic nerve of Necturus (Orkand et al., 1966). The simplest explanation of the failure of action potential currents to flow from one type of cell to the other is their separation by the intercellular clefts filled with low-resistance fluid, which provide pathways for currents to flow out to the nerve surface without passing through the relatively high resistance of satellite cell membranes (G. M. Villegas and Villegas, 1960;R. Villegas and Villegas, 1960; R. Villegas et al., 1962, 1963; Kuffler and Potter, 1964; Nicholls and Kuffler, 1964). However, in the giant nerve fiber of the squid and other marine invertebrates, special structural links between axon and Schwann cell have been detected (G. M. Villegas and Villegas, 1976) (see Section V,B). Such structural links can be disrupted by different procedures without altering the capacity of the axon to conduct action potentials, suggesting that these sites are not involved in the propagation of action potentials, which takes place in the axolemma surface exposed to the extracellular fluid (see G. M. Villegas and R. Villegas in this volume). B. Potassium Signals

Although the ionic currents set up by the propagation of nerve impulses do not cross the glial membranes, by flowing through the extracellular spaces they do appreciably modify the electrochemical potential differences across the plasma membranes of the neuronal and glial cells delineating them (Frankenhaeuser and Hodgkin, 1956; Orkand et al., 1966; Baylor and Nicholls, 1969). Thus, the accumulation of potassium ions in the intercellular clefts produced by potassium leakage from the excited neurons is currently considered to be a nonsynaptic nonspecific neuronglia signal, except that a particular glial cell is influenced preferentially by a discrete population of neurons that are in close proximity to it (Kuffler and Nicholls, 1976). However, in the giant nerve fiber of the squid the signals from the only axon impinge upon a multicellular glial monolayer extended over its whole surface, in close proximity to the axolemma. This fact, together with the high surface-to-volume ratio characteristic of the Schwann cells in these nerve fibers, can amplify and make detectable the effects of more discrete synapsis-like specific neuron-glia signaling mech-

556

JORGE VILLEGAS

anisms that could pass unnoticed or be absent in other nerves with a different geometry. Thus, as illustrated in Fig. 2a, during the propagation of nerve impulses by the repetitively stimulated giant axon, t h e membrane potential of a neighboring Schwann cell undergoes a transient depolarizing change that gives place to a sustained hyperpolarization outlasting the duration of the axonal membrane potential change. Although the initial Schwann cell depolarization can be explained by the accumulation of potassium ions in the intercellular clefts of the nerve fiber (Frankenhaeuser and Hodgkin, 1956; Orkand et al., 19661, this process cannot account for the subsequent hyperpolarization recorded in the cell. Figure 2b shows that in a similar nerve fiber pretreated with an irreversible cholinergic receptor blocker (Section III,C) that serves to suppress the hyperpolarizing phase of the response, the Schwann cell membrane potential closely follows the

125 H r

Stim.

-20

-10

t

'i

K S c h w m n cell mernbrune potential

-60

-LO

b

-

M.P

(mvl -60

-80

'

0

I

I

I

4

8

12

Time (rninl

FIG. 2. Simultaneous intracellular recordings of the Schwann cell and axon electrical potentials in repetitively stimulated nerve fibers. (a) Upper graph: fiber used as control. (b) Lower graph: fiber pretreated with 10 a-bungarotoxin, exposed to different external potassium concentrations. M . P., Membrane potential. (After J . Villegas, 1978b. 1981 .)

AXON-SCHWANN CELL RELATIONSHIP

557

changes in external potassium concentration introduced during the experiment and gives only depolarizing responses of different amplitude following axonal repetitive stimulation. These experimental findings unequivocally show that the Schwann cell resting potential, in addition to being sensitive to changes in potassium concentration occurring in the intercellular clefts of the nerve fiber, also responds to other types of signals being generated during nerve impulse train conduction (J. Villegas, 1972, 1973). C. Synapsic-like Signals

The search for a mechanism of axon-Schwann cell interaction different from potassium release began as a consequence of not being able, in the same nerve fibers, to provoke similar Schwann cell hyperpolarizing responses by transiently increasing the external concentration of this ion (J. Villegas, 1972). In addition, the existence of some form of coupling between Schwann cells and the axon appeared to be favored by the presence of the specialized membrane regions mentioned previously, which had been compared with the postsynaptic densities observed in synapses where a chemical mechanism is responsible for intercellular coupling ( G . M. Villegas and Villegas, 1968). On the other hand, although it had already been found that both axonal conduction and synaptic transmission in the giant nerve fiber of the squid are relatively insensitive to the external application of acetylcholine and other compounds acting on the cholinergic system (Miledi, 1967; Nachmansohn, 1959; Rosenberg, 1971; Webb r t al., 1966), the question remained as to what role, if any, the acetylcholinesterase present in these nerve fibers played in their normal functioning. Thus, the possibility was explored that the long-lasting Schwann cell hyperpolarizations following the conduction of nerve impulse trains by the axon could be the electrical expression of some sort of cholinergic axon-Schwann cell relationship (J. Villegas, 1973). IV. SCHWANN CELL RESPONSES A. Cholinergic Responses

In S. sepioidea giant nerve fibers exposed to the action of different cholinergic compounds, it was found that at low (loT9M ) external concentration d-tubocurarine blocked, and eserine prolonged, the Schwann cell hyperpolarizations following the conduction of nerve impulse trains by the axon (J. Villegas, 1973). It was also found that the external applica-

558

JORGE VILLEGAS

tion of acetylcholine (lo-’ M ) and its stable analog carbamoylcholine M )to the resting nerve fibers mimicked the hyperpolarizing effects of axonal excitation on the Schwann cell membrane potential (J. Villegas, 1973). Since at the concentrations indicated above these cholinergic compounds are known to act in a specific manner on acetylcholine receptors and esterases, these results led to a series of studies aimed at detecting, characterizing, and localizing the different elements of the acetylcholine system present in the giant nerve fiber. 1. ACETYLCHOLINESTERASE ACTIVITY

The physiological findings described above made it intcrcsting to investigate in S. sepioidea giant nerve fibers the exact distribution of the cholinesterase, which had already been found in other squid species (Boell and Nachmansohn, 1940; Brzin et d.,1965). The combination of histochemistry and electron microscopy showed the presence of acetylcholinesterase activity in the plasma membranes and revealed the axolemma as the membrane exhibiting the greatest enzymatic activity. The end product of the reaction appeared to be focally distributed along the axolemma. Only random deposits of end product were observed in the axoplasm and in the Schwann cell layer ( G . M. Villegas and Villegas, 1974). The in uitro assay of acetylcholinesterase carried out at that time by G. Camejo (personal communication) on subcellular fractions isolated from these nerve fibers also showed that the enzyme activity is mainly associated with the plasma membranes. The activity in the soluble portion of the nerve fiber homogenates was rather scarce, and the axolemma-enriched fraction had a specific activity of 36 kmol per milligram of protein per hour. This activity was found to be about four times higher than that of the Schwann cell plasma membrane-enriched preparation from the same nerve fibers. The distribution pattern of the acetylcholinesterase activity in the axolemma was found to be similar to that of other enzymes, such as the adenosinetriphosphatase in the giant nerve fiber of D.p h i (Sabatini et al., 1968), and also resembled that of the special structural arrangements present at the axon-Schwann cell interface ( G . M. Villegas and Villegas, 1976).

2. ACETYLCHOLINE RECEPTORS

The effects of acetylcholine and carbamoylcholine on the Schwann cell membrane potential that appeared to reproduce the long-lasting effects of impulse train conduction by the axon, the blocking action of curare, and the prolonging effects of eserine were considered as confirming the presence of acetylcholine receptors in the Schwann cell membrane (J. Villegas, 1974). Such receptors were then characterized in the same nerve

559

AXON-SCHWANN CELL RELATIONSHIP

fibers by determining the effects of a-bungarotoxin, nicotine, and muscarine on the Schwann cell electrical potential in resting and stimulated giant fibers (J. Villegas, 1975). The evidence obtained showed that abungarotoxin irreversibly blocks the long-lasting Schwann cell hyperpolarizations following the conduction of nerve impulse trains or the external application of carbamoylcholine to the resting nerve fiber, and that d-tubocurarine protects against the irreversible action of a-bungarotoxin on the Schwann cell. Finally, it was shown that the external application of nicotine, but not of muscarine, to the resting nerve fiber is able to reproduce the long-lasting effects of acetylcholine and carbamoylcholine on the Schwann cell membrane potential. All these experimental findings are summarized in Fig. 3. Further experiments aimed at localizing at the subcellular level the acetylcholine receptor sites, by studying with electron microscope autoradiography the distribution of 1251-labeleda-bungarotoxin binding sites, were carried out in S. sepioidea giant nerve fibers (Rawlins and Villegas, 1978). The evidence obtained showed that, both in the intact nerve fiber and in axoplasm-free nerve fiber sheaths, the binding sites for 1251-labeled a-bungarotoxin appear to be concentrated along the axon-Schwann cell boundary and at the lateral surfaces of the Schwann cell delimiting the intercellular channels. 3. ACETYLCHOLINE CONTENT After the demonstration of the presence of acetylcholine receptors in the Schwann cell membrane, and of acetylcholinesterase enzymatic activity at the axon and Schwann cell surfaces, a series of experiments aimed at identifying and measuring acetylcholine by means of gas chromatogra-

I

R E P E T I T I V E A X O N A L EXCITATION

I

I

SCHWANN C E L L H Y P E R P O L A R I Z A T I O N

I

FIG.3. Pharmacological characterization of the Schwann cell hyperpolarizations following axonal excitation. (After J . Villegas, 1975, 1978a.)

560

JORGE VILLEGAS

phy-mass spectrometry were carried out in S. sepioidea nerve fiber extracts (J. Villegas and Jenden, 1979).The results obtained in two different experiments in which acetylcholine was measured in samples of intact giant nerve fibers, extruded axoplasm, and axoplasm-free nerve fiber sheaths are summarized in Table 1. Since the weight of the periaxonal sheaths of a single fiber represents only about 25% of the total weight of the intact fiber, the periaxonal cells appear to contain as much acetylcholine as the voluminous giant axon. These experimental findings settled the point on the presence of acetylcholine in the giant nerve fiber of S . SPpioidea (J. Villegas and Jenden, 1979). The outstanding and somewhat unexpected feature of the results of this study was the high (100-200 p M )acetylcholine concentration in the Schwann cell, which is about 40 times that of the axoplasm and close to the 0.35 mM level reported for individual soma of characterized cholinergic neurons of other molluscs (McCaman ~t al., 1973). The low acetylcholine levels (2-5 p M )found in the axoplasm were similar to those previously reported for other noncholinergic neurons (McCaman et al., 1973). These experimental findings suggest that the Schwann cells themselves are the main source of the acetylcholine released after the conduction of nerve impulse trains in these nerve fibers (J. Villegas and Jenden, 1979). 4. ACETYLCHOLINE SYNTHESIS

The possibility that the acetylcholine present in the Schwann cells could be synthesized within them was also explored in S . sepioidea nerve TABLE 1 ACETYLCHOLINE CONCENTRATION I N THE SCHWANN CF.I.I. A N D AXONOF S. sepioidea GIANTNERVEFIBERS" Wet weight of sample Tissue sample

(ms)

Axoplasm

3.45 (4)b 8.50 (4y I .37 (4jb 3.29 (5)'

Sheaths

Cellular water per sample

(4)

3.29 (4)b 8.10 (4)' 0.05 (4)b 0.18 (5r

Concentrations ACh (pmol me-')

1.7 2 0.7 4.1 2 0.7 4.5 t 1.S 12.9 2 1.8

ACh (pmol liter-') 2 5 107 222

-t

2

*

2

0.2 0.8 23 10

'' Experimental data are from J . Villegas and Jenden (1979). Amounts are expressed per unit wet weight of tissue sample and per volume of cellular water in the samples (number of samples in parentheses); values are mean k SEM. Single nerve fiber per sample. ' Two to four pooled nerve fibers per sample to achieve greater accuracy. 'I

561

AXON-SCHWANN CELL RELATIONSHIP

fibers (Heumann et al., 198I), by assaying acetyltransferase activity in homogenates obtained from pooled samples of whole stellar nerve, intact giant nerve fiber, extruded axoplasm, axoplasm-free giant nerve fiber sheaths, and small nerve fiber bundles. The results, summarized in Table 11, showed that the giant fiber has acetyltransferase activity, which is mainly localized in the cells of the periaxonal sheaths. The rate of acetylcholine synthesis estimated for the Schwann cells is about 3-9 times that found in axoplasm and close to the level of enzyme activity found for the intact giant nerve fiber (axon plus Schwann cell sheath). These experiments showed that the acetylcholine present in the giant nerve fiber is synthesized in it, and they indicated that the higher levels of acetylcholine found in the Schwann cell sheath can be attributed to their higher synthetic activity compared to axoplasm (Heumann et al., 1981).

5. ACETYLCHOLINE RELEASE The Schwann cells in both transected nerves and denervated muscle fibers of the frog can synthesize choline acetyltransferase and acetylcholine and release acetylcholine (Katz and Miledi, 1959; Birks et al., 1960; Miledi and Slater, 1968; Dennis, 1972; Dennis and Miledi, 1974; Bevan et al., 1973, 1976; Tucek et al., 1978). However, the behavior of the denervated Schwann cell is different from that of the nerve terminal with respect to the mechanisms responsible for acetylcholine release (Dennis and Miledi, 1974; Bevan et al., 1976). Thus, the rate of miniature end-plate potentials recorded in the denervated muscle is not increased by a rise in external potassium concentration, an increase in the tonicity of the exterTABLE 11 ACETYLTRANSFERASE ENZYMATIC ACTIVITY IN THE SCHWANN CELLA N D AXONOF S. sepioidea GIANTFIBERS

ACh synthesis (pmol min-I) Protein content" Sample type

(ms)

Per milligram

Per fiber

Axoplasm

3.4 (40) 3.5 (43) 1.0 (25) 1.3 (24)

20 20 I80 70

1.7

Sheaths

1.6

7.2 3.8

(' Number of nerve fibers is given in parentheses.

562

JORGE VILLEGAS

nal medium, a rise in external calcium concentration, or application of black widow spider venom. It was found that only strong depolarizing current pulses and hyposmotic solutions were able to induce acetylcholine release from denervated Schwann cells (Dennis and Miledi, 1974; Bcvan r t d.,1976). However, in the giant nerve fiber of S. sepiuidea the Schwann cells release acetylcholine in response to axonal excitation (J. Villegas, 1973, 1975)and to the external application of hypoosmotic and hyperosmotic test solutions, black widow spider venom, and cardiac glycosides. In addition, it has also been found that in the unstimulated nerve fiber, an apparently spontaneous release process maintains a basal level of acetylcholine concentration in the intercellular clefts that can be detected, as a transient phenomenon, immediately after the addition of eserine to the external medium at a final concentration of 10 M. The acetylcholine release induced by the nerve impulses is suppresscd by removing Caz+ from the extracellular environment with EGTA or by increasing the external Mg2' from 53 mM (control level) to 75 mM, or by decreasing external K+ from 10 mM level to 0.1 m M ;whereas depolarizing the cells in high (30 mM) potassium solutions does not induce detectable levels of acetylcholine release in these nerve fibers (A. Ramos, V. Reale, and J. Villegas, unpublished results). In a similar way, the Schwann cell cholinergic responses induced by the external application of ouabain can be suppressed by reducing the tcnsion of oxygen in the external bathing solutions. Furthermore, as will be described in Section 1V.B, the external application of M L-glutamate to the unstimulated nerve fiber induces a transient increase in acetylcholine release followed by a long-lasting reversible blockage of the release mechanism in these nerve fibers. The cholinergic nature of the adaxonal glial hyperpolarizing responses to external application of ouabain was determined by Smiley and Lieberman (1981) in the medial giant nerve fiber of the ventral nerve cord of the crayfish Procamburus clarkii. The axon satellite cells in this nerve fiber have acetylcholine receptors of the nicotinic type and become hyperpolarized in the presence of acetylcholine and cholinergic agonists (Lieberman et al., 1980). Taken all together, the experimental findings referred to above appear to indicate that the physiological properties of the mechanism responsible for the release of the acetylcholine present in the Schwann cells of the giant nerve fiber of S.sepiuideu are similar to those of the normal nerve terminal. The distribution of the cholinergic system in these nerve fibers is summarized in Fig. 4. The nature of the axonal signal that triggers the Schwann cell cholinergic response is still not known. In addition to the depolarizing effect of

AXON-SCHWANN CELL RELATIONSHIP

i A C h l 5 PM CAT

20 p m o l mg-'min-'

563

A

FIG.4. Distribution of acetylcholinesterase(AChE), acetylcholine (ACh), acetylcholine receptors (ACh-R), and acetylcholine synthesis (CAT) in the Schwann cell layer (SC), axon (A), basal lamina (BM), and endoneurium (E) in the giant nerve fiber. (After J. Villegas, 1981.)

potassium leakage from the excited axon, other factors could be released during propagation of nerve impulse trains by the axon. Since glutamate is the putative neurotransmitter of the giant fibers in the squid, the sensitivity of the Schwann cell membrane potential to glutamate has also been explored. B. Glutaminergic Responses

The external application of glutamate to the giant axon of Loligo has no appreciable effect on its membrane potential at the extrasynaptic regions of the nerve fiber, whereas it depolarizes the postsynaptic fiber at the distal giant synapse in the stellate ganglion (Miledi, 1966, 1972; Bevan et al., 1975). In the giant nerve fiber of S . sepioidea, the membrane potential of the axon remains unchanged in the presence of externally applied glutamate (lo-*, M), whereas the Schwann cells in the same nerve fiber undergo a transient hyperpolarizing membrane potential change, followed by a sustained but readily reversible depolarization (J. Villegas, 1978a,b). Further experiments carried out in the same nerve fibers by J. Villegas, V. Reale, and D. W. Wiza, which will be published in detail elsewhere, confirmed the presence of specific receptor sites for glutamate in the

564

JORGE VILLEGAS

plasma membrane of the Schwann cell. It has also been found that the initial hyperpolarizing phase of the Schwann cell membrane potential response is determined by the activation of the acetylcholine release mechanism, followed by its reversible although long-lasting blockade. Thus, as appears illustrated in Fig. 5 , it has been observed that, after the first application of glutamate, the Schwann cell responses to axonal excitation are suppressed while the external application of carbamoylcholine produces its usual hyperpolarizing effect on the Schwann cells. After a relatively long interval of time (15-30 min), the cholinergic Schwann cell responses evoked either by the nerve impulse trains or by a new application of glutamate are recovered. It also became evident that the Schwann cell membrane potential responses to externally applied glutamate are suppressed by ouabain M ) and do not take place at low Ca2+(0.9 m M ) and Mg2+(3 mM) concentrations. On the contrary, large variations in the cxternal Na' and K+ concentrations, and the external application of M tetrodotoxin, have no appreciable effects on the Schwann cell membrane potential responses to glutamate. Furthermore, a series of experiments carried out in the same nerve fibers, and aimed at detecting and characterizing the possible existence of a glutamate uptake mechanism in the plasma membrane of the Schwann cell, indicate that the Schwann cells in axoplasm-free sheaths exposed to similarly low external concentrations of ~-['~C]glutarnate in the presence of external Na +,are able to accumulate radioactive labeled material in

-20

0

10

20

30

LO

Time (rninl

FIG.5. Schwann cell membrane potential (M.P.) changes induced by successive external applications of glutamate and carbarnoylcholine (carbachol) to the unstimulated nerve fiber. Each point corresponds to the potential difference measured in a different Schwann cell along the same nerve fiber.

AXON-SCHWANN CELL RELATIONSHIP

565

their interior by means of a high-affinity uptake mechanism ( R . E. BIanco and J. Villegas, unpublished results; see also Baker and Carruthers in this volume). The presence of specific receptor sites and of an uptake mechanism for glutamate in the plasma membrane of the Schwann cell strongly suggests that this amino acid may be directly involved in the mechanism of axonSchwann cell signaling described above. The sensitivity to glutamate of the acetylcholine release mechanism present in these nerve fibers appears to give further support to such a possibility. However, no direct evidence on the composition and amino acid content of the extracellular fluid in the nerve fiber clefts under physiological conditions is yet available. V.

CELL-TO-CELL COM MUNICATlON

A. Significance of Axon-Schwann Cell Signaling

Signaling between neurons and glia by the potassium mechanism represents a nonspecific system of communication, as discussed by Kuffler and Nicholls (1976), since it is not confined to special structures and occurs along the entire length of a neuron wherever potassium is released. It has been found in different experimental preparations that such nonspecific neuronal signal may modify the metabolic activity of glial cells (Orkand et a/., 1973; Salem et a / . ,1975; Penthreath and Kai-Kai, 1982). In addition, it has been found in other preparations (Orkand and Kravitz, 1971; Schon and Kelly, 1974), that glial cells can take up y-aminobutyric acid (GABA) and then release it as a consequence of increasing the external potassium concentration. On the other hand, as referred to in Section IV,A,5, in the giant nerve fiber of the squid, Schwann cells do not release acetylcholine in response to increases in external potassium concentration (J. Villegas, 1972). Thus, further experimental evidence is needed to evaluate the effects of potassium signals from the axon on its satellite Schwann cells in these nerve fibers. Furthermore, glial cells, in addition to being sensitive to relatively large millimolar changes in the external concentration of potassium and other ionic species, are known to respond also to micromolar concentrations of chemical neurotransmitter substances and neuroactive agents (Orkand and Kravitz, 1971; J. Villegas, 1973, 1974, 1975, 1978a,b; Schon and Kelly, 1974; Kelly and Dick, 1976; J. Villegas et a / . , 1976; Currie and Dutton, 1980; Lieberman et al., 1980; Smiley and Lieberman, 1981; Currie and Kelly, 1981). In addition, while the direct consequences of potassium release by axonal activity in S . sepioidea giant fibers disappear within a

566

JORGE VILLEGAS

few seconds after the termination of a train of impulses, the consequences of acetylcholine release on the satellite cells outlast for several minutes the duration of the trains and are not accompanied by any appreciable change in the membrane potential of the neuronal element. Furthermore, the neuronal element appears to be well protected by the presence of a high level of hydrolytic enzyme activity in its own membrane, against the neurotransmitter substance for which the satellite cell has specific receptors. In a similar way, while the Schwann cell has specific receptor sites and a high affinity uptake mechanism for glutamate, the giant axon, which is known to have intracellular concentrations of this amino acid in the millimolar range, appears to have receptors for it only at the synaptic regions of the nerve fiber. These experimental findings raise the possibility that, through the distribution of receptors, enzymes, storage and release mechanisms for different specific neurotransmitter substances, neurons and glial cells may be establishing systems of cell-to-cell communication and feedback control mechanisms for their functional coupling that do not interfere with their being able to play their own specific roles in two different time domains of nerve activity. 6. Transfer of Cellular Materials

The squid giant fiber has also permitted the experimental testing of the hypothesis that proteins can be transferred from glial cells to neurons (Lasek and Tytell, 1981). It has been found that 20-40% of the labeled proteins synthesized from labeled precursors by the Schwann cells appear in the axoplasm (Lasek et al., 1974). Autoradiographic analyses of the giant fiber have shown that the Schwann cells actively incorporate amino acids into proteins (Lasek ef al., 1973). In addition, in continuously perfused giant axons, newly synthesized proteins appear in the perfusate during periods of up to 8 hr (Lasek et al., 1977; Gainer el al., 1977). A comparison between the glial transfer proteins and the proteins that constitute the axoplasm has shown substantial differences between them, and it was found also that some of the proteins transferred from the glial cells to the axon are special in that they are probably not supplied also to the axon from its cell bodies (Lasek and Tytell, 1981). The question still remains as to how these proteins are transferred from the Schwann cells to the axon. Whether the protein transfer occurs through cell-to-cell contacts involving specialized regions of the axolemrna and Schwann cell plasma membranes, such as those detected morphologically (G. M. Villegas and Villegas, 1976), is not known at present. In a similar way, the images of

AXON-SCHWANN CELL RELATIONSHIP

567

endocytic or exocytic processes occurring in the Schwann cells and axon, which have been captured by freeze-fracturing the tissue (Section IC,B; see also G. M. Villegas and R. Villegas in this volume), may also be related to such a macromolecular transfer process. However, at present this is just an attractive possibility that requires further experimental work to test its validity. VI.

CONCLUDING REMARKS

In this article we have tried to summarize some of the experimental evidence resulting from the pioneering studies of J. Z. Young, B. B. Geren-Uzman, F. 0. Schmitt, B. Frankenhaeuser, A. Hodgkin, R. Villegas, and G. M. Villegas on the use of the giant nerve fiber of the squid as an experimental tool in which to explore the relationships between the axon and its satellite Schwann cell. In doing so, we have tried to show how a continued effort to correlate structure and function has provided clues for the understanding of chemical and pharmacological findings that otherwise would have been left unexplained for a longer period of time. Such an interest in disclosing the functional organization of a nerve fiber, which in Theodor Schwann’s terms has remained stationary at an early stage of development by not having learned how to make myelin, may eventually teach us some new facts about the functioning of nerves that are every bit as important as rapid conduction of nerve impulses,

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membrane sensitive to ionic concentrations and cardiac glycosides. J. Cell Biol. 69, 19-28. Villegas, R., and Villegas, G . M. (1960). Characterization of the membranes in the giant nerve fiber of the squid. J. Gen. Physiol. 43, 73-103. Villegas, R., and Villegas, G. M. (1965). Les couches superficielles de la fibre nerveuse du calmar. Actual. Neurophysiol. 6, 55-68. Villegas, R., Caputo, C., and Villegas, L. (1962). Diffusion barriers in the squid nerve fiber. The axolemma and the Schwann layer. J . Gen. Physiol. 46, 245-255. Villegas, R., Villegas, L., G i m h e z , M., and Villegas, G. M. (1963). Schwann cell and axon electrical potential differences. Squid nerve structure and excitable membrane location. J. Gen. Physiol. 46, 1047-1064. Watson, W. E. (1974). Physiology of neuroglia. Physiol. Reu. 54, 245-270. Webb, G. D., Dettbarn, W. D., and Brzin, M . (1966). Biochemical and pharmacological aspects of the synapsis of the squid stellate ganglion. Biochem. Pharmacol. 15, 18131819.

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Index

A

Acetylcholine, in Schwann cell, 559563 release, 561-563 synthesis, 560-561 Acetylcholine receptor, in Schwann cell, 558-559, 563 Acetylcholinesterase activity, in Schwann cell, 558 Action potential, 280-288, see ako Postsynaptic potential effects of anesthetics on, 450-453 Action potential waveform, voltage clamp with, 538-544 Adenosine triphosphatase. in axoplasm, 68-70 Adenosine triphosphate axonal pH and, 264-265 calcium efflux and, 226-23 I , 237-238 chloride uptake and, 181-183, 185 potassium-free effect and, 168-170 potassium influx and, 152 sodium exchange and, 159-160 sodium extrusion and, 134-139, 148-151 sugar transport and, 118-1 19 (+)-trans-Allethrin, 500 structure, 485 Amino acid, axoplasmic, 60, 92, 93 metabolism, 96-97 state of, 95-96 Amino acid transport, 119-127 Ammonium ion, effect on axonal pH, 252255 Anesthesia early studies, 449-450 squid axon model, 445-447

ATPase, see Adenosine triphosphatase Axolemma, 7, 12-19 chemical dissection studies, 75-76 membrane-associated proteins, 76-80 Axon, see Squid giant axon Axonal transport, in squid giant axon, 4749 Axon-Schwann cell interface, 548-554 inner boundary, 549 outer boundary, 549-550 periaxonal space, 550 Axon-Schwann cell signaling electrical signals, 554-555 potassium signals, 555-557 significance of, 565-566 synapsic-like signals, 557 Axoplasm, 7- 12 Ca-binding capacity, 202-207 Ca ion concentrations in, 196-197 carbohydrates in, 60, 92-1 19 cholinergic systems in, 71 cytoskeletal proteins in, 45-47 DFPase in, 70-71 energy metabolism in, 68-70 immunological studies, 80-8 1 ions and small molecules in, 59, 60, 9293 lipids in, 59-66 Mg ion concentrations in, 196-197 molecular composition, 58-68, 92-97 nucleic acids in, 67-68 protein in, 66 regulation of pH,, 189-190 sugars in, 92 Axoplasm, extruded, 49-50 artificial medium for, 61 preparation, 132 573

574

INDEX

Axoplasmic matrix, 40-41 cross-linking elements in, 41-42 domains in, 42-45 Azide, effect on axonal pH, 257-258

B Barium ion, 362 Batrachotoxin, 493-495, 506-509 structure, 485 Bicarbonate ion, effect on axonal pH, 255257, 258-261 Binding constant, 315-321 Binding energy, 366-368

in axoplasm, 178-180 in hemolymph, 178-180 Chloride transport conductive, 180- 181 coupled to Na and K transport, 181-189 Cholinergic responses, of Schwann cell, 557-563 Cholinergic system, 71 Conductance calculation of, 338-339 kinetics, 339-341 reduction of maximums, 471-473 Cyanide effect on axonal pH, 257-258 inhibitor, of transport, 124. 151, 169 Cytochalasin B, 102

C Calcium current EPSP latency and, 542-544 presynaptic, 531-534, 535 presynaptic depolarization and, 540-542 transmitter release and, 542-544 Calcium efflux, 222-235 Calcium homeostasis, 201-242 modulation of, 240-242 steady-state maintenance, 236-242 Calcium influx, 2 1 1-222 Calcium ion fluxes across axolemma, 207-235 internal concentration, determination of, 438-439 intracellular binding, mechanisms of, 202-207 CANP, see Protease. calcium-dependent neutral Cardioactive steroids inhibitors, of sodium pump, 142-144 Na fluxes in presence of, 163-168 Carbohydrate. axoplasmic, 60, 92-1 19. see ulso Sugar axoplasmic Cesium ion, 360-362, 365, 366 Channel noise, 380-382 extrinsic, 382-385 in nerve membranes, 385-397 Chemical dissection studies, of axolemma, 75-76 Chloride ion axonal pH and, 263-264

D DepolariLation, presynaptic, calcium current and, 540-542 DFP, see Diisopropyl phosphofluoridate Dialysis, internal, 133, 250-252 cu-Dihydrograyanotoxin 11. 495 Diisopropyl phosphofluoridate, 70-71 Diisopropyl phosphofluoridate hydrolyLing enzyme, 70-7 1 2,4-Dinitrophenol effect on axonal pH. 257-258 inhibitor, of potassium influx, 151 Dye measurements of membrane potential, 429-438 dye-screening in squid axon, 437-438 potential dependence of, 429-43 I , 433438 series resistance, 431-433 E Endoneurium, fine structure, 28, 29 Endoplasmic reticulum, in squid axoplasm, 9-1 1 Energy bamers in transport, 366-368 Energy metabolism, in squid giant axon, 68-70 EPSP latency, calcium current and, 542-

544 Equilibrium noise, 375-377 Equilibrium potential, 526-530

INDEX

575 F

Lipid site of anesthetic action, 447-449 Long pore effect, 114

Fluctuation analysis, 372-374

G

Gating charge-voltage relation, 341343 Gating current calculation of, 338-339 kinetics, 339-341 Glutamate transport, 119, 124-127 GIycine transport, 119, 120-124 Grayanotoxin I, 495-500 active moieties, 498-500 structure, 485

H Hexose transport, see Sugar transport Hodgkin-Huxley analysis, 294-308 Hydration energy, 366-368

I

Image potential energy, 366-368 Ionic channel, 332-334, see also Multi-ion pore ionic free-energy profile, 366-368 modulation by chemicals, 493-507 single-channel studies, 506-507 Ionic channel block current-dependent block, 489-493 kinetic schemes of, 489-493 technological developments in, 486-487 voltage-dependent, 487-488 Ionic channel surface-charge density neutralization of, 412-413 relation to average surface-charge density, 414-415 theory, 408-412

L

Lipid, metabolism of, in axoplasm. 59-66

M Magnesium homeostasis, 197-201 Magnesium ion, intracellular, effect on sodium efflux, 146-148 Membrane birefringence, 426-427 intrinsic signals, 426-429 light scattering, 428-429 Membrane noise, physical basis of, 374385 Membrane potential, dye measurements, 429-438 Membrane surface charge, 407-418 average density, 414-415 modifications of, 415-416 neutralization, 4 12-4 13 significance of, 416-417 Microelectrode, protruding tip, pH-sensitive, 252 Microfilaments, 7, 45-47 Microinjection, 133 Microtubule, 7-9 cross-bridges on, 41-42 domain, 42-45 Microtubule-associated protein, 41-42 Mitochondria, in squid axoplasm, 9 Mobile carrier model, 112, 113, 115-117 Multi-ion pore, 355-357 blocking ions and, 357 concentration-dependent properties, 355-357 unidirectional fluxes, 357-358

N

Nerve fractions, ultrastructural analysis, 29-32 Neurofilament, 7-9 cross-bridges on, 41 domain, 42-45 Neuroplasmic lattice, 11-12 Neurotransmitter, at giant axon synapse, 526

576

INDEX

Noise analysis, 371-400, see dso specific types Nucleic acid, axoplasmic, 67-68 0

Oligomycin. inhibitor, of sodium pump, 143, 146 Optical signals, 424-439 Ouahain, inhibitor, of transport. 124, 142I44

P Pancuronium, structure, 485 Perfusion, internal, 133 Periaxonal space, 550 Permeability ratio, 315-321 PH effect on sugar transport, 108, 110, 112 glutamare transport and, 126 pH, axonal ATP and, 264-265 external Na and. 261-263 internal chloride and, 263-264 metabolic inhibitors and, 257-258 regulalion of, 258-268 weak acids and, 255-257, 258-261 weak bases and, 252-255 pH. intracellular, chloride ion and, 189190 Phforetin, 102 Phosphates, high-energy, in axoplasm, 60 Postsynaptic potential as function of presynaptic calcium current, 534-535 quanta1 nature of. 525-526 Potassium channel blocking ions, 3.59-362 concentration-dependent effects, 358359 current and concentration and, 364-365 flux studies, 362-363 mathematical model. 363-366 noise analysis, 38.5-392 unidirectional flux ratio, 365-366 Potassium current effects of anesthetics on, 454-463 mechanisms of suppression of, 464-475

Potassium influx, 150-153 ATP and, 152 external K and, 152-153 inhibitors, 151 internal Na and, 1.53 Potassium ion, chloride transport and, 185- I89 Potassium signal, in axon-Schwann cell interaction, 55.5-5.57 Protease, calcium-dependent neutral, 7173 Protein in axoplasm, 66 membrane-associated, 76-80 Protein site of anesthetic action, 447-449 Protein transfer, from Schwann cell to axon, 73-74, 566-567 Pyrethroids, 500-506 R

Rubidium ion, 359-360 S

Schwann cell, 550-554, SYY nlso AxonSchwann cell cholinergic responses, 557-563 fine structure, 19-27 ghtaminergic response, 563-565 ionic permeability, 552-554 membrane potential, 5.51 protein transfer from, 73-74 relationship with squid giant axon, 5475 67 Shot noise, 377, 378-379 Single-channel recording, 397-400 Single-channel simulation, 343-347 Single-channel studies, of chemical modulations, 506-507 Sodium channel, sec c t l w Ionic channel block chemical block, 485, 486-493 chemical modulation of, 485, 493-507 closed-channel block, 489-490 closed conformations, 334-335 energy diagram, 336-338 four-state model, 335-336 inactivation inhibitors, 485

INDEX kinetic schemes of blocks, 489-493 molecular properties, 3 15-321 noise analysis, 392-397 open-channel block, 490-493 pharmacology of. 483-510 sequential models of, 332-352 steady-state inactivation, membrane thickness and, 475-478 voltage-independent polymerization step, 347-350 Sodium current effects of anesthetics on, 454-463 mechanisms of suppression of, 464-475 Sodium-dependent calcium efflux, 23 1-235 Sodium efflux external K activation of, 139-140, 148150 extrusion, free energy source for, 134139 inhibitors, I4 1- I46 internal Na activation of, 140-141, 148151

intracellular Mg and, 146-148 uncoupled, 161-163 Sodium ion axonal pH and, 261-263 chloride transport and, 183- I85 magnesium efflux and, 198-201 Sodium pump, 134- I70 active Na-K exchange, 134-156 K-free effect, 168-170 K influx, 150-153 membrane potential and, 155-156 Na efflux, 134-150 Na: K coupling ratio, 153-156 Na-Na exchange, 156-161 reversal, 163 Square pulses, in voltage-clamp studies, 530-538 Squid blood ionic composition, 60 nonelectrolyte composition, 93 Squid giant axon, 3-33, 39-51, .see cilso Axon-Schwann cell anatomical organization, 4-6 artificial medium for, 61 axoplasm extrusion, 132 as dye-screening preparation, 437-438 effects of anesthetics on, 445-463 electrical behavior, 308-3 15

577 fine structure, 7-19, 40-47 histological organization, 6-7 internal dialysis, 133 internal perfusion, 133 microinjection, 133 protein transfer to, 73-74, 566-567 voltage-clamp studies, 454-463 Squid giant axon synapse, 519-544 anatomy, 521-522 delay at, 535-536 early electrophysiology, 523-524 equilibrium potential, 526-530 neurotransmitter, 526 transmission model, 536-538 transmission without action potentials, 524-525 voltage dependence of synaptic current, 530 State diagram, 332-334 Steady-state activation curve, shifts in, 468-47 1 Steady-state inactivation curve, shift in, 464-468 Strophanthidin, inhibitor, of sodium pump, 143-144, 163-168 Strophanthin G, see Ouabain Subplasmalemmal domain, 42 Sugar, axoplasmic, see also Carbohydrate, axoplasmic state of, 92-95 Sugar transport asymmetric, 110-118 competitive, 101-102 electrical activity and, 102-103 facilitated, 100- 101 inhibitors of, 102 kinetics, 98-102, 1I I metabolic depletion and, 103-107 models, 110-119 passive, 98-100 pH and, 108, 110. 112 regulation, 102-109, 118-1 19 temperature dependence, 109 Surface charge, see Ionic channel surfacecharge density; Membrane surface charge Synapse, see Squid giant axon synapse Synaptic current, voltage dependence of, 530 Synaptic delay, 535-536

INDEX Synaptic transmission model of, 536-538 without action potentials, 524-525

T

Temperature, sugar transport and, 109, 110 (+)-rrans-Tetramethrin, 500-506, 507, 510 structure, 485 Tetrodotoxin calcium influx and, 217-221 structure, 485 Three-site model, 363-366 Time constants, 473-474 Trans effect, 114 Transport, see d s n Amino acid transport; Sugar transport

competitive, 101- 102 energy barriers, 366-368 facilitated, 100-101 mechanisms, 265-268 passive, 98-100 Transport noise, 377-380 Two-component carrier model, 112, 113, 117-1 18

V Vanadate, inhibitor, of sodium pump, 143. 144-146, 163-168, I70 Voltage clamp using square pulses, 530-538 with action potential waveform, 538-544 Voltage-clamp currents, 288-294

Contents of Previous Volumes Volume 1 Some Considerations about the Structure of Cellular Membranes MAYNARD M. DEWEYAND LLOYDBARR The Transport of Sugars across Isolated Bacterial Membranes H. R. KABACK Galactoside Permease of Escherichia roli ADAMKEPES Sulfhydryl Groups in Membrane Structure and Function ASER ROTHSTEIN Molecular Architecture of the Mitochondrion DAVIDH. MACLENNAN Author Index-Subject Index

Volume 2 The Molecular Basis of Simple Diffusion within Biological Membranes W. R. LIEBAND W. D. STEIN The Transport of Water in Erythrocytes ROBERTE. FORSTER Ion-Translocation in Energy-Conserving Membrane Systems B. CHANCEAND M. MONTAL Structure and Biosynthesis of the Membrane Adenosine Triphosphatase of Mitochondria ALEXANDER TZAGOLOPF Mitochondria1 Compartments: A Comparison of Two Models HENRYTEDESCHI Author Index-Subject Index

Volume 3 The Na ,K -ATPase Membrane Transport System: Importance in Cellular Function ARNOLDSCHWARTZ, AND GEORGEE. LINDENMAYER, JULIUSC. ALLEN +

Biochemical and Clinical Aspects of Sarcoplasmic Reticulum Function ANTHONYMARTONOSI The Role of Periaxonal and Perineuronal Spaces in Modifying Ionic Flow across Neural Membranes w. J . ADELMAN, JR. A N D Y . PALTI Properties of the Isolated Nerve Endings DE LORES GEORGINA RODRIGUEZ DE ROBERTIS ARNAIZAND EDUARDO Transport and Discharge of Exportable Proteins in Pancreatic Exocrine Cells: In Vitro Studies J. D. JAMIESON The Movement of Water across VasopressinSensitive Epithelia RICHARD M. HAYS Active Transport of Potassium and Other Alkali Metals by the Isolated Midgut of the SilkW0I.m WILLIAMR. HARVEYAND KARL ZERAHN Author Index-Subjecr Index

Volume 4 The Genetic Control of Membrane Transport CAROLYN W. SLAYMAN Enzymic Hydrolysis of Various Components in Biomembranes and Related Systems MAHENDRA KUMARJAIN Regulation of Sugar Transport in Eukaryotic Cells HOWARDE. MORGANA N D CAROL F. WHITFIELD Secretory Events in Gastric Mucosa RICHARD P. DURBIN Author Index-Subject Index

+

Volume 5 Cation Transport in Bacteria: K , Na , and H + +

579

+

580 FRANKLIN M. HAROLDA N D KARLHEINZ ALTENWRF Pro and Contra Carrier Proteins: Sugar Transport via the Periplasmic Galactose-Binding Protein WINFRIED Boos Coupling and Energy Transfer in Active Amino Acid Transport ERICHHEINZ The Means of Distinguishing between Hydrogen Secretion and Bicarbonate Reabsorption: Theory and Applications to the Reptilian Bladder and Mammalian Kidney WILLIAM A. BRODSKYA N D THEODOREP. SCHILB Sodium and Chloride Transport across Isolated Rabbit Ileum G . SCHIJLTZ AND STANLEY PETERF. CURRAN A Macromolecular Approach to Nerve Excitation ICHIJI TASAKI A N D EMILIO CARBONE Subject Index

Volume 6 Role of Cholesterol in Biomembranes and Related Systems MAHENVRA KUMARJAiN Ionic Activities in Cells A. A. LEV A N D W. McD. ARMSTRONG Active Calcium Transpon and Ca2+-Activated ATPase in Human Red Cells H.J. SCHArZMANN The Effect of Insulin on Glucose Transport in Muscle Cells TORBEN CLAUSEN Recognition Sites for Material Transport and Information Transfer HALVORN. CHRISTENSEN Subject Index

Volume 7 Ion Transport in Plant Cells E. A. C. MACROB~IE H Ion Transport and Energy Transduction in Chloroplasts RICHARD A. DILLEYAND ROBERTT. GIAQUINTA +

CONTENTS OF

PREVIOUS VOLUMES

The Present State of the Carrier Hypothesis PAULG. LEFEVRE Ion Transport and Short-circuit Technique WARRENS. REHM Subject Index

Volume 8 Chemical and Physical Properties of Myelin Proteins M .A. MOSCARELLO The Distinction between Sequential and Simultaneous Models for Sodium and Potassium Transport P. J. GARRAHAN A N D R. P. GARAY Soluble and Membrane ATPase of Mitochondria, Chloroplasts, and Bacteria: Molecular Structure, Enzymatic Properties, and Functions RIVKAPANETA N V D. RAO SANADI Competition, Saturation, and Inhibition-Ionic Interactions Shown by Membrane Ionic Currents in Nerve. Muscle, and Bilayer Systems ROBERTJ. FRENCHA N D WILLIAM 1. ADELMAN, JR. Properties of the Glucose Transport System in the Renal Brush Border Membrane R. KlNNt Subject Index

Volume 9 The State of Water and Alkali Cations within the lntracellular Fluids: The Contribution of NMR Spectroscopy MORDECHAI SHPORERA N D MORTIMER M.CIVAN Electrostatic Potentials at Membrane-Solution Interfaces STUARTMCLAUGHLIN A Thermodynamic Treatment of Active Sodium Transport S. ROYCAPLANA N D ALVINESSIG Anaerobic Electron Transfer and Active Transport in Bacteria AND WIL N. KONINGS JOHANNES BOONSTRA Protein Kinases and Membrane Phosphorylation M. MARLENE HOSEYA N D MARIANO TAO

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CONTENTS OF PREVIOUS VOLUMES Mechanism and Physiological Significance of Calcium Transport across Mammalian Mitochondrial Membranes LEENAMELA Thyroidal Regulation of Active Sodium Transport F. ISMAIL-BEIGI Subject Index

Volume 10 Mechanochemical Properties of Membranes E. A. EVANSA N D R. M. H~CHMUTH Receptor-Mediated Protein Transport into Cells. Entry Mechanisms for Toxins, Hormones, Antibodies, Viruses, Lysosomal Hydrolases, Asialoglycoproteins, and Camer Proteins DAVIDM. NEVILLE,JR. A N D TA-MINCHANG The Regulation of Intracellular Calcium AND ERNESTOCARAFOLI MARTINCROMITON Calcium Transport and the Properties of a Calcium-Sensitive Potassium Channel in Red Cell Membranes VIRGILIOL. LEW A N D HUGOG . FERREIRA Proton-Dependent Solute Transport in Microorganisms A. A. EDDY Subject Index

Volume 11 Cell Surface Glycoprotelns: Structure, Blosynthesls, end Blologlcal Functlons The Cell Membrane-A Short Historical Perspective ASER ROTHSTEIN The Structure and Biosynthesis of Membrane GIycoproteins JENNIFER STURGESS, AND MARIOMOSCARELLO, HARRYSCHACHTER Techniques for the Analysis of Membrane Glycoproteins R. L . JULIANO Glycoprotein Membrane Enzymes JOHN R. RIORDAN AND GORDONG. FORSTNER

Membrane Glycoproteins of Enveloped Viruses RICHARD W. COMPANSAND MAURICEC. KEMP Erythrocyte Glycoproteins MICHAELJ. A. TANNER Biochemical Determinants of Cell Adhesion LLOYDA. CULP Proteolytic Modification of Cell Surface Macromolecules: Mode of Action in Stimulating Cell Growth KENNETHD. NOONAN Glycoprotein Antigens of Murine Lymphocytes MICHELLELETARTE Subject Index

Volume 12 Carrlers and Membrane Transporl Proteins Isolation of Integral Membrane Proteins and Criteria for Identifying Carrier Proteins MICHAELJ. A. TANNER The Carrier Mechanism S. B. HLADKY The Light-Driven Proton Pump of Halobacterium halobium: Mechanism and Function AND MICHAELEISENBACH S. ROY CAPLAN Erythrocyte Anion Exchange and the Band 3 Protein: Transport Kinetics and Molecular Structure PHILIPA. KNAUF The Use of Fusion Methods for the Microinjection of Animal Cells R. G. KULKAAND A. LOYTER Subject Index

Volume 13 Cellular Mechanlsms of Renal Tubular Ion Transport PART I: ION ACTIVITY AND ELEMENTAL COMPOSITION OF INTRAEPITHELIAL COMPARTMENTS Intracellular pH Regulation WALTERF. BORON Reversal of the pHi-Regulating System in a Snail Neuron R. C. THOMAS

582 How to Make and Use Double-Barreled Ion. Selective Microelectrodes THOMASZUtTHtN The Direct Measurement of K, CI, Na, and H Ions in Bullfrog Tubule Cells MAMORUFUJIMOTO, KUNlHlKO KOTERA, A N D YUTAKAMATSUMUKA lntracellular Potassium Activity Measurements in Single Proximal Tubules of Necfurus Kidney TAKAHIRO KUBOTA,BRUCEBIAGI,A N D GERHARD GIEBISCH lntracellular Ion Activity Measurements in Kidney Tubules RAJAN . KHURl lntracellular Chemical Activity of Potassium in Toad Urinary Bladder JOEL DELONGA N D MORTIMER M. C I V A N Quantitative Determination of Electrolyte Concentrations in Epithelial Tissues by Electron Microprobe Analysis RWER RICK,A m L t DORGE, RICHARD BAUER,FRANZBECK, JUNEMASON,C H H I S T I A ROLOK-, N~ A N D KLAUSTHURAU PART 11: PROPERTIES OF INTRAEPITHELIAL MEMBRANE BARRIERS IN THE KIDNEY Hormonal Modulation of Epithelial Structure JAMESB. WADE Changes in Cell Membrane Surfaces Associated with Alterations of Transepithelial Ion Movement MICHAEL KASHGARIAN The Dimensions of Membrane Barriers in Transepithelial Flow Pathways AND LARRYW. WELLING DANJ . WELLING Electrical Analysis of lntraepithelial Barriers EMILEL. BOULPAEPAND HENRYSACKIN Membrane Selectivity and Ion Activities of Mammalian Tight Epithelia SIMONA. LEWIS,NANCYK. WILLS, AND DOUGLASC . EATON

CONTENTS OF PREVIOUS VOLUMES Ion Conductances and Electrochemical Potential Differences across Membranes of Gallbladder Epithelium LUIS R ~ u s s A Kinetic Model for Ion Fluxes in the Isolated Perfused Tubule BRUCEBIACI,ERNESTOGONZAI.EZ. AND GERHARD GIEBISCH The Effects of Voltage Clamping on Ion Transport Pathways in Tight Epithelia ARTHURL. FINNA N D PAULAROGENES Tubular Permeability to Buffer Components as a Determinant of Net H Ion Fluxes G. MALNIC,V. L. COSTASILVA,S. S. CAMPIGLIA. M. DE MELLOAIRES,A N D G . GlEBlSCH Ionic Conductance of the Cell Membranes and Shunts of Necfurus Proximal Tubule AND GENJIROKIMURA KtNNtTH R. SPRING Lurninal Sodium Phosphate Cotransport as the Site of Regulation for Tubular Phosphate Reabsorption: Studies with Isolated Membrane Vesicles STOI.I., HEINIMURER,REINHARD CARL.AEVERS,ROLF KINNE, ' JtAN-PHILIPPE BONJOUR,A N D HERBERTFLEISCH The Mechanism of Coupling between Glucose Transport and Electrical Potential in the Proximal Tubule: A Study of Potential-Dependent Phlorizin Binding to Isolated Renal Microvillus Membranes PETERS. ARONSON Electrogenic and Electroneutral Na GradientDependent Transport Systems in the Renal Brush Border Membrane Vesicle BERTRAM SACKTOR PART 111: INTRAMEMBRANE CARRIERS AND ENZYMES IN TRANSEPITHELIAL TRANSPORT Sodium Cotransport Systems in the Proximal Tubule: Current Developments R. KINNE,M. BARAC,AND H. MURER ATPases and Salt Transport in the Kidney Tubule MARGARITA PEREZ-GONZALFZDE LA MANNA,FULGENCIO PROVERBIO, AND GUILLERMO WHITEMBURY

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CONTENTS OF PREVIOUS VOLUMES

Further Studies on the Potential Role of an Anion-Stimulated Mg-ATPase in Rat Proximal Tubule Proton Transport AND R. KINNE E. KINNE-SAFFRAN Renal Na+ -K+ -ATPase: Localization and Quantitation by Means of Its K +-Dependent Phosphatase Activity REINIERBEEUWKES 111 A N D SEYMOUR ROSEN Relationship between Localization of N + -K -ATPase, Cellular Fine Structure, and Reabsorptive and Secretory Electrolyte Transport STEPHENA. ERNST, CLARAV. RIDDLE,AND KARL1. KARNAKY, JR. Relevance of the Distribution of Na+ Pump Sites to Models of Fluid Transport across Epithelia JOHNW. MILLSA N D DONALDR. DIBONA Cyclic AMP in Regulation of Renal Transport: Some Basic Unsolved Questions THOMASP. DOUSA Distribution of Adenylate Cyclase Activity in the Nephron F. MOREL,D. CHABARDES, AND M. IMBERT-TEBOUL Subject Index

Permeation of Nucleosides, Nucleic Acid Bases, and Nucleotides in Animal Cells AND PETERG . W . PLAGEMANN ROBERTM. WOHLHUETER Transmembrane Transport of Small Peptides D. M. MATTHEWSAND 1. W. PAYNE Characteristics of Epithelial Transport in Insect Malpighian Tubules S. H. P. MADDRELL Subject Index

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Volume 14 Carriers and Membrane Transport Proteins Interface between Two Immiscible Liquids as a Tool for Studying Membrane Enzyme Systems L. I. B~GUSLAVSKY Criteria for the Reconstitution of Ion Transport Systems ADILE. SHAMOOA N D WILLIAMF. TIVOL The Role of Lipids in the Functioning of a Membrane Protein: The Sarcoplasmic Reticulum Calcium Pump J. P. BENNET,K. A. MCGILL,AND G. B. WARREN The Asymmetry of the Hexose Transfer System in the Human Red Cell Membrane W. F. WIDDAS

Volume 15 Molecular Mechanisms of Photoreceptor Transductlon PART I: THE ROD PHYSIOLOGICAL RESPONSE The Photocumnt and Dark Current of Retinal Rods G. MATTHEWSA N D D. A. BAYLOR Spread of Excitation and Background Adaptation in the Rod Outer Segment K.-W. YAU,T. D. LAMB,AND P. A. MCNAUGHTON Ionic Studies of Vertebrate Rods W. GEOFFREYOWENA N D VINCENTTORRE Photoreceptor Coupling: Its Mechanism and Consequences GEOFFREYH. GOLD PART 11: THE CYCLIC NUCLEOTIDE ENZYMATIC CASCADE AND CALCIUM ION First Stage of Amplification in the CyclicNucleotide Cascade of Vision LUBERTSTRYER,JAMESB. HURLEY, AND BERNARD K.-K. FUNG Rod Guanylate Cyclase Located in Axonemes DARRELLFLEISCHMAN Light Control of Cyclic-Nucleotide Concentration in the Retina THOMAS G. EBREY,PAULKILBRIDE, JAMESB. HURLEY,ROGERCALHOON, AND MOTOYUKI TSUDA Cyclic-GMP Phosphodiesterase and Calmodulin in Early-Onset Inherited Retinal Degenerations

CONTENTS OF PREVIOUS VOLUMES

G . J . CHADER,Y. P. Lru. G . AOUIRRE, R. T. FLETCHER, A N D M. T’so R. SANTOS-ANDEKSON, Control of Rod Disk Membrane Phosphodiesterase and a Model for Visual Transduction P. A. LIEBMAN A N D E. N . PLJOH, JR. Interactions of Rod Cell Proteins with the Disk Membrane: Influence of Light, Ionic Strength, and Nucleotides HERMANN KUHN Biochemical Pathways Regulating Transduction in Frog Photoreceptor Membranes M . DFRIC BOWNDS The Use of Incubated Retinas in Investigating the Effects of Calcium and Other Ions on Cyclic-Nucleotide Levels in Photoreceptors ADOLPH I . COHEN Cyclic AMP: Enrichment in Retinal Cones DEBORA8. FARBER Cyclic-Nucleotide Metabolism in Vertebrate Photoreceptors: A Remarkable Analogy and an Unraveling Enigma M. W. BITENSKY, G. L. WHEELER, AND A. YAMAZAKI, M. M. RASENICK, P. J . STEIN Cuanosine Nucleotide Metabolism in the Bovine Rod Outer Segment: Distribution of Enzymes and a Role of CTP HITOSHISHICHI Calcium Tracer Exchange in the Rods of Excised Retinas ETE Z. SZUTS The Regulation of Calcium in the Intact Retinal Rod: A Study of Light-Induced Calcium Release by the Outer Segment GEOFFREY H. COLDA N D JUANI. KORENBROT Modulation of Sodium Conductance in Photoreceptor Membranes by Calcium Ions and cGMP ROEERTT . SORBI PART 111: CALCIUM, CYCLIC NUCLEOTIDES, AND THE MEMBRANE POTENTIAL Calcium and the Mechanism of Light Adaptation in Rods AND BRUCEL. BASTIAN GORWN L. FAIN

Effects of Cyclic Nucleotides and Calcium Ions on Bufo Rods JOELE. BROWNA N D GERALDINE WALOGA The Relation between CaZ+ and Cyclic GMP in Rod Photoreceptors STUART A. LIPTON A N D JOHN E. DOWLING Limits on the Role of Rhodopsin and cGMP in the Functioning of the Vertebrate Photoreceptor SANFORD E. OSTROY, EDWARD P. MEYERTHOLtN, P ~ I E 1R. STEIN, ROBERTAA. SVOBODA, A N D MEEGAN J. WILSON [Ca2+]i Modulation of Membrane Sodium Conductance in Rod Outer Segments BURKSOAKLEYI1 A N D LAWRENCE H. PINTO Cyclic-GMP-Induced Depolarization and Increased Response Latency of Rods: Antagonism by Light H. MILLERA N D WII.L.IAM GRANI D. NICOI. PART 1V: AN EDITORIAL OVERVIEW Ca*+ and cGMP WILLIAM H. MIILGR Index

Volume 16

Electrogenic Ion Pumps PART I. DEMONSTRATION OF PUMP ELECTROGENICITY IN EUKARYOTIC CELLS Electrophysiology of the Sodium Pump in a Snail Neuron R. C. THOMAS Hyperpolarization of Frog Skeletal Muscle Fibers and of Canine Purkinje Fibers during Enhanced Na+ -K Exchange: Extracellular K + Depletion or Increased Pump Current? DAVIDC. GADSBY The Electrogenic Pump in the Plasma Membrane of Nirellu ROGERM. SPANSWICK +

585

CONTENTS OF PREVIOUS VOLUMES

Control of Electrogenesis by ATP, Mg2+, H + , and Light in Perfused Cells of Chara MASAHITAZAWAA N D TERUOSHIMMEN PART 11. THE EVIDENCE IN EPITHELIAL MEMBRANES An Electrogenic Sodium Pump in a Mammalian Tight Epithelium S. A. LEWISA N D N. K. WILLS A Coupled Electrogenic Na+-K+ Pump for Mediating Transepithelial Sodium Transport in Frog Skin ROBERTNIELSEN Transepithelial Potassium Transport in Insect Midgut by an Electrogenic Alkali Metal Ion Pump MICHAEL G . WOLFERSBERGER, WILLIAM R. HARVEY,A N D MOIRACIOFFE The ATP-Dependent Component of Gastric Acid Secretion G. SACHS,B. WALLMARK, G . SACCOMANI, E. RABON, D. R. DIBONA, A N D H. B. STEWART, T. BERGLINDH PART 111. REVERSIBILITY: ATP SYNTHESIS DRIVEN BY ELECTRIC FIELDS Effect of Electrochemical Gradients on Active H+ Transport in an Epithelium QAISAL-AWQATIA N D TROYE. DlXON Coupling between H + Entry and ATP Synthesis in Bacteria PETERC. MALONEY Net ATP Synthesis by H -ATPase Reconstituted into Liposomes YASUOKAGAWA Phosphorylation in Chloroplasts: ATP Synthesis Driven by A@ and by ApH of Artificial or Light-Generated Origin PETERGRABER +

PART I V , SOME THEORETICAL QUESTIONS Response of the Proton Motive Force to the Pulse of an Electrogenic Proton Pump ERICHHEINZ

Reaction Kinetic Analysis of Current-Voltage Relationships for Electrogenic Pumps in Neurospora and Acetabularia DIETRICHGRADMANN, ULF-PETER HANSEN,AND CLIFFORDL. SLAYMAN Some Physics of Ion Transport HAROLDI. MOROWITZ PART V. MOLECULAR MECHANISMS OF CHARGE SEPARATION An H -ATP Synthetase: A Substrate Translocation Concept I. A. KOZLOVAND V. P. SKULACHEV Proton Translocation by Cytochrome Oxidase MARTEN WIKSTROM Electrogenic Reactions of the Photochemical Reaction Center and the UbiquinoneCytochrome b/c2 Oxidoreductase P. LESLIE DU-ITON, PAUL MUELLER, DANIELP. O’KEEFE. NIGELK. PACKHAM, ROGERC. PRINCE,A N D DAVIDM .TIEDE Proton-Membrane Interactions in Chloroplast Bioenergetics R. A. DILLEY,L. J. PRCCHASKA, G. M. BAKER,N. E. TANDY,AND P. A. MILLNER Photochemical Charge Separation and Active Transport in the Purple Membrane BARRYHONK Mitochondria1 Transhydrogenase: General Principles of Functioning I . A. KOZLOV Membrane Vesicles, Electrochemical Ion Gradients, and Active Transport H. R. KABACK +

PART VI. BIOLOGICAL SIGNIFICANCE OF ELECTROGENIC ION PUMPS The Role of Electrogenic Proton Translocation in Mitochondria1 Oxidative Phosphorylation JANNAP. WEHRLE Electrogenic Reactions and Proton Pumping in Green Plant Photosynthesis WOLFGANGJUNGE

CONTENTS OF PREVIOUS VOLUMES

The Role of the Electrogenic Sodium Pump in Controlling Excitability in Nerve and Cardiac Fibers MARIOVASSAI.LE Pumps and Currents: A Biological Perspective FRANKLIN M. HAROLD Index

Volume 17 Membrane Llpldr of Prokaryotes Lipids of Prokaryotes-Structure and Distribution HOWARD GOLDFINE: Lipids of Bacteria Living in Extreme Environments THOMAS A. LANGWORTHY Lipopolysaccharides of Gram-Negative Bacteria O-rro LUDEKIZ, MARINAA. FREIJDENB~RG, VOLKER LEHMANN, CHRISGALANOS, ERNSTTH. RIETSCHEL,A N D DEREKH. SHAW Prokaryotic Polyterpenes: Phylogenetic Precursors of Sterols A N D MICHEL ROHMER GUYOURISSON Sterols in Mycoplasnia Membranes SHMUELRAZIN Regulation of Bacterial Membrane Lipid Synthesis 0. ROCK A N D CHARLES JOHN E. CRONAN, JR. Transbilayer Distribution of Lipids in Microbial Membranes SHLOMO ROITEM Lipid Phase Transitions and Regulation of Membrane Fluidity in Prokaryotes DONALDL. MEICHIOR Effects of Membrane Lipids on Transport and Enzymic Activities RONALDN. MCELHANEY Index

Volume 18 PART 1. ADENYLATE CYCLASE-RELATED RECEPTORS Hormone Receptors and the Adenylate Cyclase System: Historical Overview B. RICHARD MARTIN

The Elucidation of Some Aspects of Receptor Function by the Use of a Kinetic Approach A. M. TOLKOVSKY The P-Adrenergic Receptor: Ligand Binding Studies Illuminate the Mechanism 3f Receptor- Adenylate Cyclase Coupling JEFFREY M. STADELA N D ROBERTI . LEPKOWITZ Receptor-Mediated Stimulation and Inhibition of Adenylate Cyclase DERMOTM. F. C W P ~ K Desensitization of the Response of Adenylate Cyclase to Catecholamines JOHN P. PERKINS Hormone-Sensitive Adenylate Cyclase: Identity, Function, and Regulation of the Protein Components ELLIOTTM.Ross, STEENE. PtDERSEN, A N D VlNCtNT A. FI-ORIO The Regulation of Adenylate Cyclase by Glycoprotein Hormones BRIANA. COOKE The Activity of Adenylate Cyckase Is Regulated by the Nature of Its Lipid Environment MILESD. HOUSLAYA N D LARKYM. GORDON The Analysis of Interactions between Hormone Receptors and Adenylate Cyclase by Target Size Determinations Using Irradiation Inactivation B. RICHARD MARTIN PART 11. RECEPTORS NOT INVOLVING ADENYLATE CYCLASE Vasopressin Isoreceptors in Mammals: Relation to Cyclic AMP-Dependent and Cyclic AMP-lndependent Transduction Mechanisms S ~ K GJARD E Induction of Hormone Receptors and Responsiveness during Cellular Differentiation L. MICHAELC. LIN A N D SUZANNE BECKNER Receptors for Lysosomal Enzymes and GIycoproteins VIRGINIA SHEPHERD.PAUL SCHLESINGER, AND PHILIP STAHL The Insulin-Sensitive Hexose Transport System in Adipocytes J.-GLIEMANN A N D W. D. REES

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CONTENTS OF PREVIOUS VOLUMES Epidermal Growth Factor Receptor and Mechanisms for Animal Cell Division MANJUSRI DAS Thc Linkage between Ligand Occupation and Response of the Nicotinic Acetylcholine Receptor PALMERTAYLOR, ROBERTDALEBROWN, A N D DAVID A. JOHNSON The Interaction of Cholera Toxin with Gangliosides and the Cell Membrane SIMONVANHEYNINCEN Subject Index

Ultrastructure of Na,K-ATPase in Plasma Membrane Vesicles ELISABETH SKRIVER, ARVlD B M A U ~ S B A CAHN,D PETERLETH JBRGENSEN

Electron Microscope Analysis of Two-Dimensional Crystals of Membrane-Bound Na,KATPase ARVlD B. MAUNSBACH, ELISABETH SKRIVER, HANSHEBERT,A N D PETER LETH JBRGENSEN Organization of the Transmembrane Segments of Na,K-ATPase. Labeling of Lipid Embedded and Surface Domains of the a-Subunit and Its Tryptic Fragments with [1251]Iod0Volume 19 naphthylazide, [32P]ATP, and Photolabeled Ouabain PETER LETHJBRGENSEN, STEVEN J . D. PART I . THERMODYNAMIC ASPECTS OF KARLISH,A N D CARLOS GITLER MEMBRANE TRANSPORT Structural Studies on Lamb Kidney Na,KATPase What is a Coupled Vectorial Process? J . H. COLLINS,BLISSFORBUSH111, L. WILLIAM P. JENCKS K. LANE,E. LING,ARNOLDSCHWARTZ, The Membrane Equilibrium with Chemical Reactions AND A. (REEVES) ZOT FRIEDRICH A. SAUER Two Slightly Different a-Subunit Components of Kidney Na,K-ATPase Induced by Heat Treatment PART 11. STRUCTURAL ANALYSIS OF Na ,K-ATPase T. OHTA,M. KAWAMURA, T. AND K. HASECAWA, H. ISHIKURA, NAGANO Structural Aspects of Na,K-ATPase ROBERTL. POST Radiation Inactivation Analysis of Na,KDetergent Solubilization of Na,K-ATPase ATPase PAULOTTOLENGHI,J . CLIVEELLORY, MIKAELESMANN A N D ROGERA. KLEIN Methods for the Cleavage of the Large Subunit Stoichiometrical Binding of Ligands to L e s b of Na,K-ATPase and the Resolution of the than 160 Kilodaltons of Na,K-ATPase Peptides Produced H. MATSUI,Y. HAYASHI, HENRYRODRICUtZ, RICHARD HARKINS, A N D M. TACUCHI H. HOMAREDA, A N D JACK KYTE Selective Purification of Na,K-ATPase and The Active Site Structure of Na,K-ATPase: Ca2 ,Mg* -ATPase from Eel Electroplax Location of a Specific Fluorescein Isothiocyanate-Reactive Site L. M. AMENDE,S. P. CHOCK,A N D R. W. ALBERS CYNTHIA T. CARILLI,ROBERTA. High-Performance Gel Chromatography of FARLEY,AND LEWISC. CANTLEY Subunit Distribution of Sulfhydryl Groups and Horse Kidney Na,K-ATPase Disulfide Bonds in Renal Na,K-ATPase MAKOTONAKAO,TOSHIKONAKAO, M. KAWAMURA, T. OHTA,A N D K. TOMOKOOHNO,YOSHIHIRO FUKUSHIMA, NAGANO YUKICHI HARA,AND MASAKOARAI Native Membranes from Dog Kidney Outer Lipid Regions of Na,K-ATPase Examined with Medulla, Enriched in Na,K-ATPase, and VeFluorescent Lipid Probes A . MUCZYNSKI. WARDE. sicular in Nature KIMBERLY HARRIS,A N D WILLIAML. STAHL BLISSFORBUSHI11 +

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CONTENTS OF PREVIOUS VOLUMES

Role of Cholesterol and Other Neutral Lipids in Na.K-ATPase I . J. H . H . M. D ~ P o N TW. , H. M. PETERS.A N D S. L. BONTING

PART 111. LIGAND INTERACTIONS: CARDIAC GLYCOSIDES AND IONS Cardiotonic Steroid Binding to Na.K-ATPase Bi.iss FORBUSH Ill Binding of Monovalent Cations to the Na,KATPdse M. YAMAGUCHI, J . SAKAMOTO, A N D Y. TONOMURA Half-of-the-Sites Reactivity of Na,K-ATPase Examined by the Accessibility of Vanadate and ATP into Enzyme-Ouabain Complexes OTTO HANSEN Binding of Rb and ADP to a Potassium-Like Form of Na,K-ATPase A ND PAULOITOLENGHI J ~ R G EJENSEN N Side-Dependent Ion Effects on the Rate of Ouabain Binding to Reconstituted Human Red Cell Ghosts H. H. BODtMANN, T. J . CALLAHAN, H. RIXHMANN, A N D J . F. HOFFMAN lntracellular Sodium Enhancement of Ouabain Binding to Na,K-ATPase and the Development of Ciycoside Actions TAI A K ~ K AKYOSUKE , TEMMA.A N D SATOSHIYAMAMOTO Lithium-Catalyzed Ouabain Binding to Canine Kidney Na,K-ATPase GEORGER. HtNUtRSON Ouabain Binding and Na,K-ATPase in Resealed Human Red Cell Ghosts D. G. SHOEMAKER A N D P. K. LAUF Stereoelectronic Interaction between Cardiotonic Steroids and Na,K-ATPase: Molecular Mechanism of Digitalis Action F. DirrRiCH, P. BERLIN,K. KOPKE,A N D K. R. H. REPKE Use of Prophet and MMS-X Computer Graphics in the Study of the Cardiac Steroid Receptor Site of Na,K-ATPase DOUGLASC. DWIGHTS . FULLERTON. ROHRER,KHAISI.AHMED.ARTHURH. L. FROM,EITAROKITATSUJI, AND TAMBOUE DEFFO +

Photoaffinity Labeling of the Ouabain Binding Site of Na.K-ATPase CLIFFORDC. HALL.A N D ARNOLDE. RUOHO New Ouabain Derivatives to Covalently Label the Digitalis Binding Site BERNARD Rossi. MAURICE GOH.DNER. GILLESPONZIO.CHRISTIAN HIRTH,A N D MICHELLAZDUNSKI Ouabain Sensitivity: Diversity and Disparities JOHNS . WILLISA N D J. CLivt ELLONY PART IV: LIGAND INTERACTIONS: NUCLEOTIDES, VANADATE, AND PHOSPHORY LATION Ligand Interactions with the Substrate Site of Na,K-ATPase: Nucleotides, Vanadate, and Phosphorylation Jens G. Norby Conformational Changes .of Na,K-ATPase Necessary for Transport LEWIS C. CANTLEY.CYNTHIA T. CARILLI,RODERICL. SMITH.A N D DAVID PERLMAN On rhe Mechanism behind the Ability of Na,K-ATPase to Discriminate between Na+ and K + JENS CHR.SKOU Characteristics of the Electric Eel Na,K-ATPase Phosphoprotein YODA ATSUNOBU YODAA N D SHIZUKO Sulfhydryl Groups of Na,K-ATPase: Effects of N-Ethylrnaleimide on Phosphorylation from ATP in the Presence of Na+ + MgZ+ MIKAELESMANNA N D IRENAKLODOS Alternative Pathways of Phosphorylation of Na,K-ATPase Regulated by Na+ Ions on Both Sides of the Plasma Membrane HORSTWALTER Structurally Different Nucleotide Binding Sites in Na,K-ATPase HERMANN KOEPSELLA N D DORISOLLIO Study of Na,K-ATPase with ATP Analogs PAULS, WILHELM SCHONtR, HARTMUT ENGINH. SERPERSU,GEROLD REMPETERS, ROSEMARIE PATZELTWBNCZLER,A N D MARIONHASSELBERG Affinity Labeling Studies of the ATP Binding Site of Canine Kidney Na,K-ATPase

CONTENTS OF PREVIOUS VOLUMES

JAMES B. COOPER,CARLJOHNSON,AND CHARLESG. WINTER 3’P[180]NMRKinetic Analysis of ‘*O Exchange Reaction between P, and H 2 0 Catalyzed by Na,K-ATPase A. STEPHENDAHMSAND JOELLE E. MIARA

PART V. CONFORMATIONAL CHANGES, STRUCTUREIFUNCTION, AND ACTIVE SITE PROBES Principal Conformations of the a-Subunit and Ion Translocation PETER L. J0RGENSEN Magnesium-Induced Conformational Changes in Na,K-ATPase S. L. BONTING, H. G. P. SWARTS,W. H. M. PETERS,F. M. A. H. A N D J. I . H. SCHUURMANS STEKHOVEN, H. M. DE PONT Rubidium Movements in Vesicles Reconstituted with Na,K-ATPase, Measured in the Absence of ATP and Pi, in the Presence of Either Ligand, and in the Presence of Both Ligands: Role of the “Occluded State” in Allowing for the Control of the Direction of Ion Movements S . J. D. KARLISHA N D W. D. STEIN Eosin: A Fluorescent Probe of ATP Binding to Na,K-ATPase I. C. SKOUAND MIKAELESMANN Interaction of Divalent Cations with Fluorescein-Labeled Na,K-ATPase MARCIASTEINBERG, JAMESG . C. SEN KAPAKOS,AND PARIMAL Cation Activation of Na,K-ATPase after Treatment with Thimerosal MANISHA D. MONEAND JACKH. KAPLAN Alteration of Conformational Equilibria in Na,K-ATPase by Glutaraldehyde Treatment DAVIDM. CHIPMAN,E. ELHANANY, R. BERGER,AND A. LEV Conformational Transition between ADP-Sensitive Phosphoenzyme and Potassium-Sensitive Phosphoenzyme KAZUYATANIGUCHI, KUNIAKISUZUKI, AND SHOICHI IIDA

589 Relation between Red Cell Membrane Na,KATPase and Band 3 ERICT. FOSSELA N D A. K. SOLOMON PART V1. REACTION MECHANISM AND KINETIC ANALYSIS Kinetic Analyses and the Reaction Mechanism of the Na,K-ATPase JOSEPHD. ROBINSON Evidence for Parallel Pathways of Phosphwnryme Formation in the Mechanism of ATP Hydrolysis by Elecrrophorus Na,KATPase JEFFREYP. FROEHLICH, ANN S . HOFIBS, AND R. WAYNEALBERS Evaluation of the Reaction Mechanism of the Sodium Pump by Steady-State Kinetics JOHNR. SACHS Kinetic Evidence in Favor of a Consecutive Model of the Sodium Pump D. A. EISNERAND D. E. RICHARDS Kinetic Models of Na-Dependent Phosphorylation of Na,K-ATPase from Rat Brain STANLEYJ. DONALDM. FOSTER, RUSSELL,A N D KHALILAHMED Reinvestigation of the Sequence of Sensitivity of Phosphoenzyme of Na,K-ATPase to ADP and K + during the Presteady State of the Phosphorylation by ATP A N D M. NAKAO Y. FUKUSHIMA Interaction of Na+ , K + , and ATP with Na,KATPase P. J. GARRAHAN, R. Rossi, AND A. F. RECA Sodium Ion Discharge from Pig Kidney Na,KATPase YUKICHIHARAAND MAKOTONAKAO ADP Sensitivity of the Native and Oligomycin-Treated Na,K-ATPase ANNS . HOBBS,R. WAYNEALBERS,AND JEFFREYP. FROEHLICH Three (at Least) Consecutive Phosphointermediates of Na-ATPase 1. KLODOS,J. G . N0RBY, A N D N. 0. CHRISTIANSEN Aspects of the Presteady State Hydrolysis of ATP by Na,K-ATPase A. G. LOWEAND L. A. REEVE Identity of the Na Activation Sites in ATPase

590

CONTENTS OF PREVIOUS VOLUMES

with the K Activation Sites in p-Nitrophenylphosphatase L. A. PAKODI. J. F. PINCUS, L. D.J. SORCt, A N D s. R JOSEPHSON, SIMON On the Existence of Two Distinct Hydrolysis Cycles for Na.K-ATPase with Only One Active Substrate Site 1GOR w. PLtSNtK Kinetic Analysis of the Effects of Na+ and K + on Na,K-ATPase LISELOTTE PLESNCKA N D IGOR W. PLESNER Divalent Cations and Conformational States of Na.K- ATPase J u s t p H L). ROBINSON PART VII. ION TRANSLOCATION AND REACTION MECHANISM Na.K-ATPase: Reaction Mechanisms and Ion Translocating Steps PAULD t W E ~ K Existence and Role of Occluded-Ion Forms of Na.K-ATPase 1. M. GLYNNA N D D. E. RICHARDS Na and K Fluxes Mediated by ATP-Free and ATP- Activated Na.K- ATPase in Liposomes BEATRICE M. ANNtK Sidedness of Cations and ATP Interactions wlth the Sodium Pump L. BEAU& A N D R. DIPOLO Sidedness of Sodium Interactions with the Sodium Pump in the Absence of K + RHODABLOSTEIN Magnesium Dependence of Sodium Pump-Mediated Sodium Transport in Intact Human Red Cells P. W. FLATMAN A N D V. L. LEW K -Independent Active Transport of Na by Na,K-ATPase MICHAELFORGACA N D GILBERTCHIN ADP-ATP Exchange in Internally Dialyzed Squid Giant Axons PAULDE WEtR, G t R D A E. AND BREITWIESER. BRIANG. KENNEDY, H.GILBERTSMITH Sodium Pump-Catalyzed ATP-ADP Exchange in Red Blood Cells: The Effects of Intracellular and Extracellular Na and K Ions JACK H. KAPLAN +

+

Ouabain-Sensitive ATP-ADP Exchange and Na-ATPase of Resealed Red Cell Ghosts J . D. CAVIERES Effect of Internal Adenine Nucleotides on Sodium Pump-Catalyzed Na-Na and Na-K Exchanges B R I A NG . KENNEDY, GORML U N N A, N D JOSEPH F. HOFFMAN Na/K Pump in Inside-Out Vesicles Utilizing ATP Synthesized at the Membrane W. MERCER.BEVERLEY E. RORKRT A N D PHILIP B. DUNHAM FARQUHARSON, Anion-Coupled Na Efflux Mediated by the Na/K Pump in Human Red Blood Cells S. DISSINGA N D J. F. HOFFMAN Effect of Trypsin Digestion on the Kinetic Behavior of the Na/K Pump in Intact Erythrocytes DONNAL. KROPP Sodium Movement and ATP Hydrolysis in Basolateral Plasma Membrane Vesicles from Proximal Tubular Cells of Rat Kidney F . PROVERBIO. T. P K O V ~ R B IAON D, R . MAR(N Stoichiometry of the Electrogenic Na Pump in Barnacle Muscle: Simultaneous Measurement of Na Efflux and Membrane Current M. T. NELSONA N D W. J. LEDERER PART VIII. BIOSYNTHESIS, MULTIPLE FORMS. AND IMMUNOLOGY Regulation of Na,K-ATPase by Its Biosynthesis and Turnover NORMAN1. K A R I N A N D JOHN s. COOK Biosynthesis of Na,K-ATPase in MDCK Cells J. SHERMAN, T. MORIMOTO.A N D D. D. SABATINI Possible Functional Differences between the Two Na,K-ATPases of the Brain KATHLEEN J. SWEADNER Antigenic Properties of the cx, p, and y Subunits of Na,K-ATPase WILLIAMBALL,JR., JOHNH. COLLINS, L. K. LANE,AND ARNOLDSCHWARTZ Antibodies to Na,K-ATPase: Characterization and Use in Cell-Free Synthesis Studies ALICIAMCDONOUGH,ANDREWHIAT, AND ISIWREEDELMAN Immunoreactivity of the a- and a(+)-Subunits

CONTENTS OF PREVIOUS VOLUMES

of Na, K-ATPase in Different Organs and Species IRENEV. GERARD D. SCHELLENBERG, PECH,A N D WILLIAML. STAHL Role of Na+ and Ca* + Fluxes in Terminal Differentiation of Murine Erythroleukemia Cells I. G. MACARA, R. D. SMITH,AND LEWIS C. CANTLEY NaiK Pumps and Passive K + Transport in Large and Small Reticulocytes of Anemic Low- and High-Potassium Sheep P. K. LAUFA N D G. VALET Enhancement of Biosynthesis of Na,K-ATPase in the Toad Urinary Bladder by Aldosterone But Not T3 K . GEERING, M. GIRARDET, C. BRON, A N D B. C. ROSSIER J.-P. KRAEHENBUHL, Na,K-ATPase Activity in Rat Nephron Segments: Effect of Low-Potassium Diet and Thyroid Deficiency LAL C. GARGA N D C. CRAIGTISHER Axonal Transport of Na,K-ATPase in Optic Nerve of Hamster SUSANC. SPECHT PART IX. Na,K-ATPase AND POSITIVE INOTROPY; ENDOGENOUS GLYCOSIDES Positive Inotropic Action of Digitalis and Endogenous Factors: Na,K-ATPase and Positive Inotropy; “Endogenous Glycosides” ARNOLDSCHWARTZ Endogenous Glycoside-Like Substances GARNERT. HAUPERT,JR. Monovalent Cation Transport and Mechanisms of Digitalis-Induced lnotropy THOMASW. SMITHAND WILLIAMH. BARRY Effects of Sodium Pump Inhibition on Contraction in Sheep Cardiac F’urkinje Fibers D. A. EISNER,W. J. LEDERER,AND R. D. VAUGHAN-JONES Quantitative Evaluation of [3H]Ouabain Binding to Contracting Heart Muscle, Positive Inotropy, Na,K-ATPase Inhibition, and 86Rb Uptake in Several Species ERLANDERDMANN, LINDSAYBROWN, KARLWERDAN,AND WOLFGANG KRAWIETZ +

Contractile Force Effects of Low Concentrations of Ouabain in Isolated Guinea Pig, Rabbit, Cat, and Rat Atria and Ventricles GUNTERGRUPP.INGRIDL. GRUPP.J. GHYSEL-BURTON, T. GODFRAIND, A. DE POVER,AND ARNOLDSCHWARTZ Difference of Digitalis Binding to Na,KATPase and Sarcolemma Membranes I. KUROBANE, D. L. NANDI,A N D G .T. OKITA Pharmacological and Biochemical Studies on the Digitalis Receptor: A Two-Site Hypothesis for Positive Inotropic Action INGRID L. GRUPP, ARNOLDSCHWARTZ, ROBERTJ. ADAMS,TREVORPOWELL, GUNTERGRUPP,AND E. T. WALLICK Hypothesis for the Mechanism of Stimulation of the Na/K Pump by Cardiac G l y c o s i d e s Role of Endogenous Digitalis-Like Factor T. GODFRAIND, G. CASTANEDAJ. GHYSEL-BURTON, AND HERNANDEZ, A. DE POVER Immunochemical Approaches to the Isolation of an Endogenous Digoxin-like Factor KENNETHA. GRUBER,JANICEM. AND VARDAMAN M. WHITAKER, BUCKALEW, JR. Demonstration of a Humoral Na/K Pump Inhibitor in Experimental Low-Renin Hypertension MOTILALPAMNANI, STEPHENHUOT, DAVIDCLOUGH,JAMESBUGGY,AND FRANCISJ. HADDY Absence of Ouabain-Like Activity of the Na,K-ATPase Inhibitor in Guinea Pig Brain Extract GEORGER. KRACKE Brain Na,K-ATPase: Regulation by Norepinephrine and an Endogenous Inhibitor ALANC. SWANN Inhibitory and Stimulatory Effects of Vanadate on Sodium Pump of Cultured Heart Cells from Different Species KARLWERDAN,GERHARD BAURIEDEL, WOLFGANG KRAWIETZ,AND ERLAND ERDMANN Endogenous Inhibitor of Na,K-ATPase: “Endodigin” K. R. WHITMER,D. EPPS, AND ARNOLD SCHWARTZ

CONTENTS OF PREVIOUS VOLUMES

PART X. PHYSIOLOGY AND PATHOPHYSIOLOGY OF THE Na/K PUMP Disorders in Molecular Assemblies for Na Transport in Essential Hypertension MITZYL. CANESSA, NOHMAC. ADRACNA, ISABELBIZE,HAROLD A N D DANIEL C. TOSTESON SOLOMON, The Na-K Cotransport System in Essential Hypertension C. NAZARET, A N D P. R. P. GARAY, HANNAERT Loss of Na,K-ATPase Activity ‘during Cataract Formation in Lens c. StN A N D DOUGLASR . PARIMAL PFEIFPER NaiK Pump: Effect of Obesity and Nutritional State M. DtLuISt, P. U S H E R , A N D J . FLIER Decreased Na.K-ATPase Activity in Erythrocyte Membranes and Intact Erythrocytes from Obese Man DAVIDM Mom. IWAR KLIMES,A N U RANVIL L. CLARK Functionally Abnormal Na/K Pump in Erythrocytes from a Morbidly Obese Subject J. FLIER,P. USHER, A N D M. DELUISF. Specific Insulin Binding to Purified Na,KATPase Associated with Rapid Activation of the Enzyrnc JULIEE. M. MCCEXKH Mechanism for Cholinergic Stimulation of Sodium Pump in Rat Submandibular Gland A N D AMARKK. SEN DAVIDI. STEWART Evidence for an Aldosterone-Mediated, NaDependent Activation of Na,K-ATPase in the Cortical Collecting Tubule KEVINJ. PEITY, JUHA P. KOKKO,AND DIANAMARVEK Vanadate and Somatostatin Having Divergent Effects on Pancreatic Islet Na,K-ATPase K t N J l IKEJlRl A N D SEYMOUR R. LEVIN Phosphorylation of a Kidney Preparation of Na,K-ATPase by the Catalytic Subunit of CAMP-Dependent Protein Kinase SVtN MARDH Modulation of Na,K-ATPase Activity in Rat Brain by Adenosine 3’,5’-Monophosphate A N D AMARK. RUSSELLB. LINGHAM

sBN

Stimulation and Inhibition by Plasma of Ouabain-Sensitive Sodium Efflux in Human Red Blood Cells A. R. CHIPPERFIELD Inhibition of the Na Pump by Cytoplasmic Calcium in Intact Red Cells A. M. BROWNA N D V . L. L t w Involvement of Calmodulin in the Inhibition of Na,K-ATPase by Ouabain E , T. PIASCIK, LlONtL 0. L E L I ~ V RM. J. D. P O ~ E RE., T . WALLICK.A N D ARNOLDSCHWARTZ Index

Volume 20 PART I. FREQUENCY DOMAIN ANALYSIS OF ION TKANSPORT Fluctuation Analysis of Apical Sodium Transport

T. HOSHIKO Impedance Analysis of Necturus Gallbladder Epithelium Using Extra- and Intracellular Microelectrodes J. J. LIM, C. KOTTRA, A N D E. FRBMTER L. KAMPMANN. Membrane Area Changes Associated with Proton Secretion in Turtle Urinary Bladder Studied Using Impedance Analysis Techniques A N D TROY E. DIXON CHRISCLAUSEN Mechanisms of lon Transport by the Mammalian Colon Revealed by Frequency Domain Analysis Techniques N. K. WILLS Analysis of Ion Transport Using Frequency Domain Measurements SIMON A. Lewis AN[) WII.I.IAMP. ALLES Use of Potassium Depolarization to Study Apical Transport Properties in Epithelia LAWRENCEG. P A L M E R PART I I . USE OF ANTIBODIES T O EPlTH ELI A L M EM BRAN E PROTEINS Biosynthesis of Na+,K+-ATPasein Am phibian Epithelial Cells B. C . ROSSIER

CONTENTS OF PREVIOUS VOLUMES

Use of Antibodies in the Study of Na+,K+ATPase Biosynthesis and Structure ALICIAA. MCDONOUGH Encounters with Monoclonal Antibodies to Na+,K+-ATPase MICHAEL KASHGARIAN, DANIEL BIEMESDERFER, AND BLISSFORBUSH Ill Monoclonal Antibodies a s Probes of Epithelial Cell Polarity GEORGEK. OJAKIAN A N D DORISA. HERZLINCER Immunolabeling of Frozen Thin Sections and Its Application to the Study of the Biogenesis of Epithelial Cell Plasma Membranes IVANEMANUILOV IVANOV, HEIDE PLESKEN,DAVIDD. SABATINI, AND MICHAEL J. RINDLER Development of Antibodies to Apical Membrane Constituents Associated with the Action of Vasopressin JAMESB. WADE,VICTORIA GUCKIAN, AND INGEBORG KOEPPEN Molecular Modification of Renal Brush Border Maltase with Age: Monoclonal Antibody-Specific Forms of the Enzyme BERTRAMSACKTOR A N D UZI REISS PART 111. BIOCHEMICAL CHARACTERIZATION OF TRANSPORT PROTEINS Sodium-~-Glucose Cotransport System: Biochemical Analysis of Active Sites R. KINNE,M. E. M. DA CRUZ,A N D J. T. LIN Probing Molecular Characteristics of Ion Transport Proteins DARRELL D. FANESTII., RALPHJ. KESSLER,A N D CHUNSIKPARK Aldosterone-Induced Proteins in Renal Epithelia MALCOLM Cox A N D MICHAEL GEHEB

Development of an Isolation Procedure for Brush Border Membrane of an Electrically Tight Epithelium: Rabbit Distal Colon MICHAELC. GUSTINA N D DAVIDB. P. GOODMAN Index Volume 21

ION CHANNELS: MOLECULAR AND PHYSIOLOGICAL ASPECTS Ionic Selectivity of Channels at the End Plate PETERH. BARRYA N D PETERW. GAGE Gating of Channels in Nerve and Muscle: A Stochastic Approach RICHARDHORN The Potassium Channel of Sarcoplasmic Reticulum CHRISTOPHER MILLER,JOANE. BELL, A N D ANAMARIAGARCIA Measuring the Properties of Single Channels in Cell Membranes H.-A. KOLB Kinetics of Movement in Narrow Channels DAVIDG. LEVITT Structure and Selectivity of Porin Channels R. BENZ Channels in the Junctions between Cells WERNERR. LOEWENSTEIN Channels across Epithelial Cell Layers SIMONA. LEWIS,JOHNW. HANRAHAN,A N D W. VANDRIESSCHE Water Movement through Membrane Channels ALANFINKELSTEIN Channels with Multiple Conformational States: Interrelations with Carriers and Pumps P. LAUGER Ion Movements in Gramicidin Channels S. B. HLADKYA N D D. A. HAYDON

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