International Review of Neurobiology Volume 24

INTERNATIONAL REVIEW OF

Neurobiology VOLUME 24

Editorial Board W. Ross ADEY

SEYMOUR KE~V

JCLICS AXELROD

KEII H KILLAM

Ross RALDESSARINI

CON AN

SIR ROGER B.L\NNISTER

ABELLAITHA

FLOYD BLOOM

BORISL F . B E D ~ V

DANIEL. Boi'E'r

PAVL MANDELL

PHILLIPBRADLEY

HUMPHRY OSMONI)

JOSE

DELGADO

KOR~ETSKY

RODOLFOPAOLETT1

SIRJOHX E(:CI,LS

SOl.OMON SNYDEK

JOEI+ ELKS

STEPHENSZARA

H. J .

JOHN VANE

EYSESCK

KJELL Fort

MARAT \!+RlANlAN

Bo HOLMSrEr)r

RICHARDW'I'Ar-r

PAULJ A Y S S E : ~

OLI71 k

. Z~ AN GWI LI.

INTERNATIONAL REVIEW OF

Neurobiology Edited by JOHN R. SMYTHIES Deportment of Psychiatry ond The Neurosciences Program University of Alabomo Medical Center Birminghorn, Alabomo

RONALD J. BRADLEY The Neurosciences Program University of Alabama Medical Center Birrninghom, Alobomo

VOLUME 24

ACADEMIC PRESS A Subsidiary of Harcourt Brace Jovonovich, Publishers

New York London Pork Son Diego Son Francisco SBo Paul0 Sydney Tokyo Toronto

COPYRIGHT @ 1983, BY ACADEMIC PRESS, INC. ALL RIGHTS RESERVED. NO PART OF THIS PUBLICATION MAY BE REPRODUCED OR TRANSMITTED IN ANY F O R M OR BY ANY MEANS, ELECTRONIC OR MECHANICAL, INCLUDING PHOTOCOPY, RECORDING, OR ANY INFORMATION STORAGE AND RETRIEVAL SYSTEM, WITHOUT PERMISSION IN WRITING FROM THE PUBLISHER.

ACADEMIC PRESS, I N C . 111 Fifth Avenue, New York. New York 10003

United Kingdom Edition published by ACADEMIC PRESS, I N C . ( L O N D O N ) LTD. 24/28Oval Road, London N W l 7DX

LIBRARY OF

59- 13822

CONGRESS CATALOG CARD NUMBER:

I S B N 0-12-366824-7 PRINTED IN THE UNITED STATES OF AMERICA

83 84 8s 86

9 816 5 4 3 2 1

CONTENTS ..................................................................... CONTRIBUTORS

ix

Antiacetylcholine Receptor Antibodies and Myasthenia Gravis

BERNARD W. FULPIUS 1. Introduction . . . . . . . . . . . . . . 11. Pathogenicity of Circulating Anti-nAcChR Antibodies 111. Assays for Circulating Anti-n

................... 1 ...................... 2 ................... 5

IV. Anti-nAcChR Antibody Concentration in Different Forms of Myasthenia Gravis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Anti-nAcChR Antibodies in the Cerebrospinal Fluid ....................... VI. Antigenic Determinants on nAcChR . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

7 10

12 13

Pharmacology of Barbiturates: Electrophysiological and Neurochemical Studies

MAX WILLOW

AND

GRAHAMA. R.JOHNSTON

I. Introduction ..... .............................. 11. Neuropharmacolog 111. Biochemical and Neurochemical Studies . .................... 1V. Conclusions ............................................................ References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

15 16 34

44 45

lmrnunodetection of Endorphins and Enkephalins: A Search for Reliability

ALEJANDRO BAYON.WILLIAM J. SHOEMAKER, JACQUELINE F. McGiwY. AND FLOYI)BLOOM

........................................................... .............................. .............................. IV. Is Immunodetection Reliable? ............................................ I. Introduction

11. Tissue Processing, Extraction, and Handling 111. Identification, Quantitation, and Localization

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

51 53 62 87 88

On the Sacred Disease: The Neurochemistry of Epilepsy

0. CARTERSNEADi n I. Introduction

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

11. Epilepsy: The Diversity of the Problem

94 94

vi

CONTENTS

111. Neurophysiology of Seizures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I\'. Models . . . . . . . . . . . . . . . . .................... V. Neurotransmitters and Ot Seizures .......... VI. Developmental Considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ...................... References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

95 99 106 150 152 152

Biochemical and Electrophysiological Characteristics of Mammalian GABA Receptors SrZLVATORE J. ENNA A N D JOEL

P.

GALLACHER

I. Introduction . . . . . . 11. Electrophysiological

I l l . Biochemical Studies IV. Summary and Conclusions.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

181 182 192 204 2O(i

Synaptic Mechanisms and Circuitry Involved in Motoneuron Control during Sleep

MICHAELH. CHASE I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

213

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

215

IV. Motoneuron Membrane Potential during

akefulness . . . .

during Active Sleep . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V1. Central Control Mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Concluding Remarks .......................... ......... VIII. Summary Statements . . .............. References . . . . . . . . . . . .............. .....................

232 240 251 257

Recent Developments in the Structure and Function of the Acetylcholine Receptor

F. J .

B.4RRANTES

1. Introduction .... ..................... 11. The AChR Molecule . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Biosynthesis of the AChR .... ....................... IV. Three-Dimensional Topography of the AChR in the Membrane . . . . . . . . . . . V. In Search of the Functional Role of the Nonreceptor pProteins . . . . . . . . . . . Vl. The Ion-Translocation Function in Membrane-Bound AChR . . . . . . . . . . . . . . VII. Summary and Perspectives . . ................................ References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

259 261 272

279 290 301 329 33 1

CONTENTS Characterization of

a,-

vii

and ap-Adrenergic Receptors

DAVID B . BYLUNDAND DAVIDc. U'PRICHARD ............................... ........................ .................. 111. ap-Adrenergic Receptors ......................... IV. Summary and Conclusions .............................................. References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I. Introduction

11. a,-Adrenergic Receptors

344 354 420 422

Ontogeneis of the Axolemma and Axoglial Relafionships in Myelinated Fibers: Electrophysiological and Freeze-Fracture Correlates of Membrane Plasticity

STEPHENG. WAXMAN. JOEL A. BLACK,AND ROBERTE. FOSTER I. Introduction . . . . . . . . . . .

434 437 440 Development of the Optic Nerve Freeze-Fracture Structure of Myelinated Axons ........................... 449 Freeze-Fracture Studies on Myelin Development in Optic Nerve Axons . . . . . 461 Differentiation of the Axon Membrane in the Absence of Myelin .......... 475 Concluding Comments ................................................. 479 References ............................................................ 48 1

11. Specificity in Myelination

111. IV. V. VI. VII.

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

INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . CONTENTS OF RECENT VOLUMES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

485 49 1

This Page Intentionally Left Blank

CONTRIBUTORS Numbers in parentheses indicate the pages on which the authors' contributions begin.

F. J. BARRANTES, Max-Planck-Institutfur Biophysikalische Chemie, Gottingen-Nikolausberg, Federal Republic of Germany (259) ALEJANDROBAY ON,^ Departamento de Neurociencias, Centro de Investigaciones en Fisiologza Celular, Universidad Nacional Autonoma de Mexico, Mexico D.F., Mexico (51)

A. BLACK,^ Department of Neurology, Stanford University School of Medicine, and Veterans Administration Medical Center, Palo Alto, California 94304 (433)

JOEL

FLOYDBLOOM,A. V. Davis Center f o r Behavioral Neurobiology, The Salk Institute, San Diego, California 92138 (51) DAVIDB. BYLUND, Department of Pharmacology, School of Medicine, University of Missouri at Columbia, Columbia, Missouri 65212 (343) MICHAELH. CHASE,Brain Research Institute and Departments of Physiology and Anatomy, School of Medicine, University of California, Los Angeles, Los Angeles, California 90024 (213)

SALVATORE J. E N N A ,Departments of Pharmacology and of Neurobiology and Anatomy, University of Texas Medical School, Houston, Texas 77025 (181)

ROBERTE. FOSTER,Neurotoxicology Branch, U S . Army Medical Research Institute of Chemical Defense, Aberdeen Proving Ground, Maryland 21010 (433) BERNARDW. FULPIUS, Department of Biochemistry, University of Geneva, Geneva, Switzerland (1) JOEL P. GALLAGHER,Department of of Texas Medical Branch, Galveston,

Pharmacology and Toxicology, University Texas 77550 (181)

GRAHAM A. R. JOHNSTON, Department of Pharmacology, University of Sydney, New South Wales 2006, Australia (15) 'Present address: Instituto de Investigaciones Biomedicas, Universidad Nacional Autonoma de Mexico, Apartado Postal 70-228, 04510 Mexico D. F., Mexico. 2Present address: Department of Biological Sciences, Northern Illinois University, DeKalb, Illinois 60115. ix

X

(:ONTRI BUTORS

JACQI'ELINE E MCCINTE'. A . C'. Dailis Ceiiter f o r Behavioral Neurobiology, The Sulk institute, Sari Diego, Califoritin 92138 (51)

WILLIAM J. SHOEh1.lKF.R. A . .'I Dnsis Cenlerfor Behaz~ioral(Vepi1-obiolqgy,l h e Salk l,isfitute, Sail Diego, Califbrrtia 92138 (51)

0. C : i \ R T t R SXEAI)1 1 1 . Department (fPedintiicr arid Neurology, Unzuersity of Alubama i n Birmingham Sciiool of Medicine, Bi?ini@ani, Alabama 35294 (93)

INTERNATIONAL REVIEW OF

Neurobiology VOLUME 24

This Page Intentionally Left Blank

ANT IAC ETYLCHOLlNE RECEPTOR ANT I BODIES AN D MYASTHENIA GRAVIS By Bernard W. Fulpiur Dopa~tmontof B k h o r n i r t y

Univonity of Gomvo

Gonova, Switurlond

I. Introduction

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

11. Pathogenicity of Circulating Anti-nAcChR Antibodies

111.

IV. V.

VI.

A. Pathogenic Mechanisms of Free Antibodie B. Pathogenic Mechanisms of Complexed An Assays for Circulating Anti-nAcChR Antibodies ......................... A. Immunoprecipitation Assa-rs ........................................ B. Inhibition Assays ....................................... ..... Anti-nAcChR Antibody Concentration in Different Forms of Myasthenia Gravis .................................................. Anti-nAcChR Antibodies in the Cerebrospinal Fluid ..................... A. Antibody Origin ................................................... B. Cross-Reactivity with Brain nAcChR ................................ C. Antibody Pathogenicity ............................................ Antigenic Determinants on nAcChR ....................... A. Torpedo Fish nAcChR ......... ....................... B. Human Skeletal Muscle nAcChR.. .................................. References ................................... ...............

1 2 3 4

5

6 6

7 10 10 11

I1 12 12 13 13

1. lntrodwtion

Myasthenia gravis is a human muscle disease characterized by weaknes8sand abnormal fatigability of voluntary muscles with recovery of motor power on resting, as well as positive response to treatment with ant icholinesterase drugs. T h e basic defect appears to consist in a reduction of available nicotinic acetylcholine receptors (nAcChR) at neiiromuscular junctions, brought about by an antibody-mediated autoimmune reaction. The elucidation of this defect has followed detailed studies of the molecular organization of nicotinic cholinergic synapses, made possible by the development and the application of a set of tools, the neurotoxins from elapid snake venoms, used to identify specifically nAcChR.' I n this context, one should emphasize two particular neurotoxins of very wide use: ( I ) a-bungarotoxin (a-BuTx) mainly used as an iodinated derivative to label specifically nAcChR because of its extremely slow dissociation rate, and (2) a-cobra, linked to Sepharose beads, mainly used to purify nAcChR by affinity chromatography. 1 INTERNATIONAI. REVIEW O F NEIIROBIOLOGY. VOL. 24

Cnpynght 0 1983 by Acadcmx Press, Inr ,411 righis of rcproduction in any form reserved ISBN 0-12-366824-7

2

BERNARD W. FULPIUS

Obviously both humoral and cellular immunity to nAcChR are implicated in the pathogenesis of myasthenia gravis. There is little evidence about their relative role in the defect of the neuromuscular transmission, although humoral immunity, in the form of anti-nAcChR antibodies alone or in conjunction with complement factors, would be sufficient to cause a reduction of available receptors at neuromuscular junctions. As exhaustive reviews on the subject, including data on experimental autoimmune myasthenia gravis, have been published (Lindstrom, 1979; Vincent, 1980; Drachman, 1981), we shall restrict ourselves to a discussion of some of the basic questions one may ask in an effort to understand the mode of action of anti-nAcChR antibodies in this disease. 1. How can these specific antibodies be pathogenic? This question pertains to the diffusion of immunoglobulins in compartments other than the vascular one, the interaction of immunoglobulins with the antigen, and the consequences of complex formation with the receptor. 2. How can these specific antibodies be detected? This question is related to the heterogeneity of the population of anti-nAcChR antibodies and the limitations of the different methods at disposal to test them. 3. To what extent does the concentration of circulating anti-nAcChR reflect the severity of myasthenia gravis? This question must be raised because a lack of correlation between the measured titers and the observed clinical status has been noticed. 4. Why d o these specific antibodies occur within the cerebrospinal fluid? This question concerns the origin, central or peripheral, of these antibodies, their possible specificity for brain nAcChR, and their pathogenicity. 5 . What is known of the structure of nAcChR? This question refers to the receptor considered as the autoantigen in myasthenia gravis. More specifically, it deals with the existence of different antigenic determinants, involved or not in the primary autoimmune reaction.

il. Pathogenicity of Circulating Anti-nAcChR Antibodies

According to Lefvert et al. (1978), the synthesis of anti-nAcChR antibodies is triggered by antigenic stimuli, and the antibodies are not a primary cause of myasthenia gravis. These authors postulate that the early release of nAcChR (probably from damaged endplates or myoid cells within the thymus) could act as the primary antigenic stimulus. This

ANTIACETYLCHOLINE RECEPTOR ANTIBODIES

3

is quite possible, although nothing is known of the factors that would have led to synaptic or cellular damages. As an alternative proposal, one should consider a more complex sequence of events. Namely, the neoplastic development of an anti-nAcChR antibody-producing clone could initiate locally complement activation on the postsynaptic membrane and cause the liberation of nAcChR-containingfragments into the surrounding tissue and circulation. This would then present the receptor as well as asher antigens from skeletal muscle tissue to the immune system. Additional antibody responses would be generated, among others, to normally unexposed or inaccessible nAcChR antigenic determinants? In any case, anti-nAcChR antibodies are present, often in large amounts, in the vascular compartment. A. PATHOGENICMECHANISMS OF FREEANTIBODIES

j[n order to cause synaptic dysfunction, antibodies must leave the vascular compartment, diffuse into the extracellular space, enter the narrow synaptic cleft, and reach the receptor molecules located at the top of the postsynaptic folds. To test the accessibility of nAcChRs located in the neuromuscular junctions, complexes made of 12sI-labeleda-BuTx covidently coupled to unspecific IgG were injected into mice (Zurn and Fullpius, 1976). In such experimental conditions, the region of nerve terminals appeared labeled. This was considered by the authors as a sufficient proof that molecules of about 150,000 MW could indeed enter the synaptic cleft. This result was confirmed by another set of experiments in which 1251-labeledanti-a-cobra toxin antibodies injected into mice that had received sublethal doses of a-cobra toxin were shown to reach the toxin molecules bound to the nerve terminals. Circulating anti-nAcChR antibodies can be pathogenic by interacting in situ with the receptor, hence impairing its specific role in synaptic transmission. According to Engel et al. (1977), the resulting synaptic dysfunction can be caused by three different mechanisms: (1) an alteration of the turnover of nAcChR due to a decrease in the rate of synthesis or an increase in the rate of degradation, (2) a complement-mediated

* This sequence of events might also explain why about one-quarter of myasthenic patients have serum antibodies directed against skeletal muscle tissue determinants other than nAcChR located at the level of the sarcoplasmic reticulum, as well as on the musclelike cells of the thymus. According to Feltkamp (1978), the antibodies directed against these antigens, different from nAcChR, do not seem to contribute to the pathogenesis of myasthenia gravis. They seem to be related more to thymomas, even in absence of myastherda, than to myasthenia gravis itself.

4

BERNARD W. FULPIUS

muscle membrane destruction, or (3) a blockade of the acetylcholine binding site on the receptor, Each of these mechanisms leads to a diminution of available and functional nAcChRs on the postsynaptic membrane. All three mechanisms may be involved in myasthenia gravis, but their relative role in the pathogenesis of the disease is still matter of controversy. By comparison with anti-nAcChR antibodies of the IgG type, much less is known about those of the IgM type. It is even uncertain whether molecules of large size (MW 900,000) enter the synaptic cleft. The only indication in that direction is given by the localization of IgM derivatives within the region of nerve terminals. This is observed in an experiment in which complexes of 12SI-labeleda-BuTx covalently coupled to unspecific IgM were injected in viuo into mice, an experimental condition in which the possible release of small amounts of 1251-labeleda-BuTx through enzymatic hydrolysis cannot be entirely excluded (Zurn and Fulpius, 1976). There is no evidence yet for a pathogenic role for anti-nAcChR antibodies of the IgM type. For example, when tested on muscle cells in culture, these immunoglobulins do not induce an increased rate of receptor degradation (F. Clementi, unpublished observation). The question of the pathogenicity of antibodies of the IgM type is, however, of importance in view of the well-known prevalence of these immunoglobulins in the early phase of an immunization procedure and considering the observation by Lefvert et al. (1978). These authors studied three patients with a relatively short duration of myasthenic symptoms. When the patients were examined for the first time, there were no detectable anti-nAcChR antibodies of the IgG type. There were, however, antibodies of the IgM type detectable in two patients. Later on, in all three patients, IgG anti-nAcChR antibodies appeared, whereas IgM antibodies decreased in concentration. This IgM-IgG pattern was interpreted by the authors as an indication that the synthesis of anti-nAcChR antibody was triggered by antigenic stimuli. B. PATHOGENIC MECHANISMS OF COMPLEXED ANTIBODIES Circulating anti-nAcChR antibodies could be pathogenic in another manner, namely, as immune complexes, because any humoral antibody immune response eventually involves the formation of such complexes. As such, they would cause less specific damages than free-anti-nAcChR antibodies, since receptors for these complexes are known to occur in several anatomic areas leading, in those places, to immune complex de-

ANTIACETYLCHOLINE RECEPTOR ANTIBODIES

5

positon and injury. The possible existence of such complexes raises several questions of importance in connection with the pathogenesis of myasthenia gravis. (a) How are they formed? (b) What is their size? (c) To what extent do they activate the complement system? (d) What is their clearing system? These questions must be related to several factors (Williams, 1981): (1) the quality and immunoglobulin class of antibody involved. For example, immune complexes comprising IgM antibodies are larger and more rapidly cleared than those formed from IgG antibodies; (2) the relative quantities of antigen and antibody present. For example, soluble, and hence circulating, immune complexes are formed in situations of antigen excess, whereas increasing precipitation out of complexes occurs when the relative quantities of antigen and antibody approach equivalence. I n addition, immune complexes composed of more than one antigen molecule and cross-linked lattice-wise by several bivalent IgG molecules are often capable of effective complement pathway activation, the same being true for immune complexes composed of IgM antibodies; (3) the reticuloendothelial system; (4)the presence of Fc receptors on a number of circulating blood elements; and ( 5 ) the presence of actual receptors for activated complement components or Fc portions of immunoglobulins in various tissue sites. Unfortunately, very little is known of immune complexes in myasthenia gravis, although very sophisticated methods have been developed for a quantitative estimation, in several pathological conditions, of such complex levels in serum or other body fluids. Their existence has been, however, suggested by reports of anticomplementary and C l q binding activities in myasthenic sera and has been confirmed recently by more elaborated methods (Barkas et al., 1981). One should emphasize that the methods used so far are nonspecific in that all complexes, whatever the antigen involved, will be detected. Methods for the detection of specific immune complexes containing nAcChR-derived material are therefore needed. 111. Assays for Circulating Anti-nAcChR Antibodies

T h e design of assays for anti-nAcChR antibodies is rather complicated since there exist several clones of antibodies directed against nAcChR, each of them being specific for an antigenic determinant, but not all of them being pathogenic. At present, circulating anti-nAcChR antibodies are identified by several different methods, all of which depend on cu-BuTx for their specificity.

6

BERNARD W. FULPIUS

A. IMMUNOPRECIPITATION ASSAYS

Immunoprecipitation assays are the most widely used assays (Appel et al., 1975; Lindstrom, 1977; Monnier and Fulpius, 1977). They require first labeling the receptor protein with 1251-labeleda-BuTx. AntinAcChR antibodies that combine with the toxin-receptor complexes obtained in this manner are precipitated by adding the appropriate anti-human IgC or IgM serum3 and are found in about 90% of patients with myasthenia gravis. This type of assay, however, underestimates the actual amount of anti-nAcChR antibodies that circulate free in the serum because '251-labeled a-BuTx bound to nAcChR may alter or sterically occlude antigenic determinants recognized by two particular subpopulations of anti-nAcChR antibodies: (1) those that cannot bind to the receptor when a-BuTx is already bound and (2)those that block specifically the binding of acetylcholine, the natural ligand, to the receptor (Dwyer P t al., 1979). These two subpopulations are not mutually exclusive since the first one is not necessarily specific for the acetylcholine binding site. Although both subpopulations could induce a reduction of the receptor density, and hence the myasthenic neuromuscular deficiency, by mechanisms which imply binding to the receptor on the postsynaptic membrane followed by receptor internalization and/or complement-mediated membrane destruction, the second subpopulation deserves special attention because it could act also by causing only an immunopharmacologic blockade, a potentially operative mechanism already envisioned for myasthenia gravis (Simpson, 1960). Antibodies of group 1 are revealed by a modification of the precipitation assay. They have been reported to be the only anti-nAcChR antibodies present in one of the myasthenic patients studied by Dwyer et al. (1979).

B. INHIBITION ASSAYS lnhibition assays are used to detect antibodies which block the binding of acetylcholine a n d o r a-BuTx to nAcChR; they consist of a quantitative evaluation of the competition between antibodies and a-BuTx for binding to nAcChR from different sources. Antibodies directed against the binding site of the receptor were first Protein A can also be used for precipitating complexes made of toxin-labeled nAcChR and IgC. In this case, however, one has to remember that IgG of the subclass 3 does not react with protein A and then will escape detection. This is of importance since it is known that, in certain myasthenic patients, a large proportion of anti-nAcChR antibodies belong to that subclass (Lefvert t t 01.. 1981).

ANTIACETYLCHOLINE RECEPTOR ANTIBODIES

7

described by Almon et al. (1974) who used detergent-extracted nAcChR from denervated rat muscle. According to the reports of several authors, the proportion of myasthenic patients with antibodies directed against the toxin binding site varies from 7 to 60%. There is no correlation between their concentration and the concentration of antibodies directed against other sites on nAcChR (Bender et al., 1975; Mittag et al., 1976; Lefvert and Bergstrom, 1977, 1978; Vincent and Newsom-Davis, 1979; Lefvert et al., 1981). It is difficult to evaluate quantitatively the inhibition by antibodies of toxin binding to nAcChR because anti-nAcChR antibodies might also inhibit toxin fixation by steric mechanisms (when they are directed against sites adjacent to the toxin binding site) or by allosteric mechanisms (when they are directed against remote portions of the receptor molecule, a situation likely to happen whenever the receptor is solubilized by detergent and hence looses its native conformation). Appropriate inhibition assays must therefore meet the following conditions: (1) The receptor must be in its native environment as is the case in cells or intact membrane fragments, and (2) cross-reactivity with human antinAcChR antibodies should be restricted to the ligand binding site as seems to be the case with Torpedo electric organs (Vincent, 1980) and cultured chicken muscle cells (Fulpius et al., 1980b). Further, difficulties will be encountered while performing inhibition assays, namely, to obtain sufficient amounts of antigen, to reach an adequately high sensitivity, and to purify large amounts of high-titer IgG from myasthenic patients. Finally, as the cholinergic binding site is not equal to the toxin binding site, it is necessary to test, in an additional step, acetylcholine or another small cholinergic ligand in order to assay specifically the nicotinic cholinergic nature of the inhibition by antibodies (Fulpius et al., 1981). It follows that the development of suitable specificimmunodiagnostic methods for detecting antibodies directed against the cholinergic binding site is still needed. In this respect, one should consider of great potential value the recent development of monoclonal antibodies directed against the cholinergic binding site of the receptor (Jameset al., 1890).

IV. Anti-nAcChR Antibody Concentration in Different Forms of Myadhenia Gmvis

The discovery of the existence in myasthenia gravis of autoantibodies pathognomonic for the disease originally raised a great deal of interest among clinicians in consideration of the following question: Would the

8

BERNARD W. FULPIUS

level of measured anti-nAcChR antibodies be related to the patient’s clinical status? This question is of fundamental importance. As a matter of fact, a positive correlation would permit control of the evolution of the disease, predict the occurrence of relapses, and test whether the therapy has been appropriately selected. Unfortunately, most of the data published in this context were disappointing: T h e reported antibody titers corresponded only loosely with the patient’s clinical status (Almon et al., 1974; Appel et at., 1975; Bender et al., 1975; Lindstrom et al., 1976; Mittag et al., 1976; Ito et al., 1978; Bradley et al., 1979; Roses et al., 1981). In particular, many patients who appeared to be in complete clinical remission had titers well within the range of those with active disease (Lefvert et d ,1978), and about 10%of myasthenic patients had no detectable antibody (Lindstrom, 1977). This obvious lack of correlation can be explained in several ways: 1. T h e immunoprecipitation assay used in most of the studies does not detect all kinds of anti-nAcChR antibodies. 2. The anti-nAcChR antibodies that are detected in this manner are not necessarily those which are pathogenic. 3. Usually, the immunoprecipitation assay is carried on to detect class G immunoglobulins. 4. T h e level of circulating antibodies does not necessarily reflect the level existing in the vicinity of the receptor. 5. An uptake of antibodies at affected end plates might significantly deplete the circulation of appreciable amounts of anti-nAcChR antibodies. 6. Anti-nAcChR antibodies circulating as immune complexes escape detection. 7. T h e half-life of class G immunoglobulins shows large variations according to the four subclasses known.4 8. Sera are not always taken from patients according to the same protocol. 9. There are differences in the susceptibility to proteolysis among different antibody subpopulations. IgG sublcasses 1 , 2, 3, and 4 differ in several respects. IgG 3, in particular, has a strong tendency to aggregate and form complexes, a high susceptibility to proteolysis, and a rapid turnover (half-life of 7 days) when compared to that of the other subclasses (halflife of 2 1 days). In this context two reports of Lefvert pf al. (1978, 1981)are of considerable interest. According to these authors, anti-nAcChR antibodies have half-lives shorter than 8 days, and most of those detected in myasthenic patients by both imrnunoprecipitation and inhibition assays belong to either subclass 1 or 3.

ANTIACETYLCHOLINE RECEPTOR ANTIBODIES

9

T h e comparison of titers from one patient to another is difficult because of the use, in assays performed in different laboratories, of antigens from different sources. As a matter of fact, the cross-reactivity of myasthenic serum anti-nAcChR antibodies with nAcChR from different mammalian muscle extracts is highly variable. For example, antinAcChR titers against rat extracts are always lower than those against human muscle; in some cases they are even undetectable (Savage Marengo et al., 1979; McAdams and Roses, 1980). This explains why most researchers currently agree that human muscle is the most reliable source of antigen for determining anti-nAcChR in human sera, but even so, there are still difficulties in selecting the source of antigen. This is due mainly to the following two reasons: (1) There are still differences between the various muscles of a same species. For example, there are indications that ocular muscle nAcChR has some determinants distinct from those present on limb muscle nAcChR and vice versa. This finding is of importance in view of the fact that patients with predominant ocular symptoms represent the population with the lowest mean titer of antinAcChR antibodies when the assay is performed with nAcChR extracted from limb muscle (Lindstrom et al., 1976; Ito et al., 1978). (2) There are differences in the reactivity of myasthenia gravis sera toward junctional or extrajunctional receptors? higher titers being obtained with extrajunctional nAcChR (Weinberg and Hall, 1979). It follows that much more information related to the pathogenesis of myasthenia gravis could be obtained from anti-nAcChR antibody determinations, provided that: 1. Some form of standardization be always realized with, for example, sera of well-stablished activity. 2. Myasthenic sera be tested according to more than one assay in order to measure different subpopulations of anti-nAcChR antibodies. In this respect one should mention the report by Lefvert et al. (198 1) of the occurrence of anti-nAcChR antibodies competing for the ligand binding site in 50% of the myasthenic patients studied. All of the patients with these antibodies were severely ill, an indication that such antibodies might have a more disturbing effect on the neuromuscular function than those directed against other sites on the receptor. The junctional receptors are exclusively present in innervated muscles, whereas the extrajunctionalreceptors also present in innervated muscles, happen to exist in especially large amounts in denervated muscles, a preparation often used by biochemists to increase the yield of nAcChR.

10

BERNARD W. FULPIUS

3. An assay specific for pathogenic anti-nAcChR antibodies be developed. 4. Assays with sensitivites higher than those presently used be available. V. Anti-nAcChR Antibodies in the Cerebrospinal Fluid

Antibodies directed against nAcChR from human skeletal muscle and tested by a conventional immunoprecipitation assay were originally detected by Lefvert and Pirskanen (1977) in the cerebrospinal fluid of 9 out of 12 myasthenic patients. This finding raises three intriguing questions: 1. Are these antibodies synthesized locally or does their appearance in the cerebrospinal fluid result from a passive leakage through the blood-brain barrier! 2. Do these antibodies cross-react with central nervous system nAcChR? 3. Do these antibodies alter the synaptic transmission within the central nervous system?

'4.ANTIBODY ORIGIN T h e study of antibody origin implies, in addition to the assay for specific antibodies, the use of an appropriate test to assess the integrity of the blood-brain barrier, because any damage at that level would allow anti-nAcChR antibodies to gain access to the cerebrospinal fluid by passive leakage from the serum. T h e data available on this question bring rather conflicting evidence. On one side, data from Keesey et al. (1978) favor a passive leakage from the serum in view of the observed cerebrospinal fluid :serum ratios for the concentration of albumin and antinAcChR antibodies. On the other side, data from Lefvert et al. (1978) favor a local synthesis of anti-nAcChR antibodies since the cerebrospinal fluid : serum ratio for the concentration of IgG is normal and that for anti-nAcChR antibodies increased. Further studies on this question are expected. In particular, more specific information on the different subpopulations of anti-nAcChR antibodies existing within the cerebrospinal fluid is needed. T h e problem is complicated since the concentration of anti-nAcChR antibodies in the cerebrospinal fluid is lower than that in the serum by a factor of about

ANTIACETYLCHOLINE RECEPTOR ANTIBODIES

11

100. This makes it hard to perform tests that are sensitive enough to detect these antibodies and casts some doubt on a pathogenic action for them, at such low concentrations (Keesey et al., 1978).

B. CROSS-REACTIVITY WITH BRAIN nAcChR The possible Occurrence within the central nervous system of a cholinergic receptor of the nicotinic type is usually studied by using a-BuTx as a probe. The data pertaining to that question have been reviewed by Oswald and Freeman (1981). These authors conclude from their analysis that there are a-BuTx binding sites in the mammalian central nervous system which are located on nAcChR proteins similar to those of the muscle and electroplaque. It should be remembered, however, that the amount of information concerning the molecular properties of these a-BuTx binding proteins of neural origin is especially limited. This is especially true of the immunological characterization of the receptor, which is restricted to a few conflicting reports on the crossreactivity between anti-Torpedo sera and mammalian brain a-BuTx binding protein and a sole publication on the respective antigenic properties of muscle and brain nAcChR of human origin (Fontana et al., 1979). According to these authors, the two receptors have different antigenic determinants, a conclusion based on the following observation: A myasthenic serum of high titer against muscle nAcChR shows a much lower titer when tested against brain receptor, whereas three epileptic sera with antibodies against brain n AcChR show lower titers when tested against the muscle receptor. Such a result awaits confirmation. It emphasizes that specific assays must be used to test the relative reactivity toward brain andlor muscle receptor of the anti-nAcChR antibodies found in the cerebrospinal fluid.

C. ANTIBODY PATHOGENICITY Thre is some evidence for an involvement of the central nervous system in myasthenia gravis, namely, alterations of the hypothalamopituitary axis, electroencephalographic abnormalities, psychiatric symptoms, and a reduction of rapid eye movement sleep. According to Papazian (1976), this latter finding might indicate a disturbance of central cholinergic pathways. This is of great interest in view of the existence of anti-nAcChR antibodies of nicotinic specificity in the cerebrospinal fluid and the possibility for them to diffuse to neuroneuronal synapses and

12

BERNARD W. FULPIUS

intereact with the receptors located on the postsynaptic membrane. However, almost nothing is known of an impairment of central cholinergic synapses by these antibodies. The only report on this subject describes the induction of electroencephalographic abnormalities in rabbits by microinjection of human myasthenic serum into the caudate nucleus (Fontana et al., 1978). A comparison of antibody titers within the cerebrospinal fluid with central measurable alterations in man, and the possible effect, in this respect, of an immunosuppressive therapy should bring valuable information. VI. Antigenic Determinants on nAcChR

The nicotinic acetylcholine receptor is a protein complex embedded within the postsynaptic membrane of skeletal muscle cells. It follows that in situ not all antigenic determinants of the receptor are accessible and can interact with circulating immunoglobulins. Conversely, in myasthenia gravis, not all circulating anti-nAcChR antibodies are necessarily pathogenic since the formation of some of them might well have been triggered by parts of nAcChR normally not accessible and released in the course of the destruction of the postsynaptic membrane. Most of our information on nAcChR structure comes from studies performed with nAcChR from fish electric organs. A short review of the present state of knowledge concerning this latter receptor might give an idea of the complexity of the problems regarding the relative role of different antigenic determinants in the pathogeny of myasthenia gravis. A. TORPEDO FISHnAcChR Exhaustive reviews on torpedo fish nAcChR have been published (Fulpius et al., 1980a; Vincent, 1980). The receptor is a pentameric protein complex of about 270,000 daltons embedded within the membrane. Is quaternary structure is a p p y 6 , the two a polypeptide chains being identical. Only a chains bind the cholinergic ligands. Accordingly, there are two agonist or antagonist binding sites on each nAcChR complex of 270,000 daltons. The protein complex does not seem to show any symmetry because each subunit contains oligosaccharides of unknown size and must therefore face the extracellular space. The complex may, however, form dimers which are covalently linked together by a disulfide bridge between 6 chains. According to electron microscopy studies, the receptor protrudes about 50 A from the lipid matrix into the extracellu-

ANTIACETYLCHOLINE RECEPTOR ANTIBODIES

13

lar space. T h e pure receptor protein is readily available in large quantities. This has permitted thorough biochemical studies. For example, it has been shown by Raftery et al. (1980) that the four subunits have distinct but homologous amino acid sequences, in the first 56 N-terminal acids sequenced so far.

SKELETAL MUSCLEnAcChR B. HUMAN Significant differences have been reported between various mammalian muscle nAcChRs and their counterparts in fish electroplaques. There is, however, not much information related to the molecular structure of the receptor from human skeletal muscle. This is mainly due to obvious difficulties encountered in obtaining sufficient quantities of muscle with a minimum of tissue autolysis. T h e problem is further complicated by the variability of motor innervation inherent in lower leg muscles suffering from ischemia, a factor known to be linked to a more or less pronounced proliferation or extrajunctional receptors. Most of our knowledge on human nAcChR comes from the report of Stephenson et al. (198l), according to which the receptor has the following characteristics: (1)a specific activity for a-BuTx similar to that of nAcChRs purified from other sources; (2)a sedimentation coefficient of 9 S but no evidence for the existence of a dimerized 13 S form; (3)a microheterogeneity of the carbohydrate residues; (4) an original subunit pattern with two major protein bands of 42,000 and 66,000 daltons, the acetylcholine binding subunit being of the type common to all nAcChRs. In addition, the authors have observed that immunization of rabbits with this preparation generates low titers of the corresponding anti-nAcChR antibodies and does not cause experimental autoimmune myasthenia gravis. By comparison with the information available on Torpedo nAcChR and experimental autoimmune myasthenia gravis, many more studies are needed to better characterize the antigenic determinants of the human nAcChR in order to progress in the understanding of the immune response in myasthenia gravis. References

Almon, R. R., Andrews, C. G., and Appel, S. H. (1974). Science 186, 55-57. Appel, S. H., Almon, R. R., and Levy, N. (1975). N . Engl. J . Med. 293, 760-761. Barkas, T., Boyle, R. S., and Behan, P. 0. (1981).J. Clzn. Lab. Imrnunot. 5, 27-30. Bender, A. N., Ringel, S. P., Engel, W. K., Daniels, M. P., and Vogel, Z. (1975). Lancet 1, 607-609.

14

BERNARD W. FULPIUS

Bradley, R. J., Dwyer, D., Morley, B. J., Robinson G., Kemp, G . E., and Oh, S. J. (1979). Prog. Brain Res. 49, 441-448. Drachrnan, D. B. (1981). Annu. Rev. d\’eurosci. 4, 195-225. Dwyer, D. S., Bradley, R. J., Oh, S. J., and Kernp, G. E. (1979). Clin. Exp. Immunol. 37, 448-45 1 . Engel, A. G , , Lambert, E. H., and Howard, F. M. (1977). Mayo Clin. Proc. 52, 267-280. Feltkarnp, T. E. W. (1978).In “Neurology” (W. A. den Hartog Jager, G . W. Bruyn, and A. P. J. Heijstee, eds.), pp. 81-89. Excerpta Medica, Amsterdam. Fontana, A,, Fulpius, B. W., and Grob, P. J. (1978). Doc. Ophthalmol. 17, 35-43. Fontana, A.. Fulpius, B. W., and Cuenoud, S. (1979). Adv. Cytopharmcol. 3, 287-292. Fulpius, B. W., Bersinger, N. A., James, R. W., and Schwendimann, B. (1980a).In “Receptors for Neurotransmitters, Hormones and Pheromones in Insects” (D. B. Satelle, L. M. Hall, and J. G . Hildebrand, eds.), p p 3-15. ElseviedNorth-Holland, Amsterdam. Fulpius, B. W., Miskin, R., and Reich. E. (1980b). Proc. Natl. Acad. Sci. U.S.A. 77, 43264330. Fulpius, B. W., Lefvert, A. K., Cuenoud, S., and Mourey, A. (1981). Ann. N.Y. Acnd. Sri. 377,305-315. Ito, Y., Miledi, R., Molenaar, P. C., Newsorn-Davis, J.. Polak, R. L., and Vincent, A. (1978). I n “The Biochemistry of Myasthenia Gravis and Muscular Dystrophy” (G. G. Lunt and R. M. Marchbanks, eds.), pp. 89- 110. Academic Press, New YorWLondon. James, R. W., Kato, A. C., Rey, M.-J., and Fulpius, B. W. (1980).FEBS Lett. 120, 145-148. Keesey, J. C., Tourtelotte, W. W., Hermann, C., Jr., Andrews, J. M., and Lindstrom, J. (1978). Lancet 1, 777. Lefvert, A. K., and Bergstrom, K. (1977). Eur-.J. Clin. Inirest. 7, 115-119. Lefvert, A . K., and Bergstrom, K. (1978). Srund. J. Immunol. 8, 525-533. Lefvert, A. K., and Pirskanen, R. (1977). Loncef 2, 351-352. Lefvert, A. K., Bergstrom, K., Matell, G., Osterman, P. O., and Pirskanen, R. (1978).J. h’Purol., .Veurosurg. Pqchiatry 41, 394-403. Lefvert, A. K., Cuenoud, S., and Fulpius, B. W. (198l).J. Neuroimrnunol. 1, 125-135. Lindstrom. J. M . (1977). Clin.Immunoi. Immunopnthol. 7 , 36-43. Lindstrom, J. M. (1979).Adv. Immutwl. 27, 1-50. Lindstrorn, J. M., Seybold, M. E., Lennon, V. A., Whittngham, S., and Duane, D. (1976). N e u r d o a 26, 1054-1059. McAdams, M. W., and Roses, A. D. (1980). Ann. Neuroi. 8,61-66. Mittag, T., Kornfeld, P., Tormay, A., and Woo,C. (1976).N. Engl. J . Med. 294, 691-694. Monnier, V. M., and Fulpius, B. W. (1977). C h . Exp. Immuml. 29, 16-22. Oswald, R. E., and Freeman, J. A. (1981). Neuroscience 6, 1- 14. Papazian, 0. (1976). Neuroiogy 26, 311-316. Raftery, M. A., Hunkapiller, M. W., Strader, C. D., and Hood, L. E. (1980). S c i m e 208, 1454-1457. Roses, A. D., Olanow, C. W., McAdams, M. W., and Lane, R. J. M. (1981). N m r o l o o 31, 220-224. Savage Marengo, T., Harrison, R., Lunt, G. G., and Behan, P. 0.(1979). Lalvet 1,442. Sirnpson, J. A. (1960). Srott. ,\Zed. J . 5, 419-436. Stephenson, E A,, Harrison, R., and Lunt, G. (1981). Eur.J. Eiochm. 115, 91-97. Vincent, A. (1980). Physiol. Rev. 60, 756-824. Vincent, A., and Newsom- Davis, J. (1979).Adv. Cytophnnacol. 3, 267-278. Weinberg, C. B., and Hall, Z. W. (1979). Proc. Natl. Acad. Sci. U.S.A. 7 6 , 504-508. Wiliiams, R. C., Jr. (1981). Annu. Rmr. .Wed. 32, 13-28. Zurn, A. D., and Fulpius, B. W. (1976). C h . Exp. Immuiwl. 24, 9-17.

PHARMACOLOGY OF BARBITURATES: ELECTROPHYSIOLOGICAL AND NEUROCHEMICAL STUDIES By Max Willow* and Graham A. R. Johnrtont

* Doporlment of Pharmacology Schaol of Medicine University of Washington

Seattle, Washington and

t

Doporlrnent of Pharmacology Univonity of Sydney

New South Wales, Aurfmlio

I. Introduction ......................................................... 11. Neuropharmacological Studies ......................................... A. General Effects of Barbiturates on Synaptic Transmission ............. B. Effects of Barbiturates on Axonal Conduction ........................ C. F'resynaptic Actions of Barbiturates ................................. D. Effects of Barbiturates on Transmitter Action in Vertebrate Central Neurons ................................................... E. Effects of Barbiturates on Neuronal Membrane Properties.. ........... F. Neuropharmacology of Convulsant Barbiturates ...................... 111. Biochemical and Neurochemical Studies ................................ A. Effects of Barbiturates on Mitochondria1 Respiration .................. B. Effects of Barbiturates on Transmitter Release and Reuptake .......... C. Effects of Barbiturates on the Binding of Neurotransmitters to Receptor -Ionophore Complexes .......... .................... IV. Conclusions .......................................................... References ................. ......................................

15 16 16

21 22 24 32 33 34 34 35 41 44 45

I. Introduction

Barbituric acid was h s t synthesized by Baeyer in 1864, and this date marks the birth of an era that has witnessed the production of over 2500 derivatives. The first barbiturate introduced into clinical medicine (1903) was barbital, a long-acting sedative-hypnotic agent. Phenobarbital was marketed in 1912 for use in the treatment of certain forms of epilepsy. The use of ultra-short-acting barbiturates as intravenous anesthetics began in the early 1930s, and thiopental, in particular, gained rapid popularity following its introduction in 1935. While many of the sedative-hypnotic barbiturates have been superseded following 15 INTERNATIONAL REVIEW OF NEUROBIOLOGY. VOL. 24

Copyright 8 1983 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-366824-7

16

MAX WILLOW A N D GRAHAM A. R. JOHNSTON

TABLE I MOLECCLAR STRUCTURE OF SOME C-5 SUBSTITUTED BARHITURATES

Name

Principal action

R'

RZ ~

Methohexitone Thiopentone Butobarbitone Secobarbitone Amylobarbitone Pentobarbitone Phenobarbitone Barbitone CHEB 3M2B

Anesthetic Anesthetic Sedativehypnotic Sedativehypnotic Sedativelhypnotic Sedativelhypnotic Sedative/anticonvulsant Sedativelanticonvulsant Convulsant Convulsant

Ally1 Ethyl Ethyl Ally1 Ethyl Ethyl Ethyl Ethyl Ethyl Ethyl

l-Methyl-2-pentynyl 1-Methyl butyl 1-Methyl propyl I-Methyl butyl 3-Methyl butyl 1-Methyl butyl Phenyl Ethyl 2-Cyclohexylidene 3-Methylbut-2-enyl

X ~

0 S 0 0 0 0 0 0 0 0

the discovery of the benzodiazepines, barbiturates still maintain an important role in therapeutics, especially in their use as anesthetics and antiepileptics. The molecular structure of many of the barbiturates discussed in this article is shown in Table I. In general, only the effects of barbiturates on neuronal systems in vertebrate species will be discussed here. Detailed accounts of the pharmacological actions of barbiturates on invertebrate neurons can be found elsewhere (Barker, 1975a,b,c; Prichard, 1980; Wilson et al., 1980). \I. NeumpharmMobgical Studies

A. GENERAL EFFECTSOF BARBITURATES ON SYNAPTIC TRANSMISSION

The most detailed studies on the effects of barbiturates on synaptic transmission have been performed on the spinal monosynaptic reflex. Eccles ( 1946) demonstrated that the excitatory postsynaptic potential (EPSP) recorded from ventral roots in decerebrate cats was depressed by pentobarbital (40 mg/kg, iv) and concluded that the block of the monosynaptic reflex was largely due to an increased stability of the soma membrane. Brooks and Eccles (1947) showed that pentobarbital (30 mg/kg, iv) depressed the orthodromically induced focal potential in the

PHARMACOLOGY OF BARBITURATES

17

spinal cord of decerebrate cats. With increasing concentrations, the synaptic potential was progressively depressed, until finally the monosynpatic pathway was completely blocked. Although they observed that pentobarbital (in high doses) was capable of depressing propagation in the afferent presynaptic terminals, blockage of synaptic transmission was thought to be due to a stabilization of the soma membrane which prevented the discharge of impulses initiated by normally effective synaptic potentials. Shapovalov (1963) showed that pentobarbital (20-30 mg/kg, iv) and hexobarbital ( 15-20 mg/kg, iv) depressed EPSPs recorded intracellularly from cat spinal motoneurons without altering resting membrane potential and axonal impulse conduction. A contemporary study by Somjen and Gill (1963) demonstrated that thiopental (30-50 mg/kg, iv) blocked the transmission of the monosynaptic reflex in the cat and the rat, as seen by a depression of the EPSP without alteration of the resting membrane potential. In a parallel study, Somjen (1963) showed that when EPSPs were reduced to 10% of control amplitude following the administration of thiopental (65 mg/kg, iv), impulse conduction in presynaptic terminals was unimpaired. He concluded that the most probable explanation for the depression of synaptic potentials was a nonspecific stabilization of the soma membrane or a decrease in the amount of transmitter released per presynaptic impulse. Ldyning et al. (1964) examined the effects of the short-acting barbiturate thiamylal sodium on the monosynaptic reflex in lightly anesthetized cats. Using intracellular recording techniques, they showed that intravenous administration of this drug (10 mg/kg) decreased the EPSP, without altering the spike potential, resting potential, and accommodation of the motoneurons. When potentials evoked by a volley applied to afferent nerves were recorded extracellularly at the dorsal root entry in the motor nucleus, and from the ventral root, it was found that thiamyal sodium reduced (a) the initial negative spike recorded in the motor nucleus, (b) the focal synaptic potential, and (c) the ventral root potential. It was concluded that the reduced EPSP was due mainly to thiamylal acting on different nerve terminals resulting in a decrease in transmitter release or to a reduced sensitivity of postsynaptic membrane receptor sites to the transmitter. T h e actions of pentobarbital and thiopental on monosynaptic EPSPs in cat spinal motoneurons were examined by Weakly (1969). He showed that these drugs, when administered intravenously (10 mg/ kg), significantly depressed the monosynaptic reflex discharge of triceps sural motoneurons. Both drugs reduced the mean quantum content by about 25% without altering the average amplitude of the unit EPSP. It was concluded in agreement with Ldyning et al. (1964) that depression of monosynaptic reflex transmission by thiopental and pentobarbital was

18

MAX WILLOW AND GRAHAM A. R. JOHNSTON

due to a reduction in the average amount of transmitter released by group l a afferent impulses. A reduction of excitatory synaptic transmission has been demonstrated at a number of other sites in the CNS. Galindo (1969) examined the effects of pentobarbital on synaptic transmission in the cuneate nucleus of decerebrate cats. Cuneate neurons were excited (a) by various stimuli including a jet of air applied to hairs, a weight applied on the skin, or the movement of a joint; (b) electrical pulses applied to peripheral nerves; and (c) stimulation of nerve endings in the dorsal column. I n each case, pentobarbital (2-30 mg/kg, iv) reduced synaptic transmission in response to these stimuli. Microelectrophoretic administration (40200 nA) of pentobarbital (0.2 M, pH 9.5) produced similar effects. I n addition, pentobarbital administered electrophoretically (40 nA) significantly reduced the firing rate of cuneate neurons excited by glutamate (60 nA). Nicoll (1972) examined the effects of pentobarbital, hexobarbital, and other anesthetic agents on synaptic excitation and inhibition in the olfactory bulb of the rabbit. Pentobarbital (3-30 mg/kg, iv) prolonged the granule cell inhibition of mitral cells while having little effect on synaptic excitation of granule cells and antidromic field potentials invading mitral cell dendrites. Relatively large doses of pentobarbital (40-70 mg/kg, iv) were needed to significantly depress synaptic excitation of granule cells and antidromic invasion of mitral cell dendrites. Richards ( 1972) showed that pentobarbital (0.05-0.25 mM) depressed the EPSP component of the evoked field potential recorded in an in nitro preparation of guinea pig olfactory cortex. T h e population spikes that were superimposed on the EPSP were reduced in amplitude and frequency with these concentrations of pentobarbital, indicating a failure of transmission through the cortical relay. It was concluded that pentobarbital produces its effects (a) by reducing the amount of transmitter released from presynaptic nerve terminals in response to the afferent volley or (b) by reducing the sensitivity of the postsynaptic membrane to the released transmitter. Gordon et nl. (1973) have demonstrated that synaptic transmission in the mossy fiber pathway of the cat cerebellum is depressed by thiopental (0.5-8.0 mg/kg, iv), whereas transmission in the climbing pathway is enhanced. T h e depression of' excitatory synaptic transmission is exemplified by the abolition of the axon discharge of granule cells, and as a consequence, Purkinje cells are unable to respond to the tibia1 nerve stimulation via the mossy fibers. Barbiturates have also been shown to exert depressant effects on excitatory synaptic transmission at various sites other than the CNS. Larrabee and Posternak (1952) have shown that concentrations of pentobarbital (0.2-0.5 miM) that depress postsynaptic responses in cat stellate ganglia following stimulation of the preganglionic fiber, have no

PHARMACOLOGY OF BARBITURATES

19

effect on impulse conduction in postsynaptic fibers. A similar selective action of barbiturates on synaptic transmission in the superior cervical ganglion from a number of species has been observed by Quillam and his collaborators (Brown and Quillam, 1964a,b; Elliott and Quillam, 1964; Quillam and Shand, 1964). Synaptic transmission at the vertebrate neuromuscular junction is also impaired by barbiturates. Thesleff (1956) demonstrated that pentobarbital (0.6 mM) blocked neuromuscular transmission in the sartorius nerve-muscle preparation of the frog. This effect was characterized by an increase in the electrical threshold of the muscle membrane and a reduced action potential. While the dose of pentobarbital required to produce these effects was five times greater than the mean hypnotic dose for pentobarbital in the frog, Thesleff (1956) concluded that the anesthetic activity of pentobarbital may be due to a reduction of the sodium conductance change in the cell membrane of certain neurons in the central nervous system. Adams ( I 976) examined in detail the effects of amylobarbital, methohexital, and thiopental on the physiology of voltage-clamped end plates of frog sartorius muscles. In the presence of barbiturates (80 p M ) , the conductance change evoked by electrophoretic carbachol was reduced by a prepulse of carbachol. This desensitization disappeared exponentially with a time constant of 150-200 msec. All barbiturates tested (0.4-1.2 mM) produced an increased rate of decay of nerve-evoked end-plate currents. In addition, thiopental, in a dose-dependent manner, depressed conductance changes produced by bath-applied agonists (choline, carbachol, and tetramethylammonium bromide). Adams ( 1976) also observed that the concentrations of barbiturates required to depress the bath agonist response are much greater than the kinetically determined dissociation constant for binding to active receptor-channel complexes. It was concluded that the depressant effects of barbiturates on synaptic transmission at the frog end plate were mainly mediated by a blockage of open end-plate receptor channels. While emphasis has been placed on the depressant effects of barbiturates on excitatory synaptic transmission, these compounds also enhance synaptic inhibition at a number of sites. Larson and Major (1970) showed that hexobarbital (10 mg/kg, iv) markedly prolonged the time course of the recurrent inhibitory postsynaptic potential (IPSP) in cat spinal motoneurons. This effect is not attributable to a prolonged Renshaw cell discharge since equivalent closes have been shown to shorten the Renshaw cell discharge (Eccles et ul., 1956). As mentioned above, Nicoll (1972) demonstrated that doses of barbiturates that are without effect on the EPSP in the olfactory bulb markedly prolong postsynaptic inhibition. Nicoll et al. (1975) demonstrated that pentobarbital (10-33 mg/kg, iv) markedly prolonged the time course of the IPSP recorded in cat hip-

20

MAX WILLOW AND GRAHAM A. R. JOHNSTON

pocampal pyramidal neurons. In addition, pentobarbital (10 mg/kg, iv) prolonged both the evoked and spontaneous unitary IPSPs. Barbiturates appear to enhance postsynaptic inhibition at a number of other sites in the CNS. Bloedel and Roberts (1969) examined various aspects of cerebellar physiology before and after the administration of pentobarbital in decerebrated and spinalized cats. Pentobarbital (15 mg/kg, iv) enhanced the postsynaptic inhibition of Purkinje neurons by basket cells. This result was confirmed by Eccles et al. (1971) using thiosecobarbital(l0-40 mg/kg, iv). It was concluded that in the unanesthetized cat cerebellum there is a higher level of Purkinje cell excitability, probably due to a domination of the excitatory action of parallel fibers over the inhibitory action of basket cells. Scholfield (1977) demonstrated that pentobarbital (100 p M ) prolonged the evoked IPSP recorded in guinea pig olfactory slices in vitro. The evoked EPSP was depressed with higher concentrations of pentobarbital. Barbiturates also enhance the recurrent inhibition of cortical pyramidal tract neurons (Veselyuneneet al., 1971; Steriade et al., 1974) and the recurrent inhibition of thalamic relay neurons (Bremer, 1970).All of the inhibitory pathways referred to above are thought to release y-aminobutyric acid (GABA), with the exception of recurrent inhibition of spinal motoneurons, which is mediated by glycine (Curtis and Johnston, 1974; KrnjeviC, 1974). Eccles and Malcolm ( 1946) first demonstrated that pentobarbital greatly prolongs the decay of the dorsal root potential in the isolated frog spinal cord. This observation has been confirmed in many studies both in the frog (Schmidt, 1963, 1964; Grinnell, 1966; Richens, 1969; Nicoll, 1975a) and in the cat (Lloyd, 1952; Eccles et al., 1963). The synaptic depolarization of primary afferents, which underlies the dorsal root potential, is considered to result in reduced release of excitatory transmitter from the primary afferents (Eccles, 1964), and GABA is thought to be the depolarizing transmitter responsible for this presynaptic inhibitory process (Levy, 1977). Eccles et al. (1963) demonstrated that presynaptic inhibition of monosynaptic reflexes in the cat is prolonged to the same extent as the dorsal root potential by pentobarbital and thiamylal. In addition, picrotoxin (0.3 mg/kg, iv) antagonized the actions of pentobarbital (10 mg/kg, iv) in increasing and prolonging the presynaptic inhibition of the monosynaptic reflex. Presynaptic inhibition is also enhanced by phenobarbital (Miyahara et al., 1966). Nicoll (1975a) demonstrated that concentrations of pentobarbital as low as 5 pM could prolong the dorsal root potential in the isolated frog spinal cord preparation. In addition, pentobarbital (40 p M ) depolarized primary afferent fibers. This depolarizing action was approximately equipotent to that of GABA, and was blocked by picrotoxin and bicuculline. These actions of

PHARMACOLOGY OF BARBITURATES

21

pentobarbital were also seen with amylobarbital, thiopental, and barbital. Pentobarbital (10- 15 mg/kg, iv) enhanced surface potentials (similar to dorsal root potentials) recorded on the surface of the cuneate nucleus of the cat (Banna and Jabbur, 1969). This was associated with an increased excitability of presynaptic terminals, although a depression of excitability was observed with higher doses (25 mg/kg, iv). Rudomin (1966) demonstrated that pentobarbital (10 mg/kg, iv). enhanced primary afferent depolarization in the solitary nucleus of the cat. In general, the studies cited above suggest that excitatory synaptic transmission is depressed by barbiturates, whereas inhibitory synaptic transmission, especially that mediated by GABA, is enhanced. OF BARBITURATES ON AXONAL CONDUCTION B. EFFECTS

It is now generally accepted that barbiturates do not greatly affect the conduction of impulses along axons at concentrations likely to be present during anesthesia (approximately 100-200 p M for pentobarbital; see Fisher et al., 1948; Jori et al., 1970; Richards, 1972; Saubermann et al., 1974). Heinbecker and Bartley (1940) and Schoepfle (1957) demonstrated a local anesthetic-like action of barbiturates in blocking excitation in peripheral nerves. Later studies using voltage-clamp techniques showed that millimolar concentrations of pentobarbital and thiopental decreased and prolonged sodium conductance in lobster giant axons (Blaustein, 1968) and squid giant axons (Narahashi et al., 1969). The potassium conductance is reduced but not significantly prolonged by barbiturates in these preparations. Barbiturates have a more rapid onset of action and greater potency in producing these effects when they are applied intracellularly as opposed to external application (Narahashi et al., 197 1; Frazier et al., 1975). It has been suggested that the un-ionized form of the barbiturate molecule is responsible for the block in conduction (Krupp et al., 1969; Narahashi et al., 197 1). Of interest is the finding that there is no apparent difference in the time course of action between clinically short-acting and long-acting barbiturates in blocking axonal conduction (Frazier et al., 1975). While barbiturates exert local anesthetic actions on nerve fibers of large diameter at only relatively high concentrations, smaller myelinated and nonmyelinated fibers may be more susceptible to lower concentrations of these agents. This proposal was first suggested by Frank and Sanders (1963) and later by Seeman (1972). Staiman and Seeman (1974) have provided some experimental confirmation of this hypothesis. They demonstrated that pentobarbital produced a 50% block in conduction in

22

M A X WILLOW AND GRAHAM A. R. .JOHNSTON

phrenic nerves at 400 pX1, whereas a concentration of 800 g M was required to produce similar effects on large sciatic nerve fibers. Thus, a slight reduction in the amplitude of the action potential (which could possibly occur at anesthetic concentrations) in small-diameter fibers may be of importance in altering the amount of transmitter released from nerve terminals in the presence of barbiturates.

c. PRESYSAPrIC: ACTIOSSOF BARBITURATES Brooks and Eccles ( 1947) demonstrated that barbiturates depressed the focally recorded presynaptic volley associated with the spinal monosynaptic reflex, but dismissed this as a primary site of action because it was seen only with high doses. However, in later studies, Ldyning ut “1. (1964) and Richens (1969) suggested that this action on the primary afferent terminals was the only effect that could adequately account for the depression of the EPSP. Weakly (1 969) established a selective presynaptic action by demonstrating that subanesthetic doses of thiopental and pentobarbital (10 mgkg, iv) reduced the mean quantum content of the unitary EPSP in spinal motoneurons by about 2576, without altering the average amplitude of the unitary EPSP. I n addition, these concentrations of barbiturates did not alter the input resistance of the motoneuron or the strength-duration relationship. Nicoll (1980) has suggested that barbiturates may exert their presynaptic action by (a) decreasing the size and/or blocking terminal invasion of the action potential or (b) directly interfering with the transmitter release mechanism, possibly through an action on calcium fluxes or metabolism. Ldyning Pf nl. (1964) showed that barbiturates reduced the amplitude of the action potential invading primary afferent terminals and suggested that barbiturates were acting like local anesthetics. Such an action would be expected to depress terminal excitability. Galindo (1969) reported a reduction of the excitability of primary afferent terminals on the cuneate nucleus. Nicoll (1975a) demonstrated that the barbiturate depolarization of primary afferent terminals in the isolated frog spinal cord is associated with an increase in terminal excitability. This result does not exclude a local anesthetic action of pentobarbital which may be masked by its action on primary afferent depolarization. Indeed, with higher concentrations (> 1 mM), the local anesthetic action of pentobarbitai predominates (Nicoll, 1975a). At synapses other than those involved with the monosynaptic reflex pathway, the presynaptic actions of barbiturates are variable. Richards ( 1 972) found no change in the size of the presynaptic spike or in the

PHARMACOLOGY OF BARBITURATES

23

excitability of presynaptic fibers with concentrations of pentobarbital (0.25 mM) that depress excitatory transmission in the olfactory cortex. Scholfield and Harvey (1975) demonstrated that pentobarbital exerted a selective depressant action on synaptic potentials compared to action potentials in isolated guinea pig olfactory slices, with rather high concentrations (>1 mM) required to cause a 50% depression of the amplitude of the action potential. Nicoll(l972) has suggested that the depression of the olfactory bulb EPSP by hexobarbital(40-90 mg/kg, iv) may be due in part to a presynaptic action, since the size of the presynaptic dendritic response is concomitantly reduced. The vertebrate neuromuscular junction has been another site where the effects of barbiturates on transmitter release have been examined. Barbiturates have been shown to increase the quantal content of the end-plate potential (EPP) following nerve stimulation (Quastel et al., 1972; Thomson and Turkanis, 1973; Seyama and Narahashi, 1975; Proctor and Weakly, 1976). This increase in release has been attributed to the prolongation of the presynaptic potential since the amplitude of the presynaptic spike is unaffected or even reduced by barbiturates (Thomson and Turkanis, 1973). Barbiturates also increase the frequency of spontaneous miniature end-plate potentials (MEPP) at the vertebrate neuromuscular junction (Quastel et al., 1971, 1972; Westmoreland et al., 1971; Thomson and Turkanis, 1973). On the other hand, barbiturates depress the frequency of MEPPs at the crustacean neuromuscular junction (Iravani, 1965). A recent study by Pincus and Insler (1981) suggests that the effects of barbiturates on transmitter release at the frog neuromuscular junction may depend largely on the calcium content of the bathing medium during periods of evoked release. Both phenobarbital and the convulsant barbiturate 5-ethyl-5-(2’-cyclohexylidene-ethyl)-barbituricacid (CHEB) increased the quantal content of the EPP and the amplitude of the EPP in Ca2+-deficientRinger’s solution. I n contrast, both drugs depressed the amplitude of the EPP without altering quantal content when normal Ringer’s media (Ca2+concentration = 1.8 mM) was employed. T h e variable effects of barbiturates on transmitter release at different synapses may be attributable to differences in the release mechanism (e.g., degree of calcium dependency) in addition to anatomical factors such as the presence of presynaptic inhibitory inputs on nerve terminals (e.g., primary afferent terminals). It may be more relevant to study the effects of barbiturates on transmitter release at central synapses rather than peripheral synapses, despite the technical difficulties involved in measuring release from CNS neurons. T h e puzzling finding that CHEB and phenobarbital exert simi-

24

MAX WILLOW A N D GRAHAM A. R. JOHNSTON

lar actions on release at the neuromuscular junction (Pincus and Insler, 1981)suggest that these actions may be unrelated to the pharmacological effect of these drugs observed in ztivo.

D. EFFECTS OF BARBITURATES ON TRANSMITTER ACTIONI N VERTEBRATE NEURONS CENTRAL 1 . Efects .f Barbiturates on Responses Evoked by Excitatory Transmittn Substnnces Most of the studies examining the interaction of barbiturates with putative excitatory transmitters in the CNS have utilized electrophoretic techniques in which the firing rate of neurons has been recorded extracellularly . Krnjevic and Phillis ( 1 963) reported that systemic administration of barbiturates produced a substantial and prolonged reduction in the firing of cat cerebral cortical neurons by acetylcholine (ACh). T h e cells examined (Betz cells) appeared to have a population of muscarinic cholinergic receptors upon them. Thiopental and hexobarbital (0.5- 10 mg/kg, iv) abolished the responses of cat caudate neurons to electrophoretically ejected ACh (Bloom et al., 1965). Curtis and Ryall (1966) showed that systemically administered pentobarbital reduced the frequency of firing of cat Renshaw cells in response to ACh, n-butyrylcholine, nicotine, and acetyl-P-methylcholine. Pericruciate cortical neurons stimulated by electrophoretic ACh are also sensitive to various barbiturates, including pentobarbital, diallylbarbital, and methylthioethyl-2-pentyl-thiobarbiturate(Crawford and Curtis, 1966; Crawford, 1970). Catchlove et al. (1972) demonstrated a depression of ACh-evoked responses of deep pericruciate neurons by methohexital(3 mg/kg, iv) and suggested that barbiturates, like dinitrophenol, act by inhibiting aerobic mitochondria1 metabolism. T h e firing of rat brainstem neurons by ACh was depressed by systemic or electrophoretic administration of pentobarbital (Bradley and Dray, 1973). Duggan et nl. (1974) concluded that the nicotinic response of cells in the paramedian reticular nucleus of the cat was more sensitive to the depressant actions of barbiturates than was the muscarinic response. Adams (1976) has shown that barbiturates have more potent effects in depressing the response to exogenously applied ACh at the neuromuscular junction than their action on the rise phase of the end-plate current (EPC). Since the opening of ionic channels during the EPC is very fast, it was proposed that barbiturates do not have sufficient time to enter and block during the rising phase of the EPC. O n the other hand, the rising phase of the response to

PHARMACOLOGY OF BARBITURATES

25

exogenous ACh follows a much slower time course, and it was suggested that barbiturates could block open channels during this period, resulting in a diminished response. I n addition, barbiturates have been shown to depress the excitatory effects of acidic amino acids on the firing rate of various neurons. Crawford and Curtis (1966) demonstrated that the firing rate of deep pyramidal cells (of the cat pericruciate cortex) by electrophoretic DLhomocysteic acid is depressed by systemic and electrophoretic administration of barbiturates. Phillis and Tebecis (1967) showed that pentobarbital (2 mg/kg, iv) reduced the responses of cat thalamic neurons to L-glutamate (discharge frequency reduced to about two-thirds of the control magnitude). Pentobarbital, when applied electrophoretically or administered systemically (20 mg/kg, iv), depressed the firing rate of cat cuneate nucleus cells in response to L-glutamate (Galindo, 1969). Of interest is the fmding that doses of barbiturates that depressed the Achevoked firing of rat brainstem neurons had little effect on glutamateevoked responses (Bradley and Dray, 1973). Nicoll (1 975b) has shown that pentobarbital (20- 100 p M ) depressed the depolarization of frog motoneurons by glutamate. Barbiturates have also been shown to depress the glutamate-evoked depolarization of mouse spinal neurons grown in tissue culture (MacDonald and Barker, 1979; Ransom and Barker, 1975). I n addition, barbiturates also depressed the glutamateinduced conductance increases in spinal motoneurons (MacDonald and Barker, 1979). Richards and Smaje (1976) observed that pentobarbital (0.1-0.3 mM) consistently depressed the excitatory actions of L-glutamate on prepiriform cortical cells in vztro. Responses of various neurons, excited by serotonin (5-HT), have also been shown to be sensitive to barbiturates. Roberts and Straughan (1967) have shown that the systemic injection of small quantities of thiopental selectively and reversibly reduced the sensitivity of cat cortical neurons to excitation by 5-HT when at the same time the response to glutamate was unaffected. Johnson et al. (1969) similarly demonstrated a depression of 5-HT and norepinephrine-evoked firing of cortical neurons in the cat. In addition, it was noted that the number of cells excited by norepinephrine in barbiturate anesthetized animals was markedly less than the number of cells excited in unanesthetized or N20-halothane anesthetized animals.

2. The Effects of Barbiturates an Responses Evoked by Inhibitory Transmitter Substances T h e inhibitory effects of 5-HT (Roberts and Straughan, 1967; Johnson et al., 1969; Tebecis and DiMaria, 1972), norepinephrine

26

MAX WILLOW AND GRAHAM A. R. JOHNSTON

(Bloom P t nl., 1965; Johnson et nl., 1969), dopamine (Bloom et al., 1965; Tebecis and DiMaria, 1972), and ACh (Bloom et ai., 1965) are, in general, little affected by moderate amounts of barbiturates. On the other hand, a great deal of attention has recently been focused on the effects of barbiturates on responses to exogenously applied GABA in a variety of i n uiiw and in vitro preparations. Nicoll (1975b) showed that pentobarbital (20 p M ) increased the amplitude and duration of the GABA-mediated hyperpolarization of frog motoneurons. At higher concentrations (200 pLzI),pentobarbital caused a direct hyperpolarization of frog motoneurons. At these concentrations the amplitude of the GABA response was depressed and markedly prolonged. Bowery and Dray (1976) demonstrated a reversal by barbiturates of the bicuculline methochloride (BMC) antagonism of the GABAmediated depolarization of the isolated superior cervical ganglion of the rat and the inhibition by GABA of the firing of medullary neurons. Of particular interest was the finding that pentobarbital did not potentiate these responses to GABA in the absence of BMC. This finding was questioned by Curtis and Lodge (1977), who unequivocally demonstrated that the response of doral horn interneurons in the cat to electrophoretic GABA was enhanced by pentobarbital (also administered electrophoretically in cats anesthetized by a-chloralose or urethane). In agreement with Bowery and Dray (1976), they showed that pentobarbital partially reversed the antagonism by BMC of the inhibition of cell firing by GABA. In a later study, Lodge and Curtis (1978) showed that in the unanesthetized decerebrate cat, pentobarbital (15 mg/kg, iv) increased the time course of recovery of dorsal horn interneurons stimulated by GABA, without altering the time course of recovery of cell firing by glycine. Evans (1979) showed that pentobarbital (10-80 p M ) enhanced the depolarization of immature rat dorsal root fibers by GABA, a process which appeared to be bicuculline insensitive. On the other hand, higher concentrations o f pentobarbital (50- 160 p M ) produced a bicucullinesensitive depolarization of dorsal root fibers, characterized by a slower onset and offset than that produced by GABA. It was suggested that bicuculline could antagonize the GABA-like actions of pentobarbital but could not antagonize the enhancement of GABA by pentobarbital. Nicoll could enhance the GABA(1978) showed that pentobarbital (100 pFLI\I) mediated depolarization of frog sympathetic ganglion cells, but the effects of bicuculline were not investigated in this study. Recently, Connors (198 1) has examined the actions of pentobarbital on neurons of dorsal root ganglia from adult rats. Pentobarbital (40-200 FM, bath applied) enhanced the GABA-mediated transient inward current into ganglion cells voltage clamped at their resting potential. In cells which were not

PHARMACOLOGY OF BARBITURATES

27

voltage clamped, concentrations of 40 and 200 p M pentobarbital enhanced the GABA-induced depolarization of such cells, and in addition, enhanced the increase in conductance due to GABA. In all of these experiments, GABA was bath applied. Increasing the concentration of pentobarbital to 1 mM resulted in an attenuation of GABA responses. Concentrations of pentobarbital that enhanced GABA responses (40200 p M ) did not alter the resting membrane potential or conductance. However, at 1 mM, pentobarbital produced a small depolarization (<2 mV) associated with very small increases in membrane conductance. Action potentials, evoked by the passing of current intracellularly, were unaffected by 100 pM pentobarbital. At 1 mM pentobarbital, the amplitude and the rate of rise of the action potential were depressed. In addition, the threshold current and voltage were increased. In a number of cells, pentobarbital (100 p M ) reduced the number of spikes evoked by pulses (30 msec) of depolarizing current. I n a series of elegant studies, Barker and his colleagues have examined the effects of barbiturates on transmitter responses (particularly GABA and glutamate) and membrane properties on cultured mouse CNS neurons. Ransom and Barker (1975) examined the action of pentobarbital (applied electrophoretically) on the responses of cultured mouse spinal neurons to GABA and glutamate (also applied electrophoretically). Pentobarbital depressed the glutamate-evoked depolarization of neurons, whereas the response to GABA was markedly prolonged in the presence of the drug. The half-time for restoration of control conductances following electrophoresis of GABA was increased from 2.5 sec to 3.9 sec following administration of pentobarbital. In addition, pentobarbital altered the polarity of the GABA response of spinal neurons by accentuating a secondary depolarizing component of the response. I n addition to these effects on transmitter responses, pentobarbital also produced a direct decrease in membrane resistance by 5-15% in about 80% of cells tested, an effect accompanied by a small hyperpolarizing action (1-3 mV). I n a following study, Ransom and Barker (1976) showed that pentobarbital (0.1-0.2 mM, applied by diffusion) prolonged the conductance change induced by an electrophoretic pulse of GABA (30 nA, 0.5 sec) applied to the somas of mouse spinal neurons grown in tissue culture. T h e time course of conductance decay was prolonged by a mean of 67% in the presence of pentobarbital, and was accompanied by only slight changes in resting membrane potential and input resistance of spinal neurons. The time course of conductance decay for GABA was often characterized by an early rapid phase followed by a slow late phase. I n neurons that exhibited a biphasic increase in GABA-induced conductance, both phases were prolonged by pen-

28

MAX WILLOW AND GRAHAM A. R. JOHNSTON

tobarbital (0.1-0.2 m&f). The change in membrane potential produced by brief electrophoretic pulses of GABA on spinal neurons was accompanied by a secondary depolarizing component in the presence of pentobarbital. T h e depolarization of mouse spinal neurons by electrophoretic application of glutamate (10 nA) was depressed by 45% in the presence of 0.1-0.2 mbf pentobarbital. Pentobarbital did not generally alter the time course of glycine-induced increases in membrane conductance in cells that exhibited an increase in the half-decay time of the GABAinduced increase in membrane conductance in the presence of the drug. I n a more detailed study, Barker and Ransom (1978) examined the actions of pentobarbital on mammalian central neurons grown in tissue culture. When added to the bathing medium, pentobarbital (0.1 mM) abolished all spontaneous synaptic activity in spinal cord or cerebellarbrainstem neurons. While remaining spontaneously inactive in the presence of the drug, these cells could be excited when depolarized to threshold by injected current. Pentobarbital, administered by diffusion (0.1 mhl) or electrophoresis (50 nA) produced an increase in membrane conductance, the magnitude being considerably less, and the time course or conductance change slower than that induced by GABA. The direct action of pentobarbital was blocked by diffusion of a 1 mM solution of picrotoxin into the bathing medium. In addition, pentobarbital reversed the antagonism of GABA responses by picrotoxin at concentrations that did not directly affect membrane properties. With sufficient currents of pentobarbital, GABA responses were actually potentiated in the presence of picrotoxin before the barbiturate-induced increase in membrane conductance was observed. T h e inversion potential for the pentobarbital-induced change in membrane properties was dependent on the concentration of chloride ions in the bathing medium, and was similar to the dependence of GABA conductance changes for the presence of this ion. In addition to its direct actions on membrane conductance, pentobarbital enhanced the conductance increases following the administration of GABA. T h e decay in GABA conductance changes was markedly slowed by pentobarbital. T h e GABA dose (current)response (both voltage and conductance response) curve was shifted to the left in the presence of pentobarbital. The potentiating effects of pentobarbital on GABA-mediated events were relatively specific for GABA, since responses to glycine and p-alanine were unaffected. Pentobarbital directly depressed the depolarization of neurons by glutamate or indirectly depressed glutamate responses by increasing membrane conductance. The direct depressant action of pentobarbital on glutamate responses could be seen at concentrations of the drug that did not in-

PHARMACOLOGY OF BARBITURATES

29

crease membrane conductance. Barker and Ransom (1978) suggested that the direct depression of glutamate responses by pentobarbital was as sensitive as the enhancement of GABA responses by this drug. In some experiments, electrophoresis of glutamate produced rapid events in addition to the primary postsynaptic response, and these events may be due to glutamate-evoked efRux of transmitter from terminals presynaptic in relation to the recorded neuron. At concentrations that did not directly alter membrane conductance, pentobarbital abolished these events suggesting that transmitter release in this preparation may be sensitive to barbiturates. MacDonald and Barker (1979) have described qualitative and quantitative differences in the pharmacological actions of phenobarbital and pentobarbital on cultured mouse spinal neurons. T h e spontaneous activity in such cultures was characterized by an abundance of excitatory and inhibitory postsynaptic potentials, random firing of action potentials, and occasional short period bursts of action potentials. In the presence of 0.2 mM phenobarbital, this pattern was basically unaltered except that the neuronal firing rate was slightly increased and the frequency of the short burst of action potentials was decreased. I n contrast, the same concentration of pentobarbital abolished all observable synaptic activity. When picrotoxin (0.08 mM) was added to the bathing medium, the activity of the neurons was characterized by random, abrupt depolarizations associated with bursts of action potentials (paroxysmal depolarizing events). In the presence of 0.2 mM phenobarbital, these paroxysmal events were abolished, and the spontaneous activity of neurons was similar to that observed in the control medium or in the presence of phenobarbital. Electrophoretic application of pentobarbital (300 nA), but not phenobarbital (300 nA), resulted in membrane hyperpolarization and an increase in membrane conductance. Picrotoxin and penicillin antagonized these direct membrane actions of pentobarbital at concentrations which do not alter neuronal membrane properties in the absence of added drugs. T h e amplitude of the GABA-induced hyperpolarization was augmented by the electrophoretic application of both pentobarbital and phenobarbital, although higher currents of phenobarbital were required than those for pentobarbital. The time course of the GABA response was prolonged by pentobarbital in a dose-dependent manner, whereas there was no such alteration in the presence of phenobarbital. Both drugs depressed the depolarization of spinal neurons by glutamate, but phenobarbital appeared to be two to three times weaker than pentobarbital in this respect. Similar differences in pharmacological actions between the anticonvulsant, mephobarbital, and the anesthetic barbiturates, secobarbital and racemic 1,3-dimethyl-butethyI barbituric acid

30

MAX WILLOW AND GRAHAM A. R. JOHNSTON

have also been described (MacDonald and Barker, 1979). Schulz and MacDonald ( 1981) have recently examined the comparative actions of phenobarbital and pentobarbital on a number of amino acid-mediated events in cultured mouse spinal cord neurons. This study differed from that bv MacDonald and Barker (1979) in that barbiturates were applied (a) directly to the bathing medium or (b) by local miniperfusion of individualized neurons, rather than by electrophoretic application. Pentobarbital and phenobarbital both abolished bicuculline-evoked paroxysmal depolarizing events (EDWsof 150 and 170 p h f , respectively). Both barbiturates were effective in augmenting GABA-mediated hyperpolarizations (EDSosof 50 and 75 p L U ,respectively). Pentobarbital was shown to be more potent than phenobarbital in abolishing the spontaneous activity recorded in these cultured cells. Similarly, pentobarbital produced direct GABA-like increases in membrane conductance at much lower concentrations ( 100-750 pill) compared to phenobarbital (1000-4000 pi\f).Schulz and MacDonald (1981) suggested that the latter two effects of pentobarbital (i.e., abolition ofspontaneous activity possiblv due to a reduction in transmitter release and a direct increase in membrane conductance) may be partly responsible (in addition to the enhancement of GABA responses and reduction in glutamate-mediated excitation) for the anesthetic properties of this drug. On the other hand, the relative lack of effect of phenobarbital on (a) the spontaneous activity of neurons and (b) the neuronal membrane conductance, may explain why this agent does not possess anesthetic properties at concentrations relevant to its anticonvulsant actions. Using fluctuation analysis, Barker and McBurney (1979) examined the effects of electrophoretically applied phenobarbital on the properties of conductance channels activated by GABA in cultured mouse spinal neurons. The amplitude of the single channel conductance, y , can be estimated from GABA-induced changes in membrane current. T h e estimate of single channel conductance in the presence of phenobarbital was similar to control values. In order to examine the mean duration of “opening” of GABA-activated channels, Barker and McBurney ( 1979) obtained analyses of membrane current fluctuations recorded during the plateau phases of GABA responses and transformed these fluctuations into power density spectra. T h e mean channel-open time, T , is calculated from a resolution of these current fluctuations. In the presence of phenobarbital the duration of the unitary conductance activated by GABA is prolonged approximately fivefold. In addition, phenobarbital prolonged the time constant of decay of a population of spontaneous synaptic currents while not affecting the amplitude or rise time of these currents. These currents may be due to activation of postsynaptic mem-

PHARMACOLOGY OF BARBITURATES

31

brane receptors following the spontaneous release of quanta1 GABA in this preparation (Barker and McBurney, 1979). T h e pharmacological effects of the (+) and (-) isomers of pentobarbital have been examined on cultured mouse spinal neurons (Huang and Barker, 1980). In a dose-dependent manner, (+) pentobarbital depolarized the membrane potential in 62% of cells studied when administered by pressure from glass pipettes. In 34% of cells, (+) pentobarbital evoked excitatory synaptic potentials despite the presence of a high concentration of Mg2+ (10 mM). In a small minority of cells, (+) pentobarbital produced complex alterations in membrane potential and conductance, suggesting that different types of spinal neurons exhibit different drug sensitivities (Huang and Barker, 1980). I n 1 1 of 30 cells studied, (-) pentobarbital hyperpolarized spinal neurons and increased the membrane conductance. The concomitant application of both isomers resulted in a marked attenuation of the excitatory responses seen with (+) pentobarbital alone. The GABA-induced increase in the membrane conductance of spinal neurons was greatly enhanced by (-) pentobarbital in 18 of 24 neurons studied. I n 3 neurons studied, (+) pentobarbital potentiated the amino acid response to a similar degree, and to a lesser degree on another 3 cells. In 20 cells, (+) pentobarbital did not affect the GABAevoked increase in membrane conductance. Huang and Barker (1980) concluded that the primary action of the (+) isomer was excitatory and that of the (-) isomer was inhibitory. Mathers and Barker (1980) showed that like GABA, (-) pentobarbital can directly open membrane channels in cultured mouse spinal neurons. T h e mean channel-open time, T , of (-) pentobarbital-activated channels was five times that of GABAactivated channels. Nicoll and Wojtowicz (1980) have recently examined the effects of pentobarbital and related compounds on the response of frog motoneurons to exogenously applied GABA and glutamate. Pentobarbital (140 p M ) and phenobarbital (800 p M ) produced a half-maximal enhancement of the GABA-mediated hyperpolarization. A 50% reduction in the glutamate-evoked depolarization was obtained at similar concentrations. Pentobarbital reversed the noncompetitive antagonism of GABA responses by picrotoxin, without altering the bicuculline methiodide antagonism of GABA. Pentobarbital was also shown to directly hyperpolarize motoneurons in a similar manner to GABA. Bicuculline and picrotoxin both abolished this action. T h e threshold concentration for the pentobarbital-mediated hyperpolarization (100 p M ) was about 10 times greater than the concentration required to enhance GABA-mediated responses. Higher concentrations of pentobarbital (0.4-2.0 m M ) reduced the antidromic field potential, suggest-

32

MAX WILLOW A N D GRAHAM A. R. JOHNSTON

ing that motoneuron excitability was depressed. It was suggested that this action was due to a GABA-mimetic action of pentobarbital, rather than a local anesthetic action, since the reduction in excitability was attenuated by picrotoxin (at concentrations which did not alter excitability in the absence of barbiturates). Barker and Mathers (1981) have recently suggested that the two fundamental properties of pentobarbital related to GABA receptors are (1) a direct activation of chloride conductance channels which may not necessarily be involved with the engagement of the GABA recognition site, and (2) potentiation of GABA responses involving a prolongation of channel lifetime. This increase in channel lifetime may be related to the increase in apparent affinity of membrane binding sites for GABA in the presence of pentobarbital, as has been recently demonstrated by Willow and Johnston (1980); see Section II1,C for a more detailed discussion. ox NEURONAL MEMBRANE PROPERTIES E. EFFECTSOF BARBITURATES

From in vivo studies, there appear to be conflicting reports on the effects of barbiturates on the membrane properties of spinal motoneurons. Eccles (1946) and Brooks and Eccles (1947) observed that barbiturates reduced the amplitude of the antidromic field potential and suggested that a decrease in the excitability of the soma membrane could account for the depression of the monosynaptic reflex. Later studies (Sasaki and Otani, 1962; Shapovalov, 1963; Somjen and Gill, 1963) confirmed the observation that the excitability of the motoneuron was reduced in the presence of barbiturates. However, LByning et al. (1964) and Weakly (1969) observed a reduction in the amplitude of the motoneuron EPSP without any changes in the membrane potential or excitability of the soma. These differences in the effects of barbiturates on neuronal membrane excitability are probably related to the variations in the doses employed. Ldyning et al. (1964) and Weakly (1969) used doses of barbiturates (10 mg/kg) that d o not consistently induce anesthesia in cats. A number of in uitro studies have demonstrated consistent effects of barbiturates on the membrane conductance and potential of neurons at concentrations ( 100-200 piV for pentobarbital, see Richards, 1972) found in the brains of laboratory animals during anesthesia. These effects include the depolarization of dorsal root fibers in the immature rat spinal cord (Evans, 19’79), hyperpolarization of frog motoneurons (Nicoll and Wojtowicz, 1980), and hyperpolarization of cultured mouse spinal neurons associated with an increase in membrane conductance

PHARMACOLOGY OF BARBITURATES

33

(Barker and Ransom, 1978; MacDonald and Barker, 1979; Schulz and MacDonald, 1981).

F. NEUROPHARMACOLOGY OF CONVULSANT BARBITURATES Convulsant barbiturates were initially described by Dox and Yoder (1922) when 5-benzyl-5-ethyl barbituric acid was shown to produce a tetanus-like syndrome in laboratory animals prior to hypnosis. T h e synthesis of (+)-5-(1,3-dimethylbuty1)-5-ethylbarbituric acid and a description of its excitatory actions were published during the next decade (Swanson, 1934; Swanson and Chen, 1939). The convulsant properties of the (+) stereoisomers of pentobarbital (Kleiderer and Shonle, 1934) and methohexital (Gibson et al., 1959) have also been described. T h e cyclic barbiturate CHEB was first synthesized by Velluz et al. (1951), and several recent investigations have examined the pharmacology of this agent. Downes and Williams (1969) examined the effects of CHEB on the monosynaptic (2N)reflex recorded from spinal ventral roots in the unanesthetized cat. The amplitude of the monosynaptic spike was significantly increased following the systemic administration of CHEB (0.5 mg/kg, iv). The effects of CHEB on the presynaptic inhibition of spinal reflexes were also examined, but the results were too variable and inconsistent to draw any positive conclusion. I n a later study, Downes and Franz (1971) studied the effects of CHEB (1-2 mglkg, iv) on dorsal root ganglion cells in unanesthetized spinal cats. Using intracellular recording techniques, they showed that such doses of CHEB depolarized the dorsal root ganglion cells (which had been isolated from the spinal cord by severance of the dorsal roots), in association with a reduced firing threshold and fall in membrane resistance. In addition, CHEB elicited discharges from the central end of the severed dorsal roots, which was attributable to a depolarization of intraspinal primary afferent terminals. This depolarizing action of CHEB has also been described by Nicoll (1980) and Andrews et al. (198 1). Unlike the depolarization of primary afferent fibers by pentobarbital (Nicoll, 1975a; Evans, 1979), the CHEB-evoked depolarization is unaffected by the GABA antagonists, bicuculline and picrotoxin (Nicoll, 1980; Andrews et al., 1981). I n addition, CHEB depolarized motoneurons in the frog spinal cord (whereas pentobarbital has a hyperpolarizing action) in the presence of Mg2+ used to block synaptic transmission (Nicoll, 1980). Nicoll(l980) has suggested that this direct depolarizing action could explain the increase in the size of the monosynaptic spike observed by Downes and Williams (1969).

34

MAX WILLOW AND GRAHAM A . R. JOHNSTON

A pharmacological property common to convulsant and depressant barbiturates is their ability to enhance the inhibitory actions of GABA on feline dorsal horn interneurons 27) ~ ~ Z U O (Lodge, 1979). This action was considered to be of little significance in determining the pharmacology of the convulsant barbiturates, since CHEB, like picrotoxin and bicucculline, reduced the dorsal root potential (Lodge, 1979). Crawford (1969) found that when administered electrophoretically, the convulsant barbiturate 5-ethyl-5-(1,3-dimethyLbutyl)barbituric acid depressed the responses of cortical neurons to the excitatory actions of ACh and DL-homocysteic acid, a result qualitatively similar to the action of depressant barbiturates. In addition, the action of glutamate on frog motoneurons is depressed by CHEB (Nicoll, 1980). The effects of the excitatory stereoisomer of pentobarbital on the properties of cultured mouse spinal neurons have been discussed elsewhere in this article (see Section II,D), and the neurochemical pharmacology of convulsant barbiturates will be covered below (Section 111,B).

111. Biochemical and Neurochemical Studies

A.

EFFECTS O F B A R B I T U R A T E S ON MITOCHONDRIAL

RESPIRATION

Nearly 50 years ago, Quastel and Wheatley (1932) examined the effects of a number of anesthetics (including barbiturates) on the respiration of slices of guinea pig brain. They showed that phenobarbital (approximately 5 m'21) and other barbiturates caused a 5070 inhibition of oxygen uptake when glutamate was the substrate and caused no inhibition when succinate was oxidized. Ernster et 01. (1955) suggested that amylobarbital may be a specific inhibitor of the mitochondria1 NADH oxidase system. They showed that at a final concentration of 1.8 mM, am ylobarbital produced complete inhibition of the oxidation of NADlinked substrates and partial inhibition of the oxidation of succinate or of the phosphorylation coupled to succinate in rat liver mitochondria. It was suggested that amylobarbital exerted its effects between the NADlinked dehydrogenases and the point of convergence of the NADH oxidase and succinate oxidase systems. Although Chance (1956) and o b e r g ( 1 96 1) claimed that barbiturates act between NADH and NADH dehydrogenase in the respiratory chain, more recent evidence supports the view that inhibition occurs on the oxygen side of the flavoprotein and cytochromes c and c1 or cytochrome b at the time when investigators

PHARMACOLOGY OF BARBITURATES

35

placed cytochrome b on a side path of the respiratory chain. Using the NADH-cytochrome c oxidoreductase complex preparation, Hatefi et al. (1961) showed that while cytochromes b and c l were oxidized in the presence of amylobarbital, flavoprotein remained reduced. The structural requirements for the barbiturate-like inhibition of the NADH oxidase system have been examined in beef heart submitochondrial particles (Cowger et d.,1962). It was shown that the essential structure consisted of a nonspecific hydrocarbon group attached to an amide, carbamide, or barbituric acid; these compounds have the -CO-NHgroup in common. Barbituric acid was not inhibitory, and, in general, a lengthening of one or of both alkyl side chains at C-5 increased the inhibition. B. EFFECTS OF BARBITURATES ON TRANSMITTER RELEASE A N D REUPTAKE

To date, the most detailed neurochemical studies involving barbiturates have examined their effects on neurotransmitter release in uitro. Kalant and Grose (1967) showed that pentobarbital exerted biphasic effects on the potassium-evoked release of endogenous ACh from guinea pig cortical slices. In the presence of 5 mM KCl, the steady efflux of ACh was increased by low concentrations (10 p M ) of pentobarbital and depressed by higher concentrations (400 p M ) . When efflux was measured in the presence of 27 mM KCl, similar effects were observed, but the magnitudes of the increase and decrease in release were greater. Grewaal and Quastel (1973) examined the effects of amylobarbital and thiopental on the efflux of [14C]ACh from rat brain cortex slices. With increasing concentrations of both drugs (0.25- 1.0 mM), the release of [14C]ACh evoked by 31 mM KCl was progressively inhibited. On the other hand, release evoked by 4 mM KCl was only depressed with higher concentrations of barbiturates ( 1 mM). In addition, when calcium was omitted from the incubation medium, the release of [14C]ACh(in the presence of 4 mA4 KC1) was also depressed by amylobarbital and thiopental (0.25-0.50 mM). Thiopental (0.25-0.50 mM) but not amylobarbital (0.25-0.50 mM) depressed ACh release evoked by protoveratrine (10 p M ) . The inhibition of [14C]ACh release (evoked by 3 1 mM KCl) by barbiturates was thought to be due to an increase in the release of mitochondria1 Ca2+,which in turn could presumably alter Na+ and K+ fluxes across the neuronal membrane (Grewaal and Quastel, 1973). Carmichael and Israel (1975) examined the effects of barbiturates on the release of [3H]norepinephrine (NA) and [I4C]ACh(from rat brain cortical slices) evoked by electrical field stimulation (25 Hz, 0.5 msec, 25

36

MAX WILLOW AND GRAHAM A. R. JOHNSTON

V) applied over a 3-min period. Pentobarbital (300 p M ) and phenobarbital (1 mM) significantly inhibited ACh release (35 and 45%, respectively). Higher concentrations of both drugs were needed to inhibit pH]NA release to a similar extent. Nonstimulated release of NA or ACh was unaffected by pentobarbital (300 phi) or phenobarbital (1 mhl). Richter and Waller (1977) showed that pentobarbital (50 mg/kg, ip) significantly inhibited the KC1 (50 mM)-evoked efflux of the ['*C]ACh from minislices derived from rat cerebral cortex, hippocampus, striatum, or pons-medulla regions. Similar inhibitory actions were observed when pentobarbital (100 p M ) was administered in vitro. Pentobarbital, administered in uiz~oor in vitro, did not modify ACh release in the presence of 5 m M KCl. In a later study, Richter and Werling (1979) examined the effects of a variety of barbiturates on the K+-stimulated efflux of PH]ACh from rat midbrain minislices. In the presence of 5 mM KCl, the efflux of PH]ACh was unaffected by thiamylal(500 p M ) ,methohexital (500 p,CI), secobarbital (1 mXI), amylobarbital (1 mM), pentobarbital (1 mM), and phenobarbital (5 mM). When the concentration of KCI in the incubation medium was increased to 50 mM, PH]ACh release was inhibited (50%) by much lower concentrations of these drugs (e.g., thiamylal, 92 +\I; pentobarbital, 297 p M ) . Simon and his co-workers (Simon and Kuhar, 1975; Simon et al., 1976) have examined the effects of pentobarbital on high-affinity [3HJcholine uptake into rat hippocampal synaptosomes. [3H]Choline uptake was reduced by 30-65o/c when animals had been treated with pentobarbital (65 mg/kg, ip) 30 min prior to sacrifice. pH]Glutamate and [SHItyrosine uptake were unaffected by similar drug treatment. On the other hand, pentobarbital (0.2-1.0 mM, in vitro) did not alter ~Hlcholineuptake. T h e effect of pentobarbital on the efflux of rH]NA has been studied in rat cerebral cortical slices (Lidbrink and Farnebo, 1973) and mouse forebrain synaptosomes (Haycock ~t al., 1977) using a variety of stimuli to evoke release. Lidbrink and Farnebo (1973) observed a lack of effect of pentobarbital (100 p M ) on the efflux of rH]NA evoked by electrical field stimulation (10 Hz, 12 nA, 2-msec pulse). They did note, however, that at this concentration pentobarbital significantly inhibited [3H]NA uptake into cortical slices. On the other hand, Haycock et al. (1977) demonstrated a significant inhibition of E3H]NA and [14C]GABA efflux by pentobarbital (200 p M ) when KCI (50 mM) or veratridine (100 p M ) were used to evoke release. This group suggested that pentobarbital may affect release through an effect on depolarization-triggered Ca2+influx, since [3H]NA release was unaltered by pentobarbital (200 prM) in the presence of the artificial Ca2+ ionophore A23 187.

PHARMACOLOGY OF BARBITURATES

37

Cutler and his colleagues (Cutler and Dudzinski, 1974; Cutler et al., 1974) have examined the effects of barbiturates on amino acid transport in rat brain cortical slices. Cutler and Dudzinski (1974) examined the sensitivity of rH]GABA and [x4C]glutamateuptake and release processes to pentobarbital. Pentobarbital, tested at two concentrations (0.1 and 5.0 mM), depressed GABA uptake by 24 and 58%, respectively, with little effect on glutamate uptake. The spontaneous efflux of rH]GABA was depressed by 26% (100 p M ) and 48% (5 mM). Pentobarbital at these concentrations exerted only minor inhibitory effects (14%at 5 mM) on the spontaneous efflux of [14C]glutamate.T h e release of CJHIGABA evoked by electrical field stimulation (rectangular pulse, 10 nA, 5 msec, 80 Hz) was enhanced by 46% in the presence of 5 mM pentobarbital. However, [14C]glutamate release was only slightly increased (13%)with this concentraton. Both GABA and glutamate release were unaffected by lower concentrations of pentobarbital (100 pM).I n a parallel study, Cutler et al. (1974) ,examined the effects of various barbiturates on rH]GABA release from rat brain cortical slices. They showed that pentobarbital (0.01- 1 .O mM) inhibited the spontaneous efflux of rH]GABA and K+-evoked release of [3H]GABA from rat brain cortical slices in a concentration-dependent manner. Amylobarbital produced a similar inhibition at higher concentrations than pentobarbital (0.1-5.0 mM), whereas hexobarbital ( 1 mM) was inactive. Spontaneous glutamate release was only slightly inhibited by high concentrations of pentobarbital ( 1 mM). Pentobarbital (0.1 mM) and amylobarbital (0.1 mM) inhibited the ouabain (10 p M ) -evoked efflux of PHIGABA, but phenobarbital (5 mM) did not inhibit efflux. When rH]GABA release was evoked by various concentrations of KCI (3.5-60 mM), amylobarbital (0.9 mM) depressed release at each concentration tested. At 60-mM KCl (efflux maximum at this concentration), concentrations of amylobarbital as low as 50 p M exerted an inhibitory effect on efflux. In addition to these findings, Cutler et al. ( 1 974) showed that high concentrations of pentobarbital (2 mM) increased the Michaelis constant (K,) for both low- and high-affinity GABA uptake without altering maximal uptake (VmaX).T h e inhibitory effects of barbiturates on potassium- and ouabain-stimulated efflux were interpreted as a result of inhibition of carrier-mediated membrane transport of GABA (Cutler et al., 1974). Jessell and Richards (1977) examined the effects of pentobarbital on rH]GABA uptake and release in rabbit hippocampal slices. In contrast to the findings of Cutler et al. (1974),they found that high concentrations of pentobarbital (1 mM) did not alter low- or high-affinity GABA uptake. On the other hand, K+-evoked efflux of THlGABA was inhibited by lower concentrations of pentobarbital (200 p M ) . Olsen et al. (1978) ex-

38

MAX WILLOW AND GRAHAM A. R. JOHNSTON

amined the effects of various psychoactive substances (including diethyl barbituric acid) on fH]GABA binding, uptake, release, and synthesis an nitro. Diethyl barbituric acid (600 p M ) inhibited uptake (presumably high affinity) by 50%, and did not affect K+ (55 mM) -evoked release of PHIGABA, L-glutamic acid decarboxylase (GAD) activity, or THIGABA binding at the highest concentrations of drug tested (100 p M ) . Recently, Cutler and Young (1979) examined the effects of various barbiturates on the release of endogenous amino acids from rat cortical slices. They showed that pentobarbital (1 mill) inhibited the release of GABA and glutamic acid (evoked by 60 mM KCI), whereas lower concentrations (0.1 m,\f) were without effect. T h e efflux of amino acids evoked by and veratridine (5 p M ) was unaffected by pentobarbiouabain (10 pAM) tal and a variety of other barbiturates (1 mM). Amylobarbital (1 mM) produced a similar depression of K+-evoked efflux to pentobarbital (1 m,\l), but secobarbital (1 mLV)and phenobarbital (1 mM) had no inhibitory action. Minchin (1980) recently showed that K+ (40 mM) -evoked release of rH]GABA from rat cortical slices was inhibited by 50% in the presence of thiopental (100 p M ) , pentobarbital (200 p M ) , and methohexital (600 pLtZ). Collins (1980) examined the effects of pentobarbital (0.01- 1.0 mM) on the evoked release of endogenous aspartate, GABA, and taurine from rat olfactory cortical slices. Release was evoked by electrical stimulation of the lateral olfactory tract (pulse width, 100 psec, supramaximal voltage, frequency of 4 stimuli/min for 20 min) at a site distant from the cortical cup. In addition, the effects of pentobarbital (0.02- 1.0 mM) on the uptake of fH]GABA by olfactory cortical slices were also measured. Preincubation and perfusion of preparations with pentobarbital (0.011.O m,M) resulted in a concentration-dependent inhibition in the evoked (lateral olfactory tract stimulation) release of aspartate and taurine. Under these conditions, the release of GABA was significantly increased in the presence of up to 250 pXf pentobarbital, but was reduced at 1000 ,uM. Picrotoxin (15 p.V) did not reverse the depressant actions of 100 p M pentobarbital on aspartate release, whether release was evoked directly or indirectly by lateral olfactory tract stimulation. Picrotoxin (15 p M ) partially reversed the increased GABA release evoked by lateral olfactory tract stimulation in the presence of pentobarbital ( 1000 p M ) . Pentobarbital (100 p1M)did not alter GABA release in preparations that were directly stimulated. Pentobarbital (20-750 p M ) did not affect [3H]CABA uptake by olfactory cortical slices. Collins ( 1980) suggested that the reduction in the evoked release of aspartate in the presence of pentobarbital (100 p M ) may be due to (a) a reduction in the changes in calcium permeability which accompany depolarization of presynaptic

PHARMACOLOGY OF BARBITURATES

39

terminals or (b) a prolongation of the presynaptic inhibitory actions of released GABA which may depolarize the terminals of lateral olfactory tract fibers. The mechanism by which pentobarbital potentiated GABA release upon stimulation of the lateral olfactory tract is unclear, but Collins ( 1980) suggested that synaptic activation is a necessary prerequisite on the basis that direct stimulation of slices does not alter GABA release. Willow et nl. (1980) have demonstrated that anesthetic and convulsant barbiturates exert differing actions on the K+-evoked release of preloaded ~-pH]aspartaticacid from rat cerebral cortical minislices. At concentrations of 100 p M , the anesthetic barbiturates pentobarbital and amylobarbital significantly reduced the spontaneous efflux of D-[~H]aspartate (K+ concentration = 4.75 mM), whereas at similar concentrations the convulsant barbiturates, CHEB, 5-ethyl-5-(1’,3’-dimethylbutyl) barbituric acid (DMBB), and 5-ethyl-5(3-methylbut-2’-enyl) barbituric acid (3M2B) enhanced the spontaneous efflux of this amino acid. T h e efflux of ~-pH]aspartateevoked in the presence of 44.7 mM KCl was also depressed by the anesthetic barbiturates (at 100 p M ) , whereas CHEB and 3M2B increased efflux and DMBB had no significant effect. Willow and Johnston (1979) demonstrated that anesthetic barbiturates stimulate the activity of a synaptosomal Ca2+-activatedATPase, whereas convulsant barbiturates inhibit activity. It has been previously suggested that this enzyme may be involved in the active efflux of calcium ions from presynaptic terminals (Robinson, 1978). A calcium-pumping role for this enzyme is supported by the observations that (a) Ca2+-ATPaseactivity has been measured in synaptic plasma membranes (Sobue et al., 1979) and (b) calcium efflux under normal physiological conditions may be mediated through an ATP-dependent process (Di Polo and Beauge, 1979) rather than a previously described sodium-calcium countertransport mechanism (Blaustein and Hodgkin, 1969). The effects of anesthetic and convulsant barbiturates on the efiux of ~-pH]aspartate (Willow et nl., 1980) are consistent with the hypothesis that barbiturates may act to alter the activity of the plasma membrane calcium pump, and hence either decrease (anesthetics) or increase (convulsants) the cytoplasmic concentration of calcium ions within nerve terminals (Willow and Johnston, 1979). The anomalous finding that the convulsant barbiturate DMBB was without effect on the evoked efflux of D-aspartate may be due in part to other actions of this drug; for example, convulsant barbiturates may, like pentobarbital and thiopental, inhibit the entry of calcium ions into synaptosomes during depolarization (Blaustein and Ector, 1975; see section below for detailed discussion of this action). Willow et nl. (1980) have suggested that the effects of barbiturates on spontaneous

40

MAX WILLOW AND GRAHAM A. R. JOHNSTON

release may provide a better test for the involvement of a membrane calcium pump underlying these actions, since calcium influx into synaptosomes in a low [K’] medium is apparently insensitive to barbiturates (Blaustein and Ector, 1975). Furthermore, since unstimulated terminals probably d o not accumulate significant amounts of calcium, it is likely that a membrane calcium pump is important in regulating cytoplasmic levels of calcium ions in the absence of nerve terminal depolarization. When terminals are depolarized, it is likely that mitochondrial (Rahamimoff et al., 1975; Nicholls, 1978) and nonmitochondrial (Baker, 1972; Blaustein P! al., 1978a.b; Politoff et al., 1974; Harris, 1981) calcium uptake systems play an important role in the sequestration of intracellular calcium. Blaustein and Ector (1975) examined the effects of some barbiturates on %az+ uptake into synaptosomes in the presence of KC1 (71 mM), veratridine (75 p.V), or gramicidin D (10 pg/ml). Pentobarbital (0.4-0.5 milt) and thiopental(O.2 miM) inhibited 45Ca2+uptake in the presence of all three depolarizing agents, whereas phenobarbital (0.9 mM was without effect. In addition, the convulsant (+) isomer of DMBB ( 1 mM) also inhibited ‘%azf uptake in the presence of 71 mil4 KCl. T h e inhibitory effects of barbiturates on *Ca2+ uptake were thought to contribute to the effects these substances exert on synaptic transmission. Leslie el al. (1979) also demonstrated that pentobarbital (0.3 mM) significantly inhibited the K+-evoked entry of %a2+ into synaptosomes isolated from a variety of species. In synaptosomes isolated from rats tolerant to phenobarbital, acute pentobarbital (0.3 miM) challenge did not alter the depolarizationinduced influx of W a Z +(Leslie ~t ul., 1980). A recent study has shown that pentobarbital (0.1- 1 .O mM) stimulates the initial rate of Ca2+transport into rat brain mitochondria (Willow and Bygrave, 1982). On the other hand, the Na+-induced efflux of preloaded 4sCa2+is unaffected by pentobarbital ( 1 mA4) but is significantly inhibited by the local anesthetic dibucaine. In contrast, Pincus and Hsiao (1981) have demonstrated that phenobarbital (0.2 mM) lower the content of mitochondrial Caz+.T h e differences in the effects may be accounted for in the methods used to measure the accumulation of 45Ca2+in mitochondria. Willow and Bygrave (1982) employed the Ruthenium red-EGTA quench technique, which specifically measures Caz+ transport by the inner mitochondrial membrane protein carrier (Reed and Bygrave, 1974, 1975). I n contrast, Pincus and Hsiao (1981) measured the total “upake” of Caz+ by mitochondria, of which a considerable fraction may have been due to processes not related to the Ca2+transport system (i.e., binding of CaZ+to phospholipids; see Scarpa and Azzone, 1969).

PHARMACOLOGY OF BARBITURATES

41

C. EFFECTS OF BARBITURATES ON THE BINDING OF NEUROTRANSMITTERS TO RECEPTOR-IONOPHORE COMPLEXES I n the past two to three years several advances have been made in elucidating the molecular mechanisms by which barbiturates enhance synaptic inhibtions in the CNS, particularly those mediated by GABA. We have recently demonstrated, for the first time, an enhancement by pentobarbital of the binding of GABA to rat brain synaptosomal membranes (Willow and Johnston, 1980). Previous attempts to demonstrate a barbiturate enhancement of GABA binding had been unsuccessful (Enna and Snyder, 1976; Peck et al., 1976; Olsen et al., 1979), probably attributable to the methods used to prepare synaptosomal mernbranes. I n these studies, GABA binding had been optimized by removal of endogenous inhibitors using procedures such as high-frequency homogenization, freezing and thawing, and detergent extraction of the membranes. These endogenous inhibitors include GABA itself (Napias et al., 1980), phospholipids (Johnston and Kennedy, 1978),a 15,000-MW protein (Guidotti et al., 1978; Toffanoet al., 1978; Massotti and Guidotti, 1980; Massotti et al., 1981), and other low molecular weight substances (Yoneda and Kuriyama, 1980). In demonstrating an enhancement of GABA binding by pentobarbital, particular attention was focused on preparing a synaptosomal fraction that probably retained many of these endogenous inhibitors (while removing most of the endogenous GABA and Na+), in order to preserve a membrane environment which closely resembles that encountered in vivo (Willow and Johnston, 1980; Johnston and Willow, 1981). The pentobarbital enhancement of GABA binding is concentration dependent (Willow and Johnston, 1980, 198la; Johnston and Willow, 1981) and appears to be due to an increase in the affinity of the high-aflinity GABA recognition site for GABA (Willow and Johnston, 1980, 1981a; Johnston and Willow, 1981; Willow, 1981). Picrotoxin, which is thought to act at GABA-activated chloride ionophores (Takeuchi and Takeuchi, 1969; Ticku et al., 1978), abolished the enhancement of GABA binding by pentobarbital (Willow and Johnston, 1980, 1981a; Johnston and Willow, 1981; Willow, 1981). This finding is in agreement with the observations that picrotoxin blocks the enhancement of GABA responses in immature rat dorsal root fibers (Evans, 1979), frog motoneurons (Nicoll and Wojtowicz, 1980), and cultured spinal neurons (Barker and Ransom, 1978). this result, together with the finding that anesthetic barbiturates displace radiolabeled dihydropicrotoxinin (a biologically active analog of picrotoxinin) from rat brain Pz + P3 membranes (Ticku and Olsen, 1978), suggests that bar-

42

MAX WILLOW AND GRAHAM A. R. JOHNSTON

biturates may enhance GABA responses by binding to a site closely linked to chloride ionophores associated with GABA receptors. In contrast, bicuculline methochloride, which is thought to inhibit GABA responses by interacting with the GABA recognition site, does not alter the enhancement of GABA binding by pentobarbital (Willow, 1981) consistent with a number of electrophysiologicaI observations (Curtis and Lodge, 1977; Evans, 1979; Nicoll and Wojtowicz, 1980). Kinetic studies have revealed that pentobarbital (100 yA4) slows the dissociation of GABA from rat brain synaptosomal binding sites (Willow and Johnston, 1981b). T h e dissociation of GABA from crude synaptosomal membranes is best described by a biphasic process. T h e initial rapid phase of GABA dissociation followed by the slower secondary phase probably represents the dissociation of GABA from low- and high-affinity sites, respectively (Olsen, 1980). T h e dissociation rate constant ( K - , ) for the slower phase was significantly reduced in the presence o f 100 p,\f pentobarbital. T h e reduction of the GABA dissociation rate constant in the presence of pentobarbital can alone account for the increased affinity of GABA for its higher affinity binding site in the presence of this agent, as determined in steady-state radioligand binding studies (Willow and Johnston, 1980; Willow, 1981). Furthermore, the reduction in the rate of GABA dissociation in the presence of pentobarbital may account for (a) the prolongation of GABA-activated conductance changes by barbiturates in cultured mammalian spinal neurons (Ransom and Barker, 1976; Barker and Ransom, 1978), and (b) the observation that the recovery of firing of feline dorsal horne interneurons following electrophoretic application of GABA zn Z J ~ V O is significantly prolonged following systemic or electrophoretic administration of pentobarbital (Lodge and Curtis, 1978). In addition to pentobarbital, other barbiturates enhance GABA binding to synaptosomal membranes (Willow and Johnston, 1981a; Willow et NI., 1981). Methohexital and thiopental, agents used clinically as intravenous anesthetics, were shown to be the most potent of those barbiturates tested so far (Willow at d ,1981) in enhancing GABA binding (50% maximal enhancement at 5.5 2 0.3 and 6.2 2 0.4 p M , respectively). Phenobarbital and barbital (EC5, values = 88 f. 6.0 and 101 6.1 y M , respectively) were less potent than pentobarbital (EC,, = 33.0 2 2.0 p;ll) in their ability to enhance GABA binding (Willow et al., 1981), whereas barbituric acid (at concentrations as high as 5 mM) was without affect. ‘The ability of anesthetic/sedative/anticonvulsantbarbiturates to enhance GABA binding correlates well with the ability of these agents to displace [3H]phenobarbital from a single class of low-affinity/highdensity binding sites in crude synaptosomal rat brain membranes (Wil-

*

PHARMACOLOGY OF BARBITURATES

43

low et al., 1981). This binding site is apparently distinct from GABA recognition sites, diazepam binding sites, and also picrotoxin binding sites. Picrotoxinin, which abolishes the enhancement of GABA binding by barbiturates (Willow and Johnston, 1980, 1981a; Willow, 1981), does not affect phenobarbital binding to synaptosomal membranes (Willow et al., 1981). However this does not necessarily preclude an interaction between barbiturates that bind to the phenobarbital binding site, and ligands that bind to the picrotoxinin binding site (Ticku and Olsen, 1978; Ticku et al., 1978). It has been proposed that picrotoxinin may interact with the C-5 side chains without displacing the entire barbiturate molecule (Willow et al., 1981). Binding to the phenobarbital binding site per se may only require the presence of the barbituric acid ring (since barbituric acid displaces THIphenobarbital from synaptosomal binding sites), whereas pharmacological activity and interaction with picrotoxinin-like ligands requires the presence of C-5 substitutes on the barbituric acid ring. Thus, the ability of barbiturates to enhance GABA binding (by altering receptor affinity) at concentrations consistent with those measured in the brains of laboratory animals during anesthesia (Richards, 1979, supports electrophysiological studies that have shown that the enhancement of GABA responses by barbiturates is mediated by an increase in receptor affinity (Evans, 1979; Nicoll and Wojtowicz, 1980). Olsen (1981) has demonstrated a barbiturate enhancement of GABA binding which is anion dependent and characterized by an apparent increase in the number of high-affinity GABA binding sites. In contrast to our findings, the method of membrane preparation is not such a critical factor in the demonstration of this effect. The concentrations of barbiturates required to produce this effect are three- to fivefold higher than concentrations that enhance GABA binding by altering receptor affinity (Olsen, 1981). It is not clear whether this anion-dependent enhancement of GABA binding by barbiturates is related to the potentiation of GABA responses observed in viva and in vitro, since an increased GABA binding density (Olsen, 1981) would not account for the prolongation of GABAactivated chloride channels (see Barker and Mathers, 198 1) or necessarily enhance GABAergic transmission if spare receptors exist for GABA. It will be of considerable interest to examine the relationship between barbiturate binding sites that mediate the enhancement of GABA binding (Willow and Johnston, 1980; Olsen, 1981) and those sites that are involved in the competitive inhibition of rH]dihydropicrotoxinin binding by barbiturates (Ticku and Olsen, 1978). I n addition to their ability to enhance GABA responses, barbiturates have also been shown to possess direct GABA-mimetic actions at higher

44

MAX WILLOW AND GRAHAM A. R. JOHNSTON

concentrations (Evans, 1979; Nicoll and Wojtowicz, 1980; Barker and Mathers, 1981; Schulz and MacDonald, 1981). There is now some evidence that these two actions of barbiturates are mediated at two separate sites (Willow and Johnston, 198lc). In extensively washed synaptosomal membranes, pentobarbital enhanced GABA binding in a dosedependent manner (12.5-500 p M ) ; at concentrations greater than 500 P M , this enhancement was progressively reversed toward control levels of GABA binding (Willow and Johnston, 1981~).T h e effect of pentobarbital, seen at higher concentrations, may reflect a GABA-mimetic action, since similar concentrations enhanced rH]diazepam binding to extensively washed membranes, an effect antagonized by both bicuculline methochloride and picrotoxinin. This effect of pentobarbital on diazepam binding is similar to that produced by GABA and various GABA analogs (Tallman et al., 1978; Chiu and Rosenberg, 1979; Dudai, 1979; Karobath and Sperk, 1979) and is in agreement with previous studies demonstrating a barbiturate enhancement of diazepam binding to membranes of neuronal origin (Leeb-Lundberg et al., 1980; Skolnick et al., 1980; Ticku, 1981). When the crude synaptosomal membranes were incubated in the presence of the nonionic detergent Triton X-100 (0.5% v/v, 30 min at 37"C), the enhancement of GABA binding by low concentrations of pentobarbital was abolished, whereas at higher concentrations GABA binding was progressively inhibited. This finding suggests that the site mediating the GABA-mimetic action is retained. T h e loss of barbiturate enhancement of GABA binding following Triton X-100 incubation is in agreement with the observation that a class of barbiturate binding sites, possibly involved in the enhancement of GABA binding, is solubilized following treatment with this detergent (Willow et al., 1981).

IV. Conclusions

A number of conclusions can be made concerning the mechanisms of actions of depressant barbiturates. It appears that barbiturates depress excitatory synaptic transmission at a number of CNS synapses. This action may be due in part to a reduction in the amount of excitatory transmitter released from nerve terminals or to an interaction of the drug with postsynaptic neurotransmitter receptors (e.g., glutamate). T h e reduction in transmitter release may be related to the effects barbiturates exert on various systems involved in the maintenance of low intracellular levels of ionized calcium (e.g., membrane calcium channels,

PHARMACOLOGY OF BARBITURATES

45

membrane-bound calcium pump, mitochondria1 and nonmitochondrial sequestration systems). T h e depression of transmitter release by barbiturates is unlikely to be mediated by actions on the conduction of impulses along axons at pharmacologically relevant concentrations. There is now considerable evidence to demonstrate that barbiturates enhance and prolong synaptic inhibitions in the CNS mediated by GABA. Electrophysiological and neurochemical studies suggest that the enhancement of GABA responses are due to an alteration in the affinity of the GABA receptor for its ligand following an interaction of the barbiturate with a binding site closely associated with picrotoxinin binding sites. I n addition, barbiturates may also exert direct transmitter-like effects in opening chloride ionophores. T h e mechanism of action of convulsant barbiturates is less well understood. While sharing many properties in common with depressants, convulsant barbiturates appear to have potent depolarizing actions at concentrations below 100 F M . This action, and the ability of these agents to increase spontaneous and evoked transmitter release, may be important factors contributing to their excitant properties.

Acknowledgment

We would like to thank Professor D. R. Curtis, Mike M. Tamkun, and Scott Sherman for their assistance in the preparation of this article.

References

Adams, P. R. (1976).J . Physiol. (London) 260, 531-552. Andrews, P. R., Evans, R. H., Johnston, G. A. R., and Willow, M. (1981). Expm'entia 37, 172- 174. Baker, P. F. (1972). Prog. Biophys. Mol. Biol. 24, 177-223. Banna, N. R., and Jabbur, S. J. (1969). Int. J. Neurophannacol. 8, 299-307. Barker, J. L. (1975a). Brain Res. 92, 35-55. Barker, J. L. (1975b). Brain Res. 93, 77-90. Prog. Anesthesiol. 1, 135-156. Barker, J. L. (1975~). Barker, J. L., and McBurney, R. N. (1979). Proc. R. Sac. London. Ser. B 206, 319-327. Barker, J. L., and Mathers, D. A. (1981). Trends Neurosci. 4, 10-13. Barker, J. L., and Ransom, B. R. (1978).J . Physiol. (Londm) 280, 355-372. Blaustein, M. P. (1968).J. Gen. Physiol. 51, 293-307. Blaustein, M. P., and Ector, A. C. (1975). Mol. Pharmacol. 11, 369-378. Blaustein, M. P., and Hodgkin, A. L. (1969). J . Physiol. (London) 200, 497-527.

46

MAX WILLOW A N D GRAHAM A. R . JOHNSTON

Blaustein, M. P., Ratzlaff, R. W..Kendrick, N. C., and Schweitzer, E. S. (1978a).J. Gen. Phy.no1. 72, 15-4 1. Blaustein, M. P., Ratzalaff, R. W., and Schweitzer, E. S. (1978b).J. Gen. Physiol. 72,43-66. Bloedel, J. R., and Roberts, W. J. (1969).J. Seurophysiol. 32, 75-84. Bloom, F. E.. Costa, E., and Salmoiraghi, G. C. (1965).J. Phnrmnrol. E s p . Thrr. 150, 244252. Bowery, N. G., and Dray, A. (1976). Sntzirr (London) 264, 276-278. Bradley, P. B., and Dray. A. (1973). H r . J. Phrir~ncird.48, 212-224. Bremer. F. (1970). Elertrornrrphnlugr. Clin. Sczrrophysiol. 28, 1- 16. Brooks, C . McC., and Eccles, J . C. (1947).5. Neurophy.sio1. 10, 349-360. Brown, D. A.. and Quillam, J. P. (1964a). B r . , ] . Pharfnurol. Chrmother. 23, 241-256. Brown, D. .4.. and Quillam. J. P. (1964b).Br. J. Phnrimrol. Chanothcr. 23, 257-272. Carmichael, F. J., and Israel, M. (1975).J. Pharinnrol. Exp. Ther. 193, 824-834. Catchlove, R. H. F., Krjevic, K., and Maretic, H. (1972). Can. J . Physiol. Phnrmarol. 50, 11 11-1114. Chance, B. (1956).In “Enzymes: Units of Biological Structure and Function” (0.H. Gaebler, ed.), pp. 447-463. Academic Press, New York. Chiu, T. H., and Kosenberg. H. C. (1979). E i i r . J . Phnrinnrol. 56, 337-345. Collins. G. G. S. (1980). Brain Res. 190, 517-528. Cctnnors, B. 14’. (1981).Bmiri Re.7. 207, 357-369. Cowger, M. L., Labee, R. F., and Mackler, B. (1962). .4wh. Biorhrm. Biophps. 96, 583-587. Crawford, J . M. (1969). Brain Res. 12, 485-489. l o g31-46. ~ Crawford, J. M. (1970). , ~ n i r r ~ / i n r ~ n r i r o9, Crawford, J. M., and Curtis, D. R. (1966).J. Physiol. ( h J l l d O f l ) 186, 121-138. Curtis. D. R., and Johnston, G. A. R. (1974). Ergrb. Physiol., B i d . Chem. Exp. Phnrmnkol. 69, 97-188. Curtis, D. R., and Lodge, D. (1977). S r i t i ~ w(Lotidon) 270, 543-544. Curtis, D. R., and Ryall, R. W. (1966). Ernin Res. 2, 49-65. Cutler, R. W. P.. and Dudzinski, D. S. (1974). Brnitr Hes. 67, 546-548. Cutler, R. M’.P., and Young, J. (1979). dVeurorhcm.Krs. 4, 319-329. Cutler, R. W. P., Markowitz, D., and Dudzinski, D. S. (1974). Brain Re.\. 81, 189-197. Di Polo, R., and Beauge, L. (1979). .Vatwe ( f . O f l d 0 1 2 ) 278, 271-273. Downes, H., and Franz, D. K . (1971).J. Phatwu-ol. Exp. Thcr. 179, 660-670. Downes, H., and Williams, J. K. (1969).J.Phnrmnrol. Esp. thrr. 168, 283-289. Dox, A . W., and Yoder, L. (1922).J. Am. Chrni. Sor. 44, 1141-1145. Dray, A., and Bradley, P. B. (1976). L3r.J. Phnrnurrd. 48, 212-224. Dudai, Y. (1979). Hrcrrri Rrs. 167, 422-425. Duggan, A. W., Headley, P. M., and Lodge. D. (1974). Hr. J . Phnrmnrol. 54, 23-31. Eccles, J. C. (1946).J. .Yeti Eccles, J. C. (1964). “The ses.” Springer-Verlag, Berlin. Eccles, J. C . , and Malcolm, J . L. (1946).J. Seurophysiol. 9, 139-160. Eccles, J . C., Eccles, R. M., and Fatt, P. (1956).J. Phyiol. (Luizdon) 131, 154-169. Eccles, J. C.. Schmidt, R., and Willis, W. D. (1963).J.Physiul. (Loridon) 168, 500-530. Eccles, .J. C.. Faber. D. S., and Tabovikova, H. (1971). Brain RCS.25, 335-356. Elliott, R. C., and Quillam. J. P. (1964). B r . J . Phnrmnrol. Chmother. 23, 222-240. Enna, S. J.. and Snyder, S. H. (1976).J. .Veurorhem. 26, 221-224. Ernster, L., Jalling, O., Low, H., and Lindberg, 0.(1955).Ercp. Cell. Rrs., Suppl. 3, 124- 132. Evans, R. H. (1979). Brairi Rrr. 171, 113-120. Fisher, R. S., Walker. J. T. and Plunimer, C. W. (1948)..41n.J. Clirr. Pnthul. 18, 462-469. Frank, G. B., and Sanders, H. D. (1963). Br. J . Phnrmricol. Chrmother. 21, 1-9.

PHARMACOLOGY OF BARBITURATES

47

Frazier, D. T., Murayama, K., Abbott, N. J., and Narahashi, T. (1975). Eur. J . Pharmacol. 32, 102-107. Galindo, A. (1969).J. Pharmacol. Exp. Ther. 169, 185-195. Gibson, W. R., Doran, W. J., Wood, W. C., and Swanson, E. E. (1959).J. Pharmacol. Exp. Ther. 125, 23-27. Gordon, M., Rubia, F. J., and Strata, P. (1973). E@. Brain Res. 17, 50-62. Grewaal, D. S., and Quastel, J. H. (1973). Biochem. J. 132, 1-14. Grinnell, A. D. (1966).J. Physiol (London) 182, 612-648. Guidotti, A., Toffano, G., and Costa, E. (1978). Nature (London) 275, 553-555. Harris, R. A. (1981). Biochem. Pharmacol. 30, 3209-3215. Hatefi, Y., Jurtshuk, P., and Haavik, A. G. (1961). Arch. Biochem. Biophys. 94, 148-155. Haycock, J. W., Levy, W. B., and Cotman, C. W. (1977). Biochem. Pharmacol. 26, 159-161. Heinbecker, P., and Bartley, S. H. (1940).J. Neurophysiol. 3, 210-236. Huang, L. Y. M., and Barker, J. L. (1980). Science 207, 195-197. Iravani, J. (1965). Naunyn-Schmiedebergs Arch. Exp. Pathol. P h a m k o l . 251, 375-395. Jessell, T. M., and Richards, C. D. (1977).J. Physiol. (London) 269, 42P-44P. Johnson, E. S., Roberts, M. H. T., and Straughan, D. W. (1969).J. Physiol. (London) 203, 261-280. Johnston, G. A. R.,and Kennedy, S. M. E. (1978).1n “Amino Acids as Chemical Transmitters” (F. Fonnum, ed.), pp. 507-516. Plenum, New York. Johnston, G . A. R., and Willow, M. (1981). Adv. Biochem. Psychopharmacol. 26, 191-198. Jori, A,, Bianchetti, A,, and Prestini, P. E. (1970). Biochem. Pharmucol. 19, 2687-2694. Kalant, H., and Grose, W. (1967).J. P h a m c o l . Exp. Ther. 158, 386-393. Karobath, M., and Sperk, G. (1979). Proc. Natl. Acad. Sci. U.S.A. 76, 1004-1006. Kleiderer, E. C., and Shonle, H. A. (1934).J. Am. Chem. SOC. 56, 1772-1774. Krnjevic, K. (1974). Physiol. Rm. 54,418-540. Krnjevit, K., and Phillis, J. W. (1963).J. Physiol. (London) 166, 296-327. Krupp, P., Bianchi, C. P., and Suarez-Kurtz, G. (1969).J. Pharm. Pharmacol. 21, 763768. Larrabee, M. G., and Posternak, J. M. (1952).J. Neurophysiol. 15,91-114. Larson, M. D., and Major, M. A. (1970). Brain Res. 21, 309-31 1. Leeb-Lundberg, F., Snowman, A., and Olsen, R. W. (1980).Proc. Natl. Acad. Sci. U.S.A. 77, 7468-7472. Leslie, S. W., Elrod, S. V., Coleman, R., and Belknap, J. K. (1979). Biochem. P h a m c o l . 28, 1437-1440. Leslie, S. W., Friedman, M. B., Wilcox, R. E., and Elrod, S. V. (1980). Brain Res. 185, 409-417. Levy, R. A. (1977). Prog. Neurobiol. 9, 211-267. Lidbrink, P., and Farnebo, L. 0. (1973). Neuropharmacology 12, 1087-1095. Lloyd, P. D. C. (1952). Cold Spring Harbor Symp. Quant. Biol. 17, 203-219. Lodge, D. (1979). Clin. Expl. Pharmacol. Physiol. 6, 686. Lodge, D., and Curtis, D. R. (1978). Neurosci. Lett. 8, 125-129. Ldyning, Y., Oshima, T., and Yokota, T. (1964).J. Neurophysiol. 27, 408-428. MacDonald, R. L., and Barker, J. L. (1979). Neurology 29,432-447. Massotti, M., and Guidotti, A. (1980). Life Sci. 27, 847-854. Massotti, M., Guidotti, A., and Costa, E. (198l).J. Neurosci. 1, 409-418. Mathers, D. A., and Barker, J. L. (1980). Science 200, 775-777. Minchin, M. C. W. (1980). Br.J. Pharmacol. 68, 131P. Miyahara, J. T., Esplin, D. W., and Zablocka, B. (1966). J . Pharmucol. Exp. Ther. 154, 118-127.

48

MAX WILLOW AND GRAHAM A. R. JOHNSTON

Napias. C., Bergman, M. O., Van Ness, M. O., Greenlee, D., and Olsen, R. W. (1980). Life Sri. 27, 1001-1011. Narahashi, T., Moore, J. W., and Poston, R. N. (1969).J. h‘eurohiol. 1, 3-22. Narahashi, T., Frazier, D. T., Deguchik, T., Cleaves, C. A., and Ernan, M. C. (1971). J . Phor~rtocd.EX^. Thm. 177, 25-33. Nicholls, D. G. (1978).Binrhrm.j. 170, 511-522. Nicoll, R. A. (1972).,]. Phyiol. (Luizdon) 223, 803-814. Nicoll. R. A. (1975a). Pror. Sa!l. &ad. Sri. C,‘.S.*4. 72, 1460-1463. Nicoll, R. A. (1975b). Bi.ain Res. 96, 119-123. Nicoll, R. A. (1978). Srienre 199, 451-452. Nicoll, R. A. (1980). Haiidb. Psyhopharmnrol. 12, 187-234. Nicoll, R. A., and Wojtowicz, J. M. (1980). Braitt Res. 191, 225-237. Nicoll, R. A., Eccles, J. C., Oshima, T., and Rubia, F. (1975). Nature ( . h d O i / ) 258,625-627. Oberg, K. E. (1961).Exp. Cell Re.i. 24, 163-164. Olsen, R. W. (1980). I,? “Psychopharmacology and Biochemistry of Neurotransmitter Receptors” (H. 1. Yamamura, R. W. Olsen, and E. Usdin. eds.), pp. 537-550. Am. Elsevier, New York. Olsen, R. W. (198l).J. SPtirurhem. 37, 1-13. Olsen, R. W., Ticku, hl. K., Van Ness, P. C., and Greenlee, D. (1978). Bruit/ r p s . 139, 277 -294. Olsen, R. W., Ticku, M. K., Greenlee. D., and Van Ness, M. 0. (1979). In “GABANeurotransmitters” (P. Krogsgaard-Larsen, J. Scheel-Kruger, and H. Kofod, eds.), pp. 165- 178. Munksgaard, Copenhagen. Peck, E. J., Miller, A. L., and Lester, B. R. (1976). Brain RP.i. Bull. 1, 595-597. Phillis, J. W., and Tebecis, A. K. (1967). LiJe Sri. 6, 1621-1625. Pincus, J. H., and Hsiao, K. (1981).Bruiii Reg. 217, 119-127. Pincus, J . H., and Insler, N. F. (1981). Bruin Res. 213, 127-137. Politoff, A. L., Rose, S., and Pappas, A. D. (1974).J. Cell Bid. 61, 818-823. Prichard, J. W. (1980). . 4 h . .Veurol. 27, 505-522. Proctor, W. R., and Weakly, J. N. (1976).J. PIiwiol. (London) 258, 257-268. Quastel, D. M. J.. Hackett, J. T., and Cook, J. D. (1971). Scieriw 172, 1034-1036. Quastel, D. M. J., Hackett, J. T., and Okamoto, K. (1972). Can. ,I. Phy.&l. Pharmacol. 50, 279-284. Quastel, J. H., and Wheatley, A. H. M. (1932). Biorhem. J. 26, 725-744. Quiltam, J. P.. and Shand, D. G. (1964). Hr. J . Pharmarol. ChemothPr. 23, 273-284. Rahamimoff, R., Erulkar, S. D., Alnaes, E., Meiri, H., Rothschenker, S., and Rahamimoff, H. (1975). Cold Spring Harbor Symrzp. Quatrt. Bid. 40, 107-1 16. Ransom, B. R., and Barker, J. L. (1975). A’ottdre (London) 254, 703-705. Ransom, B. R., and Barker, J. L.(1976). Brain Res. 114, 530-535. Reed, K. C., and Bygrave, F. L. (1974). Biorhem. J. 140, 143-155. Reed, K. C., and Bygrave, F. L. (1975). Aital. Biorhm. 67, 44-54. Richards, C. D. (1972).J. PIiwiol. (London) 227, 749-767. Richards. C . D., and Smaje, J. C. (1976). Br. J. Pharmnrol. 58, 347-357. Richens, A. (1969). Br. J . Pharmarol. 36, 294-311. Richter, J. A., and Waller, M. B. (1977). Biorhem. P h a r m r o l . 26, 609-615. Richter, J. A., and Werling, L. L. (1979).J. Seworhem. 32, 935-941. Roberts, M. H. T., and Straughan, D. W. (1967).J. Phyiol. (London) 193, 269-294. Robinson, J. D. (1978). FEES Lett. 87, 261-264. Rudomin, P. (1966). Brain Res. 2, 181-183. Sasaki, K.. and Otani, J. (1962).Jp)i.,I. Physiol. 12, 383-396.

PHARMACOLOGY OF BARBITURATES

49

Saubermann, A. J., Gallagher, M. L., and Hedley-Whyte, J. (1974). Anesthesiology 40, 41-51. Scarpa, A., and Azzone, G. F. (1969). Biochim. Biophys. Acta 173, 78-85. Schmidt, R. F. (1963). Pfluegers Arch. Gesamte Physiol. Menschen Tiere 277, 325-346. Schmidt, R. F. (1964). Prog. Brain Res. 12, 119- 134. Schoepfle, G. M. (1957). Fed. Proc., Fed. Am. SOC.Exp. Biol. 16, 114. Scholfield, C. N. (1977). Br. J . Pharmacol. 59, 507P. Scholfield, C. N., and Harvey, J. N. (1975).J. Pharmacol. Exp. Ther. 195, 522-531. Schulz, D. W., and MacDonald, R. L. (1981). Brain Res. 209, 177-188. Seeman, P. (1972). Pharmacol. Rev. 24, 583-655. Seyama, I., and Narahashi, T. (1975).J. Pharmacol. Exp. Ther. 192, 95-105. Shapovalov, A. L. (1963). Fed. Proc., Fed. Am. SOC. Exp. B i d . 23, Trans. Suppl., 113-1 16. Simon, J. R., and Kuhar, M. J. (1975). Nature (Lolidon) 235, 162-163. Simon, J. R.,Atweh, S., and Kuhar, M. J. (1976).J. Neurochem. 26, 909-922. Skolnick, P., Paul, S. M., and Barker, J. L. (1980). Eur. J. Pharmacol. 65, 125-127. Sobue, K., Ichida, S., Yoshida, H., Yamazaki, R., and Kakiuchi, S. (1979). FEBS Lett. 99, 199-202. Somjen, G. G. (1963).J. Pharmacol. Exp. Ther. 140, 396-402. Somjen, G. G., and Gill, M. (1963).J. Pharmacol. Exp. Ther. 140, 19-30. Staiman, A., and Seeman, P. (1974). Can. J . Physiol. P h a m c o l . 52, 535-550. Steriade, M., Wyzinski, P., and Halle, J. (1974). In “Basic Sleep Mechanisms” (0.PetreQuadens and J. Schlag, eds.), pp 143-203. Academic Press, New York. Swanson, E. E. (1934). Proc. SOC.Exp. B i d . Med. 31, 963-964. Swanson, E. E., and Chen, K. K. (1939). Q. J . Pharm. Pharmacol. 12, 657-660. Takeuchi, A., and Takeuchi, N. (1969).J. Physiol. (London) 205,377-391. Tallman, J. F., Thomas, J. W., and Gallager, D. W. (1978).Nature (London) 274, 383-385. Tebecis, A. K., and DiMana, A. (1972). Exp. Brain Res. 14, 480-490. Thesleff, S. (1956). Acta Physiol. Scaiid. 37, 335-349. Thomson, T. D., and Turkanis, S. A. (1973). Br. J. Pharmacol. 48,48-58. Ticku, M. K. (1983). Biochem. Pharmacol. 30, 1573-1579. Ticku, M. K., and Olsen, R. W. (1978). Life Sci. 22, 1643-1651. Ticku, M. K., Ban, M., and Olsen, R. W. (1978). Mol. Pharmacol. 14, 391-402. Toffano, G. Guidotti, A., and Costa, E. (1978).Proc. Natl. Acad. Sci. U.S.A. 75,4024-4028. Velluz, L., Mathieu, J., and Jequier, R. (1951). Ann. Pharm. Fr. 9, 271-275. Veselyunene, M. A., Gutrnan, A. M., and Lesene, V. A. (1971).Fannakol. Toksikol. ( K i n , ) 5, 520-522. Weakly, J. N. (1969).J. Physiol. (London) 204, 63-77. Westmoreland, B. F., Ward, D., and Johns, T. R. (1971). Bruin Res. 26, 465-468. Willow, M. (1981). Brain Res. 220, 427-431. Willow, M., and Bygrave, F. L. (1982).J. Neurochem. 39, 557-562. Willow, M., and Johnston, G. A. R. (1979). Neurosci. Lett. 14, 361-364. Willow, M., and Johnston, G. A. R. (1980). Neurosci. Lett. 18, 323-327. Willow, M., and Johnston, G. A. R. (1981a).J. Neurosci. 1, 364-367. Willow, M., and Jchnston, G. A. R. (1981b). Neurosci. Lett. 23, 71-74. Willow, M., and Johnston, G. A. R. (1981c).J. Neurochem. 37, 1291-1294. Willow, M., Bornstein, J. C., and Johnston, G. A. R. (1980). Neurosci. Lett. 18, 185-190. Willow, M., Morgan, I. G., and Johnston, G. A. R. (1981). Neurosci. Lett. 24, 301-306. Wilson, W. A,, Zbicz, K. L., and Cote, I. W. (1980). Adv. Neurol. 27, 533-540. Yoneda, Y., and Kunyama, K. (1980). Nature (London) 285, 670-673.

This Page Intentionally Left Blank

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS: A SEARCH FOR RELIABILITY’ By Aleiandro Bayonz, William J. Shoemaker, Jacqueline F. McGinty, and Floyd Bloom A. V. Davis Center for Behavioral Neurobiology The Salk Institute San Diego, Colifornia

..........

I. Introduction . . . . . . .

...........

1. Introduction

T h e isolation and identification of the endogeneous opioid peptides called enkephalins (Hughes et al., 1975) and the discovery of the biologically and chemically related endorphins (Li et aE., 1965; Bradbury et al., 1975; Guillemin et al., 1976) have generated one of the fastest growing fields in current neurobiological research? Investigations of the role played by these opiate-like substances in nervous function have confronted scientists with methodological probThe literature survey in this review was completed in early 1981. address: Instituto de Investigaciones Biomedicas, Universidad Nacional Autonoma de Mexico, Apartado Postal 70-228, 04510 Mexico D.F., Mexico. P-Lipotropin (P-LPH) contains the sequences of P-endorphin (P-LPH, 61-91), y-endorphin (P-LPH, 61 -77); a-endorphin (P-LPH, 61-76), and methionine5-enkephalin (P-LPH, 6 1-65). Leucines-enkephalin differs from methionine5-enkephalin in the carboxy-terminal amino acid. The term mdorphiiis commonly refers to P-endorphin and the naturally occurring p-endorphin fragments other than enkephalin, that is, a- and y-endorphin and C’ fragment @-LPH, 61-87). In this article endorphin, and endorphin immunorenctivity, designate P-endorphin, P-LPH, and the 31-K proopiomelanocortin for the practical reason that our radioimmunoassay reads all these molecules to the same molar extent. Similarly,enkehpalins will designate both methionine- and leucine-enkephalin (Met- and Leu-enkephalins, respectively).

* PrPsnit

51 INTERNATIONAL REVIEW OF NEUROBIOLOGY, VOL. 24

Copyright 0 1983 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-366824-7

52

ALEJANDRO BAYON

et al.

lems of identification and quantitation that demand more than traditional neurochemical expertise. This is due chiefly to the greater chemical complexity of peptides compared to classical neurotransmitters and because peptide levels in nervous tissues are very low. T h e twin demands of high specificity and high sensitivity can be met by methods based on stereomolecular recognition; that is, the molecule-to-molecule interaction that occurs in antigen-antibody recognition. Thus, for most of us, immunological detection has become the major tool in understanding the actions of biologically active peptides, since it discriminates molecules on the basis of chemical structure rather than biological actions. Most important, immunodetection has provided the possibility of using the same specific instrument for studying these peptides from both histological and biochemical aspects. Early studies have already indicated the power of the immunological approach in contributing to the study of the opioid peptides. Both radioimmunoassay (RIA) and immunocytochemistry (ICC) have been utilized in studies of the origin (Mains et al., 1977; Yang et a/., 1978), developmental relationships (Bayon et a/., 1979a), neural distribution and cellular localization (Bloom et a/., 1978a; Rossier et al., 1977a, Watson et al., 1977, 1978; Elde et al., 1976; Hokfelt ef al., 1977; Simantov et al., 1977a; Sar et al., 1978; Yang ~t a/., 1977; Kobayashi et al., 1978; Bayonet a / . , 1980b), neuronal transport and release (Bayon et al., 1978, 1980a, 1981; Iversen et al., 1978), and possible neural pathways (Cuello and Paxinos, 1978; Uhl et al., 1978; Gros et a/., 1978b; Rossier et al., 1979) of opioid peptides. I n this article, we seek to demonstrate the usefulness, as well as the limitations, of immunological methods as reflected in the study of the opioid peptides in our laboratory. We suggest this article may guide readers in two ways: first, for developing new methodology, and second, for evaluating the existing literature on endorphins and enkephalins and the burgeoning data expected in the 1980s on these and other neuropeptides. We shall approach the problems of immunodetection methodology beginning with the preparative steps of extraction and stabilization, commonly required in RIA and ICC. These initial procedures are emphasized here because we feel that they are critical in determining the outcome of the subsequent immunodependent detection. Following the technical considerations of tissue handling, special emphasis will be given to testing the sensitivity and the specificity of antisera in the experimental conditions used in RIA and ICC. Finally, the differences and similarities between RIA and ICC are discussed with regard to our own experience, which indicates that studies using a combination of RIA and ICC are needed in order to approach a valid interpretation.

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

53

r

0'

0

I

I

2

3 4 TIME (hr)

I

I

5

6

FIG. 1. Postmortem degradation of enkephalin in mice brains. One group of mice was killed and brains were homogenized in 4.5 ml water and incubated at 25°C for the indicated times ( 0 4 ) .Incubation was stopped by addition of 0.5 ml of 1.0 N HCl. The second group of mice was killed and the intact body remained undisturbed at room temperature (0-0). Brains were excised at the time indicated and homogenized in 5 ml of 0.1 N HC1. Met- and Leu-enkephalin levels were determined by RIA; data are expressed as the sum of Met- and Leu-enkephalin levels. (From Childers and Snyder, 1979.)

It. Tissue Processing, Extraction, and Handling

A. WITHIN-TISSUE STABILIZATION T h e study of postmortem changes of enkephalins proviczs the basis for choosing the procedures used to extract these peptides from tissues. Only recently has direct evidence been obtained as to the remarkable postmortem stability of tissue enkephalins (Childers and Snyder, 1979; also unpublished observations in our laboratory). When mice are killed and remain undisturbed, the brain levels of enkephalins d o not change significantly for 6 hr, contrasting with the fast degradation observed in homogenates of otherwise identically treated brains (Fig. 1). However, this apparent stability might be preceded by rapid fluctuations occurring at the time of sacrifice. I n this regard, there is contradictory evidence indicating both that enkephalin values are higher in microwaveirradiated animals than in decapitated animals (Yang et d,1977; Rossier et al., 1977b) and that no significant differences are found after either sacrificing procedure (Gros et al., 1978a). I n addition, it is uncertain whether microwave irradiation increases enkephalin levels in dead ani-

54

ALEJANDRO BAYON

I s F L t x S c L OF

et al.

TABLE I PROCEI)L'RLS OF KILLISG A S U TISSUE EXTRACTION ON ESRWHALISLEVELS I S K.AT S T R I A T V M ~ Radioimmunoassay (nmoYgm tissue)

Procedures

Met-en k

Leu-enk

Decapitation and Tris-buffer extraction (4) Decapitation and 0.1 .Y HCl extraction (12) Microwave irradiation and 0.1 *Y HCI extraction (10)

0.14 ? 0.04 1.20 ? 0.15

0.27 ? 0.03

1.51 2 0.33

" Rats were assigned in two groups for decapitation or microwave irradiation, respectively. Immediately after killing the striata were dissected on ice, homogenized in cold 0.05 .\I T'ris buffer or 0.1 HCI. Radioimmunoassay for Met-enk and Leu-enk and radioreceptor assay for endorphins were performed on the same extracts. (Values are means 2 SEM; number of experiments in parentheses. Partial reproduction from Gros ct id., 1978a.)

mals (Simantov rt al., 1977b; Yang ~t ai., 1977). However, most of the evidence consistently indicates that enkephalins are very stable within tissue, however, disruptive handling of the tissue destabilizes these peptides-probably through exposure to proteases not normally able to encounter the peptides. Thus, homogenization of tissues has been shown to promote enkephalin degradation when carried out in neutral buffers, but not in protease-inactivating concentrated acids (Rossier et al., 1977b; Gross P t nl., 1978a) (see Table I). In addition, we have gathered evidence indicating that dissection, slicing, incubation, or superfusion can produce a drop in the tissue enkephalin content (Fig. 2). During superfusions (Iversen rt ul., 1978; Bayon et n l . , 1978), and also probably during incubations, the decrease of enkephalin levels in the sliced tissue is mostly due to leakage of peptides followed by extracellular degradation, since addition of bacitracin to the perfusing medium partially protects the enkephalin released into the perfusate but does not prevent the lowering of the tissue level (Table 11). These findings further support the idea that intracellular enkephalins are stable as long as they remain inside the tissue. Studies on the tissue stability of endorphin-like immunoreactive substances present similar problems. Our first experience indicated that brain endorphin-immunoreactivity (IR) was stable after decapitation, yielding values similar to those obtained after microwave irradiation, provided that the tissue was heat inactivated prior to homogenization (see below) (Rossier et oi., 1977b). These results disagree with the report of Cheung et 01. (1977) who found that heat inactivation was not critical

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

55

(4)

(81

T

T

-L

L

I

(4)

I

n W

-

I

FIG. 2. Loss of enkephalin immunoreactive material in the rat globus pallidus as a consequence of slicing and incubation. Tissue slicing was performed at 200-pm intervals; incubations were for 60 min in Krebs-bicarbonate (oxygenated, with 0.1 % bovine serum albumin added). Enkephalin extraction and assay were as described in Iversen et al. (1978). 1 Unit = 1 ng of Leu-enkephalin immunoequivalents (expressed this way to allow comparison with Table 11); bars represent the means +. SEM with the number of experiments in parentheses. (From A. Bayon, F. Dray, W. Shoemaker, and F. E. Bloom, unpublished results.)

(using the radioreceptor assay and acetone-formic acid extraction). Shortly afterward, Liotta et al. (1978) reported on the postmortem stability of P-endorphin and P-lipotropin (P-LPH) in rat anterior pituitaries, the results of which showed that 6 hrs after decapitation, less than 20% of P-LPH had been degraded (at least, in part, to &endorphin; Table 111). The need for tissue integrity to preserve endorphin stability was

56

ALEJANDRO BAYON

et al.

TABLE I1

TISSVE CON-rEsr

OF- EXKEPHALIN IXIMUSOREAC.TIVITY I N RA.r

Conditions Freshly dissected At end of 40-min control superfusion" At end of 40-min superfusion in calcium-free medium A t end of 40-min superfusion with control medium without bacitracin

GLOBUS PALLIDUS

Number of experiments

Tissue enkephalin (units per globus pallidus sample)

4 10

11.32 '-c 0.46 4.62 & 0.38

4

4.06

?

0.52

4

4.17

rt

0.53

Tissue was cross-cut at 200-pm intervals and super-perfused at 37°C. Control medium was a modified Krebs-bicarbonate, oxygenated and containing bovine serum albumin 0.1%. Bacitracin concentration was 30 p g ml-'. 1 unit = 1 ng Leu-enkephalinimmunoequivalents. (For additional details refer to the original paper by Iversen et a/., 1978.)

indicated by the observation that freezing and thawing of the glands produced more than an 80% drop in the endorphin levels. Although postmortem conditions could differ in pituitary and in brain, the stability of endorphin IR in brain is also suggested by the apparently greater lability of P-LPH in pituitary than in brain homogenates (Graf et al.,

1979). TABLE 111 EFFECIOF TEMPERATKRE A N D FREEZE-THA~VIN(; O N IMMUNOREACTIVEP-LPH .ASU P-ESDORPHIN (EP) CONCESTRATIOSS IN RATPARS DISTALIS" Handling prior to homogenization

I>

P-LPHb

P-EP

LPH/EF

None 6 hr at room temperature Freeze, thaw Freeze, thaw, 6 hr at room temperature

6 6 6 6

15.9 12.1 2.9 c0.05

0.3 1.6 11.5 1.3

53.00 7.60 0.25 -

" Twenty-four freshly removed rat anterior pituitaries were arbitrarily divided into four pools of six each. Each pool was submitted to one of the following treatments: immediately homogenized; maintained at room temperature for 6 hr and then homogenized; frozen on dry ice and completely thawed prior to homogenization: and frozen and thawed and then left at room temperature for 6 hr prior to homogenization. All tissue pools in this experiment were homogenized in 0.2 ,MHCl. P-LPH and P-endorphin were separated by gel filtration. Immunoreactivity was quantified by using a p-endorphin antiserum that cross-reacts with P-LPH on an equimolar basis. (Modified from Liotta rt nl., 1978.) * pmoYlOO pg protein. Molar ratios.

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

57

Several attempts have been made to allow tissue handling without detriment to peptide stability. Although microwave fixation has been reported as satisfactory for whole brain (Yang et al., 1977; Rossier et al., 1977b; Gros et al., 1978a), its variable detrimental effect on anatomical structures makes it difficult to use it for studies requiring fine regional dissections. Furthermore, reproducible microwave inactivation requires a narrow range of animal sizes, and favors mice or small rats with no prior neurosurgical manipulation. Also, homogeneous, effective irradiation was difficult to achieve, that is, the amount of radiation required for good generalized heat coagulation of cortical regions and pituitary usually destroyed the diencephalon (Rossier et al., 1977b). As an alternative to this method, we have reported that in uiuo perfusion of the brain with mild paraformaldehyde fixative does not modify the amount of endorphins (Bayon et al., 1980a,b) or enkephalins (Hughes et al., 1977; Bayon et al., 1980a,b) extracted from the brain (but see Section III,B,l). Beside perfusion with fixative, the use of heat coagulation has been attempted to stabilize these peptides in the tissue. Boiling of brain tissue in Ringer’s solution or phosphate buffers destroys most of its enkephalin and endorphin content when gross pieces are processed (boiling in concentrated acid solution will be discussed later). However, during studies on prenatal development of the opioid peptide systems (Bayon et al., 1979a) where rapid brain perfusion is difficult, we observed that 2-3-min incubations of the whole head in excess volume of boiling Ringer’s stabilized the endorphin and enkephalin levels in the undissected whole brain and facilitated the anatomical dissection by coagulating the tissue and preserving the structures (normally of jelly-like consistency, the heated tissue acquires a “hard-boiled egg” firmness). The small size of these brains and their incipient skull could contribute to the effectiveness of this procedure in an embryo compared to an adult, but the differences in prenatal brain biochemistry may also play a role in the success of this procedure. When dissection of the tissue is not necessary or experimentally useful (e.g., blood, cerebrospinal fluid, tissue culture, or whole organs), stabilization of the enkephalin or endorphin levels can be more easily performed simultaneously with extraction, as will be discussed in the next section.

B. EXTRACTION A N D PROCESSING

Stability an.d recovery are the main considerations for an extraction procedure. Since disruption of tissue is a preliminary step in extraction, the instability of enkephalins in brain extracts, serum, and cerebrospinal

38

ALEJANDRO BAYON

et nl.

fluid (Hambrook rt cd., 1976; Meek rt nl., 1977; Miller et nl., 1977) was soon identified as the major problem for satisfactory recoveries. In contrast to enkephalins, @-endorphin is relatively stable in plasma, with no significant degradation after 30 min in rat serum diluted 1 : 10 at 37°C; on the other hand, in cerbrospinal fluid after intraventricular injection in the rhesus monkey, @-endorphin has a half-life of 1 hr (Rossier ut nl., 1977b; A4. K. Ommaya, F. E. Bloom, J. Rossier, and R. Guillemin, unpublished observations). In brain extracts, however, degradation of endorphins is faster than in plasma or CSF. Several attempts have been made, with variable success, to inhibit the responsible peptidases. Nearly complete inhibition of enkephalin degradation by brain peptidases has been achieved with millimolar concentrations of bacitracin (Miller rt nl., 1977; Musacchio rt nl., 1979), o-phenanthroline (Meek et d . , 1977; Musacchio et nl., 1979), and puromycin (Barclay and Phillips, 1978; Musacchio ut d . , 1979). Even mixtures of synthetic peptides have been used to protect enkephalins (Henderson rt a(., 1978). Inhibition of endorphin-degrading peptidases seems to be more difficult to achieve, although bacitracin plus the serine proteinase inhibitor BOC-i>Phe-Pro-Arg-H have been shown to offer some protection (Graf rt d.,1979). Peptidase inhibitors have been used only secondarily for opioid peptide protection (see Udenfriend et nl., 1979), probably because of the greater simplicity and reliability of protease denaturing procedures. Substantial enkephalin degradation occurs in buffered homogenates during the time between their preparation and heat deproteinization (Gros rt d . , 1978a). Alternatively, homogenization of fresh tissue in strong acid alone (0.1 h' HCl) seems to protect the enkephaiins (Gros et cli., 1978a; Miller rt d., 1978). Endorphins are protected by homogenization of fresh tissue in HCI but not in acetic acid solutions (Liotta ei nl., 1978; Rossier ut nl., 1977b). Yet some proteases have been shown to be active i n brain extracts even at pH 3.5 (Marks et d., 1974). In our current experience, a more effective procedure to protect these peptides is to inactivate the proteases before extraction in nonhomogenized tissue (see Section 11,A). Placing tissue samples in boiling 2 A' acetic acid for 15-30 min prior to homogenization prevents any significant loss of endorphin or enkephalin immunoreactivity. Although boiling in acetic acid causes the tissues to sweli, it inactivates the proteases fast enough to bring the brain a-endorphin values below the sensitivity of its assay, indicating that the usually measured levels of this &endorphin metabolite reflect endorphin degradation after disruption of the tissue. In earlier experiments (Rossier rt cil., 1977b) we found lower enkephalin values in brains after decapitation and boiling than after microwave irradiation. How-

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

59

ever, the fact that the brains were cut in several pieces after boiling explains the lower levels obtained. Liotta et al. (1978) have reported that homogenization in 0.2 N HC1, but not in 1 N boiling acetic acid, prevents degradation of P-LPH and P-endorphin in rat pars distalis. However, in their procedure tissue is not boiled prior to but rather after homogenization, and in those conditions endorphin degradation is to be expected. Different stabilization procedures and extractant mixtures have been used to extract endorphins and enkephalins, and the values obtained in several brain regions seem to agree. The possible extraction problems encountered in earlier reports have now been resolved (Simantov and Snyder, 1976; see also comments by Yang et al., 1977). Table IV summarizes and compares equivalent or alternative steps from the most commonly used extraction protocols. Current extraction procedures using reverse-phase resins also allow concentration of samples with low enkephalin content, making it possible to detect them in serum or CSF (Clement-Jones et al., 1980). Although most of the procedures in Table IV can be used for both enkephalins and endorphins, a stabilization and fractionation procedure has been recently described by Watson et al. (1979) that separates enkephalins from P-LPH in the first extraction step, precipitating the latter after freezing the tissue and homogenizing in ice-cold 0.4 N perchloric acid. In practice, the negligible cross-reactivity between the sera used for enkephalin and endorphin RIAs makes their separation unnecessary. A final consideration about the stability of opioid peptides during extraction is related to the sensitivity of certain amino acid residues, such as cysteine, methionine, and tyrosine, to oxidation. The use of antioxidants does not seem to be an absolute necessity in routine handling of enkephalins and endorphins. However, we have found it useful to add 0.1 5% mercaptoethanol when storing (at -20°C) tissue extracts containing enkephalin for long periods of time. Protection may also be advisable during certain chromatographic procedures (Rubinstein et al., 1977). Similarly, efficiency in the recovery of endogenous peptides from the tissue can only be estimated by comparison of the values obtained by different methods, since internal standards can only compensate for losses occurring after the initial extraction step. Acidic solutions alone and in combination with polar organic solvents have been widely used yielding, in general, satisfactory recoveries of internal standards of endorphins and enkephalins (see references in Table IV). But concentrated trichloroacetic or perchloric acid solutions (Fig. 3; see also Rossier et al., 1977b; Watson et al., 1979) may not fully solubilize molecular weight opioid peptides and cannot be used in general extraction method. For purposes other than tissue extraction, endorphins and enkephalins

60

ALEJANDRO BAYON

et al.

TABLE I V OUTIJNEOF BASICEXTRACTION PROCEDURES FOR ENDORPHINS A N D ENKEPHALINS Homogenization of tissue samples (extractant)"

HCI 0.1-0.2 A'* (ice cold)

~~

Elztrunattun M neuhalizatimz of ertrartant. Parttal punjicntzm

Acid-acetoned

Centrifugation (4°C)e Supernatant recovered

SPpnration of soluble peptides ~

Acetic acid 1-2 N e (boiling)

~

Adsorption on Amberlite XAD-2 elution 90% methanol, evaporation, redissolved'

Neutralization (strong base is used)#

Lyophylization and resuspension (in assay buffer)h

Acetone removed under stream of Nd or concentration zn vacuak, resuspension

Second centrifugation usually needed prior to assay ~

~

Homogenization in buffers has been discarded because of enkephalin degradation (see Miller al., 1978, and text). * Hughes et al. (1977); Miller et a1 (1978); Liotta et a / . (1978). Heat deproteinization may follow (Gros et al., 1978a). Rossier et al. (1977b); Bayon et al. (1978). When tissue is not boiled in acid before homogenization or fixed by microwave irradiation (Yang t r al.. 1978). homogenization in acetic acid leads to lower recoveries of endorphins (Liotta et al., 1978) and enkephalins (Miller et al., 1978). d T h e tissue is frozen prior to homogenization. The acid-acetone composition vanes: 0.2 M HCI-Acetone, 1 : 3 (Rubinstein ct al., 1977); acetone-HCI-H,O, 40:1 :5 (Smyth and Zakarian, 1979). In subsequent procedures, buffers are added with 0.01% thiodiglycoi and 0.0018 phenylmethyl-sulfonyl fluoride (a protease inhibitor), Rubinstein et al. (1977). Centrifugal force used varies from 1000 g, 30 min (Rossier et al., 1977b) to 50,000 g, 60 min (Miller et al., 1978). Filtration of supernatants in glass wool has also been used (Miller et al., 1978). This method has only been routinely used with enkephalins (Hughes et al., 1977). Adsorption in anionic-exchange resins has also been employed (Akil et al., 1978a). Rossier et al. (1977b): Akil el al. (1978a); Liotta et al. (1978). Miller et al. (1978); Bayon el al. (1978). Rubinstein et al. (1977). Lipids can be extracted with ethyl acetate-ether, 3 : 1. Smyth and Zakarian (1979).

PI

'

may also be safely handled when dissolved in aqueous buffers. The enkephalins, but not the endorphins, are soluble in organic solvents (e.g., methanol). Drying of the samples should be avoided when small amounts of these peptides are processed, since binding to glass and to several types of plastic has been observed. Siliconization of glassware does not completely prevent the loss of material, and bound peptides can only be brought back into solution with detergent-containing buffers (e.g., sodium dodecyl sulfate, 0.1 %). Whenever possible we process endorphin- and/or enkephalin-containing samples in polypropylene tubes

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

+I +I

+

.c

u I-

8- ENDORPHIN ( ng / brain )

61

-

ENKEPHALIN ( U-Enk/ brain )

FIG.3. Extraction of opioid peptides. Rats were killed by microwave irradiation (4 sec); their brains were removed, placed in 16 ml of the extraction media tested, and homogenized with a Brinkmann Polytron at setting 6 for 10 sec. After centrifugration (1000 g x 1 hr) supernatants were frozen overnight, thawed, and treated as follows: 1 N acetic acid extracts were neutralized to pH 7.5 with 1 N NaOH supplemented with 0.2 M Na2HP04,frozen overnight, thawed, and centrifuged (1OOOg x 1 hr). The clear supernatant was then processed in the RIAs. PBS supernatants were directly assayed. Five percent TCA supernatants were extracted three times with two volumes of diethylether and then adjusted to pH 7.5 with 0.2 M NazHP04.HC1-methanol (methanol, 90%; 0.1 N HCI, 10%) was evaporated and the residue diluted to the original volume with phosphate-buffered saline. After centrifugation (1000 g x 1 hr) the supernatant was assayed. U-Enk, Units equivalent to 1 ng of Leu-enkephalin. (From Rossier et al., 1977b.)

and in buffers containing bovine serum albumin (0.01%) and Triton X-100 (0.1 %), since we have observed that these precautions minimize losses (Guillemin et al., 1977). When tissue extracts are subjected to chromatographic separation, eluents like those employed during tissue stabilization have been frequently used to prevent unwanted interactions of these peptides with the support (resins, gels, etc.) and other components of the extracts. For example, enkephalin elution is delayed in commercial gel filtration media (Sephadex, Biogel-P) when nonchaotropic buffers are used; furthermore, small amounts of synthetic P-endorphin may interact with large tissue components. Strongly acidic or chaotropic solutions normalize the chromatographic permeation of both endorphin and enkephalin thus improving their recovery (Austen et al., 1977; Rossier et al., 1977a; Rubinstein et al., 1977; Liotta et al., 1978; Hughes et al., 1977; Yang et al., 1978). Similarly, permeation in dextran-coated controlled-pore glass beads requires elution in the presence of detergent to allow for a satisfactory recovery of endorphins (Bayon et al., 1979b). Ion exchange and

62

ALEJANDRO BAYON

et al.

adsorption resins have been used mainly to “clean-up” enkephalin- and endorphin-containing extracts (Hughes el d., 1977; Akil et al., 1978a), and more rarely for identification purposes (Smyth and Zakarian, 1979). Recovery yields from this process are rarely reported; when coupled with thin-layer chromatography (TLC), the overall enkephalin recovery can be as low as 30% (Henderson Pf nl., 1978). I n our own experience, recovery of enkephalins in the picogram (pg) level from silica gel plates ranges between 60 and 90%. I n spite of this relative disadvantage of TLC, it is frequently used because of its high resolving power for enkephalins (Hughes et u l . , 1977; Gros at nl., 1978a; Yang ut nl., 1979). High-pressure liquid chromatography (HPLC) was used to fractionate endorphins for the first time by Burgus and Rivier (1976). Other groups quickly adopted it for the fractionation and identification of endorphins and enkephalins because of its high resolving power and excellent recoveries (90-lOOrC) (Meek and Bohan, 1978; Rubinstein ri al., 1977; Bayon et ul., 1978). However, elution conditions have to be carefully designed to attain these high recoveries: Higher temperatures, deRivier, unpublished observations; A. Bayon, R. Burgus, and tergents F. E. Bloom, unpublished observations), or tertiary gradients (Rubinstein ut ul., 1977) are required. Electrophoresis in polyacrylamide-SDS gels has been of utmost importance in the study of the relationships among the endorphins and possible endorphin precursors (Mains and Eipper, 1978; Roberts et al., 1978). Endorphin yields higher than 80% have been attained by elution of gel slices; commonly the procedure consists of overnight incubation (37°C) in solutions containing detergents and chaotropic agents (but see also Yoshimi rt nl., 1978). All these fractionation methods have been used not only to separate and identify the various molecular species that contribute to the total immunoreactive endorphin- or enkephalin-like materials, but also to remove tissue components that may interfere with their assay. However, as will be discussed in Section 111, these fractionation methods may, by themselves, also introduce assay-interfering materials.

u.

111. IdentifKation. Quantitation, and Localization

A. E STA B LI S H I N G

sEN S I TI V I T Y 4 N D sPECI FI c I T

Those researchers with experience in radioimmunoassay understand its power and limitations. Without this experience one may find unex-

pected difficulties in interpreting certain RIA data related to the detec-

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

63

tion (presence or absence) and identification of the material measured by the assay. I n RIA, detection and identification are directly related to the problems of sensitivity and specificity, respectively. 1. Sensitiuity Leaving aside the discrepancies on the mathematical formulations and conceptual uses of the term sensitivity (see the papers by Berson, by Ekins, and by others, in Margoulies, 1969), in practice it is defined as the minimum amount of unlabeled antigen producing a statistically significant displacement of trace in RIA (Midgley et al., 1969). This definition implies that RIA, in contrast to most chemical assays, does not involve a “blank value,” that is, a change in maximum binding of the tracer antigen produced by factors other than competing antigens. In most other types of chemical assays, the blank value is usually independent of the sample tested, consisting usually of interfering components of the reagents. In RIA, however, the blank value is produced by sample components that frequently are unknown and that may interfere only in a particular assay system, that is, the sample may not produce a blank in a different RIA with similar specificity. Once the presence of a blank effect is suspected it has to be identified and its cause investigated. T h e most common feature to lead one to suspect a blank effect is the lack of agreement of sample antigen levels when tested at different doses. Moreover, the blank effect manifests itself by producing higher antigen readings at the lower doses tested. Direct identification of a blank effect is usually done by plotting the levels of antigen measured by the RIA at several volumes of the same tested sample; a blank problem is indicated by a straight line that does not intersect the origin? Figure 4 illustrates two examples of blank effects observed in our enkephalin RIAs of mice pituitaries and tissue perfusates. While assaying mice pituitaries (Fig. 4A), w e believed we had found the existence of a blank-producing factor because trace displacements produced by material extracted from the samples with chloroform accounted for the positive blank value. However, an interfering agent has not been positively identified. Conversely, bicarbonate used in superfusing medium was first shown to increase the maximum binding of When a plot of RIA values versus volume of assay sample results in a curved instead of a straight line, it suggests that complicating factors are present in the assay. The most common factors researchers must deal with are antigen and antibody heterogeneity, both of which are discussed below in great detail. Other factors are the pH and salt effects on the dissociation of the antigen-antibody complex and degradation of antibody or antigen. For a more detailed discussion of blank effects, see Ekins and Newman (1970) and Nugent and Mayes (1970).

64

ALEJANDRO BAYON

Ioc

et al.

20

A

50

1

2

M I C E PITUITARIES/ ML OF EXTRACT

5

i ''VOLUME PERFUSATE IN R I A (PI)

'

FIG. 4. Blank effects produced by some biological samples in a Leu-enkephalin RIA. (A) Positive blank effect due to decrease in maximum binding. Deproteinized (1 N acetic acid) 2-mi extracts from 1, 2, and 5 mice pituitaries were lyophylized, resuspended, centrifuged, and dissolved in 1 ml ofassay buffer, and its enkephalin content determined from 20-*1(-0-) and lOO-pl(-O-)aliquots in duplicates. The blank value is independent not only of the volume assayed (20- 100 pl), but also of the concentration of the extracts. Here, the

straight-line plots intercept the ordinate above the origin, giving a positive blank value to be subtracted from tissue values. (From R. Azad, A. Mauss, T. M. Vargo, A. Bayon, W. Shoemaker, and F. E. Bloom, unpublished results.) (B) Negative blank effect due to increase in maximum binding. I n this case RIA of different volumes of a Krebs-bicarbonate perfusate of rat globus pallidus slices are used as an example. T h e enkephalin values were determined from a standard curve not containing Krebs-bicarbonate solution. Results are means of triplicates ? SEM. The numerical value of the blank is the intercept with the ordinate, here a negative number-this value can be substracted algebraically from the RIA sample values to vield a correct value.

trace in our enkephalin RIA, and only later was this demonstrated to be a blank effect (Fig. 4B); for routine purposes the problem was solved by adding bicarbonate to the standard curves (Iversen et al., 1978). Since identification of the sample components producing a blank is not always feasible, fractionation methods are used to remove these ma-

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

65

terials. We have observed that in the absence of sample, effluents from several commercially available brands of reverse-phase HPLC supports raise the baseline of chromatograms in collected fractions measured by an enkephalin RIA (see Bayon et al., 1978). I n this regard, it has been reported that support media used in chromatography as well as residues from solvents may interfere in an additive manner in RIA systems (de Souza et al., 1970). These types of effects can be compensated for by processing the standards in the same way as the samples. An alternative to the removal of the interfering material is the assessment of blank values by using samples depleted of the antigen. However, extraction of the antigen has been reported to be unreliable because the interfering material was also removed (de Souza et al., 1970). On the other hand, hypophysectomy has been used to produce endorphin-free serum samples for use as a blank since plasma has been reported to interfere with endorphin RIAs (Ghazarossianet al., 1978; Akilet al., 197813). Even here, one cannot be certain that the pituitary is the sole source of &endorphin, making attainment of a true blank impractical.

2. Specajicity When obtaining a positive reading in RIA, the assayist needs verification that the substance detected is in fact the substance sought. The cross-reacting material can have the same antigenic determinants (e.g., a common amino acid sequence) as the immunizing antigen, and it will have the same afhity constant for the antibody (Kd). When crossreacting material possesses merely a similar antigenic determinant, the affiity constant will be ordinarily different from true standards. This situation is analyzed by plots of bound tracer versus antigen. Identical affinity constants of two substances yield parallel dilution curves in a RIA. However, parallelism in these plots is not sufficient by itself to demonstrate the presence of identical immunological determinants in two tested substances. Theoretical calculations (see Midgley et al., 1969) show that even 10-fold differences in the affiity constants of two crossreactants for the antibodies in the RIA may still yield curves that d o not depart significantly from parallelism. For example, the high molecular weight “immunoreactive &endorphin” found in extracts of human placenta has been shown to be a fragment of immunoglobulin G Uulliard et al., 1980). T h e hazards of interpreting RIA results (such as that referenced above) have led some to depend only on chemical characterization (e.g., column chromatography, disk gel electrophoresis, and sequencing of labeled peptides) (see Seidah et al., 1978); however, few neurobiologists have these intense chemical techniques available and must rely on the less technically demanding immunodetection systems.

66

ALEJANDRO BAYON

et al.

The unreliability of the parallelism criterion is exemplified in cases of peptide degradation. Thus, it was recently reported that endorphin immunoreactivity, detected by RIA, is released from mast cells. It was subsequently demonstrated that all of this immunoreactive endorphin can be accounted for by a heat- and acid-sensitive proteolytic activity that destroys the iodine-labeled peptide (P-endorphin trace) during the course of the RIA. Not surprisingly, this false IR parallels the RIA standard curve. When the samples were boiled prior to assay, the proteolytic activity was inactivated resulting in stable trace and no endorphin IR (Di Augustine et al., 1980). Similarly, lack of parallelism between a standard curve and the dilution curve of a biological extract has been claimed as an indicator of the existence of a different affinity constant for the cross-reacting material (Ekins and Newman 1970). However, lack of parallelism is not a safe indicator of the absence of a common sequence in the standard and test materials, since different tertiary structures within the molecule may substantially modify the affinity for the specific antibody. Biphasic or asymmetric dilution curves are as common and usually easier to detect than lack of parallelism. Both are related to the presence of a heterogenous antibody population in the antiserum used in RIA. Heterogeneity in antibodies present in sera is not uncommon and is probably due to multiple antigenic determinants present during immunization. This heterogeneity might be uncovered when there are multiple antigenic sites in the samples (see Ekins and Newman, 1970). We have reported that myelin basic protein runs parallel to P-endorphin in the standard curve of our RIA (Fig. 5). Furthermore, immunocytochemical staining of some myelinated fibers was observed with an endorphin antiserum (Fig. 6; Bloom et al., 1978a). T h e presence of at least two types of antibodies in the antiserum was uncovered only when preincubation of the serum with purified myelin basic protein selectively blocked this unwanted staining, whereas preincubation with synthetic &endorphin abolished both the specific and the spurious immunoreactivity. Thus, one antibody population responsible for the myelin staining reads an unexpected common determinant in P-endorphin and myelin basic protein; another antibody population is highly specific for endorphin, with negligible cross-reactivity for myelin basic protein. Biphasic dilution curves differ from asymmetric curves in that different antibody populations are revealed by their degree of af€inity for the sample antigens and by their differing affinity for separate immunological determinants. Thus, certain antigen-related substances will not completely displace the tracer antigen. An example of this behavior has been documented by Ross et al. (1978), where peptide fragments lacking the

67

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

100

m" \

m 50

s

0.01

0.I

1.0 QUANTITY

10

100

FIG. 5. Cross-reactivity of P-endorphin antiserum RB263 with myelin basic protein (taken from Bloom et al., 1979). Note the brain extract curve is asymmetric compared to P-endorphin; this is due to the myelin basic protein present in the extract. BIB,,, Isotope bound/maximal bound (in absence of competing antigen). (0-0), P-Endorphin (nghube); (A-A), brain extract (mg wet weighdtube); (W-W), myelin basic protein (pg/tube).

free amino terminal of P-endorphin could not displace more than 70% of this tracer indicating the presence of a subpopulation of antibodies only reading the amino end of P-endorphin in this RIA (Fig. 7). T h e use of an impure tracer preparation with different affinities for the antibodies in the antiserum also leads to asymmetric curves. Gros et al. (1978a) have reported the separation by TLC of monoiodinated and diodinated enkephalins, the latter showing a much lower binding to their specific antisera (Table V). Similarly, we have found at least two different immunoreactive species of iodine-labeled synthetic P-endorphin after fractionation by reverse phase HPLC (Fig. 8). Thus, immunological identity cannot be established with certainty when employing a single RIA. Moreover, even combined use of RIA and receptor binding activity can be misleading-an unidentified factor interfering with both assays for methionine5 (Met)-enkephalin has been detected in brain (Takahashi et al., 1979). Houck et al. (1980) based their identification of a P-endorphin precursor from human placenta on both RIA and receptor binding assays. As we have discussed above, the substance identified is actually IgG( Julliard et al., 1980). Assaying the same sample in t w o RIA systems differing in antisera specificity or tracer antigen increases the probabilities of making a correct identification of the tested material. This approach has been used, coupled with HPLC frac-

FIG.6. Immunoperoxidase staining with RB263 of hypothalamic endorphin-containing fibers (A) and of cross-reactive myelinated axons of the cingulate gyrus of the rat (B). For details, see Bloom ef a/. (1978b).

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

100

69

-

80 60

-

40 -

s Ic)

p!

U

I 10 [PEPTIDE] nM

0.I

100

FIG.7. Competition of p,-endorphin (P-E),P,-endorphin (2-3l), &-endorphin ( 10-3l), and P,-LPH with ['z511p-endorphinfor binding to the antiserum raised to P-endorphin. (From Ross et al., 1978.) Note the biphasic curves (not completely drawn out) indicating lack of complete displacement of the trace.

TABLE V CHROMATOGRAPHIC DATAA N D RADIOIMMUNOLOGICAL PARAMETERS OF IODINATED TRACERS Bound radioactivity

ICSO of homologous unlabeled enkephalns (nM)

Productsb

RP

Met-enk Monoiodo Met-enk Diiodo Met-enk Monoiodo Met-enkd (modified) Diiodo Met-enkd (modified) Leu-enk Monoiodo Leu-enk Diiodo Leu-enk

0.61 0.38 0.42

-

-

80 30

0.57 -

0.70

0

(%)

0.75 0.85 0.72 0.78

0

~

-

-

87 62

0.55 -

~

Chromatography on silica gel TLC plates using n-BuOH : HOAc : HzO (4 : 1 : 1) as solvent. Met-enkephalin was iodinated with the HzOz-lactoperoxidase method; Leuenkephalin was iodinated with the chloramine-T method. For additional details see the original paper by Gros et al. (1978a). Calculated for 1 : 600 final dilution of antisera in the presence of 17,000 dpm of tracer. Products of secondary reaction in halogenation process. a

70

ALEJANDRO BAYON

et al.

c

P

0 X

n Y) N

Y

Q

a

FRACTION

NUMBER

FIG. 8. HPLC separation of the iodination products of @-endorphinand specific immunoprecipitation of the fractions with a p-endorphin antiserum. Synthetic @-endorphin was iodinated by the chloramine-T method and free lZsI removed by gel filtration in Sephddex G-25. An aliquot of the excluded peak was injected in a p-Bondapak-CN column (Waters Associates, Boston, Massachusetts) and eluted with a linear gradient of acetonitrile in 0.01 .\I ammonium acetate (pH adjusted to 4.0 with acetic acid). The flow rate was 2 niymin and 1-ml fractions were collyted in polypropylene tubes. Fractions were counted for radioactivity (-0-) and the specific antibody binding monitored in alternate tubes by double antibody precipitation in quadruplicate. @-Endorphinantiserum was used at a final dilution of 1 : 700 (expressed as bound/free counts, -O-). A similar binding profile was obtained at a final dilution of 1 : 7000 (not shown), but binding was proportionally higher in the peak eluting around fraction 20 than in the peak eluting around fraction 9. (From A. Bayon. R. Burgus, R. Guillemin. and F. E. Bloom, unpublished results.)

tionation, in the identification of Met- and leucine5 (Leu)-enkephalin in globus pallidus extracts and perfusates (Fig. 9; Bayon et nl., 1978). Similarly, coupling RIA with gel filtration procedures has become a routine procedure to improve the selectivity of immunodetection for endorphin and its possible precursors (see, for example, Gramsch et al., 1980). FIL. 9. HPLC fractionation of an extract from freshly dissected globus pallidus tissue. ‘I-he p-Bondapak-CN column was calibrated with unlabeled synthetic standards. (ME), Met-enkephalin; (LE), Leu-enkephalin. The other standards are named as fragments of porcine 0-lipotropin (P-LPH) or by using the single-letter amino acid nomenclature (Y, T y r : G, Gly; F, Phe; M. Met; L, Leu). Their retention times (arrows) were determined by

RETENTION TIME (MIN 1

2

4

6 W

J

A

?

a

10

20

30

40

FRACTION NUMBER absorption at 210 nm. The flow rate was kept at 2 mYmin and 400-plfractions of the effluent were collected. (A) Radioimmunoassay using both Met-enkephalin and Leuenkephalin antisera were performed with each fraction of the chromatograms (at two dilutions in duplicates). The values are expressed in terms of ng of Met-enkephalin ( 0 ) that would produce an equivalent immunodisplacement in the Met-enkephalin assay and, analogously, ng of Leuenkephalin (0)producing an equivalent displacement in the Leuenkephalin assay. The total immunoreactivity is that contained in the globus pallidus of one rat (the overall recovery being 96%).The content of Met-enkephalin and Leu-enkephalin in their respective peaks was estimated from the two radioimmunoassays and agreed within 5% variation. (B) Radioactivity profiles of samples, similar to those in (A), that were extracted in the presence of either tritiated Met-enkephalin (0)or tritiated Leu-enkephalin (0)(10,000 cpm per sample). The radioactivity is expressed as percentage of the cpm injected into the column. The overall recovery of radioactivity for either tracer was >go%. The radioimmunoassay profiles of these chromatograms were comparable to those in (A). (Taken from Bayon et al., 1978.)

‘TABLE V I RIA SPECIFICITY OF SOMEANTISERA RAISED A G A I N RENKEPHAI.INS~ I,

-.J

Fs

Peptide (antigen)*

Animal species and conjugateC

Met-enk

Rabbit succinyl Met-en k + succinyl hemocyanin to polylysine; CDI Rabbit hemocyanin; CDI Rabbit hemocyanin; glutaraldehyde Rabbit hemocyanin; CDI

Leu-enk Met-enk

Leu-enk

Met-enk

Rabbit ovalbumin; CDI

Tracer antigen and labeling methodd [3 H ]Met-enk

Cross-reactivity with’

Sensitivity

Other enkephalins (‘% )

Endorphins

p

pmol

10

a and

[3H]Leu-enk

5 2 pmol

10

a a n d p ()

[3H]Met-enk

5 10 pmol

<0.5

@-End(neg)

[3H]Leu-enk

5 1 pmol

c0.5

p-End (neg)

IIMer-enk (LP)

5 2 0 fmol

5

[lZ5

5 10

(< 1 %,)

a and p (0.19)

Reference Yang PI al. ( 1977)

Yang rt (11. (1977) Childers c/ nl. (1977, 1978) Childers et al. (1977, 1978) Cros rt al. (1978a)

Leu-enk Met-enk Leu-enk Met-enk Leu-enk Leu-enk

~

Rabbit ovalbumin; CDI Rabbit BSA; glutaraldehyde Rabbit BSA; glutaraldehyde Rabbit BSA; glutaraldehyde Rabbit BSA; glutaraldehyde Rabbit BSA; glutaraldehyde

a and

p (0.1%)

5 2 0 fmol

7

10 fmol

<1

p-End (neg)

10 fmol

<1

p-End (neg)

1 pmol

<0.2

p and a (0.2-0.4%)

[SH]Leu-enk

1 pmol

5

p and a (0.2-0.4%)

[12sI]Leu-enk (ChT)

2 fmol

3

p and a (<0.1%)

[12sI]Leu-enk (ChT) [1z51]Met-enk (ChT) [1z51]Leu-enk (ChT [SH]Met-enk

Gros ci al. (1978a) Miller et al. (1978) Miller et al. ( 1978) Akil et al. (1978a) Akil et al. (1978a) Rossier et al. (1977a)

a These RIA data exemplify various types of antisera specificity for the enkephalins in relation to immunization procedures, preparation of tracer antigen, and assay sensitivity. They d o not include all RIAs currently in use nor the specificity of antisera characterized for purposes of immunoprecipitation or immunocytochemistry. * Abbreviations: p-end, P-endorphin; a-end, a-endorphin; Met-enk, MeP-enkephalin; Leu-enk, Leu5-enkephalin. The coupled macromolecule is given first, the coupling agent last. In most cases conjugates were emulsified with complete Freunds adjuvant. Details on the immunization protocols usually appear in the references. Abbreviations: CDI, 1-ethyl-3-(3-dimethyl-aminopropyl)carbodiimide; BSA, bovine serum albumin. Iodination methods: ChT, chloramine-T method; LP, lactoperoxidase method. When not explicitly stated in the references, it was calculated from RIA data or from reported standard curves at 20% displacement of the bound tracer. Neg, Negligible (numerical data are not given).

’

74

ALEJANDRO BAYON

et al.

A gamut of antisera for both endorphins and enkephalins has been raised and tested in different laboratories (Tables VI and VII). Although most of the reported anti-Met-enkephalin and anti-Leu-enkephalin sera are highly selective for the particular pentapeptide, they all cross-react to a small extent with the other peptide. This cross-reactivity is a minor problem for their quantitation by RIA, but is a major and limiting factor for ICC studies, as will be discussed later. However, the loose specificity of an antiserum raised against Met-enkephalin that also recognizes the amino end of Leu-enkephalin has helped to detect possible enkephalin precursors found in the adrenal medulla and the striatum. These peptides are amino or carboxy extensions of the earlier discovered pentapeptides (Stern et al., 1979). Endorphin antisera have been obtained that show high specificity for particular segments of the @-LPH molecule. Although most of those in frequent use recognize the carboxy terminal of this molecule, some have been reported to read its aminoterminal half or the amino end of @-endorphin.T h e use of these newer antisera will increase the power of the immunological methods to study the enkephalins and endorphins. B . I MhlVSOC 1TOCHEMISTRI'

L'EKSUS

h DIOI hlhlUNOASSAT

1 . '11ethodological Coiiipatibility

The possibility of using the same immunological tool in histological and quantitative chemical methods has encouraged attempts to correlate the cellular distribution of endorphins and enkephalins with their regional levels in nervous tissue (Bloom et al., 1978a; Rossier et al., 1977a; Bayon et al., 1980a,b). Based on previous concepts and experience in dealing with this problem (Sternberger, 1973), our first investigations indicated the requirements and limitations involved in comparing results obtained from both approaches. Since tissue fixation is required for the preservation of histological structures, the effects of this fixation must be suitable for the immunocytochemical visualization of these peptides. T h e fixation should produce minimal damage to the antigens in the tissue and allow permeation of the antibodies into the cells; in addition, the fixation should be compatible with the stability and efficiency of recovery needed during the extraction procedures that precede the RIAs. This compromise has been met by using a rapid and mild fixation of the brain, consisting of transcardial perfusion (in 90 sec) of 150-200 ml of 5% paraformaldehyde in phosphate-buffered saline (pH 7.4), followed by a short (1-3 hr) postfixation period in the same fixative and a stepwise equilibration of the tissue into 18%sucrose in phosphate-buffered saline.

TABLE VII RIA SPECIFICITY OF SOME ANTISERA RAISED AGAINST ENDORPHINS~ Peptide (antigen)*

Animal species and conjugate'

Tracer antigen and labeling methodd

Sensitivity' (Pg)

P,-LPH

Rabbit

[12sI@h-LPH LP

s 100

P,-End

Guinea pig yh globulin; CDI

[lZsI@,-End LP

5 100

P,-End

Rabbit BSA; BDB

[12sI]P-End

50

[12sI]a-End ChT

50

Purified [1z511p,-LPH ChT

500

[1251@,-LPH(1-47) LP

50

a-End

P,-LPH

P,-LPH

Rabbit bovine thyroglohulin; BDB Rabbit

(same antiserum as above)

(purified) P-LPH)

(Po-LPH, 1-47)

Cross-reactivity with related peptides and possible antigenic determinant' P,-LPH (10% w/w-not parallel); &,-end, P-MSH (<0.3%);y-end, Met-enk, Leu-enk (neg) P,-End (100%);Ph-LPH (10% w/w); Met-enk (neg) determinant P-end (6-15) Po-LPH, 3 I-K proopiocortin (full molar cross-reactivity) Met-enk, y- and a-end (neg) determinant p-end (20-27) stabilized by its (6-19) sequence P-End (0.05%); P-LPH (0.08%); Met-enk (neg) determinant a-end (10-16) P-LPH (1-47) and (1-65) show higher affinity than P,-LPH, but only produce partial trace displacement a subpopulation of antibodies reads only the carboxy terminus of P-LPH; another subpopulation reads the amino end of P-LPH (interpreted otherwise in the original paper) P-LPH (50% w/w); P,-LPH (10%) p-, a - , and y-end, a (<0.1%) and P-MSH

Reference Rao and Li (1977)

Li et al. (1977)

Guillemin et al. (1977)

Guillemin et al. (1977) Desranleau et al. (1972)

Lissitsky t t a1 (1978)

(continued)

Peptide (antigen)*

Animal species and conjugateC

Tracer antigen and labeling methodd

Sensitivity'

(PFJ

P,,-End

Hnbhzt amino cellulose NaNQ

[1251]@,,-End Ch?'

a-End

Ritbhit bovine thyroglobulin; CDI Rabbit unconjugated

[Iz5

<50

[1251&3,-LPH ChT

25-75

Ph-LPH

I]a-end ChT

1400

Cross-reactivity with related peptides and possible antigenic determinant' P-End (6-3 1) and P,,-LPH produce partial displacement of tracer. Enkephalins (<0.001% wlw) a-end (<0.04%) a subpopulation of antibodies is directed against &end (10-3) with a contribution of the (20-27) sequence (See Fig. 7) P-End (14- 18% w/w); P-LPH ( 4 4 % ) ; enkephalins (<0.004%) determinant IS amino region of a-end Antiserum was purified by affinity chromatography to remove p-end binding sites. P,,-LPH (8%);MSH, a-, P-, y-end (neg) full crossreactivity with y-LPH; P-LPH (1-58) determinant is the amino terminus of P-LPH (A similar antiserum for P,,-LPH has been reported by Krieger ct nl., 1977b.)

Reference Ross rt a1 (1978)

Ross rt 01. (1978)

Krieger rt nl. (1977a, 1979)

&-End

&End (C fragment)

Rabbit (crude porcine ACTH preparation) Rabbit bovine y globulin

~

4 -rl

[1251]&,-end (purified) ChT

5

[1Z51]P,-end

<150

~

&-LPH (5.5% w/w); a-end, enkephalins (<0.1%)

~

determinant is carboxy terminus of p-end C’ fragment, i.e., P-end (1-26) (25% w/w) P,-LPH (16%),Met-enk (neg) N-acyl C and C‘ fragments show identical cross-reactivity than their parent molecules) determinant carboxy terminus of P-end ~

~

~

~

~~

Yoshimi et al. (1978)

Smyth and Zakarian (1979)

~

~

These RIA data exemplify various types of antisera specificity for different segments of P-LPH in relation to immunization procedures, preparation of tracer antigen, and assay sensitivity. They do not include all RIAs currently in use nor the specificity of antisera characterized for purposes of immunoprecipitation or immunocytochemistry. * Abbreviations: P-LPH, p-lipotropin; p-end, P-endorphin; a-end, a-endorphin; MSH, melanocyte-stimulating hormone. Subscripts: h, human; 0, ovine; p, porcine; c, camel. The coupled macromolecule is given first, the coupling agent last. In most cases conjugates were emulsified with complete Freund’s adjuvant. Details on the immunization protocols usually appear in the references. Abbreviations: CDI, l-ethyl-3-(3-dimethyl-aminopropyl)carbodiimide; BDB, bis-diazotized benzidine; BSA, bovine serum albumin. Iodination methods: ChT, chloramine-T method; LP, lactoperoxidase method. When not explicitly stated in the references, it was calculated from RIA data or from reported standard curves at 20% displacement of the bound tracer. Neg, Negligible (numerical data are not given).

78

ALEJANDRO BAYON

et al.

TABLE V l I I EFFECXOF B R A I X FIXA nos os EXTWACT.AHLE EXWKPHIN ~ N ESK EPH .~LIN I\I\IL-.UORE.~C:TI\.ITI

Leus-enkephalin-IR ( p g h g tissue)a

Endorphin-IR (pg/mg tissuey Rat brain region Hypothalamus (ventral) Septa1 area Periaqueductal gray matier

D

Fresh tissue

Fixed tissue

286 t 73 97% 6

163 58 9 0 2 17

*

Fresh tissue

Fixed tissue

55 t 5 60 4

53 t 9 54 2 9

*

51 t 7 42* 3 27 t 3 24 ? 3 _ ~ a Mean of five deternunations % SEM Endorphin and enkephahn IR were measured from the same tissue extracts. (For details on method of fixation and extraction see text and Bacon c t a1 , 1980a ) _

_

~~

~

~

~

T h e success of this procedure as a preparative step for the ICC of endorphins and enkephalins has been reported (Bloomrt al., 1978a,b). As a tissue stabilization method it does not significantly modify the amount of enkephalin extracted from brain. Brain endorphin IR is also efficiently recovered, except in the hypothalamus where there is a nonsignificant but consistent decrease in its measurable levels (Table VIII; Bayon et al., 1980a). Although w e may assume that this decrease in hypothalamic endorphin content could be due to attachment of endorphins to the tissue during the process of fixation, susceptibility of the endorphin-IR material to the fixative has to be taken into account. Although it is possible to perform RIA and ICC on identically prepared tissue samples, there are independent problems that should be considered. On the one hand, we have dissection and extraction of tissues, which are part of the routine protocol for RIA. On the other hand, we shall refer to their counterpart in ICC, namely, accessibility of the ligands in the tissue to the antisera. A dissection procedure following classical anatomical regions may or may not produce boundaries corresponding to the zones of distribution of immunoreactive material. Thus, although the correlation of RIA and ICC data from both endorphins and enkephalins in mammalian brain has been satisfactory when using this type of dissection (Rossier ei ul., 1977a; Bloom Pt nl., 1978a), the correlation obtained in avian brain was even better since the regional dissection was designed following the gross anatomical areas that intensely stain using ICC (Bayon r t ul., 1980b). One obvious conclusion to be drawn from these results is that the morpholog-

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

79

ical mapping of the substance is the ideal guide to a dissection scheme. This is also exemplified by the studies on endorphin levels in the hypothalamus (Rossier et al., 1977a), and the enkephalin distribution within the amygdaloid complex and the bed nucleus of the stria terminalis (Gross et al., 1978b); in these cases the ICC data were obtained before quantitative anatomical distribution was studied. On the other hand, the use of regional RIA in screening tests for subsequent ICC studies may sometimes be advantageous, although the RIA will frequently overlook isolated and small spots of high IR content. For example, enkephalin RIA values in the hippocampus and neighboring areas would predict, and, in fact, positively correlate with the immunohistochemical staining in this area (Fig. 10). However, we also found (Bloom et al., 197813) small and highly localized groups of enkephalincontaining cells in the dorsal subiculum, whose existence could not be suspected based on RIA measurements. T h e problem of verifying that all the antigens read by ICC can be extracted and measured in the RIA is of special interest. Immunoreactive material may not be fully extracted and measured by RIA because of solubility or stability problems, but is nevertheless read in the ICC; this elusive possibility often cannot be completely ruled out. However, its likelihood is minimized by using different extractant mixtures; in preliminary experiments we have not found significant differences in enkephalin-1R levels when extracting fixed tissue with 1 N acetic acid with or without 0.1% Triton X-100 added. The requirements for cell penetration by the antisera remain obscure. It is generally assumed that membranes constitute the limiting barrier to the antibody, and consequently, detergents such as Triton X-100 are widely used in the incubation media. However, work performed in our laboratory (J. McGinty, unpublished results) and by others has shown that physical variables (i.e., increased area of contact between the tissue and the incubation media by using free-floating sections instead of adhering them on glass slides) markedly influence the number and types of cells and processes immunodetected. The different methods of antibody exposure also produce differences which are more apparent than real; thus free-floating sections, normally thicker than slide-mounted samples, are exposed for two surfaces and therefore will always show more than twice the density of stained structures, although a strict quantitative estimate of staining yield is difficult to attain. Now that the basis for methodological compatibility has been established, comparison of immunological specificity in RIA and ICC can be considered. Antisera normally contain a heterogeneous population of antibodies with various degrees of specificity and antigen-binding capac-

FIG. 10. Distribution of enkephalin immunoreactivity in hippocampal areas, subiculum, and entorhinal cortex. (A) Leuenkephalin RIA equivalents for bilateral hippocampi in 200-gm rats. Mean of five animals f SEM. (From A. Bayon, unpublished.) (B) Immunocytochemical localization of enkephalin IR in the hippocampal mossy fiber system of a kainic acidtreated (1 pglpliter) rat. Note dense fiber distribution in dentate hilus and CA3 field. Calibration bar = 500pm. FD, Fascia dentata; 1-4, CA fields of hippocampus. Inset: Enkephalin-IR dentate granule cell. Calibration bar = 15 p m .

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

81

ities (see Section III,A,2). Therefore, different subpopulations of antibodies might intervene to different extents in the RIA reactions and in the ICC process. It has been our experience that some antisera suitable for RIA cannot be successfully used for ICC (the opposite case can rarely be observed); we have also found that using different antisera for ICC and RIA may lead to discrepant results. Furthermore, even when using a single antiserum for both RIA and ICC, one cannot be assured of its identical performance in both methods. T h e main practical reason why the antisera specificity obtained in RIA conditions may not apply to ICC resides in the widely different antiserum dilutions frequently required in the two methods; at the higher antiserum concentrations used in ICC, antibodies not easily detected by RIA may significantly intervene in the reaction (Swaab et al., 1977). This problem is aggravated by the differences in the validation tests used in RIA and ICC. As shown in Table IX, these tests are not equivalent in every respect, although to some extent they are complementary. Thus RIA specificity tests do not define the specificity of an antiserum for ICC (see also Swaab et al., 1977). Several possible links between ICC and RIA specificities can be envisaged at this point: Immunoprecipitation may serve, in principle, to compare the specificity of an antiserum under ICC and RIA conditions since it uses low antibody dilutions as in ICC, and cross-reactivity studies may be performed with the same test tube procedures used in RIA. As an example, precipitation of tritium-labeled Met-enkephalin by our Leuenkephalin antiserum is negligible in RIA conditions (Leu-enkephalin bound = 15%; final antiserum dilution, 1 : 3500). However, when a final antiserum dilution of 1 : 500 is used, both tritiated enkephalins are precipitated to a similar extent (30-40%). Similarly, the cross-reactivity of Met-enkephalin in this assay increases two- to threefold at this higher concentration of antiserum. Aside from the implications about the possible heterogeneity of this antiserum, these observations would predict a lower selectivity of the antiserum in ICC than in RIA. T h e use of model systems for ICC as the one developed by Stumph et nl. (1974) and Raamsdonk et al. (1977) offers more attractive possibilities: Thin longitudinal sections of SDS-polyacrylamide gels of tissue extracts can be stained after fixation by an indirect peroxidase method allowing identification and quantification of tissue antigens as in RIA but under quasi-ICC conditions. However, one of its obvious limitations, as in immunoprecipitation, is that the loss of cellular structure and compartmentation appears to be the more complex source of differences between ICC and RIA, as will be noted later on. Other model systems, such as hormones bound to sepharose beads that are subsequently adhered to

Property tested

RIA

ICC

Nonimmunological binding

Tat-substitution of preimmune serum for antiserum (tests nonspecific aii/tgeii binding) Rmtl/--residual binding is defined as nonspecific

Test-substitution of preimmune serum for antiserum (tests nonspecific antivruin binding) Result-residual staining is defined as nonspecific

Antibody -antigen specificity (crossreactivity of antigenrelated species)

Tus/-RIA displacement curves with antigenrelated cross-reacting species ( B E vs log ligand) Result--full displacement of tracer antigen yielding symmetrical curves suggests common or closely related antigenic determinants in antigen and cross-reactant; quantitates extent of cross-reactivity

Trst-adsorption of antiserum with antigenrelated cross-reacting species.

Not detected in routine cross-reactivity tests using antigen as a tracer. Does not affect RIA and is not a source of artifacts

7est-adsorption of antiserum with the homologous antigen

Antiserum recognition of nonhomologous antigens (antibody subpopulation binds nonhomologous tissue components)

Result-blockade of staining equal to that produced by adsorption with antigen suggests common or closely related antigenic determinants; not quantative.

Result-blockade of staining related to the antigenic determinants; residual staining due to antibody populations binding to nonhomologous antigens in tissue Antiserum recognition of homologous antigens (a) separate antibody subpopulations recognize different determinants in the homologous antigen

(a) Tes/-RIA displacement curves with f r a g ments of the antigen

(a) Tust-adsorption of antiserum with fragments of the antigen

Result-incomplete tracer displacement with plateau of curve at higher ligand concentrations

Result-incomplete blockade of staining when compared to that obtained with the antigen; might affect intensity, elements stained or both

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

83

TABLE IX (continued) Property tested (b) separate antibody subpopulations recognize an antigenic determinant to different extents (affinity)

RIA

ICC

(b) Test-RIA displacement curves with unpredicted crossreactants

(b) Test-adsorption of antiserum with unpredicted cross-reactants

Result-full displacement of tracer yielding assymetric or biphasic curves

Result-usually blockade of staining comparable to that attained with antigen adsorption is observed; when affinities or amount of the antibody subpopulations are widely different incomplete blockade is observed; might affect staining intensity, elements stained or both

glass slides (Swaab and Pool, 1975), have even more limited potential. It is worth referring here to the first application of this latter technique in the cellular localization of enkephalins. Several groups (Elde et al., 1976; Hokfelt et al., 1977; Saret al., 1978) have found that, due to the small but measurable cross-reactivity of their antisera for Met- and Leuenkephalin, either antiserum could label the same cells in immunocytochemistry. This technical limitation did not allow the localization of the two enkephalins in the same or different cells. This limitation can be reduced, but not eliminated, by decreasing the effective crossreactivity of the two antigen-antiserum systems. However, Larsson et al. (1979) have claimed, based on ICC experiments and model systems, that the two enkephalins are located in different neurons. Since they have not completely eliminated the cross-reactivity of their antisera, their results show, at most, that some cells are enriched in one of the enkephalins. The identification of a high molecular weight peptide containing the sequences of both Met- and Leu-enkephalin (Kimura et al., 1980) adds further doubt to this postulated cellular separation of the two enkephalin systems. An alternative solution could be provided by the use of monoclonal antibodies (Kohler et al., 1978), but this more sophisticated preparation has only recently been introduced to the study of opioid peptides. However, preliminary experiences in this laboratory indicate that problems unrelated to antibody heterogeneity will persist when

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

85

using these new tools: Some monoclonal antibodies show peptide binding in solution but give no specific tissue staining. Even compatible preparation and processing of the tissues and good correlation of specificity-validation tests between RIA and ICC will not guarantee reliability of the immunodetection data. The chemical reactions in the tissue are surely much more complex than in the assay tube. This is true, not only in relation to the cellular barriers and compartments, but also for the differing kinetics and stoichiometry of the reactions that probably are not occurring in solution.

2. Changes in Staining Intensity and in R I A Values An operant, practical validition of the complementarity of RIA and ICC would require the correlation of changes of content or distribution of endorphins and enkephalins after perturbation of the system. We have induced peptide changes (Bayon et al., 1980a) by using the antimitotic agent, colchicine, which disrupts microtubules and impairs axonal transport, consequently producing an accumulation of peptide in their neuronal perikarya (Inoue and Sato, 1967; Schmitt, 1968). The increase of enkephalin IR in ventral hypothalamus observed both by RIA and ICC techniques after intracerebroventricular or intracisternal injection of colchicine is documented in Fig. 11. Enkephalin perikarya were visualized only in colchicine-treated animals, and colchicine treatment increases the assayable enkephalin about 50%. The-hypothalamus is rich in enkephalin-containing cell bodies, and there is evidence indicating that some of them may give rise to a relatively long fiber tract reachFIG. 11. Changes induced by colchicine treatment in endorphin- or enkephalincontaining neuronal systems in rat brain. (A and B). Endorphin-immunoreactive perikarya in the ventral hypothalamus (area of the arcuate nucleus) in control (A) and colchicinetreated (25 p g 125 PI) rats (B). In treated animals there is a more intense, but diffuse staining. Also, extractable endorphin immunoreactivity was higher in colchicine-treated animals (362 2 74 pg of endorphin immunoequivalentdmg tissue) than in saline-injected controls (163 +. 58). (C and D). Enkephalin immunoreactivity in the rat globus pallidus at the interface with the caudate nucleus in control (C) and colchicine-treated animals (D). A more intense fiber staining as well as previously undetected perikarya (arrows) are visualized in colchicine-treated animals. However, extractable enkephalin levels in the pallidum are not significantly different in treated (291 24 pg of enkephalin immunoequivalents pg/mg tissue) compared to control animals (333 30). CN, Caudate nucleus. (E and F) Enkephalin IR in the hypothalamus. The area of the paraventricular nucleus 48 hr after lateral ventricular injection of saline (E) or colchicine (25 pg/25 p1) (F). Colchicine treatment reveals thicker fibers, as well as perikarya not observed in the controls. Arrow indicates a fine enkephalin IR fiber. There is a concomitant increase in the extractable enkephalin of the hypothalamus due to colchicine treatment. Controls, 53 9 pg; colchicine treated, 77 2 3 pg. Leu-enkephalin equivalentslmg tissue (Bar = 200 pm). (Further details in Bayon et al., 1980a.)

*

*

86

ALEJANDRO BAYON

et al.

ing the pituitary stalk and the posterior lobe (Rossier et al., 1979). T h e observation that assayable enkephalin IR in the posterior pituitary decreases after colchicine treatment in the same animals where hypothalamic enkephalin is augmented is in agreement with the idea that the enhanced enkephalin ICC staining is due to the peptide accumulation measured by RIA. In spite of the usefulness of colchicine to induce a “pile-up” of peptides in the neuronal somata, it has undesirable side effects that may impair the RIA-ICC correlations. It has been observed that colchicine treatment, in conditions similar to those employed in this study, disrupts cytoplasmic membranes (Hindeland-Gertner et nl., 1976). This could affect the compartmentation of peptides, modifying their intracellular distribution, but could also improve their availability to the antisera used in the ICC reaction. T h e ventral hypothalamus contains the only group of endorphin perikarya yet observed in the brain, both in colchicinetreated and normal animals. Colchicine treatment induces a local increase in assayable endorphin IR. But after colchicine, these cells are swollen and the staining-although more intense than in control animals-has lost its granular appearance and produces a more diffuse appearance (Fig. 11). T h e diffuse aspect of the endorphin staining can be understood on the basis of this colchicine side effect. In addition, recent studies in our laboratory (McGinty et al., 1981) have explored another side effect of colchicine treatment: Its selective neurotoxicity toward granule cells of the hippocampus (Goldschmidt and Steward, 1980) markedly reduces the number of mossy fibers exhibiting enkephalin IR. In a few instances ICC observations do not correlate with the RIA measurements in colchicine-treated rats (Fig. 1 1). Although the causes f& these discrepancies have not been elucidated, some of the problems described in the preceding section could partially explain them. A clear example of lack of correlation between changes measured by these two immunodetection methods occurs in the rat striaturn. Colchicine treatment induces a marked intensification of enkephalin IR in the globus pallidus of the rat, however there is no measurable change in the RIA levels (Fig. 11). As we have discussed elsewhere (Bayon et al., 1980a), a localized redistribution of enkephalin within the globus pallidus could partly account for this discrepancy. This possibility is supported by the finding of enkephalin-containing perikarya in this region, near the interface with the caudate nucleus. However, the existence of a strio-pallidal enkephalin pathway, postulated by Cuello and Paxinos (1978), would predict a decrease of enkephalin content in the pallidum after colchicine

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

87

treatment along with an increase in the striatum content; this latter increase is observed by ICC in our colchicine-treated rats, but no change is measured by RIA. T h e data now available indicate the possibility of both local (Schwartz et al., 1978; Costa et al., 1978; Bayon et al., 1979b) and common (Cuello and Paxinos, 1978) enkephalin systems between the caudate and globus pallidus. A heuristic explanation for the lack of correlation between ICC and RIA data after colchicine treatment in both the globus pallidus and the caudate nucleus would rest on the assumption that localized, more than distant, redistributions of enkephalin would enhance cytochemical visualization without having a noticeable effect on regional content of enkephalin IR in the caudate and the globus pallidus. Given these unresolved problems, our preliminary experience indicates that extreme caution is needed when deriving conclusions from changes observed in endorphin or enkephalin immunoreactivity using a single immunodetection method. Only when a reasonable correlation is observed in the results obtained by RIA and ICC can solid inferences be made. IV. Is lmmunodetection Reliable?

Our current concepts of endorphin-enkephalin neurobiology have been built in large part by immunological methodology, both by its power and its limitations. But, as we have discussed, the limitations can lead one to doubt the positive results on a RIA or hold back conviction when finding no detectable staining on the tissue section. the confidence or caution we exercise is in direct relationship to our insight of immunodetection methods. T h e consequences are not trivial. Given the number of ways the presence of a peptide can be obscured in ICC, or artifactually show u p in RIA, perhaps more caution would be in order. Is there no end to this perversity of uncertainty? Fortunately, there is much one can d o to extract the most power from immunodetection systems. T h e use of several different antibodies directed against different antigenic determinants of the same molecule is extremely helpful for both RIA and ICC. I n fact, this approach has recently been adapted for use with molecular genetic techniques (Lerner et al., 1981) to provide a systematic array of antigen determinants. Serious application of the specificity tests outlined above also will prevent falling into methodological pitfalls. I n addition, the knowledge that antibody specificity requirements and the degree of uncertainty are different for RIA and ICC will assure that “rude awakenings” are kept to a minimum.

88

ALEJANDRO BAYON

et al.

Another improvement now being used is the utilization of HPLC and other highly efficient separation techniques coupled with traditional RIA on the resultant fractions. Such “hybridizations” of other methodologies with immunodetection could result in even greater potentiation of the power of immunodetection methods. Considering the enormous contributions the “art” of immunodetection has made to our understanding of biological processes, one can only be optimistic about the future of these techniques.

Acknowledgments

We thank Ms. Nancy Callahan for excellent clerical assistance. A. B. received a postdoctoral fellowship from the Sloan Foundation and acknowledges support from CONAC y ‘l-, Mexico. W.I.S. and F.E.B. acknowledge support of N.I.D.A. Grant 01785. References

Akil, H., Watson, S. J., Berger, P. A., and Barchas, J. D. (1978a). Adz!. Biochum. Psychqhanncol. 18, 125-139. Akil, H., Watson, S. J., Levy, R. M., and Barchas, J. D. (1978b). Zu “Characteristics and Function of Opioids” (J. M. van Ree and L. Terenius, eds.), pp. 123-134. Elsevied North-Holland, Amsterdam. Austen, B. M., Smyth, D. G., and Snell, C. R. (1977). ’Vature (London) 269, 619-621. Barclay, R. K., and Phillips. M. A. (1978). Biorhern. Biophys. Rrs. Commun. 81, 1119-1 123. Bayon, A., Rossier, J., Maws, A., Bloom, F. E., Iversen, L. L., Ling, N., and Guillemin, R. (1978). Proc. ,Vatl. Acad. Sci. U.S.A. 75, 3503-3506. Bayon, A., Shoemaker, W. J., Bloom, F. E., Maws, A., and Guillernin, R. (1979a).Brain Kes. 179,93-101. Bayon, A., Shoemaker, W. J., Milner, R. J., Azad, R., and Bloom, F. E. (1979b). Abstr. Soc. h’mrosci. 5, 1760. Bayon, A., Koda, L., Battenberg, E., and Bloom, F. E. (1980a). Brain Res. 183, 103- 111. Bayon, A., Koda, L., Battenberg, E., Azad, R., and Bloom, F. E. (1980b). Nruro.sci. Lutt. 16, 75-80. Bayon, A.. Lugo, L.. Drucker-Colin, R., Shoemaker, W., Azad, R., and Bloom, F. E. (1981). il‘eurosci. Lutt. 24, 65-70. Bloom, F., Battenberg, E., Rossier, J.. Ling, N., and Guillernin, R. (1978a). Proc. Nut/. Accrd. Sci. L!.S..4.75, 1591-1595. Bloom, F. E., Rossier, J.. Battenberg, E. L. F., Bayon, A,, French, E., Henriksen. S. J., Siggins, G. R., Segal, D., Browne, R., Ling, N., and Guillemin, R. (1978b). Adz!. BiochPm. Psyhopharmacol. 18, 89- 109. Bloom, F. E., Rossier, J., Battenberg, E. L. F., Bayon, A,, French, E., Henriksen, S. J.. Siggins, G. R., Ling, N., and Guillemin, R. (1979). In “Endorphins in Mental Health Research” (E. Usdin, W. E. Bunney, Jr., and N . S. Kline, eds.), pp. 17-29. Macrnillan, &ew YorWLondon.

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

89

Bradbury, A. F., Smyth, D. G., and Snell, C. B. (1975).In “Peptides: Chemistry, Structure and Biology” (R. Walter and J. Meienhofer, eds.), pp. 609-615. Ann Arbor Sci. Publ., Ann Arbor, Michigan. Burgus, R., and Rivier, J. (1976). Pept. 1976, Proc. Eur. Pept. Symp., 14th, 1976 pp. 85-94. Cheung, A. L., Stavinoha, W. B., and Goldstein, A. (1977). L f e Sci. 20, 1149- 1155. Childers, S. R., and Snyder, S. H. (1979).I n “Endorphins in Mental Health Research” (E. Usdin, W. E. Bunney, Jr., and N. S. Kline, eds.), pp. 181-188. Macmillan, New YorW London. Childers, S. R., Simantov, R., and Snyder, S. H. (1977). Eur. J . Pharmacol. 46, 289-293. Childers, S. R., Schwarcz, R., Coyle, J. T., and Snyder, S. H. (1978). Adu. Biochem. Psychopharmacol. 18, 161- 173. Clement-Jones, V., Lowry, P. J., Rees, L. H., and Besser, G. M. (1980).J. Endocrinol. 86, 23 1-243. Costa, E., Fratta, W., Hong, J. S., Moroni, F., and Yang, H.-Y.T. (1978). Adu. Biochem. Psychopharmacol. 18, 217-226. Cuello, A. C., and Paxinos, G. (1978).Nature (London) 271, 178-180. de Souza, M. L. A., Williamson, H. O., Moody, L. O., and Diczfalusy, E. (1970). SteroidAssay Protein Binding, Karolinska Symp. Res. Methods Reprod. Endocrinol., 2nd, 1970 pp. 171187. Desranleau, R., Gilerdeau, C., and Chretien, M. (1972). Endocrinology 91, 1004-1007. Di Augustine, R. P., Lazarus, L. H., Jahnke, G., Khan, M. N., Erisman, M. D, and Linnoila, R. I. (1980). Lge Sci. 27,2663-2668. Ekins, R., and Newman, B. (1970). Steroid Assay Protein Binding, Karolinska Symp. Res. Methods Reprod. E n d o c r i d . , 2nd, 1970 pp. 1 1 -36. Elde, R., Hokfelt, T., Johannson, O., and Terenius, L. (1976). Neuroscience 5, 349-355. Ghazarossian, V., Dent, R. R., Ross, M., Cox, B. M., and Goldstein, A. (1978). I n “Characteristics and Function of Opioids” (J. M. van Ree and L. Terenius, eds.), pp. 273-274. Elsevier/North-Holland, Amsterdam. Goldschmidt, R., and Steward, 0. (1980). Proc. Natl. Acad. Sci. U.S.A. 77(5), 3047-3051. Graf, L., Kenessey, A., Bajusz, S., Patthy, A., Ronai, A. Z., and Berzetei, I. (1979). I n “Endorphins in Mental Health Research” (E. Usdin, W. E. Bunney, Jr., and N. S. Kline, eds.), pp. 189-202. Macmillan, New YorWLondon. Gramsch, C., Kleber, G., Hollt, V., Pasi, A., Mehraein, P., and Hertz, A. (1980).Brain Res. 192, 109-119. Gros, C., Pradelles, P., Rougeot, C., Bepolin, O., Dray, F., Fournie-Zaluski, M. C., Roques, B. P., Pollard, H., Llorens-Cortes, C., and Schwartz, J. C. (1978a).J. Neurochem. 31, 29-39. Gros, C., Pradelles, P., Humbert, J., Dray, F., Le Gal La Sale, G., and Ben-Ari, Y. (1978b). Neurosci. Lett. 10, 193- 196. Guillemin, R., Ling, N., and Burgus, R. (1976). C . R. Hebd. Seances Acad. Sci., Ser. D 282, 783-785. Guillemin, R., Ling, N., and Vargo, T. M. (1977).Biochem. Biophys. Res. Commun. 77, 361366. Hambrook, J. M., Morgan, B. A., Rance, M. J., and Smith, C. F. C. (1976).Nature (London) 262, 782-783. Henderson, G., Hughes, J., and Kosterlitz, H. W. (1978). Nature (London) 271, 677-679. Hindelang-Gertner, C., Stoeckel, M. E., Porte, A., and Stutinsky, F. (1976). Cell Tkme Res. 170, 17-41. Hokfelt, T., Elde, R., Johannson, O., Terenius, L., and Stein, L. (1977). Neurosci. Lett. 5, 25-3 1 .

90

ALEJANDRO BAYON

et al.

Houck, J. C., Kimball, C., Chang, C., Pedigo, N. W., and Yamamura, H. I. (1980).Science 207, 78-80. Hughes, J., Smith, -1. W.,Kosterlitz, H. W., Fothergill, L. A., Morgan, B. A., and Morris, H. R. (1975). S n t r t r ~( L u d m ) 258, 577-579. Hughes, J . , Kosterlitz, H. W., and Smith, T. W. (1977). BI-.J . Phnrmnrol. 61, 639-647. Inoue, S . , and Sato, H. (1967).J. Gun. P h y . ~ i ~50, / . 259-288. Iversen, L. I,., Iversen, S. D., Bloom, F. E., Vargo, T. M., and Guillemin, R. (1978).N n t w e (London) 271, 679-681. Julliard, J. H., Shibasaki, T., Ling, N., and Guillemin, R. (1980). Srieim 208, 183-185. Kimura, S., Lecois, R. V., Stern, A. S., Rossier, J., Stein, S., and Udenfriend, S. (1980).Pror. S a / l . .4cad. Sci. C‘.S...I. 77, 1681-1685. Kobayashi, R. M.. Palkovits. M., Miller, R. J., Chang, K.-J., and Cuatrecasas, P. (1978).Ljfi Sri. 22, 527-530. Kohler, G., Hengartner, H., and Shulman, M. T. (1978). E w . J . Inrmunol. 8, 82-88. Krieger. D. T., Liotta, A., and Li, C. H. (1977a). Lift Sri. 21, 1771-1778. Krieger, D. T., Liotta, A , , and Toshihiro, S. (1979). 112 “Endorphins in Mental Health Research” (E. Usdin, W. E. Bunney, Jr., and N. S. Kline, eds.), pp. 561-568. MacmilIan, New YorWLondon. Larsson, L.-I.. Childers, S., and Snyder, S. (1979).Soturr (Loitdon) 292, 407-410. Lerner, R. A.. Green, K . , Alexander, H., Liu, F.-T., Sutcliffe, J. G., and Shinnick, T. M. (1981).Pruc. h h t l . .4ccid. Sri. I.‘.S..4. 78, 3403-3407. Li, C. H., Barnafi, L., Chretien, M.. and Chung, D. (1965). A’uturr (Loridon) 206, 10931094. Li, C. H . , Rao, A. J.. Donnen, B. A., and Yamashiro, D. (1977). Biochem. Biophjs. Rcy. Conriniiii. 75, 576-580. Liotta, A. S., Suda, T., and Krieger, D. T. (1978).Pi-or. ‘Vntl. Aced. Sci. C’.S.A. 75, 29502954. Lissitskp,J. C., Morin, O., Dupont, A., Labrie, F., Seidah, N. G., Chretien, M., Lis, M., and Coy, D. H . (1978). L f i Sri. 22, 1715-1722. McGinty, J., Gozes, I., and Bloom, F. E. (1981). Sor. ,Veuro.sri. 7, 915. Mains, R. E., and Eipper. B. ,4. (1978).J. Biol. C I t m . 253, 651-655. Mains, R. E., Eipper, B., and Ling, N. (1977). Pror. Soti. .4cnd. Sci. C . S . A . 74, 3014-3018. Slargoulies. M. (1969). “Protein and Polypeptide Hormones,” Int. Congr. Ser. 161. Excerpta Med., Amsterdam. Marks, N.,Galoyan, A., Grynbaum, A., and Lajtha, A. (1974).J. A’rrtrorheiri. 22, 735-739. Meek, J . L., and Bohan, T. P. (1978). A4dr3.Biorltem. Psychophnnnnrol. 18, 141-147. Meek, J. L.. Yang, H.-Y.T., and Costa, E. (1977). , ~ t r t r o ~ ~ h n m r16, o / /151-154. ~~ Midgley, A . R., J r . , Niswender, G. D., and Rebar, R. W. (1969).I~nmurton.~.scty C;onndotwphiit.s, I“infohitskrr Sytnp. Re\. .\lethods Repi-(~d. Etrdocriiiol., Is/, 1969 pp. 163- 184. Miller, R . J . , Chang, K.-J., and Cuatrecasas, P. (1977). Hiorhein. Biophy. Rrs. C / J ~ A W Z 74, U,~. 1311-1317. Miller, R. J., Chang, K.-J., Cooper, B., and Cuatrecasas, P. (1978). J . B i d . Chein. 253, 531-538. Musacchio,J. M.. Puig, M. M..and Craviso, G. L. (1979).Irt “Endorphins in Mental Health Research” (E. Usdin, W. E. Bunnev, Jr., and N. S. Kline, eds.), pp. 219-232. Macmillan, New YorWLondon. Nugent, C. A , , and Mayes, D. (1970). S / ~ r o i dA u q Pr-oturii Binding, I\lro.oliitska Syntp. R P S . .\lrthor/.s Reprod. Eiidorriiio/.. ?rid, 1970 pp. 257-274. Raamsdonk, W., Pool, C. W.. and Heyting, C. (1977).J. Iinmrtno/. dlrthods 17, 337-348.

IMMUNODETECTION OF ENDORPHINS AND ENKEPHALINS

91

Rao, A. J.. and Li, C. H. (1977). Znt. J . Pept. Protein Res. 10, 167-171. Roberts, J. L., Phillips, M., Rosa, P. A., and Herbert, E. (1978). Biochemist? 17, 36093618. Ross, M., Ghazarossian, V., Cox, B. M., and Goldstein, A. (1978). L;fe Sci. 22, 11231130. Rossier, J., Vargo, T. M., Minick, S., Ling, N., Bloom, F. E., and Guillemin, R. (1977a).Proc. Natl. Acad. Sci. U.S.A. 74, 5162-5165. Rossier, J., Bayon, A., Vargo, T. M., Ling, N., Guillemin, R., and Bloom, F. E. (1977b).Life Sci. 21,. 847-852. Rossier, J., Battenberg, E., Pittman, Q., Bayon, A., Koda, L., Miller, R., Guillemin, R., and Bloom, F. (1979). Nature (London) 277, 653-655. Rubinstein, M., Stein, S., Gerber, L. D., and Udenfriend, S. (1977). Proc. Natl. Acad. Sci. U.S.A. 74, 3052-3055. Sar, M., Stumpf, W. R., Miller, R. J., Chang, K.-J., and Cuatrecasas, P. (1978).J. Comp. Neurol. 182, 17-38. Schmitt, F. 0. (1968).Neurosci. Res. Program Bull. 6, 119-144. Schwartz, J. C., Pollard, H., Llorens, C., Malfroy, B., Gros, C., Pradelles, P., and Dray, F. (1978).Adv. Biochm. Psychvpharmacol. 18, 245-246. Seidah, N. G., Gianoulakis, C., Crine, P., Lis, M., Banjannet, S., Routhier, R., and Chretien, M. (1978). Proc. Natl. Acad. Sci. U.S.A. 75, 3153-3157. Simantov, R., and Snyder, S. H. (1976).I n “Opiates and Endogenous Opiate Peptides” (H. Kosterlitz, ed.), pp. 41 -48. ElseviedNorth-Holland, Amsterdam. Simantov, R., Kuhar, M. J., Uhl, G. R., and Snyder, S. H. (1977a). Proc. Natl. Acad. Sci. U.S.A. 74, 2167-2171. Simantov, R., Childers, S. R., and Snyder, S. H . (1977b).Brain Res. 135, 358-367. Smyth, D. G., and Zakarian, S. (1979). In “Endorphins in Mental Health Research” (E. Usdin, W. E. Bunney, Jr,, and N. S. Kline, eds.), pp. 84-92. Macmillan, New YorW London. Stern, A. S., Lewis, R. V., Kimura, S., Rossier, J., Gerber, L. D., Brink, L., Stein, S., and Udenfriend, S. (1979). Proc. Natl. Acad. Sci. U.S.A. 76, 6680-6683. Sternberger, L. A. (1973).Zn “Electron Microscopy of Enzymes” (M. A. Hayat, ed.), Vol. 1, pp. 150- 191. Van Nostrand-Reinhold, Princeton, New Jersey. Stumph, W. E., Elgin, S. C. R., and Hood, L. (1974).J. Immunol. 113, 1752-1756. Swaab, D. F., and Pool, C. W. (1975).J. Endocrid. 66, 263-271. Swaab, D. F., Pool, C. W., and Van Leeuwen, F. W. (1977). J. Histochern. Cytocheni. 25, 388-391. Takahashi, M., Kaneto, H., Ueno, E., Watanabe, J., Koida, M., Ogawa, H., and Yajima, H. (1979).Jpn.J . Pharmacol. 29, 203-209. Udenfriend, S., Rubinstein, M., and Stein, S. (1979). In “Endorphins in Mental Health Research” (E. Usdin, W. E. Bunney, Jr., and N. S. Kline, eds.), pp. 119-130. MacmilIan, New YorWLondon. Uhl, G. R., Kuhar, M. J., and Snyder, S. H. (1978).Brain Res. 149, 223-228. Watson, S. J., Barchas, J. D., and Li, C. H. (1977). Proc. Natl. Acad. Sci. U.S.A. 74, 51555158. Watson, S. J., Akil, H., Richard, C. W., and Barchas, J. D. (1978). Nature (London) 275, 226-228. Watson, S. J., Akil, H., and Barchas, J. D. (1979). In “Endorphins in Mental Health Research’’ (E. Usdin, W. E. Bunney, Jr., and N. S. Kline, eds.), pp. 30-44. Macmillan, New Yor WLondon.

92

ALEJANDRO BAYON

et al.

Yang, H.-Y. T., Hong, J . S., and Costa, E. (1977). Meurtghannacology 16, 303-307. Yang, H.-Y. T., Fratta, W., Hong, J. S., Digiulio, A. M., and Costa. E. (1978). NeurnpharmncoloF 17, 433-438. Yang, H.-Y. T., Hong, J. S., Fratra, W., and Costa, E. (1979). I n “Endorphins in Mental Health Research’ (E. Usdin, W. E. Bunney, Jr.. and N. S. Kline, eds.), pp. 235-241. Mamillan, New York/London. Yoshirni, H., Matsukura, S., Sueoka, S., Fukase, M., Yokota, M., Hirata, Y., and Imura, H. (1978). Lifi Sci. 22, 2189-2195.

ON THE SACRED DISEASE: THE NEUROCHEMISTRY OF EPILEPSY1 0. Carter Snead Ill The University of Alabama in Birmingham School of Medicine Birmingham, Alabama

I. Introduction . . . . . . . . . . . . . . . ... . . . . .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Epilepsy: The Diversity of the Problem . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Neurophysiology of Seizures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Models ..................................... ............ A. Generalized Convulsive Seizure Models . . . . B. Generalized Absence Models . . . .. . . ... .. . . C. Generalized Myoclonic Models . . .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Partial-Seizure Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Neurotransmitters and Other Neuroactive Substances in Seizures . . . . . . . . . A. Catecholamines ................................................... B. Serotonin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . . .. C. Acetylcholine .. . . . .. D. y-Aminobutyric Acid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Other Inhibitory Amino Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. Excitatory Amino Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . G. Cyclic Nucleotides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . .. . . . H. Peptides .. ..... ... .. ... ....... . . .... ..... ... . . ... ... ... ... .. . . .. . VI. Developmental Considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

.

.

. .

94 94 95 99 101 102 104 104 106 107 116 122 128 140 141 142 143 150 152 152

It seems to me that the disease is no more divine than any other. It has a natural cause just as other diseases have. Hippocrates, 400 BC (OLeary and Goldring, 1976) When the transmitter substances in the brain are known . . . and when the chemical environment of the nerve cell . . . are better understood, the neurologist may feel less bewildered by the problem of epilepsy than he is today. Sir Charles Symonds (1959) Since the sequence of potentials . . . are, on the basis of existing evidence, most likely to represent successive waves of excitatory and inhibitory synaptic actions, the question of the identity of the synaptic pathways becomes paramount. Ayala ~t a/. (1973) Supported in part by NINCDS grant K07NS00484-02. 93 INTERNATIONAL REVIEW OF NEUROBIOLOGY, VOL. 24

Copyright 8 1983 by Academic Press. Inc. All rights of reproduction in any form reserved. ISBN 0-12-366824-7

94

0. CARTER SNEAD I11

1. Introduction

Scientists have been in search of Hippocrates’ “natural cause” of epilepsy, the sacred disease (Temkin, 197 l ) , for centuries. Experimental work in the neurophysiology of epilepsy was initiated late in the nineteenth and early in the twentieth century (Fitzsch and Hitzig, 1870; Cybulski, 1914). T h e suggestion that inhibitory mechanisms play a major role in the pathogenesis of seizures was put forth by Adrian in 1936 (Adrian, 1936). In 1954, Florey provided the first evidence for a neurochemical mechanism for this inhibition with the demonstration of an “inhibitory (I) factor” in brain extract, which proved to be inhibitory to crayfish stretch receptors. Florey’s I factor was subsequently identified as y-aminobutyric acid (GABA) (Bazemore r i nl., 1957), a compound now generally acknowledged to be a major inhibitory neurotransmitter in brain. Since the classic paper of Symonds ( 1 959) on excitation and inhibition in epilepsy, the various research strategies in this group of diseases have focused more and more on basic excitatory and inhibitory mechanisms in seizures on a cellular as well as a whole-brain level. In order to understand the complex neurochemistry of seizure states, it is first necessary to define precisely the meaning of the terms seizure and epilepsy and review the neurophysiology of these aberrant neuronal systems. I shall then discuss various animal models of seizures and following that, shall review the neurochemistry of epilepsy in the context of neurotransmitters and neuroactive agents in animal models, anticonvulsant drug interaction, and the clinical seizure state. T h e membrane biology and metabolic aspect of seizures will not be covered since comprehensive reviews of these aspects of epilepsy have been recently published (Delgado-Escueta and Horan, 1980; A. L. Miller, 1981). What follows then, will hopefully supplement rather than reiterate previous comprehensive reviews of the subject (Lovell, 1971; Meldrum, 1975; Maynert, 1969; Maynert et nl., 1975; Emson, 1975; Singh and Huot, 1973).

II. Epilepsy: The Diversity of the Problem

Epilepsy is not a single disease entity. Rather, this term refers to a group of disorders which have in common the clinical phenomenon of the seizure and the cellular phenomenon of a paroxymal discharge of neurons. A seizure, on the other hand, is an episodic event with a distinct beginning and end. T h e seizure is involuntary and often associated with

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

95

TABLE I CLASSIFICATION OF SEIZURES International

Clinical

Generalized seizures Convulsive Absence M yoclonic

Grand ma1 Petit ma1 Myoclonic Infantile spasms

Partial seizures" With elementary symptomatology Partial motor Partial sensory With complex symptomatology Partial complex

Focal motor Focal sensory Psychomotor or temporal lobe

~

(I

With or without secondary generalization.

postseizure impairment: the postictal state. The international system of classification classifies the epilepsies as either generalized or partial depending on whether the electrical seizure activity involves the whole brain at its outset or is focal and whether or not consciousness is impaired. (Gasteaut, 1970; Dreifuss et al., 1981). Alternatively, seizures may be classified in terms of their clinical manifestations. These two systems are listed in Table I. A perusal of this table will evidence the clinical heterogeneity of seizure disorders. Similarly, there are a host of underlying neuropathologic processes that may give rise to the human condition known as epilepsy in one of its many forms. The clinical experience would thus seem to indicate that any search for a single neurochemical mechanism for seizures will be fruitless indeed. It is much more likely that there are multiple complex neurochemical mechanisms at work in any seizure disorder. One must bear in mind this diversity of clinical seizure types when critically evaluating experimental data on seizures. The precise type of seizure to which the experiments apply has to be specified before one can ascertain the relevance of the data to a particular type of clinical seizure.

111. Neurophysiology of Seizures

Prior to delving into neurochemical mechanisms it behooves one to know just what neurophysiologic events are taking place. T h e common underlying cellular event in any seizure is presumably a paroxysmal

96

0. CARTER SNEAD 111

neuronal discharge. This massive discharge of nerve cells is recorded on the electroencephalogram (EEG) as a spike and slow wave which may be recorded during a clinical seizure (Fig. 1). T h e patient with a seizure disorder may also show interictal spikes between paroxysms (Fig. 2). T h e cellular correlate of the interictal spike and the spike and slow wave has been extensively investigated and reviewed (Ayala et al., 1973; Prince, 1978; Goldensohn and Ward, 1975; Schwartzkroin and Wyler, 1980). In 1964, Matsumoto and Ajmone Marsan described the chronology of events in cells made epileptiform by topical penicillin application. T h e penicillin focus has been utilized as a model of partial seizures since its initial description by Walker and Johnson in 1945. T h e sequence of events described by Matsumoto and Ajmone Marsan (1964) are also common to other models of partial-seizure disorders such as freezing (Goldensohn and Purpura, 1963),strychnine application (Li, 1959), and electrical stimulation, as well as topical application of metal (Goldensohn and Ward, 1975). T h e experiments of Matsumoto and Ajmone Marsan were done with acutely elicited penicillin epileptogenic foci in the cerebral cortex of cats. Penicillin application resulted in paroxysmal discharges as recorded from cortical EEG. T h e intracellular correlate of this synchronous cortical discharge was a large 20-50-mV depolarization that lasted from 50 to 100 msec. This intense intracellular depolarization correlated with the surface cortical paroxysmal discharge in 96% of cells and was named the paroxysmal depolurizing shqt (PDS). T h e PDS would terminate spontaneously when the membrane repolarized and then

1

5

.

1

,

0L ~ I Sec

FIG. 1. Electroencephalographic recording of a seizure in a 10-year-old child demonstrating the Occurrence of spike and slow wave activity during a clinical seizure. The child’s seizure was manifested by tonic posturing. A refers to an ear lead; odd and even numbers refer to left and right, respectively; Ep, prefrontal; C, central; 0, occipital; and T, temporal areas.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

97

5OpV I Sec

FIG. 2. Interictal spike (arrows) as recorded in the EEG of a 4-year-old child with a partial complex seizure disorder. No clinical seizure activity was evident during this recording. See legend to Fig. 1 for key to abbreviations.

hyperpolarized. The hyperpolarization phenomenon during which no PDS could occur was termed the after-hyperpolarization potential (AHP). The epileptic spike in humans correlates with the PDS, while the cellular correlate of the slow wave is the AHP. The AHP is presumably responsible for cessation of the PDS and thus for prevention of progression of this hypersynchronous discharge to overt behavioral seizure activity (Ayala et al., 1970) (Fig. 3). However, as the AHP decreases in duration, the PDS increases in frequency, and a tonic seizure occurs when the AHP disappears. Since the PDS-AHP complex appears to be the cellular substrate of seizure activity, a great deal of effort has gone into elucidating its mechanism. Subsequent to the basic description of the PDS, it has been demonstrated (Prince, 1968; Matsumoto et al., 1969) that this phenomenon can be elicited in cortex or hippocampus by orthodromic but not intracellular stimulation. This has led to the hypothesis (Prince, 1968, 1978) that the PDS is a giant excitatory postsynaptic potential (EPSP). This giant EPSP is thought to be produced by recurrent excitatory circuits that result from increased synaptic drive (Dichter and Spencer, 1969; Ayala et al., 1970).Although nonsynaptic factors such as axonal discharges may play a role in the generation of the PDS (Prince, 1978), recent experiments examining the behavior of the PDS under current and voltage

98

0. CARTER SNEAD 111

FIG.3. A schematic diagram of relations between cortical discharges and both intracellular and extracellular activity in a penicillin epileptic focus in cats. Isolated interictal discharges and tonic-clinic ictal activity is shown. [Reproduced from Ayala rt a / . (1970) with permission.]

change conditions as a function of membrane potential provide strong evidence that the PDS is in fact a function of synaptic driving force (Johnston and Brown, 1981) and probably represents a giant synaptic potential. Equally as important as the PDS is the mechanism of the AHP, since this event is intimately involved in the termination of the PDS and prevention of recurrent PDS with progression to tonic seizure activity (Matsumoto and Ajmone Marsan, 1964). T h e AHP has not been as well investigated as the PDS, but there are some data to support the thesis that this is a persistent inhibitory postsynaptic potential (IPSP) (Prince, 1979). However, Alger and Nicoll (1980) have presented evidence that the AHP may be an intrinsic inhibitory potassium potential mediated by calcium, although this finding has been challenged (Schwartzkroin and Stafstrom, 1980). In spite of the fact that the PDS-AHP complex is strikingly similar from seizure model to seizure model and has been demonstrated to occur in human neurons involved in epileptogenesis (Prince and Wong, 1981), there is still a quantum leap from a single cell which shows paroxysmal depolarization shifts to the clinical seizure in humans. T h e question thus arises of how the cellular events described above progress to cause a behavioral seizure. Obviously, there are a number of critical factors involved which include the anatomy, physiology, and neurochemical make-up of cells within the brain which are capable of producing epileptiform discharges, changes in intrinsic properties of the

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

99

neuronal and synaptic membrane, and the ionic microenvironment. In the remainder of this article, I shall attempt to review these factors in terms of the neurochemistry involved, particularly in regard to excitation and inhibition, and relate the neurochemistry to various experimental models of epilepsy currently in use. I shall also touch upon some developmental aspects of experimental seizure research since epilepsy is seen so frequently in the developing nervous system.

IV. Models

One rational research approach to the clinical heterogeneity of epilepsy has been the development of a large number of experimental models that provide insight into the basic mechanisms of seizure disorders, serve for screening purposes to assess anticonvulsant efficacy of drugs, and elucidate basic neurobiological mechanisms which at times are unrelated to,seizures. The ideal experimental model should meet the following criteria (Wada, 1977). 1. Precise experimental control should be exerted over the anatomy and size of the epileptogenic lesion. 2. Pathologic changes should be minimal. 3. Seizures should be controllable in terms of precipitation and chronologies. 4. A spontaneous, chronic seizure should develop over time. These criteria are best applied to animal models (Table 11),but there are in vitro models that have also proved useful. The simplest of these is the central nervous system (CNS) neuron in culture. This experimental system consists of dissociated fetal mouse spinal cord neurons maintained in culture for several months. The individual cultured neurons are visible under phase contrast optics and are suitable for long-term intracellular recording associated with extracellular iontophoresis of various neurotransmitter agonist and antagonist drugs. These spinal cord neurons are said to possess receptors for several neurotransmitters and to have electrophysiological and pharmacological properties similar to neurons in other central nervous system preparations (Gruol et al., 1980; Barker and Ransom, 1978a,b; Ransom et al., 1977a,b). The next level of complexity in model systems is the mammalian brain slice preparation, since this is obviously a multineuronal and therefore multisynaptic preparation. Both cortical (Halpern, 1972; Courtney

100

0. CARTER SNEAD I11

TABLE I1 EXPERIMENTAL MODELSOF SEIZURES Generalized convulsive

Generalized absence

Generalized myoclonic

Partial seizures

Electroconvulsive shock Chemical convulsants Pentylenetetrazol, picrotoxin, bicuculline Genetic Audiogenic seizure-mouse Photomyoclonus-baboon (Pnpzo pupio) Mongolian gerbil, epileptic fowl, canine model Miscellaneous-0, toxicity, drug withdrawal Electrical stimulation-thalamus stellate or spinal trigeminal ganglion Chemical convulsants Penicillin, y-hydroxybutyric acid, leucine-enkephalin, pentylenetetrazol, convulsant metals (bilateral or subcortical) Genetic Papio papio; mutant mouse tottering Miscellaneous-fiash evoked after discharge Genetic Papio pap’o Chemical convulsants y-Hydroxybutyric acid a-Choralose; subcortical picrotoxin; kynurenines; muscimol 1,1,l-Trichloro-2,2-bis(p-chlorophenyl)ethane Electrical-kindling Chemical convulsants Alumina gel, penicillin, cobalt, iron, kainic acid Genetic Beagle dog, mutant mouse totterer Miscellaneous-freezing

and Prince, 1977) and hippocampal (Dingledine and Gjerstad, 1979; Schwartzkroin and Prince, 1977. Yamamoto, 1972) slices have been used. Recently, in vitro brain slice techniques have been applied to the study of epileptogenesis in neocortical slices prepared from biopsy specimens of human epileptic brain (Prince and Wong, 1981). Another hippocampal isolation technique that may prove extremely useful is that of transplanting the fetal rat hippocampus to the anterior chamber of the eye. These transplants have been shown to grow and

SACRED

DISEASE:

NEUROCHEMISTRY OF EPILEPSY

101

form the intrinsic excitatory and inhibitory circuitry typical of hippocampus (Olson et al., 1977). These transplants are capable of responding to a number of epileptogenic stimuli such as electrical stimulation, penicillin superfusion, and cobalt iontophoresis and show the PDS-AHP activity common to seizing neurons (Freedman et al., 1979; Hoffer et al., 1977b). The whole-animal model of seizures is the most complex in terms of relating neurochemical and neurophysiologic data to causation and treatment, yet these experimental models have yielded a wealth of information on the neurochemistry of epilepsy. These models are best classified in the same terms as human seizures, that is, generalized and pal'tial (Table 11). Once the behavioral and anticonvulsant pharmacologic profile is thus defined, many of these models can be further subdivided into electrical stimulation, chemical convulsants, genetic models, and the inevitable miscellaneous classification.

MODELS A. GENERALIZED CONVULSIVE SEIZURE

1. Electroconvulsive Shock Electroconvulsive shock (ECS) is one of the oldest seizure models arising from the work of Fitzsch and Hitzig (1870) and Albertoni (1882). The techniques vary with regard to type of stimulation, electrode placement and stimulus frequency and duration (Swinyard, 1949, 1972; Swinyard et al., 1952), but basically the brain receives a maximal or supramaximal electrical stimulus which results in a generalized convulsive seizure.

2. Chemical Convulsants Since the description of camphor as a chemical convulsant (Wiedemann, 1877), a variety of chemical convulsants have come into use (Stone, 1972). Perhaps the most commonly used systemic chemical convulsant today is pentylenetetrazol (PTZ), which has been in use since 1926 (Hildebrandt, 1926). Other chemical convulsants capable of producing generalized convulsive seizure activity (Swinyard et al., 1952; Stone, 1970) are picrotoxin (Hahn and Okerdorf, 1962; Chusid and Kopeloff, 1969; Horton et al., 1976), thiosemicarbazide (Tews and Stone, 1965; De Vanzo et al., 1961; Murakami et al., 1976); bicuculline (Meldrum and Horton, 1971; Worms and Lloyd, 1978), DL-allylglycine (Ashton and Wauquier, 1979; Schneider et al., 1960; McFarland and Wolner, 1965), pyidoxal phosphate (Ebadi and Klangkalya, 1978; Kousoumdjiam and Ebaai, 1981), and benzene (Contreras et al., 1975).

102

0. CARTER SNEAD 111

3. Getietir LLlocirls Nature has provided us with a wide spectrum of convulsive disorders that occur in animals (Consroe and Edmonds, 1979; Jobe and Laird, 1981) most of which are characterized by stimulus-evoked, generalized convulsive seizures. The oldest of these models is the audiogenic seizure which was first studied in rats (Seyfried, 1979). However, since the discovery (Hall, 1947) of a striking genetic difference in susceptibility between the C57 and DBA inbred mice strains to sound-induced seizure activity, the model in this species has been one of the most intensively studied phenotypes in behavioral genetics (Maxson and Cowen, 1976; Sprott and Staats, 1975) and has provided a wealth of data on the neurochemical substrate of paroxysmal disorders. Auditory-evoked seizures also occur in rabbit (Antonitis et nl., 1954; Hohenboken and Nelhaus, 1970), but this model is not as well investigated as the mouse. Other commonly used genetic models of generalized convulsive seizures are photic-induced seizures in the baboon (Killam et nl., 1967; Stark et al., 1970; Killam, 1976; Naquet and Meldrum, 1972), stimulus-evoked seizures in the Mongolian gerbil (Kaplan, 1975; Thiessen et nl., 1968; Kaplan and Miezejeski, 1972; Loskota and Lomax, 1975; Rudeen et nl., 1980) and mouse (Kohsaka et nl., 1978), photic-induced seizures in epileptic fowl (Johnson et nl., 1979, 1981), and spontaneous seizures in the beagle dog (Edmonds rt al., 1979).

4. i\liscellcinroits ,\lodels There are a few miscellaneous models that have been used for drug screening and elucidation of mechanisms of pathogenesis of generalized convulsive seizures. These include barbiturate (Essig, 1967, 1972) and ethanol (Goldstein, 1972; Gibbins et al., 1971; Essig and Laurn, 1968; Walker and Zornetzer, 1974) withdrawal syndromes and hyperbaric oxygen (Lambertsen, 1966; Wood, 1972).

B. GENERALIZED ABSENCE MODELS 1. Electticul Stinzulation A protracted single shock or repetitive shocks delivered to specific thalamic nuclei in monkey produces seizures that bear a pharmacologic, behavioral, and electrical resemblance to petit ma1 seizures (Steriades, 1974; Pohl et nl., 1980). Electrical stimulation of the stellate ganglion (Esplin and Zablocka, 1969) and of the spinal trigeminal nucleus (Fromm and Kohli, 1972) form the basis for models which are pharmacologically similar to petit mal seizures.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

103

2 . Chemical Convulsants Systemic chemical convulsants which produce a petit ma1 seizure state include large doses of intramuscular penicillin in cat (Guberman et al., 1975; Fisher and Prince, 1977; Testa and Gloor, 1974; Prince and Farrell, 1969; Gloor and Testa, 1974; Quesney and Gloor, 1978), inhalation of toluene (Contreras et al., 1975), parenteral administration of y-hydroxybutyrate (Snead et al., 1976; Snead, 1978a,b,c), and intraventricular administration of the opioid peptide leucine enkephalin (Snead and Bearden, 1980a, 1981). Although PTZ does not produce behavioral absence seizures, the clonic seizures induced by this compound in rodents have been used extensively as a pharmacological model of the petit ma1 seizure (Krall et al., 1978; Jenney and Pfeiffer, 1956; Chen et al., 1963; Esplin and Curto, 1957; Myslobodsky, 1976). Generalized absence seizures have also been produced with topical application of chemical convulsants, usually bilaterally or subcortically. Bilateral cortical or cerebellar application of cobalt (Grimm et al., 1969; Cereghino and Dow, 1970; Marcus et al., 1972) or conjugated estrogens (Marcus et al., 1966), or subcortical implantation of aluminum oxide (Guerrero-Figueroa et al., 1963) has been used in this fashion to produce petit ma1 seizures. 3 . Genetic Models

Although the clinical manifestation of the photic-induced myoclonic seizure in the baboon seems to be a generalized convulsive or generalized myoclonic seizure, the pharmacologic characteristics of this event have led to its use as a screen for anti-petit-ma1 drugs (Woodbury, 1972). T h e only genetic model of generalized absence seizures in which the seizures are genuine spontaneous behavioral absence attacks is the mutant mouse tottering (tg, chromosome 8, autosomal recessive) where the absence seizures coexist with focal motor seizures (Noebels and Sidman, 1979). 4. Miscellaneous Models

The flash evoked after discharge (FEAD) is a hypersynchronous burst of slow waves or slow wave and spike complexes produced from the visual cortex of rat (Shearer et al., 1976; Kimura, 1962; King et al., 1980; King and Burnham, 1980) and rabbit (Myslobodsky, 1976) by presentation of a single light flash stimulus. Both convulsants and anticonvulsants with a predominantly thalamic level of action are effective in modulating the FEAD in rat. Anti-petit-ma1 anticonvulsants suppress this response, whereas PTZ enhances it.

104

0. CARTER SNEAD 111

C. GENERALIZED MYOCLONICMODELS T h e generalized muscle jerks associated with myoclonic seizures may or may not be associated with rhythmic cortical EEG discharges (Grinker et al., 1938; Halliday, 1967). Although myoclonic jerks may occur in the course of experimental absence seizures, for example, the y-hydroxybutyrate model (Snead et al., 1976; Snead, 1978a) or the leucine-enkephalin-treated rat (Snead and Bearden, 1980a). Pure myoclonic seizure models are few. Adrian and Moruzzi (1939) demonstrated that chloralose produced a stimulus-sensitive myoclonic seizure state. Any stimulus, whether auditory, photic, somatic, or electrical, produces myoclonic jerks in these animals (Alvord and Fuentes, 1954). Picrotoxin administered into the caudate nucleus produces contralateral focal myoclonus; however, this phenomenon was observed only if the sensorimotor cortex was damaged indicating that the latter structure was necessary for manifestation of myoclonus in this model (Tarsy P t nl., 1978). Hwang and Van Woert (1978) have recently demonstrated (p,p '-DDT) produces that 1,1,1-trichloro-2,2-bis(p-chlorophenyl)ethane spontaneous and stimulus-sensitive myoclonus in mice and rats. Experimental myoclonus has also been produced in guinea pigs and rats by intraperitoneal 5-hydroxytryptophan which is exacerbated by pretreatment with 5,7-dihydroxytryptamine (Klawans et al., 1973; Stewart et a/., 1976). Kynurenines, neuroactive metabolites of tryptophan, also produce myoclonic seizures in hind legs of mice (Lapin, 1981).

MODELS D. PAKTIAL-SEIZURE The vast majority of partial-seizure models are dependent on focal application of some noxious stimuli, for example, electrical, chemical, or traumatic. The nature of the seizure, that is, whether it is partial with elemental or complex symptomatology, depends on which part of the brain is the recipient of the noxious stimuli. Hence, partial motor seizures would involve the sensorimotor cortex and partial complex seizures would involve the hippocampus or some other part of the limbic system. 1. Electrical Stimulation: Kindling Goddard (1967) and Goddard et al. (1969) have shown that brief application of a nonpolarizing electrical stimulus to discrete regions of mammalian brain produces a permanent alteration in function such that

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

105

animals respond to each burst with behavioral convulsions. The seizures progress from focal motor or partial complex to generalized convulsive (Racine, 1972). Since the original description of this model over 14 years ago, there have been a number of variations on the theme (Cain, 1979; Frenk and Yitzhaky 1981) including chemical kindling with repeated administration of subconvulsant doses of chemical convulsants (McMillen and Isaac, 1978a; Vosu and Wise, 1975; Wasterlain and Jonec, 1980b). A variety of kindling paradigms have shed much light on underlying neurochemical influences in both partial and generalized convulsive seizures (Wada, 1977; Sato and Nakashima, 1976; Engel et al., 1981). 2. Chemical Convulsants T h e two chemical convulsants most commonly used to produce partial seizures in animals are alumina cream and penicillin. Both of these are applied topically to cortex and both have been used to elucidate the cellular events described above that are common to all seizures. T h e alumina hydroxide method for inducing recurrent partial motor seizures in monkey was first described by Kopeloff et al. (1942). This model is now a standard one for spontaneous partial simple seizures (Lockard et al., 1975; Chusid and Kopeloff, 1969). Application of the alumina cream to hippocampus results in partial complex seizures (Perryman et al., 1980; Soper et al., 1978). The alumina gel is most effective in primates and results in significant disruption of cortical cytoarchitecture (Ward, 1972). The penicillin model of partial seizures, developed in 1945 by Walker and Johnson, is effective in many species and produces little damage (Edmonds and Stark, 1974; Okada, 1971; Stark et al., 1974; Edmonds et al., 1974). Other metals which produce partial-seizure states when applied topically include cobalt (Dow et al., 1972; Dewar et al., 1972; Joseph and Emson, 1976; Matani, 1967a,b; Emson and Joseph, 1975) and iron (Willmore et al., 1978). Another convulsant agent that has proved useful for inducing partial complex seizures in animals when given either intracerebrally or systemically in kainic acid. This substance is a rigid analog of L-glutamic acid (Kizer et al., 1978; Nadler, 1979), which when injected into brain causes degeneration of neurons with cell bodies in the vicinity of the injection but not of axons of passage or of termination from distant neurons (Coyle and Schwartz, 1976; Coyle et al., 1978; Schwartz and Coyle, 1977; McGeer and McGeer, 1976). Injection of kainic acid into the amygdala produces prolonged partial complex seizures with rather severe damage in ipsilateral structures (Zaczek et al., 1978a; Ben-Ari et al., 1978, 1979, 1980; Meniniet al., 1980; Schwartz et al., 1978; Pisa et al., 1980). Systemi-

106

0. CARTER SNEAD 111

cally administered kainic acid also results in partial seizures often with secondary generalization to generalized convulsive seizure activity (Olney et nl., 1974; Stone and Javid, 1980; Liebmanetnl., 1980; Lothman et nl., 1981; Lothman and Collins, 1981). 3. G n i e t i c .\lodels

T h e only genetic models of partial seizures are the beagle dog (Edmonds et nl., 1979) and the mutant totterer mouse (Noebels and Sidman, 1979; Noebels, 1979). In the canine model some animals have “minimal” seizures which are inherited, spontaneous, and bear behavioral resemblance to partial complex seizures. In the mouse model, focal motor seizures occur spontaneously but in association with generalized absence attacks.

4. ~ \ I w c P l l n v I Y o l t J T h e production of partial seizures by freezing has been observed since 1883 (Lewin, 1972). This technique reliably produces discrete, active foci of epileptic discharges which are analogous to those seen with alumina gel or cobalt (Stalmaster and Hanna, 1972; Hori et nl., 1979; Lewin p t nl., 1969).

V. Neurotransmittersand Other Neuroactive Substances in Seizures

There are several lines of evidence that may implicate a particular compound in brain in the precipitation, exacerbation, or alleviation of pathologic seizure states. These include the effect of pharmacologic manipulation of said compound or systems that synthesize, secrete, or degrade it on seizure threshold in various seizure models, the overt EEG and behavioral effect of the substance itself given either systemically or introcerebroventricularly (ICV), or a correlation of any change in endogenous brain levels or biological activity of the compound with the therapeutic effect of anticonvulsant drugs. Clinical evidence for involvement of a particular neuroactive agent in the causation of epilepsy may include data from postmortem brain or brain obtained at surgery such as that done for intractable seizures (Prince and Wong, 1981). Other clinical data may come from human cerebrospinal fluid studies. These usually consist of measurement of the neurotransmitter or its metabolites. Finally, clinical involvement of a specific neurotransmitter in epilepsy may be inferred from studies of the anticonvulsant efficacy of drugs known to affect that neurotransmitter system in viuo (Snead,

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

107

1981a). In the following section I shall review these various aspects of the neurochemistry of seizures for a number of compounds in brain, some of which clearly meet the rigorous criteria R. Cooper et al., 1978; Werman, 1966) required for a compound to be considered a neurotransmitter and some neurotransmitter candidates that have not fulfilled those criteria, for example, peptides.

u.

A. CATECHOLAMINES 1. Anatomy

There is a marked difference in the organization of central catecholamine neuronal systems between dopaminergic (DA) and noradrenergic (NA) systems. The dopamine system has the most neurons, a discrete topography, and a restricted terminal distribution area, whereas the converse is true for norepinephrine (Moore and Bloom, 1978, 1979). T h e anatomy of norepinephrine- and dopamine-containing neurons has been extensively reviewed (Moore and Bloom, 1978, 1979; Lindvall and Bjorklund, 1978; Lindvall, 1979) and will be given cursory treatment here. Dahlstrom and Fuxe ( 1 964) first described the distribution of catecholamine-containing neurons in the lower brainstem and classified them into 12 groups designated A1 - 12. Since then, additional groups have been described bringing the total to 15 (Fuxe et al., 1969; Bjorklund and Nobin, 1973; Hokfelt et al., 1975; HalaSz et al., 1977). The central DA projection system is outlined in Table 111. Excluding the retinal dopamine system, there are six projection systems. These include the mesostriatal or mesencephalic system, with the primary cells of origin in the pars cornpacta and the pars lateralis of the substantia nigra, and the ventral tegmental area projecting to the caudate-putamen complex; the mesocortical projecting from the substantia nigra to septum, olfactory tubercle, amygdala, and entorhinal cortex; the periventricular; incertohypothalamic; tuberohypophysial; and periglomerular system. T h e major norepinephrine (NA)-containing cell group is the locus coeruleus, which is located in the brainstem reticular formation. NA fibers are projected via the dorsal tegmental and median forebrain bundle to neocortex, hippocampus, and thalamus. The neocortical NA neurons terminate in the molecular layer with their highest density in the cingulate cortex. The DA cortical innervation differs from NA in that the dopamine neurons are restricted to frontal lobe and entorhinal areas. T h e amygdala receives both DA and NA afferents, whereas the

108

0. CARTER SNEAD I11

TABLE 111 CESTRAL CATECHOLAAIINERGIC PROJECTIONS" Catecholamine Dopamine

System Nigrostriatal

Substantia nigra

Mesocortical

Substantia nigra Ventral tegmental area Periaqueductal gray and caudal thalamus Zona incerta and periventricular hypothalamus Arcuate and periventricular hypothalamic nuclei Olfactory bulb Locus coeruleus

Periventricular

Incertohypothalamic Tuberoh ypoph ysial

Norepinephrine

Cells of origin

Periglomerular Locus coeruleus

Lateral tegmental

Dorsal motor vagus, nucleus tractus solitarius, lateral tegmentum

Projection Caudate putamen, globus pallidus, amygdala, nucleus accumbens Septum, amygdala, piriform cortex, supra- and ventral entorhinal cortex Medial thalamus, hypothalamus, periaqueductal gray Zona incerta, septum, hypothalamus Median eminence, pituitary

Olfactory glomeruli Raphe, cerebellum, hippocampus, thalamus (geniculate bodies), isocortex, hypothalamus, brainstem, spinal cord Spinal cord, brainstem, hypothalamus, basal telencephalon

" Modified from Moore and Bloom (1979) and Lindvall (1979). catecholaminergic innervation of hippocampus is strictly NA, with the highest NA fiber density in the region of the dentate gyrus. T h e specific thalamic nuclei are innervated by the locus coeruleus (LC), while the medial and midline t-halamic nuclei get both N A and DA innervation. Other norepinephrine-containingcells lying outside the locus coeruleus project to the fornix, ventral part of stria terminalis, and septa1 nuclei. Descending catecholaminergic projections form the bulbospinal system, which projects from the medulla to ventral and dorsal horns and the sympathetic system. There is also a descending N A pathway to the dorsal motor nucleus of the vagus, nucleus of the solitary tract and cord. Although the bulbospinal tract is primarily NA, there is evidence for DA fibers as well (Reid et al., 1975; Magnusson, 1973; Commissiong and Neff, 1979).

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

109

2. Physiology As with many other neuroactive agents discussed below, it is difficult with the catecholamines to resolve the effects of NA and DA into simple inhibitory and excitatory actions. This is because of multiple subtypes of DA (Langer, 1977; Starke et al., 1977; Skirboll et al., 1979; Costal1 and Naylor, 1981; Cools and Van Rossum, 1980; Groves et al., 1975; Kebabian and Calne, 1979; DeBelleroche and Bradford, 1978) and NA receptors (Hoffman and Lefkowicz, 1980: Minneman and Molinoff, 1980; Palacios and Kuhar, 1980; Schultzetal., 1981; Minnemanetal., 198 1) each with varying sensitivity, and in some cases different electrophysiologic responses to agonist action. An additional complicating factor is that stimulation of a system with one effect, say inhibition, may result in an opposite effect, excitation, due to “disinhibition” of an inhibitory interneuron (E. Roberts, 1974, 1976). There is evidence that dopamine has both excitatory (Kitai et al., 1976; York, 1976, 1979; Zarzecki et al., 1976; Conner, 1970) and inhibitory (Stone, 1976; Spencer and Havilicek, 1974; Bloom et al., 1965; Bunney and Aghajanian, 1973; Ben-Ari and Kelly, 1976) actions. Most of the studies that have examined the electrophysiology of DA systems have utilized the nigrostriatal projection (Moore and Bloom, 1978). Electrical stimulation of the substantia nigra leads to excitation of a large number of neurons in the caudate (York, 1979), but there is another population of caudate neurons which receives inhibitory input from the nigra. Similarly, there are cells in the caudate-putamen both excited and inhibited by iontophoresis of dopamine. Neurons of the basolateral amygdala appear to be inhibited by dopamine (Ben-Ari and Kelly, 19’76), as do those of frontal and cingulate cortex (Bunney and Aghajanian, 1977). The physiologic action of norepinephrine is clearer than that of dopamine in that it appears to be uniformly inhibitory throughout brain (Moore and Bloom, 1979; Bloom, 1979). Electrical stimulation of the locus coeruleus produces inhibitory responses in the cerebellum (Hoffer et al., 1973; Siggins et al., 1971, 1976), hippocampus (Segal and Bloom, 1974), septum (Segal, 1976), and thalamus (Nakai and Takaori, 1974). Thus the general physiologic properties of NA pathways based on studies of LC projections to cerebellum, hippocampus, thalamus, and cortex indicate that the circuits are inhibitory, slow in onset, prolonged in action and mediated through 0 receptors (Bloom, 1979). 3. Role in Seizure Models

a. Generalized Convulsive. Since the observation by Chen et al. (1954) that reserpine lowered the dose of PTZ needed to produce tonic exten-

110

0. CARTER SNEAD 111

sion of hindlimbs in mice, presumably by depleting the brain of biogenic amines, there has been a great deal of research effort toward elucidating a role for the cateholamines in the pathogenesis of the epilepsies (Maynert rt nl., 1975; Moskowitz and Wurtman, 1975). Since that initial report, depletion of catecholamines by a variety of methods has been shown to increase susceptibility to generalized convulsive seizures induced by PTZ and ECS (Azzaro et al., 1972; DeSchaepdryver et al., 1962; Jones and Roberts, 1968; Pfeifer and Galambos, 1967; Swinyard et al., 1964; Kilian and Frey, 1973; London and Buterbaugh, 1978; Koslow and Roth, 1971; Jobe et al., 1974). Inhibitors of monoamine oxidase (MAO) or administration of the catecholamine precursor L-DOPA have been reported to raise the electroconvulsive threshold (Kilian and Frey, 1973; Chen et cil., 1968; Prockop et al., 1959) as does apomorphine, a dopamine agonist (McKenzie and Soroko, 1972), although this compound exacerbates PTZ-induced seizures (Soroko and McKenzie, 1970). Both norepinephrine and dopamine when given ICV decrease the threshold for ECS; however, if the resultant hypothermia is corrected, this reduction in seizure threshold does not occur (Browning and Maynert, 1978b). Conversely, ICV administration of dopamine or norepinephrine to animals pretreated with catecholamine-depleting agents is anticonvulsant (Stull et al., 1973, 1977). In spite of the anticonvulsant effect of ICV dopamine and the suggestion of some that dopamine may be important (DeSchaedryver et al., 1962; Azarro et al., 1972), the preponderance of data suggests that norepinephrine is the more important of the two catecholamines in determining seizure threshold to PTZ- or electroshock-induced seizures (Wenger rt al., 1973; Jobe rt al., 1974; Kilian and Frey, 1973; Doteuchi and Costa, 1973). Although Spencer and Turner ( 1969) showed that amphetamines potentiated PTZ seizures and suggested that this was a dopaminergic effect, more extensive studies by Riffee and Gerald (1976) show that the effect of amphetamines on seizures is a complex issue involving a- and p-noradrenergic systems. In the genetic models of generalized convulsive seizures, there is an increased concentration of NA in the hemispheres of epileptic fowl with decreased brain DA; however the seizure susceptibility of this model is not changed by either increasing brain DA or by a- or P-adrenergic blockade (Johnson rt al., 1979, 1981). There is no significant difference in either NA or DA between the epileptic beagle dog brain and the seizure-resistant beagle (Edmonds rt al., 1979). In the Mongolian gerbil, whole-brain concentration and striatal uptake of DA is no different from control (Cox and Lomax, 1976). In the audiogenic-seizure-pronemouse there is a developmental difference in catecholamine storage and synthe-

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

111

sis between seizure-prone and seizure-resistant animals. T h e seizureprone mice show a lower accumulation of NA at the age of maximal seizure (Kellog, 1976). The gradual resistance to seizures that occurs with age in these animals is associated with an increase in turnover of DA and NA (Shaywitz et al., 1978). However, there is no difference in the high-affiity transport mechanism of dopamine in the seizure-prone vs seizure-resistant animal (Bondy et al., 1979). The dopamine agonist apomorphine has a protective effect against seizures in the Mongolian gerbil (Cox and Lomax, 1976), DBN2 mice (Anlezark and Meldrum, 1975), and photosensitive baboon (Meldrum et al., 1975; Anlezark et al., 1978), but amphetamine and propanolol are not anticonvulsant in the baboon (Brailowsky and Naquet, 1976; Anlezark et al., 1978). Introcerebroventricular NA but not DA is anticonvulsant in the baboon, and single doses of reserpine also protect against seizures in this model (Altshuler et al., 1976). Chronic depletion of catecholamines has the same effect in the genetic models as it does in the electroshock and chemical convulsant models in that it exacerbates audiogenic seizures in mouse Uobe et al., 1973a,b; Boggan and Seiden, 1971) and produces photosensitivity in previously insensitive baboons (Altshuler et al., 1976). One technique that has been used extensively over the past few years is to study catecholaminergic mechanisms in generalized convulsive as well as partial-seizure models by the use of 6-hydroxydopamine. This is an isomer of NA which, when injected into discrete areas of brain, produces specific degeneration of catecholaminergic neurons (Javoy et d., 1976; Kostrzewa and Jacobowitz, 1974; Wills et al., 1976). By pretreating the animal with certain compounds, one can selectively protect either DA or NA systems from the neurotoxic actions of 6-hydroxydopamine (Pycock et al., 1980; Breese and Taylor, 1971). This technique exacerbates audiogenic (Bourn et al., 1972, 1977; Lehmann, 1977), electroshock (Browning and Maynert, 1978a; Browning and Simonten, 1978; Quattrone et al., 1978; Oishi et al., 1979), PTZ (Tabakoff et al., 1978; Mason and Corcoran, 1979a,b; Corcoran et al., 1973), ethanol withdrawal (Chu, 1978), and barbiturate withdrawal seizures (Morgan, 1976). The change in seizure threshold produced by 6-hydroxydopamine has been attributed more to NA depletion than DA, since protection against NA depletion results in no change in seizure threshold to electroshock in the 6-hydroxydopamine-treated animal (Quattrone et al., 1978). T h e strongest evidence in this regard comes from Mason and Corcoran (1979b) who, with selective depletion of NA and DA in the hippocampus and nigrostriatal pathway, showed that NA-depleted animals had a lower threshold to PTZ with more severe

112

0.CARTER SNEAD 111

seizures. Electroshock also produced more severe seizures in the NAdepleted animals, whereas depletion of DA to 15% of control produced no change in threshold to PTZ or electroschock seizure. Generalized seizures themselves may produce changes in catecholaminergic systems. Ischemia-induced seizures produce reduction in brain levels of NA and DA (Welch et al., 1978), whereas PTZ (McMillen and Isaac, 1978a) and electroshock (Kerty et al., 1969) produce increased turnover of catecholamines in CNS. Electroconvulsive seizure has also been associated with a decrease in both high- and lowaffinity @-adrenergicbinding sites (Bergstrom and Kellar, 1979; Deakin e f al., 1981) with no change in DA or a-NA receptors (Deakin et al., 1981). b. Generalized Absence. There are not as many experimental data to support a role for catecholaminergic mechanisms in generalized absence as in generalized convulsive seizures. Amphetamine suppresses the FEAD model of absence seizures, whereas apomorphine has no effect. T h e FEAD is exacerbated by the dopamine antagonist pimozide and the a-adrenergic antagonist phenoxybenzamine (King and Burnham, 1980). T h e GHB model of absence seizure is associated with marked changes in DA metabolism in brain. GHB produces elevated DA and 3,4-dihydroxyphenylacetic acid (DOPAC) levels in the striatum with a decrease in neuronal firing of DA neurons (Roth, 1976). Because of the prominence of this biochemical effect of GHB ,a cause-effect relationship between the dopaminergic and EEG effects of GHB has been suggested (Walters and Roth, 1972; Snead, 1978d), although evidence in support of this thesis has been contradictory (Snead, 1978d; Godschalk et al., 1977). Recent evidence, however, has indicated that there is a significant difference of time course, dose response, and pharmacologic effect of anticonvulsants between the EEG and DA effects of GHB (Snead, 1980; Snead, 1982). This makes a cause-effect relationship unlikely. r . Myoclotiir. There are no data on the role of catecholamines in myoclonic seizures other than the dopaminergic studies mentioned in conjunction with the GHB absence model, which does have a myoclonic component (Snead, 1978a). d. Pnrtinl Seizures. It is of some interest that given the wealth of literature regarding cellular mechanisms of partial seizures induced by chemical convulsants such as penicillin, alumina gel, and cobalt, there are very few data regarding the contribution of catecholamines to the pathogenesis of seizures in these models (Dow et al., 1974). Adrenergic input has been shown to modulate epileptiform activity in an inhibitory fashion in the intraocular hippocampal transplant model

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

113

(Freedman et al., 1979). Intrastriatal kainic acid, which produces partial complex seizures (Pisa et al., 1980), is associated with increased DA cell firing, and increased DA synthesis and turnover (Baraszkoet al., 1981), but the relationship of these effects to seizure activity has not been addressed. The model of partial seizure for which there is the most data regarding catecholamines is the kindling model (Engel et al., 1981). Chemical kindling may be associated with cellular mechanisms of epileptiform discharge similar to those described above for the penicillin model (Oliver et al., 1980). Electrical kindling is associated with a number of changes in catecholaminergic systems. It appears to produce decreased concentration of NA in whole brain (Engel and Sharpless, 1977; Sato and Nakashima, 1975), as well as in hippocampus, midbrain, limbic lobes, and frontal cortex (Callaghan and Schwark, 1979). There is also a kindling-induced depletion of dopamine in the rat amygdala on the kindled side (Engel and Sharpless, 1977),but apparently no change anywhere else (Callaghan and Schwark, 1979). Kindling is also associated with a decrease in tyrosine hydroxylase activity in amygdala (Farjo and Blackwood, 1978), increased brain turnover of DA and NA (Wilkinson and Halpern, 1979a), and a decrease in high- and low-affinity binding sites for dopamine in the amygdala (Gee et al., 1979), although some authors have found no change in DA receptor binding in kindled animals (Ashton et al., 1980). Pharmacologic reduction of catecholamine function with tyrosine hydroxylase inhibition (reserpine), blockade of DA or P-NA receptors, or treatment with 6-hydroxydopamine all facilitate kindling (Wilkinson and Halpern, 197913; Corcoran et al., 1974; Racine et al., 1979; Arnold et al., 1973; Sato and Nakashima, 1976; McIntyre et al., 1979; Corcoran and Wada, 1979; Mohr and Corcoran, 1981). There are data to support both DA (Wilkinson and Halpern, 1979b; Corcoran et al., 1974) and NA (McIntyre et al., 1979) in the facilitation of kindling by 6-hydroxydopamine suggesting that both are probably involved in a complex way. Kindling is also facilitated by lesions of the stria terminalis (Engel and Katzmann, 1977), but is unaffected by the DA agonists apomorphine or pimozide or by the a blockers, phenoxybenzamine and clonidine (Callaghan and Schwark, 1979; Babington and Wedeking, 1973; Ashton et al., 1980; Sato et al., 1980). Electrical kindling is antagonized by the tricyclic antidepressants, cocaine, and amphetamine (Sato et al., 1980; Callaghan and Schwark, 1979).Chemical kindling with carbachol is inhibited by ICV dopamine and NA (Wasterlain and Jonec, 1980a; Jonec and Wasterlain, 1981; 0. C. Snead, unpublished observations).

114

0. CARTER SNEAD 111

4. Interaction with A n ticow zlulsa nt Drugs Phenytoin, which is an anticonvulsant useful against generalized convulsive, myoclonic, and partial seizures, has a number of interactions with central catecholaminergic systems (Woodbury, 1980). This drug produces a noncompetitive inhibition of norepinephrine uptake in normal synaptosomes, but stimulates NA uptake in anoxic conditions (Hadfield, 1972; Hadfield and Boykin, 1974; Weinberger et al., 1976). Phenytoin is also associated with a decreased uptake of dopamine in caudate putamen and hypothalamus (Hadfield, 1972) and both a competitive and noncompetitive inhibition of dopamine uptake in nigral synaptosome preparations (Hadfield and Rigby, 1976). Phenytoin also inhibits monoamine oxidase (Azzaro and Gutrecht, 1975; Azzaro et al., 1973), produces a dose-dependent inhibition of apomorphine and aniphetamine-induced circling (Elliott et nl., 1977), is associated with a 100% increase in NA concentration in mouse cerebellum (Fry and Ciarlone, 1981), and decreases the therapeutic efficacy of L-DOPA in Parkinson’s disease (Mendez et al., 1975). T h e therapeutic efficacy of phenytoin and phenobarbital, another anticonvulsant drug with the same clinical spectrum, is potentiated by both L-DOPA and amphetamine, unchanged by apomorphine (Kleinrok et al., 1980), and antagonized by 6-hydroxydopamine (Browning and Simonton, 1978). Trimethadione, a n antiabsence drug, has been shown to produce a decrease in the DA metabolite homovanillic acid (HVA) in CSF, but has little effect on PTZ-induced changes in CSF catecholamine metabolite concentrations (McMillen and Isaac, 1978a). The anticonvulsant efficacy of ethosuximide, another antiabsence drug, in the GHB petit ma1 model is diminished by DA receptor blockers and tyrosine hydroxylase inhibition, but not by depletion of NA (Klunk and Ferendelli, 1980). Ethosuximide also attenuates the GHB-induced elevation of striatal dopamine (Snead, 1980). T h e benzodiazepine anticonvulsant clonazepam is effective in generalized absence and myoclonic seizures. This drug has been demonstrated to inhibit L-DOPA and amphetamine but not apomorphine-induced behavior in guinea pig (Weiner et nl., 1977b). Clonazepam and diazepam also counteract the elevation of brain HVA produced by DA receptor blockade either by intensifying GABAergic transmission or via direct action on the benzodiazepine receptor (Keller et al., 1976). Acetazolamide is a carbonic anhydrase inhibitor which possesses anticonvulsant activity against generalized seizure disorders. T h e anticonvulsant efficacy of this compound is decreased by reserpine (Torchiana p t af., 1973; Gray et al., 1958, 1963; Gray and Rauh, 1964, 1967;

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

115

1974; Rudzik and Mennear, 1966). This effect of reserpine is apparently due more to depletion of NA than DA (Torchiana et al., 1973; Browning and Simonton, 1978; Gray and h u h , 1967, 1971). Carbamazepine, an anticonvulsant most effective against partial complex seizures, produces inhibition of uptake and release of NA from brain synaptosomes without a change in regional concentration of dopamine (Purdy et al., 1977; Westerink et al., 1977). The anticonvulsant activity of carbamazepine is significantly diminished by 6-hydroxydopamine (Quattrone and Samanin, 1977; Quattrone et al., 1978). 5. Clinical Cmrelates a. CSF Studies. The measurement of CSF concentration of neurotransmitters and their metabolites in normal and pathologic states may provide a clue to neural function; however, a number of precautions should be taken in interpreting such data. These include ventriculolumbar concentration gradients, conditions of CSF collection, drugs the patient is receiving, and whether the CSF concentration of neurotransmitter and metabolites really reflects the CNS activity of that substance (J. H. Wood, 1980; Garelis and Sourkes, 1974; Garelisetal., 1974; Moiret al., 1970; Sjostrom et al., 1975; Sourkes, 1973; Vecht et al., 1975). The subject of catecholamine metabolites in CSF has been recently reviewed (Wood and Brooks, 1980).The major metabolite of dopamine, HVA, has been reported to be both decreased (Shaywitz et al., 1975; Barolin and Hornykiewicz, 1967; Papeschi et al., 1972) in seizure patients and unchanged from control groups (Garelis and Sourkes, 1974; Laxer et al., 1979; Reynolds et al., 1975; Livrea, 1976; Livrea et al., 1976). No differences have been reported for the NA metabolite 3-methoxy-4-hydroxyphenylethylene glycol (MHPG) in CSF of seizure versus nonseizure patients (Laxer et al., 1979).Seizure patients undergoing cerebellar stimulation for control of their epilepsy have a higher CSF concentration of NA associated with stimulation (Wood et al., 1977b),but the increase does not correlate with control of seizures (Van Buren et al., 1978). Cerebellar stimulation does not change CSF HVA concentration (Wood and Brooks, 1980). 6 . Drugs that Affect Catecholamines. Other clinical lines of evidence for catecholamine involvement in epilepsy lie in the anticonvulsant efficacy of the amphetamines, and the proconvulsant activity of the tricyclic antidepressants and dopamine receptor antagonists. Amphetamines are sometimes useful in nocturnal generalized convulsive as well as generalized absence seizures (Cook and Dole, 1942; Logothetis, 1955; Cohen and Myerson, 1938; Livingston et al., 1973,

116

0. CARTER SNEAD 111

1974; Menkes, 1973). There are sporadic case reports concerning the clinical anticonvulsant efficacy of L-DOPA (Cools et al., 1975), although

more extensive controlled trials have not confirmed this impression (Chadwick et al., 1978). The non-MA0 inhibitor antidepressants, that is, the tricyclics, experimentally lower the seizure threshold (Wallach et al., 1969) and in clinical practice may precipitate seizures at therapeutic doses (Trimble, 1978; Pineda and Russell, 1974). Similarly, the psychotropic drugs which are dopaminergic blockers, the phenothiazines and butyrophenones, are also epileptogenic (Itil and Soldatos, 1980; Logothetis, 1967; Borenstein et al., 1962). Catecholamine depletion with reserpine is also known to aggrevate seizure disorders clinically (Pallister, 1959).

B. SEROTONIN 1. Anutomj

The main source of serotonin (5-HT) in the forebrain comes from cells lying in or near the midline of raphe regions of the pons and upper brainstem (Table IV). The major serotinergic innervation to the telencephalon and diencephalon is provided by the dorsal and median raphe nuclei (Azrnitia, 1979). The dorsal raphe forebrain tract and median raphe forebrain tract run in the median forebrain bundle (MFB); and the dorsal raphe arcuate tract, periventricular tract, and cortical tract traverse a route external to the MFB. The dorsal raphe projects primarily to cortex, basal ganglia, thalamus, hippocampus, amygdala, hypothalamus, and septum, while the median raphe projects to the TABLE IV CEHTRAL SEROTINERCIC PROJECTION" Cells of origin

Dorsal raphe (DR)

System

DR cortical tract DR arcuate Periventricular tract Forebrain tract

Median raphe (MR)

MR forebrain tract Raphe medial tract

' From Azmiua (1978).

Projection Cortex, basal ganglia Thalamus (ventrolateral geniculate), Substantia nigra Midline thalamus, periventricular nucleus Amygdala, basal ganglia, hippocampus, septum Preoptic area, olfactory bulb, hippocampus, septum Mamillary body, interpenduncular nucleus

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

117

preoptic area, olfactory bulb, hippocampus, and septum. The serotinergic projections to amygdala are exclusively from dorsal raphe forebrain tract. The main projection to the septa1 hippocampal complex is the median raphe forebrain tract. The dorsal raphe cortical tract projects to the temperoparietal cortex, while the median raphe cortical tract projects to frontal, cirgulate, and entorhinal cortex. 2. Physiology The discrete location of serotinergic neurons in the midbrain raphe area has permitted a number of recording and iontophoretic experiments. The serotonin (5-HT) neurons in this area have a slow, spontaneous rate of firing which decreases with rapid eye movement (REM) sleep (Aghajanian, 1972; McGinty and Harper, 1976). Serotinergic projections are inhibitory to thalamus (Bloom et ad., 1972), amygdala (Wang and Aghajanian, 1977),septum (Segal, 1974), and hippocampus (Segaf, 1975). 3. Role in Models Much of the work cited above in the course of discussion of catecholaminergic mechanisms in seizures also applies to serotonin. Much of the experimental evidence that appears to include or exclude serotonin’s role in various seizure states is clouded by the use of drugs, such as reserpine, which are rather nonspecific in terms of which monoamine they affect. The use of other pharmacologic techniques such as p-cholorophenylalanine (pCPA) to deplete 5-HT, specific serotonin agonists and antagonists, and lesioning of serotinergic pathways and the raphe nuclei have helped to clarify those issues (Essman, 1978). a. Generalized Seizures. The effect of serotonin antagonists or agonists on seizure thresholds vanes according to the model used. When Chen et al. (1954) reported the effect of reserpine in decreasing the threshold to ECS, it was unclear as to what role, if any, serotonin played in this phenomenon. Azzaro et al. ( 1972), utilizing serotonin antagonists and precursors, concluded that both catecholamines and serotonin were required for this phenomenon. Further evidence for serotinergic modulation of ECS threshold came from experiments demonstrating that inhibition of sertonin synthesis with pCPA or serotinergic receptor blockade with crypropheptadine resulted in lowering the threshold to ECS (Koe and Weissman, 1966, 1968; Van Riezen, 1972; Kilian and Frey, 1973; Wenger et al., 1973; Chen et al., 1968), while 5-hydroxytryptophan (5HTP), a serotonin-releasing agent decreases the severity of electrically induced seizures (Kilian and Frey, 1973; Buterbaugh, 1977; Fuller et al., 1974; Przegalinski, 1976a). However, there are some conflicting data

118

0. CARTER SNEAD 111

that tend to refute involvement of serotonin in modulation of the electroconvulsive threshold Uobe et al., 1974; Rudzik and Johnson, 1970; Heymans et al., 1964). The most compelling experimental work along these lines has utilized selective destruction of serotinergic neurons with 5,7-dihydroxytryptamine and lesioning of the raphe nuclei (Crunelli et al., 1979). This selective lesioning of serotinergic pathways had no effect on electroconvulsive shock threshold nor did administration of serotinergic drugs. A decrease in threshold was produced by pCPA, but these authors proposed a nonserotinergic action of pCPA to explain this. Other authors have also shown no change in electroconvulsive threshold with raphe iesions (Quattrone ~t al., 1978). Introcerebroventricular serotonin has little or no effect on electroshock seizures if the hypothermia associated with these seizures is eliminated (Browning and Maynert, 1978b). The evidence for anticonvulsant effect of serotonin in PTZ-induced generalized convulsive seizures is more clear cut. Depletion of serotonin lowers the threshold for PTZ seizures (Kilian and Frey, 1973; Przegalinski, 1975; Alexander and Kopeloff, 1970). Administration of 5-HTP in conjunction with a peripheral decarboxylase inhibitor raises the PTZ threshold as does ICV serotonin (De la Torre and Mullan, 1970; De la Torre et al., 1970; Przegalinski, 1975; Kobinger, 1958; Schlesinger et al., 1969). Pentylenetetrazol-induced seizures are reduced in severity by raphe stimulation, and this is blocked by serotonin antagonists (Kovacs and Zoll, 1974; Wilkinson and Halpern, 1975). Serotonin has been extensively investigated in genetic seizure models. Jobe et 01. ( 1 9 7 3 ~have ) shown that pCPA treatment results in an increased intensity of audiogenic seizures in mice which is blocked if brain serotonin is increased with iproniazid and 5-HTP. Introcerebroventricular 5-HTP also decreases the intensity of audiogenic seizures (Schlesinger et al., 1969). Other authors have provided further evidence that audiogenic seizures are exacerbated with depletion and antagonized by elevation of 5-HT in brain (Lehmann, 1967; Alexander and Alexander, 1976). The C57-B1 seizure-resistant mouse appears to have a greater capacity for serotonin synthesis by virtue of higher brain activity of tryptophan h ydroxylase than the DBA/l seizure-prone animal (Ginsburg and Sze, 1975). The effect of serotonin in audiogenic seizures may be species specific, since in some strains pCPA results in little or no change in seizure threshold (Alexander et al., 1971; Alexander and Kopeloff, 1976). Although there is a decreased concentration of 5-HT in the hemispheres of epileptic fowl, increasing the brain level does not alter seizure

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

119

susceptibility (Johnson et al., 1979, 1981). The brain concentration of serotonin in epileptic beagle dog is normal (Edmonds et al., 1979). Administration of 5-HTP protects baboons with photosensitive epilepsy from seizures (Wada et al., 1972) and also blocks imipramine-induced aggrevation of seizure in this model (Trimble et al., 1977); however, ICV serotonin and systemic tryptophan do not affect seizure susceptibility in this model (Altshuler et al., 1976). The latter authors concluded from their experiments that cerebral concentrations of serotonin do not serve a major regulatory role in photosensitivity of the baboon. The major difficulty in interpreting these data concerning serotonin in baboon photomyoclonus is the lack of corresponding direct biochemical data with regard to 5-HT in brain. Seizures induced by hyperbaric oxygen are prevented by serotonin (Essman, 1978) and are associated with decreased brain levels of serotonin (Faiman and Heble, 1966; Huggins and Nelson, 1975; Tunnicliff et al., 1973). As with the catecholamines, generalized seizure per se may result in changes in concentration of 5-HT in brain. Although there is some evidence to the contrary (Bonnycastle et al., 1957), the preponderance of data suggest that both ECS- and PTZ-induced seizures result in lowered brain serotonin (Garratini and Valzelli, 1957; Breitner et al., 1961, 1964; Tagliamonte et al., 1972; Diaz, 1970; Essman, 1968, 1978; Kety et al., 1969; Bertaccini, 1959). 6 . Generalized Absence. There are little or no data concerning serotinergic mechanisms in the various models of absence seizures. Gamma hydroxybutyrate has been reported to produce an increase in striatal and mesolimbic 5-HT (Waldmeier and Fehr, 1978),but there are no data as to a possible relation between this and the epileptogenic action of the drug. c. Myoclonic. Although there is some evidence that 5-HT receptor stimulation may result in myoclonus (Klawans et al., 1973; Stewart et al., 1976; Growdon, 1977), drugs that enhance serotinergic activity reduce, and serotonin antagonists enhance, p,p '-DDT-induced myoclonic seizures. Serotonin concentrations are increased in midbrain and cerebellum and 5-Hydroxyindole acetic acid (HIAA) levels are increased diffusely throughout brain in this model (Hwang and Van Woert, 1978). d. Partial Seizures. As with the catecholamines, most of the data concerning 5-HT in partial seizures relate to the kindling model. Colasanti and Craig (1973) have provided some data using the cobalt model and have shown that 5-HT levels and turnover are not altered in the seizure focus. Unilateral injection of another toxin, tungstic acid, into the hip-

120

0. CARTER SNEAD 111

pocampus produces partial seizures associated with a regional increase of 5-HT in the damaged hippocampus as well as in the brainstem (Okada, 1973). There are a number of data to suggest that 5-HT has inhibiting properties in amygdaloid kindling (Kovacs and Zoll, 1974; Zoll et al., 1976). Facilitation of amygdaloid kindling is produced by destructive lesions of serotinergic pathways (Racineet al., 1979),whereas stimulation of the raphe seems to retard progression of the kindling process (Kovacs and Zoll, 1974; Siege1 and Murphy, 1979). However, there is some contradictory data that suggests that 5-HT agonists and precursors exacerbate, and antagonists decrease, the kindling process (Ashton et al., 1980). These same authors have shown no change in the number of 5-HT receptor sites in multiple brain regions of kindled versus nonkindled animals. 4. Interaction uith Anticonvulsants Phenytoin produces an increase in brain 5-HT concentration (Bonnycastle et d., 1962; Chase et al., 1970; Anderson et al., 1962; Jenner et nl., 1975; Fry and Ciarlone, 1981).Single-dose phenytoin has no demonstrable effect of 5-HT synthesis, whereas chronic administration results in increased synthesis of 5-HT but does not affect uptake (Green and Grahame-Smith, 1975). Phenobarbital is associated with a decrease in 5-HT turnover in brain at high doses (Lidbrink et nl., 1974). Serotonin facilitates the anticonvulsant activity of phenobarbital, but 5-HT antagonists do not alter the anticonvulsant dose requirement of this drug (Meyer and Frey, 1973; Przegalinski, 1975). The anti-petit-ma1 drug, trimethadione, produces an increase in brain 5-HT turnover with an increased rate of synthesis which is antagonized by PTZ (Diaz, 1974).The benzodiazepines decrease serotonin turnover (Wise et al., 1972). Clonazepam produces a dosedependent increase in 5-HT and its major metabolite, 5-HIAA, that is much more potent than that seen with diazepam or phenytoin (Jenner et al., 1975; Fennessy and Lee, 1972; Chase et al., 1970; Fernstrom et al., 1974),but is ineffective against 5-HTP-induced myoclonus in guinea pig (Weiner et al., 1977a). Valproic acid is an anticonvulsant which is clinically useful against generalized absence and myoclonic seizures. This drug produces an increase in brain tryptophan and 5-HIAA, but not in 5-HT levels. Valproic acid also potentiates the antimyoclonic action of chlorimipramine in the p,p’-DDT animal model of myoclonic seizures (Hwang and Van Woert, 1979).

SACRED

DISEASE: NEUROCHEMISTRY OF EPILEPSY

121

Raphe lesions do not affect the anticonvulsant efficacy of phenytoin, phenobarbital or carbamazepine (Quattrone et al., 1978). The carbonic anhydrase inhibitors, although thought to exert their effects via catecholaminergic mechanisms (see above), do have some effect on serotonin. These drugs produce elevated brain 5-HIAA, and their anticonvulsant efficacy is decreased with a corresponding decrease in brain serotonin (Przegalinski, 1976a,b). 5. Clinical Correlates a. CSF Studies. The measure of 5-HIAA, the major metabolite of 5-HT, in CSF has been used extensively to implicate serotonin in the pathogenesis of a number of disease processes involving the CNS (Bowers et al., 1969). The reservations outlined above for CSF studies of catecholamines perhaps apply even more to experimental data on CSF 5-HIAA since a significant percentage of this metabolite found in CSF is contributed from spinal cord (Bulat, 1977; Garelis and Sourkes, 1973; Sjostrom et al., 1975; Wier et al., 1973). That said, there are a number of clinical studies of CSF levels of 5-HIAA in seizure states. Shaywitz et al., (1975) have demonstrated low levels of HIAA in probenecid-treated seizure patients. The decrease was unrelated to age or anticonvulsant medication. Although Shaywitz’sfindings have been confirmed by some (Wood and Brooks, 1980), other investigators (Chadwick et al., 1975a, 1977a) have shown normal 5-HIAA levels in untreated seizure patients and elevated CSF 5-HIAA concentrations with phenytoin and phenobarbital treatment; however, Chadwick and colleagues did not utilize probenecid loading, so it is difficult to compare his data to those of Shaywitz. Laxer et al. ( 1979)have made yet a different observation in that they found no significant difference in CSF 5-HIAA between seizure and nonseizure or treated vs untreated seizure patients. There is general agreement however that CSF 5-HIAA is lowered in myoclonic seizure disorders (see below). b. Drug Studies. Although L-tryptophan has been tried unsuccessfully in generalized convulsive and partial-seizure disorders (Chadwick et al., 1978),there is considerable evidence for the clinical usefulness of the serotonin precursor, 5-HTP, in various myoclonic disorders. The initial work along these lines came from Lhermitte et al. (1971, 1972), who showed that 5-HTP had a beneficial effect on two patients suffering from intention myoclonus. This entity, closely akin to a myoclonic seizure disorder, refers to an involuntary, arhythmic jerking of muscles which is induced by willed muscle activity as well as by auditory, visual, tactile, and emotional stimuli (Lance and Adams, 1963). Since Lhermitte’s orig-

122

0. CARTER SNEAD 111

inal papers, a number of authors have confirmed this observation using 5-HTP in conjunction with a peripheral decarboxylase inhibitor (Chadwicket al., 1974, 1975b; Lhermitte et al., 1975; Guilleminault et nl., 1973; Magnussen et al., 1978; De Lean et nl., 1976; De Lean, 1977; Van Woert and Sethy, 1975). These findings have been extended to other types of neurologic disorders associated with myoclonus including seizures (Van Woert et nl., 1977; Growdon et al., 1976; Magnussen et al., 1977; Chadwick et nl., 1977b). Although there is some variation of opinion in terms of improving the functional state of these patients with 5-HTP (Thal et nl., 1979), there is near unanimity concerning its effectiveness in reducing the frequency and severity of myoclonic jerks, as well as decreasing the electrical paroxysms that sometimes accompany them. There is also agreement among these authors that CSF 5-HIAA is low in patients with myoclonus and is elevated by drugs that are effective against the myoclonus such as 5-HTP or some other anticonvulsant (Fahn, 1978), although a low CSF 5-HIAA does not predict a positive clinical response to 5-HTP (Thal et al., 1979). C. ACETYLCHOLINE 1. Anatomj Current knowledge concerning the anatomic distribution of cholinergic systems in brain (Table V) is based on histochemical studies of acetylcholinesterase (ACHE) (Lewis and Shute, 1967, 1978; Koelle, 1949,1954; Jacobowitz and Palkovits, 1974; Shute and Lewis, 1967) and TABLE V CENTRAL CHOLINERGIC PATH\\'AYS" Cells of origin

Tract

Projection ~

Pontine reticular formation Midbrain reticular formation

Ventral tegmental pathway, dorsal tegrnental pathway

Globus pallidus Septum Amygdala Caudate-putamen a

From Lewis and Shute (1978).

~~

Cerebellum Lateral preoptic area, globus pallidus, midbrain reticular formation Cortex Hippocampus Am ygdala Caudate- putamen

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

123

choline acetylase (Storm-Mathisen, 1977; Palkovits et al., 1974; Kobayashi et al., 1975; Brownstein et al., 1975), as well as on direct measurement of acetylcholine (ACH) (Hoover et al., 1978) and regional receptor binding studies of muscarinic (Kobayashi et al., 1978) and nicotinic (Morley et al., 1977, 1979) receptors. The topography of cholinergic pathways in brain is not as well known or discrete as that of catecholaminergic or serotinergic projections. The first forebrain cholinergic pathway identified was that arising from the medial septa1 and diagonal band nuclei and projecting to the hippocampus and dentate gyrus (Lewis et al., 1964, 1967). The cholinergic projections in the amygdala are both intrinsic and projected from the lateral preoptic nuclei. There is substantial cholinergicinput to cerebellum, basal ganglia, and thalamus. Cholinergic neurons are also intrinsic to cerebral cortex, forming circuits in deeper cortical layers and synapsing with pyramidal cells of layer V. These axons also ascend to layer I (Hebb et al., 1963; Krnjevic and Silver, 1965).

2. Physiology Acetylcholine is excitatory to some cortical (Krnjevic and Reinhardt, 1979; Krnjevic and Phillis, 1963; Ferguson and Jasper, 197l), cerebellar (Crawford et al., 1966), reticular formation (Velasco et al., 1981a), hippocampal (Briscoe and Straughan, 1966), and caudate (Bloom et al., 1965) neurons. There is also evidence for inhibition of superficial cortical (Randit et al., 1964; Phyllis and York, 1967), hippocampal (Valentino and Dingledine, 198l), medullary, and hypothalamic neurons (Bradley, 1968; Salmoiraghi and Steiner, 1963; Bradley et al., 1966) by ACH. Although it has been hypothesized that this difference in excitation and inhibition may be due to whether nicotinic or muscarinic receptors are being stimulated (Maynert et al., 1975; Bradley et al., 1966), the issue of CNS nicotinic cholinergic pharmacology is controversial (Morley et al., 1979).Some have suggested that nicotine produces its neurophysiologic and neuropharmacologic action on brain by noncholinergic mechanisms (Abood et al., 1979). 3. Role in Models The use of pharmacologic manipulation of CNS cholinergic systems to investigate a role for ACH in various seizure models is complicated by (1) the fact that ACH itself clearly has convulsant properties in brain (Karczmar, 1979) and (2) the apparent coexistence of both muscarinic acetylcholine receptors (mACHR) and nicotinic acetylcholine receptors (nACHR) in brain (Morley et al., 1977, 1979).

I24

0.CARTER SNEAD I11

In 1949 Kristiansen and Curtois showed that the combination of ACH and physostigmine, an anticholinesterase drug, when applied to isolated cortex resulted in electrical seizure activity. Since that time, the isolated cortex has been shown to be supersensitive to the epileptogenic effects of ACH (Echlin and Battista, 1963), possibly secondary to decreased activity of ACHE (Rosenberg and Echlin, 1965). Generalized convulsive seizures usually result from local application or systemic administration of ACH or its agonists (Grossman, 1963; Whalstrom, 1978; Cools et al., 1975; Guerrero-Figueroaet al., 1964; Haley and McCormick, 1957; Feldberg and Sherwood, 1954; Lynch et al., 1971; Crawford et al., 1966; Ferguson and Cornblath, 1975; Tan, 1977; Baker and Benedict, 1968; Babb et al., 1973), precursors (Wahlstrom, 1978; Hanbrich et al., 1975), and ACHE inhibitors (Bokums and Elliott, 1968; Girgis, 1978; Cetesia and Jasper, 1966; Sie et al., 1965; Van Meter et al., 1978; Wills, 1970; Baker and Benedict, 1968). The production of seizures in isolated cortex by ACHE inhibitors has been postulated to be due to natural release of ACH in the presence of low ACHE (Celesia and Jasper, 1966; Ferguson and Jasper, 1971). Although most of the data above deal with muscarinic agonists, nicotine, when given ICV, also produces seizure activity as do the nicotinic antagonists d-tubocurarine and a-bungarotoxin (Cohen et al., 1981; Abood et al., 1979; Ashorabi et al., 1979). Acetylcholine muscarinic antagonists have both convulsant and anticonvulsant effects (Longo, 1956, 1966; Karczmar, 1979). Atropine induces EEG hypersynchrony which is abolished by physostigmine (Longo, 1966), blocks electrical spiking that results from local applications of ACH (Miller et at., 1!340), and reverses ACHE-induced generalized convulsive seizure activity (Essig et al., 1950; Longo et al., 1960). Scopolamine has similar effects but is more potent (Longo, 1956). Both scopolamine and atropine produce epileptiform activity when applied topically to cortex (Tan et al., 1978; Daniels and Spehlman, 1973; Dudar and Szerb, 1969). a. Generalized Convulsive. Acetylcholinesterase inhibitors lower the threshold for ECS (Baker, 1965; Baker and Benedict, 1968). Both ECSand PTZ-induced seizures are associated with decreased brain acetylcholine (Longoni et al., 1976a,b; Pedata et al., 1976; Essman, 1972) and an increased efflux of acetylcholine from brain (Gardner and Webster, 1977; Hemsworth and Neal, 1968; Beselin et al., 1965). Pentylenetetrazol also is associated with an increase in sodium-dependent, high-affinity choline uptake in brain which can be blocked in the hippocampus with leisioning of cholinergic septohippocampal tracts (Simon et al., 1976). The effect of physostigmine on ECS threshold is dose dependent with

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

125

low doses increasing, and high doses decreasing, the threshold (Chen et al., 1968; Williams and Russell, 1941; Ikonomoff, 1970). Atropine is not effective against seizures induced by PTZ and ECS (Longo, 1966; Funderburk and Case, 1951). In the genetic models of generalized convulsive seizures, atropine reduces audiogenic seizures in rats, whereas physostigmine exacerbates those seizures in a dose-dependent fashion (Humphrey, 1942). The audiogenic-seizure-prone mice and posture-induced seizure-prone eb mice both show increased brain concentration of ACH compared with seizure-resistant animals (Naruse et al., 1960; Takahashi et al., 1961; Kurokawa et al., 1963; Pryor, 1968). However, younger seizure-resistant animals in these strains have higher levels of ACH than the older seizure-prone animals (Naruse et al., 1960; Schlesingeret al., 1965; Fink, 1966; Reeves, 1966). Neither atropine nor scopolamine have any effect one way or another on photogenic seizures in baboons (Meldrum et al., 1970). b. Generalized Absence. There are few data concerning ACH and absence seizure models. GHB produces a significant increase in acetylcholine in rat brain which is specific for cortex and midbrain (Giarman and Schmidt, 1963; Sethy et al., 1976)) and a decrease in high-affinity choline uptake (Simon et al., 1976), but these changes are apparently unrelated to the EEG and behavioral effects of GHB (0.C. Snead, unpublished observations; Davies, 1978; Sethy et al., 1976). Carbachol, when given ICV in a dose of 1 p g produces seizure activity that is petitmal-like in terms of absence behavior and blockade by anti-petit-ma1 anticonvulsants (Snead, 198 lb); however, this is a very dose-specific effect with higher doses producing generalized convulsive seizures. c. Generalized Myoclonic. There are very few data on cholinergic mechanisms in myoclonic seizure models. Neither physostigmine nor benztropine have any effect on myoclonic activity in thep,p '-DDT myoclonic model (Hwang and Van Woert, 1978). d. Partial Seizures. Atropine is ineffective in the penicillin model of partial seizures (Longo 1966; Goodman and Lebovits, 1980). Experimental alumina gel seizure foci show increased ACHE activity as do the mirror foci. The latter return to normal with removal of the primary seizure focus (Pope et al., 1947; Guerrero-Figueroa et al., 1964). The partial seizures that result from cobalt implantation are antagonized by cholinergic agonists and facilitated by cholinergic antagonists (Hoover et al., 1977). Carbachol also exacerbates partial seizures from alumina gel foci (Velasco et al., 1981b). Kainic acid produces a decrease in the activity of choline acetyltransferase as well as the density of MACHRs (Zaczek et al., 1978), but to

126

0. CARTER SNEAD 111

date there is no correlation between this biochemical effect of kainic acid and its epileptogenic action. Experiments indicating that cholinomimetics produce seizure and hypersynchronous activity in the hippocampal explant which is blocked by muscarinic antagonists suggest that cholinergic input modulates epileptiform activity in this model (Freedman ri nl., 1979). There are a fair amount of data available concerning cholinergic mechanisms in kindling. Part of the reason for interest in the role of ACH in the kindling process lies in the fact that it is possible to produce the kindling paradigm with repeated aministration of either the cholinergic agonist carbachoi (Vosu and Wise, 1975; Wasterlain and Jonec, 1980a) or the ACHE inhibitor physostigmine (Girgis, 1980, 1981). This cholinergic kindling appears to be mediated via muscarinic receptors (Wasterlain and Jonec, 1980b; Jonec and Wasterlain, 1981; Wasterlain and Jonec, 1981). Electrical kindling is associated with neuronal supersensitivity to ACH which correlates with progression of the kindling process (Burchfiel ut nl., 1979). Arnold et al. (1973) have reported that atropine significantly retarded the progression of electrically kindled seizures, but others have been unable to duplicate this finding (Wada, 1977; Corcoran et al., 1976; Ashton et al., 1980), and there is one report (Fitz and McNamara, 1979) that atropine activates interictal spiking in kindled animals. Joy et al. (1981) have shown no change in kindling with chronic cholinesterase inhibition. Electrical kindling produces no change in choline acetylase or ACHE activity or choline uptake, but is associated with an apparent decrease in muscarinic binding sites in dentate and amygdaia (McNarnara, 1978; Dasheiff et al., 1981; Byrne et al., 1980). 4. Ititemctioii zrith Antironvulsa nt Drugs

Low doses of phenytoin are associated with an increased production of ACH by brain slices and parasympathetic ganglia, whereas high-dose phenytoin has an opposite effect (Woodbury and Kemp, 1971; McLennon and Elliott, 1951). Phenytoin produces a decrease in concentration of ACH in rat brain as well as the cerebral cortex of epileptic guinea pig, but has no effect on the seizure-resistant strain of the latter (Agarwal and Bhargava, 1964; Bianchi et at., 1975; Domino and Olds, 1972). Phenytoin also decreases the amplitude of cholinergic EPSPs and increases the amplitude of K+-dependent IPSP without affecting the ClF-dependent IPSP in invertebrate neurons (Ayala et al., 1977). Phenobarbital blocks cholinomimetic-induced seizures in the hippocampal explant (Freedman ut ai., 1979). Barbiturates have been reported to block the excitatory action of ACH on brainstem and cortical

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

127

neurons (Bradley and Dray, 1973; Crawford, 1970; Crawford and Curtis, 1966; Prichard, 1980) but have no effect on neocortical cells (Smaje, 1976). They also decrease the amplitude of cholinergic' EPSPs without affecting either K+- or C1--mediated IPSP (Barker and Gainer, 1973). The data concerning the effect of barbiturates in ACH metabolism in brain varies according to which barbiturate is used. Schultz (1943) has shown that single, anesthetizing doses of barbiturates do not change ACHE activity in brain but chronic dosing results in a decrease in this activity. Pentobarbital produces an increased level of ACH in brain with a decreased rate of synthesis in hippocampus and cortex (Nordberg and Sundwall, 1977a,b; Holmstedt et al., 1963; Richter and Crossland, 1949). Trimethadione in studies on the postsynaptic cholinergic receptor in Aplysia depresses the Na+-mediated EPSP and the C1--mediated IPSP but has no effect on K+-mediated IPSP (King and Kreisman, 1981). This anticonvulsant also depresses the miniature end plate potentials at the neuromuscular junction of frog indicating some action of ACHRs (Alderdice and McMillan, 1980). Trimethadione has no effect on choline acetylase or ACHE activity and does not prevent PTZ-induced increase in ACH synthesis (Torda and Wolff, 1947). Diazepam produces increased levels of ACH in striatum, hemispheres, and hippocampus of rat, mouse, and guinea pig; causes a decreased turnover of ACH in cortex and midbrain; but has no effect on choline content (Killiam and Suria, 1980; Consoloet al., 1975; Zsilla et al., 1976; Cheney et al., 1977; Uhl and Snyder, 1976). Diazepam also antagonizes the hyperactivity syndrome in mice produced by anticholinergic drugs (Soubrie et al., 1976). Another benzodiazepine anticonvulsant, clonazepam, is effective against seizures induced by ACHE inhibitors (Lipp, 1972, 1973, 1974) and suppresses atropine-induced spikes in the limbic system (Herink et al., 1980).

5. Clinical Correlates Acetylcholine in CSF is difficult to measure because of the difficulty of available assay procedures and the instability of ACH. This compound has been shown to be present in CSF (Duvoisin and Dettbarn, 1967) and has been reported to be elevated in seizures (Tower and McEarchem, 1949; Tower and Elliott, 1952; Schain, 1960); however, the relation of this finding to actual CNS cholinergic function in these patients is not known. ACHE activity is said to be increased in epileptic foci clinically (Pope et al., 1947; Pappins and Elliott, 1958), while ACHE inhibitors have been reported to both exacerbate and facilitate the management of absence seizures (Williams and Russell, 1941 ; Ikonomoff, 1970; Mazars et al., 1966). Although Wolff (1956) has reported the effec-

128

0. CARTER SNEAD 111

tiveness of atropine and scopolamine in generalized convulsive and absence seizures, there has been very little clinical confirmation of this observation (Millichap ei al., 1968). There are a number of reports of an increased frequency of seizure disorders among patients with myasthenia gravis with an equally high incidence of EEG abnormalities (Hokkanen and Toivakka, 1969; Hoefer et nl., 1958; Fearnsides, 1915; Mortier et al., 1971; Snead et al., 1980). T h e cause of this increased incidence of seizure is unknown, but the following lines of evidence suggest that antibody to nACHR may play a role: (1) T h e CNS is affected in experimentally induced myasthenia (Fulpius et al., 1977), (2) antibody to nACHR has been found in CSF of some myasthenic patients (Lefvert et al., 1978), and (3) nACHR antibody has been reported to be present in the sera of patients with seizures and IgA deficiency but no myasthenia (Fontana et nl., 1978a,b; Fontana and Grob, 1979). Evidence against this thesis lies in the fact that sera containing nACHR antibody from patients with seizures is not epileptogenic when given ICV to rats nor is electric fish anti-nACHR antibody (Cohen et al., 1981). In addition, the sera from myasthenic patients with seizures does not interact with CNS a-bungarotoxin binding sites (Morley and Snead, 1979).

D. ~-AMINOBUTYRIC ACID 1. Anatomny

Extensive regional distribution studies of 7-aminobutyric acid (GABA) and the enzymes involved in its metabolism have been done in human, rhesus monkey, and other species (Fahn and Cote, 1968; Miller and Pitts, 1967; Kanazawa et al., 1973; Fonnum et al., 1974; Roberts, 1978; Walaas and Fonnum, 1979). y-Aminobutyric acid is present in brain, spinal cord, and retina, but not in peripheral nerve. In brain, it occurs predominantly in gray matter with the highest concentration in human occurring in globus pallidus and substantia nigra (Table VI). The next highest amounts are found in hypothalamus and nucleus accumbens. Glutamic acid decarboxylase (GAD) is the enzyme responsible for formation of GABA from glutamic acid. Glutamic acid decarboxylase activity parallels GABA concentrations in human (Table VI) and other species, with the highest activity occurring in hypothalamus, limbic system, and globus pallidus as demonstrated by elegant neuroanatomical and immunocytochemical techniques (Roberts, 1978). Because of the close correlation between GAD activity and GABA levels in brain, GAD is used as a marker enzyme to identify GABAergic neuronal pathways.

129

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

TABLE VI

REGIONAL DISTRIBUTION OF GABA MARKERS IN MONKEYBRAIN"

Brain region

[3H]GABA receptor binding (fmoYmg protein)

Cerebral cortex Frontal Temporal Occipital Hippocampus Amygdala Hypothalamus Thalamus Caudate Putamen Globus pallidus Substantia nigra Cerebellum

1.53 0.67 2.03 0.74 1.40 0.41 1.02 2.52 2.37 2.00 1.14 1.72

GABA (pmoYgm)

GAD activity (pmol CO,/gm protein/H)

2.10

70

2.68

84 9.9 14.7

-

6.19 2.68 3.20 3.62 9.54 9.70 2.03

-

20 12.2 18.4 18.4 36.4 35.1 9.7

GABA uptake (fmoYmg protein)

607 739 533 390 847 787 224 339 299 852 587 229

" From Enna (1981) and Fahn (1976). Although the data in human studies is not complete, the animal data suggests that little or no correlation exists between other enzymes in the GABA metabolic pathway and the concentration of GABA in discrete areas of brain (Fahn, 1976). Although GABA levels in cerebellar cortex are less than half those in globus pallidus and substantia nigra, much interest has centered about the metabolism, distribution, and function of GABA in cerebellum. T h e reasons for this interest are twofold: First, other neurotransmitters such as dopamine and serotonin are conspicuous by their low levels or absence in cerebellum, whereas GABA is present in clearly definable amounts. Second, the output of the cerebellum is solely inhibitory (Eccles et al., 1976) via the Purkinje cells that utilize GABA as their inhibitory neurotransmitter (It0 and Yoshida, 1964; Ito et al., 1964; Obata and Takeda, 1969). Immunocytochemical techniques that enable visualization of GAD at light and electron microscopic levels have thus demonstrated well-defined GABAergic pathways in the cerebellar Purkinje system which terminate in subcellular nuclei and Deiters lateral vestibular nucleus. To date, such techniques have also demonstrated GABAergic pathways from caudate, putamen, or globus pallidus to substantia nigra and possibly from the stria medullaris to the lateral habenular nucleus (Fonnum etal., 1974; Roberts, 1978; Galeet al., 1977; Hattoriet al., 1973; Van der Heyden et al., 1979).

130

.

0. CARTER SNEAD 111

2. Phyaologv y-Aminobutyric acid is generally thought to exert an inhibitory effect physiologically throughout the CNS (E. Roberts, 1974; Krnjevic, 1976; Matthews et nl., 1981b). This amino acid alters membrane conductance to chloride ions with the membrane potential staying near the resting level in mammalian cerebral cortex (Krnjevic and Schwartz, 1967; Dreifuss et nl., 1969), Dieter’s nucleus (Obata et al., 1970; Ten Bruggencate and Engberg, 1971; Curtiset nl., 1970a; Ito and Yoshida, 1964), and cerebellum (It0 rt al., 1964). Since GABA is thus considered to be a primary inhibitory neurotransmitter in brain, any derangement of inhibitory states in the nervous system which might lead to a chronic seizure state could presumably involve GABAergic neurons in some way. Hence, there has been considerable interest over the years concerning the role of GABA in the pathogenesis of epilepsy that has led to a voluminous literature on the subject (Wood, 1975; Tower, 1976; Meldrum, 1975; Emson, 1975; Snead, 1981a) which exceeds that concerning any other neuroactive substance and epilepsy. 3. Role in iWodel.y A central role for GABA in the pathophysiology of a variety of seizure types is inferred by the fact that generalized convulsive seizures always occur secondary to impairment of GABA synthesis (Horton and Meldrum, 1973; Metcalf, 1979; Beart and Bilal, 1979; Killam and Bain, 1957; Killam, 1957; Horton et al., 1978; Roa et nl., 1964; Wood and Peesker, 1974; Murakami ef nl., 1976; Ashton and Wauquier, 1979; Loscher, 1979), release (Curtis at nl., 1973), receptor binding (Mathers and Barker, 1981; Matthews et al., 1981a; Curtis et nl., 1970b, 1971b; Macdonald and Barker, 1977; Worms and Lloyd, 1978; Olsen et al., 1976; Enna, 1981), or a block in GABA-mediated changes in C1- conductance (Olsen ef al., 1978, 1980; Horton et al., 1976). Furthermore, GABA per se has been shown to have anticonvulsant properties in a number of experimental seizure states (Kubrin and Seifter, 1966; Bed et al., 1961; Wood et nl., 1963; Ballantine, 1963; Feher et al., 1965; Guerrero-Figueroa et nl., 1964; Crighel, 1966). In view of the above, there has been considerable interest in elucidating more precisely the role of GABA in the epilepsies (Meldrum, 1975). T h e pharmacologic techniques utilized toward this end include the use of GABA agonists, mainly muscimol, baclofen [P-(p-ch1oro)phenyl GABA], T H I P [4,5,6,7tetrahydroisoxoazolo(5-4-c)pyridine], 3-aminopropane sulfphonic acid (3-APS), a n d SL 76002 [[a-(chloro-4-phenyl)fluoro-5-hydroxy-2benzilidene amino]-4-butyramide] (Morselli et al., 1980; Meldrum, 1978, 1979, 1981; Meldrumetni., 1980; Lloyd and Worms, 1981; Slater

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

131

1980; Enna and Maggi, 1979; Lloyd et al., 1981); GABA uptake inhibitors such as nipecotic acid, 2,4-diaminobutyric acid; and p-alanine (Meldrum, 1978; Frey et al., 1979; Taberner and Roberts, 1978); and inhibitors of GABA catabolism. The latter technique involves compounds which inhibit the enzyme that catalyzes the formation of succinic semialdehyde from GABA, GABA transaminase (GABA-T). The most frequently used GABA-T inhibitors in seizure models are ethanolamineO-sulfate (EOS), y-vinyl GABA (GVG), y-acetylenic GABA (GAG), aminooxyacetic acid (AOAA), and valproic acid (Metcalf, 1979; Meldrum, 1978; Loscher, 1980a). Since inhibition of GABA synthesis and GABA receptor blockade result in seizures, these techniques are rarely used in elucidating the role of GABAergic systems in seizure models. u. Generalized Convulsive Seizures. T h e GABA-T inhibitor AOAA has been shown to produce an increase in the threshold for ECS and PTZ seizures (Kleinrok et al., 1980; Kuriyama et al., 1966; Wood and Peesker, 1975; Wallach, 1961; De Vanzo et al., 1961; Loscher and Frey, 1978). This anticonvulsant effect correlates with the increase in GABA (Loscher and Frey, 1978), but some authors (Wood and Peesker, 1975) feel that the explanation for the anticonvulsant action of AOAA is more complex than simply its inhibition of GABA-T. Since AOAA is not a pure GABA-T inhibitor, efforts to develop an irreversible, specific GABA-T inhibitor have led to the compounds y-vinyl GABA (4-amino-hex-5enoic acid), y-acetylenic GABA (4-amino-hex-5-ynoic acid) (Jung et al., 1977a,b), and ethanolamine-o-sulfate (Fowler and John, 1972; Pycock and Horton, 1978), although there is some doubt about the specificity of GVG and GAG (Perry et al., 1979). GAG and GVG are associated with an increased duration of PTZ seizure, and they abolish the postictal state in most models but d o not alter the PTZ threshold (Myslobodsky et al., 1979). However, other workers have shown that these drugs produce an increased threshold to both PTZ and ECS seizure (Loscher, 1980b). There is also some evidence that these drugs may exert anticonvulsant activity independent of GABA-T inhibition (Kendall et al., 1981). Paradoxically, either single-dose (Schecter and Grove, 1979) or chronic (Bachus et al., 1980) GAG administration can result in spontaneous seizures. The issue of relation of brain GABA levels to convulsant or anticonvulsant effects of various drugs has generated some contradictory data. Gale and Iadarola (1980; Iadarola and Gale, 1980) using denervated substantia nigra have shown that the time of maximal anticonvulsant effect of GABA-T inhibitors correlates with elevation of GABA in nerve terminals. Although Gale and Iadarola did not directly measure synaptosomal GABA, others have done so demonstrating the feasibility of the technique and showing that synaptosomal concentrations of rt al.,

132

0. CARTER SNEAD 111

GABA may well be affected significantly by convulsant drugs in the absence of any corresponding change in whole-brain GABA levels (Wood et al., 1979; Kontro et al., 1980; Matsuda et al., 1979). Inhibitors of high-affinity GABA uptake also produce an elevated threshold for ECS and PTZ seizures which parallels their in vitro potency, but this change in threshold is not associated with a change in whole-brain GABA concentration. However, no symptosomal data were obtained (Frey et al., 1979). The GABA agonist muscimol apparently has anticonvulsant activity against PTZ and ECS seizures (Meldrum, 1981; Bernard et al., 1980; Matthews and McCafferty, 1979; Collins, 1980), although there is not universal agreement concerning this issue (Worms et al., 1979). Other GABA agonists effective in raising the seizure threshold in those models of generalized convulsive seizures include cetyl GABA (Frey and Loscher, 1980) and baclofen. The latter has been reported to be both effective (Benedito and Leite, 198 1) and ineffective (Bernard et al., 1980) in the ECS model, but is generally considered ineffective against the PTZ model of generalized convulsive seizures (Benedito and Leite, 1981; Bernard et al., 1980). A single ECS seizure produces no change in brain GABA levels (McCandlesset al., 1979), receptor binding, or GAD activity (Atterwillet al., 1980), but repeated ECS results in decreased GABA turnover with an increased GABA level in nucleus accumbens and caudate 24 hr postictally and increased GAD activity (Atterwill et al., 1980; Green et al., 1978; Gold and Roth, 1979). These data contrast with those of E. S. Essman and Essman (1980) which show a decrease in synaptosomal uptake of GABA with an increase in GABA receptor density after ECS. Pentylenetetrazol has been shown to interfere with GABA-mediated inhibition (Macdonald and Barker, 1977) by reversibly antagonizing GABA-evoked ISPSs in cultured mammalian spinal cord neurons in a dose-dependent fashion, as well as by blocking GABA-induced changes in membrane conductance (Macdonald and Barker, 1978a). In the genetic models of generalized convulsive seizures, the GABA-T inhibitors uniformly protect against audiogenic seizures in mouse, an action associated with a parallel increase in brain GABA (Anlezark et al., 1976; Schechter et al., 1977a,b; Schechter and Grove, 1980; Simler et al., 1973; Horton et al., 1977). Also protective in this model are inhibitors of high-affinity GABA uptake (Horton et al., 1979), as well as the GABA agonists muscimol (Anlezark et al., 1977; Meldrum et al., 1980; Worms et al., 1979; Meldrum, 1981), THIP, and baclofen (Meldrum et d., 1980). In the genetic models of generalized convulsive seizures there is no difference in the high-affinity transport system for

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

133

GABA between the seizure-prone and seizure-resistant strain of mouse (Bondy et al., 1979). The various GABA-T inhibitors have been found effective against photomyoclonic seizures in the baboon (Meldrum and Horton, 1978; Naquet and Meldrum, 1972); however, muscimol exacerbates seizures in this model (Pedley et al., 1979; Meldrum, 1981). In the beagle dog model of generalized convulsive seizures there is no difference in concentration of GABA in seizure versus nonseizure animals (Edmonds et al., 1979). Repeated audiogenic seizures in mice produce a decrease in GABA in basal ganglia, cerebellum, cerebral cortex, hippocampus, and amygdala (Ciesielski et al., 1981). Generalized convulsive seizures associated with hyperbaric oxygen are accompanied by decreased GAD activity with a concomitant drop in brain GABA (J. D. Wood, 1980; Wood and Peesker, 1975). There is disagreement as to whether the drop in GABA is the cause or the effect of seizures in this model since elevation of GABA by the GABA-T inhibitor, gabaculline, does not protect against the seizures (Faiman et al., 1980); however, other GABA-T inhibitors such as AOAA (Wood and Peesker, 1975)and the GABA agonist 3-APS (Fariello and Golden, 1980; Adembri et al., 1974) are effective against hyperbaric oxygen seizures. GABA-T inhibitors and GABA agonists are also effective against alcohol (Cooper et al., 1979) and barbital withdrawal seizures (Benedito and Reite, 1981). b. Generalized Absence. Contrary to generalized convulsive and partial seizures, augmentation of GABAergic transmission appears to exacerbate many models of generalized absence seizures (Myslobodsky, 1976). The GABA agonists muscimol and imidozole acetate produce an increase in the occurrence of FEADs (King, 1979).Similarly, 3-APS has an activating effect on the penicillin model of petit ma1 (Fariello and Golden, 1980; Fariello et al., 1981). Furthermore, baclofen and muscimol produce, potentiate, or induce absence-like seizure activity (Scotti de Carolis and Massotti, 1978; Pedley et al., 1979), and GABA-T inhibitors potentiate PTZ-induced spike-wave seizures (Myslobodsky et al., 1979). y-Hydroxybutyrate (GHB), a metabolite of GABA (Roth and Giarman, 1969; Gold and Roth, 1977) is known to produce petit-mal-like seizures (see models). The mechanism by which this epileptogenic effect is exerted is not known, but GHB has been classified as a GABA agonist (Meldrum et al., 1980; Meldrum, 1981), and so it could conceivably be acting through GABAergic systems. The evidence for a GABAergic effect of GHB is as follows: 1. Some of the pharmacologic effect of GHB can be reversed by picrotoxin (Roth and Nowycky, 1977).

134

0. CARTER SNEAD 111

2. T h e EEG effects of GHB are said to be similar to those of muscimol and baclofen (Scotti d e Carolis and Massotti, 1978; Pedley et d., 1979). 3. The effects of microiontophoretically applied GHB on the neuronal activity of rabbit sensorimotor cortex is similar to that of GABA and is reversed by bicuculline (Kozhechkin, 1980, 1981). On the other hand, GHB has some neurophysiologic actions directly opposite to those of GABA and its agonists (Scholes, 1966; Crawford and Curtis, 1964; Osorio and Davidoff, 1978; Walters et al., 1978); GHB has no affinity for the GABA receptor (Enna and Maggi, 1979; Lloyd and Dreksler, 1979), and the neurophysiologic effects of GHB on dopaminergic neurons are insensitive to GABA antagonists (Olpe and Koella, 1979). In spite of the questions concerning GABA involvement in GHB induced absence seizures, the other evidence enumerated above has led to the hypothesis that petit ma1 seizures may result from hyperfunction or paroxysmal activity of inhibitory (i.e., GABA) pathways (King, 1979; Fromm et al., 1980). c. Gmemlized Mvoclonzc Seizures. T h e GABA-T inhibitors AOAA and valproic acid have effects onp,p '-DDT-induced myoclonic seizures similar to those of picrotoxin in that all these compounds slightly decrease the latency to seizures (Hwang and Van Woert, 1978). Relatively high doses of muscimol consistently produce myoclonic seizures in mice which are blocked with THIP, baclofen, and GHB (Menon, 1981; Menon and Vivonia, 1981). Paradoxically, myoclonus is a prominent feature of GHB-induced seizures (Snead et al., 1976; Snead, 1978a). Tarsy et al., (1978) have demonstrated that focal myoclonus could be produced by striatal injection of GABA antagonists. This myoclonic activity is dependent on the presence of cortical damage. d. Partin1 Seizures. Penicillin has been demonstrated to antagonize GABA-induced depolarization of dorsal root terminals and also the inhibitory action of GABA on cortical and spinal neurons in a fashion similar to that for PTZ (Macdonald and Barker, 1977; Hochner et al., 1976; Davidoff, 1972a; Van Duijn et d.,1973; Curtis et al., 1972; Hill et al., 1973, 1976). y-Aminobutyric acid and GAD activity and GABA uptake and release are all decreased in penicillin epileptic foci (Gottesfeld and Elazar, 1972, 1975; Beart and Johnston, 1973; Cutler and Young, 1979). Penicillin focal seizures are antagonized by 3-APS (Fariello and Golden, 1980), muscimol (Collins, 1980), SL 76002 (Lloyd et al., 1981), and AOAA (Collins and Mehta, 1978). y-Aminobutyric acid and GABA uptake are also reduced in cobalt foci (Balcar et al., 1978; Craig and Hartman, 1973; Emson and Joseph, 1975; Koyama, 1972; Van Gelder

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

135

and Curtois, 1972), and AOAA and 3-APS reduce spike frequency in this model of partial seizures as well (Emson, 1976; Adembri et al., 1974; Fariello and Golden, 1980). GABAergic systems also seem to be profoundly disturbed in the alumina1 gel focus with GABA, GAD activity, receptor binding, and CSF GABA all decreased over the lesioned hemisphere. There is also a diminution in the number of GAD containing neurons over the lesioned side (Ribak et al., 1979). Kainic acid treatment is associated with a decrease in GABA binding (Zaczek et al., 1978) and decreased concentration of GABA in brain (Coyle and Schwartz, 1976; Ando et al., 1979). The seizures produced by kainic acid are antagonized by GABA agonists and GABA-T inhibitors (Stone and Javid, 1980; Liebman et al., 1980; Fuller and Olney, 1979; Lenicque et al., 1979; Fariello and Golden, 1980; Bernard et al., 1980).I n contrast to changes in GABA seen in other models of partial seizures, kindling produces no change in GABA receptor binding (Ashton et al., 1980), high-affinity transport (Bondy et al., 1979), or sensitivity of neurons to GABA (Burchfield et al., 1979). y-Aminobutyric acid transferase inhibitors have been shown to decrease the severity of kindled seizures (LaSalle, 1980; Myslobodsky et al., 1979; Myslobodsky and Vallenstein, 1980), but have little or no protective effect against seizures in the focal freezing model of partial seizures (Hori et al., 1979). There is no change in GABA levels in epileptogenic freeze foci (Lewin, 1972; Berl et al., 1959). 4. Interaction with Anticmvulsant drugs The evidence for the effect of phenytoin on GABA metabolism in brain is contradictory. Some authors (Hitchcock and Gabra-Sanders, 1977; Patsalos and Lascelles, 1981) have observed a decrease in GABA concentration in hypothalamus and cerebellum with no change in the activity of GABA-T (Loscher, 1980b) or GABA release (Abdul-Ghani et ai., 1981), whereas others (Vernadakis and Woodbury, 1960; Saad et al., 1972; Loscher and Frey, 1977) have shown increased concentrations of brain GABA in phenytoin-treated animals. GABA uptake has been reported to be increased by phenytoin (Weinberger et al., 1976), but this varies in convulsed and nonconvulsed animals (W. B. Essman and Essman, 1980). Phenytoin prolongs GABA-mediated inhibition (Ayala et al., 1977; Deisz and Lux, 1977), enhances presynaptic inhibition in amphibian spinal cord, and prolongs recurrent inhibition in pyramidal tract cells (Davidoff, 1972b; Raabe and Ayala, 1976). Furthermore, phenytoin is effective against seizures induced by GAD inhibition (Ashton and Wauqujer, 1979; Loscher, 1979). T h e interaction of barbiturates with GABAergic systems has, until recently, been a confusing issue because of discrepant data concerning

136

0.CARTER SNEAD 111

anticonvulsant and anesthetic barbiturates (Macdonald and Barker, 1979b). Much of the earlier work was directed at the anesthetic barbiturate, pentobarbital (Ransom and Barker, 1975, 1976; Nicoll, 1975; Barker and Ransom, 1978a), rather than the anticonvulsant, phenobarbital (Gallagher et al., 1981). It has been postulated (Evans, 1979; Macdonald and Barker, 1978b) that phenobarbital acts primarily at the bicuculline-sensitive GABA receptor, whereas pentobarbital acts at both bicuculline-sensitive and -insensitive sites. Recent work (Macdonald and Barker, 1978b, 1979a,b; Schulz and MacDonald, 1981) indicates most of the anticonvulsant effects of barbiturates are associated with GABA receptor responses, while anesthetic actions are seen with an increase in chloride conductance. A model of the GABA-benzodiazepine receptor-chloride ionophore complex is shown in Fig. 2. Pentobarbital and phenobarbital at anesthetic doses apparently act on chloride conductance, while lower, anticonvulsant doses of phenobarbital augment the response at the GABA receptor. Phenobarbital has no effect on GABA release (Abdul-Ghani et al., 1981)or GABA-T activity (Loscher, 1980b); however, it does produce an increase in GABA uptake (W. B. Essman and Essman, 1980) and an increase in brain GABA (Patsalos and Lascelles, 1981; Saad et al., 1972; Ho, 1980) although there are conflicting data concerning the last point (Ferrari and Arnold, 1961; Crossland and Turnbull, 1972; Tsuji et al., 1963).Phenobarbital is effective against seizures produced by picrotoxin (Horton et d.,1976) and GAD inhibitors (Ashton and Wauquier, 1979; Loscher, 1979). The benzodiazepine (BDZ) anticonvulsants are especially interesting because of their close relationship to GABA function (Fig. 4). Over the past few years there has been mounting evidence for an intimate interaction of GABA and the benzodiazepines at a receptor protein complex containing receptor sites for GABA, benzodiazepine, picrotoxinin (barbiturates?), and the chloride ionophore (Fig. 4) (Haefely et al., 1975; Costa et al., 1975; Waddington, 1978; Montarolo et al., 1979; Squires and Braestrup, 1977; Mohler and Okada, 1977; Braestrup et al., 1977; Olsen, 1981; Leeb-Lundberg et al., 1981; Tallman et al., 1980). y-Aminobutyric acid and GABA-mimetic drugs potentiate the binding of benzodiazepines to the receptor complex (Martin and Candy, 1978; Williams and Risely, 1979). y-Aminobutyric acid analogs enhance BDZ binding proportionately to their potency of binding at the GAB 4 receptor (Braestrup et al., 1979; Greenlee et al., 1978; Krogsgaard-Larsen et al., 1979).The interaction of benzodiazepines on GABA binding is not as well established (Olsen, 1981; Costa et al., 1978) with a number of authors (Mohler and Okada, 1978; Olsen et al., 1978; Andrews and Johnston, 1979) finding that BDZ produces no change in GABA bind-

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

137

a

? a W 0

W

I1I

W

za

W N

5

0

:z: CAGE CONVULSANTS

W

m

ANXIOLYTIC PYRAZOLO-PYRIDINE CONVULSANT

BEN 20 DIA 2 E P INE

FIG. 4. Theoretical model of the GABA receptor-ionophore complex, including receptors for convulsant and anticonvulsant drugs. The complex contains three drug binding sites, the GABA receptor, the picrotoxinin-barbiturate receptor, and the benzodiazepine receptor, as well as the chloride ion channel. This ion channel may be a separate component or part of one of the drug-binding proteins. GABA function (opening of the chloride channel) may be modulated by drugs binding to the other two receptor sites, either endogenous ligands or exogenous substances. These drugs can be either positive (depressants) or negative (excitants) modulators of GABA chloride ionophores. [Reproduced from Olsen and Leeb-Lundberg (1981) with permission.]

ing, whereas one group (Guidotti et al., 1978) showed enhancement of GABA binding by BDZ. I n addition to the binding data, there are a number of neurophysiologic lines of evidence that BDZ may augment GABAergic transmission (Schlosser and Franco, 1979a): The BDZs enhance presynaptic inhibition in spinal cord (Polc et al., 1974; Schlosser, 1971), reduce the firing rate of cerebellar Perkinje cells (Curtis et al., 1976a; Pieri and Haefely, 1976), reverse the effects of GABA antagonists (Curtis et al., 1976b), and potentiate GABA-mediated inhibition (Nistri and Constanti, 1978; Schlosser and Franco, 1979b). The pharmacologic spectrum of the benzodiazepines is broad, varying from antianxiety to anticonvulsant to convulsant (Schlosser and Franco, 1979a). There are recent data that relate BDZ anticonvulsant activity (Paul et al., 1979), as well as that of barbiturates (Leeb-Lundberget al., 1981; Ticku, 1981), to BDZ receptor binding, with at least one group (Lippa et al., 1980) advocating a specific BDZ subgroup of receptors with anticonvulsant properties. There is some evidence (Syapin and Rickman, 1981) that PTZ may be a BDZ receptor antagonist.

138

0. CARTER SNEAD 111

Currently, no endogenous ligand has been firmly established for the BDZ receptor, although a naturally occurring protein which inhibits BDZ binding has been proposed (Guidotti et al., 1978; Toffano et al., 1978) as a possibility. Other candidates for this ligand include inosine, hypoxanthine, nicotinamide (OBrien et al., 198l), and P-carbolines. T h e latter group of compounds occur naturally and antagonize BDZ binding (O'Brien et al., 1981; Muller et al., 1981; Barker et al., 1981; Buckholtz, 1980; Braestrup et al., 1980; Nielsen et al., 1979; Turner and Hirsch, 1980; Rommelspacher et al., 1980). T h e carbolines are of particular interest in the context of seizures because they lower the seizure threshold to PTZ and antagonize the anticonvulsant action of diazepam (Oakley and Jones, 1980; Skolnick et al., 1981; Cowen et al., 1981). Valproic acid is another anticonvulsant that may have GABAergic activity. This compound was initially thought to act primarily as a GABA-T inhibitor with increased GABA being responsible for its anticonvulsant action (Simler et al., 1973; Horton et al., 1976; Bruni and Wilder, 1979); however, the biochemical effect of valproic acid is much more complex than this (Kerwin and Taberner, 1981). Valproic acid is a weak competitive inhibitor of GABA-T and succinic semialdehyde dehydrogenase, but produces a stronger, noncompetitive inhibition of NADPH-dependent aldehyde reductase, the enzyme responsible for the formation of GHB in brain (Whittle and Turner, 1978, 1981). Valproic acid also potentiates GABA-mediated postsynaptic inhibitor responses (Macdonald and Bergey, 1979; Kerwin and Olpe, 1980; Hayashi and Negishi, 1979). Ethosuximide and trimethadione have no effect on brain GABA levels (Nahorski, 1972; Ferrari and Arnold, 1961), although these antipetit-ma1 drugs may have a slight inhibitory effect on GABA-T (AbdulGhani et 01.. 1981). Ethosuximide is not effective against seizures produced by GAD inhibitors, but trimethadione is (Ashton and Wauquier, 1979). 5 . Ch u r n 1 Co?-relateb

T h e first clinical evidence for involvement of GABAergic systems in seizures came with the demonstration that severe dietary deficiency of B, vitamins results in seizures which can be controlled by B6 supplements (Snyderman et al., 1953; Scriver, 1976; Hunt et al., 1954; Coursin, 1954, 1955). It has been suggested (Tower, 1956, 1976) that the explanation €or these seizures in B6 deficiency relates to the function of vitamin B6 in the form of pyridoxal 5'-phosphate as a coenzyme for GAD. Lott et nl. (1978) have demonstrated decreased GABA content in frontal and occipital cortex with normal GAD activity in a patient who had pyridoxine-

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

139

dependent seizures. Unfortunately, the GABA and GAD content of deep structures was not determined and four of five patients used for control values of GABA and GAD had severe primary central nervous system disease. I n view of the fact that there are a number of other enzyme systems in brain in addition to GAD which utilize pyridoxal 5'-phosphate as a coenzyme, the conclusion that GAD alone is involved in Be-deficient and -dependent seizures may be erroneous. Although there are conflicting data over GABA concentration in human epileptic foci (Van Gelder et al., 1972; Perry and Hansen, 1981; Perry et al., 1975), there are recent data suggesting decreased GAD activity in human seizure foci, particularly those secondary to tumor. Also, there is a decreased number of GABA binding sites in these seizure foci with no change in GABA-T activity (Lloyd et al., 1981). The latter information is more reliable than GABA levels, since GABA changes drastically in tissue, either from biopsy or postmortem, very quickly after tissue removal (Love11et al., 1963; Alderman and Schellenberger, 1974). Since GABA in CSF is also quite unstable (Grossman et al., 1980), any CSF GABA studies in epilepsy must be well controlled to be valid (J. H. Wood, 1980). The CSF data available in seizure patients suggest that CSF GABA is significantly decreased in these patients (Enna et al., 1977; Wood et al., 1979; Wood and Brooks, 1980). A variety of therapeutic modalities addressed to correcting a presumed GABA deficiency in seizure disorders have been attempted. An early report of successful treatment of seizures with GABA (Tower, 1960) has not been duplicated. Cerebellar stimulation has been used in seizure disorders (Cooper, 1973, I. S. Cooper et al., 1973, 1976, 1978; Grabow et al., 1974; Lockard et al., 1979) based on the following rationale: (1) Cerebellar mechanisms may be involved in experimental seizures Uulien and Laxer, 1974; Dow et al., 1962; Reiner et al., 1967); ( 2 ) some anticonvulsants produce an increase in firing rate of cerebellar Purkinje cells (Halpern and Julien, 1972a,b; Puro and Woodward, 1973); and 3) cerebellar stimulation would be expected to cause GABA to be released with increased Purkinje cell firing (Obata et al., 1967; Obata and Takeda, 1969). However, recent evidence has disproved the latter thesis. Transfolial cerebellar stimulation, in fact, is associated with suppression of cerebellar Purkinje cell activity (Dauth et al., 1978) and a decrease in CSF GABA concentration associated with no change or deterioration in seizure control (Wood et al., 1977a; Van Buren et al., 1978; Strain et al., 1979). To date, the main pharmacologic GABAergic agent used in seizures is valproic acid, although there is a question, as discussed above, concerning its actual mode of action. The GABA agonist muscimol has proved

140

0. CARTER SNEAD 111

too toxic for clinical use (Chase and Tamminga, 1979); however, SL 76002 has shown promise as an anticonvulsant in a preliminary clinical trial (Lloyd et al., 1981; Lloyd and Worms, 1981; Bartholiniet al., 1979).

E. OTHERINHIBITORY AMINOACIDS 1. Glycine Although glycine is known to be a major inhibitory neurotransmitter in spinal cord (Aprison and Nadi, 1978; Pycock and Kerwin, 1981), it is also found in brain where it may also serve as a neurotransmitter, particularly in the substantia nigra (Shank and Aprison, 1970; Pycock and Kerwin, 1981; Pycock et al., 1981; Palacios et al., 1981). Glycine appears to be inhibitory on cortical neurons as it is on spinal neurons (Curtis et al., 1968; Johnson at al., 1979; Johnson et al., 1970; Marciani et al., 1980). The specific glycinergic antagonist, strychnine, produces severe generalized convulsive seizures by virtue of its blockade of glycine receptors throughout the neuraxis (Bradley et al., 1953; Curtis, 1969; Curtis et al., 1967, 1971a; Esplin and Woodbury, 1961; Krall, 1980). In spite of its inhibitory actions, there are few clinical or basic experimental data dealing with glycine in the epilepsies. Glycine has anticonvulsant activity in the audiogenic seizure model in mouse comparable to that of GABA (Laird and Huxtable, 1978). Strychnine has been used clinically in children with an inborn error of glycine metabolism, nonketotic hyperglycinemia, in an attempt to treat the seizures and psychomotor retardation but with varying success (Van Wendt et al., 1980; Arneson et al., 1979). Glycine has been shown to be markedly elevated in brain epileptogenic foci in some seizure patients (Perry and Hansen, 1981).

2. Taurine Taurine (2-aminoethane sulfonic acid) is found in large amounts in the CNS in a subcellular localization, has inhibitory properties electrophysiologically, and has been proposed as an inhibitory neurotransmitter (Guidotti, 1978; Bonaventure et al., 1974; Phillis, 1978; Rassin, 1981). Taurine has been reported to be decreased in cobalt (Van Gelder, 1972; Joseph and Emson, 1976; Emson and Joseph, 1975), penicillin (Mutani et al., 1978), and human epileptic foci (Van Gelder et al., 1972). Conversely, administration of taurine is reported to protect against seizures in a variety of models of partial seizures and also audiogenic seizures (Izumi et al., 1973, 1974; Mutani et al., 1974a,b, 1978;

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

141

Laird and Huxtable, 1978). The results of taurine in clinical trials have been equivocal showing a possible beneficial effect of large doses in partial-seizure disorders but with development of tolerance (Van Gelder, 1975, 1978a; Van Gelder et al., 1972; Mutani et al., 1978; Takahashi and Nakone, 1978). Whether taurine exerts its anticonvulsant effect by a direct inhibitory action or by altering the concentration of the amino acids GABA and/or glutamic acid (Van Gelder, 1978a,b) is unknown.

F. EXCITATORY AMINOACIDS

The two major excitatory amino acids in brain are L-glutamic and L-aspartic acid (Johnston, 1978, 1979; Johnson, 1972; Curtis et al., 1960). Glutamic acid is the most abundant amino acid in the CNS, with the highest concentration in nucleus accumbens, cerebellum, hippocampus, and cortex (Nitsch, 1981). There is some evidence that both aspartate (Nitsch, 1981; Nitsch et al., 1979; Nadler et al., 1976; Curtis, 1979) and glutamate (Costa et al., 1979; Shank and Aprison, 1979; Kuhar and Snyder, 1970; Logan and Snyder, 1972; Nitsch, 1981; Storm-Mathisen and Opsahl, 1978) are excitatory neurotransmitters. There appear to be receptors for L-glutamate (Johnston, 1979; De Robertis and Fiszer de Plazas, 1976; P. J. Roberts, 1974; Michelis et al., 1974), L-aspartate (Fiszer de Plazas and De Robertis, 1976), kainate, and N-methy1-D-aspartate (Evans and Watkins, 1981; McLennon et al., 1981; Roberts, 1981) in brain which could mediate such an excitatory action. The ubiquitous distribution and diverse functions of glutamic acid, for example, incorporation into protein, peptides, fatty acid, and GABA synthesis, and regulation of ammonia formation and excretion, make it difficult to define a precise role for this excitatory compound in the pathogenesis of seizures. Introcerebroventricular glutamate is known to be convulsant (Delgado et al., 1971), and the elevation of glutamic acid produced by GAD inhibitors may play a role in the seizures caused by these compounds (see above). The level of glutamic acid correlates with epileptogenicity of the cobalt and freeze foci, although little or no change in glutamate uptake occurs in those models nor is there a change in glutamate concentration in the secondary foci (Pumain and Chauvet, 1975; Pumain et al., 1978; Berl et al., 1959; Dodd and Bradford, 1976; Emson and Joseph, 1975; Koyama, 1972; Coutinho-Netto et al., 1981). There are conflicting data on glutamate levels in human epileptic foci (Emson, 1978; Perry et al., 1975; Perry and Hansen, 198 1; Van Gelder et al., 1972).

142

0. CARTER SNEAD I l l

T h e excitatory effects of glutamate and aspartate are antagonized by phenytoin, benzodiazepines (Stone, 198l), and phenobarbital (Richards, 1976). In summary, L-glutamate could be involved in the pathogenesis of seizures in one or more of three ways: 1. T h e ratio of glutamate to GABA could be critical with the flux from glutamic acid to GABA determining the excitatory or inhibitory tone of the system (Emson, 1978; Van Gelder, 1981). 2. Glutamic acid is clearly excitatory, and in high enough concentrations could be epileptogenic. 3. The role of glutamate in ammonia elimination (Shank and Aprison, 1979; Meister, 1979) could be crucial since elevated ammonia at the cellular level can result in seizure activity (Iles and Jack, 1980; Roberts, 1980). G. CI-CLIC NUCLEOTIDES Cyclic nucleotides are formed by the action of adenylate and guanylate cyclase, which convert ATP to cyclic adenosine 3' ,5'monophosphate (cyclic AMP) and GTP to cyclic guanosine 3',5'monophosphate (cyclic GMP). Since Sutherland et al. showed that these compounds act as an intracellular mediator for the regulatory actions of hormones on non-neuronal tissue (see Robison et nl., 1971), there has been a great deal of work concerning the role of these compunds in CNS pharmacology and physiology. Cyclic AMP systems are associated with postsynaptic neuronal sites and appear to be involved in inhibitory neurotransmission for noradrenergic, dopaminergic, and serotinergic pathways. Cyclic GMP systems, perhaps less well understood, are also postsynaptic, possibly presynaptic, and are involved in responses to the excitatory neurotransmitters, glutamate and ACH (Daly, 1975, 1977; Greengard, 1976). The first evidence for involvement of cyclic nucleotides in seizures came in 197 1 when Sattin reported increased levels of cyclic AMP associated with ECS seizures. This finding was confirmed and extended to PTZ-induced seizures by others (Ma0 at al., 1974a; Folbergrova, 1975; Berti et al., 1976; Lust et al., 1976) who showed an elevation in cyclic GMP as well. The increase in cyclic GMP precedes the onset of the seizure, whereas cyclic AMP increased only after onset of the paroxysmal discharge (Ferrendelli and Kinscherf, 1977; Lust et al., 1981). Those drugs which are effective against these seizures prevent the changes in

SACRED

DISEASE: NEUROCHEMISTRY

OF EPILEPSY

143

cyclic nucleotides as well, possibly by suppressing the seizure per se (Palmer et al., 1979; Lust et al., 1981; Ferrendelli and Kinscherf, 1977). Pentylenetetrazol seizures are exacerbated by pharmacologic manipulation which blocks the elevated cyclic AMP; however, antagonism of the rise in cyclic GMP has no effect on seizure threshold. This suggests that the cyclic GMP may be convulsant and the cyclic AMP anticonvulsant (Gross and Ferrendelli, 1979). Additional evidence to support this thesis is that cyclic GMP has the same effect as cholinomimetics in initiating seizure and hypersynchronous discharge in the hippocampal explant model of partial seizures (Freedman et al., 1979; Hoffer et al., 1977a,b). Furthermore, there is an inverse relationship between GAB A and cyclic GMP in cerebellum following anticonvulsant drug administration (Ma0 et al., 1974a,b, 1975). The data concerning cyclic nucleotide concentrations in the CSF of seizure patients is contradictory (Myllya et al., 1975; Brooks et al., 1976; Tsang et al., 1976; Wood and Brooks, 1980), but anticonvulsant treatment does apparently alter cyclic nucleotide concentration in CSF (Wood and Brooks, 1980).

H. PEPTIDES 1. Opiate Peptides The first peptides reported to be produced and released by mammalian neurons were oxytocin and vasopressin. Subsequently,more than 20 peptides have been identified as neurotransmitter candidates in brain. Many of these peptides are located in both brain and gut (Ambinder and Schuster, 1979; Gershon and Erde, 1981). Most prominent among the brain-gut peptides are the pentapeptides, leucine and methionine enkephalin, which are endogenous ligands in brain for the opiate receptor and which may play a natural role in analgesia and nociception (Beaumount and Hughes, 1979). The enkephalins and &endorphin, another naturally occurring opiate peptide in brain, are also potent epileptogenic compounds. These compounds are predominantly inhibitors in their effects neurophysiologically (North, 1979; Carette, 1981). Leucine and methionine enkephalin have been demonstrated to produce paroxysmal spiking in the EEG of rats when given intraventricularly (Urca et al., 1977, Frenket al., 1978a)in doses from 25-100 p g (Fig. 5). This epileptiform activity is associated with immobility and myoclonic jerks and is aborted by naloxone, a specific opiate antagonist. The electrical spiking begins subcortically and can be produced by injection of enkephalin into the forebrain dorsomedial nucleus of thalamus but not

FIG.5. EEG changes produced by 1OOpg leucine enkephalin. Inserts show a faster time trace. RF and RP are right frontal and parietal; LF and LP, left frontal and parietal leads. Animal was immobile during this paroxysmal electrical activity. T h e first change in the EEG occurred within 1 min of ICV administration and was a proxysm of high voltage spikes at a frequency of 7-9 cycles per second (cps) lasting 30-40 sec. This tapered off to 2-3 cps slow wave activity which was followed by 20-25 sec of low-voltage fast activity. A few seconds of 1 cps high-voltage slow wave activity then built u p to 25-30 sec of a second paroxysm of spike and slow wave activity followed by 15-20 sec of low-voltage fast activity. Finally, there was a prolonged period of high-voltage single spikes occurring at a rate of 1 paroxysm per 5 sec. [Reproduced from Snead and Bearden (1980a) with permission.]

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

145

in the periaqueductal gray. The site at which the enkephalins exert their epileptogenic action thus differs pharmacologically as well as anatomically from that which is responsible for their analgesic effects (Frenk et al., 1978b). The EEG and behavioral abnormalities produced by enkephalins are not overcome by standard anticonvulsants,but are aborted by anti-petit-ma1 drugs (Snead and Bearden, 1980a). Higher doses of naloxone are required to abort this seizure activity than the analgesia produced by the enkephalins. /?-Endorphin (BE) is a 3 1 amino acid peptide which is derived from a larger peptide called /?-lipotropin (P-LPH). /?-Endorphin is found in very high concentrations in the thalamus, dorsal and ventral preoptic nuclei, hypothalamus, and pituitary (Bloom et al., 1978) and has a high affinity for the opiate receptor. This compound has analgesic and epileptogenic properties similar to those outlined for the enkephalins, but BE is a much more potent compound in its epileptic actions. The seizures induced by ICV administration may represent partial complex seizures (Henriksen et al., 1978), but the published EEG response of rat to endorphin differs from that observed for leucine enkephalin (Snead and Bearden, 1981). The relatively high dose of naloxone required to block the epileptic effect of leucine enkephalin and BE suggests that this property of the enkephalins and endorphins differs in its site of action from the analgesia produced by these compounds. This raises the basic question as to whether there might be a specific group of opiate receptors responsible for the seizure activity produced by the opiate peptides. Pharmacologic evidence for disparate sites of action of the opiates has been developed (Martinet al., 1976; Gilbert and Martin, 1976)which suggests the presence of at least three types of opiate receptors in brain. These receptors have been named for the drugs that gave rise to the distinction: p (morphine), K (ethylketocyclazocine)(EKZ),and u (SKF 10,047).There is some biochemical evidence for such opiate receptor heterogeneity (Lord et al., 19’77;Kosterlitz and Lelie, 1978; Wuster et al., 1978, 1981; Kosterlitz et al., 1980; Rothman and Westfall, 1981; Zhang and Pasternak, 1981),although the presence of K receptors in rat brain has recently been questioned (Hiller and Simon, 1979). However, another opioid peptide, dynorphin, has recently been proposed as the endogenous ligand for the K receptor (Chavkin et al., 1982; Quiron and Pert, 1981). The p receptor is thought to be responsible for the analgesic actions of the opiates and requires the lowest doses of naloxone for reversal, hence the dose-response curve of naloxone against leucine enkephalin-induced seizures would indicate that some receptor other than the p receptor is responsible for the epileptogenic action of this substance. However, re-

146

0. CARTER SNEAD I11

cent evidence has accumulated to suggest the presence of at least two populations of p receptors, one inhibitory and one excitatory (Pasternak et al., 1980;Jacquet, 1980). There also appears to be a group of receptors selective for P-endorphin, the c receptors (Wiister et al., 1981), which are found in highest concentration in striatum and thalamus and lowest concentration in brainstem (Law et aZ., 1979). T h e evidence to date thus suggests six possible groups of opiate receptors. These receptors and their presumed ligands are listed in Table VII. A detailed summary of the evidence supporting multiple opiate receptors can be found in the paper of Chang et nl. (1979). We have recently demonstrated that each opiate receptor agonist produces a distinct receptor-specific seizure with leucine enkephalin inducing the most consistent, stereotyped electrical paroxysm with absence behavior (Snead and Bearden 1981). Morphine produces both absence and generalized seizures, whereas EKZ, SKF 10,047, and P-endorphin all produce distinct hippocampal and amygdaloid spiking. Each opiate agonist-induced seizure type has a different anticonvulsant profile and a different naloxone dose-response curve suggesting that opiatergic systems in brain (Herkenham and Pert, 1980; Goodman et al., 1980; Gall et ul., 1981) may play a role in a variety of seizure types. Naloxone has been shown to be effective in the GHB model of petit ma1 (Snead and Bearden, 1980b), but preliminary data d o not support the efficacy of this compound in other types of clinical or experimental seizures (Schreiber, 1979; Hardy P t al., 1980; Frenk et al., 1979; Montplaisir ~t af., 198 1); however, this may have to d o with dosages used since much higher doses of naloxone are required to overcome experimental seizures than the analgesic effects of the opiates and the type of seizure disorder in which it was tried. TABLE VII OPIATE RECEPTORS A N D LIGANDS Receptors

Ligands

p-Excitatory

Morphine

p -I nhibi tory

Morphine P-Endorphin n-Ala*- and Leus-enkephalin, Leus- and Mets-enkephalin SKF 10,047 Ethylketocyclazocine

c

8 U K

EEG effect of ligand Seizure: absence and generalized convulsive Seizure Seizure partial complex Seizure generalized absence

Seizure partial complex

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

147

2. Other Brain-Gut Peptides In addition to the enkephalins, there are a number of other peptides common to gut and brain (Pearse, 1978). Gastrin or gastrin-like immunoreactivity has been found in human and animal brain with a predominance in cerebral cortex. The concentration of this substance in cerebral gray matter was 200 nglgm dry weight, which is quite high (Vanderhaegen et al., 1975). There are no published data on the EEG effects of gastrin and its function on brain is unknown. Vasoactive intestinal peptides (VIP) and cholecystokinin (CCK) peptides have been found in the brain of all mammalian species including humans (Dockray, 1976, 1980; Dockray et al., 1978; Rehfeld, 1975; Emson et al., 1980a; Fuxe et al., 1977). CCK-8 occurs in the cerebral cortex in concentrations of 1 nmoYgm tissue making it the most abundant of all the gut-brain peptides found in brain (Rehfeld, 1978a). T h e octapeptide CCK-8 has also been found to occur in subcortical brain regions (Goltermann et al., 1980) and has been demonstrated to have a vesicular localization and calcium-dependent release in rat brain (Emson et al., 1980b). Cholecystokinin receptors in brain have also been identified (Saito et al., 1980). T h e function of CCK-8 in brain is unknown, although an interaction with the opiate peptides and possible role in analgesia have been suggested (Zetler, 1979, 1980a). There have also been recently published data to support a role of this compound in satiety (Della-Fera and Bailey, 1979; Madison, 1977; Morley, 1980). There are some electrophysiologic experiments that suggest that the carboxy terminal sequence of CCK is the effective part of the molecule, since CCK-4 is as potent as CCK-8 in exciting hippocampal neurons (Emson et al., 1980a). Interestingly, the C-terminal sequence of gastrin is identical to CCK-4. T h e amphibian skin decapeptide, caerulein, and the chemically related C-terminal octapeptide of cholecystokinin (CCK-8), after subcutaneous administration to mice, both delay the onset and retard the development of toxic effects of convulsants such as strychnine, PTZ, bicuculline, and picrotoxin. They also increase the seizure threshold doses of intravenously infused PTZ and picrotoxin. I n this regard, both peptides are at least equipotent with diazepam (Zetler, 1980b). Vasoactive intestinal peptide is found in highest concentration in cerebral cortex, hypothalamic nuclei, amygdala, hippocampus, and striatum (Said et al., 1980). Like CCK, this peptide has been localized to synaptic vesicles (Emson et al., 1978), is released by physiologic stimuli (Giachetti et al., 1977), and is often mentioned as a candidate for neurotransmitter status. There are no structure-activity data on this peptide. It has an excitatory effect on cortical neurons, but the data regarding EEG effects of this

148

0. CARTER SNEAD 111

compound are sparse, referring only to ‘‘arousal’’in dogs receiving the peptide intra-arterially (Said et al., 1980). 3. Adrenocorticotropic Hormone

Human adrenocorticotropic hormone (ACTH) is a 39 amino acid polypeptide (Bennett, 1979; Donald, 1980) whose function in regulation of the adrenal gland was first elucidated in 1916 by Smith. Over the past 15 years there have been a number of advances in the biochemistry and endocrinology of ACTH. The advances in these two areas have been accelerated within the past 5 years with the explosion of interest and research related to neuropeptides which crystallized with the demonstration of the presence of opiate receptor and its endogenous ligands in brain (Way, 1979). The surge of interest in ACTH among neuroscientists has resulted both from the demonstration of its presence in brain outside the pituitary (Krieger and Liotta, 1979) and from the fact that there is now evidence that ACTH may exert effects in brain independent of its putative endocrine function. Adrenocorticotropic hormone and P-lipotropin (P-LPH) both derive from a common precursor molecule with a molecular weight of 30,000, proopiocortin (Rubenstein et al., 1977). Both P-LPH and ACTH may be enzymatically cleaved into smaller, biologically active peptides: &endorphin from P-LPH, and melanocyte-stimulating hormone (MSH) from ACTH (Gramsch et al., 1980). The cell bodies containing these peptides are within the area of the arcuate neurons of the hypothalamus with projections to many brain areas including the limbic system, thalamus, and brainstem (Watson et al., 1978; Krieger and Liotta, 1979; Pelletier et al., 1980). There is also evidence that @endorphin and ACTH are secreted concomitantly by the pituitary (Guillemin et al., 1979; Hollt et al., 1978). Further evidence for an intimate interrelationship between ACTH and opiate peptides in their diverse effects on the CNS concerns the affinity of ACTH and its fragments for the opiate receptor (Terenius, 1975; Terenius et al., 1975) and the involvement of ACTH in stress analgesia (Lewis et al., 1980). Much of the evidence concerning an action of ACTH on brain independent of its endocrine effect centers about structure-activity relationships of various ACTH peptide fragments as regards their ability to stimulate the adrenals versus their ability to produce changes in behavior. The observation that ACTH could influence behavior was first made in 1953 by Mirsky et al. These behavioral changes have since been intensively studied and extensively reviewed (Beckwith and Sandman, 1978; de Weid and Gispen, 1979; Bolus, 1979; de Weid, 1977a,b) and will be

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

149

mentioned only briefly here. Adrenocorticotropic hormone has been found to have an effect on acquisition and extinction of a conditioned avoidance response, facilitate reversal learning, increase resistance to a complex brightness discrimination test, reverse COz retrograde amnesia, and induce excessive grooming followed by stretching and yawning movements. A number of ACTH analogs have been synthesized and tested for behavioral and adrenal activity (de Weid et al., 1975; Wiegant et al., 1977). The peptide fragments of ACTH4-lo and ACTH1-lo are devoid of the corticotropic activity of ACTH1+, or ACTHl-z4, yet are behaviorally active. Microinjections of ACTHl-lo and ACTH4-lo into the posterior thalamus produce behavioral effects, whereas systemic administration produces no change (de Weid and Gispen, 1979). The tetrapeptide ACTHI-, contains the essential elements required for the behavioral effect of ACTH analogs. There is now substantial electrophysiologic evidence supporting an effect of ACTH on electrical activity of brain. Adrenocorticotropic hormone has been demonstrated to activate hypothalamic and midbrain cells (Steineretal., 1969; Van Delft and Kitay, 1972) and also to enhance spinal cord neuron excitability (Nicolou, 1967). Urban et al. (1974) have demonstrated that the fragment ACTH4-lo, devoid of corticotrophic activity, has an activating effect on hippocampal theta rhythm in dogs and also in rats (Urban and de Weid, 1976). Adrenocorticotropic hormone fragments have also been given to normal human subjects with EEG monitoring. These fragments were demonstrated to suppress stimulus-induced EEG synchrony with the ACTH4-lo fragment being the most potent (Endroczi et al., 1970). The observation that ACTH and cortisone had an effect upon the EEG in humans was first made in the 1940s (Thorn et al., 1949; Hoefer and Glaser, 1950; McQuarrie et aE., 1942). In 1950, Klein and Livingston reported dramatic improvement in the EEG and clinical status of six seizure patients treated with ACTH. The anticonvulsant efficacy of ACTH and adrenal cortical steroids is paradoxical in view of the seizures that are occasionally precipitated by these compounds when they are used for other clinical disorders (Glaser, 1953; Wayne, 1954; Woodbury and Vernadakis, 1966; Ehlers and Killam, 1979). In 1958, Sore1 and Dusaucy-Bauloye described remarkable clinical improvement and disappearance of the gross EEG abnormality in children with infantile spasms treated with ACTH. The syndrome of infantile spasms is a myoclonic seizure disorder associated with a grossly abnormal EEG (hypsarrhythmia) which occurs only in very young children and which has an inordinately high morbidity and mortality (Lacy and Penry, 1976;

150

0. CARTER SNEAD I11

Jeavons and Bower, 1964; Jeavons et a/., 1973). T h e beneficial effect of ACTH in this disease was subsequently confirmed by many authors (Stamps et a/., 1959; Harris, 1964; Pollack et al., 1979). Prednisone too has been shown effective in infantile spasms (Hrachovy et al., 1979). Although there are no controlled studies comparing the efficacy of prednisone to that of ACTH in infantile spasms, the literature cited above suggests that ACTH is more efficacious. Although the mode of action of ACTH in its anticonvulsant effect against seizures is unknown, there are experimental (Torda and Wolff, 1952) and clinical (Willig et a/., 1979; Crosley et a/., 1980) data that indicate this is an extra adrenal action of ACTH; however, one published study using ACTH,-lo showed no effect of this fragment on the EEG of patients with seizures (Willig and Lagenstein, 1980). An additional clinical observation which is relevant to our proposal is that some authors (Low, 1958; Millichap and Bickford, 1962) have noted an inverse relation between response of children with seizures to ACTH and the age of the patient. This along with the age specificity of infantile spasms should be borne in mind when considering the human and animal experimental data concerning basic neurochemical and neuroph ysiologic effects of ACTH outlined above, since all those data relate to adults.

Vi. Development01 Considerations

Since seizures are more prevalent in children than adults, the neurochemistry and physiology of the developing brain is apropos to a discussion of the neurochemistry of epilepsy. The developing brain is probably a beter experimental model of clinical childhood seizure states because of dynamic neurochemical and neurophysiologic processes which may contribute to its selective vulnerability to one or another convulsant stimuli. There is a large volume of literature that deals with the ontogeny of neurotransmission in the developing nervous system (Coyle, 1977). Modulating mechanisms and feedback regulations of neuronal functions develop from 4 to 10 days postnatally in the rat with respect to dopaminergic and cholinergic systems in striatum (Kellogg and Wennerstrom, 1974; Keller et al., 1973; Fibiger et al., 1970; Coyle and Henry, 1976; Baez et nl., 1976). y-Aminobutyric acid levels in rat at day 15 of gestation are 19% of the adult level and increase to 60% of adult concentration by birth. Between birth and 1 week postnatally, GABA concentrations are constant followed by a linear increase to adult

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

151

levels by 28 days. In contrast, GAD, although present at 15 days gestation, is only 12% of adult activity at birth and reaches adult levels of activity at 28 days of age. The development of GABA uptake activity is biphasic, peaking between 7 and 10 days of age and dropping two- to three-fold adult levels (Coyle and Enna, 1976), but the neuronal calcium-dependent release system for GABA develops in a progressive fashion similar to that observed for the biogenic amines. This suggests that the GABA stored from enhanced GABA uptake observed in the younger animals is not stored in a functionally releasable pool (Redburn et al., 1978). The absolute amounts of both endorphin and enkephalin, as well as tissue protein, increase with age in all regions studied but the concentrations, expressed on a protein basis, reveal interesting differences. Between embryonic day 20 and postnatal day 6, the concentration of endorphin decreases in all regions with the greatest decrease (about 50%) occurring in the corpus striatum. This continues decreasing to very low levels at adulthood. The corpus striatum shows the highest endorphin concentration in the brain before birth, whereas the hypothalamus is the richest in the adult. The concentration of enkephalin does not significantly change from embryonic day 20 to postnatal day 6 in any of the regions studied; enkephalin concentration remained almost constant after birth except for a marked increase (about threefold) in the region containing the preoptic areas and the septum. The corpus striatum contained the highest concentration of enkephalin in both the embryonic and adult rat. In contrast to the .brain, the pituitary concentrations of both endorphin and enkephalin are constant from embryonic day 20 to postnatal day 6; both peptides subsequently increased severalfold by adulthood (Bayon et al., 1979). Neurophysiologically, there appears to be a differential rate of development of excitatory versus inhibitory neuronal processes in brain with earlier appearance of inhibitory synapses (Purpura, 1969; Crain and Bornstein, 1974; Purpura et al., 1964). This is manifested by differences in the electrical activity of brain as monitored by EEG and also by differing responses to standard convulsive agents such as electroshock, pentylenetetrazol, strychnine, and picrotoxin. These differences consist of varying behavioral and electrical responses based on age as well as differing sensitivities to these agents as measured by convulsant dose or CDSo (Vernadakis and Woodbury, 1969). During the first postnatal week in rat, the electrocorticogram shows irregular, intermittent, low-amplitude activity. From days 7-10 there is a tendency toward rhythmicity with a progressive increase in amplitude up to day 36 when the adult pattern is reached (Crain, 1952; Bradley et al., 1960; Deza and Eidelberg, 1967;

152

0. CARTER SNEAD 111

Mares et al., 1979). T h e immature rodent brain is less vulnerable to development of kindling than the adult brain (Moshe et aE., 1981). Although extensive ontogeny data are not available concerning human brain, the available data from lower species suggest that the explanation for the higher incidence and protean clinical manifestations of epilepsy in childhood may be due to complex aberrations in the development of one or more neurochemical and/or neurophysiologic systems in brain.

VII. Conclusion

It should be apparent to anyone who has persevered and read this far, that a single neurochemical etiology of epilepsy is unlikely. Although elegant unitary hypotheses have been advanced in an attempt to explain the phenomenon of the seizure (Delgado-Escueta and Horan, 1980; Roberts, 1980), such an approach is made difficult because of the heterogeneity of seizure states as evidenced by the multiplicity of clinical seizures (Table I) and seizure models (Table 11). Also mitigating against a unitary concept of epilepsy are the complex interactions that exist between central neurotransmitters, neuromodulators, and their systems via both multisynaptic well-defined circuitry and point to point, ill-defined local circuitry. A final complicating factor, since so many seizures occur in children, is that of brain development with neurochemical and neurophysiologic ontogeny, myelination, and dendritic arborization all contributing to selective vulnerability or resistance to a particular form of seizure. These observations are not to imply, however, that no answer concerning basic mechanisms and causes of seizure disorders will ever be forthcoming. On the contrary, the extensive bibliography referenced herein is illustrative of an enormous amount of investigation that is ongoing. Rather, the search for causes and treatment of the human condition known as epilepsy will continue to require an integrated, multidisciplinary neurobiological approach. In this way, as the secrets of the “mindful brain” (Edelman and Mountcastle, 1978)are unraveled, so will be the mysteries of “seleniazetai” (Ross, 1978).

References

Abdul-Ghani, A., Coutinho-Netto, J., Druce, D., and Bradford, H . F. (1981). B i o c h a . Pharmacol. 30, 363-368. A M , C. G., Lowry, K., Tornetsko, A,, and MacNeil, M. (1979). Arch. Int. Pharmacodyn. Ther. 237, 213-299.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

153

Adembri, G. A., Bartolini, A., Bartolini, R., Giotti, A., and Zilletti, C. (1974). Br. J . Pharmacol. 52,439-440. Adrian, E. D. (1936).J. Physiol. (Lmdon) 88, 127-161. Adrian, E. D., and Moruzzi, G. (1939).J. Physiol. (Ladon,) 97, 153-199. Agarwal, S. L., and Bhargava, V. (1964). Indian J . Med. Res. 52, 1179-1182. Aghajanian, G. K. (1972). Fed. Proc., Fed. Am. Soc. Exp. Biol. 31, 91-96. Albertoni, P. (1882). Arch.fur Exp. Pathol. Pharmacol. 15, 248-288. Alderdice, M. T., and McMillan, J. E. (1980). Fed. Proc., Fed. Am. Soc. Ex$. Biol. 39, 409. Alderman, J. C., and Shellenberger, M. K. (1974).J. Neurochem. 22,937-940. Alexander, G. J., and Alexander, R. B. (1976). Res. Commun. Psychol., Psychiatry Behav. 1, 105- 114. Alexander, G. J., and Kopeloff, L. M. (1970). Bruin Res. 22, 231-235. Alexander, G. J., and Kopeloff, L. M. (1976). Res. Commun. Chem. Pathol. Pharmucol. 14, 437-448. Alexander, G. J., Kopeloff, L. M., and Alexander, R. B. (1971). L$”e Sci. 10, 877-882. Alger, B. E., and Nicoll, R. A. (1980). Science 210, 1122-1124. Altshuler, H. L., Killam, E. K., and Killam, K. F. (1976).J. Pharmacol. Exp. Ther. 196, 156- 166. Alvord, E. C., and Fuentes, M. G. F. (1954).Am.J. Physiol. 176, 253-261. Ambinder, R. F., and Schuster, M. M. (1979). Gastroenterology 77, 1132-1 140. Anderson, E. G., Markowitz, S. D., and Boungcastle, D. D. (1962).J. Pharmacol. Exp. Ther. 136, 179-182. Ando, N., Simon, J. R., and Roth,R. H. (1979).J. Neurochem. 32, 623-625. Andrews, P. R., and Johnston, G. A. (1979). Biochem. Pharmacol. 28, 2697-2702. Anlezark, G. M., and Meldrum, B. S. (1975). Br. J . Phurmacol. 53, 419-426. Anlezark, G. M., Horton, R. W., Meldrum, B. S., and Sawaga, M. C. B. (1976). Biochem. Pharmacol. 25, 413-417. Anlezark, G. M., Collins, J. F., and Meldrum, B. S. (1977). Neurosci. Lett. 7, 337-340. Anlezark, G. M., Horton, R. W., and Meldrum, B. S. (1978).Adu. Biochem. Psychopharmacol. 19,383-388. Antonitis, J. J., Carary, D. D., Swain, P. B., and Cohen, C. J. (1954).J. Hered. 45,279-284. Aprison, M. H., and Nadi, N. S. (1978). I n “Amino Acids as Chemical Transmitters” (F. Fonnum, ed.), pp. 531-570. Plenum, New York. Arneson, D., Chjien, L. T., Chance, P., and Wilvoy, R. S. (1979). Pediatrics 63, 369-373. Arnold, P. S., Racine, R. J., and Wise, R. A. (1973). Exp. Neurol. 40,457-470. Ashorabi, R. B., Guha, D., and Pradhan, S. N. (1979). Psychopharmacology (Berlin) 64, 345-353. Ashton, D., and Wauquier, A. (1979). Pharmacol., Biochem. Behuv. 11, 221-226. Ashton, D., Leysen, J. E., and Wauquier, A. (1980). Life Sn’. 27, 1547-1556. Atterwill, C. K., Batts, C., and Bloomfield, M. R. (1980).J. Pharm. Pharmacol. 33,329-331. Ayala, G. F., Matsumoto, H., and Gummit, R. J. (1970).J. Neurophysiol. 33, 73-85. Ayala, G. F., Dichter, M., Gummit, R. J., Matsumoto, H., and Spencer, W. A. (1973). Bruin Res. 52, 1-17. Ayala, G. F., Johnston, D., Lin, S., and Dichter, H. N. (1977). Brain Res. 121, 259-270. Azmitia, E. C. (1978).In “Handbook of Psychopharmacology” (L. L. Iversen, S. D. Iversen, and S. H. Snyder, eds.), Vol. 9, pp. 223-314. Plenum, New York. Azzaro, A. J., and Gutrecht, J. A. (1975). Neurology 25, 378-385. Azzaro, A. J., Wenger, G. R., Craig, C. R., and Stitzel, R. E. (1972).J. Pharmucol. Exp. Ther. 180, 558-568. Azzaro, A. J., Gutrecht, J. A., and Smith, D. J. (1973).Biochem. Pharmacol. 22,2718-2729. Babb, T. L., Babb, M., Mahnke, J. H., and Verseano, M. (1973). Int. J . Neurol. 8, 198-210. Babington, R. G., and Wedeking, P. W. (1973). Pharmacol., Biochem. Behav. 1, 461-467.

154

0.CARTER SNEAD 111

Bachus, S. E.. Young, A. B., and Valenstein, E. S. (1980). Soc. ,Veiwmci. .4bstr. 6, 545. Baez, L. A,, Eskridge, N. K., and Schein, R. (1976). E u r . J . Phartnacol. 36, 155-161. Baker, M’.W.(1965)..4rch. I t / / . Phcomaco+. Thm. 155, 273-281. Baker, W. W.. and Benedict, F. (1968). Itit. J. A‘etirophnrrtmcol. 7, 135-147. Ralcar, V. J., Pumain, R., Mark, J., Borg, J., and Mandel. P. (1978). Bruin Rex. 154, 182185. Ballantine, E. (1963).Co/loq. I n / . C. .\. R . S . 112, 447-451. Baraszko, J. J.. Bannon, M. J., Bunney. B. S., and Roth, R. H. (1981).J. Phnrmarol. Exp. Thm. 216, 289-293. Barker, J. L., and Gainer, H. (1973). Scirirre 182, 720-722. Barker, J. L., and Ransom, B. R. (1978a).J. Phyiol. (Lonllou) 280, 331-354. Barker, J . L., and Ransom, B. R. (1978h).J. Ph?.jiol. (London) 280, 355-372. Barker, S. .4., Harrison, R. E. W., Monti, J. A., Brown, G. B., and Christian, S. T. (1981). Riorhein. Phoi-nmrol. 36, 9- 17. Barolin. G. S., and Hornykiewitz, 0. (1967). II‘i~ri.K/in. Il’ochmschr. 79, 815-818. Bartholini, C., Scatton, B., Zivkovic, B., and Lloyd, K. G. (1979). I n “GABAIVeurotrarismitters” (P. Kragsgaard-Larsen, J., Scheelakrueger, and H. Kofod, eds.), pp. 326-339. Academic Press, New York. Bayon, A., Shoemaker, W. J.. Bloom, F. E., Mauss, A,, and Guilleimin, R. (1979).Brain Res. 179,93-98. Bazemore. ‘4. W., Elliott, A. C., and Florev, E. (1957).J. h‘rurochttn. 1, 334-342. Beart, P. M., and Bilal, K. (1979). Bic~hem.Phnrmcirol. 72, 346-351. Beaumont, A., and Hughes, J. H. (1979).,4nrru. Rn1. Pharm. Toxicol. 19, 245-267. Beckwith, B. E., and Sandman, C. A. (1978). A’eurosci. Bfhazr. Rev. 2, 31 1-345. Beart, P. M.. and Johnston, G. F. R. (1973).J. Seurorhpm. 20, 319-324. Ben-Ari, B. Y., and Kelly, J. S. (1976).J. Physiol. (Londo,i) 256, 1-21. Ben-Ari. B. Y..Lagowska. Y., Le Gal La Salle, G., Tremhlay, E., Otterien, 0. P., and Naquet, R. (1978).Ezir. J. Phai-mnrol. 52, 419-420. Ben-Ari, B. Y., Lagowska, Y., Tremhlay, E., and Le Gal La Salle, G. (1979). Brain Rex. 163, 176- 179. Ben-Ari, B. Y., Tremhlay, E., Ohersen, 0. P., and Meldrum, B. S. (1980). Bruin RPS.191, 79-97. Benedito, M. A . C.. and Leite, J. R. (1981). Exp. Xuurol. 72, 346-351. . 247-295. Bennett, H. P. J . (1979). Phnnrincol. R P Z ~30, Bergstriim, D. X., and Kellar, K. J. (1979). .\’attire (Loridon) 178, 531-537. Berl, S.. Purpura, D. Q.. Girado, C., and Waelsch, H. (1959).J. Nrwrorheni. 4, 311-317. Berl, S., Takagaki, G., and Purpura, D. P. (196l).J. A‘eurorhem. 7, 198-209. Bernard, P. S.. Sohiski. R. E., and Dawwn, K. M . (1980). Brain Re.?. Bull. 5, Suppl. 2, 519-523. Bertaccini, G. (1959).J. .\~tiroch~in. 4, 217-222. Berti, F., Bernareggi, V., Folco, G. C.. Fumagalli, R., and Paoletti, R. (1976)..4dlt. Hiorhem. Psschophn rrrrcirol. 15, 367 -378. Beselin, D., Polak, R. L., and Sproul, D. H. (1965).J. Phssiol. (London) 181, 308-316. Bkanchi, C., Beemi. L., and Bertelli, A. (1975). h’eiiruphartnnco1og-i 14, 327-332. Bjorklund. .4., and Nobin, A. (1973). Rrairi Reg. 83, 531-537. Bloom, F. E. (1979). Itr “Catecholamines: Basic and Clinical Frontiers” (E. Usdin, I. J. Kopin, and J. Bachus, eds.), pp. 609-611. Pergamon, Oxford. Bloom, F. E.. Costa, E., and Salmonaghi, G. C. (1965).J. Pharmacol. Exp. Thw. 150, 244252. Bloom, F. E., Hoffer, B. J., Siggins, G. R., Barker, J. L., and Nicoli, R. A. (1972).Fed. Pror., Fed. Am. Sor. Exp. B i d . 31, 97-106.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

155

Bloom, F. E., Rossier, J., Bottenberg, E. L. F., Bayon, A., French, E., Henriksen, S. J., Siggins, G. R., Segal, D., Browne, R., Ling, N., and Guillemin, R. (1978).Adu. Biochem. Psychopharmacol. 18, 89-98. Boggan, W. O., and Seiden, L. S. (1971). Physiol. Behau. 6, 215-217. Bohus, B. (1979). Pharmacology 18, 113. Bokums, J. A., and Elliott, H. W. (1968).Pharmacology 1, 98-110. Bonaventure, N., Wioland, N., and Mandel, P. (1974). Brain Res. 80, 281-289. Bondy, S. C., Tepper, J. M., and Bettis, D. B. (1979). Neurochem. Res. 4, 755-761. Bonnycastle, D. D., Giarman, N. J., and Paasonem, M. K. (1957). Br. J . Pharmacol. Chemother. 12,228-231. Bonnycasde, D. D., Bonnycastle, M. F., and Anderson, E. G. (1962).J. Phannacol. E x f . Ther. 135, 17-20. Borenstein, P., Dabbah, M., and Bles, G. (1962). Ann. Med-Psychol. 120, 133-137. Bourn, W. M., Chin, L., and Picchioni, A. L. (1972).J. Pharm. Phurmacol. 14, 913-914. Bourn, W. M., Chin, L., and Picchioni, A. L. (1977). Life Sci. 21, 701-706. Bowers, M. B., jr., Heninger, G. R., and Gerbode, E (1969). Znt. J . Neuropharmacol. 8, 255-262. Bradley, K., Easton, D. M., and Ecdes, J. C. (1953).J. Physiol. (Lundmr) 122,474-488. Bradley, P. B. (1968). Int. REV.Neurobiol. 11, 1-56. Bradley, P. B., and Dray, A. (1973). Br. J. Pharmacol. 48, 212-214. Bradley, P. B., Eayrs, J. T.,and Schmalback, K. (1960).Electroencephalogr. Clin. Neurophysiol. 12,467-474. Bradley, P. B., Dhawan, B. N., and Wolstencroft, J. H. (1966).J. Physiol. (London) 183, 658 -674. Braestrup, C., Albrechtsess, R., and Squires, R. F. (1977). Nature (London) 269, 702-704. Braestrup, C., Neilsen, M., Krogsgaard-Larsen, P., and Falch, E. (1979). Nature (London) 280,331-333. Braestrup, C., Nielsen, M., and Olsen, C. E. (1980). Proc. Natl. Acad. Sci. U.S.A. 77, 22882292. Brailowsky, S., and Naquet, R. (1976). Epilepsia 17, 271-275. Breese, G. R., and Taylor, T. D. (1971). Br. J. Pharmacol. 42, 88-99. Breitner, C., Picchioni, A,, Chin, L., and Burton, L. E. (1961). Dis. Neru. Syst. 22, 93-96. Breitner, C., Picchioni, A., and Chin, L. (1964).J. Neuropsychiatq 5, 153-158. Briscoe, T. J., and Straughan, D. W. (1966).J. Physiol. ( L a d a ) 183, 341-359. Brooks, B. R., Wood, J. H., Sude, J., and Engel, W. K. (1976). Trans. Am. Neurol. Assoc. 101, 22 1-222. Browning, R. A., and Maynert, E. W. (1978a). Eur. J. Pharmacol. 50, 97-101. Browning, R. A,, and Maynert, E. W. (1978b). Neuropharmacology 17,649-653. Browning, R. A,, and Simonton, R. L. (1978). Life Sn’. 22, 1921-1930. Brownstein, M., Kobayashi, R. M., Palkovits, M., and Saavedra, J. M. (1975).J. Neurochem. 24,35-38. Bruni, J., and Wilder, B. J. (1979).Arch. Neurol. (Chicago) 36, 393-398. Buckholtz, N. S. (1980). Lqe Sn‘. 27, 893-903. Bulat, M. (1977). Brain Res. 122, 388-391. Bunney, B. S., and Aghajanian, G. K. (1973).In “Frontiers in Catecholamine Research” (E. Usdin and S. Snyder, eds.), pp. 957-962. Raven Press, New York. Bunney, B. S., and Aghajanian, G. K. (1977). I n “Pharmacology of Non-Striatal Dopaminergic Neurons” (E. Costa, M. Trabucchi, and G. C. Gessa, eds.), pp. 65-70. Raven Press, New York. Burchfiel, J. C., Duchoway, M. S., and Duffy, F. H. (1979). Sciplice 204, 1096-1098. Buterbaugh, C. G. (1977). Neuropharmacology 16, 707-709.

156

0.CARTER SNEAD 111

Byrne, M. C., Gottlieb, R., and McNamara, J. 0. (1980). Exp. ,ivurnl. 69, 85-98. Cain, D. P. (1979). Exp. .Vpurol. 66, 319-329. Callaghan, D. A,, and Schwark, W. S. (1979). Neurophnnnacolo,gy 18, 541-545. Carette, B. (1981). A’europeptides 1(4), 283-291. Celesia. G. G.. and Jasper, H. H. (1966). Areurology 16, 1053-1063. Cereghino, J. J., and Dow, R. S. (1970). Epilepsia 11,413-421. Chadwick, D., Reynolds, E. H., and Marsden, C. D. (1974). Lanret 2, 111-112. Chadwick, D., Jenner, P., and Reynolds, E. H. (1975a). Lamet 1,474-476. Chadwick, D., Harris, R., Jenner, P., Reynolds, E. H., and Marsden, C. D. (1975b).Lanret 2, 434-435. Chadwick, D., Jenner, P., and Ryenolds, E. H. (1977a). Ann. Neurol. 1, 218-224. Chadwick, D., Hailett, N., Harris, Z., Jenner, P., Reynolds, E. H., and Marsden, C. D. (1977b). Brain 100,455-487. Chadwick, D., Trimble, M., Jenner, P., Driver, M. V., and Reynolds, E. H. (1978).Epilepsia 19, 3- 10. Chang, K. J., Cooper, B. R., Hazum. E., and Cuatrecasas, P. (1979). Md. Pharrnacol. 16, 91- 104. Chase, T. N., and Tamminga, C. A. (1979). I n “GABA-Neurotransmitters” (P. Krogsgaard-Larsen, J., Scheelakrueger, and H. Kofod, eds.), pp. 283-294. Academic Press, New York. Chase, T. N., Katz, R. I., and Kopin, I. J. (1970). Neurophamrologl, 9, 103- 108. Chavkin, C., James, I. F., and Goldstein, A. (1982).Sciozce 215,413-415. Chen, G., Ensor, C. R., and Bohner, B. (1954). Proc. Sor. Exp. B i d . Med. 86, 507-510. Chen, G., Westan, J. K., and Bratton, J. C. (1963). Epiiepsiu 4, 66-76. Chen, G., Ensor, C. R., and Bohner, B. (1968).Arch. lnt. P h a m a c o d p . Ther. 172,183-218. Cheney, D. L.. Lefevere, H. F., and Racagni, G. (1975). Neurophamacologl, 14, 801-809. Cheney, D. L., Moroni, F., Malthe-Sovenssen, D., and Costa, E. (1977). In “Cholinergic Mechanisms and Psychopharmacology” (D. J. Jenden, ed.), pp. 551-563. Plenum, New York. Chu, N. (1978). Epilepsiu 19, 603-609. Chusid, J. G . , and Kopeloff, L. M. (1969). Epilepsia 10, 239-262. Ciesielski, L., Simler, S., and Mandel, P. (1981). Neurncha. Res. 6, 267-273. Cohen, B., and Myerson, A. (1938). Am. J. P.ychiatIy 95,371-393. Cohen, S . L., Morley, B. J.. and Snead, 0. C. (1981). Prog. Neuro-Psychopharmacol. 5, 383388. Colasanti, B. K., and Craig, C. R. (1973). Neuropharmacology 12, 221-231. Collins, R. C. (1980). “Vturology 30, 575-581. Collins, R. C., and Hehta, S. (1978). Brain Res. 157, 311-320. Comrnissiong, J. W., and Neff, N. H. (1979). Biochm. Phamacol. 28, 1569-1573. Conner, J. D. (1970).J. Physiol. (Lortdon) 208, 691-703. Consolo, S . , Garrattini, S., and Ladinsky, H. (1975). Adu. Biorhem. Psychopharmacol. 14, 63-80. Consroe, P., and Edmonds, H. L. (1979).Fed. Proc., Fed. Am. SOC.Exp. B i d . 38,2397-2398. Contreras, C. M., Gonzalez-Estrada, T., Zaucbozo, D., and Fernandez-Guardiola, A. (1975). Electronicephalogr. Clin.iVmrophysio1. 46, 290-301. Cook, G. H., and Dole, J. A. (1942). Dis. N m . Syst. 3, 366-370. Cools, A. R., and Van Rossurn, J. M. (1980). Lifp Sci. 27, 1237-1253. Cools, A. R., Hendriks, G., and Korten, J. (1975). J. Neural. Tralzsm. 36, 91-105. Cooper, B. R., Viik, K., Ferris, R. M., and White, H. L. (1979).J. P h a m c o l . Exp. Ther. 209, 396-403. Cooper, I. S . (1973). Lancet 1, 206.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

157

Cooper, I. S., and Gilman, S. (1973).I n “Neural Organization and its Relevance to Prosthetics” (W. Fields, ed.), pp. 371-375. Intercontinental Publ., New York. Cooper, I. S., Amin, I., and Gilman, S. (1973). Trans. Am. Neurol. Assoc. 98, 192-196. Cooper, I. S., Amin, I., Riklan, M., Walter, J. M., and Poon, T. P. (1976). Arch. Neurol. (Chicago) 33,559-570. Cooper, I. S., Amin, I., Upton, A,, Ricklan, M., Watkins, S., and McLellan, C. (1978).Appl. Neurophysiol. 40, 124- 134. Cooper, J. R., Bloom, F. E., and Roth, R. H. (1978). In “The Biochemical Basis of Neuropharmacology,” 3rd ed., pp. 37-45. Oxford Univ. Press, LondodNew York. Corcoran, M. E., and Wada, J. A. (1979).Life Sci. 24, 791-800. Cocoran, M. E., Fibiger, H. C., McGeer, E. G., and Wada, J. A. (1973).J. Pharm. Pharmacol. 25,497-499. Corcoran, M. E., Fibiger, H. C., McCaughram, J. A., and Wada, J. A. (1974).Exp. Neurol. 45, 118-133. Corcoran, M. E., Wada, J. A., Wake, A., and Urstad, H. (1976).Exp. Neurol. 51,271-275. Costa, E., Guidotti, A., and Mao, C. C. (1975).Adv.Biochem. Psychophannacol. 14, 131-151. Costa, E., Guidotti, A., and Toffana, G. (1978).Br. J . Psychiatry 133, 239-242. Costa, E., Guidotti, A., Moroni, F., and Peralta, E. (1979).In “Glutamic Acid: Advances in Biochemistry and Physiology” (L. J. Filer, M. R. Kare, S. Garattini, and W. A. Reynolds, eds.), pp. 151-161. Raven Press, New York. Costall, B., and Naylor, R. J. (1981).Life Sci. 28, 215-229. Coursin, D. B. (1954).JAMA,J. Am. Med. Assoc. 154, 406-408. Coursin, D. B. (1955).Am. J . Dis. Child. 90, 344-348. Courtney, K. R., and Prince, D. A. (1977),Brain Res. 127, 191-196. Coutinho-Netto, J., Abdul-Ghani, A. S., Collins, J. F., and Bradford, H. F. (1981).Epilepsia 22, 289-296. Cowen, P. J., Green, A. R., and Netto, J. (1981).Nature (London) 290, 54-55. Cox, B., and Lomax, P. (1976).Pharmacol., Biochem. Behav. 4, 263-267. Coyle, J. T. (1977).Znt. Rev. Neurobiol. 20, 65-103. Coyle, J. T., and Enna, S. J. (1976).Brain Res. 111, 119-127. Coyle, J. T., and Henry, D. (1976).J. Neurochem. 21, 61-66. Coyle, J. T., and Schwartz, R. (1976).Nature ( L a d a ) 263, 244-246. Coyle, J. T., Molliver, M. E., and Kuhar, M. J. (1978).J. Comp. Neurol. 180, 301-323. Craig, C. R., and Hartman, E. R. (1973).Epilepsia 14,409-414. Crain, S. M. (1952). Proc. SOC. Exp. Biol. Med. 81, 49-55. Crain, S. M., and Bornstein, M. B. (1974).Brain Res. 68, 351-360. Crawford, J. M. (1970).Neuropharmacology 9, 31-46. Crawford, J. M., and Curtis, D. R. (1964).Br. J . Pharmacol. 23, 313-329. Crawford, J. M., and Curtis, D. R. (1966).J. Physiol. (Ladon) 186, 121-138. Crawford, J. M., Curtis, D. R., Vourhoens, P. E., and Wilson, J. V. (1966).J. Physiol. (Londa) 186, 139- 165. Crighel, E. (1966).Epilepsia 7, 283-290. Crosley, C. J., Richman, R. A., and Thorpy, M. J. (1980).Ann. Neurol. 8, 220. Crossland, J., and Turnbull, M. J. (1972).Neuropharmacology 11, 733-738. Crunelli, V., Berasconi, S., and Samann, R. (1979).Psychofhamcology 66, 79-85. Curtis, D. R. (1969).Prog. Brain Res. 31, 171-189. Curtis, D. R. (1979).In “Glutamic Acid: Advances in Biochemistry and Physiology” (L. J. Filer, M. R. Kare, S. Garattini, and W. A. Reynolds, eds.), pp. 163-175. Raven Press, New York. Curtis, D. R., Phillis, J. W., and Watkins, J. C. (1960).J. Physiol. (Londa) 150, 656-682. Curtis, D. R., Hosli, G., and Johnston, G. A. R. (1967).Nature (Ladon) 215, 1502-1503.

158

0. CARTER SNEAD I11

Curtis, D. R., Hosli, G., a n d Johnston, G. A. R. (1968). Exb. Brain Res. 6, 1-18. Curtis, D. R.. Duggan, A. W., and Felix, D. (1970a). Brain Res. 23, 117-120. Curtis, D. R., Duggan. A. W., Felix, D.. and Johnston, G. A. R. (1970b). Nature ( L o n d m ) 226, 1222-1230. Curtis, D. R.. Duggan, A. W., and Johnston, G. A. R. (1971a).Exp.Brain Res. 12,547-565. Curtis, D. R., Duggan, A. W., Felix, D., and Johnston, G. A. R. (1971b). Bmin Res. 32, 69-96. Curtis, D. R.. Came. C . J . A., Johnston, G. A. R., McCulloch, R. M., and McClachlan, R. M. (1972). Brain Res. 43, 242-245. Curtis, D. R., Felix, D., Game, C. J. A., a n d McCullough, R. M. (1973). Brain Res. 51, 358- 362. Curtis, D. R., Lodge, D., Johnston, G. A. R., and Brad, C. J. (1976a). Bruin RPS.118, 344-347. Curtis, D. R., Game, C. J. A., a n d Lodge, D. (1976b).B r . J . Phnrmacol. 56, 307-311. Cutler, R. W. P., a n d Young, J. (1979).Brain Res. 170, 157-163. Cybulski, N. (1914). Bull. Int. Acad. Sco.. Cmcovie SYTB pp. 776-781. Diihlstrom, A,, a n d Fuxe, K. (1964).Actn Physioi. Scand. 62, Suppl. 247, 1-36. Daly, J. W. (1975).Biorhem. Phnrmarol. 24, 159-164. Daly, J . W. (1977). Int. Rev. 2’Veirrobiol.20, 105-168. Daniels, J. C., a n d Spehlman, R. (1973). Electroencephalogr. Clin. Neurophysiol. 34, 83-87. Dasheiff, R. M., Byrne, M. C., Patrone, V., a n d McNamara, J . 0. (1981). Bruin Res. 206, 233-238. Dauth, G. W., Dell, S., a n d Gillman, S. (1978).A’euroloa 28, 654-660. Davidoff, R. A. (1972a).Brain Res. 45, 638-642. Davidoff, R. A. (l972b). Trarzs. Am. .\‘eurol. As~or.97, 193-196. Davies, J. A. (1978).Psjchophnnnarology 60, 67-72. Deakin, J. F. W., Owen, F., Cross, A. J., a n d Dashwood, M. J. (1981).Psyhopharmacology 73, 345-349. DeBelleroche, J., a n d Bradford, N. F. (1978).Brain RPS.142, 53-68. Deisz, R. A,, a n d Lux, H. D. (1977). Meicrosci. Lett. 5, 199-203. De la Torre, J. C., a n d Mullan, S. (1970).J. Pharm. Pharmucol. 22, 858-859. De la Torre, J. C., Kawanga, H. M., and Mullan, S. (1970).Arch.Int. Pharmacodyn. Ther. 188, 298-304. De Lean, J. (1977). S. Eng1.J. Med. 296, 1414-1415. De Lean, J., Richardson, J. C., and Hornykiewicz, 0. (1976).Neurology 26,863-868. Delgado, J. M. R., De Feudis, F. V., a n d Bellido, I. (1971). Commun. Behav. Biol. 5,347-357. Delgado-Escueta, A. V., and Horan, M. P. (1980).In “Antiepileptic Drugs: Mechanisms of Action” (G. H . Glaser, J. K. Penry, and D. M. Woodbury, eds.), pp. 85-126. Raven Press, New York. Della-Fera, M. A., a n d Bailey, C. A. (1979).Science 206, 471-474. De Robertis, E., and Fiszer de Plazas, S. (1976).J. Neurochem. 26, 1237-1243. De Schaepdryver, A,, Pietle, Y., a n d Delaunois, A. (1962).Arch.Int. Phanacodyn. Ther. 140, 358-367. De Vanzo, J. P., Greig, M. E., and Cronin, M. A. (1961).Am. J . Physiol. 201, 833-837. Dewar, A. J., Dow, R. C., a n d McQueen, J. K. (1972). Epilepsla 13,552-560. de Weid, D. (1977a).Ann. N.).’.Acad. Sci. 297,262-267. De Weid, D. (1977b).Life Sri. 20, 195-204. de Weid, D., a n d Gispen, W. H. (1979).In “Peptides in Neurobiology” (H. Gainer, ed.), pp. 397-448. Raven Press, New York. d e Weid, D., Witter, A., and Greven, H . M. (1975).Biocha. Phannacol. 24, 1463-1470. Deza. L., a n d Eidelberg, E. (1967).Exp. Neurol. 16,425-474.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

159

Diaz, P. M. (1970). L f e Sci. 9, 831-840. Diaz, P. M. (1974). Neuropharmacology 13, 615-621. Dichter, M., and Spencer, W. A. (1969).J . Neurophysiol. 32,663-686. Dingledine, R., and Gjerstad, L. (1979). Brain Res. 168, 205-209. Dockray, G. J. (1976).Nature (London)264, 568-570. Dockray, G. J. (1980). Brain Res. 188, 155-165. Dockray, G. J., Gregory, R. A,, Hutchinson, J. B., Hanis, J. I., and Runswick, M. J. (1978). Nature (London) 274, 711-713. Dodd, P. R., and Bradford, H. F. (1976).Brain Res. 111, 377-388. Domino, E. F., and Olds, M. E. (1972). Psychopharmucologza 23, 1-16. Donald, R. A. (1980). Clin. Endocrinol. (Oxford) 12, 491-524. Doteuchi, M., and Costa, E. (1973). Neuropharmacology 12, 1059-1072. Dow, R. C., McQueen, J. K., and Townsend, H. R. A. (1972).Epilepsia 13,459-465. Dow, R. C., Hill, A. G., and McQueen, J. K. (1974).Br. J. Pharmacol. 52, 135. Dow, R. S., Fernandez-Guardiola, A,, and Manni, E. (1962). Electroencephalogr. Clin. Neurophysiol. 14, 383-3 98. Dreifuss, F. E., Bancaud, J., Henriksen, O., Rubio-Donnadieu, F., Seino, M., and Penry, J . K. (1981).Epilepsia 22,489-501. Dreifuss, J. J., Kelly, J. S., and Krujevie, K. (1969).Exp. Brain Res. 9, 137-154. Dudar, J. D., and Szerb, J. C. (1969).J. Physiol. (London) 203, 741-762. Duvoisin, R. C., and Dettbran, W. D. (1967). Neurology 17, 1077-1081. Ebadi, M., and Klangkalya, B. (1978). Neuropharmacology 18, 301-307. Eccles, J. C., Ito, M., and Szendgothai,J. (1967). “The Cerebellum as a Neuronal Machine. “Springer-Verlag, New YorWBerlin. Echlin, L. A., and Battista, A. (1963).Arch. Neurol. (Chicago) 9, 154-170. Edelman, G. M., and Mountcastle, V. B. (1978). “The Mindful Brain.” MIT Press, Boston, Massachusetts. Edmonds, H. C., Hegreberg, G. A., Van Gelder, N. M., Sylvester, D. M., Clemmons, R. M., and Chatburn, C. B. (1979). Fed. Proc., Fed. Am. SOC.Exp. Biol. 38, 2424-2428. Edmonds, H. L., and Stark, C. G. (1974). Neuropharmacology 13,269-277. Edmonds, H. L., Stark, C. G., and Hollinger, M. A. (1974). Exp. Neurol. 45, 377-386. Ehlers, C. L., and Killam, E. K. (1979). Electroencephalogr. Clin. Neurophysiol. 47, 404-412. Elliott, P. N. C., Jenner, P., Chadwick, D., Reynolds, E., and Marsden, C. D. (1977).J. Pharm. Pharmacol. 29, 41-43. Emson, P. C. (1975).Int. J. Biochem. 6, 689-694. Emson, P. C. (1976).J.Neurochem. 27, 1489-1494. Emson, P. C. (1978). I n “Taurine and Neurological Disorders” (A. Barbeau and R. J. Huxtable, eds.), pp. 319-338. Raven Press, New York. Emson, P. C., and Joseph, M. H. (1975).Brain Res. 93, 91-110. Emson, P. C., Fahran-Krug, J., Schaffalitzky, D. E., Muckadell, 0. B., Jessell, T. M., and Iversen, C. C. (1978).Brain Res. 143, 184-194. Emson, P. C., Hunt, S. P., Rehfeld, J. F., and Gahrenkrug, J. (1980a). Adv. Biochem. Psychopharmacol. 22, 63-70. Emson, P. C., Lee, C. M., and Rehfeld, J. F. (1980b). L f e Sci. 26, 2157-2165. Endroczi, E., Lissak, K., and Felcete, T. (1970). Prog. Brain Res. 32, 254-256. Engel, J., and Katzmann, R. (1977). Brain Res. 122, 137-142. Engel, J., and Sharpless, N. S. (1977).Brain Res. 136, 381-386. Engel, J. Ackermann, R. F., Caldecott-Hazard, S., and Kuhl, D. E. (1981).I n “Kindling” (J. A. Wada, ed.), pp. 193-21 1 . Academic Press, New York. Enna, S. J. (1981).B i o c h a . Pharmacol. 30, 907-913. Enna, S. J., and Maggi, A. (1979).Life Sci. 24, 1727-1738.

160

0.CARTER SNEAD 111

Enna, S. J., Wood, J. H., andSnyder, S. H. (1977).J. Neurochem. 28, 1121-1124. Esplin, D. W., and Cuno, E. M. (1957).J. P h a m c o l . Exp. Ther. 121,457-467. EkpIin, D. W., and Woodbury, D. W. (1961).Scieme 133, 1426-1427. Esplin, D. W., and Zablocka, B. (1969). Epilepsia 10, 193-210. Essig, C. F. (1967).Epilepsia 8, 21-30. Essig, C. F. (1972). In “Experimental Models of Epilepsy-A Manual for the laboratory Worker” (D. P. Purpura,J. K.Penry, D. Tower, D. M. Woodbury, and R. Walter, eds.), pp. 496-508. Raven Press, New York. Essig, C. F., and Laurn, R. C. (1968). Arch. Neurol. (Chicago) 18, 626-632. Essig, C. F., Hampson, J. L., McCauley, A., and Himwick,H. E. (1950).J. Neurophysiol. 13, 269-275. Essman, E. S., and Essman, W. B. (1980). Brain Res. Bull. 5, Suppl. 2, 209-211. Essman, W. B. (1968).Arch.Int. Phawnacodyn. Ther. 171, 159-173. Essman, W.B. (1972). Semin. Psychiatly 4, 67-79. Essman, W. B. (1578).In “Serotonin in Health and Disease,” Vol. 111, pp. 317-401. Spectrum Publications, New York. Essman, W. B., and Essman, E. J. (1980). Brain Res. Bull. 5, Suppl. 2, 821-824. Evans. R. H. (1979).Brain Res. 171, 113-120. Evans, R. H., and Watkins, J. C. (1981).Lge Sci. 28, 1303-1308. Fahn, S. (1976).I n “GABA in Nervous System Function” (E. Roberts, T. N. Chase, and D. B. Tower, eds.), pp. 169-186. Raven Press, New York. Fahn, S. (1978).N. Eng1.J. ,Wed. 299, 313-314. Fahn, S., and Cote, L. J. (1968).J. Nmrochtm. 15, 209-213. Faiman, M. D., and Heble, A. R. (1966). L f e Sci. 5, 2225-2234. Faiman, M. D., Haga, K., Zempel, J. A., and Schower, R. L. (1980). Brain Res. Bull. 5, Suppl. 2, 789-792. Fariello, R. G., and Golden, G. T. (1980). Brain Res. Bull. 5, Suppl. 2, 691-699. FarieUo, R. G., Golden, G. T., and Black, J. A. (1981).Epilepsia 22, 217-224. Farjo, I. B., and Blackwood. R. H. R. (1978). Bratn Res. 153,423-426. Fearnsides, E. G. (1915). Proc. R. Soc. ,ifed. 9, 47-49. Feher, O., Halasz, P., and Mechler, F. (1965).Epilefsia 6, 47-53. Feldberg, W., and Sherwood, S. G. (1954).J. Phjsiol. (Ladon) 123, 148-167. Fennessy, M. R., and Lee, J. R. (1972). Arch. Int. Pharmarodyn. Ther. 197, 37-44. Ferguson, J. H., and Cornblath, D. R. (1975).Exp. Neurol. 46, 302-314. Ferguson, J. H., and Jasper, H. H. (1971).Electromephulogr. Clin. Neurophysiol. 30, 377390. Fernstrom, J. D., Shabshelowitz, H., and Faller, D. V. (1974).L$e Sri. 15, 1577-1584. Ferrari, R. A., and Arnold, A. (1961).Biochim. Biophys. Acta 52, 361-367. Ferrendelli, J. A., and Kinscherf, D. A. (1977).Epilepsia 18, 525-531. Fibiger, H. C., Lytle, C. D., and Campbell, B. A. (1970).J. Cmp. Physiol. Psychol. 72, 384-388. Fink, M. (1966).J. Nmi. Ment. Dir. 24, 475-484. Fisher, R. S., and Prince, D. (1977).Electroencephalogr. Clin. Nmrophysiol. 42, 625-635. Fiszer de Plazas, S., and De Robertis, E. (1976).J. Neurochem. 27, 889-894. Fitz, J. G., and McNamara, J. 0. (1979).Brain Res. 178, 117-127. Fitzsch, G., and Hitzig, E. (1870). Arch. Anat. Physiol. Anat. Abt., 37, 300-332. Florey, E. (1954).Arch. Int. Physiol. 62, 33-47. Folbergrova, J. (1975). Brain Res. 92, 165-169. Fonnum, F., Grofova, I., RInvik, E., Storm-Mathisen, J., and Walberg, F. (1974).Brain Res. 71,77-92.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

161

Fontana, A., and Grob, P. H. (1979).J . Neural. 220, 297-301. Fontana, A., Fulpius, B. W., and Cuenoud, S. (1978a). Schweiz. Med. Wochenschr. 108, 1307-1310. Fontana, A., Joller, H., Skvaril, F., and Grob, P. H. (1978b).J. Neural., Neurosurg. Psychiatry 41,593-597. Fowler, L. J., and John, R. A. (1972). Bi0chem.J. 130,569-573. Freedman, R., Taylor, D. A., Seiger, A., Olson, C., and Hoffer, B. J. (1979).Ann. Neural. 6, 281-295. Frenk, H., and Yitzhak, J. (1981). Exp. Neural. 71,487-496. Frenk, H., Urca, G., and Liebeskind, J. C. (1978a). Brain Res. 147, 327-337. Frenk, H., McCarty, B. C., and Liebeskind, J. C. (1978b). Sciace 200, 335-337. Frenk, H., Engel, J., Ackermann, R. F., Shavit, Y., and Liebeskind, J. C. (1979).Brain Res. 167,435-444. Frey, H. H., and Loscher, W. (1980). Neurophamcology 19, 217-220. Frey, H. H., Popp, C., and Loscher, W. (1979). Neuropharmuculogy 18, 581-590. Fromm, G., and Kohli, C. M. (1972). Neurology (Minneapolk) 22, 1012-1020. Fromm, G. H., Glass, J. D., Chattha, A. S., Martinez, A. J., and Silverman, M. (1980). Neurology 30, 126-131. Fry, B. W., and Ciarlone, A. E. (1981). Neuropharmacology 20, 623-625. Fuller, R. W., Perry, K. W., and Molloy, B. B. (1974).L$e Sci. 35, 1161-1171. Fuller, T. A., and Olney, J. W. (1975). Lqe Sn’. 34, 1793-1798. Fulpius, B. W., Fontana, A., and Cuenoud, S. (1977). Lancet 2, 350-351. Funderburk, W. H., and Case, T. J. (1951). Electroencephulogr. Clin. Neurophysiol. 3, 213223. Fuxe, K., Hokfelt, T., and Unghstedt, U. (1969).Zn “Metabolism of Amines in the Brain” (G. Hooper, ed.), pp. 10-22. Macmillan, New York London. Fuxe, K., Hokfelt, T., Said, S. I., and Mutt, 0. (1977). Neurosci. Lett. 5, 241-247. Gale, K., and Iadarola, M. J. (1980). Science 208, 288-291. Gale, K., Hong, J., and Guidotti, A. (1977). Bruin Res. 136, 371-375. Gall, C., Brecha, N., Karten, H. J., and Chang, K. (1981).J. Cmnp. Neural. 198, 335-350. Gallagher, J. P., Inokuchi, H., Nakamura, J., and Schinnick, P. (1981). Neuropharmcology 20,427-433. Garattini, S., and Valzelli, L. (1957). In “Psychotropic Drugs” (S. Garattini and V. Ghetti, eds.), pp. 428-436. Elsevier, Amsterdam. Gardner, C. R., and Webster, R. A. (1977). Eur. J. Pharmacol. 42, 247-256. Garelis, E., and Sourkes, T. L. (1973).J . Neural., Neurosurg. Psychiatry 36, 625-629. Garelis, E., and Sourkes, T. L. (1974). J. Neural., Neurosurg. Psychiatq 37, 704-710. Garelis, E., Young, S. N., Lal, S., and Sourkes, T. L. (1974). Brain Res. 79, 1-8. Gastaut, H. (1970). Epilepsia 11, 102-113. Gee, K. W., Hollinger, N. A., Boyer, J. F., and Killan, E. K. (1979). Exp. Neural. 66, 771-777. Gershon, M. D., and Erde, S. M. (1981). Gastroenterology 80, 1571-1594. Giachetti, A., Said, S. J., Reynolds, R. C., and Komiges, F. C. (1977). Proc. Natl. Arud. Sci. U.S.A. 74, 3424-3432. Giarman, N. J., and Schmidt, K. F. (1963). Br. J . Pharmacol. 20, 563-568. Gibbins, R. J., Kalant, H., Le Blanc, A. E., and Clark, W. (1971). Psychopharmacologia 19, 95- 104. Gilbert, P. E., and Martin, W. R. (1976).J. Phannocol. Exp. Ther. 198, 66-82. Ginsburg, B. E., and Sze, P. Y. (1975). NIDA Res. Mmogr. Ser. 6, 85-95. Girgis, M. (1978). Epilepsia 19, 521-530.

162

0.CARTER SNEAD 111

Girgis, Y. (1980).Esp. Smrol. 70, 121-125. Girgis, M. (1981).Electrorncrphnlop. Clin. A’mrophjsiol. 51, 417-425. Glaser, G. H . (1953). Epilrpsia 2, 7-15, Gloor, P., a n d Testa, G. (1974). EIrctroorrg/dp-. Clin. Smr-ophyiol. 36, 499-5 15. Godschalk, M., Djotjic, M. R., a n d Bonta, I. L. (1977).J. Pharm. Pharnacol. 29, 605-611. Goddard, G. V. (1967). SntitrP 214, 1020-1021. Goddard. G. V., McIntyre, D. C.. and Leech, C . K. (1969).Exp. A’curol. 25, 295-330. Gold, B. I., a n d Roth, R. H. (1977).J. Smrorhmvi. 28, 1069-1073. Gold, B. I., a n d Roth, R. H. (1979).j. Sntrorhrnr. 32, 883-888. Goldensohn, E. S., a n d Purpura, D. P. (1963). Science 139, 840-842. Goldensohn, E. S., and Ward, A. A. (1975).In “The Nervous System” (D. B. Tower ed.), Vol. 2, pp. 249-260. Raven Press, New York. Goldstein, D. B. (1972).J. Phai-wrcol.,Exp. Thm. 183, 14-27. Goltermann, N. R.. Rehfeld, J. F., and Peterson, H. P. (1980).J. Nrurorhem. 35, 479-485. Goodman. J. H., and Lebovitz. R. M. (1980).Sor. Srurotci. Ahstr. 6, 401. Goodman, R. R., Snyder, S. H., Kuhar, M. J., a n d Young, W. S. (1980). Soc. ,Vrurmci. Ah.j/r. 5, 523. Gottesfeld, Z., a n d Elazar, Z. (1972). .Vatwe (London) 240, 478-479. Gottesfeld. Z., and Elazar, Z. (1975). Br-aiir K P S . 84, 346-350. Grabow. J. D.. Ekersold, M. J., Albers, J. W., a n d Schimra, E. M. (1974).iMn~ Clin. Pror. 49, 759-774. Gramsch. C., Kleber, G., Hollt, V., Pasi, A,, Mehraein, P., and Hem, A. (1980).Brairr KIT. 192, 109- 119. Gray, W. D., a n d Rauh, C.E. (1964).J. Pharntarol. Exp. Thrr. 163, 43 1-438. Gray, W. D., and Rauh, C. E. (1967).J. Phrirmnrol. Exp. Thpr. 155, 127-134. Gray, W. D., a n d Rauh, C. E. (1971).J. Phrrrmcrrol. Exp. T k . 177, 206-218. Gray, W. D., and Raugh, C. E. (1974).Eur. J. Pharmncol. 28,42-54. Gray, W. D., Rauh, C. E., Osterberg. A. C., and Lipchuck, L. M. (1958).J. Phnrmncol. Exp. Thm 124, 149-155. Gray, W. D., Rauh, C. E.. a n d Shanahan, R. W. (1963).J. Phnrmacol. Exp. Thrr. 139, 350-357. Green, A . R.. a n d Grahame-Smith, D. G. (1975). Arriiropharmnco/og?.14, 107-113. Green, A. R., Peralta, E., Hong, J. S., Mao, C . C., Atterwill, C. K., and Costa, E. (1978). J. .Yrut-oc/irin. 31, 607-61 1. Greengard, P. (1976).S n t u r r (London) 260, 101-108. Greenlee, P. V., Van Ness, P. C., a n d Olsen, P. W. (1978).J . ,\?purorhem. 31, 933-938. Grimm. R. J., Frazee,J. G., Bell, C . C., Kawasaki, T., a n d Dow, R. S. (1969).1nt.J. Nenrol. 7, 126-140. Grinker, R. R., Serota, H., and Stein, S. I. (1938).Arch. h‘eurol. Pqrhdr! 40, 968-980. Gross, R. A,, and Ferrendelli, J. A. (1979).Ann. ,\‘mrol. 6, 296-301. Grossman, M. H., Hare, T. A., Mangam, B., Glaeser, B. S.. and Wood, J. H. (1 980). Brain Res. 182, 99-106. Grossman, S. P. (1963). Scienrr 142, 409-41 1. Groves, P. M., Wilson, C . J., Young, W. J., a n d Rebec, G. U . (1975).ScicnrP 190, 522-529. Growdon, J. H. (1977). A’~u~-oIog?. 27, 1074-1077. Growdon, J. H.. Young, R. R., and Shahani, B. T. (1976).Nrurolom 26, 1135-1140. Gruol, D. LA., Barker, J. G., a n d Smith, T . G. (1980).Braiii Res. 198, 323-332. Guberman, A., Gloor, P., and Sherwin, A. L. (1975).Seurolog?.25, 758-764. Guerrero-Figueroa, R., Barros, A., d e Balbain Verster, F., and Heath, Z. G. (1963).,4rch. Srurol. fChirngo)9, 297-306.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

163

Guerrero-Figueroa, R., Verster, F., Barros, A., and Heath, R. G. (1964). Epileflsia 5, 140155. Guidotti, A. (1978).I n “Taurine and Neurological Disorders” (A. Barbean and R. J. Huxtable, eds.), pp. 237-248. Raven Press, New York. Guidotti, A., Toffano, G., and Costa, E. (1978). Nature (London) 275, 553-555. Guillemin, R., Vargo, T., Rossier,J., Minick, S., Ling, N., Rivier, C., Vale, W., and Bloom, F. (1979). Science 202, 1367-1369. Guilleminault, C., Tharp, B. R., and Cousin, D. (1973).J. Neurol. Sci. 18,435-441. Hadfield, M. G. (1972). Arch. N a m l . (Chicago) 26, 78-84. Hadfield, M. G., and Boykin, M. E. (1974). Res. Commun. Chem. Pathol. Pharmacol. 7,209212. Hadfield, M. G., and Rigby, W. F. C. (1976). Biochem. Pharmacol. 25, 2752-2754. Haefely, W., Kulcsas, A., Mohler, H., Pieri, C., Tola, P., and Shaffuer, R. (1975). Adu. Biochem. Psychophamacol. 14, 131-151. Hahn, F., and Oberdorf, A. (1962).Arch. Int. Pharmacodyn. Ther. 135, 9-30. Halasz, N., Ljungdahl, A., Hokfelt, T., Johansson, O., Goldstein, M., Park, D., and Bikerfield, P. (1977). Brain Res. 126, 455-474. Haley, T. J., and McCormick, W. G. (1957). Br. J . Pharmacol. Chemother. 12, 12-15. Hall, C. S. (1947).J. Hered. 38, 2-6. Halliday, A. M. (1967). Brain 90, 241-284. Halpern, L., and Julian, R. (1972a). Epilepsia 13, 377-385. Halpern, L., and Julian. R. (1972b). Epilepsia 13, 387-400. Halpern, L. M. (1972).I n “Experimental Models of Epilepsy-A Manual for the Laboratory Worker” (D. P. Purpura, J. K. Penry, D. Tower, D. M. Woodbury, and R. Walter, eds.), pp. 197-222. Raven Press, New York. Hanbrich, D. R., Wang, P. F. L., Clody, D. E., and Wedeking, P. W. (1975). Life Sci. 17, 975-980. Hardy, C., Panksepp, J., Rossi, J., and Zolovick, A. J. (1980). Brain Res. 194, 293-297. Harris, R. (1964).Arch. Dis. Child. 39, 564-568. Hattori, T., McGeer, P. G., Fibiger, A. C., and McGeer, E. G. (1973). Brain Res. 54, 103114. Hayashi, T.. and Negishi, K. (1979). Brain Res. 175, 271-276. Hebb, C. O., Krnjevic, K., and Silver, A. (1963). Nature ( L a d o n ) 198, 692. Hemsworth, B. A., and Neal, M. J. (1968). Br. J . Pharmarol. Chemother. 32, 543-550. Henriksen, S. J., Bloom, F. E., McCoy, F., Ling, N., and Guillemin, R. (1978). Proc. Natl. Acad. Sci. U.S.A. 75, 5221-5225. Herink, J., Fusck, J., and Huding, V. (1980). Act. Neru. Super. 22, 209-210. Herkenham, M., and Pert, C. B. (1980). Proc. Natl. Acad. Sci. U.S.A. 77, 5532-5540. Heymans, C., Schaepdryver, A. F., Delaunois, A. L., and Piette, Y. (1964).Med. Maatsschr. 18, 220. Hildebrandt, F. (1926). Naunyn-Schmiedebergs Arch. Ex$. Pathol. Pharmakol. 116, 100- 109. Hill, R. G., Simmonds, M. F., and Straughn, D. W. (1973). Br. J . Pharmacol. 49, 37-51. Hill, R. G., Simmonds, M. F., and Straughn, D. W. (1976). Br. J. Pharmucol. 56, 9-19. Hiller, J. M., and Simon, E. J. (1979). Eur. J . Pharmacol. 40, 389-398. Hitchcock, E., and Gabra-Sanders, T. ( 1977).J. Neurol., Neurosurg. Psychiatry 40, 565569. Ho, I. K. (1980). Brain Res. Bull. 5, Suppl. 2, 913-917, Hochner, B., Spira, M. E., and Werman, R. (1976).BrainRes. 107, 85-103. Hoefer, P. F. A., and Glaser, G. H. (1950). Pror. First Clin. ACTH C a J 1, 536-542. Hoefer, P. F. A., Aranow, A., and Rowland, C. P. (1958).Arch. Neurol. (Chicago) 80, 10-17.

164

0. CARTER SNEAD 111

Hoffer, B. J., Siggins, G. R., Oliver, A. P., and Bloom, F. E. (1973).J. Pharmucol. Exp. Ther. 184,553-569. Hoffer, B. J., Seiger, A., Freedman, R., Olson, L., and Taylor, D. (1977a). Brain Res. 119, 107-132. Hoffer, B. J.. Seiger, A,, Taylor, D., Olson, L., and Freedman, R. (1977b). Exp. Nturol. 54, 233-250. Hoffman, B. B., and Lefkowitz, R. J. (1980). IV. Eng1.J. Mrd. 302, 1390-1396. Hohenboken, W. D.. and Nelhaus, G. (1970).J. Hered. 61, 107-112. Hokfelt, T., Halasz, N., Ljungdahl, A,, Johansson, O., Goldstein, M., and Park, D. (1975). Neurosci. Lett. 1, 85-90. Hokkanen, E., and Tiovakka, E. (1969). Acta Neurol. Scand. 45, 556-567. Hollt, V., Przewlocki, R.,and Herz, A. (1978). Naunll.n-Schmieriebergrg’sArch,. Pharmacol. 303, 171- 177. Holmstedt, B., Lundgren, G., and Sundwall, A. (1963). Lifp Sn’. 10, 731-736. Hoover, D. B., Craig, C. R., and Colasanti, B. K. (1977). Exp. Brain Res. 29, 501-513. Hoover, D. B., Muth, E. A., and Jacobowitz, D. M. (1978). Bruin Reg. 153, 295-306. Hori, M., Ito, T., Yoshida, K., and Shimizu. M. (1979). Epilepsiu 20, 25-36. Horton, R. W., and Meldrum, B. S. (1973). Br. J . Phamacol. 49, 52-63. Horton, R. W., Meldrum, B. S., Sawaya, M. C. B., and Stephenson, J. D. (1976). Eur. J. Pharmacol. 40, 101-106. Horton. R. W., Anlezark, G. M., Sawaya, M. C. B., and Meldrum, B. S. (1977). Eur. J. Pharmacol. 41, 387-397. Horton, R. W., Chapman, A. G., and Meldrum, B. S. (1978).J. Neurochm. 30, 1501-1504. Horton, R. W., Chapman, A. G., and Meldrum, B. S. (1979a).J. Neurochem. 33, 745-749. Horton, R. W., Collins, J. F., Anlezark, G. M., and Meldrum, B. S. (1979b). Eur. J . Pharmacol. 59, 75-83. Hrachovy, R. A., Frost, J. D., Kellaway, R., and Zion, T. (1979). Epilepsia 20, 403-411. Huggins, A. K., and Nelson, D. R. (1975).J. Neurochem. 25, 117-121. Humphrey, G . (1942).J. C m p . Physiol. Psvchol. 33, 315-323. Hunt, A. D., Stokes, J., McCrory, W. W., and Stroud, H. H. (1954).Pediatrics 13, 140-145. Hwang, E. C., and Van Woert, M. H. (1978). Neurology 28, 1020-1025. Hwang, E. C., and Van Woert, M. H . (1979). Neuropharmacology 18, 391-397. Iadarola, M. J., and Gale, K. (1980). Adv. Epileptol., Epilepsy Int. Symp., l l t h , 1979 pp. 449-455. Ikonomnff, S. I. (1970). I3r.J. Pqchiatq 117,679-680. Iles, J. F., and Jack, J. H. B. (1980). Brain 103, 555-578. ltal, T. M., and Soldatos, C. (198O).JAMA, J. Am. Med. Asoc. 244, 1460- 1463. Ito, M.. and Yoshida, M. (1964).Expm‘entia 20, 515-516. Ito, M., Yoshida, M., and Obata, K. (1964). Experimtia 20,575-576. Izumi, K., Donaldson, J., Minnich, J., and Barbeau, A. (1973). Can.J. Physiol. Pharmacol. 51, 885-889. Izumi, K., Igisu, H., and Fukuda, T. (1974).Brain Reg. 76, 171-173. Jacobowitz, D. M., and Pdkovits, M. (1974). J. Comp. Nmrol. 157, 13-28. Jacquet, Y. (1980). S c i m e 210, 95-97. Javoy, F., Sotelo, C., Herbert, A., and Agid, Y. (1976). Brain Res. 102,201-215. Jeavons, P. M., and Bower, B. D. (1964). “Infantile Spasms.” Heinemann, London. Jeavons, P. M., Bower, B. D., and Dimitrakoudi, M. (1973). Epllepsia 14, 153-160. Jenner, P., Chadwick, D., Reynolds, E.H., and Marsden, C. D. (1975).]. Pharm. Pharmacol. 27, 707-710. Jenney. E. H., and Pfeiffer, C. C. (1956). Ann. N.Y. Acad. Sn’. 64,679-689.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

165

Jobe, P. C., Picchioni, A. L., and Chin, L. (1973a).J. Pharmacol. Exp. Ther. 184, 1-10. Jobe, P. C., Picchioni, A. L., and Chin, L. (1973b).J. P h r m . Pharmacol. 25,830-831. Jobe, P. C., Picchioni, A. L., and Chin, L. (1973~). LiJe Sci. 13, 1-13. Jobe, P. C., Stull, R. E., and Geiger, P. F. (1974). Neuropharmarology 13, 961-968. Jobe, P. C., and Laird, H. E. (1981).Biochem. Pharmacol. 30,3137-3144. Johnston, D., and Brown, T. H. (1981). Science 211, 294-297. Johnston, D. D., Davies, H. L., and Crawford, R. D. (1979). Fed. Proc., Fed. Am. SOC.Exp. Biol. 38, 2417-2423. Johnson, D. D., Jaju, A. T., Ness, L., Richardson, J. S., and Crawford, R. D. (1981). Can.J. Physiol. Pharmacol. 59, 144- 149. Johnson, E. S., Roberts, M. H. T., and Straughn, D. W. (1970). Br. J . Pharmacol. 38, 659-666. Johnson, J. L. (1972).Brain Res. 37, 1-19. Johnston, G. A. R. (1978). In “Receptors in Pharmacology” R. Smythies and R. J. Bradley, eds.), pp. 295-333. Dekker, New York. Johnston, G. A. R. (1979). I n “Glutamic Acid; Advances in Biochemistry and Physiology” (L. J. Files, M. R. Kane, S. Garattini, and W. A. Reynolds, eds.), pp. 177-185. Raven Press, New York. Jonec, V., and Wasterlain, C. G. (1981). Neurology 31(2), 157. Jones, B. J., and Roberts, D. J. (1968). Br. J . Pharmacol. 34, 27-31. Joseph, M. H., and Emson, P. C. (1976).J. Neurochem. 27, 1495-1501. Joy, R. M., Stark, C. G., Gordon, C. S., Peterson, S. L., and Albenson, T. E. (1981). Exp. Neurol. 73, 588-594. Julien, R. M., and Laxer, M. D. (1974). Electroencephalogr. Clin. Neurophysiol. 37, 123127. Jung, M. J., Lippert, B., Metcalf, B. W., Schecter, P. J., Bohler, P., and Sjoerdsma, A. (1977a).J . Neurochem. 28, 717-723. Jung, M. J., Lippert, P., Metcalf, B. W., Bohler, P., and Schecter, P. J. (1977b).J. Neurochem. 29,797-802. Kanazawa, I., Miyata, Y., Toyokura, Y., and Otsuka, M. (1973). Brain Res. 51, 363-365. Kaplan, H. (1975). LiJe Sci. 17, 693-698. Kaplan, H., and Miezejeski, C. (1972).J. Comp. Physiol. Psychol. 81, 267-273. Karczmar, A. G. (1979). I n “Brain Acetylcholine and Neuropsychiatric Disease” (K. L. Davis and P. A. Berger, eds.), pp. 265-310. Plenum, New York. Kebabian, J. W., and Calne, D. B. (1979). Nature (London) 277, 93-96. Keller, H. H., Bartholini, G., and Pletschen, A. (1973). Brain Res. 64,371-377. Keller, H. H., Schatfner, R., and Haefel, W. (1976).Naunyn-Schmiedeberg’s Arch. Pharmacol. 294, 1-7. Kellogg, C. (1976). Brain Res. 106, 87-103. Kellogg, C., and Wennerstrom, C. (1974). Brain Res. 79, 451-458. Kendall, D. A., Fox, D. A., and Enna, S. J. (1981). Neuropharmacology 20,351-355. Kerwin, R. W., and Olpe, H. R. (1980). Br. J . Pharmacol. 70, 76. Kerwin, R. W., and Taberner, P. V. (1981). Gen. Pharmacol. 12, 71-75. Kety, S. S., Javoy F., Thievry, A., Julou, &,, and Glowinski, J. (1969). Proc. Natl. Acad. Sci. U.S.A. 58, 1249-1254. Kilian, M., and Frey, H. H. (1973). Neurqbharmacology 12,681-692. Killam, E. K. (1976). Med. Proc. 35, 2264-2269. Killam, E. K., and Suria, A. (1980).In “Antiepileptic Drugs: Mechanisms of Action” (G. H. Glaser, J. K. Penry, and D. M.Woodbury, eds.), pp. 597-615. Raven Press, New York. Killam, K. F. (1957).J. Pharmacol. Exp. Ther. 119, 263-271.

u.

166

0. CARTER SNEAD 111

Killam, K. F., and Bain, J. A. (1957).J. Pharmacol. Exp. Ther. 119, 255-262. Killam, K. F., Killam, E. K., and Naquet, R. I. (1967).Electroencephalogr. Clin. Neuruphysiol. 22,497-513. Kimura. D. (1962). Elertroencephalogr. Clin. LV~urophysiol.14, 115- 122. King, G. A. (1979). Seurophnrmacolog?. 18,47-55. King, G . A., and Burnham, W. M. (1980). Psychopharmacology 69, 281-285. King, G. A., Burnham, W. M.. and Livingston, K. E. (1980). Epilepsia 21, 531-539. King, W. M., and Kreisman, N. R. (1981). Brain Res. 209, 221-226. Kitai, S. T., Sugimori, M., and Kocsis, J. D. (1976). Exp. Brain Res. 24, 351-363. Kizer, J. S., Nerneroff, C. B., and Youngblood, W. W. (1978). Phannacol. Rev. 29,301-318. Klawans, H. L., Jr., Goetz, C., and Weiner, W. J. (1973). Neurology (Minneap.) 23, 12341240. Klein, R., and Livingston, S. (1950).J. Pediatr. 37, 733-740. Kleinrok, Z., Czuczwar, S. J.. and Kozicka, M. (1980). Ep’lepsia 21, 519-529. Klunk, W.E., and Ferendelli, J. A. (1980). Neurolog 30,421. Kobayashi. R. M., Brownstein, M., Saavedra, J . M., and Palkovits, M. (1975).J. Neu?-ochem. 24,637-640. Kobayashi, R. M.. Palkovits, M., Hruska, R. E., Roth-Schild, R., and Yamamura, H. I. (1978). Brnin Kv.\. 154, 13-23. Arch. Exp. Pathol. Pltarmakol. 233, 589-566. Kobinger, W.(1958). ,~‘ntinsn-SclimiPdPb~~s Koe, B. K., and Weissman, A. (1966).J. Pharmcol. Exp. Ther. 154, 499-516. Koe, B. K., and Weissman, A. (1968).Adzr. Pharmacol. 6B, 29-47. Koelle, G. B. (1949). Proc. Soc. Exp. Biol. .\.led. 7, 617-622. Koelle, G . B. (1954). I. Comp. Neurol. 100, 211-228. Kohsaka, M., Hiramatsu, M:, and Mori, A. (1978).Adz!. Biochum. P.\ychopharmacol. 19, 389392. Kontro, P., Marnek, K. M., and Oja, S. S . (1980). Brain Res. 184, 129-141. Kopeloff, L. M., Banera, J. E., and Kopeloff, N. (1942). Am. J. Psychiatry 98, 881-902. Koslow, S. H., and Roth, C. J. (1971).J. Pharmacol. Exp. T k . 176, 711-717. Kosterlitz, H. W., and Leslie, F. M. (1978). Br. J . Pharmacol. 64, 607-614. Kosterlitz, H. W., Lord, J. A. H., Paterson, S. J., and Waterfield, A. A. (1980). Br. J. Phiinnacol. 68, 333-342. Kostrzewa. R. M., and Jacobowitz. D. M. (1974). Pharmacol. R a ! . 26, 200-288. Koujoumdjian, J. C., and Ebadi, M. (198l).J. Neurochem. 36, 251-257. Kovacs, D. A., and 2011, J. G. (1974). Brain R P ~70, . 165-169. Koyarna, I. (1972). Can. J. PhySiol. Pharmacol. 50, 740-752. Kozhechkin, S. N. (1980). Bull. Exp. Biol. .\led. (Engl. Transl.) 88, 1293-1296. Kozhechkin, S. N. (1981). Arih. Int. P h a r m r o d y . Ther. 250, 242-253. Krall, R. I . (1980). I n “Antiepileptic Drugs: Mechanisms of Action” (G. H. Glaser, J. K. Penry, and D. M. Woodbury, eds.), pp. 233-303. Raven Press, New York. Krali, R. I., Perry. J. K.,White, B. G., Kupferberg, H. J., and Swinyard, E. A. (1978). Ejniepsin 19, 409-428. Krieger, D. L., and Liotta, A. S. (1979). Siimce 205, 366-374. Kristiansen, K., and Cunois, G. (1949). Electroewephalogr. Clin. h’europhysiol. 1, 265-272. Krnjevic, K. (1976). I n “GABA in Nervous System Function” (E. Roberts, T . N. Chase, and D. B. Tower, eds.), pp. 269-281. Raven Press, New York. Krnjevic, K., and Phillis, J. W. (1963).j. Phvsiol. (London) 165, 274-304. Krnjevic, K., and Reinhardt, W. (1979). Science 206, 1321-1324. Krnjevic, K., and Schwartz, S. (1967). Exp. Brain Res. 3, 320-336. Krnjevic, K.,and Silver, A. (1965).J. Anat. 99, 711-719.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

167

Krogsgaard-Larsen, P., Hjeds, H., Curtis, D. R., Lodge, P., and Johnston, G. A. R. (1979).J. Neurochem. 32, 1717-1724. Kubrin, S., and Seifter, J. (1966).J . Pharmacol. Exp. Ther. 154, 646-651. Kuhar, M. J., and Snyder, S. H. (1970).J. Pharmacol. Exp. Ther. 171, 141-152. Kuriyama. K., Roberts, E., and Rubinstein, M. K. (1966).Biochem.Phamacol. 15,221-236 Kurokawa, M., Machiyama, Y., and Kato, M. (1963).J . Neurochem. 10, 341-348. Lacy, J. R., and Penry, J. K. (1976). “Infantile Spasms.” Raven Press, New York. Laird, H. E., and Huxtable, R. J. (1978). In “Taurine and Neurological Disorders” (A Barbeau and R. J. Huxtable, eds.), pp. 339-357. Raven Press, New York. Lambertsen, C. J. (1966). In “Fundamentals of Hyperbaric Medicine,” pp. 12-20. Natl Acad. Sci.-Nat. Res. Counc., Washington, D.C. Lance, J. W., and Adams, Z. D. (1963).Brain 86, 111-136. Langer, S. Z. (1977).Br. J. Pharmacol. 60,481-497. Lapin, I. P. (1981). Epilepsia 22, 257-265. LaSalle, G. (1980). Can.J. Physiol. Pharmacol. 58, 7-1 1. Law, P., Loh, H., and Li, C. (1979).Proc. Natl. Acad. Sci. U.S.A. 76, 5455-5461. Laxer, K. D., Sourkes, T. L., Fang, T. Y., Young, S. N., Gauthier, S. G., and Missala, K. (1979). Neurology 29, 1157-1161. Leeb-Lundberg, F., Snowman, A., and Olsen, R. W. (1981).Eur.J. Phamacol. 72, 125-129. Lefvert, A. K., Bergstrom, K., Matell, G., Osterman, P. O., and Pirskanen, R. (1978).J. Neurol., Neurosurg. Psychiatry 41, 394-403. Lehmann, A. (1967).L$e Sci. 6, 1423-1431. Lehmann, A. (1977).Life Sci. 20,2047-2056. Lenicque, P. M., Welpierre, J., and Cohen, Y. (1979).Psychopharmacology 66, 51-53. Lewin, E. (1972). In “Experimental Models of epilepsy-A Manual for the Laboratory Worker” (D. P. Purpura, J. K. Penry, D. Tower, D. M. Woodbury, and R. Walter, eds.), pp. 13-49. Raven Press, New York. Lewin, E., Charles, G., and McCrimmon, A. (1969). Neurology 19, 565-569. Lewis, J. W., Cannon, J. T., and Liebeskind, J. C. (1980).Science 208, 623-624. Lewis, P. R., and Shute, C. C. D. (1967).Brain 90, 521-540. Lewis, P. R., and Shute, C. C. D. (1978). In “Handbook of Psychopharmacology” (L. L. Iversen, S. D. Iversen, and S. H. Snyder, eds.), Vol. 9, pp. 315-355. Plenum, New York. Lewis, P. R., Shute, C. C. D., and Silver, A. (1964).J. Physiol. (London)172, 9-10. Lewis, P. R., Shute, C. C. D., and Silver, A. (1967).J. Physiol. (London) 191, 215-224. Lhermitte, F., Peterfalvi, M., Marteau, R., Gazengel, J., and Serdara, M. (197 1). Rev. Neurol. 124, 21-31. Lhermitte, F., Marteau, R., and Degos, C. F. (1972).Rev. Neurol. 126, 107-114. Lhermitte, F., Degos, C. F., and Marteau, R. (1975).Nouv. Presse Med. 4, 31. Li, C. L. (1959).J. Neurophysiol. 22,436-450. Lidbrink, P., Corrodi, H., and Fuxe, H. (1974).Eur. J . Pharmucol. 26, 35-40. Liebman, J. M., Pastor, G., Bernard, P. S., and Saelens, J. K. (1980). Life Sci. 27, 19911998. Lindvall, 0.(1979).I n “The Neurobiology of Dopamine” (A. S. Horn, J. Korf, and B. H. C. Westerink, eds.), pp. 3 19-342. Lindvall, O., and Bjorklund, A. (1978). In “Handbook of Psychopharmacology” (L. L. Iversen, S. D. Iversen, and S. H. Snyder, eds.), Vol. 9, pp. 139-231. Plenum, New York. Lipp, J. A. (1972). Electroencephalogr. Clin. Neurophysiol. 32, 557-565. Lipp, J. A. (1973).Arch. Int. Pharmacodyn. Ther. 202, 244-251.

168

0.CARTER SNEAD 111

Lipp, J. A. (1974). Arch. Int. Pharmacodjn. T l m . 210,49-54. Lippa, A. S., Klepener, C. A,:Bensor, D. I., Critichett, D. J., Sano, M. C., and Beer, B. (1980). Brain Res. Bull. 5, Suppl. 2, 861-865. Livingston, S., Berman, W., and Pauli, L. L. (1973). Pediatrics 52, 753-754. Livingston, S., Berman, W., and Pauli, L. L. (1974). Pediatrics 53, 952-953. Livrea, P. (1976). Acta Neurol 31, 580-600. Livrea, P., Direda, L., and Papagna, G. (1976). Acta Neurol. 31, 632-636. Lloyd, K. G., and Dreksler, S. (1979). Brain Res. 163, 77-87. Lloyd, K. G., and Worms, P. (1981). Adv. Biochem. Psychqbhannacol. 29, 59-67. Lloyd, K. G., Munari, C., Worms, P., Bossi, C., Bancaud, J., Talairach, J., and Morselli, P. C. (1981). Adv. B i o c h n . Psychqbharmacol. 26, 199-206. Lockard, J. S., Uhlir, V., Ducharme, L. C., Farquhar, J. A., and Huntsman, B. J. (1975). Epikpsia 16,301-317. Lockard, J. S., Ojemann, G. A., Congdon, W. C., and Ducharme, L. L. (1979).Epilepsia 20, 223-234. Logan, W. J., and Snyder, S. H. (1972). Brain Res. 42, 413-431. Logothetis, J. (1955). il$urology 5, 236-241. Logothetis, J . (1967). Neurolog). 17,869-877. London, E. D., and Buterbaugh, G. G. (1978).]. Pharmacol. Exp. Ther. 206,81-90. Longo, V. G. (1956).]. P a h a m c o l . 116, 198-208. Longo, V. G. (1966). Pharmacol. Rev. 18, 965-996. Longo, V. G., Nechmansoa, D., and Bovet, D. (1960). Arch. Int. Pharmacodyn. Ther. 123, 282-290. Longoni, R., Mulas, A,, Pepeu, I. M., and Pepeu, G. (1976a). Eur. J . Pharmacol. 40, 329332. Logoni, R., Mulas, A., Novak, B. O., Pepeu, I. M., and Pepeu, G. (1976b).Neurophamcolo a 15,283-286. Lord, J. A. H., Waterfield, A. A , , Hughes, J., and Kosterlitz, H. W. (1977).Nature ( L a d o n ) 267,495-499. Loscher, W. (1979). Biochem. Phannacol. 28, 1397-1407. Loscher, W. (1980a). Naunyn-Schmiedeberg’.r Arch. Pharmacol. 315, 119- 128. Loscher, W. (1980b). Arch. In!. Pharmacodyn. Ther. 243, 48-55. Liischer, W., and Frey, H. H. (1977). Naunyn-Schmiedebergs Arch. Pharmakol. 269,263-296. Liischer, W., and Frey, H. H. (1978). Biochem. Pharmacol. 27, 103-108. Loskota, W. J., and Lomax, P. (1975). Elecfromephalogr. Clin. Neurophysiol. 38, 597-604. Lothman, E. W., and Collins, R. C. (1981). Brain Res. 218, 299-318. Lothman, E. W., Collins, R. C., and Ferendelli, J. A. (1981). Neurology 31, 806-812. Lott, 1. T., Coulomke, T., DiPaolo, R. V., Richardson, E. P., and Levy, H. L. (1978). iVeurolog). 28,47-54. Lovell, R. A. (1971). In “Handbook of Neurochemistry” (A. Lajtha, ed.), Vol. 6, pp. 63102. Plenum, New York. Lovell, R. A., ELliott, S. I., and Elliott, K. A. C. (1963).]. Neurochem. 10, 479-488. Low, N. L. (1958).Pedintrics 22, 1165-1174. Lust, W. D., Goldberg, W. D., and Passoneau, J. V. (1976).J. Neurochm. 26, 5-10. Lust, W. D., Fuessner, G. K., Passoneau, J. V., and McCandless, 0. W. (1981). In “Neuropharmacology of Central Nervous System and Behavioral Disorders” (G. C. Palmer, ed.), pp. 407-431. Academic Press, New York. Lynch, G., Matthews, D. A., Mosco, S., Parks, T., and Cotman, I. (1971). Brain Rts. 42, 311-318. McCandless, D. W., Feussner, G. W., Lust, W. D., and Passoneau,J. V. (1979).J. Ne-urochem. 32,743-753.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

169

Macdonald, R. L., and Barker, J. C. (1977). Nature (London) 267,720-721. Macdonald, R. L., and Barker, J. C. (1978a). Neurology 28, 325-330. Macdonald, R. L., and Barker, J. C. (1978b). Science 200, 775-777. Macdonald, R. L., and Barker, J. C. (1979a).Neurology 29,432-447. Macdonald, R. L., and Barker, G. K. (1979b). Brain Res. 170, 558-562. Macdonald, R. L., and Barker, J. L. (1977). Ann. Neurol. 2, 264. Macdonald, R. L., and Bergey, G. K. (1979). Brain Res. 190, 558-562. MacFarland, D., and Wolner, A. (1965). Life Sci. 4, 1587-1590. McGeer, E. G., and McGeer, P. L. (1976).J. Neurochem. 26, 65-76. McGinty, D. J., and Harper, R. M. (1976). Brain Res. 101, 569-575. McIntyre, D. C., Saari, M., and Pappas, B. A. (1979). Exp. Neurol. 63, 527-544. McKenzie, G. M., and Soroko, F. E. (1972).J . Phann. Pharmacol. 24,696-701. McLennon, H., and Elliott, K. A. C. (1951).J. Pharmacol. Exp. Ther. 123, 35-43. McLennon, H., Hicks, T. P., and Hall, J. G. (1981). Adv. B i o c h a . Psychopharmacol. 29, 2 13-221. McMillen, B. A,, and Isaac, L. (1978a). Biochem. Pharmacol. 27, 1815-1820. McMillen, B. A., and Isaac, L. (1978b). Brain Res. 150,424-430. McNamara, J. 0. (1978). Brain Res. 154, 415-420. McQuarrie, J., Anderson, J. A., and Ziegler, M. R. (1942).J. Clin. Endocrinol. 2,406-412. Madison, S. (1977). Physiol. Behav. 19, 819-830. Magnussen, I., Dupont, E., Hansen, A., Prange, A., and DeFine Olivarius, B. (1977). Acta Neurol. Scand. 55,251-252. Magnussen, I., Dupont, E., Engbaek, F., and DeFine Olivarius, B. (1978). Acta Neurol. Scand. 57,289-294. Magnusson, T. (1973). Naunyn-Schmiedeberg’s Arch. Pharmacol. 278, 13-22. Mao, C. C., Guidotti, A., and Costa, E. (1974a). Brain Res. 79, 510-514. Mao, C. C., Guidotti, A., and Costa, E. (1974b). Mol. Pharmacol. 10, 736-745. Mao, C. C., Guidotti, A., and Landis, S . (1975). Brain Res. 90, 335-339. Mardani, M. G., Stanziona, P., Cherubini, E., and Bernardi, G. (1980). Neurosci. Lett. 18, 169-172. Marcus, E. M., Watson, C. W., and Goldman, P. C. (1966). Arch. Neurol. (Chicago) 15, 52 1-532. Marcus, E. M., Fullerton, A., Losh, E., and Bowler, R. (1972). Epilepsia 13,343-344. Mares, P., Zouhar, A., and Brbzek, G. (1979). Act. Neru. Super. 21, 218-225. Martin, I. C., and Candy, J. M. (1978). Neuropharmacology 17, 993-998. Martin, W. R., Eades, C. G., Thompson, J. A., Hppla, R. E., and Gilbert, P. E. (1976). J . Pharmacol. Exp. Ther. 197, 517-532. Mason, S. T., and Corcoran, M. E. (1979a). Science 203, 1265-1266. Mason, S. T., and Corcoran, M. E. (197913). Brain Res. 170,497-507. Mathers, D. A., and Barker, J. G. (1981). Brain Res. 204, 242-247. Matsuda, M., Abe, M., Hoshino, M., and Sakurai, T. (1979). B i o c h a . Pharmacol. 28,27852789. Matsumoto, H., and Ajmone Marsan, C. (1964). Exp. Neurol. 9, 286-304. Matsumoto, H., Azola, G. F., and Gummit, R.J. (1969).J. Neurophysiol. 32, 688-703. Matthews, W. D., and McCafferty, G. P. (1979). Neuropharmacology 18, 885-889. Matthews, W. D., Intoccia, A. P., Osbourne, V. L., and McCafferty, G. P. (1981a). Eur. J. Pharmacol. 69, 249-254. Matthews, W. D., McCafferty, G. P., and Setler, P. E. (1981b). Neuropharmacology 20, 561565. Maxson, S. C., and Cowen, J. S. (1976). Physiol. Behav. 16, 623-629. Maynert, E. W. (1969). Epilepsia 10, 145-162.

170

0. CARTER SNEAD 111

Maynert, E. W., Marczynski, T. J., and Browning, R. A. (1975). A&. h’rurol. 13, 79-147. . 114, 214-215. Mazars, T., Mazars, G., and Piut, C. (1966). R ~ JNeurol. Meister, A. (1979). I n “Glutamic Acid: Advances in Biochemistry and Physiology” (L. J. Filer, M.R. Kane, S. Garchini, and W. A. Reynolds, eds.), pp. 69-84. Raven Press, New York. Meldrum, B. S . (1975). Ir7f. R m . Seirrobiol. 17, 1-36. Meldrum, B. S. (1978). Lnnret 2, 304-306. Meldrurn, B. S. (1979). I n “GABA-Neurotransmitters” (P. Krogsgaard-Larsen, J. Scheelakrueger, and H. Kofod, eds.), pp. 390-405. Academic Press. New York. Meldrum, B. S. (1981). .4dv. Biorhem. Pswhophnrmaroi. 26, 207-217. Meldrum. B. S., and Horton, R. W. (1971). Brnin Res. 35, 419-436. Meldrurn, B. S., and Horton, R. W. (1978). P.iyhophnrmacolo~59, 47-50. Meldrum, B. S., Naquet, R., and Balzano, E. (1970). Elecfroenrephalogr. Clin. Neurophysiol. 28,449-458. Meldrum, B. S., Anlezark, G., and Trimble, M. (1975). E u r . ] . Phannroi. 32, 203-213. Meldrum, B. S . , Pedley, T.. Horton, R.. Anlezark, G., and Franks, A. (1980). Brain Res. Hid/. 5, Suppl. 2, 685-690. Mendez, J . S., Cotzias, G.C., Mena, I., and Papavasilion, P. S. (1975).Arch. LVeuro/.(Chicago) 32,44-46. Menini, C., Meldrum, B. S., Riche. D., Silva-Compte, C., and Stutzrnann, J. M. (1980). A n t i . Srrc,-o/. 8, 501-509. Menkes, J. H. (1973). Pediafrirs 53, 952. Menon, M. K. (1981). Lije Sri. 28, 2865-2868. l o ~ ~441-444. Menon. M . K., and Vivonia, C. A. (1981). . ~ e u r o p h a r , ~ z a c o 20, Metcalf, B. W. (1979). Hiochem. Phnrtnacol. 28, 1705-1712. Meyer, H., and Frey, H. H. (1973). ,Veuropharnincolo~ 12, 939-947. Michelis, E. K., Michelis, M. L., and Boyarsky, C. C. (1974). Biochim. Biqpliys. Acta 367, 338-348. Miller, -4. L. (1981). I,!/. R e ) . S ~ ~ r ( ~ b i22, o 1 .47-82. Miller, A. L.. and Pitts, F. N.,Jr. (1967). .Yeir~-orheini~tr~ 14, 579-584. Miller. F. R., Stavrakp. G. W., and Wmton, G. A. (1940).J. Seurvphy.&/. 3, 131-138. Miller, R. J. (1981). Phnnnncol. Thw. 12, 73-108. Millichap, J. G.. and Bickford, R. G. (1962)../AdM, J . Am. ,\.led. A.tror. 182, 523-528. Millichap. J. G., Pitchford, G. L., and Millichap, M. G. (1968). Proc. Sor. Exp. B i d . M e d . 127, 1187-1190. Minneman, K. P., and Molinoff, P. B. (1980). Biorhem. Phnrmrol. 29, 1317-1323. Minneman, K. P., Pittman, R. N., and Molinoff. P. B. (1981). ,4nnu. Rat. Nmrosri. 4, 419-462. Mirsky, A., Miller, R., Stein, M. (1953). P.syhusom. ‘\fed. 15, 574-579. Mohler, H., and Okada, T. (1977). Science 198, 849-851. Mohler, H., and Okada, T. (1978). ‘\fo/. Phnnnnrol. 14, 256-2155, Mohr. E., and Corcoran, M. E. (1981). Esp. LYeuro/.72, 507-511. Moir, A. T. B., Ashcroft, G. W., Crawford, T. B. B., Eccleston, D., and Guldberg, H . C. (1970). HI-nin 93, 357-368. Montaroio, P. G.. Raschi. F., and Struata, P. (1979). Brnin Res. 162, 358-362. Montplaisir, J., Saint-Hilaire, J. M., Walsh. J. T., Laverdiere, M., and Bouvier, G. (1981). .Veuro/og 31, 350-352. Moore, R. Y.,and Bloom, F. E. (1978). Annu. Rnr. ,\’eurwi. 1, 129-160. Moore, R. Y., and Bloom, F. E. (1979). 14uuu. RRJ.,\’curosri. 2, 113-168. Morgan, W. W. (1976). Expetimtia 32, 489-499. Morley, B. J., and Snead, 0. C. (1979). Tron.c.r l m . SOC.Nmrochem. 11, 188.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

171

Morley, B. J., Lorden, J. F., Brown, G. B., Kemp, G. E., and Bradley, R. J. (1977).BrainRes. 134, 161-166. Morley, B. J., Kemp, G. E., and Salvatera, P. (1979). L f e Sci. 24, 859-872. Morley, J. E. (1980). Life Sci. 27, 355-367. Morselli, P. C., Bossi, G., Henry, J. F., Zarifian, E., and Bartholini, G. (1980).Brain Res. Bull. 5, Suppl. 2, 411-414. Mortier, W., Schenk, K., and Kleu, G. (1971).Nmenarzt 42,498-501. Moshe, S . L., Sharpless, N. S., and Kaplan, J. (1981). Brain Res. 211, 190-193. Moskowitz, M. A,, and Wurtman, R. J. (1975). N . Engl. J. Med. 293, 332-338. Muller, W. E., Fehske, K. J., Borbe, H. O., Wollert, U., Nanz, C., and Rommelispacher, H. (1981). Pharmacol., Biochem. Behav. 14, 693-699. Murakami, Y., Abe, M., and Murakami, K. (1976).J. Neurochem. 26, 655-656. Mutani, R. (1967a). Epilepsia 8, 73-92. Mutani, R. (1967b).Epilepsia 8, 223-240. Mutani, R., Bergamini, L., Fairello, R., and Delsedime, M. (1974a). Brain Res. 70, 170173. Mutani, R., Bergamini, L., Delsedime, M., and Durelli, L. (1974b). Brain Res. 79, 330332. Mutani, R., Bergamini, L., and Durelli, L. (1978).In “Taurine and Neurological Disorders” (A. Barbeau and R. J. Huxtable, eds.), pp. 359-373. Raven Press, New York. Myllya, U. U., Keikkoner, E.R., Vapaatalo, H., and Hokkanen, E. (1975). Eur. Neural. 13, 123- 130. Myslobodsky, M. S. (1976). “Petit Ma1 Epilepsy.” Academic Press, New York. Myslobodsky, M. S., and Vallenstein, E. S. (1980). Epilepsia 21, 163-175. Myslobodsky, M. S., Ackermann, R. F., and Engel, J. (1979).Phannacol.,Biochem. Behav. 11, 265-27 1. Nadler, J. V. (1979).Life Sci. 24,289-300. Nadler, J. V., Vaca, K. W., White, W. F., Lynch, G. S., and Cotman, C. W. (1976). Nature ( L m d m ) 260, 538-540. Nahorski, S. R. (1972).J.Neurochem. 19, 1937-1946. Nakai, Y., and Takaori, S. (1974). Brain Res. 71, 47-60. Naquet, R., and Meldrum, B. S. (1972).In “Experimental Models of Epilepsy-A Manual for the Laboratory Worker” (D. P. Purpura, J. K. Penry, D. Tower, D. M. Woodbury, and R. Walter, eds.), pp. 374-406. Raven Press, New York. Naruse, H., Kato, M., Kurokawa, M., Haba, R., and Yabe, T. (1960). J . Neurocha. 5, 359-369. Nicoll, R. A. (1975). Proc. Natl. Acad. Sci. U.S.A. 72, 1460-1463. Nicolou, N. (1967). Foliu Med. (Plovdiv) 9,249-254. Nielsen, M., Gredal, O., and Braestrup, C. (1979). Life Sci. 25, 679-686. Nistri, A., and Constanti, A. (1978). Neuropharmacology 17, 127-135. Nitsch, C. (1981). Adv. Biochem. Psychophamcol. 29, 97-104. Nitsch, C., Kim, J. K., Shimada, C., and Okada, Y. (1979). Neurosci. Lett. 11, 295-299. Noebels, J. L. (1979). Fed. Proc., Fed. Am. SOC.Exp. Biol. 38, 2405-2410. Noebels, J. L., and Sidman, R. L. (1979). Science 204, 1334-1336. Nordberg, A., and Sundwall, A. (1977a).Acta Physiol. Scand. 99, 336-344. Nordberg, A., and Sundwall, A. (1977b).In “Cholinergenic Mechanisms and Psychopharmacology” (D. J. Jenden, ed.), pp. 629-641. Plenum, New York. North, R. A. (1979). Life Sci. 24, 1527-1546. Oakley, N. R., and Jones, B. J. (1980). Eur. J. Pharmucol. 69, 381-382. Obata, K., and Takeda, K. (1969).J.Neurochem. 16, 1043-1047. Obata, K., Ito, M., &hi, R., and Sato, N. (1967). Exp. Brain Res. 4,43-57.

172

0. CARTER SNEAD 111

Obata, K., Takeda, K., and Shinozaki, H. (1970). Exp. Brain Res. 11,327-342. OBrien, R. A., Schlosser, W., Spirt, N. M., Franco, S. M., Horst, W. D., Polo, P., and Bouetti, E. P. (1981). Life Sn‘. 29, 75-82. Oishi, R., Suenaga, N., and Fukuda, T. (1979). Phunnacol., Biochem. Behav. 10, 57-61. OLeary, J. L., and Goldring, S. (1976). “Science and Epilepsy,” Raven Press, New York, p. 16. Okada, K. (1971).J . Neurophysiol. Exp. Neurd. 30, 120-134. Okada, K. (1973). Psychiatr. Neurd. Jpn. 75, 397-408. Oliver, A. P., Hoffer, B. J., and Wyatt, R. J. (1980). Science 210, 1264-1265. Olney, J. W., Rhea, V., and Ho, 0. G. (1974). Bruin Res. 77, 507-512. Olpe, H. R., and Koella, W. P. (1979). Eur. J . Phurtnucol. 53, 359-364. Olsen, R. W. (198l).J. Neurochern. 37(1), 1-13. Olsen, R. W., and Leeb-Lundberg, F. (1981). Adv. Biochm. Psychopharmucol. 26, 93-102. Olsen, R. W., Ban, M., and Miller, T. (1976). Bruin Res. 102, 283-299. Olsen, R. W., Ticku, M. K., Van Ness, P. C., and Greenlee, D. (1978). Brain Res. 139, 277-294. Olsen, R. W., Leeb-Lundberg, F., and Napias, C. (1980). Brain Kes. Bull. 5, Suppl. 2, 2 17-22 1. Olson, L., Freedman, R., Seiger, A., and Hoffer, B. (1977). Bratn Res. 119,87-106. Osorio, I . , and Davidoff, R. A. (1978). Ann. Neurol. 6, 111-1 16. Palacios, J. M., and Kuhar, M. J. (1980). Science 208, 1378-1380. Palacios, J. M., Wamsley, J. K.. Zarbin, M. A., and Kuhan, M. J. (1981). Adv. Biochem. Psychophannacol. 29, 445-451, Palkovits, M., Saavedra, J. M., Kobayashi. R. M., and Brownstein, M. (1974). Brain Res. 79, 443-450. Pallister, P. D. (1959). Rocky M t . Med. J. 56, 45-50. Palmer, G. C., Jones, D. J., Medina, M. A., and Stavinoha, W. B. (1979). Epilepsia 20, 95- 104. Papeschi, P., Molina-Negro, P., Sourkes, T. L., and Giuseppe, E. (1972). Neurology 22, 1151- 1159. Pappins, H. M., and Elliott, K. A. C. (1958).]. Appl. Psychol. 12, 319-323. Pasternak, G. W., Childers, S. R., and Snyder, S. H. (1980). Science 208, 514-517. Patsalos, P. N., and Lascelles, P. T. (1981). J . Neurocheni. 36(2), 688-695. Paul, S. M., Syapin, P. J., Paugh, B. A., Moncada, V., and Skolnick, P. (1979). Nature ( L a d o n ) 281,688-689. Pearse, A. G. E. (1978). In “Centrally Acting Peptides” (J. Hughes, ed.), pp. 49-57. Univ. Park Press, Baltimore, Maryland. Pedata, F., Mules, A,, Pepeu, I. M.. and Pepeu, G. (1976). Eur. J . Pharmacol. 40,329-332. Pedley, T. A., Horton, R. W.,and Meldrum, B. S. (1979). Epilcpsia 20,409-416. Pelletier, G . , LeClerc, R., Saavedra, J. M., Browntein, M. J., Vandry, H., Ferland, L., and Labria, F. (1980). Brain Res. 192, 435-443. Perry, T. L., and Hansen, S. (1981). Neurology 31, 871-876. Perry, T. L., Hansen, S., Kennedy, J., Wada, J. A,, and Thompson, G. B. (1975). Arch. Neurol. (Chicago) 32, 752-754. Perry, T. L., Kish, S. J., and Hansen, S. (1979).J . Neurochem. 32, 1641-1645. Perryman, K. M., Babb, T. L., Finch, D. M., Brown, W. J., and Crandall, P. H. (1980). Epi&p& 21,479-487. Pfeifer, A. K., and Galambos, E. (1967). Arch. Int. Phurtnucodyn. Ther. 165, 201-211. Phillis, J. W. (1978). In “Taurine and Neurological Disorders” (A. Barbeau and R. J. Huxtable, eds.), pp. 289-303. Raven Press, New York.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

173

Phillis, J. W., and York, D. H. (1967). Bruin Res. 5, 517-520. Pieri, C., and Haefely, W. (1976). Naunyn-Schmiedeberg’s Arch. Phannacol. 296, 1-4. Pineda, M. R., and Russell, S. R. (1974). Dis. N m . Syst. 35, 322-323. Pisa, M., Sanberg, P. R., Corcoran, M. E., and Fibiger, H. C. (1980). Brain Res. 200, 481-487. Pohl, M., Mares, P., and Fischer, J. (1980). Act. Neru. Super. 22(3), 214-215. Polc, P., Mohler, H., and Haefely, W. (1974). Naunyn-Schmeideberg’s Arch. Phannacol. 284, 3 19-397. Pollack, M. A., Zion, T. E., and Kellaway, P. (1979). Epilepsiu 20, 255-262. Pope, A., Morris, A. A., Jasper, H., Elliott, K. A. C., and Penfield, W. (1947).Res. Pub1.Assoc. Res. N m . M a t . Dis. 26, 218-231. Prichard, J. W. (1980).In “Antiepileptic Drugs: Mechanisms of Action” (G. H. Glaser, J. K. Penry, and D. M. Woodbury, eds.), pp. 505-522. Raven Press, New York. Prince, D. A. (1968). Exp. Neurol. 21, 467-485. Prince, D. A. (1978).Annu. Rev. Neurosci. 1, 395-415. Prince, D. A., and Farrell, D. (1969). Neurology 19, 309-310. Prince, D. A., and Wong, R. K. S. (1981). Bruin Res. 210, 323-333. Prockop, D. J., Shore, P. A., and Brodie, B. B. (1959). Expmkntia 15, 145-147. Pryor, G. T. (1968). Lije Sn’. 7, 867-874. Przegalinski, E. (1975). Pol. J . Pharmucol. 27, 195-199. Przegalinski, E. (1976a). Pol. J . Phannacol. 28, 143-155. Przegalinski, E. (197613).Arch. Immunol. Ther. Exp. 24, 821-827. Pumain, R., and Chauvel, P. (1975)..Exp.Bruin Res. 23, Suppl. 167. Pumain, R., Louvel, J., and Chauvel, P. (1978).In “Iontophoresis and Transmitter Mechanism in the Mammalian Central Nervous System” (R. W. Ryalt and J. S. Kelley, eds.), pp. 38 1-383. ElseviedNorth-Holland, Amsterdam. Purdy, R. E., Julien, R. M., Fairhurst, A. S., and Terry, M. D. (1977).Epilepsiu 18,251-257. Puro, D. G., and Woodward, D. J. (1973). Neuropharmucology 12,433-440. Purpura, D. P. (1969).In “Basic Mechanisms of the Epilepsies” (J. H. Jasper, A. A. Ward, and A. Pope, eds.), p. 481. Little, Brown, Boston. Purpura, D. P., Shoter, R. J., and Housepian, E. M. (1964). Prog. Bruin Res. 4, 187-197. Pycock, C. J., and Horton, R. (1978). B i o c h a . Phurmucol. 27, 1827- 1830. Pycock, C. J., and Kerwin, T. W. (1981). Life Sci. 28, 2679-2686. Pycock, C. J., Carter, C. J., and Kenvin, R. W. (1980).J. Neurocha. 34, 91-99. Pycock, C. J., Dawbarn, P., and Kerwin, R. W. (1981). Adv. B i o c h a . Psychqphurmacol. 29, 77-87. Quattrone, A., and Samanin, R. (1977). Eur. J . Pharmacol. 41, 333-336. Quattrone, A., Crunelli, V., and Samanin, R. (1978).Neurophamcology 17, 643-647. Quesney, L. F., and Gloor, P. (1978). Epile$sia 19, 35-45. Quiron, R., and Pert, C. B. (1981). Eur. J . Phamcol. 76,467-468. Raabe, W., and Ayala, G. F. (1976). Bruin Res. 195, 597-601. Racine, R. J. (1972). Electroacephulop. Clan. Neurophysiol. 32, 281-294. Racine, R. J., Burnham, W. M., and Livingston, K. (1979). Can.J. Neurol. Sci. 6,47-51. Randit, M., Siminoff, R., and Straughan, D. W. (1964). Exp. Neurol. 9, 236-242. Ransom, B. R., and Barker, J. L. (1975). Nature (London) 254,703-705. Ransom, B. R., and Barker, J. L. (1976). Bruin Res. 114, 530-535. Ransom, B. R., Neale, E., Henkant, M., Bullock, P. N., and Nelson, P. G. (1977a). J . Neurophysiol. 40, 1132-1 150. Ransom, B. R., Christian, C. N., Bullock, P. N., and Nelson, P. G. (1977b).J Neurophysiol. 40, 1151-1162.

174

0. CARTER SNEAD 111

Rassin, D. K. (1981). ,4dri. Riorltrm. Psjrhophnrntnrol. 29, 127- 134. Redhurn, D. A., Brooke, J., Ferkeny, J., and Heller, A. (1978). Brain Kes. 152, 51 1-517. Reeves, C. (1966). Psyrhol. Bull. 65, 325-335. Rehfeld, J. F. (1978a).J. B i d . Chem. 253, 4020-4022. Rehfeld, J. F. (1978h).J. Biol. Chem. 253, 4022-4027. Reid, J. L., Zivin, J. A,, Foppen, F. H., and Kopin, 1. J. (1975). Lifr Sci. 16, 975-984. Reiner, G. R., Grimm, R. J., and Dow, R. S. (1967). Elrrtroenrephalogr.Clin. Npurophyiol. 23, 456-462. Reynolds, E. H., Chadwick, D., and Jenner, P. (1975).J. Neurol. Sri. 26,605-615. Rihak, C. E., Harris, A. B., Vaughn, J. E., and Roberts, E. (1979).Scimir~205, 211-214. Richards, C. D. (1976). In “Biochemistry and Neurology” (H. F. Bradford and C. 0. Marsden, eds.), pp. 185- 193. Academic Press, New YorWLondon. Richter, D., and Crossland, J. (1949). .din. J . Phyiul. 159, 247-255. Riffee, W. H., and Gerald, M. C. (1976). .Vrziruphnrmarolo,p 15, 677-682. Roa. P. D., Tews, J . K., and Stone, W. E. (1964). Biochrin. Pharinacol. 13, 477-487. Roberts, E. (1974). Biorhrin. Pharmarol. 23, 2637-2649. Roberts. E. (1976). 111 “GABA in Nervous System Function” (E. Roberts, T. N. Chase, and D. B. Tower, eds.), pp. 514-540. Raven Press, New York. Roberts, E. (1978). I i t “Interactions Between Putative Neurotransmitters in the Brain” (S. Garattini, J . F. Pujol, and R. Samanin, eds.), pp. 89-107. Raven Press, New York. Roberts. E. (1980). I I I “Antiepileptic Drugs: Mechanisms of Action” (G. H. Glaser, J. K. Penry, and D. M. Woodbury, eds.), pp. 667-713. Raven Press, New York. Roberts, P. J. (1974). .Vntrtre (London) 252, 399-401. Roberts, P. J. (1981). ‘4d71.Biochrrn. P.sjrhopharmnro1. 29, 379-386. Rohison, G. A., Butcher, F. R. W., and Sutherland, E. W. (1971). “Cyclic AMP.” Academic Press, New York. Rommelspacher, H., Namz, C., Borbe, H. 0..Fehske, K. J., Muller, W. E., and Wolbert, U. (1980). ,~niiirun~ir-Sr/t~rPirrb~g’s .4rrh. Pharnmrol. 314, 94- 100. Rosenherg, P., and Echlin, F. A. (1965). A’rurolog~ 15, 700-707. Ross, J. M. (1978). Drri. .\led. Child Srui-ol. 20, 677-678. Roth, R. H. (1976).Pharinncol. Thrr., Part B 2, 71-88. Roth, R. H . , and Giarman, N. J. (1969). Hiorhem. Pharmarol. 18, 247-250. Roth, R. H., and Nowycky, M. C. (1977). Biochon. Pharmarol. 26, 2079-2082. Rothman, R. B., and Westfall, T. C. (1981). Ei1r.J. Phnrinnrol. 72, 365-368. Ruhenstein, 34.. Stein, S., and Udenfriend, S. (1977). Pror. h‘atl. A d . Sci. U.S.A. 75, 669-678. Rudeen, P. K., Philo, R. C., and Symmes, S. K. (1980). Epilqpsia 21, 149-154. Rudzik. A. D., and Johnson, G . A. (1970).111 “Amphetamines and Related Compounds” (E. Costa and S. Garattini, eds.), pp. 715-728. Raven Press, New York. Rudzik, A. D., and Mennear, J. H. (1966). Lifv Sri. 5, 747-756. Saad, S. F., Elmasry, A. M., and Scott, P. M. (1972). Eur. J. Phnrincirol, 17, 386-392. Said, S. I., Giachetti, A,, and Nicosia, J. (1980). ‘4dv. Riochem. P,sjrhophnnnnrol. 22, 75-84. Saito, A., Srandaran, H., Goldfine, I. D., and Williams, J. A. (1980). Srirncr 208, 11551157. Salmoiraghi, G. C., and Steiner, F. A. (1963).J. A‘nirophjsiol. 26, 581-597. Sato, M., and Nakashima, T. (1975). Con. J . Srurol. Sci. 2, 439-446. Sato, M., and Nakashima, T. (1976). I I I “Kindling” (J. Wada, ed.), pp. 103- 116. Raven Press, New York. Sato, M., Tomoda, T., Hikkesa, N., and Otsuki, S. (1980). Epilepxia 21, 497-507. Sattin, A. (1971).J. rVrttrorhrc. 18, 1087-1096.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

175

Schain, R. J. (1960). Yale J. Biol. Med. 33, 15-36. Schechter, P. J., and Grove, J. (1979). Brain Res. Bull. 4, 712-723. Schechter, P. J., and Grove, J. (1980). Brain Res. Bull. 5, Suppl. 2, 627-631. Schechter, P. J., Tranier, Y., Jung, M., and Sjoerdsma, A. (1977a).J. Pharmacol. Exp. Ther. 201,606-612. Schechter, P. J., Tranier, Y., Jung, M. J., and Bohlew, P. (1977b). Eur. J . Pharmacol. 45, 3 19-328. Schlesinger, K., Boggan, W., and Freedman, D. (1965). Life Sn‘. 4, 2345-2351. Schlesinger, K., Stavnes, K., and Boggan, W. 0. (1969). Psychopharmacdogia 15,226-231. Schlosser, W. (1971). Arch. Znt. Pharmacodyn. Ther. 194, 93-102. Schlosser, W., and Franco, S. (1979a).J. Pharmacol. Exp. Ther. 211, 290-295. Schlosser, W., and Franco, S. (197913). Neuropharmacology 18, 377-381. Schneider, J. H. R., Cassir, H. R., and Chordikian, F. (1960).J . Biol. C h a . 235, 1437-1440. Scholes, N. W. (1966).]. Pharmacol. Exp. Ther. 153, 128-132. Schreiber, R. A. (1979). Psychopharmacology 66, 205-206. Schultz, F. (1943).J. Physiol. (London) 102, 269-273. Schultz, J. E., Siggins, G. R., Schocker, F. W., Tiirck, M., and Bloom, F. E. (1981). J . Pharmacol. Exp. Ther. 216, 28-38. Schulz, D. W., and MacDonald, R. L. (1981). Bruin Res. 209, 177-188. Schwartz, R., and Coyle, J. T. (1977). Brain Res. 127, 235-249. Schwartz, R., Zaczek, R., and Coyle, J. T. (1978). Eur. J. Pharmacol. 50, 209-220. Schwartzkroin, P. A,, and Prince, D. A. (1977). Ann. Neurol. 1, 463-469. Schwartzkroin, P. A., and Stafstrom, C. E. (1980). Science 210, 1125-1 126. Schwartzkroin, P. A., and Wyler, A. R. (1980). Ann. Neurol. 7, 95-107. Scotti de Carolis, A., and Massotti, M. (1978). Prog. Neuro-Psychophamcol. 2, 431-442. Scriver, C. B. (1976). Am.]. Dis. Child. 113, 109-114. Segal, M. (1974). Life Sci. 14, 1345-1351. Segal, M. (1975). Brain Res. 94, 115-131. Segal, M. (1976).J. Physiol. ( L m d a ) 261,617-631. Segal, M., and Bloom, F. E. (1974). Brain Res. 72, 99-114. Sethy, V. H., Roth, R. H., Walters, J. R., Marini, J., and Van Woert, M. H. (1976). NaunynSchmiedeberg’s Arch. Pharmacol. 295, 9- 14. Seyfried, T. N. (1979). Fed. Proc., Fed. Am. SOC.Exp. Biol. 38, 2399-2404. Shank, R. P., and Aprison, M. H. (1979).Zn “Glutamic Acid: Advances in Biochemistry and Physiology” (L. J. Files, M. R. Kare, S. Garattini, and W. A. Reynolds, eds.), p. 139150. Raven Press, New York. Shaywitz, B. A., Cohen, D. J., and Bowers, M. B. (1975). Neurology 25, 72-79. Shaywitz, B. A., Yager, R. D., and Gordon, J. W. (1978). Dm.Psychobiol. 11, 243-250. Shearer, D. E., Fleming, D. E., and Bigler, E. D. (1976). Epilepsia 17,429-435. Shute, C. C. D., and Lewis, P. R. (1967). Brain 90, 497-520. Sie, G., Jasper, H., and Wolfe, L. (1965). Electroencephalogr. Clin. Neurophysiol. 18, 206. Siege], J., and Murphy, G. J. (1979). Brain Res. 174, 337-340. Siggins, G. R., Hoffer, B. J., Oliver, A. P., and Bloom, F. E. (1971). Nature 233,481-483. Siggins, G . R., Henriksen, S. J., and Landis, S. C. (1976). Brain Res. 114, 53-65. Simler, S., Ciesielski, L., Maitre, M., Randrian Arisoa, H., and Mandel, P. (1973). Biochem. Pharmacol. 22, 1701-1708. Simon, J. R., Atweh, S., and Huhan, M. J. (1976).J. Neurochem. 26, 909-922. Singh, P., and Huot, J. (1973). Znt. Encycl. Pharmacol. Ther., Sect. 19 2, 427-504. Sjostrom, R., Ekstedt, J., and Anggard, E. (1975). J. Neurol., Neurosurg. Psychiatry 38, 666-668.

176

0. CARTER SNEAD 111

Skirboll, L. R., Grace, A. A., and Bunney, B. S. (1979). Science 206, 80-82. Skolnick, P., Paul, S., Crawley, J., Rice, K., Barker, S., Weber, R., Cain, M., and Cook, J. (198 1). Eur. J. Phannacol: 69, 525-527. Slater, P., Lee, L. A., Longman, D. A., and Crossman, A. R. (1980).J. Phamacol. Methodc 3, 39-49. Srnaje, J. C. (1976). Br. J. Phhannarol. 58, 359-366. Smith, R. E. (1916). Science 44,280-282. Snead, 0. C. (1978a). Neurology 28,636-642. Snead, 0. C. (197813).Neurdogy 28,643-648. Neurology 28, 1172-1 178. Snead, 0. C. (1978~). Snead, 0. C. (1978d). Nmrobgy 28, 1179- 1 182. Snead, 0. C. (1980). Neurosci. Abstr. 6, 12. Snead, 0. C. (1981a). I n “Brain-Behavior Relationships” R. Merikanas, ed.), pp. 113153. Lexington Books, D. C. Heath and Co.,Lexington, Massachusetts. Snead, 0. C. (1981b). Neurology 30(2), 143. Snead, 0. C. (1982). Smrophannocology 21, 539-543. Snead, 0. C., and Bearden, L. J. (1980a). Science 210, 1031-1033. Snead, 0. C., and Bearden, L. J. (1980b). Neurology 30,832-838. Snead, 0. C., and Bearden, L. J. (1981). Neuroloa 31(2), 157. Snead, 0. C., Yu, R. K., and Huttenlocher, P. R. (1976). Neurology (Minneap.) 26,51-56. Snead, 0.C., Benton, J. W., Dwyer, D., Morley, B. J., Kemp, G. E., Bradley, R. J., and Oh, S. J. (1980). hreuroloa 30, 732-739. Snyderman, S. E., Holt, L. E.. Canetero, R., and Jacobs, K. (1953).J. Clzn. Nutr. 1, 200207. Soper, H. V., Strain, G. M., Babb, T. G., Lieb, J. P., and Crandell, P. H. (1978). Exp. Neurol. 62,99-121. Sorel, L., Dusaucy-Bauloye, A. (1958). Arta Neurol. Belg. 58, 130-136. Soroko, F. E., and McKenzie, F. E. (1970). Phannocologist, 12, 253. Soubrie, P.. Simon, P., and Boisier, J. R. (1976). Neuropharmcology 15, 773-776. Sourkes, T. L. (1973).J. Neural Transm. 34, 153-157. Spencer, H. H., and Havilicek, V. (1974). Can. J . Phyiol. Phamacol. 52, 808-813. Spencer, P. S. J., and Turner, T. A. R. (1969). Br. J. Pharmacol. 37, 94-103. . 5, 27-81. Sprott, R. C., and Staats, M. (1975). B ~ h a vGenet. Squires, R., and Braestrup, C. (1977). h‘ature ( L a d m i ) 266, 732-734. Stidmaster, R. M., and Hanna, G. R. (1972). Epilepia 13, 313-324. Stamps, F., Gibbs, E. L., Rosenthal, I. M., and Gibbs, F. A. (1959).JAMA,J. Am. Med. Ascoc. 171,408-413. Stark, L. G., Killam, K. F., and Killam, E. K. (1970).J. Phamacol. Exp. Ther. 173, 125-132. Stark, L. G., Edmonds, H. L., and Keesling, P. (1974). Neurapharmarology 13, 261-267. Starke, K., Taube, H. D., and Borowski, E. (1977). Biochcm. Pharmarol. 26, 259-268. Steiner, F. A., Rof, A., and Akert, K. (1969). Brain Reg. 12, 74-85. Steriades, M. (1974). Electroencephalogr. Clin. h’europhysiol. 37, 247-263. Stewart, R. M., Growdar, J. H., Cancian, D., and Baldessarini, R. J. (1976). Neuroloai 26, 690-692. Stone, T. W. (1976). B r . J . Pharmarol. 56, 101-110. Stone, T. W. (1981). Adz!. Biochem. Psyhapharmarol. 29, 223-230. Stone, W. E. (1970). Phannacology 3, 367-370. Stone, W. E. (1972). In “Experimental Models of Epilepsy-A Manual for the Laboratory Worker” (D. P. Purpura, J. K. Penry, D. Torver, D. M. Woodbury, and R. Walker, eds.), pp. 407-432. Raven Press, New York.

u.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

177

Stone, W. E., and Javid, M. J. (1980). Arch. Int. Pharmacodyn. Ther. 243, 56-65. Storm-Mathisen, J. (1977). Prog. Neurobiol. 8, 119-181. Storm-Mathisen, J., and Opsahl, M. W. (1978).Neurosci. Lett. 9, 65-70. Strain, G. M., Babb, T. L., Soper, H. U., Perryman, K. M., Lieb, J. P., and Crandall, P. H. (1979). Epilepsia 20, 651-664. Stull, R. E., Jobe, P. C., Geiger, P. F., and Ferguson, G. G. (1973).J. Pharm. Pharmacol. 25, 842 -844. Stull, R. E., Jobe, P. C., and Geiger, P. F. (1977).J. Phann. Pharmacol. 29, 8-11. Swinyard, E. A. (1949). Am. J. Physiol. 156, 163-169. Swinyard, E. A. (1972).In “Experimental Models of Epilepsy-A Manual for the Laboratory Worker” (D. P. Purpura, J. K., Penry, D. Tower, D. M. Woodbury, and R. Walker, eds.), pp. 433-458. Raven Press, New York. Swinyard, E. A., Brown, W. C., and Goodman, L. S. (1952).J. Pharmacol. Exp. Ther. 106, 319-330. Swinyard, E. A., Boson, F. C. B., and Goodman, L. S. (1964).J . Pharmacol. Exp. Ther. 131, 73-84. Syapin, P. J., and Rickman, D. ,W. (1981). Eur. J. Pharmacol. 72, 117-120. Symonds, C. (1959). Brain 82, 133-146. Tabakoff, B., Yama, R. F., and Ritzrnan, R. (1978). Science 200,449-451. Taberner, P. V., and Roberts, F. (1978).Eur. J . Pharmacol. 52, 281-286. Tagliamonte, A., Tagliamonte, P., Di Chiara, G., Gessa, R., and Gessa, G. L. (1972).J. Neurocha. 19, 1509-1512. Takahashi, R., and Nakone, Y. (1978). In “Taurine and Neurological Disorders” (A. Barbeau and R. J. Huxtable, ed~.),pp. 375-385. Raven Press, New York. Takahashi, R., Nasu, T., Tamura, T., and Kariye, T. (1961).J . Neurochem. 7, 103-112. Tallman, J. F., T a d , S. M., Skolnick, P., and Gallagher, D. W. (1980). Science 207,274-281. Tan, U. (1977). Electroencephalogr. Clin. Neurophysiol. 42, 252-258. Tan, U., Senyuva, F., and Marangoz, C. (1978). Epilepsia 19,223-232. Tarsy, D., Pycock, C. J., Meldrum, B. S., and Marsden, C. D. (1978).Brain 101, 143-162. Temkin, 0. (1971). “The Falling Sickness,” pp. 3-27. Johns Hopkins Press, Baltimore, Maryland. Ten Bruggencate, G., and Engberg, I. (1971).Brain Res. 25,431-448. Terenius, L. (1975).J. Pharm. Pharmacol. 27, 450-457. Terenius, L., Gispen, W. H., and de Wied, D. (1975). Eur. J . Pharmacol. 33, 395-400. Testa, G., and Gloor, P. (1974). Electroencephalogr. Clin. Neurophysiol. 36, 517-524. Tews, J. K., and Stone, W. D. (1965).Prog. Brain Rcs. 16, 135-163. Thal, L. J., Sharpless, N. S., Wolfson, L., and Kauman, R. (1979).Ann. Neural. 7, 570-576. Thiessen, D. D., Lindzey, G., and Friend, H. C. (1968). Psychosom. Sci. 11, 227-228. Thorn, G. W., Forsham, P. H., Bennett, C. C., Roche, M., Reiss, R. S., Slessor, F., Flink, E. B., Somerville, W. (1949). Tram. A. Amer. Phys. 62, 233. Ticku, M. K. (1981).Biocha. Phurmacol. 30, 1573-1579. Toffano, G., Guidotti, A., and Costa, E. (1978).Proc. Natl. Acad. Sci. U.S.A. 75,4024-4028. Torchiana, M. L., Lotti, V. J., and Stone, C. A. (1973).Eur. J . Pharmacol. 21, 343-345. Torda, C., and Wolff, H. G. (1947). Am. J . Physiol. 151, 345-354. Torda, C., and Wolff, H. G. (1952).Am. J . Physiol. 168, 406-415. Tower, D. B. (1956).Am. J . Clin. Nutr. 4, 329-345. Tower, D. B. (1960). In “Inhibition in the Nervous System and Gamma-Amino Butyric Acid’ (E. Roberts, ed.), pp. 562-578. Pergamon, Oxford. Tower, D. B. (1976).In “GABA in Nervous System Function” (E. Roberts, T. N. Chase, and D. B. Tower, eds.), pp. 461-476. Raven Press, New York.

178

0.CARTER SNEAD 111

Tower, D. B., and Elliott, K. A. C. (1952).J. AppI. Phjsiol. 4,669-676. Tower, D. B., and McEarchern, D. (1949a). Can. J . Res. 27, 120-131. Tower. D. B., and McEarchern, D. (1949b). C a n . J . Re.y. Sert. E 27, 120-131. Trimble, 51. (1978). Epilppsia 19, 241-250. Trimble, M., Anlezark, G., and Meldrum, B. (1977). Psjrhopharmarologj 51, 159-164. Tsang, D., Lal, S., Sourkes, T. G., Ford, R. M., and Abonoff, A. (1976).J. NeuroI., Neurosurg. P.yhirrtt;y 39, 1186- 1190. Tsuji, H., Balagot, R. C., and Sadove, M. S. (l963).JA.\fAA,J. Am. Med. Assor. 183,659-661. Tunnicliff, G., Urton, M., and Wood, J. D. (1973). Biochmn. Phrmacol. 22, 501-505. Turner, S. S., and Hirsch, J. D. (1980). Suture (London) 288, 609-610. Uhl, G. R., and Snyder, S. H. (1976). Lije Sri. 19, 1827-1832. Urban, I., and de Weid, D. (1976). Exp. Brain Res. 24, 325-337. Urban, I., Lopes da Silva. F. H., Storm van Leeuwren, W., and d e Wied, D. (1974). Brain Res. 69,361 -370. C‘rca, G., Frenk, H., Leibeskind, J. C., and Taylor, A. N. (1977). Sczetzce 197, 83-86. Valentino, R. J., and Dingledine, R. (1981).J. Smrosci. 1, 784-792. Van Ruren, J. M., Wood, J. H., Oakley, J., and Hambrecht, F. (1978).J. Neurosurg. 48, 407-416. Van Delft, A. M. L., and Kitay, J. I. (1972). Sruror~idocrinology9, 188-196. Vanderhaegen. J. J., Signeau, J. C., and Gepts, W. (1975). Nature (London) 257, 604-605. Van der Heyden. J. A. M., Kloet, E. R.. Korf, J.. and Bersteeg, D. H. G. (1979). J . ,Yr tirvchein. 33, 857 -86 1. Van Dugn, H., Schwartzkroin, P. A., and Prince, D. A. (1973). Brain Res. 53, 470-476. Van Gelder, N . M. (1972). Brain Re$. 47, 73-77. Van Gelder, N. M. (1975). Brain Res. 94, 297-306. Van Gelder, N. M. (1978a). Can. J. Phyiol. Phannacol. 56, 362-374. Van Gelder, N. M. (1978b). f n “Taurine and Neurological Disorders” (A. Barbeau and R. J. Huxtable, eds.), pp. 387-399. Raven Press, New York. Van Gelder, N . M. (1981). Adz!. Bioclietn. Psjrhopharniarol. 29, 115-125. Van Gelder, N. M., and Curtois, A. (1972). Brain Res. 43,477-484. Van Gelder, N. M., Sherwin, A. L.. and Rasmussen, T. (1972). Brain Rrs. 40, 385-397. Van Meter, W. G., Karczmar. A. G., and Fiscus, R. R. (1978). Arch. I n t . Phat7nacodyii. Thn.. 23, 249-260. Van Riezen, H. (1972). Arrh. I u t . Pharmnrdjn. Ther. 198, 256-269. Van Wendt, L., Simila, S., Saukkonen, A., and Kavisto, M. (1980).Prdiatrics 65, 1166-1 169. Van Woeri, hl. H . , and Sethy, V. H. (1975). S e u r o l o ~25, 135-140. Van Woert, M. H.. Rosenbaum, D., Howieson, J., and Bowrers, M. B., Jr. (1977).N . Engl. J . .\.f~d.296, 70-75. Vecht, C . J.. Van Woerkom, T. C. A. M., Teelken, A. W., and Minderhoud, J. M. (1975). .-1rrh. Siwrol. (Chicago) 32, 792-797. Velasco, M., Velasco, F., Romo, R., and Estrada-Villanueva, F. (1981a). Exp. R’eurol. 72, 318-331. Velasco, M., Velasco, F., Romo, R., and Martinez, A. (1981b). Exp. Nnirnl. 72, 332-345. Vernadakis, .4., and Woodbury, D. M. (1960). In “Inhibition in the Nervous System and Gamma-Amino Butyric Acid’ (E. Roberts, ed.), pp. 242-248. Pergamon, Oxford. Vernadakis, A , , and Woodbury, D. M. (1969). Epilrpsia 10, 163-170. Vosu, H., and Wise, R. A. (1975). Behail. B i d . 13,491-495. Wada, J. A. (1977). .4rrh. .VPurul. (Chirago) 34, 389-395. Wada, J. A., Balzamo, E., Meldrum, B. S., and Naquet, R. (1972). Elrctromicephalogr. Clin. .Vno-oph,s.tinl. 33, 520-526.

SACRED DISEASE: NEUROCHEMISTRY OF EPILEPSY

179

Waddington, J. L. (1978). Eur. J . Pharmacol. 51,417-422. Wahlstrom, G. (1978). Eur. J. Pharmucol. 51, 219-227. Walaas, I., and Fonnum, F. (1979). In “GABA-Neurotransmitters” (P. Korgsgaard-Larsen, J. Scheelakrueger, and H. Kofod, eds.), pp. 60-73. Academic Press, New York. Waldmeier, P. C., and Fehr, B. (1978). E u r . 1 . Pharmacol. 49, 177-184. Walker, A. E., and Johnson, H. C. (1945).Arch. Surg. (Chicago) 50, 68-73. Walker, D. W., and Zornetzer, S. F. (1974). Electroencephalogr. Clin. Neurophysiol. 36, 233243. Wallach, D. P. (1961). B i o c h a . Pharmucol. 5, 323-331. Wallach, M. B., Winters, W. D., Mandrell, A. J., and Spooner, C. E. (1969). Electroacephalogr. Clin. Neurophysiol. 27, 563-573. Walters, R. J., and Roth, R. H. (1972). Biochem. Pharmacol. 21, 21 1-221. Walters, R. J., Lakoski, J. M., and Eng, N. (1978). SOC. Neurosci. Abstr. 4, 285. Wang, R. Y., and Aghajanian, G. F. (1977). Brain Res. 120, 85-102. Ward, A. A. (1972). In “Experimental Models of Epilepsy-A Model for the Laboratory Worker” (D. P. Purpura, J. K. Penry, D. Tower, D. M. Woodbury, and R. Walter, eds.), pp. 13-36. Raven Press, New York. Wasterlain, C. G., and Jonec, V. (1980a). SOC.Nmrosci. Abstr. 6, 112. Wasterlain, C. G., and Jonec, V. (1980b). Life Sci. 26, 387-391. Wasterlain, C. G., and Jonec, V. (1981). Exp. Neurol. 73, 595-599. Watson, S. J., Richard, C. W., and Barchas, J. D. (1978). Science 200, 1180-1183. Way, E. L. (1979).Adv. B i o c h . Psychopharmacol. 20, 3-10. Wayne, H. L. (1954).J. Clin. Endocnnol. Metub. 14, 1039-1046. Weinberger, J., Nichlas, W. J., and Berl, S. (1976). Neurology 26, 162-166. Weiner, W. J., Goetz, C., Nausieda, P. A., and Klawans, H. L. (1977a).Eur.1. Pharmacol. 46, 21-24. Weiner, W. J., Goetz, C., Nausieda, P. A., and Klawans, H. L. (1977b).Life Sci. 21,901-906. Welch, K. M. A., Wang, T. P. F., and Chabi, E. (1978). Ann. Neurol. 3, 152-155. Wenger, G. R., Stitzel, R. E., and Craig, C. R. (1973). Neuropharmacology 12,693-703. Werman, R. (1966). Comp. B i o c h . Physiol. 18,745-766. Westerink, B. H. C., Lejeune, B., Korf, J., and Van Pragg, H. M. (1977).Eur. J. Pharmacol. 42, 179-190. Whittle, S. R., and Turner, A. J. (1978).J. Neurochem. 31, 1453-1459. Whittle, S. R., and Turner, A. J. (1981). Biochim. Biophys. Acta 657, 94-105. Wiedemann, C. (1877). Arch. Exp. Pathol. Pharmakol. 6, 216-232. Wiegant, V. M., Gispen, W. H., Terenius, L., and d e Wied, D. (1977). Psychoneuroadocrinology (Oxford) 2,63-70. Wier, R. L., Chase, T. N., Ng, L. K. Y., and Kopin, I. J. (1973). Brain Res. 52, 409-412. Wilkinson, D. M., and Halpern, L. M. (1975). Proc. West. Pharmucol. SOC. 18, 146-147. Wilkinson, D. M., and Halpern, L. M. (1979a). Neuropharmacology 18,219-222. Wilkinson, D. M., and Halpern, L. M. (1979b).J. Pharmacol. Exp. Ther. 211, 151-158. Williams, D., and Russell, W. R. (1941). Lancet 1, 476-479. Williams, M., and Risley, E. A. (1979). Life Sci. 24, 833-844. Willig, R. P., and Lagenstein, I. (1980). Monatsschr. Kinderheilkd. 128, 100-105. Willig, R. P., Lagenstein, I., and Iffland, E. (1979). Maatsschr. Kinderheilkd. 126, 191-197. Willis, G. L., Singer, G., and Evans, B. K. (1976). Pharmacol., B i o c h a . Behau. 5, 207-213. Willmore, L. J., Sypert, G. W., Mumson, J. B., and Hurd, R. W. (1978). Science 200, 150 1 - 1503. Wills, J. H. (1970). Int. Encycl. Pharmacol. Ther. 1, 345-469. Wise, C. D., Burger, B. D., and Stein, L. (1972). Science 177, 180-183.

180

0. CARTER SNEAD 111

K’ochmschr. 81, 1350-1360. Wolff, V. H. (1956). D k h . Wood, J. D. (1972). I n “Experimental Models of Epilepsy-A Manual for the Laboratory Worker” (D. P. Purpura, J. K. Penry, D. Tower, D. M. Woodbury, and R. Walter, eds.), pp. 459-476. Raven Press, New York. Wood, J. D. (1975). Prog. Neurobiol. 15, Part 1, 79-95. Wood, J. D. (1980). Brain Res. Bull. 5, Suppl. 2, 777-780. Wood,J . D., and Peesker, S. J. (1974).J . A3eurochem. 23, 703-712. Wood, J. D., and Peesker, S. J. (1975).J . iVeurochem. 25, 277-282. Wood, J. D., Watson, W. J., and Clydesdale, E. M. (1963).J. Neurochm. 10, 625-633. Wood, J. D., Russell, M. P., Kurylo, E., and Newstead, J. D. (1979).J. Neurochem. 33,61-68. Wood, J . H . (1980). Seurolog?’ 30, 645-651. Wood, J . H . , and Brooks, B. R. (1980). 1n “Neurobiology of Cerebrospinal Fluid’ (J. H. Wood, ed.), Vol. 1, pp. 259-278. Plenum, New York. Wood, J. H., Glaeser, B. S., Hare, J. A., Side, J., Brooks, B. R., and Van Buren, J. M. (1977a).J. Nncrosurg. 47, 502-589. Wood, J. H., Ziegler, M. G., Cake, C. R., Sode, J., Brooks, B. R., and Van Buren, J. M. (1977b).Neurosurgq 1, 260-265. Wood, J . H., Hare, T. A., Glaeser, B. S., Ballenger, J. C., and Post, R. M. (1979).Neurology 29, 1203-1208. Woodbury, D. M. (1972).Zn “Experimental Models of Epilepsy-A Manual for the Laboratory Worker” (D. P. Purpura, J. K. Penry, D. Tower, D. M. Woodbury, and R. Walter, eds.), pp. 558-599. Raven Press, New York. Woodbury, D. M. (1980). In “Antiepileptic Drugs: Mechanisms of Action” (G. H. Glaser, J. K. Penry, and D. M. Woodbury, eds.), pp. 447-472. Raven Press, New York. Woodbury, D. M., and Kemp, J. W. (1971). Psychiatr., Neurol., Neurochir. 74, 91-115. Woodbury. D. M., and Vernadakis, A. (1966). ,Method< Harm. Res. 5, 1-57. Worms, P., and Lloyd, K. G. (1978). Eur. J . Phannacol. 51, 55-58. Worms, P., Depoortere, H., and Lloyd, K. G. (1979). Life Sci. 25, 607-614. Wiister, M., Schultz, R., and Herz, A. (1978). Neurosci. Lett. 15, 193-201. Wiister, M., Schultz, R., and Herz, A. (1981). Biochem. Pharmacol. 30, 1883-1887. Yamamoto, C. (1972). Exp. Neurol. 35, 154- 164. York, D. H. (1976). Brain Res. 20, 233-249. York, D. H. (1979).Zn ”The Neurobiology of Dopamine” (A. S. Horn, J. Korf, and B. H. C. Westerlink. eds.), pp. 395-415. Academic Press, New York. Zarecki, P., Blake, D. J., and Somjeu, G. G. (1976). Brain Res. 115, 257-272. Zaczek, R., Nelson, M. F., and Chyle, J. T. (1978). Eur. J. Pharmacol. 52, 323-327. Zetler, G. (1979). Eur. J . Pharmacol. 60, 67-77, Zetler, G. (1980a). Neurophannacology 19,415-422. Zetler, G. (1980b). Eur. J. Phannacol. 65, 297-300. Zhang, A. Z., and Pasternak, G. W. (1981). Life Sn’. 29,843-851. Zoll, J. G., Kovacs, D. A., and Lineham, D. T. (1976). SM. Neurosci. A h . 1,270. Zsilla, G . , Cheney, D. L., and Costa, E. (1976). h‘auny~i-Schmiedeb~g’s Arch. Pharmacol. 294, 25 1-255.

BIOCHEMICAL AND ELECTROPHYSIOLOGICAL CHARACTERISTICS OF MAMMALIAN GABA RECEPTORS By Salvatore J. Enna Dopaltments of Pharmacology and of Nourobiology and Anatomy Univorrity of Texas Medical School Houston, Toxar

and Joel P. Gallagher Dopartment of Pharmacology and Toxicology University of Toxar Medical Branch Galvorton, Toxar

I. Introduction ........................................................ 11. Electrophysiological Studies . . .............. ............ A. Inhibition: The Physiological Product of GABA Receptor Activation ... B. Types of GABA Receptors ......................................... C. Pharmacological Implications ...... 111. Biochemical Studies .................. A. GABA Receptor Binding .......... B. Regulation of Recognition Site Binding .............................. C. Classification of GABA Receptors ................................... D. Solubilization of GABA Receptors.. ................................. IV. Summary and Conclusions ............................................. References .....................................

181 182 182 188

196 201 203 204

1. Introduction

The identification of y-aminobutyric acid (GABA) in the vetebrate central nervous system was made by Roberts and Frankel (1950), Udenfriend (1950), and Awapara (1950). In the succeeding 30 years, data have accumulated to suggest that this amino acid may serve as one of the more important neurotransmitters in mammalian brain and spinal cord. While early work concentrated on characterizing the manner in which this substance is synthesized, catabolized, stored, released, and accumulated in brain tissue, in more recent years there has been a shift in emphasis to identifying and defining the synaptic receptor sites for GABA. Indeed, there was much skepticism about a neurotransmitter 181 INTERNATIONAL REVIEW OF NEUROBIOLOGY, VOL. 24

Copyright 8 1989 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-366824-7

182

SALVATORE J. E N N A A N D JOEL P. GALLAGHER

role for GABA prior to the discovery of agents capable of antagonizing, relatively specifically, the electrophysiological effects of this substance (Curtis at al., 197 la). During the past decade the tempo of research on GABA transmission has increased dramatically. Evidence has been found to link alterations in GABA function with the symptoms of a variety of Iieuropsydiialr-ic disorders (Enna, 1980). Furthermore, discoveries have been made implicating GABA and its receptors in the mechanism of action of a variety of clinically effective agents, such as anxiolytics, hypnotics, muscle relaxants, and anticonvulsants (Krogsgaard-Larsen et al., 1979). Moreover, knowledge in this area advanced to a stage where it is now possible to design drugs that will rather selectively modify the GABA system (DeFeudis, 1981; Enna, 1981a). Many of these agents are currently undergoing clinical trials. The aim of the present communication is to summarize the results of recent biochemical and electrophysiological studies directed toward characterizing GABA receptors. Because of the rapid developments in this field, a comprehensive treatise was not possible. Rather, emphasis is placed on highlighting and discussing selected topics that the authors feel represent some of the more exciting advances in this area. Readers wishing to obtain more information on this subject, or on other aspects of GABA transmission and pharmacology, are urged to consult any of a number of other reviews and monographs (Krogsgaard-Larsen and Falch, 1981; Macdonald and Young, 1981; DeFeudis, 1981; Enna and DeFrance, 1980; Roberts, 1956, 1979; Curtis, 1979a; Enna, 1981a,b; Krogsgaard-Larsen et al., 1979; Mandel and DeFeudis, 1979; Roberts et al., 1976; Costa et nl., 1981).

II. Eledrophysiological Studies

Although a great deal of electrophysiological data have been obtained using invertebrate preparations, the scope of this review will be limited to results derived from mammalian studies. For a discussion of GABA receptors in invertebrate and nonmammalian vertebrates, see the review by Nistri and Constanti (1979). A. INHIBITION: THEPHYSIOLOGICAL PRODUCT OF GABA RFCFPTOKACTIVATION The first indication that GABA may be a neurotransmitter in the vertebrate central nervous system (CNS) came from studies in which

CHARACTERISTICS OF MAMMALIAN

GABA RECEPTORS

183

topically applied solutions of the amino acid exerted inhibitory effects on brain electrical activity (Hayashi and Nagei, 1956; Hayashi and Suhara, 1956). Furthermore, work with convulsant hydrazides (Killam and Bain, 1957; Killam, 1957) seemed to confirm this hypothesis. As a result of these and other reports, it is now felt that the majority of inhibition observed in the vertebrate CNS is mediated by GABA. In general, GABA-induced inhibitions are thought to occur by way of two different mechanisms, one of which is characterized as postsynaptic inhibition (Eccles, 1964; Curtis, 1979b) and the other as presynaptic inhibition (Frank and Fourtes, 1957; Eccles et al., 1962, 1963; Eccles, 1964, 1969; Nicoll and Alger, 1979; Curtis, 1979; Curtis et al., 1959). 1. Postsynaptic Inhibition GABA produces inhibition at a variety of different sites within the mammalian central, and possibly peripheral, nervous systems. Using an intact preparation, a postsynaptic inhibitory action resulting from iontophoretically applied GABA was found in cat spinal motoneurons and dorsal and ventral horn interneurons (Curtis et al., 1968). Evidence was subsequently provided to indicate that this inhibitory effect was due to an increase in chloride, and possibly potassium, conductance (Curtis et al., 1968; ten Bruggencate and Engberg, 1968). A similar postsynaptic inhibitory action for GABA in mammalian cerebral cortex was first demonstrated by Krnjevic and his colleagues (1966; Krnjevic and Schwartz, 1966, 1697). These experiments revealed a hyperpolarizing action for GABA that was associated with a marked increase in membrane conductance. Ionotrophoretic application of GABA mimicked the inhibitory postsynaptic potential evoked by epicortical stimulation. Importantly, both hyperpolarizing responses were reversed to depolarizations by the intracellular injection of chloride. Krnjevic ( 1974) summarized the results of several investigations (Dreifuss et al., 1969; Kelly et al., 1969; Obata et al., 1970; ten Bruggencate and Sonnhof, 1972; Curtis and Johnston, 1974a) and concluded that the universal action of GABA on vertebrate central neurons appears to be a membrane hyperpolarization brought on by an increase in chloride, and possibly potassium, permeability.

2. Presynaptic Inhibition During this same period the phenomenon of presynaptic inhibition was being investigated in spinal cord using extracellular techniques. Several of these studies suggested that GABA was a likely candidate as the transmitter for this phenomenon. However, this contention is still somewhat controversial (Krnjevic and Morris, 1974; Kriz et al., 1974; Krnjevic, 1976; Levy, 1977; Ryall, 1978; Davidoff, 1981). Nevertheless,

184

SALVATORE J. ENNA AND JOEL P. GALLACHER

an impressive amount of evidence has been accumulated to suggest that GABA is capable of causing a depolarization of primary afferent neuron terminals in the spinal cord. This action, which presumably occurs by way of an axoaxonic synapse, results in a decreased release of transmitter from the depolarized afferent (Eccles et al., 1963; Schmidt, 1971; Davidson and Southwick, 1971; Nicoll and Alger, 1979). These in uiuo experiments provided functional evidence that GABA can act as an inhibitory neurotransmitter in the mammalian CNS. However, because of the difficulties associated with in uiuo intracellular recording techniques, as well as the limitations imposed upon a mechanistic interpretation of recordings obtained from intact tissue, various in vitro peripheral and central preparations were developed. These in uitro systems have facilitated a quantitative analysis of the interaction of GABA with its receptor at the level of the cell membrane. 3. In Vitro LVfodeL~

Peripheral models have historically been used to investigate the actions of central neurotransmitters. For example, the nicotinic action of acetylcholine was first defined at the skeletal neuromuscular junction, whereas the muscarinic actions of this substance were defined in guinea pig ileum. The rabbit ear artery has been a popular system for studying the actions of noradrenaline. Following deGroat’s observations ( 1970, 1972) that GABA depolarizes mammalian sympathetic, parasympathetic, and sensory ganglia, various laboratories began to use these tissues as peripheral models of central GABA receptors. While the function of these receptors is still unknown, their presence on these peripheral neurons may be related to their common embryonic origin from neural crest tissue, although there are exceptions (Nicoll and Alger, 1979). The question of whether these peripheral GABA receptors serve a similar inhibitory function to those found in the CNS is currently being debated (Koketsu et al., 1974; Kato et al., 1978; Bowery and Hudson, 1979; Brown and Higgins, 1979). Nonetheless, the possibility that these neurotransmitter receptors may not be functional is not without precedent. Thus, no function has yet been ascribed to amnionic acetylcholine receptors (Cuthbert, 1962) or to fetal cardiac cholinergic receptors (Burn, 1954), and yet both of these preparations have served as useful models for studying these receptor systems. Using extracellular techniques, Bowery and Brown (1972, 1974) defined the potency of GABA and related amino acids to depolarize rat sympathetic ganglia and compared these results with the activity of GABA in other test preparations such as dog blood pressure (Stanton and Woodhouse, 1960), crayfish stretch receptor (Edwards and KuWer,

CHARACTERISTICS OF MAMMALIAN

GABA RECEPTORS

185

1959; McGeer et al., 196l), crayfish neuromuscular junction (Dudel, 1965; Robbins, 1959), toad spinal cord (Curtis et al., 1961), cat motoneurons (Curtis and Watkins, 1960), and mammalian cortex (Purpura et al., 1959). Based on their studies it appeared that the rat sympathetic ganglion GABA receptor was similar in many respects to that found at crustacean neuromuscular junctions, although in some ways it also resembled GABA receptors identified in the vertebrate central nervous system. A finding common to all these preparations was that the GABA action could be selectively antagonized by the convulsants picrotoxin and bicuculline, whereas strychnine, a glycine receptor antagonist, was much weaker in this regard. The use of these convulsants to characterize GABA receptors was suggested by the work of others (Curtis et al., 1971a,b;Johnston et d.,1972; Kelly and Renaud, 1973; Curtis and Johnston, 1974a,b).Another common feature of these various preparations was that, although GABA depolarized some and hyperpolarized others, the membrane change was invariably due to an increase in chloride conductance. Using intracellular techniques, Adams and Brown (1973, 1975) identified the ionic mechanism for the GABA response on rat sympathetic ganglia. They showed that, depending upon the resting membrane potential, GABA could either depolarize or hyperpolarize the ganglia, and that the usual response (depolarization) was due to an efflux of chloride ion. Bowery et al. (1980)have suggested the presence of a GABA receptor on the terminals of rat sympathetic ganglia and mammalian central nervous system neurons. This GABA receptor appears to be unique in that it is insensitive to bicuculline and does not produce a membrane potential change (Bowery et al., 1980). Also it appears to be most effectively activated by the GABA analog, p-chlorophenyl GABA (baclofen), a substance which appears to be inactive at bicuculline-sensitiveGABA receptors. The molecular mechanism whereby activation of this baclofensensitive, bicuculline-resistant GABA receptor decreases transmitter output is presently unknown (Dunlap and Fischbach, 1981; Dunlap, 1981). Intracellular studies have also been conducted with rat (Feltz and Rasminsky, 1974; Deschenes et al., 1976; Deschenes and Feltz, 1976; Obata, 1974) and cat (Gallagher et al., 1974, 1975a,b, 1978) dorsal root (sensory) ganglia. Feltz’s studies with the rat dorsal root ganglion were performed in uivo, whereas Obata’s were conducted on neurons in culture. All studies with the cat ganglion were performed in uitro. Similar conclusions were reached by all of these investigators and their findings were reminiscent of the results obtained with rat sympathetic ganglia (Adams and Brown, 1975). That is, GABA appears to depolarize the

186

SALVATORE J. E N N A A N D JOEL P. GALLAGHER

membrane of dorsal root ganglion somata, and this depolarization is primarily associated with an increased chloride conductance. Although structure-activity studies were performed only with the cat dorsal root ganglia (Gallagher et ni., 1978), the relative potencies of these substances were not identical to those reported for either the pre- or postsynaptic GABA receptors on rat sympathetic ganglia (Bowery and Brown, 1974; Bowery et al., 1980). Furthermore, no baclofen-sensitive, bicucullineresistant GABA receptors were found on cat dorsal root ganglion. Indeed, the relative activity of various GABA analogs (Nakamura et al., 1981) was comparable to that found in rat brain membranes using ligand binding assays (Greenlee ei nl., 1978). This correlation supports the suggestion that dorsal root ganglia may be a more appropriate model for brain cortical GABA receptors and receptors at primary afferent terminals than are GABA receptors associated with rat sympathetic ganglia. On the other hand, sympathetic ganglion GABA receptors appear to be more similar to those found on motoneurons and interneurons. T h e GABA-induced depolarization in cat parasympathetic pelvic vesical ganglion originally found by deGroat (1970) using an extracellular technique has been reinvestigated by intracellular recordings (Mayer et al., 1981). This approach revealed that GABA produces a biphasic (depolarization followed by hyperpolarization) response. Furthermore, the hyperpolarizing effect was associated with a conductance decrease. No other GABA-induced membrane change in the mammalian CNS has been found in association with a conductance decrease, although such an action has been recorded from crayfish muscle fibers (Dudel, 1979). T h e ionic mechanism and pharmacologic profile of this GABA response is unknown. Neurons in culture also offer certain technical advantages for electrophysiological analysis (Fischbach and Nelson, 1979). T h e main disadvantage, however, is the difficulty in precisely determining the type and/or stage of differentiation of the cell being studied. As a result, data are not easily compared with those obtained using adult neurons in zriuo or in zutro. Nevertheless, cell culture has been a valuable technique for studying GABA receptors. A variety of tissues have been studied in culture to define the action of GABA, with much of the work utilizing techniques developed with mouse and chick spinal cord cells and dorsal root ganglia (Nelson, 1975; Ransom and Nelson, 1975; Ransom and Barker, 1976; Ransom et al., 1977; Fischbach, 1972; Choi et al., 1977; Dunlap and Fischbach, 1978). T h e results indicate that there may be at least two types of GABA receptors on these neurons. When activated, one type induces a membrane polarization (either hyperpolarization or depolarization) associated with

CHARACTERISTICS OF MAMMALIAN

GABA

RECEPTORS

187

an increase in chloride conductance. This chloride channel-coupled GABA receptor is blocked by either bicuculline or picrotoxin. The other population of GABA receptors (Dunlap and Fischbach, 1978, 1981; Dunlap, 1981) is found only on certain types of cultured dorsal root ganglia and does not appear to be associated with a chloride channel. Activation of this site causes a selective decrease in a voltage-sensitive calcium conductance. This receptor is resistant to bicuculline and appears to be similar to the presynaptic receptor described by Bowery et al. ( 1980). One advantage of cultured neurons is that they make possible the use of more refined electrophysiological methodology. For example, by studying neurons in culture Barker and his co-workers have contributed a great deal of basic information regarding the interaction of GABA with its receptor and the subsequent elementary events occurring at the associated chloride channels. This information has been obtained using procedures such as fluctuation analysis (McBurney and Barker, 1978; Barker and McBurney, 1979a,b; Mathers and Barker, 1980; Barker et al., 1981, 1982; Barker and Mathers, 1981) and the extracellular patch clamp technique (Mathers et al., 1981; Mathers and Barker, 1983). Two additional in vitro systems have been utilized to characterize the electrophysiologicalproperties of GAB A receptors. The first of these is a brain slice preparation. This technique has many of the advantages of the in viuo preparation without some of the technical difficulties.Several different slice preparations have been studied using both extracellular (Brown and Galvan, 1979; Hayes and Simmonds, 1978; Simmonds, 1978, 1980a,b, 1981) and intracellular (Schwartzkroin, 1975; Langmoen et al., 1978; Alger and Nicoll, 1979; Thalman et al., 1979; Wong et al., 1979; Assaf et al., 1981; Brown and Scholfield, 1979; Pickles, 1979) recording techniques. Early in uiuo intracellular studies suggested that GABA-induced inhibition in supraspinal regions (cortex, cerebellum) was due to a hyperpolarization of the cell membrane. However, using in uitro techniques, it has recently been shown that GABA is able to depolarize neurons in the olfactory cortex (Brown and Scholfield, 1979; Pickles, 1979) and to produce biphasic responses in the hippocampus (Alger and Nicoll, 1979; Thalmann et al., 1979; Langmoen et al., 1978; Assaf et al., 1981). All of these actions are bicuculline and picrotoxin sensitive and appear to be partially mediated by chloride. The fact that only hyperpolarizing responses had been previously reported using in viuo preparations (Andersen et al., 1964a,b; Eccles et al., 1977; Kandel et al., 1961; but see Krnjevic, 1981) may be due to the less-negative resting potentials obtained in these studies. Clearly, quantitative electrophysiologicalanalysis

188

SALVATORE J. ENNA AND JOEL P. GALLAGHER

using brain slices is a useful tool for further research in this area (Dingledine et al., 1980). A second in uitm preparation utilized immature rat spinal cord, with extracellular recordings taken from spinal root fibers (Evans, 1978, 1979, 1980; Allan et al., 1980). Results obtained with this system suggest that GABA receptors located on root fibers, which are nonsynaptic, may differ from those which mediate spontaneous activity. However, since both types of GABA receptors are bicuculline sensitive, the present way to discriminate between these two sites is by the use of selective GABAreceptor agonists such as THIP [4,5,6,?-tetradydroisoxazolo(5,4c)pyridin-3-01] or (+)-cis-3-aminocyclopentane-1-carboxylic acid. These two compounds are at least 20 times more effective than GABA as agonists on the synaptic receptors, whereas they are less active than GABA in depolarizing root fibers. However, it could be argued that this preparation shares certain disadvantages with the tissue culture system. That is, since with immature spinal cord the GABA receptors are in early stages of development, the response may not be the same as those observed in adult preparations. B. TYPES OF GABA RECEPTORS Based upon these electrophysiological investigations it appears that there may be at least two different kinds of GABA-receptor complexes. One of these consists of a recognition site coupled to an ion channel, activation of which leads to an influx (hyperpolarization) or efflux (depolarization) of chloride. Both effects cause inhibition, the type depending upon the neuronal localization of the receptor (pre- or postsynaptic). The direction in which chloride flux occurs depends upon the chloride gradient across the cell membrane at the time the receptor is activated. The intracellular concentration of chloride can vary from neuron to neuron, or possibly even within different parts of the same neuron, whereas the extracellular concentration of chloride is more constant. While the precise mechanism for maintaining the intracellular chloride content of an excitable cell is unknown, inward- (Keynes, 1962a,b; Eccles et nl., 1964a,b)and outward- (Boistel and Fatt, 1958; Fatt, 1960) directed pumps have been proposed. In addition to this basic unit of a GABA recognition site-chloride channel, various modulatory subunits may also be present at or near the recognition site or channel. The relationship of these various subunits to the basic complex ultimately determines the result of GABA receptor activation. A second type of GABA receptor appears to consist of a recognition

CHARACTERISTICS OF MAMMALIAN

GABA RECEPTORS

189

site linked to a calcium channel. The pharmacological specificity of this site appears to differ from that associated with chloride channels. Activation of this receptor leads to an inactivation of calcium channels which, while not altering membrane polarity, may modify cellular activity. The pharmacology of this GABA recognition site is now undergoing investigation (Bowery et al., 1981; Dunlap, 1981). Biochemical studies have suggested the existence of a third type of GABA receptor, one of which may function as a regulator of GABA release (Brennan et al., 1981). This site appears to be situated on GABA terminals and when sufficient GABA has been released into the synaptic cleft, excess transmitter activates this autoreceptor, thereby modifying release. Since the ionic mechanism associated with this action has not yet been defined, it is conceivable that the receptor could be similar to one of the above models. In any event, the possible existence of such a site offers another target for designing more selective GABAergic drugs. C. PHARMACOLOGICAL IMPLICATIONS

The possible existence of more than one type of GABA receptor makes it more feasable to develop a variety of therapeutically useful agents. That is, since GABA receptors are distributed so ubiquitously throughout the central nervous system, it would be difficult, if not impossible, to develop drugs capable of activating or inhibiting GABAergic transmission in select brain regions if all were pharmacologically and functionally identical. The therapeutic advantages of having multiple receptor subtypes is evident from the advances achieved by being able to selectively modify distinct receptor populations for the cholinergic, adrenergic, and histamine systems. With this point in mind, the following section was prepared to briefly review the pharmacology of GABA receptors. It must be stressed, however, that, from a pharmacological standpoint, the concept of multiple GABA receptors is still somewhat theoretical. Much more concrete evidence will be necessary to more precisely define the functional, and anatomical, differences between the electrophysiologically distinct GABA receptors. Using electrophysiological analysis of mammalian systems, GABA antagonists are usually classified as selective, rather than competitive or noncompetitive (Johnston, 1978). However, a few studies have attempted to characterize the interaction of these drugs. The results of one intracellular investigation suggested that both bicuculline and picrotoxin act noncompetitively, and probably at different sites (Gallagher et al., 1978). Other investigators have reached a similar conclusion (Bowery

190

SALVATORE J . ENNA AND JOEL P. GALLAGHER

and Brown, 1974). However, Simmonds (1980a) suggested that bicuculline antagonizes GABA in a competitive manner in the cuneate nucleus, whereas picrotoxin appears to be noncompetitive. Furthermore, Simmonds (1980a) reported that penicillin is also a noncompetitive GABA antagonist, selectively blocking the chloride channel at a site different fi-om either bicuculline 01-pici.otoxin (Pickles and Siniiiiorids, 1980). Thus, there is as yet no general agreement as to the precise sites and manner in which GABA antagonists exert their effect. This apparent lack of a competitive antagonist has hindered the classification of GABA receptors using electrophysiological techniques since, without such drugs, it is difficult to selectively distinguish bicuculline-sensitive GABA receptors from those that are not blocked by this alkaloid (Bowery et al., 1980; Dunlap, 1981). On the other hand, direct-acting GABA receptor agonists may be useful in addressing the issue of multiple receptors. Thus, GABA agonist structure-activity studies have been an important method for classifying these receptors (Johnston et al., 1979). Using receptor ligand binding assays (see below), numerous studies have been conducted to define the structural specificity of these sites with agonists (KrogsgaardLarsen et nl., 1975; Krogsgaard-Larsen, 1978; Enna et al., 1979; Olsen, 1976; Olsen et nl., 1978). However, there have only been a few reports (Nakamura et al., 1982; Barker and Mathers, 1981) of studies using intracellular techniques to analyze a comparable series of compounds in intact tissue. Because of slight variations in the rank order of potencies observed in these intact preparations (cat dorsal root ganglion and spinal cord neurons in culture), it appears that there may be subtle differences in the nature of these GABA receptors. Meldrum (1981) has also attempted to classify GABA receptors using agoriists. For example, it appears that bicuculline-resistant GABA receptors are especially sensitive to baclofen, whereas bicuculline-sensitive GABA receptors are not. Thus, this GABA analog may be useful for characterizing the pharmacology of this chloride-independent GABA receptor (Bowery et al., 1981). The other class of receptors are baclofen resistant, bicuculline sensitive, and are coupled to a chloride channel. This group may be further subdivided into different categories based upon their association with various modulatory subunits. It has been proposed that one of these subunits may be linked to a benzodiazepine receptor. Thus, agonists such as muscimol mimick the action of GABA in facilitating the binding of labeled benzodiazepines to membrane preparations, whereas substances such as T H I P and piperidine-4-sulfonic acid do not activate benzodiazepine binding, but rather antagonize muscimol in this regard (Braestrup et al., 1979;

CHARACTERISTICS OF MAMMALIAN

GABA RECEPTORS

191

Maurer, 1979). Other electrophysiologically active agonists such as 3-aminopropane sulfonic acid and isoguvacine seem to be partial agonists with respect to the GABA receptor-mediated facilitation of benzodiazepine binding. In light of these biochemical findings it is not surprising that the results of electrophysiological experiments do not consistently demonstrate an interaction between benzodiazepines and GABA. For example, benzodiazepines facilitate GABA responses recorded from spinal cord (Eccles et al., 1963) and cuneate nucleus (Polc and Haefely, 1976) in uiuo as well as from spinal cord neurons in tissue culture (Choi et al., 1977; Macdonald and Barker, 1978; Study and Barker, 1983) and slices of cuneate nucleus (Simmonds 1980b). On the other hand, benzodiazepines have also been reported to antagonize GABA responses recorded from cerebellar explants (Gahwihler, 1976), cerebellar purkinje cells (Curtis et al., 1976), and vestibular and cerebellar neurons (Steiner and Felix, 1976). Although it would appear that the action of benzodiazepines may vary with respect to both the neuron under investigation and the type of GABA receptor present, Study and Barker (1983) have shown that the facilitatory action of these drugs is due to their ability to increase the frequency of chloride channel opening, rather than to an effect on the single channel conductance or at the level of the recognition site. Four groups of agents (excluding the benzodiazepines) have been reported to facilitate the action of GABA. These substances do not appear to act directly at the recognition site or ion channel associated with the receptor, but rather activate these sites by enhancing the activity of endogenous GABA. Included in these groups are substances that promote the release of endogenous GABA, agents that inhibit the enzymatic catabolism of GABA, and substances that inhibit GABA uptake into neuronal or glial elements (Enna and Maggi, 1979). Few electrophysiological studies have been conducted with these drugs since their mechanisms are difficult to analyze using this approach. However, the importance of the glial uptake system in terminating the action of GABA and the involvement of this uptake process in receptor desensitization have recently been examined (Gallagher et al., 1981). Further investigations concerning the degree to which desensitization may contribute to terminating the action of GABA are needed (Krnjevic, 1981). The barbiturates appear to represent a fourth group of drugs that indirectly facilitate the action of GABA. These drugs have been proposed to prolong the lifetime of chloride channels, some of which may be associated with the GABA receptor complex (Mathers and Barker, 1980; Simmonds, 1981). Moreover, biochemical studies suggest that

192

SALVATORE J. ENNA A N D JOEL P. GALLAGHER

barbiturates may also influence the attachment of GABA to its recognition site (Johnston and Willow, 1981). There are various agents that may indirectly, and rather nonspecifically, alter GABA responses by acting on a portion of the neuronal membrane not coupled to the GABA receptor complex. Drugs such as the neuroleptics (Higashi et d.,1981) and some diuretics, the latter perhaps acting on a chloride-pump mechanisms (Gallagher et al., 1980),may be placed in this class. Further investigations into the manner in which these drugs influence membrane integrity may yield a better understanding of the GABA receptor.

111. Biochemical Studies

As discussed above, the initial data used to characterize the GABA receptor were derived from electrophysiological studies. While fruitful, this approach was limited with regard to defining the molecular properties of this receptor site. For this, investigators had to approach the question using biochemical techniques. However, prior to 1970, the required technology was not available for undertaking such studies. During the past decade a number of new methodologies have been developed enabling neuroscientists to study receptors at the molecular level. Of these, one of the most useful has been receptor ligand binding assays (Yamamura et al., 1978). T h e ability to selectively tag receptors with radiolabeled substances has opened the way for a more detailed characterization of these sites. Among other things, these assays have made it possible to map the anatomical localization of receptors (Kuhar, 1978), to define the kinetic characteristics of these sites, and to establish structure-activity requirements. Moreover, the use of ligand binding assays has led to a better understanding of the way in which receptor sites are regulated and how drugs may influence these processes. It must be borne in mind, however, that binding assays are not a measure of functional activity. Thus, before theories based on these data can be fully accepted, more physiological techniques must be used to test the hypothesis. Nevertheless, these biochemical assays have provided a wealth of information about the possible constituents of neurotransmitter receptors, which in turn has led to a better understanding of the molecular events that determine receptor activity. Numerous biochemical studies have been conducted on GABA receptor binding sites. I n the following sections an attempt will be made to summarize these data with the aim of providing some insights into the

CHARACTERISTICS OF MAMMALIAN

GABA RECEPTORS

193

pharmacological and molecular properties of this membrane constituent.

A. GABA RECEPTORBINDING Attempts to identify GABA binding components in mammalian tissue can be traced back to the 1960s (Sano and Roberts, 1963; Varon et al., 1965). Using [14C]GABA,these investigators and others (Strasberg and Elliott, 1967) were able to identify a sodium-dependent binding site for this substance in brain membranes. While the precise nature of this site was unknown, subsequent work has suggested that it may represent a sodium-dependent transport system for this amino acid (Enna and Snyder, 1975). Because the concentration of neurotransmitter receptors in brain tissue is quite limited, biochemical identification of synaptic GABA receptors had to await the development of radioactive ligands having a greater specific activity than can be achieved with 14C. Using [3H]GABA, Peck and his colleagues (1973) were the first to report a binding site in rat cerebellar membranes that had the pharmacological profile of a GABA receptor. Thus, the binding was present in the absence of sodium and was competitively inhibited by bicuculline, the receptor antagonist. Using a similar technique Zukin ~f al. (1974) described a method for labeling GABA receptors in various brain regions. This report was quickly followed by a series of studies which refined the assay still further and more completely characterized the [3H]GABA binding site (Enna and Snyder, 1975, 1976, 1977a,b; Enna et al., 1975, 1976a,b,c, 1977a,b; Coyle and Enna, 1976).As a result of these studies it appeared that there were at least two different sites to which [3H]GABA could attach in mammalian brain tissue. Binding to one was dependent upon the presence of sodium ion and was blocked by agents that inhibit GABA transport (Enna and Snyder, 1975).The other site was most apparent in tissue that had been previously frozen and to which binding most readily occurred in the absence of sodium. [3H]GABAattachment to this latter site was most potently inhibited by amino acids and drugs that had been shown in electrophysiological studies to be active as GABA receptor agonists and antagonists (Enna and Snyder, 1975, 1977a). Based on the pharmacological specificities of these sites it was concluded that the sodium-dependent component represented a neuronal or glial transport site for GABA, whereas the Sodium-independent binding represented attachment to the synaptic GABA receptor. [SH]GABA receptor binding assays conducted in brain tissue that

194

SALVATORE J . ENNA A N D JOEL P. GALLAGHER

had been preincubated with the nonionic detergent Triton X-100 revealed that there were at least two separate components to the sodiumindependent binding site (Enna and Snyder, 1977a). These components were differentiated on the basis of their kinetic characteristics. Thus, one had a relatively high affinity ( K c , = 16 nM), but low capacity (B,,, = 600 fmoYmg protein), whereas the other was a low-affinity (Kd = 130 nM) high-capacity (5 pmoYmg protein) site. Pharmacological analysis of these two sites suggested that they had similar profiles, indicating that these receptors differed only with regard to their affinity constants and relative concentrations. However, more recent work indicates that these sites may also be pharmacologically and functionally distinct entities. GABA receptor antagonists have also been used to characterize this site. Thus, Mohler and Okada (1977, 1978) identified a binding site for [3H]bicuculline methiodide. While from a pharmacological standpoint this receptor was similar to the sodium-independent [3H]GABA site, some interesting differences were noted. For example, although GABA agonists were capable of inhibiting attachment to the [3H]bicuculline site, both their absolute and relative potencies differed somewhat from those found using the [3H]GABA binding assay. This prompted these investigators to suggest that there might be two conformations for the GABA receptor, one of which is preferentially labeled by agonists and the other by antagonists. Thus, while both radioligands attach to the same receptor complex, the binding sites themselves may be separate and distinct entities. Alternatively, it was also pointed out that these data could be interpreted as indicating that bicuculline labels only a certain population of GABA receptors, whereas [3H]GABA labels all sites, both bicuculline sensitive and insensitive. Further evidence for this was provided by the finding that the regional distribution of bicuculline-labeled sites differed slightly from the regional distribution of [3H]GABA binding sites (Mohler and Okada, 1977). Moreover, unlike [3H]GABA binding, the attachment of [3H]bicuculline was not markedly influenced by freezing and thawing (Mohler and Okada, 1978). GABA receptors have also been labeled with the anatagonist [3H]adihydropicrotoxinin ([3H]DHP) (Ticku et al., 1978a,b; Ticku and Olsen, 1978; Olsen et ctl., 1980). As opposed to [3H]GABA and [3H]bicuculline binding, it appears that [3H]DHP does not label the transmitter recognition site on this complex. Rather, Z3H]DHP binding seems to be more closely associated with the chloride channel. Evidence for this conclusion is based on the finding that drugs and amino acids known to activate and inhibit the GABA receptor recognition site have little effect on [3H]DHP binding, whereas substances thought to act by way of the receptor-linked chloride channel more potently interfere with [3H]DHP attachment

CHARACTERISTICS OF MAMMALIAN

GABA

RECEPTORS

195

(Ticku et al., 1978a; Ticku and Olsen, 1979). Interestingly, data have been accumulated to suggest that this [3H]DHP binding site may represent the molecular entity through which some barbiturates exert their effect (Olsen and Enna, 1983). Other agonists ligands have also been used to label GABA receptors. These include [3H]muscimol (Beaumont et al., 1978; Snodgrass, 1978; Williams and Risley, 1979; Yoneda and Kuriyama, 1980), [3H]piperidine-4-sulfonic acid (Krogsgaard-Larsen et al., 198 l ) , and r3H]THIP (Falch and Krogsgaard-Larsen, 1982). Evidence to date would seem to suggest that all three of these electrophysiologically active GABA receptor agonists attach to the same component on brain membranes as [3H]GABA itself-.Whereas, because of its higher affinity, some have preferred to use [3H]muscimol routinely to study GABA receptors, caution must be exercised in generalizing data obtained with this ligand, or any of the other radioactive analogs, because of the possibility that they may be labeling a select population of receptors. Although there have been hints that one or more of these analogs may be markers for subpopulations of GABA receptors, conclusive data are still lacking. Nevertheless, since this possibility does exist, it may be more prudent to refer to these binding components as muscimol or THIP binding sites rather than to use the generic expression GABA receptor binding site. Using [3H]baclofen, Hill and Bowery (1981) have described a GABA receptor binding site that is insensitive to bicuculline. This site is present on peripheral autonomic terminals and in mammalian brain slices. Moreover, it differs from the binding site labeled with [3H]GABA or its tritiated analogs in that it is dependent on the presence of divalent cations, and the majority of the classical, electrophysiologically active, GABA receptor agonists are inactive in inhibiting attachment to this site. With respect to function, it has been pIopused that this baclofensensitive GABA receptor is localized on the nerve terminals of nonGABAergic neurons, with activation of this site diminishing neurotransmitter release (Bowery et al., 1980). Thus, binding assays have been developed that are capable of labeling at least four different sites, all of which seem to be related to GABA neurotransmission. One is a sodium-dependent site that appears to be a component of the neuronal or glial transport system for GABA. A second, and the most comprehensively studied, is a sodium-independent component that would appear to represent the recognition site on the GABA receptor complex. This element can be labeled with the agonists [3H]GABA, [3H]piperidine-4-sulfonic acid, r3Hlmuscimol, and [3H]THIP, as well as with [3H]bicuculline, an antagonist. A third constituent is labeled with r3H]DHP, and while this site would appear to be a

196

SALVATORE J. ENNA AND JOEL P. GALLAGHER

part of the GABA receptor complex, [3H]DHP appears to attach to a component of the ion channel rather than to the recognition site. The fourth site is labeled with r3H]baclofen and appears to be distinctly different from the sodium-independent binding site anatomically, functionally, and pharmacologically. With the continued development of GABA receptor agonists and antagonists it seems likely that the future will bring more radioligands with which to further define these binding sites.

B.

REGULATION OF

RECOGNITION

SITEBINDING

Several mechanisms have evolved to enable neuronal tissue to maintain a fairly constant level of activity under a variety of conditions. While many of these entail modification of presynaptic activity such as transmitter synthesis and release, changes in postsynaptic processes also play a role in the homeostatic control of neurotransmission (Enna and Strada, 1983). For example, denervation heightens the sensitivity of receptors located on postsynaptic elements (Creese et nl., 1977). In some cases this appears to be brought about by an increase in the number of recognition sites for the deficient neurotransmitter. Conversely, an overabundance of transmitter, such as occurs during chronic treatment with some drugs, leads to a decreased receptor sensitivity (Kendall et al., 1981). Conceivably, there are many ways in which the cell can increase or decrease receptor number and activity. The most obvious is by modifying the rate at which the receptors are synthesized or degraded. Another would be to expose or sequester receptor sites that are already present on the membrane. A third possibility would be to modify the attraction (affinity) between the receptor and the transmitter by an allosteric alteration of the recognition site. And finally, receptor activity could be changed by altering the coupling between the receptor site and some intracellular process, such as a cyclic nucleotide system, without changing the number or characteristics of the recognition sites themselves (Enna and Strada, 1983). The following section reviews some of the data accumulated with regard to factors that influence GABA receptor binding and activity. 1. Drug Treatment

As has been observed with other neurotransmitter receptors in mammalian CNS, chronic administration of drugs that activate GABA receptors leads to a reduction in the number of binding sites for this substance (Ferkany et al., 1980; Ferkany and Enna, 1980). Thus it has

CHARACTERISTICS OF MAMMALIAN

GABA RECEPTORS

197

been reported that chronic (15 days) treatment with direct-acting receptor agonists such as THIP, or inhibitors of GABA-transaminase such as y-acetylenic GABA, leads to a significant decline in the number of GABA receptors in the corpus striatum. Kinetic analysis revealed that this decrease in receptor binding was due entirely to a loss of receptor sites, with no change in affinity. Moreover, several days of drug treatment were necessary before a significant alteration in binding could be observed, suggesting that the number of binding sites was diminished because of a decrease in receptor synthesis. Interestingly, however, no change in receptor binding was noted in other brain regions such as the cerebral cortex (Ferkany et al., 1981). This finding indicates that the GABA receptors in different brain areas have a differential sensitivity to this type of regulation. Thus it could be argued that striatal receptors have a much more rapid rate of turnover than those in the cortex, or that the mechanisms which control receptor number differ in the two brain regions. GABA receptor binding has also been reported to be modified by drugs not thought to influence directly the recognition site (Maggi and Enna, 1980). For example, long-term (21 days) treatment with lithium resulted in a 60% reduction in GABA receptor binding in the corpus striatum and a 40% reduction in the hypothalamus. N o significant changes in receptor binding were noted in the cerebral cortex, cerebellum, or hippocampus. Once again, the striatal GABA receptors appear to be especially vulnerable to modification. As before, the changes in GABA receptor binding were entirely due to a decrease in the number of sites. Since lithium does not appear to interact directly with the GABA receptor, it seems likely that the lithium-induced change in binding was secondary to an increase in GABAergic activity in these brain regions. Other substances, such as ethanol and opiates, have also been reported to modify GABA receptor binding (Reggianiet al., 1980; Tran et al., 1981; Ticku and Huffman, 1980).As with lithium, these changes are probably secondary phenomena. In all cases, however, it has yet to be demonstrated whether these alterations in GABA binding lead to a modification in the functional activity of the GABA system. However, since GABA receptors are found in virtually every region of the CNS (Enna, 1981a), it seems reasonable to assume that a modification in recognition site binding could have subtle, if not dramatic, effects on brain function.

2. Hormones Recent studies have shown that brain neurotransmitter receptor binding and activity are subject to hormonal control (Hruska and Sil-

I98

SALVATORE J . ENNA A N D JOEL P. GALLAGHER

bergeld, 1980). These hormone effects are most evident in situations where the nervous system is responding to a change in conditions, such as during stress or drug treatment (Enna et al., 1983). Moreover, receptor modifications for a number of neurotransmitter systems have abo been found in association with aging (Enna and Strong, 1981). It is not surprising, therefore, to discover that GABA receptors are modified under these conditions as well. Thus it has been reported that the number of midbrain GABA receptors increases within 72 hr after adrenalectomy in the rat (Kendall ut af., 1982a). In addition, removal of the adrenals causes an increase in GABA receptor binding 1 to 2 weeks later. O n the other hand, hypophysectomv decreases midbrain GABA receptor binding but is not reversed by corticosteroid treatment, indicating that these receptors are probably being modified by pituitary rather than steroid hormones. Further evidence for this hypothesis was supplied by the finding that ACTH treatment, like adrenalectomy, causes an increase in both midbrain and striatal binding for [?H]GABA (Kendall et al., 1982a). As observed with drug treatment, striatal GABA receptors appear to be particularly sensitive to modification by ACTH in that, under the conditions of these experiments, no significant change in binding was noted in other brain regions (cerebral cortex, hippocampus, cerebellum, and brainstem). Kinetic analysis showed that the ACTH-induced increase in GABA receptor binding was due to the appearance of a high-capacity, low-ahity site. Whether ACTH, and possibly other hormones, increases the production of new recognition sites, or is capable of exposing sites that are already present, is unknown. GABA receptor recognition site binding and activity also appear to be influenced by aging (Kendall et al., 1982b; Lippa et nl., 1981). Thus, the number, and affinity, of GABA receptor sites in rat brain cerebral cortex, midbrain, and cerebellum are significantly less in aged (28 months) rats compared to mature (6 months) animals. While the functional significance of these findings is unclear, it is curious that, in a separate study, the electrophysiological response to iontophoretically applied muscimol was enhanced in the older animals (Lippa et al., 1981) rather than decreased. However, a decline in the number of receptor recognition sites may not necessarily be indicative of a decrease in receptor activity. That is, it is conceivable that an increase in the coupling between the remaining sites and postreceptor phenomena may be sufficient to overcome a deficiency in receptors. Alternatively, since the binding assay was conducted using [3HJGABA, and the electrophysiological study performed with muscimol, these data may point to differences in the receptor sites for these t w o ligands. Thus it is possible that only

CHARACTERISTICS OF MAMMALIAN

GABA RECEPTORS

199

muscimol-resistant GABA receptors decline with age, but muscimolsensitive sites are unaffected or enhanced. In any event, results such as these should serve as a warning against making facile assumptions about the functional consequences of a receptor binding site alteration. Age-related changes in receptor number or function may be due to a number of factors. The most obvious of these is that the loss in receptor sites is a consequence of cell death. Moreover, it is also possible that the number of recognition sites is changed in an intact cell due to a breakdown in the machinery necessary for receptor synthesis. Then too, since it appears that hormones can modify receptor number, and since there are changes in the circulating levels of hormones with advancing years, some age-related alterations in receptor binding may be secondary to modifications in endocrine function. Based on the data accumulated thus far, it would seem most prudent to assume that all of these factors are contributory.

3. Other Regulators Most investigations aimed at understanding the regulation of GABA receptors have concentrated on acute changes in these sites. Perhaps the first evidence that the biochemical properties of GABA receptors could be rapidly modified was provided by the finding that Triton X-100 was capable of revealing a higher afhity binding component (Enna and Snyder, 1977a). The possible significance of this finding is that the membrane contains several populations of GABA receptors having differing affinities for the transmitter and that the type of receptor available may vary with conditions. If this is the case, then the cell is endowed with the capacity to rapidly alter its sensitivity to GABA. Also, by defining the membrane components that determine which receptors are present at any given time, it may be possible to develop drugs capable of modifying the sensitivity of the tissue to GABA. This notion that the number and affinity of GABA receptor recognition sites can be rapidly regulated at the receptor level has received a great deal of support in recent years. Thus it has been reported that certain anxiolytics and hypnotics are capable of altering GABA receptors in vitro while not directly competing for the recognition site. For example, under the proper assay conditions, benzodiazepines have been reported to increase the amount of [3H]GABAbound to brain membranes (Guidotti et al., 1978; Toffano ct al., 1978), an increase which is reminiscent of that seen after treating tissue with Triton X-100. Indeed, if the brain membranes have been previously exposed to the detergent, benzodiazepines no longer enhance binding; this finding suggests that the two substances may be acting in a similar manner. If the benzodiazepines

200

SALVATORE J. ENNA A N D JOEL P. GALLAGHER

activate GABA binding in uiuo, then the ability to expose a set of higher afFinity GABA receptors to the endogenous ligand could be the mechanism whereby these drugs facilitate GABAergic transmission (Olsen and Enna, 1983). Numerous studies have revealed the presence of constituents in brain tissue that are capable of modifying GABA receptor binding (Massottiet nl., 1981; Johnston and Kennedy, 1978; Mazzari et al., 1980). These include phospholipids, peptides, and an assortment of low molecular weight substances. Guidotti and his colleagues (Massotti et al., 1981) have provided evidence to suggest that the benzodiazepines enhance GABA binding by displacing a peptide that they have termed GABA-modulin. These investigators propose that GABA-modulin is a normal constituent of the GABA receptor complex that regulates the availability of recognition sites. GABA receptor binding is also enhanced by barbiturates (Willow and Johnston, 1980, 1981). The mechanism of this enhancement appears to differ from that observed with the benzodiazepines. Thus, the pentobarbital-induced increase in binding is blocked by picrotoxin, suggesting that at least some barbiturates may influence GABA receptor activity by an action at the ion channel. Interestingly, while picrotoxin blocks the action of pentobarbital on GABA binding, by itself it does not influence the attachment of [SH]GABAto the recognition site. These data are in accord with the results obtained from binding studies using [3H]DHPas a ligand (Ticku and Olsen, 1978). As discussed previously, this radioactive substance appears to be a specific label for the chloride channels associated with GABA receptors, and it has been found that among other substances, the barbiturates are capable of inhibiting attachment to this site. This suggests that drugs interacting at the channel may influence the state of the recognition site. Biochemical studies have also demonstrated a possible link between GABA receptor recognition sites and benzodiazepine receptors (Tallman et nl., 1978; Karobath and Sperk, 1979). Thus, activation of GABA receptors by some agonists leads to an increase in benzodiazepine binding in brain membranes. This enhancement in binding appears to be due to an increase in the affinity of the receptor site for benzodiazepines with no apparent change in the number of drug receptors. Whereas there is still some debate as to whether all GABA receptors are associated with a benzodiazepine site, it seems likely that all benzodiazepine receptors are coupled to a GABA receptor (Maggi et d., 1980). Taken together these data indicate that the GABA receptor is a complex entity composed of at least three major components. One compo-

CHARACTERISTICS OF MAMMALIAN

GABA RECEPTORS

20 1

nent is the recognition site for GABA, the second is an associated ion channel, and the third may be a portion of the benzodiazepine receptor. In addition, the recognition site may possess two different conformations (agonist and antagonist), the affinities of which may be regulated by endogenous modulators. Furthermore, it appears that activation of the recognition site influences not only the ion channel, but may also modify the benzodiazepine site, and that some channel-active substances can reciprocally modify recognition site binding as well. Accordingly, the GABA receptor should be envisioned as a dynamic entity composed of a number of interacting segments all of which act in concert to produce the appropriate response following activation.

C. CLASSIFICATION OF GABA RECEPTORS As with electrophysiological studies (see Section II,B), biochemical assays have suggested the presence of pharmacologically and functionally distinct GABA receptors. A better definition of these sites will make it possible to devise more selective GABA receptor agonists and antagonists in a manner analogous to what has been accomplished for the noradrenergic, cholinergic, and histaminergic systems (Yamamura and Enna, 1981). Several studies have suggested that the high- and low-affinity GABA binding sites represent distinct subsets of receptors (Browner et al., 1981; Palados et al., 1979; Braestrup et al., 1980; Guidotti et al., 1979). Although earlier investigations indicated that these two sites had similar pharmacological profiles (Enna and Snyder, 1977a; Enna et al., 1979), the more recent work indicates that they may be anatomically and functionally distinct. Thus, Guidotti and co-workers demonstrated that the ratio of high- to low-affinity GABA binding sites varies significantly throughout the rat brain (Guidotti et al., 1979). These workers found that while the concentration of high-affinity sites was fairly constant (approximately 1 pmoi/mg protein) in five different regions of the brain, there was up to a 10-fold difference in the concentration of low-affiity sites. Thus, the region having the highest ratio of high- to low-affiity sites was the substantia nigra, where the concentrations were about equal. In the cerebellum, however, there are almost 10 times more lowaf€inity than high-affinity sites. These workers suggested that the higher affinity component may represent the receptor that is most associated with GABAergic transmission since, among other observations, lesion experiments appeared to affect only this component. Further evidence that the high- and low-affinity binding sites differ

202

SALVATORE J. ENNA AND JOEL P. GALLAGHER

was provided by studies examining the influence of GABA agonists on benzodiazepine binding. Thus, while a variety of GABA analogs are electrophysiologically active as agonists and are able to displace specifically bound [3H]GABA,only certain members of this class are capable of activating benzodiazepine receptor binding (Braestrup et nl., 1980; Ferkany et ( I / . , 1981). For example, GABA, muscimol, and kojic amine are GABA receptor agonists capable of increasing benzodiazepine receptor binding, whereas, under similar incubation conditions, agonists such as THIP, piperidine-4-sulfonic acid, and imidazoleacetic acid are virtually without affect on the benzodiazepine site. This suggests either that not all GABA receptors are coupled to benzodiazepine recognition sites, or that the portion of the recognition site coupled to the chloride channel may be pharmacologically different from that linked to benzodiazepine receptors. In either case, the data are consistant with the concept of heterogeneitv in GABA receptor binding sites. Further evidence of pharmacological and functional differences in these components was provided by the finding that only the low-affinity binding site appears to be associated with the benzodiazepine receptor (Palacioset al., 1979; Braestrupet al., 1980; Browner et nl., 1981). Thus it has been demonstrated that GABA agonists are fully capable of activating benzodiazepine binding under conditions where the high-afhity receptor is apparently masked. Moreover, Browner et al. (198 1) discovered that certain chaotropic agents, such as ammonium thiocyanate, selectively abolish GABA binding to the higher a f h i t y component and yet thiocyanate does not interfere with the ability of GABA to activate benzodiazepine receptors. While not conclusive, the results of these studies support the notion that the high- and low-affinity GABA binding sites differ, not only kinetically, but in the extent of their association with benzodiazepine receptors. Another possible difference in GABA receptor binding sites was observed by examining the potency of bicuculline to inhibit C3H]GABA binding in various brain regions (Browner rt al., 1981). In this study it was found that bicuculline is some five times more potent in inhibiting GABA binding in the midbrain and cerebral cortex than in the corpus striatum and cerebellum. This differential sensitivity may be a biochemical correlate of the electrophysiological data indicating that not all GABA receptors are equally sensitive to this antagonist (Curtis and Felix, 1971; Krnjevic, 1974). Other approaches have also yielded data indicating the existence of' functionally distinct GABA receptors. Thus it has been reported (Brown and Higgins, 1979; Mitchell and Martin, 1978; Snodgrass, 1978; Brennan et nl., 1981) that GABA is capable of inhibiting its own potassium-

CHARACTERISTICS OF MAMMALIAN

GABA RECEPTORS

203

stimulated release from rat brain synaptosomes. This discovery led to the suggestion that there may be a class of GABA receptors located presynaptically on GABAergic terminals (autoreceptors) which modulate transmitter release. Analysis of this site reveals that its pharmacological specificity is similar to that found for the GABA binding site, including the fact that it is sensitive to blockade by bicuculline. As discussed previously (Section III,A), Bowery (Bowery et al., 1980; Hill and Bowery, 1981) has discovered a GABA receptor that modulates the release of other neurotransmitters. While these data would seem to suggest the presence of axoaxonic synapses in brain, it is unclear at present whether this action is the consequence of a GABA-induced depolarization as observed in spinal cord. This site differs from the autoreceptor in several respects. One is that this receptor is located on other neurotransmitter terminals, whereas the autoreceptor is on GABA terminals. Another difference is that the one site is sensitive to activation by baclofen, insensitive to bicuculline, and is calcium-dependent, whereas the autoreceptor is blocked by bicuculline and not influenced by baclofen. Thus, several lines of evidence suggest the existence of multiple types of GABA receptors. Not only are there kinetically distinct sodiumindependent binding sites, these sites have a different regional distribution, and they appear to be functionally distinct with only the lower affinity component appearing to have an association with benzodiazepine receptors. Moreover, these two binding sites are differentially sensitive with regard to the effects of denervation and the action of chaotropic agents. Furthermore, it would appear that GABA binding sites can be distinguished on the basis of their sensitivity to bicuculline and on their pharmacological specificity with regard to activation of benzodiazepine receptors. GABA receptors may also be defined on the basis of anatomical localization and function. Some postsynaptic sites regulate chloride channels in addition to benzodiazepine receptors, whereas presynaptic receptors regulate the release of GABA and other neurotransmitter agents. Because of the likelihood of pharmacological differences among these various populations, it may be possible to develop substances that will selectively modify one function without affecting the others. D. SOLUBILIZATION OF GABA RECEPTORS

A number of investigators have attempted to solubilize GABA receptor binding sites in an attempt to characterize these entities more

204

SALVATORE J. ENNA AND JOEL P. GALLAGHER

completely (Yousufiet al., 1979; Gavish et al., 1979; Chude, 1979; Asano and Ogasawara, 1980; Gavish and Snyder, 1981; Massotti et al., 1981). For these studies, solubilizing agents such as sodium deoxycholate, lysolecithin, and Triton X-100 have been used. In all cases the solubilized fraction has been shown to contain a [3H]GABA or [3H]muscimolbinding component possessing a pharmacological specificity quite similar to that found in brain membranes. While in some studies this solubilized component has been shown to possess both a GABA and benzodiazepine binding site (Gavish et al., 1979; Gavish and Snyder, 1981),other studies have been able to separate these two entities (Yousufi et al., 1979; Massotti et al., 1981). This has led to some debate as to whether the GABA and benzodiazepine recognition sites are part of the same molecular entity or whether they are two different constituents of a supramolecular complex. The most recent data would seem to favor the latter conclusion, although further studies will be necessary to resolve this issue. There is general agreement, however, that these purification procedures are extracting the sodium-independent GABA binding site, which is an important initial step in characterizing the molecular properties of this receptor. It will be interesting to discover whether this solubilized site also contains the chloride channel, or whether the channel component is distinct from the recognition site. Furthermore, it is impossible to ascertain at this time whether the solubilized preparation contains only postsynaptic receptors or whether presynaptic sites are also being isolated. Since autoreceptors appear to be pharmacologically different from the postsynaptic GABA binding sites, it is conceivable that the molecular properties of the two components may differ. The paucity of neurotransmitter receptors in mammalian CNS is a limiting factor in purification studies, making it seem likely that progress in this area will be slow. However, further efforts should be encouraged since the precise determination of the molecular characteristics of GABA receptors depends on success in this area.

IV. Summary and Conclusions

The concept that GABA is a neurotransmitter in the mammalian CNS is supported by both electrophysiological and biochemical data. Whereas the electrophysiologicalstudies are essential for demonstrating a specific functional response to GABA, the biochemical approach is useful for characterizing the molecular properties of this site. As a result

CHARACTERISTICS OF MAMMALIAN

GABA RECEPTORS

205

of these studies the concept of the GABA receptor has progressed from a simple model of a single recognition site associated with a chloride channel to a more complex structure having a variety of interacting components. Thus, both electrophysiological and biochemical data support the existence of at least two pharmacologically distinct types of GABA receptors, based on the sensitivity to bicuculline. Also, anatomically, there appear to be two different types of receptors, those located postsynaptically on the soma or dendrites of a neighboring cell and those found presynaptically on GABAergic and other neurotransmitter terminals. From biochemical studies it appears that the GABA receptor may be composed of at least three distinct interacting components. One of these, the recognition site, may exist in two conformations, with one preferring agonists and the other having a higher afkity for antagonists. Ion channels may be considered a second component, with some of these regulating the passage of chloride ion, whereas others may be associated with calcium transport. The third major element of GABA receptors appears to be a benzodiazepine recognition site, although only a certain population of GABA receptors may be endowed with this property. In addition to these, the GABA receptor complex appears to contain substances that modulate the recognition site by influencing the availability of higher affinity binding proteins. It would appear therefore that changes affecting any one of these constituents can influence the characteristics of the others. While increasing the complexity of the system, this arrangement makes for a more sensitive and adaptable receptor mechanism. Thus the GABA receptor can be envisioned as a supramolecular complex of interacting sites, all of which contribute to the functional expression of receptor activation. Because of this complexity, GABA receptors can theoretically be modified in a variety of ways by drug treatment or disease. Accordingly, it may be possible to develop selective agonists and antagonists that may act at one of the basic components, as well as agents that may alter the receptor modulators. Conversely, a disorder of any of these entities may result in an alteration of GABA receptor function, which in turn could contribute to the symptoms of a variety of neuropsychiatric disorders. The possible existence of pharmacologically and functionally distinct GABA receptors opens avenues for further drug development. For example, it may be possible to design drugs that will activate or inhibit only those receptors coupled to a transmitter release process without influencing those associated with chloride channels. In this regard, receptor purification studies will be invaluable in helping to better classify these sites.

206

SALVATORE J . ENNA A N D JOEL P. GALLAGHER

Acknowledgments

Preparation of this article was made possible, in part, by support provided by U S . Public Health Service grants NS-13803, NS-16228, and NS-00335, a Research Career Development Award (S.J.E.). We thank Ms. Doris Rayford and Ms. Martha Myers for their excellent secretarial assistance.

References

Adams, P. R.. and Brown, D. A. (1973). BI-.J. Phoimacol. 47, 639P-640P. Adams, P. R., and Brown, D. A. (1975).J . Physiosiol. (London) 250, 85-120. Alger, B. E., and Nicoll, R. A. (1979). n’ature (London) 281, 315-317. Allan, R. D., Evans, R. H., and Johnston, G. A. R. (1980). Br. J . Phnnnacol. 70, 609-615. Andersen, P., Eccles, J. C., Schmidt, R. F., and Yokota, T. (1964a). J. Nezirophysiol. 27, 78-91. Andersen, P., Eccles, J . C., Schmidt, R. F., and Yokota, T. (1964b).J. il’euruphysiol. 27, 92- 106. Asano, T., and Ogasawara, N . (1980). Lifv Sci. 26, 1131-1137. Assaf, S. Y., Crunelli, V., and Kelly, J. S. (1981).111 “Amino Acid Neurotransmitters” (F. V. DeFeudis and P. Mandel, eds.), pp. 239-248. Raven Press, New York. Awapara, J. (1950). Fed. Proc., F P ~,4m. . Sor. Exp. Biol. 9, 148. Barker, J. L., and McBurney, R. N. (1979a). Sature (London) 277, 234-236. Barker, J. L., and McBurney, R. N. (1979b). Pror. R. Soc. London, Srr. B 206, 319-327. Barker, J. L., and Mathers, D. A. (1981). Srieiwe 212, 358-361. Barker, J. L., MacDonald, J. F., Mathers, D. A., McBurney, R. N., and Oertel, W. (1981).In “Amino Acid Neurotransmitters” (P. Mandel and F. V. DeFeudis, eds.), pp. 281-293. Raven Press, New York. Barker, J. L., McBurney. R. N.. and MacDonald, J. F. (1982). J . Physiol. (London) 322, 365-387. Beaumont, K., Chilton, W. S., Yaniamura, H. I., and Enna, S. J. (1978). Braiii Res. 148, 153-162. Boistel, J.. and Fatt, P. (1958).J. PhvSol. ( L o n d o ~144, ) 176-191. Bowery, N. G., and Brown, D. A. (1972). Br..J. Phurinucol. 45, 16OP-161P. Bowery, N. G., and Brown, D. A. (1974). B r . J . Pharmacol. 50, 205-218. Bowery, N. G., and Hudson, A. L. (1979). Br. J . Phannucol. 66, 108P. Bowery, N. G., Doble, A., Hill, D. R., Hudson, A. L., Turnbull, M. J.. and Warrington, R. (1981).In “Amino Acid Neurotransnritters” (F. V. DeFeudis and P. Mandel, eds.), pp. 333-341. Raven Press, New York. Bowery, N. G., Hill, D. R., Hudson, A. 0..Doble, A., Middlemiss, D.N., Shaw, J., and Turnbull, M. (1980). ilhturt (London) 283, 92-94. Braestrup, C., Nielsen, M.. Krogsgaard-Larsen, P., and Falch, E. (1979). Nature (Lotdon) 280, 331-333. Braestrup, C., Nielsen, M., Krogsgaard-Larsen, P., and Falch, E. (1980). I n “Receptors for Neurotransmitters and Peptide Hormones” (G. Pepeu, M. J. Kuhar, and S. J. Enna, eds.), pp. 301-312. Raven Press, New York.

CHARACTERISTICS OF MAMMALIAN

GABA

RECEPTORS

207

Brennan, M. J. W., Cantrill, R. C., and Krogsgaard-Larsen, P. (1981). Adv. Biochem. Psychopharmacol. 26, 157-167. Brown, D. A., and Gaivan, M. (1979). Br. J . Pharmacol. 65, 347-353. Brown, D. A,, and Higgins, A. J. (1979). Br. J. Pharmacol. 66, 108P-109P. Brown, D. A., and Scholfield, C. N. (1979). B r . J . Pharmacol. 65,339-345. Browner, M., Ferkany, J. W., and Enna, S. J. (1981).J . Neurosci. 1, 514-518. Burn, J. H. (1954).Pharmucol. Rm. 6, 107-112. Choi, D. W., Farb, D. H., and Fischbach, G. D. (1977). Nature ( L m d a ) 269,342-344. Chude, 0. (1979).J . Neurochem. 33, 621-629. Costa, E., DiChiara, G., and Gessa, C., eds. (1981). “GABA and Benzodiazepine Receptors,” Adv. Biochem. Psychopharmacol., Vol. 26. Raven Press, New York. Coyle, J. T., and Enna, S. J. (1976). Brain Res. 111, 119-133. Creese, I., Burt, D. R., and Snyder, S. H. (1977). Science 197, 596-598. Curtis, D. R. (1979a). In “GABA-Neurotransmitters” (P. Krogsgaard-Larsen, J. ScheelKruger and H. Kofod, eds.), pp. 17-27. Munksgaard, Copenhagen. Curtis, D. R. (1979b). In “Neurotransmitters” (P. Simon, ed.), pp. 281-298. Pergamon, Oxford. Curtis, D. R., and Felix, D. (1971). Brain Res. 34, 301-321. Curtis, D. R., and Johnston, G. A. R. (1974a). Ergeb. Phyiol., Biol. Chem. Ex$. Pharmakol. 69, 97- 188. Curtis, D. R., and Johnston, G. A. R. (1974b).In “Neuropoisons” (L. L. Simpson and D. R. Curtis, eds.), pp. 207-248. Plenum, New York. Curtis, D. R., and Watkins, J. C . (1960).J. Neurochem. 6, 117-141. Curtis, D. R., Phillis, J. W., and Watkins, J. C. (1959).J. Physiol. ( L m d a ) 146, 185-203. Curtis, D. R., Phillis, J. W., and Watkins, J. C. (1961). Br. J . Pharmacol. Chemother. 16, 262-283. Curtis, D. R., Hosli, L., Johnston, G. A. R., and Johnston, I. H. (1968). Exp. Brain Res. 5, 235-258. Curtis, D. R., Duggan, A., Felix, D., and Johnston, G. A. R. (197 la). Brain Res. 32,69-96. Curtis, D. R., Duggan, A. W., Felix, D., Johnston, G. A. R., and McLennan, H. (1971b). Brain Res. 33, 57-73. Curtis, D. R., Lodge, D., Johnston, G. A. R., and Brand, S. J. (1976). Brain Res. 118, 344-347. Cuthbert, A. W. (1962). Br. J . Phurnacol. Chemotfw. 18, 550-562. Davidoff, R. A. (198l).In “Amino Acid Neurotransmitters” (F. V. DeFeudis and P. Mandel, eds.), pp. 249-255. Raven Press, New York. Davidson, N., and Southwick, C. A. P. (1971). Brain Res. 44, 63-71. DeFeudis, F. V. (1981).DrugDeu. Res. 1, 93-105. deCroat, W. C. (1970).J. Pharmacol. Exp. Ther. 172, 384-396. deCroat, W. C. (1972). Brain Res. 38, 429-432. Deschenes, M., and Feltz, P. (1976). Brain Res. 118, 494-499. Deschenes, M., Feltz, P., and Lamour, Y. (1976). Brain Res. 118, 486-493. Dingledine, R., Dodd, J., and Kelly, J. S. (1980).J . Neurosci. Methods 1, 323-362. Dreifuss, J. J., Kelly, J. S., and Krnjevic, K. (1969). Brain Res. 9, 137-154. Dudel, J. (1965). Pjuegers Arch. Gesamte Physiol. Menschrm Tiere 283, 104-118. Dudel, J. (1979).J . Physiol. (Park) 75, 597-600. Dunlap, K. (1981). Br. J. Pharmacol. 74, 579-585. Dunlap, K., and Fischbach, G. D. (1978). Nature (Ladon) 276, 837-838. Dunlap, K., and Fischbach, G. D. (1981).J . Physiol. ( L m d a ) 317, 519-535. Eccles, J. C. (1964). “The Physiology of Synapses.” Springer-Verlag, BerIidNew York.

208

SALVATORE J. ENNA AND JOEL P. GALLAGHER

Eccles, J. C. (1969). “The Inbibitory Pathways of the Central Nervous System.” Thomas, Springfield, Illinois. Eccles, J . C., Schmidt, R. F., and Willis, W. D. (1962).J. Phystol. (London) 161, 282-297. Eccles, J. C., Schmidt, R. F., and Willis, W. D. (1963).J. Phystol. (Lolidon) 168, 500-530. Eccles, J. C., Eccles, R. M., and Ito, M. (1964a). P r w . R. Sac. h 7 2 d ~ 1 ,Ser. B 160, 181-196. Eccles,J. C., Eccles, R. M., and Ito, M. (1964b).Proc. R. Sor. Londoii, Sm. B 160,197-2 10. Eccles, J . C., Nicoll, R. A,, Oshima, T., and Rubioa, F. J. (1977). Pror. R. Soc. Lorulon, Srr. B 198,345-361. Edwards, C., and Kuf€ier. S. W. (1959).5. Seurochetn. 4, 19-30. Enna, S. J. (1980). Can. J. Neurol. Sri. 7 , 257-259. Enna, S. J. (1981a). I N ”Neuropharmacology of Central Nervous System and Behavioral Disorders” (G. C. Palmer, ed.), pp. 507-537. Academic Press, New York. Enna, S. J. (1981b). Biorhm. Phurmarol. 30, 907-913. Enna. S. J., and DeFrance, J. F. (1980). I n “Neurotransmitter Receptors” (S. J. Enna and H. 1. Yamamura, eds.), Part 1, pp. 43-63. Chapman & Hall, London. Enna, S. J., and Maggi, A. (1979). Life Sri. 24, 1717-1738. Enna, S. J.. and Snyder, S. H. (1975). Bruit, RPS. 100, 81-97. Enna, S. J., and Snyder, S. H. (1976). Brain Rrs. 115, 174-179. Enna, S. J.. and Snyder, S. H. (1977a). .Llol. Pharmnrol. 13, 442-453. Enna, S. J., and Snyder. S. H. (1977b).J . ,Veurochem. 28, 857-860. Enna, S. J., and Strada, S. J. (1983). In “Clinical Neurosciences” (R. Rosenberg, R. Grossman, S. Schochet, E. R. Heinz, and W. Willis, eds.). Churchill-Livingstone, Edinburgh/New York (in press). Enna, S. J., and Strong, R. (1981). It) “Brain Neurotransmitters and Receptors in Aging and Age-Related Disorders” (S. J. Enna, T. Samajski, and B. Beer, eds.), pp. 133-142. Raven Press, New York. Enna, S. J., Kuhar, M. J., and Snyder, S. H. (1975). Bruin Re.s. 93, 168-173. Enna, S. J., Yamamura, H. I., and Snyder, S. H. (1976a). Brain Res. 101, 177-183. Enna, S. J.. Bird, E. D., Bennett, J. P., Bylund, D. B., Iversen, L. L., and Snyder, S. H. (1976b). Brain Res. 116, 531-537. Enna, S. J., Bird, E. D., Bennett, J. P., Bylund, D. B., Yamamura, H. I., Iversen, L. L., and Snyder, S. H. (1976~).N. Engl. J . “Mrd. 294, 1305-1309. Enna, S. J., Collins, J. F., and Snyder, S. H. (1977a). Bruin Res. 124, 185-190. Enna, S. J., Ferkany, J. W., and Krogsgaard-Larsen, P. (1979). In “GABANeurotransmitters” (P. Krogasgaard-Larsen, J. Scheel-Kruger, and H. Kofod, eds.), pp. 191-200. Munksgaard, Copenhagen. Enna, S. J., Yamamura, H. I., Bennett, J. P., Creese, I., Burt, D. R., Bylund, D. B., and Snyder, S. H. (1977b).J. Neurorhem. 28, 233-236. Enna, S. J., Kendall, D. A,, and Duman, R. (1983). In “Neuropeptide and Hormone R. Sladek and M. Modulation of Brain Function, Homeostasis and Behavior” Ordp. eds.), in press. Evans, R. H. (1978).Br.J. Pharmarol. 62, 171-176. Evans, R. H. (1979). Bruin Re.\. 171, 113-120. Evans, R. H. (1980).J. PhJsiol. (Lmidoti) 298, 25-35. Falch, E., and Krogsgaard-Larsen, P. (1982).J. ,Veurorhem. 38, 1123- 1129. Fatt, P. (1960). In “Inhibition in the Nervous System and Gamma-aminobutyric Acid” (E. Roberts, ed.), pp. 104- 114. Raven Press, N e w York. Feltz, P., and Rasminsky, M. (1974). ,Veuropharrnnrolog); 13, 553-563. Ferkany. J. W., and Enna, S. J. (1980). L f e Sri. 27, 143-149.

u.

CHARACTERISTICS OF MAMMALIAN

GABA

RECEPTORS

209

Ferkany, J. W., Strong, R., and Enna, S. J. (1980).J . Neurochem. 34, 247-249. Ferkany, J. W., Andree, T. H., Clarke, D. E., and Enna, S. J. (1981).Neuropharmacology 20, 1177-1182. Fischbach, G. D. (1972).Dw.Bid. 28, 407-529. Fischbach, G. D., and Nelson, P. G. (1979).I n “Handbook of Physiology” (J. M. Brookhart, V. B. Mountcastle, and E. R. Kandel, eds.), Sect. 1 , Vol. I, pp. 719-774. Am. Physiol. Soc., Washington, D.C. Frank, K., and Fourtes, M. G. F. (1957).Fed. Proc., Fed. Am. SOL. Exp. Bid. 16, 39-40. Gahwihler, B. H. (1976). Brain Res. 107, 176-179. Gallagher, J. P., Higashi, H., and Nishi, S. (1974). Int. Congr. Physiol. Sci., 26th 11, 453. Gallagher, J. P., Higashi, H., and Nishi, S. (1975a).Fed. Proc., Fed. Am. SOC.Exp. Bid. 34, 112. Gallagher, J. P., Shinnick-Gallagher, P., and Nishi, S. (1975b). Pharmacolo@t 17, 446. Gallagher, J. P., Higashi, H., and Nishi, S. (1978).J. Physiol. (London) 275, 263-282. Gallagher, J. P., Nakamura, J., and Shinnick-Gallagher, P. (1980).Int. Congr. Physiol. Soc., 28th 14, 1490. Gallagher, J. P., Nakamura, J., and Shinnick-Gallagher, P. (1981). Soc. Neurosci. 7, 320. Gavish, M., and Snyder, S. H. (1981). Proc. Natl. Acad. Sci. U.S.A. 78, 1939-1942. Gavish, M., Chang, S. L., and Snyder, S. H. (1979).Life Sci. 25, 783-790. Greenlee, D. V., van Ness, P. C., and Olsen, R. W. (1978).J. Neurochem. 31, 933-938. Guidotti, A,, Toffano, G., and Costa, E. (1978). Nature ( L d m ) 257, 553-555. Guidotti, A., Gale, K., Suria, A., and Toffano, G. (1979). Bruin Res. 172, 566-571. Hayashi, T., and Nagai, K. (1956). Int. Congr. Physiol. Soc., loth p. 410. Hayashi, T., and Suhara, R. (1956). Int. Congr. Physiol. Soc., loth p. 410. Hayes, A. G., and Simmonds, M. A. (1978). Br. J . Pharmacol. 63,503-507. Higashi, H., Inokuchi, H., Nishi, S., Inanaga, K., and Gallagher, J. P. (1981). Brain Res. 222, 103-117. Hill, D. R., and Bowery, N. G. (1981).Nature (London) 290, 149-152. Hruska, R. E., and Silbergeld, E. K. (1980). Science 208, 1466-1468. Johnston, G. A. R. (1978).Annu. Rm. Pharmacol. 18, 269-289. Johnston, G. A. R., and Kennedy, S. M. E. (1978).I n “Amino Acids as Chemical Transmitters’’ (F. Fonnum, ed.), pp. 507-516. Plenum, New York. Johnston, G. A. R., and Willow, M. (1981).Adv. Biochem. Psychophamcol. 26, 191-198. Johnston, G. A. R., Beart, P. M., Curtis, D. R., Game, C. J. A., McCulloch, R. M., and Maclachlan, R. M. (1972). Nature New Bid. 240, 219. Johnston, G. A. R., Allan, R. D., Kennedy, S. M. E., and Twitchin, B. (1979).I n “GABANeurotransmitters” (P. Krogsgaard-Larsen, J. Scheel-Kruger, and H. Kofod, eds.), pp. 149- 164. Munksgaard, Copenhagen. Kandel, E. R., Spencer, W. A., and Brinley, F. J. (1961).J. Neurophysiol. 24, 225-241. Karobath, M., and Sperk, G. (1979). Proc. Nutl. Acud. Sci. U.S.A. 76, 1004-1006. Kato, E., Kuba, K., and Koketsu, K. (1978). Brain Res. 153, 398-402. Kelly, J. S., and Renaud, L. P. (1973).Br. J . Pharmacol. 48, 369-386. Kelly, J. S., Krnjevic, K., Morris, M. E., and Yim, G. F. W. (1969).Exp. Brain Res. 7, 1 1-3 1 . Kendall, D. A., Stancel, G. M., and Enna, S. J. (1981). Science 211, 1183-1185. Kendall, D. A,, McEwen, B. S., and Enna, S. J. (1982a). Brain Res. 236, 365-374. Kendall, D. A., Strong, R., and Enna, S. J. (1982b).I n “Cellular and Molecular Mechanisms of Aging in the Nervous System” (G. Gilogamo, E. Giacobini, and A. Vernadakis, eds.). Raven Press, New York pp. 211-221. Keynes, R. D. (1962a).J. Physiol. (London) 163, 16P.

210

SALVATORE J. ENKA A N D JOEL P. GALLAGHER

Keynes, R. D. (1962b). IH/.Cougt-. Yhj.siu/. Sri.. 12th 1, 563-564. Killam, K. F. (1957).J. f‘hamm-d. Esp. Tlro-. 119, 163-271. Killam. K. F.. and Bain, J. A. ( 1 9 5 7 ~ 3Phnrtncicd. . E Y ~Tho.. . 119, 255-262. Koketsu, K.. Shoji, T., and Yamamoto, K. (1974). Expcrinitin 30, 382-383. Kriz, N., Sykova, E., Ujec, E., a n d Vyklicky, L. (1974).J. Phyiol. (Lotdoti) 238, 1-15. Krnkevic, K. (1974). Ph\-sio/. K n i . 54, 518-540. Krnjevic, K. (1976).It/ “GABA in Nervous System Function” (E. Roberts, T. N. Chase, and D. €5. Tower, eds.), pp. 269-281. Raven Press, New York. Krnjevic, K. (1981). I t ( “Amino Acid Neurotransmitters (F. V. DeFeudis and P. Mandel, eds.). pp. 231 -237. Raven Press, New York. Krnjevic, K., and Morris, M. E. (1974). Cott. J. Phsjiol. Phnrtacol. 52, 852-871. Kmjevic, K., and Schwartz, S. (1966). . Y o t i ~ t -(Lotrduti) ~ 211, 1372-1374. Krnjevic, K., a n d Schwartz, S. (1967). hi).B m i n R m 3, 320-336. Krnjevic, K., Randic, M., a n d Straughan, D. W. (1966).J. Physisiol. (Lordon)184, 49-77. Krogsgaard-Larsen, P. (1978). Itr “Amino Acids as Chemical Transmitters” (F. Fonnum, ed.). pp. 305-321. Plenum, New York. Krogsgaard-hrsen, P., and Falch. E. (1981). .\lo/. Cr/l B i o e h i . 38, 129- 146. Krogsgaard-Larsen, P., Johnston, G. A. R., Curtis. D. R., Game, C. J. A,, and McCullough, R. M. (1975).J. .Ywroc/rtwz. 25, 803-809. Krogsgaard-Larsen, P., Scheel-Kruger, J . , and Kofod, H., eds. (1979). “GABANeurotransmitters.” Munksgaard, Copenhagen. Krogsgaard-Larsen, P., Snowman, A., Lummis, S. C., a n d Olsen, R. W. (1981). J . .Yc,trrorlwm. 37, 40 1-409. Kuhar, J. J. (1978).Iri “Neurotransmitter Receptor Binding” (H. I. Yamamura, S. J. Enna, and M. J. Kuhar, eds.), pp. 113-126. Raven Press, New York. Langmoen, I . A., Anderson, P., Gyerstad. L., Mosfeldt, L. A., and Ganes, T . (1978).Actu I-’h!.do/. Srcllirl. 102, 27. Lev)’, R. A. (1977). f’I.fJg. sV/’tIrObiO/. 9, 21 1-267. Lippa. A. S., Critchett, D. J., Ehlert, F., Yamamura, H. I., Enna, S. J., and Bartus, R. T. .4git/g 2, 3-8. (198 1 ). .Y~UJ-O/J~U/. .IlcBurne,. R. X., and Barker, J. L. (1978). .Yutiiw (Lotidoon) 274, 596-597. Macdonald, R., and Barker, J. L. (1978). . Y d i w (Lutzdotr) 271, 563-564. Macdonald, R., a n d Young, A. B. (1981). .\fd. C d / . Rioclirm. 38, 147-162. McGeer. E. G., McGeer, P. L., a n d McLennon, H. (196l).J. ,Vcrtrochm. 18, 36-49. Maggi, .\., and Enna, S. U . (1980).j. ~YtfII‘(JChfvtf,234, 888-892. Maggi, A . , Satinover, J., Oberdorfer, M., Mann, E., a n d Enna, S. J. (1980).Rrnin Rt.c. Bull. 5, 167-171. Mandel, P.. and DeFeudis, F. V., eds. (1979). “GABA-Biochemistry and CNS Function.” Plenum, N e w York. Massotti, ?&.. Guiclotti, .4.. and Costa, E. (1981)..]. ,Vvuro\ri. 1, 409-418. Mathers, D. A., and Barker, J. L. (1980). Scinzcr 209, 507-509. Mathers, D. A , , and Bai-ker, J. L,(1983). I t t t . KO$. .Yrut-obio/. (in press). Mathers. D. A., Jackson, M. B., Lecar, H., a n d Barker, J. L. (1981). Biophjs. J . 33, 14a. Maurer, R. (1979). S m r o w i . Lctt. 12, 65-68. Mayer. M. I... Higashi. H., Shinnick-Gallagher. P., and Gallagher, J. P. (1981). B m i n Rts. 222,204-208. iMazzari, S., Leon, A., Massotti, M. Guidotti, A,, a n d Costa, E. (1980). I n “Psychopharmacolog and Biochemistry of Neurotransmitter Receptors” (H. I. Yamamura. R. W. Olsen, a n d E. Usdin. eds.), pp. 607-615. Elsevier/North-Holland, Amsterdam. Meldrum, B. (1981). .4&. Biorhmt. P.~~rhophatt/lrrro/. 26, 207-217.

CHARACTERISTICS OF MAMMALIAN

GABA

RECEPTORS

21 1

Mitchell, P. R., and Martin, I. L. (1978). Nature (London) 274, 904-905. Mohler, H., and Okada, T. (1977). Nature (London) 267, 65-67. Mohler, H., and Okada, T. (1978). Mol. Pharmacol. 14, 256-265. Nakamura, J., Gallagher, J. P., and Shinnick-Gallagher, P. (1982). Abstr. Commun., Adz!. Phamtacol., Proc. Int. Congr., Sth, 1981 p. 1441. Nelson, P. G. (1975). Cold Spring Harbor Symp. Quunt. Biol. 40, 359-371. Nicoll, R. A., and Alger, B. E. (1979). Znt. R m . Neurobiol. 21, 219-258. Nistri, A,, and Constanti, A. (1979). Prog. Neurobiol. 13, 117-235. Obata, K. (1974). Brain Res. 73, 71-88. Obata, K., Takeda, K., and Shinozaki, H. (1970). Exp. Brain Res. 11, 327-342. Olsen, R. N. (1976).112 “GABA in Nervous System Function” (E. Roberts, T. N. Chase, and D. B. Tower, eds.), pp. 287-304. Raven Press, New York. Olsen, R. N., and Enna, S. J. (1983). In “Anxiolytics: Neurochemical, Behavioral, and Clinical Perspectives” (J. B. Malick, S. J. Enna, and H. I. Yamamura, eds.), pp. 55-76. Raven Press, New York. Olsen, R. N., Greenlee, D., VanNess, P., and Ticku, M. K. (1978). I n “Amino Acids as Chemical Transmitters” (F. Fonnum, ed.), pp. 467-486. Raven Press, New York. Olsen, R. W., Leeb-Lundberg, F. and Napias, C. (1980). Bruin Res. Bull. 5,217-221. Palacios, J. M., Niehoff, D. L., and Kuhar, M. J. (1979). Brain Res. 179, 390-395. Peck, E. J., Schaefer, J. M., and Clark, J. H. (1973). Biochem. Biophjs. Res. Comrnun. 52, 394-400. Pickles, H. (1979). Br. J. Pharmacol. 65, 223-228. Pickles, H., and Simmonds, M. A. (1980). Neuropharmacology 19, 35-38. Polc, P., and Haefely, W. (1976). ~ a u ~ ~ y t ~ - S c h m ~ ~ dArch. e b mPharmucol. gs 294, 121-131. Purpura, D. P., Girado, M., Smith, T. G., Callan, D. A., and Grundfest, H. (1959). J. Neurochem. 3, 238-268. Ransom, B. R., and Barker, J. L. (1976). Bruin Res. 114, 530-535. Ransom, B. R., and Nelson, P. G. (1975). Handb. Psyhopharmacol. 2, 101-127. Ransom, B. R., Bullock, P. N., and Nelson, P. G. (1977).]. Neurophysiol. 40, 1163-1177. Reggiani, A., Barbaccia, M. L., Spano, P. F., and Trabucchi, M. (1980). Psychopharmacology 67,261-264. Robbins, J. (1959).J. Physiol. (Lmdon) 148, 39-50. Roberts, E. (1956).Prog. Neurobiol. 1, 11-25. Roberts, E. (1979). Adu. Pharmacol. Ther. 2, 43-65. Roberts, E., and Frankel, S. (1950). Fed. Proc., Fed. Am. Soc. Exp. Biol. 9, 219. Roberts, E., Chase, T. N., and Tower, D. B., eds. (1976). “GABA in Nervous System Function.” Raven Press, New York. Ryall, R. W. (1978). Trends Neurosci. 1, 164-166. Sano, K., and Roberts, E. (1963). Biochem. Pharmacol. 12, 489-502. Schmidt, R. F. (1971). Ergeb. Phyiol., Biol. Chem. E x f . Pharmakol. 63, 20-101. Schwartzkroin, P. A. (1975). Brain Res. 85,423-436. Simmonds, M. A. (1978). Br. J . Pharmacol. 63, 495-502. Simmonds, M. A. (1980a). Neur@harmacology 19, 39-45. Simmonds, M. A. (1980b). Nature (London) 284, 558-560. Simmonds, M. A. (1981). Br. J . Pharmacol. 73, 739-747. Snodgrass, S. R. (1978). Nature (London) 273, 392-394. Stanton, H. C., and Woodhouse, F. H. (1960).J. Pharmacol. Exp. Thm. 128, 233-242. Steiner, F. A,, and Felix, D. (1976). Nature (London) 260, 346-347. Strasberg, P., and Elliott, K. A. C. (1967). Can.]. Biochem. 45, 1795-1807. Study, R. E., and Barker, J. L. (1983). Proc. Natl. Acad. Sci. U.S.A. (in press).

212

SALVATORE J. ENNA A N D JOEL P . CALLAGHER

Tallman. J. F., Thomas, J. W., and Gallagher, D. W. (1978). Natut-e (London) 274,383-385. ten Bruggencate, G., and Engberg, I. (1968). Brain Res. 11, 446-450. ten Bruggencate, G., and Sonnhof, U. (1972). PJIuegms Arch. 334, 240-252. Thalman, R. H . , Peck, E. G., and Ayala, G. F. (1979).Proc. Soc. Nfurosci. 5, 747. Ticku, M. K., and Huffman, R. D. (1980).Eur.J. Pharmcol. 68, 97-106. Ticku, M. K., and Olsen, R. W. (1978). L f e Sci. 22, 1643-1652. Ticku, M. K., and Olsen, R. W. (1979). Seurophnrmacolog?. 18, 315-318. Ticku, M. K., Ban, M., and Olsen, R. W. (1978a). Jfol. Phnrmarol. 14, 391-402. Ticku, M. K., Van Ness, P. C., Haycock, J. W., Levy, W. B., and Olsen, R. W. (1978b).Broin R u . 150, 642-647. Toffano, G., Guidotti, A., and Costa, E. (1978). Proc. S a f i . Acad. Sn‘. U.S.A. 75,4024-4028. Tran, V. T., Snyder, S. H., Major, L. F., and Hawley, R. J. (1981).Ann. Neurol. 9,289-292. Udenfriend, S. (1950). Fed. Proc., Fed. A m . Sor. Exp. Biol. 9, 240. Varon, S., Weinstein, H., Kakefuda, T., and Roberts, E. (1965). Biochrm. Phaimacol. 14, 12 13- 1224. Williams. M., and Risley, E. A. (1979).j.Seurochunr. 32, 713-718. Willow, M., and Johnston, G. A. R. (1980). A4‘mrosci.Left. 18, 323-327. Willow, M., and Johnston, G. A. R. (1981).J.Smrosci. 1, 364-367. Wong, R. K. S . . Prince. D. A., and Basbaurn, A. I. (1979).Proc. Natl. Acad. Sci. U.S.A. 76, 986-990. Yamarnura, H. I., and Enna, S. J.. eds. (1981). “Neurotransmitter Receptors,” Part 2. Chapman & Hall, London. Yamamura, H. I., Enna, S. J., and Kuhar, M. J., eds. (1978). “Neurotransmitter Receptor Binding.” Raven Press, New York. Yoneda, Y., and Kuriyarna, K. (1980).Brain Rex 197, 554-560. Yousufi, M. A., Thomas, J. W., and Tallman, J . F. (1979). Life Sci. 25,463-470. Zukin, S. R., Young, A. B., and Snyder, S. Y. (1974). Proc. Natl. Acad. Sci. U.S.A. 71, 4802-4807.

SYNAPTIC MECHANISMS AND CIRCUITRY INVOLVED IN MOTONEURON CONTROL DURING SLEEP Michael H. Chase Brain Research Institute and Departments of Physiology and Anatomy School of Medicine University of California, Lor Angeles

Lor Angeler, California

I. Introduction . . . . . . . . .

leep and Wakefulness.. .

A. Brainstem anisms ......................................... A. Extracellular Reflex Studies

VIII. Summary Statements

240

..........

1. Introduction

The following is a review of the central neural control of motoneuron activity during sleep and wakefulness in the cat.' Extracellular and intracellular experiments are discussed to elucidate the basic motor control mechanisms that operate during wakefulness (W), quiet sleep (QS), and active sleep (AS). The data have been slightly reordered in this article, with respect to their publication dates, so that a comprehensive presentation could be developed within a specific thematic framework. Throughout this article the term motoneuron designates alpha motoneuron

213 INTERNATIONAL REVIEW OF NEUROBIOLOGY, VOL. 24

Copyright 0 1983 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-366824-7

214

MICHAEL H . CHASE

II. Overview

In the 1950s, sleep was revealed to consist of two separate states whose attendant physiological and mental processes were different not only from each other, but also from wakefulness as well (cf. Jouvet, 1967; Moruzzi, 1972). In large measure, the intervening years have been spent cataloging these differences. Studies have been carried out dealing with EEG patterns, muscle tone, memory retention, dream recall, heart rate, and respiratory activity, to list just a few of the many processes that have revealed state dependencies (Chase, 1972). I n spite of numerous investigations, the neuronal circuitry responsible for the initiation and maintenance of the sleep states, and for the generation of statedependent patterns of activity, have escaped detection. There is an enormous gap between our descriptive knowledge of state-dependent phenomena and our understanding of the mechanisms that control them. For example, although we may know when and where the pontogeniculate-occipital (PGO) spikes of AS arise, we are not aware of the precise neuronal processes that initiate this or any other statedependent pattern of physiological activity. Presumably such patterns are based upon unique discharge in neuronal systems which occurs specifically during a given sleep state. And yet, with few exceptions, it has been difficult even to develop a consensus that there is sleep-selective discharge within any specific group of central nervous system neurons. Where then, might there be a strategically efficacious beginning to the investigation of the behavioral states of sleep and their correlated patterns of physiological activity? It would seem, since the states of sleep are behaviors that can be described (and even defined) on the basis of somatomotor activity, that an analysis of state-dependent changes in motor processes might lead to an understanding of the key mechanisms responsible for these states. At the very least, a line of investigation of this nature has a clearly discernible point of origin-atonia during ASthat is unambiguous. T h e research strategy of my laboratory is to backtrack along known pathways and processes from the state-dependent physiological process of somatomotor atonia during AS to the central mechanisms that control this phenomenon. At every stage of the evolving exploration, even though the terminus is unknown, each successive study can be founded on a linked sequence of data based on this fundamental physiological component of AS. We expect that an understanding of the basic mechanisms underlying the atonia of AS will shed light on explorations of the foundations of this state, and also indirectly highlight the basis of motor control and other state-dependent phenomena during QS and W as well.

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

215

The progression of our studies of the mechanisms responsible for somatomotor atonia during AS, and related patterns of motor control during QS and W, began with an extracellular examination of the spontaneous state-related amplitude fluctuations of a simple monosynaptic reflex. The ensuing data served as the basis for the discovery of reticular response-reversal (described in following sections), which was an important motor control “signpost” leading us from the periphery to the brainstem reticular formation. Our extracellular studies were then recapitulated by intracellular experiments performed in the unanesthetized, undrugged, normally respiring chronic cat; these intracellularly derived data established the foundation for the development of a model of state-dependent motor control that is described in the last section of this article.

111. Somatic Reflex Activity during Sleep and Wakefulness

The somatic reflex is a basic component of an animal’s behavior. In our initial exploration of motor control mechanisms during the behaviors of sleep and wakefulness, an analysis of reflex modulation was considered to be a logical avenue to pursue. There were also a number of very specific reasons for choosing to study somatic reflex activity. At an extracellular level of analysis the somatic reflex is easily recorded in the chronic freely moving cat as a discrete time-locked event (i.e., contraction of a somatic muscle) following afferent stimulation (Chase, 1974). The fluctuations in reflex amplitude are consistent across states, between animals, and over time (Chase, 1974). Moreover, the factors that modulate its activity can be analyzed in a relatively straightforward fashion. Previous studies had demonstrated changes in reflex excitability at the level of the spinal cord during sleep and wakefulness (Pompeiano, 1967); we were interested in determining whether a monosynaptic reflex, organized within the brainstem, evidenced similar variations in activity. I n our initial study we examined the amplitude fluctuations of the brainstem jaw-closing masseteric reflex (Chase et al., 1968). T h e monosynaptic connection of this reflex is located entirely within the pons (Figs. 1 and 4). Afferent fibers whose cell bodies lie within the mesencephalic V nucleus link proprioceptive receptors within the masseter muscle to trigeminal motoneurons; when these motoneurons discharge, the masseter muscle contracts. In our experiments we induced the masseteric reflex by direct electrical excitation of sensory cells within the

216

MICHAEL H . CHASE

asseter nerve

7 1 -----Interlor

---- Mvlohvoid

nerve

dental nerve

Masreteric reflex

FIG. 1 . Reflex activity of the masseter musculature (A‘) was obtained by electrical stimulation of the ipsilateral mesencephalic V nucleus (A). T h e bipolar recording electrodes within the masseter musculature are not shown. Calibration line: 10 msec; 200 p V .

mesencephalic V nucleus. T h e monosynaptic reflex response was recorded either from the masseter nerve or from the muscle itself. (In our subsequent intracellular studies, we recorded directly from masseter motoneurons.) In the freely moving adult cat, the mean amplitude of the masseteric reflex was largest during active W, smallest during AS, and of an intermediate size during QS (Chase et al., 1968). This general overview, however, does not reflect many of the dynamic changes that occurred within each state, for intrastate fluctuations in reflex amplitude were timelocked to phasic patterns of central neural activity and the animal’s behavior. During wakefulness, analyses were undertaken (1) when the cat was immobile as well as mobile and (2) when there were no eye movements, few eye movements, and many eye movements. In general, the amplitude of the masseteric reflex was positively correlated with the degree of behavioral and ocular activity. When the cat was awake, but quiet and immobile, the reflex was small, compared to its amplitude when the animal was mobile or hyperexcited. Against a background of W, an increased amplitude of response occurred in conjunction with episodes

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

217

of rapid eye movements (REMs). Waking periods that comprised few eye movements were correlated with a reduction in response (Fig. 2). On the other hand, when there were REMs during AS, the reflex amplitude was suppressed phasically even beyond the tonically reduced level present during AS without REMs. The preceding variations in brainstem monosynaptic reflex activity were found, in general, to be comparable to changes in excitability reported for spinal cord reflexes (Pompeiano, 1967). Thus, at all levels of the neuraxis, the principal and most dramatic aspect of these statedependent changes is the almost complete abolition of reflex responsiveness which occurs exclusively during AS. I n order to follow our overall research strategy, we felt that the next step would be to obtain precise information concerning the mechanisms responsible for the suppression of reflex activity during AS. We asked the following key questions: Is the reduction in reflex amplitude and the atonia of AS due to the advent of inhibitory postsynaptic potentials (IPSPs), or a result of the withdrawal of excitatory postsynaptic potentials (EPSPs)? When there is motor activity (twitches) during AS, is this achieved by the advent of EPSPs, or the withdrawal of IPSPs? I n order to attend to these questions, we developed two chronic cat preparations in which it was possible to monitor the membrane potential activity of identified trigeminal and lumbar motoneurons during W, QS, and AS. We then examined the state-dependent synaptic mechanisms controlling motoneuron activity within the brainstem and spinal cord. WAKEFULNESS

W

0

ACTIVE SLEEP

53

5a

40

x

*O

5

\

!-j10 LI W

U

NO

FEW MANY NO FEW MANY EYE MOVEMENTS

FIG. 2. This figure depicts the amplitude changes of the masseteric reflex during the eye movement and no-eye movement periods which accompany wakefulness and active sleep. The amplitude of the reflex was plotted on an arbitrary but relative scale. During wakefulness, in conjunction with an increase in the frequency of eye movements, the masseteric reflex was augmented. During periods of rapid eye movement of active sleep, the reflex was maximally depressed. Thus, as the density of eye movements increased during wakefulness, the masseteric reflex increased in amplitude. As the density of eye movements increased during active sleep, the amplitude of the masseteric reflex decreased. (Reprinted from Chase, 1974.)

218

MICHAEL H . CHASE

IV. Motoneuron Membrane Potential during Sleep and Wakefulness

A. BRAINSTEM MOTONEURONS i n the preceding section, the brainstem monosynaptic jaw-closing reflex (ie., the masseteric reflex) was described as being depressed during AS as compared to QS or W, and further depressed during bursts of REMS within AS. As the first in a series of related studies, a determina-

FIL. 3. Schematic diagram of a chronic cat with its head in a precise stereotaxic plane to allow for the localization of subcortical nuclei and to assure stability for intracellular recording. The reference electrode (A) is located in the temporal muscle. T h e intracellular electrode carrier is shown with a micropipette (B) being lowered by a microdrive (C) through a hole to penetrate through the cerebellum in order to record from trigeminal jaw-closer motoneurons within the motor V nucleus. (D) is a cable connecting permanently placed electrodes with stimulating and recording equipment. (E) is a hole in the acrylic resin and calvarium that allows electrodes to be placed in cerebral structures, acutely, for the duration of an experimental session. (Reprinted from Chase t t al., 1980.)

219

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

n

l -

6

CTr

/JL 4.0 48

16

14

12

10

8

6

4

PE

0

A2

4

P2

0 A2 4

6

:m

1

4rnrn

FIG. 4. (A) Sagittal sections of brainstem showing relationship beween various brainstem structures and the intracellular micropipette located in the motor V nucleus. The recording and stimulating electrodes are drawn approximately to scale. (B) Anatomical and electrophysiological relationship between the mesencephalic V nucleus and the motor V nucleus. 7N, Facial nerve; BC, brachium conjunctivum; FTC, central tegmental field; FTG, gigantocellular tegmental field; FTL, lateral tegmental field; IC, inferior colliculus; INP, nucleus interpositus; PG, pontine gray; RN, red nucleus; SC, superior colliculus. (Reprinted from Chase et d.,1980.)

tion was made of the membrane potential variations of trigeminal motoneurons during sleep and wakefulness. Prior to presenting the results of this study, however, the basic procedure for obtaining intracellular records from brainstem motoneurons of intact, unanesthetized, undrugged, normally respiring cats during both sleep and wakefulness will be synopsized. This procedure is described in detail in Chase et al. (1980). Each cat is first anesthetized with sodium pentobarbital and fixed in a heavy-duty stereotaxic instrument (David Kopf Instruments) (Fig. 3). Screw electrodes are inserted in the calvarium over the marginal and sigmoid gyri to monitor EEG activity. A bipolar strut electrode is placed in the mesencephalic V nucleus (Fig. 4B) to induce the trigeminal jawclosing reflex. Another bipolar strut electrode is positioned in the lateral geniculate body to monitor PGO activity. Insulated wires from the cortical screw and bipolar strut electrodes are soldered to a 20-pin female

220

MICHAEL H . CHASE

connector, which is bonded to the calvarium with acrylic resin. Previously established methodologies are employed for monitoring behavioral states and inducing the masseteric reflex in the chronic cat (Chase and McGinty, 1970; Chase et al., 1968). Two additional procedures are performed during surgery to permit intracellular recording from trigeminal motoneurons (Figs. 3 and 4A). The first entails the drilling of a hole in the calvarium to allow for the subsequent insertion of a micropipette into trigeminal motoneurons. Additional holes are made overlying other structures (e.g., the nucleus pontis oralis, the nucleus gigantocellularis) that are to be explored with micro- or macroelectrodes (Fig. 3E). T h e dura is removed and the openings are filled with bone wax. These access holes allow one to stimulate or record from various sites with electrodes attached to standard stereotaxic manipulators during the course of experimentation. T h e second procedure involves the attachment by acrylic resin of plastic (or steel) tubes to the calvarium. One tube is placed in front of and the other behind the head plug (Fig. 3). T h e tubes are designed to receive four calibrated steel bars (two to each side) that can be fixed, in turn, to the stereotaxic apparatus by a Kopf 880 semichronic headholder. After the acrylic resin sets, the four holding bars are removed and the animal is returned to its home cage to recover from these surgical procedures. On the day of experimentation each cat is placed in the stereotaxic apparatus by inserting the four steel bars into the tubes attached to the calvarium. The bars are then fixed to the stereotaxic frame. Thus, the calvarium is returned to a standard stereotaxic orientation and is held rigidly in the stereotaxic frame without the animal experiencing either pressure or pain. A male connector-cable assembly is attached to the female connector plug in order to record EEG and PGO activity and deliver stimuli to the mesencephalic V nucleus. Fine-wire electrodes (I-mm tip exposure, 0.005 in. diameter) are placed in the masseter muscle ipsilateral to the mesencephalic V electrode [to record electromyographic (EMG) activity and stimulate nerve terminals], in the neck musculature (to record EMG activity), and in the skin at the lateral borders of the eyes (to monitor EOG activity). T h e reference electrode for intracellular recording consists of an Ag-AgC1 wire that is coiled and inserted, under local Xylocaine anesthesia, into the temporal muscle by reflecting the skin next to the acrylic mound (Fig. 3A). T h e bone wax plug in the calvarium overlying the cerebellum is taken out and tissue regrowth, when present, is removed. A glass micropipette is then lowered, with a stepping microdrive, into the trigeminal motor nucleus (Figs. 3,4A). For intracellular recording, a micropipette is filled with either 3-M KCl or 2-M potassium citrate; tip

22 1

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

resistances range from 10 to 20 m a . For extracellular field-potential analysis, a micropipette filled with 2-M NaCl (1 to 3 M a resistance) is employed. Stimulation of the mesencephalic V nucleus evokes in the ipsilateral trigeminal motor nucleus a negative monosynaptic field potential with a latency of approximately 0.6 msec and an amplitude of 2-6 mV (Fig. 5B).Stimulation of the ipsilateral masseter nerve induces an antidromic negative field potential of 1-4 mV with a latency of 0.6-0.8 msec. B ANTIDROMIC field potential 6,)

+

ORTHODROMIC DEPTH (mm)

15.8

field potential

+> II

16.2

C Dorsol

'"'"1

6

16.6

17.0 W E

9-

0-

17.4

+

17.8

L

18.2

Ventral

I

,

,

,

,

,

,

05

10

15

20

25

30

AMPLITUDE

,

3.5mV JlmV 2 msec

FIG. 5. Depth analysis of trigeminal motor nucleus. Comparison of field potentials evoked by antidromic and orthodromic stimulation. (A) Schematic stimulation and recording paradigm. Antidromic excitation of trigeminal motoneurons was achieved by bipolar stimulation of the masseter nerve via two hypodermic needles inserted into the masseter muscle (Sl). Orthodromic excitation was induced by stimulation of the mesencephalic V nucleus by a fixed bipolar strut electrode (S2). Extracellular field potentials were recorded by micropipettes filled with 2-M NaCl having a tip resistance of 1-3 M a . (B) Dorsoventral distribution of antidromic and orthodromic field potentials recorded within the motor V nucleus. When a micropipette was lowered in a dorsoventral direction, a characteristic negative potential could be localized within the stereotaxically defined region of the motor V nucleus. This circumscribed region was precisely the site in which trigeminal motoneurons could be impaled. Numbers in (B) and (C) indicate the depth in millimeters from the surface of the cerebellum to the tip of the microelectrode, which was oriented at a 30" angle to the frontal plane. (C) Graphic presentation of the peak amplitude of the antidromic (0-0) and orthodromic (0-0) field potentials as a function of depth from the surface of the cerebellum. (Reprinted from Chase et nl., 1980.)

222

MICHAEL H . CHASE

Trigeminal motoneurons, which innervate the jaw-closing muscles (jawcloser motoneurons), are impaled in the region where the field potential is recorded. T h e motoneurons are identified by the presence of a monosynaptic EPSP which arises with a latency of 0.5-0.7 msec following stimulation of the ipsilateral mesencephalic V nucleus. Masseter motoneurons are distinguished by the occurrence of antidromic spikes evoked by stimulation of the ipsilateral masseter nerve. Standard procedures are used €or intracelluiar recording (Chase ct nl., 1980). Experiments are performed on each animal from 1 to 3 successive days. Each animal is reintroduced into the experimental paradigm numerous times over a 2- to 3-month period. 1. Trari.dion f r o m klkkejulness to Quiet Sleep

When the cat was awake and resting quietly [i.e., drowsy (Chase and Sterman, 1967)], passage into QS was accompanied either by a slight increase or no discernible change in the level of membrane potential. T h e principal determining factors appeared to be the prior degree of background muscle tone and the level of arousal. Thus, when the animal was alert or actively moving, or when the tonic EMG activity was at a high level, the subsequent transition to QS was accompanied by hyperpolarization. When the transition to QS was spanned by an extended intervening period of drowsiness, only very slight or no hyperpolarization was noted (Fig. 6). However, in most cases, the membrane potential level remained relatively constant, €or quiet W (i.e., drowsiness) usually was the forerunner of QS. T h e change in behavioral state was also reflected by variations in spike potential activity; that is, the sustained discharge of W, when present, first decreased during QS and then was replaced by brief bursts of spikes (Fig. 6). On a number of occasions we were able to record spike potentials of a masseter motoneuron that were followed by unitary EMG activity with a one-to-one ratio at a fixed latency of about 1 msec. The insets A and B of Fig. 6 depict action potentials that were recorded intracellularly from a masseter motoneuron along with simultaneous records of the potentials, recorded extracellularly, of the masseter muscle fiber group that was innervated by this specific motoneuron. T h e transition from QS to aroused W was accompanied by membrane depolarization (Fig. 7). T h e degree of depolarization was positively correlated with the level of arousal, as indicated in the initial 15-sec period of W in Fig. 7 and the subsequent 15-sec epoch when there occurred an increase in neck muscle activity and membrane depolarization. When quiet W replaced QS, little or no change in membrane potential was noted.

FIG. 6. Intracellular recording from a masseter motoneuron. This figure illustrates the gradual increase in membrane potential and decrease in spike occurrence as the animal changed behavioral state from wakefulness to quiet sleep. An oscilloscopic picture of the activity of an individual masseter motor unit is shown in the inset, which portrays the time-locked relationship between each motoneuron spike potential and an action potential of a masseter muscle unit at increasingly faster time bases (A and B, 1-4). In ( 5 ) , the motoneuron spike is presented at higher gain (spike amplitude, 40 mV). Membrane potential band pass on polygraphic record: low gain, DC to 35 Hz; high gain, DC to 0.5 Hz. Top EEG trace, frontal cortex; bottom EEG trace, marginal cortex. (Reprinted from Chase et al., 1980.)

224

MICHAEL H . CHASE

QUIET SLEEP

WAKEFULNESS

EEG

~

EEG

M

EOG

J

~

J

&

+

W

J

~

W

N

W

W

’

,

*

NECK

-

-

-

.--I

-

-- .

EMG

t

=PJ

------j.€ -50

----~

TRIGEMINAL . MOTONEURON POTENTIAL

----’

-

-_--

--.--

___-

.

- 70 mV 20 SEC

’

Fit.. 7. lntracellular recording from a trigeminal jaw-closer motoneuron: change in

membrane potential during quiet sleep compared to wakefulness. When an extended period of quiet sleep was followed by sustained wakefulness, membrane depolarization occurred. The degree of depolarization was positively correlated with the level of arousal and muscular activity, as portrayed in the middle of the figure when a brief increase in neck EMG activity was correlated with a time-locked decrease in membrane polarization. Membrane potential band pass on polygraphic record: DC to 0.1 Hz. Top EEG trace, frontal cortex; bottom EEG trace, marginal cortex (Reprinted from Chase o f ( I / . . 1980.) Continuous Record OUIET SLEEP

ACTIVE SLEEP

WAKEFULNESS

EEG PGO

EoG

NECK

EMG-.

-____J(mw*(m((m(l

MEMBRbNE--

QUIET SLEEP

Ib

18

19

20

ACTIVE SLEEP

21

L -

P

23

.-~.-.-

24

25

L

Z6

?IT

~.Am- 6

~

$-

iI

FIG. 8. Intracellular recording from a trigeminal jaw-closer motoneuron: correlation of membrane potential and state changes. The membrane potential increased rather abruptly at 3.5 min in conjunction with the decrease in neck muscle tone and transition from quiet to active sleep. At 12.5 min the membrane depolarized and the animal awakened. After the animal passed into quiet sleep again, a brief, aborted episode of active sleep occurred at 25.5 min that was accompanied by a phasic period of hyperpolarization. One minute later, the animal once again entered active sleep and the membrane potential increased. EEG trace, marginal cortex; membrane potential band pass on polygraphic record, DC: to 0.1 Hz. (Reprinted from Chase et nl., 1980.)

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

225

2. Transition from Quiet Sleep to Active Sleep Motoneurons were hyper polarized when AS was compared to QS (Figs. 8 and 9). The degree of hyperpolarization varied from 2 to 10 mV, its development paralleled the various ways in which AS emerges from QS. For example, although the onset of AS is demarcated by EEG desynchronization and a reduction in muscle tone, these indices are not always present at the same time, and either may precede the other as the animal enters into the AS state. Moreover, EEG desynchronization and EMG suppression may develop slowly or rapidly. Accordingly, hyperpolarization was correlated in some cases with relatively abrupt EEG desynchronization and EMG suppression (Fig. 8, 3- to 4-min time marks), whereas at other times it continued to develop beyond the period of the initial onset of EMG suppression (Fig. 8, 26- to 27-min time marks). When the EMG of the muscle units innervated by the recorded motoneurons was monitored, a perfect correlation was observed (Fig. 6). As the animal changed state from QS to AS, there was an accompanying cessation of spontaneous spike activity (Fig. 9). During AS, isolated or short bursts of spike potentials were occasionally observed in conjunction with facial twitches and rapid eye movements (REMs).

3. Transatioli from Active Sleep to Wakefulness T h e transition from AS to W most often takes place in 1 to 2 sec. In all cases, when the animal awoke from AS, the membrane potential rapidly depolarized (Fig. 8, 12- to 13- and 25- to 26-min time marks). T h e degree of depolarization was equal to or exceded the level maintained during the preceding episode of QS. 4. Summary

These data demonstrate that the membrane potential of trigeminal jaw-closer motoneurons is strongly hyperpolarized during AS compared to QS and W, and is only slightly hyperpolarized or remains at the same potential level when QS is compared to W. This pattern of membrane potential modulation reflects variations in the extracellularly monitored amplitude of the jaw-closing reflex during sleep and wakefulness; that is, the reflex is depressed during AS while its amplitude is only slightly reduced during QS compared to W (Chase et al., 1968). B. SPINAL CORDMOTONEURONS

Extracellularly, at the level of the spinal cord, the modulation of tonic and reflexively induced somatomotor activity has been well studied and

MEMBRANE-----'-POTENTIAL SPIKE ACTIVITY

-_ - _-

....--%--,P---,

L

LIIL--nM.AaL

-75

-.

.

-

--

-

-, -.,

- _-_---1-80,

-+ 10 sec

F I ~ .9. . Intracellular recording from a trigeminal jaw-closer motoneuron: correlation of spike activity and membrane potential with change of state from quiet to active sleep. Note the abrupt cessation of spike activity coincident with membrane hyperpolarization at the transition from quiet to active sleep. Spike activity (amplitude = 52 mV) portrayed via a window discriminator and Schmitt trigger. (Reprinted from Chase cf d..1980.)

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

227

reviewed in detail by Pompeiano (1967) and Steriade and Hobson (1976). On the basis of EMG indices, the level of activity is practically constant across the states of quiet Wand QS; however, during AS, there is a marked reduction in tone and reflex activity. Two lines of evidence have indicated that this profound reduction in somatomotor activity is the result of a decrease in the excitability of motoneurons. First, there is a decrease in the amplitude of the recurrent discharge of motoneurons (monitored electromyographically) in the deafferented hindlimb (Gassel et al., 1965). Second, there is a decrease in the motor response to direct electrical stimulation of the ventral horn (Kubota and Kidokoro, 1965; Morrison and Pompeiano, 1965). These results suggest the occurrence of motoneuron hyperpolarization during AS. However, solely on the basis of these data, it is also possible that disfacilitation could cause the reported decrease in excitability. We therefore decided to determine the underlying causal process by utilizing intracellular recording techniques, as we had previously done at the level of the brainstem. In order to record intracellularly from spinal cord motoneurons in the chronic cat, we combined procedures developed for brainstem intracellular recording (Chase et al., 1980) with new techniques designed to immobilize portions of the vertebral column and antidromically identify spinal cord motoneurons. These methodologies are outlined below, portrayed in Figs. 10 and 11 , and described in detail in Morales et al. (198 1). Each animal is first anesthetized and then prepared for the implantation of electrodes and immobilization of the head region in the identical fashion described in Section IV,A. While these procedures are underway, with appropriate surgical intervention, hard plastic (Delrin R) clamps, sculpted to follow the vertebral contours, are placed around L3, L4, and L6. The tips of the clamps pass ventrally around the vertebral body, as shown in Fig. 10 (inset). Stainless steel screws are placed in the articular processes of L3, L4, L5, and L6. The screws and clamps provide a secure foundation for the subsequent application of acrylic resin, which bridges all exposed vertebral surfaces. T h e spinous process of L5 is resected, and a small hole (2 mm in diameter) is drilled in the caudal portion of the lamina of L5, 1.5 mm lateral to the midline. T h e underlying dura matter is cut, and the hole is filled with bone wax. One steel holding bar is placed between the L3 and L4 clamps, and another is positioned immediately rostra1 to the L6 clamp. Both holding bars project transversely to the vertebral axis (Fig. 10, inset). The bars are designed to be attached at a later date to a second stereotaxic frame which is oriented back to back with respect to the stereotaxic frame holding the cat’s head (Fig. 10). T h e exposed vertebrae, clamps, and

228

MICHAEL H. CHASE

FIG. 10. Diagrammatic illustration of the chronic cat preparation used to record intracellularlv from spinal cord motoneurons. The intracellular electrode carrier is shown with a micropipette (A) being lowered by a microdrive (B) through a hole in the dorsal lamina of L5 in order to record from cells in the ventral horn (L7-S1 motonuclei). (C) is a cable connecting permanently placed electrodes with stimulating and recording equipment. Inset depicts one of the clamp and bar configurations used to immobilize the vertebral column. Head fixation has been described previously. (Reprinted from Morales pt a/., 1981.)

holding bars are bound together with acrylic resin. Only the area above the lamina of L5 is left uncovered. Preceding the initial experimental session, chronic hind limb nerve electrodes are implanted while the cat is briefly anesthetized with 4% halothane (Fig. 11). The insulated wires emerging from these electrodes are passed subcutaneously to exit in the region of L3 to L5 where they are affixed to the acrylic mound and capped except during experimentation.

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

229

On the following day, the initial experimental session begins by first immobilizing the animal's head and vertebral column via the chronically implanted restraining devices (Fig. 10). EEG, EOG, and EMG activity are monitored in the fashion previously described for intracellular recording from the brainstem of the chronic cat. The animal is removed from the stereotaxic frame after experimentation during the day, fed and exer-

-

1 rnsec

l~rnv

05z e c

L7

FIG. 11. Diagrammatic section of lumbar spinal cord showing relationships between the recording micropipette, motoneuron pool, and peripheral nerves (including the stimulating cuff electrodes). The strategy of neuronal identification is depicted by the intracellular records shown in (A-D). In (A), suprathreshold stimulation of the common peroneal nerve results in an antidromicaily activated potential. By lowering the stimulus intensity below threshold for excitation of the peripheral motor axons, either an orthodromic spike (B) or EPSP (C) is induced. The latency of these potentials, their configuration, and the relatively low intensity of the initiating stimulus reflects their monosynaptic la origin. (D) shows an IPSP which arises following low-intensity stimulation of the tibia1 nerve. In (A'), (B'),and (C') are high-gain, time-expanded records corresponding to (A), (B), and (C). The latency of the antidromic spike is 1.6 msec, and the conduction velocity of the motor axon is 84 m/sec. The conduction velocity of the afferent fibers is 90- 100 d s e c , based on a synaptic delay and central conduction time of 0.5 to 0.7 msec. The latency of the IPSP is 2.1 msec. Peripheral nerve stimulation: (A) 0.9 V, 0.4 msec; (B) 0.7 V, 0.4 msec; (D) 0.9 V, 0.5 msec. (Reprinted from Morales et al., 1980.)

230

MICHAEL H . CHASE

cised, and returned to its cage. Experimental sessions covering successive days are employed on numerous occasions spread out over a period of two to three months. Utilizing this preparation, intracellular activity was recorded from antidromically identified lumbar motoneurons. When neurons were penetrated during W or QS, the mean resting potential was 65 mV SD t 6.7 mV); the range was 55 to 75 mV. T h e mean antidromic spike amplitude was 6 9 mV with a range from 55 to 95 mV. Cells were rejected for analysis when their initial membrane potential was less than 55 mV. Recording periods ranged from 12 min to 2.5 hr; their mean duration was 40 min. All records included for analysis contained multiple state transitions. In some motoneurons we were able to monitor the membrane potential throughout two to three consecutive sleep-waking cycles (ix., with each cycle comprising a sequence of W + QS + AS -+ W. During W, periods of membrane depolarization as well as spike activity (which is not reproduced in the filtered polygraphic record) occurred in conjunction with phasic movements of the ipsilateral leg (Fig. 12, e.g., at 27 and 34 min). N o dramatic change in the level of membrane polarization was present when quiet W was compared to QS (Fig. 12). However, there was a slight increase in the level of polarization accompanied by a reduction in muscle tone when active or aroused W was compared to QS. Overall, the level of motoneuron membrane polarization did not exhibit dramatic fluctuations between W and QS. Essentially, the intracellular data provided a good reflection of extracellularly monitored patterns of somatomotor activity during W and QS (Chase, 1974, 1978; Pompeiano, 1967). When the animal passed from QS to AS, there occurred hyperpolarization of the motoneuron membrane (Figs. 12 and 13). T h e average magnitude of hyperpolarization was 6.7 mV with a range of 4 to 10 mV. When the animal awoke from AS, the level of polarization always decreased (Figs. 12 and 13). Comparable results have been obtained by others (Glenn P t d.,1978). During periods of REMs, there were further increases in hyperpolarization. Occasionally, when there were twitches of the leg, spike potentials were generated. As discussed at the beginning of this section, motoneuron hyperpolarization had been hypothesized on the basis of extracellular studies to be responsible for the depression of tonic activity of spinal cord motoneurons during AS. However, by intracelluiarly monitoring the degree of membrane polarization, we were able to demonstrate that, in fact, spinal cord motoneurons, as well as brainstem motoneurons, do become hyperpolarized during AS compared with W or QS.

Continuous Record

E OG

-65

MEMBRANE LUMBAR MOTONEURON POTENTIAL ---u"--

-70

- 75mV 2

(MIN)

' 4

'

6

'

b

' 1 0 ' 1 ~ ' 1 4 ' I 6 ' 1 ~ ' 2 b ' 2 2 ' 2 4 ' 2 6 ' 2 8 ' 3 b ' 3 2 ' 3 ~ ' 3 6 ' 3 8

I' 1I-

-65 -70 -75mV 40

42

44

46

48

50

52

54

56

58

60 1 HR

62

64

66

68

70

72

74

76

FIG. 12. Intracellular record from a lumbar motoneuron during sleep and wakefulness: correlation of membrane potential and behavioral state. This neuron was recorded from for 1 hr and 17 min; during this period the animal exhibited numerous cycles of wakefulness and quiet deep. There were no striking changes in the mean level of polarization between these two states except when phasic periods of membrane depolarization with accompanying spike generation occurred during wakefulness (spike activity is not evident because the DC record, for display purposes, was passed through a 0.1-cps high-frequency polygraphic filter). After recording 56 min, the animal entered into a brief episode of active sleep which was accompanied by a corresponding period of membrane hyperpolarization. The intracellular record was obtained with a potassium citrate-filled micropipette. T h e first and second polygraph traces are those of EEG activity recorded from left and right frontal-parietal cortex, respectively. (Reprinted from Morales and Chase, 1978.)

232

MICHAEL H . CHASE QUIET SLEEP

.

CL"

ACTIVE SLEEP

WAKEFULNESS

..-

-

.

. I I

EMG

I

. . 1 L. 1.1

.I

I100pv

.-55

MEMBRANE POTENTIAL

_-

LUMBAR MOTONEURON I^_

,-

A' NEC-K -BICEPS

FEMORISILYMICL

-

.-:

20 sec

FIG. 13. Intracellular record from a lumbar motoneuron during sleep and wakefulness: correlation of membrane potential and behavioral state. This figure highlights membrane hyperpolarization which accompanies active sleep. Hyperpolarization commenced prior to the cessation of muscle tone. which was accompanied by a further and rather sharp increase in membrane polarization [(A), and shown oscilloscopically at higher gain and expanded time base in (A')]. At the termination of active sleep, the membrane depolarized coincident with the resumption of muscle tone and behavioral awakening (B, B'). Note the brief periods of depolarization during active sleep and wakefulness, which were accompanied by phasic increases in muscle activity (i.e., muscular twitches during active sleep and leg movements during wakefulness). Spike potentials often occurred during these periods of depolarization but are not evident in this figure because the DC record was passed through a 0. I-cps high-frequency polygraphic filter. This motoneuron was recorded for 28 min; the traces shown were obtained 12 min after the cell was impaled. The first and second polygraph traces are those of EEG activity recorded from left and right frontalparietal cortex, respectively. (Reprinted from Morales and Chase, 1978.)

V. Synaptic Mechanisms Responsible for Motoneuron Hyperpolarization during Active Sleep

The two basic mechanisms which may be responsible for motoneuron membrane hyperpolarization during AS are postsynaptic inhibition and/or the disfacilitation of tonically active presynaptic elements. Both

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

233

mechanisms may be operating continuously, be superimposed upon one another at certain times, or act individually for restricted periods.

A. BRAINSTEM MOTONEURONS The synaptic mechanisms responsible for modulating trigeminal motoneuron activity during sleep and wakefulness are described in this section. Emphasis is placed on the processes responsible for the previously reported decrease in somatic reflex activity and the hyperpolarization of motoneurons during AS. T h e data that will be presented indicate that hyperpolarization of trigeminal motoneurons is caused, at least in part, by postsynaptic inhibition. This conclusion is based on a confluence of information obtained by studying various indices of synaptic activity. First, the characteristics of the antidromic spike were examined during QS and AS. Two types of antidromic blockade were observed during the transition into AS. The first type was characterized by a progressive slowing of invasion of the soma-dendritic (SD) spike, as well as by its fragmentation and an increase in its time-to-peak (Chandler et al., 19803). Eventually, as the AS state developed, only an initial segment (IS) spike of 10-25 mV remained. A second type of antidromic spike blockade was characterized by complete failure of spike invasion into the initial segment (Fig. 14). As the animal’s state changed from QS to AS and the membrane hyperpolarized, antidromic spikes could still be elicited for a brief period (Fig. 14A,B). At this point in time, the block of the full spike was all or none in character; persistent block was maintained when the animal progressed further into AS (Fig. 14C,C’). During the transition into AS, antidromic spikes exhibited a decrease in peak potential (Chandler et al., 1980a). The decrease in antidromic spike summit level appeared to be independent of the degree of membrane hyperpolarization. In the acute cat preparation Llinas and Terzuolo (1964) found a comparable decrease in the antidromic spike during hyperpolarization induced by reticular stimulation. They concluded that hyperpolarization of motoneurons was accompanied .by an increase in membrane conductance. Our results, including a decrease in amplitude and an increase in the decay rate of the falling phase of the trigeminal Ia-induced monosynaptic EPSP during AS (Fig. 15), indicate that membrane hyperpolarization during AS is due to an increase in membrane conductance (Chandler et al., 1980a). This conclusion, of course, does not exclude the possibility that disfacilitation may also be present.

Q U I E T SLEEP E P

E

G

*

G

O

A C T I V E SLEEP W W

V

T

U I 1.b 4 6 -4-L

EOG E M G NECK

-

T

-__Lxc-

:

-

'

5sec

IlOO~V

'

A

C

L

_c_

1

-.V m O -.-l--~- . . . - . -

4 msec

FIG. 14. Development of complete block of antidromic spike potentials in a masseter motoneuron recorded intracellularly with a potassium citrate electrode during transition from quiet to active sleep. (A and B) Oscilloscopic traces of antidromic spikes recorded during the period of active sleep shown in (C'). Note that the antidromic spike is indicated on the polygraph record by the use of a Schmitt trigger to give a pulse output synchronous with the antidromic spike. This is labeled as the antidromic unit in the polygraphic trace. Membrane potential band pass on polygraph record was DC to 0.5 Hz. Antidromic stimulus parameters, 1 Hz, 10 V, 0.05 msec. (Reprinted from Chandler ef n l . , 1980a.)

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

B

A QUIET SLEEP

235

ACTIVE SLEEP

C

11

I

;

I

0

I

I

2

I

3

I

I

4

I

I

5

msec

FIG. 15. Increase in rate of decay and decrease in peak amplitude of an averaged EPSP during active sleep as compared to quiet sleep. (A and B) Averaged EPSP (n = 25) recorded in a jaw-closer motoneuron with a potassium citrate electrode during quiet (resting potential -67 mV) and active sleep (resting potential -72 mV). Arrows indicate onset of stimulus to mesencephalic V nucleus. (C) Semilogarithmic plot of time course of decay for EPSP recorded during quiet (0)and active sleep (0).(Reprinted from Chandler et al., 1980a.)

However, when taken in concert, the data suggest that postsynaptic inhibition of masseter motoneurons occurs during AS and that this process is responsible for the tonic reduction in reflex amplitude and increase in polarization that we reported previously (Chase et al., 1974, 1980).

B. SPINAL CORDMOTONEURONS Having demonstrated that lumbar motoneurons are hyperpolarized during AS compared to QS or W (Morales and Chase, 1978),we turned our attention to an exploration of the extent to which postsynaptic inhibition of these motoneurons during AS can account for reflex depression and atonia. The results of these experiments, which are described below, demonstrate that hyperpolarization of lumbar motoneurons during AS is due to postsynaptic inhibition and that postsynaptic inhibition is also responsible for the phasic episodes of decreased motoneuron excitability and augmented hyperpolarization accompanying bursts of REMs during this state (Morales and Chase, 1978, 1981).

236

MICHAEL H. CHASE

1 . Antidromic Field Potential The antidromic field potential amplitude was depressed throughout the entire period of AS, indicating a decrease in excitability of populations of spinal cord motoneurons during this state (Morales and Chase, 1981). In addition, phasic suppression was observed during intense bursts of REMs, indicating an increase in “suppressor” influences. In a comparable study, the antidromic field potential of the trigeminal motor nucleus was found to be relatively smaller during AS compared to QS or W (Chandler et al., 1980a).

2. A ntidroiiik Actim Potential T h e IS-SD delay was lengthened when the animal entered AS, further reflecting the conclusion that motoneuron excitability is depressed during this state (Fig. 16). During bursts of REMs, in association with an increase in membrane polarization, the IS-SD delay was further prolonged and there were phasic changes in the peak amplitude of the antidromic spike. These variations in IS and SD spikes are indirect evidence of an increase in conductance of the soma membrane (Brock et al., 1953; Coombs ei al., 1957; Krnjevic et al., 1977; Llinas and Terzuolo, 1964); similar changes have been reported during postsynaptic inhibi-

0.2 msec I lomv 0.5Ksec FIG. 16. Antidromic spikes and their electrically differentiated records during quiet (A, A’) and active sleep without rapid eye movements (B, B’). T h e IS-SD delay is more prominent during active sleep than during quiet sleep. Electrical differentiation of the antidromic spike revealed this phenomenon even more clearly as the delay in transmission between the IS and SD components of the antidromic spike became quite evident. (A’) Electrical differentiation of‘ (A); (B‘) electrical differentiation of (B). Time constant, 0.7 msec. Peripheral nerve (tibial) stimulation, 3 V, 0.1 msec. (Reprinted from Morales and Chase, 1981.)

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

237

tion arising from excitation of an antagonist nerve (Curtis and Eccles, 1959) or during stimulation of the reticular formation Uankwoska, 1968; Llinas and Terzuolo, 1964).

3. Action Potential Generation by Intrasomatic Stimulation Rheobasic currents during QS were comparable to those during W, whereas strong depression of excitability was found when AS was compared to QS (Table I). Episodes of REMs during AS were accompanied by a further phasic suppression of excitability. Although elevated thresholds were always accompanied by hyperpolarization, it alone cannot completely account for the increase in rheobasic current (Morales and Chase, 1981).An analysis of the data from rheohasic determinations and input resistance indicate that current flow is “shunted” by an increase in membrane conductance during AS; moreover, hyperpolarization and increased conductance appear to contribute to the observed increase in rheobasic current. TABLE I THRESHOLD CURRENTS (DURATION 20 MSEC) FOR 14 LUMBAR MOTONEURONS DURING QUIET SLEEP A N D ACTIVE SLEEP WITHOUT RAPID EYE MOVEMENTS“ Percentage increase

Nanoamps Neuron 1 2 3 4 5 6 7 8 9 10 11 12 13 14

AS

(V

x 100)

6.0 8.5 3.7 7.5 7.0 4.0 6.0 5.5 4.5 8.0 5.9 7.0 1.5.0 7.5

9.0 25.0 17.5 9.0 9.5 10.0 9.0 14.0 11.0 10.0 9.5 11.0 19.0 10.3

50 194 366 20 36 150 50 154 144 25 61 57 27 37

X = 6.86 SD = 2.75

X = 12.41 SD = 4.82

x = 98b SD = 96.3

Each number represents the average of 10 determinations in each state. Paired t test,p < ,001. (Reprinted from Morales and Chase, 1981.)

238

MICHAEL H . CHASE

4. Orthodromic Actimtion T h e amplitude of the monosynaptic EPSP was smaller during AS compared to QS (Fig. 17). T h e fact that during AS there was a decrease in excitability and an increase in membrane conductance, together with motoneuron hyperpolarization, clearly indicate that the EPSP amplitude is depressed by a mechanism of postsynaptic inhibition (Burke, 1967; Cook and Cangiano, 1972; Curtis and Eccles, 1959). Phasic periods of further suppression of EPSP amplitude were observed during intense bursts of REMs (Fig. 17). These periods were generally accompanied by hyperpolarization of the postsynaptic membrane, an increase in conductance, and a decrease in excitability. Thus, it is evident that phasic postsynaptic inhibitory processes affect the size of the EPSP during bursts of REMs. Reductions in EPSP amplitude that were not accompanied by detectable membrane potential changes occasionally were present, indicating the possibility of presynaptic or dendritic inhibition; however, intrasomatic penetrations do not allow one to clearly differentiate between these t w o mechanisms.

5 . ,\lotoneurou Input Resistance There was no difference in membrane input resistance when QS was compared to W. During AS, there was a striking decrease in resistance as

0

C y--J--ymv

1msec

F I ~ .17. . Averaged EPSP activity ( n = 32) during quiet sleep (A), active sleep without rapid eye movements (REMs) (B), and active sleep with REMs (C). Note the decrease in amplitude when active sleep without REMs is compared to quiet sleep and a further decrease during active sleep with REMs. T h e averaged peak amplitude decreased 19%) during active sleep without REMs and 37% during active sleep with REMs. Peripheral nerve (common peroneal) stimulation, 0.5 V, 0.3 msec. (Reprinted from Morales and Chase, 1981.)

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

239

TABLE I1 MEMBRANEINPUTRESISTANCE OF 1 1 CELLS DURING THE STATES OF QUIET A N D ACTIVE SLEEPWITHOUT RAPIDEYEMOVEMENTS" SLEEP Percentage decrease

Megohms AS

(-

QS

x 100)

Neuron

QS

1 2 3 4 5 6

1.25 2.80 1.21 0.53 1.50 4.30 1.25 1.05 1.69 2.90 1.42

0.97 1.95 0.43 0.21 0.46 1.44 0.80 0.76 1.33 1.75 0.92

22.4 30.3 64.4 60.3 69.3 69.5 36.0 27.6 21.3 39.6 35.2

X = 1.81 SD = 1.08

X = 1.00 SD = 0.55

X = 43.2b SD = 18.8

7 8 9 10 11

a

Each number is an average of 10 determinations in each state.

* Paired t test,p < .05. (Reprinted from Morales and Chase, 1981.) shown in Table 11. The resistance decrease during AS ranged from 70 to 2 1%, and was statistically significant in a paired t test ( p < 0.05). During AS with REMs, there were frequent phasic decreases in the voltage drop produced by the same current, indicating a continuously changing membrane input resistance. These findings are entirely consistent with the hypothesis that postsynaptic mechanisms are promoting motoneuron inhibition (Araki and Terzuolo, 1962; Burke and Rudomin, 1977; Gustafsson and Lipski, 1980; Smith et al., 1967). Anomalous rectification cannot account for the decrease in absolute amplitude of both the antidromic and direct spike during AS because these changes are absent during direct hyperpolarization of the motoneuron with current injection, whereas they are present during conditions of postsynaptic inhibition (Frank and Fuortes, 1956; Nelson and Frank, 1967). In addition, hyperpolarizing currents have a negligible effect on monosynaptic EPSP amplitude (Coombs et al., 1955; Gallego et al., 1979; Nelson and Frank, 1967; Shapovalov and Kurchavyi, 1974; Werman and Carlen, 1976), which we found to be significantly reduced during AS. Although it is possible that disfacilitation could be present during AS, it cannot be directly demonstrated when recording from a postsynaptic cell that is already subjected to strong postsynaptic inhibitory input. In summary,

240

MICHAEL H . CHASE

the data indicate that the principal, and probably sufficient, spinal cord mechanism underlying h ypotonia and hyporeflexia during AS is postsynaptic inhibition.

VI. Central Control Mechanisms

A. EXTRACELLULAR REFLEXSTUDIES In conjunction with the preceding studies of the spontaneous modulation of reflex excitability and motoneuron membrane potential during sleep and wakefulness, we examined the central neural control of these somatomotor processes. Two measures of somatomotor excitability were employed to document the effect of conditioning stimulation during sleep and wakefulness. Accordingly, in conjunction with stimulation of the reticular formation, an analysis was undertaken of the amplitude fluctuations of the trigeminal jaw-closing reflex, recorded electromyographically, and the membrane potential activity of trigeminal motoneurons, recorded intracellularly. Our experiments were designed to reveal the fashion in which the reticular formation controls somatomotor activity during W, QS, and AS. We reasoned that the excitation of a central neural area that is responsible for maintaining motor control throughout a particular behavioral state would be expected to yield patterns of somatic activity which normally accompany that state. Thus, if a neural area promotes motor activity that is comparable to that which occurs during a given state, this might indicate that it was a key area responsible for maintaining not only the motor aspects of the state, but possibly the state itself as well. A neural area that induces a contrasting pattern of motor activity is likely to be functionally related to specialized activities, and not be primarily involved with the maintenance of the state or its tonic behavioral components. We concentrated our analysis on the reticular core of the brainstem on the basis of its importance in sensory, motor, and integrative activities (Hobson and Brazier, 1980). These studies began with an investigation of the state-dependent modulation of the masseteric reflex by examining its control by reticular sites located at mesencephalic, pontine, and medullary levels of the brainstem. Very simply put, w e wanted to determine whether the reflex response following stimulation of various reticular regions remained the same or varied when the animal changed behavioral state. We expected

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

24 1

that the degree of control would differ only in potency or degree in a state-dependent fashion; that is, whatever modulatory effects were exerted during wakefulness would persist, to a greater or lesser degree, during the sleep states. To our surprise, we found this to be the exception, not the rule. Excitation of most areas yielded diametrically opposite response patterns during AS, compared with those that were induced during W or QS (Chase, 1978). 1 . Mesencephalon: Mesodiencephalic Junction a. Pattmns of RejZex Modulation and Their Topographic Distribution. T h e pattern of reflex modulation obtained with a standard 20-msec conditioning-test interval from a total of 51 stimulation sites during W, QS, and AS is summarized in Fig. 18. The position of each site is represented on a schematic brainstem cross-section on the left side of the figure. The effects of high (1.0 mA) and low (0.2 mA) conditioning currents are presented on the right. Reflex facilitation was observed during W and QS from sites in a region corresponding to the rostra1 extent of the mesencephalic reticular formation; however, the majority of sites were ineffective in modifying the reflex amplitude (Fig. 18). Only minor differences between W and QS were observed when reflex facilitation occurred during these states. As soon as the animal entered AS, a striking change emerged in the effect of mesodiencephalic conditioning stimulation. During AS, every site tested at 1.0 mA evoked suppression of the masseteric reflex. The effective suppressor region extended throughout the entire extent of the mesodiencephalic junction. With low conditioning current (0.2 mA) a few of the previously effective sites failed to modify reflex excitability: however, facilitation with either low or high conditioning current was never observed. b. Response Thresholds. Increasing or decreasing the amplitude of the conditioning current during any state led to an increase or decrease, respectively, in the magnitude of the induced response, but it never changed the direction of the response. Higher current was required to produce an effect (most often facilitation) during W and QS than was necessary to induce an effect (always suppression) during AS (Wills and Chase, 1979). Consequently, with low conditioning current, many sites that yielded reflex modulation during AS were ineffective in producing reflex facilitation during other states. c. Time Course of Response. The predominant reflex response at this level of the neuraxis (i.e., state-dependent suppression of the masseteric reflex that emerged only during active sleep) began at 10 to 15 msec, was maximum at 20 msec, and continued until 30 msec. Stimulation during

+

3

+

o

+

/n

+

m

0

0

s9v

7 :

I

0

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

243

W and QS produced reflex facilitation with a peak at a conditioning-test latency of 20 msec. 2. Pons: Pontomesmephalic Junction a. Patterns of R$ex Modulation and Their Topographic Distribution. T h e distribution of the 40 stimulation sites examined at this level of the brainstem are presented on a schematic section of the pontomesencephalic junction shown on the left of Fig. 19. The patterns of response at a standard conditioning-test interval of 20 msec are summarized on the right. During W and QS, stimulation induced either reflex facilitation or was ineffective. A well-defined locus for the facilitatory effect was found in the vicinity of the nuclei pontis oralis and reticularis mesencephali (Fig. 19A,B). This region was surrounded by an area that produced no modification of reflex excitability during these states. When the animal was in AS, stimulation of almost all sites led to reflex suppression. The effective region encompassed not only the reticular nuclei but also extended into the central gray, and included the superior colliculus. Generally, the effect of a lowered conditioning current was to decrease the magnitude of reflex suppression and to reduce the size of the effective region. b. Response Thresholds. T h e response threshold was comparable during W and QS; most sites required a relatively high current to induce facilitation during these states. T h e few low-threshold sites that were found were clustered in the general vicinity of the reticular nuclei. However, during AS there were many sites with very low current thresholds which were widely distributed throughout the tectum, central gray, and ventral tegmentum. In practically all cases, higher conditioning current was required to induce reflex facilitation during W and QS than was necessary to promote reflex suppression during AS. c. Time Course of Response. During W and QS, facilitation usually began at a short conditioning-test interval of 5 to 10 msec and reached a peak at approximately 20 msec. During AS, profound reflex suppression consistently occurred between 10 and 30 msec. A slight degree of facilitation was usually present between 5 and 10 msec. From a small number of sites another pattern of response was observed which consisted of prominent reflex suppression during AS with virtually no effect during W or QS. In these instances, the time course of suppression during AS paralleled that described above for other sites during this state. I n summary, throughout this region there was remarkably consistent suppression of reflex activity during AS at a conditioning-test interval of 10 to 30 msec.

Pontomesencephafic Junction

QS

W

0

5

10

J

AS

1

0

5

10

5

10

0

5

I0

,

mm

A=Sites Examined +=Facilitation -=Suppression

=No Effect

J

FIL. 19. Summary of state-dependent effects of pontotnesencephalic stimulation on masseteric reflex excitability during wakefulness (W), quiet sleep (QS), and active sleep (AS). On the left is a schematic cross-section of the pontomesencephalic junction. Stimulation sites are indicated by (A).The reflex response following the application of two levels of conditioning current (1.0 and 0.2 mA) at a conditioning-test latency of 20 msec are presented. A (+) indicates an increase of 50% or greater in the conditioned (test)reflex amplitude; a ( - ) indicates a 50% or greater reduction in amplitude. Note that all effective sites for reflex modulation yielded only facilitation during both wakefulness and quiet sleep, and only suppression during active sleep. BC, Brachium conjunctivum; BP, brachiurn pontis: FCT, central tegmental tract; IP, nucleus interpenduncularis; LM, medial lemniscus; MES, nucleus of the mesencephalic tract of V NPO, nucleus pontis oralis; NRM, nucleus reticularis mesencephali; P, pyramidal tract: SC, superior colliculus; SCG, stratum griseum centrale; TOL, tectoolivary tract; TS, tectospinal tract. (Reprinted from Wills and Chase, 1979.)

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

245

3. Medulla

Conditioning-test stimulation of medullary sites led to two distinct effects consisting of short-latency responses, which seldom extended beyond 5 msec, and long-latency responses, which lasted from 10 to 40 msec with a peak at approximately 20 msec. The short-latency responses were identical throughout W, QS, and AS, whereas the long-latency effects changed as a function of state. T h e threshold for the induction of long-latency responses was markedly lower than was the threshold for the short-latency responses. For these reasons, only the effects induced at the longer conditioning-test interval will be discussed. In addition, since no consistent pattern of effect was found following stimulation of lateral medullary sites, responses from this region will not be described at the present time. [See Chase and Wills (1979) for a complete presentation of all data.] During W and QS, conditioning stimulation of medial medullary loci reliably produced reflex facilitation which lasted approximately 30 msec and peaked at 20 msec. During AS, the identical stimulus produced reflex suppression with a time course that was a mirror image of the facilitatory pattern.

4. Summary At these three brainstem levels, conditioning stimuli, when effective during W or QS, facilitated the masseteric reflex at conditioning-test intervals of approximately 10 to 30 msec. T h e peak of this facilitatory effect occurred at latencies of 15 to 25 msec. T h e principal sites that produced facilitation lie in the medial half of the brainstem throughout its rostral-caudal extent in the mesencephalon, pons, and medulla. Within this core of reticular tissue are the nuclei reticularis mesencephali, pontis oralis and caudalis, gigantocellularis, and parvocellularis. Facilitation of the masseteric reflex from these sites is replaced during AS by reflex suppression at approximately the same conditioning-test intervals with a peak effect at a comparable latency. This phenomenon of somatomotor facilitation during W and QS, that is replaced by inhibition during AS, has been called reticular response-reversal (Chase and Babb, 1973). T h e induction of reflex suppression during AS was not restricted to reticular regions but was obtained throughout all brainstem regions studied, that is, from medial and lateral sites within the mesencephalon, pons, and medulla. The minimal amount of current sufficient for producing reflex suppression during AS was less than that required to produce facilitation during W or QS. I n addition, suppression during AS was obtained from almost every site in the brainstem, whereas only a portion of the sites also produced facilitation during W and QS.

246 B.

MICHAEL H . CHASE

INTRACELLULAK

STCDIES

1. Trigemi~inlLlfotoneurons

An examination of cellular mechanisms was deemed essential in order to differentiate the processes underlying the pattern of statedependent reticular response-reversal described in the previous section. We sought to determine whether reflex facilitation during W and QS is due primarily to the advent of EPSPs and/or to the withdrawal of inhibition, and whether reflex suppression during AS results from a process of postsynaptic inhibition and/or disfacilitation [see Chandler et al., (1980b) for a detailed report]. These intracellular investigations, which essentially recapitulated our extracellular studies, are diagrammatically presented in Fig. 20. T h e neuronal connections between the various stimulation sites and the Mosseter m

n

Semilunar ganglion

Inf alveolar n s3 -rl-

FIL. 20. General schematic diagram of paradigms for analyzing the effects of stimulation of the pontornesencephalic reticular formation (PMRF) on jaw-closer motoneurons during sleep and wakefulness. Trigeminal jaw-closer motoneurons were identified by antidromic invasion of the motoneurons, which was achieved by stimulation of motor axons (S,), and by orthodromic excitation of motoneurons, which was achieved by stimulation via Stimulaa permanently fixed bipolar strut electrode in the mesencephalic V nucleus tion of the inferior alveolar nerve (&) was carried out in order to obtain a control for inhibitory synaptic input produced by stimulation of the PMRF (S,) in the area depicted by the transverse section of the brainstem at A2. (Reprinted from Chandler ~t a/., 1980b.)

(s).

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

A

B

247

C

15 r n ~ N. pontis oralis

10 msec

FIG. 2 1. Intracellular potentials recorded from a trigeminal jaw-closer motoneuron induced by PMRF stimulation (N. pontis oralis) during wakefulness (A), quiet sleep (B), and active sleep (C). PMRF stimulation induced a predominantly depolarizing potential during wakefulness and quiet sleep, and a hyperpolarizing potential during active sleep. All records are single sweeps and were obtained with a KCI electrode from the same cell with a membrane potential of -50 mV. Bottom trace in (C)is extracellularrecord. N. pontis oralis: four pulses (500 Hz), 4 V, 0.1 msec. (Reprinted from Chander et al., 1980b.)

motor nucleus of the Vth nerve are shown in this figure. A transverse section of the brainstem at A2 and the reticular area in which an electrode to deliver conditioning stimulation was located are also portrayed. During W and QS, stimulation in the vicinity of the nucleus pontis oralis (one to four pulses, 500 cps) evoked a depolarizing potential with an amplitude of 1-3 mV, a duration of 20 to 80 msec, and a peak latency of approximately 30 msec (Fig. 21A,B). There were no qualitative differences between the configuration of the potential induced in trigeminal motoneurons during W and that of QS. During AS, the identical stimulation parameters used during W and QS produced a hyperpolarizing potential in these motoneurons (Fig. 21C). The latency of the hyperpolarizing potential ranged from 10 to 15 msec. Its peak amplitude of 3-7 mV occurred at a latency of 20-30 msec. The duration of this potential extended from 20 to 30 msec. In a previous section of this article it was pointed out that during W and QS the masseteric reflex is facilitated by nucleus pontis oralis stimulation at conditioning-test intervals of approximately 25 msec, and suppressed during AS by the identical stimulus throughout the same conditioning period (Chase and Babb, 1973). Figure 22 illustrates the cellular correlates of this phenomenon of reticular response-reversal, that is, depolarization during W and QS and hyperpolarization during AS. To determine if these induced potentials during W, QS, and AS were actually EPSPs or IPSPs, as opposed to potentials generated by a mechanism of disinhibition or disfacilitation, current was passed through the recording electrode to alter the cell's membrane potential (Fig. 23B-E). An increase in hyperpolarizing current produced an increase in the amplitude of the depolarizing potential during QS and W. This suggests that it is an EPSP (Eccles, 1957). During AS the application of depolariz-

248

MICHAEL H . CHASE

Wakefulness A

Quiet Sleep G

Mesencephalic V.

Active Sleep M

L1 2 0 r n ~ B

H

N

C

I

0

0

J

P

E

K

0

F

L

R

N. pontis oralis

10 mSec

F[<.. 22. Intracellular potentials recorded from a trigeminal jaw-closer motoneuron: effect of conditioning stimulation of the N . pontis oralis on mesencephalic V-induced activity at various conditioning-test intervals during sleep and wakefulness. Records in (A-L) were obtained from a cell with a membrane potential of -50 mV, whereas records in (M-R) are from a different cell with a resting potential of -54 mV. (A, G) Control EPSPs induced by stimulation of trigeminal mesencephalic nucleus. (B, H) Control synaptic potentials induced by stimulation of N. pontis oralis. The test trigeminal mesencephalic EPSP was facilitated when evoked during the period of the N. pontis oralis-induced depolarization during wakefulness (C-F) and quiet sleep (ILL). Note the production of an action potential in (C, D, 1, and J). (M) is control action potential induced by mesencephalic V stimulation. ( N ) Control synaptic potential induced by stimulation of N . pontis oralis during active sleep. Note suppression of the action potential during the hyperpolarizing phase of the synaptic potential of N. pontis oralis origin (0,P). Bottom traces in (G, H, M and N) are extracellular records. Records taken with a KCI electrode. Mesencephalic V nucleus: 4 V, 0.1 msec in (A-L); two pulses (500 Hz), 5 V, 0.1 msec in (M-R). N. pontis oralis: four pulses (500 Hz), 4 V,0.1 msec in (B-F) and (H-L); four pulses (500 Hz), 10 V, 0.1 msec in (N-R). The 5-mV calibration applies to all records except (M) where a 20-mV calibration applies. (Reprinted from Chandler rt nl., 1980b.)

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

249

Wakefulness B.

+&A

C. +6nA

A. Control

N. pontis oralis D. -4nA

E. -6nA

Active Sleep G. t5nA

F.

N. pontis oralis

-

H. +7nA

Control

I. -5nA

J. -7nA

10 msec

FIG. 23. Intracellular potentials recorded from a trigeminal jaw-closer motoneuron: effect of polarizing currents on synaptic potentials induced by stimulation of the N. pontis oralis during wakefulness and active sleep. Records in (A-E) were obtained from a motoneuron with a membrane potential of - 50 mV, records in (F-J) were obtained from a different motoneuron with a membrane potential of -50 mV. (A) and (F) are control synaptic potentials. Effects of polarizing currents on the synaptic potentials are shown in (B-E) and (G-J). Records are the superposition of several traces obtained with potassium citrate electrodes. Bottom traces in (A) and (F) are extracellular records. N. pontis oralis: four pulses (500 Hz), 8 V, 0.1 msec in (A-E); four pulses (500 Hz), 6 V, 0.1 msec in (F-J). (Reprinted from Chandler et nl., 1980h.)

ing current through the recording electrode increased the amplitude of the hyperpolarizing potential, whereas hyperpolarizing current decreased its amplitude and occasionally reversed its polarity (Fig. 23G-J). This response of the hyperpolarizing potential during AS to intracellular current injection demonstrates that it is an IPSP (Eccles, 1957). A likely candidate for mediating the IPSP induced by .stimulation of the nucleus pontis oralis during AS (and possibly the atonia of AS) is the medullary reticular formation (Chase and Willis, 1979; Pompeiano, 1967; Wills and Chase, 1979). Anatomical (Edwards, 1975; Graybiel, 1977) and electrophysiological (It0 et al., 1970; Mancia et al., 1974) evidence suggests the existence of projections which are excitatory from the pontomesencephalic reticular formation to the medullary reticular formation. Stimulation of the medullary reticular formation produces in-

250

MICHAEL H . CHASE

hibition of brainstem reflex activity (Chase and Wills, 1979). Inhibitory projections to cervical motoneurons from the nuclei reticularis ventralis and gigantocellularis have also been observed (Peterson et d.,1978). These data and o u r preceding experiments provided the impetus for an exploration of the interactions between cells in and in the vicinity of the nucleus pontis oralis and the medullary reticular formation, and an examination of the latter’s role in promoting reflex suppression and somatomotor atonia during AS. Accordingly, extracellular stimulation was applied to the nucleus pontis oralis and intracellular records were obtained from medullary neurons during sleep and wakefulness, as described in the following section.

2. .Iledullary Retrcu Iar Netiroris Intracellular records were obtained from 70 reticular neurons located within and in the vicinity of the nucleus reticuiaris gigantocellularis (NGC). Neurons were penetrated during W, QS, and AS. N o change in the animal’s state occurred during the period of intracellular recording in 5 1 neurons. State transitions were present while monitoring the activity of the remaining 19 cells. During transitions between W a n d QS, there was no definitive change in the level of membrane potential (Fig. 24A). In contrast, during AS the membrane potential invariably depolarized by 3 to 14 mV (8.9 5 3.5 mV) (Fig. 24B). Tonic membrane depolarization was observed only when AS occurred; it was maintained throughout this entire state of sleep. There was a positive correlation between the onset and duration of membrane depolarization and the onset and duration of AS. In addition, when spike activity was present during the transition period prior to AS, the frequency of its discharge not only increased at the onset of AS concurrent with membrane potential depolarization, but it was also maintained for the duration of the state (Fig. 25). Thus, there was an inverse relationship during AS between the spike activity of medullary neurons, which was generated tonically, and the discharge of motoneurons, which was suppressed tonically (Chase et al., 1980; Morales and Chase, 1978). A functional interpretation of these data, combined with that of the literature described below, suggest that NGC neurons may be responsible for the maintenance of tonic somatomotor inhibition during AS. In the acute cat preparation, stimulation of the medial bulbar reticular formation results in the suppression of EMG activity that is produced either reflexively or by excitation of the motor cortex (Magoun and Rhines, 1946). In experiments on anesthetized cats, stimulation in this region induces postsynaptic inhibitory potentials in flexor and extensor

25 1

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

A

WAKEFULNESS

EEG

EMG MEMBRANE

NECK

QUIET SLEEP

1 -

f'

RETICULAR

IlOO/lV

- 50

NEURON

-60

7

POTENTIAL

B

MEMBRANE POTENTIAL

-70mV

ACTIVE SLEEP

QUIET SLEEP

-

-

RETICULAR NEURON

- 40 - 50 -M)T'V

I

30 SEC

I

FIG.24. Intracellular records from two neurons of the nucleus reticularis gigantocellularis. This figure illustrates the maintenance of an unchanging membrane potential during wakefulness compared with quiet sleep (A). When the animal changed behavioral state from quiet sleep to active sleep, the membrane potential gradually began to depolarize (B). When the active sleep state was firmly established, the membrane potential stabilized at a level approximately 10 mV more depolarized than during quiet sleep (B). Membrane potential band pass on polygraphic record: DC to 0.5 Hz. (Reprinted from Chase et al., 1981.)

motoneurons (Jankowska et al., 1968). In chronic extracellular experiments, certain medullary neurons discharge selectively only during AS and are relatively silent even in the freely moving preparation (Kanamori et al., 1980; Netick et al., 1977). These convergent data lines indicate that medullary neurons are linked to generalized inhibitory functions during AS, suggest that they may be responsible for specific patterns of motoneuron inhibition during W, and exclude them from responsibility for the promotion of somatomotor excitation during AS and W.

VII. Concluding Remarks

We have previously proposed a model of state-dependent motor control wherein there is excitatory input to the NGC during AS that originates from cells within or in the vicinity of the nucleus pontis oralis (i.e., the pontomesencephalic reticular formation) (Chase, 1976, 1980; Chase

QUIET SLEEP EEG EOG P

ACTIVE SLEEP

-

I '-,1kp

-.

1

NECK

EMG P W

c IlOOpV

1-40mV

I

30 SEC

I

FIG. 2 5 . Intracellular record from a neuron of the nucleus reticularis gigantocellularis: correlation of spike activity and memhrane potential with change of state from quiet to active sleep. Note the gradual increase in spike activity coincident with memhrane depolarization at the transition from quiet sleep to active sleep. Spike activity portrayed via a window discriminator and Schmitt trigger. Membrane potential hand pass on polygraphic record: DC to 0.5 Hz.(Reprinted from Chase ef ul., 1981.)

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

253

et al., 198 1). We suggest that during AS, tonic somatomotor inhibition is

mediated either directly or indirectly by the medullary reticular formation which has been selectively activated by pontomesencephalic neurons (e.g., the nucleus pontis oralis) during this state. An as yet unidentified gating mechanism, or perhaps reciprocal feedback circuitry, operating selectively during AS, would then be responsible for the establishment of a link from the rostra1 reticular site to the caudal reticular site. During W and QS the link is not expected to be functional; the pontomesencephalic reticular formation would then be able to exert its well-documented motor facilitatory effects (reviewed by Chase, 1980). We have also suggested that in the absence of pontomesencephalic reticular discharge during AS, the suppression of tonic motor activity would not take place due to the lack of gigantocellular activation (Chase, 1976, 1980). In accord with one aspect of this model, lesions in the pontomesencephalic region have been shown to abolish the tonic suppression of somatomotor activity during AS (Henley and Morrison, 1974; Jouvet, 1967). T h e model is further supported by the finding in our study (Chase et al., 1981) and in another (Mancia et al., 1971) of a preponderance of EPSPs, rather than IPSPs, in medial bulbar neurons following stimulation in the region of the nucleus pontis oralis. An anatomical basis for these responses is present in the work of Sakai et al., (1979), who injected horseradish peroxidase into the medial bulbar region and found retrograde labeled cells within our zone of pontomesencephalic stimulation, thus demonstrating a direct monosynaptic connection linking these two areas. The synaptic basis of reticular response-reversal also accords with the model. During W a prominent EPSP is induced in trigeminal (masseter) motoneurons by stimulation of the nucleus pontis oralis. During AS the EPSP is superceded by an IPSP. These responses mimic spontaneously generated state-dependent patterns of motoneuron excitation and inhibition during W and AS, respectively (Chase and Willis, 1979; Wills and Chase, 1979). We suggest that discharge of neurons of the nucleus pontis oralis is converted to an eventual inhibitory drive by its relay through the medulla. Consequently, we have proposed that neurons of the “reticular activating system” act in concert with a neuronal population of the lower brainstem to promote motor inhibition during AS (Chase, 1980). We have not previously drawn particular attention to one intriguing aspect of these studies, namely, that almost without exception, every brain region that we have examined yields IPSPs in masseter motoneurons and reflex suppression during AS. This includes cortical as well as subcortical sites (see also Chase and McGinty, 1970; Chase, 1980). There appears to be a functional redefinition of central neural systems

2 54

MICHAEL H . CHASE

during this state, since motoneuron membrane hyperpolarization and the suppression of somatic reflex activity arise following excitation of almost all areas, even those that are facilitatory to motor activity during W or QS, as well as those that lack a motor control function during any other state. T h e reversal from motor facilitation to motor suppression in widely disparate regions, the pervasiveness of its occurrence, and the low threshold for its appearance suggest that a common fundamental reorganization of the neural control of motor systems occurs during AS. We conclude that the entire parenchyma of the brain is functionally reorganized during this state, which results in the establishment of functionally potent neuronal circuitry promoting motor suppression. What might be the significance of the emergence of widespread motor suppressor control emanating from diverse brainstem areas during AS? Its function surely, at least in part, is to promote motor suppression and, we suggest, to maintain AS by blocking or masking facilitatory motor responses that would otherwise result from the high levels of neuronal discharge in diverse sensory and integrative sites which occur during this state (Steriade and Hobson, 1976). In this manner, brainstern neuronal circuitry may be functionally altered during AS to “contain” the “arousing” effects of the enhanced discharge rates of widespread regions with diverse responsibilities. An activated nervous system is apparently necessary to perform whatever functions are subserved either during or by the state of AS. Thus, in addition to functions which are carried out by the discharge of neurons in practically all brain regions during AS; their activity, pari passu, may indirectly promote the inhibition of motoneurons. On the basis of today’s knowledge the model, as constructed, is most likely sound, but its strength may be deceptive, for some essential aspects have not been dealt with either in the literature or by our experiments. As with any model, its chief value lies in guiding or generating concepts and providing research strategies to determine their validity. T h e model proposed in this article will undoubtedly be replaced by another as new information is added to the store. However, we do have confidence in the validity of the data which serves as its foundation. I n toto, these data constitute a demonstration of the powerful, dynamic properties of central neural functions and a dependence on behavioral state for their mode of action. VIII. Summary Statements

1. Spontaneous amplitude fluctuations of the brainstem monosynaptic trigeminal jaw-closing reflex were examined in the freely moving chronic cat during wakefulness, quiet sleep, and active sleep. T h e largest

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

255

amplitude responses occurred during active wakefulness; they decreased in size during quiet sleep. The lowest amplitude responses occurred during active sleep. 2. A chronic cat preparation was developed in order to record intracellularly from identified trigeminal motoneurons for prolonged period of time throughout the states of sleep and wakefulness. 3. The membrane potential of trigeminal motoneurons exhibited fluctuations that were correlated with changes in the animal's behavioral state. The fundamental pattern consisted of ( a )slight hyperpolarization during quiet sleep, compared to arousal or alert wakefulness, (b) little if any hyperpolarization during quiet sleep compared to quiet wakefulness, and ( c ) dramatic hyperpolarization when active sleep was compared to quiet sleep. Sustained spike activity of trigeminal motoneurons, when present during wakefulness, decreased in frequency or tended to occur in bursts when the animal was in quiet sleep. During active sleep, activity ceased except for a few isolated spikes or short-duration bursts of action potentials. 4. Based on an analysis of antidromically induced spike potentials and monosynaptically induced postsynaptic potentials, it was concluded that postsynaptic inhibition of trigeminal motoneurons during active sleep acts to suppress somatic reflex activity and produce muscular atonia. 5 . A companion study of the membrane potential of lumbar motoneurons in the chronic, unanesthetized, undrugged, normally respiring cat was performed during sleep and wakefulness. 6. The antidromic field potential, antidromic and orthodromic spike, EPSP, membrane input resistance, and rheobasic current of lumbar motoneurons were studied during sleep and wakefulness. No change in motoneuron excitability occurred when quiet wakefulness was compared to quiet sleep. Postsynaptic inhibition resulted in decrease in excitability during active sleep. Further phasic decreases in excitability, also d u e to postsynaptic inhibition, occurred during active sleep in conjunction with clusters of rapid eye movements. 7. The mesencephalon, pons, and medulla were explored in a conditioning-test paradigm in an attempt to find a site where electrical stimulation induced a pattern of somatomotor reflex and motoneuron membrane potential modulation comparable to that which occurs spontaneously during sleep and wakefulness. In unanesthetized, freely moving cats during wakefulness and quiet sleep, electrical stimulation within and in the vicinity of the nucleus pontis oralis produced facilitation of the masseteric reflex, whereas during active sleep the identical stimulus resulted in potent suppression of the reflex. This phenomenon was termed reticular response-reversal.

256

MICHAEL H . CHASE

8. An omnipotent and omnipresent suppressive action on somatic reflex activity, which was present only during active sleep, originated from mesencephalic, pontine, and medullary levels of the brainstem. This effect was not localized to specific neuronal groups or fiber pathways. Consequently, it appears that the functional sequellae of brainstem activity is redirected during active sleep so as to promote somatomotor suppression selectively during this state. 9. A chronic intracellular recording technique was developed to analyze the synaptic basis for reticular response-reversal. During wakefulness and quiet sleep, stimulation of the nucleus pontis oralis induced a depolarizing potential in trigeminal motoneurons. The identical stimulus that was employed during wakefulness and quiet sleep produced a hyperpolarizing potential in these same cells during active sleep. T h e depolarizing potential of wakefulness and quiet sleep was characterized as an excitatory postsynaptic potential, whereas the hyperpolarizing potential of active sleep was characterized as an inhibitory postsynaptic potential. 10. A model of motor control during sleep and wakefulness was developed, a key element of which was that both spontaneous somatomotor inhibition during active sleep and inhibition induced by the nucleus pontis oralis during this state may be mediated by the nucleus reticularis gigantocellularis. To begin an exploration of this hypothesis, intracellular records were obtained in the chronic cat from gigantocellular neurons during sleep and wakefulness. When wakefulness and quiet sleep were compared to active sleep, the membrane potential level of these neurons gradually decreased; a depolarized membrane potential was maintained tonically and selectively throughout active sleep. These data support the concept that gigantocellular neurons assist in the generation of somatomotor atonia during active sleep, and indicate that this state-dependent inhibitory function may be controlled by the nucleus pontis oralis as part of the general phenomenon of reticular response-reversal. 11. These findings and our model indicate the possibility that the excitability of the cells in and within the vicinity of the nucleus reticularis gigantocellularis may increase during active sleep due to the opening of a neural gate during this state. Accordingly, activity arising in, or coursing through, the brainstem would be effective in suppressing reflex excitability throughout active sleep due to neuronal circuitry that was state-selectively patent, thus allowing a link to be formed between rostra1 and caudal reticular regions. (During wakefulness and quiet sleep we suggest that the gate is “normally closed” and motor facilitation ensues.) It is also proposed that this gate functions to promote motor suppression

MECHANISMS OF MOTONEURON CONTROL DURING SLEEP

257

during active sleep by “containing” the effects of the very high rates of discharge of brainstem motor facilitatory and arousal systems by activating inhibitory regions, thus allowing the active sleep state to be sustained.

Acknowledgments

The research reported in this article was supported by Public Health Service grant NS-09999, National Science Foundation grants BNS-79-12897 and INT-77-22299, and the Japan Society for the Promotion of Science.

References

Araki, T., and Terzuolo, C. A. (1962).J. Neurophysiol. 25, 722-789. Brock, L. G., Coombs, J. S., and Eccles, J. C. (1953).J. Physiol. (London) 122, 429-461. Burke, R. E. (1967).J. Neurqhysiol. 30, 1114-1137. Burke, R. E., and Rudomin, P. (1977).In “Handbook of Physiology” (E. R. Kandel, eds.), 2nd ed., Sect. I, Vol. 11, pp. 877-944. Am. Physiol. SOC.,Washington, D.C. Chandler, S. H., Chase, M. H., and Nakamura, Y. (1980a).J. Neurophysiol. 44, 359-371. Chandler, S. H., Nakamura, Y., and Chase, M. H. (1980b).J. Neurophysiol. 44,372-382. Chase, M. H., ed. (1972). “The Sleeping Brain, Perspectives in the Brain Sciences,” Vol. 1 . Brain Inf. Serv/Brain Res. Inst., University of California, Los Angeles. Chase, M. H. (1974). In “Basic Sleep Mechanisms” (0.Petre-Quadens and J. D. Schlag, eds.), pp. 249-267. Academic Press, New York. Chase, M. H. (1976).I n “Mechanisms in Transmission for Signals for Conscious Behavior” (T. Desiraju, ed.), pp. 99- 121. Elsevier, Amsterdam. Chase, M. H. (1978). SOL. Neurosci. Symp. 3, 33-65. Chase, M. H. (1980).In “The Reticular Formation Revisited’ (J. A. Hobson and M. A. B. Brazier, eds.), pp. 449-472. Raven Press, New York. Chase, M. H., and Babb, M. (1973). Bruin Res. 59, 421-426. Chase, M. H., and McGinty, D. J. (1970).Brain Res. 19, 117-126. Chase, M. H., and Sterman, M. B. (1967).Bruin Res. 5, 319-329. Chase, M. H., and Wills, N. (1979).Exp. Neurol. 64, 118-131. Chase, M. H., McGinty, D. J., and Sterman, M. B. (1968). Experientia 24, 47-48. Chase, M. H., Chandler, S. H., and Nakamura, Y. (1980).J. Neurophysiol. 44, 349-358. Chase, M. H., Enomoto, E., Toshiki, M., Nakamura, Y., and Taira, M. (1981).Exp. Neural. 71, 226-233. Cook, W. A,, Jr., and Cangiano, A. (1972).J. Neurophysiol. 35, 389-403. Coombs, J. S., Eccles, J. C., and Fatt, P. (1955).J. Physiol. (Lmdma) 130, 374-395. Coombs, J. S., Curtis, D. R., and Eccles, J. C. (1957).J. Physiol. (London) 139, 198-231. Curtis, D. R., and Eccles, J. C. (1959).J. Physiol. (London) 145, 529-546. Eccles, J. C. (1957). “Physiology of Nerve Cells.” Johns Hopkins Press, Baltimore, Maryland. Edwards, S. (1975).J. Comp. Neural. 161, 341-358.

258

MICHAEL H . CHASE

Frank. K., and Fuortes, hl. G. F. (1956).J. Phlsiol. (Lmidon) 134, 451-470. Gallego, R., Kurio, hl., Nuriez, R., and Snider, W. D. (1979).J . Physiol. (Lorrdou) 291, 191-205. ( ;assel, . . . hl. M., hlarchiafava, P. L., and Pompeiano, 0. (1965)..4rrh. Ztnl. Biol. 103, 25-44. Glenn, L. L., Foutz, A. S . , and Dement, W. C. (1978). S l c ~ p1, 199-204. Graybiel, A . (1977).J. Comp. Srrirol. 175, 37-78. Gustafsson, B.. and Lipski, J. (1980). Brain RPS. 181, 61-74. Henley, K.. and Morrison, A. R. (1974).d4rtn‘Yrurobid. E.xp. 34, 215-232. Hobson. J . A,, and Brazier, M. A. B., eds. (1980). “The Reticular Formation Revisited.” Raven Press, New York. Ito, M., Udo. M., and Mano. N. (1970).J. .\‘europhjsiol. 33, 210-226. .Jankowska, E.. Lund, S., Lundberg, A., and Pompeiano, 0. (1968). Arch. Ztnl. B i d . 106, 124- 140. Jouvet, M. (1967).Phyiol. Rml. 47, 117-177. Kanamori, N.. Sakai, K., and Jouvet, M. (1980). Brain RPS.189, 251-255. Krnjevic, K., Puil, E., and Werman, R. (1977). ( h i . J . Phjsiol. Phormucol. 55, 658-669. Kubota, K., and Kidokoro, Y. (1965).J/m. J. Physiol. 16, 217-226. Llinas. R.. and Terzuolo, C. A. (1964).J. Srurophjsiol. 27, 579-591. Magoun. H. W., and Rhines, R. (1946).,1. SPurophjsiol. 9, 165-171. Mancia. A., Mariotti, M., and Spreafico, R. (1974).B w i n RPS.80, 41-51. Mancia, %I.. Grantyn, A., Broggi, G., and Margnelli, M. (1971).Brniin Re.s. 33, 491-494. Morales, F. R., and Chase, M. H. (1978). Erp. .Vmrol. 62, 821-827. Morales, F. R.. and Chase, %I. H. (1981). Braiit Re.5. 225, 279-295. . 27, 355-362. Morales. F. R., Schadt, J., and Chase, M. H. (1981). P h p i ~ l Behurt. Morrison, A. R.. and Pompeiano. 0. (1965)..4rrh. Itnl. Biol. 103, 517-537. Moruzzi, G. (1972). Ergeb. Physiol., Biol. C h m . Exp. Phnrmakol. 64, 1- 165. Nelson, P. G., and Frank, K. (1967).J. .vfnrrophysio/. 30, 1097- 1113. Netick, A,. Orem, J., and Dement, W. (1977). B m i u Res. 120, 197-207. Peterson, B. W., Pitts. N. G . , Fukushima, K., and Mackel, R. (1978). Ex/). Brai7i Rvs. 32, 47 1-489. Pornpeiano, 0. (1967). Res. Puhl.-Awr. Re.\. Sei14. ,\ferit. Dls. 45, 351-423. Sakai, K., Sastre, J., Salvert. D.. Touret, M., Khyama, M.. and Jouvet, M. (1979). Brain RPS. 176,233-254. Shapovalov, A . J . , and Kruchavyi, G. C. (1974).Brain Res. 82, 49-67. Smith, T. G., Wuerker, R. B., and Frank, K. (1967).J. xVPurophyid. 30, 1072- 1096. Steriade, M., and Hobson. J. A. (1976). Prug. ,‘Veurobio/. 6, 155-156. U‘erman, R., and Carlen, P. L. (1976). B m i n R P . ~112, . 395-401. Wills, N., and Chase, M. H. (1979). Exp. L\cur-o/. 64, 98- 117.

RECENT DEVELOPMENTS IN THE STRUCTURE AND FUNCTION OF THE ACETYLCHOLINE RECEPTOR By F. J. Barmntes

Max-Planck-lnrtitut fur BiophyrikalircheChemie Gottingen-Nikolausberg, Federal Republic of Germany

I. Introduction ........................................................ The Acetylcholine Receptor: An Archetype? . . . . . . . . . . . . . . . . . . . . . . . . 11. The AChR Molecule . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . , . . . . . . . . . . . . . . . . . . A. Size, Molecular Weight, and Hydrodynamic Properties . . . . . . B. AChR Subunits and Their Stoichiometry , . . . . . . . . . . . . . . . . . . . . . . . . . . C. Primary Structure of the AChR Protein . . . . . . . . . . . . . . . . . . . .. . . . . . . . . D. Secondary Structure of the AChR Protein.. . . . . . . . . . . . . . . . . . . . . . . . . . E. Charge of the AChR . . . . . . . . . . . . . . . . . . ....................... F. Immunochemical Structure of the AChR . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Biosynthesis of the AChR.. . . . . . . . . . . .. ... ... .. . . . . .. . .. IV. Three-Dimensional Topography of the embrane . . . . . . . . . . A. The AChR Oligomeric Structure . . ................... B. Topography of AChR Subunits in t .....,............. V. In Search of the Functional Role of the Nonreceptor v Proteins A. Early Attempts at Assigning a Role to the v Proteins . . . . . . . . . . . . . . . . . B. Influence of the v Proteins on AChR Freedom of Motion.. . . . C. Translational Dynamics of the AChR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. The Possible Participation of Thiol Reactions in v Protein-AChR Interactions ...................................................... E. Topographical Localization of the v Proteins and Its Relationship to Receptor Structure.. . . . ............................ VI. The Ion-Translocation Func -Bound AChR., . . . . . . . . . . . A. Some Problems and The . . . . ... ... ... .. .. .. . .. ... . . B. Flux Dose-Response Curves and Channel Activation . . . . . . . . . . . . . . C. Flux Dose-Response Curves, Binding Equilibria, and Desensitization .. D. New Views on the Beha erated Channels . . . . . . . . . . . . VII. Summary and Perspectives ........................... A. The Coupling between I and Biochemical Techniques ........................................ B. Structural Counterparts of Gating and State of Ligation . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

259 260 26 1 26 1 264 269 270 27 1 27 1 272 279 281 283 290 291 292 295 295 296 30 1 302 304 305 313 329 329 330 33 1

1. Introduction

This article is concerned with recent advances in some areas of research on the nicotinic acetylcholine receptor (AChR). Emphasis is put on the structure and dynamics of the AChR in the membrane, the corre259 INTERNATIONAL REVIEW OF NEUROBIOLOGY, VOL. 24

Copyright 0 1983 by Academic Press, Inc. All rights of reproduction in any form reserved.

260

F. .J. BARRANTES

lation of structure with the ion gating properties as assessed by electrical or flux measurements, and the influence of nonreceptor proteins on AChR properties. T h e article is not intended to give comprehensive coverage of the field. T h e reader is referred to Karlin (1980) and Changeux (198 1) for more general overviews of biochemical aspects; to Fambrough (1979) for the biological and developmental aspects; to Patrick and Berman (1980) for discussion of the developmental studies on the receptor; and to Steinbach (1980), Gage (1976), and Adams (1981) for discussion of the electrophysiological aspects of the AChR. T h e important issue of the coupling between ligand binding and physiological response is not dealt with here, though the two events receive separate treatment in the discussion on the kinetics of channel gating in flux or electrophysiological experiments and their implications in the interpretation of the ligand-binding mechanisms. T h e reason for not developing the above topic in greater detail at this stage lies in the preeminence awarded by the reviewer to the use of kinetic treatment only in a functionally significant context. This should be our frame of reference when attempting to interpret kinetic data of any kind, but most particularly when the experimental conditions depart from the native ones to any substantial degree, as is still the case with most of the studies on AChRligand interactions it? nitro. Furthermore, the physiological frame of reference itself is undergoing radical revision at present, as will hopefully become apparent in the present article. Reducing dogmas to provisional status is part of the healthy exercise involved in the establishment of improved formulations.

THEACETYLCHOLINE RECEPTOR: AN ARCHETYPE? T h e AChR protein is by far the best characterized neurotransmitter receptor. This relatively advanced state of knowledge owes much to the unique characteristics of the nicotinic cholinergic system; Nature’s (fortuitous?) choice of acetylcholine as an ubiquitous transmitter substance is matched by the extreme abundance of AChR in certain biological sources. In many cases this abundance is coupled with a particular amenability of the source, especially in the peripheral nervous system, to experimental work. At first glance such characteristics would appear to make the AChR a logical source of introduction in the study of neurotransmitter receptors, but a deeper insight into the AChR system shows it to be inappropriate for this purpose. T h e peculiar organization of the AChR, for instance, constitutes the exception rather than the rule in current views on surface receptors and membrane proteins in general.

DEVELOPMENTS I N STRUCTURE AND FUNCTION

26 1

Furthermore, many properties of the nicotinic AChR have little if any applicability to its most immediate relative, the muscarinic AChR. Nevertheless, in a different (and more practical) sense the nicotinic AChR system does provide a fruitful source of generalizations which can be extended not only to other neurotransmitter receptors, but to many if not most integral membrane proteins. The legacy of research on the nicotinic AChR to other related fields probably lies not in the specific factual information gained but in the experience accumulated by the many successes and pitfalls in this area. The actual findings on the structure and function of the AChR may prove to be to a great extent irrelevant to the understanding of say, the insulin receptor, but the carry-over from the former field is certainly imposing new postures and accelerating the advance of knowledge on the less accessible, less abundant and more complicated systems. One useful example is the impetus provided by recent advances on the ion gating properties of the AChR-controlled channel, the theoretical and practical implications of which are not necessarily restricted to other neurotransmitter systems but are felt more amply in chemically and electrically excitable membranes. Another example can be seen in the way concepts initiated by studies of AChRs in the rather infrequent myasthenia gravis and related syndromes are spreading into the wider field of autoimmune diseases. The AChR still is, and probably will remain, a useful archetype in many branches of science.

11. The AChR Molecule

A. SIZE,MOLECULAR WEIGHT,A N D HYDRODYNAMIC PROPERTIES Crude or purified, membrane-bound or detergent-solubilized AChR can be reduced to a minimal molecular form upon cleavage of disulfide bonds by reducing agents. This is the monomer of about 250,000 MW (see below). The monomer is not the only molecular species present in membrane-bound or detergent-solubilized AChR preparations. I n the absence of reducing agents, higher oligomeric species can be directly visualized in the native membrane (Heuser and Salpeter, 1979; Barrantes, 1982a,b; Zingsheim et al., 1982b) or observed in sedimentation experiments (Edelstein et al., 1975; Gibson et al., 1976; Chang and Bock, 1977; Vandlen and Raftery, 1979). The AChR monomer has a sedimentation coefficient in the region of 9 S (8.6 S, Reynolds and Karlin, 1978; 9.25 S, Ruechel et al., 1981) and a Stokes radius of about 7 nm (Meunieret

262

F. J . BARRANTES

al., 1971, 1972a,b; Raftery et al., 1971, 1972). T h e discrepancy between the theoretically expected mass for a molecule with such a Stokes radius (about 500,000 MW, globular shape) and the experimentally found value can be accounted for in terms of the high detergent-binding capacity, which may vitiate many hydrodynamic measurements, and more importantly, by the asymmetric shape of the AChR monomer (see below). Molecular weight determinations of the 9-S monomer yield values of 320,000-350,000 by gel filtration methods (Meunier et al., 1972a); 230,000 (Hucho and Changeux, 1973) to 400,000 (Chang et al., 1977) by cross-linking in SDS; and 330,000 (Edelstein et al., 1975), 190,000 (Gibson et al., 1976), and 250,000 (Reynolds and Karlin, 1978) by sedimentation velocity and equilibrium centrifugation measurements. Osmometry produced figures of 270,000 MW (Martinez-Carrion et a/., 1975), though the proportion of monomer was not reported. Calculations based on electron microscopy and X-ray diffraction studies yielded estimates of 250,000 to 370,000 (Ross ct al., 1977; Klymkowsky and Stroud, 1979) for the membrane-bound AChR. From their low-angle, neutron-scattering studies of Triton-solubilized AChR. Wise et al. (1979) deduced a molecular weight of 240,000 40,000. Lo et nl. (1982) have recently used an improved version of the radiation inactivation method to study the AChR in its native membranebound state. An homogeneous population of molecules, defined as the a-neurotoxin binding target, was found in Torpudo, cat denervated muscle, and chick embryonic muscle. T h e apparent molecular weight determined by this method is 300,000. This is within the range established for the 9-S, monomeric AChR by analytical centrifugation (Reynolds and Karlin, 1978; Ruche1 el al., 1981). Table I lists molecular weight estimates of the AChR from several species. T h e ~wiuirieof the AChR monomer has been calculated with various techniques. Electron microscopy and X-ray diffraction yield values of 450 nm3 (Rosset al., 1977) and 380 ? 51-347 t 47 nm3 (Klymkowsky and Stroud, 1979). T h e volume of the hydrated portion of the membrane-bound molecule protruding into the extracellular space is 80-120 nm3 (Zingsheim et al., 1982b), which corresponds to a nonnegligible mass (- 100,000 MW). Neutron diffraction data provide a value of 305 nm3 for detergent-solubilized monomer (Wise et al., 1979). T h e radiation-inactivation method has recently yielded an estimate of 350 nm3 for the membrane-bound monomer (Lo et al., 1982). A translational diffusion constant,D,,,,, , of 2.95 X lo-' cm2 sec-' was found for the detergent-solubilized monomer using laser light scattering techniques (Doster et al., 1980). Mixtures of monomers and dimers in detergent solution could not be resolved into two distinct diffusion species and yielded average values of 2.7 X lo-' cmz sec-I (Doster et al., +_

263

DEVELOPMENTS IN STRUCTURE AND FUNCTION

TABLE I MOLECULAR WEIGHTDETERMINATIONS OF THE AChR MOLECULE

Source

Method

T.marmorata

Sedimentation Gel filtration Sedimentation Laser scattering (Doppler)/ Sedimentation Gel electrophoresis Radiation inactivation

electric organ

T. californica

E. electrim electric organ

Membrane osmometry Sedimentation Sedimenta tion Sedimentation

Molecular weight ( X 10-8)

Speciesa

Reference

500-2000 500- 1000 330-1300

Di- and oligomers Di- and oligomers Mono, oligomers

Mile& et al. (1971) Molinoff et al. ( 1972) Edelstein et al. (1975)

298 330-350 300

Monomer Monomer Monomer

Doster et al. (1980) Riichel et al. (1981) Lo et al. ( 1 982)

270 f 30

Monomer

330,660 190-330 250,500

Mono-, dimer Monomer Monomer, dimer

Martinez-Carrion et al. (1975) Edelstein et al. (1975) Gibson et al. (1976) Reynolds and Karlin ( 1976) Ross et al. (1977)

370

Electron microscopy Electron microscopy, X-ray diffraction

250-310

Monomer

Neutron scattering Neutron scattering Amino acid sequence

240 2 40 500 255

Monomer Dimer Monomer

Sedimentation Cross-linking

320,470 2302 15

Mono, dimer (?) Monomer

Cross-linking Cross-linking Light scattering/ Sedimentation

Monomer

260 250 287

f

8

Monomer Monomer

Meunier et al. (1972a) Hucho and Changeux (1973) Biesecker (1973) Lindstrom et al. (1976)

Monomer

Riichel et al. ( 1981)

?

Chang et al. (1977) Chang et al. (1977) Dolly and Barnard (1975) Bradley et at. (1976) Colquhoun and Rang (1976) Dolly and Barnard (1977) Lo et al. (1982) Lo et al. (1982)

N. brasiliensis electric organ

Cross-linking Sedimentation

400 530

Mammalian muscle

Gel filtration

430

?

Gel filtration Gel filtration

390 300

Monomer

Gel-filtration

370

?

Cross-linking Radiation-inactivation

*

270 20 300

Klymkowsky and Stroud (1979) Wise et al. (1979) Wise et al. ( 1981a) Strader et al. (1980a)

Dimer

?

Monomer Monomer

The species attributed to the apparent molecular weights are only tentative.

264

F. J. BARRANTES

1980). The similarity of the values for pure monomer and monomer + dimer mixtures in solution is also found in membrane-bound AChR using the fluorescence recovery after photobleaching technique (FRAP). Criado et al. (1982) reported lateral diffusion coefficients in the order of 2x cm2 sec-' for both pure AChR monomer and dimer in reconstituted phosphatidylcholine and phosphatidylcholine + cholesterol membranes. Already in early gel filtration experiments (Meunier et al., 1971, 1972a,b) the anomalous behavior of the AChR protein suggested departures from simple globular geometries. Subsequent hydrodynamic studies substantiated this view and provided estimates about the shape of the monomer in detergent micelles. Combining sedimentation equilibrium and velocity measurements, Reynolds and Karlin (1978) came to the conclusion that the molecule behaved as a rigid ellipsoid of revolution with axial ratios greater than 10. Neutron-scattering experiments from the same laboratory (Wise et a/., 1979) enabled the contribution of the detergent to be determined by contrast matching (i.e., blanking out the contribution of the bound detergent) and yielded a radius of gyration of 4.6 +- 0.1 for the monomer in Triton X-100. This is compatible with an oblate ellipsoid with an axial ratio of 1 :4, but other shapes could not be ruled out. Ideally, one would like to determine the shape of the AChR in its natural membrane environment, not in a detergent micelle. Progress is being made in this direction, and will be discussed below.

B. ACHR SUBUNITS A N D THEIR STOICHIOMETRT The AChR from Torpedo species is at present the most extensively characterized case. Acetylcholine receptors from other sources such as Electrophorus and skeletal muscle were for many years believed to differ from that from Torpedinidae in gross qualities such as subunit composition (see, e.g., review in Heidemann and Changeux, 1978). More recent studies have dissipated this notion, showing that the differences were due to proteolysis (Lindstrom et aE., 1980b). T h e following description of some of the molecular properties of the receptor is exemplified with data from Torpedo AChR, except where explicitly stated to the contrary. T h e AChR monomer is composed of two quasi-identical subunits (a,M , = 38,000), and three additional subunits (,B, M, = 50,000; y , M , = 57,000; and 6 , M , = 64,000) (Weill et al., 1974; Raftery et al., 1975; Chang and Bock, 1977; Hucho et al., 1976). Homologies are apparent among all subunits (see below). General acceptance of this subunit structure has involved a lengthy debate. Cross-linking studies with suberidimate led

DEVELOPMENTS IN STRUCTURE AND FUNCTION

265

Hucho and Changeux (1973) to postulate a pentameric structure, although the subunits were not identified as given above. Raftery et al. (1976) suggested an octameric structure of the type a&yS. Changeux and co-workers reported until recently (e.g., Cartaud et al., 1981) AChR preparations containing mainly one subunit (achain). The experimental observations (Sobel et al., 1977) gave rise to a hexamer model consisting exclusively of a chains (Heidmann and Changeux, 1978). It was argued that the other subunits represented impurities which copurified with the a subunit. Studies on subunit composition yielding only one polypeptide chain can be accounted for on several grounds. I n very early reports, the choice of analytical procedures was not always adequate (Klett et al., 1973), but the failure to observe the now accepted four-band pattern in SDS-polyacrylamide gel electrophoresis is to be attributed mainly to the action of endogenous proteases, which degrade the AChR subunits with differing voracity. In recent times the observation of a single AChR chain (the a subunit) in Torpedo preparations (Sobel et al., 1977, 1978) is almost certainly explained by the higher susceptibility of the other subunits to degradation by the endogenous Ca+-activated proteases (Vandlen et al., 1979). This is also manifested in the extensive proteolysis of such subunits by exogenous proteases. Proteolytic cleavage of purified (Bartfield and Fuchs, 1979) or membrane-bound (Klymkowsky et al., 1980; Strader and Raftery, 1980; Conti-Tronconi et al., 1982) AChR by trypsin or papain (Lindstrom et al., 1980a,b) is well documented. Lindstrom and co-workers (1979a,b, 1980a,b) have made a substantial contribution to clarification of this point. I n addition, the lack of observation of the y subunit and the associated variability in the electrophoretic pattern of the AChR under denaturing conditions are also partly due to the different thermal sensitivities of the individual chains, a factor which has only recently been noticed (Barrantes et al., 1980; Sobel et al., 1980; Strader et al., 1980a). The first report on the correct subunit composition (Weill et al., 1974) also included a description of the apparent molecular weights of each subunit as given above (Table 11).The exact molecular weights are lower (F.J. Barrantes, unpublished observations), partly because of the anomalous migrational properties inherent in the presence of carbohydrate moieties on all subunits (see below) and partly because of the molecular weight standards used to date. Wide variations in relative mobilities of the subunits are also apparent as a function of alkylation (Froehner and Rafto, 1979; Lindstrom et al., 1980b; Barrantes, 1982a) and gel crosslinking (Froehner and Rafto, 1979; Strader et al., 1980b). Apparent variations attributable to analytical procedures can amount to 15%. In addition, minor variations in apparent molecular weights of the AChR

266

F. J. BARRANTES

SPECIFIC ACTWIT\

Source Elecfrophumy electricus

AND

Specific activity (nmol neurotoxinhg protein) 4.5 5.4 8 11 5-6.5

Trrrppdo caifornirn

TABLE 11 POLYPEPTIDE COMPOSITION OF PURIFIED AChR‘

Apparent subunit molecular weightsb ( 10-3)

References

44 (50,95-100) 45, 54 (90) (32) 40, 47, 53 (1 10)

Biesecker (1973) Hucho and Changeux (1973) Karlin and Cowburn (1973); Karlin et al. (1976) Klett et a/. (1973) Chang ( 1974); Penn et al. (1976) Meunier et a/. (1974); Sobel et al. (1977) Lindstrom and Patrick (1974) Patrick et al. (1975) Lindstrom et al. (1980b)

4w (37) 42, 49

6.7

43, 48, 53 (90)

7.5 3.6

41, 42, 60 48, 53, 60 (110) 41, 50, 55, 62

6.3 8 10 12 8

(26), 35, 42 39, 48, 58, 64 40. 49, 60, 67 40, 48, 62, 66 40, 48, 59, 67 43, 52, 58, 63

10.6 8-9

44, 53, 60, 65 38, 50, 57, 64 38, 47, 57, 68 40, 50. 60, 65

9 1.7 5 8 7-9

42 45, 50 37, 49, 74 (93, 148)

Schmidt and Raftery (1973) Weill et a / . (1974) Raftery et al. (1974) Eldefrawi et a,/. (1975b) Hucho rf al. (1976) Chang and Bock (1977) Froehner et al. (1977a); Froehner and Rafto (1979) Nathanson and Hall (1979) Lindstrom et al. (1978, 1980b) Bartfeld and Fuchs (1979) VanOlen et 01. (1 979)

-

40 41, 51, 59, 64

Potter (1973) Heilbronn and Mattson (1974) Gordon ~t al. (1974) Eldefrawi et al. (1975a) Sobel et a/. (1977, 1978) Claudio and Raftery (1977)

10- 12

40, 50, 61, 81

Rubsamen et

34, 36, 39, 44 (70) 40, 48, 60, 64

Ong and Brady (1974) Claudio and Raftery (1977)

2.7

28, 38, 45

Schmidt and Raftery (1972)

9

43, 52, 59, 64 44. 48, 58, 65

Claudio and Raftery (1977) Chang et al. ( I 977)

2.2

33. 43 (38, 51)

Ishikawa ~t a/. (1980)

12.5 -

-

nE.

(1978)

267

DEVELOPMENTS IN STRUCTURE AND FUNCTION

TABLE I1 (continued)

Source

Specific activity (nmol neurotoxidmg protein)

Rat skeletal muscle, normal, junctional region extrajunctional region

0.2 -

Rat skeletal muscle, denervated, extrajunctional

9

Cat skeletal muscle, denervated

6

Human skeletal muscle (necropsy) BC3H-1 mouse clonal cell line (neoplasmderived)

45,49,5 1,56,62 (110)

0.5

10 Foetal calf myoblasts (primary culture)

Apparent subunit molecular weightsb ( 10-8)

3.7

0.5-3.5 2.6

References Brockes and Hall (1975a) Nathanson and Hall (1979) Brockes and Hall (1975a)

45,49,51,56,62(110)

Froehner et al. (1977b)

41

Dolly and Barnard (1975, 1977) Shorr et al. (1978)

41 42,49, 55, 58

Merlie et al. (1978) Gotti et ul. (1982)

42, 66 (58, 85)

Stephenson et al. (1981)

44, 53, 65 (72) 42, 46, 48, 60

Boulter and Patrick (1977) Merlie and Sebbane (1981)

~~

Only a comparative value should be attributed to the apparent M , values, given wide variations in the procedures employed. Values in parentheses refer to minor polypeptide components. Nonstandard electrophoretic conditions were used. (I

subunits among different biological species and in the course of development may also occur (see, e.g., Lindstrom et al., 1979b; Merlie et al., 1982), but extensive comparative studies are still lacking. The subunit composition of purified AChR preparations is listed in Table 11. 1 . Endogenous Prosthetic Groups in the AChR Protein

The AChR molecule is a glycoprotein. This has been shown for AChR purified from Electrophorus (Meunier et al., 1974), denervated rat diaphragm (Brockes and Hall, 1975a,b,c), cat skeletal muscle (Dolly and Barnard, 1977), fetal calf muscle (Merlie et al., 1978), Torpedo electric

268

F. J. BARRANTES

tissue (Mattson and Heilbronn, 1975), and the BC3H-1 notifusing cell line (Schubert et al., 1974; Boulter and Patrick, 1977). All individual AChR subunits possess glyco moieties (Raftery et al., 1976; Vandlen et nl., 1979; Lindstrom et nl., 1979a). Mannose, galactose, N-acetyl wgalactosamine, and glucose have been identified. T h e 6 subunit reacts with phytohemagglutinin A, but the a subunit does not (Wennogle and Changeux, 1980). No relationship has as yet emerged between any known structural or functional property of the adult, mature AChR and its carbohydrate moiety (Wonnacott et al.; 1980; Criado and Barrantes, 1982), and the antigenicity of the AChR is not affected by elimination of such a moiety (Wonnacott e t a / . , 1980). T h e native a subunit of the AChR in the clonal muscle cell line BC3H-1 has a single N-asparagine-linked oligosaccharide of the “high mannose” or “simple type” (Merlie et al., 1982) which is susceptible to endo-A‘-acetylglucosaminidase type-H cleavage. T h e mature Torpedo subunits are not noticeably degraded by endoglycosidases of the D type (Criado and Barrantes, 1982). When synthesized in vitro in the presence of tunicamycin to inhibit glycosylation, the production of the a subunit is only slightly diminished, but the assembly into an a-bungarotoxin binding species is notably reduced, possibly by inhibiting transport of the subunits to the Golgi apparatus (Merlie et a/.,1982). Purified AChR appears to contain about 10 phosphate residues per monomer (Reynolds and Karlin, 1978), and Vandlen et al. (1979) have detected approximately seven residues of O-phosphoserine per AChR molecule present in all of the subnits. However, the a subunit is not phosphorylated in zfitro, whereas the y and 6 subunits are (Gordon et nl., 1977, 1980; Teichberg et al., 1977). T h e state of phosphorylation appears to affect physicochemical properties of the AChR such as charge (see below). Solubility in detergents (Saitoh and Changeux, 1980) and susceptibility to heat inactivation (Saitoh ef al., 1979) are also affected by the state of phosphorylation. T h e AChR from immature, neonatal Torpedo can be phosphorylated i?? vitro at the y and 6 subunits by endogenous protein kinases (Saitoh and Changeux, 1981). T h e phosphorylation state of the AChR changes during ontogenetic development, adult AChR being more phosphorylated than its immature counterpart (Saitoh and Changeux, 1981). Other forms of O-substituted serine (about 23) and threonine (about 20) residues also occur, most likely as glycosylated residues. Acetylcholine receptor protein solubilized in 1 % cholate and purified in 0.2% cholate (i.e., below the critical micellar concentration of the detergent) contains variable amounts of 12-24 carbon-saturated and

DEVELOPMENTS IN STRUCTURE AND FUNCTION

269

-unsaturated fatty acids at concentrations of 7-75 nmol lipid/mg protein (Lindstrom et al., 1979a). The presence of residual endogenous lipids in purified AChR was reported to influence the binding properties of the receptor (Chang and Bock, 1979). T h e importance of lipids in the preservation of the ion translocation and affinity transition properties of the AChR will be dealt with further on.

2. Subunit Stoichiometry T h e currently accepted molar ratio of the AChR subunits appears to be a&&, that is, 2 : 1 : 1 : 1 (Reynolds and Karlin, 1978; Damle and Karlin, 1978; Lindstrom et al., 1979a; Raftery et al., 1980a; Strader et al., 1980a; Gonzalez-Ros et al., 1981). This stoichiometry applies equally to membrane-bound or solubilized, purified AChR. The five subunits are tightly associated in the monomer; rather drastic treatments like 4 M urea (Lindstrom et al., 1976) or extensive proteolytic degradation with papain (Lindstrom et al., 1980c) do not dissociate them. That all subunits appear to transverse the membrane can be deduced from various structural, biochemical, immunocytological, and physicochemical studies, as will be illustrated when discussing the threedimensional structure of the AChR.

C. PRIMARY STRUCTURE OF THE ACHR PROTEIN The sequence of the first 54 amino acid residues from the amino terminus of the four subunits has been determined (Raftery et al., 1980a; Strader et al., 1980a). This was preceded by preliminary reports on the sequence of the (Y subunit (Hunkapiller et al., 1979; Devilliers-Thiery et al., 1979). Raftery et al. (1980a) further observed amino acid sequence homologies between the amino termini of the subunits and suggested their early common evolutionary origin from a same ancestral gene, followed by divergence through gene duplication. Earlier peptide mapping of the whole subunits suggested differences in sequence and lack of homology between the AChR chains (Froehner and Rafto, 1979; Nathanson and Hall, 1979; Lindstrom et al., 1979a). Though initial immunological studies did not appear to indicate homologies either (Claudio and Raftery, 1977; Lindstrom et al., 1978, 1979b), more recent studies have revealed them clearly (Tzartos and Lindstrom, 1980; Tzartos et aE., 1981). I n the case of the (Y subunit, at least the 25 amino-terminal amino acid residues appear to be the same in both chains (Devilliers-Thiery et al., 1979; Hunkapiller et al., 1979).

270

F. J. BARRANTES

D. SECONDARY STRUCTURE OF THE ACHR PROTEIN Application of the treatments of Chou and Fasman (1974, 1978) and Lim (1974) to the limited sequences available has resulted in the calculation of the secondary structure predicted for the first 54 amino acid residues in the amino-terminal region (Fig. 1). 1.17 C&F

i =..Q

L IM

:

: m i1.13. . d 1.16

-

.

m

i

: j

1.21

1.20;

A m / (1.13j 1,17;

j

?SEQUENCE

i

:

;

1.12

j

il.05

i

;

j

1

1.16 A

i

11.89)

i

j

:

1.12:

;

1.i

1.22

1.11

I

oM=N i

1.11

:

;!zUw!Q

i ' .,,.,p21 llkQW!WmWUW 1.11 1.21'

1.19

;

i 1.19

~1.15 i1.26

j

i.11

m i

MODEL

1.19

S V M E O T L L S V L F . - E T Y N P K V R P A O T V G D K V T V R V G L T L T N L L I LNE K I EEMRTNV

i LIM

1.14

i 1.23 1:131.57 1.04 .82 1.19 1.30.99>30 36 .97j .88 q2/ ' U ! I W f i ~ Q ^ L Q W f i 2 n / v "

-,

DSEOUENCE

C&F

;

1.24

S E H E T R L V A N L L . . E N Y N K V I R P V E H H T H F V D I T V G L R L I O L I S ~ K I E V NI ~V E T N V

! 1.13 MODEL

i (1.111

.4

MODEL A

LIM

1.18 i

1.17

"V!Zl!lUQW

Yvvvy : Y m 1 1.17 i i 1.14 1:471,57 1.19 1.04 1.15 1.301.13)301'.14 .90! 1.14 ' QQQQWUW A~LQWWWVV~ WAAQW (1.11)

QSEQUENCE

C&F

1.14

j-..8A:

'

;

:

1.21 1,221.57: 1.08 ~

~

g

g

1.17

'

1.17

j

m l

~

1.44i o ,1.05 p

~

"17 1.621 @&UQWUQ.f;,& ~llaVVW,Q~Q%QWQW E N E E ~ R L I E ~ L L- G- D Y D K ~I ~ P A K T L D HI D~ V T ~ K L T L T N LI S < N E M E { A L T T N V i

1.13 i

: 1.14

$30

:

1'.12

i

i

1.11;

j

.95i

C&F

i

1.08

;

LlM

j

1.16

/

1'49 1.14 1'1:1

1.57 ISLKETDETLTSNV

5

io

ib

20

25

j,

35

40

45

50

55

FIG. 1. Application of the algorithms of Chou and Fasman (1978) (C & F) and of Lim (1974) to the amino terminal sequences of the a,0, y and 6 AChR subunits determined by Raftery et 01. (1980a). The values above the corresponding a-helical segments

(m),

&pleated sheets (m), and p turns ( A ) indicate the conformational parameter, P , predicted for the given conformation. &-Helices are predicted when the P values exceed 1.03for a sequence of six or more amino acids; values above 1.05 are needed for 0 sheets, and 0 turns require P > 1.0. (From Guy, 1981.)

DEVELOPMENTS I N STRUCTURE AND FUNCTION

27 1

E. CHARGE OF THE ACHR

Acetylcholine receptor from electric tissue shows a broad band at about pH 5 upon isoelectric focusing (Raftery et al., 1971; Biesecker, 1973; Eledefrawi and Eldefrawi, 1973a).Small differences between junctional and extrajunctional AChR from skeletal muscle have been reported (Brockes and Hall, 1975b; Nathanson and Hall, 1979), and two isoelectric species exist in Electrophurus (Teichberg and Changeux, 1976). The differences in charge appear to be accompanied by differences in immunological properties of the AChR (Almon and Appel, 1975; Weinberg and Hall, 1979). Charge differences between neonatal and adult Torpedo AChR have been reported (Saitoh and Changeux, 1981) and interpreted as resulting from the phosphorylation in the course of ontogenetic development.

F. IMMUNOCHEMICAL STRUCTURE OF THE ACHR Antibodies of different degrees of specificity (antitotal AChR, antisubunit, etc.) have been successfully used to map regions of the receptor molecule inaccessible to other techniques such as affinity labeling or chemical modification. The structural information obtained by immunochemical methods complements and extends that obtained with the latter technique. More than 50% of the anti-AChR in an antiserum to native AChR is directed against a special region of the macromolecule, termed the main immunogenic region (MIR; Tzartos and Lindstrom, 1980). This region consists of at least two adjacent determinants located outside the ACh recognition site on the a subunits, exposed to the extracellular milieu. It appears to be phylogenetically preserved, since antibodies against it cross-react between species and passively transfer the acute form of experimental autoimmune myasthenia gravis (EAMG). In spite of their proximity to the recognition site, these antibodies impair neither the binding of the a toxin (Gullick et al., 1981) nor the agonist-induced 22Na+ flux mediated by the AChR (Lindstrom et al., 1981b). Species crossreactivity is not an exclusive property of this type of antibody, though. Antibodies to other parts of the AChR can also cross-react between species and elicit the EAMG (Lindstrom et al., 1978, 1979b, 1981a). Monoclonal antibodies against the recognition site of the AChR are available (Gomez et al., 1979). The “library” of antibodies developed by Lindstrom and co-workers (1981b) also includes some types which inhibit noncompetitively the 2zNa+flux. Further types are able to discern be-

272

F. J. BARRANTES

tween AChR monomer and dimer, respectively (Conti-Tronconi et al., 1981). Other monoclonal antibodies can cross-link the a subunits of two adjacent monomers (leading to AChR aggregation), or the two a-subunits within the monomer (Conti-Tronconi et al., 1981; Tzartos et ai., 1981). More than 70 different types of monoclonal antibodies have been made available to date, 40 against Electrophorus AChR (Tzartos et al., 1981). These have enabled the mapping of nine immunogenic regions on the AChR, listed in Table 111. As can be seen in this table, the MIR is present in various species as a strongly conserved feature. Other regions are mapped, 3.5 to 7.0 nm distant from the MIR, some formed by continuous and others by discontinuous amino acid sequences. T h e significance of the immunological approach in the study of the AChR in terms of its medical implications has been the subject of extensive experimental work and reviewing. T h e reader is referred to recent reviews (Lindstrom and Dau, 1980; Lindstrom, 1979; Fuchs, 1979; Vincent, 1980; Lindstrom et al., 1981a,b) for a comprehensive treatment of the subject. Other derivations of the immunochemical approach for the understanding of the topography of the AChR are discussed below in relation to the vectorial orientation of the receptor in the membrane.

111. Biosynthcsisof the AChR

T h e synthesis of AChR and the regulation of its levels are complex and multivariate phenomena. Though it is currently accepted that the steady-state levels of AChR in skeletal muscle are regulated by the electrical activity of the muscle cell, little is known about the processes mediating between the sensing of the electrical activity and protein synthesis. When muscles are denervated, receptor synthesis increases (Axelson and Thesleff, 1959); upon reinnervation, AChR levels return to normal (Miledi, 1960). In the absence of neural inputs, AChR synthesis can be mimicked in muscle cell culture where it appears to be regulated by muscle activity. As contractility appears, AChR levels decline (Prives et nE., 1976). T h e rate of synthesis can be modulated by Na+-channel ligands. Veratridine, which activates the channel, diminishes AChR synthesis (Betz and Changeux, 1979), whereas blockage of Na+ channels by tetrodotoxin leads to increased synthesis (Catterall and Niremberg, 1973). T h e relationship between Na+-channel activation or blockage and AChR synthesis is not known, but it has been hypothesized that common pathways could link the signaling of synthesis at the sarcoplasmic reticulum and the excitation-contraction coupling (Pezzementi and

DEVELOPMENTS IN STRUCTURE AND FUNCTION

273

Schmidt, 1981). These authors produced the depletion of Ca2+from the sarcoplasmic reticulum with low concentrations of the drug ryanodine, a plant alkaloid, and a concomitant decrease in AChR synthesis. High (millimolar) concentrations of the drug enhanced synthesis by 200%. The intracellular Ca2+content appears, therefore, to play a messenger role in AChR synthesis and excitation-contraction coupling. The rise in cytoplasmic Ca2+not only triggers contraction, but shuts off AChR synthesis. Calcium ion release would therefore be the last step common to both phenomena (Pezzementi and Schmidt, 1981). But extracellular Ca2+ concentrations also influence AChR synthesis (McManaman et al., 1981) in a manner that appears to contradict the hypothesis of Betz and Changeux (1979). T h e few examples presented above serve to illustrate the difficulties encountered in the study of AChR biosynthesis in the intact tissue or cell. Fortunately, molecular biology techniques are currently available for the study of this process in simpler model systems, and rapid advances are evident in the development and characterization of cell-free, heterologous systems for the translation of the messenger RNAs coding for the individual AChR subunits. Progress is also being made in the cloning of their genes. The study of the AChR biosynthesis in heteroiogous cellular systems has also been tackled recently in Xenopus oocytes. This section deals with these recent developments. One of the first successful attempts to obtain cell-free synthesis of AChR constituents was reported by Mendez et al. (1980). These authors isolated ribonucleic acid from the electric organ of Torpedo calfornica and purified a fraction enriched in polyadenylated species. This poly(A+)RNA was assayed for its ability to direct protein synthesis in reticulocyte lysates. It was calculated that about 2.4% of the poly(A+)-RNAcoded for AChR peptides. T h e translational products obtained in the cell-free system, identified by immunoprecipitation with antisera against denatured AChR, had apparent molecular weights of 60,000, 51,000, 49,000, 41,000, and 35,000, that is, lower than those of the mature Torpedo AChR subunits (see Table 11). The results suggest that several different messenger RNAs code for the AChR subunits, and that the precursor forms already possess structural features recognized by the antisera prepared against native receptor. However, the capacity to recognize a-bungarotoxin is not present in the precursors synthesized in the reticulocyte lysate. Merlie et al. (1981) were able to fractionate cytoplasmic and membrane-bound polyribosomes from the clonal mouse cell line BC3H-1 and to study the in vitro synthesis of the a chain of the AChR. Both types of polyribosomes were found active in protein synthesis upon

Properties of corresponding mAbs Species specificity (number of mAbs cross-reacting ~0.1% with acetylcholine receptor from each species)

TorM

2

Antigenic regions identified on intact eel acetylcholine receptor A. MIR

B. Site(s) for mAbs 55, 56, 58, and 61

Number of mAbs

23

4

p& Subunit specificity

electric organ

Fetal calf muscle

Rat muscle

2 1: a n t i a 2: unknown

18

7

5

2

1

1

Anti-cx

Properties of antigenic regions Located on the extracellular surface of the a subunit. Distinct from the toxin-binding site. Probably formed by a continuous sequence of amino acids. Homologous to MIR on torpedo and mammalian acetylcholine receptor; therefore, a strongly conserved structural feature. Probably formed by a prominent bend in the a polypeptide. Site($ on the a subunit remote from the MIR and the toxin-binding site. Formed by continuous sequences of amino acids.

N

C. Site for mAb 26

1

Unknown

0

0

0

D. Region for mAbs 25, 29, 33, 34, and 52

5

Unknown

0

0

0

E. Region for mAbs 57 and 59

2

Anti+

1

0

0

F. Region for mAbs 30, 32,43, and 48

4

1: anti+ 3: unknown

1 1

0 0

0 0

G. Site for mAb 53

1

Anti6

0

0

0

H. Site for mAb 60

1

Anti4

0

0

0

I. Site for mAb 54

1

Unknown

1

0

0

-J

cn

Located between the MIR and region D, less than 35 A from each. Probably formed by noncontinuous sequences of amino acids. Species specific. Located less than 7 0 8, from the center of the MIR. Probably formed by noncontinuous amino acids. Species specific. Located on the p subunit remote from region F. Formed by a continuous sequence of amino acids. Located at least in part on the p subunit. Probably formed by noncontinuous amino acids. Located on the 6 subunit, but near MIR on (Y subunit. Species specific. Located on the 6 subunit remote from region H. Formed by a continuous sequence of amino acids. Species specific. Unknown location. Probably formed by noncontinuous amino acids.

a From Tzartos et al. (1981). Nine immunogenic regions (A-I), including the main immunogenic region (MIR) are detected in AChR from Electrophorus electricus with monoclonal antibodies (mAbs) and their properties listed.

276

F. J. BARRANTES

incubation with nuclease-treated rabbit reticulocyte lysates, but they produce two different sets of proteins. T h e membrane-bound type is the one responsible for the synthesis of two protein species of M , = 39,000 and 42,000, respectively, both immunologically homologous to the mature a chain of the AChR. T h e authors interpret the two protein species as corresponding to the nonglycosylated and glycosylated forms of the polypeptide. In support of this assertion, they digested the mature a subunit with endoglycosidase H and observed the reduction of the apparent molecular weight from 42,000 to 39,000. T h e site of glycosylation appears to be close to the NH2 terminus of the polypeptide. More recently, Merlie et al. (1982) have studied in greater detail the role of glycosylation in AChR biosynthesis. They showed that the a subunit synthesized in BC3 H- 1 cells has a single N-asparagine-linked oligosaccharide of the “simple” or “high mannose” type. T h e “complex” type of oligosaccharide appears to be absent from the a subunit (Merlie et al., 198 1, 1982). T h e process of assembly of the a subunits in a form active for recognizing the a toxins is completed within 30 min and is only 30% efficient. When glycosylation is inhibited by tunicamycin, the synthesis of the a subunits is diminished only slightly, but the latter assembly process is drastically reduced. In a related study, Merlie and Sebbane (1981) employed the same clonal cell line and two antisera, one against a-bungarotoxin, which precipitates receptors having bound toxin, and one against denatured AChR. T h e sera were used to analyze the biosynthetic products of the BCH3-I cell at early stages. Newly synthesized AChR requires approximately 15-30 min before it acquires a-bungarotoxin binding capacity, indicating the existence of a pool of inactive AChR precursors. This pool precedes the one defined by Devreotes et al. (1977) as a cytoplasmic precursor pool of the AChR which is transferred to the plasmalemma after about 2 hr. Fambrough and Devreotes (1978)had also shown that a large proportion of the newly synthesized AChR is located in areas of the cytoplasm morphologically corresponding to the Golgi apparatus. Merlie and Sebbane suggest that their early precursor represents the pre-Golgi form of the AChR. Where and when a-toxin ability is acquired is still not known. I n the interval between completion of polypeptide chain synthesis and acquisition of toxin binding capacity, the AChR subunits might be transferred from the rough to the smooth endoplasmic reticulum, undergoing modification of their carbohydrate side chain, and assembling into the mature oligomeric structure. T h e fact that the latter process and acquisition of toxin recognition ability are simultaneously hampered in other systems (see below) suggests a correlation between these t w o phenomena. Furthermore, taking into consideration the multiplicity of

DEVELOPMENTS IN STRUCTURE A N D FUNCTION

277

contacts that the a-neurotoxin-AChR complexes appear to exhibit (Chicheportiche et al., 1975),and the results of cross-linking studies with photoaffinity toxin labels (Witzemann and Raftery, 1978; Raftery et al., 1979, 1980b; Oswald and Changeux, 1982), it is likely that ( a ) not only the a chain, which carries the agonist recognition site, is involved in antagonist binding, but other subunits also contribute to such binding, and (b) the ontogenetic development of toxin binding ability is intimately coupled with the correct stereochemicalassembly of the subunits into the final quaternary structure of the AChR. Anderson and Blobel (1981) have used immunoprecipitation with subunit-specific antisera to identify the four Tmpedo califmnica AChR polypeptide chains synthesized in a cell-free system. Each subunit appears to be synthesized independently and the primary translation products obtained in dog pancreas microsomes have apparent M, of 38,000, 50,000, 49,000, and 60,000, respectively. Integration of each chain into the microsomal membrane is coupled with a cotranslational process analogous to that found in viral membrane glycoproteins (Katz et al., 1977). Upon cotranslational incubation with the rough microsomal membranes, the glycosylated forms of each subunit were obtained, with their molecular weights approaching those of their mature counterparts. Extensive trypsinization of these glycosylated forms reduced them to membrane-protected fragments (still in the form of glycopeptides) with M , of approximately 35,000, 37,000, 45,000, and 44,000. The trypsin sensitivity appears to increase in proportion to the molecular weights of the subunits. The authors concluded that each chain spans the membrane at least once. The in nitro-synthesized products do not appear to possess a-bungarotoxin binding capacity nor do they seem to have the ability to assemble into the mature oligomeric structure. The study of Anderson and Blobel (1981), in being able to identify the precursors with subunit-specific antibodies, clarifies some of the questions left open in the work of Mendez et al. (1980). However, the difference in molecular weights between precursor and mature subunits is not totally accounted for by the common proposal of Mendez et al. (1980) and Anderson and Blobel (1981) involving glycosylation. If one takes into account that the carbohydrate moiety of the subunits amounts to 4-796 of their weight (Lindstrom et ad., 1979a) or half these values (Vandlen et al., 1979),then it is necessary to invoke additional posttranslational modifications of the polypeptides. Estimates of the molecular weight of the AChR (Table I) and of its subunits (Table 11) differ so widely from laboratory to laboratory, however, that although the apparent differences are no doubt real, and due possibly to posttranslational processing, not too much significance should be given to the absolute

278

F. J . BARRANTES

differences until accurate molecular weight estimates of the mature subunits in their aglycosylated form are available. In addition, the presence of a fifth peptide recognizable by the immunosera against AChR in the reticulocyte synthetic products (Mendez et al., 1980) merits further investigation. I n terms of the biosynthetic mechanism involved, Anderson and Blobel (198 1) suggest that the AChR polypeptides are also synthesized on membrane-bound polysomes in vzvo. T h e fact that the glycosylated microsomal products still do not match the molecular weights of their mature counterparts leads to the obvious postulation of a subsequent posttranslational modification of the carbohydrate moiety in the Golgi apparatus. T h e lack of observation of oligomeric assembly is puzzling, and Anderson and Blobel (198 1) offer an interesting explanation which invokes lateral diffusion of the subunits on the plane of the membrane. Given the small diameter of the microsomal membranes used, the inability of the system to form the oligomeric AChR could result from the physical difficulty involved in accommodating more than one polysome per subunit on the same microsomal membrane. Sumikawa et al. (198 1) have used a reticulocyte lysate system like the one successfully employed by Mendez et al. (1980), but they only obtained three (M, = 40,000, 51,000, and 59,000) of the polypeptide chains found by the latter authors. They also confirmed Mendez et al.’s (1980) results concerning the inability of the in vitro products to bind a-neurotoxin. Better results were obtained by injecting Torpedo marmorata mRNAs into Xenopus oocytes. This system has been extensively characterized (see, e.g., Gurdon, 1974) in its capacity to undertake posttranslational processes like glycosylation, sequestration, peptide cleavage, and secretion of biosynthetic products. Sumikawa et al. (1981) produced evidence indicating the synthesis of the four polypeptide subunits of the AChR in the Xenopus oocyte, and their assembly into the 9-S monomeric AChR species. T h e multisubunit product is also glycosylated and is mostly associated with a particulate fraction, suggesting that it has reached the membrane-bound (though not necessarily the plasmalemma) form in the oocyte. Finally, the ability of the AChR monomer to bind a-bungarotoxin could be demonstrated by these authors in the heterologous system. This points to the importance of posttranslational modifications in the assembly of functionally “mature” AChR molecules since they successfully occur in a cellular but not in a cell-free translation system. Studies along the lines of the ones reviewed in this section will certainly contribute to our understanding of the biosynthetic pathways followed by the early precursors of the AChR chains, which otherwise share

DEVELOPMENTS IN STRUCTURE AND FUNCTION

279

enough homologies to fully justify Raftery et aZ.’s (1980a) interesting hypothesis of a common ancestral gene. It is already clear that each of the AChR subunits is synthesized from a separate mRNA. Other important avenues opened by the detailed characterization of these precursors and their intracytoplasmicfate relate to the time and place of acquisition of structural and functional properties by the AChR. Perhaps a certain degree of chronological overlapping will be found to occur between the assembly of the subunit precursors into the mature AChR oligomer and the “learning” of the ligand recognition ability. Merlie et al. (1982) quote unpublished observations from their laboratory indicating that the acquisition of a-toxin binding activity and physical association of the subunits occur simultaneously. This does not rule out the possibility that certain “protopatic” functions are already being displayed by the immature precursors. The subtle discriminating abilities of mature AChRsthe recognition property-contrast with the cation-selective but otherwise unspecific channel gating property of the AChR. It is conceivable that the latter function is established earlier in the ontogenetic process. Other possible offsprings of present efforts to isolate AChR mRNAs are connected with their use as templates to obtain DNA sequences and cloning. This will not only result in an acceleration of sequence analysis of the subunits, of which the first 54 amino-terminal side chains have been determined by conventional Edman degradation techniques (Raftery et al., 1980a),but may open new insights into the primary structure of the ACh recognition site, the peptide segments making up the channel, etc. Substantial progress is being made in the cloning of genes for Torpedo AChR. An ordered library of circular DNA sequences made from Torpedo mRNA has been compiled recently at the Salk Institute. A clone having the sequence appropriate for the y subunit has been identified (Ballivet et al., 1982) and the sequencing of other subunits is in progress (M. Ballivet, J. Patrick, and S. Heinemann, personal communication).

IV. Three-Dimensional Topography of the AChR in the Membrane

The various biochemical and physicochemical properties which qualify the nicotinic AChR as an intrinsic membrane glycoprotein (see Barrantes, 1979) materialize in the structural characteristics of the macromolecule inserted in its native membrane environment. The morphology of the AChR has been the subject of several studies involving electron microscope and small-angle X-ray diffraction (Cartaud et al.,

280

F. J . BARRANTES

1973, 1978, 1980, 1981; Nickel and Potter, 1973; Ross et al., 1977; Klymkowsky and Stroud, 1979; Heuser and Salpeter, 1979;Zingsheim et d., 1980, 1982a,b; SobelPtol., 1980; Klymkowskyet al., 1980; Kistler and Stroud, 1981; Kistler ef nl., 1982). T h e biochemical (Huang, 1979; Tarrab-Hazdai et nl., 1980; Strader and Raftery, 1980; Wennogle and Changeux, 1980; Wennogle et al., 1981; Conti-Tronconi et nl., 1982) and immunohistochemical (Karlin et af., 1978, 1979; Tarrab-Hazdai et al., 1978; Strader et nl., 1979) investigations addressed to the problem of the vectorial sidedness of the AChR protein or to its putative transmembrane nature find their morphological counterpart in such studies, which have been covered in more than one recent review (Heidmann and Changeux, 1978; Barrantes, 1979; Karlin, 1980; Changeux, 1981). For this reason, only a few of the more essential and more recent features will be outlined here. The morphology of the AChR emerging from studies of the macromolecule in solution is that of an asymmetric body, with a radius of gyration of about 4.6 nm for the monomer of M,. = 250,000, as determined by neutron scattering (Wise et nl., 1979), or 4.3 nm computed from the values of sedimentation experiments (Reynolds and Karlin, 1978). T h e interpretation of the latter parameter in light of the mass weight and hydrodynamic properties of the macromolecule does not yield a common picture as regards shape and dimensions. This ambiguity is not unique to the AChR, but also applies to other integral membrane proteins extracted from their native microenvironment and maintained in micellar form with the aid of detergents. In the case of the AChR an additional factor has to be taken into account, namely, the peculiar supramolecular organization of this protein in its natural membrane, densely packed in assemblies having more than 10,000 units/pm2. Obviously, the protein-protein and lipid-protein interactions present under such conditions, and the vectorial topography of the protein are no longer preserved in the micellar state, and this defficiency is likely to be accompanied by concomitant departures from what could be qualified as the native structure of the membrane-bound protein. Electron microscopy provides us with two-dimensional projections of the receptor in AChR-rich membranes. In studying these projections one benefits from the fact that the macromolecule is oriented with respect to the observer because of the mere circumstance of being inserted in a two-dimensional domain (the membrane) which in turn lies flat on the support. The collapsed membranes, flattened on the grid, are covered with the “naturally” oriented projections of the AChR along its main molecular axis. Under appropriate conditions, the edges of the flattened sacs offer the complementary side-view of the protein. A disk-

DEVELOPMENTS I N STRUCTURE AND FUNCTION

28 1

shaped particle of 7-8 nm in diameter with a central pit is the typical image rendered by the end-on views. At the maximal resolution attainable to date (1.8 nm), the most relevant piece of information so far obtained on the AChR monomer as a whole is that it appears to lack symmetry on the plane of the membrane (Zingsheim et al., 1980). This is compatible with the subunit stoichiometry (see above) and has obvious implications as regards the search for structural counterparts of some functional properties, for example, cooperative behavior. The concept of pseudosymmetry as applied to the AChR molecule (e.g., Gotti et al., 1982) appears in this context unfounded; there is neither structural, biochemical, nor functional data to support this view. Three distinct regions have been identified around the central pit of the AChR in end-on views by image averaging of minimal-electron-dose micrographs (Zingsheim et al., 1980),a result recently confirmed by others (Brisson, 1980; Kisder and Stroud, 1981). In the axis normal to the membrane surface, the AChR extends about 5.5 nm toward the extracellular space and about 1.5 nm toward the cytoplasmic compartment (Klymkowsky and Stroud, 1979; Zingsheim et al., 1982b). This corresponds to a non-negligible mass (- 100,000 MW) of the AChR protruding toward the extracellular space, a factor to be taken into account when considering the putative location of voltage-dependent binding sites (e.g., for voltage-sensitive, charged local anesthetics). After removal of the nonreceptor v peptide ( M , = 43,000) from the cytoplasmic face of the membrane, projections of the AChR toward the inner side of the membrane are observed, and a different profile of the AChR particle becomes visible (Barrantes, 1982a). Thus, the AChR monomer displays asymmetry along axes both parallel and perpendicular to the membrane plane. A. THEACHR OLICOMERIC STRUCTURE T h e AChR exists in the membrane as closely, but not maximally, packed units. Ordered arrays occur exceptionally (Ross et al., 1977; Cartaud et d., 1978), and the type of symmetry displayed in the ordered lattices varies (see review in Barrantes, 1979). The predominant disordered particles are expected to occur at much lower densities than the corresponding ordered lattices. I n spite of this, packing density of the AChR in the membrane is high with more than 10,000 particles or 20,000 toxin sites per p m 2 (see Changeux, 1981), a fact which has precluded the identification of the supramolecular organization of the AChR in native membranes until recently. For this reason, determina-

282

F. J. BARRANTES

tion of structural properties of the various AChR molecular forms has relied on measurements carried out on detergent-solubilized protein. Most of the information on this subject stems from sucrose gradient and analytical centrifugation experiments. I n Torpedinidae, a mixture of 9 S (monomer)and 13 S (dimer) appears to be the predominant molecular species detected in such experiments [see Vandlen and Raftery (1979) for a comparative study], but it has long since been known that multiples of such forms occur (Edelstein et al., 1975; Gibson et al., 1976; Chang and Bock, 1977; Vandlen and Raftery, 1979). However, a series of studies (Chang and Bock, 1977; Hamilton et ul., 1977, 1979) has singled out the monomer-dimer components and the inference has been made that the 13-S species constitutes the “native” one in the membrane. This led, in turn, to several structural studies aimed at the identification of the dimer and to reconstitution experiments directed toward the description of the gating properties of monomers and dimers. Attempts at the latter, analyzed in the discussion on AChR channel properties, have so far failed to come u p with any functional characteristic of the dimer supporting its putative role as the native species. T h e emphasis on the monomer-dimer components has likewise been reflected in recent structural studies. Wise et nl. (1981), for instance, reported the observation of Triton X-100 solubilized AChR monomers and dimers by electron microscopy and neutron scattering. Cartaud et al. ( 1980) reincorporated these two molecular forms into artificial lipid vesicles at high lipid : protein ratios, and described the morphological appearance of single and double particles compatible with the masses of AChR monomer and dimer, but in their statistical treatment of the data they neglected higher oligomeric forms of the AChR present in their preparations. Kistler and Stroud (1981) attempted the use of image reconstruction techniques on tubular structures formed from AChR membranes by a procedure developed by Brisson (1980). Although dimeric AChR preparations were examined, the technique failed to reveal structures other than the monomer (Kistler and Stroud, 1981). The contention that monomers and dimers suffice to describe the oligomeric equilibria of the AChR is not necessarily supported by two structural studies (Heuser and Salpeter, 1979; Barrantes, 1982a), which reveal other forms of organization of the AChR. T h e two works have in common the avoidance of the use of detergents for the observation of AChR in the membrane. Heuser and Salpeter (1979) used a rapidfreezing technique to observe AChR particles in the postsynaptic membrane of fresh electrolytes after freeze-fracture, deep-etching, and rotary replication. They described particle doublets, tetramers, and higher aggregational forms, often arranged in rows. Barrantes (1982a) turned

DEVELOPMENTS I N STRUCTURE AND FUNCTION

283

to a combination of ( a ) depletion of nonreceptor peptides by alkaline treatment (Neubig et al., 1979) and ( b ) a mild physical shearing of the resulting fragile membranes, leading to direct observation in the membrpze remnants of various oligomeric forms of the AChR. T h e 9-S monomeric receptor predominated in reduced membranes, the 13-S species in normal or alkylated membranes, and higher oligomeric forms, both in normal or oxidized membranes, could be correlated with the AChR species revealed by gradient centrifugation. T h e correlation of these AChR oligomeric forms with the presence or absence of the v proteins is considered in the succeeding section. An alternative approach for the study of the oligomeric organization of the AChR is based on low-dose electron microscopy and subsequent single-particle image averaging of the AChR. We have initially used the combination of these two techniques to produce the average image of the internally asymmetric membrane-bound AChR monomer at 1.8 nm resolution (Zingsheim et al., 1980). The work has now been extended to the investigation of structural differences between reduced and normal membranes, making apparent the disulfide-bonded dimers at a resolution of about 2 nm (Zingsheim et al., 1982b), and to the localization of the a subunits (Zingsheim et al., 1982a) as reviewed below.

B. TOPOGRAPHY OF ACHR SUBUNITS IN

THE

MEMBRANE

More recently, efforts have been directed toward the localization of individual AChR subunits. Hartig and Raftery (1977), for instance, employed lactoperoxidase-catalyzed iodination of sealed AChR vesicles to show the exposure of the a, /3, and y AChR subunits to the external synaptic surface (Fig. 2). Raftery et al. (1980b) have recently reviewed a series of studies from their laboratory in which photochemical probes were used to map regions of contact between the cholinergic recognition site on the a subunit and adjacent subunits. Thus, Witzemann and Raftery (1978) synthesized photolabile a-bungarotoxin derivatives which reacted covalently upon light irradiation with the a and 6 subunits from the outside of the vesicles. Long cross-linkers reacted only with the 6 subunit when monomeric AChR preparations were tested and with the 6-6 disulfide-bonded subunit on dimeric AChR preparations. Short cross-linking toxins labeled the a subunit in addition to the 6 or 6-6 in monomeric and dimeric AChR preparations, respectively. Oswald and Changeux (1982) used ultraviolet irradiation to cross-link native a-bungarotoxin with the AChR. T h e major cross-linking occurred with the a and 6 chains, and to a lesser extent with the y subunit. Tarrab-

284

F. J . BARRANTES

NORMAL (sea1ed.v protein -rich. right-side out) extrac&lu&r

mody-

( lactoperoxidase iodination(Q1

intramembranous mae_ty_

\

proteolysis I b c ) a.py6 antisubunit antibodies(di1 I glycosylated subunitslef J cross-linking with a toxinsld

1

ALKALINE TREATED

ALKALINE + SAPONIN TREATED

(sea1ed.v protein -depleted)

(holey. v protein -depleted)

SAPONIN TREATED (holey. v protein present)

_cy&plasmic- f a c i n g m t y ; antisubunit specific antibodtesiJ1.a,p.y.6 anti -v protein monospecific antibodyikj. v lactoperoxidase iodination"] v tannic acid staining(m1. v

FIG. 2. Vectorial sidedness of AChR and nonreceptor u-proteins in the membrane. Exposure of all subunits to the rxtunral face of the membrane was demonstrated by the techniques indicated by (a) Hartig and Raftery (1977); (b) Strader and Raftery (1980), Wennogle and Changeux (1980), Wennogle ef nf. (1981); (c) Conti-Tronconi ut nl. (1982); (d) Klymkowsky and Stroud (1979). Gullickrtal. (1981); (e) Rafteryulnl. (1976), Vandlen ut al. (1979); (f) Lindstrom rf (11. (19794; (g)Witzemann and Raftery (1978), Karlin ut a!. (1978), Hucho (1979), Oswald and Changeux (1982); and (j) Froehner (1981). Lipidrmhcddrd domains have been detected in the Q subunits with iodonapthylazide (h) Tarrab-Hazdai P / 01. (1980). Tarrab-Hazdai and Goldfarb (1982); and in the /3 and y subunits with pyrene sulfonylazide (i) Sator rf d . ( 1979). Cslo,+lcsmzl--fnring moieties have been detected in all AChR subunits by trypsinolysis of permeabilized membranes (Wennogle and Changeux, 1980; Strader and Raftery, 1980) and by immunochemical techFroehner (198 I)]. Similarly, the cytoplasmic exposure niques as indicated in the figure of the v-proteins has been reported (k) Barrantes (1982a); (I) St. John ut 01. (1982). Identification of cytoplasmic densities (m) which disappear after alkaline treatment was carried out by Sealock (1982) and Cartaud r / 01. (1981).

[c)

Hazdai et al. (1 980) employed one of the lipophilic compounds propounded by Gitler (see Rercovici and Gitler, 1978), iodonaphtyl- 1 azide, which partitions favorably in the hydrocarbon interior of the membrane bilayer. T h e compound labeled only the a subunit from inside the membrane (Fig. 2). On the other hand, a nitrene-

DEVELOPMENTS IN STRUCTURE AND FUNCTION

285

photogenerated pyrene sulfphonylazide appears to react with two different subunits, the @ and the y , from the lipid bilayer (Sator et al., 1979) (Fig. 2). Tarrab-Hazdai and Goldfarb (1982) suggested that the different labeling patterns could arise from the 1000-fold higher bulk concentrations of the probe used in the latter case; the possibility of differing reactivities of the two compounds was also considered (the labeling studies are diagrammatically presented in Fig. 2). Controlled proteolysis has also been used as a means of determining the exposure and sidedness of the AChR subunits. Strader and Raftery ( 1980) concluded from their trypsinolysis study that all subunits were accessible both from the inside and the outside of sealed, right-side out vesicles. From the rates of tryptic degradation they also suggested that the larger subunits were more exposed and the a-subunits less exposed to the cytoplasmic side of the membrane. Wennogle and Changeux ( 1 980) reached similar conclusions, although their preparations showed considerable proteolysis prior to the enzymatic tests. In a subsequent publication, in which precautions against endogenous protease activities were taken, Wennogle et al. (1981) identified a 16,000-MW fragment of the 6 chain. T h e fragment is still capable of cross-linking nonreduced AChR monomers into the 13-S dimers and presumably corresponds to the C-terminal of the 6 chain exposed to the cytoplasmic side of the membrane. Figure 2 summarizes these findings diagrammatically. Froehner (198 1) has recently used anti-AChR antisera in combination with chemical modification studies to disclose exposed and hidden antigenic determinants of the AChR in its membrane-bound form. T h e a , p, and 6 chains appear to possess exposed antigenic determinants. Alkaline extraction, which removes essentially the nonreceptor v proteins (see Section V) uncovers hidden determinants on a , y , and 6, presumably on the C-terminal regions of the polypeptides, exposed to the cytoplasmic compartment. Saponin treatment, which permeabilizes the vesicles without extracting membrane proteins, makes latent sites accessible on the Q and 6 subunits, also from the cytoplasmic phase. When both treatments are combined, determinants on the p, y , and 6 chains are exposed, which are not apparent when only one of these treatments is used. Figure 2 schematically shows the progressive uncovering of latent antigenic sites (Froehner, 1981) on the AChR molecule by alkaline and saponin treatments and the topographical relationship of these sites with the v proteins (Barrantes, 1982a,b). Another series of studies provides indirect evidence on the topography of the AChR subunits, as a consequence of the glycoprotein nature of the AChR individual chains. Since the sugar moiety in membrane glycoproteins is almost invariably facing the extracellular milieu, and

286

F. J. BARRANTES

recent studies have demonstrated carbohydrates on all AChR subunits (Raftery ~t al., 1976; Vandlen et al., 1979; Lindstrom et al., 1979a), the presence of these endogenous markers indicates that all subunits are at least in part exposed at the intersynaptic cleft. Further studies are likely to be developed on the basis of labeling procedures, since the carbohydrates constitute a suitable point on which appropriate probes could be tagged. Probably the 6 subunit, because of the additional “marker” provided by the 6-6 disulfide bond (see Fig. 3) will be the subunit most easily accessible to such an approach. 1. Putative Topog-raphs of t h S Subunits

As mentioned above, the combination of low-dose scanning transmission electron microscopy and single-particle image averaging techniques can be used to study the structural organization of the AChR. The comparison of native membranes with those subjected to cleavage of the disulfide bonds by reducing agents has recently yielded the average image of the dimeric form of the membrane-bound AChR (Fig. 4). It consists of the pairwise association of two nearest neighbor AChR particles. Although the resolution affordable in this case is not as high as that

V&!+Y

Carbohydrate

FIG. 3. Schematic representation of two S subunits (.LIP= 64,000) joined by a 8-6 disulfide bond occurring at the C-terminal portion of the chains, which also carries the phosphorylation site (Pi) and the cleavage points (50,000, 49,000, and 47,000) leading to stable tryptic fragments. The C-terminus faces the ctyoplasmic side of the membrane (Wennogle ct al., 1981). A bilayer-embedded domain has not been directly identified in this subunit as is the case with the a,@, and y subunits (see Fig. 2). The carbohydrate moiety is hypothetically drawn attached to the Asn, and Am, residues by analogy with other membrane glycoproteins.

DEVELOPMENTS IN STRUCTURE AND FUNCTION

monomer

287

dimer

FIG. 4.Contour plots obtained by single-particle image averaging of negatively stained AChR membranes reduced with dithiothreitol (a) and native (b). Low-dose electron micrographs taken with a scanning transmission microscope. The plots are presented at a resolution of 6.4 nm. The shaded areas correspond to the most significant density increments in the stain-excluding regions. The lack of orientationally related neighboring particles in the reduced, monomeric preparation (a) contrasts with the presence of an oriented, adjacent particle in the native membrane (b). The presence of a disulfidebonded dimer does not exclude the presence of higher oligomers. (From Zingheim et at., iga2b.)

used to study the internal structure of the monomer (Zingsheim et al., 1980), the differences between the monomer and the dimer are apparent, and the presumptive location of the disulfide-bonded 6-6 contact can be inferred. Under the reduction conditions employed, resulting in total cleavage of the disulfide bonds, the pairwise association of the adjacent particles disappears (Fig. 4). The study of Zingsheim et al. (1982b) reinforces once more the advantages of working with the membrane-bound receptor as compared to the detergent-solubilized AChR dimers used in a recent study by Holtzman et al. (1982). These authors prepared biotinyl derivatives of a toxin displaying about one-fourth of the affinityof the native (Y toxin for its binding site. Biotin exhibits extremely high affinities for avidin, a cylindrical tetrameric molecule of about 5.5 x 4.1 nm. Holtzman et al. (1982) studied the ternary complexes of detergent-solubilized AChR, biotinyl-toxin, and avidin by low-dose electron microscopy. The tendency of the multivalent avidin molecule to form aggregates with itself and to cross-link receptors was diminished by succinylation; the occurrence of “monsters” of AChR clustered by multivalent avidin oligomers was thus minimized but not completely avoided. The angles subtended by the avidin molecule peripherally tagging the biotinylated toxin on the AChR molecule and the axis connecting the two monomers in the dimer

288

F. J. BARRANTES

displayed four peaks; when only the dimers carrying two avidins were considered, t w o peaks were observed at 60-80” and 140-160”, respectively. From this information the relative positions of the a (see below) and 6 chains in the dimer were inferred. Unfortunately, the interpretation of these angles suffers from serious handicaps due to the uncertainty concerning the orientation of the AChR particles on the support (given the randomization of views of detergent-solubilized AChR) and the choice of paired AChR particles of unusually large intercenter spacings, defined as end-on views based on observation of their central pit. Tilting experiments with solubilized AChR indicate that visualization of the typical “rosette” appearance of the particle and its pit does not suffice to define an axial end-on view of the AChR, since the central pit remains unchanged for tilting angles as large as 30” (Zingsheim et al., 1982b). 2. Putative Location of the P Subunit Knowledge of the location of the 6-6 bond linking two monomers in a dimer provides a useful landmark for establishing the relative topography of other AChR subunits. Wise et al. (1981) used the oxidizing reagent diamide to produce oligomeric AChR forms and isolated the trimeric species in Triton X-100 by gradient centrifugation. The higher oligomerii forms constitute 70% of the diamide-treated preparation, 50% of which is in turn trimer. Assuming that the distances between joined monomers in a trimer represent 6-6 and P-P cross-links, measurement of these two lengths and the one distended by the free ends of the trimer, together with the interior angle formed by the “arms” of the triad, led these authors to conclude that the P-P bond is separated from the 6-6 bond by an angle of 50-80”. From this they inferred the positions of the /3 and 6 subunits, respectively. T h e conclusions are not selfevident, however, because the location of the P-P angle need not be identical with the angle defined by the centers of gravity of these two subunits. Thus, Wise et al. (1981) had to resort to external information on the location of other subunits. They used data on the tentative location of the a chains (see below), which in turn is also subject to criticism. The location of the P subunit therefore remains an open question. 3. Putative Locataori of the a Subunits

The fact that the a subunits carry the recognition site for agonists and antagonists can be exploited in an attempt to localize these chains by structural methods. Using the reaction of the external tag avidin on biotinylated, a-toxin-labeled AChR, which was described in the preceding sections, Holtzman rt al. (1982) calculated the angles separating the a subunits. They deduced that one toxin site is located in the range of 45

DEVELOPMENTS IN STRUCTURE AND FUNCTION

289

to 85" and another at about 100" further from the 6-6 cross-link axis. By further assuming that ( a )the two a chains in a monomer are arranged so that they are superimposable by rotation around the axis of the pit; (b) they subtend an angle at the pit proportional to their mass; and (c) they are separated by an angle of about loo", the conclusion was reached that the a chains cannot be contiguous, and their tentative position was calculated. A more recent approach has also made use of the a toxin to localize the a subunits. I n this case, the direct localization of the toxin site on the AChR molecule was attempted. Although a toxins are relatively small polypeptides in comparison to the AChR macromolecule ( M , = 8000 and 250,000, respectively), the quasi-irreversible attachment of a single native toxin molecule to the recognition site on each a chain produces a significant increase in the mass contributing to the average image of the AChR, roughly one-quarter of the mass per a subunit. This is so because only one-half of the total AChR molecule is made apparent as stainexcluding volume (Zingsheim et al., 1982b). If one then obtains a difference map between the toxin-tagged and untreated membranes, two statistically significant peaks of stain-excluding density appear (Fig. 5 ) . One of them ( a l ) is adjacent to the region where the 6 subunit has tentatively been localized (see above). The second peak (az)is located diametrically across, about 5 nm apart from the 6 subunit and also from a1 (Fig. 5 ) . The basic outcome of the structural study is that the two a chains exhibit contacts with a different set of subunits. The different local environments of the two subunits carrying the recognition sites provide a structural basis for rationalizing the nonequivalent ability of each a chain to react with a h i t y reagents (Delegeane and McNamee, 1980) or with other cholinergic ligands (Sine and Taylor, 1981). T h e structural asymmetry of the local environment of the a subunits is a further manifestation of the overall asymmetric nature ofthe AChR monomer.

4. Tentative Assignment of All AChR Subunits in an Azimuthal Arrangement In recent electron microscope studies (Zingsheim et al., 1982a,b) it has been possible to ascertain the sidedness of the membrane-bound AChR particles with respect to the substrate as well as the position of three (a1, a 2 ,6) of the five subunits deduced with the aid of endogenous or directly contacting (i-e., without intervening cross-linking arms) tags. Direct information on the position of the /3 and y subunits is not as yet available, but structural information on the topography of the Y proteins with respect to the AChR particle (see Section V) and biochemical data can be obtained to tentatively locate t h e y subunit. T h e relative position of the /3 subunit, still inaccessible to direct labeling, can only be inferred

290

F. J. BARRANTES

- - - - .-. . ,/-. -. - ,’, - - -. /,--. ,5’ \ I /

/

/

/ /

/

I \

I

-

-

_

/

\

.

\

\

\

I

\

\

/

I

\

\

/ ,

/

’ I

\

FIG. 5. Direct localization of the a-toxin binding sites and, by inference, putative location of the a subunits. The average images of membrane-bound AChR tagged with native a-bungarotoxin (a) and the corresponding control without toxin (b) are digitally subtracted to yield a difference map (a-b). The standard error of the difference is plotted (ae), the lowest error occurs in the regions of maximal protein density. Two regions show increased stain-excluding mass (bright areas) upon toxin binding. These two regions have been designated a 1 and a q ,as shown diagrammatically below, in an axial projection perpendicular to the membrane plane. Resolution in (a) and (b) is 1.8 nm. (From Zingsheim ct ai., 1982a.)

by default. T h e complete azimuthal arrangement of all AChR subunits in a projection perpendicular to the membrane plane can be presented (Fig. 6b). V. In Search of the Functional Role of the Nonreceptor Y Proteins

Rapidly growing interest is being shown in a class of nonreceptor proteins originally described as a polypeptide having an apparent molecular weight of about 43,000 (the 43-K protein; Sobel et al., 1977;

DEVELOPMENTS IN STRUCTURE AND FUNCTION

29 1

Neubig et al., 1979),the v peptide of Hamilton et al. (1979), the v doublet of Barrantes et al. (1980), or the 43,000-dalton proteins of Gysin et al. (1981). In this section we shall not only see that this class of polypeptides is a predominant component of highly purified AChR membranes, but also that it is ranked as the second most abundant protein localized in the cholinergic synapse and intimately related to the AChR protein itself. A. EARLY ATTEMPTSAT ASSIGNINGA ROLETO THE v PROTEINS Sobel et al. (1977) described a membrane preparation consisting exclusively of one receptor subunit, the M , 40,000 (the a subunit), and a nonreceptor 43,000 polypeptide (the 43-K protein). They subsequently reported (Sobel et al., 1978) that the fluorescence of quinacrine associated with this polypeptide was sensitive to histrionicotoxin, a ligand characteristic of the nicotinic ionic channel (Eldefrawi et al., 1977; Masukawa and Albuquerque, 1978). Their conclusion was that the 43-K protein possessed the recognition site for local anesthetics and, by inference, constituted the ionic channel (Sobel et al., 1978). Blanchard and Raftery (1979) prepared an azido derivative of procaine amide, which upon photolysis in the presence of AChR membranes weakly labeled the AChR subunits, the 5 band (presumably the (Na’, K+)-ATPase heavy chain of Lindstrom et al., 1979a),and more conspicuously, a broad band in the region of the v proteins (Blanchard and Raftery, 1979). Incorporation of the label into the a chain of the AChR could be prevented by the presence of carbamolycholine (Carb). This was presumably due to the binding of the local anesthetic analog to the AChR recognition site, as is usually the case with this type of ligand at high concentrations. The probe associated with the v proteins was refractive to histrionicotoxin. Neubig et al. (1979) used alkaline extraction of nonreceptor peptides and similar criteria to demonstrate that elimination of the v proteins altered neither the local anesthetic binding nor the ion flux properties of the receptor membranes, a result since confirmed by other laboratories (Elliot et al., 1980; Lindstrom et al., 1980a). It has further been documented that elimination of the v proteins is not manifested in changes of the AChR-mediated ion translocation, as judged by single channel measurements in reconstituted planar bilayers (Boheim et al., 1981). The reduction of the receptor polypeptide composition to only the (Y subunit has also been shown to result from proteolytic degradation of the other peptide chains, the pentameric nature of the receptor protein now being more firmly established (see, e.g., Lindstrom et al., 198Oc; Karlin, 1980).The “functional orphanage” into which the Y proteins fell at this stage was particularly surprising in view of its conspicuous pres-

292

F. J . BARRANTES

ence in partially degraded membranes, of which it forms a major constituent (see above). Although Sobel et al. (1977) had reported that the 43-K protein did not comigrate with actin in gel electrophoresis, Hamilton et al. (1979) stated that the peptide had the same electrophoretic properties as rabbit and T. californica actins, and Karlin et al. ( 1979) reported immunological evidence on actin-v protein cross-reactivity. However, Strader et al. (1980b) clearly demonstrated the separability of the two proteins in gel electrophoresis. Barrantes (1982a) has also shown conditions under which the v proteins cannot be extracted from the AChR membranes, whereas actin can be. Conventional polyacrylamide gel electrophoresis reveals microheterogeneity of the v proteins, which can be resolved into a doublet (Barrantes et nl., 1980; Klymkowsky et al., 1980). Isoelectric focusing also makes the microheterogeneity apparent, with at least five different species of v being observed (Saitoh and Changeux, 1980). Combination of the t w o techniques enabled the identification of three major v protein species: v l ,v2, and v3, with various isoelectric subspecies (Gysin et al., 1981). T h e latter two polypeptides are the most abundant ones and are mainly associated with the soluble fraction of the Torpedo electrocyte. v 1 is the membrane-associated species. All are extracted by the alkali stripping procedure (Neubig et al., 1979), and tryptic fingerprinting suggests that the three are separate gene products (Gysin et al., 1981).

B. INFLUENCE OF

THE

v PROTEINS

Oh'

ACHR FREEDOM OF MOTION

I n an electron spin resonance study Rousselet and Devaux (1977) showed the rotational immobility of the AChR protein in native membranes, and later these authors reported on the appearance of rotational motion upon alkaline extraction of nonreceptor peripheral peptides (Rousselet Pt nl., 1979). I n a morphological study it was shown that extraction of the Y proteins gives rise to alterations in the receptor packing habit (Barrantes et d.,1980). These observations were related to an increased freedom of motion of the receptor molecules, and it was further proposed that the Y proteins play a role in processes like synapse formation during ontogenesis, receptor clustering, and stabilization of the adult synapse. Lo et al. (1980) found enhanced rotational freedom of the receptor in membranes depleted of the v proteins by studying the phosphorescence anisotropy of AChR labeled with an erythrosine derivatives of a-bungarotoxin. They also reported the restoration of AChR

DEVELOPMENTS I N STRUCTURE AND FUNCTION

293

rotational immobility upon reassociation with the alkaline extract, as confirmed by Rousselt et al. (1981). Using the same technique and eosin derivatives of the a toxin, Bartholdi et al. (198 1) were able to observe a jinite rotational correlation time of 12-26 psec associated with the AChR in membranes depleted of the v proteins. The same rotational correlation time was found in dithiothreitol-reduced membranes, though not in normal or N-ethylmaleimide-alkylatedmembranes. The experimentally observed relaxation time agreed with that expected for the 9-S AChR monomer of M , 250,000. In the electron spin resonance study of Rousselet et al. (1979), an upper limit of about 1 msec could be set on the rotational immobilization of the receptor in normal membranes. No information on the distribution of oligomeric AChR forms was given in the latter publication, and one can infer that the preparations consisted of the usual mixture of monomers, dimers, and higher oligomeric species. More recently, Rousselet et al. (1981) have produced a study in greater detail in which the spin probe maleimido-2,2,5,5-tetramethyl-l-pyrrolidinyloxyl was attached to extensively proteolyzed membrane [following the preparative procedure of Sobel et al. (1977)l consisting almost exclusively of the AChR a subunit ( M , = 40,000) and the v proteins. The spin probe labeled predominantly the v proteins under normal conditions and the Q subunit of the AChR in membranes successively alkylated with N-ethylmaleimide, reduced with dithiothreitol, and submitted to alkaline extraction. This bears relationship to the work of Lindstrom et al. (198Oc),in which proteolytically degraded AChR maintained almost unaltered low-resolution morphology, flux-mediating ability, etc. (see Section VI). T h e electron spin resonance work of Rousselet et al. (1981) complements such information, indicating that the AChR reduced to only the (Y subunit is relatively immobile in the membrane, displaying rotational correlation times of several hundred microseconds. This is compatible with the spectrum of relaxation times found by Bartholdi et al. (1981) in native membranes (made u p of AChR monomers, dimers, and higher oligomeric forms). However, extraction of the v proteins leads to the observation of rotational correlation times of 100- 120 psec (Rousselet et al., 1981) or 10-26 psec (Bartholdi et al., 1981) and the acquisition of temperature sensitivity. Thus, the latter authors only observed the 10-26-psec rotational relaxation time above 37°C after depletion of the v proteins, or at room temperature after converting all AChR forms into the monomer. Since the 10-26 psec relaxation time, well resolved by the phosphorescence anisotropy experiments, agrees with the expected rotational motion of the AChR monomer (see above), the

294

F. J. BARRANTES

100- 120-psec time observed by Rousselet ef a/. (198 1) can be interpreted as resulting from the predominance of species heavier than the monomer.' Another aspect of one of Rousselet et al. 's ( 198 1) studies concerns the effect of exogenous lipid addition on the morphology and dynamics of AChR aggregates in the membrane. In otherwise untreated membranes, addition of phospholipids leads to a mosaic appearance of the membrane, which displays AChR particle-free areas and particle clusters. This could suggest that larger AChR aggregates predominate in the presence of the u proteins, in agreement with the conclusions of Barrantes (1982a,b). On the other hand, addition of exogenous phospholipids to u-protein-depleted membranes produces a dispersed distribution of AChR particles in the plane of the membrane, but cholesterol + phospholipid addition maintains the clustered particle appearance of the native membrane. T h e cited authors interpreted the latter results as indicative of the influence of cholesterol on AChR organization. Cherry uf al. (1980), for instance, have shown cholesterolinduced protein segregation in bacteriorhodopsin-phospholipid vesicles. Popot P t ( I / . (1978), Schindler and Quast (1980), and Marsh and Barrantes (1978) have also found preferential association of AChR with cholesterol-rich monolayers, effects of cholesterol on cooperativity, and immobilization of the substantial fraction of cholesterol in AChR membranes, respectively. On the contrary, in applying the polyene antibiotic filipin, which produces membrane lesions in cholesterol-rich areas, Nakajima and Bridgman (1981) found no evidence of high cholesterol content in the postsynaptic region of frog neuromuscular junctions. They also disregard all other evidence on the interactions of cholesterol and AChR, and argue that the AChR-rich membranes used for lipid analysis could have been contaminated. This is unlikely to be correct; it is also questionable whether the filipin technique is sensitive enough to produce lesions visible with the resolution afforded by conventional electron microscopy in AChR-rich areas: T h e interstices between the densely packed AChR particles are only a few angstrom units wide. In a more recent study (Rousselet P/ nl., 1982), the results of Bartholdi PI al. (1981) have been challenged, on the assumption that the 10-25 ysec rotational relaxation times observed by the latter authors result from segmental motions of phosphorescent toxins. This criticism is invalidated by the observation of the same relaxation times in multiple, randomly labeled .4ClrR ( M . Criado, T. Jovin, and F. Barrantes, in preparation), that is, circumventing the use of toxin. Furthermore. the theory used by Rousselet et nl. (1982) to develop their arguments is not applicable to anisotropic systems like the AChR membranes, especially when there is no internal consistence of the parameters derived from the spectra of T* (Thomas uf nl., 1976).

DEVELOPMENTS I N STRUCTURE AND FUNCTION

295

DYNAMICS OF THE ACHR C. TRANSLATIONAL The static picture captured by electron microscope techniques which lel to the postulation of increased freedom of motion of the AChR in these modified membranes (Barrantes P t al., 1980) implicitly pointed to a type of motion different from the rotational diffusion reviewed in the preceeding paragraphs, namely, the translational diffusion of the receptor protein. This has been the subject of various studies in adult and embryonic muscle cells by means of the fluorescence recovery after photobleaching (FRAP) technique. Essentially, these studies have shown that the AChR in the adult synapse and in the patches found in developing muscle is translationally immobile, whereas the diffusely located AChR population has a diffusion coefficientD,of about 5 X lo-" cm2/sec (Axelrod et al., 1976, 1978a,b).Po0 (1982) has recently challenged these observations, reporting that the D,of the mobile AChR population in Xmopus embryonic muscle membranes is 2.6 X 1O-O cm2/sec,i.e., 50-fold higher than the values of Axelrod et al. (1976, 1978a,b). More recently Criado et nl. (1982) have studied the lateral diffusion of the AChR reconstituted in pure lipid bilayers, and found values of D, in the range of cm2/secfor both the AChR monomer and dimer above the lipid phase transition temperature in dimyristoyl-phosphatidylcholine membranes. Additional multiple component recoveries with D,of less than 5 x lo-" cm2/secwere found below the lipid phase transition. The presence of a cholesteryl ester did not markedly affect the diffusional properties of the reconstituted AChR protein, which was incorporated at very low concentrations in the artificial membranes. Under the same experimental conditions, a fluorescent-labeled phosphatidylethanolamine diffused with D tof 8 X cm2/sec(Criado et al., 1982). Thus it is clear that in the low concentration limit neither the 9-S AChR monomer nor the 13-S dimer devoid of peripheral nonreceptor peptides encounter hindrance to lateral diffusion on the part of the lipid bilayer itself. The situation is radically different in native membranes, in which the densely packed AChR assemblies are almost totally anchored and translational and rotational diffusion is inhibited.

PARTICIPATION OF THIOL REACTIONS D. THEPOSSIBLE I N v PROTEIN-ACHR INTERACTIONS Depletion of the v proteins does not hinder the thiol-dependent interconversion of receptor. However, the presence or absence of the v proteins seems to affect the native redox equilibrium: v protein-depleted

296

F. J. BARRANTES

membranes show partial conversion of dimers to monomers (M. Criado and F. Barrantes, unpublished) and a slight shift to higher oligomeric states upon reoxidation as compared to standard membranes. Conversely, membranes in which the dimeric species predominate (the “NEM membranes”) are refractive to depletion of the v protein by alkaline extraction (Barrantes, 1982a,b). On the basis of (a) the richness in thiol groups of the v proteins (Sobel et al., 1978); (b) the observed changes in particle packing habits upon their extraction from normal membranes (Barrantes et al., 1980); (c) the AChR particle tendency to be closely packed in reduced, alkaline-extracted membranes; and ( d ) the inability of alkaline treatment to extract the v proteins from NEMmembranes, Barrantes (1982a) proposed that the “unknown protein” of Chang and Bock (1977) with ‘‘unusually low redox potential and a close proximity to the receptor” could materialize in the v proteins. It was suggested that these proteins play the role of a “redox buffering system,” normally impairing the high-order associations of AChR monomers. Upon depletion of the v proteins (by alkaline extraction or salicilate treatment) these constraints would disappear, and the receptor units would be able to associate into higher oligomeric forms (Barrantes, 198213). Ruchel et nl. (1981), on the other hand, explicitly reject the possible participation of any protein other than the AChR itself in the formation of receptor oligomers. Their conclusion is based on the observation of monomer-dimer interconversions of purified AChR in various detergents. However, in spite of claims to the contrary, their preparations of “purified” protein d o not conform to the criteria currently used to define purified AChR: T h e ATPase heavy chain ( M , about 90,000) is present and partial proteolysis noticeable (Ruche1 et al., 1981). More importantly, the presence of a conspicuous band in the M, 43,000 region is apparent in their preparation. T h e finding of this polypeptide in afiity-chromatography-purified AChR raises the question of the strength of the association between the v proteins [a most likely candidate for the material in Ruchel et al.’s (1981) work] and the AChR protein. Furthermore, since their conclusions are based on the assumption that no peptides other than the AChR protein are involved in receptorreceptor interactions, the presence of the v proteins could be taken as evidence to the contrary of their conclusions. E.

T O P O G R A P H I C A L LOCALIZATION OF THE v PROTEINS A N D RELATIONSHIP TO RECEPTOR STRUCTURE

ITS

T h e insensitivity of the v proteins to proteolytic degradation when trypsinization is applied to intact AChR membranes, a condition which

DEVELOPMENTS IN STRUCTURE AND FUNCTION

297

results in the proteolysis of the a , p, and 6 AChR subunits (Wennogle and Changeux, 1980), was one of the first pieces of evidence suggesting a “cytoplasmic-facing” location of the u proteins. Sonication of the vesicles in the presence of trypsin resulted in degradation of the v proteins (Wennogle and Changeux, 1980). Such evidence has recently been challenged (Conti-Tronconi et al., 1982) in experiments in which trypsin digestion of AChR membranes was performed either from the inside or the outside of vesicles. I n the former case, trypsin was loaded into the sealed microsacs. Conti-Tronconi et al. (1982) found more extensive degradation of the v proteins when the proteolysis was undertaken from the outside. They used, however, AChR membranes which had already been partly depleted of the v proteins by alkali stripping; the results are, therefore, not strictly comparable. Histochemical evidence (Sealock, 1980, 1982; Sealock and Kavookjian, 1980) also suggests a cytoplasmic, peripheral exposure of the v-proteins. Thin sections of AChR membranes negatively contrasted with tannic acid showed projections toward both the extracellular and cytoplasmic compartments, extending about 6.5 nm from the center of the bilayer. A major portion of the cytoplasmic projections disappeared upon removal of the peripheral proteins by alkaline treatment (Sealock, 1982). T h e immunofluorescence experiments of Froehner et al. (1981) showed the synaptic localization of the total alkaline-extractable proteins from Tmpedo AChR membranes (predominantly made up of the v proteins) in Tmpedo electrocytes and mammalian muscle. Barrantes (1982a) treated AChR membranes with saponin, which is known to form complexes with cholesterol (Lucy and Glauert, 1964), using a procedure reported to permeabilize the vesicles to molecules larger than M , 450,000 without altering binding properties or the polypeptide composition of the AChR membranes (St. John et al., 1982). Using monospecific anti-u-protein antibodies, it was shown that the antigenic determinants of the v proteins in the membrane are mainly accessible from the “ctyoplasmic” side of AChR vesicles (Fig. 2). I n agreement with these results, saponin-permeabilized vesicles show lactoperoxidase-mediated iodination of the v proteins from the inside of the membranes (St. John et al., 1982). Barrantes (1982a) also described a class of AChR particles, visible after depletion of the v proteins, having a different (i.e., lower) electron density profile and slightly different dimensions (Neugebauer and Zingsheim, 1982) than those of the normal particles. This was attributed to the “unmasking” of a normally occluded portion of the receptor protein upon removal of the v proteins. Given the asymmetric shape of the AChR and the above information on the cytoplasmic topology of the v proteins, the low-electron-density AChR particles were interpreted as

298

F. J. BARRANTES

corresponding to the end-on, in-plane view of the cytoplasmic-facing part of the AChR molecule. This would represent the mass of the prolate ellipsoid, proposed by Wise et al. (1979), made apparent after depletion of the mass normally obliterating this view. The recent findings of Froehner (1981) offer additional insight into this interpretation. This author exposed buried or latent sites in AChR membranes by alkaline extraction, saponin treatment, or a combination of these two procedures. Saponin treatment, which permeabilizes the otherwise right-side out, sealed vesicles, makes latent sites accessible on the a and 6 subunits of the AChR. Depletion of the v proteins uncovers determinants on the a,y , and 6 subunits. T h e combined treatments disclose new sites on p, y , and 6, which are not revealed by either treatment individually (Figs. 2 and 6). Taking advantage of the richness in sulfhydryl groups of the polypeptides in AChR membranes, a smaller radiolabeled compound such as [,‘\‘-3H]ethylmaleimide can be used to learn about the topographical relationship between the AChR protein and the v proteins. Depletion of the latter makes sulfhydryl groups available on the subunit of the AChR (F. J. Barrantes, unpublished observations). The simplest interpretation of these observations is that the v proteins occlude a limited mass of the AChR from the alkylating reagent, presumably an area of close contact between the two proteins (see below). T h e above information is mainly concerned with the location of the v proteins on an axis perpendicular to the membrane surface and with surface topography relationships between the AChR and the v proteins. Additional information on the vectorial sidedness of the v peptides, that is, on their topography in a plane parallel to the membrane (and relative to the end-on view of the AChR particle), can be produced from the combined application of minimal electron dose scanning transmission electron microscopy (STEM) and single-particle averaging techniques, as applied by Zingsheim ef al. (1980) for the obtainment of the azieuuge two-dimensional projection of the AChR molecule in the membrane. The procedure for locating the v proteins is essentially the same, except that in addition to the individualization of the AChR protein in normal membranes (let us call this set A ) , the procedure is repeated on v-protein-depleted membranes (set B ) and the A -B difference carried out to obtain a diffiwriu map. This map outlines the profile of the mass corresponding to the extracted v proteins (Fig. 6). T h e shaded area in Fig. 6 depicts the nvpr-agp low-resolution topography of the v proteins, making clear that the bulk of the membrane-associated M,. 43,000 proteins is located peripherally with respect to the center of the AChR particle. When this piece of information on the two-dimensional topography is combined with ( a ) that of Froehner (1981) on the exposure of

DEVELOPMENTS IN STRUCTURE AND FUNCTION

a

299

b

FIG. 6. Location of the v proteins and azymuthal distribution of AChR subunits. (a) Difference map of single-particle image averaging between native and alkali-stripped membranes yields the aueruge low resolution topography of the u proteins in AChR membranes. An axial projection perpendicular to the membrane plane is shown. The mass of protein depleted from the membranes (dark-shaded area) is located peripherally with respect to the AChR particle (lighter shaded area). Except for the indicated areas, the environment of the AChR remains featureless after alkaline extraction, and the AChR particle itself is not affected. (From H. P. Zingsheim, D.-C. Neugebauer, and F. J. Barrantes, unpublished observations.) (b) Azymuthal distribution of the AChR subunits and their relationship with the peripheral u proteins. Combined alkaline and saponin treatments make antigenic determinants available in the p, y, and 6 subunits (Froehner, 1981). Removal of the v proteins uncovers reactive sulfhydryl groups mainly in they subunit and to a lesser extent in the 6 and a chains (F. J. Barrantes, unpublished). Since the location of the 6 chain has been established (Zingsheim et ul., 1982a), and its relative position with respect to the two different a chains (a1. a e )determined (see Fig. 4), these results can be combined to infer the relative position of the y subunit with respect to the others and that of the v proteins in close contact with the y, 6, and a chains in an AChR particle. Since removal of the u proteins does not uncover reactive groups on the /3 subunit, its projection is drawn without overlap with the v proteins. The choice of dimers to illustrate this point is arbitrary. One can arrive at these topographical assignations only because the mientation of the AChR particles is known (Zingsheim et al., 1982a). The azymuthal distribution of four out of the five subunits can then be established; that of the p follows by default.

the AChR determinants upon extraction of the v proteins ( b ) that of Barrantes (1982a,b), St. John et al. (1982), and Sealock (1982) showing the cytoplasmic exposure of the polypeptides in the AChR membranes, and (c) the appearance of reactive groups on they subunit upon depletion of the v-proteins (F. J . Barrantes, unpublished), it is possible to outline a tentative profile of the membrane-associated v proteins and their putative location with respect to the AChR protein (Fig. 6).

300

F. J. BARRANTES

In summary, the v proteins 1. appear in all likelihood to be exposed toward the inner, cytoplasmic side of the AChR membrane, eccentrically located with respect to the AChR protein; 2. establish closest contact with the AChR protein presumably in a region covering part of the al,y , and 6 subunits; 3. participate, directly or indirectly, in the thiol-dependent receptor aggregational states; 4. influence the freedom of motion of the AChR protein, both in terms of its rotational and translational mobility; and 5. influence the susceptibility of the AChR to thermal denaturation (Saitoh ~f al., 1979) or enzymatic attack (Klymkowsky et al., 1980; Wennogle and Changeux, 1980).

Additional roles of the v protein(s) cannot be excluded at present, specially in view of their apparent microheterogeneity. One particular line of research on this family of synaptic proteins is likely to receive considerable attention: the study of the relationship between the u proteins and cytoskeletal-basal lamina components. The putative presence of F-actin outlining the synaptic region (Halletal., 1981) or more specifically in the postsynaptic densities, and its adjacence to the intramembranous particles observed in deep-etched electron micrographs of central synapses (Gulley and Reese, 1981) may have its counterpart in the Turprdo electrocyte postsynaptic membrane, as some ultrastructural studies appear to indicate (Rosenbluth, 1974, 1975; Heuser and Salpeter, 1979; Sealock, 1980, 1982; Cartaud et nl., 1981). Axelrod ~t al. (1978a), however, found no influence of cytoskeletal perturbing agents on the lateral diffusion of immobilized receptors. Electrophysiological experiments of Anwyl and Narahashi (1979) gave no correlation between the breakdown of microtubules by cytochalasin B and colchicine treatment and alterations of the ACh potentials. T h e concomitant lack of extraction of cytoskeletal components and of a certain proportion of AChR by a short treatment with 0.5% Triton X-100 led Prives et al. (1982) to conclude that the cytoskeleton is responsible for the anchoring of the clustered AChR. T h e experimental evidence produced is still preliminary and cannot be used to draw definitive cause-effect relationships, especially in view of the fact that detergents (at higher concentrations and acting for longer periods) also fail to extract a sizable proportion of the AChR even in the absence of cytoskeletal components. Some additional cytoskeletal proteins have been considered. For instance, vinculin, a protein first described in smooth muscle, has been found to delineate the same overall region as AChR clusters in myotubes in culture, inter-

DEVELOPMENTS IN STRUCTURE AND FUNCTION

30 1

digitating with AChR-rich areas. Vinculin is considered to further contact with actin-containing microfilaments (Bloch and Geiger, 1980). Analogies between the v proteins and ankryn or a-actinin have been drawn (Cartaud et al., 1981), but the list of cytoplasmic proteins interacting with actin is expanding so rapidly (see reviews in Schliwa, 1981, and Craig and Pollard, 1982) that more factual information is needed before the connections between the v proteins, the postsynaptic densities, the subsynaptic web, the cytoskeletal proteins, and the basal lamina are able to be unraveled. VI. The lon-Translocation Function in Mernbrane-Bound AChR

As with the pharmacological view of the AChR some decades ago, the second if not most important property of the AChR-the translocation of ions across the membrane-can at present only be defined in terms of its function; it has not yet been possible to identify the chemical structure related to that function. In this section we shall consider in vitro kinetic measurements of AChR-controlled ion flux in AChR membrane fragments. T h e cases where these measurements have been undertaken in whole cells (Sine and Taylor, 1979, 1980, 1981) will not be discussed here, since they constitute the exception to the wealth of the information, and have recently been reviewed by Karlin (1980). “Native” AChR-rich membrane vesicles from Electrophmus (Kasai and Changeux, 1971a,b; Hess and Andrews, 1977; Hesset al., 1978; 1979; Cashetal., 1980) or Tmpedo (Hazelbauer and Changeux, 1974; Popotet al., 1976; Sugiyamaet al., 1976; Andreasen and McNamee, 1977; Miller et al., 1978; Neubig et al., 1979; Moore et al., 1979a,b; Bernhardt and Neumann, 1978; Neubig and Cohen, 1980; Hess et al., 1981) display agonist-induced permeability for cations. The pharmacological properties of this phenomenon are qualitatively, though not quantitatively, similar to those of the living cell. Experiments make use either of the uptake or the release of a permeant ion (influx or efflux measurements, respectively). T h e first developed and still most commonly used assay is based on the uptake of a radioactive cation as measured by a filtration technique, introduced by Kasai and Changeux (1971a). Flame emission spectroscopy has also been used to quantitate trace amounts of Li+, Na+, and K+ (Ramseyer et al., 1981; Bernhardt et al., 1981). Another recent approach, developed by Raftery and co-workers, consists of the measurement of thalium influx by following its quenching of a fluorescent probe previously introduced into sealed AChR-rich vesicles (Moore and Raftery, 1979, 1980; Wu et al., 1981).

302

F. J. BARRANTES

A. SOMEPROBLEMS A X D THEIR ALLEVIATION All measurements on native membranes reported to date suffer from the same handicaps, the first and most important of these being inherent in the distribution of receptors in the membranes. Thus in Electrophorus native membrane fragments, the commonly measured AChR-mediated flux represents only a small percentage of the total expected flux and is obscured by nonspecific ionic fluxes unresponsive to agonists. Similarly, in native Tol-pedo vesicles the AChR density is so high that the flux equilibrates certainly more rapidly than the fastest sampling rates of the conventional filtration technique permit. There are at present two alternative solutions to these related problems: ( 1 ) T h e application of methods affording higher time resolution. T h e above-mentioned fluorescence technique (Moore and Raftery, 1979, 1980) employing fast kinetic instrumentation (stopped-flow) is to date the most direct and elegant case, where both mixing and spectrophotometric detection occur in a millisecond time scale. Alternatively, mixing can be accomplished in the upper millisecond time domain and the reaction can be terminated by chemical or physical means; detection of the reaction products and/or reacting species is accomplished subsequently on a slower time scale. This is the principle of the quenched-flow technique (Hess et nl., 1979, 1981, 1982; Neubig and Cohen, 1980). (2) The reduction of the number of receptors per “fluxing volume.” T h e following section considers these two approaches. Flux responses determined by the conventional filtration techniques imply taking measurements at times of 10-30 sec (depending on the skill of the experimenter and instrumentation) and upward. Considering the density of receptors per unit area (see Section IV) in native AChR membranes, the diameter of a sealed, average-sized vesicle (0.6 p m ) and its internal volume ( liter), an uptake of lo9 ions sec-’ per vesicle has been calculated (Moore and Raftery, 1980). That is, lo6 ions pass through each channel per second in a vesicle with lo4 channels. It is not surprising, then, that even the initial points of the assay cannot provide a direct measure of receptor activation, since the flux has to a large extent equilibrated within a few seconds even for low agonist concentrations and has ceased for high agonist concentrations: 86Rbf flux, for instance, is completed within 50 msec at 1 mM Carb (Hess et al., 1982). This does not imply that the slow, integrated flux measurements are uninformative, but rather that they are restricted to a narrow set of conditions-in particular they allow the exploration of only low agonist concentration domains-and that they possess limited and only comparative value (e.g., assessment of relative potencies/efficacies of ligands).

-

DEVELOPMENTS IN STRUCTURE AND FUNCTION

303

A second important problem is that of the contribution of AChRmediated flux to the total observable ionic exchange. First, not all vesicles contain functional receptors, especially when Electrophorus tissue is used. Hess et al. (1975) indicated that as much as 85% of the AChR-rich vesicles prepared from that source were “unspecific,”in the sense that they are not affected by cholinergic ligands (i.e., either lacking receptors or containing inactive AChR). Second, not all native AChR-rich vesicles are sealed, and a substantial proportion of the measured flux corresponds to passive leakage of the ion in question. The unstimulated 22Na+-flux(i.e., in the absence of agonist) is not described by a single rate constant, but a “significant” proportion of the ion is released with a half-time in excess of 60 min (Neubig and Cohen, 1980). Kim and Hess (1981) calculated that 30-35% of the slow exchange of ions is not related to AChRmediated flux, and equilibrates in about 20 min. Hess and co-workers have contributed to overcoming these difficulties first by introducing a procedure to subfractionate vesicles into sealed and leaky ones (Hess and Andrews, 1977). This is accomplished by loading the vesicles with Cs, and subsequently separating the heavier, Cs-retaining microsacs in a density gradient. More recently, Bernhardt et al. (1981) introduced further improvements to this type of technique, using Li+ as a nonradioactive tracer ion incorporated into the vesicles by pulses of agonist. Second, Kim and Hess (1981) have characterized the relative contributions of AChR-mediated fluxes and the unspecific ones occurring in the excitable vesicles themselves, and have therefore been able to account for what they qualify as the underestimation of the rates in the previous flux measurements of Kasai and Changeux (1971a,b). Kim and Hess (1981) also use this information to reject the attribution of diminished flux amplitudes to desensitization phenomena (Popot et al., 1976). A third difficulty concerns the heterogeneity of the vesicle population in terms of size, shape, number of active receptors, their distribution, and, above all, the inability to control such parameters. Most recently, attempts have been made to purify the conventional AChR vesicles attending to some of these parameters (Flanagan et al., 1976; Jeng et al., 1981; Johansson et al., 1981). The effort invested in these attempts is amply justified, since quantitative flux measurements under defined, controllable conditions in native vesicles are still able to provide the closest in vitro approximation to AChR-mediated ion translocation in vivo. Formal treatment of the inhomogeneity problem mentioned above can be found in a series of works by Bernhardt and Neumann (1978, 1980a,b).Bernhardt and Neumann (1978) first represented the number of activable channels as a continuously varying density. A fixed number of open channels per vesicle, and not a statistical distribution of gating

304

F. J. BARRANTES

units, modeled the activation phenomenon. Subsequently (Bernhardt and Neumann, 1980a,b), these authors developed more general analytical expressions for flux rates and amplitudes from and into sealed vesicular structures of different size and number of channels, covering both activation and inactivation phenomena. Experimental parameters such as the average volume of the vesicles (i),initial concentration of tracer ion (C,) in a collection of vesicles, “density” of the vesicles (D,i.e., number of vesicleshnit volume) in a suspension of volume V, mean number of channels per vesicle ( i )and , their variance (a2), were formally related to the overall flux [,f (t)] and its time-dependent amplitude factor by expressions of the type: ( t ) = D V C,ieXp{-t?K(t) [ l where

K

- CT2/2tiK(t)]}

(1)

is an integrated amplitude factor,

depending in turn on k, the rate constant for ion transport through each channel, and ( Y ( T ) ,the time-dependent fraction of channels open at time t in a given vesicle. The essential outcome of their treatment is that flux rates are directly proportional to the average number of channels per vesicle and inversely proportional to the volume of the vesicle, whereas the flux amplitudes depend on the average volume of the vesicle. Bernhardt and Neumann (1980a) also question the need to invoke the concept of spare receptors, as used by Moore et al. (1979a) or Neubig and Cohen (1980) when inhibition of a fraction of receptors is accomplished by antagonist blockage. Moore et 01. (1979a) observed inhibition of flux only after about 70% of the ACh receptors were inactivated by toxin.

B. F i x x DOSE-RESPONSE CURVES A N D CHANNEL ACTIVATION One of the ultimate goals of flux measurements is the establishment of the dependence of flux rates on the concentration of ligands which activate (agonists) or inactivate (antagonists) the system. In order to have access to the high agonist concentration ranges essential for establishing adequate dose-response curves, fast mixingldetection methods are necessary. T h e application of the quenched-flow technique has provided progress in this direction. Neubig and Cohen (1980) used a doublemixing quenched-flow instrument which allows the mixing of membranes and agonist and subsequently the quenching of the reaction by

DEVELOPMENTS I N STRUCTURE AND FUNCTION

305

addition of high concentrations of both a competitive (2 mM curare) and a noncompetitive (0.2 mM meproadifen) antagonist. Cash and Hess (1980) used 30 mM curare to quench the agonist-AChR reaction, which is described by a single-exponential process in the range 0.1-0.3 mM carbamoylcholine (Neubig and Cohen, 1980). Maximal flux rates, however, cannot as yet be reliably measured, even with this technique: I n the presence of high agonist concentrations the 22Na+is released from native membranes within 24 msec, that is, before the first observable time (Neubig and Cohen, 1980). These authors had to resort to extensive inactivation (> 70%) of receptors with a-bungarotoxin in order to gain access to the time regime of the quenched-flow technique. The considerable improvement of the quenched-flow method over the conventional integrated flux measurements becomes apparent when one compares the preliminary dose-response curves so far obtained. Values of 600 p M (Neubig and Cohen, 1980) characterize the [L],,, for carbamoylcholine in quenched-flow experiments, whereas values of 15-30 p M result from the slow flux measurements (Popot et al., 1976; Miller et al., 1978; Moore et al., 1979b; Neubig and Cohen, 1980). T h e maximal rate of Na+ that could be measured with the quenched-flow technique is 37 sec-' (Aoshima et al., 1981), 84 sec-' (Aoshima et al., 1980), 65 sec-' (Neubig and Cohen, 1980),and more recently, 310 sec-' (Hess et nl., 1982). The advantages of simultaneous mixing and spectroscopic detection in terms of time resolution should be noted. Rates of 1500 sec-' have been measured with this technique (Moore and Raftery, 1980) in native membranes, and rates of 487 sec-' have been measured in reconstituted vesicles (Wu et al., 1981) at saturating Carb concentrations. I n the case of the quenched-flow experiments, and under the assumption that all a toxin sites are involved in ion transport, the flux rates obtained indicate that there are about 3500 Na ions flowing per a-toxin site per sec, (7000 ions/AChR monomer), that is, only 8% of the maximal rates for the same ion in chick muscle (Catterall, 1975) and even less than this value if the transport expected from single-channel measurements in Tmpedo (Schindler and Quast, 1980; Boheim et al., 1981) is employed. Some of the factors considered above may intensify the underestimation, but it is obvious that values of tIl2 of 10 msec for ssRb+ flux at l-mM Carb (Hess et al., 1982) leave little room for maneuvre. C. FLUXDOSE-RESPONSE CURVES, BINDING EQUILIBRIA, A N D DESENSITIZATION Dose-response curves furnished from rapid-flux measurements are still scarce, but general trends are already emerging. They confirm the

306

F. J. BARRANTES

need to revise the significance of the rather extensive data accumulated on the apparent equilibrium binding parameters of various cholinergic drugs (Table IV). T h e most widely used assay, the inhibition of a-toxin rate of association, usually yields two extreme values (designated t + 0 and t + = in Table IV) for the time-dependent inhibition of toxin rates. T h e interpretation of the apparent [L],., values is of course modeldependent, but intuitively the coincidence of the t -+ 3fl parameter obtained by this method and the corresponding values resulting from equilibrium dialysis or the like is clear: Prolonged exposure to agonists stabilizes the AChR in a state(s) D characterized by its (their) high affinity for the ligand, about two orders of magnitude higher than the corresponding resting state(s)R (Table IV). Whatever the exact interpretation of the nonequilibrium [~5],,~ values (t + 0) is, their assignation to a low-affinity form of the AChR remains unaltered. T h e contribution of the more recent rapid-flux measurements in the context of binding mechanisms is that they challenge the tenability of interpretations ascribing this particular low-affinity form of the AChR to the form leading to channel activation. This stems mainly from the discrepancy between the [L],,, (t + 0) values and the corresponding [L],, figures obtained in the fast-flux measurements. Neubig and Cohen (1980), for instance, report [ L ] , , ( t + 0) values of 0.6 mhf for Carb and 0.2 mM for phenyltrimethylammonium, a partial agonist. Inspection of Table IV clearly indicates that these values depart substantially from the aforementioned [[L],, assigned to the low-affinity state of the AChR. A further difficulty arises when attempting to correlate flux data with fractional occupancy of binding sites in the light of reaction mechanisms currently accepted for the action of agonists in uizro. T h e number of agonist molecules involved in the physiological gating phenomenon appears to be close to two or larger [see Colquhoun (1979), Gage (1976), Steinbach, (1980), and Adams (198 I), for discussion of the subject]. T h e Hill coefficient for Carb found in rapid-flux experiments is close to two (Neubig and Cohen, 1980), and the need to consider a reaction mechanism involving a biliganded, low-affinity state of the AChR preceding the final activated (open) state appears justified. T h e number of unknown variables still present in models like the one shown in Fig. 7 is large, and one should exercise caution in using such multiparameter models even in their provisional character of useful working hypotheses. Simpler reaction mechanisms based on the Katz and Thesleff (1957) type of cyclic scheme (Weiland P t al., 1977; Barrantes, 1978; Boyd and Cohen, 1980) have been used to account for the kinetics of binding of Carb, ACh, or suberyldicholine, and the in ~ ~ i t rinactivation o processes ascribed to the desensitization phenomenon. Although the cyclic mechanisms involving

DEVELOPMENTS IN STRUCTURE AND FUNCTION

307

FIG. 7. Multistate model of AChR-agonist interactions, including (1) binding of two agonist molecules (A) to AChR in resting state, R (preexisting ligand binding and associated with closed channel), or desensitized state, D, and (2) isomerizations between the two states and between the closed-channel and activated, open-channel conformation R*. The new intermediate low-afFinity state postulated by Neubig and Cohen (1980), R,, is included. The model is a development of previous work (Weiland et al., 1977; Barrantes, 1978; Boyd and Cohen, 1980; Heidmann and Changeux, 1979; see also Barrantes, 1979) based, in turn, on early schemes (Changeux et al., 1976; Katz and Thesleff, 1957).

only monoliganded AChR states or even simpler versions can be satisfactorily modified by the addition of a rapid isomerization step to include the channel activation (Bonner et al., 1976; Barrantes, 1978), and yield parameters for the activation step compatible with those observed in vivo, such mechanisms are less tenable when biliganded receptor forms are incorporated and the in vitro rapid-flux data are considered. Fractional occupancy of binding sites by competitive antagonists has been used as a means to determine the number of agonist molecules needed to activate ionic flux. The results are still contradictory. Lindstrom et al. (1980a) and Anholt et al. (1981) observed a linear diminution of activable sites upon titration of AChR vesicles with a toxin and interpreted this result to indicate that monoliganded receptor (AR in Fig. 7) controls channel opening. If the view currently accepted by electrophysiologists were correct (see, e.g., Adams, 1981; Colquhoun, 1979; Steinbach, 1980),and biliganded AChR (A2R) determined gating, the concentration dependence of toxin inhibition would be hyperbolic instead of linear. This is precisely what the results of Sine and Taylor (1980, 1981) show. Reduction and subsequent alkylation of the AChR “freezes” the receptor in a very low-affinity state whose [L],, values for agonists mimic those found with electrophysiological techniques (Walker et al., 198la; Barrantes, 1980). Walker et al. (1981a) further proposed that the reduction reaction could involve attack of thiol groups in the AChR channel itself. But aside from these special cases, the low-affinity state of the AChR detected in binding experiments (Table IV) does not appear to correspond in a direct manner to the one leading to channel activation.

ZP'O LP'O 8000 20 52-6'5

POO'O 6ZO'O (.'?I)Z'O P("Y) 7110.0

WOO 800'0

P20.0 ED'0-lO'O

190

S'Z 91'0 O'Z60'0

0'1

ROO'O 0' F 800.0

co 0 m

15.0

PO00 PIO'O 0'2

ozno 890'0

unon wn 50.0 S'O SO'O W'O S' I

W V

Carb

24 20 40 22 1.4 1.9 1.5

30-50 20 20 20 8

35-121 70- 148

0.5 0.4 5 50 60 0.1 0.05-0.12 0.5 0.5 0.5 0.02 58-170 3.5 6.3 2.3 3.0 11-36 18-40

Electroph Elerhoph Electrophonrs Electmphonrs Electmphm Electroph Torpedo mannorata Torpedo mamorafa Torpedo marmordu Torpedo marmorata Torpedo mannoraka Turpedo mannoraka Tapedo mamorata Torpedo califmica Torpedo colifmnua Torpedo califmua Torpedo m m o r a f a Torpedo caliiornica Torpedo californua Rat diaphragm (denervated) Rat diaphragm (denervated) Rat diaphragm Cat leg muscle (denervated) BC3H-1 cell line BC H-1 cell line

Membrane Membrane Membrane Membrane Purified Purified Membrane Membrane Membrane Membranes Triton extract Membranes Membranes Membranes Membranes Membranes Membranes Membranes Purified Crude homogenate Triton extract Homogenate Membranes Cultured cells Cultured cells

Elerhoph Electrophonrs

Membranes Membranes Membranes Membranes Purified Membranes Membranes Membranes Cultured cells Homogenate Triton extract Membranes Membranes

1 (a-BuTX)

I (a-Bum) I (a-BuTX) I (a-BuTX) 1 ([aJHICT) I (a-BuTX) I (a-Bum) R (a-Bum) R (a.BuTX) R (a-BuTX) R (a-BuTX) 1 (Nala u toxin) I ( N a p a toxin)

Bulger and Hess (1973) Fu ef al. (1974) Weher and Changeux (1974) Kasai and Changeux (1971h) Meunier el d.(1974) Meunier and Changeux (1973) Cohen et al. (1974) Weber and Changeux (1974) Griinhagen and Changeux (1976) Franklin and Potter (1972) Eldefrawi and Eldefrawi (1973a,b) Bonner el al. (1976) Neubig and Cohen ( I 979) Quast rt al. (1978) Blanchard el al. (1979) Weiland el 01. (1977) Barrantes (1980) Walkerrf 01. (1981a) Raftery et al. (1976) Colquhoun and Rang (1976) Colquhoun and Rang (1976) Colquhoun and Rang (1976) Barnard ef al. (1977) Sine and Taylor (1979) Sine and Taylor (1980)

I (a-BuTX) E (WID=+) 1 ( N a p a toxin) E (PHlDeW E (rHIDeca) I (Naja a toxin) E (PHIDe4 F (quinacrine) I (Nala a toxin) 1 (a-BuTX) I (a-Bum) I (a-BuTX) I (PHI-a-toxin)

Bulger and Hess (1973) Fu rf al. (1974) Weber and Cbangeux (1974) Kasai and Changeux (1971b) Meunier and Changeux (1973) Weber and Changeux (1974) Weber and Changeux (1974) Criinhagen and Changeux (1976) Sine and Taylor (1979) Colquhoun and Rang (1976) Colquhoun and Rang (1976) Barnard el 01. (1977) Barrantes (1980)

I (rH1Deca) I (Naja a toxin) I (PHID-) I (PHIDeca) I (PHlDeca) F (dansyl-C,-choline) I (Naja a toxin) F (quinacrine)

F (intrinsic)

K (PHICarh)

Deca

34-50

0.7

8 0.9 0.8 1.3 0.02 1 0.8 0.74 0.6 22 2.1 8.6 3.0 50

Eltcfroph

Elerhophmus Elerhoph Torpedo Topedo mannorata Torpedo mannoraka BC3H-1 cell Line Rat diaphragm (denervated) Rat diaphragm (denervated) Cat leg muscle (denervated) Torpedo manmala

(continued)

PZ'O 8F'O 22.0

vo 2'0 p(".Y) I ' O - F E O O EZ'O LI'O LI n

(1.Y)S'I-FO

6E'O

zo LI'O Z (1 3.L-P 0%

2

m

6X HI1

06 09 Ot z9 19

SubCh

0.5-0.8 0.31 7.7 2 4.6 27.5 I 0.4-0.97 0.13

01 w

0.04 0.045

Cat leg muscle (denervated) Rat diaphragm

0.55

Rat diaphragm

0.4-0.7 0.38 0.033 -c 0.006 0.3 0.01 0.15-0.24 0.0004-0.004 0.17 0.01 (0.005. 0.025)

c.

PTA

35 111

1.6 26.2

64 427

0.2-2.5 81.7

BC3H-I cell line Tmprdo rnlifrrnim Tmpfdo mammala

BC3H-1 cell line Torpedo mammala BC3H-1 cell line Torpedo cahfmira BC3H-I cell line Tmpede mannoratn T. caiif0rnUa T. mbii. Tmppdo califmnjca BC3H-1 cell line

Membrane Purified (junctional) Purified (extrajunctional) Cultured ceUs Membranes Membranes Cultured cells Membranes Cultured cells Membranes Cultured cells Membranes

I (a-BuTX) I (a-BuTX)

Barnard e/ a/. (1977) Brockes and Hall (1975h)

I (a-BuTX)

Brockes and Hall (1975b)

1 ( N a p a toxin)

I (Naja a toxin) E ([aH]d-TC) 1 (Naja a toxin) I (a-CT) 1 (Naja a toxin) I ( N a p a toxin) I (NQ@ a toxin) I (Naja a toxin)

Sine and Taylor (1979) Weiland and Taylor (1979) Neubig and Cohen (1979) Sine and Taylor (1981) Barrantes (1976, 1978) Sine and Taylor (1979) Weiland and Taylor (1979) Sine and Taylor (1980) Neubig and Cohen (1979)

Membranes Cultured cells

I (Naja a toxin) I (Naja a toxin)

Weiland and Taylor (1979) Sine and Taylor (1979)

Membranes Cultured cells

I (Naja a toxin) 1 (Na@ a toxin)

Weiland and Taylor (1979) Sine and Taylor (1979)

Nicotine Torpedo calt/nrnira

BC3H-I cell line

The term equilibrium dissociation constant is reserved for determinations employing equilibrium measurements. [Lb.,are the half-concentrations for 1 -+ 0 and t + co, respectively (sometimes referred to as K , , , "protection constant"). Abbreviations used are: ACh, acetylcholine; C a h , carbamoylcholine; D e a , decamethonium; Hexa, hexamethonium; I-TC, d-tubocurarine; BuTX, a-bungarotoxin; Naja a toxin, a-neurotoxin from Naja naja siomeruis; a-CT, a-cobrotoxin (N.n.atra). SubCh, suberyldicholine; PTA, phenyltrimethylarnmonium. Methcds: 1, inhibition of binding of given ligand; E, direct determination of equilibrium dissociation constant; F, fluorescence of specified ligand; K , indirect determination of equilibrium constant from kinetic parameters. Gibson (1976) fitted his data to an allosteric model and calculated the equilibrium dissociation constant for the two states, T and R.

312

F. J. BARRANTES

Neubig and Cohen ( 1980) ascribed the “conventional” low-affinity state to a biliganded form, A,R,, different from the native resting (activable) biliganded receptor, A2R,which ought to precede channel gating (A2R* in Fig. 7). I have included the corresponding monoliganded species in the scheme in Fig. 7 not because of symmetry considerations but attending to current electrophysiological data to be discussed below. One interesting property of Neubig and Cohen’s (1980) low-affinity R, state is that it will contribute to the apparent inactivation (desensitization) processes (Fig. 7). When only the biliganded AChR is considered (AR,is not allowed to exist), desensitization ought to be apparent only at high agonist concentrations, at which the maximal response will be diminished. Under such conditions desensitization would show u p as a biphasic phenomenon. Responses at low ligand concentration should not be affected. If on the other hand the monoliganded form of this lowaffinity AChR state (AR,) manifests itself, two phases of desensitization should be apparent at all ligand concentrations. Only one desensitization phase was originally observed in similar measurements undertaken in EIectrophorus membranes (Aoshima et al., 1981), but in more recent studies (Walkeret al., 1981b; Hess et al., 1982) two inactivation processes were found in Tmpedo membranes. T h e rapid inactivation process proceeds with a rate of 2 sec-’, and the slow inactivation step superceeds it with a rate of 0.12 sec-’ in the presence of Carb. Bonner et nl. (1976) first characterized this slow phase of desensitization with rapid kinetic methods and reported values of 0.16 and 0.37 sec-* for this process with ACh and Carb in Torpedo membranes. Later, Barrantes (1978) found values of 1.2- 1.5 sec-’ with suberyldicholine, which may correspond to the faster desensitization process. These values also show close resemblance to those described more recently by Sakmann et al. (1980) (to be discussed later) in their study of desensitization by patch-clamp techniques. Only one very slow desensitization phase is apparent in reduced Tmpedo AChR (Walker et al., 1981a). Recovery from desensitization is a slower process occurring over seconds to minutes, depending on various parameters (see Adams, 1981; Cohen and Boyd, 1979; Boyd and Cohen, 1980). In the absence of agonist, the majority of receptors should exist in the resting, activable conformation R (Fig. 7). At equilibrium, the R c, D balance is characterized by a net predominance of R over D. T h e R/D equilibrium constant measured in various systems indicates values between 10% (Weiland and Taylor, 1979) and 20-25% (Heidmann and Changeux, 1978; Barrantes, 1978; Boyd and Cohen, 1980) desensitized AChR preexisting ligand binding.

DEVELOPMENTS I N STRUCTURE AND FUNCTION

D. NEWVIEWSON

THE

313

BEHAVIOR OF ACH-OPERATED CHANNELS

T h e patch-clamp technique has allowed the direct observation of the currents flowing through individual ionic channels. Since its first application to ACh-activated extrasynaptic channels (Neher and Sakmann, 1976a), glutamate (Patlak et al., 1979; Cull-Candy and Parker, 1982), GABA (Bormann et al., 1981), Na+ (Sigworth and Neher, 1980; Horn et al., 1981), Ca2+-activatedchannels (Marty, 1981; Hamill, 1981; Pallota et al., 1981; Colquhoun et al., 1981), K+ channels of the anomalous (inward current) rectifier type (Ohmori et al., 1981; Fukushima, 1981), and Ca+ channels (Lux and Nagy, 1981; Lux et al., 1981; Fenwick et al., 1982) have been explored with this technique. Many other neurotransmitter, hormonal, ionic, or voltage-sensitive systems are certain candidates for the immediate future. It is possible to distinguish three periods in the evolution of findings from the patch-clamp methodology. The initial period, which extends to 1979, appeared to provide experimental confirmation of many of the hypotheses on AChR-channel properties implicit in the theory behind noise or perturbation techniques, transforming essential assumptions of the macroscopic methods into observable variables. The picture which emerged from all available techniques was convergent on a simple view of the ion gating process: Channels behaved as two-state switchlike devices, open or closed. The second period (the present one) coincides with the availability of several methodological refinements (Neher et al., 1978; Sigworth and Neher, 1980; Horne and Patlak, 1980; Neher, 1982; Hamill et al., 1981), enabling the observation of new phenomena in AChR channels. The simplicity of the all-or-none phenomenon gives way to a more complicated, subtle structure underlying the “apparent” single-channel event. I n fact, what was believed not long ago to be the unitary channel response is now recognized as a composite of open and closed states. Additional discrete open states have also been found. All these findings question the tenability of some of the classical views on the AChR gating. A brief description of the third and future stage in this evolution will follow after presentation of the characteristic elements of AChR channel activity. 1. Quantitative Kinetic Parameters Associated with Single-Current Recordings Let us start with a brief phenomenological description of patch-clamp recordings. A typical trace usually appears as rapidly flickering pulselike current events (Fig. 8). The silent intervals during which channels are closed can be easily distinguished from those in which one or a few are

314

F. J. BARRANTES

gaps

40 rnsec Flc.. 8. Patch-clamp recording from a rat muscle cell in culture (“myoball”) in the presence of 5-p.if i\Ch at -70 mV membrane potential. A “classical” single-current event of the all-or-none open-close type is exemplified by the rightmost current trace. The beginning of the trace shows several short closures (gaps) within the apparent open period, which is redefined as an N-burst (see text), a composite of open and closed states. Colquhoun and Sakmann (1981) found at least three time constants of gap durations (rt 45 psec, representing 77% of the events; T , 350 psec representing 21%;and T~ < I msec representing 3%). Notice the higher noise in the open-channel state, a feature characterized by Sigworth (1982). The original record was provided by Dr. 0. P. Hamill.

-

-

open when the number of activated channels is kept low. This is accomplished in practice by working in the low agonist-concentration domain and ( ( 1 ) by choosing a membrane area with low receptor density, as in extrasynaptic regions (Neher and Sakmann, 19’76a,b), (b) by using quasi-irreversible antagonists to eliminate most of the receptors, or (c) by exploring high concentration domains, but it is then necessary to allow agonist-induced desensitization to occur before individual channel gating can be resolved (Sakmann rt al., 1980). When the number of contributing channels is small, opening and closure of a given channel will take place most likely before any other individual channel opens. On probabilistic grounds the distribution of the conducting and silent intervals can then be attributed to open and closed states of a single unit, whose gating is rare and apparently random. Although the stochastic model of chemical reactions specifies that the transition from a given state to another state is determined by the transition probability per unit time,ptr,iridr/mdeiitly of the number of molecules which happen to be in either state I or j (see, e g . , Zwolinski and Eyring, 1947), the technical difficulties arising when the number of active units is large still impede the analysis of traces from many contributing channels. Assuming the

DEVELOPMENTS IN STRUCTURE AND FUNCTION

315

independence of channels, the transitions between the nonconducting and the conducting states are treated as Markov processes. The duration of each state (dwell time) is exponentially distributed in time. The inherent advantages of dealing with a single operating unit are made apparent in the fact that independent and direct estimates of the individual rate constants for channel opening and closing can be obtained from such distributions. For comparison, in a “macroscopic” measurement (e.g., noise or relaxation experiment), and in the simplest case of a two-state first-order (A c, B) reaction, the relaxation rate at which the system strives toward equilibrium is given by

that is, a composite of the individual probabilities determining the rate constants that obtain. The dwell times displayed repeatedly and sequentially by a single channel reproduce in their random behavior that of an assembly of many channels simultaneously driven out from equilibrium; the comparison drawn by Ehrenstein et al. (1974) with a radioactive decay curve is a useful analogy. Under certain circumstances the individual current events are grouped into bursts whose average duration depends on agonist concentration (Fig. 8). This type of burst will henceforth be called N-burst2 because it occurs under apparently normal circumstances (Nelson and Sachs, 1979; Colquhoun and Sakmann, 1981) and is to be distinguished from B-bursts ( B for blockers, following Colquhoun and Sakmann, 1981) and D-bursts, observed with desensitizing agonist concentrations (Sakmann et al., 1980). The significance of the different types of bursts will be clarified below. The square-like pulse events (“single channels”) are

* Burst is a term coined by Neher and Steinbach (1978) when describing the modification of the usual squarelike shape of single-current pulses occurring in the presence of local anesthetic substances. TheN-burst is defined as a group of openings (believed to arise from activation of a single AChR channel) separated by interruptions (gaps) whose duration is short relative to the total length of the burst. N - is meant to imply normal; an alternative philological root is found in Narhschlag, the German word for backlash, as first used by Neher and Sakmann, to describe the multiple gating phenomenon. Nelson and Sachs (1979) first reported the occurrence of this phenomenon in embryonic muscle cells (myoballs) in culture. The term Nachschlag was used only subsequently (e.g., Adams, 198 1; Lester, 1982) and referred to Nelson and Sachs but without clarification of its origins or significance. How normal and widespread the phenomenon is awaits further experimentation. It also appears to occur with acetylcholine (in addition to suberyldicholine) and in the junctional region of adult muscle (in addition to myoballs) (B. Sakmann, personal communica tion).

316

F. J. BARRANTES

“compressed” in the slow records and appear as spikes. Bursts, in turn, can be seen to be grouped into a higher order association-clusters of bursts-when long strings of records are examined (see Fig. 1 1 ) . T h e separation of bursts and interburst intervals within a cluster and the distances between clusters and intercluster intervals also vary with the agonist concentration (Sakmann et al., 1980). Quantitative parameters can be defined for these “aggregates” in a manner analogous to those used for the single-channel traces; tb is the burst duration; t i the interburst interval (see Fig. 1 1 ) ; accordingly, cb is the cluster duration and c , the intercluster interval. Typical time scales of busts and clusters are a few hundred milliseconds and seconds, respectively. These parameters have been been used to study the kinetics of channel blockage and desensitization, as will be illustrated. Recently, short interruptions within an individual single-channel pulse could also be resolved (Colquhoun and Sakmann, 1981). These silent periods within the apparent open state of the channel are termed gaps and have average durations of tens to hundreds of microseconds (Fig. 8). T h e gaps complete the presently available repertoire of channel phenomena amenable to observation with improved patch-clamp instrumentation (Hamill et al., 198 1). Having named the individual features distinguishable to date in the microsecond-second time domains, we can now illustrate the way the corresponding average kinetic parameters are derived and some of the information to be gained from these. T h e basic assumption here is that the single-current events are exponentially distributed random variables. Thus analysis of a large number of events (gaps, bursts, etc.), or their corresponding interevent durations can most conveniently be done in the form of the distribution of their dwell times as a function of the number of events. In view of the stochastic nature of the events, exponential fitting of the distributions will yield mean (average) time constants of such distributions, as expected for Poisson processes. We shall also see that in some cases the analysis yields a sum of exponential terms; a collection of Poisson phenomena is then assumed. the more extensively characterized of these time (decay) constants are T, (mean channel-open time) and 7, (mean channel-closed time, Fig. 9), but analogous mean times can be defined for the other channel-associated parameters. T h e relationship between the experimentally determined T, and T, and the rate constants for channel closing and opening, respectively (a and p), is of course model-dependent. For a two-state reaction scheme of the type

317

DEVELOPMENTS IN STRUCTURE AND FUNCTION

u)

c C

$

0

50

0

loo

100

50

to (rnsec)

5

tc (rnsec)

20

50

[AChl PM

FIG.9. Distribution of the durations of open (to) and closed (te)states of current pulses during D-bursts and dependence of their reciprocals on agonist concentration. (a,b) Sequences of D-bursts occurring in the presence of 10-pM ACh (12"C, -130 mV) were analyzed for dwell times in the open and closed states. The distribution of intervals in each state was fitted with single exponential functions (. . .) yielding mean open T, and mean closed T~ times of 10.6 and 18 msec, respectively. (c) Double-logarithmic plot of the reciprocal of mean open (open symbols), T,, and close (closed symbols), T,, time constants as a function of agonist concentration. T , is noticeably concentration dependent, whereas 7" is not. (From Sakmann et a/.,1980.)

the mean dwell time in state 0 is given by ( L 1 ) - l , and the mean dwell time in state C is (kJ1. That is, the mean period spent in a given state is given by the elementary rate constant leading away from that state. Generalization of this to other more complicated reaction mechanisms is possible (Colquhoun and Hawkes, 1977; Neher and Steinbach, 1978); the reciprocal of the mean dwell time in a given state is the sum of all reaction rates depopulating (i.e., leaving) that state: kin

= T~ =

(5)

n

I n the case of the binding-isomerization models which are currently taken as the most plausible ones, such as the two-binding two-isomerization scheme below (see also Fig. 7): K,

R

+ A-

ki k-1

KZ

RA

A k-a

RA,

.B

a

R+A,

(6)

the reciprocal of the mean open or close times are related to the rate constants for channel opening and closing by obviously more compli-

318

F. J . BARRANTES

cated relationships than in the simpler two-state model above:

a (7) (8 ) I h , = p‘ = {[A]k,/([A]k, + k 1 ) } , p (assuming k l = 2 k 2 , k-, = 2 k 1 , and a + p e k, + kl). Sakmann et nl. (1980) studied the behavior of AChR channels exposed to desensitizing agonist concentrations, and within a limited ACh concentration domain they could follow the behavior of mean open and closed times as illustrated in Fig. 9c. T h e intersection of the two reciprocal time constants T~ and I, occurred at about 22-pM (-90-mV holding potential) and 17-pM (- 130-mV) ACh. This intersection can be defined as the concentration of agonist, at a given membrane potential, at which the apparent rates of channel opening and closing are equal, that is, an “apparent” equilibrium isomerization constant results (which is less prone to depend on modeling assumptions). ( ~ “ ) - l= a’is independent of agonist concentration and becomes slower with membrane hyperpolarization; ( ~ ~ ) -=l p’ depends on agonist concentration, this dependence being described in a first approximation by a power law with an exponent of 1.4. This has been interpreted as being consistent with reaction schemes involving the binding of two agonist molecules preceding channel opening. U T , , = a’ =

2. Fin r Structurr .f f l i p “Siriglr”-CzLrmit Pulse: The Multiplr Gcitiiig Piirnomenon

Improved instrumental resolution has recently enabled the observation of short closure (gaps) occurring within the apparent open channel state in the sole presence of agonist, even at low agonist concentration. There are various types of gaps, attending to their mean time distributions. The shorter gaps so far observed have a mean duration of about 50 psec. In addition, Colquhoun and Sakmann (1981) observed gaps of about 10 times longer duration in muscle endplates, which presumably correspond to the longer gaps described by Nelson and Sachs (1979) in myoballs. These constitute the intermediate population (7, 0.7 msec) of the former authors. T h e briefness of the “short” gaps led Colquhoun and Sakmann (1981) to infer that they are unlikely to be associated with t w o independent channel activations. Instead, the suggestion was made that they could arise from rapid open-close transitions of the channel (multiple gating) occurring during the time for which the AChR remains occupied by the agonist, as previously considered on statistical grounds by Colquhoun and Hawkes (1977). Three other interpretations of the presence of gaps were considered by Colquhoun and Sakmann (1981): ( a ) blocking by the agonist, as previously observed by Adams and Sakmann (1978) in the case of decamethonium; (6) blocking by an unknown L-

DEVELOPMENTS IN STRUCTURE AND FUNCTION

319

endogenous substance; and ( c ) blocking by a mechanism related to ion permeation. Hypothesis (a) was rejected and the others not considered further. Channel-blocking mechanisms will be dealt with later. T h e occurrence of the intermediate type of gaps was not elaborated. Gaps are electrically silent events. Their significance remains to be evaluated, but one can already advance that these transient closures within the apparent open state will not be a mere addition to the phenomenological repertoire of channel parameters. Instead, these substructures will probably constitute manifestations of an electrically silent AChR state apparently different from the resting state. Observation of the gaps forces a redefinition o’f the parameters previously used to derive the channel lifetime: What was previously considered to be the channel open state now becomes a composite of open and closed states. Thus, the apparent length of the “open” channel state previously used to derive the rate constant for channel closing ti in a macroscopic measurement (e.g., noise or relaxation experiment) now becomes a mean N-burst duration, which includes both open and “closed within the open” phenomena [the open state($ + gaps].

3 . Heterogeneous Distribution .f Mean Open Times: The N-Burst The first level of supraorganization of the individual current pulses is the burst. T h e heterogeneity of N-burst lengths manifests itself in the histogram of apparent open times (Fig. 3 of Colquhoun and Sakmann, 198 l ) , which needs to be fitted with at least two exponentials. An N-burst is defined as a series of openings interrupted by gaps that are less than some critical duration (0.5-3 msec). Since this length is very much smaller than the mean intervals betweefi current pulses assumed to be independent, the N-burst duration is an operationally well-defined quantity. The two time constants fitting the N-burst distributions were rf = 0.15 msec (25%)and T, = 10 msec (75%)with 20 nM suberyldicholine in the perisynaptic region of frog neuromuscular region (Colquhoun and Sakmann, 1981). The slow time constant is the one which would be apparent in noise or relaxation analysis as a single component of about 10 msec, from which false values for the mean channel lifetime would be derived. The true channel open time is clearly shorter, after taking into account the temporarily closed periods (gaps), for the mean open time is the mean number of openings per N-burst ( i i ) multiplied by the mean length of an opening plus (ii - 1) times the mean gap duration (&). Applying this notion to the sequential scheme [Eq. (S)]the mean number of openings per N-burst is given by

320

F. J. BARRANTES

where the mean length of a gap is

and the resulting “true” mean open time is given by

T h e actual open lifetime is of course inaccessible as yet, since one cannot exclude the possibility of openings lasting less than the currently available resolution of the technique (a few microseconds). In any event, using the figures of Colquhoun and Sakmann (1981) mentioned above, the average N-burst consists of roughly four openings in quick succession, so the mean channel-open time would be about one fourth of the observed 10 msec (i.e., only 2.5 msec). Estimates of the rate constants for channel opening (p) and for ligand dissociation ( k - 2 ) can be obtained from the measurement of the mean gap duration and from the mean number of gaps per N-burst as given above. Values of /3 = 10,00015,000 sec-’ and k2 = 2000 sec-’ have been derived with suberyldicholine in adult muscle (Colquhoun and Sakmann, 1981). It is of particular interest to analyze these new findings in the light of theoretical considerations (Colquhoun, 1979; Colquhoun and Hawkes, 1977, 1981) in relation to the stochastic properties of channel behavior. One of the above properties concerns the equilibrium distribution of states, here listed for the two-agonist binding scheme [Eq. (6)]:

where [A] is the agonist concentration and the K’s are equilibrium dissociation constants [Eq. (6)]. Using currently accepted values for the various rate constants of agonist action obtained from macroscopic measurements, Colquhoun and Hawkes (1981) were able to express these equilibrium state probabilities in actual lifetimes spent in each state, from which it became apparent that, for independent binding steps with l and at ligand concentrations apparent K, values of about 30 ~ h (ACh), below this value (but still reasonably high to open about 5% of the channels) the channel spends most of the time in the biliganded activated (open) state R*A2 (see Fig. 7). However, 97% of all single occupancies would not be followed by channel opening via R*Az (though there is a

DEVELOPMENTS IN STRUCTURE AND FUNCTION

32 1

0.7% chance of opening via R*A). This is because dissociation of the agonist occurs before a second ligand has a chance to bind: a closed AChR channel with only one agonist bound has a higher probability (97%)of loosing the ligand than of reaching the biliganded state. There is only a roughly 2.5% chance that monoliganded receptors become doubly liganded (which does not mean that they necessarily open thereafter). However, once the biliganded state is reached, channel opening has almost the same probability as the dissociation of the ligand. This implies that doubly occupied channels may continue to open several times after the ligand concentration has fallen to zero. I n addition, whenever the states RA or RA2 are reached (in the presence of finite agonist concentrations) there is the possibility of any number (0 to m) of RA + RA2 oscillations (state transitions) before leaving to either R or R*A2. Here one finds a possible explanation of the N-burst (the multiple gating, Nachschlag) phenomenon. Although the idea is tempting, it is still premature to make it of general applicability. 4. Implications of the Assignation of the Rate-Limiting Step(s)

An outcome of the above finding (which is of course modeldependent) is its contradiction of the widely accepted assumption that agonist binding steps are faster than the close-open isomerization. If the multiple gating hypothesis were correct, the possibility should be entertained that the initial reaction steps in Eq. (6) proceed slower than the channel gating kinetics, constituting the rate-limiting steps in the activation pathway. A possible advantage of such constraints was discussed by Barrantes (1980) in relation to the affinity states of the AChR. In fact, two sets of experimental data in which the association rates of agonist binding have been measured in vitro with fast kinetic techniques yield similar on-rates of 1-2 X lo7 M-' sec-'. In one case (Neumann and Chang, 1976) AChR binding to T. califmica purified AChR was measured by absorption techniques, using murexide as an indicator of ACh binding. In a second case (Barrantes, 1978) the intrinsic fluorescence of membrane-bound AChR from T. marmorata was monitored to follow SubCh binding. These figures probably represent an overall, complex rate parameter involving some faster elementary steps. Off-rate constants were on the order of 10-3000 sec-'. These figures contrast with those obtained with a fluorescent agonist, NBD-choline, with which rates of 1.2-40 x lo* M-' sec-' have been reported (Prinz et al., 1980). The latter values would certainly make the rate-limiting character of the binding step(s) untenable. It is interesting to note that using essentially different types of arguments and experimental observations Lands et al.

322

F. J. BARRANTES

(1981) have recently calculated binding rates of 2-5 x lo7i t V 1 sec-' for ACh at the living end plate. 5. Chntinel Hi&wg.eiieity in Embl-rotiic n ~ i dDenerz1nted Muscle Denervation hypersensitivity is a well-known phenomenon that follows the sectioning of the motor innervation. One manifestation of this pathological situation is the spread of sensitivity to agonists, normally localized at the end-plate region, to the whole of the sarcolemmal surface. This phenomenon is accompanied by the appearance of newly synthesized AChR molecules at the surface membrane, and the normal density of AChRs of extrajunctional areas increases dramatically (200%) within a few days, depending on the species (see Fambrough, 1979). The gating properties of these AChR channels have been characterized by fluctuation analysis (Katz and Miledi, 1972; Neher and Sakmann, 1976b; Dryer et nl., 1976). Basically, the extrajunctional type of channel appear to have smaller conductance and slower kinetics (longer mean open time) than the corresponding mature junctional type (Gage and Hamill, 1980). The norrti(iI ontogenetic development of AChR channel gating properties in rat muscle cells appears to recapitulate the pathological situation found in the denervation of adult muscle. T h e analogy extends to the distributional habits of the AChR, initially diffusely located, mobile and with high turnover, and localized, relatively immobile and metabolically stable at subsequent stages of development. Diffusely located AChRs in embryonic muscle cells display initially the gating characteristics of the extrajunctional channels in adult tissue (Michler and Sakmann, 1980). Neonatal muscle cells still show the extrajunctional type of gating during the initial stages of synapse formation, but subsequently, a progressive transformation to the junctional type occurs (Sakmann and Brenner, 1978; Steinbach et nl., 1979; Michler and Sakmann, 1980). Both types of channels coexist at intermediate stages of development. Recent single-current recordings from embryonic rat muscle cells (Hamill a1 01.. 1981; Hamill and Sakmann, 1981) and from denervated adult frog muscle (Sakmann et d.,1980) have extended the description of the two types of channels occurring before innervation and after denervation, respectively. Most electrical properties of the two channel types are essentially similar (reversal potential, cation selectivity, and pharmacological specificity). T h e crucial question as to what leads to the conversion and stabilization of the junctional type or the reappearance of the extrajunctional type remains unanswered (for discussion of this subject see Fambrough, 1979). Figure 10 shows current traces obtained with the patch-clamp technique from the sarcolemma of 14-day-old rat

323

DEVELOPMENTS IN STRUCTURE AND FUNCTION

y

l.l-T T S W7rVf -

4.2 2.8

100

msec

FIG. 10. Two types of AChR channels coexist in embryonic muscle: the junctional type (J), with a main conductance level of about 50 pS and a short lifetime, and the extrajunc-

tional type (E), of longer duration and smaller conductance (-35 pS). A discrete sublevel (S) occurs in bothjunctional and extrajunctionalchannels. Notice the nonintegral nature of the sublevel S in comparison to the main levels J or E. (From Hamill and Sakmann, 1981.)

“myoballs” (i.e., myotubes in cell culture 4 days after treatment with colchicine). Two types of AChR channels coexist within a very small radius of noninnervated sarcolemmal membrane (the one from which the patch pipette records): Ajunctional J type, by analogy to the one seen in adult end-plate region, and an extrajunctional E type. T h e J type (Fig. 10) is characterized by a main conductance of about 50 pS and a short lifetime; the E type has a longer duration but a smaller conductance (35 pS) Hamill and Sakmann, 1981). 6 . Additional Alien Phenomena That Challenge the Open-Close Switch: The Sublevels

The preceding paragraphs document the recently available information questioning the tenability of some of the assumptions of noise, relaxation, and patch-clamp data in terms of a simple two-state, open-shut mechanism. The channel heterogeneity observed in embryonic muscle does not contradict accepted dogma: Different channels coexisting spatially could each show gating modalities characteristic of extrasynaptic and synaptic type of receptors. This finding is reasonable enough; but Hamill and Sakmann (198 1) showed that in addition to these two gating habits about 10% of the channels display a more complex behavior. Aside from the main current levels of J- and E-type channels, both types share a sublevel of lower conductance (Fig. 10). Some characteristics of the main and substates have been summarized by Hamill (1982): (i) the conductance of all states follows Gaussian distributions, (ii) all states display ohmic behavior, (iii) conductances for all states have a weak temperature dependence (Qlo 1.2) whereas the average lifetimes are more strongly temperature-dependent (Qlo 3 3), and (iv) all states display similar ionic selectivity for alkali cations. The probability of the open channel adopting the sublevel is almost negligible under normal conditions. It increases with membrane hyper-

-

324

F. J. BARRANTES

polarization and upon decreasing the temperature. When the AChR channel has reached the sublevel, the probabilities of switching to the resting or to the main level are approximately equal, an experimental observation reminiscent of the statistical property of biliganded AChR in a two-binding, two isomerization scheme (Colquhoun and Hawkes, 1981). Hamill and Sakmann (1981) offered two alternative explanations for the multiple conductance states: subunit rearrangement or different aggregational forms of the AChR subunits. 7. Eloqiieiit Sileme: Deserwitization ,tlcmured by Loiig-Li-oed Electrically S i l m t I ti terunls The amount of membrane depolarization elicited by an agonist at the cholinergic synapse depends on the nature of such a ligand and its concentration at the synaptic cleft. Upon prolonged exposure to the agonist, however, the depolarization is not maintained, but a slower and spontaneous repolarization supervenes. This phenomenon, termed “desensitization” by Thesleff (1955), implies that in spite of the persistent presence of the agonist, the average number of channels in the open state decreases. T h e kinetics of this phenomenon is different from those of the open-close transitions of the individual receptor-controlled channels (see Gage, 1976). T h e first attempts to formalize the desensitization phenomenon within the context of AChR-agonst interactions date to the work of Katz and Thesleff (1957). They postulated the conversion of the active agonist-receptor complex into an inactive, desensitized state via a comparatively slow transition of the active complex involving dissociation of the bound agonist. Many more complex reaction mechanisms have since been invoked, the now-classical cyclic scheme of Katz and Thesleff (1957) having provided a useful frame of reference on which much of the current in uitro work has developed (Weiland et al., 1977; Barrantes, 1978; Neubig and Cohen, 1980; Cohen and Boyd, 1979; Boyd and Cohen, 1980; Heidemann and Changeux, 1979). As we have seen, a variety of AChR states leading to the activated AChR-channel complex are associated with the shut-channel conformations (e.g., R, RA, RA2in Fig. 7). These states are electrically silent and at first glance are indistinguishable from one another in a patch-clamp recording. T h e complicating addition of other states associated with closed channels would appear to make the situation worse, since their contribution could not be distinguished from that of the other electrically silent forms (i.e., having the same resting current value) in a patchclamp recording. Sakmann et nl. (1980)had only one clue for interrogating these silent domains of agonist action: the time ranges in which the desensitization

DEVELOPMENTS IN STRUCTURE AND FUNCTION

325

D

1

1 sec

ti

I ”

tb

D BURS1

CLUSTER OF BURSTS

FIG. 1 1 . Supraorganization of current events into D-bursts and clusters in the presence of desensitizing agonist concentrations. Immediately after contacting the cell surface, hyperactivity is manifested by gating of several AChR channels [at least four levels are detected (upper trace)]. After a few seconds, desensitization occurs (D). The lower trace shows bursts of current (D-bursts) hypothetically arising from resensitization of AChR in the biliganded, desensitized state, AID (see Fig. 7) to reach the activated state, A,R*, for the duration of the burst, tb. Termination of the D burst is assumed to be caused by a rapid desensitization, whose rate is measured by the distribution of interburst intervals, ti. D-bursts, in turn, are grouped into clusters, whose time distribution yields information on a second, slower desensitization process (see text and Sakmann et al., 1980).

phenomenon manifests itself in “macroscopic” recordings. They first observed that when the extrajunctional area of frog denervated muscle was exposed to high agonist concentration in the patch pipette an initial hyperactivity of open channels could be recorded (up to four current levels were resolved, as shown in Fig. 11). The activity faded within seconds, as the desensitization onset progressed. Subsequently, bursts of activity reappeared at irregular intervals (Fig. 11). Elementary currents within these bursts appear to have the same characteristics as those recorded under nondesensitizing agonist concentrations, as if the openclose transition were independent of the desensitization phenomenon (Fig. 9). This bears a relationship to hypotheses linking desensitization with channel blockage by the agonist (e.g., Adams, 1975a). Sakmann et al. (1980) specifically rejected a channel blocking mechanism, and analyzed the duration of bursts (which we have defined above as D-bursts) and of the intervals between bursts of single current traces. They also measured the duration of the supraorganization of D-bursts, the clusters, and the mean duration of the intercluster intervals (ci). The reciprocal of the mean durations of D-bursts [(tb)-l]and clusters [(ti)-’]increase as a function of agonist concentration, that is, they become shorter as the rate of desensitization augments. Assuming that the sequence of D-bursts and D-burst-intervals represents the conversion to and from a rapidly desensitizing state, rates of 2

326

F. J . BARRANTES

and 5 sec-’ were calculated, respectively, from data obtained at 20 p M ACh. At the same concentration, the ci and c, values were used to derive rate constants leading to and from a second, slowly attainable desensitized state (0.2 and (0.03 sec-I). The latter figures are strikingly similar to those found by Bonner et nl. (1976), Barrantes (1 978), Heidmann and Changeux (1978, 1979), and Quast ~f nl. (1978) in rapid kinetic studies of agonist-induced state transitions measured by fluorescence spectroscopy. It should be stressed that the interpretation of burst-cluster phenomena in terms of the desensitization process(es) is the most plausible and interesting hypothesis, but data covering more than one agonist concentration are needed to make the generalization valid. 8. The B-Butst Silent (nonconducting) states other than the resting (R) or desensitized state (D) of the AChR can be defined when attempting to interpret the action of some local anesthetic drugs on channel kinetics. T h e most straightforward hypothesis (Adams, 1975c, 1976, 1977) is that in addition to the open-close transitions, occlusion (plugging) of the open channel occurs in the presence of barbiturates, procaine, lidocaine derivatives, or quinacrine. This hypothesis is not universally accepted (Katz and Miledi, 1980) but it still offers the simplest explanation of most, if not all, the experimentally observed alterations of channel kinetics, consisting of the appearance of an extra relaxation time constant in noise or voltage jump experiments (Steinbach, 1968a,b; Kordai, 1970) together with a modification of the preexisting time constant. T h e effects of the lidocaine derivative QX-222 have been studied with the patch-clamp technique (Neher and Steinbach, 1878), and a channel-plugging mechanism also emerged as the most plausible one. In the presence of QX-222 channel activity is abnormal; B-bursts characterize the pathological blocked state. A sequential scheme of the type

QX-222

A I

1

PL

fL

ff

b

C-0-B

was postulated by Neher and Steinbach (1978) to describe the blocking reaction. T h e open channel state 0 has two possible routes to decay; a is the “normal” route, [see Eq. (4)], in which the AChR reverses to the closed conformation C. The additional route consists of the plugging of the open channnel by the blocker, leading to the B state via f. This

DEVELOPMENTS IN STRUCTURE AND FUNCTION

32’1

forward rate for blocking is proportional to blocker concentration, [B]. Applying Eq. (5) yields

t, = (a +f[BI)

(14)

where i0 is the (apparent) mean dwell time of the open-channel state. Its reciprocal is linearly proportional to [B]. Two closed intervals are also observed. One corresponds to the “true” closed dwell time (the time spent in the C state by a normal channel), and the other, which is much faster (for QX-222), is the blocked interval. Fast voltage-dependent channel blockers like QX-222 or procaine produce biphasic MEPC decays, and voltage-jump relaxation or noise analysis equally show altered, multiphasic kinetics. Substances like quinacrine produce slowly reversible block of the AChR channel (Adams and Feltz, 1977, 1980a,b; Tsai et ul., 1979), similar to that seen with barbiturates (Adams, 1976). Slowly reversible blockers give rise to bursts that do not resemble those observed with QX-222; the kinetics within the B-burst are apparently normal, and the interburst interval is l/Nb seconds long (N being the number of channels in the patch). The different blocking modalities reflect underlying differences in ufinities of the substances for the channel. Thus, the mean gap duration in a B-burst is given by

ig = (([Blf)/a}b-’

= ([B]/K,)

(Y-’

(15)

where K, is the equilibrium dissociation constant blf of the blocker for the open-channel state. Furthermore, the mean number of openings per B-burst is

(1

+ ([Blf/a)){a + iBlf1-l

(16)

Combining Eqs. (15) and (16) yields the mean B-burst duration in the case of a blocker showing resolvable gaps:

I n all above cases Eq. (13) is applied and [B] is the blocker concentration. An interesting offspring of the more extensive work on local anesthetic blockage of AChR channelsis the series of studies on the effect of permeant ions in channel kinetics. The basic issue of these studies concerns the possibility that the same ions that permeate the channel are capable of impeding channel closure upon binding to sites inside the

328

F. J . BARRANTES

channel (Van Helden el al., 1977; Ascher et al., 1978; Gage and Van Helden, 1979; Marchais and Marty, 1979; Adamset al., 1981).T h e problem is similar to that of channel blockage by local anesthetics in that the a h i t y of the ion determines the dwell time on its "binding" site. One outcome of this spreading hypothesis is that channel gating is not independent of the nature of the permeant cation (Adams et al., 1981). 9. Chanwl Behnziior as a Diagnostic Tool of Reaction hiiechanisms It has been stressed throughout that the recently observed gating phenomena call for a redefinition of some views of AChR channel kinetics. Already the qualitative observation of some of the AChR gating modes in patch-clamp records makes the need for a reappraisal evident. T h e observation of N-bursts in the presence of low agonist concentrations, for instance, indicates that at least three AChR states are involved in the gating reaction. The two-state mechanism is automatically excluded, though the number of states indicated by the N-burst is not dependent on a particular model. When association of bursts is apparent, then at least four states can be postulated. More generally, the number of open states must be as large as the number of states observed. Since the dwell time of a given state is the reciprocal of the sum of rate constants leading away from that state, if only one open state is observed in a control experiment and addition of a drug decreases the lifetime of this state, it is possible to infer that the drug in question depopulates that state (e.g., Neher and Steinbach, 1978). On the other hand, if the openstate lifetime is found to be independent of drug concentration, then channel closing cannot be associated with binding of that ligand. This was the case, for instance, with the observations of Sakmann et al. (1980) on the relative insensitivity of i, on agonist concentration, leading to the postulation that the D-bursts were not the result of channel blockage but of desensitization, a phenomenon attributed to isomm'zntions of the AChR and not to binding steps (see Fig. 7). Exploring the action of blocking substances, further diagnostic elements concerning the affinity of the drug for the AChR channel can be deduced from inspection of patch-clamp recordings. Thus 1 . Drugs having relatively high affinities result in long-lived blocked states [B in Eq. (13)Jrelative to the open state 0: Channel currents will display faster kinetics, but the channel conductance remains unaltered. 2. If the blocked state is in a time range comparable to that of the open channel lifetime, B-bursts result. Channel conductance is not affected. Affinities of the blockers are not particularly high in this case. 3. Low affinity blocking drugs yield brief blocked states, manifested

DEVELOPMENTS I N STRUCTURE AND FUNCTION

329

in a longer apparent channel lifetime in patch-clamp records with a smaller conductance than corresponding controls; the apparent conductance is a composite of the blocked and unblocked open states. 4. Permeating ions are a category apart; they constitute ultrafast blocking ligands, the dwell times of the blocked state being in the lower microsecond time domain; hence they will pass undetected in (present) patch-clamp records.

VII. Summary and Perspectives

A. THECOUPLING BETWEEN ELECTROPHYSIOLOGICAL A N D BIOCHEMICAL TECHNIQUES The description of the AChR gating function is no longer restricted to electrophysiological measurements, but has recently entered a new phase by the successful reconstitution of the channel in planar bilayers (Schindler and Quast, 1980; Nelson et al., 1980; Boheim et al., 1981). T h e reader is referred to the work of Montal et al. (198 l), Anholt (198l), and Briley and Changeux (1978) for reviews on reconstitution of the AChR. T h e merit of the above preliminary phenomenological descriptions of channel gating properties in reconstituted systems should be judged in the light of the unavailability of such information in the living electrolyte. It appears that the Torpedo AChR channel is voltageinsensitive and that it responds to cholinergic agonists in a manner similar to the neuromuscular junction. The agonist concentrations eliciting responses are extremely low, and probably different experimental conditions will be called for in order to characterize fully the dose-response curve. T h e strategy of Schindler and Quast (1980) offers the possibility of correlating channel behavior with physical properties amenable to characterization in the intermediate, monolayer state such as cohesive energy of the planar bilayer (“surface pressure”). Thus, the physical state of the final bilayer can be adjusted to that of the monolayer and ultimately to that of the parental AChR vesicle, thus matching the thermodynamic state of the host membrane. The method introduced by Schindler, from which all others have derived, also offers the unique advantage of being able to control the amount of receptor (at the monolayer stage) which will finally be incorporated in the planar bilayer. I n Nelson et al. (1980) the possibility of combining flux measurements in reconstituted vesicles and electrical properties in the bilayer is suggested. This complementation might alleviate the limitations of the integrated flux measurements, especially in relation to the time resolution problems

330

F. J. BARRANTES

of the latter. T h e work of Boheim et 01. (1981) constitutes the first attempt to study the AChR gating behavior in a defined lipid environment. A pure synthetic lipid, having well-defined chemical and physical properties was employed. Furthermore, AChR preparations of increasing degree of purity were compared, monomers and dimers were found not to differ in gating properties, and the lack of influence of the nonreceptor v proteins (see above) on channel activity was shown. The study of Boheim ef d.(198 1) also provided the first comparison of gating modalities of muscle and electric organ AChR channels. Channel behavior in planar bilayers resembled in some instances the burst and cluster phenomena observed with the patch-clamp technique. These strategies define the trends that might prove more fruitful in the future for investigating the contribution of the AChR polypeptide components, nonreceptor peptides, lipid classes, and environmental factors to AChR channel properties. Given the remarkable technical advantages of patch-clamp recordings, making now possible the excision of small membrane fragments from the intact cell, the control of the sidedness of this membrane patch, and the ability to rapidly modify the environment of the membrane by superfusion of any of its t w o faces (Hamill rt a/., 1981), the major contribution of bilayer measurements should reside in their successful combination with adequate chemical dissection of AChR building blocks and other individual molecular constituents of the postsynaptic membrane. Furthermore, it could well be that in some instances planar bilayer experiments offer a more apt framework than the corresponding electrophysiological measurements for the formulation of certain problems, given the greater ability to control a complex set of multivariate parameters and detailed knowledge of geometrical constraints, diffusion barriers, exact composition of reaction partners, and state of aggregation of the AChR, etc, under defined in ziitr-o conditions. Knowledge of these parameters in the membrane area under patchclamp or even in the excised membrane patch is still difficult to obtain. In attempting to resolve some of these problems, the art of reconstitution may be on its way to achieving the restitutio ad integrum of the AChR system.

B. STRUCTURAL COUSTERPARTS OF GATING A N D STATEOF LIGATION The supramolecular arrangement of the AChR in the synapse, a subject of theoretical considerations in the past (see Changeux et al., 1967a,b; Changeux and Podleski, 1968; Levitzki, 1974), makes it unlikely that the activation of any one AChR could pass undetected or have

DEVELOPMENTS IN STRUCTURE AND FUNCTION

33 1

no consequences on the adjacent units. A major challenge of future studies (the third stage in the evolution of patch-clamp) will be to attempt the simultaneous description of the individual channel behavior, identifying microscopic states and the kinetics of their transitions, and the population behavior resulting from the recruitment of many such individual channels upon agonist action. An important link between structure and function would thus be established. The observation of discrete conductance states of the AChR other than the main open state (the “sublevels,” Fig. 10) raises hopes of being able to identify structural forms associated with a given kinetic state. Aside from the possibility of different subunit arrangements to account for the occurrence of conductance sublevels (Hamill and Sakmann, 198l), different oligomm’c states of the AChR might underlie this phenomenon. This is the type of hypothesis which may be more easily tested in the planar lipid bilayers. The conventional flux experiments do not appear adequate for this purpose. Whereas the prospective line of research considered above puts weight on the structural counterparts .f gating, another important avenue for the future concerns the correlation between the latter function and the state of ligation of the AChR. The possibility that the fast opening events ( T ~ 150 psec) detected in addition to the main longer openings ( T ~ 10 msec) correspond to the monoliganded AR* state (see Fig. 7) has been considered (Colquhoun and Sakmann, 1981). Although almost negligible in probabilistic terms, one could hope that all conducting receptor species R* will become apparent and adequately characterized under appropriate conditions (e.g., driving force leading away from equilibrium in perturbation experiments). Finally, the heterogeneity of gap lifetimes within an N-burst (Colquhoun and Sakmann, 1981) also merits further investigation, since characterization of these short-lived events might lead to identification of the otherwise silent, closed AChR states that are normally the daily bread of the biochemist.

-

-

References

Adams, J., Nonner, W., Dwyer, T. M., and Hille, B. (1981).J. Gen. Physiol. 78,593-615. Adams, P. R. (1975a).PJluegers Arch. 360, 135-144. Adams, P. R. (1975b).J. Physiol. (London) 246, 61P-63P. Adams, P. R. (1976).J . Physiol. (London) 260, 531-552. Adams, P. R. (1977).J.Physiol. ( L a d o n ) 268, 291-318. Adams, P. R. (1981).J . Memh. Biol. 58, 161-174. Adams, P. R., and Feltz, A. (1977).Nature (London) 269, 609-611.

332

F. J. BARRANTES

Adarns, P. R., and Feltz, A. (1980a).J. Physiol. (London) 306, 261-281. Adarns, P. R., and Feltz, A. (1980b).J. Physiol. (London) 306, 283-306. Adarns, P. R., and Sakmann, B. (1978). Prcz. Natl. Acad. Sci. U.S.A. 75, 2994-2998. Alrnon, R. R., and Appel, S. H. (1975). Biochim. Biophys. Acta 393, 66-77. Anderson, D. J., and Blobel, G. (1981). Pi-or. Satl. Acad. Sci. U.S.A. 78, 5598-5602. Andreasen, T.J., and McNamee, M. G. (1977). BiorhPm. Biophys. Rex Commun.79,958-965. Anholt, R. (1981). Trend.$Biorhem. Sri. 6, 288-291. Anholt, R., Lindstrom, J., and Montal, M. (1981).J. Rial. Chem. 256, 4377-4387. Anwyl, R., and Narahashi, T. (1979). Br. J. Pharmarol. 65, 483-488. Aoshirna, H., Cash, D. J., and Hess, G. P. (1980). Biochem. Biothys. Res. Commun. 92, 896904. Aoshima, H., Cash, D. J., and Hess, G. P. (1981). Biochmistn; 20, 3467-3474. Asher, P., Martp, A., and Neidle, T. 0. (1978).J. Physiol. (London) 278, 177-206. Axelrod, D., Ravdin, P., Koppel, D. E., Schlessinger, J.. Webb, W. W., Elson, E. L., and Podleski, T. R. (1976). Proc. Null. Arad. So‘. U.S.A. 73,4594-4598. Axelrod, D., Ravdin, P. M., and Podleski, T. R. (1978a). Eiochim. Biophys. Acta 511, 23-38. Axelrod, D., Wright, A., Webb, W., and Horitz, A. (1978b). Biochemist? 17, 3604-3609. Axelson, J., and Thesleff, S. (1959).J. Phyiol. (Londott) 147, 178-193. Ballivet, M., Patrick, J., Lee, J . . and Heinernan, S. (1982). Pro(. N u t / . Acnd. Sci. U.S.A. 79, 4466. Barnard, E. A., Coates, V., Dolly, J. 0..and Mallick, B. (1977).Cell Biol. In/.Rep. I, 99-106. Barrantes. F. J. (1976). Eiochem. Eiophy. Res. Commun. 72, 479-488. Barrantes, F. J. (1978).J. ,210l. Biol. 124, 1-26. Barrantes, F. J . (1979).Annu. R m . Biophy. Bioeng. 8, 287-321. Barrantes, F. J. (1980). Biochmistry 19, 2957-2965. Barrantes, F. J. (1982a).J. Cell Biol. 92, 60-68. Barrantes, F. J . (1982b). In “Neuroreceptors” (F. Hucho, ed.), pp. 315-328. de Gruyter, BerlidNew York. Barrantes, F. J., Neugebauer, D.-C., and Zingsheim, H. P. (1980). FEES Let!. 112, 73-78. Bartfeld, D., and Fuchs, S. (1979). Biochem. Biophjs. Res. Commun. 89, 513-519. Bartholdi, M., Barrantes, F. J., and Jovin, T. M. (1981). Eur. J. Eiochem. 120, 389-397. Bercovici, T., and Gitler, C. (1978). Bzochmkt? 17, 1484- 1489. Bernhardt, J., and Neumann, E. (1978). Pror. Natl. Acad. Sci. U.S.A. 75, 3756-3760. Bernhardt, J., and Neumann, E. (1980a). Nncruchem. lnt. 2, 243-250. Bernhardt, J., and Neurnann, E. (1980b).J. Thmr. B i d . 86, 649-661. Bernhardt, J., Moss, K. M., Luckinger, R. M., and Neumann, E. (1981). FEES Lett. 134, 245-248. Betz. H., and Changeux, J.-P. (1979). il’ature (Lurduu) 278, 749-452. Biesecker, G. (1973). Biochemist? 12, 4403-4409. Blanchard, S. G., and Raftery, M. A. (1979). Proc. Nad. tfcad. Sci. C . S . A . 76, 81-85. Blanchard, S. G., Quast, U., Reed, K., Lee, T., Schimerlik, M. I., Vandlen, R., Claudio, T., Strader, C., Moore, H. P. H., and Raftery, M. A. (1979). Biochemist? 18, 1875-1883. Bloch. R. J., and Geiger, B. (1980). Cell 21, 25-35. Boheirn, G., Hanke, W., Barrantes, F. J., Eibl, H., Sakmann, B., Fels, G., and Maelicke, A. (1981). Proc. ,Vatl. Arad. Sci. U.S.A. 78, 3586-3590. Bonner, R., Barrantes, F. J., and Jovin, T. M. (1976). Nature ( L a d o u ) 263, 429-431. Bormann, J., Hamill, 0. P., and Sakmann, B. (1981). Pfluggers Arch. 391, R29. Boulter, J., and Patrick, J. (1977). Eiochmis/? 16, 4900-4908. Boyd, N. D., and Cohen, J. B. (1980). Biochemist? 19, 5353-5358. Bradley, R. J., Howell, J. H., Romine, W. O., Carl, G. F., and Kemp, G. E. (1976). Biorhem. Biophys. Res. Cmnmun. 68, 577-584.

DEVELOPMENTS IN STRUCTURE AND FUNCTION

333

Briley, M., and Changeux, J. -P. (1977). Int. Rev. Neurobiol. 20, 31-63. Brisson, A. (1980). These Doctorat d’Etat, Grenoble, France. Brockes, J. P., and Hall, Z. W. (1975a). Biochemistry 14, 2092-2100. Brockes, J. P., and Hall, Z. W. (1975b).Biochemistry 14, 2101-2106. Brockes, J. P., and Hall, Z. W. (1975~ ). Proc. Natl. Acad. Sci. U.S.A. 72, 1368-1372. Bulger, J. E., and Hess, G. P. (1973). Biochem. Biqbhys. Res. C m m u n . 54, 677-684. Cartaud, J., Benedetti, L., Cohen, J. B., Meunier, J. C., and Changeux, J.-P. (1973).FEBS Lett. 33, 109-113. Cartaud, J., Benedetti, L., Sobei, A., and Changeux, J.-P. (1978).J . Cell Sci. 29, 313-337. Cartaud, J., Popot, J.-L., and Changeux, J.-P. (1980). FEBS Lett. 121, 327-332. Cartaud, J., Sobel, A., Rousselet, A., Devaux, P. F., and Changeux, J.-P. (1981).J. Cell B i d . 90,418-426. Cash, D. H., and Hess, G. P. (1980).Proc. Natl. Acad. Sci. U.S.A. 77, 842-846. Cash, D. J., Aoshima, H., and Hess, G. P. (1980).Biochem. Biophys. Res. Commun. 95, 10101016. Catterall, W. A. (1975).J. B i d . Chem. 250, 1776-1781. Catterall, W. A,, and Niremberg, M. A. (1973).Proc. Natl. Acad. Sci. U.S.A. 70, 3759-3763. Chang, H. W. (1974).Proc. Natl. Acad. Sci. U.S.A. 71, 2113-2117. Chang, H. W., and Bock, E. (1977). Biochemistry 16,4513-4520. Chang, H. W., and Bock, E. (1979). Biochemistry 18, 172-179. Chang, H. W., and Neumann, E. (1976). Proc. Natl. Acad. Sci. U.S.A. 73, 3364-3368. Chang, R. S. L., Potter, L. T., and Smith, D. S. (1977). Tissue Cell 9, 623-644. Changeux, J.-P. (1981). Haruey Lect. 75, 85-254. Changeux, J.-P., and Podleski, T. R. (1968). Proc. Natl. Acad. Sci. U.S.A. 59, 944-950. Changeux,J.-P., Thiery, J., Tung, Y., and Kittel, C. (1967a).Proc.Natl. Acad. Sci. U.S.A. 57, 335-341. Changeux, J.-P., Podleski, T. R., and Wofsy, L. (1967b). Proc. Natl. Acad. Sci. U.S.A. 58, 2063-2070. Changeux, J.-P., Benedetti, L., Bourgeois, J. P., Brisson, A., Cartaud, J., Devaux, P., Griinhagen, H. H., Moreau, M., Popot, J. L., Sobel, A., and Weber, M. (1976). Cold Spring Harbor Symp. Quant. Biol. 40, 203-210. Cherry, R. J., Miiller, U., Holenstein, C., and Heyn, M. P. (1980).Biochim. Biophys. Acta 596, 145- 151. Chicheportiche, R., Vincent, J.-P., Kopeyan, C., Schweitz, H., and Lazdunski, M. (1975). Biochemistry 14, 208 1-2091. Chou, P. Y., and Fasman, G. D. (1974). Biochemistry 18, 172-179. Chou, P. Y., and Fasman, G. D. (1978). Annu. Rev. Biochem. 47, 251-276. Claudio, T., and Raftery, M. A. (1977).Arch. Biochem. Biqbhys. 181, 484-489. Cohen, J. B., and Boyd, N. D. (1979). In “Catalysis in Chemistry and Biochemistry” (B. Pullman, ed.), pp. 255-279. Reidel Publ., Dordrecht, Netherlands. Cohen, J. B., Weber, M., and Changeux, J.-P. (1974). Mol. Pharmucol. 12, 519-535. Colquhoun, D. (1979).I n “The Receptors: A Comprehensive Treatise” (D. O’Brien, ed.), pp. 93- 142. Plenum, New York/London. Colquhoun, D., and Hawkes, A. G. (1977).Proc.R. Soc. London, Ser. B 199, 231-262. Colquhoun, D., and Hawkes, A. G. (1981).Proc. R. Soc. London, Ser. B 211,205-235. Colquhoun, D., and Rang, H. P. (1976). Mol. Pharmacol. 12, 519-535. Colquhoun, D., and Sakmann, B. (1981). Nature (London) 294,464-466. Colquhoun, D., Neher, E., Reuter, H., and Stevens, C. F. (1981). Nature (London) 294, 752-754. Conti-Tronconi, B., Tzartos, S., and Lindstrom, J. (1981). Biochmistry 20, 2181-2191. Conti-Tronconi, B., Dunn, S. M. J., and Raftery, M. A. (1982). Biochemistry 21, 893-899.

334

F . J. BAKKANTES

Craig, S. W., and Pollard, T. D. (1982). TJ-oI~,\ Biorhrzn. Sri. 7, 88-92. Criado, M.. and Barrantes, F. J. (1982). 2Ymrorhcrri.I f t t . 4, 289-3112, Criado, M., Vaz, W.. Barrantes, F. J., and Jovin, T. M. (1982). Biorhumktj?; (in press). Cull-Candy, S. G., and Parker, I. (1982).Suture (Lomfon) 295, 410-412. Damle, V. N . , and Karlin, A. (1978). Biorhcnhtq 17, 2039-2045. Damle, V. N.,Hamilton, S., Valderrama, R., and Karlin, A. (1976).Phnrmnrologst 18, 146. Delegeane, A . M., and McNamee, M. G. (1980).Biochelbtry 19, 890-895. Devilliers-Thiery, A., Changeux, J.-P., Parotaud, P., and Strosberg, A. D. (1979).FEBS Lett. 104,99-105. Devreotes. P. N.,Gardner, J. M., and Fambrough, D. M. (1977). Cell 10, 365-373. Dolly. J . O., and Barnard, E. (1975).FEBS Lett. 57, 267-271. Dolly, J. O., and Barnard, E. (1977).Hiorhmistq 16, 5053-5060. Doster, W., Hess, B., Watters, D., and Maelicke, A. (1980). FEBS Lrtt. 113, 312-314. Dreyer, F.,Walther, C., and Peper, K. (1976).Ppueg..v Arch. 366, 1-9. Edelstein, S., Beyer, W. B., Eldefrawi, A . T., and Eldefrawi, M. E. (1975).J. Hiol. Chum. 250, 61 0 1-6106. Ehrenstein, G.. Blumenthal, K.. latorre, R., and Leccar, H. (1974).J . Gen. Phyiol. 63, 707-721. Eldefrawi, A. T., Eldefrawi, M. E., Albuquerque. E. X., Oliveira, A. C., Mansour, N., Adler, hi., Daly, J. LV., Brown, G. B., Burgermeister, W. B., and Witkop, B. (1977). Prrir. .Yotl. .4md. Sri. l ’ . S . , 4 . 74, 2172-2176. Eldefrawi. M. E.. and Eldefrawi, A. T. (1973a)..4rrh. Biurhum. Biriphys. 159, 362-373. Eldefrawi, M. E.. and Eldefrawi. A. T. (1973b).Biochrm. Phur-mnrol. 22, 3145-3150. Eldefrawi, M. E., and Eldefrawi, A. T. (1977). Rucpt. Rwognition Ser. A 4, 73-84. Eldefrdwi, M . E., Eldefrawi, A. T., Gilmour, L. P., and OBrien, R. D. (1971a). iZfol. Pho,mticol. 7, 420-428. Eldefrawi, M. E., Eldefrawi, A. T.. and O’Brien, R. D. (1971b).Pror. Nutl. Acad. Sci. 1I.S.A. 68, 1047-1050. Eldefrawi, M. E., Eldefrawi, A. T., and Shamoo, A. E. (1975a). .4nn. N.1: A d . Sri. 264, 183-202. Eldefrawi, M. E., Eldefrawi, A. T., and Wilson, D. B. (1975b). Eiorhrrnirtry 14, 4304-4310. Elliot, J., Blanchard, S. G . . Wu, W., Miller, J., Strader, C. D., Hartig, P., Moore, H.-P., Racs, J., and Raftery. M. A. (1980). Biochein. J . 185, 667-677. Fambrough, D. M. (1979). Phviol. Rev. 59, 165-227. Fambrough, D. M., and Devreotes, P. M. (1978).J. C d l Biol. 76, 237-244. Fenwick. E. M.. Marty, -4., and Neher, E. (1982).J. P/ty.riol. (London) 331, 599-635. Flanagan, S. D., Barondes, S. H., and Taylor, P. (1976).J. Hiol. Chem. 251, 858-865. Franklin. G . I . , and Potter, L. T. (1972). FEBS Lett. 28, 101- 106. Froehner, S. C. (1981).Biochumist?;~20, 4905-4915. Froehner. S. C., and Rafto, S. (1979). B i o c h u m i q 18, 301-307. Froehner, S. C., Karlin, A , , and Hall, A. W. (1977a). Pwr.. AYutl. Acucl. Sci. 1T.S.A. 74, 4685-4688. Froehner, S. C., Reiness, C. G., and Hall, 2. W. (1977b).J. B i d . Chem. 252, 8589-8596. Froehner, S. C., Buldbrandsen, V., Hyman, C., Jeng, A. Y., Neubig, R. R., and Cohen, J. B. (1981). Pror. A’uti. Arud. Sci. 1 ’ S . A . 78, 5230-5234. Fu, J. L., DOnner, D. B., and Hess, G. P. (1974). Eiorhem. Biophys. Res. Commun. 60, 1072-1080. Fuchs, S. (1979). Cum. Top. .lficrobiol. I m r i i u n d . 85, 1-29. Fukushima, Y. (1981). A’uture (London) 294, 368-371. Gage, P. W. (1976). P/iy.siol. Rnt. 56, 177-247.

DEVELOPMENTS IN STRUCTURE AND FUNCTION

335

Gage, P. W., and Harnill, 0. P. (1980).J . Physiol. (Londcm) 298, 525-538. Gage, P. W., and Van Helden, F. (1979).J. Physiol. (London) 288, 509-528. Gibson, R. E. (1976). Biochemistry 15, 3890-3901. Gibson, R. E., OBrien, R., Edelstein, S. J., and Thompson, W. R. (1976). Biochaistry 15, 2377-2383. Gomez, C., Richman, D., Berman, P., Burres, S., Aranson, B., and Fitch, F. (1979).B i o c h h . Biophys. Res. Commun. 88, 575-582. Gonzalez-Ros, J. M., Paraschos, A., Farach, M. C., and Martinez-Carrion, M. (1981). Bivchim. Biophys. Acta 643, 407-420. Gordon, A. S., Bandini, G., and Hucho, R. (1974). FEBS Lett. 47, 204-208. Gordon, A. S., Davis, C. G., and Diamond, I. (1977). Proc. Nutl. Acad. Sci. U.S.A. 74, 263-267. Gordon, A. S., Davis, C. G., Milfay, D., Kaur, J., and Diamond, I. (1980). Biochim. Biophys. Actu 600, 421-431. Gotti, C., Conti-Tronconi, B. M., and Raftery, M. A. (1982). Biochemistry 21, 3148-3154. Griinhagen, H.-H., and Changeux, J.-P. (1976).J. Mol. B i d . 106, 497-516. Gulley, R. L., and Reese, T. S. (1981).J. Cell Biol. 91, 298-302. Gullick, W. J., Tzartos, S., and Lindstrom, J. (1981). Biochemistry 20, 2173-2180. Gurdon, J. B. (1974). “The Control of Gene Expression in Animal Development.” Oxford Univ. Press (Clarendon), LondodNew York. Guy, H. R. (1981). Cell. Mol. Neurobiol. 1, 231-258. Gysin, R., Wirth, M., and Flanagan, S. D. (1981).J. B i d . Chem. 256, 11373- 11376. Hall, Z. W., Lubit, B. W., and Schwartz, J. H. (1981).J. Cell B i d . 90, 789-792. Harnill, 0. P. (1981).J. Physiol. (London) 319, 97-98P. Hamill, 0. P. (1982).Zn “Neuroreceptors” (F. Hucho, ed.), pp, 233-242. de Gruyter, Berlin New York. Hamill, O., and Sakmann, B. (1981). Nature (London) 294, 462-464. Hamill, O., Marty, A., Neher, E., Sakmann, B., and Sigworth, F. J. (1981). PJluegers Arch. 391, 85-100. Hamilton, S. L., McLaughlin, M., and Karlin, A. (1977).Biochem. Bivphys. Res. Commun. 79, 692-699. Hamilton, S. L., McLaughlin, M., and Karlin, A. (1979). Biochemistry 18, 155-163. Hartig, P. R., and Raftery, M. A. (1977). Biochtm. Bivphys. Res. Commun. 78, 16-22. Hazelbauer, J., and Changeux, J.-P. (1974). Proc. Nutl. Acud. Sci. U.S.A. 71, 1479-1483. Heidmann, T., and Changeux, J.-P. (1978). Annu. Rev. Biochem. 47, 371-441. Heidmann, T., and Changeux, J-P. (1979). Eur. J . Biochem. 94, 255-279. Heilbronn, E., and Mattson, C. (1974).J. Neurochem. 22, 315-317. Hess, G. P., and Andrews, J. P. (1977). Proc. Nutl. Acud. Sci. U.S.A. 74, 482-486. Hess, G. P., Andrews, J. P., Struve, G. E., and Coornbs, S. E. (1975). Proc. Nutl. Acad. Sci. U.S.A. 72,4371-4375. Hess, G. P., Lipkowitz, S., and Struve, G. E. (1978). Proc. Natl. Acad. Sci. U.S.A. 75, 1703- 1707. Hess, G. P., Cash, D. J., and Aoshima, H. (1979). Nature ( L m d o n ) 282, 329-331. Hess, G. P., Aoshima, H., Cash, D. J., and Lenchitz, B. (1981).Proc.Nutl. Acud. Sci. U.S.A. 78, 1361-1365. Hess, G . P., Pasquale, E. B., Walker, J. W., and McNarnee, M. G. (1982). Proc. Nutl. Acud. Sci. U.S.A. 79, 963-967. Heuser, J. E., and Salpeter, S. R. (1979).J. Cell B i d . 82, 150-173. Holtzrnan, E., Wise, D., Wall, J., and Karlin, A. (1982). Prvc. Nutl. Acad. Sci. U.S.A. 79, 310-314.

336

F. J. BARRANTES

Horne, R., and Patlak, J. (1980). Pror. S a t l . Arad. Sci. U.S.A. 77, 6930-6934. Huang, L. (1979). FEBS Lett. 102,9-12. Hucho, F. (1979). FEBS Le/t. 103,27-32. Hucho, F., and Changeux, J.-P. (1973). FEBS Let/. 38, 11-15. Hucho, F., Layer, P., Kiefer, H . R., and Bandini, G. (1976). Proc. Nntl. Acnd. Sci. U.S.A. 73, 2624-2628. Hunkapiller, M. W., Strader, C. D., Hood, L., and Raftery, M. A. (1979).Biochon. Riophys. Krs. Conitnuti. 91, 164-169. Ishikawa, Y., Yoshida, H., and Tamiya, N. (198O).J. Biochm. (Tokyo) 87, 313-321. Jeng, A. Y., St. John, P. A,, and Cohen, J. B. (1981). Biorhzm. Biophys. Acta 646,411-421. Johansson, G., Gsyn, R., and Flanagan, S. D. (1981).J. Biol. C h m . 256, 9126-9135. l Rat. 6, 191-260. Karlin, A. (1980). C ~ l SUI$ Karlin, A,, and Cowburn. D. (1973). P ~ o cSntl. . Acad. Sri. U.S.A. 70, 3636-3640. Karlin, A., Weill, C., NcNamee, M., and Valderrama, R. (1976). &id Spring Harbor Symp. Quant. Biol. 40, 203-210. Karlin, A,, Holtzman, E., Valderrama, R., Damle, V., Hsu, K., and Reyes, F. E. (1978).J. Cdi Biol. 76, 577-592. Karlin, A . , Damle, V., Hamilton, S., McLaughlin, M., Valderrama, R., and Wise, D. (1979). .4d71.(:ytr$hnrinnrol. 3, 183- 188. Kasai, M., and Changeux, J.-P. (1971a).J. Jfenibr. B i d . 6, 1-23. Kasai, M., and Changeux, J.-P. (1971b).J. d l m b r . B i d . 6, 58-80. Katz, B., and Miledi, R. (1972).J. Phyiol. (London) 224, 665-699. Katz, B., and Miledi, R. (1980).I n “Ontogenesis and Functional Mechanisms of Peripheral Synapses. INSERM Symp. 13,” (J.Taxi, ed.), pp. 171-178. Elsevier, AmsterdadNew York. Katz, B., and Thesleff, S. (1957).J. Phyiol. (Loitdoti) 138, 63-80. Katz, F. M., Rothman, J. E., Lingappa, V. R., Blobel, G., and Lodish, H. F. (1977). Proc. S a t l . clrad. Sci. 1’.S..4. 74, 3278-3282. Kim, P. S., and Hess, G. P. (1981).J. .\fmibr. Bioi. 58, 203-21 1. Kistler, J., and Stroud, R. M. (1981). Proc-. Sn/l. Acnd. Sri. 1 ’ S . A . 78, 3678-3682. Kistler, J., Stroud, R. M., Klymkowsky, M.W., Lalancette, R., and Fairclough, R. H. (1982). Bioplin. J. 37, 371-383. Klett, R. P., Fulpius, B. W., Cooper, D., Smith, M., Reich, E., and Posani, L. D. (1973).J. Hiol. Chrm. 248, 6841-6853. Klymkowsky, M. W., and Stroud, R. M. (1979).J. .\foi. B i d . 128, 319-334. Klymkowsky, M. W., Heuser. J, E., and Stroud, R. M. (1980).J. Crll Biol. 85, 823-838. Kordai, M. (1970).j. Phjsiol. (London) 209, 689-699. Lands, B. R., Salpeler. E. E., and Salpeter, M. M. (1981). Proc. iVnt1. A m d . Sri. 11.S.A. 78, 7200-7204. Lester, H. S. (1982). Sntio-e (Lundon) 294, 398-399. Levitzki, A. (1974). J. Theor. B i d . 44, 367-372. Lim, V. I . (1974).J. .\fol. B i d . 88, 873-894. Lindstrom, .J. (1979). ,4drl. Imniirnol. 27, 1-50. Lindstrom, J.. and Dau, P. (1980). ,4nnfr. R m . Phrirmnrol. Tosirol. 20, 337-362. Lindstrom, J., and Patrick, J. ( 1974). I n “Synaptic Transmission and Nerve Interaction” (W. L. Bennett, ed.), pp. 191-216. Raven press, New York. Lindstrom, J.. Lennon, V., Seybold, M., and Whittingham, S. (1976). Anti. N.1’. A m d . Sci. 274,254-274. Lindstrom, J., Einarson, B., and Merlie, J. (1978). Proc. S a t l . Arrid. Sci. U.S.A. 75,769-773. Lindstrom, J., Merlie, J., and Yogeeswaran, G. (1979a). B i o c h r m h t ~18, 4465-4470.

DEVELOPMENTS IN STRUCTURE AND FUNCTION

337

Lindstrom, J., Walter, B., and Einarson, B. (1979b). Biochemistry 18,4470-4480. Lindstrom, J., Anholt, R., Einarson, B., Engel, A,, Osame, M., and Montal, M. (1980a). J . B i d . Chem. 255, 8340-8350. Lindstrom, J., Cooper, J., and Tzartos, S. (1980b). Biochemktry 19, 1454-1458. Lindstrom, J., Gullick, W., Conti-Tronconi, B., and Ellisman, M. ( 1 9 8 0 ~)Biochemistry . 19, 479 1-4795. Lindstrom, J., Einarson, B., and Tzartos, S. (1981a). I n “Methods in Enzymology” (J. J. Langone and H. Van Vunakis, eds.),Vol. 74, pp. 432-460. Academic Press, New York. Lindstrom, J., Tzartos, S., and Gullick, B. (1981b). Ann. N.Y. A c d . Sci. 377, 1-19. Lo, M. M. S., Garland, P. B., Lamprecht, J., and Barnard, E. A. (1980). FEBS Lett. 111, 407-412. Lo, M. M. S., Barnard, E. A., and Dolly, J. 0. (1982). Biochemistry 21, 2210-2217. Lucy, J. A., and Glauert, A. M. (1974).J. Mol. B i d . 8, 727-748. Lux, H. D., and Nagy, K. (1981). Pfluegers Arch. 391,252-254. Lux, H. D., Neher, E., and Marty, A. (1981).Pfluegprs Arch. 389, 293-295. McManaman, J. L., Blosser, J. C., and Appel, S. H. (1981).J. Netrrosci. 1, 771-776. Marchais, D., and Marty, A. (1979).J. Physiol. ( L m d h ) 297, 9-45. Marsh, D., and Barrantes, F. J. (1978). Proc. Natl. Arad. Sri. U.S.A. 75,4329-4333. Martinez-Carrion, M., Sator, V., and Raftery, M. A. (1975).Biorhem. Biophys. Res. Cominun. 65, 129-137. Marty, A. (1978).J . Physiol. ( L a d a ) 278, 237-250. Marty, A. (1981).J. Physiol. ( L o n d a ) 291, 497-500. Masukawa, L. M., and Albuquerque, E. X. (1978).J. Gen. Physiol. 72, 351-367. Mattson, C., and Heilbronn, E. (1975).J. Neurochem. 25, 899-901. Mendez, B., Valenzuela, P., Martial, J. A., and Baxter, J. D. (1980). Science 209,695-697. Merlie, J. P., and Sebbane, R. (198l).J. B i d . Chem. 256, 3605-3608. Merlie, J. P., Changeux, J.-P., and Gros, F. (1978).J. Biol. Chem. 253, 2882-2891. Merlie, J. P., Hofler, J. G. C., and Sebbane, R. (1981).]. B i d . C h a . 256, 6995-6999. Merlie, J. P., Sebbane, R., Tzartos, S., and Lindstrom, J. (1982).J. Bid. Chem. 257, 2694270 1 . Meunier, J. C., and Changeux, J.-P. (1973). FEBS Lett. 32, 143- 148. Meunier, J. C., Olsen, R. W., Menez, A., Morgat, J. L., Fromageot, P., Ronseray, A. M., Boquet, P., and Changeux, J.-P. (1971). C.R. Hebd. Seances Acad. Sci., Ser. D 273, 595-598. Meunier, J. C., Olsen, R. W., and Changeux, J.-P. (1972a). FEBS Lett. 24, 63-68. Meunier, J . C., Olsen, R. W., Menez, A., Fromageot, P., Boquet, P., and Changeux, J.-P. (1972b). Biochemistry 11, 1200-1210. Meunier, J. C., Sealock, R., Olsen, R. W., and Changeux, J.-P. (1974). Eur. J . Biorhem. 45, 371-394. Michler, A., and Sakmann, B. (1980). Dev. B i d . 80, 1-17. Miledi, R. (1960).]. Physiol. (London) 154, 190-205. Miledi, R., Molinoff, P., and Potter, L. T. (1971). Nature (London) 229, 554-557. Miller, D. L., Moore, H. P., Hartig, P. R., and Raftery, M. A. (1978). Biorhem. Biophys. Res. Commurr. 85,632-640. Molinoff, P. B., and Potter, L. T. (1972). Adu. Biochem. Psychophamncol. 6, 111-134. Montal, M., Darszon, A., and Schindler, H. (1981). 4. Rev. Biophys. 14, 1-79. Moody, T., Schmidt, J., and Raftery, M. A. (1973). Biochem. Biophys. Res. Commun. 53, 761-772. Moore, H.-P.H., and Raftery, M. A. (1979). Biochemistry 18, 1862-1867. Moore, H.-P.H., and Raftery, M. A. (1980).Proc. Natl. Acad. Sci. U.S.A. 77, 4509-4513.

338

F. J. BARRANTES

Moore, H.-P.H., Hartig, P. R., and Raftery, M. A. (1979a).Proc. ,\ntl. Acnd. Sci. C'.S.A. 76, 6265-6269. Moore. H.-P.H., Hartig, P. R.. Wu, W.C. S., and Raftery, M. A. (1979b). Bioclrrm. B i o p l y . Re.\. c / J r r / t r / / L r / . 88, 735-743. Nakajinia, Y.,and Bridgman. P. C. (1981).,]. Cull B i d . 88, 453-458. Nathanson. S . M..and Hall, 2. M'. (1979). Biochrrrti.\tr:y 18, 3392-3401. Neher, E. (1982). Itr "Techniques in Cellular Physiology" (P. F. Baker, ed.). Elsevier, Amsterdam. Neher. E., and Sakmann, B. (1976a). A'n/ur-c (Lorzdorr) 260, 799-802. Neher, E. and Sakmann, B. (1976b).J. Pltjtrol. (Lotrdorr) 258, 705-729. Neher, E., and Steinbach, J. H. (1978).J. P/tjsio/. (Lortdon) 277, 153-176. Neher, E., Sakmann, B., and Stinbdch, J. H. (1978). Pjluqyr.5 Arch. 375, 219-228. Kelson. D. J., and Sachs, F. (1979). A\n/rrvc (Lortdott) 282, 861-863. Nelson, N.,Anholt, R., Lindstrom, J., and Montal, M. (1980). Proc. Nntl. A d . Sci. U . S . A . 77, 3057-3061. Neubig, R., and Cohen. J . B. (1979). Bioc/twri.sfr~18, 5464-5475. 19, 2770-2779. Neubig, R., and Cohen, J. B. (1980). Bioclwrriist~~ Neubig. R. R., Krodel, E. K., Boyd, N. D., and Cohen. J. B. (1979). Proc. "v'otl. Acnd. Sci. (..S..-1. 76, 690-694. Neugebauer. D.-C., and Zingsheim, H. P. (1982). Bioclrint. Biophjs. .4c/rr 684, 272-276. Neumann, E., and Chang, H. W. (1976). Proe. ,VntI. .4crid. Sei. l'.S..4. 73, 3994-3998. Nickel, E., and Potter, L. (1973). Br-oitz RP\. 57, 508-517. O'Brien. R. D.. and Gibson, R. E. (1975). ,4r-c/r. Bioc/rrrrr. Biop/tj,\. 169, 458-463. Ohmori, H.. Yoshida, S., and Hagiwara, S. (1981). Proc. A'ntl. .4ccrd. Sci. U.S..4. 78, 49604 964. Ong. I). E., and Brady, R. N. (1974). Bioc/trmi.\tr;y 13, 2822. Oswald. R. E., and Changeux, J.-P. (1982). FEBS Lett. 139, 225-229. Pallota, B. S.. Magleby, K. L., and Barnett, J . N. (1981). S n t i r r r (Lorrrlon) 293, 471-474. Patlak. J. B., Gration. K. A. F.. and Usherwood, P. K . R. (1979). Nntrcrr (Lonrlow) 278, 64 3 - 645. Patrick. J., and Berman, P. W. (1980). Cr// Srotj. R n f . 6, 157-190. Patrick, J., Boulter. J., and O'Brien, J . C. (1975). Bioclirtn. Bif+/rj\. RP\. ( : f J r f / m l d f l . 64, 219. Penn, A. S. , Chang. H. W., Lovelace, R. E., Niemi, W.,and Miranda, A. (1976).Arr11.N.1: z 4 c r ~ dScr. . 274, 354-376. Pezzementi, L., and Schmidt, J . (198l).J. B i d . C h m . 256, 12651-12654. Poo, M - X f . (1982). S'otitr(T (Lotrclort) 295, 332-334. Popot, J. L., Sugiyarna, H., and Changeux, J.-P. (1976).J. .\fd.Bid. 106, 469-483. Popot, J . L., Deniel, R. A., Sobel, A , , Van Deenen, L., and Changeux, J.-P, (1978).E w . J . BioOrmt. 85, 27-42. Potter, L. (1973). I u "Drug Receptors'' (H. P. Rang, ed.), pp. 295-312. Macmillan, New YorkiLondon. Prinz, H., Jiirss, R.. and Maelicke. A. (1980). AYwroc/wrrl.Irrt. 2, 251-256. Prives. J., Silnian, I., and Amsterdam, .4. (1976). Crll 7, 514-516. Prives. J., Fulton. A . B.. Penman, S., Daniels, M. P., and Christian, C. N . (1982).J. Cu/lRiol. 92,23 1-236. Quast, IJ., Schimerlik. M. I., Lee, T.. M'itzemann, V., Blanchard, S. G., and Raftery, M. A. (1978). Biorherrrktr~ 17, 2405-2414. Quast, U . , Schimerlik, M. I., and Raftery, M.A. (1979). Biochrrnistq 18, 1891-1901. Raftery, M. A., Schmidt, J . . Clark, D. G., and Wolcott, R. G. (1971). Biochum. Biophjs. RPS. Cotrrrriuri. 45, 22- 1629.

DEVELOPMENTS IN STRUCTURE AND FUNCTION

339

Raftery, M. A., Schmidt, J., and Clark, D. G. (1972). Arch. Biochem. Biophys. 152,882-886. Raftery, M. A., Vandlen, R., Michaelson, D., Bode, J., Moody, T., Chao, Y., Reed, K., Deutsch, J., and Duguid, J. (1974).J. Supramol. Struct. 2, 582-592. Raftery, M. A., Bode, J., Vandlen, R., Michaelson, D., Deutsch, J., Moody, T., Ross, M. J., and Stroud, R. M. (1975). I n “Protein-ligand Interactions” (H. Sund and G. Blauer, eds.), pp. 328-352. de Gruyter, Berlin. Raftery, M. A,, Vandlen, R. L., Reed, K. L., and Lee, T. (1976). Cold Spring Harbor Symp. Quant. Biol. 40, 193-202. Raftery, M. A., Blanchard, S. G., Elliott,J., Hartig, P., Moore, H., Quast, U., Schimerlik, M. I., Witzemann, V., and Wu, W. (1979). Adv. Cytopharmacol. 3, 159-182. Raftery, M. A., Hunkapiller, M. W., Strader, C. D., and Hood, L. E. (1980a). Science 208, 1454- 1457. Raftery, M. A., Witzemann, V., and Blanchard, S. G. (1980b). Ann. N.Y. Acad. Sci. 346, 458-474. Ramseyer, G. O., Morrison, G. H., Aoshima, H., and Hess, G. P. (1981).A m l . Biochem. 115, 34-41. Reynolds, J. A., and Karlin, A. (1978). Biochernistly 17, 2035-2038. Rosenbluth, J. (1974).J. Cell Biol. 62, 755-766. Rosenbluth, J. (1975).J. Neurocytol. 4,697-712. Ross, M. J., Klymkowsky, M. W., Agard, D. A., and Stroud, R. M. (1977).J. Mol. Biol. 116, 635-659. Rousselet, A., and Devaux, P. F. (1977). Biochem. Biophys. Res. Commun. 78, 448-454. Rousselet, A., Cartaud,J., and Devaux, P. F. (1979). C.R. Hebd. SeancesAcad. Sci., Ser. D 289, 461-463. Rousselet, A,, Cartaud, J., and Devaux, P. F. (1981). Biochim. Biophys. Acta 648, 169-185. Rousselet, A., Cartaud, J., Devaux, P. F., and Changeux, J. P. (1982).E M B O J . 1,439-445. Rubsamen, H., ELdefrawi, A. T., Eldefrawi, M. E., and Hess, G. P. (1978).Biochemistry 17, 3818-3826. Riichel, R., Watters, D., and Maelicke, A. (1981). Eur.J. Biochem. 119, 215-223. Saitoh, T., and Changeux, J.-P. (1980). Eur. J. Blochem. 105, 51-62. Saitoh, T., and Changeux, J.-P, (1981). Proc. Natl. Acad. Sci. U.S.A. 78, 4430-4434. Saitoh, T., Wennogle, L., and Changeux, J.-P. (1979). FEBS Lett. 108, 489-494. Sakmann, B., and Brenner, €3. R. (1978). Nature (London) 276,401-402. Sakmann, B., Patlak, J., and Neher, E. (1980). Nature (London) 286, 71-73. Sator, V. S., Gonzalez-Ros, J. M., Calvo-Fernandez, P., and Martinez-Carrion, M. (1979). Biochemistry 18, 1200-1206. Schiebler, W., and Hucho, F. (1978). Eur. J. Biochem. 85, 55-63. Schindler, H., and Quast, U. (1980). Proc. Natl. Acad. Sci. U.S.A. 77, 3052-3056. Schliwa, M. (1981). Cell 25, 587-590. Schmidt, J., and Raftery, M. A. (1972). Biochem. Biophys. Res. Commun. 49, 572-578. Schmidt, J., and Raftery, M. A. (1973). Biochemishy 12, 852-856. Schubert, D., Harris, A., Devine, C. E.,and Heinemann, S. (1974).J. CellBiol. 61,398-413. Sealock, R. (1980). Brain Res. 199, 267-281. Sealock, R. (1982).J. Cell Biol. 92, 514-522. Sealock, R., and Kavookjian, A. (1980). Brain Re$. 190, 81-93. Shorr, R. G., Dolly, J., and Barnard, E. A. (1978). Nature (London) 274, 283-284. Sigworth, F. J. (1982). Biophys. J . 37, 309a. Sigworth, F. J., and Neher, E. (1980). Nature (London) 287,447-449. Sine, S., and Taylor, P. (1979).J. Biol. Chem. 254, 3315-3325. Sine, S., and Taylor, P. (1980).J. Biol. Chem. 255, 10144-10156.

340

F. J. BARRANTES

Sine, S., and Taylor, P. (1981).J. Biol. Chem. 256, 6692-6699. Sobel, A , , Weber, M., and Changeux, J.-P. (1977). Ertr. J . Biochem. 80, 215-224. Sobel. A,, Heidmann, T., Hofler, J., and Changeux, J.-P. (1978).Proc.Natl. Acnd. Sci. U.S.A. 75, 510-514. Sobel, A.. Heidmann, T., Cartaud, J., and Changeux, J.-P. (1980). Eur. J . B i o c h m . 110, 13-33. Steinbach, A. B. (1968).j. Gm. P h s d . 52, 144-161. Steinbach, J. H. (1980). Cell S u g . Rezl. 6, 119-156. Steinbach, J. H., Merlie, J., Heinemann, S., and Block, R. (1979). Proc. Nntl. Acnd. Sci. C'.S..4. 76, 3547-3551. Stephenson, F. A,. Harrison, R., and Lunt, G. G. (1981). Eur.J. Biochm. 115, 91-97. St. John, P. A,, Froehner, S. C., Goodenough, D. A,, and Cohen, J. B. (1982).J. Ce/l. Biol. 92,333-342. Strader, C. D., and Raftery, M. A. (1980).Proc. iVntl. Acud. Sri. U.S.A. 77, 5807-5811. Strader, C. D., Revel, J. P., and Raftery, M. A. (1979).J. Cell Biol. 83,499-510. Strader, C. D., Hunkapiller, M. W.,Hood, L. E., and Raftery, M. A. (1980a). 117 "Psychopharmacology of Neurotransmitter Receptors" (H. 1. Yamamura, R. Olsen, and E. Usdin, eds.), pp. 35-46. Elsevier, Amsterdam. Strader. C. D., Lazarides, E.. and Raftery, M. A. (1980b).Biochm.Biophys. Res. Commun. 92, 365-373. Sugiyama, H., and Changeux, J.-P. (1975).Eirr. J . Biorhm. 55, 505-515. Sugiyama, H., Popot, J. L., and Changeux, J.-P. (1976).]. Mol. Biol. 106, 485-496. Sumikawa, K., Houghton, M., Emtage, J. S., Richards, B. M., and Barnard, E. A. (1981). A'ature (Loudon) 292, 862-864. Tarrab-Hazdai, R., and Goldfarb, V. (1982).Eur. J . Biochem. 121, 545-551. Tarrab-Hazdai, R., Geiger, B., Fuchs, S., and Amsterdam, A. (1978).Proc. Natl. Acnd. Sci. l'.S..4. 75, 2497-2501. Tarrab-Hazdai, R., Bercovici, T., Goldfarb, V., and Gitler, C. (1980).J. Biol. Chem. 255, 1204-1209. Teichberg, V., and Changeux, J.-P. (1976). FEBS Lett. 67, 264-268. Teichberg, V., Sobel, A., and Changeux, J.-P. (1977).Nature (London)267, 540-542. Thesleff, S. (1955).Acta PhTsiol. Scnnd. 34, 218-231, 386-392. Thomas, D. D., Dalton, L. R., and Hyde, J. S. (1976).J. Chem. P h y . 65, 3006-3024. Tsai, M. C., Oliveira, A. C., Albuquerque, E. X., Eldefrawi, M. E., and Eldefrawi, A. T. (1979).,\fol. P h a m c o l . 16, 382-392. Tzartos, S. J., and Lindstrom, J. (1980).Proc. 'Vntl. Acad. Sci. U.S.A. 77, 755-759. Tzanos, S. J., Rand, D. E., Einarson, B. L., and Lindstrom, J. (1981).J. Biol. Chenz. 256, 8635-8645. Vandlen, R. L., and Raftery, M. A. (1979).Arch. Biochm. Biophys. 197, 503-515. Vandlen, R. L., Wu, W.C.-S., Eisenach, J. C., and Raftery, M. A. (1979). Biochemistry 10, 1845-1854. Van Helden, D. F., Hamill. 0. P., and Gage, P. W. (1977).Nature (London) 269, 111-113. Vincent, A. (1980). Ph?.siol.R e , . 60, 756-824. Walker, J. W., Lukas, R. J.. and McNamee, M. (1981a).Biochemist? 20, 2191-2199. Walker, J. W., McNamee, M. G., Pasquale, E., Cash, D. J., and Hess, G. P. (198lb).Biochem. Biophys. Res. Commun. 100, 86-90. Weber, M., and Changeux, J.-P. (1974). .tfol. Pharmacol. 10, 15-34. Weiland, G., and Taylor, P. (1979). Mol. Phnnnacol. 15, 197-212. Weiland, G., Georgia, B., Lappi, S., Chignell, C. F., and Taylor, P. (1977).J. B i d . Chem. 252, 7648-7656.

DEVELOPMENTS IN STRUCTURE AND FUNCTION

34 1

Weill, C. L., McNamee, M. G., and Karlin, A. (1974). Biorhzm. Biophys. Res. Commun. 61, 997- 1003. Weinberg, C. G., and Hall, Z. (1979). Pror. Natl. Acad. Sci. U.S.A. 76, 504-508. Wennogle, L., and Changeux, J.-P. (1980). Eur. J . Biochem. 106, 381-393. Wennogle, L., Oswald, R., Saitoh, T., and Changeux, J.-P. (1981). Biochemistry 20, 24922497. Wise, D. S., Karlin, A,, and Schoenborn, B. (1979). Biophys. J. 28, 473-496. Wise, D. S., Schoenborn, B. P., and Karlin, A. (1981a).J. B i d . C h a . 256, 4124-4126. Wise, D. S., Wall, J., and Karlin, A. (1981b).J. Biol. C h a . 256, 12624-12627. Witzemann, V., and Raftery, M. A. (1978). Biochem. Biophys. Res. Commun. 85, 623-631. Wonnacott, S., Harrison, R., Lunt, G. G., and Barkas, T. (1980). Eur. J. Biochem. 108, 62 1-629. Wu, W. C.-S., Moore, H.-P.H., and Raftery, M. A. (1981). Proc. Natl. Acad. Sci. U.S.A. 78, 775-779. Zingsheim, H. P., Neugebauer, D.-C., Barrantes, F. J., and Frank, J. (1980). Pror. Natl. Acad. Sci. U.S.A. 77, 952-956. Zingsheim, H. P., Neugebauer, D.-C., Frank, J., Hanicke, W., and Barrantes, F. J. (1982a). EMBO J . 1,541-547. Zingsheim, H. P., Barrantes, F. J., Frank, J., Hanicke, W., and Neugebauer, D.-C. (198213). Nature (London) 299, 8 1-84. Zwolinski, B. J., and Eyring, H. (1947).J. Am. Chem. SOC.69, 2702-2707.

This Page Intentionally Left Blank

CHARACTERIZATION OF ( ~ 1 -AND (~2-ADRENERGIC RECE PTORS By D a v i d B. Bylund Deportment of Pharmacology School of Medicine University of Missouri Columbia, Missouri and David

C. U’Prichard

Department of Pharmacology School of Medicine Northwestern University Chicago, Illinois and Department of Neurobiology and Physiology College of Arts and Sciences Northwestern university Evonrton, Illinois

I. Introduction ......................................................... A. Receptors for Epinephrine and Norepinephrine ...................... B. Pharmacological Subdivision of a-Adrenergic Receptors ............... C. Radioligand Binding Studies ....................................... 11. a,-Adrenergic Receptors ................... ......... A. Characterization by Radioligand Binding ............................ B. Effector Systems Coupled to a,-Adrenergic Receptors ................ C. Regulation of a,-Adrenergic Receptors .............................. D. Solubilization of a,-Adrenergic Receptors ............................ 111. a,-Adrenergic Receptors .............................................. ................ A. Characterization by Radioligand Binding B. Effector Systems Coupled to a,-Adrenergic Receptors ................ C. Comparison of Agonist and Antagonist Binding: Toward a Kinetic Model of a,-Receptor Function ........................................... D. Regulation of a,-Adrenergic Receptors .............................. E. Localization of a,-Adrenergic Receptors ............................. F. Solubilization of a,-Adrenergic Receptors ............................ IV. Summary and Conclusions ............................................. References ...........................................................

344 344 345 349 354 3 54 364 368 376 376 377 398 404 410 417 419 420 422

343 INTERNATIONAL REVIEW OF NEUROBIOLOGY, VOL. 24

Copyright 0 1983 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-366824-7

344

DAVID

B. BYLUND

AND DAVID

c.

U’PRICHARD

1. Introduction

A. RECEPTORS FOR EPINEPHRINE A N D NOREPINEPHRINE The catecholamines are important regulatory compounds in the body and produce a variety of physiological effects. T h e actions of norepinephrine, a peripheral sympathetic and central neurotransmitter, and epinephrine, an adrenal hormone and putative central neurotransmitter, are mediated through adrenergic receptors. A large number of drugs also produce their effects by interacting with these receptors. T h e concept of adrenergic receptors originated with Langley (1905). Extrapolating from his work on the action of nicotine and curare on muscle, he suggested that epinephrine also exerts its effects by interacting with “receptive substances.” Dale (1906) made use of this concept to explain the differential effects of the ergot alkaloids on smooth muscle. He raised the possibility that the receptors at myoneural junctions could be of t w o types: one mediating the excitatory (motor) actions of norepinephrine and the other mediating the inhibitory actions of epinephrine. This functional receptor subclassification, based on the idea that adrenergic receptors could be considered to be of two classesthose whose actions result in excitation of the effector cells and those whose actions result in inhibition of the effector cells-competed with an alternative suggestion that two opposing transmitter substances (Sympathin E and Sympathin I) competed antagonistically at the same receptive site (Cannon and Rosenblueth, 1937). Then Ahlquist (1948) made a major advance by suggesting that the subtypes of adrenergic receptors could be differentiated on the basis of their pharmacology and not on the basis of their function. He studied the effects of five catecholamines (norepinephrine, methylnorepinephrine, epinephrine, methylepinephrine, and isoproterenol) on eight different physiological functions (nasal vasoconstriction, contraction of the uterus and the nictitating membrane, dilation of the pupil, relaxation of the gut, nasal vasodilation, relaxation of the uterus, and myocardial stimulation) and clearly showed that the order of potency for the catecholamines for the first five functions was markedly different from the order of potency for the remaining three functions. He attributed this difference to an actual difference in the receptors involved and suggested that they be called a-and P-adrenergic receptors, respectively. Inherent in these data is the concept that a single tissue, such as vascular smooth muscle, could contain both subtypes of adrenergic receptors which would mediate different functions. At the time of Ahlquist’s experiments the known adrenergic antagonists, such as phenoxybenzamine, phentolamine, and various ergot

CHARACTERIZATION OF a1- AND ff2-ADRENERGIC RECEPTORS

345

derivatives, appeared to block only a-adrenergic receptor responses, whereas specific p-receptor antagonists such as dichloroisoproterenol and propranolol were not developed for another 10 years. More recently Lands et al. (1967) proposed the subdivision of P-adrenergic receptors into P1 and P2 subtypes. This was also a pharmacological differentiation based on a comparison of the relative potencies of 12 agonists in several isolated organ systems. This subclassification of /3 receptors has been substantiated both by the development of subtype selective antagonists and by direct binding studies (Minneman et al., 1981).

B. PHARMACOLOGICAL SUBDIVISION OF a-ADRENERGIC RECEPTORS In contrast to the simple subclassification of P-adrenergic receptors by pharmacological criteria, attempts to correctly subdivide the a-adrenergic receptors have followed a more circuitous route. The initial subclassification was based on the presumed anatomic or topographic localization of the receptor. The idea that the terminals of noradrenergic axons contain functionally important a-adrenergic receptors stemmed from early work of Brown and Gillespie (1957) who reported that phenoxybenzamine, an a-adrenergic antagonist, increased the overflow of norepinephrine elicited by nerve stimulation in the perfused cat spleen. Although this data was originally misinterpreted, as early as 1959 Furchgott suggested the possibility that these results were due to an increase in neurotransmitter release from the nerve ending. Langer et al. ( 197 1) and Starke (197 1) suggested that presynaptic a-adrenergic receptors were responsible for the increase in norepinephrine overflow seen with a-adrenergic blockers and for the decrease seen with a-adrenergic agonists. The existence of these presynaptic or autoreceptors for norepinephrine which modulate its release is now widely accepted (Starke, 1977; Langer, 1977), although a presynaptic location for the receptors which mediate this function has not yet been conclusively proved. The observation that phenoxybenzamine was 30 times more potent in antagonizing autoreceptor function as compared to the classical postsynaptic receptor led Langer (1974) to suggest that autoreceptors and postsynaptic receptors are not identical and that they should be referred to as a2and a l ,respectively. This anatomic subdivision of a-adrenergic receptors (a,-receptors on presynaptic terminals) has not proved particularly useful because of considerable evidence suggesting that receptors having similar pharmacological properties to the autoreceptors are not located presynaptically. Inhibition of adenylate cyclase activity and subsequent physiological responses have been shown to be mediated

346

DAVID B. BYLUND AND DAVID

c. U’PRICHARD

through a-adrenergic receptors, which would be classified pharmacologically as azreceptors in both human platelets (Grant and Scrutton, 1979; Lasch and Jakobs, 1979; Hsu et al., 1979) and human adipocytes (Burns et al., 1981; Lafontan and Berlan, 1980; Kather and Simon, 1981). The human adipocyte also contains a receptors which would be pharmacologically classified as a1 receptors and can be differentiated from the az receptors by both functional and radioligand binding studies (Burns P t nl., 1981). Similarly, vascular smooth muscle, in addition to containing the classical postsynaptic a l receptor, also appears to contain postsynaptic receptors which are pharmacologically similar to az receptors (Drew and Whiting, 1979; Hamilton and Reid, 1980; Ruffolo et al., 1980; Timmermans et al., 1979). I n the rat submandibular gland azadrenergic receptors appear following reserpine treatment or denervation (Bylund and Martinez, 1980), and these receptors are localized postsynaptically (Bylund and Martinez, 1981). Nonpresynaptic azadrenergic receptors also have been characterized in pancreatic islets (Nakaki P t a!., 1981) and undifferentiated neuroblastoma X glioma hybrid cells (Sabol and Nirenberg, 1979a; Kahn et al., 1982). Even before most of the above data were available, Berthelsen and Pettinger ( 1977) realized the limitations of the anatomic subdivision of a-adrenergic receptors and suggested instead that they be classified on the basis of their function. According to this classification scheme, aladrenergic receptors are excitatory and az-adrenergic receptors are inhibitory. While this classification scheme certainly has merit and does tend to subdivide a receptors into groups that have similar pharmacological characteristics, it also has a number of drawbacks. As these authors pointed out in their paper, “the designation of a receptor as inhibitory or excitatory is an arbitrary one which is limited by our ignorance of the more complex molecular mechanism of each receptor’s actions” (Berthelsen and Pettinger, 1977). For example, several a2 responses could be considered excitatory, rather than inhibitory. The inhibition by a-adrenergic agonists of the release of catecholamines from adrenergic terminals may be due to activation of the sodium pump, which results in enhancement of calcium efflux (Cohen et al., 1980) and/or activation of guanylate cyclase (Pelayo et al., 1978; O’Dea and Zatz, 1976). Thus, the functional or physiological subclassification of subtypes of a-adrenergic receptors has been replaced by a more useful pharmacologic definition, just as the original subdivision of adrenergic receptors into excitatory and inhibitory types was replaced by the pharmacologic subdivision into a and /3 receptors. The third attempt to subclassify a-adrenergic receptors was based on what might be thought of as a biochemical approach. Wikberg (1979)

CHARACTERIZATION OF a1- AND a2-ADRENERGIC RECEPTORS

347

and Fain and Garcia-Sainz (1980) suggested that the at receptors mediate effects secondary to an elevation in intracellular calcium and involve an increased turnover of phosphatidylinositol, while a2-adrenergic effects are mediated by the inhibition of adenylate cyclase. While a subclassification based on biochemical mechanisms may eventually prove to be useful, at the present time the attempt to do this is limited by our relative ignorance of the actual mechanisms for a-adrenergic receptors in most tissues. Table I summarizes some of the biochemical responses that have been observed following a-adrenergic receptor stimulation. It would appear that there are a variety of biochemical mechanisms inTABLE I BIOCHEMICAL RESPONSES MEDIATEDBY Q-ADRENERGIC RECEPTORS Biochemical responses

Tissue

References

Q1

Phosphatidylinosital turnover (increase in cytosol Ca*+?) Phosphatidylinosital turnover Phosphorylase activation Potassium release Potassium release

Adipocyte

Burns et al. (198 1)

Aorta Liver Submandibular gland Parotid gland

Vilalobos-Molina et al. (1982) El-Refai et al. ( 1979) Bylund et al. (1982a) It0 et al. (1982)

Kidney cells (MDCK)

Levine and Moskowitz (1979)

Pineal gland Submandibular gland

Pelayo et al. (1978) ODea and Zatz (1976) Cohen et al. (1980)

Platelet

Grant and Scrutton (1979)

Adipocyte

Burns et al. (1981)

Pancreatic islets

Nakaki et al. (1981) Sabol and Nirenberg (1979a)

Adenylate cyclase inhibition Adenylate cyclase inhibition

Neuroblastorna X glial hybrid Liver Thyroid gland

Adenylate cyclase inhibition Intestinal secretion inhibitionb

Renal cortex Jejunum

4,) Promotes phospholipid deacylation Presynaptid" Guanylate cyclase activation (inhibits norepinephrine release) Na,K-ATPase activation (promotes calcium efflux, inhibits norepinephrine release) Q2

Adenylate cyclase inhibition (promotes aggregation) Adenylate cyclase inhibition (inhibits lypolysis) Adenylate cyclase inhibition (inhibits insulin release) Adenylate cyclase inhibition

Jard et al. (1981) Desmedt (1980) Yamashita et al. (1980) Woodcock et al. (1980) Nakaki et al. (1982)

Some evidence indicating the receptor is of the a2subtype. Some evidence indicating that this response is not mediated by cyclic AMP.

348

DAVID B. BYLUND AND DAVID

c . U’PRICHARD

volved. This conclusion is consistent with recent evidence that P-adrenergic receptors may have biochemical functions or mechanisms that are independent of the activation of adenylate cyclase (Maguire and Erdos, 1980; Hirata et al., 1979). For example, the mechanism of a-adrenergic inhibition of neurotransmitter release is not known, but one of the suggested hypotheses involves calcium. T h e biochemical subclassification would tend to indicate an a1 subtype rather than the pharmacologic a2 subtype (Cohen et al., 1980). T h e ultimate subclassification of adrenergic receptors will probably be based on the primary structure of the macromolecules involved. However, until that information is available it would appear prudent to use a pharmacologic subclassification scheme to subdivide a-adrenergic receptors. This is consistent with the primary division of adrenergic receptors into a and p types, as well as the p, versus P2 subclassification, both of which are pharmacologic schemes. The validity of a pharmacologic subclassification scheme can be determined by comparing the potencies of a variety of a-adrenergic agonists and antagonists in a variety of systems. A summary of some of these data is given in Tables I1 and 111. These tables compare the potencies or affinities of several a-adrenergic drugs in mediating or inhibiting physiologic responses, or in inhibiting the binding of selective a-adrenergic radioligands. The data have been normalized by dividing the potency of each drug by the potency of norepinephrine for agonists and that of phentolamine for antagonists. A ratio less than 1.0 means the drug is more potent (lower K , ) than the reference drug, whereas a ratio greater than 1.O means that it is less potent (higher K , ) . It is important to note that essentially the same conclusions are drawn from both the physiologic studies and radioligand binding studies. In both cases epinephrine tends to be slightly more potent than norepinephrine and is nonselective for the two subtypes. Phenylephrine is at selective, while clonidine is a2 selective as can be seen from the ratio of clonidine to phenylephrine, which tends to be about 1 for a1systems, but much less than 1 for a2systems. For the antagonists, prazosin is a , selective, while yohimbine is somewhat a2selective. T h e ratio of yohimbine to prazosin clearly differentiates the two subtypes and is a much better indicator than the clonidine : phenylephrine ratio. Rauwolscine may be even more a2 selective than yohimbine (Tanaka and Starke, 1980). I n addition to these physiologic and binding studies, there are several tissues such as the adipocytes in which two different a-adrenergic biochemical responses can be measured. These studies also support the pharmacological selectivity of clonidine and yohimbine as a,-selective agents and prazosin as an a,-selective agent (Burns et al., 1981).

CHARACTERIZATION OF (Y1- AND ff2-ADRENERGIC RECEPTORS

349

C. RADIOLIGAND BINDINGSTUDIES The successful labeling of a-adrenergic receptors using the radioligand binding technique, in which a drug of high specific radioactivity binds in a reversible manner to the receptor recognition site, was developed subsequent to labeling of /3-adrenergic receptors. In 1976 Lefkowitz took advantage of the fact that hydrogenation of naturally occurring ergot alkaloids results in compounds which are highly potent and selective a-adrenergic antagonists. The product of the catalytic reduction of a-ergocryptine with tritium gas, PHIdihydroergocryptine (DHEC), specifically labeled a-adrenergic receptors in the rabbit uterus (Williams and Lefkowitz, 1976). Independently, Snyder and collaborators used [3H]clonidine and rH]WB4 101 [2-[(2’,6’-dimethoxy) phenoxyethanolamino]methylbenzodioxan]to label a-adrenergic receptors in the brain (Greenberget al., 1976). These investigators felt that the agonist [3 Hlclonidine would preferentially label presynaptic receptors on noradrenergic nerve terminals in the brain. However, it was found that [3H]clonidine binding in brains from animals treated with the neurotoxin, 6-hydroxydopamine, was slightly increased rather than decreased as compared to control brains (U’Prichardet al., 1977a).The binding of the antagonist [3H]WB4101 was also increased slightly. These data were interpreted as an indication of denervation supersensitivity and that both ligands were labeling postsynaptic receptors. Since there were differences in the absolute affinities of agonists and antagonists at the binding sites labeled by the two ligands, it was felt that brain a-adrenergic receptors existed in two distinct but interchangeable conformational states (Greenberget al., 1976; U’Prichard et al., 1977a; Greenberg and Snyder, 1978). However, subsequent investigations in several laboratories have led to the conclusion that [3H]WB4101 selectively labels central aladrenergic receptors and rH]clonidine selectively labels a2 receptors (U’Prichard et al., 1979b; U’Prichard and Snyder, 1979; Tanaka and Starke, 1980). After an initial negative report (Davis et al., 1978) rH]DHEC was also shown to label a-adrenergic receptors in the brain. It became apparent that PHIDHEC labels both al-and az-adrenergic receptors with equal affinity in the brain (Peroutka et al., 1978; U’Prichard et al., 1978b, Miach et al., 1978). This general relationship of differential a-ligand binding to a1and az receptors has also been shown to apply to many peripheral tissues as well. A number of other radioligands have been used to label a-adrenergic receptors. These include [3H]epinephrine and [3H]norepinephrine in the brain (U’Prichard and Snyder, 1977a) and liver (El-Refai et al., 1979). [3H]Dihydroazapetine has been used in the rat vas deferens (Ruffolo et al., 1976). More recently, [3H]-

Agonist affinity ratio'

'Tissue

Epinephrine

Phenylephrine

&-Methyl norepinephrine

Clonidine

Oxymetazoline

Clonidine : phenylephrine

Reference

a, Receptors

w in 0

Physiologic studies Guinea pig aorta Rabbit pulmonary artery Rabbit aorta Guinea pig ileum Rabbit jejunum Rat kidney Radioligand binding studies Rat brain ([SH]WB4101)

0.83 0.22 1.4 0.14 0.5 0.72

8.3 4.3 5.0 0.25 6.2 1.5

4.0 12.4

0.59

2.6

6.8

0.43

0.024

0.17

Rat brain ([3H]prazosin)

0.37

2.2

2.7

0.30

0.036

0.14

Rat lung (PHlWB4101)

1.8

68

4.4

0.45

0.28

Rat lung (rHJprazosin) Rat submandibular gland Hlprazosin)

0.48

5.2

27

0.34

0.38

0.07

U'Prichard rt al. (1977a) Perry and U'Prichard (1982) Latifpour and Bylund (1981) Latifpour (1981)

0.52

2.3

1.22

0.021

0.53

Bylund rt al. (1982a)

(r

16

4.3 5.6

-

5.1

62 6.7 25 5 40 9.4

5.0 1.5 0.2 0.03 0.01 -

7.5 1.6 5.0 20 6.5 6.3

Wikberg (1979) Starke rt nf. (1975) Wikberg (1979) Wikberg (1979) Boudier ~t a/. (1975) Schmitz rt al. (1981)

Rat submandibular gland (PHlWB4 101) Rat kidney (rH]prazosin) Rat liver (PHIprazosin) Rat liver ([3H]norepinephrine) at Receptors Physiologic studies Guinea pig ileum Rabbit pulmonary artery Radioligand binding studies Rat brain ([3H]clonidine) Bovine brain (PH]rauwolscine)

W u1 c1

6.4 1.o 2.0

0.47 0.88 0.62

-

1.10 0.72 0.1

0.085 -

0.17 0.72 0.05

Bylund et al. (l982a) Schmitz et al. (1981) Geynet et al. (1981)

-

0.11

Geynet et al. (1981)

1.2

28

-

3.2

0.29 0.16

112 140

0.31 0.67

0.1 1 0.83

0.59 0.26

0.0010 0.0059

Wikberg (1979) Starke et al. (1975)

0.35

16

0.94

0.34

0.11

0.021

U’Prichard et al. (1977a)

0.32

0.57

0.81

0.013

0.003

0.023

Perry and UPrichard (1982)

Human platelet (PHIyohimbine) Human platelet (PHIp-aminoclonidine)

0.2 1

2.6

1.7

0. I4

0.017

0.054

Daiguji et al. 71981a)

0.37

6.0

-

0.47

0.41

0.068

UPrichard et al. (1982a)

Rat submandibular gland ( [3H]clonidine)

1.1

13.8

0.50

0.17

0.050

Bylund and Martinez ( 1980)

Rat lung [(neonatal) [3H]yohimbine]

1.2

0.11

1.o

0.0069

Latifpour et al. (1982)

10

16 ~~

a

5.6 20

Affinity ( K , ) of drug/a&ity

(K,) of norepinephrine.

0.82

Antagonist affinity ratio" Tissue

Prazosin

Yohimbine

Yohimbine : prazosin

Reference

Q1

Physiologic studies Rat anococcugeus muscle Rabbit aorta Rat kidney Radioligand binding studies Rat brain (rHlWB4101) Rat lung (FHIprazosin) Rat lung (['H]WB4101) Human lung (PHIprazosin) Rat submandibular gland ([3H]prazosin) Rat submandibular gland ([3H]WB4101) Rat kidney (rH1prazosin)

0.32 0.83

20 45

62 54

0.20

87

440

42 80 462 152 480 1130 36

470 1700 1500 12,000 2400 3600 1600

0.089 0.046 0.31 0.012 0.20 0.31 0.023

Doxey et al. (1977) Sheys and Green (1972); Cavero rt a/. (1978) Schmitz et a/. (198 1) U'Prichard et al. ( 1 9 7 8 ~ ) Latifpour and Bylund (1981) Latifpour and Bylund (1981) Barnes et a/. (1980) Bylund et al. (1982a) Bylund et a/. (1982a) Schmitz et al. (1981)

ff2

01

01 01

Physiologic studies Rat vas deferens Radioligand binding studies Human platelet (PHIclonidine) Human platelet (PHIyohimbine) Human adipocyte (PHIyohimbine) Bovine brain (PHIepinephrine) Bovine brain (PH]rauwolscine) Rat submandibular gland ([SHIclonidine) Rat kidney ( [3H]yohimbine) Rat lung (neonatal ([3H]yohimbine) Neuroblastoma (PHIyohimbine) Rat brain (cortex) (rHlyohimbine) Rat brain (striatum) (PHIyohimbine) Rat brain (cortex) (rH1clonidine) Rat brain (striatum) ([3H]clonidine) a

Affinity (K,) drug/affinity (K,) phentolamine.

>60 142 100 350 4000 125 1440 5.0 1.54 1.08 3.8 0.91 352 142

1.6 0.23 0.22 0.83 11 0.63 17 0.67 0.28 0.20 0.50 0.12 15 7.1

(0.03 0.0016 0.0022 0.0024 0.0027 0.0050 0.012 0.13 0.18 0.19 0.13 0.13 0.043 0.050

Doxey et al. (1977) Shattil et al. (198 1) Daiguji et al. (1981a) T h a r p et al. (1981) Perry and U'Prichard (1982) Perry and U'Prichard (1982) Bylund and Martinez (1980) Schmitz et al. (1981) Latifpour et al. (1982) Kahn et al. (1982) D. B. Bylund (unpublished) D. B. Bylund (unpublished) D. B. Bylund (unpublished) D. B. Bylund (unpublished)

354

DAVID B. BYLUND AND DAVID

c.

U'PRICHARD

prazosin, [3H]p-aminoclonidine (PAC), [3H]phentolamine, VHIyohimbine, [3H]rauwolscine, and [1251]BE-2254(HEAT) also have been used to label a-adrenergic receptors. It is the purpose of this article to review recent contributions to the characterization of a,- and a,-adrenergic receptors. Several previous reviews are available which deal in part with various areas of this subject (Lefkowitz, 1978; Kunos, 1978; Hoffman and Lefkowitz, 1980a,b; U'Prichard, 1981; Wood et al., 1979a). We have focused on those studies which deal with a 1 -and a,-adrenergic receptor subtypes and have emphasized the regulation and the relationship of binding and function.

II. a,-Adrenergic Receptors

A.

C H A K A C I'EKI/ATION

I31 R A D I O L I G 4 N D

BINDING

1. A n t a g o m t RadiolzgandT a. [3H]Ij'BBJIOl and r H ] p r a z o ~ n .The potent a,-adrenergic antagonists, WB4101 and prazosin, have been used as radioligands to label receptor binding sites in both the brain (Table IV) and in a variety of peripheral tissues (Table V). T h e K , values obtained in various laboratories for both CJHlWB4101 and VHIprazosin binding in the brain are remarkably consistent and, with a few exceptions, range between 0.14 and 0.31 n'tf. T h e K , values reported for peripheral tissues are also relatively consistent, although the variation is somewhat larger. T h e majority of the values lie between 0.1 and 0.9 niM. T h e reportedB,,, values for the various tissues are generally in the range of about 20-400 pmol/ gm prot and are similar to other catecholamine receptors such as the P-adrenergic and dopamine receptors (Maguire et al., 1977; Seeman, 1980). In contrast to the relatively good agreement among tissues and laboratoriey for K,, and B,,, values for a,-receptor binding, the K, for inhibitors that have been reported in the literature are much more variable. Table VI presents data for the inhibition of rHlWB4101 binding by various a-adrenergic drugs in four tissues from the rat. Although the K , values vary by as much as 10-fold or more, in each of the four tissues the pharmacological profile of the labeled receptor is characteristic of an a,-receptor subtype. It is generally accepted that rHlWB4lOl selectively labels a, receptors in CNS tissue, but its selectivity in peripheral tissues has been questioned. For example, in the rat uterus, [3H]WB4101 appears to bind to both a1and a , receptors with similar a h i t i e s (Hoffman

CHARACTERIZATION OF 0 1 - AND (Y2-ADRENERGIC RECEPTORS

355

and Lefkowitz, 1980d). In the human platelet, which lacks detectable a , receptors, [3H]WB4101 appears to bind potently to a2 receptors, although the K , value is still about 10-fold greater than in tissues where it labelsa, receptors (Daigujiet al., 1981a). In the platelet the& values for prazosin and yohimbine in inhibiting rHIWB4 101 binding are about 3000 and 5 nM, respectively, confirming the a2 characteristics of the binding (D. B. Bylund, unpublished). In the bovine retina, rHlWB4101 also appears to label a2 receptors (Bittiger et al., 1980). Due partially to this lack of specificity of [3H]WB4101in at least some tissues, rH]prazosin is now generally used for the study of a,-adrenergic receptors. The Ki values summarized in Table VII clearly indicate that t Hlprazosin labels an a,-receptor binding site. However, as was the case with [3H]WB4101, there is a considerable variation in the Ki values among tissues and species. Although it is generally assumed that rHlWB4101 and THIprazosin label the same population of al-adrenergic receptors, this has not yet been demonstrated due to the lack of studies which have compared the binding of both ligands under identical conditions. In the rat submandibular gland, lung, and cortex where this comparison was made, we have found that the Ki values for drugs in inhibiting the binding of the two radioligands were similar but not identical (Tables I1 and 111). In the submandibular gland, the B,,, value for rH]prazosin was 61% higher than that for THlWB4101 (Table V). Furthermore, in the rat lung the B,,,, for [3H]prazosinwas more than double that found for rHlWB4101 (Table V). The importance of these discrepancies is not yet clear. In addition to the human platelet and the bovine retina, several other tissues apparently lack a,-adrenergic receptors. These tissues include neuroblastoma glioma X hybrid cells (Haga and Haga, 1981; Kahn et al., 1982), possibly human subcutaneous adipose tissue (Burns et al., 1981; Tharp et al., 1981), and rat and pig cerebral microvessels (Harik et al., 1980). On the other hand, a,-adrenergic receptor binding was detected in the bovine cerebral microvessels (Peroutka et al., 1980). The monovalent cations, such as lithium and sodium, and the guanine nucleotides appear to have little effect on the binding of p Hlantagonists to a,-adrenergic receptors in most tissues, although these agents markedly affect the binding of radioligands to a2receptors. In the calf and rat cerebral cortex, the specific binding of [3H]WB4101 and rH]prazosin, respectively, was not significantly altered by guanine nucleotides at concentrations up to 1 mM (U’Prichard and Snyder, 1978a; Perry and U’Prichard, 1983). Sodium ,lithium, or potassium ions at concentrations up to 150 mM did not alter rHlWB4101 binding in the calf cortex (Greenberg et al., 1978). In addition, no effect of guanine nuc-

TABLE IV RAI>IOLIGANL) BINUING TO CENTRAL (Y~-AI)HENEKGI(: RECEPTOKS" Bm,,

W

01 Q,

Brain region

Species

Radioligand

pmol gm prot

100 82 t 17 77 rfr 3

Whole brain Whole brainb*= Whole brainb Whole brainb

Rat Rat Rat Rat

WB4101 DHEC Prazosin Prazosin

Whole brainb Whole braind Cerebral cortex Cerebral cortex Cerebral cortex Hypothalamus Hypothalamuse Thalamus

Rat Rat Rat Rat Rat Rat Rat Rat

Prazosin WB4101 WB4101 BE2254g BE2254' WB4101 DHEC WB4101

pmol gm tissue

11

80 rfr 75 rfr 84 rfr 210 180 rfr

*

2 9 9 26

20 3.2 4.1

85 t 4

h.,,

(nM) 0.48 4.7 rfr 1.3 0.26 t 0.08 0.28 rfr 0.04 0.26 rfr 0.08 0.16 rfr 0.02 0.14 rfr 0.01 0.08 rfr 0.014 0.10 rfr 0.03 0.28 1.07 0.32 rfr 0.06

Reference U'Prichard PI al. (1977a) Miach el al. (1978) Miach rt al. (1980) Greengrass and Bremner (1979) Morns et al. (1980) U'Prichard P I al. ( 1979a) U'Prichard rt al. (1979a) Engel and Hoyer (1981) Glossmann rt al. (1981) Neethling et al. (1981) Neethling ~t al. (1981) U'Pnchard et al. (1980b)

Midbrain Hippocampal gyrus' Dentate gyrus' Cerebral cortex Cerebral cortex Hippocampus Corpus striatum Cerebral cortex Frontal cortex Caudate nucleus Pons

Rat Rat Rat Mouse Mouse Mouse Mouse Calf Calf Calf Calf

WB4101 WB4101 WB4101 WB4101 WB4101 WB4101 WB4101 WB4101 WB4101 WB4101 WB4101

58 169? 14 269 ? 7 66 230 48 3 10

0.17

Gheyouche et al. (1980) Crutcher and Davis (1980) Crutcher and Davis (1980) Rehavi et al. (1980a,b) Rehavi et al. (1980a,b) Rehavi et al. (1980a,b) Rehavi et al. (1980a,b) Lyon and Randall (1980) U'Prichard et al. (1977b) U'Prichard et al. (1977b) U'Prichard et al. (1977b)

3

6.5 5.2 4.9 2.9

k

3 0.25 2.9 0.27 4.3 0.26 f 0.03 0.3 1 0.3 1 0.29

1.0

~

~

~

~

~

Unless otherwise noted the assay temperature was 23-25"C, the tissue preparation was a crude particulate fraction, and tritium was the isotope. Values given are means f SEM. Less cerebellum. In the presence of 0.1 p M yohimbine. Less cortex. Nonspecific binding defined by 0.1 p M prazosin. 'Assays temperature 30°C. le51 used as isotope. a

nm,,

Ds

w?

m

pmol gm prot

~~

Tissue

Species

Tritiated ligand

Adipocyte Heart, ventricle Heart, left ventricle Heart Aorta Aorta Aorta Irisb Livee Livef Livef Lung Lung

Human Rat Rat Guinea pig Rabbit

WB4101 WB4101 WB4101 Prazosin WB4101 WB4101 Prazosin WB4101 Prazosin Norepinephrine Prazosin Prazosin Prazosin

cow cow

Rabbit Rat Rat Rat Human Guinea pig

303 f 46 52 2 4 28 ? 1 65 ? 26 162 ? 6 134 ? 5 730 436 If: 75 340 ? 70 760 ? 40 600 47 ? 7

pmol gm tissue

2.57

?

0.14

2.0

?

0.6

ti,, (11

nr)

0.86 ? 0.07 1.2 f 0.1 0.18 0.53 f 0.17 0.84 ? 0.07 1.67 t 0.33 0.66 ? 0.16 2.3 0.05 138 ? 60 0.15 ? 0.02 1.75 0.20 ? 0.05

Reference Burns rt (11. (1981) Torda rt d.( 1981) Yamada rt 01. (1980b) Karliner ef nl. (1979) Fuder r / nl. (1981) Rosendorff ct a/. (1981) Rosendorff rt nl. (1981) Taft rt ul. (1980) Hoffman rt al. (1981a) Geynet rt nl. (1981) Ceynet P / al. (1981) Barnes et al. (1980) Barnes rt a/. (1979)

Lung Lung Lung Lung Lung Sublingual gland Submandibular gland Submandibular gland Parotid gland Uteruse Uterus Kidney cw

g

Rat Rat Rat Rat Dog Rat Rat

WB4101 WB4101 DHEC Prazosin Prazosin Prazosin Prazosin

57 f 2 51 f 3 60 f 7 126 f 2 22 57 ? 11

Rat

WB4101

Rat Rabbit Rabbit Rat

Prazosin DHEC Prazosin Prazosin

3.2 t 0.6 8.7 h 0.06

0.33 f 0.03 1.2 f 0.1 1.7 f 0.3 0.11 +. 0.01 0.48 0.43 ? 0.16 0.43 h 0.06

Latifpour and Bylund (1981) Torda et al. (198 1) Latifpour and Bylund (1981) Latifpour (1981) Hasegawa and Townley (1981) Martinez et al. (198213) Bylund et al. (1982a)

96 f 10

5.3

0.37 f 0.04

Bylund et al. (1982a)

13 f 1 25 29 ? 7 57 f 6

0.43

0.78 rt 0.32 4.7 0.5 ? 0.15 0.85 ? 0.05

Ito el al. (1982) Hoffman and Lefkowitz (1980~) Lavin et al. ( I98 1) Schmitz et al. (1981)

rt

0.7

Unless otherwise noted the assay temperature was 23-25°C and the tissue preparation was a crude particular fraction. Values given are mean f SEM. * Microsomal fraction. Plasma membranes. Chronic obstructive airway disease. 17% of total DHEC binding.

360

DAVID B. BYLUND A N D D A V I D

c. U’PRICHARD

TABLE VI I S H I ~ I T I OOF N [3H]WB4101 BINDING I N RAT TISSUES

Drug Agonists (-)-Epinephrine (-)-Norepinephrine Oxymetazoline ( - f-Phenylephrine (+)-Norepinephrine Clonidine Antagonists Phentolamine Prazosin WB4101 Yohi mbi ne

Brain“

Heartb

590 1000

90 723

200

1170

1800

369

8500 480

24 2600 67,000 430 3.6 0.49 0.6 480

Lung

110

50

5.3 0.18 0.20 48 1

1.6 0.5 740

Submandihular glandd

64 136 10 764 13,500 150

0.71 0.22 0.23 800

U’Prichard ut ai. (1977a). (1980h). Latifpour and Bylund (1981). Bylund et rrl. (1982a).

* Yamada P I 01.

leotides was found on the binding affinities of agonists for rat liver and brain a l receptors labeled with THIDHEC, rHlWB4101, or [3H]prazosin (Hoffman Pt nl., 1980a; U’Prichard and Snyder, 1978a; Perry and U’Prichard, 1983). However, it has been reported that guanine nucleotides decrease the affinity of epinephrine at rat cardiac a 1 receptors labeled by [3H]WB4101 (Yamada ff nl., 1980~) and decrease the binding of [3H]norepinephrine to rat hepatic a1receptors (Geynet el al., 1981). The binding of [3H]prazosin to rat brain membranes was not altered by the presence of 150 nM NaCl, although sodium did alter the affinity of agonists in inhibiting [3H]prazosin binding (Glossmann and Hornung, 1980a). It appears that sodium may decrease the affinity of a1 agonists but may increase the affinity of agagonists at the receptor site labeled by [3H]prazosin (Glossmann and Hornung, 1980a). [3H]WB4101 appears to be useful in characterizing receptors using the autoradiographic technique. In the rat brain, HlWB4 101 binding to slide-mounted tissue sections had all the characteristics associated with a, receptors, and thus was used for light microscopic autoradiographic localization of these receptors (Young and Kuhar, 1980). [3 HIPhentolamine was used in autoradiographic study of the urinary bladder of the rat, although specific binding to a-adrenergic receptors was not demonstrated (Jonaset al., 1980).

TABLE V I I INHIBITION OF rH]PRAZOSIN BINDING

Drug Agonists (-)-Epinephrine (-)-Norepinephrine Oxymetazoline ( -)-Phenylephrine (+)-Norepinephrine Clonidine Antagonists Phentolamine Prazosin WB4101 Yohimbine a

Miach et al. (1980).

Rat brain"

1440 5615

193,000 2315 304 0.3 3.8 8900

* Greengrass and Bremner (1979). Bylund et al. (1982a). Latifpour (1981). Barnes et al. (1980). Barnes ~t al. (1979).

Rat brainb

600 900 23 1400 43,000 340

0.1 1.o 1000

Rat submandibular gland'

40 77 1.6 174 2200 94 0.98 0.32 0.17 450

Rat lungd

262 400 327 2085

Human lunge

460 590

690 1400

46,000

110,000 110,000 2,200

365 9 0.38 750

Guinea pig lung'

27 0.33 4100

6.6 0.13 0.80 3100

362

DAVID B . BYLUND A N D DAVID c . U'PRICHARD

a- 1-Adrenergic receptor binding studies have also proved useful in understanding the side effects of some psychoactive drugs. T h e affinities of the tricyclic antidepressant drugs for a ,-adrenergic receptor sites labeled by [3H]WB4101 in the brain correlate well with the capacity of these drugs to relieve psychomotor agitation and to induce sedation and hypotension (U'Prichard et a(., 1978a). Similarly, the relative affinity of neuroleptic drugs for rHIWB4101 binding sites provides an index of the relative propensities of these drugs for eliciting autonomic side effects such as orthostatic hypotension and sedation (Peroutka rt al., 1977). In a study of the metabolites of neuroleptic drugs, there was also an excellent correlation between the potencies of the metabolites at the a 1 receptor as determined by binding studies and their potency as established by clinical and animal studies (Bylund, 1981). h. ['251]BE-225-f].In binding studies where the yield of receptorcontaining tissue is necessarily limited, for example, using cell and tissue cultures, i t is especially useful to have a radioiodinated ligand. [Iz5I]BE2254, 2-[~-(4-hydroxphenyl)ethylaminomethyl]tetralone, also called [1251]HEAT,an aminotetralone derivative has recently been synthesized with a specific activity of 2000 Cdmmol and shown to label brain a 1 receptors quite specifically with a K , in the 0.1 nM range (Engel and Hover, 1981; Glossmann el ul., 1981). r . [3H]Dili~~~ro~).gocr~ptitie. As a radioligand for the specific study of a,-adrenergic- receptors, HIDHEC suffers from the disadvantage of labeling a , and a2receptors with about equal affinity. Several techniques have been developed to overcome this difficulty and permit the selective study of the a-receptor subtypes. One method involves the use of complex computational techniques with computer modeling of inhibition curves of prazosin against rH]DHEC (Hoffman et al., 1979). Using this method the percentage of a 1 and a , receptors in half of a given tissue sample is determined from inhibition studies, and the total number of a receptors is determined by HIDHEC saturation analysis on the other half of the tissue sample. T h e density of, and inhibitor affinity at, the two receptor subtypes is then calculated. The other method has several variations, but basically involves the use of a subtype-selective drug at a specified concentration. T h e concentration is chosen so that essentially all of radioligand binding at one of the subtypes will be inhibited, while binding at the other subtype will not be affected (Miach P t nl., 1978). In the best docutnented example, B,,,, and K,)values in rabbit uterine membranes were obtained in the presence and absence of 100-nM prazosin. The B,,,, in the absence of prazosin was taken to represent the total adrenergic receptor population, whereas that in the presence of prazosin was defined as a,-adrenergic receptors. T h e contribution of

r

CHARACTERIZATION OF (Y1- AND (Y2-ADRENERGIC RECEPTORS

363

al-adrenergic receptors was then calculated by the difference (Hoffman and Lefkowitz, 1980~). Using this method, the density of a , sites labeled by [3H]DHEC was 38 fmoVmg prot, which is in fair agreement with 29 fmoVmg prot of a1sites labeled by [3H]prazosin(Lavin et al., 1981; there is a typographical error in this paper which gives the density of pH]prazosin sites as 19 fmoVmg prot rather than the correct value of 29). The estimate of the density of a2receptors using rH]DHEC was 80% higher (129 fmoumg prot) than that obtained from rHlyohimbine binding (72 fmoVmg prot). The reason for this rather large difference is not presently understood. Using a similar approach, the total a-, al-, and a2adrenergic receptor sites in the rat hypothalamus were assayed using 100 n M phentolamine, prazosin, and clonidine, respectively, to define specific binding (Neethling et al., 1981). The values obtained in this manner agreed reasonably well with the values obtained using [3 HlWB4 101 and p Hlclonidine to assay a , and a2 receptors separately (Haga and Haga, 1980). Thus, while THIDHEC can be used to study a , receptors, it is not clear that the use of these techniques has any advantages over the more direct approach using subtype specific radioligands, such as THlprazosin or [1251]BE-2254. d. Multiple Afinity States of Alpha-1 Receptms. In contrast to the a2and P-adrenergic receptor systems, where there is strong evidence for the existence of high- and low-affinity agonist states, there is thus far little evidence that a,-adrenergic receptors have multiple affinity states. The inability to demonstrate 3H-labeled agonist binding to a , receptors at low ligand concentrations suggests the absence of a high-affinity agonist state. The lack of an effect of guanine nucleotides on agonist inhibition of 3H-labeled antagonist binding in almost all a1systems studied might also be taken as evidence of a high-affinity agonist state, although it is probably more properly interpreted as a result of a , receptors not being coupled to adenylate cyclase. There is, however, a recent report of [3H]norepinephrine binding to putative a , receptors in rat liver plasma membranes which is decreased by GTP (Geynetet al., 1981). The reports of regulation by sodium ions of agonist affinity in inhibiting central [3 Hlprazosin and [1251]BE-2254binding indicate that in some ionic media multiple affinity states do exist, although no attempt has yet been made to fit them to a kinetic model of a,-receptor function. The Rosenthal analyses of p HlWB4 101 and THlprazosin saturation data are generally monophasic indicating a single class site, although in a few instances biphasic Rosenthal plots have been reported (U’Prichard et al., 1979a; Lyon and Randall, 1980; Weinreich et al., 1980; Rehavi et al., 1980b). In some of these cases the low-affinity component may be due to the binding of [3H]WB4101to a2 receptors (see above) or to calcium ion

364

DAVID B. BYLUND A N D DAVID

c. U'PRICHARD

channels (Atlas and Adler, 1981) rather than to a low-affinity state of the a , receptor.

2. Agonist RadioligaridA To date, H-labeled agonist binding to a,-adrenergic receptors has been reported only in a single tissue and only by one laboratory (Geynet et al., 1981). I n general, VHIepinephrine and [3H]norepinephrine appear to bind specifically to a2-adrenergic receptors (see below), which can be interpreted as indicating that the a , receptors lack a high-affinity agonist conformation. Thus, the binding of these agonists to a , receptors is of too low affinity to be observed with the present techniques. It is certainly reasonable to think that eventually a, agonists with sufficiently high affinity will be found which can be used as radioligands.

B . EFFECTOR SYSTEMS COUPLED TO a,-ADRENERGIC RECEPTORS 1. Adipocytes During the past several years, the adipocyte has emerged as one of the better systems for the study of a-adrenergic receptor binding and function. Adipocytes from both the human and hamster appear to have a modest level of a,-adrenergic receptor binding (Table VIII), although one laboratory has been unable to observe &,-receptor binding in human subcutaneous adipose tissue. T h e density of a,-adrenergic receptors appears to be somewhat variable among subjects or among animals, although generally it is in the range of 50 to 300 fmol/mg prot. TABLE V I I I CI~-ADRENERGIC RECEPTOR BINDING I N ADIPOSETISSUE

Species ~~

Tissue type

Tritated ligand

Bin,, (fmoVmg prot)

Properitoneal Subcutaneous Subcutaneous White fat White fat

WB4101 Prazosin DHEC DHEC DHEC

300" 290 lod 2off

Reference

~

Human Human Human Hamster Hamster

67d

Burns et al. (1981) Wrightut 01. (1981) Tharp t t al. (1981) Pecquery and Giudicelli (1980) Garcia-Sainz et al. (1980b)

Range was 175-450 fmoUmg tissue.

* N o binding to al receptors was detectable. 15-25% of total DHEC binding was estimated to be to a, receptors. 0-30% of total DHEC binding was estimated to he to a1receptors.

CHARACTERIZATION OF (Y1- AND Q2-ADRENERGIC RECEPTORS

365

Simulation of a,-adrenergic receptors in the adipocyte appears to be coupled to increased turnover of the phosphatidylinositol and phosphatidic acid. It has been proposed that the increased turnover of these phospholipids in many tissues results in an increase in cytoplasmic calcium ion concentration due to the uptake of extracellular calcium and the release of bound calcium (Michell, 1975; Fain and Garcia-Sainz, 1980). The evidence for the a1nature of the increased turnover of these phospholipids is based on the selective inhibition of the epinephrinestimulated 32Pincorporation. The potency order of prazosin and yohimbine in reducing 32 P incorporation into phosphatidylinositol and phosphatidic acid is clearly that of an a,-mediated process (Burnsetal., 1981). Thus, while the adipocyte appears to have both a receptor binding site that can be labeled by radioligands and a biochemical function that has a specificity for a ,-receptor processes) this does not yet constitute proof that the receptors which are labeled in binding studies are identical to those receptors responsible for the biochemical effect. At the present time, neither the molecular mechanism whereby a,receptor stimulation results in an increase in the turnover of phospholipids nor the link between the increased turnover and calcium ion movement is known. In the rat adipocyte a,-adrenergic stimulation results both in an increase in phosphatidylinositol turnover and in the inactivation of glycogen synthase (Garcia-Sainz et al., 1980a). The divalent cation ionophore A23 187 can also inactivate glycogen synthase (Lawrence and Larner, 1977, 1978). These and other data suggest that turnover of phosphatidylinositol is involved in some fashion in the gating or mobilization of calcium. It has, in fact, been suggested that phosphatidic acid itself is a calcium ionophore (Putney et al., 1980; Harris et al., 1981). However, a causal relationship between the a,-mediated increase in the turnover of phosphatidylinositol and the elevation of cytosolic calcium is not yet established.

2. Salivary Glands The rat salivary glands have also proved to be a useful tissue for the investigation of relationship between receptor binding and function. The rat has three salivary glands. The parotid is a serous gland which releases amylase followingP-adrenergic stimulation. The submandibular is a mixed serous and mucous gland to which is attached the mucous sublingual gland. Both the parotid (Batzri et al., 1973) and the submandibular glands (Martinez et al., 1976))but not the sublingual gland (Martinez et al., 1982b), release potassium in response to an a-adrenergic stimulation. As would be expected, but contrary to a previous report (Arnett and Davis, 1979), the al-adrenergic receptor subtype mediates

366

DAVID B. BYLUND A N D DAVID c . U’PRICHARD

this potassium release response in the submandibular glands (Bylund et al., 1982a). Both norepinephrine and the a1 agonist methoxamine stimulate the release of potassium, while prazosin, but not yohimbine, is able to block the norepinephrine-stimulated release. Similarly, in the rat parotid gland, the K , values for a series of eight adrenergic drugs in inhibiting pH]prazosin binding correlated significantly ( r = 0.85, p < 0.01) with the potency to stimulate or inhibit potassium (Itoet al., 1982). Several groups have studied the binding of HIDHEC to various preparations of rat salivary glands. [3 HIDihydroergocryptine binding has been characterized in both the parotid (Strittmatter et al., 1977) and the submandibular glands (Pointon and Banerjee, 1979), although in neither study was a differentiation attempted between a 1 and ag subtypes. In another study of rH]DHEC binding to submandibular glands, it was concluded that the a receptors labeled by the radioligands were of the az subtype (Arnett and Davis, 1979). This conclusion is difficult to reconcile with the finding that WB4101 was equally potent with yohimbine in inhibiting the binding of the radioligand and that prazosin was only eightfold less potent than these antagonists. Furthermore, significant binding of neither [3H]clonidine nor [3H]yohimbine has been demonstrated in adult rat submandibular gland (Bylund and Martinez, 1980; Pimoule et al., 1980; Bylund et al., 1982a). Thus, the delineation of the receptor subtype(s) to which [3H]DHEC binds in the submandibular glands is not yet clear, and the resolution of the question will await the application of the techniques described above for the study of subtype specific receptor binding using VHIDHEC. Alpha- 1-adrenergic receptors in the rat submandibular and sublingual glands have been studied using [3H]WB4101 and rH]prazosin. As can be seen from Tables VI and VII, the submandibular gland shows 1500-3000-fold specificity of prazosin as compared to yohimbine for these radioligands. T h e density of binding sites in the sublingual gland is only about one-third of that found in the submandibular gland (Table V), which is consistent with the lack of measurable norepinephrinestimulated potassium release in the sublingual gland slices (Martinez ct al., 1982b). Unilateral ligation of the main excretory duct of the rat submandibular gland results in a progressive decrease in gland weight due mainly to acinar cell atrophy. Based on [3H]WB4101 binding, the density of a l receptor binding sites in the ligated glands were approximately onethird that found in the contralateral control glands (Martinez rt al., 1982a). These data suggest that the majority of the al-adrenergic receptors in the submandibular gland are located on acinar cells.

r

CHARACTERIZATION OF (Xi- AND a2-ADRENERGIC RECEPTORS

367

3. Liver

Several groups have studied the binding of radioligands of a-adrenergic receptors in the rat liver, as compared to the effect of a-adrenergic drugs on glycogen phosphorylase activity. It is generally agreed that HIDHEC labels predominantly a ,-adrenergic receptor sites in this tissue and that the activation of glycogen phosphorylase is an a,-mediated process. Two laboratories obtained a good correlation between the potencies of a-adrenergic agonists and antagonists in inhibiting rH]DHEC binding in plasma membranes and the potency of agonists (antagonists) in activating (inhibiting) glycogen phosphorylase (Hoffman et al., 1980b; Aggerbeck et al., 1980). Another group found a better correlation between PHlepinephrine binding and phosphorylase activity than between [3H]DHEC binding and phosphorylase activity (El-Refai et al., 1979) and in addition, presented limited evidence that [3H]epinephrinewas binding to at receptors (El-Refai and Exton, 1980). Geynet et al. (1981) have presented evidence suggesting that [3H]prazosin and [3 Hlnorepinephrine label distinct a,-receptor binding sites, whereas rH]DHEC labels both sites. They speculate that the site labeled by rHJnorepinephrine, which has a higher af€inity for agonists, is the physiologically active form of the receptor. On the other hand, Hoffman et al. (198lb) found that [3H]epinephrine at low concentrations binds selectively to a2 receptors. The discrepancy appears to lie partly in the different concentrations of Hlcatecholamine used in the experiments, although an adequate resolution of these conflicting data is not obvious.

r

r

4. Central N m o u s System In some CNS tissues a,-receptor activation appears to be coupled to an increase in the production or levels of cyclic nucleotides. In both the brain and the spinal cord, a,-adrenergic agonists increased the tissue content of cyclic AMP, an effect which appears to be dependent upon calcium (Schwabeet al., 1978). However, few studies have attempted to correlate binding characteristics of a ,-receptors with brain adenylate cyclase activity or cyclic AMP levels. In the rat cerebral cortex a significant potency correlation ( r = 0.87) was found between rHlWB4101 sites and a-adrenergic-mediated cyclic AMP accumulation in brain slices (Davis et al., 1978). These data suggest that [3H]WB4101may bind to the membrane receptor sites mediating the adrenergic accumulation of cyclic AMP in this tissue. Other evidence suggests that this a, response is probably mediated by the interaction of calcium with calmodulindependent adenylate cyclase and by analogy with the peripheral system

368

DAVID B. BYLUND AND DAVID

c. U’PRICHARD

cited above, may result from an increased phosphatidylinositol metabolism to generate phosphatidic acid and elevate intracellular calcium levels. C. REGULATION OF CY,-ADRENERGIC RECEPTORS Physiological and pharmacological regulation of receptor density as measured by the number of receptor binding sites is emerging as an important area of study. I n at least some instances a change in receptor density has been strongly implicated as the important factor in supersensitive and subsensitive responses to hormones and neurotransmitters. T h e levels of adrenergic receptors can be altered by a number of stimuli (Bylund, 1979) and are often inversely related to the effective concentration of norepinephrine in the synapse. For example, up-regulation of receptor number which is frequently associated with functional supersensitivity appears to result from a decrease in the level of norepinephrine or from chronic receptor blockade by appropriate antagonists. Conversely, a down-regulation in receptor number and functional subsensitivity is often associated with an increase in the level of norepinephrine or with chronic administration of an adrenergic agonist. This type of regulation is termed homologous regulation because it seems to be the direct result of changes in the amount of neurotransmitter or hormone available for binding to the receptor. By contrast, regulation of adrenergic receptors and their function by other hormones or regulatory agents is termed heterologous regulation. Adrenergic receptors also undergo changes during alterations in physiological conditions such as development, as well as in some pathological states. In order to assess the significance of changes in receptor number, it is critical that biochemical and physiological determinants be studied in addition to receptor binding. For instance, in the rat submandibular gland surgical denervation or reserpine treatment results in a doubling of &receptor number which is associated with a marked increase in the accumulation of cyclic AMP in response to isoproterenol (Bylund et al., 1981). However, the isoproterenol-stimulated adenylate cyclase activity is not altered, and the increased cyclic AMP levels are actually due to a decrease in the activity of phosphodiesterase. Thus, the change in receptor number does not appear to be physiologically important. Although considerable progress has been made in the past several years in the area of a,-adrenergic regulation, most of this work has been limited to receptor binding studies, and in the future much more emphasis must be placed on the corre-

CHARACTERIZATION OF CX1- AND a2-ADRENERGIC RECEPTORS

369

lation of receptor binding data with physiological and biochemical alterations. 1. Up-Regulatim In several systems the number of a,-adrenergic receptors (as labeled by [3H]WB4101) is increased following drug treatments or procedures designed to lower the effective concentration of norepinephrine at the synapse. A summary of these data is given in Table IX. Chemical sympathectomy in the central nervous system of the rat with the neurotoxin 6-hydroxydopamine drastically reduces norepinephrine levels in certain brain regions due to a destruction of the nerve terminals. The drug reserpine also markedly lowers norepinephrine levels. This reduction in neurotransmitter is accompanied by, and may result in, a modest increase in the levels of a,-adrenergic receptors. Chronic treatment with amitriptyline, a potent a ,-adrenergic antagonist, causes a similar increase in the density of receptors. In the submandibular gland of the rat an up-regulation in a-adrenergic receptors is seen following a surgical denervation or norepinephrine depletion following chronic reserpine treatment. None of the studies referenced in Table X found a significant change in the K , value for [3H]WB4101. It is of interest that in these experiments, the magnitude of the changes in a,-receptor levels appears to be generally less than that for either p- or a2-adrenergic receptors. For example, in the submandibular gland, norepinephrine depletion results in approximately a 10-fold increase in a2-adrenergic receptors (Bylund and Martinez, 1980, 1981). In the brain neonatal 6-hydroxydopamine treatment results in a 16% increase in the density of a , receptors, but causes a 40% increase in the number of P-adrenergic receptors as determined by [3H]dihydroalprenolol binding (D. B. Bylund and w. 0. Shekim, unpublished). However, in general the changes are similar in magnitude. It also should be pointed out that there are multiple examples of the lack of change of a , receptors following treatments similar to those in Table IX. I n the salivary gland initial attempts have been made to correlate changes in receptor binding with the release of potassium induced by a1 agonists. I n one study the density of a,-adrenergic receptors increased 55% following 7 days of reserpine treatment, while the release of potassium increased 56 and 37% in response to norepinephrine and methoxamine, respectively (Bylund et al., 1981). Thus, in this case the increase in receptor binding and the biochemical effect were approximately equal, suggesting a possible causal relationship. However, other workers have failed to fmd an increase in potassium release following

Tissue“

Treatment

H,,,,,for [9H]WB4101 (percentage of control)* 124

Submandibular Submandibular Forebrain Forebrain Forebrain (mouse) Cerebral cortex Cerebral cortex

Canglionectomy Reserpine Reserpine Amitriptyline Amitriptyline Neonatal 6-hydroxydopa

116

Midbrain

Neonatal 6-hydroxydopa

109

Frontal cortex Thalamus

Dorsal bundle lesion Dorsal bundle lesion

7

146 151

129 149

6-W ydroxydopamine

~

180 177

155

Norepinephrine (percentage of control)

17

174 120

3 21

Reference Bylund rf nl. (1981) Bylund ct nl. (1981) U’Prichard and Snyder (1978b) U’Prichard rt (11. (197813) Rehavi uf al. (1980a) U’Prichard et a/. ( 1979a) D. B. Bylund and W. 0. Shekim (unpublished) D. B. Bylund and W. 0.Shekim (unpublished) U’Prichard et (11. ( 1980b) U’Prichard rf al. (1980b)

~~~~

Tissues were from the rat unless otherwise indicated. * B,,,, values were calculated from saturation experiments. In all cases, the value for treated animals was significantly different from control (p < 0.05). The first column is percentage calculated from the data expressed as pmoVgm tissue and the second column from the data expressed as pmol/gm protein. a

37 1

CHARACTERIZATION OF al- AND QI~-ADRENERGICRECEPTORS

TABLE X BINDING CHARACTERISTICS OF P H ] P R A Z O S I N T O R A T LUNGMEMBRANES AT DIFFERENT TIMES OF THE DAY

Time

No. of animals

B,,, (fmoYmg prot)

K D (nM)

132 f 3 125 f 4 119* 2 129 6

0.105 f 0.004 0.109 0.012 0.110 f 0.012 0.130 f 0.011

126 2 2

0.114 2 0.006

2 A.M. 2

P.M.

8

P.M.

6 6 6 5

Mean

23

8 A.M.

*

*

surgical denervation of the submandibular gland (Arnett and Davis, 1979) and the parotid gland (de Peusner et al., 1979). In contrast to findings (see below) in which an increased level of the agonist decreases a,-adrenergic response and receptor binding, it appears that under certain conditions a brief pretreatment with an a agonist can increase the response and density of receptors (Hata et al., 1980a,b).In the rat vas deferens, the contractile response to epinephrine was significantly enhanced by preexposing the tissue to 25 p M epinephrine for 10 min. This treatment increased the maximal contractile response by 39%, but did not affect the EDm.The number of al receptors, as measured by [3H]WB4101,increased 27% following the epinephrine treatment. The increased contractile response was blocked by phentolamine but not by propranolol, suggesting a specific a-adrenergic phenomenon. There is preliminary evidence that chronic lithium treatment increases a,-adrenergic receptor binding. Rats that had been on a lithium diet for 3 or 5 weeks had 28 and 17% increases, respectively, in the binding of rHlWB4101 when assayed at a single concentration. This increase appeared to be due to a B,,, change with little change in K , (Rosenblatt et al., 1979).

2 . Down-Regulation The term desensitization is used to describe the phenomenon of reduced cellular responsiveness to an agonist following a previous exposure to that agonist. Down-regulation of receptor number often accompanies desensitization and in at least some systems seems to be an important mechanism for producing the desensitized state. Desensitization of a-adrenergic receptors has been studied in rat parotid acinar cells. In these experiments the release of potassium in response to epinephrine in cells that had been preexposed to epinephrine was reduced compared to

372

DAVID B. BYLUND AND DAVID c . U’PRICHARD

control cells. Since there was concurrent reduction in rH]DHEC binding to membranes prepared from these cells, it was concluded that the a-adrenergic desensitization was mediated, at least in part, by a downregulation in receptor number (Strittmatter et al., 1977; Davis et al., 1980). These studies are hard to interpret in the context of the present article since the binding of E3H]DHEC to a , and a 2 receptors was not differentiated. Indirect evidence in support of the ability of a 1 receptors to be down-regulated includes observations of reductions in rat cerebellar rH]WB4 101 sites after dorsal bundle lesion, which elevates cerebellar norepinephrine by 50% (U’Prichard et al., 1980b), and reductions in rat heart and vas deferens rHlWB4 101 binding after chronic immobilization stress, which accelerates norepinephrine turnover and release (U’Prichard and Kvetnansky, 1980).

3. Phjsiologxal Regulation a. Ontogmy T h e ontogeny of a,-adrenergic receptors has been described in four tissues: heart, brain, submandibular gland, and lung. Figure 1 illustrates the change in the density of a,-adrenergic receptors during early postnatal development. In addition, norepinephrine levels are given over the same time period for all four tissues. For the submandibular gland the development of the biochemical effect (potassium release) is also given. There were no significant changes in K , ) values noted in any of the four tissues during development. In the mouse heart, a , receptors labeled by [3H]WB4101 are significantly higher during the first 2 weeks of life than they are in either the fetus or the adult (Yamada et al., 1980b). These data were interpreted to suggest that the receptors mature prior to the development of the sympathetic innervation and then decrease markedly with the innervation of the tissue. It is of interest to note that between 2 and 3 weeks, when there is the greatest increase in norepinephrine levels, the number of a,adrenergic receptors is decreasing. This may indicate that the receptors are down-regulating in response to the increased norepinephrine levels. T h e levels of a,-adrenergic receptors Hlprazosin binding) in the rat brain increase relatively uniformly to a peak at 3 weeks of age and then drop off slightly to adult levels. T h e concentration of norepinephrine increases at a somewhat slower rate and even by 3 weeks is only about 75% of adult levels. However, a direct comparison of norepinephrine levels and receptor binding in whole brain is of doubtful significance since most of the norepinephrine is in the hypothalamus, while the a,-adrenergic receptor binding reflects the predominance of cerebral cortical tissue.

(r

CHARACTERIZATION OF (Y1- AND CQ-ADRENERGIC RECEPTORS

373

-I

W

I

>

W

-I

W

z

X

CI

U I m

E

I

a

W

z E

W K 0

z

AGE (days) FIG. 1. Comparison of a,-adrenergic receptor density (BmaX)and norepinephrine levels in four tissues during development. For the rat submandibulargland, a comparison with epinephrine-stimulated K+ release is also included as an indication of al-receptor function.

374

DAVID B. BYLUND AND DAVID

c.

U’PRICHARD

T h e a ,-adrenergic receptors ([3 Hlprazosin binding) in the rat submandibular gland are similar to those in the rat brain in that they are barely detectable at birth and show the greatest increase between 1 and 3 weeks of age (Bylund rt nl., 1982b). T h e al-stimulated release of potassium is also very low at birth, but then increases to adult levels between 1 and 3 weeks of age (Martinez and Camden, 1982). Similarly, the greatest increase in norepinephrine levels also occurs between 1 and 3 weeks of age (Kuzuyact al., 1980). Thus, in this tissue there is a remarkably good correlation between the innervation as measured by norepinephrine levels, the development of receptor binding sites, and the development of the biochemical response. In the rat lung, the density of a,-adrenergic receptor binding sites ([3H]prazosin binding) is moderately high at birth and then slowly increases to a peak at 2 1 days which is approximately 50% higher than the level at birth (Latifpour, 1981). Norepinephrine levels in the rat lung also peak at about 3 weeks at a level approximately three times that at birth (Gardey-Levassort et nl., 1981). T h e comparison of the development of a,-adrenergic receptors in these four tissues fails to reveal a consistent pattern, but rather indicates that each tissue has a unique developmental pattern which is presumably consistent with the function of the cells and tissues involved. b. Circcicliu?i Rhythm. T h e possibility of a circadian variation in a l adrenergic receptor binding has been investigated in the rat brain and lung. A circadian rhythm was found in the rat brain with a variation of 30-40% in binding of [3H]WB4101 over a 24-hr period (Kafka et ul., 1981). Saturation analyses conducted at the time of maximal and minimal binding indicated the change in receptor binding was due to a change in B,,,, and not in K,. In addition, both the wave-form of the circadian rhythm and the time of maximal binding were dependent on the month in which the assays were conducted, suggesting that a seasonal rhythm may also be present. If confirmed, these observations have serious implications for design and conduct of experiments intended to investigate changes in receptor levels. As a minimum, the experiments must be conducted at the same time each day. However, even this may not be sufficient since chronic drug treatment appears to modify the circadian rhythm in the rat brain (Wirz-Justice et al., 1980). T h e physiological significance of this circadian rhythm cannot be ascertained, since there were no concomitant studies on the functional effects of the receptor-agonist interaction. By contrast, no circadian variations were found in the binding of pH]prazosin to rat lung membranes (Latifpour, 1981). In these experiments rats were sacrificed over a 3-day period every 6 hr, and the K , , and

CHARACTERIZATION OF a1- AND a2-ADRENERGIC RECEPTORS

375

B,,, were determined by saturation analyses. The maximum deviation of B,,, at any time point from the mean of all time points was only 5 % . These results suggest that rat lung a,-adrenergic receptors do not undergo any circadian changes; these results also illustrate that highly reproducible data can be obtained in carefully controlled experiments (Table X). 4 . Pathological Regulation

There is increasing evidence that variations in receptor characteristics may be the result of, result in, or be associated with certain disease states. For the a,-adrenergic receptor, the pathological state that has received the most attention is hypertension. The results of these studies are summarized in Table XI. In the heart of the deoxycorticosterone/salt hypertensive rat, the density of a ,-adrenergic receptors as determined by rHlWB4101 was significantly reduced in the ventricles but not in the atria (Yamada et al., 1980d). Cardiac a-adrenergic receptors were also found to be decreased using the subtype nonselective ligand HIDHEC (Woodcock and Johnston 1980).The density of a,-adrenergic receptors was similarly reduced in the kidney using both the spontaneously hyperactive rat and the deoxycorticalstarone/saltmodels of hypertension (Yamada et al., 1980d; U’Prichard et al., 1979b).In the brain the levels of a1 adrenergic receptors are generally increased (U’Prichard et al., 1979b), although this is dependent upon the particular brain region studied. For instance, significant increases in B,,, were found in both the cerebral cortex (Yamada et al., 1980d) and the midbrain (Gheyouche et al., 1980),although no change was found in other brain regions includ-

r

ALTERATIONS IN a

l

-

TABLE XI A RECEPTORS ~ ~ IN~ ANIMAL ~ ~ MODELS ~ OF~ HYPERTENSION ~ Bmax

Tissue

Hypertensive model

Heart Kidney Kidney Cerebral cortex Brain Midbrain

Deoxycorticosteronehalt Deoxycorticosteronekalt Spontaneously hypertensive Deoxycorticosteronehalt Spontaneously hypertensive Spontaneously hypertensive

for

[3H]WB4101 (percentage of control)’ 67 71 74

62 75 125

119 135

Reference Yamada et al. ( 1980d) Yamada et al. (1980d) U’Prichard et al. (1978b) Yamada et al. (1980d) U’Prichard et al. (1978b) Gheyouche et al. (1980)

B,,, values were calculated from saturation experiments. The first column is percentage calculated from the data expressed as pmol/gm tissue and the second column from the data expressed as pmoVmg protein. (I

376

DAVID B. BYLUND AND DAVID

c . U’PRICHARD

ing the hippocampus, hypothalamus/thalamus, cerebellum, and brainstem (Yamadaet ul., 1980d; Cantor et al., 1981). While these results suggest that a ,-adrenergic receptors may have a role in hypertension, it is not clear even in the animal models whether the alterations in receptor number are involved in the etiology of the hypertension or are only a consequence of other compensatory changes. D.

SOLUBILIZATION OF

QI I-ADRENERGIC

RECEPTORS

The solubilization and purification of the a,-adrenergic receptor will ultimately be necessary in order to have an accurate understanding of its characteristics and function. There are two reports of a,-adrenergic receptors that have been solubilized in the active form. Using the detergent digitonin a- and P-adrenergic receptors were solubilized from hepatic plasma membrane and were then identified by the binding of [3H]WB4101 and ~H]dihydroalprenololand characterized (Wood P t nl., 1979b). These workers were then able to separate the solubilized receptors by affinity chromatography and concluded that the active a- and P-adrenergic binding sites d o not simultaneously reside on the same macromolecule. Unfortunately, only 3 to 10% of the receptors present in particulate preparation were solubilized, and those solubilized receptors had a 2- to 8-fold decrease in affinity for adrenergic drugs. Solubilized hepatic a 1 receptors have been purified about 500-fold using affinity chromatography (Graham et al., 1982). T h e specificity of binding to these receptors is similar to that of the membrane-bound receptors as determined by the inhibition of PHlprazosin binding. However, the K , for [3H]prazosin (16 n,M) in saturation experiments is about 30-fold higher than in membranes. Taking a different approach, Guellaen et al. (1979, 1982) first prelabeled the receptor with the irreversible a antagonist [3H]phenoxybenzamine, and then solubilized with 2% SDS or 0.5% Lubrol PX. These workers concluded that the rat liver a 1receptor has a molecular weight of 96,000 and is composed of at least two subunits. The binding site (at least for [3H]phenoxybenzamine) is on a 44,800-dalton subunit.

111. cuZ-AdrenergicReceptors

In general, the a2receptor and its relation to the coupled response of adenylate cyclase inhibition has been much better characterized than the

CHARACTERIZATION OF (Y1- AND (Y2-ADRENERGIC RECEPTORS

377

receptor, especially in well-defined homogeneous cell populations such as human platelets, adipocytes, and rodent neural cell lines in culture. The a , receptor appears to occur in two or more states, primarily differentiated with respect to their affinity for agonists. According to the model of a,-receptor function proposed by Lefkowitz and co-workers (Hoffman et al., 1980a), antagonists recognize both states of the receptor with equally high affinity, whereas agonists have high affinity for only one of the states. Thus, in typical experimental situations, labeled agonists bind to a subset of the total a,-receptor population which is determined by labeled antagonist binding. The data are generally consistent with the Lefkowitz model, but recurrent inconsistencies are illustrated which tend to suggest that the model is an oversimplification. This section of the article will deal with the characteristics of antagonist and agonist binding sites representing a, receptors, the comparison of agonist and antagonist binding within the framework of regulation of different affinity states of the receptor, and the relationship of this regulation to transduction of the a,-receptor response. Regulation, localization, and solubilization of a2 receptors will then be considered.

a

A. CHARACTERIZATION BY RADIOLIGAND BINDING 1. Antagonist Radioligands a. [3H]Dihydro~g~cTyptine. The first tissue in which VHIDHEC binding to a receptors was demonstrated was rabbit uterus (Williams and Lefkowitz, 1976), which has been estimated to contain predominantly (80%) a , receptors (Hoffman and Lefkowitz, 1980b). Subsequently, [3 HIDHEC binding was characterized in human and rabbit platelets (Kafka et al., 1977; Newman et al., 1978; Tsai and Lefkowitz, 1978), which appear to contain exclusively a, receptors, and in rat and bovine brain membranes (Greenberg and Snyder, 1978; Peroutka et al., 1978), which have roughly equal amounts of a1 and a, receptors depending upon the specific brain region. Although the K Dof brain [3H]DHECbinding was reported to be about 1.0 nM (Greenberg and Snyder, 1978), other early studies of VHIDHEC binding gave generally higher and quite variable KD values, ranging from 3-35 nM (L. T. Williams and Lefkowitz, 1976; Kafka et al., 1977; R. S. Williams and Lefkowitz, 1978). To some extent, K , values appeared elevated because receptor concentrations in small assay volumes were quite high (Williams and Lefkowitz, 1976; Kafka et al., 1977). More recent studies of VHIDHEC binding to a receptors in many different tissues have employed lower receptor concentrations and demonstrate considerable agreement for the affinity of

378

DAVID

B. BYLUND

A N D DAVID

c:. U’PRICHAKD

[3H]DHEC at a-receptor binding sites, with K, values in the range of 1-3 n2W. [3 H]Dihydroergocryptine labels both a , and a,-receptors. In several studies, the number of PHIDHEC sites was found to be approximately equal to the sum of sites labeled by ligands more specific for the a 1and a , receptors, respectively, such as PHIWB-4101 and [3H]clonidine in rat brain and peripheral tissues (U’Prichard and Snyder, 1979; Neethling et ul., 1981). I n retrospect, these studies can be criticized because the a,receptor ligand used, [3H]clonidine, is an (partial) agonist and as such labels only a subpopulation of the a,-receptor sites (rather than the total a,-receptor pool), thereby underestimating a,-receptor density. In a more recent study (Lavin et ul., 1981),the number of a , receptors labeled by E3H]DHEC in the rabbit uterus was 80% higher than the number labeled by Hlyohimbine (see Section II,A,l ,b). Similarly, in platelets the number of PHIDHEC sites (all az)is 65% higher than PHIyohimbine sites (Hoffman et ul., 1982). On the other hand, in the rabbit uterus good agreement was found between the affinities of various antagonists at a2 sites labeled by [3H]yohimbine and by PHIDHEC. In order to determine reliable affinities for the a , and a, subtypes using PHIDHEC, a 100-fold selectivity of a competitor is required along with sophisticated computer analysis of the data (Lavin et ul., 1981). Furthermore, it is assumed that [3H]DHEC labels a1 and a , receptors with equal affinity. In almost all tissues studied, Rosenthal plots of [3 HIDHEC saturation data appear linear, which is consistent with this assumption. However, the data generated with this ligand are generally quite scatter prone, and few studies have attempted rigorously to demonstrate that in the same membrane preparation PHIDHEC has equal affinity for a 1 and a2 receptors. One early examination of [3 HIDHEC saturation in bovine cortex membranes in the presence of receptor-saturating concentrations of unlabeled a1and a , competitors did suggest that PHIDHEC is equipotent at the two receptors (Peroutka rt ul., 1978). Another, as yet unreplicated, analysis of saturation of rat heart [3H]DHEC binding in the absence and presence of the a,-specific antagonist competitor ARC 239 indicated that [3 HIDHEC exhibited “positive homotropic cooperativity” of binding at myocardial a,, but not a l , receptors (Guicheney lit nl., 1978). Despite the limitations of PHIDHEC as a radioligand, such as its nonselectivitv and its poor binding to a receptors on intact cells such as human platelets (Alexander et al., 1978), and the current availability of yohimbine isomers as more selective a,-receptor antagonist radioligands (Motulsky and Insel, 1982), PHIDHEC has been a useful tool in the analysis of a,-receptor function in human platelets, which contain a pure

r

CHARACTERIZATION OF CY1- AND ff2-ADRENERGIC RECEPTORS

379

population of a, receptors. Platelet [3 HIDHEC binding, extensively studied by Lefkowitz and colleagues, is homogeneous with respect to antagonist competitors, but heterogeneous with respect to agonists. That is, the pseudo-Hill coefficients (nH)for antagonists are about 1.0, but the n H values for agonists are significantly less than 1.0 (Tsai and Lefkowitz, 1979). The favored explanation was that [3 HIDHEC binds to distinct conformations of the az receptor which recognized agonists, but not antagonists, with different affinity, rather than that [3H]DHEC exhibits negative cooperative interactions with the receptor. Sodium affects the interactions of agonists at the platelet rH]DHEC site by reducing their apparent affinity (right shift of curve). T h e extent of this right shift correlates with the known intrinsic activities of a series of a-receptor agents in a fashion reminiscent of the “sodium shift” at opiate receptors (Tsai and Lefkowitz, 1978). Alpha receptor antagonists interacting at the platelet rH]DHEC site are unaffected by Na+. Divalent cations such as Mg2+ selectively increase agonist affinities at these sites. These data are analogous to the existence of conformations of the P-adrenergic receptor with high and low affinity for agonists, whose equilibrium is modulated by guanosine di- and triphosphates (Maguire et al., 1977). T h e platelet az receptor, like the P receptor, is coupled to adenylate cyclase, but in an inhibitory manner. GTP and the nonhydrolyzable analog guanyl-5’-yl imidodiphosphate [Gpp(NH)por GMP-PNP] shift competition curves to the right for agonists but not antagonists at platelet THIDHEC sites, whereas corresponding adenine nucleotides are ineffective. In addition, in the presence of guanine nucleotides, agonist n H values approach 1.0, suggesting increased homogeneity of agonist interactions at the receptor (Tsai and Lefkowitz, 1979). In more recent studies, the techniques of computer modeling of platelet rH]DHEC competition curves indicate that agonist interactions at this site are best fit to a two-affinity-state model, designated az(H) and a,(L) (high and low affinity for agonists, respectively). Hoffman et al. (1980a) demonstrated that antagonists such as phentolamine have equal affinity for aZ(H)and a,(L). Relatively more a2(H) are found in the presence of the full agonist (-)-epinephrine than the partial agonist methoxamine. Guanine nucleotides at saturating concentrations (100 p M ) shift all the receptors into the az(L)state. The inability of partial agonists to generate as many a , ( H ) explains the apparent lesser effect of guanine nucleotides on partial agonist competition curves compared to those of full agonists. In somewhat less elaborate studies, the azcomponent of [3H]DHEC binding to rat liver membranes was also shown to be GTP-sensitive (Hoffman et al., 1980a). These experiments strongly indicate that the platelet a , receptor is coupled to adenylate cyclase via an intermediate membrane protein(s) with a site of at-

380

DAVID B . BYLUND A N D DAVID c . U’PRICHARD

tachment for GTP, in a manner analogous to /3 receptors that stimulate adenylate cyclase activity. HIDihydroergocryptine has also been shown to label a2-receptor sites on membranes from a neuroblastoma X glioma hybrid clonal cell line NG 108-15 with high affinity ( K , = 1.8 nM). At these sites, agonist affinities are affected by GTP in the same manner as in platelets (Haga and Haga, 1981). 6. [3H]Ebhim6ineand ~ H ] R a u w o l s c i n T ~ .h e alkaloid yohimbine is an a antagonist with selective potency for agreceptors in pharmacological experiments (Hedler et al., 1981). [3H]Yohimbine binding to a2receptors has been demonstrated in membranes from a variety of tissues (Table XII). In saturation studies, the KD of PHIyohimbine ranges from about 0.3 to 11 nil4 depending on species, tissue, buffer, and assay conditions. The affinity increases about threefold in most tissues when glycylglycine buffer is used rather than Tris-HC1 or Tris-NaC1 buffers. In glyclyglycine buffer, the KD for human and pig tissues is about 0.35 nM, while in rat and guinea pig tissues it is around 1.5 nM. For most a2 receptors, the data obtained using P Hlyohimbine are much superior to the data with PHIDHEC. There is good concurrence about the basic properties of PHIyohimbine binding to human platelet membranes in five different laboratories (Daiguji et al., 1981a; Motulskyrt al., 1980; Mukherjee, 1981; Smith and Limbird, 1981; Garcia-Sevilla et al., 198lb). Both kinetic and equilibrium experiments performed at 25 or 37°C indicate that rHlyohimbine binds to a single population of sites with the same affnity over a 0.1-10 nM concentration range. T h e pharmacological properties of platelet P Hlyohimbine sites determined from competition analyses are very similar to those of platelet [3 HIDHEC sites and indicate an a2-receptor interaction, with a catecholamine potency order (-)-epinephrine > (-)-norepinephrine % (-)-isoproterenol, selectively high affinity of (-)catecholamine isomers, and yohimbine generally about 500- 1000 times more potent than prazosin (see Table XIII). As with platelet PHIDHEC sites, agonist competition curves have nHvalues significantly less than 1.0. However, competition curves for imidazoline partial agonists are somewhat steeper and antagonists interact in a homogeneous manner at platelet [3H]yohimbine sites with n H of about 1.0. Guanine nucleotides right shift and Mg2+ left shift agonist and partial agonist competition curves, so that the apparent effect of GTP is more pronounced in the presence of Mg2+ (Fig. 2). Hoffman et al. (1982) have suggested that, as for platelet [3 HIDHEC binding, agonist competition at platelet PHIyohimbine sites best fits a two-site model, and that in the absence of nucleotide, both catecholamine full agonists and imidazoline partial agonists (intrinsic activity determined using the response of inhibition of

TABLE XI1 [3H]YOHIMBINEBINDING TO aZ RECEPTORS ~~

Tissue"

Species

pmoVgm protein

Platelet Platelet Platelet Platelet Platelet Platelet Platelet Platelef Adipocytes" Adipocytes' Cerebral cortex Cerebellum Uterus Cerebral cortex Cerebral cortex Liver Lung (neonatal) Kidney Kidney Cerebral cortex Cerebellum Corpus striatum Neuroblastoma X glioma

Human Human Human Human Human Human Human Human Human Human Human Human Rabbit Pig Guinea pig Rat Rat Rat Rat Rat Rat Rat

182 ? 29 334 ? 161 191 f 23 188 2 12 138 -+ 13 422 f 22 265 2 12

Rat-mouse

pmollgm tissue

K , (nM)

Buffefl

Reference

1 1 1 1 1 2 1 1 2 1 2 2 1 2 2

2

Daiguji et al. (1981a) Motulsky et al. (1980) Mukherjee (1981) Garcia-Sevilla et al. (198lb) Hoffman et al. (1982) D. B. Bylund (unpublished) Smith and Limbird (1981) Motulsky et al. (1980) Burns et al. (1982a) Tharp et al. (1981) D. B. Bylund (unpublished) D. B. Bylund (unpublished) Lavin et al. (1981) Harris et al. (1983) D. B. Bylund (unpublished) Hoffman et al. (1981a) Latifpour (1981) Yamada et al. (1980a) Schmitz et al. (1981) D. B. Bylund (unpublished) D. B. Bylund (unpublished) D. B. Bylund (unpublished)

1

Kahn et al. (1982)

170 f 10 120 2 16 37 t 4 106 f 6

7.5 f 1.0 2.5 I0.3 6.8 f 0.6

1.25 2 0.10 2.8 2 0.9 1.7 2 0.2 3.0 f 0.1 1.5 f 0.1 0.38 k 0.01 5.7 f 0.4 2.7 f 0.7 0.39 2 0.02 3.9 2 2.4 0.46 ? 0.05 0.33 2 0.03 llf5 0.27 f 0.02 1.7 f 0.3 5 1.53 f 0.11 0.83 f 0.08 7.4g 2.2 f 0.3 1.1 f 0.1 1.3 rL- 0.1

258 f 83

22,600 f 5,OOod

9.1 2 1.1

*

543 99 145 f 34 201 f 18 54 f 5 72 f 19 167 ? 21 95 f 9 110 f 21 304 2 28

207 f 41d

122 1 3.3 2 0.3 8.3 ? 1.0 5.5 2 0.2

4.8 f 0.2

Membrane preparation, unless otherwise noted. Buffers: 1, Tris; 2, glycylglycine; 3, sodium potassium phosphate. Intact platelets. Number of sites per cell. Properitoneal tissue. Subcutaneous tissue. Determined by kinetic analysis.

2 3 3 2

2

382

DAVID B. BYLUND A N D DAVID A

2

c. U'PRICHARD

Conhol

eo-

E

j

60-

4020-

n

0-

-

100

2

:!I [40

P

#

20 -I

O 1 0

9

8

7

6 -log [PENTOCAMNI (Mi

FIL. 2. Inhibition of [3H]yohimbine binding (0.2 n N ) to platelet membranes at 25°C (60 min) by (-)-epinephrine, p-aminoclonidine, and phentolamine in the presence or absence of GTP and MgC12. Points represent mean data from four to six experiments.

platelet adenylate cyclase) cause 60-70% of the receptors to be in the az(H) state. These authors found a correlation between intrinsic activity

and the ratio of affinities of agonists for a 2 ( H ) and a 2 ( L ) ,so that the K , , values for (-)-epinephrine were 11 nM (H) and 520 nM (L) with a ratio of 47.3, whereas the corresponding values for clonidine (59% intrinsic activity) were 9 nM (H) and 110 nM (L) with a ratio of 12.2. Unlike the

CHARACTERIZATION OF al- AND Q2-ADRENERGIC RECEPTORS

383

corresponding analysis of frog erythrocyte /3 receptors (Kent et al., 1980), intrinsic activity of agonists at platelet a2 receptors labeled with [3 Hlyohimbine did not correlate with the number of receptors in the a2(H) state. Similarly, submaximal concentrations of Gpp(NH)p appeared to reduce the afhity of (-)-epinephrine at a2(H)sites, rather than simply convert a2(H)to a2(L),as was found for the corresponding p-receptor analysis (Kent et al., 1980). In some respects, platelet p Hlyohimbine and ["HIDHEC sites differ. As noted earlier (Section III,A,l ,a), p HIDHEC consistently labels more platelet sites than PHlyohimbine (Daiguji et al., 1981a; Motulsky and Insel, 1982; Hoffman et al., 1982),a fkding that would not be predicted from the Hoffman and Lefkowitz (1980a) model, since both drugs, as antagonists, should label equally well both a2(H)and a 2 ( L )states of the receptor. The excess rH]DHEC binding does not appear to be to a1 receptors or serotonin transport sites. Earlier studies had shown that neither metal cations nor guanine nucleotides affected platelet THIDHEC binding. However, Mg2+concentrations as low as 1.0 mM increased the KD of p Hlyohimbine without apparently changing the B,,, (Daiguji et al., 1981a), and 50- to 100-mM Na+ decreased the KD and increased the B,,, of [3H]yohimbinebinding to solubilized membranes (L. E. Limbird, personal communication). Although these results cannot be explained at the present time, there is the possibility of some selectivity in the potency of the antagonist [3H]yohimbinefor different states of the a2 receptor (see Section 111,C). Another cell type that lacks a1 receptors but contains a2 receptors and the associated response of adenylate cyclase inhibition is the cultured hybrid neural cell NG 108-15 (Kahn et al., 1982). THlYohimbine binds in an apparently monophasic fashion to NG 108-15 membrane sites that have the pharmacological properties of at receptors and closely resemble platelet at receptors (Table XIII) except that yohimbine itself is about eightfold less potent in saturation and inhibition experiments, and the yohimbine/prazosin potency ratio is higher at NG 108-15 compared to human platelet sites by two orders of magnitude (Table 111). THlYohimbine labels about 30,000 a2 receptors per NG 108-15 cell, which is about the same density of receptors per platelet (c.a. 100-200 receptors per cell) when the calculations are made on the basis of receptors per unit surface area of cell plasma membrane. As with platelets, PHIyohimbine sites on NG 108-15 membranes exhibit heterogeneity with respect to agonist competitors (nH = 0.5-0.7) but not antagonists, and GTP and Gpp(NH)p at high concentrations (50 p M ) right shift and steepen agonist competition curves, supporting the general concept of the existence of (H) and (L) states of the a2receptor on these neural cells

384

DAVID B. BYLUND AND DAVID

I S H I R I T I O S OF

TABLE XI11 3H-LABEI,ED ALOSISTBINDING TO (I-ADRENERGIL RECEPTORS"

r3 HlEpinephrine (ICm, n.tf)

Drug Agonists (-)-Epinephrine (-)-Norepinephrine ( +)-Norepinephrine ( -)-lsoproterenol Imidazolines p-Aminoclonidine Clonidine Antagonists Phentolamine Yohimbine WB4101

Prazosin

c. U'PRICHARD

Platelet

3.3

14 410 1,500

3.3 3.8 11 19 20,000

NG-108

P H ] p Aminoclonidine (lCm,ni\J) Platelet

NG-108

[3H]Yohimbine V C W ,nM) Platelet

NG-108

11 6.1 32 270

2.4 6.5 250 800

5.4 3.0 90 60

87 420 6,800 29,000

250 390 1,900 29,000

4.0 32

3.4 3.1

1.3 30

34 60

52 48

65

140 3 50 5500

2.9 17 26 29,000

22 36 120 2,100

4.3 1.o

2.8 430

39 8 16 42

" Assays were performed on membranes from outdated platelets at 25°C. 1.0-mM MgCI, was present in EPI and PAC assays. Ligand concentrations were VHIepinephrine 1.0 nhl; [3H]p-aminoclonidine 0.6 nM; [3H]yohimbine 0.2 nJf . IC,, values are the mean of four to eight experiments.

(Kahn ei nl., 1982). Interestingly low (1.0-10 p L b f )concentrations of nucleotides seem to increase agonist affinities somewhat at [3 Hlyohimbine sites in membranes when residual endogenous divalent cations are not removed by treatment with a chelator. The imidazolines, clonidine and p-aminoclonidine, are partial agonists with respect to inhibiting NG 108-15 adenylate cyclase (Kahn et a/., 1982; Atlas and Sabol, 1981), showing, as in platelets, about 50% of the efficacy of full (catecholamine) agonists. T h e potencies of agonists and antagonists in general corresponded well at NG 108-15 r3H]yohimbine sites and the cyclase response. The bindinglcyclase potency ratio for agonists, K,Jk', , may be an index of the efficiency of coupling of the NG 108-15 a2 receptor to adenylate cyclase, by analogy to the K D / K a c ratio t for p receptors (Maguire et al., 1977). T h e K,IKi ratio was 1.0 for full agonists, but only 0.1 for the partial agonists clonidine and p-aminoclonidine (Kahn et al., 1982). Although these values are meaningful in a relative sense, as absolute indices they are suspect because both binding and cyclase assays w e r e performed in a cell-free system. A truer estimate of efficiency of coupling would involve determinations of

CHARACTERIZATION OF al- AND (Y~-ADRENERGICRECEPTORS

385

agonist potencies in intact cells at [3H]yohimbine binding sites and in decreasing cellular CAMPlevels. It is of interest that the pharmacologic characteristics of the human and rat a2receptors labeled by [3 Hlyohimbine are different, particularly with respect to the potencies of yohimbine and prazosin. I n human tissues, yohimbine is much more potent than prazosin, with yohimbine/ prazosin ratios about 0.002, whereas in rat tissues including NG 108-15 cells, yohimbine is only slightly more potent than prazosin, with ratios about 0.15. In glycylglycine buffer, the KD for PHIyohimbine binding is 0.32 to 0.46 nM in human tissues, whereas it is between 1.1 and 2.2 nM in rat. I n the fist report of PHlyohimbine binding in a rat tissue, the authors were concerned about the relatively high potency of prazosin and therefore did not claim that it represented binding to at receptors (Yamada et al., 1980a). However, according to the current definition of a2 receptors (see Section l,B and Table 111) and more extensive pharmacologic studies (Latifpour, 1981), it is correct to refer to rH]yohimbine binding in the rat and neuroblastoma NG 108-15 as a2.Whether this pharmacologic difference represents more than a species difference is not yet known. Rauwolscine is a diastereoisomer of yohimbine that is equipotent with yohimbine at a2 receptors in both functional (Hedler et al., 1981) and binding (Tanaka and Starke, 1980) studies, but is about 50 times less potent than yohimbine as an a,-receptor antagonist (Hedler et al., 1981). Thus, rauwolscine exhibits much greater selectivity toward the a2receptor than yohimbine, and potentially would be a more useful ligand than yohimbine in tissues containing a mixed population of a1 and a2 receptors since yohimbine only shows about a 10-fold preference for a preceptors in many tissues (Hedler et al., 1981; U’Prichard et al., 1977a). Two other yohimbine isomers, corynanthine and ajalmacine, are selectively potent at a1 receptors (Tanaka and Starke, 1980). [3H]Rauwolscine labels a2 receptors in human platelets (Motulsky and Insel, 1982) and rat and bovine cerebral cortex (Perry and U’Prichard, 1981, 1983). In platelets, [3H]rauwolscine and rH]yohimbine label the same number of a2 receptors, but the K, and nonspecific binding of [3 H]rauwolscine are somewhat greater, indicating that in this tissue it is no improvement over THIyohimbine. In bovine cortex membranes, [3 H]rauwolscine binding exhibits higher a h i t y in Na+containing buffers and in Na+/K+ phosphate buffer labels a single population of sites over a 0.1-15 nM concentration range with a KD of 1-2 nM from kinetic, saturation, or competition studies. The pharmacological properties of [3H]rauwolscine binding are those of an a2 receptor, with a yohimbine/prazosin potency ratio of about 0.003 (yohimbine Ki of

386

DAVID B . BYLUND AND DAVID

c. U'PRICHARD

4 n'\f). As with other a,-receptor systems, only agonists interact at the cortex rH]rauwolscine site in a heterogeneous manner (nH = 0.5-0.7), and as in platelets, both Na+ (60 m'tf) and GTP (100 p M ) right shift agonist competition curves, but had no effect on antagonist interactions. These data strongly indicate that the brain a , receptor also exists in (H) and (L) states, differentiated primarily with respect to agonists (Perry and U'Prichard, 1983). Some effects of cations and nucleotides on ~H]rauwolscinebinding are inconsistent with the hypothesis that antagonists have invariant affinity at all a,-receptor states (Perry and U'Prichard, 1983). T h e addition of NaCl increases [3 H]rauwolscine binding with a maximum effect at 50 m,\f (ED50= 5.0 mL\f),while higher Na+ concentrations bring binding back down to control levels. Chloride salts of other monovalent cations only decrease [3H]rauwolscine binding. GTP at low concentrations (0.1 nLllto 0.1 pc\I) also increases [3H]rauwolscine binding, even in the presence of 60 m.\l Na+. This effect is reproduced by Gpp(NH)p and GDP, but not adenine nucleotides and higher guanine nucleotide concentrations which inhibit p H]rauwolscine binding. Divalent cations, which appear to favor the formation of a z ( H ) decreased the binding of [3H]rauwolscine, with Mn2+being more potent than Mg2+or Ca2+.GTP at low concentrations produces a decrease in the ability of (-)epinephrine, and an increase in the ability of yohimbine, to inhibit [3 H]rauwolscine binding. These data suggest that [3 H]rauwolscine and other antagonists could have somewhat different affinities for different states of the a z receptor, and these differences might be detected in the presence of agents which alter the equilibrium between different receptor states (Perry and U'Prichard, 1983). Indeed, when [3H]rauwolscinebinding isotherms in phosphate buffer are extended to a ligand concentration of 50 nL\f,two sites are labeled with K , values of about 1 and 30 n,\l, respectively. The higher KIl value corresponds quite closely to the K , of rauwolscine at cortex sites labeled with [3 Hlagonists. c. [3H]L~\~~rzd~>. The ergoline derivative lisuride has a high affinity at brain a , receptors labeled with [3H]rauwolscine (Perry and U'Prichard, 1983), and it appears to have sufficient selectivity for the a, receptor compared to the a , receptor so that [3H]lisuride in appropriate conditions will specifically identify rat cortex a2 receptors (Battaglia and Titeler, 1980). T h e pharmacological profile of P Hllisuride binding indicates that the ligand, which is probably an antagonist at this receptor, labels both a 2 ( H )and a,(L). As with other antagonist ligands, GTP selectively lowers the affinities of agonist competitors. The usefulness of this ligand seems limited because, in common with other ergots, it also interacts with high affinity at other brain monoamine receptors.

CHARACTERIZATION OF

a1- AND

(Y~-ADRENERGICRECEPTORS

387

d. Intact-Cell Binding. I n homogeneous cell systems, it is advantageous to be able to label receptors on intact cells in order to relate binding to response in a more physiological situation. I n intact cell-binding studies, rHlyohimbine appears to label a2 receptors much better than rH]DHEC. The K , and B,,, of [3H]yohimbine are the same in intact platelet cells and platelet membrane preparations (Motulsky et al., 1980). In competition studies, antagonists had roughly the same potency in both preparations, but agonists were much weaker in inhibiting binding to intact cells and exhibited much steeper competition curves, with n H values approaching 1.O (Motulsky et al., 1980). Furthermore, the potency of (-)-epinephrine at Hlyohimbine sites on intact cells is equivalent to its potency at the membrane sites when assayed in a physiological (Hank’s) buffer (M. Daiguji and D. C. U’Prichard, unpublished). Like other adenylate cyclase-coupled receptors, a2 receptors would be expected to appear predominantly in a low-affiity state when assayed in intact-cell preparations because intracellular GTP (established to be about 10 p M ) would very rapidly destabilize high-affinity (complexed) states of the receptor (see below). In other words, the turnover of the high-affinity state of the receptor is exceedingly rapid under normal physiological conditions, but cell lysis and elimination of endogenous GTP “freezes” the a2 receptor in the proportion of (H) to (L) states existing in the membrane at that time. Thereafter, equilibrium between (H) and (L) states can be manipulated by exposing the membrane to receptor and N-site (nucleotide-binding regulatory protein) ligands, including guanine nucleotides, Mg2+, and more speculatively, Na+. Although the agonist epinephrine has the same potency at [3 Hlyohimbine sites on platelet membranes (in Hank’s buffer) and intact cells, GTP will only further reduce the potency of epinephrine in the membrane preparation, indicating that GTP modulat.es agonist affinities at the receptor by acting at a site on the inner surface of the plasma membrane (M. Daiguji and D. C . U’Prichard, unpublished).

2. Agonist Radioligands a. [3H]Epinephrine and [3H]Norepinephrine. Despite earlier difficulties in establishing receptor specificity of [3H]catecholamine binding, these radioligands have in the past few years proved to be viable and important probes for examining a,-receptor function. According to Lefkowitz, one would expect a [3 Hlcatecholamine at low concentrations to label selectively the (H) state of the a2 receptor and with increasing concentrations to identify an increasing amount of a2(L).Thus, Rosenthal transformations of [3H]catecholaminesaturation isotherms would be

388

DAVID B. BYLUND AND DAVID c . U’PRICHARD

expected to be curvilinear over a wide enough range of ligand concentration. Over a more restricted concentration range, however, Rosenthal plots would appear more linear, and the B,,, value derived from these plots would be more an approximation of the number of a2(H)than of the total number of a2 receptors in a tissue. Initial studies utilized racemic (t)-[3H]epinephrineof low specific activity (10-15 Ci/mmol), as well as (-)-PHInorepinephrine (20-40 Ci/mmol) (U’Prichard and Snyder, 1977a). More recently, the active isomer (-)-f Hlepinephrine at high specific activity (80- 120 Ci/mmol) has become available. Data obtained in the same tissue with these different [3 Hlcatecholamine preparations have been generally quite similar (U’Prichard et al., 1980a). Because labeled and unlabeled (Table 11) norepinephrine exhibit lower affinity than epinephrine for a2 receptors in most tissues, the use of [3H]norepinephrine has been very limited compared to [3H]epinephrine. Because of expense and technical difficulties, many fewer az-receptor-containing tissues have been examined with [3H]catecholamines compared to [3H]imidazolines. Care must be taken to prevent interference through binding of the ligand to p receptors since under typical experimental conditions [3H]epinephrine has selectively high potency for the p2 subtype (U’Prichard and Snyder, 1977b; U’Prichard et al., 1978~). On the other hand, at low ~H]catecholamineconcentrations, interference from a ,-receptor interactions is generally insignificant, apparently because the a,receptor does not usually occur in a high-affinity state (see Section II,A,2). Important precautions must be taken to minimize oxidation of the ligand and to include a large excess of pyrocatechol (0.1-3.0 m;M) to inhibit as much catechol-directed nonspecific binding as possible. [3H]Catecholamines have been used to label a2receptors in rat and bovine brain regions, human platelets, rat liver, and NG 108-15 cells (U’Prichard and Snyder, 1977a, 1978a; U’Prichard et al., 1983; Hoffman et nl., 1980b; Smith and Limbird, 1981; Kahn et nl., 1982). The characteristics of binding of [3H]epinephrine in these tissues is very similar. Generally, monophasic Rosenthal plots of saturation data are observed, giving equilibrium K , values of 1- 10 nM, although dissociation in the absence of nucleotides is biphasic at several incubation temperatures (U’Prichard and Snyder, 197713; U’Prichard et al., 1983). An exception is the dissociation of [3H]epinephrine from NG 108-15 membrane a2 receptors, which appears monophasic (Kahn et al., 1982). In human platelet and NG 108-15 cell membranes, the B,,,, for [3H]epinephrine binding is only 30-60% of the B,,, for an a2antagonist radioligand, which would be expected if at the fairly low concentrations used E3H]epinephrine was selectively labeling a2(H).

CHARACTERIZATION OF (Y1- AND (Y2-ADRENERGIC RECEPTORS

r

389

Analysis of Hlepinephrine competition in these tissues (Table XIII) shows that the ligand labels a, receptors, with yohimbine 100-1000 times more potent than prazosin. Competition studies also show that Hlepinephrine (in the 1-5 nM range) selectively labels az(H). Catecholamines compete in an a,-receptor potency order with (-)epinephrine and (-)-norepinephrine usually having Ki values in the 1- 10 nM range. Imidazoline partial agonists are also about 10-fold more potent competitors at Hlepinephrine sites than at [3 Hlantagonist sites (Hoffman et al., 1980b; U’Prichard et al., 1983), except in cerebral cortex membranes where these drugs have equally high potency (U’Prichard, 1980; Perry and U’Prichard, 1983). On the other hand, some antagonists such as phentolamine have equal potency at Hlepinephrine and [3 Hlantagonist sites, while other antagonists, including yohimbine, WB4101, piperoxan, and prazosin are 10-50 times less potent at [3H]epinephrine sites (U’Prichard et al., 1983). At Hlepinephrine sites, unlike [3 Hlantagonist sites, agonist and antagonist competitors are not discriminated in terms of nHvalues. In brain tissue and NG 108-15 cells, all drugs competed at az-receptor sites labeled with [3H]epinephrine, with n Hvalues of about 1.O (U’Prichard and Snyder, 1977a; Kahn et al., 1982), indicating that at the concentration used Hlepinephrine was almost exclusively labeling aZ(H)sites. However, in platelet membranes the n H values of agonists and antagonists are less than 1.0 at rH1epinephrine sites (U’Prichard et al., 1983). Binding of [3 Hlepinephrine and Hlnorepinephrine to a, receptors has been directly compared only in bovine cortex and rat liver (U’Prichard and Snyder, 1977a; El-Refai et al., 1979). In these tissues, both catecholamines appear to label an identical population of sites, although ( -)-[3H]norepinephrine had only two-thirds the affinity of (*)Hlepinephrine in bovine cortex in saturation experiments (U’Prichard and Snyder, 1977a). A detailed examination of the thermodynamic aspects of agonist and antagonist interactions with the a, receptor has not been undertaken in the same manner as for the p receptor (Weiland et al., 1979). However, the limited information available suggests that for brain az receptors at least, agonist and antagonist interactions can be discriminated on the basis of temperature effects. When equilibrium binding of [3H]epinephrine and [3H]norepinephrine to bovine cortex membranes was measured at 37,25, and 4”C, a decrease in the K , (increasing affinity) of the agonist ligands was observed with decreasing temperature, along with a decrease in the Ki values of agonist competitors and an increase in antagonist Ki values at these sites (U’Prichard and Snyder, 1977a). The number of sites labeled by Hlepinephrine and [3 Hlnorepinephrine was

r

r

r

r

r

r

r

390

DAVID B. BYLUND A N D DAVID c . U’PRICHARD

not affected. On the other hand, the A’,) of the antagonist [3H]rauwolscine at cy2 receptors in the same tissue preparation was identical at 4 and 25”C, whether derived from kinetic or equilibrium experiments (Perry and U’Prichard, 1983). In &receptor systems such as the turkey erythrocyte, agonist interactions appear to be entropy-driven, whereas antagonist interactions are enthalpy-driven (Weiland et nl., 1979). Similar considerations may well apply to a2 receptors. Monovalent cations inhibit [3H]epinephrine and [3H]norepinephrine binding to brain a2 receptors (Greenberg et al., 1978), as would be expected from their ability to reduce agonist potencies at [3H]DHEC sites (Tsai and Lefkowitz, 1978; U’Prichard and Snyder, 1978b). Sodium and lithium are equally active in this regard, whereas larger ions such as K+ and Cs+ are less effective. T h e a,-receptor model proposed by Lefkowitz and co-workers would also predict that guanine nucleotides directly inhibit [3H]catecholamine binding to a2 receptors if these ligands are predominantly labeling a2(H).Nucleotides inhibit steady-state binding of [3H]epinephrine and [3 Hlnorepinephrine at bovine cortex a2 receptors (U’Prichard and Snyder, 1978b) and also [3H]epinephrine binding at a 2 receptors in rat cortex (U’Prichard and Snyder, 1980), NG 108-15 cell membranes (Kahn ot cil., 1982), and human platelets (U’Prichard ct ol., 1983). The potency order of nucleotides is quite consistent in different tissues: Gpp(NH)p 2 GTP = GDP > ITP > ATP = CTP = ATP > GMP (CT’Prichard ot 01.. 1983). For PHIepinephrine, the apparent ED,, of Cpp(NH)p and GTP is 1- 10 p.11 in neural tissue, and nucleotides seem more potent in bovine as compared to rat cortex membranes (U’Prichard and Snyder, 1980). Gpp(NH)p and GTP are potent inhibitors of [”]epinephrine binding to platelet a2 receptors with EDso values about 0.1 pL21 (U’Prichard ~t nl., 1983). In brain membranes, GTP increases the apparent K,)of [3H]epinephrine with no change inB,,, and accelerates both association and dissociation when added at the onset of labeling, or after steady state was achieved (U’Prichard and Snyder, 1978b). These effects of GTP on agonist binding parallel results obtained with radioagonist binding to a variety of other cyclase-coupled receptors (Lin P t nl., 1977; Williams and Lefkowitz, 1977a; Blume, 1978). However, the observed effects of GTP are not completely consistent with those predicted by the model. If PHIepinephrine labels a 2 ( H )and GTP simply alters the equilibrium between a,(H) and a2(L),then one would predict that in saturation experiments, GTP would reduce the B,,,, of [3Hlepinephrine binding without changing the K , of Hlepinephrine at residual (H) states being labeled. Likewise, if the two phases of [3H]epinephrine dissociation were taken to represent interactions with

CHARACTERIZATION OF

al-AND

(~2-ADRENERGICRECEPTORS

39 1

az(H)and a Z ( L )sites, the presence of GTP during labeling of the receptor should alter the relative amounts of the rapidly and slowly dissociating components of [3H]epinephrine binding, but not change the k - , values for each phase of dissociation. In both cases, the observed effects were opposite to those predicted by the theory. The interactive effects of guanine nucleotides and divalent cations on [3H]epinephrine binding to a2 receptors are complex and appear to be somewhat different for brain and platelet a2 receptors, although these differences may be simply explained by the different capacity of membranes from different cells to sequester cations. In platelet membranes, which are prepared by routine lysis in an EDTA-containing buffer, Mg2+ increases [3H]epinephrine binding (U’Prichard et al., 1983), an effect which parallels the left shift in the epinephrine competition curve at [3H]yohimbinesites in the presence of Mg2+(Fig. 3). On the other hand, Mg2+does not increase Hlepinephrine binding to cortex membranes prepared in the absence of EDTA (U’Prichard and Snyder, 1980),or to NG 108-15 cell membranes prepared in the presence or absence of

\ 40

- ‘* -

:

-30-

< 01

\* \

\

*\

FIG. 3. Effect of 1.0 mM MgCl, on the saturation characteristics of (-)rH]epinephrine binding to platelet membrane a) receptors (Rosenthal plot). (0),N o MgC1,; ( O ) , 1 mM MgCl,.

392

DAVID B . BYLUND AND DAVID

c . U’PRICHARD

EDTA (D. J. Kahn and D. C. U’Prichard, unpublished). Whereas in platelet membranes the ability of GTP and Gpp(NH)p to inhibit PHIepinephrine binding is enhanced in the presence of Mg2+, in brain membranes Mg2+and Ca2+not only reduce the ability of GTP to inhibit binding, but in the presence of these ions (1.O mM), low concentrations of GTP actually increase Hlepinephrine binding (U’Prichard and Snyder, 1980). Similarly, in NG 108-15 membranes prepared in the absence of EDTA, GTP and GDP (0.1-10 p M ) enhance [3H]epinephrine binding before reducing binding at higher concentrations (Kahn ~t nl., 1982). However, in NG 108-15 membranes prepared in the presence of EDTA, GTP and GDP only inhibit [3H]epinephrine binding (D. J. Kahn and D. C. U’Prichard, unpublished). These data indicate that in brain and NG 108-15 membranes, unlike platelet membranes, divalent cations antagonize the effects of GTP and GDP on agonist interactions with the az receptor. A more detailed analysis of PHIepinephrine binding to platelet aZ receptors indicates that there may be two high-affinity states of the receptor at which rH]epinephrine interacts,\ as well as the a 2 ( L ) state (U’Prichard uf nl., 1983; Mitrius and U’Prichard, 1983). In the presence of 1.0-mhf MgC12, the Rosenthal plot of [3H]epinephrine binding appears slightly curvilinear over the concentration range (up to 40 nM) of [3H)epinephrine used (Fig. 3). T h e K , of the linear portion of the curve is approximately 6 nltf corresponding to the K, value (11 nM) for the a z ( H )site derived from analysis of epinephrine inhibition of rH]yohirnbine binding data (Hoffman et GI., 1982). However, the apparent deviation from linearity cannot be due to [3H]epinephrine binding to the a,(L) state ( K D= 520 niV, Hoffman et nl., 1982) since theoretical calculations show a significant (10%)deviation from linearity only at concentrations of PHIepinephrine in excess of 150 nLU,which is well above the 40 n‘bf used. These saturation data (Fig. 3) are too scattered at the higher radioligand concentrations to estimate a K, value for the lower affinity site. However,K, values can be calculated from kinetic data. T h e association of VHIepinephrine to platelet a 2 receptors is monophasic, but the dissociation is biphasic. If one assumes that the biphasic dissociation corresponds to two affinity states [however other models, such as one based on bivalent ligand hypothesis (Minton, 1981) are just as reasonable], K , values of 2 and 35 nltf can be derived. Omission of Mg2+ reduced the number of [3 Hlepinephrine sites, reduced the overall affinity of [3H]epinephrine by a factor of 2, and eliminated the curvilinearity of the Rosenthal plot, such that observable [3 Hlepinephrine binding was to a single population of receptors with a K, of 12.8 nM (Fig. 3). Similarly, with the addition of 10 F M GTP in the presence of 1.0 mA4 Mgz+, the Rosenthal plot of PHlepinephrine binding was linear, with a some-

r

CHARACTERIZATION OF ff 1- AND ff2-ADRENERGIC RECEPTORS

393

what reduced B,,, and a K , of 30 nM (U’Prichard et al., 1983). The omission of Mg2+or the presence of GTP reduced the absolute amount of the slowly dissociating component of Hlepinephrine binding, but did not change the k-, values for the two components. U’Prichard and co-workers suggest that [3 Hlepinephrine, in addition to labeling in platelet membranes a2(H) ( K D = 30-40 nM) and perhaps a small portion of a2(L)( K , = 300-500 nM), may label another state that, by analogy to agonist interactions with muscarinic cholinergic receptors (Birdsall et al., 1978; Ehlert et al., 1980), can be called the “super high” state of the receptor [a2(SH)with KD values in the range of 2-6 nM]. T h e “a2(H)” conformation of the platelet receptor ascertained from computer modeling of [3H]yohimbine competition curves (Hoffman et al., 1982) could be a composite of a2(SH) and a2(H), as it is doubtful whether such analysis of competition curves in this system could significantly differentiate a three-state from a two-state receptor model. Unfortunately, [3H]epinephrine binding data at NG 108-15 a2receptors is also not accurate enough to resolve putative a 2 ( S H ) and a2(H)states in that system. b. [3H]Clmidine and [3H]p-Aminoclmidine.Although the imidazoline ligands have been used much more extensively than [3H]epinephrine, these ligands label sites representing high-affinity state(s) of the a2 receptor whose properties are very similar to those sites labeled by Hlepinephrine. [3 HIImidazoline binding will be considered, therefore, in less detail. In common with most other imidazolines, clonidine and p-aminoclonidine exhibit partial agonist activity in several assays of a2receptor function (Starke et al., 1974; Kahn et al., 1982; Atlas and Sabol, 198 1). [3 HIClonidine and H]p-aminoclonidine have been widely used to label a2 receptor sites (Table XIV). Interestingly, the presence of a2 receptors in some classical sympathetically innervated tissues such as rat heart and vas deferens has not yet been conclusively demonstrated using [3 Hlclonidine or H]p-aminoclonidine, although this has been a goal in many laboratories. There is a fair degree of consistency as to the characteristics of [3H]imidazoline binding in all these tissues throughout many laboratories (Table XIV). [3 HIImidazoline-specific binding is saturable with K , values usually in the 1-3 nM range and B,,, values from 20 to 460 pmol/gm prot. Rosenthal plots are generally reported as monophasic, but in some tissues there are increasing reports of curvilinearity (Atlas and Sabol, 198 l ; Garcia-Sevillaet al., 1981b; U’Prichard et al., 1983; Braunwalder et al., 1981). [3H]Imidazoline appears to bind to high-afiity states of the a2 receptor since the K i of prazosin is generally 100-1000 times greater than the K i of yohimbine, and (-)catecholamines have K i values in the 1-10 nM range. Furthermore, the binding is generally directly inhibitable by guanine nucleotides.

r

r

r

13 ,,,a\ '

'Tissue Whole hrain" Cerebral cortex Cerebral cortex Cerebellum Corpus striatum Spinal cord Submandibular Submandibular Submandibular Submandibular Sublingual Neuroblastonia X glioma Neuroblastoma X glioma Kidney Ileum Cerebral cortex Retina Cerebral cortex Platelets Platelets Adipocytes Adipocytes a

Species

fritiated ligantl

K,,

pnioYgm protein

Ra t Rat

Clonid i n r

Rat

K dt

Clonidine Clonidine Clonidine Clonidine Clonidine Clonicline Clonidine PAC Clonidine

266 t 37 235 t 19 66 t 7 72 2 5 205 1 27 2 3 150 t 25 378 t 2 460 t 33 287 t 69

Rat- mouse

Clonidine

81; 155

Rat-mouse Guinea pig Guinea pig Pig Bovine Bovine Human Human Human Human

PA c Clonidine Clonidine PAC Clonidine Clonidine Clonidine Clonidine Clonidine PAC

105 t 42

Rat Rat Rat R dt Rat?

Ratd Rat"

P'4 C'

Minus cerebellum.

* Sites per cell.

Chronic reserpine administration.

34 85 t 7 34 1

pmoligm tissue

( I1'\f )

14 13 t 2 14 t 1 4.5 2 0.6 4.6 f 0.5

5.8 0.87 t 0.25 1.8 t 0.1 1.9 2 0.2 1.6 t 0.8 2.1 t 0.2 2.4 t 0.2 1.9 t 0.3 3.7 t 0.5 2.8 t 0.2 2.1 t 0.4

U'Prichard 1.t i l l . (1977a) Rouot and Snyder (1979) D. B. Bylund (unpublished) D. B. Bylund (unpublished) D. B. Bylund (unpublished) Jones vt d. (1982) Pimoule PI ci/. (1980) Bylund and Martinez (1980) Bylund PI rrl. (1982a) Bylund P / I ] / . (1982a) Martinez I,/ I ] / . (1982b)

1.7; 33

Atlas and Sabol ( 1 98 1)

6.2 t 0.9 17 t 3 202 1 9.9 t 1.3

8600 t 23OOb 22 t 2 4.2 +- 0.3 8

35 t 3 64 t 4 348 t 10 166 t 26

Two-weeks old. p-Aminoclonidine.

1.8 ? 0.6 9.0 t 0.8 2.1 1.2 t 0.1 0.32 1.o 5.0 t 0.5 24 t 2 3.9 t 0.2 0.49 t 0.04

Reference

Kahn rt i l l . (1982) Summers (1980) Tanaka and Starke (1979) Harris ct nl. (1983) Bittiger PI nl. (1980) U'Prichard and Snyder (1979) Garcia-Sevilla rt a/. ( 198 1b) Shattil r / 01. (1981) Berlan and Lafontan (1980) Burns et al. (1981)

CHARACTERIZATION OF (Y1- AND a2-ADRENERGIC RECEPTORS

395

In the few tissues where [3H]catecholamine and [3 Hlimidazoline binding to a2 receptors have been directly compared, such as bovine cortex (U’Prichard and Snyder, 1977a), rat cortex (U’Prichard et al., 1979a), human platelets (U’Prichard et al., 1983), and NG 108-15 cells (Kahn et al., 1982), it appears that [3H]clonidine and r H ] p aminoclonidine over restricted concentration ranges label the same high-affinity sites of the a2 receptor as PHIepinephrine, but generally bind to a smaller proportion of these sites. This might suggest that intrinsic activity of agonists at the a2 receptor can be related to the extent of formation of high-affinity states, as is the case for /3 receptors (Kent et al., 1980). However, the analysis of H-labeled antagonist-agonist interactions by Hoffman et al. (1982) indicates that there is no such correlation. In EDTA-treated platelet membranes, [3 H]p-aminoclonidine saturation in the presence of 1.0 mM M 8 + is more markedly biphasic than that of Hlepinephrine. If the curvilinear Rosenthal plot is interpreted to represent two binding sites, KD values for the two sites can be estimated. Approximate KD values of 0.3-0.7 nM and 3-10 nM were obtained by drawing straight lines through the linear portions of the curve (U’Prichard et al., 1983). By this type of analysis, neither site seems equivalent to the KD of p-aminoclonidine at the a 2 ( L ) state as determined by [3H]antagonist binding which is 80-100 nM (Fig. 2). However, this method of determining KD values underestimates the KD for the low-affinity (high KD) component (Minneman et al., 1979). A more careful analysis using the graphical technique of limiting slope coupled with replotting of derived curves suggests that the data from several experiments are best fit to two sites having KD values of 0.3-0.5 nM and 10-17 nM. On the other hand, [3H]p-aminoclonidine dissociation, like rHIepinephrine, is biphasic, and if the two-site model is assumed, KD values from kinetic experiments are 0.2 nM and 2-3 nM (Mitrius and U’Prichard, 1983). Thus, while there is some evidence from [3 H]p-aminoclonidine binding to platelet membranes which can be interpreted to indicate SH and H states of the a2 receptor, much more experimental evidence is needed to establish the model. I n the absence of M 8 + , or in the presence of 10 p M GTP, the Rosenthal plot of [3H]p-aminoclonidine binding is linear, with a KD value of 3-10 nM. Neither treatment alters the kl values of each component of rH]paminoclonidine binding. The number of sites labeled by ?HIP-aminoclonidine is equivalent to 50% of the a,-receptor sites labeled by [3 Hlyohimbine, whereas the full agonist [3H]epinephrine labels 80% of the rH]yohimbine sites (Mitrius and U’Prichard, 1983). Competition studies show that H]p-aminoclonidine labels the same

r

r

396

DAVID

B. BYLUND

A N D DAVID

c.

U’PRICHARD

r

high-affinity sites of the platelet a2 receptor as Hlepinephrine (Table XIII). For both ligands, the i z H values of all competitors are less than 1.0, indicating that even at the low ligand concentrations used, [3H]epinephrine and [3H]p-aminoclonidine may be labeling more than one state of the receptor. In the presence of 1.0 p M GTP, IC,, values for agonists at ~H]p-aminoclonidineand VHIepinephrine sites increase two- to threefold, but IC,,, values for the antagonist yohimbine decrease two- to threefold. In neither case is there a significant change in n H values. PHJClonidine binding to the platelet a2 receptor has not yet been studied in quite as much detail, but the available evidence is that it binds to high-affinity a,-receptor sites, which constitute 40-60% of the total receptor population measured with antagonist radioligands. These sites are also nucleotide-sensitive (Garcia-Sevilla et al., 198lb). In NG 108-15 cell membranes, [3 H]p-aminoclonidine appears also to label the same high-ahity a2-receptor sites as VHIepinephrine (Table XIII), but as with [3H]epinephrine binding in this system, putative SH and H states cannot be resolved. At low concentrations, [3H]paminoclonidine saturation and dissociation are monophasic (Kahn et nl., 1982). Guanine nucleotide influences are similar for both ligands, with a bimodal effect of GTP and GDP that was converted to unimodel inhibition of binding after pretreatment of membranes with EDTA, whereas Gpp(NH)p in both conditions only inhibited binding (Kahn et al., 1982; D . J. Kahn and D. C. U’Prichard, unpublished). The tissue wherein rH]clonidine binding to a2 receptors has been most extensively examined has been the rat brain, especially the cerebral cortex (U’Prichard et al., 1979a; Glossmann and Presek, 1979; Glossmann and Hornung, 1980b; Summers et al., 1980; Braunwalder et ~ l . ,1981). Early studies of Hlclonidine binding used low-specificactivity radioligand and could only demonstrate monophasic saturation and dissociation. This represents high-affinity a,-receptor sites (catecholamine K , values 5-20 nlM), but was not recognized as such (U’Prichard et nl., 1977a). With the advent of higher specific activity ligand, saturation and dissociation were seen to be biphasic (U’Prichard et nl., 1979a; Vetulani et ol., 1979). In competition studies, agonists were more potent at the higher affinity component, while some antagonists were less potent (U’Prichard et al., 1979a). These data were interpreted as possibly representing different populations of a2receptors on the basis of different regional distribution throughout the rat brain, and difTerent regulation in 6-hydroxydopamine-treated animals (U’Prichard et al., 1979a). At present, a more likely interpretation of these data is that [3H]clonidine in the brain labels two states of the same receptor population, but the complexity of brain tissue and other evidence suggesting

r

CHARACTERIZATION OF a1- AND Q2-ADRENERGIC RECEPTORS

397

the existence of both pre- and postsynaptic a2 receptors (which may be coupled to different effectors, or to the same effector with varying efficiency) will make interpretation of a2-agonist-binding data difficult for some time to come. Glossmann and colleagues have identified four or five affinity states of the rat cortex a, receptor labeled with [3H]clonidine on the basis of different thermal stability and Mg2+ sensitivity (Glossmann and Presek, 1979; Glossmann and Hornung, 1980b). The more slowly dissociating phase of cortical [3 Hlclonidine binding is selectively inhibited by guanine nucleotides by reducing the number of sites without altering the K , of Hlclonidine. Guanine nucleotides do not appear to affect the other component of binding, and the K-, values for the two phases of [3H]clonidine binding are not changed by the presence of nucleotides and divalent cations (Rouot et al., 1980). However, Mg2+increases [3H]clonidinebinding to the highest affinity state of the receptor in EDTA-treated membranes (Glossmann and Presek, 1979; Glossmann and Hornung, 198Oc),and more recent evidence indicates that 10 mM M 8 + will increase the number of putative (SH) states labeled by Hlclonidine in cortical membranes not pretreated with EDTA (Salama et al., 1982). It is clear from these and similar data that the extent of clearance of endogenous ligands for the a2receptor and its associated N site from the membranes prior to binding assay plays a critical role in the type and extent of interaction of ions and nucleotides with 3H-labeled agonist a2 receptor sites. In the presence of 1.0 mM divalent cations, GTP increases [3 Hlclonidine binding in cortex (U’Prichard and Snyder, 1980; Rouot et al., 1980), while in the presence of chelating agents, GTP becomes more potent in inhibiting pH]clonidine binding (U’Prichard and Snyder, 1980). Sodium, unlike GTP, reduced rH]clonidine binding in rat cortex (Rouot et al., 1980). In summary, if we compare the characteristics of Hlclonidine binding itself, and of full agonist (catecholamine)competition at sites labeled by rHIclonidine (U’Prichard et al., 1979a) and H]rauwolscine (Perry and U’Prichard, 1983),it seems probable that the cerebral cortex a2receptor exists in multiple affinity states, but the distribution of these states among different cortical a,-receptor populations is still a matter for conjecture. Glossmann and Hornung (1980b) noted that the antagonist prazosin had a lower affinity for the highest affinity state of the rat cortex a2 receptor labeled with Hlclonidine and that prazosin interactions with Hlclonidine sites were apparently noncompetitive in nature. More recently, 10 mM Mg2+ was observed to reduce the affinity of some antagonists such as piperoxan, yohimbine, WB410 1, and prazosin as much as sixfold at rat cortex rHIclonidine binding sites, whereas the affinity of other antagonists such as phentolamine was unaffected (Salama et al.,

r

r

r

r

r

r

398

DAVID B . BYLUND AND DAVID

c . U’PRICHARD

1982). hssuming that Mg2+was increasing the ratio of putative SH to H components of [3H]clonidine binding, these data were taken as evidence that the former group of antagonists exhibited preferential affinity for a 2 ( H )compared to a2(SH),whereas phentolamine had equal affinity at both states. a2 Receptors on membranes from human adipocytes from properitorieal adipose tissue have also been characterized using [3 HIPaminoclonidine (Burns rt d.,1981, 1982a). In these studies, [3H]paminoclonidine exhibited high affinity ( K E )of 0.5 to 1.2 nM), and the number of az receptors labeled by [3 H]p-aminoclonidine was approximately 50% of the az receptors labeled by VHIyohimbine. Adipocytes from this source exhibit responses to catecholamines of stimulation of adenylate cyclase activity and glycerol production, inhibition of adenylate cyclase and glycerol production, and stimulation of phosphatidylinositol turnover, which were shown to be associated with p , a z , and a , receptors, respectively (Burns et al., 1981). In other studies concerning adipocyte receptors, analysis of competition of [3 HIDHEC binding by subtype-selective antagonists indicated that the great majority of a receptors were of the a2 subtype in adipocytes from human subcutaneous adipose tissue (Hoffman rt ol., 1979, 1980a) and in hamster adipocytes (Garcia-Sainz P t nl., 1980b). r . [ 3 H ] G ~ i a n f u c u ~Tr .h e tritium-labeled a 2agonist and antihypertensive drug guanfacine has also been used in radioligand binding studies. [3H]Guaiifacine bound to rat brain membranes with a K , of 4 nM and with properties characteristic of a2receptor binding sites (Timmermans P f d., 1982). However, its usefulness may be limited by a relatively high amount of nonspecific binding.

The initial cellular response to activation of aZ receptors in all cell systems s o far examined is inhibition of plasma membrane adenylate cyclase and a consequent decrease in intracellular CAMPlevels. Thus, the a2 receptor falls into the recently recognized large class of receptors that are coupled to adenylate cyclase in an inhibitory manner, including muscarinic cholinergic, opiate, adenosine A1 and dopamine D2 receptors, and receptors on adipocytes for nicotinic acid and prostaglandins of the E series (Jakobs, 1979). T h e mechanisms of coupling of these receptors to adenylate cyclase have recently been discussed by Jakobs ( 1979) and Limbird (1981), and many authors have compared the effects of nucleotides and other agents on the receptor-mediated response of inhibition and stimulation of adenylate cyclase. T h e sequence of events relat-

CHARACTERIZATION OF a1- AND (Y2-ADRENERGIC RECEPTORS

399

ing receptors to adenylate cyclase stimulation in relatively pure cell membranes (e.g., p receptors in frog and turkey erythrocytes and mouse S49 lymphoma cells, and glucagon receptors in rat hepatocytes) has been well described. I n these systems, receptor-mediated stimulation of the enzyme is dependent on the presence of GTP acting at intermediary N protein(s). Stimulation of cyclase is “turned off’ by the hydrolysis of GTP to GDP which is catalyzed by GTPase activity associated with N. Thus, GTP can be viewed as the primary cellular ligand for adenylate cyclase stimulation, and receptor agonists activate cyclase in a GTP-dependent manner by accelerating the GTP-GDP-GTP cycle (Ross and Gillman, 1980). Nonhydrolyzable analogs of GTP, for example, Gpp(NH)p, Gpp(NH,)p, and GTPyS, activate adenylate cyclase in an apparently irreversible manner (Londos ~t al., 1977), behaving similarly to GTP in the presence of cholera toxin, which inhibits GTPase activity and catalyzes the ADP-ribosylation of N proteins (Moss and Vaughan, 1977; Gill and Meren, 1978). These GTP analogs stimulate the enzyme after an initial lag period which is thought to be the time needed for release of inactive GDP from N; this lag period is decreased in the presence of agonists that stimulate cyclase (Birnbaumer et al., 1980). Beta agonists also stimulate GTPase activity in membranes probably by both directly increasing the V,,, and by accelerating GDP-GTP exchange (Limbird, 1981). T h e basic characteristic of receptors (including a,-adrenergic receptors) that are negatively coupled to adenylate cyclase is that stimulation of these receptors causes a decrease in the V,,, of the enzyme without changing the K , for the Mg . ATP substrate or for free Mg2+. This response, like stimulation of cyclase, requires GTP, but in higher concentrations (> 1.0 p M ) and is either dependent on, or merely facilitated by, Na+. It has been suggested that inhibition of cyclase by activation of these receptors may be due directly to increases in GTPase activity (Limbird, 1981; Koski and Klee, 1981). Rodbell and co-workers made the fundamentally important observation that in adipocytes and other tissues GTP itself has a bimodal effect on adenylate cyclase activity, stimulating at low (10- 100 nM) and inhibiting at high (1.0- 10 p M ) concentrations. T h e effects of inhibitory receptors also require GTP to be in the high concentration range (Londos et al., 1978). Rodbell (1980) has suggested that coupling proteins for inhibitory receptor systems ( N i )are physically different from the coupling proteins for stimulatory systems (Ns). I n cells such as adipocytes, where a bimodal GTP function has been demonstrated, agents such as cholera toxin, mercurials, divalent cations,, and trypsin selectively abolish one or the other phase of GTP function, and GTP has very different potencies at each phase (Cooper et al., 1979).

400

DAVID B . BYLUND AND DAVID c . U'PRICHARD

Inhibition of adenylate cyclase by a2-receptor agonists has been shown to occur in plat'elets (Jakobs et al., 1978a,b; Jakobs and Schultz, 1979), human and hamster adipocytes (Burns and Langley, 1975; Aktories et al., 1980, 1981; Jakobs and Aktories, 1981; Garcia-Sainz et al., 1980b), rat hepatocytes (Jard et nl., 1981), and cultured rodent neural cells such as the NG 108-15 hybrid (Sabol and Nirenberg, 1979a). I n other tissues such as arterial smooth muscle (Anderson, 1973; Buonassisi and Venter, 1976), rat myocytes (Watanabe et a/., 1977) and glial cultures (van Calker et nl., 1980; McCarthy and deVellis, 1978), parathyroid cells (Brown et d.,1978), and pancreatic islet /3 cells (Nakaki et ul., 1981), there is also evidence, albeit less substantial, that a2 receptors mediate inhibition of adenylate cyclase. T h e classical pharmacological response associated with a2receptors is inhibition of the release of norepinephrine into central adrenergic and peripheral sympathetic synapses (Langer, 1974). Although the pharmacological characteristics of this response are very similar to those of the cyclase inhibitory response (Starke, 1981), there is as yet no evidence to show that in nerve terminals, inhibition of adenylate cyclase causes decreased release of norepinephrine. However, the Occurrence of a lag time between a2-receptor occupation and altered norepinephrine release might support the existence of a second messenger for this response (Story Pt al., 1981). a,-Receptor function in the brain and autonomic ganglia has also been defined electrophysiologically. a2 Receptors mediate hyperpolarization and decreased firing of norepinephrine-containing cells in the locus coeruleus (Svensson et nl., 1975) and hyperpolarization of postganglionic cells causing slow IPSP (Tokimasa et al., 1981). In many cells, a2-receptor activation causes an increase in cGMP levels (Anderson, 1973) that is presumably mediated via increased free intracellular calcium. T h e relationship between this response and inhibition of adenylate cyclase is unclear. a-2-Receptor-mediated inhibition of adenylate cyclase has not yet been conclusively demonstrated in brain membranes, probably because brain monoamine receptors generally are very inefficiently coupled to cyclase in cell-free preparations (Maguire et nl., 1977). In other cells, differences can be detected in a2 receptor coupling to cyclase depending on whether activity is measured in intact cells or lysates. Stimulation of a2 receptors in intact platelets or NG 108-15 cells reduces CAMPlevels by as much as 90% (Lichtstein et al., 1979), but in cell lysates adenylate cyclase activity is rarely reduced by at agonists by more than 50% at maximally effective concentrations (Jakobs, 1979). I n lysates from cells containing a2 receptors, imidazolines such as clonidine are partial agonists, with the exception of platelet membranes where clonidine has no efficacy uakobs,

CHARACTERIZATION OF

al-AND

GQ-ADRENERGIC RECEPTORS

40 1

1978). However, in intact platelets, clonidine will reduce cAMP levels elevated by PGIz, which indicates that it exhibits some efficacy in a more efficiently coupled system (Lenox et al., 1980). Some further characteristics of the a2-receptor response in three well-studied systems are discussed below. 1. Human Platelets Activation of human platelet a2receptors stimulates the aggregation response (BartheI and Markwardt, 1974), lowers cAMP levels in intact cells (Kafka et al., 1977), and decreases adenylate cyclase activity in membrane preparations (Salzman and Neri, 1969). Jakobs and colleagues (1978a) found that inhibition of basal platelet cyclase activity, or activity stimulated by fluoride, PGE, , or adenosine, is GTP dependent in membranes. A concentration of GTP of 1.0 p M or more appears to be necessary (Jakobs and Schultz, 1979). While epinephrine could inhibit the twofold increase in cyclase activity caused by GTP in cholera toxintreated platelet membranes, no az-agonist inhibition of Gpp(NH)p- or GTPyS-stimulated cyclase was observed, and therefore the authors suggested that activation of az receptors might decrease cyclase activity by directly stimulating GTPase (Jakobs and Schultz, 1979). Steer and Wood (1979) found that in platelet membranes purified by the method of Barber and Jamieson (1970), epinephrine or GTP alone could only inhibit cyclase activity very slightly, whereas epinephrine plus GTP had a synergistic effect to inhibit activity by a maximum of 50%. Guanosine triphosphate has a bimodal effect on adenylate cyclase, with concentrations below 1.O pM stimulating and higher concentrations inhibiting. On the other hand, stimulation of cyclase by PGE appeared to be GTP independent. PGE, reduces the lag time before the onset of Gpp(NH)p stimulation. In the presence of Gpp(NH)p, no a2response is observed, but if membranes are preactivated with Gpp(NH)p, both GTP and epinephrine could inhibit the enhanced cyclase activity. These data together suggest the occurrence of distinct Ni and N, sites in platelets (Steer and Wood, 1979). I n further studies (Steer and Wood, 1981), Na+ and other monovalent cations in purified membranes reduced basal and PGE-stimulated cyclase (Na" > Li+ > K+),but did notchange the K , for Mg * ATP or Mg2+. a-2-Receptor-mediated inhibition of adenylate cyclase in this preparation was not Na+ dependent, in that the ion did not alter the maximal fractional inhibition caused by epinephrine. However, Na+ did increase the K , of epinephrine and K , of PGE, suggesting that here it may have a generalized uncoupling effect. As in adipocytes (Jakobs and Aktories, 1981), low Mn2+ was found to preferentially un-

402

DAVID B. BYLUND AND DAVID

c. U’PRICHARD

couple inhibitory receptors and thus 1.0 m,Lf MnZ+ attenuated azreceptor inhibition of cyclase without affecting PGE stimulation. Since 1.0 mL\l Mn2+did not, when compared with 6.0 miM Mg2+ (which supports the inhibitory response), alter the characteristics of epinephrine or Gpp(NH)p interactions at platelet a2-receptor sites labeled with PHlyohimbine, it was concluded that Mn2+ uncouples the system by affecting N-C, and not R-N, interactions (Hoffman et nl., 1981b). 2.

Adiporite\

In hamster adipocytes, catecholamines inhibit CAMP and glycerol production via an a,-receptor interaction, and in this system clonidine is a potent and efficacious agonist (Garcia-Sainz et nl., 1980a). Jakobs and colleagues have extensively characterized the a2 and other inhibitory responses in membranes from hamster adipocyte ghosts. Basal adenylate cyclase activity is inhibited by az agonists [ K , = 3.0 p M for (-)epinephrine and 10 pL\l for (-)-norepinephrine], E series prostaglandins, and nicotinic acid, all of which are antilipolytics, while cyclase is stimulated b) ACTH and p agonists. Inhibition is GTP dependent in the range 1- 10 pL\I GTP, and GTP alone inhibits activity. Sodium stimulates the GTP-dependent component of activity eightfold (unlike platelets, see above), and expression of inhibitory receptor activity was also Na+ dependent in this system. Inhibitory receptor responses were abolished partially by fluoride (10 mill) and completely by Gpp(NH)p (Aktories ~t nl., 1980). As in platelets, increasing Mn2+concentration from 0.05 to 1.O mdLlcompletely abolished the inhibitory effects of GTP and a2 agonists on hamster adipocyte cyclase, and the ED, for Mn2+in uncoupling the inhibitory response (again suggested to involve N-C, not R-N, interactions) was 0.1-0.2 mA\d,whereas Mg2+uncoupled at 10-20-mM concentrations (Jakobs and Aktories, 1981). T h e potency order of monovalent cations in increasing GTP (10 pM)-inhibited enzyme, or in decreasing @agonist or ACTH-stimulated enzyme, was Na+ > Li+ > K+. As in platelets, Na’ appeared to increase the K , of the stimulatory hormone ACTH. Guanyl-5’-yl imidodiphosphate, like GTP, lowered cyclase activity, but did not support a2-receptor inhibition, and in the presence of Gpp(NH)p, cation stimulation was less pronounced (Aktories et nl., 1981). The authors reached the important conclusion that monovalent cations interact at the N site(s) to accentuate N,-mediated, and impair N,-mediated, effects. T h e same group has recently shown that in purified rat liver membranes, cations support GTP-stimulated cyclase (Li+ > Na+ > K+), and this activity is inhibited via a2-receptor interactions (Jard ut ~ 1 . .198 1). In human adipocytes catecholamines have historically been known to

CHARACTERIZATION OF (Y1- A N D (Y2-ADRENERGIC RECEPTORS

403

inhibit cAMP production and lipolysis through an a receptor as well as stimulate cAMP production and lipolysis through a p receptor (Robison et al., 1972). Epinephrine, which activates both a 2 and /3 receptors, increases cAMP levels only about eightfold above basal. Stimulation of only p receptors (either by isoproterenol or epinephrine plus yohimbine) increases cAMP levels about 100-fold, whereas a2receptor activation (clonidine or epinephrine plus propranolol) decreases basal cAMP levels 40-50% (Burns et al., 1981, 1982a). Thus the @stimulated activity is more effectively inhibited by a2 receptors than is the basal activity. Prazosin at concentrations as high as 10 p M is ineffective in inhibiting the a2 response. Similar effects of a2inhibition are seen on adenylate cyclase activity and lipolysis. Other investigators have found that a2 inhibition of membrane basal or PTH-stimulated adenylate cyclase achieves a maximum effect of 30-50% decrease (Kather et al., 1980), with Ki values for (-)-epinephrine and (-)a-methylnorepinephrine of 2.0 and 7.0 p M , respectively. Forskolin has been shown to activate adenylate cyclase in membrane preparations and to increase cyclic AMP levels in intact cells of a variety of tissues (Seamon et al., 1981). The effects of forskolin appear not to be mediated by N, (Seamon and Daly, 1981), although the precise locus of action of forskolin remains unclear (Forte et al., 1982). Forskolin caused a dose-dependent 100-fold increase in the intracellular concentration of cyclic AMP and a 6-fold increase in glycerol release in the human adipocyte. Alpha-2-adrenergic activation (epinephrine plus propranolol) significantly inhibited forskolin-stimulated cyclic AMP accumulation and glycerol release, shifting the dose-response curves approximately 8-fold and 5-fold to the right, respectively (Burns et al., 1982b). It appears that forskolin will be a useful tool in elucidating the mechanism of action of the a,-adrenergic receptor. 3. NG 108-15 Cells Mouse neuroblastoma X rat glioma hybrid cells NG 108-15 ( 108CC15) in culture have “cholinergic” characteristics in that they take

u p choline and synthesize and release acetylcholine (dibutyryl CAMPdifferentiated cells) when stimulated (Hamprecht, 1977). These cells contain receptors for PGEl, adenosine (A2), secretin, and glucagon that stimulate adenylate cyclase, and a2-adrenergic, &opiate, muscarinic, and somatostatin receptors that inhibit cyclase (Propst et al., 1979). Activation of a2 receptors in NG 108-15 membranes reduces basal and PGE,stimulated cyclase activity, and the inhibition occurs without any observable lag time. a2, opiate, and muscarinic inhibition of cyclase in NG 108-15 membranes is GTP dependent (Blume et al., 1979; Kahn et al.,

404

DAVID B. BYLUND AND DAVID

c.

U’PRICHARD

1982). The K, values for (-)-epinephrine and (-)-norepinephrine are 0.2-0.5 pXI, similar to other a2-receptor systems discussed above. Recently, a bimodal effect of Gpp(NH)p on cyclase activity in NG 108-15 membranes has been seen, with very low Gpp(NH)p concentrations inhibiting the activity (Propst and Hamprecht, 1981). This has also been found with cyclase activity in rat hippocampal membranes (Girardot et al., 1983). Alpha-:! agonists also inhibit cholera toxin-stimulated cyclase. These findings are adduced to support the existence of separate N, and N, in NG 108-15 membranes (Propst and Hamprecht, 1981). Analysis of the coupling of a2 and other receptors to adenylate cyclase may be inherently more difficult in neural tissue than in platelets or adipocytes because there is abundant evidence that the Ca-calmodulin complex is an essential component of receptor-N-cyclase coupling in nerve cells, and that nucleotide activation (Brostrom et al., 1978, 1981; Partington et a/., 1980; Toscano et al., 1979; Wilkening et al., 1980; Brandt et al., 1980) and inhibition (Girardot et nl., 1983) of adenylate cyclase is calmodulin dependent. Calcium has a bimodal effect on NG 108-15 adenyiate cyclase. Low (
c. COMPARISON OF AGONIST.4ND ANTAGONIST BINDING: TOWARD A KINETICMODELOF a2-RECEPTOR FUNCTION

Is it possible from consideration of the above binding and cyclase data to suggest a kinetic model for a,-receptor coupling to adenylate cyclase that takes into account all observed interactions of agonist, antagonist, guanine nucleotides, and metal cations? At a certain level of analysis, a model of a,-receptor function has been proposed which is similar in all essential respects to the “dynamic receptor a h i t y ” model of stimulatory p receptor function (Hoffman and Lefiowitz, 1980a). T h e latter model presents a coherent, integrated picture of the kinetics of (3-receptor (R) activation of adenylate cyclase (C), through the sequential interaction of guanine nucleotide-binding regulatory subunits (N) with R and C. T h e data to develop this model have come mainly from receptor binding (Hoffman and Lefkowitz, 1980a) and cyclase reconstitution (Ross and Gilman, 1980) studies. A preexisting equilibrium between R and an R-N complex is shifted in the direction of R-N when agonist or

CHARACTERIZATION OF

cq-

AND (YZ-ADRENERGIC RECEPTORS

405

hormone (H) combines with R, leading to the predominance of a ternary H-R-N complex (DeLean et al., 1980). Uncomplexed R is equivalent to the low-affinity state R(L), while R-N is a high-affinity state R(H), as these states have been analyzed in studies of heterogeneous agonist competition curves at antagonist radioligand binding sites (Hoffman and Lefkowitz, 1980a). Antagonists have the same affinity for R(L) and R(H). Negative heterotropic cooperativity exists between R and N, so that GTP, the endogenous ligand for N, decreases agonist affinities at R by converting R(H) to R(L) (i.e., R-N to R), while H, through the formation of H-R-N, decreases the affinity of N for nucleotides and thereby facilitates the exchange of inactive GDP for GTP. By combining with N, GTP dissociates R from N, and GTP-liganded N complexes with the catalytic moiety (C) in a stable manner. The N-C complex represents “active” cyclase (C*). Activation of cyclase is “turned off’by the hydrolysis of GTP to GDP, catalyzed by GTPase activity associated with N. Magnesium is proposed to stabilize the H-R-N complex by interacting with the N component, and thus, in contrast to GTP, favors the R(H) state. T h e above interactions can be represented schematically in simplified (nonkinetic) form:

A large body of evidence, especially derived from, but not limited to, platelet studies, supports a similar basic model for a2-receptor function. Labeled antagonists interact with a single population of a,-receptor sites which agonists recognize with two affinities. Thus, antagonists label a,(H) and a 2 ( L ) ,which are discriminated only by agonist, and not antagonist, competitors. Guanosine triphosphate, which is a prerequisite for a2-receptor adenylate cyclase coupling, right shifts and steepens agonist competition curves (Fig. 2), indicating that GTP is shifting equilibrium in favor of a2(L).Magnesium ion left shifts agonist competition curves and makes them shallower (Fig. 2), suggesting an increase in the a2(H) state. Partial agonists exhibit steeper competition curves at antagonist sites since they are less able to induce the formation of the H-R-N complex, or az(H), which is reflected as a lower number of,

406

DAVID B. BYLUND AND DAVID c . U'PRICHARD

a n d o r affinity at, a z ( H ) compared to a z ( L )states. T h e formation of the H-R-N complex is a prerequisite for coupling of the a2 receptor to adenylate cyclase. T h e characteristics of, and influences on, labeled agonist binding sites parallel very well agonist interactions at the antagonist site. Over low concentration ranges, agonists preferentially label az(H) states, that is, agonist competitors have much higher affinity at these binding sites (Table XIII), and in many tissues the number of agonist sites is 30-70% of the number of antagonist sites. Over broader concentration ranges, Rosenthal plots of labeled agonist binding tend to be nonlinear, presumably because the agonist has different affinities for crz(H)and a z ( L ) .Guanine nucleotides decrease agonist binding and accelerate dissociation or decrease the proportion of slowly dissociating agonist. Conversely, M$+ generally increase agonist binding and increases the proportion of slowly dissociating agonist. Imidazoline partial agonists label fewer high affinity sites than do full agonists, but the sites that partial agonists label have the same a 2 ( H ) characteristics. It is not yet clear if the kinetics of' coupling of a z and /3 receptors to adenylate cyclase are so similar, why occupation of the az receptor should lead to a less active state of the enzyme. The most likely explanation is that the constituents of N are different for the two receptors. It is intriguing that Gpp(NH)pappears to dissociate the az-receptor H-R-N complex as effectively as GTP, yet a2 agonists will not inhibit Gpp(NH)p-stimulated cyclase. This has raised the suggestion that az agonists may inhibit cyclase by in some way directly stimulating GTPase activity, although observed increases in GTPase activity (Koski and Klee, 1981) may be equally due to increased GDP release from N and thereby increased GTP cycling. It is also curious that az inhibition of (GTP plus cholera toxin) stimulated cyclase is observed, when GTPase is inhibited by the toxin. There is as yet no direct evidence that az agonists facilitate GDP release from N. Figure 4 is another representation of the agonist-induced coupling of the azreceptor to adenylate cyclase. A role for sodium at a constituent of N is postulated, since Na' in all systems at least facilitates coupling, and affects agonist interactions at the receptor in a manner similar to GTP. However, it must be recognized that the precise role and locus of action of Na+ is by no means as clear as that of GTP. Some findings are not currently explained by the above model of a,-receptor function. First, H-labeled agonist binding studies using both platelet and brain membranes have been interpreted to suggest the presence of two or more states of the receptor with high affinity for agonists, in addition to the low affinity state (U'Prichard et nl., 1979a,

CHARACTERIZATION OF (Y1- AND (Y2-ADRENERGIC RECEPTORS

407

* Less Aclive Stole? FIG. 4. Sequential model of a2-receptor coupling to adenylate cyclase in cell membranes. See text for explanation of symbols.

1983; Rouot et al., 1980; Glossmann and Hornung, 1980b). The receptor state with highest affinity for agonists [az(SH)]is Mg2+ dependent and is analogous to the (SH) state of the muscarinic cholinergic receptor (Birdsall et al., 1978; Ehlert et al., 1980) which is also negatively coupled to adenylate cyclase (Jakobs et al., 1979). How could this (SH) state be incorporated into a kinetic model of receptor coupling? Maguire and colleagues have suggested that for p receptors the ternary complex can occur in Mg2+-boundand unbound forms (Cech et al., 1980). It is possible that the putative (SH) state is equivalent to H-R-N Mg2+,while (H) is equivalent to H-R-N. Aggregation of N subunits can be affected by Mg2+(J. K. Northup, personal communication), and R may bind with different affinities for N protein monomers or dimers, which would be reflected in different dissociation constants for H-R. Preliminary evidence suggests that there may be an increase in (SH) states of the a2 receptor in platelet membranes where a,-receptor coupling to adenylate cyclase is lost, and in membranes from NG 108-15 cells exposed for a short time to the agonist (-)-epinephrine (15-30 min), when the potency of epinephrine in inhibiting adenylate cyclase is decreased (Mitrius and U’Prichard, 1983; Kahn and U’Prichard, 1983). This might indicate that (SH) states are uncoupled from the catalytic moiety of the enzyme. The second finding not explained by the model is that affinities of some antagonists appear to be different at different states of the receptor. In human platelet and NG 108-15 cell membranes, most antagonists are more potent competitors at a2 receptors labeled by Hlantagonists than at [3H]agonistsites (Table XIII). Likewise in bovine cerebral cortex membranes, a comparison of drug affinities at [3 Hlepinephrine and [3 H]rauwolscine sites (Fig. 5 ) shows that all full (catecholamine)agonists exhibit, to the same extent, higher affinity for the VHIagonist site. Partial (imidazoline) agonists are also more potent inhibitors of [3 Hlagonist binding, but to a lesser extent. While some antagonists such as

r

408

DAVID B. BYLUND AND DAVID c . U’PRICHARD

Fic,. 5 . Comparison of affinities of competitors at a,-receptor sites in bovine frontal cortex labeled by the agonist [SH]epinephrine and the antagonist [aH]rauwolscine. Full agonists (0):1, (-)-epinephrine; 2, (-)a-methyl-norepinephrine; 3, (-)-norepinephrine; 4, (-)-phenylephrine; 5 , (+)-norepinephrine; 6, (-)-isoproterenol. Partial agonists (0): 7, oxymetazoline; 8, p-aminoclonidine; 9, clonidine. Antagonists (0):10, phentolamine; 11, phenoxybenzamine; 12, dihydroergocryptine; 13, rauwolscine; 14, yohimbine; 15, rnianserin; 16, WB4101; 17, piperoxan; 18, prazosin; 19, propranolol.

phentolamine have equal affinity at a2-receptor sites labeled by VHIrauwolscine and rHIepinephrine, most antagonists are more potent competitors at ~H]rauwolscinesites. Additional evidence for differential antagonist affinities is that MgZf,which shifts receptors to high-affinity states, decreases the affinity of the antagonist rHlyohimbine at platelet a2 receptors (Daiguji et al., 1981a) and the affinities of antagonist competitors at brain cY2-receptorsites labeled with [3H]clonidine(Glossmann and Hornung, 1980b; Salama et al., 1982). Conversely, both Na+ and guanine nucleotides, which shift receptors to low-affinity states, increase the affinity of [3H]yohimbine and antagonist competitors at platelet a2 receptors (U’Prichard et al., 1983; Limbird et ul., 1982; Mooney et al., 1982) and affinity of [3H]rauwolscine and antagonist competitors at brain a2receptors (Perry and U’Prichard, 1983). These data suggest that the rank order of affinity of different receptor states for an antagonist is the obverse of the order of affinity of these states for agonists. If the antagonists Hlyohimbine and H]rauwolscine had different affinities for different receptor states, one would predict multiphasic binding

r

CHARACTERIZATION OF CYl- AND (Y2-ADRENERGIC RECEPTORS

409

isotherms, rather than the typical monophasic isotherms observed. However, in most experiments these ligands are used over somewhat restricted concentration ranges. One would also predict that [3H]yohimbine and [3H]rauwolscinein typical experiments would not observably label all up receptors, but select somewhat for the ap(L) state. Several laboratories have noticed that in platelet membranes, [3H]yohimbine labels fewer az-receptor sites than another az antagonist [3H]DHEC (Daiguji et al., 1981a; Motulsky and Insel, 1982). The Hoffman and Lefkowitz (1980a) model of az-receptor function can be modified to accommodate some of these data along lines originally suggested by Colquhoun (1973), who postulated that a drug could act as an antagonist either if it has equal affinity for “active” and “inactive’’ states of the receptor or if it had a higher a h i t y for the “inactive” [aa(L)]state. This can be kinetically represented for a,(H) and az(L) state interactions with agonists and antagonists in terms of the ability of ligands to stabilize or destabilize the formation of a ternary complex, as suggested by DeLean et aE. (1980); X + R-

K

-X-R+

N

where X is either an agonist or antagonist ligand. The formation of R-N in the absence or presence of ligand is determined by the equilibrium association constants,M and L. Agonists bind with higher affinity to R-N (K’ > K) and stabilize R-N (L > M). An antagonist may have the same affinity for R and R-N (K’ + K) and neither stabilize nor destabilize R-N ( L = M) (e.g., phentolamine), or it may bind with higher affinity (K‘ < K) to R than to R-N, thus destabilizing R-N ( L < M) (e.g., yohimbine, rauwolscine). A third finding which is at variance with the proposed model is that in some tissues, only [3H]agonist binding can be demonstrated under a variety of incubation conditions (changes in buffer and GTP and Mg2+ concentration). For example, in the rat submandibular gland (reserpinized or denervated), there is good rHIclonidine and pH]paminoclonidine binding, but no [3 Hlyohimbine binding (Bylund and

410

DAVID B. BYLUND AND DAVID

c. U'PRICHARD

Martinez, 1980, 1981; Bylund et al., 1982a), whereas in the neonatal rat lung, VHlyohimbine and ~Hlrauwolscinelabel a large number of sites, but [3H]clonidine and HIP-aminoclonidine binding could not be observed (Latifpourrt al., 1982). It could be argued that in the lung, almost all of the receptors are in the (L) state and thus binding of 3H-labeled agonists would not be expected. If this were the case, then 3H-labeled agonist binding should have been observed in the presence of Mg2+, which would stabilize the (H) state. Furthermore, in the neonatal lung, Gpp(NH)p shifts to the right the curve of norepinephrine inhibition of [3 Hlyohimbine, indicating that [3H]yohimbine is labeling at least to some extent the proposed (H) state of the receptor (Latifpour et al., 1982). A fourth unresolved issue concerns the differences in divalent cation-guanine nucleotide interactions at platelet and brain a2-receptor sites. At the former, Mg2+ facilitates the inhibition of high-affinity agonist binding by nucleotides (Fig. 2), whereas at the latter, Mg2+ and other divalent cations antagonize nucleotide effects on agonist binding, especially those of hydrolyzable nucleotides. T h e antagonism can be partially explained by the ability of divalent cations to accelerate nucleotide metabolism in the presence of brain membranes (Mallat and Hamon, 1982; Hamon et al., 1982), but some of these brain-specific interactions, such as increased agonist binding with low concentrations of GTP and Mg2+ present, may result from phosphorylation reactions specific to brain membranes, or may have to do with calmodulin influences on the receptor-cyclase system in neural tissue.

r

D. REGULATION OF (Yz-ADRENERGICRECEPTORS

1. Homologou~Regulation a . L'p-Regttlntiou. In common with most other hormone and neurotransmitter receptors, a2receptors appear to be regulated in such a manner as to compensate for changes in agonist concentration or presynaptic input. For a2 receptors, up-regulation is, in general, operationally defined as an increase in binding site number indicating an increase in receptor density. Only rarely have concomitant increases in receptor function been measured. Up-regulation has been shown to occur either as a result of diminished presynaptic function (e.g., depletion of norepinephrine with reserpine treatment, or chemical or surgical denervation) or chronic occupation of the receptor by an antagonist. An important caveat concerning a2 receptor up-regulation studies is that many of them have utilized only H-labeled agonist ligands and thus

CHARACTERIZATION OF 0 1- AND ff2-ADRENERGIC RECEPTORS

4 11

increases in the number of 3H-labeledagonist sites can be interpreted as shifts in equilibrium toward high-affinity states of the receptor without necessarily involving any change in the total receptor population. Early studies indicated that chemical denervation of rat brain noradrenergic neurones with intracerebroventricular 6-hydroxydopamine (6-OHDA) increased the number of brain a,-receptor sites labeled with [3H]clonidine(U’Prichardet al., 1977a). It was later shown that the same treatment increased the number of higher affinity pH]clonidine-labeled sites in 15 brain areas, with no change in the number of lower afhity p Hlclonidine sites (U’Prichard et al., 1979a). Chronic reserpine treatment also increased with the number of rat cortex a2 receptors labeled by [3H]epinephrine, with no change in [3H]epinephrine affinity (U’Prichard and Snyder, 1978a),as did 6-OHDA treatment (U’Prichard et al., 1979a). A specific lesion of the ascending dorsal noradrenergic bundle in rats also increased p H]clonidine binding in several forebrain areas (U’Prichard et al., 1980b). The rat submandibular gland has been a useful tissue for the study of a2 up-regulation, since no significant [3 Hlclonidine or Hlyohimbine binding is observed in membranes from normal glands. Chronic reserpine treatment or surgical denervation induces the appearance of a2 receptors in membranes from this gland (Bylund and Martinez, 1980, 1981; Pimoule et al., 1980; Bylund et al., 1982a). These a2receptors are labeled by [3H]clonidineand [3 H]p-aminoclonidine and are located postsynaptically. This phenomenon may represent a true de novo generation of postsynaptic a2 receptors in the tissue as a result of interruption of neuronal input. Interestingly, the adjoining sublingual gland, which receives very limited sympathetic input (the submandibular, by contrast is richly innervated), normally has a high level of a2receptors, and the level is further increased by reserpine treatment (Martinez et al., 1982b). Significant increases in [3H]clonidine binding in the rat submandibular gland occur within a few hours after the onset of reserpine administration and then decrease rapidly following termination of reserpine treatment (Fig. 6), indicating that expression of a2 receptors in this tissue is particularly adaptable (Bylund et al., 1982a). Although chronic occupation of a2receptors with a potent antagonist such as mianserin leads to an increase of a2-receptor function (Cerrito and Raiteri, 1981), few data showing a concomitant increase in a 2 receptor number are yet available. Treatment with yohimbine for 3 days reportedly increases the B,,, of pH]clonidine in rat cortical membranes (Johnson et al., 1980),and 7 days of yohimbine administration results in a modest increase in a2 receptors in the rat submandibular gland (Bylund et al., 1982a).

r

412

DAVID B. BYLUND AND DAVID c . U'PRICHARD

6.0-

(9)

,

2 4 6 DAYS OF RESERPINE TREATMENT

I

*

4

DAYS AFTER END OF TREATMENT

FIG. 6. The effect of the duration of reserpine treatment on the apparent density of a,-adrenergic receptor binding sites in submandibular gland. Rats were treated daily with reserpine (0.5 mglkg, i.p.) and then sacrificed 24 hr after the last injection (except for the first two time points which were 6 and 12 hr following a single injection). Other rats were treated with reserpine for 7 days, and sacrificed 2, 3, or 5 days after the last injection. Values given are means ? SEM.

6. DoziwRegulation. Receptor down-regulation is believed to be a regulatory response of the cell when it is exposed over a period of time to higher than normal concentrations of agonist. Loss of responsiveness to the agonist can be due to two events which may be associated: uncoupling of the receptor from its effector; and loss of receptors from the cell membrane concomitant with receptor internalization. Perkins and colleagues have shown for p receptors that there is a concomitant reduction in receptor number and receptor-related response during exposure to agonist. However, the loss of response precedes loss of receptors, because the first phase of desensitization involves uncoupling of the receptor from adenylate cyclase [also evidenced by conversion of (H) to (L) states of the receptor] before there is any change in receptor number (Harden rt al., 1979; Su ~t al., 1980). Approaches toward examining the mechanisms of a,-receptor down-regulation involve the exposure to agonists of a,-receptor-containing cells such as platelets, NG 108- 15 cells, and adipocytes, and examining the effects on a,-receptor function and number following treatment with antidepressants, which increase agonist concentration in the synapse by inhibiting norepinephrine reuptake. An early study indicated that incubation of intact human platelets with 100 pM (-)-epinephrine gradually reduced by 50% the number of

CHARACTERIZATION OF

AND fX2-ADRENERGIC RECEPTORS

413

rH]DHEC a2-receptor sites over a 4-hr period (Cooper et al., 1978), suggesting agonist-induced down-regulation. Over the time of exposure, epinephrine gradually induced total refractoriness of the aggregating response to catecholamines. However, more recently, Insel and colleagues have found that decreases in [3H]yohimbinebinding after a 4-hr exposure to 100 p M (-)-epinephrine are due to competition at the [3 Hlyohimbine sites by epinephrine taken up by platelets and retained even after extensive membrane washing. Thus in membranes from epinephrine-exposed platelets, the K, of [3 Hlyohimbine was increased with no change in B,,,, and the assay of these membranes in the presence of Na+ and GTP completely restored platelet Hlyohimbine binding (Karliner et al., 1982). A major problem with the human platelet as a model system for examining a2-receptor down-regulation is that activation of the platelet a2 receptor alters the characteristics of platelets in fundamental ways by stimulating the aggregation response. Regulation of NG 108-15 cell a2 receptors as a function of exposure to (-)-epinephrine can be studied in intact cells, grown in a serum-free defined medium at 37°C (Kahn and U’Prichard, 1983). Within the first 30 min of exposure to 10 p M (-)-epinephrine, there appears to be a shift toward high-affinity states of the receptor (increased B,,, of [3H]epinephrine and [3H]p-aminoclonidine, and decreased IC50 of epinephrine and p-aminoclonidine at [3H]rauwolscine sites) with no change in the total receptor population (B,,, of rH]rauwolscine) in Gpp(NH)p-treated membranes from exposed cells. More prolonged exposure of the cells to 10 pM (-)-epinephrine causes apparently parallel decreases in the B,,, of rH]rauwolscine, [3H]epinephrine, and H]paminoclonidine. However, although epinephrine exposure does not change the KD of [3H]rauwolscine at residual sites, it causes a timedependent increase in the KD of THlepinephrine and rH]paminoclonidine and an increase in the ICJo of epinephrine and p-aminoclonidine competing at H]rauwolscine sites. Thus prolonged exposure to epinephrine causes a selective loss of high-afhity states of the NG 108-15 a2receptor, as is observed for /3 receptors (Suet al., 1980; Kent et al., 1980). The ED50 for (-)-epinephrine in causing a reduction in [3H]rauwolscinesites after 8 hr exposure was 1.0 nM, similar to the potency of epinephrine at high-affinity states of the receptor. The initial increase in 3H-labeled agonist sites at shorter exposure times may represent uncoupling of the a2 receptor from adenylate cyclase, with the induction of more (SH) states (Kahn and U’Prichard, 1983). Unlike @receptor systems, exposure to agonist does not alter the maximum a2-receptor response [inhibition of adenylate cyclase activity in washed membranes with 100 pM (-)-epinephrine] but causes a twofold reduc-

r

r

414

DAVID B. BYLUND A N D DAVID

c. U’PRICHARD

tion in the apparent potency of (-)-epinephrine in inhibiting cyclase, indicating that N G 108-15 cells have “spare” a2 receptors in addition to “spare” opiate receptors (Fantozzi at al., 1981). Another phenomenon associated with exposure of NG 108-15 cells to agonists for negatively coupled receptors, but not seen with P-receptor systems, is a gradual increase in basal adenylate cyclase activity (Sabol and Nirenberg, 1979b), which may result from decreased ability of inhibitory N proteins to couple to C. Our experiments indicate that epinephrine exposure can increase basal cyclase at early exposure times (Kahn and U’Prichard, 1983). Similar a,-receptor desensitization experiments have been performed with isolated human properitoneal adipocytes by incubation for 3 hr at 37°C with 10 pLVf(-)-epinephrine (in the presence of propranolol to block p receptors). Although exposure to (-)-epinephrine reduced the B,,, of [3H]p-aminoclonidine by 4356, neither the B,,,, of [3H]yohimbine nor the ability of 10 p.M epinephrine (plus propranolol) to lower CAMPlevels in the cells was affected (Burns at af., 1982a). It is possible that adipocytes also have spare a2 receptors, and that for these cells a 3-hr exposure was not long enough to observe the second phase of desensitization, that is, a decrease in total receptor number. When rats under mild restraint were chronically infused with a-methylDOPA (which is metabolized to the at agonist amethylnorepinephrine) via the jugular vein for 72 hr, binding of both [3H]rauwolscine and [3H]p-aminoclonidine to cerebral cortex membranes was decreased by 30 and 50%, respectively. Infusion of the partial az-agonist clonidine for 72 hr had a bimodal effect on rat cortex [3H]p-aminoclonidine sites, with lowest concentrations of clonidine increasing ? H jp-aminoclonidine binding 50% above control, whereas higher doses decreased ? H]p-aminoclonidine binding 60 5% below control values. On the other hand, rat cortex [3H]rauwolscine binding was not affected at any clonidine concentration (U’Prichard et al., 1981). This dose-dependent regulation of a2-receptor high-affinity states with no change in total receptor population following a fixed time of agonist exposure may parallel the time-dependent bimodal regulation of agonist binding with a fixed concentration of agonist seen in N G 108-15 cells. Chronic treatment of rats with tricyclic and other antidepressant drugs, in addition to down-regulating brain PI receptors and the associated adenylate cyclase response, causes a functional desensitization of central and peripheral a2receptors (Crews and Smith, 1978; McMillen et 01.. 1980; Spyraki and Fibiger, 1980), as well as a functional a,-receptor supersensitivity (Menkes and Aghajanian, 1981). Binding studies are somewhat contradictory. Antidepressant treatment reduced the B,,, of

CHARACTERIZATION OF (Y1- AND (Y2-ADRENERGIC RECEPTORS

415

[3H]clonidine binding in brain regions other than cortex (Smith et al., 198l), whereas abbreviated (3 days) treatment with amphetamine, iprindole, or desmethylimipramine increased the number of higher affinity a2 sites in rat cortex membranes labeled with ?HIP-aminoclonidine or [3H]clonidine(Johnson et al., 1980; Reisine et al., 1980).In the absence of concomitant assessment of brain a2-receptor function, it is difficult to determine the significance of these binding changes. Chronic administration of various tricyclic antidepressant drugs to human patients appears to down-regulate platelet a2 receptors (decreased B,,, of ?HIclonidine) (Garcia-Sevilla et al., 198la,c). Chronic immobilization stress, which accelerates turnover of brain norepinephrine, down-regulates rat brain p receptors and increases the number of [3 Hlclonidine a2-receptor sites in cerebral cortex membranes, but decreases the affinity of [3H]clonidine and the number of ?HIclonidine sites in other brain areas (U’Prichard and Kvetnansky, 1980). 2. Heterologous Regulation

Numerous studies have now demonstrated that a2-receptor number can be influenced by other hormones. Estrogen treatment of immature female rabbits causes a three- to fourfold increase in the number of a2 receptors in uterine membranes, compared to tissues from progesterone-dominant animals, and in estrogen-primed tissue the a-receptor (contraction) response predominates over the &receptor response (relaxation) (Williamsand Lefkowitz, 1977b; Roberts et al., 1977). Estrogen treatment also decreases the contractile ED5, for norepinephrine compared to uteri from untreated animals, suggesting the presence of spare a2 receptors (Roberts et al., 1981). After 24 hr in organ culture, rabbit myometrium is more sensitive to a-adrenergic stimulation in vitro, and membranes from cultured myometrium have a threefold increase in a-receptor number, but no change in p receptors, suggesting that uterine a2receptors are under tonic inhibitory control in vivo (Cornett et al., 1981). Estrogen treatment has the opposite effect on rabbit platelet a2receptors, decreasing the density of PHIDHEC sites by 40-50% and diminishing the aggregation response to epinephrine (Roberts et al., 1979; Elliott et al., 1980). On the other hand, platelet membranes enriched or depleted in cholesterol, a membrane-stabilizingagent, exhibited increased or reduced responses to (-)-epinephrine, respectively, which was unaccompanied by any change in the number of a2 receptors labeled by the antagonist CJHIDHEC (Insel et al., 1978). Unfortunately, in this study a2-agonist binding parameters were not examined.

416

DAVID B. BYLUND AND DAVID

c.

U’PRICHARD

Activation of brain p receptors appears to lead to an increase in the number of high-affinity sites of the a 2 receptor in cerebral cortical membranes, either in vitro, when rat cortical slices were incubated with isoproterenol (Maggiet al., 1980),or in vivo, when isoproterenol is constantly infused intracerebroventricularlyfor up to 7 days via an Alzet minipump (Wang and U’Prichard, 1980). In the former studies, coincubation with the p antagonist sotalol prevented the isoproterenol-induced increase in H]p-aminoclonidine binding, whereas in the latter experiments, the increase in [3H]p-aminoclonidine sites was dependent on the time of infusion and amount of isoproterenol infused (Wang and U’Prichard, 1980). These data suggest that in membranes of rat cortical neurons, p and a2 receptors may be reciprocally regulated under some conditions.

r

3. Physiological Regulation The regulation of a2-receptor binding in response to normal physiologic changes has not been given much attention. Preliminary studies suggest that the number of sites labeled by [3H]yohimbine on human platelets varies during the normal female menstrual cycle (D. B. Bylund, unpublished). On the other hand, factors such as age, gender, and season do not appear to alter the density of platelet a2 receptors to any great extent. Many additional studies are needed in this area, at least as controls for the numerous studies on the regulation of human platelet a2 receptors in various pathologic states (Section III,D,4). The ontogeny of a2 receptors has been investigated in rat submandibular gland and brain. In the submandibular gland, the level of a2 receptors as determined by Hlclonidine and [3H]p-aminoclonidine binding at birth is relatively high (7 pmoVgm tissue) and then increases markedly during the first 2 weeks of life (Bylund et al., 1982b). Thereafter, the binding decreases such that at 6 weeks it approaches the very low level observed in glands from adult animals. By comparison, the level of [3H]clonidine binding in rat brain is low at birth, increases severalfold during the first 3 weeks, and then decreases slightly to adult levels (Morris et al., 1980).

r

4. Patholopal Regulation

The existence of a2receptor on human platelets, an easily obtainable tissue, has led to investigations of possible abnormal number, function, or regulation of a2 receptors in pathological states. Thus, platelets from patients with essential thrombocytopenia, which fail to aggregate or release serotonin in response to epinephrine, contained less than 50% of the normal complement of a2-receptor sites as measured with rH]DHEC (Kaywin et al., 1978).

CHARACTERIZATION OF

al-A N D

(Y~-ADRENERGICRECEPTORS

417

The hypofunction of central noradrenergic systems, believed to underlie the pathology of endogenous depression, may be associated with an abnormal increase in a,-receptor function, which in turn may be reflected as an increase in total receptor number or an enhanced coupling to adenylate cyclase. If this abnormality had a genetic basis, it could be expressed also in platelets as well as brain tissue. Although the results are conflicting, several laboratories have examined platelet a , receptors in unmedicated depressed patients. Smith and colleagues observed an increase in platelet [3H]clonidinebinding in a small group of depressed patients (Garcia-Sevilla et al., 198lc), whereas Meltzer and co-workers have observed no changes in the K , or B,,, of THIyohimbine binding to a , receptors in platelet membranes from unipoiar or bipolar depressives (Daiguji et al., 1981b). Basal platelet CAMPproduction was reported to be reduced in samples from male schizophrenic patients, but no change in platelet a,-receptor number or function (percentage inhibition of adenylate cyclase) was observed (Kafka et al., 1979). However, no change in p Hlyohimbine binding was found in unmedicated schizophrenics,but there was an increase in B,,, in a small group of patients termed schizoaffective (U’Prichard et al., 1982). In another study, no alteration in platelet THIyohimbine binding was found when five patients with Parkinson’s disease were compared to appropriate controls (D. Bylund, unpublished). Since centrally acting antihypertensive drugs such as clonidine and a-methylnorepinephrine (formed from a-methylDOPA) are potent a,receptor agonists, animal models of hypertension have been examined for possible a,-receptor changes. In genetically hypertensive (SHR) rats, a 35% increase in the number of a,-receptor sites labeled with pH] clonidine in hypothalamic membranes has been reported (Morris et al., 1981). However, THIyohimbine binding is unaltered in most tissues of the deoxycorticosterone/salthypertensive rat (Yamada et al., 1980a).

E. LOCALIZATION OF (Y2-ADRENERGIC RECEPTORS a, Receptors were originally defined in terms of their capacity to modulate norepinephrine release and were presupposed to occur on presynaptic terminal membranes. However, the Occurrence of presynaptic a , receptors in brain and peripheral tissues by means of binding studies has not yet been demonstrated, due perhaps in large part to two complicating factors. First, if presynaptic and postsynaptic a , receptors occur in the same tissue and are pharmacologically indistinguishable, attempts to establish the presynaptic location of receptors involving de-

418

DAVID B . BYLUND A N D DAVID

c . U’PRICHARD

nervation of the noradrenergic input will be bedevilled by the probability of postsynaptic receptor supersensitivity masking the loss of presynaptic receptors. T h e issue is compounded by the necessity of waiting at least 2 to 4 weeks after denervation to allow complete phagocytic removal of presynaptic membranes which could, of course, still be labeled with receptor ligands even though presynaptic function has long been lost. Second, studies published so far have almost exclusively involved the use of as-agonist radioligands, although changes in the number of 3H-labeled agonist sites do not necessarily reflect changes in the az-receptor density of the tissue. T h e equation of at receptors with a presynaptic location has become steadily weakened in recent years with the discovery of a 2 receptors on noninnervated cells such as platelets and isolated adipocytes, and by more recent pharmacological evidence that adrenergic vasoconstriction involves vascular postjunctional a 2 , as well as a l , receptors (Timmermans and van Zwieten, 1980; Kobinger and Pichler, 1981; Ruffolort al., 1980). Alphae receptors have been identified on bovine aorta membranes (Rosendorff et al., 1981). Some authors have gone so far as to suggest that at adrenergic neuroeffector junctions, postsynaptic (possibly a e )receptors may have trans-synaptic effects on terminal norepinephrine uptake and release (Manukhin and Volina, 1979). With the above caveats in mind, VHIclonidine binding studies indicate that rat brain a2 receptors are predominantly “postsynaptic,” in that they are not located on noradrenergic terminals (U’Prichard et al., 1977a, 1979a). After a dorsal noradrenergic bundle lesion, however, rH]clonidine binding was decreased in amygdala and septum, suggesting that in these brain areas there might be a greater prevalence of presynaptic receptors (U’Prichard et al., 1980b). Similarly, az-receptor sites when present, appear to be postsynaptic in rat submandibular gland (Bylund and Martinez, 1981) and kidney (McPherson and Summers, 1982). There has been some controversy over the occurrence and location of rat heart a 2receptors. Guicheney and co-workers obtained evidence for an a,-receptor component of tH]DHEC binding to rat heart membranes and referred to this component as “presynaptic,” but did not demonstrate its location (Guicheney et a/., 1978). Langer and colleagues observed that ventricular HIDHEC binding was decreased 60% after chemical sympathectomy, but a pharmacological differentiation of a 1 and a2 components of rH]DHEC binding was not performed (Briley et nl., 1979; Story rt a/., 1979). In contrast, several laboratories have found no evidence of a2-receptor binding to rat heart membranes using r H ] clonidine (U’Prichard and Snyder, 1979), rH]yohimbine, or VHIDHEC (R. J. Lefkowitz, personal communication).

r

CHARACTERIZATION OF al- AND (Y~-ADRENERGICRECEPTORS

4 19

Another approach to the localization of a2receptors in the brain has been the labeling of slide-mounted brain tissue sections with rH]paminoclonidine, and subsequent processing for autoradiography. Tritiated p-aminoclonidine binding in these conditions was shown to be saturable, a2 receptor specific, and highly concentrated in certain brain regions such as locus coeruleus and nucleus tracti solitarii where a2 receptors have been demonstrated electrophysiologically (Young and Kuhar, 1979, 1980). The autoradiographic distribution of az-and opiate receptor sites was found to be very similar in many brain regions (Young and Kuhar, 1980), which is of interest in view of the coexistence of a2 and &opiate receptors in many neuroblastoma cell lines (Hamprecht, 1977; D. J. Kahn and D. E. U'Prichard, unpublished). An intriguing finding in this connection is that intravenous clonidine infusion in rats causes biphasic alterations in the binding to cortex membranes of the &opiate agonist [3H]d-Alad-Leu-enkephalin, which exactly parallel the changes in H]p-aminoclonidine binding (U'Prichard et d.,198 1).

r

F. SOLUBILIZATION OF QZ-ADRENERGJC RECEPTORS

Human platelet a2 receptors have been successfully solubilized by Smith and Limbird (1981) using the detergent digitonin. The characteristics of [3H]yohimbinebinding ( K DandB,,,) are the same for membranes and solubilized preparations. The solubilized a2receptors labeled with [3H]yohimbineappear to be in the low-affinity state, with ICsOvalues for agonist competitors equal to the IC,, values in membrane preparations in the presence of 100 p M Gpp(NH)p. Thus, solubilization of platelet membranes with digitonin appears to dissociate R-N complexes. On the other hand, prelabeling membrane-bound receptors with the agonist VHIepinephrine confers some stability on the ternary complex during solubilization, since high-affinity, guanine nucleotide-dissociable [3 Hlepinephrine binding was retained after digitonin treatment. [3H]Epinephrine-labeled receptors sedimented more rapidly through a continuous sucrose gradient after solubilization than did [3H]yohimbine-labeledreceptors, suggesting that the agonist-receptor complex had a larger protein mass. These results support the notion that high-affinity agonist binding represents the ternary H-R-N complex, whereas after solubilization antagonist binding is to the free R species. Essentially, similar results have been obtained by Lefkowitz and colleagues (Michel et d.,1981). Recently, clonidinep-isothiocyanatehas been used as an affinity label for platelet a2-adrenergic receptors (Atlas and Steer, 1982). Vipoxin, a

420

DAVID

B. BYLUND

AND DAVID

c. U’PRICHARD

protein from Russell’s viper venom, appears to have a high affinity for a,-receptor sites, and the binding is essentially irreversible (Freedman and Snyder, 1981). These two probes, particularly in their radioactive form, should be very useful in the further study of a , receptors in areas such as regulation and purification.

IV. Summary and Conclusions

Within the short period of 5 years, the availability of a variety of specific radioligands has allowed the resolution of a1- and a , -adrenergic receptor populations in many different tissues and enabled researchers to begin investigations of the mechanisms of regulation and coupling of a l and a, receptors to their different cellular effector systems. Binding data have demonstrated that the pharmacological properties of each type of a receptor are, in general, similar across tissues and species, although there are some differences in the relative affinities of antagonist drugs. Further attempts to subclassify a1 and a , receptors may be expected in the future. T h e historical development of the interpretation of pH]clonidine binding is of interest in this regard. [3H]Clonidine was proposed to label the “agonist state” of the a receptor, and then to label a2 receptors. It is now thought that it labels the agonist state of a, receptors. Might it actually label a subpopulation of a , receptors or just the agonist state of that subpopulation? Alpha-1 receptors by and large appear to occur in a single-affinity state with respect to both agonists and antagonists. By comparison, a , receptors may exist in multiple-afbnity states reflecting the ability of the a, binding site protein to complex to additional membrane proteins which themselves are receptors for the physiological substrates GTP, Na+, Mgz+, and possibly Ca2+-calmodulin. Binding studies have also strongly indicated that a, receptors in most, if not all, tissues are probably coupled in an inhibitory manner to adenylate cyclase, as has been demonstrated in platelets, adipocytes, and NG 108-15 cells. Clearly the present status of a-receptor research has left many questions unresolved. We still have no idea what membrane effector system and associated second messenger is coupled to the a 1 receptor. T h e prevailing belief is that Ca2+ and the membrane Ca2+ channel fulfill these roles. However, others have suggested that phosphoinositide turnover represents the proximal receptor response, and indeed a membrane-bound phospholipase C may play an analogous role to adenylate cyclase for other adrenergic receptors (Putney et al., 1980). There is,

CHARACTERIZATION OF

a1-AND

a2-ADRENERGIC RECEPTORS

42 1

however, some evidence that in some situations a1receptors may directly stimulate adenylate cyclase, and guanine nucleotide modulation of agonist affinities at a1-receptor sites has been reported. The significance of these data and reported modulatory effects of Na+ at a1 receptors (Glossmann and Presek, 1979; Glossmann et al., 1981) is still to be resolved. Nothing is known about the mechanisms of regulation of al receptors, although both up- and down-regulation of a1 receptor have been demonstrated. In this regard, the ability to label and study a1 receptors on cells in culture would be particularly useful. With regard to a2receptors, it is still not clear how many affinity states exist and what their role is in terms of the kinetics of a2-receptor coupling to adenylate cyclase. In particular, the interaction of the proposed (SH) state of the receptor with the catalytic enzyme moiety is unresolved. Some questions concerning a2-receptor coupling and function are common to all inhibitory receptors: 1. What is the protein subunit composition of the regulatory proteins (Ni) associated with inhibitory receptors (Rodbell, 1980)? Are they analogous to the subunits of N, (Northup et al., 1980)? 2. What are the similarities and differences between Ni and N,? 3. How does activation of inhibitory receptors result in decreased cyclase activity?

Other questions related to a2 receptors concern whether there are indeed sequential phases of desensitization involving uncoupling and loss of receptor protein. For neural a , receptors, future studies must determine what the role of calmodulin is in the coupling and regulation of a , receptors. More generally, it is unclear at what membrane site Na+ interacts to regulate agonist affinities at the receptor and in particular if the site is associated with the Ni complex. Clearly, the ability to measure a,-receptor populations and the associated response in homogeneous cell populations has been particularly advantageous. By analogy to the mouse lymphoma S49 cell @-receptorsystem, the development of selection pressures to produce mutants of a,-receptor-containing cultured neural cells deficient in one or other aspect of the receptor-effector system would lead to great advances. Indeed, receptor-deficient variants of NG 108-15 cells are currently being developed in several laboratories. For these and other studies, the development of radio-iodinated a2receptor probes would prove very beneficial, particularly to resolve decisively the issue of the existence of nerve terminal “autoreceptors,” whose density would be expected to be very low.

422

DAVID B . BYLUND A N D DAVID

c . U’PRICHARD

The successful solubilization of platelet az receptors is sure to be followed by solubilization of az receptors in other tissues. Over the next few years, it is reasonable to expect both a 1 and az receptors to be purified at least to the extent that specific receptor antibodies can be produced by monoclonal techniques. One would also predict that reconstitution studies along the lines developed by Gilman and colleagues for p receptors (Ross and Gilman, 1980) will resolve the issue of the uniqueness of Ni for inhibitory receptors. These investigations and others concerning the mechanisms of coupling and regulation of a-adrenergic receptors have great therapeutic relevance in view of the widespread use of &-receptor agents such as prazosin, clonidine, and a-methyIDOPA in the treatment of hypertension, together with the likely development of new therapies for endogenous depression based on az-adrenergic receptors.

References

Aggerbeck, M., Guellaen, G., and Hanoune, J. (1980). Riochrm. Phannncol. 29, 643-645. .4hlquist, R. P. (1948). A m . J. Phyiol. 153. 586-600. Aktories, K., Schultz, G., and Jakobs, K. H . (1980). Saicriyti-.Sr~hmir~lr6~~’.i Arch. Pharmncol. 312, 167-173. Aklories, K., Schultz, G., and Jakobs, K . H. (1981). Bzochim. Bi0phy.v. Arta 676, 59-67. Alexander, R. W.. Cooper, B., and Handin, R. 1. (1978).J. Clin. I n i w t . 61, 1136- 1144. A n d e r s o n , R. (1973). Artn Physiol. Scarid. 87, 84-95. Amett, C. D., and Davis, J. N. (1979).J. Phnrmnrol. Exp. Ther. 211, 394-400. Atlas, D., and Adler, M. (1981).Proc. A‘ntl. .4rad. Sri. Cr.S.A. 78, 1237-1241. Atlas, D., a n d Sabol, S. L. (1981). Eur. J . Biochtm. 113, 521-529. Atlas, D., a n d Steer, M. L. (1982). Proc. .Vatl. Acad. Sri. U.S.A. 79, 1378-1382. Barber, A. J., and Jamieson, G. A. (1970).J. Biol. C h m . 245,6357-6365. Barnes, P. J., Karliner, J. S., Hamilton, C. A., and Dollery, C. T. (1979). Lqr Sci. 25, 1207- 1214. Barnes, P. J.. Karliner, J. S., a n d Dollery, C. T. (1980). Clin. Sri. 58, 457-461. Barthel, W., and Markwardt, F. (1974). Biorhem. Phnrmnrol. 23, 37-45. Battaglia, G., and Titeler, M. (1980). .Yrut-osri. A6.ctr. 6, 852. Batzri, S., Selinger, Z., Schramm, M., and Robinovitch, M. R. (1973).J. Biol. Chem. 248, 361-368. Berlan, M., a n d Lafontan, M. (1980). Etrr. J. Phai-tmcol. 67, 481-484. Berthelsen, S., and Pettinger, W. A. (1977). Life Sri. 21, 595-606. Birdsall, IV. J . M., Burgen, A. S. V., and Hulme, E. C. (1978).illol. Pharmncol. 14,723-736. Birnbaumer, L., Bearer, C. F., a n d Iyengar, R. (1980).J. Riol. Chem. 255, 3552-3557. Bittiger, H., Heid, J., and Wigger, N. (1980). Nature (London) 287, 647-649. Blume, A. J. (1978). Lifr Sri. 22, 1843-1852. Blume. A. J., Lichtstein, D., a n d Boone, G. (1979). Proc. A‘atl. Arad. Sri. L’.S.A. 76, 56265630.

CHARACTERIZATION OF (Y1- AND (Y2-ADRENERGIC RECEPTORS

423

Boudier, H. S., deBoer, J., Smeets, G., Lien, E. J., and van Rossum, J. (1975). Life Sn‘. 17, 377. Brandt, M., Buchen, C., and Hamprecht, B. (1980).J. Neurochem. 34,643-651. Braunwalder, A., Stone, G., and Lovell, R. A. (198ll.J. Neurochem. 37, 70-78. Briley, M. S., Langer, S. Z., and Story, D. F. (1979). Br. J. Pharmacol. 66, 9OP. Brostrom, M. A., Brostrom, C. O., and Wolff, D. J. (1978). Arch. Biochem. Biophys. 191, 341-350. Brostrom, M. A., Brostrom, C. 0.. Huang, S. C., and Wolff, D. J. (1981).Mol. Pharmucol. 20, 59-67. Brown, E. M., Hurwitz, S. H., and Aurbach, G. D. (1978). Endocrinology 103, 893-899. Brown, G. L., and Gillespie, J. S. (1957).J. Physiol. (London) 138,81-102. Buonassisi, V., and Venter, J. C. (1976). Proc. Natl. Acad. Sci. U.S.A. 73, 1612-1616. Burns, T. W., and Langley, P. E. (1975).J. Cyclic Nucleotide Res. 1, 921-328. Burns, T. W., Langley, P. E., Terry, B. E., Bylund, D. B., Hoffman, B. B., Tharp, M. D., Lefkowitz, R. J., Garcia-Sainz,J. A,, and Fain, J. N. (1981).J. Clin. Invest. 67,467-475. Burns, T. W., Langley, P. E., Terry, B. E., and Bylund, D. B. (1982a). Metab., Clin. Exp. 31, 288-293. Burns, T. W., Langley, P. E., Terry, B. E., Bylund, D. B., and Forte, L. R. (1982b).LifeSn’. 31,815-821. Bylund, D. B. (1979).In “Modulators, Mediators, and Specifiers in Brain Function” (Y. H. Ehrlich, J. Volavka, L. G. Davis, and E. G. Brunngraber, eds.), p. 133. Plenum, New York. Bylund, D. B. (1981).J . Pharmacol. Ex$. Ther. 217, 81-86. Bylund, D. B., and Martinez, J. R. (1980). Nature (London) 285, 229-230. Bylund, D. B., and Martinez, J. R. (1981).J . Neurosci. 1, 1003-1007. Bylund, D. B., Forte, L. R., Morgan, D. W., and Martinez, J. R. (1981).J. Pharmacol. Exp. Ther. 218, 134-141. Bylund, D. B., Martinez, J. R., and Pierce, D. L. (1982a). Mol. Pharmacol. 21, 27-35. Bylund, D. B., Martinez, J. R., Camden, J., and Jones, S. B. (198213). Arch. Oral Biol. 27,945-950. Cannon, W. B., and Rosenblueth, A. (1937). “Autonomic Neuroeffector Systems.” MacmilIan, New York. Cantor, E. H., Abraham, S., and Spector, S. (1981). Life Sn’. 28, 519-526. Cavero, I., Fenard, S., Gomein, R., Lefevre, F., and Roach, A. G. (1978).Eur.J. Pharmacol. 49,254. Cech, S. Y., Broaddus, W. C., and Maguire, M. E. (1980). Mol. Cell. Biochem. 33, 67-92. Cerrito, F., and Raiteri, M. (1981). Eur.J. Pharmacol. 70, 425-426. Cohen, J., Eckstein, L., and Gutman, Y. (1980). Br. J . Pharmacol. 71, 135-142. Colquhoun, D. (1973). In “Drug Receptors” (H. P. Rang, ed.), pp. 149-182. Univ. Park Press, Baltimore, Maryland. Cooper, B., Handin, R. I., Young, L. H., and Alexander, R. W. (1978).Nature (London) 274, 703-706. Cooper, D. M. F., Schlegel, W., Lin, M. C., and Rodbell, M. (1979).J . Biol. Chem. 254, 8927-8931. Cornett, L. E., Goldfien, A,, and Roberts, J. M. (1981). Nature (London) 292, 623-625. Crews, F. T., and Smith, C. G. (1978). S c i m e 202, 322-324. Crutcher, K. A., and Davis, J. N. (1980). Brain Res. 182, 107-117. Daiguji, M., Meltzer, H. Y., and UPrichard, D. C. (1981a).Life Sn’. 28, 2705-2717. Daiguji, M., Meltzer, H. Y., Tong, C., U’Prichard, D. C., Young, M., and Kravitz, H. (1981b).Life Sn’. 29, 2059-2064.

424

DAVID B. BYLUND A N D DAVID

c. U'PRICHARD

Dale, H. H. (1906).J. Phyiol. (London) 34, 163-206. Davis, J. N., Arnett, C. D., Hoyler, E., Stalvey, L. P., Daly, J. W., and Skolnick, P. (1978). Brain Rrs. 159, 125-135. Davis, J. N., Olender, E., Maury, W., and McDaniel, R. (1980).2Mol. Pharmacol. 18, 356361. DeLean, A., Stadel, J . M., and Lefkowitz, R. J. (1980).J . B i d . Chrm. 255, 7108-7117. d e Peusner, I . C. W.. Perec, C. J., and Stefano, F. J. E. (1979).i V a u n v ~ f - S c l f m i e d e Arch. b~~s Pharmcol. 308,217-221. Desmedt, D. H. (1980). Biochtm. Phnrinncol. 29, 1966-1968. Doxey, J. C., Smith, C. F. C., and Walker, J. M. (1977).Br. J . Pharmacol. 60, 91-96. Drew, G. M., and Whiting, S. B. (1979). Br. J . Phnrmacol. 67, 207-215. Ehlert. F. J., Roeske, W. R., and Yamamura, H. I. (1980).J. Sziprnmul. Struct. 14, 149-162. Elliott, J. M., Peters, J. R., and Grahame-Smith, D. G. (1980). Eur.J. Pharmacol. 66,21-30. El-Refai, M. F., and Exton, J. H. (1980). Eur. J. Pharmacol. 62, 201-204. El-Refai, M. F., Blackmore, P. F., and Exton, J. H. (1979).J . B i d . Chrm. 254, 4375-4386. Engel, G., and Hoyer, D. (1981). Eur. J. Pharmacol. 7 3 , 221-224. Fain, J. N., and Garcia-Sainz, J. A. (1980). Life Sci. 26, 1183-1 194. Fantozzi, R., Mullikin-Kilpatrick, D., and Blume, A. 1. (1981)..Zlol. Pharmacol. 20, 8-15. Forte, L. R., Bylund, D. B:, and Zahler, W. L. (1982).Ftd. Proc., Fed. A m . SOC.Exp. B i d . 41, 1471. Freedman, J. E., and Snyder, S. H. (1981).J. Biol. CIwni. 256, 13172-13179. Fuder, H., Nelson, W. L., Miller, D. D., and Patil, P. N. (1981).J . P h n m f Z C C J [ . Exp. Thw. 217, 1-9. Garcia-Sainz, J. A., Hasler, A. K., and Fain, J. N. (1980a). Biochvm. Phnrmarol. 29, 33303333. Garcia-Sainz, J. A., Hoffman, B. B., Li, S.-Y., Lefkowitz, R. J., and Fain, J. N. (1980b). Lift Scz. 27, 953-961. Garcia-Sevilla, J. A., Zis, A. P., Zelnick, T. C., and Smith, C. B. (1981a).Eur. J . Pharmacol. 69, 121-123. Garcia-Sevilla, J. A., Hollingsworth, P. J., and Smith, C. B. (1981b). Eur. J. Pharmacol. 74, 329-34 1 . Garcia-Sevilla, J. A., Zis, A. P., Hollingsworth, J. P., Greden, J. F., and Smith, C. B. (198 lc). ?4rrh. Gen. Pqrhiati? 38, 1327-1333. Gardey-Levassort, C., Richard, M.-O., delauture, D., Thiroux, G., and Olive, G. (1981). Life Sn. 28, 331-337. Geynet, P., Ferry, N., Borsodi, A., and Hanoune, J. (1981).Biochrm. Pharmnrol. 30, 16651675. Gheyouche, R., LeFur, G., Colotte, O., Burgevin, M. C, and Uzan, A. (1980).,]. Pharm. Plinrmacol. 32, 366-368. Gill, D. M.,and Meren, R. (1978). Pror. Satl. Arad. Sri. U.S.A. 75, 3050-3054. Girardot. J . M., Kempf, J., and Cooper, D. M. F. (1983).J. h'eurorhem. (in press). Glossmann, H., and Hornung, R. (1980a). Eur. J. Pharmacol. 61, 407-408. Glossmann, H.. and Hornung, R. (1980b). . ~ a u , i ~ i i - S r h m i e ~ rArch. b ~ ~ sPhaimacol. 312, 105- 106. Glossmann, H., and Hornung. R. (19ROc). h h u r i ~ i f - S c h m i e d e b wArch. ~ ~ Pharmnrol. 314, 101-109. Glossmann, H., and Presek, P. (1979). ,~ciuri?n-Srh.miedebergs'4rch. Pharmncol. 306, 67-73. Glossmann, H., Lubbecke, F., Bellemann, P., and Presek, P. (1981).Eur. J . Pharmacol. 75, 149-153. Graham, R. M., Hess, H., and Homcy, C. J. (1982). Proc. LVatl.Acad. Sci. U.S.,4. 79, 21862190.

CHARACTERIZATION OF CY1- AND CY2-ADRENERGIC RECEPTORS

425

Grant, J. A,, and Scrutton, M. C. (1979). Nature (London)277, 659-661. Greenberg, D. A,, and Snyder, S. H. (1978). Mol. Pharmacol. 14, 38-49. Greenberg, D. A,, U’Prichard, D. C., and Snyder, S. H. (1976). Life Sci. 19,69-76. Greenberg, D. A,, U’Prichard, D. C., Sheehan, P.,and Snyder, S. H. (1978).Brain Res. 140, 378-384. Greengrass, P., and Bremner, R. (1979). Eur. J . Pharmacol. 55, 323-326. Guellaen, G., Aggerbeck, M., and Hanoune, J. (1979).J. Biol. C h a . 254, 10761-10768. Guellaen, G., Goodhardt, M., Barouki, R., and Hanoune, J. (1982).Eiochern.Pharmacol. 31, 2817-2820. Guicheney, P., Garay, R. P., Levey-Marchal, C., and Meyer, P. (1978). Proc. Natl. Acad. Sci. U.S.A. 75, 6285-6289. Haga, T., and Haga, K. (1980). Life Sci. 26, 211-218. Haga, T., and Haga, K. (1981).J. Neurochm. 36, 1152-1159. Hamilton, C. A., and Reid, J. L. (1980). Br. J . Pharmacol. 70, 63-64. Hamon, M., Mallat, M., El Mestikawy, S., and Pasquier, A. (1982).J. Neurocha. 38, 162172. Hamprecht, B. (1977). Znt. Rev. Cytol. 49, 99-170. Harden, T. K., Su, Y. F., and Perkins, J. P. (1979).J. Cyclic Nucleotide Res. 5, 99-106. Harik, S. I., Virendra, K. S., Wetherbee, J. R., Warren, R. H., and Banerjee, S. P. (1980). Eur. J. Pharmacol. 61, 207-208. Harris, R. A., Schmidt, J., Hitzemann, B. A., and Hitzemann, R. J. (1981). Science 212, 1290. Harris, R. A., Fenner, D., Feller, D., Sieckman, G., Lloyd, S., Mitchell, M., Dexter, J. D., Tumbleson, M. E., and Bylund, D. B. (1983). Pharmacol., Eiochem. Behau. 18, 363367. Hasegawa, M., and Townley, R. (1981). Fed. Proc., Fed. Am. Soc. Exp. Bid. 40, 246. Hata, F., Takeyasu, K., Uchida, S., and Yoshida, H. (1980a). Eur. J . Pharmacol. 67, 193199. Hata, F., Uchida, S., Takeyasu, K., Ishida, H., and Yoshida, H. (1980b).Jpn.J. Pharmacol. 30, 570-574. Hedler, L., Stamm, G., Weitzell, R., and Starke, K. (1981). Eur. J. Pharmacol. 70, 43-52. Hiram, F., Strittmatter, W. J., and Axelrod, J. (1979). Proc. Natl. Acad. Sci. U.S.A. 76, 368-372. Hoffman, B. B., and Lefkowitz, R. J. (1980a).z4nnu.Rev. Pharmacol. Toxicol. 20, 581-608. Hoffman, B. B., and Lefkowitz, R. J. (1980b). N. Engl. J. Med. 302, 1390-1396. Hoffman, B. B., and Lefkowitz, R. J. (1980~). Biocha. Pharmacol. 29, 452-454. Hoffman, B. B., and Lefkowitz, R. J. (1980d).Eiochem.Pharmacol. 29, 1537-1541. Hoffman, B. B., DeLean, A., Wood, C. L., Shoc:ken, D. D., and Lefkowitz, R. J. (1979).Lfe Sci. 24, 1739-1746. Hoffman, B. B., Mullikin-Kilpatrick, D., and Lefkowitz, R. J. (1980a).J. Bid. Chem. 255, 4645-4652. Hoffman, B. B., Michel, T., Mullikin-Kilpatrick, D., Lefkowitz, R. J., Tolbert, M. E. M., Gilman, H., and Fain, J. N. N. (1980b). Proc. Natl. Acad. Sci. U.S.A. 77, 4569-4573. Hoffman, B. B., Dukes, D. F., and Lefkowitz, R.J. (1981a). Life Sci. 28, 265-272. Hoffman, B. B., Yim, S., Tsai, B. S., and Lefkowitz, R. J. (1981b). Biochem. Biophys. Res. Commun. 100, 724-731. Hoffman, B. B., Michel, T., Brenneman, T. B., and Lefkowitz, R. J. (1982). Endocrinology 110,926-932. Hsu, C. Y., Knapp, D. R., and Halushka, P. V. (1979).J. Pharmacol. Exp. Ther. 208, 366370. Insel, P. A., Nirenberg, P., Turnbull, J., and Shattil, S. J. (1978).Biochemistr~17,5269-5274.

426

DAVID B. BYLUND A N D DAVID

c. U'PRICHARD

lto, H., Hoopes, M . T., Baum, B. J., and Roth, G. S. (1982). Biorhrin. Pharmarol. 31,

567-573. Jakobs, K. H. (1978). Snirrru (Lui~doit)274, 819-820. Jakobs, K. H. (1979). '\fo/.CelI. Etidocriirol. 16, 147-156. Jakobs, K. H.. and Aktories, K. (1981). Biorhiin. Biophjs. .4rt0 676, 51-58. rlrrh. Phnnnncol. 310, 121-127. Jakobs. K. H., and Schultz, G. (1979).Snccrrjir-Srhnriedc6rr~~~ Jakobs, K. H., S a w , W.,and Schultz, G. (1978a). FEBS Lull. 85, 167-170. Jakobs, K. H., S a w , W.,and Schultz, G. (1978b). Jlol. Pharfnctco/. 14, 1073-1078. Jakobs, K. H., Aktories, K., and Schultz, G . (1979). h'ntirrTrt-Sc/imirdr~~~'.\ A w h . Phnrmncol. 310, 113- 119. Jard, S.. Cantau, B.. and Jakobs, K. H. (198l).J. B i d . Chtin. 256, 2603-2606. Johnson, R. W., Reisine, T., Spotnitz, S., Wiecb, N., Ursillo, R., and Yamamura, H. I. (1980). Eur. ./. Phcrrmrtcol. 67, 123- 127. Jonas, I)., Moritz, F., Jenner, S., and Baurngarten, H. G. (1980). C'rol. In/. 35, 47-62. 21, 191-195. Jones, D. J.. Kendall, D. E., and Enna, S. J. (1982). "Vrzirophnr~narolo~ Kafka, M. S., Tallman, J. F., and Smith, C. C. (1977). Lqr Sri. 21, 1429-1438. Kafkd, M. S., van Kammen, D. P., and Bunney, W. E. (1979). Am. J . Phsjchintr? 136, 685-687. Kafka, M .S., Wirz-Justice, A., and Naber, D. (1981). Brrtin Rex. 207, 409-419. Kahn, D. J., and U'Prichard, D. C. (1983).J. CTrlir 2Vrtrlro/iduRus. (in press). Kahn, D. J., Mitrius, J. C., and U'Prichard, D. C. (1982). ,\fol. Phrtrtnneol. 21, 17-26. Karliner, J. S.. Barnes, P., Hamilton, C. A., and Dollerp, C. T. (1979).Biorhm. Riophpy. RPS. C o m i n i i r t . 90, 142- 149. Karliner, J . S., Motulsky, H. J., and Insel, P. A. (1982). .\fol. Phnrmrtrol. 21, 36-43. Kather, H.. and Simon, B. (1981).Eirr. J . Clcn. I n z v s / . 11, 1 1 1-1 14. / . 345-348. Kather, H.. Pries, J., Schrader, V., and Simon, B. (1980). Ertr. J. O'lin. I n m ~ . ~10, Kavwin, P., McDonough, M., Insel, P. A., and Shattil, S. J. (1978). A'. Engl. J . ,Lfrd. 299, 505 - 509. Kent, R. S.. DeLean, A., and Lefkowitz. R. J. (1980).*\lo/. Phnrmncol. 17, 14-23. Kobinger, W'., and Pichler, L. (1981). Eur. J . Phurrnnrol. 73, 313-321. Koski. G.. and Klee, U'..4. (1981). Pror. .Vat/. .4rnd. Sri. l'.S..4. 78, 4185-4189. Kunos, G. (1978).,dnrirt. Rn,.Phnrirrnrol. Tfn'irol. 18, 291-31 I . Kuzuya. H., Ikeno. T., Ikeno, K., Nemoto, K., and Hashimoto, S. (1980).Arch. O r n l R i d . 25,31-36. Lafontan, Sf.,and Berlan. M. (1980). Eitr. J . Phnrmnrol. 66, 87-93. Lands, A. M.,McAuliff, 4 . A., Luduena, F. P., and Brown, T. G. (1967).N n t c ~ r r(London) 214, 597-398. Langer, S. Z. (1974).Bitrehuni. Phnrmrrro/. 23, 1793- 1800. Langer, S. 2. (1977).Bi-. J . Phnrmnrol. 60, 481-497. C . Corrgr. Phy.riol. Langer, S. Z., Adler, D., Enero, M. A , , and Stefano, F. J. E. (1971).P ~ C JInt. Scr. 25, 335. Langlev, J. N. (1905).J. Phjs-rol. (Londorr) 33, 374-413. ,4rch. Phnrmnrol. 306, 119-125. Lasch, P., and Jakobs, K. H. (1979). ~\~t~u,rjir-Srliirriud~brr~'s Latifpour, J. (198 1). Ph.D. Dissertation, University of Missouri, Columbia. Latifpour. J., and Bylund, D. B. (1981). Riorhrni. Phorinnrol. 30, 2623-2625. latifpour, J., Jones, S. B., and Bvlund. D. B. (1982).,]. Phnrinncol. Ezp. Thrr. 223, 606611. Lavin, I'.N.. Hoffman, B. B., and Lefkowitz, R. J. (1981). ,ifo/. Phnrmneo/. 20, 28-34. Lawrence, J . C., Jr., and Larner, J. (1977). i\JoI. Phnrrncicol. 13, 1060-1075. Lawrence, J. C.. Jr.. and Larner, J. (1978). .\fol. Pharnmrol. 14, 1079-1091.

CHARACTERIZATION OF a l - AND (Y~-ADRENERGICRECEPTORS

427

Lefkowitz, R. J. (1978). Fed. Proc., Fed. Am. Sor. Exp. B i d . 37, 123-129. Lenox, R. H., McMains, C. L., and Van Riper, D. A. (1980). Neurosci. Soc. Abstr. 6, 252. Levine, L., and Moskowitz, M. A. (1979).Proc. Natl. Acad. Sci. U.S.A. 76, 6632-6636. Lichtstein, D., Boone, G., and Blume, A. (1979).J. Cyclic Nucleotide Res. 5, 367-375. Limbird, L. E. (1981).Bi0chem.J. 195, 1-13. Linibird, L. E., Speck, J. L., and Smith, S. K. (1982). Mol. Phannacol. 41, 607-619. Lin, M. C., Nicosia, S., Lad, P. M., and Rodbell, M. (1977).J. Bid. Chem. 252,2790-2792. Londos, C., Lin, M. C., Welton, A. F., Lad, P. M., and Rodbell, M. (1977).J.Biol. Chem. 252, 5180-5182. Londos, C., Cooper, D. M. F., Schlegel, W., and Rodbell, M. (1978). Proc. Natl. Acad. Sci. U.S.A. 75, 5362-5366. Lyon, T. F., and Randall, W. C. (1980). Life Sci. 26, 1121-1129. McCarthy, K. D., and de Vellis, J. (1978).J. Cylic Nucleotide Res. 4, 15-26. McMillen, B. A., Warnack, W., German, D. C., and Shore, P. A. (1980).Eur.J. Pharmacol. 61,239-246. McPherson, G. A., and Summers, R. J. (1982). Biochem. Pharmacol. 31, 583-587. Maggi, A., U’Prichard, D. C., and Enna, S. J. (1980). S c z m e 207, 645-647. Maguire, M. E., and Erdos, J. J. (1980).J. Bid. Chem. 255, 1030-1035. Maguire, M. E., Ross, E. M., and Gilman, A. G. (1977).Adv. Cyclic Nucleotide Res. 8, 1-83. Mallat, M., and Hamon, M. (1982).J. Nmrochem. 38, 151-161. Manukhin, B. N., and Volina, E. V. (1979).Biochem. Phannacol. 28, 2037-2044. Martinez, J. R., and Camden, J. (1982). Arch. Oral Bid. 27, 939-944. Martinez, J . R., Quissell, D. O., and Giles, M. (1976).J. Pharmacol. Exp. Ther. 198,385-394. Martinez, J. R., Bylund, D. B., and Cassity, N. (1982a). Arch. Oral Biol. 27, 443-450. Martinez, J. R., Bylund, D. B., and Camden, J. (198213). Naunyn-Schmiedebmg’s Arch. Pharmacol. 318, 313-318. Menkes, D. B., and Aghajanian, G. K. (1981). Eur. J . Pharmacol. 74, 27-35. Miach, P. J., Dausse, J.-P., and Meyer, P. (1978). Nature (London) 274, 492-494. Miach, P. J., Dausse, J.-P., Cardot, A., and Meyer, P. (1980). Naunyn-Schmiedeberg’s Arch. Pharmacol. 312, 23-26. Michel, T., Hoffman, B. B., Lefkowitz, R. J., and Caron, M. G. (1981).Biochem.Biophys. Res. Commun. 100, 1131-1136. Michell, R. H. (1975). Biochim. BiOphys. Acta 415, 81-147. Minneman, K. P., Hegstrand, L. R., and Molinoff, P. B. (1979).Mol. Pharmacol. 16,34-46. Minneman, K. P., Pittman, R. N., and Molinoff, P. B. (1981). Annu. Rev. Neurosci. 4, 419-461. Minton, A. P. (1981). Mol. Pharmacol. 19, 1-14. Mitrius, J. C., and U’Prichard, D. C. (1983).J. B i d . Chem. (in press). Mooney, J. J., Horne, W. C., Handin, R. I., Schildkraut, J. J., and Alexander, R. W. (1982). Mol. Pharmacol. 21, 600-608. Morris, M. J.. Dausse, J., Devynck, M., and Meyer, P. (1980). In “Biogenic Amines in Development” (H. Parvez and S. Parvez, eds.), pp. 24 1-261. ElseviedNorth-Holland Biomedical Press, Amsterdam. Morris, M. J., Derynck, M. A., Woodcock, E. A., Johnston, C. I., and Meyer, P. (1981). Hypertension 3,516-520. Moss, J., and Vaughan, M. (1977). Proc. Natl. Acad. Sci. U.S.A. 74, 4396-4400. Motulsky, H. J., and Insel, P. A. (1982).Biochem. Pharmacol. 31, 2591-2597. Motulsky, H. J., Shattil, S. J., and Insel, P. A. (1980). Biochem. Biophys. Res. Commun. 97, 1562- 1570. Mukherjee, A. (1981). Biochim. Biophys. Acta 676, 148-154.

428

DAVID B. BYLUND AND DAVID

c. U’PRICHARD

Nakaki, T., Nakadate, T.. Ishii, K., and Kato, R. (1981).J. Phnrmacol. Exp. Ther. 216, 607-6 12. Nakaki, T., Nakadate, T., Yamamoto, S., and Kato, R. (1982).J . Pharrnncol. Exp. Ther. 220,637-641. Neethling, A. C., McCarthy, B. W., and Taljaard, J. J. F. (1981). Biochrm. Pharinucol. 30, 565- 569. Neaman, K. D., Williams, L. T., Bishopric, N. H., and Lefkowitz, R. J. (1978).J. Clin. Invest. 61,395-402. Northup. J. K., Sternweis, P. C., Smigel, M. D., Schleifer, L. S., Ross, E. M., and Gilman, A. G . (1980). Proc. A’atl. Amd. Sci. U.S.A. 77, 6516-6520. O’Dea, R. F., and Zatz, M. (1976).Proc. 9 a t l . Acad. Sci. b’.S..4. 73, 3398-3402. Partington, C. R., Edwards, M. W., and Daly, J. W. (1980).J. Neurochrin. 34, 76-82. Pecquery, R.. and Giudicelli. Y. (1980). FEBS L ~ t f116, . 85-94. Pelayo, F., Dabocovich, M. L., and Langer, S. Z. (1978).Nufure (London) 274, 76-78. Peroutka. S. J., U’Prichard, D. C., Greenberg, D. A., and Snyder, S. H. (1977).Neuropharrnacolog?: 16, 549-556. Peroutka, S. j., Greenberg, D. A., U’Prichard, D. C., and Snyder, S. H. (1978). Mol. Phannacol. 14, 403-412. Peroutka, S. J., Moskowitz. M. A,, Reinhard,J . F., Jr., and Snyder, S. H. (1980).Science 208, 610-61 2. Perry, B. D., and U’Prichard, D. C. (1981).Eur. J. Phunnncol. 76, 461-464. Perry, B. D., and U’Prichard, D. C. (1983). Mol. Pharmacol. (in press). Pimoule, C., Briley, M. S., and Langer, S. Z. (1980).Eus. J . Pharmacol. 63, 85-87. Pointon, S. E., and Banerjee, S. P. (1979). Biochim. Biophjs. Acta 584, 231-241. Propst, E,and Hamprecht, B. (198l).J. Neurochem. 36, 580-588. Propst, F., Moroder, L., Wunsch, E., and Hamprecht, B. (1979).J. Nmrochern. 32, 14951500. Putney, J. W., Jr., Weiss, S. J., Van De Walle, C. M.,and Haddas, R. A. (1980). Nature (Lundun) 284, 345-347. Rehavi, M., Rarnot, O., Yavetz, B., and Sokolovsky, M. (1980a).Bruin Res. 194, 443-453. Rehavi, M., Yavetz, B., Ramot, 0..and Sokolovsky, M. (1980b).Life Sn‘. 26, 615-621. Reisine, T. D., U’Prichard, D. C., Wiech, N. L., Ursillo, R. C., and Yamamura, H. I. (1980). Brain Reg. 188, 587-592. Roberts, J. M.,Insel, P. A., Goldfien, R. D., and Goldfien, A. (1977).Natttrr (London)270, 624-62 5. Roberts, J. M., Goldfien, R. D., Tsuchiya, A. M., Goldfien, A., and Insel, P. A. (1979). Endocriiwlogy 104, 722-728. Roberts, J. M.,Insel, P. A., and Goldfien, A. (1981).dfol. Phnrnzncol. 20, 52-58. Robison, G. A., Langley, P. E., and Burns, T. W. (1972).Biorhm. Phannacol. 21, 589-592. Rodbell, M. (1980).Nafure (London)284, 17-22. Rosenblatt, J. E., Pert, C. B.,Tallman, J. F., Pert, A., and Bunney, W. E., Jr. (1979).Brain Res. 160, 186-191. Rosendorff, C., U’fiichard, D. C., and Hurwitz, M. L. (1981).Bm.cRes. Cardiol. 76, 536539. Ross, E. M., and Gilman, A. G. (1980).Annu. R m . Biochm. 49, 533-564. Rouot, B., and Snyder, S. H. (1979).L f e Sci. 25, 769-774. Rouot, B., U’Prichard, D. C., and Snyder, S. H. (198O).J. Neurochm. 34, 374-384. Ruffolo, R. R., Jr., Fowble, J. W., Miller, D. D., and Patil, P. N. (1976).Proc. Natl. A d . Sci. U.S.A. 73, 2730-2734.

CHARACTERIZATION OF

(YI-

ANI) OL2-ADRENERGIC RECEPTORS

429

Ruffolo, R. R., Jr., Yaden, E. L., and Waddell, J. E. (1980).J. Pharmacol. Exp. Ther. 213, 557-561. Sabol, S. L., and Nirenberg, M. (1979a).J. Biol. Chem. 254, 1913-1920. Sahol, S. L., and Nirenberg, M. (1979b).J. Biol. Chem. 254, 1921-1926. Salama, A. I., Lin, L. L., Repp, L. D., and U’Prichard, D. C. (1982).LifeSci. 30, 1305-131 1. Salzman, E. W., and Neri, L. L. (1969). Nature (London) 224, 609-610. Schmitz, J. M., Graham, R. M., Sagalowsky, A , , and Pettinger, W. A. (1981).J. Ph,armarol. ExP. Thm. 219, 400-406. Schwabe, U., Ohga, Y., and Daly, J. W. (1978).Naunyn-Schmie&bmg’sArch. Pharmacol. 302, 141-151. Seamon, K., and Daly, J. W. (1981).J. Biol. Cliem. 256, 9799-9801. Seamon, K. B., Padgett, W., and Daly, J. W. (1981). Proc. Natl. Acad. Sn’. U.S.A. 78, 33633367. Seeman, P. (1980). Pharmacol. Rev. 32, 229-3 13. Shattil, S. J., McDonough, M., Turnbull, J., and Insel, P. A. (1981). Mol. Pharmacol. 19, 179-183. Sheys, E. M., and Green, R. D. (1972).J. Pharmacol. Exp. Ther. 180, 317-325. Smith, C. B., Garcia-Sevilla, J. A., and Hollingsworth, P. J. (1981). Brain Res. 210,413-418. Smith, S. K., and Limbird, L. E. (1981). Proc. Natl. Acad. Sci. U.S.A. 78, 4026-4030. Spyraki, C., and Fibiger, H. C. (1980). Life Sci. 27, 1863-1867. Starke, K. (1971). Natunuirrmchaften 58, 420. Starke, K. (1977). Rev. Physiol., Biorhem. Pharmacol. 77, 1-124. Starke, K. (1981). Rev. Physiol., Biochem. Pharmacol. 88, 199-236. Starke, K., Endo, T., and Tabue, H. D. (1975). Nature (London) 254, 440-441. Starke, K., Montel, H., Gayk, W., and Merker, R. (1974). Naunyn-Schmiedeberg’s Arch. Pharmacol. 285, 133-150. Steer, M. L., and Wood,A. (1979). J . Biol. Chem. 254, 10791-10797. Steer, M. L., and Wood, A. (1981).J. Biol. Chem. 256, 9990-9993. Story, D. D., Briley, M. S., and Langer, S. 2. (1979). Eur. J . Pharmacol. 57, 423-426. Story, D. F., McCulloch, M. W., Rand, M. J., and Standford-Starr, C. A. (1981). Nature (London) 293,62-65. Strittmatter, W. J., Davis, J., and Lef’kowitz, R. J. (1977).J. Biol. Chem. 252, 5472-5477. Su, Y. F., Harden, T. K., and Perkins, J. P. (1980).J. Biol. Chem. 255, 7410-7419. Summers, R. J. (1980). Br. J . Pharmarol. 71, 57-63. Summers, R. J., Jarrott, B., and Louis, W. J. (1980). Eur. J . Pharmacol. 66, 233-241. Svensson, T. H., Bunney, B. S., and Aghajanian, G. K. (1975). Brain Res. 92, 291-306. Taft, W. C., Jr., Abdel-Latif, A. A., and Akhtar, R. A. (1980). Biochem. Phurmcof. 29, 2713-2720. Tanaka, T., and Starke, K. (1979). Naunyn-Sch.miedeberg’s Arch. Pharmacol. 309, 207-2 15. Tanaka, T., and Starke, K. (1980). Eur. J . Pharmacol. 63, 191-194. Tharp, M. D., Hoffman, B. B., and Lefkowitz, R. J. (1981).J. Clin. Endocriml. Metab. 52, 709-7 14. Timmermans, P. B. M. W. M., and van Zwieten, P. A. (1980). Eur. J . Pharmacol. 63, 199202. Timmermans, P. B. M. W. M., Kwa, H. Y., and van Zwieten, P. A. (1979). NaunynSchmiedeberg’s Arch. Pharmacol. 1110, 189- 193. Timmermans, P. B. M. W. M., Schoop, A. M. C., and van Zwieten, P. A. (1982). Biochem. Pharmacol. 31, 899-905. Taokimasa, T., Morita, K., and North, A. (1981). Nature (London) 294, 162-163.

430

DAVID B. BYLUND AND DAVID c . U’PRICHARD

Torda, T., Yamaguchi, l., Hirata, F., Kopin, 1. J., and Axelrod, J. (1981).J. Phrcrnioml. Exp. T h r . 216, 334-338. Toscano. W. A , , Westcott, K. R.. LaPort, D. C., and Storm, D. R. ( 1 979). Proc. Nntl. ‘4cad. Sci. C‘.S..4. 76, 5582-5586. ’Ikai. B. S., and Lefkowitz, R. J. (1978). LVv[.Phartttnrol. 14, 540-548. Tsai, B. S., and Lefkowitz, R. J. (1979). .\lo/. Phnrmrrrol. 16, 61-68. U’Prichard, D. C. (1980). I n “Ergot Compounds and Brain Function” (M. Goldstein, D. B. Calne, A . Lieberman, and M. D. Thorner. eds.), pp. 103-1 15. Raven Press, New York. U’Prichard, D. C. (1981). AVurrwfrattsm.Rerep/. 1, 131-179. U’Prichard, D. C., and Kvetnansky, R. (1980). In “Catecholamines and Stress: Recent Advances” (E. Usdin, R. Kvetnanskv, and 1. J. Kopin, eds.), pp. 299-308. Elsevier/ North-Holland, Amsterdam. U’Prichard. D. C., and Snyder, S. H. (1977a).J. Biol. C / w n . 252, 6450-6463. UPrichard, D. C.. and Snyder, S. H. (397713).h h t u r u (London) 270, 261-263. U’Prichard, D. C . , and Snyder, S. H. (1978a). E u r . J . Phnrnincol. 51, 145-155. U’Prichard, D. C., and Snyder, S. H. (1978b).J. Biol. Chetn. 253, 3444-3452. P 24, 79-88. C’Prichard, D. C., and Snyder, S. H. (1979). L ~ Sri. U’Prichard, D. C., and Snyder, S. H. (1980).J. jV’nrrorhrm. 34, 385-394. Phnnnacol. 13,454U’Prichard, D. C.. Greenberg, D. A , , and Snyder, S. H. (1977a). IV~(JI. 473. U’Prichard, D. C.. Greenberg, D. A., Sheehan, P. P., and Snyder, S. H. (1977b). Rrrcin Res. 138, 151-158. U’Prichard, D. C., Greenberg, D. A., Sheehan, P. P., and Snyder, S. H. (1978a). Science 199, 197-198. U’Prichard, D. C., Charness, M. E., Robertson, D., and Snyder, S. I€. (1978b). Eur. J . Phnrttmrd. 50, 87-89. U‘Prichard, D. C., Bylund, D. B., and Snyder, S. H. (1978c).J. Bid. Chrm. 253,5090-5102. U’Prichard, D. C., Bechtel, W. D., Rouot. B. hi., and Snyder, S. H. (1979a). i\fol. Phrcrmurol. 16,47-60. U’Prichard. D. C., Greenberg, D. A , , and Snyder, S. H. (1979b). Itr “Nervous System and Hypertension” (P. Meyer and H. Schmitt, eds.), pp. 38-48. Wiley, New York. U‘Prichard. D. C., Mitrius, J. C., Kahn, D. J., and Daiguji, M. (1980a). In “Psychopharmacologv and Biochemistry of Neurotransmitter Receptors” (H. I. Yamamura, R. W. Olsen, and E. Usdin, eds.), pp. 247-259. Elsevier/North-Holland, Amsterdam. U’Prichard, D. C., Reisine, T. D., Mason, S. T., Fibiger, H. C., and Yamamura, H. I. (1980b). B ~ n i nRex. 187, 143-154. U‘Prichard, D. C., Wang, C. H., and Freed, C. R. (1981). Ntwrosci. Sor. Ab.\tr. 7, 427. U’Prichard, D. C., Mitrius, J. C., Kahn, D. J.. and Perry, B. D. (1983a). I n “Molecular Pharmacology of Neutoransmitter Receptor Systems” (C. T. Segawa, H. I. Yamamura, and K. Kuriyama, eds.) pp. 53-72. Raven Press, New York. U’Prichard, D. C., Daiguji, M.,Tong, C., Mitrius, J. C., and Meltzer, H. Y. (1983b). In “Biological Markers in Psychiatry and Neurology” (I. Hanin and E. Usdin, eds.). pp. 205-217. Pergamon, Oxford. van Calker, D., Muller. M., and Hamprecht, B. (1980). Pror. A’ntl. Arad. Sci. C1.S.A. 77, 6907-691 1. Vetulani, J., Nielsen, M.. Pilc, A., and Golembiowska-Nikitin, K. (1979). Eur. J. Pticirmacol. 58, 95-96. Villalobos-Molina, R., Uc, M., Hong, E., and Garcia-Sainz, J . (1982). J. Pharmarol. Exp. Ther. 222, 258-261. Wang, C. H., and U’Prichard, D. C. (1980). Neurosci. Soc. Abstr. 6, 1.

CHARACTERIZATION OF (Y1- AND (Y2-ADRENERGIC RECEPTORS

431

Watanabe, A. M., Hathaway, D. R., Besch, H. R., Farmer, B. B., and Harris, R. A. (1977). Circ. Res. 40, 596-602. Weiland, G. A., Minneman, K. P., and Molinoff, P. B. (1979). Nature (London) 281, 114117. Weinreich, P., Deck, J., and Seeman, P. (1980). Biochm. Pharmacol. 29, 1869- 1870. Wikberg, J. E. S. (1979). Acta Physiol. Scand., Suppl. 468. Wilkening, D., Sabol, S. L., and Nirenberg, M. (1980). Brain Res. 189, 459-466. Williams, L. T., and Lefkowitz, R. J. (1976). Science 192, 791-793. Williams, L. T., and Lefkowitz, R. J. (1977a).J. Biol. Chem. 252, 7207-7213. Williams, L. T., and Lefkowitz, R. J. (1977b).J. Clin. Invest. 60, 815-818. Williams, R. S., and Lefkowitz, R. J. (1978). Czrc. Res. 43, 721-727. Wirz-Justice, A., Kafka, M. S., Naber, D., and Wehr, T. A. (1980). L f e Sci. 27, 341-347. Wood, C. L., Arnett, C. D., Clarke, W. R., Tsai, B. S., and Lefkowitz, R. J. (1979a). Biochm. P h a m c o l . 28, 1277-1282. Wood, C. L., Caron, M. G., and Lefkowitz, R. J. (1979b). Biochm. Biophys. Res. Commun. 88, 1-8. Woodcock, E. A., and Johnston, C. I. (1980). Hypertension 2, 156-161. Woodcock, E. A., Johnston, C. I., and Olsson, C. A. (1980). J. Cyclic Nucleotide Res. 6 , 261-269. Wright, E. E., Ackerman, G. E., and Simpson, E. R. (1981). Fed. Proc., Fed. Am. SOC.Exp. Biol. 40, 1822. Yamada, S., Yamamura, H. I., and Roeske, W. R. (1980a). Life Sci. 27, 2405-2416. Yamada, S., Yamamura, H. I., and Roeske, W. R. (1980b).J. Pharmacol. Exp. Ther. 215, 176- 185. Yamada, S., Yamamura, H. I., and Roeske, W. R. (1980~).Eur. J. Pharmacol. 63, 239-241. Yamada, S., Yamamura, H. I., and Roeske, W. R. (1980d). E u r . J . Pharmacol. 68,217-221. Yamashita, K., Yamashita, S., and Aiyoshi, Y. (1980). Life Sn'. 27, 1127-1130. Young, W. S., and Kuhar, M. J. (1979). Eur. J . Pharmacol. 59, 317-319. Young, W. S., and Kuhar, M. J. (1980). Proc. Natl. Acad. Sci. U.S.A. 77, 1696-1700.

This Page Intentionally Left Blank

ONTOGENESIS OF THE AXOLEMMA AND AXOGLIAL 0NS H IPS IN M YE LINATE D F IBERS: RE LAT1 E LECTROPHYSIOLOGICAL AND FREEZE-FRACTURE CORRELATES OF MEMBRANE PLASTICITY By Stephen G. Waxman and Joel A. Black* Department of Neurology Stanford University School of Medicine and Veterans Administration Medical Center Palo AI~O,Ca.lifornia

and Robert E. Foster Neurotoxicology and Experimental Therapeutics Branch

U.S. A m y Medical Research Institute Aberdeen Proving Ground, Maryland

I. Introduction ......................................................... 11. Specificity in Myelination

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

111. Development of the A. Introduction ... B. Electrophysiology IV. Freeze-Fracture Structure of Myelinated Axons . . . . . . . . . . . . . . . . . . . . . . . . . A. Introduction to Freeze-Fracture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Freeze-Fracture of Myelinated Axons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Freeze-Fracture of Adult Rat Retinal V.. Freeze-Fracture Studies on Myelin Devel A. Introduction to Myelin Development . . B. Premyelinated Axolemma . . . . . . . . . . . .. . . . . . .. . . . .. .. . . . . . . . . . .. . . . C. Axolemmal Changes Associated with Glial Ensheathment . . . . . . . . . . D. Myelinated Axolemma . . . . . . . . . . . . . . . E. Aberrant Axoglial Association . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. Summary of Developmental Changes in Axon Membrane.. . . . . . . . . . . . VI. Differentiation of the Axon Membrane in the Absence of VII. Concluding Comments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . ........................... .......... .

I

.

.

.

.

.

.

.

.

434 437 440 440 441 449 449 45 1 452 461 461 463 465 470 473 474 475 479 481

* Present address: Department of Biological Sciences, Northern Illinois University, DeKalb, Illinois 601 15. 433 INTERNATIONAL REVIEW OF NEUROBIOLOGY, VOL. 24

Copyright 6 1983 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-366824-7

434

STEPHEN G. WAXMAN

et al.

1. Introduction

Although it is well known that myelinated nerve fibers are differentiated, at the light microscopic level, into regions covered by myelin (internodes) and regions devoid of myelin (the nodes of Ranvier), it has only been in the past few years that details of axon membrane differentiation itself have been studied. It is now becoming apparent that the axon membrane (axolemma) exhibits an elegant differentiation in terms of its macromolecular architecture, and that nodal and internodal regions of this membrane can be shown to be different by morphological, electrophysiological, and pharmacological techniques. This differentiation of the mature axolemma into nodal and internodal domains with distinct properties has been recently reviewed in a number of articles (Waxman and Foster, 1980; Ritchie and Chiu, 1981; Rosenbluth, 1981a). A previous article (Waxman and Foster, 1980) reviewed cytochemical aspects of the plasticity of developing axon membranes. T h e present article reviews studies from our laboratory dealing with electrophysiological and freeze-fracture aspects of axon membrane reorganization during ontogenesis of the trunk of the mammalian myelinated fiber. It is the purpose of this article to discuss the development of the axon membrane of the myelinated fiber, and to demonstrate that the axolemma exhibits a high degree of plasticity during development, with significant changes in axon membrane structure being related to association with glial cells. Action potentials in myelinated fibers are generally considered as being conducted in a saltatory manner, with the impulse traveling discontinuously along the axon. T h e distribution of ionic channels in the axon membrane of mammalian myelinated fibers has been shown to be nonuniform, with markedly different densities of sodium and potassium channels in the node and internode, respectively. Voltage-clamp (Conti et ul., 1976) and PHIsaxitoxin binding studies (Ritchie and Rogart, 1977) suggest a high density (5,000- 12,000/pmZ)of sodium channels in the mammalian nodal axolemma. In contrast, the density of sodium channels in the internodal axolemma (beneath the myelin sheath) is quite low (<25/pm2),possibly too low to support impulse electrogenesis (Ritchie and Rogart, 1977; Chiu and Ritchie, 1981). Moreover, cytochemical studies (Waxman and Quick, 1977, 1978; Waxman et nl., 1978) also differentiate nodal from internodal membrane and are consistent with a high density of sodium channels in the nodal membrane. Voltage-sensitive potassium conductance appears to be low at mammalian nodes of Ranvier (Chiu P t al., 1979; Brismar, 1979, 1980; Kocsis and Waxman, 1980, 1981; Sherratt et ul., 1980; Bostock et al., 1981).There is,

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

435

however, evidence for the presence of potassium channels in the internodal axolemma. The primary support for this concept came from the elegant voltage-clamp studies of Chiu and Ritchie (1980), who showed the appearance of voltage-sensitive potassium conductance in mammalian myelinated fibers following acute disruption of myelin. Voltageclamp studies on the internodal membrane confirmed the presence of potassium conductance in the internodal axolemma (Chiu and Ritchie, 1981, 1982). Potential clamp studies on diphtheria toxin-demyelinated fibers show that, even in the case of paranodal demyelination which is barely detectable at the light microscopic level, potassium conductance appears (Brismar, 1981). Studies on alloxan-diabetic rats (Brismar, 1979) and on fibers demyelinated with diphtheria toxin (Sherratt et al., 1980) are also consistent with the concept that, in demyelinated mammalian fibers, potassium conductance is present in the region which was formerly internodal membrane. In addition, studies on regenerating mammalian fibers suggest that potassium conductance is present in the regenerating axolemma (Ritchie, 1982a,b; Kocsis et al., 1982a,b). Thus, the picture that emerges for normal myelinated axolemma is one of a complementary distribution of ionic channels, with sodium channels present in high density in the nodal axolemma and potassium channels present primarily in the internodal axolemma (Ritchie and Chiu, 1981; Waxman, 1982). This distribution of sodium channels is consistent with the saltatory nature of impulse conduction in myelinated fibers (Huxley and Stampfli, 1949). In contrast to myelinated axons, most nonmyelinated fibers display a clearly different axolemmal architecture and mode of conduction. Impulse conduction in mammalian nonmyelinated fibers has classically been considered to occur in a continuous manner (as noted in Section VI of this article; however, some specialized nonmyelinated fibers exhibit a heterogeneous membrane structure, with focal axolemmal specializations, suggesting the possibility of nonuniform conduction in these fibers). T h e depolarization phase of impulse electrogenesis in mammalian nonmyelinated axons is dependent upon voltage-sensitive sodium channels, the density of which has been estimated as 110/pm2in rabbit vagus nerve (Ritchie et al., 1.976).Repolarization depends in part on the presence of potassium channels; the contribution of potassium channels to impulse electrogenesis has been demonstrated in mammalian peripheral (Sherratt et al., 1980; Bostocket al., 1981) and central (Kocsiset al., 1981; Preston et al., 1983) nonmyelinated fibers, although the density of these channels has not been established. T h e distribution of sodium channels in most nonmyelinated axolemma is apparently homogeneous. Cytochemical studies of nonmyelinated C-fibers (Waxman and Quick, 1977) do not indicate regions

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

437

of high sodium channel density, nor do freeze-fracture images of most nonmyelinated axons reveal aggregations of particles similar to those seen in myelinated fibers (Black et al., 1981). By way of a brief introduction to freeze-fracture electron microscopy and as an example of the spatial homogeneity of typical mammalian nonmyelinated axolemma, Fig. l a is representative of the relatively uniform structure of nonmyelinated axon membrane. Nonsynaptic regions of cerebellar parallel fibers are shown. These central nonmyelinated fibers are known to conduct at a relatively slow conduction velocity, with potassium conductance, as well as sodium conductance, playing a role in action potential electrogenesis (Kocsis et al., 1981). Intramembranous particles are observed in greater density on the P-fracture face (Fig. lb; 794/pm2) than on the E-fracture face (Fig. lc; 207/pm2); moreover, the particles show no apparent ordered pattern of distribution (Black et al., 1981). This density and distribution of particles within most regions of the nonmyelinated axolemma will be contrasted later in this article with that observed in nodal and internodal regions of myelinated fibers and with that found in some specialized nonmyelinated regions. While the morphological correlates of sodium and potassium channels have not yet been identified with certainty, the axon membrane of the myelinated fibers exhibits a morphological differentiation which parallels that predicted from the electrophysiological and pharmacological studies. This ultrastructural specialization is clearly apparent in cytochemical (Quick and Waxman, 1977; Waxman, 1977) and freezefracture (Rosenbluth, 1976; Kristol et ol., 1978) studies. This article will discuss the development of morphological differentiation of the myelinated axon membrane as seen by freeze-fracture, a method that permits study of quantitative as well as qualitative changes in membrane structure. II. SpecifKity in Myelination

It was initially suggested by Duncan (1934) that during development myelination was directly dependent on axonal diameter, occurring when FIG. 1. Freeze-fracture replica of nonmyelinated axons from the cerebellar cortex. (a) Low-power electron micrograph of E- and P-fracture faces (eA and PA, respectively) of several cerebellar parallel fibers. The P- and E-faces display asymmetry of particle distribu. P-faces and (c) E-faces of the parallel fiber axolemma tion. Scale bar: 1 pm. ~ 4 3 , 3 0 0 (b) are illustrated at increased magnification. The P-face has a moderate density of particles; few particles are evident on the E-face. Scale bar: 0.25 p m . X 132,500. (Modified from Black et a/., 198 1.)

438

STEPHEN G. WAXMAN

et al.

a “critical diameter” was achieved by the axon. However, this view has not been substantiated by more recent studies. It is now apparent that while fibers smaller than 1 p m are rarely myelinated in the peripheral nervous system (PNS), axons with diameters as small as 0.2 p m are often myelinated in the central nervous system (CNS) (eg., Waxman and Bennett, 1972; Waxman and Swadlow, 1977). A number of studies (Fleischauer and Wartenberg, 1967; Matthews and Duncan, 1971; Fraher, 1972; Waxman and Swadlow, 1977) indicate that there is an overlap in the diameters of myelinated and unmyelinated fibers at all stages of development, in both central and peripheral nervous system. This raises the important question as to the nature of the signal which initiates myelination. Simpson and Young (1945) showed, in cross-union experiments carried out in peripheral nerve, that the axon signals whether or not myelination will proceed. This result was extended by Weinberg and Spencer (1976) and Aguayo ef al. (1976), who demonstrated with autoradiography that Schwann cells in cross-anastomosed myelinated and nonmyelinated nerves are multipotential. These experiments indicate that it is the axon that determines whether myelination will occur. It has also been shown that axons are mitogenic for Schwann cells (Wood and Bunge, 1975) and that Schwann cell proliferation requires contact with neurons and may be mediated by glycoproteins located on the axonal surface (Salzer PI nl., 1980). Since glial proliferation and axonal ensheathment occur before myelination, it must be stressed that the signals for glial proliferation, ensheathment of axons, and myelination may not be the same. Myelination depends 011 processes which are much more complex than a binary decision as to whether to produce myelin or not. There is a high degree of specificity in the interaction between axon and myelinforming cell, with myelin sheath characteristics matched to axonal size and type. For example, myelin thickness is not invariant but, on the contrary, is correlated with fiber diameter. In general, the number of myelin lamellae increases monotonically with fiber diameter (Friede and Samorajski, 1968; Bishop et d.,1971; Waxman and Swadlow, 1977), although there are different ratios between myelin thickness and diameter for different fiber types (Williams and Wendell-Smith, 1971). As pointed out by Rushton (1951), the ratio between myelin thickness and fiber diameter is, for many fibers, close to the value which would maximize conduction velocity. Another example of specificity in myelination is provided by examination of internode distance. In peripheral nerve and some central myelinated tracts, there is an approximately linear relationship between

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

439

internode distance and fiber diameter (Thomas and Young, 1949; Hess and Young, 1952; Gutrecht and Dyck, 1970; McDonald and Ohlrich, 197 1). As predicted by Huxley and Stampfli (1949), and demonstrated by the computations of Brill et al. (1977), internodal distances in peripheral nerve fall close to the values which maximize conduction velocity for any given diameter. However, in some areas in the central nervous system, the ratio of internode distance to diameter is considerably smaller than in normal peripheral nerve (Waxman, 1970, 1972; Waxman and Melker, 1971). In some cases, the reduced internode distance : diameter ratios are consistent with electrophysiologically demonstrated delay line function of the axons (Waxman, 1972; Meszler et al., 1974). In other systems, such as reduced internode distances in preterminal fibers (Zenker, 1964; Waxman, 1970,1972) or the reduced internode distances close to the bifurcation leading to the dorsal root ganglion cells (It0 and Takahashi, 1960), the reduction in internode distance occurs proximal to a region of impedance mismatch. In each of these situations, it would be expected that reduction in the internode distance should be reflected by an increase in the amount of current available for impulse propagation (Revenko et al., 1973; Waxman and Brill, 1978). Thus, with respect to both myelin thickness and internode distance, the geometry of the myelin sheath is not fixed, but appears to be matched to functional requirements of the axon. This suggests a highly specific interaction between the axon and the myelin-forming cell, so as to achieve the appropriate pattern of myelination. There are numerous other examples of the highly ordered and specific nature of myelination along the trajectory of individual axons. The axon initial segment (Palay et al., 1968; Peters et al., 1968; Waxman and Quick, 1978) is a specialized region of the axon, which functions as the trigger zone for impulse initiation in motoneurons (Coombs et al., 1957; Fuortes et al., 1957). This axon region exhibits a diameter similar to that of adjacent myelinated regions but is devoid of myelin. Several specialized axons, such as the electrocyte fibibers in Sternarchus albifrons (Bennett, 1970; Waxman et al., 1972) and some specialized preterminal axons in the central nervous system (Waxman, 1972), have nonmyelinated regions that are considerably longer than normal nodes of Ranvier and are intercalated at specific sites along otherwise normally myelinated axons. Thus, it appears that there is a high degree of specificity in the relationship between axon and myelin-forming cell, with the pattern of myelination varying in different types of axons, and even at different regions along single axons (Waxman, 1972; Waxman and Swadlow, 1977; Spencer and Weinberg, 1978). Moreover, the axon appears to

440

STEPHEN G. WAXMAN

et al.

serve as the frame of reference with respect to the location and geometrical properties of the myelin segments. Observations on regenerating Sternarchus electrocyte axons (Waxman and Anderson, 1980) were interpreted as suggesting that axonal differentiation occurs as the primary event, with the Schwann cell probing the axon surface and forming myelin in areas specified by the axon. Moreover, it appears that some aspects on axon membrane differentiation in Sternarchus are stable, and are maintained following demyelination (Quick and Waxman, 1978). T h e mechanisms determining the formation of myelin in specific axon regions are not well understood. It is not unreasonable to expect that they may depend, at least in part, on contact-mediated mechanisms involving the axon surface. It should be recalled in this context, however, that the pattern of myelination varies during growth with, e.g., internode distances increasing with age (Vizoso and Young, 1948; Gutrecht and Dyck, 1970). Also, in remyelinated fibers, nodes of Ranvier are more closely spaced than in prepathological fibers (Sanders and Whitteridge, 1946; Gledhill et al., 1973); thus, nodes in some cases are formed at sites which were previously internodal. Nevertheless, the nodes formed along remyelinated fibers develop normal cytochemical (Weiner et al., 1980) and saxitoxin-binding characteristics (Ritchie et al., 198l), and there is evidence that remyelination restores secure conduction (Smith et nl., 198l),which suggests that the newly formed nodes recapitulate (in terms of sodium channel distribution) a relatively normal structure. Evidence that the axon membrane exhibits a high degree of regional differentiation with respect to intrinsic membrane proteins, that this differentiation is an inherent property of the axon membrane, and that the axon membrane exhibits plasticity with respect to its regional differentiation is discussed below.

111. Development of the Optic Nerve

A. INTRODUCTION An excellent model for correlative electrophysiological and morphological studies on the development of myelinated axons is provided by the rat optic nerve (Black et al., 1982a; Foster et al., 1982). This tract presents a population of axons in which the end points of development are nearly uniform with respect to myelination: At birth and until approximately 6 days postnatal, the axons of the optic nerve remain completelv nonmyelinated (Skoff et al., 1976a,b). By adulthood, on the other

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

441

hand, virtually 100% of the axons are myelinated (Forrester and Peters, 1967). Moreover, the available evidence indicates that, at birth, rat optic nerves probably have reached their synaptic targets (Lund and Bunt, 1976). Thus, by studying the isolated optic nerve, it is possible to examine the development of the axon trunk prior to, during, and following myelination. Figure 2a-d illustrates representative fields of axons, cut in transverse section, from 2-, 8-, and 16-day-old and adult rats, respectively. As seen in these figures, development of the optic nerve is characterized by an increase in axonal diameter and an increase in the percentage of myelinated fibers. Neonatal optic nerve axons are approximately 0.14 p m in diameter. T h e axons are separated by a relatively wide and electron-lucent extracellular space which contains few glial processes. Axon diameter increases with age so that, in the adult, the mean axon diameter is approximately 0.77 pm. In the mature optic nerve, essentially all of the axons are myelinated, and the area between axons is filled with glial elements, leaving only a relatively small extracellular space. Figure 3 illustrates graphically the time course of development of optic nerve axon diameter and the time course of myelination. Myelination (defined by the presence of at least one layer of compact myelin) begins at approximately the sixth postnatal day (Fig. 3b, open triangles) and proceeds relatively rapidly, so that approximately 85% of optic nerve fibers are myelinated by 28 days of age. Axonal diameter increases relatively slowly prior to the onset of myelination (filled circles in Fig. 3a). In particular, the mean diameter of optic nerve axons remains at approximately 0.2 p m until myelination begins. With the onset of myelination, there is an increase in diameter (compare filled squares which correspond to myelinated axons, to filled circles, which correspond to nonmyelinated axons at 11, 16, and 28 days in Fig. 3a). It should be noted in this context that there is evidence (for review, see Aguayo et al., 1979) for a local effect of myelin-forming cells on the diameter of axons.

B. ELECTROPHYSIOLOGY OF DEVELOPING OPTICNERVE 1. Conduction Velocity T h e development of conduction velocity of optic nerve axons is shown as a function of postnatal age in Fig. 4 (Foster et al., 1982). Points corresponding to longest latency peaks (slowest conducting fibers) from a littermate series are connected (filled circles). Squares and triangles represent faster components of multiple-peak compound action potentials, and Xs represent data from nonlittermate animals. There is a relatively

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

443

steady, monotonic increase in conduction velocity with age. At birth, the conduction velocity of the nonmyelinated optic nerve fibers is approximately 0.2 m/sec, and the fibers display a single-component compound action potential. During the period in which myelination commences (6- 16 day points), conduction velocity does not increase markedly. However, the single-component compound action potential evolves into a compound action potential with multiple peaks. When myelination is complete (adult), conduction velocity varies between 3 and 30 m/sec, and the compound action potential has the three-component waveform characteristic of the mature optic nerve (Foster et al., 1982). It is notable that, as illustrated in Fig. 4,conduction velocity increases during development, but is not strictly related to fiber diameter. Conduction velocity rises monotonically from birth but at a rate which can not be accounted for by the increase in fiber diameter between 0 and 8 days, prior to myelination or the development of oligodendroglia. I n particular, during this period, mean axonal diameter increases from 0.14 to 0.2 pm. This increase in diameter would account, if all other conditions were assumed to be equal, for an increase of approximately 20% in conduction velocity (Waxman and Bennett, 1972), rather than the 100% increase that was observed. The compound action potential matures from a simple triphasic response with a single negative peak at birth to a polyphasic potential in the adult, and myelination appears to be correlated with the development of multiple peaks in the compound action potential rather than with a discrete stepwise increase in conduction velocity (Foster et al., 1982). These data are similar to those of Cullheim and Ulfhake (1979), who have reported that there are changes in the relationship between conduction velocity and diameter with increasing age in peripheral myelinated fibers of the cat. The data are compatible with the hypothesis that changes other than an increase in diameter (e.g., changes in ionic channel kinetics or density, resting potential, or in the characteristics of the extracellular millieu or glia in the optic nerve) may have played a role in the change in conduction velocity. FIG.2. Electron micrograph of optic nerves from 2-, 8-, and 16-day-old and adult rats. (a) Portion of optic nerve from 2-day-old Long Evans rat. All of the axons (A) in 2-day-old nerve are nonmyelinated and are of a fairly uniform diameter (-0.2 pm). Microtubules are prominent within the axoplasm. (b) Optic nerve from 8-day-old Long Evans rat. Oligodendrocytic processes (C) encompass certain axons (Ae),while other axons (A) remain without glial association. (c) Section of optic nerve from 16-day-old Long Evans rat. Some axons have acquired multiple wrappings of glial processes. Oligodendrocytic processes (G) are seen amidst fibers (A) that have not acquired myelin. Scale bar for (a-c): 0.5 pm. X43,OOO. (d) Optic nerve from adult Long Evans rat. Compact myelin (M) has formed on virtually all of the axons (A). Scale bar: 0.5 pm. ~ 2 6 , 5 0 0 (Modified . from Black et al., 1982a.)

444

STEPHEN G . WAXMAN

et al.

0

1.0

* 0.2

0

Postnatal Age (days) FIG. 3. Myelination and diameter change with development of the optic nerve. (a) Change in mean diameter (ordinate: 0 and W ) plotted versus age (abscissa: days). Nonmyelinated fibers (0)have smaller diameters than myelinated axons (m). Bars indicate standard errors. (b) Percentage of myelination (ordinate: A) plotted versus age (abscissa: days). Myelination begins at 6-8 days of age, and by 4 weeks -85% of the fibers are myelinated. (From Black t t ul., 1982a.)

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

445

30I

.

I

0 I

3-

,

0

I

J'

03-

0.1

-

AGE (days) FIG.4. Graph showing development of conduction velocity in the optic nerve. Conduction velocity (mfsec) is plotted as a function of age (days). The data points at each age are from a single optic nerve with four exceptions: data points from nonlittermate optic nerves (x)are included at 0 days (neonatal), 2 days, 28 days, and adult. The filled circles ( 0 )plot the longest latency response from the optic nerves of the littermates. The open squares (a), open triangles (A), and filled squares (W) plot the conduction velocity of the faster peaks from the compound action potential of the littermate series. With increasing age, the compound action potential changes from a single negativity to multiple negativities, and finally to the adult 3-negativity form. Note the monotonic increase in conduction velocity. (From Foster et al., 1982.)

As noted below, development of the recovery cycle in these fibers also suggests that processes other than the production of myelin around the fibers may play a role in the development of adult conduction properties.

2 . Development

of Recovery Cycle

The recovery cycle of many axons is characterized by the absolute refractory period (during which the fiber is inexcitable), which is followed by the relative refractory period (increased threshold with decreased conduction velocity), and, in some axons, a supernormal period (decreased threshold and increased conduction velocity). The recovery

446

STEPHEN G. WAXMAN

et al.

cycle has been discussed in detail by Waxman and Swadlow (1977). It is interesting that the duration of the refractory period (Paintal, 1967; Swadlow and Waxman, 1976) and of the supernormal period (Swadlow and Waxman, 1976) are inversely related to conduction velocity, and therefore presumably to fiber diameter. Figure 5 shows the recovery curve for rat optic nerve at three ages (neonatal, filled circles; 6 days, open squares; adult longest latency component, filled triangles). At all stages of postnatal development, the triphasic recovery cycle was observed. However, the time course of the recovery cycle changes significantly with development of the optic nerve (Foster P t ol., 1982). Notably,

\\

n

8

Y

?5Z

9

U

ISI (msec) ID +

7.0

O

L

1

FIG.5. Changes in the excitability cycle of the optic nerve with development. Three and adult n3 (longest latency, A). The ages are illustrated: neonatal (0);6 days (0); percentage latency change [latency to a single test stimuludlatency to the same (test) stimulus preceded by a conditioning shock] is plotted as a function of the interstimulus interval ( 1 9 ) between the conditioning and test stimuli. A latency shift in the + direction (points above the abscissa) indicates refractoriness. A latency shift in the - direction (points below the abscissa) indicates the response is supernormal with respect to conduction velocity. Oscilloscope traces at upper right illustrate the supernormal conduction of the neonatal optic nerve using a 75-msec IS]. Scale bar: 1 msec. (From Foster u/ nl., 1982.)

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

447

FIG. 6. Ionic mechanisms underlying conduction in the neonatal optic nerve. Compound action potentials are shown. (a) Response of the nerve to 1 p M tetrodotoxin (TTX). TTX abolishes the evoked potential suggesting that sodium is the primary current carrying ion. Cont, Control response; TTX, response with 1 pM TTX in bathing solution. (b) Test for calcium conductances in a neonatal optic nerve. In the normal bathing solution the nerve responds normally (Cont). In a barium-substituted, sodium-free Tris solution the response of the nerve was abolished (Bat+).After 20 min in the normal bathing solution the nerve recovered (Recovery). These data suggest that a voltage-sensitive calcium conductance is not present in the developing axon trunks at this stage of development of the optic nerve. Calibration pulses = 1 mV/1 msec. (From Foster et al., 1982.)

the refractory period decreased substantially by 6 days of age, that is, prior to any myelination. Since there was only a small increase in diameter during this time, changes in axonal size alone probably do not account for this change in the recovery cycle. Thus, analysis of the recovery cycle suggests, as does analysis of the development of conduction velocity, that conduction properties of optic nerve axons change prior to myelination and do not depend exclusively upon fiber diameter.

3. Ionic Basis of Action Potential Propagation Impulse conduction in the optic nerve appears to depend on sodium conductance at all postnatal ages. As illustrated in Fig. 6a, tetrodotoxin, a specific inhibitor of voltage-dependent sodium conductance, abolishes the compound action potential in optic nerve at all postnatal ages. Figure 6b shows the results of another experiment, which tested whether active calcium currents might contribute to the action potential of axons in the neonatal optic nerve. When the bath sodium was replaced with the impermeant cation Tris, conduction was blocked. Subsequently, when most of the Tris was replaced isoosmotically with barium, which is known to permeate calcium channels and suppress active potassium conductances (Werman and Grundfest, 1961), no signs of a conducted, divalent cation-dependent action potential could be detected. These results suggest that the axon trunks s f developing, premyelinated fibers depend on sodium conductance for impulse propagation (Foster et al.,

448

STEPHEN G. WAXMAN

et al.

1982). Moreover, there was no evidence for calcium-dependent impulse propagation in the developing axon trunks; this mechanism of conduction has recently been demonstrated in the distal parts (presumably the growing tips) of developing axons (Strichartz rt al., 1980) and regenerating axons (Meiri et al., 1981). Notably, the contribution of potassium conductance to impulse electrogenesis appears to attenuate with myelination of the more rapidly conducting fibers in the optic nerve. As shown in Fig. 7a, in totally nonmyelinated axons (2 days postnatal; conduction velocity = 0.2 m/sec) the amplitude of the compound action potential was increased and the recovery phase prolonged by the application of 0.5 mM 4aminopyridine (4-AP). Thus, sodium and potassium conductances both contribute to action potential electrogenesis prior to myelination. At the onset of myelination (e.g., 8 days), the compound action potential consisted of t w o components (conduction velocity = 0.4 and 0.6 m/sec). 1

a

b

C

FIG. 7. The response of the optic nerve to electrical stimulation examined at three different ages using 0.5 m.W 4-aminopyridine (4-AP) as a pharmacological probe for potassium conductance. The traces illustrated were matched as closely as possible for conduction distance. The response of the nerve in the normal artificial CSF bath is illustrated by the series labeled CONT at 2 days (a), 8 days (b), and 28 days (c). Note the transformation of the potential from single to multiple negativities with increasing age. The effect of 4-AP on the axons of the optic nerve is illustrated b7- the series of traces labeled 4-AP. 4-AP has a complex effect on the potentials recorded from optic nerves of all ages, suggesting that the axonal membrane develops potassium conductance mechanisms early in life. However, the response to 4-AP is attenuated in rapidly conducting myelinated fibers in the adult. Calibration bar; 2 msec. (From Foster et nl.. 1980a.)

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

449

While both components of the compound action potential were blocked by tetrodotoxin, the slow component was considerably more sensitive to 4-AP (Fig. 7b), suggesting that, as myelination began, voltage-dependent potassium conductance became less important with respect to action potential electrogenesis. Figure 7c shows the compound action potential from 28-day postnatal optic nerve, in which 85% of the fibers have myelinated. At this time, the action potential exhibits two fast (and definitely associated with myelinated fibers) components (3.9 and 7.3 m/sec) and a slow component (1.8 d s e c ) . As in younger optic nerve, all components of the compound action potential were abolished by tetrodotoxin. However, only the slower components were effected by 4-AP. Myelination in the rat optic nerve thus appears to be accompanied by a loss of sensitivity to 4-AP (with respect to action potential waveform) of the more rapidly conducting myelinated fibers. These findings suggest that during myelination the axon reorganizes from a state where both sodium and potassium conductances are present and contribute to action potential electrogenesis to a state where potassium conductance is in large part functionally masked (Waxman and Foster, 1980; Foster et al., 1980a). Such a developmental sequence is consistent with recent evidence that suggests a complementary distribution of sodium and potassium channels in mammalian myelinated fibers, the potassium channels being located primarily in the internodal axon membrane beneath the myelin sheath (Chiu and Ritchie, 1980; Ritchie and Ciu, 1981). It should be noted in this context that there may be differences in the detailed architecture between large and small fibers or fibers in different tracts. The important concept, however, that there are marked differences between nodal and internodal domains of the axon membrane appears well established. IV. Freeze-Fracture Structure of Myelinated Axons

A. INTRODUCTION TO FREEZE-FRACTURE The structural nature of biological membranes has been greatly elucidated by recent development of the freeze-fracture technique, which allows the exposure of large en face expanses of the membrane interior. Freeze-fracture usually involves the rapid freezing of an aldehyde-fixed, cryoprotected tissue, followed by freeze-cleaving the specimen under high vacuum, and coating of the exposed surface with a metal-carbon layer to form a replica (see Sleytr and Robards, 1977, and Stolinski, 1977, for review of methods). It is now generally accepted that

22 NVNXVM ‘ 3 N3Hd3J.S

STEPHEN G . WAXMAN

’1”

OSP

450

et al.

the process of freeze-fracture cleaves the membrane lipid bilayer and reveals the face of the lipid monolayer left adhering to either the protoplasm (“P-face”) or extracellular matrix (“E-face”). As demonstrated schematically in Fig. 8a,b, the view exposed by this procedure is one of a smooth background with a variable number of “particles” embedded in

FK.. 8. Schematic representation of freeze-fractured tissue. (a) The fracture plane is represented by the heavy line entering the upper plasmalemma at the arrows, traversing between the lipid bilayer, cross-fracturing through the cytoplasm, and again traversing between the lipid bilayer. Intramernbranous proteins (IMPs) are intercalated in the lipid bilayer. (b) Tissue above the fracture plane is removed by the fracturing process, and IMPs are observed as “particles” within a smooth rnonolipid background. T h e P-face is the half-membrane left adherent to the cytoplasm, while the E-face is the half-membrane immediately adjacent to the extracellular space (Ecs).

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

45 1

it. These particles, or intramembranous particles (IMPs), are generally interpreted as representing proteins or glycoproteins intercalated within the membrane (Branton, 1966; Pinto da Silva and Branton, 1970; Vailet al., 1974), and this concept is consistent with the fluid mosaic model of membrane structure (Singer and Nicolson, 1972). For most biological membranes, there is an asymmetrical distribution of particles between Pand E-fracture faces, with P-faces being moderately to heavily studded with particles and there being a scarcity of particles on E-faces.

OF MYELINATED AXONS B. FREEZE-FRACTURE

Freeze-fracture electron microscopy has been utilized extensively to describe the membrane structure of myelinated axons and associated glial cells (Livingston et al., 1973, Schnapp et al., 1976; Schnapp and Mugnaini, 1978; Rosenbluth, 1976, 1979; Kristolet al., 1977, 1978; Sandri et al., 1977; Ellisman, 1979; Wiley and Ellisman, 1980). These studies demonstrate that axonal membrane architecture is spatially heterogeneous, and that nodal, paranodal, and internodal regions each have unique patterns and distributions of IMPs. Paranodal membrane specializations have been the subject of much study (Livingston et al., 1973; Rosenbluth, 1976; Sandri et al., 1977; Schnapp and Mugnaini, 1978; Wiley and Ellisman, 1980; Tao-Cheng and Rosenbluth, 1980). Generally, the appearance of this region of axon membrane is paracrystalline, or ropelike, where axon and glial membrane are apposed, although this view may be altered by preparative techniques (e.g., fixation, cryoprotection) (Wiley and Ellisman, 1980). T h e paranodal region of the axolemma will not be further discussed in this article. In myelinated fibers, internodal membrane displays a marked asymmetrical distribution of IMPs (Kristol et al., 1978; Ellisman, 1979; Black et al., 1982a); the particle density on the P-face is = 1500/pm2,whereas the E-face particle density is 100-200/pm2. This partitioning of particles is in stark contrast to that observed for nodal axolemma, where both fracture faces contain 1500 IMPs/pm2 (Rosenbluth, 1976; Kristol et al., 1978; Ellisman, 1979; Tao-Cheng and Rosenbluth, 1980; Black et al., 1982a). Moreover, a further structural difference between nodal and internodal axolemma emerges when mean particle size (diameter) of these two regions is examined. Nodal membrane has a significantly greater percentage of large (>9.6 nm) particles than internodal membrane (Rosenbluth, 1976; Kristol et al., 1978; Ellisman, 1979; Tao-Chang and Rosenbluth, 1980; Black et al., 1982a). T h e distinction between “large” and “small” IMPs is based on criteria set forth in other studies of

-

-

452

STEPHEN G. WAXMAN

et al.

axon membrane (Kristol ~t al., 1978; Ellisman, 1979). This categorization of IMP size is obviously arbitrary but is based on data which suggest that the categories are a useful discriminator for nodal versus other axolemma. In fact, some reports have suggested that large IMPs may represent the morphological correlate of sodium channels (Rosenbluth, 1976; Kristol et n f . , 1978). A review of the evidence suggesting that large I MPs correspond to voltage-sensitive sodium channels at hodes of Ranvier is provided by Rosenbluth (1981a). Whether or not large intramembranous particles are, in fact, sodium conductance pathways, it is clear that nodal and internodal axolemma can be differentiated on the basis of both particle density and size distribution.

c. FREEZE-FRACTURE OF ADULTRAT RETINAL GANGLION CELLAXOLEMMA As noted above, axons of retinal ganglion cells offer a unique opportunity to investigate axolemmal changes during myelination (within the optic nerve segment) and to compare these membrane alterations with a portion (intraretinal segment) of the same axon which remains nonmyelinated. Retinal ganglion cells issue an axon from the vitread side of the perikaryon. This axon courses toward the optic disc as a nonmyelinated axon in the nerve fiber layer (NFL). Axons in the NFL are usually < 1 pm in diameter and travel toward the optic disc immediately subjacent to the vitreous body. T h e axons are densely packed with few if any intervening glial processes. Freeze-fracture electron microscopy of NFL axons reveals that, in the adult, there is a highly asymmetrical distribution of IMPs between the fracture faces. T h e E-face contains approximately 205 IMPs/pmZ(21% >9.6 nm), and the P-face has an IMP density of 1700/pmZ(1 1% >9.6 nm). I n this region of the trajectory of the axon, particles are generally random in distribution within E- and P-faces. T h e nerve fiber layer axons exit the retina and acquire myelin (in the adult) in the optic disc. As they exit the retina, the retinal ganglion cell axons, heretofore following fairly straight trajectories, begin a convoluted, tortuous course and become surrounded by glial processes. T h e glial envelopment of groups of axons in the optic disc is not unlike the Schwann cell envelopment of nonmyelinated axons in the PNS in that several axon profiles are associated with one glial process. T h e region of glial envelopment yields distally to the acquisition of myelin derived from oligodendrocytes in a region of heminodes of Ranvier. From this point the myelinated (adult) optic nerve continues its course to the brain.

-

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

453

As noted later in this article, some nonmyelinated fibers in the optic disc region exhibit a striking degree of spatial heterogeneity in terms of axolemmal structure. The adult optic nerve is virtually 100% myelinated. The axons of the optic nerve exhibit an average diameter (axon without myelin sheath) of 0.77 p m (20.25 F m SD; Black et al., 1982a) and have an average g-ratio (inner diameter of myelin : outer diameter of myelin) of 0.78. The size and g-ratio histograms are roughly unimodal. AS in all normal myelinated axons studied to date, optic nerve axons exhibit internodal, paranodal, and nodal regions characterized by axolemma with different morphological properties. I n thin sections of cytochemically stained (Waxman and Quick, 1977) adult axons, nodal membrane is the only area of axon membrane that exhibits ferric ion affinity. The staining is coextensive with nodal membrane alone, with little, if any, staining of adjacent paranodal or internodal membrane. Likewise, nodal membrane of optic nerve fibers exhibits specific ferric ion affinity (R. E. Foster, J. A. Black, and S. G. Waxman, unpublished observations). A remarkable characteristic of the optic nerve and other CNS myelinated tracts is clustering of nodes of Ranvier (cf. Peters et al., 1976). In some freeze-fracture replicas, as many as three adjacent nodes are present. I n thin sections, one can often find five or more nodal profiles and/or paranodal profiles in the same vicinity. The physiological consequence of the clustering of nodes in close proximity is unknown. As in the few CNS myelinated axons so far examined (Rosenbluth, 1976; Kristol et al., 1978; Schnapp and Mugnaini, 1978), optic nerve nodal axolemma (Figs. 9 and 10) is readily distinguishable from paranodal axon membrane and has unique freeze-fracture characteristics of IMP density (Table I) and size (Table 11).E-fracture faces of adult optic nerve nodes (Fig. 1lb) reveal a highly particulate membrane face. The density of E-face IMPs is 1316/pm2 (Table I), with almost 43% of the particles in the large (>9.6 nm) category (Table 11).T h e average size of an E-face IMP is 10.3 nm. P-face nodal axolemma (Fig. 1la) is likewise highly particulate (Table I), with a density of 1406 IMPs/pmZ.T h e mean size of nodal IMPs is 10.7 nm. As with E-face nodal membrane, P-face nodal membrane has many large particles; -49% of the IMPs are categorized as large (Table 11). Internodal membrane in rat optic nerve is distinct from the membrane at and adjacent to nodal membrane. The axon membrane in internodes is covered by compact myelin and does not exhibit ferric ion affinity. The freeze-fracture ultrastructure of internodal axolemma reveals a distinct difference between the E- and P-facture faces-the P-face (Fig. 12a) is highly particulate, whereas the E-face (Fig. 12b) is

454

STEPHEN G. WAXMAN P t

d.

F I G .9. Medium-power electron micrograph of a freeze-fracture replica of the P-face of a node of Ranvier (pN) from adult optic nerve. The nodal region has a high density of intramembranous particles. Terminal glial loops (G) delimit one side of this node. On the upper side of this node, a terminal loop has not been entirely fractured away and delimits this side of the nodal region. Scale bar: 1 pm. x42,OOO.

not. The E-face of internodal membrane has 104 IMPs/pm2 (Table I), with an average size of 8.6 nm. Of the E-face IMPS, only 25% are in the large category (Table 11). In contrast, the P-face of internodal membrane has 1709 IMPs/pmZ (Table I; Fig. 12a). However, the particle size distribution is similar to that of the E-face; the average size of P-face IMPS is 8.6 nm and 20% of the particles are classified as large (Table 11). The freeze-fracture observations summarized above indicate that optic nerve nodal and internodal axon membrane are similar to the

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

455

FIG. 10. Medium-power electron micrograph of a freeze-fracture replica of the E-face of a node of Ranvier (eN) from adult optic nerve. The nodal region is highly particulate. Terminal glial loops (G) are associated with a scalloped appearance on the E-face on the paranodal region (ePN). Note the IMPS that are linerally arranged in the “grooves” between adjacent glial loops (arrowheads). An astrocytic process (asterisk) is in close proximity to the nodal region. Scale bar: 1 pm. X42,OOO.

456

STEPHEN G. WAXMAN

et al.

TABLE 1 PARTICLE DENSITY PER gmZOF DEVELOPING A N D ADULT OPTICNERVE AXOLEMMA~ Fracture face Age (days)

Strain Long Evans Wistar Long Evans Wistar

I

12

Long Evans

14

Wistar

r

16

Long Evans

{

16

Wistar

28

Long Evans

14- 16

Long Evans/Wistar

.4dult

Long Evans

a

I I

Condition

PF

EF

Premyelinated Premyelinated Premyelinated Premyelinated Premyelinated Ensheathed Premyelinated Ensheathed Premyelinated Ensheathed Premyelinated Ensheathed Premyelinated MyelinatedC Myelinatedd Myelinated" Myelinatedd

512 2 50.7 553 ? 51.8 398 5 31.0 564 2 63.5 661 ? 77.8 1206 ? 105.5 588 t 95.4 730 2 56.3 431 5 17.4 929 t 96.4 629 5 29.2 924 2 83.6 599 5 29.2 1010 k 74.1 1175 2 62.2 1709 5 203.1 1406 t 152.2

125 t 16.8 124 ? 14.3 117 -+ 13.2 159 t 24.9 193 t 20.8b 104 ? 23.4b

104 t 13.gb 128 t 8.gb 67 90 t 21.3 1312 2 132.3 104 2 23.6 1316 ? 104.2

Mean 5 SEM. E-face not categorized as to premyelinated or ensheathed. Internode. Node.

other CNS myelinated axolemma that have been studied (Livingston et al., 1973; Rosenbluth, 1976; Sandri ef al., 1977; Kristol et al., 1978; Schnapp and Mugnaini, 1978). Thus, nodal membrane in the optic nerve is characterized in freeze-fracture images as a specialized region of high IMP density with many more large particles than adjacent axonal membrane. Bilateral paranodal specializations delimit nodal membrane, and the paranodal region is different from either node or internode. Development of nodal regions will be discussed below. Internodal membrane in the optic nerve of adult rats demonstrates some morphological similarities when compared to nonmyelinated axolemma in the same axon (i.e.,in the retinal nerve fiber layer) and to nonmyelinated axolemma of other neurons. Except for its covering by myelin, the internodal axolemma is cytologically indistinguishable from most regions of nonmyelinated axolemma in thin section analyses. Also like nonmyelinated axolemma, internodal optic nerve axolemma has no affinity for ferric ion staining. T h e freeze-fracture results described above also suggest a degree of morphological similarity (especially when

PARTICLE SIZEDISTRIBUTION ON PAge (days)

AND

TABLE I1 E-FRACTURE FACESOF DEVELOPING A N D ADULT RATOPTICNERVEAXOLEMMA Fracture face

Particles/pm* (19.6 nm)

Particles/pmz (>9.6 nm)

Percentage >9.6

Mean f SD (nm)

PF EF PF EF PF EF PF EF PF PF EF PF PF EF PF PF EF PF PF EF PF EF PF EF PF EF PF EF

420 101 508 109 309 79 495 147 615 1060 162 543 654 93 38 1 828 85 545 676 98 905 68 98 1 99 1 1366 78 715 751

92 24 45 15 89 38 69 12 46 146 31 45 76 11 51 101 19 84 248 30 105 22 194 32 1 343 26 69 1 565

17.9 18.8 8.1 7.7 22.4 32.9 12.2 7.7 6.9 12.1 16.2 7.7 10.4 10.5 11.6 10.9 18.6 13.3 26.9 23.3 10.4 24.7 16.5 24.5 20.1 25.3 49.2 42.9

8.2 f 2.5 7.9 f 2.6 7.3 f 2.0 7.6 f 2.0 8.6 2.1 9.0 f 2.5 7.6 +- 2.2 7.6 f 2.1 7.2 f 2.1 8.0 t 2.0 8.0 f 2.3 7.3 f 2.1 8.0 f 1.9 7.0 t 2.5 7.8 f 2.0 7.4 f 2.2 8.0 f 2.4 7.7 f 2.3 8.6 f 2.7 8.3 f 2.6 7.7 f 2.0 8.6 f 2.8 8.2 f 2.3 8.6 t 2.3 8.6 f 2.3 8.6 f 2.6 10.7 f 2.6 10.3 f 2.6

E-face not categorized as to premyelinated or ensheathed.

Internode.

Strain

2

Long Evans

2

Wistar

8

Long Evans

8

Wistar

Condition

( Premyelinated ( Premyelinated ( Premyelinated ( Premyelinated

12

Long Evans

Premyelinated Ensheathed

14

Wistar

Premye h a t e d Ensheathed

16

Long Evans

Premyelinated Ensheathed

16

Wistar

Premyelinated Ensheathed

14- 16

Long EvandWistar

14- 16

Long EvansIWistar

Adult

Long Evans

Adult

Long Evans

a

( Myelinated* [ MyelinatedC [ Myelinatedb ( Myelinatedc

Node.

*

FIG. 11. P- and E-fracture faces of paranodes and nodes of Ranvier from adult optic nerve axons. (a) The P-face of the node of Ranvier is demarcated (arrowheads) from the paranodal region, and contains a high percentage of large particles. (b) The highly particulate E-face of the node of Ranvier (arrowheads) is in contrast to the relatively particle-free paranodal region; linear aggregations of IMPS are present in grooves between glial loops (arrows). Scale bar: 0.25 pm. X 125,000.

FIG. 12. P- and E-face internodal axolemma from optic nerve of adult rat. (a) The P-face (PA) is highly particulate, and the particles appear to be randomly distributed. The myelin sheath (M) has been cross-fractured, as has the axoplasm on the left of the fiber. (b) the E-face contains few particles; compare this density to that in the nodal region (Fig. I Ib). Myelin (M) has been cross-fractured. Scale bar: 0.25 pm. X 100,000.

460

STEPHEN G . WAXMAN

et al.

contrasted with nodal membrane) between these putatively dissimilar axonal regions, since both internodal membrane and nonmyelinated membrane have a similar asymmetrical partitioning of particles between E- and P-faces (i.e., E-faces have low particle densities and P-faces have moderately high densities). This comparable characteristic is not compelling by itself since most biological membranes demonstrate such a partitioning (Branton, 1969). However, the difference in particle density between internodal and nonmyelinated axolemma is much smaller than the difference between internodal and nodal axon membrane. One difference between the nonmyelinated intraretinal segment and the optic nerve internode is the absolute number of E-face IMPs (205/pm2; 43/pm2 > 9.6 nm versus 104/pm2; 26/pm2 > 9.6 nm, respectively). It is interesting, however, that the percentage of large E-face particles is similar in these t w o types of membrane (21 and 2596, respectively). Indeed, one might expect on developmental grounds that IMP densities in nonmyelinated and internodal membrane would be relatively close, since the internodal membrane was derived from immature nonmyelinated (premyelinated) axolemma. One possibility, in fact, is that nodal and internodal domains of mature axolemma are derived by lateral diffusion of IMPs within the premyelinated membrane (Black et al., 1982a). Within the comparison between nonmyelinated and internodal axolemma there lies an enigma of significant importance. That is, nonmyelinated axolemma (e.g., rabbit vagus nerve) has been reported to contain saxitoxin-binding sites (putative sodium channels) at a concentration of 110/pm2 (Ritchie et al., 1976). T h e available evidence suggests that nonmyelinated axolemma is electrogenic, with conduction classically thought of as proceeding in a “continuous” manner. In contrast, the internodal axon membrane in rabbit sciatic nerve has been reported to contain few (<25/pm2) saxitoxin-binding sites (Ritchie and Rogart, 1977) and is not electrogenic (Chiu and Ritchie, 1981). Yet, the internodal membrane, as studied by cytochemistry and freeze-fracture, shows relatively small structural differences compared to electrogenic nonmyelinated membrane in the same and other CNS neurons. This apparent disparity, between the results of pharmacological and physiological studies (which suggest marked differences between nodal, internodal, and nonmyelinated membrane) on the one hand, and the morphological evidence presented here on the other hand, can not be resolved simply. It might be speculated that sodium channels are present in the internodal axolemma, but that in the process of myelination are changed in such a way as to make them inoperative without drastically changing their conformation as viewed by freeze-fracture (in the form of IMPs). This hypothesis, which reconciles the freeze-fracture results with the

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

461

physiological and pharmacological results, also focuses attention on the important possibility that sodium channels may be present, albeit in an altered and inactive form, in the internodal axolemma. If this hypothesis proves to be correct, then following demyelination, restoration of the ionic channel to its original structure could result in continuous conduction along the demyelinated axon. In fact, Bostock and Sears (1976, 1978) have demonstrated continuous conduction in diptheria-toxindemyelinated axons. A case can thus be made, from observations such as those of Bostock and Sears (1976, 1978) and the results noted above, for considering the hypothesis that myelinated axolemma may represent a regionally specialized derivative of nonmyelinated axolemma in which the internodal membrane remains morphologically similar to nonmyelinated axolemma, but has been functionally altered. Another possibility is that the differences in E-face IMP density between internodal and nonmyelinated axolema do, in fact, represent differences in the density of functional voltage-sensitive sodium channels. If this latter hypothesis is correct, then (assuming that internodal membrane in the optic nerve is electrically inexcitable) either the density of sodium channels in nonmyelinated membrane is just above that required to support impulse conduction, or the density of channels in the internode is close to the value necessary for electrical excitability. This latter possibility is consistant with the notion that relatively minor structural alterations could lead to continuous conduction in some demyelinated fibers. Finally, it is possible that E-face particles (or large E-face particles) do not represent a morphological correlate of sodium channels, but are related to some other membrane specialization, or that only a subset of the large IMPS are related to sodium channels. It is not possible at the present time to unequivocally distinguish between the alternatives outlined above. Nevertheless, freeze-fracture provides a clear structural marker for nodal-type membrane, and as discussed below, provides a probe of membrane structure which may be quite useful in terms of studying developmental processes.

V. Freeze-Fracture Studies on Myelin Development in Optic Nerve Axons

A. INTRODUCTION TO MYELINDEVELOPMENT Most freeze-fracture studies of the axon membrane during myelination (Wiley-Livingston and Ellisman, 1980; Tao-Chang and Rosenbluth, 1980; Rosenbluth, 1981a; Waxman et al., 1982) have focused on the

462

STEPHEN G. WAXMAN

et al.

development of regional aggregations of particles on the E-fracture faces. A major observation emerging from these studies is that patches of IMPS with a high percentage of large particles (>9.6 nm) can develop prior to the appearance of paranodal specializations. As myelin develops, the freeze-fracture pattern characteristic of paranodal axoglial junctions (transverse bands, paracrystalline axolemmal pattern) appears first in association with outermost layers of the sheath (Tao-Chang and Rosenbluth, 1980; Rosenbluth, 198la). T h e progression of axolemmal changes leading up to the appearance of specialized regions of the axon membrane (nodes, internodes) has been studied in detail by Black et al. (1982a). Their studies utilized rat optic nerve axons as a model system. As noted above, the axons of retinal ganglion cells are, at birth, nonmyelinated throughout their length (Skoff P t nl., 1976a,b), whereas, in the adult, essentially all fibers within the optic nerve segment acquire a myelin sheath (Forrester and Peters, 1967; Foster et al., 1982). This provides a tractable model system in which to monitor axolemmal changes during development of myelin along the axonal trunk. Earlier sections of this article have focused on a description of the axolemma of the intraretinal segment and the adult optic nerve segment of retinal ganglion cell axons. In the present section, developmental changes in optic nerve axonal membrane during the process of myelination are described. Figure 13 shows a schematic representation of the development of retinal ganglion cell axons. There are two major consequences of optic nerve development: (1) an increase in axonal size and (2) the development of a myelin sheath. T h e gradient of myelin development and intermediate axonal diameters that are observed between premyelinated (neonatal) and myelinated (adult) conditions are represented by the middle diagram in Fig. 13. Skoff et al. (1980) have reported that rat optic nerve myelination is not uniform along the length of the fibers, but occurs at several discrete sites along the axon and in a general rostrocaudal direction. Thus, a given axon during development may have nonmyelinated, loosely myelinated, or compactly myelinated regions. Figure 3a,b reviews the time course of the growth of rat optic nerve axons and the rate of myelination. Several important points in the development of optic nerve myelination should be reiterated. First, myelination begins at approximately the sixth postnatal day and proceeds rapidly, such that by 28 days of age -85% of the fibers are myelinated. Second, neonatal rat optic nerve axons are relatively uniform in diameter (mean ? SD = 0.14 k 0.03 pm), and diameter increases by 50% during the first 8 days (mean diameter % SD = 0.21 % 0.03 pm). Third, at the time of appearance of compact myelin, an increase in axonal

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

ROth 2 D A Y

463

Optic NervejTract

I I

! I

12

D

A Y

FIG. 13. Schematic drawing of retinal ganglion cell axons at three ages. At birth and for several days thereafter, the axon of a retinal ganglion cell is totally nonmyelinated (top). Myelination of the optic nervehact segment of the retinal ganglion cell axons begins at -6 days postnatal, and by 12 days regions of the nerve are compactly myelinated, loosely myelinated, ensheathed, or remain nonmyelinated (middle). In the adult, the optic nerve/ tract is entirely myelinated (bottom). As well as acquiring myelin, the optic nerve fibers have a fourfold increase in axonal diameter. Note that the intraretinal segment of the axon does not acquire myelin at any age. (From Black et al., 1982a.)

diameter is observed, which continues until the adult size is reached (mean 2 SD = 0.77 ? 0.18 pm). Finally, those axons that are not yet myelinated remain at 0.2-0.3-pm diameter until myelination begins.

B. PREMYELINATED AXOLEMMA T h e term premyelinated, as used in this article, refers to axons which are not yet visibly associated with glial cells. Ensheathed axons, in contrast, have established a relationship with glial cells that may contact and at least partially envelop them (Black et al., 1982a). As shown in Fig. 2a, the optic nerves of 2-day-old rats do not contain myelinated fibers, and the axons are fairly uniform in diameter and surrounded by extensive, electron-lucent extracellular space. The axolemmal structure, as revealed by freeze-fracture, of premyelinated fibers from 2-day-old rat optic nerves is demonstrated in Fig. 14. The fibers have a moderate number of particles on the P-faces, while few particles are observed on the E-faces. The IMPS appear to be randomly distributed, and are not aggregated or grouped. I n this respect, the premyelinated fibers are very similar to mature nonmyelinated axons, though the particle densities of the two are widely different.

464

STEPHEN G . WAXMAN

et al.

FIG. 14. High-magnification electron micrograph of freeze-fracture replica from 2-day-old Wistar rat optic nerve. At this stage of development, all fibers are nonmyelinated (premyelinated). The P-face (PA) is moderately particulate and has a heterogeneous population of IMPS. The E-face (eA) has few particles. On both fracture faces, the particles appear randomly distributed. Scale bar: 0.25 Fm. X 125,000.

T h e freeze-fracture image of premyelinated axon membrane from 16-day-old animals is shown in Fig. 15. Despite the 2-week difference in age, the axolemmal structure is remarkably similar to that at 2 days. Table 1 summarizes the mean IMP densities for premyelinated axolemma (2-28 days; Black et al., 1982a), and it shows that, despite some fluctuations, the density remains similar over this period of time. Mean IMP sizes for E- and P-faces of premyelinated fibers are shown in Table 11. Again there are some variations on the respective fracture faces; however, there is not a major change in the mean size of the particles. T h e conclusion that can be reached from these data, then, is that the axolemmal architecture, as revealed by conventional freeze-fracture methods, of premyelinated fibers does not change substantially prior to glial ensheathment. Since axon diameter increases 50- 100% prior to myelination and length of the axons increases, membrane structure need not, however, be regarded as static prior to myelination. T h e fact that the

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

465

FIG. 15. High-magnification electron micrograph of premyelinated axons from 16day-old Long Evans rat optic nerve. Both the P-face (PA) and E-face (eA) have IMP densities similar to respective fracture faces from 2-day-old animals (cf. Fig. 14).Scale bar: 0.25 pm. x 125,000.

membrane maintains an essentially stable particle density despite substantial increase in area supports, in fact, an ongoing process of IMP insertion. CHANGES ASSOCIATED WITH GLIAL ENSHEATHMENT C. AXOLEMMAL From about 6 days of age, some axons in the optic nerve are closely associated with oligodendroglial processes (e.g., Fig. 2b), and a few axons are myeiinated. Profiles of glial envelopment comparable to those seen in thin-section electron micrographs are rarely seen in freeze-fracture replicas of 8-day-old optic nerve, although “lingulae”of oligodendrocytic processes are observed among premyelinated fibers. However, by 12 days postnatal, many examples of axons closely associated with oligodendroglial processes are apparent. As noted above, these fibers have been termed “ensheathed” to differentiate them from premyeli-

466

STEPHEN G . WAXMAN

et al.

nated and myelinated axons. T h e fact that both premyelinated and ensheathed axons can be observed in single replicas of 12- to 28-day-old optic nerve permits a direct comparison of the axolemma of these two types of fibers, in which age, per se, is not a variable (e.g., Fig. 18a,b below). Examples of ensheathed fibers are shown in Figs. 16 and 17. Encircling glial processes are present, though compact myelin is not apparent. Concomitant with association with glial projections, ensheathed axons show t w o major structural axolemmal changes. First, there is a significant increase in the density of IMPs on the P-face of ensheathed axons as compared to premyelinated axons at the same age. And second, in certain regions of ensheathed axolemma, there are aggregations of particles on E-faces. T h e difference in IMP densities between premyelinated and ensheathed axolemma can be appreciated in Fig. 18a,b and is summarized for different ages in Table 111. T h e mean IMP size increases slightly between premyelinated and ensheathed axons, suggesting the

FIG. 16. Freeze-fracture electron micrograph of ensheathed axon from 16-day-old Long Evans rat optic nerte. The P-face (PA) of the ensheathed axon has moderately high density of IMPs. Wrappings of the ensheathing oligodendrocyte have been exposed to reveal P- (pC) and E-faces (eG). Scale bar: 0.25 pm. x 120,000.

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

467

FIG. 17. Freeze-fracture electron micrograph of ensheathed and premyelinated axons from 14-day-old Wistar rat optic nerve. An oligodendroglial process (pG) surrounds the ensheathed fiber. Note the difference in P-face particle densities between the ensheathed axons (PA) and the premyelinated fiber (asterisk) immediately adjacent to it. Scale bar: 0.25 pm. x 120,000. (Modified from Black et al., 1982a.) TABLE 111 DISTRIBUTION OF SMALL A N D LARGE PARTICLES ON P-FACES OF PREMYELINATED AND ENSHEATHED RATOPTICNERVE AXOLEMMA Age (days)

Strain

12

Long Evansn

14

Wistar

16

Long Evans"

16

Wistarb

Condition Prem yelinated Ensheathed Premyelinated Ensheathed Premyelinated Ensheathed Premyelinated Ensheathed

Total particles/pm2

Particles/pm2 ( 5 9 . 6 nm)

Particles/pme (>9.6 nm)

Mean (nm)

66 1 1206 588 730 432 929 629 924

615 1060 543 654 380 828 545 676

46 146 45 76 51 101 84 248

7.2 8.0 7.3 8.0 7.8 7.4

7.7 8.6

Difference between premyelinated and ensheathed significant at p < .005, Student's t-test. Difference between premyelinated and ensheathed significant at p < .01, Student's t-test.

STEPHEN G . WAXMAN

et al.

Fie,. 18. Freeze-fracture electron micrographs of premyelinated (a) and ensheathed (b) axons from 12-day-old Long Evans rats. The difference in particle densities on the P-faces of these fibers is apparent. Also note the increase in axonal diameter with glial ensheathment. Scale bar: 0.1 pm. x 150,000. (From Black e! al., 1982a.)

insertion of a population of relatively large P-face particles with ensheathment. It is not clear, as yet, whether this increase in P-face particles induces glial ensheathment or is a response to glial association. It is tempting, however, to suggest that there is a causal relationship between changes in axon membrane structure and glial ensheathment. Alternatively, it is possible that other mechanisms may govern these processes. Aggregations of E-face particles prior to axoglial membrane specializations have been reported in developing PNS axons by Wiley-Livingston and Ellisman (1 980) and Tao-Chang and Rosenbluth (1980) and in developing CNS axons (Waxmanetnl., 1982). An example of such a clustering of particles is shown in Fig. 19. This aggregate of IMPS contains particles in higher density than found over other parts of the axolemmal E-face. Moreover, the aggregate contains a high percentage of large

FIG. 19. Freeze-fracture electron micrograph of developing optic nerve from 16-dayold Long Evans optic nerve. An aggregation of partides (arrowheads) is observed on the generally low-particulateE-face (eA). Oligodendrocytic processes (asterisks)are associated with this axon. The P-face (PA) of an adjacent axon is also observed. Scale bar: 0.25 pm. ~ 7 9 , 2 0 0(Modified . from Waxman el al., 1982.)

470

STEPHEN G . WAXMAN

et al.

particles. It is notable that these IMP aggregations occur in regions in which edges of glial processes are apparent. It has been suggested (Wiley-Livingston and Ellisman, 1980; Tao-Chang and Rosenbluth, 1980; Waxman et al., 1982) that these aggregates may be the precursors of nodes of Ranvier. It is interesting, in this regard, that cytochemical observations also suggest the differentiation of nodal-like axon membrane prior to the formation of compact juxta-nodal myelin or wellformed axoglial junctions (Waxman and Foster, 1980).

By 14- 16 days postnatal, unambiguous examples of internodal and nodal axolemma are apparent in freeze-fracture replicas. As with adult internodal axon membrane, there is a highly asymmetrical partitioning of particles in the internodal membrane of 14- to 16-day-old animals, with an abundance of IMPs on the P-face and few particles on the E-face (Table I). There is, however, a substantial difference in the P-face density between the internodal membrane in 14- to 16-day-old versus adult axons. T h e difference can be appreciated in Fig. 20. T h e P-face IMP density difference between the two age groups is not reflected in the I M P values for respective E-faces. Table IV presents the mean density and size of IMPs for 14- to 16-day-old and adult internodal membrane. There is not a substantial change in the size distribution between 14-to 16-day-old and adult fibers for either the P- or E-face. By 14-16 days of age, some nodes of Ranvier have clearly differentiated, and are apparent in freeze-fracture replicas (e.g., Fig. 21). These nodes are similar to adult nodes in that the nodal axolemma has a relatively symmetrical distribution of IMPs. Both the E- and P-faces have a high density of particles, and these densities are similar to those obTABLE IV INTEKNODE AXOLEMMAL PAKTICLE DENSITY A N D SIZE

Age

Fracture face

Total particles/pm*

Particles/pm* ( 5 9 . 6 nm)

Particledpm* (>9.6 nm)

Mean particle size (nm)

14-16days Adult" 14-lfidays Adult

PF PF EF EF

1010 1709 90 104

905 1366 68 78

105 343 22 26

8.6 8.6 8.6

a

Significant compared to 14- 16 day at p < ,005.

7.7

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

471

FIG.20. Freeze-fracture electron micrograph of internodal region of myelinated fiber from 14- to 16-day-old optic nerve. The P-face (PA) on the internodal axolemma is highly particulate. Compare the density of particles on this internodal membrane with that of premyelinated or ensheathed axolemma (Figs. 15 and 16, respectively). The myelin sheath (M) has been cross-fractured. Inset: P-face of internodal axolemma from adult optic nerve at the same magnification. Note the increased density of IMPS. Scale bar: 0.25 pm. X 125,000. (Modified from Black et al., 1982a.)

472

STEPHEN G . WAXMAN

et al.

FIG.2 1 . Freeze-fracture electron micrograph of nodal region of myelinated fiber from 14- to 16-day-old optic nerve. The E-face of the node (eN) is highly particulate. The node is bounded on the left by the paranodal region (ePN). The axolemma in the paranodal region is scalloped by subjacent terminal glial loops. Note the large particles that are present in the paranodal region; many of the particles are linearly arranged in the grooves of the paranodal axolemma (arrowheads). Scale bar: 0.5 pm. x 120,000. (Modified from Black et al., 1982a.)

served within adult nodal membrane (Table V). In contrast, though, the mean IMP size of E- and P-faces for 14- to 16-day-old nodal membrane is substantially less than that of corresponding fracture faces of adult nodal axolemma (Table V). T h e differences summarized above between the axolemma of myelinated fibers from 14- to 16-day-old animals in comparison to adult animals indicate that the development of optic nerve axolemma does not TABLE V NODE AXOLESIMAL PARTICLE DENSITY A N D SIZE

Age

Fracture face

Total particles/pmZ

14-16days Adult 14-16 days Adult

PF PF EF EF

1175 1406 1312 1316

Particles/~mP Particles/prn2 (I: 9.6 nm) (>9.6 nm) 98 I 715 99 1 751

194 691 32 1 565

Mean particle size (nm) 8.2 10.7 8.6 10.3

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

473

cease with the acquisition of myelin and formation of nodes of Ranvier. T h e axon membrane continues to develop, the further changes occur before the axon membrane achieves its adult structure.

E. ABERRANT AXOGLIAL ASSOCIATION It should also be noted that in occasional instances, apparent errors in axon membrane architecture occur. These may represent aberrant axoglial junctions. Figure 22 shows, for example, the E-fracture face of an apparently aberrant axoglial junction from the optic disc region of an adult Long Evans rat. The axolemma surface appears indented in the region where it displays a ropelike appearance characterized by curvilinear bands approximately 30 nm apart. There is some similarity between the structure of this membrane specialization and those observed at normal paranodal axoglial junctions (Wiley and Ellisman, 1980). It should be noted, though, that these linear striations are oriented parallel to the axis of the axon, which is in contrast to the circumferential nature observed in normal axoglial junctions. The structure of this junction from an apparently normal animal is quite similar to that observed in the mutant “shiverer” mouse (Rosenbluth, 1981b). It appears that some

FIG.22. Freeze-fracture electron micrograph from optic disc region of adult Long Evans rat showing aberrant axon membrane structure. The E-face of this axon is indented and appears to form a membrane specialization. Linear striations approximately 30 nm apart are oriented parallel to the axis of the axon in the region of indentation. Scale bar: 0.5 pm. ~ 7 2 , 6 0 0 .

474

STEPHEN G. WAXMAN

et al.

aberrant axon membrane specializations occur in apparently normal animals. Whether these aberrant membrane areas are static, or are modified so as to develop more normal structure, has not yet been established.

F. SVMMARY OF DEVELOPMENTAL CHANCES I N AXONMEMBRANE On the basis of the findings summerized above, it is possible to make several statements regarding the development of the axon membrane in optic nerve: 1. Premyelinated fibers show relatively similar membrane morphology regardless of the age of the animal. 2. Concomitant with glial ensheathment, there is a striking change in axon membrane structure, that is, a marked increase in the number of particles on the P-face of the axolemma. 3. During the developmental sequence, IMPS aggregate on E-faces in regions that presumably become nodes of Ranvier; this aggregation occurs prior to the formation of compact myelin or paranodal axon-glial junctions. 4. T h e nodal axolemma continues to change after nodes form; aduIt nodal axolemma contains a similar density but different size distribution of E- and P-face particles than axolemma at 14-16 days. 5. Similarly, newly formed internodal axolemma continues to change (with an increase in P-face IMP density) with further maturation. These observations are summarized in Fig. 23 and indicate that significant membrane reorganization occurs during the development of rat optic nerve. T h e axolemma from 2-day-old rat optic nerve does not show the particle aggregations or axolemmal specializations that are observed on the fracture faces of adult myelinated fibers. Moreover, premyelinated axonal membrane from 2-day-old animals is distinct from ensheathed axolemma at 12-16 days and from adult axolemma in terms of IMP density and size distribution. From the results presented above it can be seen that the retinal ganglion cell axon trunk undergoes remarkable changes between neonatal and adult forms. T h e optic nerve axolemma from the 2 day old is uniformly particulate, and from this homogeneous beginning there is derived an end product which becomes myelinated and is differentiated into nodal and internodal membrane. T h e freeze-fracture results from developing optic nerve provide clear evidence for membrane plasticity in developing myelinated axons.

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

;; 2d

M

475

L.

FIG. 23. Summary histogram of IMP density for Long Evans axonal membranes: premyelinated/ensheathed optic nerve (ON) and myelinated ON [16d and adult (A)] (Blacket al., 1982a); myelinated ON and retinal nerve fiber layer (Nfl) (S. G. Waxman, J. A. Black, and R. E. Foster, unpublished data); other nonmyelinated (Black et al., 1981). The densities are expressed as the number of particles per pm2 for each of the samples. On the absciksa, the age of the animal from which the sample was taken is indicated (i.e., A, adult; 2d, 2 days postnatal, etc.) above the sampled area (I, internode; N, node of Ranvier; Cc, corpus callosum; Cb, cerebellar parallel fibers; Vg, vagus nerve). The open and crosshatched histogram bars are the density values for P-face axolemma, and the solid bars are the values for E-face axolemma. The cross-hatched bars are P-face densities for ensheathed axons; the open bars are P-face densities for premyelinated axolemma. The standard error of the mean is indicated as the line extending from the top of each bar. (From Black et al., 1982a.)

VI. Differentiationof the Axon Membrane in the Absence of Myelin

The development of the proximal axon segment of retinal ganglion cells (within the intraretinal nerve fiber layer) has been studied by Black et al. (1983).These intraretinal axon segments remain nonmyelinated in the adult, in contrast to the ganglion cell axon segments in the optic nerve, which as noted previously, become myelinated in the adult. Nota-

476

STEPHEN G. WAXMAN

et al.

bly, there is an increase in P-face particle density of the intraretinal nerve fiber layer axons, beginning at approximately the same time as the commencement of myelination in the distal optic nerve segments of the axons; between 8 and 16 days of age, there is a 50% increase in P-face particle density of the intraretinal axons. Nevertheless, even in neonatal animals, the axolemma of the intraretinal axon segments exhibits a different structure from the axolemma of the optic nerve segments with respect to IMP size distribution. T h e mean IMP sizes for P- and E-faces of intraretinal axons are both 7.2 nm, compared to values of 8.2 and 7.9 nm, respectively, for the corresponding faces of the axolemma in the optic nerve segment (Black et al., 1983). These data suggest, in agreement with cytochemical results (Waxman and Foster, 1980), that the axolemma even at this early developmental stage is capable of organizing itself into regions that will remain nonmyelinated and regions that will become myelinated. It should be recalled that at 2 days of age, mature oligodendrocytes have not yet differentiated in the rat optic nerve (Skoff et al., 1976a). Thus, a direct glial influence on axon membrane structure, at this stage of development, may not be involved in this aspect of axonal differentiation. Finally, it should be noted that, although the most striking differentiation of the axon membrane occurs along myelinated fibers, that is, between nodal and internodal axolemma, the axolemma of some nonmyelinated fibers can exhibit a high degree of spatial heterogeneity. One example of this spatial heterogeneity in the absence of myelin is provided by developing fibers prior to myelination. As discussed above, it has been shown, for example, that ventral root fibers prior to myelination exhibit focal regions approximately 1 p m in length, with the cytochemical characteristics of nodal membrane, at the junction between adjacent Schwann cells (Waxman and Foster, 1980). Freeze-fracture studies similarly show aggregations of E-face particles in developing myelinated axon trunks prior to the formation of compact myelin or paranodal axoglial junctions (Ellisman, 1979; Wiley-Livingston and Ellisman, 1980; Rosenbluth, 1981a; Waxman et al., 1982). Thus, during normal ontogenesis, the axolemma differentiates into regions with morphological properties similar to those of normal nodes of Ranvier, and regions devoid of nodal properties, prior to compact myelin formation. Of course, it is also well known that the growing tips of developing neurites, in regions devoid of myelin, exhibit a gradient of particle sizes and densities, and reorganize as development proceeds (Pfenninger and Bunge, 1974).

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

477

Bray et al. (1979) and Ellisman (1979) have noted, in the amyelinated axons of the dystrophic dy/dy mouse, occasional irregular membrane patches in which E- and P-face particles occur in high density. Bray et al. (1981) suggested that these particles might correspond to sites of inward membrane current generation. Physiological heterogeneity of the axon membrane in demyelinated fibers has been reported by Bostock et al. (1980) and Smith et al. (1982). In ventral root axons demyelinated with lysophosphatidyl choline, longitudinal current analysis revealed discontinuous conduction, in some cases with the presence of discrete regions of inward current termed “phi-nodes” (Bostock et al., 1980; Smith et al., 1982). Although the morphological correlate of the phi-nodes has not yet been definitively ascertained, it is possible that, in these remyelinating fibers, these discrete foci of inward current represent precursors of nodes of Ranvier. The demonstration of these spatially separated sites of inward current represents a physiological demonstration of spatial heterogeneity of nonmyelinated membrane. Finally, freeze-fracture observations of retinal ganglion cell axons in the region of transition from nonmyelinated (retinal nerve fiber layer) to myelinated (optic nerve) zones of the axons reveals a highly specialized morphological structure, in which nonmyelinated fibers exhibit a high degree of spatial heterogeneity (Black et al., 1982b). Figure 24 shows three axons from the optic disk of adult rat. The E-face particles of these axons are aggregated into apparently circumferential bands of particles that are 1-3 IMPs in width. These bands of IMPs show a distinct periodicity of approximately 0.16 pm. There is a high percentage (approximately 25%) of large (>9.6 nm) particles within these linear bands (Fig. 24, inset). The mean density of the E-face particles within this specialized axolemma, computed without consideration of the pattern of particle distribution, is 316/pm2.This value is only slightly higher than the E-face particle density (204/pm2) in the more proximal nonmyelinated region of the axons within the nerve fiber layer of the retina. Black et al. (1982b) noted that, if it is assumed that the large particles observed in the bands represent sodium channels, the nonrandom distribution of particles in optic disk fibers might be related to the mechanism of impulse electrogenesis. They further noted that the focal accumulations of E-face particles could represent foci of inward current which might play a role in facilitating conduction between nonmyelinated and myelinated segments. Irrespective of the implications for the mode of conduction, however, the findings illustrated in Fig. 24 demonstrate that, in certain spe-

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

479

cialized regions, the axolemma of nonmyelinated fibers can exhibit distinct spatial heterogeneity. Using transmission electron microscopy of thin sections stained with lead citrate and uranyl acetate, Hildebrand and Waxman (1983)observed focal patches of undercoating subjacent to the axolemma in these fibers. This morphology could be consistent with a nonuniform mode of conduction. Although in the latter case a relationship to glial processes was noted, the important point is that both methods (freeze-fracture and conventional thin-section analysis) reveal a nonuniform structure of the axolemma in these specialized regions of nonmyelinated fibers. As outlined above, observations on normal developing axons, demyelinated axons, and specialized nonmyelinated axons all indicate that spatial heterogeneity of the axolemma can occur in the absence of myelin. It thus appears that the capability to form a heterogeneous structure may be a property inherent to the axon membrane. Waxman and Foster (1980) suggested that the axon membrane differentiates into nodal and internodal regions before, or early in the process of, myelination, and speculated that this differentiation might provide a signal demarcating regions to be covered by the myelin-forming cell. The axonal demarcation hypothesis might provide a partial explanation for the specific patterns of myelination described earlier in this article.

VII. Concluding Comments

As discussed previously in this article, myelinated fibers exhibit a high degree of regional differentiation and specificity with respect to the pattern of myelin formation. There is now clear evidence for membrane plasticity during development of myelinated fibers. This, together with the physiologically demonstrated segregation of the axon membrane into nodal and internodal regions containing different populations of ionic channels, raises the important issue of how the axon membrane develops this high degree of differentiation during ontogenesis. ~

FIG. 24. Freeze-fracture electron micrograph of nonmyelinated axons with the optic disc region of adult rat retina. Note linear bands of IMPS (arrowheads), 1-3 particles wide, on the E-faces of three adjacent axons (al, a2, a3). Periodicity of bands -0.16 pm. Portions of glial processes are observed between adjacent axons. Scale bar: 0.5 pm. ~65,000.Inset: Higher magnification of E-face particles in a 1. Many particles are large (>9.6 nm). Scale bar: 0.1 pm. X 150,000.

480

STEPHEN G. WAXMAN

et al.

As discussed in detail above, a tractable experimental model system for studying this question is provided by the optic nerve of the rat. Physiological studies during development of this central white matter tract indicate that there are alterations in physiological properties which cannot be explained on the basis of changes in axonal size, or on the basis of myelination, alone. One inference that could be made from the electrophysiological observations on developing optic nerve is that there are changes in functional architecture of the axon membrane during development. T h e freeze-fracture observations indeed show striking structural changes in axon membranes of this model system during development. It is interesting that fibers prior to myelination show similar membrane morphology regardless of the age of the animal. This, together with the result noted above, that is, that conduction properties change in a manner that cannot be explained by increases in axonal diameter prior to myelination, suggests that factors other than changes in axon membrane properties may play a role in determining conduction properties prior to myelination. Alternatively, it is possible that there are changes in the membrane, but these changes are not revealed by freeze-fracture. Concomitant with association of axons with glial cells, however, there are marked changes in membrane structure. Regions which will presumably differentiate into nodes of Ranvier become apparent at this time, and the remainder of the membrane exhibits distinct developmental alterations in terms of P-face particle density. Thus, at the time of glial ensheathment, there are distinct alterations in the axon membrane morphology. These changes, while the most striking, d o not constitute the end-point of the developmental sequence. Following the formation of nodes, development continues, and there appears to be a continued process of insertion of large particles into the axon membrane. Definitive interpretation of these findings will obviously depend crucially on a deiineation of the identity of the various particle-types which are observed in the axon membrane. Studies aimed at elucidating the identity of intramembranous particles are in progress, and the results will obviously be of great value. Even prior to the delineation of the identity of IMPS, however, it now appears quite clear that the axon membrane does not exhibit a static and fixed structure, but, on the contrary, exhibits a high degree of plasticity during development. Moreover, it is notable that, in some specialized regions, the axon membrane of adult nonmyelinated fibers also exhibits a striking degree of regional specialization. As noted above, one crucially important question concerns the identity of the intramembranous particles. In this regard, it might be inquired whether potassium channels, as well as sodium channels, are

ONTOGENESIS OF AXOLEMMA AND AXOGLIAL RELATIONSHIPS

48 1

revealed by freeze-fracture. It also might be asked whether there is a gradient of channels within various parts of the axolemma, for example, within the internode. Mechanisms of ionic channel aggregation are not yet fully understood, and have obvious importance for the understanding of the organization of axonal membranes. The mechanisms by which new channels are synthesized and incorporated into the axon membrane must also be studied. Finally, the question of ionic channel distribution and turnover, and of possible reorganization, in demyelinated axon membranes must be addressed (Waxman and Ritchie, 1981). We anticipate that all of these questions will receive intensive study over the coming years, and expect that many of them will soon be answered.

Acknowledgments

The authors wish to thank Susan K. Cameron, Mary E. Smith, and Marsha S. Kantor for excellent technical assistance and Joyce Hughes for preparation of this manuscript. The studies reported in this article have been supported in part by the Medical Research Service, Veterans Administration, and by grants from the National Institutes of Health, National Multiple Sclerosis Society, and the Kroc Foundation.

References

Aguayo, A. J., Charron, L., and Bray, G. M. (1976).J . Neurocytol. 4, 395-418. Aguayo, A. J., Bray, G. M., and Perkins, S. C. (1979). Ann. N.Y. Acad. Sci. 317, 247-264. Bennett, M. V. L. (1970). Annu. Rev. Physiol. 32,471-528. Bishop, G. H., Clare, M. H., and Landau, W. M. (1971). Znt. J . Neurosri. 2, 69-78. Black, J. A., Foster, R. E., and Waxman, S. G. (1981).J. Neurocytol. 10, 981-993. Black, J. A., Foster, R. E., and Waxman, S. G. (1982a). Bruin Res. 250, 1-20. Black, J. A., Waxman, S. G., and Foster, R. E. (1982b). Cell Tissue Res. 224, 239-246. Black, J. A., Foster, R. E., and Waxman, S. G. (1983).J. Neurocytol. (in press). Bostock, H., and Sears, T. A. (1976). Nature (London) 263, 786-787. Bostock, H., and Sears, T. A. (1978).J . Physiol. (London) 280, 273-301. Bostock, H., Hall, S. M., and Smith, K. J. (1980).J. Physiol. (London) 308, 21p-23p. Bostock, H., Sears, T. A., and Sherratt, R. M. (1981).J. Physiol. (London) 313, 301-315. Branton, D. (1966). Proc. Natl. Acad. Sci. U..S.k. 55, 1048-1056. Branton, D. (1969). Annu. Rev. Plant Physiol. 20, 209-238. Bray, G., Cullen, M. J., Aguayo, A. J., and Rasminsky, M. (1979). Neurosci. Lett. 13, 203208. Bray, G., Rasminsky, M., and Aguayo, A. J. (1981). Annu. Rev. Neurosci. 4, 127-162. Brill, M. H., Waxman, S. G., Moore, J. W., and Joyner, R. W. (1977).J. Neurol., Neurosurg. Psychiatry 40, 769-774.

482

STEPHEN G . WAXMAN

et al.

Brismar, T. (1979). .4ctn Phyiol. Srnttd. 105, 384-386. Brismar, T. (1980).J. Phwiol. (Lmdon) 298, 171-184. Brismar, T. (1981). Acta Phyiol. Srntid. 113, 167-176. Chiu, S. Y., and Ritchie, J. M. (1980). S n t u r e (Loitdon) 284, 170-171. Chiu, S. Y., and Ritchie, J. M. (1981). ,4drr. Seurol. 31, 313-328. Chiu, S. Y., and Ritchie, J. M. (1982).J. Phyzal. loti do^) 322, 485-501. Chiu, S. Y., Ritchie, J. M., Rogart, R. B., and Stagg, D. (1979).J. Physiol. (London) 292, 149- 166. Conti, F., Hille, B., Neumcke, B., Nonner, W., and Stamp&, R. (1976).J. Phyiol. (London) 262,699-727. Coombs, J. S., Curtis, D. R., and Eccles, J. C. (1957).J. Phjszol. (London) 139, 232-249. Cullheim. S., and Ulfhake, B. (1979).J. Coznp. Srurol. 188, 679-686. Duncan, D. (1934). J. Comp. N ~ u r o I .60, 437-47 1. Ellisman, M. H. (1979).J. Seuror~tol.8, 719-735. Fleischauer, K.. and Wartenberg, H. (1967). Z. Z~llforsch.t\likrosk. .4nnf. 83, 568-581. Forrester, J., and Peters, A. (1967). S a t u r e (London) 214, 245-247. Foster, R. E., Connors, B. W., and Waxman, S. G. (1980a).iVVuurosri. Sor. Abstr. 6, 291. Foster, R. E.. Whalen, C. C., and Waxman, S. G. (198Ob). Srioiw 210, 661-663. Foster, R. E., Connors, B. W., and Waxman, S. G. (1982). Dn). Bmiir Rrs. 3, 371-386. Fraher, J. P. (1972).J. .4iurt. 112, 99-124. Friede, R. L.. and Samorajski, T. (1968).J. Cotup. ,Yriirol. 130, 223-232. Fuortes, M. G . F., Frank, K., and Becker, M. C. (1957).J. Gm.Phyiol. 40, 735-752. Gledhill, R. F., Harrison, B. M., and McDonald, W. 1. (1973). Nnturr (London) 244, 443444. Gutrecht, J. A,, and Dyck, P. J. (1970).J. Cornp. I\leur~rl.138, 117-130. Hess, A . , and Young, J. 2. (1952). Pror. R. Soc. London, Sm. B 140, 301. Hildebrand, C., and Waxman, S. G. (1983). Arniri RP,Y.258, 23-32. Huxlev, A. F., and Stamp& R. (1949).J. P h ~ $ o l .(Loridon) 108, 315-339. Ito. M., and Takahaski, I. (1960).Irr “Electrical Activity of Single Cells” (Y. Katsuki, ed.), pp. 159-179. lkagu Shoin Ltd., Tokyo. Kocsis. J . D., and Waxman, S. G. (1980). .Vnture (Lotidurt) 287, 348-349. Kocsis, J. D., and Waxman, S. G. (1981). ,4din. “t’mrol. 31, 299-312. Kocsis, J. D., Malenka, R. C., and Waxman, S. G. (1981). Braivt Res. 207, 321-331. Kocsis, J . D., Waxman, S. G., and Hildebrand, C. (1982a). Pror. 1nt. Cungr. N r u r o n u x . Ilk., ith, 26, 2. Kocsis, J . D., Waxman. S. G., Hildebrand, C., and Ruiz, J. (1982b).Pror. Roy. Sor. London B , 217,277-287. Kristol, C., Akert, K., Sandri, C., Wyss, U. R., Bennett, M. V. L., and Moor, H. ( 1 977). Rrnin Re\. 125, 197-212. Kristol, C., Sandri, C., and Akert, K. (1978). Brain RCS. 142, 391-400. Livingston, R. B., Pfenninger, K., Moor, H., and Akert, K. (1973). Brain K ~ s 58, . 1-24. Lund, R. D., and Bunt, A. H. (1976).J. Cotnp. “purol. 165, 247-264. McDonald, W. I., and Ohlrich, G. D. (1971).J. Annt. 110, 191-202. Matthews, M. A,, and Duncan, D. (1971).J. Comp. ,\’!purol. 142, 1-22. Meiri, H., Spira, M. E., and Parnas, I. (1981). Sririrrr 211, 709-712. Meszler, R. M., Pappas, G. D., and Bennett, M. V. L. (1974).J. N r u r o r p l . 3, 251-261. Paintal, A. S. (1967).J. Phyz’ol. (London) 193, 523-533. Palay, S. L., Sotelo, C., Peters, A., and Orkand, P. M. (1968).J. Cell B i d . 38, 193-201. Peters, A., Proskauer, C. C., and Kaiserman-Abramof, I. R. (1968).J. Cell Biol. 39, 604619.

ONTOGENESIS OF AXOLEMMA A N D AXOGLIAL RELATIONSHIPS

483

Peters, A., Palay, S. L., and Webster, H. deF. (1976). “The Fine Structure of the Nervous System: The Neurons and Supporting Cells.” Saunders, Philadelphia. Pfenninger, K., and Bunge, R. P. (1974).J. Cell Biol. 63, 180-196. Pinto da Silva, P., and Branton, D. (1970).J. Cell Biol. 45, 598-605. Preston, R. J., Waxman, S. G., and Kocsis, J. D. (1983). Exper. Neurol. (in press). Quick, D. C., and Waxman, S. G. (1977).J. Neural. Sci. 31, 1-11. Quick, D. C., and Waxman, S. G. (1978).J. Neurol. Sci. 35, 235-241. Revenko, S. V., Timin, Y. N., and Khodorov, B. I. (1973). Biojzzku 18, 1074-1078. Ritchie, J. M. (1982a).J . Physiol. (London) 322, 20P. Ritchie, J. M. (1982b). Proc. R . Sac. London, Ser. B , 215, 273-287, Ritchie, J. M., and Chiu, S. Y. (1981).Adv. Neural. 31, 329-342. Ritchie, J. M., and Rogart, R. B. (1977).Proc. Nutl. Acud. Sci. U.S.A. 74, 211-215. Ritchie, J. M., Rogart, R. B., and Strichartz, G. (1976).J. Physiol. (London) 261, 477-494. Ritchie, J. M., Rang, H. P., and Pellegrino, R. (1981). Nature (London) 294, 257-259. Rosenbluth, J. (1976).J. Neurocytol. 5, 731-745. Rosenbluth, J. (1979).Int. Congr. Ser.-Excerflta Med. 455, 200-209. Rosenbluth, J. (1981a). Adv. Neurol. 31, 69-92. Rosenbluth, J. (1981b). Bruin Res. 208, 283-297. Rushton, W. A. H. (1951).J. Physiol. (London) 115, 101-122. Salzer, J. C., Bunge, R. P., and Glaser, L. (1980).J. Cell Biol. 84, 767-778. Sanders, F. K., and Whitteridge, D. (1946).J. Physiol. (London) 105, 152-174. Sandri, C., Van Buren, J. M., and Akert, K. (1977). Prog. Bruin Res. 46, 1-380. Schnapp, B., and Mugnaini, E. (1978).I n “Physiology and Pathobiology of Axons” (S. G. Waxman, ed.), pp. 83-123. Raven Press, New York. Schnapp, B., Peracchia, C., and Mugnaini, E. (1976). Neuroscience 1, 181-190. Sherratt, R. M., Bostock, H., and Sears, T. A. (1980). Nature (London) 289, 570-572. Simpson, S. A., and Young, J. Z. (1945).J. Aizat. 79, 48-65. Singer, S. J., and Nicolson, G. L. (1972). Science 175, 720-731. Skoff, R. P., Price, D. C., and Stocks, A. (1976a).J. Comp.Neurol. 169, 291-312. Skoff, R. P., Price, D. C., and Stocks, A. (1976b).J. Camp.Neural. 169, 313-334. Skoff, R. P., Toland, D., and Nast, E. (1980).,J.Comp. Neural. 191, 237-253. Sleytr, U. B., and Robards, A. W. (1977).J. Microsc. 3, 77-100. Smith, K. J., Blakemore, W. F., and McDonald, W. I. (1981). Bruin 104, 383-404. Smith, K. J., Bostock, H., and Hall, S. M. (1982).J. Neural. Sci. 54, 13-3 1. Spencer, P. S., and Weinberg, H. J. (1978). In “Physiology and Pathobiology of Axons” (S. G. Waxman, ed.), pp. 389-406. Raven Press, New York. Stolinski, C. (1977). Mimon 8, 87-111. Strichartz, G., Small, R., Nicholson, C., Pfenninger, K. H., and Llinas, R. (1980). Neurosci. Abstr. 6, 660. Swadlow, H. A., and Waxman, S. G. (1976). Ex$. Neurol. 53, 115-127. Tao-Cheng, J. H., and Rosenbluth, J. (1980). Bruin Res. 199, 249-265. Thomas, P. K., and Young, J. Z. (1949).J. Anat. 83, 336-351. Vail, W. J., Papahadjopoulos, D., and Moscarello, M. A. (1974). Biochim. Biophys. Actu 345, 463-467. Vizoso, A. D., and Young, J. Z. (1948).J. Anut. 82, 110-135. Waxman, S. G. (1970). Nature (London) 227, 283-284. Waxman, S. G. (1972). Bruin Res. 47, 269-288. Waxman, S. G. (1977).Arch. Neurol. (Chicago) 34, 585-589. Waxman, S. G. (1982). N . Engl. J . Med. 306, 1529-1533. Waxman, S. G., and Anderson, M. J. (1980). Cell T k u e Res. 208, 343-352.

4 84

STEPHEN C . WAXMAN

et al.

Waxman, S. G., and Bennett, M. V. L. (1972). h’uturr A’rujBiol. 238, 217-219. Waxman, S. G., and Brill, M. H. (1978).J. Nrurol., il’purosurg. Psyrhintry 41, 408-417. Waxman, S. G., and Foster, R. E. (1980).Brain Res. R m . 2, 205-234. Waxman, S. G., and hlelker, R. J. (1971).Brain Res. 32, 445-448. Waxman, S. G., and Quick, D. C. (1977).J. ,l’eurol., Neurosurg. Psyrhiat? 40, 379-386. Waxman. S. G.. and Quick, D. C. (1978).Brnin Res. 144, 1-10. Waxman, S. G., and Ritchie, J. M. (1981).,4do. Nrurol. 31, 511-513. Waxman, S. G., and Swadlow, H. A. (1977).Prog. LVeurobzol.8, 297-325. Waxman, S. G., Pappas, G. D., and Bennett, M. V. L. (1972).J. CrllBiol. 53, 210-224. Waxman, S. G., Bradley, W. G., and Hartwieg, E. A. (1978).Proc.R. Sor. London, Srr. B 201, 301-308. Waxman, S. G., Black, J. A., and Foster, R. E. (1982).h’eurologj 32, 418-421. Weinberg, H . J., and Spencer, P. S. (1976).Brain Rts. 113, 363-378. Weiner. L. P., Waxman, S. G., Stohlman, S. A., and Kwan, A. (1980). Ann. Nruiol. 8, 580-584. Werman. R., and Grundfest, H. (196l).J. G m . Phvsiol. 44, 997-1027. Wiley, C . A,, and Ellisman, M. H. (1980).J. Crll B i d . 84, 261-280. Wiley-Livingston, C. A , , and Ellisman, M. H. (1980).Drv. Biol. 79, 334-355. Williams, P. L., and Wendell-Smith, C. P. (1971).J. Anat. 109, 505-526. Wood, P. M., and Bunge, R. P. (1975).lVn.‘nlure(Lmidon) 256, 662-664. Zenker, W. (1964).Z. Zdlforsrh. dfikrmk. ,4iiu/. 62, 531-545.

INDEX

A Absence seizures acetylcholine role, 125 y-aminobutyric acid in, 133- 134 serotinergic mechanisms, 119 Acetylcholine effect on equilibrium constants for cholinergic ligands, 308 receptors, see Acetylcholine receptors role in seizures, 122-128 Acetylcholine-operated channels, behavior, 313-329 B-burst, 326-328 as diagnostic tool, 328-329 in embryonic and denervated muscle, 322-323 multiple grating phenomenon, 318-319 N-burst, 319-321 open-close switching, 323-324 rate-limiting steps, 321-322 silent intervals, 324-326 Acetylcholine receptors, 259-34 1 as archetype, 260-261 cx subunit, 288-289 f3 subunit, 288 biosynthesis, 272-279 charge, 271 6 subunits, 286-288 gating related to state of ligation, 330-331 hydrodynamic properties, 26 1-264 immunochemical structure, 27 1-272 in membrane ion-translocation function, 301-329 three-dimensional topography, 279-290 molecular properties, 261-272 molecular weight and size, 261-264 v protein effect on, 292-294 oligomeric structure, 281-283 protein primary structure, 269

prosthetic groups, 267-269 secondary structure, 270 specific activity and polypeptide com position, 266-267 subunit composition, 264-269 topographic mapping, 274-275 translational dynamics, 295 .4dipocytes cx,-adrenergic receptor binding by, 364-365 a2-adrenergicbinding in, 402-403 a-Adrenergic receptors biochemical responses mediated by, 347 pharmacological subdivision, 345-349 radioligand binding studies, 349-354 cwl-Adrenergicreceptors, 343-43 1 agonist affinity ratio studies, 350 antagonist affinity ratio studies, 352 biochemical responses mediated by, 347 characterization by radioligand binding, 354-364 agonist radioligands, 364 antagonist radioligands, 354-364 multiple affinity states, 363-364 effector systems coupled to, 364-368 adipocytes, 364-365 central nervous system, 367-368 salivary glands, 365-367 regulation, 368-376 circadian rhythm, 374-375 down-regulation, 37 1-372 pathological, 375-376 physiological, 372-375 up-regulation, 369-37 1 solubilization of, 376 up-Adrenergic receptors, 343-43 1 agonist affinity ratio studies, 350 antagonist affinity ratio studies, 353 biochemical responses mediated by, 347 characterization by radioligand binding, 377-398 agonist radioligands, 387-398 antagonist radioligands, 377-387

485

486

Index

conipdrison, 404-4 10 intact-cell binding, 387 effector systems coupled to, 398-404 adipocytes, 402-403 human platelets, 401-402 N G 108- 15 cells, 403-404 localization, 117-4 19 regulation. 410-417 down-regulation, 412-4 15 heterologous, 4 15-416 pathological, 416-4 17 physiological, 4 16 up-regulation, 110-41 1 solubilization of, 4 19-420 Adrenocorticotropic hormone (ACX’H), epileptic seizures and, 148-150 Alpha motoneuron, see Motoneuron Amino acids, epileptic seizures and. 140-142 y-Aminobutyric acid, w GABA p-.4minoclonidone, as a,-adrenergic agonist. 393-398 Amylobarbitone, sti-ucture and activity, 16 Antiacetylcholine receptor antibodies antigenic determinants in, 12-13 assays for, 5-7 by immunoprecipitation, 6 by inhibition, 6-7 in cerebrospinal fluid, 10-12 of human skeletal muscle, 13 myasthenia gravis and, 1 - 14 in various disease forms, 7- 10 pathogenicity of. 2-5, 11-12 complexed antibodies. 4-5 free antibodies, 3-4 of torpedo fish, 12-13 Anticonvulsants effect on acetylcholine, 126- 127 seroronin, 120- 121 reaction with catecholamines, 114-1 15 L-Aspartic acid, seizures and, 14 1-142 Axoglial relationships to axolemma, 433-484 aberrant, 473-474 Axolemma axoglial relationship to. 433-484 aberrant, 473-474 changes of, with glial ensheathment, 465-470

myelinated, 470-473 premyelinated. 463-465 of retinal ganglion cells, freeze-fracture, 4.52-461 Axon(a1) conduction, barbiturate effect on. 21-22 membrane differentiation in myelin absence, 475-479 summary of developmental changes in, 474 myelinated, freeze-fracture, 45 1-452 optic nerve, 474 freeze-fractur-e studies of myelination in, 461-474

Barbitone, structure and activity, 16 Barbiturates biochemical and neurochemical studies. 34-44 convulsant, neuropharmacology, 33-34 effects on axonal conduction, 2 1-22 mitochondria1 respiration, 34-35 neuronal membranes, 32-33 neurotransmitter release and uptake, 35-44 synaptic transmission, 16-2 1 transmitter action, 25-32 general structures and activities, 16 neuropharmacological studies on. 16-2 1 pharmacology, 15-49 presynaptic actions, 22-24 B-burst, in acetylcholine-operated channels, 326-328 RE-2254 (compound), as al-adreriergic antagonist, 362 Brain-gut peptides. epileptic seizures and, 147-148 Brainstem motoneurons hyperpolarization during sleep, 233-235 membrane potential during sleep and wakefulness, 2 18-225 Burst, definition, 3 15 Butobarbitone, structure and activity, 16

487

Index

C Catecholaminergic projections, in CNS, table of, 108 Catecholamines anticonvulsant reaction with, 114-1 15 drugs affecting, 115-1 16 role in seizures, 107- 116 Central nervous system (CNS) al-adrenergic receptor binding in, 367-368 catecholaminergic projections, 108 cholinergic pathways, 122 serotinergic projections, 1 16 Cerebrospinal fluid (CSF) antiacetylcholine receptor antibodies in, 10-12 cross-reactivity with brain antibodies, 11 origin, 10- 11 neurotransmitters in, seizure studies, 115

serotinin in, seizures and, 121 CHEB (barbiturate), structure and activity, 16 Cholinergic ligands, equilibrium dissociation constants, 308-3 1 1 Cholinergic pathways, in central nervous system, 122 Circadian rhythm, in al-adrenergic receptor binding, 374-375 Clonidine, as a2-adrenergic agonist, 393-398 Convulsants (chemical) barbiturates, neuropharmacology, 33-34 seizures from, 101, 103 Cyclic nucleotides, epileptic seizures and, 142-143

D Decamethonium, effect on equilibrium constants for cholinergic ligands, 309 Dihydroergocr y ptine as a,-adrenergic antagonist, 362-363 as a2-adrenergic antagonist, 377-380

E Electrical stimulation of seizures, 103 by kindling, 104-105 Electroconvulsive shock, seizures from, 101 Endorphins and enkephalins identification and localization, 62-87 immunoassay versus radioimmunoassay of, 74-87 immunodetection, 5 1-92 reliability, 87-88 sensitivity, 62-74 tissue extraction, 57-62 tissue processing and handling, 53-57 Epilepsy, see also Seizures, epileptic developmental aspects, 150- 152 diversity of disorders in, 94-95 experimental models, 99- 106 neurochemistry, 93- 180 Epinephrine as az-adrenergic agonist, 387-393 receptors for, 344-345

F Freeze-fracture of axolemma, 452-461 of myelin development in optic nerve axons, 461-474 of myelinated axons, 45 1-452 technique, 449-45 1

G GABA importance as neurotransmitter, 181- 182 markers, in monkey brain, 129 seizures and, 128-140 drug interaction and, 135-140 GABA receptors, 181-212 binding, 193-196 drug effects, 196-197 hormone effects, 197-199 regulation, 196-201 biochemical aspects, 192-204 classification, 201-203

-188

Index

electrophvsiological studies. 182- 192 inhihit ion. 182- 188 t i t ivfw riiodels, 184- 188 postsvnapric, 183 presynaptic. 183- 184 pharmacological aspects. 189- 192 solubilization. 203-204 tvpes. 1x8-I XY Genetic ni0dt45 of seizures, 102, 103, 106 I 4;lutamic acid, seizures and, 141-142 Clvcine. epileptic seizures and, 140 Guanfacinc. as uy-adrenergic agonist, 398

H Hr.xamethoniurri, effecr on equilibrium constants tor cholinergic ligands. 310 Hormones. effect o n GABA receptor binding. 1!)7- 199 Human skeletal muscle, antiacetylcholine receptor antibodies of. I3 Hypertension, a I-adrenergir receptor binding and, 373-356

I fort translocation

by membrane-hound acet! Icholine receptor, 301-329 channel activation, 304-305 desensitization and binding equilibria, 305-3 12

K Kindling o f seizures. 104- I03

L

hlesencephalon. motoneuron control and, 24 1-243 klesgdiencephalic junction. motoneuron control and, 241-243 hfethohexitone, structure and aciivity, 16

Mitochondria1 respiration. bdrbiturate effects on, 34-35 hlotoneurons control during sleep, 213-258 central control mechanisms, 240-251 extracellular reflect studies on, 240-245 medulla, 245 mesencephalon, 24 1-243 pons. 243 hyperpolarization, synaptic mcchanisrris for, 232-240 intracellular studies, 246-251 medullary reticular, control studies on, 250-251 membrane potential during sleep and wakefulness, 218-232 i n brainstenr, 2 18-225 i n spinal cord, 225-23 I trigeminal, control studies on, 246-250 Musclc. acetylcholine-operated channels in, 322-323 Myasthenia gravis antiacetylcholine receptor antibodies and, 1-14 central nervous system involvernent. 11-12 Myelinated fiber, axolemma and axoglial relationships in, 433-484 hryelination, specificity in 437-440 hfyoclonic models of seizures, 104 Myoclonic seizures acetylcholine role, I25 y-aminobutyric acid and, 134 carecholamines and. 112

Lisuride. as a,-adrenergic antagonist, 386

3M2B (barbiturate). structure and actitit;, 16

Medulla, motoneuron control and, 245 hfedullary reticular neurons, control studies on. 250-251

N-burst, in acetylcholine-operated channels, 3 19-32 1 Neurocheniistry of epilepsy, 93- 180 Neuronal mentbranes, barbiturate effects on, 32-33 Ncurophysiology of epileptic seizures, 95-99

489

Inidex Neurotransmitters barbiturate effects on, 24-32 binding properties, 41-44 role in seizures, 106- 150 NG 108-15 cells, a*-adrenergic binding in, 403-404 Nicotine, effect on equilibrium constants for cholinergic ligands, 3 11 Nicotinic acetylcholine receptors, (nAcChR) reduction in myasthenia gravis, 1 Norepinephrine as a2-adrenergicagonist, 387-393 receptors for, 344-345 u proteins, 290-301 effect on acetylcholine receptors, 292-294 thiol-reaction role, 295-296 function studies, 290-301 topographical localization, relationship to acetylcholine receptor structure, 296-30 1

0 Opiate peptides receptor and ligands, 146 role in epileptic seizures, 143-146 Optic nerve axons, myelin development in, 461-474 development, 440-449 electrophysiological studies, 441 -449

P Pentobarbitone, structure and activity, 16 Peptides brain-gut type, role in epileptic seizures, 147- 148 role in epileptic seizures, 143-150 Phenyltrimethylammonium, effect on equilibrium constants for cholinergic ligands, 3 11 Platelets, ae-adrenergic receptor binding in, 401-402 Pons, motoneuron control and, 243 Pontomesencephalic junction, motoneuron control and, 243 Prazosin, as al-adrenergic antagonist, 354, 360-362 Presynaptic activity of barbiturates, 22-24

Protein A, in antibody precipitation, 6

R Rauwolscine, as a?-adrenergic antagonist, 380-386

5 Salivary glands, al-adrenergic receptor binding in, 365-367 Secobarbitone, structure and activity, 16 Seizures, epileptic absence models, 102-103 acetylcholine role, 122-128 clinical correlates, 127- 128 drug effects, 126-127 ACTH and, 148-150 y-aminobutyric acid and, 128-140 L-aspartic acid and, 141-142 catecholamine role, 107-1 16 anticonvulsants and, 114-1 15 from chemical convulsants, 101, 105-106 classification, 95 cyclic nucleotides and, 142-143 developmental aspects, 150-152 from electrical stimulation, 102 kindling, 104-105 experimental models, 99- 106 table, 100 genetic models, 102, 106 L-glutamic acid and, 141 glycine and, 140 myoclonic models, 104 neurophysiology, 95-99 neurotransmitter role, 106-150 partial-seizure models, 104-106 peptide role, 143-150 serotonin role, 116-122 anticonvulsant role, 120-121 CSF studies, 121 taurine and, 140-141 Serotonin role in seizures, 116-122 Sleep motoneuron control during, 213-258 motoneuron membrane potential during, 218-232 somatic reflex activity during, 2 15-2 17 states of, 214

490

Index

Somatic reflex activity, during sleep and wakefulness, 2 15-2 17 Spinal cord motoneurons hyperpolarization during sleep. 233-235 membrane potential, during sleep and wakefulness. 225-23 1 Suberyldicholine, effect on equilibrium constants for cholinergic ligands. 31 1 Synaptic transmission, barbiturate effects on. 16-21

T Taurinc, epileptic seizures and, 140-141 Thiol reactions, role in u proteinacetylcholine receptor interaction, '195-296 Thiopentone, structure and activity, I6 Torpedo fish. antiacetylcholine receptor antibodies of, 12-13

Transmitters in central neurons, barbiturate effects on, 24-32 Trigeminal motoneurons control mechanisms, 246-250 d-Tubocurarine. effect on equilibrium constants for cholinergic ligands, 310-311

W Wakefulness, somatic reflex activity during, 215-217 M'B 4101 (compound), as a,-adrenergic antagonist, 3.54, 360-362

Y I'ohimbine, as a,-adrenergic antagonist. 380-386

CONTENTS OF RECENT VOLUMES Volume 12

Drugs and Body Temperature Peter Lomax Pathobiology of Acute Triethyltin Intoxication R . Torack, J . Gordon, andJ. Prokop Ascending Control of Thalamic and Cortical Responsiveness M . Steriade

The Fine Structural Localization of Biogenic Monoamines in Nervous Tissue F k y d E. Bloom Brain Lesions and Amine Metabolism Robed Y. Moore Morphological and Functional Aspects of Central Monoamine Neurons Kjell Fuxe, Tomas Hakfelt, and Urban Ungqerstedt

Theories of Biological Etiology of Affective Disorders John M . Davis

Uptake and Subcellular Localization of Neurotransmitters in the Brain Solomon H. Synder, Michael J . Kuhar, Alan 1. Green, Joseph T. Coyle, and Edward G. Shaskan

Cerebral Protein Synthesis Inhibitors Block Long-Term Memory Samuel H. Barondes

Chemical Mechanisms of TransmitterReceptor Interaction John T. Garland and Jack Durell

The Mechanism of Action of Hallucinogenic Drugs on a Possible Serotonin Receptor in the Brain J . R . Smythies, F. Bmington, and R . D . Morin

The Chemical Nature of the Receptor Site-A Study in the Stereochemistry of Synaptic Mechanisms J . R. Smythies

Simple Peptides in Brain Isamu Sano

Molecular Mechanisms in Information Processing Georges Ungar

The Activating Effect of Histamine on the Central Nervous System M. Monnier, R. Sauer, and A. M . Hatt Mode of Action of Psychomotor Stimulant Drugs Jacques M . van Rossum

The Effect of Increased Functional Activity on the Protein Metabolism of the Nervous System B. Jakoubek and B. Sm$inovskj Protein Transport in Neurons Raymond J . Lasek

AUTHOR INDEX-SUBJECT INDEX

Neurochemical Correlates of Behavior M . H. Aprison and J . N. Hingtgm

Volume 13

Some Guidelines from System Science for Studying Neural Information Processing Donald 0. Walter and Martin F. Gardiner

Of Pattern and Place in Dendrites Madge E . Scheibel and Arnold 8. Scheibel

AUTHOR INDEX- SU BJECT INDEX

49 1

492

Contents of Recent Volunies

Volume 14

The Pharmacology of Thalamic and Geniculate Neurons J . W. Phillis The Axon Reaction: A Review ofthe Principal Features of Perikaryd Responses to Axon Inquiry A . R . Liebnman

C 0 2Fixation in the Nervous Tissue Szf-Chuh Chm,C Reflections on the Role of Receptor Systems for Taste and Smell John G. Sinclair Central Cholinergic Mechanism and Behavior S.A'. Pradhan and S N . Duita The Chemical Anatomy of Synaptic Mechanisms: Receptors J . R . Smpthies A U T H O R INDEX-SUBJECT INDEX

Volume 15

Projection of Forelimb Group I Muscle Afferents to the Cat Cerebral Cortex fngmar Rosin Physiological Pathways through the Vestibular Nuclei victor^] Wilson Tetrodotoxin, Saxitoxin, and Related Substances: Their Applications in Neurobiology Martin H . Eoans The Inhibitory Action of y-Aminobutyric Acid, A Probable Synaptic Transmitter Kunihiko Obata Some Aspects of Protein Metabolism ofthe Neuron Mei Satake

Chemistry and Biology of Two Proteins, S-100 and 14-3-2, Specific to the Nervous System B l a h W. Moore T h e Genesis of the EEG Rafael Elul Mathematical Identification of Brain States Applied to Classification of Drugs E R . ~ J o h n P. , Walker. D. Cawood, M Rush, and^ J. Gehnnann AUTHOR INDEX-SUBJECT INDEX

Volume 16

Model of Molecular Mechanism Able to Generate a Depolariziation-Hyperpolarization Cycle Clara Torda Antiacetylcholine Drugs: Chemistry, Stereochemistry, and Pharmacology T. D. Inch and R . W. Brimblecornbe Kryptopyrrole and Other Monopyrroles and Molecular Neurobiology Donald G . Irvine RNA Metabolism in the Brain Victor E. Shashoua

A Comparison of Cortical Functions in M a n and the Other Primates R.E . Passin~qharnand G. Ettlinger Porphyria: Theories of Etiology and Treatment H. A . Peters, D . J , Cripps, and H. K Reese SUBJECT I N D E X

Volume 17

Epilepsy and y-Aminobutyric Acid-Mediated Inhibition B. S. Meldrum

Contents of Recent Volumes

493

O n Axoplasmic Flow Liliana Lubiriska

Synaptosomal Transport Processes Giulio Leui and Maurizio Raiteri

Schizophrenia: Perchance a Dream? J. Christian Gillin and Richard J - Wyatt

Glutathione Metabolism and Some Possible Functions of Glutathione in the Nervous System Marian Orlowski and Abraham Karkowsky

SUBJECT IN D E X

Volume 18

Neurochemical Consequences of Ethanol on the Nervous System A r m K. Rawat

Integrative Properties and Design Principles of Axons Stephen G. Waxman

Octopamine and Some Related Noncatecholic Amines in Invertebrate Nervous Systems H . A . Robertson and A . V. Juotio

Biological Transmethylation Involving S-Adenosylmethionine: Development of Assay Methods and Implications for Neuropsychiatry Ross J . Baldessarini

Apormorphine: Chemistry, Pharmacology, Biochemistry F C. Calpaert, W. F M . Van Beuer? and,/. E . M . F. Leysm

Synaptochemistry of Acetylcholine Metabolism in a Cholinergic Neuron Bertalan Csillik Ion and Energy Metabolismofthe Brain at the Cellular Level Leg Hertz and Arne Schousboe Aggression and Central Neurotransmitters S. N . Pradhan A Neural Model of Attention, Reinforcement and Discrimination Learning Stephen Grossberg Marihuana, Learning, and Memory Ernest L. A be1 Neurochemical and Neuropharmacological Aspects of Depression 8.E. Leonard SUBJECT INDEX

Volume 19

Do Hippocampal Lesions Produce Amnesia in Animals? Susan D. Iversen

Thymoleptic and Neuroleptic Drug Plasma Levels in Psychiatry: Current Status Thomas B. Cooper, Georqe M. Simpson, and,J. Hillary Lee SUBJECT INDEX

Volume 20

Functional Metabolism of Brain Phospholipids G. Bnan Ansell and Sheila Spanner Isolation and Purification of the Nicotine Acetylcholine Receptor and Its Functional Reconstitution into a Membrane Environment Michael S. Bn'ley and Jean-Pierre Changeux Biochemical Aspects ofNeurotransmission in the Developing Brain Joseph T. Coyle The Formation, Degradation, and Function of Cyclic Nucleotides in the Nervous System John W. D a b Fluctuation Analysis in Ncurobiology Louis J. DeFelice

494

Contents of Recent Volumes

Peptides and Behavior Gcoyes Ungar Biochemical Transfer of Acquired Information S R . Mitrhdi, J M . Beaton. and R. ,J. Rradlfy Aminotransferase Activity in Brain M . Bentuk and A . Lajtha The Molecular Structure of Acetylcholine and Adrenergic Receptors: An All-Protein Model J . R. Smythies Structural Integration of Neuroprotease Activity E h Gabrieiescu Lipotropin and the Central Nervous System W. H . Gispen.,J. M . van Rec, and D. de Wied Tissue Fractionation in Neurobiochemistry: An AnalyticalTwloraSourceofArtifacts Pime Laduron Choline Acetyltransferase: A Review with Special Reference to Its Cellular and Subcellular Localization Jean Rossier

Presynaptic Inhibition: Transmitter and Ionic Mechanisms R. A . Nicoll and B. E. Aker Microquantitation of Neurotransmitters in Specific Areas of the Central Nervous System Juan M. Saavedra Physiology and Glia: Glial-Neuronal Interactions R. Malcolm Stewart and Roger N . Rosenberg Molecular Perspectives of Monovalent Cation Selective Transmembrane Channels Dan W. Urp Neuroleptics and Brain Self-stimulation Behavior Albert Wauquier

Volume 22 Transport and Metabolism of Glutamate and GABA in Neurons and Glial Cells Arne Schousboe Brain Intermediary Metabolism in Vbo: Changes with Carbon Dioxide, Development, and Seizures Alexander L. Miller

SUBJECT INDEX

Volume 21

Relationship of the Actions of Neuroleptic Drugs to the Pathophysiology of Tardive Dyskinesia Ross J . Baldessarini and Daniel Tarsy Soviet Literature on the Nervous System and Psychobiology of Cetacea ThedoreH. Bullockand VladimirS. Gurevich Binding and Iontophoretic Studies on Centrally Active Amino Acids-A Search for Physiological Receptors F. V. DcFeudis

N,N-Dimethyltryptamine: An Endogenous Hallucinogen Steven A . Barker, John A . Monti, and Samuel T. Christian Neurotransmitter Receptors: Neuroanatomical Localization through Autoradiography L . Charles Munin Neurotoxins as Tools in Neurobiology E. G. McGeer and P. L. McGeer Mechanisms of Synaptic Modulation William Shain and David 0. Carpenter Anatomical, Physiological, and Behavioral Aspects of Olfactory Bulbectomy in the Rat B. E. Leonard and M . Tuite

Contents of Recent Volumes The Deoxyglucose Method for the Measurement of Local Glucose Utilization and the Mapping of Local Functional Activity in the Central Nervous System Louis Sokoloff INDEX

Volume 23

495

Glucocorticoid Effects on Central Nervous Excitability and Synaptic Transmission Edward D. Hall Assessing the Functional Significance of Lesion-Induced Neuronal Plasticity Oswald Steward Dopamine Receptors in the Central Nervous System Ian Creese, A. Leslie Morrow, Stuart E . Lt$j David R . Sibley, and Mark W. Hamblin

Chemically Induced Ion Channels in Nerve Cell Membranes David A . Mathers and Jeffery L. Barker

Functional Studies of the Central CatechoIamines T. W. Robbins and B. J . Eueritt

Fluctuation of Na and K Currents in Excitable Membranes Berthold Neumcke

Studies of Human Growth Hormone Secretion in Sleep and Waking Wallace B. Mendelson

Biochemical Studies of the Excitable Membrane Sodium Channel Robert L. Barchi

Sleep Mechanisms: Biology and Control of REM Sleep Dennis J . McGinty and Rent! R . Drucker-Colin

Benzodiazepine Receptors in the Central Nervous System Phil Skolnick and Steven M . Paul Rapid Changes in Phospholipid Metabolism during Secretion and Receptor Activation F. T. Crews

INDEX

This Page Intentionally Left Blank