Functional Neuroanatomy of the Nitric Oxide System

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Author: H.W.M. Steinbusch | J. de Vente | S.R. Vincent

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FUNCTIONAL NEUROANATOMY OF THE NITRIC OXIDE SYSTEM

FUNCTIONAL NEUROANATOMY OF THE NITRIC OXIDE SYSTEM

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H A N D B O O K OF CHEMICAL NEUROANATOMY Series Editors: A. Bj6rklund and T. H6kfelt

Volume 17

FUNCTIONAL NEUROANATOMY OF THE NITRIC OXIDE SYSTEM Editors:

H.W.M. STEINBUSCH and J. DE VENTE European Graduate School of Neuroscience (EURON), Department of Psychiatry and Neuropsychology, Maastricht University, PO Box 616, 6200 MD Maastricht, The Netherlands

S.R. VINCENT Graduate Program in Neuroscience, University of British Columbia, Kinsman Laboratory of Neurological Research, Department of Psychiatry, Vancouver, BC V6T 1Z3, Canada

2000

ELSEVIER Amsterdam-

Lausanne - New York-

Oxford-

Shannon - Singapore - Tokyo

ELSEVIER SCIENCE B.V. Sara Burgerhartstraat 25 P.O. Box 211, 1000 AE Amsterdam, The Netherlands

© 2000 Elsevier Science B.V. All rights reserved.

This work is protected under copyright by Elsevier Science, and the following terms and conditions apply to its use: Photocopying Single photocopies of single chapters may be made for personal use as allowed by national copyright laws. Permission of the Publisher and payment of a fee is required for all other photocopying, including multiple or systematic copying, copying for advertising or promotional purposes, resale, and all forms of document delivery. Special rates are available for educational institutions that wish to make photocopies for non-profit educational classroom use. Permissions may be sought directly from Elsevier Science Rights & Permissions Department, PO Box 800, Oxford OX5 1DX, UK; phone: (+44) 1865 843830, fax: (+44) 1865 853333, e-mail: [email protected]. You may also contact Rights & Permissions directly through Elsevier's home page (http://www.elsevier.nl), selecting first 'Customer Support', then 'General Information', then 'Permissions Query Form'. In the USA, users may clear permissions and make payments through the Copyright Clearance Center, Inc., 222 Rosewood Drive, Danvers, MA 01923, USA; phone: (978) 7508400, fax: (978) 7504744, and in the UK through the Copyright Licensing Agency Rapid Clearance Service (CLARCS), 90 Tottenham Court Road, London W1P 0LP, UK; phone: (+44) 171 631 5555; fax: (+44) 171 631 5500. Other countries may have a local reprographic rights agency for payments. Derivative Works Tables of contents may be reproduced for internal circulation, but permission of Elsevier Science is required for external resale or distribution of such material. Permission of the Publisher is required for all other derivative works, including compilations and translations. Electronic Storage or Usage Permission of the Publisher is required to store or use electronically any material contained in this work, including any chapter or part of a chapter. Except as outlined above, no part of this work may be reproduced, stored in a retrieval system or transmitted in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, without prior written permission of the Publisher. Address permissions requests to: Elsevier Science Rights & Permissions Department, at the mail, fax and e-mail addresses noted above. Notice No responsibility is assumed by the Publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made. First edition 2000 Library of Congress Cataloging in Publication Data Functional neuroanatomy of the nitric oxide system / editors, H.W.M. Steinbusch... [et al.].- 1st ed. p. ; cm - (Handbook of chemical neuroanatomy ; v. 17) Includes index. ISBN 0-444-50285-8 (alk. paper) 1. Nitric oxide. 2. Neurochemistry. 3. Neuroanatomy. I. Steinbusch, Hendrik Wilhelm Maria, 1950- II. Series. [DNLM: 1. Nitric-Oxide Synthase-physiology. 2. Nervous System Physiology. 3. Nitric Oxide-physiology. 4. Signal Transduction. QV 126 F979 2000] QP535.N1 F86 2000 612.8'042-dc21 00-028784

I S B N 0-444-50285-8 (volume) ISBN: 0-444-90340-2 (series) The paper used in this publication meets the requirements of A N S I / N I S O Z39.48-1992 (Permanence of Paper). Printed in The Netherlands

List of Contributors J. AIJON Dpto. Biologfa Celular y Patologfa Universidad de Salamanca Avda. Alfonso X el Sabio 1 37007 Salamanca Spain

T.EC. BATTEN Institute of Cardiovascular Research University of Leeds The Worsley Building Leeds LS2 9JT UK

J.R. ALONSO Dpto. Biologfa Celular y Patologfa Universidad de Salamanca Avda. Alfonso X el Sabio 1 37007 Salamanca Spain

J.G. BRINON Dpto. Biolog/a Celular y Patologfa Universidad de Salamanca Avda. Alfonso X el Sabio 1 37007 Salamanca Spain

S. AMIR Center for Studies in Behavioral Neurobiology Concordia University 1455 de Maisonneuve Boulevard West Montreal, PQ H3G 1M8 Canada

M.S. DAVIDOFF Institute of Anatomy University of Hamburg Martinistr. 52 D-20246 Hamburg Germany

C.R. ANDERSON Department of Anatomy and Cell Biology University of Melbourne Parkville, VIC 3052 Australia R. ARI~VALO Dpto. Biologfa Celular y Patologfa Universidad de Salamanca Avda. Alfonso X el Sabio 1 37007 Salamanca Spain L. ATKINSON School of Biomedical Sciences University of Leeds The Worsley Building Leeds LS 2 9NQ UK

J. DEUCHARS School of Biomedical Sciences University of Leeds The Worsley Building Leeds LS 2 9 NQ UK J. DE VENTE European Graduate School of Neuroscience (EURON) Department of Psychiatry and Neuropsychology Maastricht University PO Box 616 6200 MD Maastricht The Netherlands

W.D. ELDRED Department of Biology Laboratory of Visual Neurobiology Boston University 5 Cummington Street Boston, MA 02215 USA J.B. FURNESS Department of Anatomy and Cell Biology University of Melbourne Parkville, VIC 3052 Australia L.J. IGNARRO Department of Molecular and Medical Pharmacology UCLA School of Medicine CHS 23-120, Box 951785 Los Angeles, CA 90095-1735 USA A. JACOBS Department of Molecular and Medical Pharmacology UCLA School of Medicine CHS 23-120, Box 951785 Los Angeles, CA 90095-1735 USA R. MIDDENDORFF Institute of Anatomy University of Hamburg Martinistr. 52 D-20246 Hamburg Germany A. PORTEROS Dpto. Biologfa Celular y Patologfa Universidad de Salamanca Avda. Alfonso X el Sabio 1 37007 Salamanca Spain

vi

N.L. SCHOLZ Northwest Fisheries Science Center Seattle, WA 98112 USA H.W.M. STEINBUSCH European Graduate School of Neuroscience (EURON) Department of Psychiatry and Neuropsychology Maastricht University PO Box 616 6200 MD Maastricht The Netherlands J.W. TRUMAN Department of Zoology University of Washington Box 351800 Seattle, WA 98195-1800 USA S.R. VINCENT Graduate Program in Neuroscience Kinsmen Laboratory of Neurological Research Department of Psychiatry University of British Columbia Vancouver, BC V6T 1Z3 Canada E. WERUAGA Dpto. Biolog/a Celular y Patologfa Universidad de Salamanca Avda. Alfonso X el Sabio 1 37007 Salamanca Spain B. WOODSIDE Center for Studies in BehaVioral Neurobiology Concordia University 1455 de Maisonneuve Boulevard West Montreal, PQ H3G 1M8 Canada

W. WU Department of Anatomy University of Hong Kong Faculty of Medicine 5 Sassoon Road Hong Kong People's Republic of China

H.M. YOUNG Department of Anatomy and Cell Biology University of Melbourne Parkville, VIC 3052 Australia

vii

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Preface This volume concerns the neuronal isoform of nitric oxide synthase (nNOS, bNOS or NOS I). Only in the first chapter, the endothelial isoform and the inducible form are briefly mentioned in an attempt to provide a more complete picture. In agreement with the goal of the Handbook Series, the focus of this volume is on the localization of NOS in the nervous system where today extensive information is present from studies on the brain, spinal cord and peripheral nervous system. This is to a considerable extent a consequence of the important discovery that NADPH-diaphorase activity is a part of NOS, which made it possible to use an earlier established histochemical staining method to visualize NOS systems. With this method brilliant staining patterns can be achieved in an easy and relatively cheap way. There are, however, certain caveats in the use of this NADPH-diaphorase method, which are discussed in more or less detail in several chapters of this volume. Nevertheless, this enzyme histochemical staining method remains a valuable and preferred tool for many researchers involved in NOS neuroanatomy. We have tried to bring together a number of reviews on different aspects of NOS, ranging from detailed descriptions of the nNOS distribution to analysis of NO signaling with the aim to provide a first major reference guide to the localization of NOS in different species. In addition we wanted to present an overview of the co-localization of other (co)transmitters with NOS. A more functional approach to NO can be achieved by the combined visualization of the localization of NOS and activated soluble guanylyl cyclase as a target for NO. Although many more direct effects of NO on a number of molecular targets, e.g. ion channels and NMDA receptors, have been reported in the literature, it remains to be demonstrated that these effects are relevant to the physiological function of NO in the nervous system. NOS is present in virtually every area throughout the central and peripheral nervous system. Over the last decade NO has emerged as an important regulator of many brain processes, and its physico-chemical properties and synthetic and release characteristics make NO an unconventional messenger molecule. It is synthesized upon demand and can diffuse over considerable distances in spite of its short half-life. The fact that NO is a free radical makes it potentially harmful to the living organism. Therefore, the expression of the genes for the different NOS isoforms as well as the metabolism of NO in the nervous system have to be rigidly controlled. This potential neurotoxic role may have been accentuated in the first decade of NO research at the expense of its physiological signaling function. It is our hope that this volume will help to further stimulate research on NO as a messenger molecule in the nervous system, which is one of the aims with the publication of this volume of the Handbook Series. We would like to express our gratitude to the authors of this volume who have taken the time and made the effort to produce such excellent chapters, as well as to the highly competent and cooperative staff at Elsevier. Maastricht, Vancouver, Lund and Stockholm, January 2000 HARRY W.M. STEINBUSCH ANDERS BJORKLUND

JAN DE VENTE

STEVEN R. VINCENT TOMAS HOKFELT ix

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Contents List of Contributors Preface

v

ix

NITRIC OXIDE SYNTHASE AND THE PRODUCTION OF NITRIC O X I D E L.J. IGNARRO AND A. JACOBS 1. 2. 3. 4. 5. II.

1 3 6 10 11

HISTOCHEMISTRY OF NITRIC OXIDE SYNTHASE IN THE CENTRAL NERVOUS SYSTEM- S.R. VINCENT 1. 2.

3. 4. 5. 6. III.

Introduction and historical perspectives NO synthase isoforms Protein structure of NO synthase isoforms Neuronal NO synthase References

Introduction Localization of nitric oxide synthase in the brain 2.1. The olfactory bulb 2.2. The cerebral cortex 2.3. The hippocampal formation 2.4. The basal forebrain 2.5. The basal ganglia 2.6. The thalamus 2.7. The superior colliculus 2.8. The auditory system 2.9. The hypothalamus 2.10. Circumventricular organs 2.11. The mesopontine tegmentum 2.12. The cerebellum 2.13. The medulla 2.14. The spinal cord Arginine metabolism in the brain Heme oxygenase Conclusions References

19 21 21 22 24 26 27 28 29 31 31 33 33 36 36 37 38 39 39 39

COMPARATIVE AND DEVELOPMENTAL NEUROANATOMICAL ASPECTS OF THE NO SYSTEM- J.R. ALONSO, g. ARI~VALO, E. WERUAGA, A. PORTEROS, J.G. BgIlqON AND J. AIJON 1. 2.

Introduction Methods to localize nitric oxide and nitric oxide synthases in brain tissue 2.1. Histological detection of NADPH-diaphorase/nitric oxide synthase

51 52 52 xi

3.

4.

5. 6. IV.

53 53 54 54 54 55 55 55 59 63 67 71 76 87 87 89 91 92 94 101

NITRIC OXIDE IN THE RETINA - W.D. ELDRED 1. 2.

3. 4.

5.

xii

2.1.1. NADPH-diaphorase histochemistry 2.1.2. Immunohistochemistry 2.1.3. In situ hybridization 2.2. Biochemical detection of nitric oxide synthase, nitric oxide production, and nitric oxide proper Results 3.1. Fish 3.1.1. Cyclostomes 3.1.2. Teleosts 3.2. Amphibians 3.3. Reptiles 3.4. Birds 3.5. Mammals 3.6. General pattern of NADPH-diaphorase expression during rat brain development Discussion 4.1. Methodological aspects 4.2. Interspecies differences in the NADPH-diaphorase distribution pattern 4.3. Implications of the NADPH-diaphorase expression during the developmental processes 4.4. NADPH-diaphorase/nitric oxide synthase distribution pattern and nitric oxide functional implications Abbreviations References

Introduction Localization of nitric oxide synthase in the retina 2.1. Methodological considerations 2.1.1. NADPH-diaphorase histochemistry 2.1.2. NOS immunocytochemistry 2.2. Isoforms of NOS in the retina 2.3. Anatomical localizations of NOS 2.3.1. Mammals 2.3.2. Lower vertebrates 2.4. Regional distribution of NOS 2.5. Efferents Development of NOS in the retina and central visual targets Biochemistry and molecular biology of NOS in the retina 4.1. NOS 4.2. Photoreceptors Function of NO in specific retinal cell types 5.1. Photoreceptors 5.2. Horizontal cells 5.3. Bipolar cells 5.4. Amacrine cells 5.5. Ganglion cells

111 111 112 112 113 113 114 114 117 120 122 122 125 125 126 128 128 129 131 131 132

6. Release of NO in the retina 7. Modulation of transmitter release by NO 8. The cGMP signal transduction pathway in retina and its modulation by NO 9. Future areas of investigation 10. Abbreviations 11. Acknowledgements 12. References V.

I32 133 133 138

I39 140 I40

NITRIC OXIDE SIGNALING IN THE HYPOTHALAMUS - B. WOODSIDE AND S, AMIR Introduction 147 NOS in the hypothalamus 147 Co-localization 149 Regulation 151 Functional considerations 157 5.1. Stress axis 157 5.2. Magnocellular neurosecretory system 159 5.2.1. Modulation of vasopressin and oxytocin release 159 5.2.2. Drinking 161 5.2.3. Reproductive behavior 161 5.3. The reproductive axis 161 5.3. I . Modulation of luteinizing hormone-releasing hormone secre161 tion 5.3.2. Modulation of prolactin release 163 5.3.3. Sexual behavior 163 5.4. Somatostatin release 163 5.5. Circadian regulation 1 64 5.6. Autonomic regulation 164 5.7. Plasticity 165 6. Conclusions 166 7. Acknowledgements 167 8. References 167

1. 2. 3. 4. 5.

VI.

NITRIC OXIDE SYSTEMS IN THE MEDULLA OBLONGATA AND THEIR INVOLVEMENT IN AUTONOMIC CONTROL - T.F.C. BATTEN, L. ATKINSON AND J. DEUCHARS 1. Introduction 2. Methods 3. Results and discussion 3.1. Anatomy of NOS in the medulla oblongata 3. I . 1. Description of the location and morphological characteristics of NOS-IR neurones in the rat medulla oblongata 3.1.2. Relative distribution of NOS-IR and NADPH-diaphorase activity 3.1.3. Co-localisation of NOS-IR or NADPH-diaphorase activity with other neuronal phenotypes in the medulla oblongata

177 177 180 180 180 187 188 ...

Xlll

3.2.

,

5. 6.

Role of nitric oxide in medullary pathways involved in autonomic functions 3.2.1. NO in the control of the cardiovascular system 3.2.2. NO in neuronal circuitry underlying control of the oesophagus 3.2.3. NO in CNS control of the stomach and large intestine 3.2.4. NO in central respiratory control 3.2.5. NO in pathways coordinating autonomic and nociceptive responses Abbreviations Acknowledgements References

199 199 204 204 205 206 207 208 208

VII. NITRIC OXIDE IN THE PERIPHERAL AUTONOMIC NERVOUS SYSTEMH.M. YOUNG, C.R. ANDERSON AND J.B. FURNESS Introduction 1.1. Brief history of the identification of NO as a peripheral neurotransmitter 1.2. General properties of NO-mediated neurotransmission 1.3. Scope of this review 2. NO in autonomic ganglia 2.1. Nitric oxide and sympathetic pre- and postganglionic neurones 2.1.1. Presence of NOS in sympathetic pre- and postganglionic neurones 2.1.2. Functionally identified subclasses of sympathetic preganglionic NOS neurones 2.1.3. Presence of NOS in parasympathetic pre- and postganglionic neurones 2.2. Nitric oxide and ganglionic transmission 3. Role of NO in the neural control of the vasculature and the heart 3.1. Role of neurally derived NO in the control of the vasculature 3.1.1. Autonomic vasodilator neurones 3.1.2. Sensory vasodilation 3.1.3. Role of neurally released NO in regulation of blood vessels summary 3.2. Role of neurally derived NO in the control of the heart 4. Role of NO in the neural control of the gastrointestinal tract 4.1. Introduction 4.2. NO is a neurotransmitter of enteric inhibitory motor neurones 4.3. Role of NO in co-transmission from enteric inhibitory motor neurones 4.4. Axo-axonal interactions involving NO 4.5. NOS interneurones within the intestine 4.6. NOS in nerve fibres innervating oesophageal motor endplates 4.7. NOS innervation of the mucosa in the stomach 4.8. NOS in intestinofugal neurones 4.9. NOS in the biliary system and pancreas 5. Role of NO in the neural control of the trachea and lower airways 5.1. Types of neurones innervating the trachea and lower airways xiv

215 215 216 216 217 217 217 219 221 223 224 224 224 228 228 229 232 232 232 233 237 237 238 238 238 239 240 240

5.2.

Pharmacology of transmission and source of NOS neurones in the trachea Role of NO in the neural control of salivary glands and other secretory tissues 6.1. Salivary glands 6.1.1. Types of neurones innervating the salivary glands 6.1.2. Role of NO in vasodilation and secretion in the salivary glands 6.2. Sweat glands 6.3. The nasal mucosa 7. Role of NO in the innervation of the adrenal medulla 7.1. Presence of NOS in neurones in the adrenal medulla 7.2. Role of neurally derived NO in the adrenal medulla 8. Overview of peripheral autonomic NO neurones 9. Acknowledgements 10. References .

241 242 242 242 242 244 244 245 245 246 248 249 249

VIII. THE NITRIC OXIDE SYSTEM IN THE UROGENITAL TRACTM.S. DAVIDOFF AND R. MIDDENDORFF 1.

2.

3. 4. IX.

The urinary tract 1.1. The upper urinary tract 1.1.1. The kidney 1.1.2. The renal pelvis 1.1.3. The ureter 1.2. The lower urinary tract 1.2.1. The urinary bladder 1.2.2. The urethra The genital tract 2.1. The male reproductive organs 2.1.1. The testis 2.1.2. The spermatozoa 2.1.3. The epididymis 2.1.4. The vas deferens 2.1.5. The seminal vesicle 2.1.6. The prostate 2.1.7. The penis 2.2. The female genital tract 2.2.1. The ovary 2.2.2. The Fallopian tube 2.2.3. The uterus 2.2.4. The vagina 2.2.5. The placenta and umbilical artery Acknowledgements References

267 267 267 277 277 279 279 281 282 282 282 286 288 290 290 291 291 292 292 294 294 297 297 301 301

RESPONSE OF NITRIC OXIDE SYNTHASE TO NEURONAL INJURY - W. WU 1. 2.

Introduction Injury-induced expression of NOS

315 316 XV

2.1.

Up-regulation of NOS expression in injured neurons 317 2.1.1. Up-regulation of NOS expression in the neurohypophyseal systern 317 2.1.2. Up-regulation of NOS expression in the nuclei of cranial nerves 317 2.1.3. Up-regulation of NOS expression in the spinal cord 317 2.1.4. Up-regulation of NOS expression in the peripheral nerve and ganglia 319 321 2.2. De novo expression of NOS in injured neurons 2.2.1. De novo expression of NOS in the cerebral cortex 321 2.2.2. De novo expression of NOS in the cerebellar cortex 321 2.2.3. De novo expression of NOS in the nuclei of cranial nerves 321 2.2.4. De novo expression of NOS in spinal motoneurons 324 2.2.5. De novo expression of NOS in nuclei associated with the long descending and ascending pathways 324 325 2.3. Time course of NOS expression in injured neurons 2.3.1. Time course of NOS expression is different in different populations of neurons 325 2.3.2. Time course of NOS expression can be different in the same population of neurons following different types of injuries 326 3. Co-expression of NOS with other injury-relevant components 326 4. Age-related expression of NOS in injured neurons 328 5. Species-related expression of NOS in injured neurons 329 6. Ultrastructure of injury-induced NOS-positive neurons 330 7. Regulation of NOS expression in injured neurons 334 7.1. Regulation of injury-induced NOS by the length of the remaining proximal axons following axotomy 334 7.2. Regulation of injury-induced NOS by peripheral nerve (PN) graft transplantation 336 7.3. Regulation of injury-induced NOS by neurotrophic factors 338 8. Mechanisms of NOS expression in injured neurons 338 8.1. Expression of injury-induced NOS in neurons is a general response to injury 339 8.2. Expression of injury-induced NOS results from the interruption of axonal transport 340 8.3. Expression of injury-induced NOS results from stimulation of NMDA receptors 340 8.4. Expression of injury-induced NOS results from deprivation of neurotrophic factors 341 Potential roles of NOS expression in neuronal degeneration and regeneration 341 342 9.1. Potential role of NOS in neuronal degeneration 342 9.1.1. Involvement of NOS in neural degeneration 343 9.1.2. Potential mechanisms of NO/NOS-mediated neurotoxicity 9.1.3. Evidence suggesting that NO/NOS is not involved in degener344 ative processes after neuronal injury 345 9.2. Potential role of NOS in neuronal regeneration 346 10. Summary 346 11. Acknowledgements 347 12. References ,

xvi

Xo

NITRIC OXIDE-cGMP SIGNALING IN THE RAT BRAIN - J. DE VENTE AND H.W.M. STEINBUSCH 1. 2. 3. 4. 5.

Introduction 355 Soluble guanylyl cyclase 356 Phosphodiesterase activity and the termination of the NO-cGMP signal 357 cGMP immunocytochemistry 358 Immunocytochemical localization of NOS and NO-mediated cGMP synthesis 359 5.1. A note on the use of brain slices 359 5.2. A note on the use of phosphodiesterase inhibitors in in vitro studies 360 5.3. NOS activity in brain slices 361 5.4. Localization of NO-mediated cGMP accumulation in brain slices 362 5.4.1. Telencephalon 362 5.4.2. Diencephalon 365 5.4.3. Mesencephalon 367 5.4.4. Cerebellum 367 5.4.5. Pons and medulla oblongata 372 5.5. Colocalization of cGMP and NOS 378 6. Localization of cGMP in the cerebellum 384 7. NO-cGMP signaling in other vertebrates 389 8. NO-cGMP signaling in the rat brain during development 390 9. Colocalization of cGMP with neurotransmitter systems in the rat brain 395 10. Abbreviations 405 11. Acknowledgements 405 12. References 406

XI.

INVERTEBRATE MODELS FOR STUDYING NO-MEDIATED SIGNALINGN. SCHOLZ AND J.W. TRUMAN 1. 2. 3.

4. 5.

6. 7. 8. 9.

Introduction Components of the NO signaling pathway in invertebrates Developmental roles of NO/cGMP signaling for invertebrates 3.1. NO signaling and proliferation control 3.2. NO sensitivity during neuronal development 3.3. Functional significance of NO/cGMP signaling The role of NO in feeding behavior The role of NO in regulating rhythmic motor networks 5.1. The gastropod feeding circuit 5.2. The crustacean stomatogastric ganglion 5.3. The crustacean cardiac network NO and nociception NO as a potential blood-borne neurohormone Acknowledgements References

Subject Index

417 418 419 419 420 427 428 429 429 430 433 435 436 438 438 443

xvii

CHAPTER I

Nitric oxide synthase and the production of nitric oxide L.J. IGNARRO AND A. JACOBS

1. INTRODUCTION AND HISTORICAL PERSPECTIVES Not too long ago, nitric oxide (NO) was viewed by many as a noxious component of polluted air over industrial cities or as a gas that could be purchased in gas cylinders for the purpose of conducting chemical experiments. The possible biological importance of NO emerged in the 1970s, when a variety of studies from different laboratories revealed that nitroso compounds and related chemical species that were suspected of decomposition or conversion to NO could stimulate the production of cyclic GMP in mammalian tissues by activating the cytosolic isoform of guanylate cyclase (DeRubertis and Craven, 1976; Arnold et al., 1977; Katsuki and Murad, 1977; Katsuki et al., 1977; Miki et al., 1977: Schultz et al., 1977; B6hme et al., 1978; Craven and DeRubertis, 1978; Murad et al., 1978; Axelsson et al., 1979; Craven et al., 1979; Kukovetz et al., 1979; Ignarro, 1989). The well known chemical effects of some of these nitro compounds, such as nitroprusside and nitroglycerin, led us to suspect that their conversion to NO might account for the mechanism of vasodilation elicited by nitroprusside, and organic nitrate esters and organic nitrite esters. Accordingly, NO was tested and found to be a potent vascular smooth muscle relaxant which caused vasorelaxation via the second messenger actions of cyclic GMP (Gruetter et al., 1979, 1980, 1981; Napoli et al., 1980). These observations were confirmed and extended by several laboratories (Ignarro et al., 1981, 1984; Axelsson et al., 1982; Keith et al., 1982; Axelsson and Andersson, 1983; Galvas and DiSalvo, 1983; Horowitz et al., 1983; Ignarro and Kadowitz, 1985; Ignarro, 1989). The discovery that NO itself is a potent relaxant helped to explain the many earlier observations that diverse nitro and nitroso compounds caused both vasorelaxation and tissue accumulation of cyclic GME The clinical awareness that certain nitrovasodilators also interfered with platelet aggregation led us to ascertain whether NO and cyclic GMP were responsible for such anti-platelet actions. NO and a series of S-nitrosothiols, which are used as NO donor agents, were found to inhibit human platelet aggregation and by mechanisms involving the second messenger actions of cyclic GMP (Mellion et al., 1981, 1983). Inhibition of platelet aggregation occurred regardless of the agent used to promote aggregation, including ADP, collagen, thrombin, or thromboxane analogs. It is of historic interest, perhaps, that the first two pharmacological actions described for NO were vasorelaxation and inhibition of platelet aggregation, now appreciated to be the two most important actions of endothelium-derived relaxing factor (EDRF). In the process of elucidating the mechanism by which nitroglycerin causes vascular smooth muscle relaxation, we found that sulfhydryl (-SH) containing compounds or thiols Handbook of Chemical Neuroanatom~; Vol. 17: Functional Neuroanatomv of the Nitric Oxide System H.W.M. Steinbusch, J. De Vente and S.R. Vincent, editors (~ 2000 Elsevier Science B.V. All rights reserved.

Ch. I

L.J. Ignarro and A. Jacobs

(cysteine, dithiothreitol) were required for the activation of cytosolic guanylate cyclase by nitroglycerin, other organic nitrate esters and some organic nitrite esters (Ignarro and Gruetter, 1980; Ignarro et al., 1980a,b). The reason for this thiol requirement was the liberation of NO from intermediate S-nitrosothiols formed, as a result of a chemical reaction between the nitrovasodilator and thiol (Ignarro et al., 1981). These studies constituted the first demonstration of the pharmacological actions of S-nitrosothiols and their characteristic property as NO donor agents. Two of these S-nitrosothiols have become widely used NO donor agents, namely S-nitroso-N-acetylpenicillamine (SNAP) and S-nitrosoglutathione (GSNO). The extraordinary high potency of nitroglycerin as a vasodilator and the finding that NO is responsible for its pharmacological effects suggested that mammals might possess an endogenous nitroglycerin, another NO donor species, or NO itself. This hypothesis was confirmed when the EDRF, discovered in 1980 (Furchgott and Zawadzki, 1980), was identified as NO in 1986-1987 (Ignarro et al., 1986a, 1987a,b, Palmer et al., 1987). This observation provided explanations for previous findings that EDRF activates guanylate cyclase (Frrstermann et al., 1986; Ignarro et al., 1986b) and inhibits platelet aggregation (Azuma et al., 1986). Subsequent experiments revealed the mechanism by which EDRF inhibits platelet aggregation and adhesion compared with the anti-platelet effect of prostacyclin (Radomski et al., 1987). Most of the early biological research on NO focused on its effects on vascular smooth muscle, platelets and cytosolic guanylate cyclase, whereas little or no attention was given to the possible influence of NO on neuronal cell function or as a neurotransmitter. Early studies revealed that certain regions of the brain were rich in cyclic GMP and cytosolic guanylate cyclase (Garthwaite, 1990). Not only NO (Miki et al., 1977) but also an unidentified low molecular weight factor from rat forebrain (Deguchi, 1977) were shown to activate guanylate cyclase, and enzyme activation was inhibited by hemoglobin. This soluble endogenous activator of guanylate cyclase was later identified as arginine (Deguchi and Yoshioka, 1982), although we had observed that arginine alone could not directly activate purified preparations of guanylate cyclase (unpublished observations). In retrospect, these original observations made by the Deguchi laboratory were well ahead of their time. We now understand that in crude soluble fractions from certain brain regions, arginine serves as the substrate for neuronal NO synthase to generate NO, which was likely responsible for the activation of guanylate cyclase in the experiments of Deguchi. This interpretation also explains the earlier finding by Ferrendelli (Ferrendelli et al., 1974) that glutamate caused a calcium-dependent stimulation of cyclic GMP accumulation in cerebellar slices. Now the stage was set for the pioneer studies of Garthwaite in 1988 that NMDA stimulated cyclic GMP accumulation in rat cerebellar cells (Garthwaite et al., 1988). This response to NMDA was associated with the release of an EDRF-like factor, which in turn acted on distinct target cells to elevate cyclic GMP levels. The EDRF-like factor was quickly shown to be NO (Moncada et al., 1991 ). Following the important observation that vascular endothelial cells could synthesize NO and citrulline from arginine by a process requiring NADPH (see Moncada et al., 1991), similar observations were made in brain tissue and the arginine-to-NO pathway was identified and elucidated as the NO synthase enzymatic pathway (Bredt and Snyder, 1989, 1990; Bredt et al., 1990). These pioneering studies showing the calcium- and calmodulin-dependent catalytic activity of NO synthase were performed with enzyme purified from rat cerebellum. The enzyme was termed neuronal NO synthase (nNOS). The discovery and characterization of the other two isoforms of NOS came later. Soon after the identification of nNOS in the brain~ many investigators began to elucidate the possible physiological and pathophysiological roles of endogenous NO in the brain. Since cellular and subcellular localizations of neurotransmitters

Neuronal NO synthase

Ch. I

often help to suggest and clarify biological function, antibodies to nNOS were developed and the tissue distribution and localization of nNOS in brain neurons as well as in peripheral neurons were demonstrated (Bredt and Snyder, 1994). Unfortunately, specific definitive roles for nNOS or NO in the brain are not well established and this is an area of extensive investigation (Bredt and Snyder, 1994). NO has been implicated in long-term potentiation in the hippocampus where it may function as a retrograde messenger in memory and learning processes. Direct evidence for specific neurotransmitter actions of NO derives from research conducted in the peripheral nervous system. In the central nervous system, although NO appears to function in synaptic transmission, the neuronal release of excess quantities of NO is associated with neurotoxicity. For example, excess glutamate release stimulates NMDA receptors and consequent NO production, which mediates the focal schemia of vascular stroke (Choi, 1988) and perhaps other CNS disorders (Bredt and Snyder, 1994). In the peripheral nervous system, NO appears to function as a neurotransmitter mediating vascular and nonvascular smooth muscle relaxation. These tissues are innervated by nonadrenergic-noncholinergic (NANC) neurons, nNOS containing neurons exist in the myenteric (Auerbach's) plexus of the gastrointestinal tract (Bredt et al., 1990; Dawson et al., 1991) and neuronal stimulation causes NO release and nonvascular smooth muscle relaxation (Bult et al., 1990; Boeckxstaens et al., 1991; Desai et al., 1991 ; Tottrup et al., 1991) associated with inhibition of peristalsis. NANC neurons exist also in the outer adventitial layers of large blood vessels, and in arteries within erectile tissue (Burnett et al., 1992). Functional, constitutive nNOS was recovered from rabbit corpus cavernosum (Bush et al., 1992a) and later localized to neurons innervating the erectile tissue (Burnett et al., 1992). In the erectile tissue (corpus cavernosum) of the penis, NO is the NANC neurotransmitter that mediates penile erection by provoking both vascular and nonvascular smooth muscle relaxation, thereby allowing the trabecular and sinusoidal vascular beds to become engorged with blood (Ignarro et al., 1990; Bush et al., 1992a,b,c; Rajfer et al., 1992; Trigo-Rocha et al., 1993).

2. NO SYNTHASE ISOFORMS After the discovery of the nNOS isoform, the inducible isoform of NOS (iNOS) was found. Early studies by Hibbs and colleagues revealed that activated rodent macrophages killed target tumor cells by arginine-dependent mechanisms, and could be inhibited by chemical analogs of arginine such as NG-methylarginine (Hibbs et al., 1987a). Other studies showed that murine macrophages activated by lipopolysaccharide or other agents could produce nitrite and nitrate, and that these latter products might be responsible for the resulting target cell cytotoxicity (Stuehr and Marietta, 1985, 1987a,b; Iyengar et al., 1987; Miwa et al., 1987). Also, arginine was identified as the precursor for nitrite and nitrate in these latter studies (Iyengar et al., 1987). None of these investigators suggested that NO was actually generated first, as the intermediate product of arginine conversion, and subsequently oxidized to nitrite and nitrate, as is now known to occur. Indeed, the first proposed enzymatic pathway for this conversion of arginine to nitrite was via arginine deiminase, which produces citrulline plus ammonia, followed by oxidation of ammonia to nitrite (Hibbs et al., 1987b). The first clue that the conversion of arginine to nitrite and nitrate might involve the intermediate formation of NO came from experiments conducted with activated rat neutrophils, which generated a vascular smooth muscle relaxing factor with the properties of NO (Rimele et al., 1988; Sturm et al., 1988). The definitive observations that arginine can be converted to NO plus

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citrulline came from experiments utilizing vascular endothelial cells (Palmer et al., 1988). It turns out, however, that the endothelial cells contain a constitutive isoform of NOS, whereas the activated macrophages and neutrophils contain a distinct inducible isoform of NOS. Research conducted in other laboratories in the early 1980s revealed that the production of nitrate in mammals could occur by pathways that were independent of intestinal microbial metabolism, and present endogenously in the animal (Green et al., 1981). Administration of lipopolysaccharide to rats led to increased urinary output of nitrate (Wagner et al., 1983). The mechanism of this effect was not understood until Stuehr and Marletta (1985) showed that lipopolysaccharide stimulated nitrate production in mice, and that this effect was probably mediated by macrophages, as lipopolysaccharide-activated macrophages were capable of generating nitrite and nitrate. Additional experiments showed that both nitrite and nitrate were derived from a common precursor, arginine (Iyengar et al., 1987). Therefore, research from several independent laboratories showed that lipopolysaccharide-treated animals and lipopolysaccharide-activated macrophages and other cell types could synthesize nitrite and nitrate from arginine. Following the demonstration that NO could be synthesized from arginine by vascular endothelial cells (Palmer et al., 1988), it became apparent that NO was likely an intermediate in the production of nitrite and nitrate from arginine by activated macrophages. These findings led to subsequent studies confirming that activated macrophages synthesized NO from arginine and that the NO was subsequently oxidized to nitrite and nitrate (Hibbs et al., 1988; Marletta et al., 1988; Stuehr et al., 1989). After the initial discovery and purification of nNOS (Bredt and Snyder, 1994), the inducible iNOS and endothelial NOS (eNOS) isoforms were characterized (F6rstermann et al., 1994; Griffith and Stuehr, 1995). Two distinct nomenclatures have been used to designate the three isoforms of NOS. One nomenclature refers to the NOS isoforms according to principal tissue distribution and the inducibility of one isoform, and these isoforms are designated as nNOS, eNOS and iNOS. Some of the cited problems with this nomenclature are that the so-called constitutive isoforms (nNOS and eNOS) can be expressionally regulated (upregulation and downregulation), and that certain isoforms can be found in locations other than those of their principal designation. This led some to propose the use of type I (nNOS), type II (iNOS) and type III (eNOS), according to the chronological order of their isolation and purification. The main problem with this latter nomenclature is the non-descriptive nature of the terms. In general, the NOS isoforms catalyze the oxidation or oxygenation of arginine to NO plus citrulline (Fig. 1). One atom of dioxygen (molecular oxygen) is incorporated into NO and a second atom of dioxygen is incorporated into citrulline (Kwon et al., 1990; Leone et al., 1991). One mole of L-arginine yields one mole of NO plus one mole of L-citrulline (Taych and Marietta, 1989; Bush et al., 1992d). This is a cytochrome P450-1ike oxidation reaction and requires NADPH as the principal electron donor. Like cytochrome P450, NOS contains heine as a prosthetic group for oxygen binding and subsequent incorporation into substrate to yield the products of the reaction (Griffith and Stuehr, 1995). An important intermediate in this enzymatic reaction is N~-hydroxyarginine (Stuehr et al., 1991; Wallace and Fukuto, 1991), which elicits pharmacological and perhaps physiological effects of its own (Buga et al., 1996). In addition to being a hemoprotein, NOS is a flavoprotein and requires both FAD (flavin adenine dinucleotide) and FMN (flavin mononucleotide) for the full expression of catalytic activity (Stuehr et al., 1991; Griffith and Stuehr, 1995). The enzyme-bound flavins function to transfer the electrons from the NADPH to the heine prosthetic group, thereby keeping the heme iron in the reduced state (ferrous: Fe 2+) to facilitate oxygen binding. Thus, NOS is a flavo-hemoprotein, one of the very few known to occur in mammals. Calmodulin, a calcium-binding protein, is also required for NOS catalytic activity (Abu-Soud

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NH

0.5 NADPH 0.5 NADP§

I

I

OH Fig. 1. The conversion of L-arginine to L-citrulline plus nitric oxide (NO) is catalyzed by the enzyme NO synthase. This conversion involves an initial 2-electron oxidation of one of the two equivalent guanidino nitrogen atoms of L-arginine to form NG-hydroxy-L-arginine, which can dissociate from the enzyme. A second, 3-electron oxidation of NG-hydroxy-L-arginine forms L-citrulline plus NO. Two molecules of dioxygen and 1.5 equivalents of NADPH are consumed in the overall enzymatic reaction.

and Stuehr, 1993; Griffith and Stuehr, 1995). Calmodulin binds free calcium to form a complex which binds to. nNOS and eNOS and functions to facilitate the transfer of electrons from enzyme-bound FAD to the heine iron. Since this transfer of electrons is essential for the expression of catalytic activity, calmodulin, and therefore calcium, acts as an on/off switch for NO biosynthesis from arginine. The iNOS isoform contains calmodulin already bound tightly as a subunit and, therefore, does not require calcium for enzyme activation (Cho et al., 1992). An additional cofactor that appears to be required for full NOS catalytic activity is tetrahydrobiopterin (Gross and Levi, 1992; Stuehr and Griffith, 1992; Griffith and Stuehr, 1995). It was speculated initially by some that the pterin may participate in the redox chemistry of the catalytic cycle. It is now believed that tetrahydrobiopterin functions in stabilizing the NOS protein rather than participating in catalysis (Giovanelli et al., 1991), and later reports added to this view based on the findings that tetrahydrobiopterin prevented the direct negative feedback effect of NO on NOS catalytic activity (Rogers and Ignarro, 1992; Griscavage et al., 1993). A summary of some of the properties of the NOS isoforms is given in Table 1. The NOS isoforms appear to catalyze an atypical, odd-numbered electron (5-electron) oxidation of L-arginine to NO plus L-citrulline (Griffith and Stuehr, 1995). In the first step of the reaction, one of the two basic guanidino nitrogen atoms of arginine undergoes a 2electron oxidation to yield NG-hydroxyarginine (typical of cytochrome P450 monooxygenase chemistry), and in the second step of the reaction, NG-hydroxyarginine undergoes a 3-electron oxidation to yield NO and citrulline (atypical of cytochrome P450). The stoichiometry of the NOS catalytic cycle has been studied extensively and often debated, but the conclusion has been reached that if NO is indeed the immediate product of the NADPH-dependent oxidation of NG-hydroxyarginine, then a 5-electron oxidation of arginine must occur. A more typical, even-numbered, 4-electron oxidation of arginine would necessarily yield HNO (nitroxyl in the protonated state) instead of NO (Fukuto et al., 1992, 1993a). Interestingly, HNO is chemically unstable and is readily oxidized to NO by numerous physiologically important oxidants (Fukuto et al., 1993b). Indeed, the pharmacological effects of HNO are virtually identical to those for NO (Fukuto et al., 1993a), and this may be due to the rapid oxidation of HNO to NO. These observations prompted additional studies aimed at elucidating the chemical nature

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TABLE 1. Properties of representative NO svnthase isoforms Property

nNOS (I)

eNOS (III)

iNOS (II)

Principal source Subunit Mr (kDa) Subcellular localization Native structure Constitutively expressed Inducible Regulation ECso Ca2+ (nM) Km arginine (IxM) Vmax (gmol min-1 mg-1) Km NG-hydroxyarginine(gM) IC5o NG-methylarginine(gM) IC5o N~-nitroarginine (IxM)

Neuronal 160 Cytosolic Dimer Yes No Ca2.`./calmodulin 300 1-4 1-3 25 1.6 0.9

Endothelial 133 Membrane-bound Dimer Yes No Ca2§ 300 1-5 0.8-1 0.9 0.2

Macrophages 130 Cytosolic Dimer Yes Yes Protein synthesis Ca2+ not required 2-20 1-2 25 7.5 200

See Griffith and Stuehr (1995) and Hobbs and Ignarro (1997) for more details.

of the NOS products. Using a sensitive chemiluminescence technique that is selective for NO (relative to HNO and other oxides of nitrogen), evidence was obtained that HNO is an intermediate in the oxidation of arginine to NO by iNOS and nNOS (Hobbs et al., 1994; Hobbs and Ignarro, 1997). These studies suggested that NOS might catalyze a 4-electron oxidation of arginine to HNO, and the HNO would be readily and rapidly oxidized under physiological conditions to NO. These observations are more consistent with the chemistry of cytochrome P450-type monooxygenases and would solve the potential problem of having to deal with atypical 5-electron oxidation reactions of arginine to yield NO directly. These observations and conclusions have been confirmed by others (Schmidt et al., 1996).

3. PROTEIN STRUCTURE OF NO SYNTHASE ISOFORMS In 1991, Bredt and co-workers reported the cDNA sequence that codes rat cerebellum nNOS, which has a molecular mass of approximately 160 kDa (Bredt et al., 1991). There is good homology of the C-terminal portion of nNOS to the C-terminal portion of iNOS and eNOS, and all NOS isoforms show close homology to NADPH cytochrome P450 reductase (Bredt et al., 1991; Sessa, 1994; Xie et al., 1994; F6rstermann and Kleinert, 1995). NOS isoforms and NADPH cytochrome P450 reductase possess very similar characteristics including consensus sequences for NADPH, FMN and FAD binding sites. Many investigators have shown high homology among the individual NOS isoforms across species (Bredt and Snyder, 1994; Griffith and Stuehr, 1995). Table 2 gives information on the cDNAs encoding the three NOS isoforms (McMillan and Masters, 1993). Fig. 2 illustrates schematically the relationships among the cofactor sequences and other sequences for NOS isoforms and cytochrome P450 reductase. The observations of sequence homology between the C-terminal portion of NOS isoforms and cytochrome P450 reductase, together with the findings that NOS possesses heme that is required for catalysis, suggested that there must be a binding site for heine. In view of the fact that the heme for cytochrome P450 reductase actually resides within a distinct and separate protein (cytochrome P450), and that the NOS protein itself is capable of catalysis,

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TABLE 2. cDNAs encoding ttm three NO svnthase isq[olvns Isoform

nNOS (I) eNOS (III) iNOS (II)

cDNA source

Brain Endothelium Macrophage Smooth muscle Hepatocyte

Predicted protein Amino acids

Mr (kDa)

1429-1433 1203-1205 1144 1147 1147

160-161 133 130-131 131 131

mRNA (kb) 10-10.5 4.05-4.8 4 4 4.5

See F6rstermann and Kleinert (1995) for more details.

the hypothesis developed that the heme may bind at the N-terminal portion of NOS (Griffith and Stuehr, 1995). This would mean that NOS is composed of a distinct reductase domain (C-terminal) and a distinct oxygenase domain (N-terminal). Initial studies revealed this to be the case for nNOS, as limited trypsin proteolysis generated an N-terminal heme-containing fragment that binds L-arginine, and a C-terminal FMN- and FAD-containing fragment that catalyzes the NADPH-dependent reduction of cytochrome c (Sheta et al., 1994). These studies suggested that nNOS is a bi-domain enzyme in which the oxygenase and reductase domains can fold and function independently of one another. Similar observations were subsequently made with the iNOS isoform (Ghosh and Stuehr, I995). In a relatively short period of time since the initial purification of nNOS from cerebellum (Bredt and Snyder, 1990; Mayer et al., 1990), a great deal has been learned about structurefunction relationships for all three NOS isoforms. As discussed above, NOS binds both arginine and NADPH, and contains heme, tetrahydrobiopterin and flavins. The two constitutive isoforms (nNOS, eNOS) bind calmodulin, whereas the inducible isoform (iNOS) contains

H Fig. 2. Primary sequence map of the three isoforms of human nitric oxide synthase (NOS) and comparison to human NADPH-cytochrome P450 reductase. All NOS isoforms are flavo-hemoproteins which utilize NADPH as a substrate for reducing equivalents. Illustrated in the diagram are binding sites for the cofactors: NADPH, FAD, FMN, and Ca2+/calmodulin. Calmodulin is constitutively bound to the enzyme in the inducible NOS isoform. Not illustrated are the residues which bind tetrahydrobiopterin. The oxygenase domain of NOS contains a conserved cysteine residue which acts as an axial ligand for the heme iron. This contrasts with the cytochrome P450 isoforms, which contain a histidine ligand.

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tightly bound calmodulin. The heine is the site of oxygen activation for both sequential monooxygenase reactions catalyzed by NOS. The heine iron must be in the reduced or ferrous (Fe 2+) state to bind oxygen, and the electrons required to achieve iron reduction are derived from NADPH. The nNOS isoform is a bi-functional enzyme comprised of an N-terminal oxygenase domain and a C-terminal reductase domain. The N-terminal oxygenase domain contains binding sites for heme, tetrahydrobiopterin and L-arginine, whereas the C-terminal reductase domain contains binding sites for FMN, FAD and NADPH (Bredt et al., 1990; McMillan and Masters, 1993, 1995: Klatt et al.. 1994: Sheta et al., 1994; Nishimura et al., 1995). The binding of the flavins to NOS is structurally analogous to the binding of flavins to NADPH cytochrome P450 reductase and related proteins (Bredt and Snyder, 1994; Griffith and Stuehr, 1995). The flavins function to transfer NADPH-derived electrons either to an external heme protein acceptor (cytochrome P450) or an internal heme prosthetic group (NOS) (Gachhui et al., 1996). Calmodulin binding to nNOS increases the rate of electron transfer from NADPH to the flavins and triggers the interdomain transfer of electrons from the flavins to the heine iron, providing a basis by which the calcium-calmodulin complex activates nNOS and eNOS (Abu-Soud and Stuehr, 1993: Gachhui et al., 1996). This explains the requirement by nNOS and eNOS of calcium-facilitated calmodulin binding in order to stimulate NOS catalytic activity. Thus, nNOS and eNOS can participate in signal transduction pathways by generating NO only in response to increases in intracellular free calcium. Similarly, prolonged calcium influx, which is characteristic of reperfused or reoxygenated ischemic tissues, can lead to neuropathological conditions attributed to excess localized NO production (Garthwaite and Boulton, 1995). The high-output production of NO by iNOS is attributed to the sustained generation of relatively large quantities of NO by cells for many hours or days. This is explained by the capacity of iNOS to bind calmodulin as a subunit at ambient levels of intracellular free calcium (Cho et al., 1992: Stevens-Truss and Marietta, 1995; Ruan et al., 1996). The nNOS and eNOS isoforms contain a unique polypeptide insert in their FMN binding domains that is not present in the iNOS isoform (Salerno et al., 1997a). This polypeptide insert may function as an autoinhibitory domain by binding to endogenous peptides that could compete with calmodulin for binding sites on nNOS and eNOS but not iNOS. Further advances in this field could lead to the development of potent peptide inhibitors of nNOS or eNOS. The eNOS isoform resembles the other NOS isoforms in possessing similar heme ligation geometry (Salerno et al., 1997b). In each isoform, the heine is 5-coordinate and retains the axial thiolate ligand. Utilization of optical difference spectroscopy (McMillan and Masters, 1993) and EPR spectroscopy (Salerno et al., 1995) on all three NOS isoforms expressed in, and purified from, stably transfected human kidney embryonic cells or from Escherichia coli has allowed a comprehensive analysis of the interactions between isoforms and probes such as substrate, intermediate and substrate analog inhibitors (Salerno et al., 1997b). This work makes it possible to map the active sites of the three NOS isoforms via analysis of various substrate analogs and. therefore, to design more isoform-specific NOS inhibitors. The eNOS isoform is distinct from nNOS and iNOS in having a particulate or membrane-bound subcellular distribution (F6rstermann et al., 1991; Pollock et al., 1991; Busconi and Michel, 1993; Feron et al., 1996). The eNOS isoform appears to undergo a complex series of covalent modifications that influence its subcellular targeting. In endothelial cells, N-myristoylation and thiopalmitoylation of eNOS represent important determinants for its subcellular localization (Busconi and Michel, 1993: Liu and Sessa, 1994; Sase and Michel, 1997). Moreover, eNOS is specifically targeted to plasmalemmal caveolae as a consequence

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of palmitoylation of eNOS (Shaul et al., 1996). The subcellular translocation of eNOS suggests that the targeting of eNOS to endothelial cell caveolae may be dynamically regulated (Robinson and Michel, 1995; Robinson et al., 1995). Plasmalemmal caveolae are signal-transducing membrane microdomains consisting of cholesterol- and glycosphingolipid-enriched domains that may function to sequester diverse membrane-targeted signaling proteins (Anderson, 1993). Caveolin is an oligomeric integral membrane protein that appears to serve as a structural scaffold within caveolae, and can interact with and inhibit numerous structurally diverse signaling proteins (Li et al., 1995, 1996; Couet et al., 1997). It appears that the caveolin-scaffolding domain can specifically and potently inhibit eNOS catalytic activity and may function as a competitive inhibitor of the allosteric activation of eNOS by calmodulin (Garcia-Cardena et al., 1996; Michel et al., 1997a). More recent work using myristoylationand palmitoylation-deficient eNOS mutants suggests that the association between eNOS and caveolin is independent of the state of acylation of eNOS and that agonist-evoked, calmodulin-dependent disruption of the caveolin-NOS complex is what attenuates caveolin-mediated tonic inhibition of eNOS catalytic activity (Feron et al., 1998). Therefore, caveolin may play a role as an eNOS chaperone by regulating NO production independently of the presence of eNOS within caveolae or its state of acylation. Information on the crystal structure of the three NOS isoforms has recently become available (Crane et al., 1997, 1998; Raman et al., 1998). Crane et al. (1997) determined the crystal structure for iNOS, which revealed an unusual fold and heme environment for stabilization of activated oxygen intermediates that are key for enzyme catalysis. An active center created by an interface allows substrate and effector molecules such as tetrahydrobiopterin and calmodulin to modulate catalysis by altering the association between domains and subunits. NOS appears to be highly conducive to multiple regulatory pathways, and the authors believe that NO may have evolved to be a very effective signaling molecule in higher organisms. Additional studies by this group (Crane et al., 1998) revealed the dynamic structure of the iNOS dimer containing binding sites for arginine substrate and tetrahydrobiopterin cofactor at the catalytic center of the enzyme protein. The authors interpreted their findings to suggest that pterin binding causes a conformational change at the central interface region to expose the heine prosthetic group for interactions with the reductase domain. Pterin binding may also have electronic influences on heine-bound oxygen. L-Arginine appears to bind to glutamic acid-371 via hydrogen bonding and interacts with heine in a hydrophobic pocket to facilitate activation of heine-bound oxygen. This interaction may help to explain the two steps of oxidation of arginine, first to NG-hydroxyarginine and second to NO plus citrulline. The crystal structure of the constitutive isoform eNOS has been recently reported (Raman et al., 1998) and the conclusions drawn by these authors are considerably different from those drawn by Crane et al. for the inducible iNOS isoform (Crane et al., 1997, 1998). Raman et al. (1998) have addressed pterin function in NOS by determining the high-resolution crystal structure of the dimeric eNOS home domain. The protein structures reveal that pterin binding is not required for dimerization, as suggested earlier (Crane et al., 1997, 1998), but rather may be critical for enzyme catalysis. Moreover, Raman et al. (1998) argue that pterin binding fails to cause any conformational changes in the enzyme protein either at the pterin binding site or anywhere else in the protein. Another difference between these two groups is the finding by Raman et al. (1998) that the bottom region of the interface between monomers contains two conserved cysteine residues, rather than a disulfide bridge (Crane et al., 1997), that are involved in coordinating a zinc atom. The function of zinc may be stabilization of the pterin binding site and facilitation of stereospecific pterin recognition. The electropositive surface around the zinc metal center may serve as a docking site for the reductase domain of NOS.

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These observations may have direct implications for understanding the molecular basis of vascular dysfunction in pterin-deficient disorders.

4. NEURONAL NO SYNTHASE The biochemical properties of constitutive neuronal NOS (nNOS) are strikingly similar to those for constitutive eNOS and inducible iNOS. The crystal structure of the nNOS dimer should prove to be closely similar to the crystal structures of eNOS and iNOS. Like the other isoforms of NOS, nNOS is a modular enzyme which consists of a flavin-containing reductase domain and a heine-containing oxygenase domain linked by a chain of amino acids containing a calmodulin binding site. Calmodulin binding to nNOS facilitates the transfer of NADPH-derived electrons from the reductase domain to the oxygenase domain, thereby resulting in the oxygenation of arginine to yield NO plus citrulline. Like the other NOS isoforms, nNOS is capable of generating superoxide anion (O_;-) from oxygen when arginine availability is limited (Miller et al., 1997). It is not only interesting but also important to recognize that nearly all organs in the mammal are innervated by neurons that release NO as a neurotransmitter, which elicit both physiological and pathophysiological actions. The nNOS is distributed to both the cytosolic and particulate or membrane-bound fractions in most peripheral neurons. The membrane-bound nNOS may be attributed to the PDZ/GLGF motif found in the NH2-terminal sequence of the nNOS protein (Brenman et al., 1996a,b). nNOS is also distributed in skeletal muscle mainly to the particulate fraction, where it is attached to the sarcolemma-dystrophin complex via the PDZ/GLGF motif and interacts with (L-syntrophin (Brenman et al., 1995). The significance of these findings is that a selective loss of sarcolemmal nNOS is associated with muscular dystrophy where the dystrophin gene is mutated (Brenman et al., 1995" Chao et al., 1996; Grozdanovic et al., 1996). Although nNOS is generally considered to be present constitutively in most neurons and other tissues, there is increasing evidence that nNOS can undergo expressional regulation. Upregulation of nNOS mRNA seems to represent a general response of neuronal cells to stress induced by a widespread diversity of physical, chemical and biological agents (F6rstermann et al., 1998). This upregulation of nNOS may represent a part of a more global expression of numerous genes, in response to cellular stress, ultimately resulting in cellular injury and apoptosis. Downregulation of nNOS may occur in response to endotoxin and cytokines that cause induction of iNOS. Since expressional changes in constitutive nNOS may lead to changes in the quantity of NO generated and released from neurons, it is important to develop a better understanding of genetic control of nNOS expression. The human nNOS gene has been mapped to the q l4-qter position of chromosome 12 (Kishimoto et al., 1992), and the genomic structure of nNOS is well documented from human brain (Hall et al., 1994; Marsden et al., 1994). The catalytic activity of nNOS is turned on in neuronal cells by an increase in the intracellular concentration of free calcium as a result of the action potential or sodium current. Calcium binding to calmodulin allows the calcium-calmodulin complex to bind to the appropriate amino acid sequence in nNOS. As discussed above, the binding of calmodulin triggers the electron transport cycle in nNOS, which results in activation of heine iron-bound molecular oxygen and consequent oxygenation of arginine to yield NO plus citrulline. Although depriving cells of flavins, heine, tetrahydrobiopterin or arginine can each lead to diminished catalytic activity, the principal physiological mode of regulation of nNOS activity 10

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is a change in the intracellular concentration of free calcium. This m e c h a n i s m affords rapid changes in the catalytic activity of nNOS, whereas expressional changes in nNOS protein levels in response to cellular stress represents a much slower and delayed or long-term regulation of nNOS activity and N O production. Therefore, rapid increases in N O production by n N O S result primarily from the firing of action potentials in the neuron. The only known m e c h a n i s m for rapid decreases in N O production by nNOS is negative feedback control of n N O S catalytic activity by N O itself (Rogers and Ignarro, 1992; Buga et al., 1993; Griscavage et al., 1993, 1994; A b u - S o u d et al., 1995; Hyun et al., 1995; Cohen et al., 1996). Evidence from experiments using e n z y m e s , isolated tissues, cell culture and in vivo experiments indicate that p h y s i o l o g i c a l l y relevant concentrations of NO directly inhibit the catalytic activity of all three NOS isoforms, and that NO plays an important autoregulatory role.

5. REFERENCES Abu-Soud HM, Stuehr DJ (1993): Nitric oxide synthases reveal a role for calmodulin in controlling electron transfer. Proc Natl Acad Sci USA 90:10769-10772. Abu-Soud HM, Wang J, Rousseau DL, Fukuto JM, Ignarro LJ, Stuehr DJ (1995): Neuronal nitric oxide synthase self-inactivates by forming a ferrous-nitrosyl complex during aerobic catalysis. J Biol Chem 270:22997-23006. Anderson RG (1993): Caveolae: where incoming and outgoing messengers meet. Ptvc Natl Acad Sci USA 90:10909-10913.

Arnold WE Mittal CK, Katsuki S, Murad F (1977): Nitric oxide activates guanylate cyclase and increases guanosine 3':5'-cyclic monophosphate levels in various tissue preparations. Proc Natl Acad Sci USA 74:3203-3207. Axelsson KL, Andersson KG (1983): Tolerance towards nitroglycerin, induced in vivo, is correlated to a reduced cGMP response and an alteration in cGMP turnover. EurJ Pharmacol 88:71-79. Axelsson KL, Wikberg JES, Andersson RGG (1979): Relationship between nitroglycerin, cyclic GMP and relaxation of vascular smooth muscle. Life Sci 24:1779-1786. Axelsson KL, Andersson RGG, Wikberg JES (1982): Vascular smooth muscle relaxation by nitro compounds: reduced relaxation and cGMP elevation in tolerant vessels and reversal of tolerance by dithiothreitol. Acta Phapvnacol Toxicol 50:350-357. Azuma H, Ishikawa M, Sekizaki S (1986): Endothelium-dependent inhibition of platelet aggregation. Br J Pharmacol 88:411-415.

Boeckxstaens GE, Pelckmans PA, Ruytjens IF, Bult H, De Man JG, Herman AG, Van Maercke YM (1991): Bioassay of nitric oxide released upon stimulation of non-adrenergic non-cholinergic nerves in the canine ileocolonic junction. Br J Pharmacol 103:1085-1091. B6hme E, Graf H, Schultz G (1978): Effects of sodium nitroprusside and other smooth muscle relaxants on cyclic GMP formation in smooth muscle and platelets. Adv Cyclic Nucleotide Res 9:131-143. Bredt DS, Snyder SH (1989): Nitric oxide mediates glutamate-linked enhancement of cGMP levels in the cerebellum. Proc Natl Acad Sci USA 86:9030-9033. Bredt DS, Snyder SH (1990): Isolation of nitric oxide synthetase, a calmodulin-requiring enzyme. Proc Natl Acad Sci USA 87:632-685. Bredt DS, Snyder SH (1994): Nitric oxide: a physiologic messenger molecule. Annu Rev Biochem 63:175-195. Bredt DS, Hwang PM, Snyder SH (1990): Localization of nitric oxide synthase indicating a neural role for nitric oxide. Nature 347:768-770. Bredt DS, Hwang PM, Glatt CE, Lowenstein C, Reed RR, Snyder SH (1991): Cloned and expressed nitric oxide synthase structurally resembles cytochrome P-450 reductase. Nature 351:713-714. Brenman JE, Chart DS, Xia H, Aldape K, Bredt DS (1995): Nitric oxide synthase complexed with dystrophin and absent from skeletal muscle sarcolemma in Duchenne muscular dystrophy. Cell 82:743-752. Brenman JE, Chao DS, Gee SH, McGee AW, Craven SE, Santillano DR, Wu Z, Huang F, Xia H, Peters ME Froehner SC, Bredt DS (1996a): Interaction of nitric oxide synthase with the postsynaptic density protein PSD-95 and alphal-syntrophin mediated by PDZ domains. Cell 84:757-767. Brenman JE, Christopherson KS, Craven SE, McGee AW, Bredt DS (1996b): Cloning and characterization of postsynaptic density 93, a nitric oxide synthase interacting protein. J Neutvsci 16:7407-7415. Buga GM, Griscavage JM, Rogers NE, Ignarro LJ (1993): Negative feedback regulation of endothelial cell function by nitric oxide. Circ Res 73:808-812. 11

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CHAPTER II

Histochemistry of nitric oxide synthase in the central nervous system S.R. VINCENT

1. INTRODUCTION Our current understanding of the distribution of the neurons that can produce nitric oxide has been built on a number of complementary technical developments. The introduction of NADPH diaphorase histochemistry in formaldehyde-fixed material led to the frequent use of this method for neuroanatomical studies even before the nature of the enzyme responsible was discovered (Scherer-Singler et al., 1983). The demonstration that the NADPH diaphorase histochemical staining is due to the activity of nitric oxide synthase (NOS) (Hope et al., 1991) quickly enabled the detailed anatomical analysis of NO-producing neurons throughout the nervous system (Vincent and Hope, 1992; Vincent and Kimura, 1992). The direct relationship between NADPH diaphorase staining and NOS expression has been well documented (Bredt et al., 1991; Dawson et al., 1991 ). The absence of neuronal expression of NOS immunoreactivity and NADPH diaphorase activity in knockout mice lacking nNOS provided definitive evidence for the specificity of this simple histochemical procedure (Huang et al., 1993). The distribution of NADPH diaphorase-positive neurons in the rat brain has been reviewed previously, together with discussions on the specificity of this method (Leigh et al., 1990; Vincent and Hope, 1992; Vincent and Kimura, 1992; Vincent, 1994; Blottner et al., 1995; Norris et al., 1995). Ultrastructural examination of NADPH diaphorase has also been described (Hope and Vincent, 1989; Rothe et al., 1998). The relationship of NADPH diaphorase staining and NOS immunohistochemistry to soluble guanylyl cyclase (Schmidt et al., 1992a) and to NO-induced cGMP-immunoreactivity has been reviewed (Southam and Garthwaite, 1993; De Vente et al., 1998). More recently, the development of a World Wide Web accessible digital atlas of NADPH diaphorase staining in the mouse brain [http://nadph.anatomy.lsumc.edu] presents an exciting new tool for the neuroscience community. The results obtained with NADPH diaphorase histochemistry have been confirmed and extended using antibodies against the various NOS isoforms (Bredt et al., 1990) as well as in situ hybridization (Bredt et al., 1991). Of particular irnportance has been the description of alternatively spliced forms of nNOS expressed in certain brain regions (Brenman et al., 1997; Eliasson et al., 1997). Another approach for localizing NOS that has been use~t is the autoradiographic localization of [3H]L-NG-nitro-arginine binding to the enzyme (Burazin and Gundlach, 1995; Kidd et al., 1995; Hara et al., 1996; Rao and Butterworth, 1996). Certain caveats must of course be kept in mind when using any of these techniques. For example, the NADPH diaphorase staining observed in cell groups lacking NOS expression Hamlhook ,{f CIwmical Neuroanatomx. Vol. 17: Functional Nettroanatomy ,!t the Nitric Oxide System H.W.M. Steinbusch, J. De Vente and S.R. Vincent. editors @ 2000 Elsevier Science B.V. All rights reserved.

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often appears to be due to technical artifacts, including fixation difficulties (Hope and Vincent, 1989; Matsumoto et al., 1993; Spessert and Layes, 1994; Buwalda et al., 1995; Gonzalez-Hernandez et al., 1996). Indeed, some have suggested running the NADPH diaphorase reaction in the presence of formaldehyde to overcome this problem (Nakos and Gossrau, 1994; Grozdanovic et al., 1995). Poor fixation has been reported to result in the NADPH diaphorase staining of astrocytes as well (Gabbott and Bacon, 1996: Kugler and Drenckhahn, 1996). Alkaline phosphatase activity can sometimes hydrolyze NADPH to NADH which can then be used as a substrate for other diaphorase reactions leading to false-positive formazan formation (Song et al., 1994; Grozdanovic and Gossrau, 1995b). Cytochrome-P450 reductase has also been suggested as a possible source for NADPH diaphorase activity, however, although some neuronal populations do express both NOS and cytochrome-P450 reductase-immunoreactivity, this enzyme does not appear to contribute to NADPH diaphorase staining (Norris et al., 1994; Young et al., 1997). Poor fixation may also contribute to false-positive staining with NOS antibodies (Wendland et al., 1994; Kugler and Drenckhahn, 1996). Finally, some neurons deep in the rat cortex have been reported to be diaphorase-positive but nNOS-immunonegative (Kharazia et al., 1994). It may be that these cells contain an isozyme of nNOS which is detected histochemically but not with the particular antibody used. There have been attempts to develop pharmacological methods to help ensure the specificity of NADPH diaphorase staining for NOS. Biochemical studies on the unfixed enzyme indicate that the NADPH diaphorase activity of nNOS is Ca 2+/calmodulin-independent (Schmidt et al., 1992b). This is consistent with observations that calcium chelators are without effect on NADPH diaphorase histochemistry (Hope and Vincent, 1989; Spessert et al., 1994). However, one group has reported that EDTA can inhibit neuronal NADPH diaphorase staining (Sancesario et al., 1993). Furthermore, studies by Morris et al. (1997) indicate that neuronal NADPH diaphorase staining is dependent on Ca2+/calmodulin and suggest that the intensity of the NADPH diaphorase staining may be related to the level of enzyme activation at the moment of tissue fixation. A number of groups have reported inhibition of NADPH diaphorase with the non-selective flavoprotein inhibitor, diphenyleneiodonium (Blottner and Baumgarten, 1992, 1995; Spessert and Claassen, 1998). NADPH diaphorase staining of formaldehyde-fixed intermediolateral spinal neurons was blocked by NG-nitro-L-arginine (L-NNA), but was still observed in the presence of NG-monomethyl-L-arginine (NMMA) and 7-nitroindazole (Blottner and Baumgarten, 1992, 1995). The NADPH diaphorase staining in the olfactory epithelium was stimulated by addition of the NOS substrate L-arginine, and was inhibited by the NOS inhibitor L-NG-nitro arginine (Dellacorte et al., 1995). Methylene blue, which is a very non-specific inhibitor of NOS, can also block NADPH diaphorase staining (Luo et al., 1995). ~-NADPH can be substituted for the physiological [3-NADPH, and this may result in a diaphorase staining more specific for NOS (Hope and Vincent, 1989; Grozdanovic and Gossrau, 1995a), although one group found that neuronal NADPH diaphorase in the olfactory bulb could not utilize ~-NADPH (Spessert et al., 1994; Spessert and Claassen, 1998). Over the past decade, the use of NADPH diaphorase histochemistry, complemented by these other methods, has allowed NO-producing neurons to be described in a wide variety of species. The comparative anatomy of the NO system in various vertebrates is reviewed in another chapter in this volume (Alonso et al. this volume). In this chapter, the distribution and characteristics of such cells in the mammalian central nervous system will be reviewed. Unless otherwise mentioned, this description is based on work in the rat brain.

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2. LOCALIZATION OF NITRIC OXIDE SYNTHASE IN THE BRAIN

2.1. THE OLFACTORY BULB Within the nasal mucosa, a dense innervation of NOS-positive fibers derived from the sphenopalatine ganglion is present (Hanazawa et al., 1993; Kulkarni et al., 1994; Lee et al., 1995; Jeon et al., 1997; Kim et al., 1997). NADPH diaphorase staining has also been reported in cells with morphology reminiscent of microvillar olfactory cells, but not in the respiratory epithelium and the sustentacular cells (Dellacorte et al., 1995). nNOS is expressed in the presumptive nervous layer of the olfactory placode and the cells that differentiate into embryonic olfactory receptor neurons (Roskams et al., 1994). However, no evidence for NOS expression was found either in the mature main olfactory epithelium nor in the vomeronasal organ, in spite of the strong diaphorase staining of the surface of the main olfactory epithelium (Kishimoto et al., 1993; Kulkarni et al., 1994; Roskams et al., 1994; Lee et al., 1995). There is intense NADPH diaphorase staining of the glomeruli of the olfactory bulb (Vincent and Kimura, 1992). In the accessory olfactory bulb the entire vomeronasal nerve and all vomeronasal glomeruli were strongly labeled, contrary to the main olfactory bulb, where only dorsomedial olfactory glomeruli displayed NADPH diaphorase activity. This glomerular NADPH diaphorase reaction is pharmacologically different from that due to NOS, and does not co-localize with NOS immunohistochemistry (Spessert et al., 1994). Thus NOS does not appear to be present in mature primary olfactory neurons. Interneurons containing NOS protein and mRNA, and exhibiting NADPH diaphorase activity have been well described in the plexiform layer of the main olfactory bulb and the granule cell layer of main and accessory olfactory bulbs (Fig. 1A,C) (Kishimoto et al., 1993). Periglomerular cells (Fig. 1B) and granule cells in the main olfactory bulb were also NOS-positive (Kishimoto et al., 1993). NADPH diaphorase-positive neurons were identified as periglomerular cells in the glomerular layer and external plexiform layer, horizontal cells in the internal plexiform layer, and granule cells and deep short-axon cells in the granule cell layer (Porteros et al., 1994). Some of these cells appear to correspond to the superficial short-axon cell described in Golgi and electron microscopic studies, and the dendrites of these cells lie within the periglomerular region and in the superficial external plexiform layer (Scott et al., 1987). In addition to NOS, these cells also express somatostatin as well as NPY and the C-terminal flanking peptide of NPY, C-PON (Scott et al., 1987; Villalba et al., 1989). Positive periglomerular cells are more frequently associated with typical than atypical glomeruli (Crespo et al., 1996). The NADPH diaphorase-positive periglomerular cells appear to form a distinct population, and do not express tyrosine hydroxylase (Samama and Boehm, 1996), calbindin Dzgk (Alonso et al., 1993), calretinin or parvalbumin (Brifirn et al., 1997). A type of short-axon cell which is also stained for NADPH diaphorase, NPY, C-PON and somatostatin, lies deep in the granule cell layer, frequently near the ventricular layer and its dendrites lie parallel to that layer (Scott et al., 1987; Villalba et al., 1989). Many NADPH-stained neurons are also found in regions known to provide centrifugal inputs to the olfactory bulb, including all subdivisions of the anterior olfactory nucleus (Fig. 1D), the anterior hippocampal rudiment, anterior and posterior levels of the piriform cortex, and the vertical and horizontal limbs of the diagonal band of Broca (Davis, 1991; Garcia-Ojeda et al., 1994).

21

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S.R. Vincent

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Fig. 1. NADPH diaphorase staining in the olfactory bulb (A) is present in periglomerular cells (B) as well as the inner plexiform layer. The granule cell layer of the accessory olfactory bulb is particularly heavily stained (A, C). Many large multipolar cells are present in the anterior olfactory nucleus lying about the intrabuibar anterior commissure. Bar indicates 500 ~tm in A, 75 I,m in B, 100 Itm in C. and 150 Itm in D.

2.2. THE CEREBRAL CORTEX Neurons expressing NOS are widely scattered throughout layers II-VI and in the subcortical white matter of the cortex (Fig. 2A,B). Although these cells represent only a very small proportion of cortical neurons (0.5-2%), they give rise to an extensive fiber network throughout the entire cortical neuropil (Vincent and Kimura, 1992: Valtschanoff et al., 1993b). An electron microscopic examination of NADPH diaphorase-stained material found that postsynaptic elements were only very rarely stained in the cortex and that most staining was in presynaptic elements (Faber-Zuschratter and Wolf, 1994). Pyramidal neurons of the rat neocortex do not normally express NADPH diaphorase reactivity (Vincent and Kimura, 1992; Kitchener et al., 1993). Some cortical NADPH diaphorase neurons are aspiny, while others possess moderate numbers of spines (Gabbott et al., 1995). Cortical NOS neurons have been shown to be GABAergic (Chesselet and Robbins, 1989; Gabbott and Bacon, 1995) 22

Histochemistry of nitric oxide svnthase in the central ner~'ous system

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a m

Fig. 2. Scattered multipolar cells are lound throughout the neocortex, giving rise to a dense fiber network that is often associated with blood vessels (arrow)(A). Similar muitipolar cells are common in the subcorticai white matter (B). Similar multipolar cells are found scattered in the ventral (C, D, E) and dorsal striatum (F). The granule cells in the islands of Calleja are very heavily stained (C, D, E). Bar indicates 50 ~m in A and E, 75 Ixm in B, C and F, and 250 g m in D.

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and to express somatostatin and NPY (Vincent et al., 1983a.c; Kowall and Beal, 1988). The intermediate filament protein peripherin is present in a discrete population of aspiny intemeurons which are stained by NADPH diaphorase or NOS immunohistochemistry in the adult rat neocortex, although the functional significance of this is unclear at present (Rhrich-Haddout et al., 1997). Most of the neurons found in the subcortical white matter in the rat express NOS (Fig. 2B; Meyer et al., 1991). Similar interstitial cells in the subcortical white matter have also been noted in the cat (Mizukawa et al., 1988b). These interstitial neurons in the subcortical white matter give rise to association fibers which project mostly to the gray matter of the overlying cytoarchitectonic area, but which may extend also over different cytoarchitectonic areas (Meyer et al., 1991 ). An association of NADPH diaphorase staining with developing barrel fields has been described in the mouse (Mitrovic and Schachner, 1996). The somatosensory barrel fields of the rat show stronger NADPH diaphorase staining than the surrounding neuropil, but positive neurons are more frequently found outside the barrels (Valtschanoff et al., 1993b; Franca and Volchan, 1995). The NADPH diaphorase activity in the neuropil of the rat visual cortex appears to vary in a unimodal circadian manner, being highest at 0600 h and decreasing between 1200 h and 1800 h (Hilbig and Punkt, 1997). Together, these observations suggest that NO may play a role in sensory processing in the cortex. The well-documented role of NO as a vascular regulator has led to suggestions that it may serve a similar role in the brain. NADPH diaphorase-positive neurons are often found in close proximity to the intracerebral vascular profiles, sending processes to the vessels and/or being directly apposed to their walls (Fig. 2A) (Estrada et al., 1993; Samama et al., 1995). Neurovascular contacts are preferentially located close to the interface between the cerebral cortex and white matter (Estrada et al., 1993). NADPH diaphorase-positive neurons often lie near branching points of the arteriolae which descend through the cerebral cortex from its pial surface. This spatial relationship is consistent with their involvement in the neural control of cortical blood flow (Regidor et al., 1993). 2.3. THE HIPPOCAMPAL FORMATION Following the original suggestion that NO may be able to function as a retrograde messenger in the brain (Garthwaite et al., 1988), much attention has focussed on the localization of NOS in the hippocampus, and its possible involvement in learning and memory. Pyramidal cells in the rat subiculum are NADPH diaphorase-positive and immunoreactive for neuronal NOS, while the adjacent cells in the CA1 region are not (Fig. 3) (Vincent and Kimura, 1992; Valtschanoff et al., 1993a; Greene et al., 1997). In the ventral subiculum, the majority of NADPH diaphorase-positive pyramidal neurons were found in the superficial cell layer (Greene et al., 1997). This distribution of NADPH diaphorase activity was mimicked by that of immunoreactivity for the neuronal isoform of NOS (Greene et al., 1997). Electrophysiological analysis indicates that NADPH diaphorase activity is preferentially found in subicular neurons exhibiting a regular spiking phenotype and is absent from the intrinsically burst-firing neurons (Greene et al., 1997). One group reported that neurons in the olfactory bulb and striatum, the cerebellar granule cells, dentate granule cells and CA1 pyramidal cells express immunoreactivity corresponding to endothelial NOS (eNOS) (Dinerman et al., 1994). This observation has led to suggestions that eNOS in CA1 pyramidal cells may play a role in hippocampal synaptic plasticity (O'Dell et al., 1994). However, in these studies both NADPH diaphorase and eNOS staining were 24

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Fig. 3. NADPH diaphorase staining in the ventral hippocampal formation is illustrated in this horizontal section (A). There is dense fiber staining in entorhinal cortex (EC), the pyramidal cells in the subiculum (S) are positively stained (B), while those in the adjacent CA1 region lack NADPH diaphorase staining (C). The granule cells of the dentate gyrus (DG) are also unstained (D). Instead basket cells (D) and other interneurons are stained throughout the hippocampus. Bar indicates 400 ~tm in A, and 200 Ixm in B, C and D.

found to be highly variable and dependent on fixation parameters (Dinerman et al., 1994; Vaid et al., 1996), and other groups have not observed any neuronal staining for eNOS (Seidel et al., 1997; Stanarius et al., 1997). Indeed, the first immunohistochemical study of NOS in the brain used an antibody which stained both endothelial cells and neurons, but did not result in staining of CA1 pyramidal cells (Bredt et al., 1990). Endothelial cells in the brain clearly show a strong NADPH diaphorase activity indicative of eNOS expression (Gabbott and Bacon, 1993), and neuronal NADPH diaphorase is completely absent in nNOS knockout mice which still express eNOS (Huang et al., 1993). These observations indicate that the suggestion that eNOS is expressed in neurons should be treated with some skepticism. Detailed double-staining experiments have demonstrated that nearly all of the NADPH diaphorase-positive neurons in the hippocampus also contain GABA (Valtschanoff et al., 25

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1993a), suggesting that they are interneurons. Within the dentate gyrus, NOS and NADPH diaphorase are found in basket cells and other interneurons, but are absent from the granule cells (Fig. 3D) (Mizukawa et al., 1989: Leigh et al., 1990: Vincent and Hope, 1992; Vincent and Kimura, 1992; Hong et al., 1993). These NADPH diaphorase neurons in the infragranular zone of the dentate gyrus may be selectively vulnerable to transient ischemic insults (Hong et al., 1993). A scattered population of NADPH diaphorase-positive neurons similar to that in cortical regions has been noted in the amygdala of various species. All nuclei of the amygdala contained subpopulations of diaphorase-positive neurons and the highest density is seen in the lateral nucleus (Mizukawa et al., 1988a: McDonald et al., 1993). Positive cells have also been noted in the claustrum in the cat (Hinova-Palova et al., 1997). 2.4. THE BASAL FOREBRAIN The granule cells of the islands of Calleja are intensely positive for NADPH diaphorase (Figs. 2 and 4) and display nNOS mRNA (Meyer et al., 1994: Sugaya and McKinney, 1994). These cells closely surround the arterioles perfusing the ventral striatum and pallidum and

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Fig. 4. Many of the magnocellular cholinergic neurons of the basal forebrain are strongly NADPH diaphorase-positive such as these in the horizontal limb of the nucleus of the diagonal band of Broca (A, B). The granule cells of the major island of Calleja are also strongly stained (,4, C). Bar indicates 100 rtm in A, and 50 l~tm in B and C.

26

Histochemistry of nitric oxide svnthase in the central ner~'ous system

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may play a role in the regulation of blood flow to specific centers of the limbic forebrain (Meyer et al., 1994). The presence of magnocellular NADPH diaphorase neurons in the basal forebrain has been well described (Brauer et al., 1991). In the rat, many NADPH diaphorase-positive cells located in the medial septum and the nucleus of the horizontal limb of the diagonal band of Broca (Fig. 4A,B) project to the hippocampus (Kinjo et al., 1989). Some NADPH diaphorasepositive, calbindin-D2sk-negative cells have also been described in the lateral septum in the guinea pig (Doutrelant-Viltart and Poulain, 1996). Double- and triple-staining experiments indicate that NADPH diaphorase labels a majority of the magnocellular cholinergic neurons in the medial septum and diagonal band nuclei (Pasqualotto and Vincent, 1991). Virtually all NADPH diaphorase-reactive magnocellular neurons in the medial septum and the vertical and horizontal limbs of the diagonal band of Broca were choline acetyltransferase-immunoreactive (Kitchener and Diamond, 1993). In the most rostral sections of the medial septum and diagonal band, approximately 70% of the choline acetyltransferase-immunoreactive neurons were NADPH diaphorase-reactive, whereas the proportion decreased progressively to about 30% at the level of the decussation of the anterior commissure (Kitchener and Diamond, 1993). A similar number of the ChAT-positive cells in these regions expressed NOS mRNA (Sugaya and McKinney, 1994). Most of these neurons also contain galanin immunoreactivity (Pasqualotto and Vincent, 1991). However, small populations of galanin-positive/diaphorase-negative or diaphorase-positive/galanin-negative cholinergic neurons were also observed. In the more caudal portions of the cholinergic basal forebrain, very few galanin or NADPH diaphorasepositive neurons were observed. Thus, galanin and NADPH diaphorase coexist in the majority of cholinergic basal forebrain neurons in the regions innervating limbic structures (Pasqualotto and Vincent, 1991). These cells also express the low-affinity p75 NGF receptor (Peng et al., 1994), but do not express the alpha 1 subunit of GABA1 receptors (Hartig et al., 1995). In the nucleus basalis magnocellularis of the cat, large cholinergic cells that project to striate cortex have been reported to contain NADPH diaphorase, and calbindin (Bickford et al., 1994). However, it is interesting to note that while NADPH diaphorase activity occurred in about one third of the basal forebrain cholinergic neurons in the rat it is found in virtually none of these neurons in the monkey, baboon or human (Ellison et al., 1987; Mesulam et al., 1989; Geula et al., 1993). Thus it is difficult to generalize regarding the role of a septohippocampal NOS system in hippocampal synaptic plasticity. 2.5. THE BASAL GANGLIA Within the striatum, NOS is expressed in a small population of medium-aspiny interneurons. These NADPH diaphorase neurons are evenly distributed throughout the striatum and nucleus accumbens (Fig. 2C-F) (Kowall et al., 1985: Vincent and Kimura, 1992). Essentially all striatal NADPH diaphorase neurons show somatostatin immunoreactivity, and most also display NPY and C-PON immunoreactivities (Vincent et al., 1982, 1983a,c; Vincent and Johansson, 1983; Villalba et al., 1988; Rushlow et al., 1995; Figueredo-Cardenas et al., 1996). These cells were initially reported to lack GAD expression (Chesselet and Robbins, 1989), although others have reported the presence of GAD67-immunoreactivity in these cells (Kubota and Kawaguchi, 1994). The giant striatal cholinergic neurons and the medium-spiny GABA output neurons do not express NADPH diaphorase activity (Vincent et al., 1983a,c; Geula et al., 1993). The striatal NOS neurons appear to receive a direct dopaminergic input (Fujiyama and Masuko, 1996) and the intensity of NADPH diaphorase staining in rat striatal neurons 27

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is decreased following systemic treatment with the D l-like dopamine receptor antagonist SCH23390, and increased by the D2-1ike antagonist eticlopride (Morris et al., 1997). These cells also express mRNA for substance P receptors (Kaneko et al., 1993) and muscarinic cholinergic receptors (Bernard et al., 1992). The NOS interneurons lack calretinin (Bennett and Bolam, 1993a: Kubota and Kawaguchi, 1994), and parvalbumin (Kita et al., 1990: Kubota and Kawaguchi, 1994), although some express calbindin D28k (Bennett and Bolam, 1993b). Similar results have been seen in accumbens neurons (Hussain and Totterdell, 1994: Hussain et al.. 1996). Calcineurin, the calmodulindependent protein phosphatase also appears to be absent from these cells (Goto et al., 1987). Most striatal NADPH diaphorase neurons show low levels of glutamate receptor NR1 immunoreactivity, although a small number of these cells do express high levels of the NMDA receptor mRNA (Price et al., 1993; Augood et al., 1994). There appears to be some controversy concerning the extent to which the huntingtin protein is expressed in the striatal NADPH diaphorase neurons (Kumar et al., 1997: Fusco et al., 1999). These cells also express high levels of Mn-superoxide dismutase (Inagaki et al., 1991), which may protect them from damage from NO and other radicals. This might be one reason for the sparing of these neurons seen in Huntington's disease (Fen'ante et al., 1985). Age-related declines in NADPH diaphorase-positive neuronal density and in neuropil staining have been noted (Kuo et al., 1997). The striatal NADPH diaphorase neurons have been characterized electrophysiologically as PLTS cells (persistent and low-threshold spike cells) (Kawaguchi, 1993). These cells fired both sodium-dependent, persistent depolarization spikes and calcium-dependent, low-threshold spikes in addition to fast spikes. It is possible that activation of these low-threshold calcium spikes may act as one trigger for NO production in these cells. 2.6. THE THALAMUS NOS-positive cell bodies are not common in the thalamus (Fig. 5). NADPH diaphorasepositive and NOS-immunoreactive perikarya are principally seen along the midline in the paraventricular (Fig. 5A), rhomboid, and central medial nuclei, and in the dorsal and ventral lateral geniculate nuclei (Fig. 5D) (Bertini and Bentivoglio, 1997). Scattered neurons are found in the lateral posterior nucleus, in the dorsal part of the medial geniculate nucleus, and in the ventromedial nucleus (Bertini and Bentivoglio, 1997). NADPH diaphorase-positive neurons are also present in the paratenial nucleus up until postnatal day 15 (Garcia-Ojeda et al., 1997). Intensely stained NADPH diaphorase neurons are restricted to the lateral half of the magnocellular division of ventral lateral geniculate (Mitrofanis, 1992; Gonzalez-Hernandez et al., 1994). There is evidence that NADPH diaphorase is present in both geniculotectal projection neurons and local-circuit GABAergic neurons in the ventral lateral geniculate nucleus (Gabbott and Bacon, 1994a,b; Gonzalez-Hernandez et al., 1994). The anteroventral and anteromedial nuclei, the midline nuclei, the anterior intralaminar nuclei, and the lateral and medial geniculate nuclei have high densities of NADPH diaphorase-positive fibers (Fig. 5) (Bertini and Bentivoglio, 1997). NADPH diaphorase is found in patches in the mediodorsal and midline thalamic nuclei of cats. These patches match acetylcholinesterase (AChE)-rich patches within the medial thalamus (Mengual et al., 1993) and appear to derive from the cholinergic NOS neurons of the laterodorsal and pedunculopontine tegmental nuclei. NADPH diaphorase fibers in the paraventricular thalamic nucleus derive from both the lateral hypothalamus and the brainstem (Otake and Ruggiero, 1995). A group of positive cells is present in the subthalamic nucleus of rat (Fig. 5C) and cat (Mizukawa et al., 1989). Subthalamic neurons also express high levels of guanylyl cyclase 28

Histochemistry of nitric oxide svnthase in the central nervous system

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Fig. 5. Although there are few NADPH diaphorase-positive neurons in the thalamus, there is an extensive fiber network, particularly in the ventrolateral anteroventral nucleus and the midline (A). Note also the very strong staining of the axons in the stria terminalis (arrow). Positive cells are found in the paraventricular nucleus (B), the subthalamic nucleus (C) and the magnocellular portion of the ventral lateral geniculate (D). Bar indicates 1 mm in A, and 200 g m in B, C and D.

(Giuili et al., 1994) and type II cGMP-dependent protein kinase mRNA (E1-Husseini et al., 1995), which suggests that NO may act in an autocrine fashion in these cells• 2.7. THE SUPERIOR COLLICULUS In the pretectal complex, NADPH diaphorase activity is restricted to neurons and terminals in the nucleus of the optic tract and the dorsal terminal nucleus (Hilbig et al., 1995). The superior 29

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Fig. 6. The tectum contains a dense network of NADPH diaphorase-positive cells and fibers found throughout both the superior (A) and inferior (B) colliculi. In the superior colliculus, cells are most common in the superficial layers (C). A cluster of cells is also found in the dorsolaterai periaqueductal gray, extending from the aqueduct (A) into the deeper layers of the superior colliculus tD). In the inferior colliculus, positive multipolar cells are present in both the external cortex (E) and in the central nuclei (F). Bar indicates 125 ttm in A and B, 60 Itm in C, E and F, and 100 I,tm in D.

colliculus displays a complex pattern of NADPH diaphorase staining (Fig. 6A) (Tenorio et al., 1995, 1996). The highest activity occurs in the stratum zonale and stratum griseum superficiale (Fig. 6C), contrasting with the pale neuropil in the stratum opticum, where only a few positive neurons are found. In the stratum griseum-intermedium, positive neurons are grouped in patches separated by narrow, NADPH diaphorase-negative bands. In the deeper layers, the neuropil is NADPH diaphorase-negative, and few neurons show enzymatic activity. 30

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In contrast, numerous neurons in the dorsolateral part of the periaqueductal gray are intensely positive and continuous with the positive neurons in the stratum album profundum (Fig. 6D) (Gonzalez-Hernandez et al., 1992). 2.8. THE AUDITORY SYSTEM NADPH diaphorase-positive neurons are very common in the auditory system (Druga and Syka, 1993). Within the cochlea, hair cells, nerve fibers and ganglion cells were labeled by nNOS immunohistochemistry as well as NADPH diaphorase histochemistry in various species (Gosepath et al., 1997; Ruan et al., 1997; Hess et al., 1998; Fessenden et al., 1999). Positive cells are also seen in the medial nucleus of the trapezoid body and the periolivary nuclei (Reuss, 1998; Fessenden et al., 1999). Many positive cells are found in the dorsal and external cortices of the inferior colliculus and in the intercollicular commissure (Fig. 6B,E,F). Stained neurons are present in the dorsal part of the suprageniculate nucleus and in the basal part of the medial division of the medial geniculate. NADPH diaphorase-positive cells are also dispersed in the auditory temporal cortex within layers II-VI (Druga and Syka, 1993). 2.9. THE HYPOTHALAMUS There is a widespread distribution of NOS-positive neurons throughout the hypothalamus, suggestive of a major role for NO in autonomic regulation. A large group of multipolar, NADPH diaphorase-positive cells is present in the lateral hypothalamus (Fig. 7C,E) (Vincent and Kimura, 1992; Vanhatalo and Soinila, 1995). In addition, positive cells have been reported in the bed nucleus of the stria terminalis, and the axons of the stria terminalis are intensely stained (Fig. 5A) (Mizukawa et al., 1989; Bruning et al., 1994; Iwase et al., 1998b; Briski and Sylvester, 1999). This suggests that a NOS pathway from the hypothalamus to the amygdala may be present. The staining of the magnocellular neurosecretory neurons has been confirmed by many groups (Fig. 7C,D) (Sagar and Ferriero, 1987; Ar6valo et al., 1992). Many hypothalamic NADPH diaphorase neurons innervate the pituitary (Vanhatalo and Soinila, 1995), and the NADPH diaphorase activity in the posterior pituitary can be markedly increased by stimulation of the hypothalamo-neurohypophyseal system with salt loading (Sagar and Ferriero, 1987). NADPH diaphorase is found in magnocellular neurons expressing both argininevasopressin and oxytocin (Calka and Block, 1993; Miyagawa et al., 1994; Sanchez et al., 1994). Virtually all PACAP-38-immunoreactive neurons in the paraventricular and supraoptic nuclei exhibited NADPH diaphorase activity (Okamura et al., 1994a). Some NADPH diaphorase neurons in the periventricular parvicellular subdivision of the paraventricular nucleus displayed somatostatin immunoreactivity (Alonso et al., 1992a,b). Many magnocellular neurosecretory NADPH diaphorase neurons express calbindin D2sk (Alonso et al., 1992a), but the hypothalamic NADPH diaphorase neurons only rarely express calretinin (Ar6valo et al., 1993). Some are also tyrosine hydroxylase-immunoreactive (Blanco et al., 1997), and many hypothalamic NADPH diaphorase neurons display NMDA R l-immunoreactivity (Bhat et al., 1995). The retinorecipient cells in the suprachiasmatic region are not NADPH diaphorase-positive (Amir et al., 1995), but NOS fibers have been reported in the suprachiasmatic region (Amir et al., 1995; Chen et al., 1997). NOS cells are reported to be a subgroup of the suprachiasmatic population characterized also by immunoreactivity to vasoactive intestinal polypeptide (Reuss et al., 1995). 31

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Fig. 7. Positive cells are common beneath the fornix (F)in the subfornical organ (A) and the organum vasculosum of the lamina terminalis above the third ventricle (3V) (B). Within the hypothalamus, an extensive group of multipolar cells is present in the lateral hypothalamus {LH) {C. E). Many of the magnocellular neurosecretory neurons in the supraoptic nucleus (SO) beside the optic tract {OT) are also well stained {C. D). In the caudal hypothalamus, an extensive cell group is present in the supramammillary nucleus (SM), and the cells in the lateral medial mammillary nucleus (LM). The medial mammillary nucleus is unstained {MM). Bar indicates 50 Ixm in A and B, 200 ~m in C, 30 ~m in D and E. and 500 lJtm in F.

Some, but not all, neurons in the ventrolateral subdivision of the ventromedial nucleus display both NADPH diaphorase and NOS immunoreactivity (Rachman et al., 1996). Double-labeling histochemistry revealed that more than 70% of the NADPH diaphorase-positive neurons in the ventromedial nucleus express estrogen receptors (Okamura et al., 1994b; Rachman et al., 1996). Indeed, treatment with estradiol benzoate for two days elevated the 32

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number of NADPH diaphorase-positive neurons in female but not male rats (Okamura et al., 1994b). In the caudal hypothalamus, the presence of NADPH diaphorase activity in the circularis and anterior and posterior fornical nuclei has been noted (Ar6valo et al., 1992). There is an extensive network of NADPH diaphorase neurons in the various nuclei of the mammillary body (Fig. 7F) (Vincent and Kimura, 1992). Some of these neurons expressed enkephalin-, cholecystokinin-, or substance P-like immunoreactivities (Yamada et al., 1996). 2.10. CIRCUMVENTRICULAR ORGANS The cells of the organum vasculosum of the lamina terminalis stain intensely for NADPH diaphorase (Fig. 7B) (Bhat et al., 1995). nNOS immunolabeling has also been reported in the two most rostrally located circumventricular organs, the organum vasculosum of the lamina terminalis and the subfornical organ (Fig. 7A), and in the latter, some coexistence with arginine vasopressin has been observed (Alto et al., 1997). However, no nNOS immunoreactivity or NADPH diaphorase could be found in the subcommissural organ or in the area postrema (Vincent and Kimura, 1992; Alm et al., 1997). Both supra- and sub-ependymally localized NADPH diaphorase neurons and fibers in the wall of the third and lateral ventricles have extensive contact with the CSF (Sancesario et al., 1996a,b; Rodrigo et al., 1997). A similar contact is seen along the spinal canal (Tang et al., 1995a,b). Scattered NADPH diaphorase-positive cells have been noted in the pineal, and positive, non-sympathetic nerve fibers are located in the pineal capsule and intraparenchymally between the pinealocytes (Lopez-Figueroa and Moller, 1996; Aim et al., 1997). NADPH diaphorase or nNOS-positive nerve fibers derived from the sphenopalatine ganglion accompany blood vessels in the choroidal stroma and the choroidal epithelial cells have also been reported to be stained (Lin et al., 1996; Szmydynger-Chodobska et al., 1996). However, some groups find the choroidal epithelial cells do not display immunoreactivity for neuronal NOS (Sancesario et al., 1996b) while other groups find nNOS immunoreactivity in the epithelium of the choroid plexus (Lin et al., 1996; Alm et al., 1997). The NADPH diaphorase activity that has been reported in choroid epithelium (Lin et al., 1996) appears distinct from that in neurons (Sancesario et al., 1993, 1996a,b). There is also evidence for eNOS in the choroidal epithelial cells (Stanarius et al., 1997). These observations are consistent with a role of NO and the cGMP system in water homeostatic mechanisms, both via the hypothalamohypophyseal system and via the production of cerebrospinal fluid. 2.11. THE MESOPONTINE TEGMENTUM There is a dense cluster of NADPH diaphorase neurons in the interpeduncular nucleus (Fig. 8) (Mizukawa et al., 1989; Leigh et al., 1990; Vincent and Kimura, 1992; Dun et al., 1994; Iwase et al., 1998a), which are also known to express NOS mRNA (Sugaya and McKinney, 1994). NO produced in these neurons may act on the terminals of the neurons projecting there from the medial habenula, since these cells express very high levels of guanylyl cyclase and cGMP-dependent protein kinase mRNA (Matsuoka et al., 1992; E1-Husseini et al., 1995). The neurons of the magnocellular nucleus of the posterior commissure are intensely stained for NADPH diaphorase (Fig. 8C) (Vincent and Kimura, 1992; Iwase et al., 1998b) and also express NOS mRNA (Iwase et al., 1998a). A small band of NADPH diaphorase-positive neurons located in the middle of the antero-posterior extent of the periaqueductal gray below the level of the aqueduct and 33

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Fig. 8. A horizontal section through the interpeduncular nucleus illustrating the distinct staining pattern of neurons in the caudal (IPC) and rostral (IPR) nuclei (A). The positive cells in the IPR are shown in parasagittal section in (B). The strongly stained cells of the magnocellular nucleus of the posterior commissure are illustrated in this parasagittal section (C). Bar indicates 50 jtm in A and B. and 100 Itm in C.

directly above the supraoculomotor gray has been described and termed the supraoculomotor cap (Carrive and Paxinos, 1994). Numerous neurons in the caudal linear, dorsal, median, supralemniscal, and pontine raphe nuclei contained both serotonin-like immunoreactivity and NADPH diaphorase activity (Fig. 9) (Johnson and Ma, 1993: Dun et al., 1994). No NADPH diaphorase activity was detected in serotonergic neurons in the medullary nuclei (Johnson and Ma, 1993; Dun et al., 1994). Tyrosine hydroxylase-like immunoreactive neurons in the substantia nigra, locus cemleus, hypothalamus, olfactory bulb, and dorsal raphe nucleus of the rat do not display NADPH 34

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Fig. 9. A horizontal section through the mesopontine tegmentum caudal to the fourth ventricle (4v) illustrates the intense staining of the neurons in the laterodorsal tegmental nuclei (A, B) and the strong staining of the domal raphe nucleus in the midline (A, C). Bar indicates 250 ~tm in ,4, and 100 It m in B and C.

diaphorase activity (Vincent and Kimura, 1992; Johnson and Ma, 1993). The NOS neurons in the dorsal mesopontine tegmental nuclei have received much attention (Petrovicky and Nemcova, 1995; Nemcova et al., 1997). Essentially all of the cholinergic neurons in the laterodorsal and pedunculopontine tegmental nuclei express very high levels of NADPH diaphorase activity (Fig. 9) (Vincent et al., 1983b; Dun et al., 1994), and 35

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expressed NOS mRNA at high levels (Sugaya and McKinney, 1994). The high expression of NOS in these cholinergic neurons is seen in many species (Geula et al., 1993). Many of these cells also express a number of neuropeptides including substance P, CRF, GRP and the atriopeptides (Vincent et al., 1986; Nemcova et al., 1997). These neurons express a variety of glutamate receptor subtypes (Inglis and Semba, 1996), but do not display calbindin-D28k immunoreactivity (Geula et al., 1993). In the rat, these cells demonstrate a mild, but continuous regression of the dendritic tree during normal aging (Lolova et al., 1997). 2.12. THE CEREBELLUM Purkinje cells do not express NOS, which instead is found in the granule cells and basket cells of the cerebellar cortex (Fig. 10A) (Vincent and Kimura, 1992; Buwalda et al., 1995). In the mouse, NADPH diaphorase staining is seen in patches of granule cells and synaptic glomeruli. The patches are robust, and seen in all lobules, and are topographically ordered with respect to the Purkinje cell compartments (Hawkes and Turner, 1994). A similar patchy NADPH diaphorase pattern is seen in rat cerebellum (Li et al., 1997). cGMP immunostaining in cerebellar slices stimulated with the NO donors, nitroprusside and SIN-1, was found in granule cells, glomeruli, fibers, Bergmann glia and in other astrocytes (De Vente et al., 1989, 1990; Southam et al., 1992). However, type I cGMP-dependent protein kinase is very highly expressed in the Purkinje cells, suggesting that these cells are an important target of NO in the cerebellum (Vincent et al., 1998). 2.13. THE MEDULLA Within the medulla, NADPH diaphorase has been found in neurons displaying glutamate and somatostatin immunoreactivities in the paramedian region, lateral reticular field, the nucleus prepositus hypoglossi and the rostral nucleus of the solitary tract (Fig. 10B,C) (Maqbool et al., 1995). Within the nucleus of the solitary tract, the majority of NADPH diaphorase-positive cells are found within the central, medial, and ventral/ventrolateral subnuclei (Fig. 10B) (Takemura et al., 1994; Maqbool et al., 1995; Krowicki et al., 1997). Fiber staining is present in the subnucleus centralis, subnucleus gelatinosus, subpostremal zone, and the medial nucleus tractus solitarius (Krowicki et al., 1997). Most of the axon terminals with NADPH diaphorase in the nucleus of the solitary tract derive from the primary afferent neurons in the nodose ganglion (Lu et al., 1994). The NOS-immunoreactive neurons in the central subnucleus of the nucleus of the solitary tract have been shown to provide a projection to the nucleus ambiguus in rabbit, connecting esophageal afferents and efferents (Gai et al., 1995). In the dorsal motor nucleus of the vagus there are positively stained cells caudal to the obex and at its most rostral extent, but not at the intermediate level (Krowicki et al., 1997). A portion of these NOS-containing neurons are preganglionic vagal neurons which project to the abdominal viscera (Krowicki et al., 1997). NADPH diaphorase-positive neurons have been described in the medial vestibular nucleus and the spinal vestibular nuclei (Shaer et al., 1996; Takemura et al., 1996; Krukoff and Khalili, 1997). NADPH diaphorase staining has also been localized in a subpopulation of vestibular efferents (Singer and Lysakowski, 1996). There are neurons in the gracile nucleus which display both NOS mRNA and NADPH diaphorase staining and this staining increases with aging (Ma et al., 1997). In the spinal trigeminal nucleus (Fig. 10D), NOS is found in interneurons located in nucleus caudalis which give rise to processes that innervate trigeminothalamic neurons (Dohrn et al., 36

Histochemistrv of nitric oxide swTthase in the central ner~'ous system

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S Fig. 10. Strong staining is present in granule cells and glomeruli in the granule cell layer, and in basket cells and parallel fibers in the molecular layer of the cerebellar cortex (A). The large Purkinje cells are not stained. In the medulla, stained cells and fibers are present in the nucleus of the solitary tract (B), the medullary raphe (C) and the spinal nucleus of the trigeminal (D). Bar indicates 20 Itin in A, 100 Itm in B and C, and 50 t~m in D.

1994). This suggests that NO may indirectly influence orofacial nociceptive processing at the level of the spinal trigeminal nucleus. Similar staining is observed in the dorsomedial spinal trigeminal nucleus oralis where NO may regulate gustatory inputs (Takemura et al., 1994). The rostral ventral medulla contains a well-defined group of NADPH diaphorase neurons that are distinct from the adrenergic neurons of the C1 group. These have only limited monosynaptic projections to the spinal cord (Iadecola et al., 1993; Maqbool et al., 1995). These NADPH diaphorase neurons are also glutamate-immunoreactive and overlap the nucleus ambiguus, the lateral reticular nucleus and the A1/C1 catecholaminergic cell groups. A few NADPH diaphorase and glutamate-immunoreactive cells are found in the paraolivary area and gigantocellular tegmental field, in the external cuneate and infratrigeminal nuclei (Maqbool et al., 1995). In the rostral ventrolateral medulla NADPH diaphorase and somatostatin-immunoreactive cells are present in the paragigantocellular nucleus. 2.14. THE SPINAL CORD Neurons which stain intensely for NOS are found in laminae I-IV and X throughout the entire spinal cord. Preganglionic sympathetic neurons in the intermediolateral cell column 37

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of the thoracic and lumbar spinal cord are also intensely stained for NOS and express NOS mRNA (Vogel et al., 1997). Preganglionic parasympathetic neurons in the sacral cord also show substantial NOS immunostaining. Scattered neurons in laminae V, VI, VII, and VIII at all cord levels are weakly positive. In addition, punctate NOS staining is found throughout laminae I, III and surrounding some large motor neurons in the ventral horn (Valtschanoff et al., 1992; Saito et al., 1994). In the upper thoracic cord, positive cells are seen in the dorsal horn, the dorsolateral funiculus and lateral spinal neurons, the central canal region (Rexed lamina X) and in preganglionic sympathetic neurons in the intermediolateral nuclei (Blottner and Baumgarten, 1992; Valtschanoff et al., 1992; Blottner et al., 1993; Spike et al., 1993; Tang et al., 1995a,b; Vogel et al., 1997). In laminae I-III of the rat dorsal horn, NADPH diaphorase-positive neurons may contain GABA, glycine and acetylcholine (Spike et al., 1993; Laing et al., 1994), but they do not express NPY or enkephalin (Laing et al., 1994). Some of the lamina X NOS neurons were reported to be retrogradely labeled from the thalamus suggesting that they may be spinothalamic neurons (Lee et al., 1993). In the intermediolateral cell column at mid-thoracic levels, about half of cholinergic neurons stained for NADPH diaphorase (Blottner and Baumgarten, 1992). In particular, NADPH diaphorase is seen in the preganglionic cholinergic neurons innervating the adrenal gland, which also express enkephalin (Blottner and Baumgarten, 1992; Holgert et al., 1995). In the sacral spinal cord of the rat, the majority of both sympathetic and parasympathetic preganglionic neurons showed staining for NADPH diaphorase (Anderson, 1992; Hamilton et al., 1995). NADPH diaphorase staining may also be present in a few motoneurons in the aged rat spinal cord and may lead these neurons to eventual death (Kanda, 1996). A similar pattern of NADPH diaphorase staining has been described in the mouse spinal cord (Bruning, 1992).

3. ARGININE METABOLISM IN THE BRAIN

The discovery of NOS, and the generation of NO and citrulline from arginine has reawakened interest in arginine metabolism in the brain. Double-staining experiments have indicated that citrulline immunoreactivity is present in a subpopulation of NADPH diaphorase-positive neurons (Pasqualotto et al., 1991). In the neuropil in the ventroposterior nucleus of the thalamus NADPH diaphorase-positive fibers are often apposed to arginine-immunoreactive astrocytes and oligodendrocytes, and NADPH diaphorase-positive endothelial cells are often adjacent to arginine-positive astrocytes (Kharazia et al., 1997). These observations suggests that arginine may be stored in supporting cells, and supplied to nearby nerve fibers or endothelial cells as substrate for NOS. Indeed, many NOS-positive neurons express the neutral and basic amino acid transporter (NBAT) which facilitates the sodium-independent transport of arginine and other amino acids (Pickel et al., 1999). In rat brain slices, the arginine uptake inhibitor L-lysine reduced the cyclic GMP response to NMDA indicating that extracellular arginine availability can influence NO production (Grima et al., 1998). The relationship of NOS-positive neurons to those expressing argininosuccinate synthetase and lyase has been described in the rat brain (Arnt-Ramos et al., 1992) and spinal cord (Nakamura, 1997) and in cat hypothalamus (Isayama et al., 1997). The results indicate that some NOS neurons possess the enzymes required to directly recycle citrulline back to arginine. In other situations NOS appears to be expressed in neurons lacking this capability, and instead, a trans-cellular pathway for arginine and citrulline metabolism appears likely to occur (Arnt-Ramos et al., 1992). 38

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4. HEME OXYGENASE Following the demonstration that nitric oxide was the physiological EDRF the suggestion was made that carbon monoxide might be a similar sort of messenger (Marks et al., 1991; Schmidt, 1992; Verma et al., 1993). Carbon monoxide is produced by heine oxygenase during the degradation of heme to bile pigments. This enzyme reaction is not regulated, and heine concentrations in cells are limited. Furthermore, although carbon monoxide can bind to the heme of soluble guanylyl cyclase, it does so in a manner distinct from that of nitric oxide, and can only weakly activate the cyclase (Burstyn et al., 1995; Kharitonov et al., 1995). Thus it is unlikely that CO plays a significant role as a physiological messenger. Instead, the properties of heme oxygenase suggest that it may regulate the activities of the hemoproteins NOS and guanylyl cyclase by degrading free heme in cells containing these enzymes (Turcanu et al., 1998). Similar conclusions have been reached based upon immunohistochemical studies of the localization of the two heme oxygenase isoforms in many of the neurons in the brain which express NOS or soluble guanylyl cyclase (Vincent et al., 1994).

5. CONCLUSIONS Work over the past decade has led to a very complete understanding of the distribution of neurons which can generate nitric oxide in the central nervous system. NOS often co-localizes with classical neurotransmitters or neuropeptides, and thus NO synthesis is likely often coupled to the release of these transmitters. In this regard, it is interesting to note that NOS is found in neurons using the excitatory transmitter glutamate or the inhibitory neurotransmitter GABA. It is also found in cholinergic neurons and other aminergic cells. NOS is present in local circuit interneurons in many brain regions; however, in some areas it is found in principal neurons with long projections throughout the brain. NOS is also found in neurosecretory neurons and cells in some circumventricular organs. Thus, as with other neurotransmitters, it is difficult to generalize regarding the role of NO in the nervous system. NOS is localized in both cell bodies and dendrites, as well as in axons and nerve terminals. Thus, NO production is probably triggered by activation of postsynaptic calcium channels and the release of intracellular calcium stores in dendrites and cell bodies, or by the opening of voltage-gated calcium channels in nerve terminals. Hopefully our knowledge of NOS localization will lead to studies in the coming decade which will define the role of NO in these various sites in the nervous system.

6. REFERENCES Alm P, Skagerberg G, Nylen A, Larsson B, Andersson KE (1997): Nitric oxide synthase and vasopressin in rat circumventricular organs. An immunohistochemical study. Exp Brain Res 117:59-66. Alonso JR, Sanchez F, Ar6valo R, Carretero J, Aij6n J, Vazquez R (1992a): CaBP D-28k and NADPH-diaphorase coexistence in the magnocellular neurosecretory nuclei. Neuroreport 3:249-252. Alonso JR, Sanchez F, Ar6valo R, Carretero J, Vazquez R, Aij6n J (1992b): Partial coexistence of NADPHdiaphorase and somatostatin in the rat hypothalamic paraventricular nucleus. Neurosci Len 148:101-104. Alonso JR, Ar6valo R, Porteros A, Brifi6n JG, Lara J, Aij6n J (1993): Calbindin D-28K and NADPH-diaphorase activity are localized in different populations of periglomerular cells in the rat olfactory bulb. J Chem Neuroanat 6:1-6. Amir S, Robinson B, Edelstein K (1995): Distribution of NADPH-diaphorase staining and light-induced Fos

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expression in the rat suprachiasmatic nucleus region supports a role for nitric oxide in the circadian system. Neuroscience 69:545-555. Anderson CR (1992): NADPH-diaphorase-positive neurons in the rat spinal cord include a subpopulation of autonomic preganglionic neurons. Neurosci Lett 139:280-284. Ar6valo R, Sanchez F, Alonso JR, Carretero J, Vazquez R. Aij6n J (1992): NADPH-diaphorase activity in the hypothalamic magnocellular neurosecretory nuclei of the rat. Brain Res Bull 28:599-603. Ar6valo R, Sanchez F, Alonso JR, Rubio M, Aijon J, Vazquez R {1993): Infrequent cellular coexistence of NADPH-diaphorase and calretinin in the neurosecretory nuclei and adjacent areas of the rat hypothalamus. J Chem Neuroanat 6:335-341. Arnt-Ramos LR, O'Brien WE, Vincent SR (1992): Immunohistochemical localization of argininosuccinate synthetase in the rat brain in relation to nitric oxide synthase-containing neurons. Neuroscience 5•:773-789. Augood SJ, McGowan EM, Emson PC ( 1994): Expression of N-methyl-D-aspartate receptor subunit NR1 messenger RNA by identified striatal somatostatin cells. Neuroscience 59:7-12. Bennett BD, Bolam JP (1993a): Characterization of calretinin-immunoreactive structures in the striatum of the rat. Brain Res 609:137-148. Bennett BD, Bolam JP (1993b): Two populations of calbindin D28k-immunoreactive neurones in the striatum of the rat. Brain Res 610:305-310. Bernard V, Normand E, Bloch B (1992): Phenotypical characterization of the rat striatal neurons expressing muscarinic receptor genes. J Neulvsci 12:3591-3600. Bertini G, Bentivoglio M (1997): Nitric oxide synthase in the adult and developing thalamus: histochemical and immunohistochemical study in the rat. J Comp Neurol 388:89-105. Bhat GK, Mahesh VB, Lamar CA, Ping L, Aguan K, Brann DW {1995): Histochemical localization of nitric oxide neurons in the hypothalamus: association with gonadotropin-releasing hormone neurons and co-localization with N-methyl-D-aspartate receptors. Neutvendocrinoiogy 62:187-197. Bickford ME, Gunluk AE, Van Horn SC, Sherman SM {1994): GABAergic projection from the basal forebrain to the visual sector of the thalamic reticular nucleus in the cat. J Comp Neurol 348:481-510. Blanco E, Jirikowski GF, Riesco JM, Juanes JA. Vazquez R (1997): Coexistence of NADPH-diaphorase with tyrosine hydroxylase in hypothalamic magnocellular neurons of the rat. Neuropeptides 3•:227-230. Blottner D, Baumgarten HG (1992): Nitric oxide synthetase (NOS)-containing sympathoadrenal cholinergic neurons of the rat IML-cell column: evidence from histochemistry, immunohistochemistry, and retrograde labeling. J Comp Neurol 316:45-55. Blottner D, Baumgarten HG (1995): L-NNA inhibits the histochemical NADPH-d reaction in rat spinal cord neurons. Histochem Cell Biol 103:379-385. Blottner D, Schmidt HH, Baumgarten HG (1993): Nitroxergic autonomic neurones in rat spinal cord. Neutvreport 4:923-926. Blottner D, Grozdanovic Z, Gossrau R (1995): Histochemistry of nitric oxide synthase in the nervous system. Histochem J 27:785-811. Brauer K, Schober A, Wolff JR, Winkelmann E. Luppa H, Luth HJ et al. (1991): Morphology of neurons in the rat basal forebrain nuclei: comparison between NADPH-diaphorase histochemistry and immunohistochemistry of glutamic acid decarboxylase, choline acetyltransferase, somatostatin and parvalbumin. J Hirnforsch 32:1-17. Bredt DS, Hwang PM, Snyder SH (1990): Localization of nitric oxide synthase indicating a neural role for nitric oxide. Nature 347:768-770. Bredt DS, Glatt CE, Hwang PM, Fotuhi M, Dawson TM, Snyder SH (1991): Nitric oxide synthase protein and mRNA are discretely localized in neuronal populations of the mammalian CNS together with NADPHdiaphorase. Neuron 7:615-624. Brenman JE, Xia H, Chao DS, Black SM. Bredt DS (1997): Regulation of neuronal nitric oxide synthase through alternative transcripts. Dev Neurosci 19:224-231. Brifi6n JG, Alonso JR, Garcia-Ojeda E, Crespo C, Ar6valo R. Aij6n J (1997): Calretinin- and parvalbuminimmunoreactive neurons in the rat main olfactory bulb do not express NADPH-diaphorase activity. J Chem Neuroanat 13:253-264. Briski KP, Sy!vester PW (1999): Site-specific induction of Fos immunoreactivity in preoptic and hypothalamic NADPH-positive neurons during glucoprivation. Neuroendocrinology 69:181-190. Bruning G (1992): Localization of NADPH-diaphorase, a histochemical marker for nitric oxide synthase, in the mouse spinal cord. Acta Histochem 93:397-401. Bruning G, Wiese S, Mayer B (1994): Nitric oxide synthase in the brain of the turtle Pseudemys scripta elegans. J Comp Neurol 348:183-206. 40

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Regidor J, Edvinsson L, Divac I (1993): NOS neurones lie near branchings of cortical arteriolae. Neuroreport 4:112-114. Reuss S (1998): Nitric oxide synthase in the auditory brain stem. Neuroreport 9:3643-3646. Reuss S, Decker K, Rosseler L, Layes E, Schollmayer A, Spessert R (1995): Nitric oxide synthase in the hypothalamic suprachiasmatic nucleus of rat: evidence from histochemistry, immunohistochemistry and Western blot; and colocalization with VIP. Brain Res 695:257-262. Rhrich-Haddout F, Klosen P, Portier MM, Horvat JC (1997): Expression of peripherin, NADPH-diaphorase and NOS in the adult rat neocortex. Neuroreport 8:3313-3316. Rodrigo J, Riveros-Moreno V, Bentura ML, Uttenthal LO, Higgs EA, Fernandez AP et al. (1997): Subcellular localization of nitric oxide synthase in the cerebral ventricular system, subfornical organ, area postrema, and blood vessels of the rat brain. J Comp Neurol 378:522-534. Roskams AJ, Bredt DS, Dawson TM, Ronnett GV (1994): Nitric oxide mediates the formation of synaptic connections in developing and regenerating olfactory receptor neurons. Neuron •3:289-299. Rothe F, Canzler U, Wolf G (1998)" Subcellular localization of the neuronal isoform of nitric oxide synthase in the rat brain: a critical evaluation. Neuroscience 83:259-269. Ruan RS, Leong SK, Yeoh KH (1997): Localization of nitric oxide synthase and NADPH-diaphorase in guinea pig and human cochleae. J Hirnforsch 38:433-441. Rushlow W, Flumerfelt BA, Naus CC (1995): Colocalization of somatostatin, neuropeptide Y, and NADPHd~aphorase in the caudate-putamen of the rat. J Comp Neurol 351:499-508. Sagar SM, Ferriero DM (1987): NADPH-diaphorase activity in the posterior pituitary: relation to neuronal function. Brain Res 400:348-352. Saito S, Kidd GJ, Trapp BD, Dawson TM, Bredt DS, Wilson DA et al. (1994): Rat spinal cord neurons contain nitric oxide synthase. Neuroscience 59:447-456. Samama B, Boehm N (1996): Ontogenesis of NADPH-diaphorase activity in the olfactory bulb of the rat. Brain Res Dev Brain Res 96:192-203. Samama B, Chateau D, Boehm N (1995): Expression of NADPH-diaphorase in the rat forebrain during development. Neurosci Lett 184:204-207. Sancesario G, Morello M, Massa R, Fusco F, Bernardi G (1993): NADPH-diaphorase activity is inhibited by EDTA in neurons but not in choroid plexus epithelium. Neurosci Lett 158:101-104. Sancesario G, Morello M, Massa R, Fusco FR, D'Angelo V, Bernardi G (1996a)" NADPH-diaphorase neurons contacting the cerebrospinal fluid in the ventricles of rat brain. J Cereb Blood Flow Metab 16:517-522. Sancesario G, Reiner A, Figueredo-Cardenas G, Morello M, Bernardi G (1996b): Differential distribution of nicotinamide adenine dinucleotide phosphate-diaphorase and neural nitric oxide synthase in the rat choroid plexus. A histochemical and immunocytochemical study. Neurosciem'e 72:365-375. Sanchez F, Alonso JR, Ar6valo R, Blanco E, Aij6n J, Vazquez R (1994): Coexistence of NADPH-diaphorase with vasopressin and oxytocin in the hypothalamic magnocellular neurosecretory nuclei of the rat. Cell Tissue Res 276:31-34. Scherer-Singler U, Vincent SR, Kimura H, McGeer EG (1983): Demonstration of a unique population of neurons with NADPH-diaphorase histochemistry. J Neurosci Methods 9:229-234. Schmidt HHHW (1992): NO., CO and .OH. Endogenous soluble guanylyl cyclase-activating factors. FEBS Lett 307:102-107. Schmidt HHHW, Gagne GD, Nakane M, Pollock JS, Miller MF, Murad F (1992a): Mapping of neural nitric oxide synthase in the rat suggests frequent co-localization with NADPH-diaphorase but not with soluble guanylyl cyclase, and novel paraneural functions for nitrinergic signal transduction. J Histochem Cytochem 40:1439-1456. Schmidt HHHW, Smith RM, Nakane M, Murad F (1992b): Ca-"+ /calmodulm-dependent NO synthase type I: a biopteroflavoprotein with Ca2+/calmodulin-independent diaphorase and reductase activities. Biochemistr3' 3•:3243-3249. Scott JW, McDonald JK, Pemberton JL (1987): Short axon cells of the rat olfactory bulb display NADPHdiaphorase activity, neuropeptide Y-like immunoreactivity, and somatostatin-like immunoreactivity. J Comp Neurol 260:378-391. Seidel B, Stanarius A, Wolf G (1997): Differential expression of neuronal and endothelial nitric oxide synthase in blood vessels of the rat brain. Neurosci Lett 239:109-112. Shaer JL, Fernandez-Rodriguez P, Martinelli GP, Holstein GR (1996): NADPH-diaphorase histochemical staining in the rat vestibular nuclei during postnatal development. Am~ NY Acad Sci 781:696-699. Singer M, Lysakowski A (1996): Nitric oxide synthase localized in a subpopulation of vestibular efferents with NADPH-diaphorase histochemistry. Ann NY Acad Sci 781:658-662.

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Song ZM, Brookes SJ, Costa M (1994): Characterization of alkaline phosphatase-reactive neurons in the guinea-pig small intestine. Neuroscience 63:1153-1167. Southam E, Garthwaite J (1993): The nitric oxide-cyclic GMP signalling pathway in rat brain. Neuropharmacology 32:1267-1277. Southam E, Morris R, Garthwaite J (1992): Sources and targets of nitric oxide in rat cerebellum. Neurosci Lett 137:241-244. Spessert R, Claassen M (1998)" Histochemical differentiation between nitric oxide synthase-related and-unrelated diaphorase activity in the rat olfactory bulb. Histochem J 30:41-50. Spessert R, Layes E (1994): Fixation conditions affect the intensity but not the pattern of NADPH-diaphorase staining as a marker for neuronal nitric oxide svnthase in rat olfactory bulb. J Histo~'hem Cvtochem 42:13091315. Spessert R, Wohlgemuth C. Reuss S. Layes E {1994) NADPH-diaphorase activity of nitric oxide synthase in the olfactory bulb: co-factor specificity and characterization regarding the interrelation to NO formation. J Histochem Cvtochem 42:569-575. Spike RC, Todd AJ, Johnston HM (1993) Coexistence of NADPH-diaphorase with GABA. glycine, and acetylcholine in rat spinal cord. J Comp Neuroi 335:320-333. Stanarius A, Topel I, Schulz S. Noack H. Wolf G (1997): Immunocytochemistry of endothelial nitric oxide synthase in the rat brain: a light and electron microscopical study using the tyramide signal amplification technique. Acta Histochem 99:411-429. Sugaya K, McKinney M (1994): Nitric oxide synthase gene expression in cholinergic neurons in the rat brain examined by combined immunocytochemistry and in situ hybridization histochemistry. Brain Res Mol Brain Res 23:111-125. Szmydynger-Chodobska J, Monfils PR. Lin AY. Rahman ME Johanson CE. Chodobski A (1996): NADPHdiaphorase histochemistry of rat choroid plexus blood vessels and epithelium. Neurosci Lett 208:179-182. Takemura M, Wakisaka S, Yoshida A, Nagase Y. Bae YC. Shigenaga Y {1994): NADPH-diaphorase in the spinal trigeminal nucleus oralis and rostral solitary tract nucleus of rats. Neuroscience 61:587-595. Takemura M, Wakisaka S. Iwase K. Yabuta NH. Nakagawa S. Chen K et al. 11996) NADPH-diaphorase in the developing rat: lower brainstem and cervical spinal cord. with special reference to the trigemino-solitary complex. J Comp Neurol 365:511-525. Tang FR, Tan CK, Ling EA (1995a)" The distribution of NADPH-d in the central grey region (lamina X) of rat upper thoracic spinal cord. J Neurocvtoi 24735-743. Tang FR, Tan CK, Ling EA (1995b) Light and electron microscopic studies of the distribution of NADPHdiaphorase in the rat upper thoracic spinal cord with special reference to the spinal autonomic region. Arch Histol Cvtol 58:493-505. Tenorio F, Giraldi-Guimaraes A. Mendez-Otero R (1995)" Developmental changes of nitric oxide synthase in the rat superior colliculus. J Neutvsci Res 42:633-637. Tenorio F, Giraldi-Guimaraes A. Mendez-Otero R {1996): Morphology of NADPH-diaphorase-positive cells in the retinoceptive layers of the developing rat superior colliculus. Int J Dev Neurosci 14" 1-10. Turcanu V, Dhouib M, Poindron P (1998): Heine oxygenase inhibits nitric oxide synthase by degrading heine: a negative feedback regulation mechanism for nitric oxide production. Tran.splant Proc 30:4184-4185. Vaid RR, Yee BK, Rawlins JN. Totterdeil S t1996~: NADPH-diaphorase reactive pyramidal neurons in Ammon's horn and the subiculum of the rat hippocampal formation. Brain Res 733:31-40. Valtschanoff JG, Weinberg RJ, Rustioni A (1992): NADPH-diaphorase in the spinal cord of rats. J Comp Neurol 321:209-222. Valtschanoff JG, Weinberg RJ, Kharazia VN. Nakane M, Schmidt HH (1993a): Neurons in rat hippocampus that synthesize nitric oxide. J Comp Neurol 331:111-121. Valtschanoff JG, Weinberg RJ, Kharazia VN, Schmidt HH. Nakane M. Rustioni A (1993b): Neurons in rat cerebral cortex that synthesize nitric oxide" NADPH-diaphorase histochemistry, NOS immunocytochemistry, and colocalization with GABA. Neurosci Lett 157:157-161. Vanhatalo S, Soinila S (1995)" Nitric oxide synthase in the hypothalamo-pituitary pathways. J Chem Neutvanat 8:165-173. Verma A, Hirsch DJ, Glatt CE. Ronnett GV, Snyder SH (1993): Carbon monoxide: a putative neural messenger [see comments] [published erratum appeared in Science 1994 Jan 7:26315143)15]. Science 259:381-384. Villalba RM, Martinez-Murillo R, Blasco I. Alvarez FJ. Rodrigo J 11988): C-PON containing neurons in the rat striatum are also positive for NADPH-diaphorase activity. A light microscopic study. Brain Res 462:359-362. Villalba RM, Rodrigo J, Alvarez FJ. Achaval M. Martinez-Murillo R (1989): Localization of C-PON immunoreac-

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tivity in the rat main olfactory bulb. Demonstration that the population of neurons containing endogenous C-PON display NADPH-diaphorase activity. Neuroscience 33:373-382. Vincent SR (1994),;. Nitric oxide: a radical neurotransmitter in the central nervous system. Prog Neurobiol 42:129160. Vincent SR, Hope BT (1992): Neurons that say NO. Trends Netuvsci 15:108-113. Vincent SR, Johansson O (1983): Striatal neurons containing both somatostatin- and avian pancreatic polypeptide (APP)-like immunoreactivities and NADPH-diaphorase activity: a light and electron microscopic study. J Comp Neurol 217:264-270. Vincent SR, Kimura H (1992): Histochemical mapping of nitric oxide synthase in the rat brain. Neuroscience 46:755-784. Vincent SR, Johansson O, Skirboll L, H6kfelt T (1982): Coexistence of somatostatin- and avian pancreatic polypeptide-like immunoreactivities in striatal neurons which are selectively stained for NADPH-diaphorase activity. Adv Biochem Psychopharmacol 33:453-462. Vincent SR, Johansson O, H6kfelt T, Skirboll L, Elde RE Terenius Let al. (1983a): NADPH-diaphorase: a selective histochemical marker for striatal neurons containing both somatostatin- and avian pancreatic polypeptide (APP)-like immunoreactivities. J Comp Neurol 217:252-263. Vincent SR, Satoh K, Armstrong DM, Fibiger HC (1983b): NADPH-diaphorase: a selective histochemical marker for the cholinergic neurons of the pontine reticular formation. Neurosci Lett 43:31-36. Vincent SR, Staines WA, Fibiger HC (1983c): Histochemical demonstration of separate populations of somatostatin and cholinergic neurons in the rat striatum. Neurosci Len 35:111-114. Vincent SR, Satoh K, Armstrong DM, Panula P, Vale W. Fibiger HC (1986): Neuropeptides and NADPH-diaphorase activity in the ascending cholinergic reticular system of the rat. Neuroscience 17:167-182. Vincent SR, Das S, Maines MD (1994): Brain heme oxygenase isoenzymes and nitric oxide synthase are co-localized in select neurons. Neuroscience 63:223-231. Vincent SR, Williams JA, Reiner PB, EI-Husseini AD (1998): Monitoring neuronal NO release in vivo in cerebellum, thalamus and hippocampus. In: Mize RR. Dawson TM, Dawson VL, Friedlander MJ (Eds), P~vgress in Brain Research. Amsterdam: Elsevier, pp 27-35. Vogel M, Luck G, Bachmann S, Blottner D (1997): NOS type-1 mRNA expression and protein localization in spinal autonomic neurons. Neuroreport 8:3389-3393. Wendland B, Schweizer FE, Ryan TA, Nakane M, Murad E Scheller RH et al. (1994): Existence of nitric oxide synthase in rat hippocampal pyramidal cells. Proc Natl Acad Sci USA 91:2151-2155. Yamada K, Emson E H6kfelt T (1996): Immunohistochemical mapping of nitric oxide synthase in the rat hypothalamus and colocalization with neuropeptides. J Chem Neutvanat 10:295-316. Young HM, O'Brien AJ, Furness JB, Ciampoli D, Hardwick JR McCabe TJ et al. (1997): Relationships between NADPH diaphorase staining and neuronal, endothelial, and inducible nitric oxide synthase and cytochrome P450 reductase immunoreactivities in guinea-pig tissues. Histochem Cell Biol 107:19-29.

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CHAPTER III

Comparative and developmental neuroanatomical aspects of the NO system J.R. ALONSO, R. AREVALO, E. WERUAGA, A. PORTEROS, J.G. BRII~ON AND J. AIJON

1. I N T R O D U C T I O N Neurons labeled with NADPH-diaphorase (NADPH-d) histochemistry are widely distributed in the brain of all vertebrates. Literally thousands of papers have been published describing the characteristics of NADPH-d-positive cells in the brain, their distribution or comparison with other neuroactive substances, and their variations after experimental or pathological conditions. There are several reasons that explain why the study of these cells has attracted so much attention. First, the relatively simple NADPH-d histochemical technique allows the demonstration of specific populations and subpopulations of neurons and fibers throughout both central and peripheral nervous systems. Positive elements are frequently stained in a Golgi-like manner with considerable detail of dendritic and axonal prolongations, thus providing valuable morphological information. Then, a new "NADPH-diaphorase system' has been progressively described in the brain in parallel with a refinement of the preexistent classifications of anatomical divisions and neural populations. The second, more important and attractive reason for the interest of neuroanatomists in this histochemical staining arose when NADPH-d was identified as a marker for nitric oxide synthase (NOS, Hope et al., 1991). This makes this staining (with the technical conditions formulated below) a robust tool to identify nitric oxide-producing elements in the brain (Vincent, 1994). Nevertheless, this identification is not total and it has to be validated in each particular case (see Section 4). In the central nervous system, all three NOS isoforms (neuronal, endothelial and immunological, or I, II and III) have been detected, the neuronal isoform being the most widely expressed, but not exclusive, in neural elements. Although the NOS-positive neurons were originally described as 'solitary active cells' (Thomas and Pearse, 1961, 1964), there are brain regions where neuronal NOS is not expressed by isolated, scattered cells but by dense populations of neurons, such as the cerebellum, the olfactory bulb, or the hypothalamic supraoptic nucleus. Endothelial NOS is constitutively expressed not only in blood vessels but also in some neural cells, occasionally in the same population as neuronal NOS (Dinerman et al., 1994). While both neuronal and endothelial isoforms are constitutively expressed in the nervous system, immunQlogical NOS can be detected in neural elements only after induction of its expression in experimental or pathological situations (Endoh et al., 1994b; Minc-Golomb et al., 1996). Whereas more and more is known about the cells containing nitric oxide, important questions about the meaning of this expression remain still unsolved. Why is NOS widely

Handbook of Chemical Neuroanatom~, Vol. 17: Functional Neuroamaomy of the Nitric Oxtde S~ ~tem H.W.M. Steinbusch, J. De Vente and S.R. Vincent. editor.,, ~ 2000 Elsevier Science B.V. All rights reserved.

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distributed in the brain but is only present in discrete populations of cells? Why is it that some cells of a particular neuronal type express NADPH-d/NOS whereas cells with similar morphology, connections, and presumably functions are NADPH-d/NOS-negative? Nitric oxide is a diffusible messenger with a wide range of spatial action, then, what is the meaning of such amounts of intermingled neurites of different NOS-positive elements in the same brain region? What is the relationship between nitric oxide synthesis in neurons, glial cells and endothelial cells? Has nitric oxide produced in astrocytes the same targets as nitric oxide produced in nearby neurons? Is nitric oxide involved in the same functions in the adult and during the development? If the nitrergic system is such a simple and conserved transmitter system, why is the NADPH-d staining pattern so variable between phylogenetically close species? Any hypothesis answering these questions has to be confronted with what is known on the distribution of nitric oxide producing systems. The aim of this chapter is to provide an overview on the anatomical aspects of the distribution of nitric oxide synthase and NADPH-d activity in the nervous system of all major groups of vertebrates and during the development of the mammalian brain. First, we comment the methodological aspects concerning the localization of these neuronal markers, then we describe the general aspects of their distribution patterns in fish, amphibians, reptiles, birds and mammals, third, we report the changes during prenatal and postnatal development in the rat brain, and finally, we comment on some functional aspects of nitric oxide derived from the fine localization of markers for their synthases. We hope that this morphological information will provide baseline data for a selection of the best experimental model as well as for the design of new experiments.

2. METHODS TO LOCALIZE NITRIC OXIDE AND NITRIC OXIDE SYNTHASES IN BRAIN TISSUE

There are two main approaches to study nitric oxide production in the brain and, subsequently, two main ways to treat tissue. Firstly, it is possible to visualize directly the expression of the NOS messenger RNA, the protein, or its related NADPH-d activity using histological techniques. On the other hand, the estimation of quantitative temporal patterns of expression (in development or under experimental conditions) is difficult to estimate with these methods. Following this approach, it is possible to observe the cells containing NOS, but it is difficult to be certain about how much enzyme they contain, how much of it is active, and how much nitric oxide is really produced. The second approach is to detect and to measure nitric oxide itself, its production, and its synthases by biochemical methods using homogenates of whole brain tissue or from defined neuroanatomical regions. Although it provides a better functional representation, the anatomical resolution is much worse; it strongly depends on the quality of brain or peripheral nerve tissue dissection, that can be improved using the micropunching technique of thick tissue slices (Palkovits and Brownstein, 1988). The histological methods, especially NADPH-d histochemistry, have been fruitfully employed during last decades to determine the cellular nitric oxide sources, therefore, in the following paragraphs we shall summarize and compare them. 2.1. HISTOLOGICAL DETECTION OF NADPH-DIAPHORASE/NITRIC OXIDE SYNTHASE The immunohistochemical localization of the protein and the in situ hybridization of its mRNA are classical methods to detect any neuroactive, protein-related molecule, as NOS is. 52

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Furthermore, NADPH-d staining has been used since the sixties to label discrete populations of neurons. Bredt et al. (1991a), using an antibody to NOS from rat brain, oligonucleotide probes of rat NOS, and NADPH-d histochemistry, demonstrated that NOS protein and message are coincident with NADPH-d activity. The histochemical staining remains the most used technique to study nitrergic patterns because of its technical simplicity and valuable morphological results. NOS immunohistochemistry and NOS mRNA in situ hybridization are increasingly employed as parallel or complementary technical approaches. In addition, new methods such as the autoradiographic localization of NOS by the demonstration of the binding of its irreversible inhibitor [-~H]t,-nitro-arginine are also available (Burazin and Gundlach, 1995).

2.1.1. NADPH-diaphorase histochemistry The histochemical staining for NADPH-d is a simple technique that is accomplished in a one-step incubation: the medium usually contains a highly soluble salt of [3-NADPH, an artificial acceptor of reducing equivalents (chromogen), a detergent to permeate tissue, and a buffer with a pH of about 7.4-8.0 (Thomas and Pearse, 1961; see also Alonso et al., 1995a). The standard chromogen for optic microscopy examination is a salt of NBT (nitroblue tetrazolium), which precipitates and turns dark-blue when reduced, transforming itself into formazan (Thomas and. Pearse, 1961; Kuonen et al., 1988; Hope and Vincent, 1989). The detergent of choice is Triton X-100, which also helps to solubilize NBT. Incubation can be developed onto air-dried sections on gelatin-coated slides or with free-floating sections. The reaction runs in darkness at 37~ in 45-90 rain, and it should be controlled under the microscope because of non-specific formazan crystal accumulation. The result of this incubation is neural elements frequently stained in a perfect Golgi-like manner, even within sections of 50-100 btm thickness. Foresights have to be taken to obtain satisfactory results, otherwise imprecise staining may result: (1) aldehyde fixation is necessary to eliminate stainings non-related to NOS, (2) excessively short or prolonged fixation (e.g. stocking in formalin of autopsy material) results in a loss of visualization of weakly stained NOS-positive elements, (3) fixed material should be properly frozen and stored at -80~ for long periods, (4) time between thawing sections and their histochemical incubation should not exceed 2 weeks, even if they are stored in buffer at 4~ (5) paraffin sections are not suitable for NADPH-d staining, and (6) incubation medium should be always freshly prepared. NADPH-d histochemistry can also be resolved successfully for electron microscopic observation, by changing the chromogen. BSPT, the synonym for 2-(2'-benzothiazolyl)-5-styril-3-(4'-phthalhydrazidyl) tetrazolium chloride, turns highly electron dense and osmiophilic when reduced by accepting protons from [3-NADPH (see Wolf et al., 1992, 1993, 1995). On the other hand, BSPT needs a higher concentration of Triton X-100 or formamide to solubilize, which, together with the incubation at 37~ renders the observation of some cell details troublesome (e.g. synaptology) because of loss of membrane integrity.

2.1.2. Immunohistochemistry The fixation protocol for the histological visualization of NADPH-d activity lined above is valid for NOS immunolocalization (W6rl et al., 1994; Alonso et al., 1995a; Buwalda et al., 1995; Gonz~ilez-Hern~indez et al., 1996; Spessert and Claassen, 1998). There are available antisera for all three isoforms of the enzyme, but immunolocalization of NOS is possible only 53

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if aldehyde fixation is not excessive (Buwalda et al., 1995). Glutaraldehyde is a good fixative for electron microscopy observation, but its use results in a clear reduction in the labeling of NOS-positive elements. Immunostaining of formaldehyde-fixed and paraffin-embedded tissue is possible, but the labeling in the antisera test in our laboratory was reduced in weakly stained elements as compared with similarly fixed, cryostat sections.

2.1.3. In situ hybridization Several authors have used in situ hybridization with radioactive labeled probes to study the distribution of NOS message in neural tissue (Bredt et al., 1991a; Kishimoto et al., 1993). These antisense oligoprobes are normally formed by 40-50 nucleotides sequences fitting with a conserved region in both human and rat. These terminal-labeled oligoprobes were employed in the first studies (Dawson et al., 1991), but thereafter, radioactive or non-radioisotopic riboprobes of higher specificity and sensitivity have been used (Endoh et al., 1994a; Iwase et al., 1998a). 2.2. BIOCHEMICAL DETECTION OF NITRIC OXIDE SYNTHASE, NITRIC OXIDE PRODUCTION, AND NITRIC OXIDE PROPER Nitric oxide is formed by the stoichiometric conversion of L-arginine to L-citrulline and NO through an oxidative-reductive pathway. Benefitting on the change in net charge of both amino acids, Bredt and Snyder (1990) developed a simple assay for NOS catalytic activity based on the conversion of [3H]L-arginine to [3H]L-citrulline followed by separation of both substances over an anion-exchange column. The possibility of detection and measure of the message and protein of NOS (or any protein) by Northern and Western blot, respectively, is obvious in most biochemical laboratories. Measuring of nitrites/nitrates of fresh tissue homogenates allows indirect detection of nitric oxide, as it is oxidized to those metabolites by different cellular pathways (see Crow and Beckman, 1995). Nitrates can be reduced to nitrites by a one-step enzymatic reaction catalyzed by nitrate reductase, using FAD and NADPH as cofactors (Bories and Bories, 1995). Nitrites are easily detected by Griess reaction (Green et al., 1982). Currently, both primary nitrites and enzymatic reduced nitrates are used as indicators of total concentration of nitric oxide in a tissue extract (Ta~kiran et al., 1997; Yamada and Nabeshima, 1997). Other methods to directly measure nitric oxide in vivo or in vitro are electronic paramagnetic resonance using nitric oxide-high-affinity nitrous complexes (Kubrina et al., 1992; Xia and Zweier, 1997) or by differential normal-pulse voltammetry using porphyrinic microsensors. With the latter technique, it has been claimed that it is possible to measure in vivo levels of extracellular nitric oxide (Desvignes et al., 1997).

3. RESULTS The number of published studies on the distribution of NADPH-d/NOS-positive cells is very varied from one group of vertebrates to another. The following results are mainly based on results and collaborations from our group carried out in teleosts (Ar6valo et al., 1995), amphibians (Gonz~ilez et al., 1996; Mufioz et al., 1996), reptiles (Smeets et al., 1997), birds (Panzica et al., 1994) and mammals (Ar6valo et al., 1992; Alonso et al., 1993, 1995b, 1998; Porteros et al., 1994; Brifi6n et al., 1998; Weruaga et al., 1998). These results are compared 54

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with other articles, usually in different species of the same group although the technical procedures are normally very similar. We describe the general characteristics of the labeling in each major vertebrate group following the main brain subdivisions. 3.1. FISH

3.1.1. Cyclostomes The presence of NADPH-d activity has been described in the central nervous system of the larval lamprey Lampetra planeri (Schober et al., 1994) and in the olfactory mucosa of the larval sea lamprey Petromyzon marinus (Zielinski et al., 1996). In the brain of larval Lampetra planeri, NADPH-d labeling is observed in the olfactory fibers and glomerular layer of the olfactory bulb. NADPH-d staining in the telencephalon is exclusively located in fibers. In the diencephalon, NADPH-d activity appears in the pineal organ, the parapineal organ, the habenula and in the subcommissural organ. NADPH-d activity is present in the giant cells of the cyclostome mesencephalon and brainstem. These cells are reticulospinal neurons located in the mesencephalon, the aqueductalisthmic region and in the anterior portion of the spinal cord. Other groups of neurons appear in the vagal region and in the spinal cord. The distribution of NADPH-d has also been studied in the brain of the adult lamprey (Pombal and Gonz~ilez, pers. commun., 1999). The distribution pattern of NADPH-d in the adult brain shows significant differences from what has been described in the larval brain. NADPH-d-positive cells appear within the pallium, the thalamus, the habenula, the hypothalamus, tectal and pretectal areas, the isthmus, the formatio reticularis of the rhombencephalon, and in the spinal cord.

3.1.2. Teleosts Using the histochemical technique, NADPH-d reactivity has been demonstrated in the brains of cyprinid teleosts: the tench Tinca tinca (Ar6valo et al., 1995) and the goldfish Carassius auratus (Brtining et al., 1995" Villani and Guarnieri, 1995); in the brain of an atherinomorph teleost, the swordtail fish Xiphophorus helleri (Anken and Rahmann, 1996); in the electrosensory and electromotor systems of a gymnotiform teleost, Apteronotus leptorhynchus (Turner and Moroz, 1995); in a salmonid teleost, the Atlantic salmon Salmo salar (Holmqvist et al., 1994) and studies restricted to the diencephalon of the rainbow trout Oncorhynchus mykiss (Schober et al., 1993), to the Mauthner cells of the tench, (Crespo et al., 1998a) and goldfish (Bell et al., 1997) and to the retina of the Atlantic salmon (Ostholm et al., 1994). Telencephalon. In the olfactory bulb of the tench, all olfactory fibers and glomeruli are positive for NADPH-d. However, NADPH-d appears to be absent from the olfactory bulb of the goldfish and the swordtail fish. Within the telencephalic hemispheres, the NADPH-d staining follows dorsoventral and rostrocaudal gradients. In all teleosts studied, NADPH-d-positive cells are seen in the area dorsalis telencephali pars centralis and the area dorsalis telencephali zona posterior (Fig. 1a,b). In addition, positive cells are observed in the area dorsalis telencephali pars lateralis in the goldfish and in the area dorsalis telencephali pars medialis in the swordtail fish and goldfish. A larger number of NADPH-d-stained cells appears in the area ventralis telencephali in tench and swordtail fish. They are found in the pars ventralis, lateralis, centralis and 55

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Fig. I. Camera lucida drawings of coronal sections from rostral (a) to caudal Ik) levels through the brain of the tench, Tinca tinca.

supracommissuralis (Fig. la). In goldfish, Villani and Guarnieri (1995) do not report positive cells in the ventral telencephalon, whereas Braining et al. (1995) find positive cells in the pars ventralis, centralis and supracommissuralis. 56

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In the telencephalon, NADPH-d-positive fibers are only described in the commissura anterior of the tench. Diencephalon. In the swordtail fish, NADPH-d-positive cells are observed within the nucleus entopeduncularis, which does not contain the enzyme in the tench. In the preoptic region, NADPH-d-positive cells are observed in the nucleus magnocellularis praeopticus (Figs. lb,c and 2a). In the epithalamus, positive neurons in the habenula are missing both in the Atlantic salmon and in the tench, whereas some are labeled in the goldfish. The thalamus contains positive cells in the nucleus centralis posterior of the goldfish and the swordtail fish. NADPH-d staining appears in the posterior tuberculum within the nucleus periventricularis tuberculi posterioris, the corpus mammillare, the nucleus tuberis posterior and the torus lateralis (Fig. l d-g). The preglomerular complex and the lobus inferior hypothalami only contain positive cells in the swordtail fish. Scarce NADPH-d staining is found in the hypothalamus of the tench. Positive cells can be seen in the nucleus periventricularis hypothalami and in the nucleus tuberis anterior (Fig. l e g). In the nucleus fasciculi longitudinalis medialis, and close to the fasciculus retroflexus and commissura posterior, positive neurons are demonstrated in the tench (Fig. l e,f) and in the swordtail fish. In the nucleus praetectalis superficialis, NADPH-d-stained cells are seen in the swordtail fish and in the goldfish. Mesencephalon. Within the tectum opticum, the largest part of the mesencephalon, a few NADPH-d-positive cells are found in the strata periventriculare, album centrale and griseum centrale. The tractus opticus shows a population of stained cells appearing between unstained fibers (Fig. l b-d). Positive cells are also observed in the torus semicircularis of all teleosts studied (Fig. 1f,g). Within the tegmentum, the nucleus nervi oculomotorii and the nucleus nervi trochlearis (Fig. 2b) are positive in the swordtail fish and in the tench. However, NOS is not observed in the nucleus nervi oculomotorii of the goldfish. Moreover, in the mesencephalon of the tench, NADPH-d-positive cells are detected in the nucleus Edinger-Westphal, in the torus longitudinalis, in the nucleus tegmentalis rostralis and in the formatio reticularis mesencephali (Fig. lf-h). In the mesencephalon of swordtail fish, labeled cells are located within the nucleus mesencephalicus nervi trigemini, the nucleus perilemniscaris, the nucleus ruber and the nucleus isthmi. Rhombencephalon. We observed a scarce NADPH-d staining in the tench cerebellum. Occasionally, some labeled cells are observed in its molecular layer (Fig. l i). In Apteronotus leptorhynchus, NADPH-d activity is present in Purkinje cell somata, mossy fiber synaptic glomeruli, granule cells, and parallel fibers both in the corpus cerebelli and in the valvula cerebelli. NADPH-d staining is also present in the Purkinje cell layer of the swordtail fish but the stained cells are not Purkinje cells. The highest number of NADPH-d-positive cells in the brain of teleosts is observed within the myelencephalon. The patterns of distribution of NADPH-d activity in all studied teleosts are very similar. NADPH-d staining is mainly located in the nuclei of the cranial nerves. Thus, we found strongly labeled neurons within the nucleus nervi trigemini, the nucleus nervi abducentis, the nucleus nervi facialis, the nucleus nervi glossopharyngei and the nucleus nervi vagi (Figs. l i - k and 2c,e). In the octaval area, NADPH-d-labeled cells are observed in the nucleus 57

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octavus anterior, the nucleus octavus magnocellularis, the nucleus octavus descendens, the nucleus octavolateralis medialis and in the population octavia secundaria (Figs. l i and 2d). In the goldfish, all these nuclei have NADPH-d staining but do not contain NOS (Brtining et al., 1995). Positive cells are also seen in the nucleus fasciculi solitarii, the radix descendens nervi trigemini, the nucleus tangentialis, the columna motoria spinalis and the formatio reticularis (Figs. l h - k and 2f). In the lobus vagi and lobus facialis, some scattered small cells are observed (Fig. lj-k). The Mauthner cells are also stained for NADPH-d. In all subdivisions of the tench brain within tracts and nerves we find NADPH-d-stained cells whose morphological aspect resembled those of oligodendrocytes. Spinal cord. NADPH-d reaction is present in many structures in the spinal cord. In the tench, several cell populations and fibers are observed in the cornu ventrale as well as in the lateral zone of the dorsal spinal cord. However, NADPH-d-stained neurons are present both in the cornu ventrale and cornu dorsale in the goldfish. In addition, many ascending and descending fibers in the white matter, and radial glial fibers are labeled in the goldfish. 3.2. AMPHIBIANS NADPH-d activity has been confirmed in the retina (Sato, 1990b), pineal organ (Sato, 1990a), hypothalamo-hypophysial system (Prasada-Rao et al., 1997), peripheral nervous system of anuran amphibians (Li et al., 1992), and in the olfactory bulb of anuran and urodele amphibians (Porteros et al., 1996). The early fragmentary data of NADPH-d activity in amphibians have become more complete with comprehensive studies on the overall distribution of NADPH-d in the brain of two anuran amphibians, Rana perezi (Mufioz et al., 1996) and Xenopus laevis (Brtining and Mayer, 1996), and in the brain of a urodele amphibian, Pleurodeles waltl (Gonz~,lez et al., 1996). Telencephalon. In the main olfactory bulb of studied amphibians, NADPH-d staining is observed in all olfactory fibers (Fig. 4a) except for Xenopus laevis where only a fraction of fibers is labeled. In the accessory olfactory bulb, vomeronasal fibers are NADPH-d-positive in urodele amphibians but they are NADPH-d-negative in anuran amphibians. NOS-immunoreactivity is not observed in the primary afferents (Porteros et al., 1996). Granule cells are NADPH-d-positive and NOS-immunopositive in the main and accessory olfactory bulb of anures (Figs. 3a and 4b), and in the main olfactory bulb of urodeles. All pallial and subpallial regions of the telencephalon posses a large population of NADPHd-positive cells (Fig. 3a-c). In the ventromedial wall of the hemispheres, NADPH-d-positive neurons are observed in anuran amphibians. They are located in the nucleus of the diagonal band of Broca and in the nucleus accumbens (Fig. 4c), as well as in the septum of all studied amphibians. NADPH-d-labeled cells are also distributed within the dorsal and ventral portions of the striatum, and in the amygdala pars lateralis (Fig. 3b,c). Diencephalon. A high NADPH-d activity has been found throughout the diencephalon of amphibians, both in its dorsal and ventral subdivisions.

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Fig. 2. Micrographs of 30 lam thick coronal sections of the tench brain (Tinca tinca) stained with NADPHdiaphorase histochemical technique. (a) Nucleus praeopticus magnocellularis. (b) Nucleus nervi trochlearis. (c) Nucleus nervi facialis. (d) Area octavolateralis. (e) Nucleus nervi glossopharyngei. (f) Spinal cord fascicles. Scale bars are 100 ~tm.

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Fig. 3. Camera lucida drawings of coronal sections from rostral (a) to caudal (j) levels through the brain of the frog, Rana pere:i.

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Dorsally in the diencephalon, moderate groups of NADPH-d-positive cells are located in the nucleus habenulae pars dorsalis and ventralis, and dense fibers are found only in the right nucleus habenulae pars dorsalis of the anures. By contrast, no NADPH-d-positive cells are found in the nucleus habenulae of urodeles, where NADPH-d-positive fibers innervate exclusively its left side component. Numerous NADPH-d-positive cells are located in the preoptic area around the recessus praeopticus, in the anterior preoptic area and in the caudal region of the preoptic area, within the nucleus magnocellularis. In the hypothalamus, large cells appear in the infundibular region, in the caudal portion of the nucleus suprachiasmaticus (Fig. 3d) and in the nucleus tuberculi posterioris. Occasionally, a few CSF-contacting cells are observed in the nucleus of the periventricular organ of Rana perezi; however, comparable cells are not found in other anurans nor in urodeles. The thalamus of the frog demonstrates a large population of NADPH-d-positive neurons located in the nuclei anterior, centralis, lateralis anterior and lateralis posterior thalami (Fig. 3d). In contrast, the urodele thalamus lacks NADPH-d-labeled neurons. The most intensely labeled areas of the thalamus correspond to visual centers, where strongly labeled fibers coming from the retina or the tectum opticum can be observed: the neuropil of Bellonci, corpus geniculatum thalamicum, and nucleus superficialis ventralis thalami. Scattered NADPH-d-labeled cells are located in the pretectal area within the nucleus commissurae posterioris and the griseum praetectale (Fig. 3e). NADPH-d-positive fibers appear in the nucleus lentiformis, in the area above the nucleus commissurae posterioris and crossing the commissura tecti and the commissura posterior. Among all studied amphibians, NADPH-d activity is found in the pituitary gland and median eminence only in the frog Rana esculenta (Prasada-Rao et al., 1997). Mesencephalon. Numerous NADPH-d-positive cell bodies are found in the mesencephalon, both in the tectum and in the tegmentum. In the tectum opticum, most labeled cells are found in all layers of the periventricular gray (Fig. 3e,f). These cells have a primary process that ascends perpendicular to the brain surface and arborizes in the deep portion of the tectal fiber layers (Fig. 4d). Occasionally, scattered NADPH-d-stained cells are found in the nucleus mesencephalicus nervi trigemini, these cells being NOS-immunonegative (Gonzfilez et al., 1996). In anuran amphibians all subnuclei of the toms semicircularis demonstrate NADPH-d-positive cell bodies as well as varicose fibers and axon terminals (Fig. 3f). The mesencephalic area of the frog that demonstrates the most strongly labeled cells is the tegmental region. They are located in the nuclei tegmentalis and the nucleus profundus mesencephali (Fig. 3e). Moderately labeled cells are found in the nucleus fasciculi longitudinalis medialis. The nucleus interpeduncularis is densely innervated by NADPH-d-labeled axons. In the isthmic region of the frog, a group of NADPH-d-positive cells is located medial to the nucleus isthmi (Fig. 4f). In this latter nucleus, a peculiar feature is a group of labeled terminals that can be related to the tecto-isthmal projection that arises from cells in the tectum opticum. Rhombencephalon. The population of NADPH-d-positive cells in the frog rhombencephalon is very abundant. The cerebellum has scattered and moderately stained cells in the molecular and granule cell layers (Figs. 3g and 4e). In the granule cell layer, NADPH-d-positive fibers are also present, they arborize profusely and have varicose swellings. The majority of labeled cells in the rhombencephalon are found in the reticular nuclei but other structures also demonstrate labeled cells: the nuclei sensori nervi trigemini, the nuclei 61

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raphe and the nucleus tractus solitarius (Fig. 3h,i). The motor nuclei of the cranial nerves in the rhombencephalon do not stain for NADPH-d. The octaval nuclei in the frog Rana perezi have stained cell bodies. In this region of the brain, among the numerous labeled fibers, the area of the nucleus olivae superioris is particularly well stained. In addition, the radix descendens nervi trigemini and the tractus solitarii have NADPH-d-positive fibers, and dispersed fibers course caudally in the ventrolateral aspect of the rhombencephalic tegmentum. A population of NADPH-d-positive cells is found in the transition zone between the rhombencephalon and the spinal cord, within the nucleus columna dorsalis. Spinal cord. A large population of densely labeled cells is found from cervical to lumbar levels of the spinal cord (Fig. 3j). Almost all fields of the gray matter have cells showing NADPH-d activity. A dense NADPH-d-positive neuropil can be seen in the dorsal field of the gray matter, principally in the cervical segments. At most caudal levels, abundant labeled fibers course in the funiculus ventralis and in the funiculus lateralis. 3.3. REPTILES The distribution of NADPH-d has been studied in the central nervous system of three species belonging to different groups within the class of Reptilia: the turtle Pseudemvs scripta elegans (Brtining et al., 1994b); the lizard Gekko gecko (Smeets et al., 1997); and the snake Trimeresurus flavoviridis (Jiang and Terashima, 1996). Other studies in reptiles are restricted to the brainstem (Luebke et al., 1992) and to the hippocampus (D~ivila et al., 1995) of lizards. Telencephalon. In the olfactory bulb, the NADPH-d staining is located in a scarce number of weakly stained cell bodies in the internal granular layer, in the inner plexiform layer and in the mitral cell layer. The NADPH-d-positive fibers are very limited in number and intensity and they are dispersed throughout all layers of the olfactory bulb in lizards. In the snake and turtle olfactory bulbs, the olfactory/vomeronasal nerve, glomeruli, and periglomerular cells in the accessory olfactory bulb are NADPH-d-positive. The cortex of turtles and lizards contains a limited number of NADPH-d-positive cell bodies and fibers located in the large-celled portion of the cortex medialis, the cortex dorsalis, the cortex lateralis anterior and the cortex lateralis posterior (Fig. 5a-c). However, the cortex of the snake is NADPH-d-negative. The basal forebrain contains numerous NADPH-d-positive cell bodies and fibers located in the nucleus olfactorius anterioris, the tuberculum olfactorium, the nucleus accumbens, the pallial thickening and mainly, in the dorsal ventricular ridge, the striatum and the amygdaloid complex (Figs. 5a-c and 6a,d). The distribution of the neuronal elements is highly heterogeneous; variations in density of cell bodies and fibers are observed not only between shell and core regions, but also between medial and lateral, as well as between rostral and caudal portions. In contrast to the numerous stained cells in the striatum, in the nucleus

Fig. 4. NADPH-d labeling in the brain of the frog, Rana perezi. (a) Intensely stained glomeruli (G) and varicose centrifugal fibers (arrows) in the main olfactory bulb. (b) NADPH-d staining pattern in the accessory olfactory bulb. Note the absence of staining in the vomeronasal fibers and glomeruli of the stratum glomerulare and the intense staining in granule cells in the stratum granulare and their dendrites in the stratum plexiforme externum. (c) Labeled cells in the diagonal band of Broca. (d) Stained monopolar cells and neuropil in the tectum opticum. (e) Weakly NADPH-d-reactive granule cells in the cerebellum. (0 Strongly stained cells in the limits of the nucleus isthmi. Scale bars: 50 ~tm in a, 100 Ism in b and e, 25 Ixm in c, d and f.

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Fig. 5. Camera lucida drawings of coronal sections from rostral (a) to caudal (tl) levels through the brain of the lizard Gekko gecko.

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accumbens of Gekko gecko only a few stained cells are observed. Such a distinction between nucleus accumbens and striatum is not clearly delimited in turtles. In the septum, NADPH-d-positive cell bodies and fibers are distributed heterogeneously. In Gekko gecko, NADPH-d staining is observed in the caudalmost portion of the nucleus septalis anterior, the nucleus septalis medialis, the nucleus septalis lateralis, the nucleus septalis dorsalis, the nucleus of the diagonal band of Broca and the fasciculus medialis telencephali (Figs. 5b-d and 6b). In Pseudemys, NADPH-d-positive cells are only described in the horizontal limb of the diagonal band of Broca. Diencephalon. In lizards, NADPH-d-stained cells are observed at the level of the commissura anterior within the area praeoptica periventricularis (Fig. 6c), dorsally to the chiasma opticum, the nucleus periventricularis hypothalami and in the dorsal portion of the area praeoptica. At postcommissural levels, stained cells occur in the nucleus periventricularis hypothalami, the nucleus medialis posterior thalami, nucleus dorsolateralis thalami pars magnocellularis, nucleus suprapeduncularis and in the area lateralis hypothalami (Fig. 5e). The staining in the diencephalon has similar features in turtles and lizards; however, notable differences are observed in the thalamus and pretectum. Thus, these regions are in Pseudemys essentially devoid of NADPH-d-stained cells whereas several distinct cell groups occur in Gekko. At caudalmost levels of the lizard diencephalon, NADPH-d-positive cells are located in the nucleus lentiformis mesencephali, the nucleus praetectalis ventralis, the nucleus dorsolateralis thalami and in the nucleus profundus mesencephali (Fig. 5e), all of them NADPH-d-negative nuclei in the turtle. Fiber staining was essentially identical in the diencephalon of the studied reptiles. NADPHd-positive fibers are demonstrated in the fasciculus medialis telencephali, the stria medullaris, the nucleus dorsolateralis thalami, the nucleus dorsomedialis thalami, the nucleus medialis thalami, the corpus geniculatum lateralis and in the ganglion habenulae. The nucleus rotundus is characterized by the almost complete lack of NADPH-d-positive fibers. The chiasma opticum and the tractus opticus contain labeled fibers. Mesencephalon. A considerable number of NADPH-d-stained cells and fibers is observed in the tectum and tegmentum mesencephali. In the optic rectum, NADPH-d-positive cells and fibers appear in all tectal layers but most are located in the periventricular zone (Figs. 5e,f and 6e). In the tegmentum, the distributions of NADPH-d in turtles and lizards are similar. NADPH-d-positive cell groups are observed in the area tegmentalis ventralis, the substantia nigra pars compacta, the retrorubral area, the nucleus laminaris of the torus semicircularis and in the nucleus intercollicularis. Additionally, stained cells are scattered throughout the formatio reticularis and in the nucleus centralis of the torus semicircularis (Fig. 5f, g). Stained fibers are widely distributed throughout the tegmentum but are especially abundant in the substantia nigra pars reticulata and around the fasciculus longitudinalis medialis. In both Gekko and Pseudemys, the mesencephalic trigeminal neurons are NADPH-d-positive, but NOS-immunonegative. Rhombencephalon. Weakly stained cells are also found scattered through the molecular and granular layers of the cerebellum in lizards. In the snake, NADPH-d staining is not observed in the cerebellum. No obvious differences are distinguished in the distribution of NADPH-d in the rhombencephalon of turtles and lizards. In both groups, NADPH-d-positive cells are present in the locus coeruleus, the reticular nuclei, the cerebellar nuclei, the nucleus vestibularis dorsolateralis, the nucleus descendens nervi trigemini, the nucleus tractus solitarii, the nucleus 65

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perihypoglossum, the tegmentum ventrolaterale and in the nucleus isthmi pars parvocellularis (Fig. 5g,h). A marked difference is observed in the nucleus raphe inferior, which in Gekko contains numerous NADPH-d-positive cells (Fig. 6t-) and in Pseudemvs is not stained. In both species, the densest plexuses of NADPH-d-positive fibers are found in the nucleus isthmi pars parvocellularis and in the locus coeruleus. A dense accumulation of NADPH-d-positive neurons is also found in the locus coeruleus of the snake Trimeresurus. Spinal cord. NADPH-d-positive cell bodies and fibers are found throughout the spinal cord of Gekko, although the number and intensity of staining are variable at different levels. The number of cells and fibers is higher at rostral levels than at caudal ones. In the snake spinal cord, NADPH-d-stained fibers are seen in the radix dorsalis, Lissauer's tract, cornu dorsale and in the funiculus dorsalis. The spinal cord was not analyzed in the turtle. 3.4. BIRDS The distribution of NADPH-d activity has been described in the central nervous system of different species: the chicken Gallus domesticus (Brtining, 1993; BrOning et al., 1994a; S~inchez et al., 1996; Von Bartheld and Schober, 1997); the Japanese quail Coturnixjaponica (Panzica et al., 1994, 1996; S~.nchez et al., 1996); the budgerigar Melopsittacus undulatus (Cozzi et al., 1997); and the zebra finch Taeniopygia guttata (Wallh~iusser-Franke et al., 1995). The general pattern of distribution of NADPH-d shows similarities in all avian species studied. Telencephalon. This region contains a wide population of NADPH-d-positive cells: only few regions of the telencephalon are totally devoid or provided with few positive elements. There are marked differences in the distribution and staining intensity of the labeling, particularly NADPH-d-positive fibers, between the different regions. The olfactory bulb contains only a few NADPH-d-positive fibers in the internal granular layer of the chicken and the quail. Some stained neurons are also observed in the granular layer of the chicken olfactory bulb. Other regions of the olfactory pathway show a medium to high density of NADPH-d-positive processes and neurons: the tuberculum olfactorium, the cortex piriformis, the nucleus taeniae, the stria medullaris and the habenula. The paleostriatal-paraolfactory lobe complex, which is the avian homologue of the mammalian striatal complex, is the region that shows the highest number of NADPH-d-positive neurons and fibers (Fig. 7a-c). The paleostriatum primitivum (Fig. 8a), the paleostriatum augmentatum (Fig. 8c) and the paleostriatum ventrale (Fig. 8b) demonstrate a large number of neurons and fibers stained for NADPH-d. Scattered labeled neurons are observed in the neostriatum and hyperstriatum where a dense network of fibers stained for NADPH-d can also be seen. NADPH-d activity is absent in the ectostriatum. Neurons and fibers are more numerous along the lines subdividing the avian telencephalic regions (lamina frontalis superior, lamina medullaris dorsalis and lamina hyperstriatica) and close to the lateral wall of the telencephalic ventricles. The NADPH-d staining in the striatal structures shows no differences in all'studied avian species.

Fig. 6. Photomicrographs of transverse sections through the brain of the lizard Gekko gecko showing NADPH-dpositive neurons and fibers in the striatum (a), the septum mediale Ib), the area praeoptica periventricularis (c), the nucleus externus amygdalae and the nucleus centralis amygdalae (d), the stratum griseum periventriculare of the tectum opticum (e), and in the nucleus raphe inferior (]3. Scale bars are 100 Itm.

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Fig. 7. Camera lucida drawings of coronal sections from rostral (a) to caudal (i) levels through the brain of the Japanese quail, Cotumix japonica.

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Fig. 8. Photomicrographs of transverse sections through the brain of the Japanese quail, Coturnirjaponica, showing NADPH-d-positive neurons and fibers in the paleostriatum primitivum (a), the paleostriatum ventrale (b), the paleostriatum augmentatum (c), the nucleus pedunculopontinus tegmenti (d), the stratum griseum periventriculare of the tectum opticum (e), and in the nucleus motorius nervi trigemini q). Scale bars are 100 ~tm. 69

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The septal region in the quail has several cell clusters embedded in an extremely dense positive neuropil mainly located in the nucleus septalis medialis, the nucleus septalis lateralis and in the nucleus commissurae pallii (Fig. 7b). In the budgerigar, the only nucleus in the septal region showing NADPH-d activity is the nucleus commissurae pallii. The hippocampus is totally devoid of NADPH-d-positive cells in the quail and in the chicken, but in the budgerigar, stained neurons are observed both in the hippocampus and in the region between the area parahippocampalis and the hippocampus. Diencephalon. The NADPH-d staining in the diencephalon is limited to a few cell clusters, and a diffuse, low-intensity network of varicose fibers. The pattern of distribution is comparable in all avian species. In the area praeoptica the positive elements are located close to the tractus septomesencephalicus, in the nucleus praeopticus dorsalis, and in the nucleus praeopticus medialis (Fig. 7b). In the epithalamus, the nucleus habenularis lateralis contains numerous NADPH-d-positive fibers, whereas in the nucleus habenularis medialis both fibers and a few NADPH-d-positive neurons can be observed. In the thalamus, NADPH-d-positive neurons are found in the nucleus dorsomedialis anterior thalami, the nucleus dorsolateralis anterior thalami, periventricularly and dorsally to the nucleus paraventricularis magnocellularis, the nucleus dorsomedialis posterior, the nucleus ovoidalis, the nucleus praetectalis medialis and around the ventral aspect of the nucleus rotundus (Fig. 7c). A high density of positive fibers is also present in the nucleus dorsolateralis anterior thalami and in the nucleus lateralis thalami. Neurons are stained for NADPH-d in the hypothalamus within the nucleus anterior medialis hypothalami and the nucleus suprachiasmaticus, pars lateralis (Fig. 7b,c). Many varicose, positive fibers appear within the decussatio supraoptica and the nucleus geniculatus lateralis, pars ventralis. The larger hypothalamic groups are located within the tuberoinfundibular regions, which are embedded in a dense network of positive fibers and puncta. These regions in the budgerigar display a smaller number of positive cells surrounded by fibers. The nucleus mammillaris shows NADPH-d-positive neurons in the chicken and the quail (Fig. 7d), whereas it is unstained or weakly stained in the budgerigar. Mesencephalon. The pattern of distribution of NADPH-d-positive neurons and fibers in the tectum opticum is quite complex. The strongest reaction and the largest number of positive neurons are observed in the stratum griseum periventricularis. In addition, scattered neurons are observed in many layers of the stratum griseum and fibrosum superficiale and in the stratum griseum centrale (Fig. 8e). Thin, punctate fibers cover all these strata. Scattered neurons are visible in the nucleus intercollicularis, which also shows positive neuropil (Fig. 7d). At mesencephalic levels, NADPH-d staining is mainly located within the nucleus pedunculopontinus tegmenti (Fig. 8d) and the area ventralis tegmentalis. Scattered neurons also appear in the substantia grisea centralis, the formatio reticularis medialis mesencephali, the nucleus mesencephalicus nervi trigemini and in the nucleus isthmoopticus. The nucleus interpeduncularis and the raphe region contain a moderately positive neuropil and a few NADPH-d-positive neurons (Fig. 7e). Rhombencephalon. In the cerebellum, the granule cells show a weak reaction, whereas the Purkinje cells are unstained (Fig. 7e-i). NADPH-d-stained fibers are present in the molecular layer as well as in the cerebellar nuclei. A wide population of NADPH-d-positive cells are located in the locus coeruleus and scattered neurons appear within the nucleus subcoeruleus dorsalis and the nucleus subcoeruleus 70

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ventralis (Fig. 7f), which are embedded in a high density neuropil formed by thin varicose fibers and puncta. Other rhombencephalic regions with NADPH-d-stained cells are the nucleus pontis medialis and lateralis, the vestibular nuclei, the nucleus paragigantocellularis, the nucleus tractus solitarii and the formatio reticularis (Fig. 7g-i). Large neurons are also clustered laterally to the fasciculus longitudinalis medialis (Fig. 7h). NADPH-d-positive fibers are also observed within the vestibular nuclei and in the nucleus tractus solitarii. A high number of positive puncta surrounding unstained cells is observed in the nucleus laminaris. The encephalic motor nuclei are generally unstained, only a few, small neurons are observed in the nucleus nervi glossopharyngei et motorius nervi vagi (Fig. 7i) and in the nucleus nervi trigemini (Fig. 8f). Spinal cord. There are no available data on distribution of NADPH-d in the avian spinal cord. 3.5. MAMMALS Detailed mapping studies of the distribution of NADPH-d and/or NOS are available for mammals (cat: Mizukawa et al., 1989; rat: Vincent and Kimura, 1992, Rodrigo et al., 1994; human: Egberongbe et al., 1994). In addition, there are studies available on the presence of NADPH-d activity restricted to different regions of the nervous system such as the main olfactory bulb (Alonso et al., 1993, 1995b, 1998; Kishimoto et al., 1993; Brifi6n et al., 1998; Weruaga et al., 1998), accessory olfactory bulb (Porteros et al., 1994), nucleus olfactorius anterior (Garcfa-Ojeda et al., 1994), cerebral cortex (Mizukawa et al., 1988), medial septal complex (Kinjo et al., 1989), striatum and nucleus accumbens (Kowall et al., 1985), amygdala (Sims and Williams, 1990), hippocampal formation (Mufson et al., 1990), hypothalamus (Arrvalo et al., 1992), caudatus putamen (Sandell et al., 1986), colliculus superior (Wallace and Fredens, 1989), nucleus pedunculopontinus (Skinner et al., 1989), pontomesencephalic region (Nakamura et al., 1988), nucleus nervi vagi (Gonz~ilez et al., 1987), reticular system (Vincent et al., 1986), and spinal cord (Valtschanoff et al., 1992; Pullen et al., 1997), among others. The distribution of NADPH-d in the mammal brain is relatively similar in all studied species, although it is important to note that most studies are in rodents, and even in relatively closely related species some differences have been found. Telencephalon. In the main olfactory bulb of rodents, a subpopulation of olfactory fibers, and a subpopulation of olfactory glomeruli, located on the dorsomedial side, are positive for NADPH-d (Fig. 10a). In contrast, in the main olfactory bulb of primates all olfactory fibers and glomeruli are NADPH-d-positive. The highest number of positive cells is observed in the glomerular layer in rodents and insectivores. Surrounding both the NADPH-d-positive and NADPH-d-negative glomeruli, there are NADPH-d-positive periglomerular cells (Figs. 9a and 10a). Only a few periglomerular cells are positive in the primate olfactory bulb. Two types of short axon cells are positive for NADPH-d in the rodent and primate olfactory bulb. They are located at the boundary of the glomerular and external plexiform layers. In contrast, in the hedgehog, the superficial short-axon cells are not stained. An other interspecies difference is the presence and relative frequency of NADPH-d-stained cells in the external plexiform layer. These cells are rare in the hedgehog, rodents and primates, whereas they are more abundant in the hamster. In the deep portion of the olfactory bulb, several types of NADPH-d-positive neurons have been described: deep short-axon cells and granule cells in all studied species, horizontal and vertical neurons in rodents and in the hedgehog, and stellate cells in primates. A discrepancy between the NADPH-d and NOS stainings has been found 71

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i Fig. 9. Camera lucida drawings of coronal sections from rostral (a) to caudal (i) levels through the brain of the adult rat, Rattus norvegicus.

in the olfactory fibers. Whereas the NADPH-d and NOS labeling is coincident in all neural types, no NOS-immunoreactivity is detected in the olfactory fibers and glomeruli (Kishimoto et al., 1993; Hopkins et al., 1996: Alonso et al., 1998; Weruaga et al., 1998). In the accessory olfactory bulb, all vomeronasal fibers and glomeruli are NADPH-dlabeled. NADPH-d-positive neurons are identified as periglomerular cells in the glomerular 72

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layer and external plexiform layer, horizontal cells in the internal plexiform layer, and granule cells and deep short-axon cells in the granule cell layer. The labeled dendrites of the granule cells form a dense neuropil in the granule cell layer, internal plexiform layer and external plexiform layer. NADPH-d-positive neurons and fibers are located in the nucleus olfactorius anterior within the lateral, dorsal, ventral, medial, posterior and external subdivisions. The transition areas of the nucleus olfactorius anterior show the same NADPH-d activity distribution patterns as those of the corresponding subdivisions. The corpus callosum is crossed by positive fibers and contains a few stained neurons (Fig. 9b-d). Large neurons strongly stained for NADPH-d are observed in the cerebral cortex (Fig. 9bh). They are located through the neocortex in layers II-VI and have been identified as Martinotti cells. The highest density of NADPH-d-positive cells in the cortex is observed at the junction between gray and white matters. The subcortical white matter also displays NADPH-d-positive neurons. NADPH-d-positive prominent, varicose cell processes are located especially in the deeper layers and in the subcortical white matter. The NADPH-d staining in the cerebral cortex is similar for all cerebral lobes and there are no observable differences between fight and left sides. The nucleus endopiriformis dorsalis and the claustrum contain NADPH-d-positive neurons (Fig. 9b,c), which present long processes that form a small plexus located dorsally along the lateral border of both nuclei. The striatum contains intensely NADPH-d-labeled neurons and the neuropil staining is dense (Figs. 9b-e and 10c). A similar staining pattern appears in the nucleus accumbens and in the tuberculum olfactorium (Fig. 9b,c). Within the tuberculum olfactorium, intense staining is observed in the islands of Calleja, in the fasciculus medialis telencephali and surrounding the islands of Calleja. The islands of Calleja contain a dense plexus of NADPH-d-positive varicose nerve fibers and small positive granular ceils. Strongly stained neurons are located in the diagonal band of Broca, where stained fibers are also present (Fig. 9c). Similar NADPH-d-labeled neurons are observed in the septum medialis and in the septum lateralis in rodents (Figs. 9c and 10b). NADPH-d-positive fibers are located in the septum lateralis and fornix. In the human brain, no NOS-immunoreactivity has been found in the septum. Groups of positive neurons and a high density of NADPH-d-positive fibers are observed in the substantia innominata and in the globus pallidus. The nucleus entopeduncularis shows a large number of positive neurons (Fig. 9c-e). Valtschanoff et al. (1993) described in detail the distribution of NADPH-d/NOS-positive neurons in the rat hippocampal formation. Positive cells are numerous in the pyramidal cell layer of the subiculum, in the stratum radiatum of the hippocampus, and in the subgranular zone of the dentate gyms (Fig. 9e-g); granule cells and pyramidal cells are reported to be NADPH-d-negative. Endoh et al. (1994a), however, have reported neuronal NOS staining in the pyramidal layer of CA1. Wallace and Fredens (1992) reported the presence of strong NADPH-d staining in the pyramidal neurons of CA1 in the mouse and Vaid et al. (1996) reported the presence of NADPH-d/NOS-positive pyramidal neurons in CA1 and in the subiculum. In the macaque monkey hippocampal formation NADPH-d/NOS-positive neurons with different morphological features appeared distributed in the molecular layer and granule cell layer of the dentate gyms and throughout all portions of the hippocampus. Most of the stained neurons have the morphological features and locations of intemeurons, whereas 73

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hippocampal pyramidal cells in the primate brain were NADPH-d/NOS-negative (Alonso et al., 2000). Studies in different species of monkeys as well as in humans indicate that the same types of NADPH-d/NOS-positive neurons are found in the hippocampal formation of several primates. There are, however, fairly substantial differences in the patterns of fiber staining. The distribution of NADPH-d in the amygdaloid complex is heterogeneous. The highest number of positive neurons is found in the nucleus medialis amygdalae. Scattered positive neurons appear in the nucleus basolateralis amygdalae, the nucleus basomedialis amygdalae and in the amygdalostriatal and amygdalopiriform transition areas (Fig. 9e). Diencephalon. Rostrally, NADPH-d-positive neurons are observed in the nucleus septohypothalamicus, the nucleus praeopticus medialis, the nucleus praeopticus anteroventralis, the nucleus praeopticus anteromedialis, the area praeoptica medialis and in the area praeoptica lateralis. In the hypothalamus, NADPH-d activity is observed in the magno- and parvo-cellular neurons of the nucleus paraventricularis, in the nucleus supraopticus, in the nucleus circularis, in the nuclei fornicalis and in the area posterioris hypothalami, as well as in a population of isolated neurons located among the different nuclei of the hypothalamus (Figs. 9d,e and 10d,e). The mammillary region shows NADPH-d-positive neurons within the ventral and dorsal portion of the nucleus paramammillaris, the ventral part of the nucleus mammillaris lateralis and the nucleus mammillaris medialis, the nucleus supramammillaris and in the decussatio supramammillaris (Fig. 9f). In the nucleus habenularis lateralis, NADPH-d-positive neurons are found in the medial and lateral regions, and labeled fibers are observed in the dorsal region. Many positive fibers are also observed in the commissura habenularis, while only a few NADPH-d-labeled axons are present in the fasciculus retroflexus. NADPH-d-positive neurons are found in the dorsomedial area of the thalamus within the nucleus paraventricularis thalami, the stria medularis and the nucleus anteroventralis thalami (Fig. 9e). Scattered neurons are also located in the nucleus laterodorsalis thalami, the nucleus paraventricularis thalami, the nucleus parafascicularis thalami, the nucleus praecommissuralis, the nucleus praetectalis olivari, the nucleus praetectalis anterior, the nucleus lateroposterioris thalami and in the nucleus geniculatus. However, only NOS-immunoreactive fibers have been demonstrated in the human thalamus. Mesencephalon. The NADPH-d staining in the colliculus superior forms a complex pattern. Neurons are mainly distributed in the superficial gray layer (Fig. 10f). Some scattered neurons also appear in the optic nerve layer and in the medial and lateral portions of the intermediate gray and white layers. Labeled fibers are distributed mainly in the intermediate gray and white layers, including the deep gray and white layers. Horizontally oriented positive fibers appear in the commissure of the colliculus superior. Stained cells and dense fiber plexus are present throughout the cortical nuclei of the colliculus inferior (Fig. 9h). These cortical nuclei also contain a dense NADPH-d-positive fiber plexus.

Fig. 10. Images of 30 ~m thick coronal sections of rodent brain labeled with NADPH-d-histochemistry: (a) from mouse (Mus musculus), and (b-f) from different rat encephalic regions IRattus norvegicus). (a) Within the olfactory bulb, diverse interneurons are NADPH-d-positive l arrows). NADPH-d-positive glomeruli (G+) are intermingled with negative ones (G-). (b) Septum mediale. (c) Caudatus putamen. (d) Nucleus paraventricularis hypothalami. (e) Area posterioris hypothalami. (f) Colliculus superior. Scale bars: 25 ~m in a, 100 gm in (b-z/'). 75

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In the nucleus nervi oculomotorii and in the fasciculus longitudinalis medialis of the cat, neurons with stained dendrites can be seen; these cells are not stained in the rat mesencephalon. NADPH-d-positive neurons are observed within the nucleus tegmentalis laterodorsalis, the nucleus interfascicularis, the nucleus paranigralis, the area tegmentalis (rostralis and ventralis), the rostral, rostrolateral and caudal subnuclei of the nucleus interpeduncularis, the nucleus Edinger-Westphal, the nucleus raphe medianus and the rhabdoid nucleus (Fig. 9g,h). A few neurons are located in the pars lateralis, compacta and reticulata of the substantia nigra. Labeled fibers are present in the substantia nigra pars compacta and pars reticulata, the nucleus interfascicularis, the nucleus interpeduncularis dorsomedialis, the nucleus raphe medianus and paramedianus, the parvocellular and magnocellular red nucleus, the area tegmentalis ventrolateralis, the nucleus profundus mesencephali and the rhabdoid nucleus. Rhombencephalon. NADHP-d staining is present in the cerebellum (Fig. 9h,i). The granule cells, basket cells and stellate cells in the inner third of the molecular layer show a weak reaction. The Purkinje cells are reported to be NOS-immunoreactive in the human cerebellum (Egberongbe et al., 1994). In contrast, in all other species studied, the Purkinje cells are unstained. In addition, the molecular layer neuropil is homogeneously stained. Stained fibers are also observed in the subcortical white matter and in the cerebellar nuclei, principally in the nucleus interpositus and nucleus lateralis. The cerebellar nuclei also contain a few weakly stained large cells. Weakly stained neurons are present in the nucleus ventralis lemnisci lateralis, the nucleus parabracchialis lateralis and medialis, the nucleus paralemniscalis, the nucleus of the trapezoid body and in the nucleus reticularis pontis caudalis. Moderately stained fibers are observed within the trapezoid body, the nucleus nervi trigemini and in the nucleus cochlearis dorsalis. No NADPH-d staining is detected in any cranial motor nuclei of the rat. However, in the cat, the cranial nuclei of the nervus facialis and nervus vagus are labeled. Moderately stained cells appear in the nucleus supragenualis, nucleus nervi hypoglossi, the nucleus raphe, the nucleus lateralis paragigantocellularis and in the nucleus reticularis medialis. The nucleus reticularis lateralis only contains scattered stained fibers. Strongly stained cells are observed in the nucleus tracti solitarii, the nucleus commissuralis of the tracti solitarii and in the nucleus ambiguus (Fig. 9i). Moderately stained neurons are scattered in the nucleus vestibularis medialis and in the nucleus paratrigeminus. Labeled fibers are located in the nucleus nervi hypoglossi, the nucleus paratrigeminus, the nucleus gracilis, the nucleus cuneatus and in the substantia gelatinosa of the nucleus spinalis trigemini. Spinal cord. Numerous NADPH-d-positive cells are identified in the superficial cornu dorsale and around the central canal at all spinal levels, and in the intermediolateral cell column at thoracic and sacral levels. Scattered cells are present in the deep cornu dorsale, cornu ventrale and white matter. Labeled fibers are observed in laminae I and II and in the intermediolateral column. 3.6. GENERAL PATTERN OF NADPH-DIAPHORASE EXPRESSION DURING RAT BRAIN DEVELOPMENT The distribution of nitrergic neurons has been studied during the development of the mammalian brain, mainly in the rat, but results are fragmentary and referred to specific 76

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brain regions. Most studies are carried out using NADPH-d histochemistry. Using the same technique, we have studied the onset and variations of nitrergic neurons during the prenatal and postnatal development of the rat brain. The size of the perikarya and the staining intensity of the NADPH-d-labeled neurons show in most cases a progressive increase until adulthood. In the initial stages, NADPH-d-positive neurons have immature morphologies, with thick and irregular processes, which become longer and more regular during the subsequent developmental stages until they reach the mature phenotype. Thus, morphologic characteristics such as size, number of cell processes, and staining intensity define the NADPH-d-positive cells at each developmental stage. In order to confirm that NADPH-d staining corresponds to nitrergic cells, NOS presence in the brains of developing animals is also analyzed by immunocytochemistry. Both NADPH-d activity and NOS immunoreactivity distribution patterns are similar in animals at the same developmental stage, but NADPH-d-stained elements demonstrate a more complete labeling, including frequently portions of the dendritic tree and lengthy axons that are not so clearly observable with NOS immunocytochemistry. Vascular endothelium is NADPH-d-stained, but it is not neuronal NOS-immunolabeled. This endothelial labeling occurs before the onset of NADPH-d activity in neural elements (is even present at E16, when NADPH-d-positive neurons are absent) and can be explained by the NADPH-d activity shown by endothelial NOS, not recognized by antineuronal NOS antibodies. The developmental pattern of NADPH-d expression is highly variable among the different brain nuclei. NADPH-d appears at the prenatal stage El7 in the telencephalon, diencephalon, mesencephalon and myelencephalon, and later on, at the postnatal day P6, in the metencephalon. Although the general pattern is an increase in the number of NADPH-d-stained cells and in their staining intensity, several nuclei display a transient expression of NADPH-d following three different models. The number of NADPH-d-positive short-axon cells in the main olfactory bulb and NADPH-d-stained cells in the gigantocellular myelencephalic area and in the nucleus praepositus hypoglossi increases progressively until a critical stage when the number of cells decreases up to the final density observed in the adult. In the deep layers of the colliculus superior and in the griseum centrale of the mesencephalon there is a gradual increase from the onset, but there is an interval during the perinatal period when there is an evident decrease of the labeled elements with respect to previous stages. The third model of transient expression of NADPH-d, observed exclusively in the nucleus parataenialis, consists of an initial increase in the number of the labeled elements until a defined stage (P10) when the number of positive cells starts to decrease until the complete depletion in the adult. Telencephalon. Throughout the development, the evolution of NADPH-d expression towards the adult pattern occurs slowly. Only two telencephalic areas lack NADPH-d activity: the globus pallidus and the cortex piriformis. Although other studies indicate that NADPH-d expression in the telencephalon begins at E 15 (Iwase et al., 1998b) we observe NADPH-d labeling for the first time at the prenatal stage E 17, when scarce NADPH-d-positive neurons appear at medial levels of the fundus striati and the caudatus putamen (Fig. 1 la), scattered within the basal telencephalon and in the medial and posterior subdivisions of the bed nucleus of the stria terminalis (Fig. 1 l b,c). Numerous NADPH-d active neurons are detected for the first time at E 18 in the medial and basomedial amygdala. A scarce population of NADPH-d-positive cells is found in the pallium ventrale and in the ansa lenticularis. Regions expressing NADPH-d activity for the first time at El9 are the nucleus olfactorius anterior, the commissura anterior, the nucleus accumbens and the substantia innominata. Occasionally isolated NADPH-d-positive cells are seen in the cortical neuroepithelium and in the cortical intermediate layer. 77

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Fig. 11. Camera lucida drawings of coronal sections from rostral (a) to caudal (g) levels through the brain of the E 17 rat embryo.

At the last prenatal developmental stage, E2 l, the onset of NADPH-d activity occurs in the anterior amygdala, and an increase in the number of NADPH-d-positive neurons and fibers is found in the telencephalon (Fig. 12a-d). NADPH-d-active elements are especially abundant in the caudatus putamen. Cortical NADPH-d active neurons display still an immature aspect with respect to the telencephalic nuclei at the same stage. NADPH-d-positive cells are detected at P0 for the first time in the taenia tecta and in the neuroepithelium close to the septal nuclei, and a weak fibrillar staining is observed in some glomeruli of the main olfactory bulb. At P 1, NADPH-d-positive short-axon cells are observed in the main olfactory bulb. Also at P1, the onset of NADPH-d activity occurs in the septum mediale. Regions demonstrating the onset of NADPH-d activity at P2 are the accessory olfactory 78

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lucida drawings of coronal sections from rostral (a) to caudal (i) levels through the brain of the

E21 rat embryo.

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Fig. 13. Camera lucida drawings of coronal sections from rostral (a) to caudal Ik) levels through the brain of the P5 postnatal rat.

bulb, the tuberculum olfactorium and the diagonal band of Broca. Weak NADPH-d-positive neurons are found in the anterior amygdala, the nucleus endopiriformis ventralis, and the region located between these nuclei and the cortex. Also at P2, a scarce NADPH-d-pos80

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Fig. 14. (a, b) NADPH-d staining in the diagonal band of Broca during rat brain development. At P4 (a) weakly stained NADPH-d-positive cells were present in the horizontal limb, and neurons containing NADPH-d are not observed in the vertical limb. At P25 (b) NADPH-d-positive cells are present in both the horizontal and vertical limb. (c, d) General view of the NADPH-d-staining pattern in the laterodorsai and dorsal pericentral tegmental nuclei at two postnatal stages. Note that in the DPTg, NADPH-d-positive neurons are scarce at both developmental stages (arrowheads), whereas in the LDTg they are scarce at PI (c) and abundant at P25 (d). Scale bars: 100 ~tm in a and b, 500 ~m in c and d.

itive neuronal population becomes visible in the radiatum, pyramidal and oriens strata of the Ammon's horn and in the white matter of the subiculum in the hippocampal formation. 81

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At P3, scarce and weak NADPH-d-positive periglomerular cells are labeled in the olfactory bulb by the first time. A strong fibrillar staining is observed in the telencephalic nuclei presenting NADPH-d activity, but not in the cerebral cortex. NADPH-d-positive cells are sparsely distributed in all layers of Ammon's horn. At this developmental stage, NADPH-d-positive neurons of cerebral cortex are also present in the external layers, as a difference to previous stages. Increases in the staining intensity, in the number of ramifications of the dendritic tree, and in the cell size of the NADPH-d-labeled elements are found in the telencephalon at P4. Although at this stage NADPH-d-positive elements demonstrate a lower degree of maturation than those observed in the adult, the general distributions of NADPH-d-labeled fibers and cells in the olfactory peduncle, the cortex, the caudatus putamen, the fundus striati, and in the basal telencephalon are similar to those of the adults. There is a noticeable increase in the density of NADPH-d-positive neurons in the cortical and septal regions. At P5, new elements expressing NADPH-d consist of weak or moderate stained fibers observed in the insula Callejae and both cells and fibers in the nucleus tractus olfactorii (Fig. 13c). After the first week of postnatal life, general increases in the staining intensity, cell size and ramification of the dendritic tree in the NADPH-d-positive neurons are seen. There is an evolution in the NADPH-d staining pattern towards the adult model in the olfactory bulb, the hippocampal formation, the septal nuclei, the amygdaloid complex, and in the cortical areas (Fig. 13a-g) and in the diagonal band of Broca (Fig. 14a,b). At P7, the nucleus entopeduncularis demonstrates moderately stained NADPH-d-positive neurons. A new group of NADPH-d-labeled neurons appears in the presubiculum and parasubiculum of the hippocampal formation at this stage. At P8, both main and accessory olfactory bulbs show lower numbers of NADPH-d-positive elements than in the adult stage; however, the morphological characteristics of the NADPHd-stained neurons included in these structures are already similar to those observed in the adult. In the nucleus olfactorius anterior, in the taenia tecta and in the commissura anterior the density and staining intensity of NADPH-d-stained neurons are similar to the definitive pattern, but their morphological characteristics are still immature. This circumstance also occurs in the caudatus putamen and in the corpus callosum. An increase in the staining intensity and in the degree of maturation of the NADPH-d-positive cells is observed in the different nuclei of the amygdaloid complex. At P8, NADPH-d-positive neurons are seen for the first time in the subfornical organ. At P9, we observe an evident increase in the size of NADPH-d-positive neurons in the nucleus olfactorius anterior, the taenia tecta, the cortex, the corpus callosum, the commissura anterior and in the caudatus putamen in comparison with previous developmental stages. At P10, NADPH-d-stained neurons are seen in the rostralmost and caudalmost levels of the caudatus putamen, where they are not detected at previous stages. Although the general pattern at P10 is very similar to that observed in the adult rat telencephalon, some structures demonstrate variations when the late postnatal stages are analyzed. Thus, in the P I5 main olfactory bulb a lower density of NADPH-d-positive granular cells but a higher density of NADPH-d-labeled short-axon cells than those found in the adult are observed, while the number of NADPH-d-active periglomerular cells is similar. The density of NADPH-d-stained cortical cells undergoes a gradual decrease from P15 onwards (Fig. 15a,b). At P20, the morphological characteristics of the NADPH-d-positive cells in the septal nuclei are more immature as compared with the adult model. At the last subadult stage analyzed, P30, the NADPH-d expression pattern in the telencephalon is similar to that in the adult, except for 82

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the higher number of NADPH-d-stained short-axon cells in the granule cell layer of the main olfactory bulb. Diencephalon. At E 17, NADPH-d-positive neurons appear in the nucleus septohypothalamicus, the nucleus mammillaris dorsalis and in the nucleus commissuraae posterioris (Fig. 1 ld). At this stage, this latter nucleus demonstrates the highest density of NADPH-d activity in the whole brain. At El8, the nucleus praeopticus medialis, located between the midline and the nucleus .septohypothalamicus presents NADPH-d-labeled cells. This neuronal population is more numerous than that of the nucleus septohypothalamicus. More caudally, NADPH-d-stained cells are detected in the area lateralis hypothalami and in the nucleus mammillaris ventralis. In the nucleus paraventricularis hypothalami, two different neuronal populations start to express NADPH-d at El8: the population located in its medial part is weakly stained and more abundant than the population located in its dorsal part. At the following prenatal stages, El9, E20, and E21, no substantial variations on the NADPH-d staining pattern are found in the diencephalon. There is only a noticeable general increase in the number of NADPH-d-positive elements and in the staining intensity in the nucleus commissurae posterioris (Fig. 12e), whereas the staining intensity remains weak in the hypothalamic nuclei. After birth (P0), the lateral part of the nucleus ventromedialis thalami and the nucleus supramammillaris demonstrate for the first time weakly labeled NADPH-d-positive neurons. A noticeable increase in the number of NADPH-d-positive cells is detected in the nucleus septohypothalamicus and in the nucleus praeopticus medialis, as well as an increase in the staining intensity of neurons in the area lateralis hypothalami. New nuclei that display NADPH-d activity at P1 are the nucleus circularis hypothalami, the area posterioris hypothalami, the nucleus tuberis medialis and the parataenialis. At P2, NADPH-d activity starts at the nucleus subthalamicus. At P4, NADPH-d-positive neurons are seen for the first time in the nucleus periventricularis hypothalami and in the nucleus centromedialis, the nucleus interanteromedialis and the nucleus romboidalis of the medial thalamus, forming a scarce and scattered neuronal population. The NADPH-d-positive neuronal populations in the area lateralis hypothalami, in the posterior part of the nucleus paraventricularis, and in the nucleus circularis, display stronger staining intensity, larger somatic volume and longer processes than at P3. At P5, new NADPH-d-positive neuronal groups occur in the nucleus mammillaris lateralis, the nucleus mammillaris dorsomedialis and in the nucleus supraopticus hypothalami (Fig. 13f). At P8, a small group of weak NADPH-d labeled neurons is observed in the nucleus supraparafascicularis and in the nucleus intralaminaris posterior thalami. At P9, thin and long fibers are observed in the habenula. At P12 we observe for the first time NADPH-d-stained fibers in the nucleus geniculatus lateralis and in the nucleus ventrolateralis thalami. At P13, we also observe NADPH-d-stained fibers in the area praetectalis anterior. From P10 to P15 there is a gradual decrease in the number of labeled cells in the nucleus parataenialis and also in the staining intensity of the remaining population of this hucleus until the complete abolition in the adult. This transient postnatal expression in the nucleus parataenialis has been previously reported (Garcfa-Ojeda et al., 1997; Gorbatyuk et al., 1997). By contrast, the number of labeled cells and their staining intensity increase progressively during the postnatal development in other diencephalic nuclei. At P20, NADPH-d-positive cells appear in the dorsal and lateral thalamus. At P25, we observe for the first time NADPH-d-stained fibers crossing the tractus mammillaris. At P30, the neuronal NADPH-d staining pattern in the diencephalon is very similar to the adult model, with numerous intense 83

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NADPH-d labeled neurons and a dense fibrillar staining in both thalamic and hypothalamic nuclei. Mesencephalon. In the mesencephalon, a highly variable NADPH-d staining pattern is observed among different nuclei throughout development. In the tegmental nuclei, a remarkable high density of NADPH-d-positive elements is detected along the prenatal period. A feature of the NADPH-d staining pattern in the mesencephalon is a decrease of the density of NADPH-d-positive neurons in the deep layers of the colliculus superior and in the griseum centrale around the first week of postnatal life. In these developmental periods we also detect an increase of perikarya volume of some neuronal groups of the tegmental nuclei, in agreement with Skinner et al. (1989). At El7, a single group of neurons demonstrates NADPH-d in the mesencephalon. It is located in the midline between the superior colliculi of both hemispheres, at the top of the Sylvian aqueduct (Fig. l 1f). At the following stage, El8, NADPH-d activity appears for the first time in the griseum centrale, in the nucleus pedunculopontinus tegmenti, in the nucleus tegmentalis dorsalis pericentralis and in the nucleus tegmentalis laterodorsalis. New mesencephalic areas expressing NADPH-d at El9 are the deep gray layer of the colliculus superior and the deep mesencephalic area. At E20, the colliculus superior, the stratum griseum intermedium of shows a scarce population of NADPH-d-positive neurons with stained proximal dendrites. At the last prenatal stage, E21, the density of NADPH-d labeled elements remains unchanged (Fig. 12f), and a few neurons located in the colliculus inferior start to express NADPH-d activity (Fig. 12g-i). At P0, the main variation with respect to previous stages was the presence of NADPH-d activity in the area tegmentalis ventralis. At P1, a few neurons in the stratum opticum of the colliculus superior are NADPH-d-stained. A continuous increase of the staining intensity, size, and density of NADPH-d-positive elements in the colliculus superior, in the griseum centrale, and in the tegmental nuclei occurs after P1 and during the first week of postnatal life (Fig. 14c,d). Cells expressing NADPH-d are firstly labeled at P4 in the bracchium colliculi inferioris. At P5 and P6, a dramatic increase of somatic volume of tegmental NADPH-d-positive neurons is observed. Also in the tegmentum, the NADPH-d-positive neurons show a more developed phenotype and stronger staining intensity than at previous stages. A considerable increase in the density of NADPH-d-stained neurons is detected in the tegmental nuclei and also in the colliculus inferior (Fig. 13g,h). After the first week of postnatal life (P7 and P8) we observe a drastic decrease on the density of NADPH-d-positive neurons in the griseum centrale and in the deep and intermediate layers of the colliculus superior (Fig. 15c,d), and the start of expression in the most superficial layers, as it has been previously reported by other authors (Gonz~ilez-Hem~indez et al., 1993). The decrease contrasts with the increase of NADPH-d-positive elements in the tegmental nuclei and colliculus inferior. The gradual decrease in the NADPH-d staining of the colliculus superior and griseum centrale stops at P10. At this developmental stage, the size and shape of the NADPH-d-labeled neurons in these structures are similar to those observed in the adult. From P11 onwards, NADPH-d labeling becomes visible in all layers of the colliculus superior. We also observe a differential distribution of NADPH-d-stained cells between

Fig. 15. (a, b) NADPH-d distribution pattern in the developing cerebral cortex of the rat. A decrease in the number of NADPH-d-positive neurons is detected between P15 la) and P30 (h). I-VI are layers of the cerebral cortex. (c, d) Transversal sections of the colliculus superior at P6 to) and P30 (d). The deep layers are intensely stained in the first postnatal stages, while in the superficial layers, NADPH-d activity increases progressively during the late postnatal development. Scale bars are 100 p.m.

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cortical and inner regions of the colliculus inferior. In the cortical region, a high density of weak and moderate NADPH-d-positive neurons and a moderate fibrillar staining are seen, whereas in the inner part, a low density of NADPH-d-active neurons is detected. At the following developmental stages (P 15, P20, P25 and adults), this NADPH-d staining pattern is maintained in the colliculus inferior. Rhombencephalon. In the metencephalon, NADPH-d activity appears later than in other brain regions, at postnatal life, but there are discrepancies in the exact onset in this region, from P1 to P14 (Giuili et al., 1994; Black et al., 1995; Li et al., 1997; Iwase et al., 1998b). We observe the first NADPH-d-positive neurons in the granular layer of the cerebellar cortex at P6. Three different NADPH-d-labeled neuronal populations are detected: the most abundant neurons are identified as granule cells, but some horizontal cells and Golgi cells are also labeled. These cells are only located in the central zone of the granule cell layer within the deep cerebellar lobes. At P8, substantial changes are observed. In addition to the labeling in the granule cell layer, NADPH-d-positive neurons are also found in both the molecular layer and the white matter layer of the cerebellar cortex. In the molecular layer, basket cells are labeled after NADPH-d histochemistry. In the white matter, medium-sized NADPH-d-positive internal cells are occasionally observed. At this stage, the NADPH-d-labeled granule cells, in addition to the central region, are distributed in the lateral part of the inner cerebellar lobes. At P11, a scarce population of large NADPH-d-positive neurons is observed in the nucleus interpositus and nucleus lateralis of the cerebellum. In the nucleus medialis of the cerebellum, NADPH-d activity appears at P12, and consists of large and weakly stained neurons located exclusively in the external portion of this nucleus (as a difference with both nuclei interpositus and lateralis, where NADPH-d-positive elements are distributed throughout all their extension). No new cerebellar layers and nuclei show NADPH-d activity from P12 onwards. Purkinje cells are unstained after NADPH-d histochemistry in all developmental stages analyzed. In the nuclei interpositus and lateralis, some NADPH-d-positive fibers are found at P12. After this developmental stage, a quick evolution towards the adult model is observed. At P13, NADPH-d-stained basket cells and granule cells are also detected in the outer cerebellar lobes. At P15, the labeling in the cerebellar nuclei and cerebellar cortex is similar to that described in previous stages. At P25, some NADPH-d-positive fibers are found in the nucleus medialis. At P30, all cerebellar nuclei present a NADPH-d staining pattern identical to those observed in the adult. Most NADPH-d-positive myelencephalic structures have NADPH-d-labeled neurons in the prenatal life. The outstanding characteristic of the NADPH-d expression in the rat myelencephalon throughout development is the partial loss of NADPH-d-positive elements in some nuclei postnatally. At El7, NADPH-d-positive neurons are detected in the raphe and trigeminal nuclei, as described by Takemura et al. (1996). In the caudalmost levels of the myelencephalon, NADPH-d-positive neurons are found in the nucleus raphe obscurus (Fig. 1 l e,f), in the nucleus spinalis trigemini, in both sides of the pyramidal decussation, and in the ventrolateral zone to the nucleus nervi hypoglossi, reaching the dorsal region of the nucleus reticularis (Fig. 1 l g). El8 is the stage of appearance of NADPH-d activity in the gigantocellular and intermediate part of the formatio reticularis. Noticeable is the large cell body size, high number of processes, and variety of cell morphologies exhibited by the NADPH-d-positive neurons in the formatio reticularis from the onset onwards. These phenotypes are not observed in any other brain nuclei throughout the prenatal life. 86

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Other myelencephalic nuclei start to express NADPH-d activity at El9: the pontine nuclei, the lateral part of the formatio reticularis and the vestibular nuclei. At El9, the NADPH-d-labeled cells described at previous stages are more mature (higher size, number and length of processes than those at El7 or El8). However, an increase in the density or staining intensity is not observed. At E20, NADPH-d-positive neuronal populations are observed in the nucleus lemniscalis, the nucleus fasciculi solitarii, the nucleus praepositus hypoglossi and in the nucleus ambiguus. Distinctive neuronal groups display for the first time NADPH-d activity at the last prenatal stage, E21: the paracentral and medullar parts of the formatio reticularis, and the nucleus reticularis paragigantocellularis dorsalis (Fig. 12g-i). At birth (P0), NADPH-d-positive fibers and NADPH-d-labeled neurons occur for the first time in the nucleus trigeminus spinalis. At P1, NADPH-d expression starts in the locus coeruleus, the parabracchialis, the nucleus gracilis, the nucleus raphe dorsalis and also in the nucleus raphe medialis. At this stage, the distribution pattern of NADPH-d-positive elements in these myelencephalic regions is similar to that observed in the adult brain, although the density and maturity of NADPH-d-stained neurons is lower than in the adult. After this developmental stage, the changes in the staining and in the morphological characteristics of NADPH-d-positive neurons occur slowly, and are more evident during the first week of postnatal life. The onset of NADPH-d activity in the nucleus paratrigeminus, and in the rostral levels of the nucleus fasciculi solitarii occurs at P4. At this same stage, we observe a weakly labeled fiber staining in the area postrema. At the following stage, P5, NADPH-d-positive cells are abundant in the raphe nuclei and in the formatio reticularis (Fig. 13h-k). We detected NADPH-d activity for the first time at this same stage in the nucleus interpenduncularis, the nucleus pontinus and in the nucleus cuneatus. After P6, a decrease in the density of NADPH-d-positive cells is observed in some myelencephalic structures. Thus, in the dorsoposterior part of the area gigantocellularis, in the formatio reticularis and in the nucleus praepositus hypoglossi, the density of NADPH-d-positive decreases until adulthood. No new regions express NADPH-d activity from P6 onwards, and most regions undergo a progressive increase in the density, staining intensity and size of NADPH-d-positive cells.

4. DISCUSSION 4.1. METHODOLOGICAL ASPECTS The NADPH-d histochemical technique using NADPH-d as substrate and tetrazolium salts as chromogen allows to localize NADPH-d activity in both fixed and unfixed brain tissue (Alonso et al., 1995a). However, the identification between NADPH-d and NOS staining patterns is only observed after aldehyde fixation. Under these conditions, the colocalization of NADPH-d with NOS protein and mRNA in the same cells (Bredt et al., 1991a) suggests that NOS catalytic activity accounts for NADPH-d staining. Moreover, transgenic knock-out models for NOS do not exhibit NADPH-d staining (Huang et al., 1993; Darius et al., 1995) while a cell line transfected with neuronal NOS cDNA expresses de novo both the enzyme and NADPH-d activity (Dawson et al., 1991). In the nervous system, NADPH-d activity and neuronal NOS-immunoreactivity widely colocalize in specific sets of well characterized cells (Bredt et al., 1991a; Hope et al., 1991; Kishimoto et al., 1993; Kugler and Drenckhahn, 1996; Brifi6n et al., 1998). Nevertheless, the exact matching between neuronal NADPH-d staining and NOS has been discussed (Hope 87

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et al., 1991). There are particular examples where NADPH-d staining is not partially or totally coincident with that for NOS, as in the adrenal gland (Afework et al., 1992), in the olfactory and vomeronasal receptors and their projecting axons (Kishimoto et al., 1993; Kulkami et al., 1994; Spessert and Layers, 1994: Brifi6n et al., 1998; Weruaga et al., 1998), cells in the cerebral cortex (Kharazia et al., 1994; Sobreviela and Mufson, 1995), and neuronal populations in specific nuclei of the spinal cord (Vizzard et al., 1995). These discrepancies have been interpreted as lack of neuronal NOS, whereas other oxidative/reductive enzymes would account for the NADPH-d activity in those cells (Kulkami et al., 1994). In fact, NADPH-d activity is present in cells containing other enzymes such as cytochrome C or cytochrome P450-reductase, which show high molecular homology with NOS (Bredt et al., 1991b; L6wenstein and Snyder, 1992; Knowles and Moncada, 1994). NADPH-d activity related with NOS is conserved after fixation with aldehydes (Matsumoto et al., 1993; Tracey et al., 1993), but this affirmation should be certain for other enzymes as well. For both immunohistochemistry and NADPH-d histochemistry, tissue fixation is crucial, but tissue treatment is more determinant for the latter technique. Cryostat or vibratome sectioning are the most widely employed and reliable methods. As a general rule, perfusion of the animal for 15 min with a physiologic-buffered solution containing 4% paraformaldehyde, and postfixation of trimmed pieces of the dissected brain in the same cold solution for 2-4 h is an optimal treatment of tissue for NADPH-d histochemistry. Relatively low concentrations of glutaraldehyde (until 0.5%) increase definition of positive fibers (Alonso et al., 1995a; Buwalda et al., 1995), but glutaraldehyde is a drawback when other techniques, such as immunohistochemistry, are planned (Matsumoto et al., 1993; Tracey et al., 1993). As well as fixative composition, fixation time is important for the reliability of the histochemical technique. Weakly stained NOS-positive neurons in brain tissue slightly fixed (shortly fixed) seem to lose NADPH-d activity (Gonz~ilez-Hern~indez et al., 1996), but a similar phenomenon occurs when the time of fixation is too long (Alonso et al., 1995a; Brifi6n et al., 1998). There are examples of contradictory or inconsistent results attributable to the fixation quality. Thus, identical results have been described with tissue fixed with both paraformaldehyde alone and mixtures of paraformaldehyde-glutaraldehyde (Valtschanoff et al., 1993), whereas other authors found remarkable differences, including the labeling of 'new' neural cells by using the glutaraldehyde-containing fixatives (Dinerman et al., 1994; Buwalda et al., 1995; Vaid et al., 1996). No brain region has elicited so many controversial results on its NADPH-d/NOS-staining as the hippocampus. Valtschanoff et al. (1993) find similar results for tissue fixed both in paraformaldehyde alone and in mixtures containing different amounts (1-2.5%) of glutaraldehyde. Importantly, they find no staining of hippocampal pyramidal neurons. By contrast, Dinerman et al. (1994) have suggested that hippocampal pyramidal cells in the rat are NADPH-d-negative after standard paraformaldehyde fixation but stain robustly following glutaraldehyde-based fixatives. They further suggest that the positive pyramidal cells contain endothelial NOS rather than neuronal NOS (Dinerman et al., 1994; Kantor et al., 1996). A similar observation has been reported in human hippocampus where pyramidal cells do not contain neuronal NOS but do demonstrate endothelial NOS (Doyle and Slater, 1997). In agreement with this observation, we find that pyramidal cells in the macaque monkey hippocampal formation (Alonso et al., 2000) were neuronal NOS-immunonegative and they do not show clear NADPH-d staining after fixation with no, low (0.1%) or high (2.5%) glutaraldehyde amounts. Of course the possibility that low amounts of neuronal NOS are expressed in principal neurons in the primate hippocampal formation but are not detectable with standard histochemical or immunohistochemical procedures cannot be entirely ruled 88

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out. In this case, the species difference may be only in the level of NADPH-d/NOS that is expressed in principal neurons. Given the high involvement of the fixation quality in the results of these techniques, it is necessary to be especially cautious when working with autopsy material since this tissue is under suboptimal fixation. In the human hippocampal formation, Sobreviela and Mufson (1995) have found that strongly NADPH-d-labeled cells are always NOS-immunoreactive, while weakly stained neurons do not. Such a difference has also been found in the mouse olfactory bulb, but the colocalization with NOS is usually accomplished when tissue fixation and manipulation are proper (Alonso et al., 1995a; Gonzfilez-Hern~.ndez et al., 1996; Weruaga et al., 1998). Glial labeling with both NOS-immunohistochemistry and NADPH-d histochemistry should be considered apart. First studies employing NADPH-d staining reported labeled glial cells in vivo and in vitro, as well as in gliomas (see Sestan and Kostovi6, 1994). With the standard fixation related below, glial staining is eliminated, although oligodendrocytes are NADPH-d-stained in teleosts (Ar6valo et al., 1995; P6rez et al., 1996; Crespo et al., 1998a). For mammals, the visualization of astrocytes is possible running the NADPH-d staining on very slightly fixed tissue. This staining is absolutely coincident with that obtained with neuronal NOS-immunolabeling, but only with special fixation and cryo-drying of the sections (Gabbott and Bacon, 1996; Kugler and Drenckhahn, 1996). Therefore, when investigating nitric oxide-synthesizing elements, it is recommendable to compare the results with other methods for the localization of NOS, and to test different fixation and incubation protocols when using different species or tissues or different developmental stages until the stainings fit completely or differences are clearly demonstrated. 4.2. INTERSPECIES DIFFERENCES IN THE NADPH-DIAPHORASE DISTRIBUTION PATTERN NADPH-d/NOS-positive elements in the central nervous system of vertebrates are distributed throughout most brain subdivisions, constituting, in general, small neuronal subpopulations within a given structure. From the location and nature of NADPH-d/NOS-stained cells it is hardly possible to extrapolate a common feature such as definite functional pathways or neurotransmitter systems that could match with the distribution of these cells. In a general overview of their distribution in the vertebrate brain, there are structures where nitrergic populations are constantly present throughout the phylogenetic scale and other brain nuclei or zones where the presence of NADPH-d/NOS is not consistently found among different vertebrate groups. In this sense, NADPH-d/NOS-positive elements have been found in all vertebrate species studied in the striatum, amygdaloid complex, medial septum, hypothalamus, midbrain tectum, nucleus spinalis trigemini, solitary tract area, and spinal cord (or their homologous regions). In other brain regions such as the olfactory bulb, cortex, thalamus, ventral tegmental area, substantia nigra, retrobulbar area and raphe nuclei, more striking differences have been detected among vertebrate classes: NADPH-d/NOS-stained cell bodies are frequently found, but not consistently throughout the phylogenetic scale. In this sense, NADPH-d/NOS-stained cell bodies are absent in the olfactory bulb of lampreys (Schober et al., 1994) and teleosts (Ar6valo et al., 1995; Brfining et al., 1995), whereas only a few, weakly stained cells are found in those of reptiles (Brfining et al., 1994b; Smeets et al., 1997) and birds (Brfining et al., 1994a; Panzica et al., 1994). By contrast, numerous stained cell bodies have been observed in the main and accessory olfactory bulbs of both anuran (Mufioz et al., 1996; Porteros et al., 1996) and urodele amphibians (Gonzfilez et al., 1996; Porteros et al., 89

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1996), as well as in rodents (Vincent and Kimura, 1992; Porteros et al., 1994), and in the main olfactory bulb of monkey (Alonso et al., 1998) and human (Brifi6n et al., 1998). The mere presence of nitrergic populations in a given structure through the phylogenetic scale does not allow to conclude evident anatomical, physiological and/or evolutionary relationships of this neuronal subsystem given the remarkable differences in their number and distribution among groups, even when comparing functionally or structurally similar brain regions. As an example, significant qualitative and quantitative interspecies variations have been found in the NADPH-d/NOS-positive cells occurring in the midbrain. Whereas in reptiles and birds the ventral tegmental area, substantia nigra and retrobulbar group contain numerous NOS-stained cell bodies, in rats they are mainly confined to the ventral tegmental area. A comparison with anamniotes is difficult, because distinct midbrain groups cannot be distinguished in lampreys (Pierre et al., 1994) and teleost fishes (Meek, 1994; Smeets and Reiner, 1994). However, in the dorsomedial part of the posterior tubercle and rostral tegmentum of amphibians, which are comparable to the ventral tegmental area and the substantia nigra of amniotes (Marfn et al., 1995), only a few NADPH-d/NOS-positive cells are found (Gonz~lez et al., 1996; Mufioz et al., 1996). In addition to these variations among groups, remarkable differences also occur within each class in specific locations. For example, NADPH-d- and NOS-positive cells were found in the thalamus of the reptile Gekko, but not in an other reptile, Pseudemys. A similar discrepancy is observed in amphibians, where a large population of stained cells is present in the thalamus of anurans (Mufioz et al., 1996), but absent in urodeles (Gonz~ilez et al., 1996). Substantial differences in density and distribution were also observed in the avian thalamus, comparing chickens (Brfining, 1993) with quails (Panzica et al., 1994). The significance of the differences must be necessarily related to the distinct organization of each brain structure in each vertebrate group or subgroup. Nevertheless, other factors must be also involved given the remarkable interspecies variability demonstrated by brain regions characterized by their similar structural and functional features along the phylogenetic scale. For example, within mammals there are important differences between NOS-positive elements in the hippocampal formation of the rat (Valtschanoff et al., 1993) with those described in different monkey species (Carboni et al., 1990; Mufson et al., 1990) and human (Sobreviela and Mufson, 1995). In addition, minor differences persist even within a same group such as primates, although the divergences are less dramatic and a relatively common NADPH-d/NOS-staining cell pattern is observed. The interspecies differences on the NADPH-d/NOS staining pattern could be explained by specific variations in the circuitry within a structurally constant region. The hippocampus and the olfactory bulb constitute clear examples for this hypothesis. Whereas the dense NADPH-d/NOS innervation of the rat hippocampal formation has one of its main sources in the cholinergic septohippocampal cells in the rat (Kinjo et al., 1989; Kitchener and Diamond, 1993), in the macaque monkey and human it has a different origin since in both species cholinergic septohippocampal cells do not contain NOS (Ellison et al., 1987; Geula et al., 1993; Alonso et al., 2000). In addition, the practical inexistence of NOS-positive fibers in the primate fimbria (Sobreviela and Mufson, 1995; Alonso et al., 2000) indicates that other non-cholinergic populations of septohippocampal neurons projecting through the fimbria are also NADPH-d/NOS-negative. Therefore, it appears that the dense innervation of NOS-stained fibers found in the primate hippocampal formation has a different origin from those observed in rodents. Concerning the olfactory bulb, the NADPH-d/NOS somal and fiber stainings show striking divergences in different vertebrate species. Even within the mammalian group, 90

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differences can be detected in the number, distribution and nature of NADPH-d/NOS-stained elements between macrosmatic and microsmatic species. Detailed analysis of similarities and differences among vertebrate groups suggests that the complexity of NADPH-d/NOS staining in the olfactory bulb could correlate to the developmental level of the olfactory structures and the importance of olfaction in the biology of that species. Thus, the primate olfactory bulb (Alonso et al., 1998; Brifi6n et al., 1998) exhibits a simplified NADPH-d/NOS staining pattern as compared to the staining in rodents (Davis, 1991; Vincent and Kimura, 1992; Alonso et al., 1993). The NADPH-d/NOS-Iabeled neurons in the primate olfactory bulb are less abundant, and show a lower morphological diversity than those described for rats and mice. Similarly, rodents have simpler NADPH-d/NOS staining patterns than the hedgehog, an insectivore with extraordinarily developed olfactory structures (Alonso et al., 1995b). Studies in other groups of vertebrates seem to confirm this hypothesis: no NADPH-d staining is observed in the secondarily simplified olfactory bulb of birds (Panzica et al., 1994), and only olfactory fibers are NADPH-d-positive in the olfactory bulb of teleosts (Ar6valo et al., 1995). Immunohistochemical and in situ hybridization studies have revealed an unique distribution for NOS that does not correlate in an exclusive manner with the distribution of any other neurotransmitter or neuroactive substance or with specific sensory, motor or integrative pathways (Bredt et al., 199 la). Nitric oxide occurs in glutamatergic cells in the cerebellar granule cells, but also in GABAergic basket cells. In the cerebral cortex, NADPH-d/NOS neurons colocalize somatostatin, neuropeptide Y and GABA, and somatostatin and neuropeptide Y in the striatum (Dawson et al., 1991), while in the nucleus pedunculopontinus tegmenti of the brainstem, NOS-positive neurons lack somatostatin and neuropeptide Y, but stain for choline acetyltransferase (Dawson et al., 1991). In the hypothalamus there are partial coexistences of NADPH-d activity with somatostatin, vasopressin, oxytocin, calbindin D-28k, calretinin, parvalbumin, or acetylcholinesterase (Alonso et al., 1992a,b; Ar6valo et al., 1993; S~inchez et al., 1994; Crespo et al., 1998b). In addition, the relationship of nitric oxide and other neuroactive substances in specific regions of the central nervous system is not a general feature throughout the phylogenetic scale. The colocalization of NADPH-d/NOS with catecholamines markers is well documented. In birds and reptiles, NADPH-d/NOS activities extensively colocalize with catecholamines in the midbrain (Panzica et al., 1996; Smeets et al., 1997). By contrast, colocalization in the midbrain of mammals is scarce (Johnson and Ma, 1993). It is also very limited in anuran amphibians (Mufioz et al., 1996) and completely absent in urodeles (Gonz~ilez et al., 1996). All these data suggest that although the coexpression of NADPH-d/NOS staining with catecholaminergic markers exists in the vertebrate brain (mainly in the midbrain), there are considerable variations between groups, and even within groups. The rate of colocalization is hitherto difficult to extrapolate with any functional significance, or with the phylogenetic evolution of the brain. 4.3. IMPLICATIONS OF THE NADPH-DIAPHORASE EXPRESSION DURING THE DEVELOPMENTAL PROCESSES During brain development, NADPH-d-stained cells are initially observed after cell bodies ceased dividing and extended their processes. NADPH-d activity is not expressed during the cell division or the cell migration process, but in late developmental stages, when the establishment of functional connections is taking place. Available data suggest a role for these nitric oxide-producing neurons in the formation of the pattern of connectivity during development. In this sense, NADPH-d expression in several brain regions correlates with the schedule of innervation. This is the case for mossy fibers in the cerebellum, and 91

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for thalamo-cortical (Bredt and Snyder, 1994) and nigro-striatal (Derer and Derer, 1993) connections. Also the spatiotemporal pattern of NADPH-d expression observed in thalamic and tectal neurons matches with the arrival of innervation to these regions (Williams et al., 1994; Bertini and Bentivoglio, 1997). NMDA receptor-mediated postsynaptic mechanisms are thought to play important roles in the process of establishment of precise connections between different areas (Cline and Constantine-Paton, 1989, 1990; Simon et al., 1992). Nitric oxide is released in a calciumdependent manner by some NADPH-d-positive cells in response to glutamate activation of NMDA receptors, which suggests that this neuroactive substance may play a role in the establishment of axonal projections, in the elimination of redundant connections and/or in the refinement of axonal synapsis in the late development (Gally et al., 1990). Experimental evidence supports this hypothesis. Nitric oxide synthesis blockade alters the normal pattern of development of connections, e.g. in the transient ipsilateral projection from the retina to the optic tectum in the chicken (Wu et al., 1994), or the complex pattern of lamination of the retinogeniculate projection in the ferret (Cramer et al., 1996). Nitric oxide is also involved in activity-dependent establishment of connections in developing and regenerating olfactory neurons (Roskams et al., 1994). Several brain nuclei display a transient expression of NADPH-d activity in critical periods of development. In the lateral geniculate nucleus, the peak of NADPH-d/NOS activity occurs at the developmental stages that require afferent activity and postsynaptic NMDA receptor activation for the establishment of the definitive pattern of innervation (Bertini and Bentivoglio, 1997). But in other regions this transient expression of NADPH-d/NOS may be due to the influence of nitric oxide in cell survival and differentiation at late developmental stages. Thus, there are evidences indicating a role for nitric oxide in the regulation of apoptosis, growth arrest and dendritic branching on cortical neurons (Bredt and Snyder, 1994; Peunova and Enikolopov, 1995; Palluy and Rigaud, 1996). 4.4. NADPH-DIAPHORASE/NITRIC OXIDE SYNTHASE DISTRIBUTION PATTERN AND NITRIC OXIDE FUNCTIONAL IMPLICATIONS Nitric oxide in the brain acts in different ways. Firstly, nitric oxide functions as retrograde messenger (see Good, 1996). In a model of brain activity such a retrograde messenger is involved in the mechanism of inducing and/or maintaining long-term memory (O'Dell et al., 1991; Chapman et al., 1992; Izumi et al., 1992). A second role for nitric oxide in the brain is more likely that of a normal neurotransmitter in that it acts upon nerve cells other than the presynaptic nerve cell although direct proximity is not a prerequisite. It also seems probable that nitric oxide in the brain plays a role in the cerebral blood supply (Raszkiewicz et al., 1992; Rosenblum, 1992). The regulatory function of nitric oxide in transmitter release seems to be correlated to the different NOS isoforms in different neuronal subsets. Knock-out mice for neuronal NOS show a substantial reduction of NMDA-mediated glutamate release, whereas in the knock-out for endothelial NOS there is a considerable reduction of GABArelease (Kano et al., 1998). Therefore, the activity of each isoform seems to be in parallel to the regulation of excitatory or inhibitory neurotransmission, and thus, the expression of distinct NOS isoforms in particular neuronal subsets could lead to distinct regulatory mechanisms in transmitter release. It has been proposed that nitric oxide regulates neurotransmitter releasing through cGMP-dependent protein phosphorylation (O'Dell et al., 1991; Hirsch et al., 1993; Meffert et al., 1994; Schuman and Madison, 1994; Garthwaite and Boulton, 1995; Kuriyama and 92

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Ohkuma, 1995; Nei et al., 1996; Bogdanov and Wtirtman, 1997). Nitric oxide has been proved to be directly involved in the releasing of corticotropin-releasing factor and histamine in the hypothalamus (Karanth et al., 1993; Prast et al., 1997), and acetylcholine in the striatum (Prast et al., 1994). Some transmitters are released after stimulation of NMDA glutamate receptors, and NOS inhibitors block the transmitter release in synaptosomes obtained from cerebral cortex and striatum (Hirsch et al., 1993; Montague et al., 1994). Other experiments using nitric oxide donors have also demonstrated the involvement of this molecule in the regulation of the releasing of luteinizing hormone, oxytocin and luteinizing hormone-releasing hormone (Aguan et al., 1996; Okere et al., 1996; Luckman et al., 1997; Rettori et al., 1997), thus acting at the neuroendocrine control level. Even, the releasing of glutamate, the transmitter that indirectly activates neuronal NOS, is also regulated through a nitric oxide- and cGMP-dependent pathway (Sistiaga et al., 1997). This mechanism may have implications in the nitric oxide-mediated neurotoxicity. The presence of NOS isoforms with specific distribution and regulated expression makes the analysis of these functional processes difficult. In this sense, neuronal NOS and endothelial NOS are present in different elements of the hippocampus (Wendland et al., 1994; Gonz~ilezHern~ndez et al., [996; Vaid et al., 1996). The presence of distinct NOS isoforms in neurons acting as pre- and post-synaptic elements can be related to the specific function of nitric oxide in each of these elements (Iadecola, 1997), given their distinct regulatory mechanisms. In addition, the stimulation and tissue manipulation carried out in these experiments may induce the expression of different NOS isoforms, that would respond in different ways to inhibitors (see Iadecola, 1997). In conclusion, establishing a functional effect for nitric oxide based upon data from the distribution pattern of nitric oxide synthesizing cells is more complex than it has usually been for other transmitter systems. This is due to the alternative pathways for nitric oxide production that can be misinterpreted by the detection system and the difficulties to localize target elements. We have described the characteristics of the NADPH-d labeling in all major groups of vertebrates and during the rat brain development. Whereas consistent patterns are usually found between different animals under the same normal conditions, the NADPH-d/NOS staining is drastically altered after experiments or diseases. It has been suggested that the overproduction of nitric oxide could be responsible for brain damage and certain neurodegenerative conditions, and it seems to be involved in the neurotoxicity of Alzheimer's disease, Huntington's disease, and cerebral ischemia (Hoffman, 1991), alcohol-induced brain damage (Lancaster et al., 1992), cerebral stroke (Nowicki et al., 1991) and in the neuropathology associated with AIDS dementia complex (Morgan et al., 1992). We have reviewed those changes in the hippocampal formation (Alonso et al., 2000), one of the brain regions where the effects of a higher variety of conditions on the NADPH-d/NOS staining have been explored. Alterations in the NADPH-d staining pattern include NADPH-d-positive staining of previously negative neuronal types after mechanical damage (Divac et al., 1993; Regidor et al., 1993) or associated with epilepsy (Talavera et al., 1997). In other cases, a reduction on the number of NADPH-d-stained elements is observed. In epilepsy such activity loss has been described as transient (Stringer and Erden, 1995) or permanent (Miettinen et al., 1995; Kotti et al., 1997), but it seems to be a long-lasting alteration in schizophrenia (Akbarian et al., 1993), ischemic damage (Hong et al., 1993), or treatments with DSP-4 neurotoxin (Zhang and Yu, 1995). Other changes in the NADPH-d/NOS staining pattern in the hippocampus include the relative sparing of NADPH-d-positive cells in comparison with NADPH-d-negative neurons after neurotoxicity induced by glutamate agonists (Wolf et al., 1993) and in Alzheimer's disease (Hyman et al., 1992), or changes in NOS activity or NOS mRNA production without affecting 93

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the number of NOS-containing neurons as reported in aging (Yamada and Nabeshima, 1998) and chronic ethanol exposure (Kimura and Brien, 1998). Changes in the NADPH-d staining pattern have been also reported in the fiber staining. There is an increase in the density of NADPH-d fiber staining as a result of aging (Sobreviela and Mufson, 1995) and a reduction in the density of NADPH-d fiber staining in Alzheimer's disease (Rebeck et al., 1993). Furthermore, NADPH-d staining can be found in previously negative glial cells following ischemia (Endoh et al., 1993, 1994b; Kato et al., 1994), in epilepsy (Lerner-Natoli et al., 1994), after mechanical or photothrombic damage (Regidor et al., 1993; Wallace and Bisland, 1994; Bidmon et al., 1998; Stojkovic et al., 1998) or intense neuronal activity (Wallace and Bisland, 1994). These changes have very different time courses (from minutes to months) and they do not affect all brain regions or neuronal populations to the same extent. This capacity for plasticity is therefore another aspect that has to be considered when the distribution pattern of NADPH-d/NOS is evaluated. What contributes to the selective vulnerability of certain subsets of NADPH-d/NOS-positive neurons will be one of the most important subjects for future investigation.

5. ABBREVIATIONS

A Ac Ad AD ADT Aid Alh Am Amb Amc Ame AMH AMT APH Apl Apm AV AVT B bci BCI BL BM BST CA1 CA2 CA3 CA 94

nucleus anterior thalami nucleus accumbens nucleus tegmentalis anterodorsalis area dorsalis amygdalae nucleus anterodorsalis thalami archistriatum intermedium, pars dorsalis area lateralis hypothalami amygdala nucleus ambiguus nucleus centralis amygdalae nucleus externus amygdalae nucleus anteromedialis hypothalami nucleus anteromedialis thalami area parahippocampalis amygdala, pars lateralis amygdala, pars medialis area ventralis amygdalae area ventralis tegmenti nucleus basalis Meynert bracchium colliculi inferioris nucleus bracchii colliculi inferioris nucleus basolateralis amygdalae nucleus basomedialis amygdalae bed nucleus of the stria terminalis hippocampus, area CA1 hippocampus, area CA2 hippocampus, area CA3 commissura anterior

Comparative and developmental neuroanatomical aspects of the NO system CAA Cans Cb CC CCb CCI CD Cgus CH CHa ci CI C1 CLI CM cmsp CO CoPL CoPM CP Cpop CPu CS Ct CTc Cu CV Cven Cx Cxd Cxla Cxmp D DB DBH DBV Dc Dd D1 DLA DLI Dlm Dlp Dm DM DMH DMTg Dp

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nucleus corticalis anterioris amygdalae commissura ansulata cerebellum corpus callosum corpus cerebelli nucleus centralis colliculi inferioris nucleus cochlearis dorsalis commissura nuclei gustatorii secundarii commissura horizontalis commissura habenularum capsula interna colliculus inferior claustrum nucleus centralis lobi inferioris hypothalami corpus mammillare columna motoria spinalis chiasma opticum nucleus corticalis posterolateralis amygdalae nucleus corticalis posteromedialis amygdalae commissura posterior commissura postoptica = commissura supraoptica caudatus putamen colliculus superior nucleus centralis thalami commissura tecti nucleus cuneatus nucleus cochlearis ventralis commissura ventralis rhombencephali cortex cortex dorsalis cortex lateralis anterior cortex medialis, pars parvocellularis dorsal field of the spinal cord nucleus of the diagonal band of Broca horizontal limb of the diagonal band of Broca vertical limb of the diagonal band of Broca area dorsalis telencephali, zona centralis area dorsalis telencephali, zona dorsalis area dorsalis telencephali, zona lateralis nucleus dorsolateralis anterior thalami nucleus diffusus lobi inferioris hypothalami nucleus dorsolateralis thalami, pars magnocellularis nucleus dorsolateralis thalami, pars parvocellularis area dorsalis telencephali, zona medialis nucleus dorsomedialis thalami nucleus dorsomedialis hypothalami nucleus tegmentalis dorsomedialis area dorsalis telencephali, zona posterior 95

Ch. III

DPT DPTg dso dsum DV DVR E EG En EW f Fi fll tim fit fmt fr FRM FStr G GC GD Gi GLD GLV GM GP GPt Gr GS HA Hab HbL HbM Hp HV IA

icj ICo IG Inf IP Is Ism Isp La LC LCa 96

J.R. Alonso et al.

nucleus dorsalis posterior thalami nucleus tegmentalis dorsalis pericentralis decussatio supraoptica decussatio supramammillaris radix descendens nervi trigemini dorsal ventricular ridge ectostriatum eminentia granularis nucleus endopiriformis nucleus Edinger-Westphal fornix fimbria hippocampi fasciculus longitudinalis lateralis fasciculus longitudinalis medialis fasciculus lateralis telencephali fasciculus medialis telencephali fasciculus retroflexus formatio reticularis mesencephali fundus striati glomerus olfactorius griseum centrale gyrus dentatus nucleus gigantocellularis reticuli nucleus geniculatus laterodorsalis nucleus geniculatus lateroventralis nucleus geniculatus medialis globus pallidus griseum praetectale nucleus gracilis nucleus gustatorius secundarius hyperstriatum accessorium habenula nucleus habenularis lateralis nucleus habenularis medialis hippocampus hyperstriatum ventrale colliculus superior, stratum album intermedium insula Callejae nucleus intercollicularis colliculus superior, stratum griseum intermedium superior colliculus superior, stratum inferius nucleus interpeduncularis nucleus isthmi nucleus isthmi, pars magnocellularis nucleus isthmi, pars parvocellularis nucleus lateralis thalami, pars anterior locus coeruleus lobus caudalis cerebelli

Comparative and developmental neuroanatomical aspects of the NO system LD LDTg 11 LLi LM lm Lm LP LRt LVII LX MC Mdd Mdv ME Me ML MLd MM MnV Mp mt N NCP NCPa NDBC NDV ne Nflm nIII NIII NIV nIV NIX-X NIX NLV NOB NPM NPT Nsa Nsd Nsi Nsl Nsol NT NTA NVI NVII

Ch. III

nucleus laterodorsalis thalami nucleus tegmentalis laterodorsalis lemniscus lateralis nucleus ventralis lemnisci lateralis lateral motor field of spinal cord lemniscus medialis nucleus lentiformis mesencephali nucleus lateroposterioris thalami nucleus reticularis lateralis lobus facialis lobus vagus nucleus mediocentralis thalami nucleus medullaris reticularis, pars dorsalis nucleus medullaris reticularis, pars ventralis eminentia mediana nucleus medialis amygdalae nucleus mammillaris lateralis nucleus mesencephalicus lateralis, pars dorsalis nucleus mammillaris medialis nucleus motorius nervi trigemini nucleus medialis posterior tractus mammillaris thalami neostriatum nucleus commissurae posterioris nucleus commissurae pallii nucleus decussationis bracchiorum conjunctivorum nucleus descendens nervi trigemini neuroepithelium nucleus fasciculi longitudinalis medialis nervus oculomotorius nucleus nervi oculomotorii nucleus nervi trochlearis nervus trochlearis nucleus nervi glossopharyngei et motorius nervi vagi nucleus nervi glossopharyngei nucleus lateralis valvulae nucleus opticus basalis nucleus profundus mesencephali nucleusposterior thalami nucleus septalis anterior nucleus septalis dorsalis nucleus septalis impar nucleus septalis lateralis nucleus fasciculi solitarii nucleus taeniae nucleus tuberis anterior nucleus nervi abducentis nucleus nervi facialis 97

Ch. III

NVIIId NVIIIv NVme nX NX NXII 0 OAd OA1 OAm OAv OI Ols OM OMa OS P Pa PA PB PC pc Pd PG PG1 PGm PHc PHd PHv pir Pir PL P1 PLs PM Pm PMD PMn PMV PnC PnO PnV PO POa POL POM Pp PP 98

J.R. Alonso et al.

nucleus octavus dorsalis nucleus octavus ventralis nucleus mesencephalicus nervi trigemini nervus vagus nucleus nervi vagi nucleus nervi hypoglossi colliculus superior, stratum opticum nucleus olfactorius, pars dorsalis anterior nucleus olfactorius, pars lateralis anterior nucleus olfactorius, pars medialis anterior nucleus olfactorius, pars ventralis anterior oliva inferior nucleus olivae superioris nucleus octavolateralis medialis nucleus octavus magnocellularis populatio octavia secundaria nucleus pontinus nucleus paraventricularis hypothalami paleostriatum augmentatum nucleus parabracchialis nucleus paracentralis thalami pedunculus cerebelli pallium dorsale area praeglomerulosa nucleus praeglomerulosus lateralis nucleus praeglomerulosus medialis nucleus periventricularis hypothalami, zona caudalis nucleus periventricularis hypothalami, zona dorsalis nucleus periventricularis hypothalami, zona ventralis tractus piramidalis cortex piriformis nucleus pontis lateralis pallium laterale nucleus perilemniscalis nucleus praeopticus magnocellularis pallium mediale nucleus praemammillaris dorsalis nucleus raphe paramedianus nucleus praemammillaris ventralis nucleus pontino-centralis reticuli nucleus pontino-oralis reticuli nucleus pontino-ventralis reticuli area praeoptica area praeoptica anterior nucleus praeopticus lateralis nucleus praeopticus medialis area praeoptica periventricularis paleostriatum primitivum

Comparative and developmental neuroanatomical aspects of the NO system PPo PPp PPTg pr PrG Prv PrXII PT Pta PTA Pth PTp PV Pv PVN PVT R

Rad Rai Ram Ras Re Ri Rm ROb ROT RPa Rpgl rs

Rti Rtp Rts Ru S SC SCd sg SGC SGFS sgl sgr SHp SI SLD SLI SLV sm SM

Ch. III

nucleus praetectalis posterior nucleus praeopticus parvocellularis, pars posterior nucleus pedunculopontinus tegmenti area fibrillaris periventricularis thalami colliculus superior, stratum griseum profundum nucleus praetectalis ventralis nucleus praepositus hypoglossi nucleus parataenialis thalami nucleus praetectalis accessorius nucleus praetectalis anterior pallial thickening nucleus periventricularis tuberculi posterioris nucleus paraventricularis posterioris thalami pallium ventrale nucleus paraventricularis paleostriatum ventrale nucleus raphe nucleus raphe dorsalis nucleus raphe inferior nucleus raphe medianus nucleus raphe superior nucleus renius thalami nucleus reticularis inferior nucleus reticularis medialis nucleus raphe obscurus nucleus rotundus nucleus reticularis paramedianus nucleus reticularis paragigantocellularis lateralis tractus rubrospinalis area reticularis intermedia area reticularis parvocellularis area reticularis superior nucleus ruber subiculum nucleus suprachiasmaticus nucleus subcoeruleus dorsalis stratum glomerulare stratum griseum centrale stratum griseum et fibrosum superficiale bulbus olfactorius accessorius, stratum glomerulare stratum granulare nucleus septohippocampalis substantia innominata septum laterale, pars dorsalis septum laterale, pars intermedia septum laterale, pars ventralis stria medullaris septum mediale 99

Ch. III

smi SN SO sol spe SPFPT spV ss

SStr st

STh Str SubB SuG SUM SV SY T tbs TD TeB Tg tgs Th TL TLa tmca Tmg TO TOL tol tpm TPO Tr.op TR TS TSc TSco TS1 TT ttb ttbc Tu tvs

V VC1 VCm Vd 100

J.R. Alonso et al.

stratum mitrale substantia nigra nucleus supraopticus tractus solitarius stratum plexiforme extemum nucleus subparafascicularis parvocellularis thalami tractus spinalis trigeminalis stratum subependimarium area substriatalis stria terminalis thalami nucleus subthalamicus striatum nucleus subbrachialis colliculus superior, stratum griseum superficiale nucleus supramammillaris nucleus spinalis trigemini sulcus ypsiloniformis nucleus tangentialis tractus bulbospinalis nucleus tegmentalis dorsalis telencephalon basalis tegmentum tractus gustatorius secundarius thalamus torus longitudinalis torus lateralis tractus mesencephalo-cerebellaris anterior torus semicircularis, nucleus magnocellularis tectum opticum nucleus tractus olfactorii lateralis tractus olfactorius lateralis tractus praectecto-mammillaris area temporo-parieto-occipitalis tractus opticus nucleus tegmentalis rostralis torus semicircularis torus semicircularis, nucleus centralis torus semicircularis, nucleus commissuralis torus semicircularis, nucleus laminaris taenia tecta tractus tectobulbaris tractus tectobulbaris cruciatus tuberculum olfactorium tractus vestibulospinalis ventriculum valvula cerebelli, pars lateralis valvula cerebelli, pars medialis area ventralis telencephali, nucleus dorsalis

C o m p a r a t i v e a n d d e v e l o p m e n t a l n e u r o a n a t o m i c a l a s p e c t s o f the N O s y s t e m Ve

area vestibularis

Vedl

n u c l e u s vestibularis d o r s o l a t e r a l i s

Vem

n u c l e u s vestibularis m e d i a l i s

Vesp

n u c l e u s vestibularis spinalis

Vev

n u c l e u s vestibularis ventralis

Vl

area ventralis t e l e n c e p h a l i , n u c l e u s lateralis

Vm

nucleus ventromedialis thalami

VMH

nucleus ventromedialis hypothalami

VT

n u c l e u s ventralis t h a l a m i

Vv

area ventralis t e l e n c e p h a l i , n u c l e u s ventralis z o n a incerta

ZI

Ch. III

6. REFERENCES Afework M, Tomlinson A, Belai A, Burnstock G (1992): Colocalization of nitric oxide synthase and NADPHdiaphorase in rat adrenal gland. Neuroreport 3:893-896. Aguan K, Mahesh VB, Ping L, Bhat G, Brann DW t1996): Evidence for a physiological role for nitric oxide in the regulation of the LH surge: effect of central administration of antisense oligonucleotides to nitric oxide synthase. Neuroendocrinology 64:449-455. Akbarian S, Bunney WE, Pot'kin SG, Wigal SB, Hagman JO, Sandman CA. Jones EG (1993): Altered distribution of nicotinamide-adenine dinucleotide phosphate-diaphorase cells in frontal lobe of schizophrenics implies disturbances of cortical development. Arch Gen Psvchiatlw 50:169-177. Alonso JR, S~inchez F, Ar6valo R, Carretero J, Vfizquez R, Aij6n J (1992a): Partial coexistence of NADPHdiaphorase and somatostatin in the rat hypothalamic paraventricular nucleus. Neurosci Lett 148:101-104. Alonso JR, SS.nchez F, Ar6valo R, Carretero J, Aij6n J. V~zquez R (1992b): Calbindin D-28k and NADPHdiaphorase coexistence in the magnocellular neurosecretory nuclei. Neuroreport 3:249-252. Alonso JR, Ar~valo R, Porteros A, Brifi6n JG. Lara J. Aij6n J (1993): Calbindin D-28k and NADPH-diaphorase activity are localized in different populations of periglomerular cells in the rat olfactory bulb. J Chem Neuroanat 6: l-6. Alonso JR, Ar6valo R, Brifi6n JG, Garcfa-Ojeda E, Porteros A, Aij6n J. (1995a): NADPH-diaphorase staining in the central nervous system. Neurosci Protoc 050-04:1-11. Alonso JR, Ar6valo R, Garcfa-Ojeda E, Porteros A, Brifi6n JG. Aij6n J (1995b): NADPH-diaphorase active and calbindin D-28k-immunoreactive neurons and fibers in the olfactory bulb of the hedgehog (Erinaceus europaeus). J Comp Neurol 351:307-327. Alonso JR, Porteros A, Crespo C, Ar6valo R, Brifi6n JG. Weruaga E. Aij6n J (1998): Chemical anatomy of the macaque monkey olfactory bulb: NADPH-diaphorase/nitric oxide synthase activity. J Comp Neurol 402:419434. Alonso JR, Pitkanen A, Amaral DG (2000): NADPH-diaphorase active neurons and fibers in the Macaca fascicularis monkey hippocampal formation. Submitted. Anken RH, Rahmann H (1996): An atlas of the distribution of NADPH-diaphorase in the brain of the highly derived swordtail fish Xiphopkorus helleri (Atherinoformes: Teleostei). J Brain Res 37:421-449. Ar6valo R, S~.nchez F, Alonso JR, Carretero J, V:~zquez R, Aij6n J (1992): NADPH-diaphorase activity in the hypothalamic magnocellular neurosecretory nuclei of the rat. Brain Res Bull 28:599-603. Ar6valo R, S~inchez F, Alonso JR, Rubio M, Aij6n J. V,~zquez R (1993): Infrequent cellular coexistence of NADPH-diaphorase and calretinin in the neurosecretory nuclei and adjacent areas of the rat hypothalamus. J Chem Neuroanat 6:335-341. Ar6valo R, Alonso JR, Garcfa-Ojeda E, Brifi6n JG, Crespo C. Aij6n J (1995): NADPH-diaphorase in the central nervous system of the tench (Tinca tinca L., 1758). J Comp Neurol 352:398-420. Bell TD, Pereda AE, Faber DS (1997): Nitric oxide synthase distribution in the goldfish Mauthner cell. Neurosci Lett 226:187-190. Bertini G, Bentivoglio M (1997): Nitric oxide synthase in the adult and developing thalamus: histochemical and immunohistochemical study in the rat. J Comp Neurol 388:89-105. Bidmon HJ, Wu J, Buchkremer-Ratzmann I, Mayer B. Witte OW, Zilles K (1998): Transient changes in the 101

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somatostatin immunohistochemistry, and silver impregnation in detecting structural and functional impairment in experimental status epilepticus. Neuroscience 80:105-117. Kowall NW, Beal ME Ferrante RJ, Martin JB (1985): Topography of nicotinamide adenine dinucleotide phosphatediaphorase staining neurons in rat striatum. Neurosci Lett 59:61-66. Kubrina LN, Caldwell WS, Mordvintcev PI, Malenkova IV. Vanin AF (1992): EPR evidence for nitric oxide production from guanidino nitrogens of L-arginine in animal tissues in vivo. Biochim Biophys Acta 1099:233237. Kugler P, Drenckhahn D (1996): Astrocytes and Bergmann glia as an important site of nitric oxide synthase I. Glia 16:165-173. Kulkarni AE Getchell TV, Getchell ML (1994): Neuronal nitric oxide synthase is localized in extrinsic nerves regulating perireceptor processes in the chemosensory nasal mucosae of rats and humans. J Comp Neurol 345:125-138. Kuonen DR, Kemp MC, Roberts PJ (1988): Demonstration and biochemical characterization of rat brain NADPHdependent diaphorase. J Neurochem 50:1017-1025. Kuriyama K, Ohkuma S (1995): Role of nitric oxide in central synaptic transmission: effects on neurotransmitter release. Jpn J Pharmacol 69:1-8. Lancaster JR, Langrehr JM, Bergonia HA, Murase N, Simmons RL, Hoffman RA (1992): EPR detection of heine and nonheme iron-containing protein nitrosylation by nitric oxide during rejection of rat heart allograft. J Biol Chem 26 7:10994-10998. Lerner-Natoli M, De Bock F, Bockaert J, Rondouin G (1994): NADPH diaphorase-positive cells in the brain after status epilepticus. Neuroreport 5:2633-2637. Li J, Chert S, Lin RC, Smith SS (1997): Cerebellar nitric oxide synthase is expressed within granule cell patches innervated by specific mossy fiber terminals: a developmental profile. Dev Neurosci 19:274-282. Li ZS, Burness JB, Young HM, Campbell G (1992): Nitric oxide synthase immunoreactivity and NADPH diaphorase enzyme activity in neurons of the gastrointestinal tract of the toad, Bufo marinus. Arch Histol Cvtol 55:333-350. L6wenstein CJ, Snyder SH (1992): Nitric oxide, a novel biologic messenger. Cell 70:705-707. Luckman SM, Huckett L, Bicknell RJ, Voisin DL, Herbison AE (1997): Up-regulation of nitric oxide synthase messenger RNA in an integrated forebrain circuit involved in oxytocin secretion. Neuroscience 77:37-48. Luebke JL, Weider JM, McCarley RW, Greene RW (1992): Distribution of NADPH-diaphorase positive somata in the brainstem of the monitor lizard Varanus exanthematicus. Neurosci Lett 148:129-132. Marfn O, Gonz~ilez A, Smeets WJAJ (1995): Evidence tot a mesolimbic pathway in anuran amphibians: a combined tract-tracing/immunohistochemical study. Neurosci Lett 190:183-186. Matsumoto T, Nakane M, Pollock JS, Kuk JE, F/3rstermann U (1993): A correlation between soluble brain nitric oxide synthase and NADPH-diaphorase activity is only seen after exposure of the tissue to fixative. Neurosci Lett 155:61-64. Meek J (1994): Catecholamines in the brains of Osteichthyes (bony fishes). In: Smeets WJAJ, Reiner A (Eds), Phylogeny and Development of Catecholamine Systems in the CNS ~( Vertebrates. Cambridge: Cambridge University Press, pp 49-76. Meffert MK, Haley JE, Schuman EM, Schulman H, Madison DV (1994): Inhibition of hippocampal heme oxygenase, nitric oxide synthase, and long-term potentiation by metalloporphyrins. Neutvn 13:1225-1233. Miettinen R, Kotti T, Halonen T, Riekkinen P (1995): NADPH diaphorase-containing nonpyramidal cells in the rat hippocampus exhibit differential sensitivity to kainic acid. Eur J Nemvsci 7:1822-1825. Minc-Golomb D, Yadid G, Tsarfaty I, Resau JH, Schwartz JP (1996): In vivo expression of inducible nitric oxide synthase in cerebellar neurons. J Neurochem 66:1504-1509. Mizukawa K, McGeer PL, Vincent ST, McGeer EG (1988): Ultrastructure of reduced nicotinamide adenine dinucleotide phosphate (NADPH) diaphorase-positive neurons in the cat cerebral cortex, amygdala and caudate nucleus. Brain Res 452:286-292. Mizukawa K, Vincent SR, McGeer PL, McGeer EG (1989): Distribution of reduced nicotine-adenine-dinucleotidephosphate-diaphorase-positive cells and fibers in the cat central nervous system. J Comp Neutvl 279:281311. Montague PR, Gancayco CD, Winn MJ, Marchase RB, Friedlander MJ (1994): Role of NO production in NMDA receptor-mediated neurotransmitter release in cerebral cortex. Science 263:973-977. Morgan MJ, Kimes AS, London ED (1992): Possible roles for nitric oxide in AIDS and associated pathology. Med Hypoth 38:189-193. Mufson EJ, Brady DR, Carey RG (1990): Reduced nicotinamide adenine dinucleotide phosphate-diaphorase 105

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CHAPTER IV

Nitric oxide in the retina W.D. ELDRED

1. I N T R O D U C T I O N In the past few years, there has been an explosion of interest in the role of nitric oxide (NO) in neural function, development and pathology. Although numerous studies have focused on the brain, many of the most detailed studies have examined the retina. In many ways, the retina is an ideal model system to study NO. The anatomy and physiology of many retinal neurons have been well characterized in a wide variety of species, and numerous detailed comparisons of the same cell types in different species have been made. This comparative approach has an advantage over many studies in brains that have analyzed many diverse neuronal systems in a limited number of mammalian species. Detailed cross-species comparisons will establish which functions of NO are universal cellular mechanisms and which are more species-specific. The retina also has the advantage that, within certain limits, all regions of a retina have similar anatomical connections and use the same physiological mechanisms for visual processing. Finally, some of the best characterized retinal model systems, such as salamanders or turtles, have large cells that are ideal for electrophysiology and have very well-developed NO signal transduction pathways. Thus, the retina provides a perfect blend of well-characterized anatomical and physiological complexity to examine the roles of NO in neural function. NO represents a new class of neurotransmitters that serve a variety of synaptic and non-synaptic functions that are not characteristic of traditional neurotransmitters. It is not like a classic neurotransmitter in that it is made on demand and not stored for vesicular release. Its action is not restricted to specific morphologically specialized sites and it can rapidly diffuse to trigger cGMP synthesis in cells that are not connected synaptically. The cGMP that is produced can in turn activate ion channels, protein kinase G or phosphodiesterases (PDEs). NO can also have direct effect on target proteins like NMDA receptors or ADP-ribosyl transferase. This review is not intended to be an exhaustive of NO in the entire visual system, but it will summarize many aspects of NO in the retina and suggest new areas for future research. For recent reviews of the role of NO in a more general ocular function and its role in retinal and ocular pathology, please see Goldstein et al. (1996) or Becquet et al. (1997).

2. L O C A L I Z A T I O N OF NITRIC OXIDE SYNTHASE IN THE RETINA Historically, the first localizations of nitric oxide synthase (NOS) in the retina were done before it was discovered. This is because NADPH-diaphorase histochemistry was first used over 40 years ago (Wislocki and Sidman, 1954) to selectively stain retinal neurons. Only Handbook of Chemical Neuroanatom3; Vol. 17: Functional Neuroanatonly o.f the :Vitric Oxide System H.W.M. Steinbusch, J. De Vente and S.R. Vincent, editors (~) 2000 Elsevier Science B.V. All rights reserved.

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much later (Vincent and Hope, 1992), did the NADPH-diaphorase activity become associated with NOS. The precise localization of NOS at the cellular and subcellular levels is critical to provide insight into mechanisms by which NO is functionally involved in the signal transduction in the retina. Although sites of NO synthesis do not necessarily correlate with the sites of action, they must be in close proximity given the short half-life of NO (Vincent, 1994). 2.1. METHODOLOGICAL CONSIDERATIONS

2.1.1. NADPH-diaphorase histochemistry Since the enzymatic activity of NOS requires a cofactor NADPH, histochemistry has been used extensively to localize NOS through the reduction of tetrazolium salts to an insoluble formazan reaction product. NADPH-diaphorase is still widely used to localize NOS in the retina because of its effectiveness and simplicity. More recently, immunocytochemistry using antibodies directed against specific isoforms of NOS has also gained wide acceptance. Both NADPH-diaphorase histochemistry and NOS immunocytochemistry each have their own advantages and disadvantages, which can lead to difficulties in the interpretations of the anatomical localizations. Although NADPH-diaphorase histochemistry is very easy and inexpensive, there are some difficulties with this method. First, there are at least six other enzymes dependent on NADPH (Schonfelder et al., 1993), such as cytochrome P450 reductase that has a 58% sequence homology with NOS. However, paraformaldehyde fixation is reported to destroy 100% of particulate diaphorase and retain 40-60% of the cytosolic fixation-resistant diaphorase that is thought to be real NOS (Matsumoto et al., 1993; Tracey et al., 1993). Another possible cross-reaction with the NADPH-diaphorase-dependent enzyme, DT-diaphorase, is shown to be inhibited by 10-100 ~M dicumarol (Riley and Workman, 1992). Such potential difficulties in correlating NOS with NADPH-diaphorase are given strong support by a comprehensive study that compares the localizations of NADPH-diaphorase staining to the immunocytochemical localizations of nNOS, eNOS, iNOS and cytochrome P450 reductase in a wide range of tissues (Young et al., 1997). These authors find that cells that are immunoreactive for any of these four NADPH-dependent enzymes also have NADPH-diaphorase in formaldehyde-fixed tissue, but that in some tissues there is NADPH-diaphorase staining when none of these enzymes can be localized immunocytochemically. There are also two variations of the NADPH-diaphorase reaction: the direct method which adds NADPH to the reaction mix, and the indirect method that depends on activity of other enzymes in the tissue itself, like glucose-6-phosphate dehydrogenase enzyme or malic enzyme to make NADPH from NADP (Sandell, 1985). Detailed examination of the literature indicates that although the direct method is easier and is reported to produce more intense staining, there is more variation in the results of different investigators using the same method, than between the indirect and direct methods. They both can produce excellent results if done well. One remaining source of ambiguity is that NADPH-diaphorase histochemistry cannot distinguish between different isoforms of NOS (Young et al., 1997), some of which may remain to be discovered. This is particularly relevant when one is trying to establish the function of NOS, in that all NOS isoforms possibly have NADPH-diaphorase activity, but they may have very distinct regulatory mechanisms and functions. There is also the question of the ultrastructural localization of NADPH-diaphorase. To localize NADPH-diaphorase at the electron microscopic level, one must use a special 112

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tetrazolium salt, to form an osmiophilic formazan deposit. At the ultrastructural level in the retina, the subsequent BSPT reaction product is localized on the endoplasmic reticulum, the nuclear membrane, mitochondria and the Golgi apparatus (Darius et al., 1995; Villani and Guarnieri, 1996). These seemingly metabolic subcellular localizations are difficult to reconcile with the known synaptic localizations of NOS using immunocytochemical methods (Haverkamp and Eldred, 1998b). However, the BSPT reaction product is not present in NOS-knockout mice (Darius et al., 1995), which argues for the involvement of NOS. It is possible that the BSPT might be localizing an alternate novel isoform of NOS, and further studies will be necessary to resolve this issue.

2.1.2. NOS immunocytochemistry In recent years, many excellent commercial antibodies against several NOS isoforms have become available. In some cases these antibodies produce the same labeling in the retina as seen with NADPH-diaphorase histochemistry (Roufail et al., 1995), while in other cases there are some differences (Perez et al., 1995; Blute et al., 1997). In many cases, species differences may prevent good labeling with NOS immunocytochemistry. Another consideration is that these antibodies are ideally directed against a single isoform of NOS, which prevents the detection of other isoforms of NOS. These methods do have the advantage that the same NOS antibodies can also be used for Western blots to more precisely biochemically characterize the antigens being labeled (Goureau et al., 1997). Two studies use immunocytochemical methods at the electron microscopic level to localize NOS in the retina (Roufail et al., 1995; Haverkamp and Eldred, 1998b), and the results of these studies will be described in detail later. 2.2. ISOFORMS OF NOS IN THE RETINA As discussed above, the NADPH-diaphorase histochemistry does not allow the specification of the NOS isoform involved. In some cases, even pharmacological studies cannot clarify the isoform involved because the high concentrations of isoform-specific NOS inhibitors needed in vivo or in vitro eliminate their isoform specificity (Wellard et al., 1995). This leaves only isoform-specific antibodies to distinguish the presence of certain NOS isoforms. The majority of retinal research focuses on the neuronal NOS (nNOS) isoform, while relatively few studies examine the presence of endothelial NOS (eNOS) or inducible NOS (iNOS). The existence of retinal nNOS is clearly established as described in the subsequent portions of this review. The presence of eNOS in the retina remains somewhat unresolved in that although Haberecht et al. (1998) report that there is no eNOS in the rabbit retina, Goureau et al. (1997) find a 135 kDa protein corresponding to eNOS in membrane fractions using Western blots and find eNOS-like immunoreactivity (-LI) in the inner nuclear layer (INL), in Mtiller cells at inner and outer limiting membranes, in the ganglion cell layer (GCL) and in the outer plexiform layer (OPL) in the chick retina. It is possible that these differences reflect species specificity in the presence of eNOS, or more likely, a difference in the recognition of an eNOS isoform by the antibodies used. However, detailed studies of eNOS in the brain indicate it is absent from neurons (Stanarius et al., 1997). The presence of the iNOS isoform is more clearly established in both normal and pathological retinas. Park et al. (1994) use primers of a conserved cofactor region of nNOS and iNOS for RT-PCR in human retina and find that the amino acid sequence of retinal nNOS has only several amino acid differences from brain NOS, and that retinal iNOS has 113

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only one amino acid difference from human chondrocyte iNOS. Both of these differences are only in the non-cofactor binding sites and indicate more than 99% homology. Their Northern blot analyses show a message for iNOS but not nNOS. L6pez-Costa et al. (1997) immunocytochemically localize iNOS in normal rat retina in labeled Mtiller cell somas in the INL and in Mtiller cell processes in OPL and IPL, in photoreceptor inner segments, in the OPL, and maybe in some horizontal cells, ganglion cells or displaced amacrine cells. Fujii et al. (1998) use in situ hybridization in chick to detect the expression of iNOS mRNA in the outer part of the photoreceptor layer and in the inner and outer parts of retinal pigment epithelium and choroid. Interestingly, Fujii et al. (1998) find that form deprivation reduces iNOS and nNOS mRNA, while only iNOS protein is reduced. This suggests that there is an ocular-specific regulation of the iNOS gene under non-pathological conditions. There is evidence that only specific pathological conditions may change iNOS expression in the retina. Patel et al. (1997) find iNOS in scattered cells in the INL and GCL, and in the IPL in normal rat retina. They report that optic tract lesions that cause severe ganglion cell loss do not increase levels of iNOS or nNOS, and they conclude that neither iNOS or nNOS have a role in cell death or degeneration. In contrast, Goureau et al. (1994a) report that stimulation with lipopolysaccharide, interferon-), and tumor necrosis factor-~ increases NO synthesis by human pigment epithelial cells, and that this effect can be blocked by selective iNOS inhibitors, cyclohexamide or transforming growth factor-J3. Stimulation with lipopolysaccharide, interferon-), and tumor necrosis factor-~ also increases NO synthesis by Mtiller cells (Goureau et al., 1994b), although each alone has no effect. In the case of Mtiller cells, they also find macrophage iNOS mRNA and the effect is again blocked by selective iNOS inhibitor, cyclohexamide and transforming growth factor-j3. These results indicate that in both human pigment epithelial cells and Mtiller cells, the expression of iNOS is transcriptionally regulated and that iNOS may be important in inflammation and uveitis in the eye. 2.3. ANATOMICAL LOCALIZATIONS OF NOS As discussed above, many of the localizations of NOS in the retina have been done using NADPH-diaphorase histochemistry which can simultaneously localize more than one isoform of NOS. This often gives a much broader localization than is seen using immunocytochemical methods. Thus, it is prudent to analyze the localization of NOS using both NADPH-diaphorase histochemistry and a combination of antibodies directed against several distinct epitopes of different NOS isozymes. In spite of these differences, there is a strong similarity in the localization of NOS in a variety of species. 2.3.1. Mammals Rats. Although the majority of localizations of NOS in the retina have been done in the rat,

there are considerable differences reported in the literature. The first detailed study done by Sandell (1985) finds NADPH-diaphorase only in sparse somas of presumptive amacrine cells in the INL, but provides little anatomical detail about the cells involved. Subsequent studies indicate that there might be two classes of amacrine cells with NADPH-diaphorase, one that is larger and heavily stained and a second population that is smaller and more weakly stained (L6pez-Costa et al., 1997). Later studies find that these amacrine cells contain GABA (Young and Vaney, 1989) and that there are three bands of NOS-LI in the IPL at strata 1 (S1), $2-$3 and at $5 (Yamamoto et al., 1993). 114

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Other labeled cells are described in the INL, some of which are classified as horizontal cells (Yamamoto et al., 1993; Huxlin, 1995; Sheng and Ignarro, 1996), Mtiller cells (Huxlin, 1995), or as bipolar or interplexiform cells (Yamamoto et al., 1993; Perez et al., 1995). The existence of NOS-LI or NADPH-diaphorase in rat photoreceptors is under debate. Although in some cases no labeled photoreceptors are described (NADPH-diaphorase - - Sandell, 1985; NOS i m m u n o c y t o c h e m i s t r y - Yamamoto et al., 1993; Perez et al., 1995; Sheng and Ignarro, 1996), in other cases labeled photoreceptors are reported (NADPH-diaphorase - - Yamamoto et al., 1993; Huxlin, 1995; Perez et al., 1995; Roufail et al., 1995; Sheng and Ignarro, 1996; NOS i m m u n o c y t o c h e m i s t r y - Roufail et al., 1995; Lrpez-Costa et al., 1997). A recent study using a combination of light and electron immunocytochemistry (Haverkamp and Eldred, 1998b) confirms the localization of nNOS-LI in photoreceptors, bipolar cells and horizontal cells in rat. Following the initial studies showing NADPH-diaphorase in amacrine cells, it is suggested that some of these amacrine cells are also displaced to the GCL (Huxlin, 1995; Perez et al., 1995; Sheng and Ignarro, 1996). A study combining retrograde labeling and ganglion cell degeneration (Huxlin and Bennett, 1995), proves that some of the diaphorase cells in the GCL are ganglion cells and that the rest are displaced amacrine cells. A number of studies (Mitrofanis, 1989; Huxlin, 1995; Perez et al., 1995; Roufail et al., 1995) indicate the presence of NADPH-diaphorase, but not nNOS-LI (Perez et al., 1995; Roufail et al., 1995; Sheng and Ignarro, 1996) in the rat retinal vasculature. This suggests that the NADPH-diaphorase in the vasculature is eNOS, as would be expected. Additionally, in both rats and humans, cell bodies and processes of both NADPH-diaphoraseand NOS-positive neurons are associated closely with the vasculature. The dendrites of these amacrine cells go into IPL and then project back to INL to form a pericapillary network. It is suggested that NO released from these amacrine cells may induce changes in the vascular diameter and modulate retinal blood flow (Roufail et al., 1995). To date, only one study has done electron immunocytochemistry to localize NOS in the rat retina. Haverkamp and Eldred (1998b) examine the localization of NOS in the OPL and find NOS in some rod bipolar and B-type horizontal cell axon terminals at rod ribbon synapses. The fact that not all rod bipolar and B-type horizontal cell axon terminals have NOS may indicate that there is more than one physiologically distinct cell type of each. It is also significant that the NOS-LI in these bipolar and horizontal cells is primarily confined to the processes of these cells in the photoreceptor terminal, with little cytoplasmic NOS in the rest of the cell body. These results indicate that NOS-LI can be selectively localized at synapses in potentially every nerve cell type involved in the OPL. Rabbits. The localization of NOS in the rabbit retina has been carefully examined and there is considerable agreement between the various reports. The initial NADPH-diaphorase studies done by Sandell (1985) find sparse well-labeled somas in the inner INL with poorly labeled processes that are assumed to be amacrine cells. A number of subsequent studies (Sagar, 1986, 1990; Vaney and Young, 1988; Osborne et al., 1993; Perez et al., 1995; Haberecht et al., 1998) clarify that there are two distinct classes of NADPH-diaphorase-positive afnacrine cells, the ND1 and ND2, that constitute about 1 in 1000 of the total number of amacrine cells. Both the ND 1 and ND2 cells contain GABA(Vaney and Young, 1988). There are ---.4000 NDI cells that are darkly stained and have well-defined dendrites up to 5 mm long (Sagar, 1990). The ND2 cells are more numerous ('~24,000 cells), but their dendrites are weakly stained and less than 100-200 Ixm long. These amacrine cells have their dendritic arborizations primarily in $3, with many varicose dendrites in other strata in the IPL (Vaney and Young, 1988). NOS immunocytochemistry (Perez et al., 1995; Haberecht et al., 1998) confirms the localization of 115

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nNOS-LI in the NDI and ND2 amacrine cells, although a small number of cells in the GCL and amacrine cell layer only have NADPH-diaphorase (Perez et al., 1995). In addition to the amacrine cells, NADPH-diaphorase is localized in the vascular endothelium, as diffuse staining in the OPL and IPL, in displaced amacrine cells, in weakly labeled large cell bodies in GCL and in photoreceptor inner segments and cone outer segments (Perez et al., 1995; Haberecht et al., 1998). The results of NOS immunocytochemistry are not as consistent. Koch et al. (1994) find NOS-LI primarily in photoreceptor ellipsoids, with some NOS-LI in photoreceptor inner segments and in cone outer segments but not rod outer segments. Although both Perez et al. (1995) and Haberecht et al. (1998) agree that NOS-LI is in ganglion cells, Haberecht et al. (1998) find NOS-LI in rod and cone inner segments and cone outer segments, while Perez et al. (1995) find no NOS-LI in photoreceptor inner segments, the OPL or in horizontal cells, but do describe its presence in bipolar and/or interplexiform cells. These divergent results may be the result of using different nNOS antibodies that differentially recognize distinct isoforms of nNOS. Cat. Several studies localize NADPH-diaphorase in the cat retina. Following the initial localization of NADPH-diaphorase in amacrine cells (Sandell, 1985) in the INL and in some cells in the GCL, subsequent studies provide more details on the specific cells involved. Studies of W~issle et al. (1987) establish that although some ganglion cells have weak NADPH-diaphorase, the more strongly labeled cells in the GCL are a heterogeneous population of displaced amacrine cells with dendritic arborizations in the lower IPL where the rod bipolar cells terminate. There are only one quarter as many labeled amacrine cells in the INL as in the GCL. In addition to the displaced amacrine cells, there are also both normally placed amacrine cells and interstitial amacrine cells with somas in the IPL. The interstitial amacrine cells arborize in the central IPL and there is a third band of labeled dendrites in the outer IPL. Double-labeling studies indicate that one out of six NADPH-diaphorase cells can accumulate the indoleamine 5,6-dihydroxytryptamine and are likely to be serotonergic, while later studies established that the NADPH-diaphorase amacrines contain GABA (Mtiller et al., 1988). Vaccaro et al. (1991) later reported that NADPH-diaphorase is in cone pedicles, the photoreceptor layer and vasculature, and confirmed that 75% of the total of 40,000 amacrine cells are located in the GCL. Monkey. The presence of NADPH-diaphorase in amacrine cells and in some cells in the GCL is first described in owl monkey, squirrel monkey and rhesus macaque by Sandell (1985), and later NOS immunocytochemistry (Haberecht et al., 1998) confirms these localizations in macaque. Roufail and Rees (1997) describe one class of diaphorase cells in Macaca fascicularis that is similar in appearance to ND2 cells in human (Provis and Mitrofanis, 1990), with somas that are predominantly in the INL although some are in the GCL. In addition, both NADPH-diaphorase (Roufail and Rees, 1997; Chen et al., 1998) and NOS-LI are localized in cone and rod inner segments (Haberecht et al., 1998). Human. Following the initial general description of NADPH-diaphorase-positive amacrine cells (Sandell, 1985), subsequent studies (Provis and Mitrofanis, 1990; Roufail et al., 1995) clearly characterize three distinct amacrine cell types in humans. The ND1 have large oval/triangular somas that are located in the INL or GCL, with strongly labeled dendrites in the middle or outer IPL. The ND2 amacrine cells have smaller, more rounded somas that are also located in the INL and give rise to stout primary dendrites in the middle of the IPL and more delicate beaded processes in the middle or outer IPL or GCL. The ND3 amacrine cells are not always seen, but they have small round somas that are predominantly in the GCL and sometimes in the INL, with poorly labeled dendrites. In a given human retina, there can be a total of 118,000 NADPH-diaphorase-positive cells, 90% of which are located in the 116

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INL (Provis and Mitrofanis, 1990). Subsequent NOS immunocytochemistry finds NOS-LI in all of these three amacrine cell types (Roufail et al., 1995). The human ND2 cells resemble the rabbit ND2 amacrine cells in morphology and density, the human ND1 cells resemble the ND1 cells in rabbit and the NDa cells in guinea pig, while the human ND3 cells appear to be a distinct new class. In addition to these amacrine cells, NADPH-diaphorase is also localized in the outer (Provis and Mitrofanis, 1990) or inner (Chen et al., 1998) segments of photoreceptors and in the retinal vasculature (Roufail et al., 1995). The NO released from these amacrine cells is postulated to cause pericytes to change the vascular diameter and modulate blood flow (Roufail et al., 1995), because the retinal arterioles and capillaries do not receive autonomic innervation (Ye et al., 1990). Hamster. Lau et al. (1994) did the only study localizing NADPH-diaphorase in the hamster retina. They find 8033 ND(g) cells in the GCL with 2-4 sparse dendrites up to 700 ~m long and 5051 ND(i) cells in the INL with weakly labeled dendrites. The arborizations of these cells are in S 1, $3 and $5 of the IPL. Following optic nerve transection there is no change in any cell numbers, and there is also no diaphorase in any retrogradely labeled ganglion cells, both of which indicate that all of the NADPH-diaphorase-positive neurons are amacrine cells. The ND(g) cells have large somas and sparse dendrites like the ND1 cells in rabbit, while the ND(i) cells resemble the ND2 cells in rabbit. Guinea pig. In the guinea pig there are two NADPH-diaphorase-positive cell types. Cobcroft et al. (1989) find 3450 NDa cells with darkly labeled somas, primarily in the INL (and a few in the GCL), that give rise to dendrites which spread several hundred microns in the middle of the IPL. They also find 4389 NDb cells with much more lightly labeled somas in the GCL that give rise to poorly defined dendrites. These investigators conclude that both cell types are amacrine cells because there is no diaphorase in ganglion cell axons, and that the ND1 cells in rabbit are like the NDa cells, but that the ND2 cells in rabbit are different than the NDb cells. Tree shrew. The results of the study by Petry and Murphy (1995) of tree shrews resemble those in other mammals in that they find NADPH-diaphorase in some amacrine cells, displaced amacrine cells, Mtiller cells and in the ellipsoids of all cone photoreceptors. However, they also make the unique finding that NADPH-diaphorase is selectively present in the myoids in the inner segments, and surrounding the nucleus in short-wavelength-sensitive blue cones and all rods. This suggests that the short-wavelength-sensitive cones have different biochemical machinery than other cones, and may more closely resemble rods. Another correlation between short-wavelength light responses and NOS is present in fish retina (Furukawa et al., 1997), and is described later in this review. 2.3.2. Lower vertebrates Birds. In the pigeon, Sato (1990), localizes NADPH-diaphorase activity in three amacrine cell types that can be distinguished on the basis of their soma size and staining intensity. The type I cells have large intensely stained somas in the inner INL that give rise to processes that ramify for up to 150 Ilm in the IPL near the IPL and INL border, or in S1 of the IPL. Type II and type III cells also have somas in the INL, but their processes are only weakly labeled and difficult to follow, although some of the processes of type III cells run toward $3 of the IPL. In addition to amacrine cells, there is NADPH-diaphorase in faintly labeled somata in the GCL and strong labeling in the inner segments of photoreceptors. Morgan et al. (1994) use NADPH-diaphorase histochemistry and find that diaphorase activity is present in the horizontal cell layer, in the OPL and in the outer segments of the

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photoreceptors in the chicken retina, but they provide little detail on the specifics of the cells involved. Although subsequent studies also localize NADPH-diaphorase or NOS-LI in the bird retina, the focus of these studies is primarily developmental or biochemical, so little additional detailed information is added about the specific cell types involved. However, these later studies do localize NADPH-diaphorase in subsets of amacrine and ganglion cells (De Carvalho et al., 1996), in the OPL, IPL and GCL, and in cell processes (Goureau et al., 1997). Goureau et al. (1997) do immunocytochemistry that localizes nNOS-LI in the GCL and in the outer limiting membrane near the tips of the Mtiller cells, and they localize eNOS-LI in the OPL, the inner INL, and especially in the GCL and ganglion cell axons. Turtles. Several studies localize nNOS in the turtle retina using a combination of NOS immunocytochemistry and NADPH-diaphorase histochemistry. Blute et al. (1997) find both NADPH-diaphorase and NOS-LI in the ellipsoids and inner segments of photoreceptors, efferents from the brain to the retina, at least three amacrine cell types and in many processes in the IPL. In optimized double-labeled preparations, all cells with NADPH-diaphorase activity also have nNOS-LI, although some somas in the ganglion cell layer only have nNOS-LI. The ND 1 amacrine cells are relatively common, and in retinal cross-sections they have large vertically oriented darkly staining pyriform-shaped somas with relatively symmetric dendritic arborizations about 300 ~tm in diameter. In small numbers of ND1 cells, in addition to these relatively conventional dendritic fields, some cells also have at least one dendrite that leaves the primary dendritic field and runs for at least 600 ~tm through the retina. A distinguishing characteristic of ND 1 amacrine cells is that the primary dendrites run for some distance in the outer IPL, before branching and arborizing deeper in the middle of the IPL and more diffusely in the inner IPL. The dendrites in the deeper portions of the IPL are studded with boutons. The ND2 amacrine cells are the most common of the three NADPH-diaphorase amacrine cell types in the turtle retina. These cells have smallish, moderately stained, rounded somas that give rise to several delicate processes, which run a short distance in the outer IPL, before branching extensively in the central IPL and giving rise to terminal processes deep in the IPL or among the somas in the GCL. The ND3 amacrine cells have large, faintly stained, flattened, oval somas that often give rise to two thick, faintly labeled primary processes that arborize in the outer IPL. The ND1 and ND2 amacrine cell types in turtle are very similar to the ND1 and ND2 (Vaney and Young, 1988) or type 1 and type 2 (Sagar, 1990) NADPH-diaphorase-positive amacrine cell types in the rabbit, and to the NDa and NDb NADPH-diaphorase-positive amacrine cell types in the guinea pig (Cobcroft et al., 1989). In addition to amacrine cells, both NADPH-diaphorase and NOS-LI are present in 7-10 well-labeled efferent fibers that exit the optic nerve head into the retina and give rise to several branches before reversing direction and then running back toward the optic nerve head. Although the efferent fibers initially run in the IPL near the GCL, the more terminal portions run in the outer third of the IPL. More detail is provided about these efferents in a subsequent section. In addition to these light microscopic studies, NOS-LI has been ultrastructurally localized in the OPL in turtle (Haverkamp and Eldred, 1998b). These workers find nNOS-LI at both the central and lateral positions at photoreceptor ribbon synapses which indicates that nNOS-LI is in both invaginating bipolar and horizontal cell processes at photoreceptor terminals. In addition, Haverkamp and Eldred (1998b) also report processes from presumptive OFF-bipolar cells with nNOS-LI that make basal synaptic contacts with photoreceptors. Thus, NO is produced by all of the major nerve cell types in the outer retina at specific synaptic contacts 118

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(Blute et al., 2000). The selective subcellular localization of strong nNOS-LI at synaptic contacts, made by cells with low or undetectable levels of nNOS-LI at the light microscopic level, suggests that NO may be utilized at many more synaptic contacts than previously realized. A similar selective localization of NOS is also seen in the IPL of the turtle retina, where some amacrine cells have NOS-LI only at presynaptic specializations and not in their general cytoplasm, while other amacrine cells have NOS-LI throughout their cytoplasm (W.D. Eldred, pers. commun., 2000). Fish. The localization of both NADPH-diaphorase and NOS-LI is examined in several species of fish. The first study of NADPH-diaphorase in the carp retina (Weiler and Kewitz, 1993) finds diaphorase in putative amacrine and ganglion cells, in one type of ON-bipolar cell and in H 1 horizontal cells. Although there is NADPH-diaphorase in cone outer segments, there is none in rod outer segments, in H2, H3 and H4 horizontal cells or in horizontal cell axon terminals. The H1 horizontal cells with NADPH-diaphorase also contain GABA. A subsequent study of carp retina (Djamgoz et al., 1996), confirms the staining seen in amacrine, ganglion and bipolar cells, and in addition finds NADPH-diaphorase in the ellipsoids of rod and cone photoreceptors, Mtiller cells and more than one type of horizontal cell. However, no staining is seen in photoreceptor outer segments. Two studies make comparative localizations of both NADPH-diaphorase and NOS-LI in the goldfish retina (Liepe et al., 1994; Villani and Guarnieri, 1996). In both studies, NOS-LI is in amacrine cells and in somas in the GCL. However, Liepe et al. (1994) also find NOS-LI in Mtiller cells, the outer limiting membrane and terminals of ON-bipolar cells, while Villani and Guarnieri (1996) find strong NOS-LI in the OPL. In both studies, the localization of NADPH-diaphorase is more extensive than NOS-LI. In addition to the localizations above, NADPH-diaphorase is also in photoreceptor ellipsoids or outer segments, H1, H2 and H3 horizontal cells and in two bands in the IPL. Liepe et al. (1994) also find both NADPH-diaphorase and NOS-LI in amacrine cells, Mtiller cells and photoreceptor ellipsoids in catfish retina as well. Amphibians. Three studies do detailed localizations of both NADPH-diaphorase and NOS-LI in the salamander retina and find similar results. The initial study by Liepe et al. (1994) finds NOS-LI around photoreceptor nuclei and in rod and cone ellipsoids, in occasional amacrine cells, somas in the GCL, and in the IPL and OPL. They also find NADPH-diaphorase in amacrine cells, in photoreceptor ellipsoids and around photoreceptor somas, in horizontal and Miiller cells, and near the inner and outer limiting membranes. Subsequent studies by Kurenni et al. (1994, 1995) confirm and extend these initial results. They find intense NADPH-diaphorase in the rod and cone photoreceptor ellipsoids, and fainter labeling in the myoid regions of photoreceptors, but no significant labeling in the outer segments, synaptic terminals or perinuclear cytoplasm of the photoreceptors. Some bipolar cell somas in the INL and their Landolt clubs are labeled, and there is variable Mtiller cell staining that is strong in distal and proximal Miiller cell processes and is weaker, fiber-like staining in the Mtiller cell somas and trunks. Finally, there is NADPH-diaphorase in dark puncta in the OPL and IPL and some labeling in ganglion cells and in the optic nerve. The corresponding NOS immunocytochemistry by Kurenni et al. (1994, 1995) confirmed the localization of nNOS in the outer segments, ellipsoids and myoids of photoreceptors, in MUller cells, and in select amacrine, horizontal and bipolar cells. Unlike many previous studies, Straznicky and Gabriel (1991) find that they could only get NADPH-diaphorase staining in the toad (Bufo) retina by using unfixed tissue. They find labeling in the inner segments of photoreceptors, some somas in the GCL, and in many bipolar cells with intensely stained Landolt clubs. They also find three types of labeled amacrine cells 119

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that, like the bipolar cells, primarily arborize in the outer portion of the IPL. One labeled amacrine cell is a narrow-field amacrine cell, the second has an oval-shaped soma with two main dendrites, while the last amacrine cell type is characterized by having a grossly eccentric dendritic tree. In the frog retina, Sato (1990) describes two populations of amacrine cells with NADPHdiaphorase activity. As is the case for other species, one population of cells is darkly stained with clearly labeled processes, while the second population is weakly labeled with poorly stained processes. The processes of the darkly labeled amacrine cells often run to $5 of the IPL or to the GCL. In addition to amacrine cells, NADPH-diaphorase is also seen in the inner segments of photoreceptors in frog. In summary, the comparative localization of nNOS in different species can indicate what is universal and what is species-specific. Although there are some differences between species, in many cases these differences are not as great as the variation between different studies of the same species. For this reason, it is likely that there are similar localizations of nNOS in all species, and that many of the reported differences are methodological. Therefore, in considering these results as a whole, there is evidence for the presence of nNOS in all cell types in the retina, but not in every cell of a given type. For example, nNOS does seem to be restricted to certain cells of a given type, i.e. some amacrine cells. Photoreceptors in particular, have potentially a very complex localization of nNOS in their outer segments, inner segments and synaptic terminals. The localization of NOS in all retinal cell types is strongly supported by the physiological and biochemical studies described later. One nearly universal observation is that, in most species, there are some amacrine cells with darkly stained somas and processes, while often larger numbers of amacrine cells have more weakly stained somas and poorly labeled processes. It is tempting to speculate that these weakly labeled amacrine cells give rise to the many synapses with NOS-LI that are found in the IPL in processes with little cytoplasmic NOS-LI. In several species, the cells with strong cytoplasmic NOS-LI or NADPH-diaphorase have large dendritic arborizations, while the more lightly labeled cells have smaller arborizations. This raises the possibility that the cells with large dendritic arbors and strong cytoplasmic NOS may release NO more diffusely, to influence large numbers of neurons, as might take place during light and dark adaptation. In contrast, the other lightly labeled cells may employ NOS only at synapses involved in specific synaptic circuits. 2.4. REGIONAL DISTRIBUTION OF NOS The precise retinal distribution of cells with NOS or NADPH-diaphorase has only been examined in a limited number of species. In the rat retina (Huxlin, 1995), there are four times more NADPH-diaphorase-positive cells in the GCL than in the INL. The ganglion cells and displaced amacrine cells in the GCL (Huxlin and Bennett, 1995) form a heterogeneous population that are relatively evenly and randomly distributed. In contrast, the amacrine cells containing NADPH-diaphorase in the INL are more regularly distributed and the somas in the GCL physically overlap with somas in the INL. The distribution of cells with NADPH-diaphorase in the rabbit retina shows a strong central to periphera] gradient, with the highest density of cells in the visual streak (Vaney and Young, 1988). There are also significant regional distributions in the specific amacrine cell types with NADPH-diaphorase. In the visual streak there are six times more ND2 cells than ND1 cells, and in the periphery there are nine times more ND2 cells than ND1 cells (Vaney and Young, 1988). Interestingly, the dendritic arborizations of most ND1 amacrine cells in the visual 120

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streak are oriented parallel to the visual streak, while those amacrine cells in the inferior periphery of the retina have dendritic arborizations oriented orthogonally to the visual streak (Mitrofanis et al., 1992). In humans (Provis and Mitrofanis, 1990), amacrine cell types with NADPH-diaphorase have relatively uniform topographical distributions with no real peak at the fovea, although the cells have a minimal density near the retinal edges and there are none in the foveola. The outer segments of cone and rod photoreceptors also show a weaker NADPH-diaphorase staining near the foveola. Lau et al. (1994) show that the cells with NADPH-diaphorase in hamster have a different distribution in the INL and GCL. The ND(g) cells in GCL are not evenly distributed, in that there are more cells in the inferior than in the superior retina, and they have a peak density 1 mm below the optic disk. In contrast, the ND(i) cells in the INL are evenly distributed with no obvious density gradient, although there is a somewhat higher density of labeled cells 1 mm superior to the optic disk. In guinea pig, there are equal numbers of two NADPH-diaphorase-positive cells types (Cobcroft et al., 1989), each with a unique retinal distribution. The somas of the NDa cells are primarily located in the INL, although a few are in the GCL. However, in both cases these NDa cells are relatively uniformly distributed in both the superior and inferior retina, although there is a decrease in density toward the retinal edge. The NDb cells are much more lightly labeled cells and have their somas in the GCL. In contrast, the NDb cells are only above the optic disk and not below, with a higher concentration of cells in an elongated area just above the optic disk near the visual streak. The distribution of NADPH-diaphorase-positive cells in the cat retina is first analyzed by W~issle et al. (1987). These workers analyze the density of labeled displaced amacrine cells using a vertical intersect through the central retina. They find an even distribution of cells with no indication of a peak in the central retina, although the lower retina has cell densities twice those in the upper retina. The distribution of NADPH-diaphorase-positive cells is later examined in more detail by mapping the distribution of labeled cells throughout the retina (Vaccaro et al., 1991). These studies largely confirmed the results of W~issle et al. (1987), in that there is a relatively uniform distribution of cells in most of the retina. However, they did find an area with the highest cell density located slightly inferior to the area centralis. In lower vertebrates, Straznicky and Gabriel (1991) find in the toad (Bufo) that the distribution of bipolar cells follows the overall cell density, with a peak in density in the retinal center and decreased density in the dorsal and ventral regions. In contrast, the amacrine cells in Bufo have the lowest density in the central retina, and the highest density in the periphery near the ciliary margin where more cells are generated throughout life. In birds, Sato (1990) analyzes the distribution of NADPH-diaphorase-positive cells in pigeon retina. He reports that there can be 5180 labeled cells located throughout the retina, and that there is only a very shallow density gradient from the central to the peripheral area of the retina. The regional distributions of NOS-containing cells have not been examined in detail in other lower vertebrates, although there are no obvious regional specializations in the distribution of any NOS-containing cells in turtle retina (Blute et al., 1997). In summary, there is considerable variation in the distribution of retinal cells with NADPH-diaphorase in various species. The ND(i) cells in the INL of hamsters and the NDa cells in the INL of guinea pigs, and most cells in rats, humans and turtles, are relatively evenly distributed throughout the majority of the retina. In contrast, there are more cells and a slight peak in labeled cell density in the inferior retina for all cells in cats, and for the ND(g) cells located in the GCL in hamster retinas. Other variations are the peaks in cell density of 121

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NADPH-diaphorase-positive amacrine cells in the visual streak in rabbits, the slight peak in amacrine cell density toward the central retina of pigeon, and the peak in labeled bipolar cells seen in the central retina in Bufo. Finally, still other variations are seen in the distributions of amacrine cells, in that the NDb cells in the GCL in guinea pigs are only above the optic disk, and that the amacrine cells with NADPH-diaphorase in Bufo reach their peak density toward the ciliary margin. Given the diversity of these distributions in different species, it is difficult to correlate these distributions with potential functions. Some of these regional differences may represent differences in the enzymes which give rise to the NADPH-diaphorase activity in certain species. The development of more selective probes for specific NOS isoforms may clarify some of these regional differences. 2.5. EFFERENTS Two studies indicate that NOS is in efferents that project from the brain to the retina. In birds, the efferents to the retina originate from the isthmo-optic nucleus and Morgan et al. (1994) report that these projections are NADPH-diaphorase-positive. They confirm that these are efferents by doing lesions of the isthmo-optic nucleus in the brain. In the bird retina, the isthmo-optic terminals are of the restricted type and they are in 2-4 cell layers in the INL, primarily in the inferior retina close to the optic nerve head and pectin. The existence of NOS-LI and NADPH-diaphorase in retinal efferents is also demonstrated in turtle (Blute et al., 1997; Haverkamp and Eldred, 1998a). Blute et al. (1997) use a combination of NOS immunocytochemistry and NADPH-diaphorase histochemistry to label 7-10 efferent fibers that exit the optic nerve head and travel in the ganglion cell axon layer before entering the IPL. Although the efferent fibers initially run in the IPL near the GCL, the more terminal portions also run in the outer third of the IPL. In tangential sections, these darkly stained processes have numerous, large boutons and in several cases these efferents run for some distance in the retina, and then arch back toward the optic nerve head. Haverkamp and Eldred (1998a) use cholera toxin B as a retrograde tracer to locate the somas in the turtle brain that give rise to these efferents. Colocalization studies of this retrograde tracer with NADPH-diaphorase histochemistry indicate that 30% of the cells with cholera toxin B-LI are also positive for NADPH-diaphorase. The location of these double-labeled cells around the locus coeruleus corresponds to the NADPH-diaphorase-positive efferent cells in the avian isthmo-optic field. In summary, the localization of NADPH-diaphorase in these efferents in birds and turtles indicates that retinal efferents may use NO to modulate the retinal function and that these efferents arise from similar regions of the brain.

3. DEVELOPMENT OF NOS IN THE RETINA AND CENTRAL VISUAL TARGETS The development of NOS has been examined using both anatomical and biochemical techniques in the retina and in several central visual targets. In all cases, there is a close correlation between NOS and synaptic development. The development of both NADPH-diaphorase and NOS-LI has been examined in the rat retina. Mitrofanis (1989) first detected NADPHdiaphorase by postnatal day 3 (P3). By P5, NADPH-diaphorase is present in the INL, and by P11 there are labeled cells with extensive processes in the IPL. This time corresponds to the time when there is the first appearance of amacrine and bipolar synapses and the beginning of electrical activity. At P25, there is a reduction in labeling of processes in the IPL, and a decrease in the diameter of labeled somas, similar to that seen in adults. Mitrofanis 122

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(1989) speculates that the staining seen at P3-P7 in the OPL may relate to the formation of horizontal cell synapses in the OPL. He further speculates that the relatively uniform distribution of NADPH-diaphorase-positive cells in the adult rat retina, is the result of a higher initial concentration of cells in the periphery, coupled with a greater subsequent expansion in the periphery. These results are confirmed and extended by Patel et al. (1997) who use a combination of nNOS and iNOS immunocytochemistry in rat retina. They find that weak nNOS-LI is present in cells in the INL at P10, that by P14 nNOS-LI is strong in the IPL and in cells in the INL, and that by P21 and P30 the number and intensity of cells with nNOS-LI is decreased, iNOS has a similar developmental time course, in that it appears by P10, rises to a maximum near P14 and declines by P16-P30. Patel et al. (1997) conclude that because NOS is detected after the peak of retinal ganglion cell death during the first postnatal week, and because axotomy did not increase either iNOS or nNOS, NOS does not play a role in cell death or degeneration. However, they do conclude that NOS may play a role in development, because it appears during the second postnatal week, which coincides with synaptogenesis in the inner retina and maturation of the vasculature. A similar developmental situation also occurs in rabbit and cat retina. In the rabbit (Mitrofanis et al., 1992), NADPH-diaphorase cells are first seen at post-conception day 28 (28PCD) in the central part of the visual streak. Only during 28-42PCD is NADPH-diaphorase present in the OPL, and by 51PCD NADPH-diaphorase is gone from the OPL and is only present in the IPL. Again, the presence of NADPH-diaphorase corresponds with the timing of the main amacrine cell synaptogenesis between 40PCD and 51PCD. Mitrofanis et al. (1992) speculate that these cells may play a developmental role, because they appear before there are light responses, and that the NADPH-diaphorase-positive cells appear so early that they must be preprogrammed and are not environmentally determined. In the cat retina, there are two times as many cells with NADPH-diaphorase at birth as in the adult (Vaccaro et al., 1991). By P10, there are many NADPH-diaphorase-positive cells with growth cones and spines and an intense continuous band of labeling in the OPL, but there is a strong reduction in NADPH-diaphorase by P25. Vaccaro et al. (1991) conclude that the most intense NADPH-diaphorase occurs during the appearance of bipolar cell synapses from late gestation to early postnatal periods, and that there is a reduction following the IPL synaptogenesis that occurs from P11 to P 15. In birds, NADPH-diaphorase is in subsets of amacrine and ganglion cells. De Carvalho et al. (1996) detect NADPH-diaphorase beginning in the 7-day-old embryo (E7), and by El2 it is present in photoreceptors, and in somas in the INL and GCL. After hatching, the NADPH-diaphorase is in photoreceptor inner segments and in somas in the INL and GCL, but not in photoreceptor outer segments. Goureau et al. (1997) expand on these earlier studies to include both eNOS and nNOS immunocytochemistry and Western blots, in addition to NADPH-diaphorase staining. When examined using both immunocytochemistry and Western blots, both nNOS and eNOS appear by E8 and by E 12-E14 both are in photoreceptors, the ONL, INL and GCL. By El8, eNOS levels peak in photoreceptors, the OPL, IPL and GCL, and levels of nNOS peak in the IPL and amacrine cells. Levels of both eNOS and nNOS decrease by 8 days after hatching, but some remains, primarily in ganglion cells. The timing of higher levels of both nNOS and eNOS closely correlates with the time of much of the active synaptic development in chick retina (Hering and Kr6ger, 1996). The role of NOS has also been examined in a number of central targets. Williams et al. (1994) are interested in the activity-dependent presynaptic mechanisms and NMDA receptor-mediated postsynaptic mechanisms that determine the connections of the retina 123

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to central visual targets like the optic tectum in chick. They find NADPH-diaphorase NOS-positive cells in the chick optic tectum at E5-E7, and that the peak staining occurs in radially oriented bipolar and stellate cells at EI2-E15. However, in eyeless animals the number and intensity of labeled bipolar cells are reduced and no stellate cells are labeled in the tectum. They also find that the expression of NADPH-diaphorase activity in the tectum coincides with the time of innervation by ganglion cell axons and that the NADPH-diaphorase activity peaks at the time of refinement of the initial pattern of connections. Williams et al. (1994) conclude that NOS expression is dependent on the presence of the ganglion cell retinal axons, and that it is likely that the developing ganglion cell axons are exposed to NO. The NO may then provide a retrograde communication to the ganglion cell axons to facilitate the development and refinement of their tectal connections. This idea is supported by the fact that NOS expression begins to decline at the end of this period of refinement. A similar increase and decline in NADPH-diaphorase are also seen in cells in the developing rat superior colliculus. Tenrrio et al. (1996) find that NADPH-diaphorase activity starts in five cell types by the end of the first postnatal week and reaches a maximum by postnatal day 15. Following this time, the NADPH-diaphorase activity decreases in three of these cell types. These authors conclude that these NADPH-diaphorase-positive neurons may play a role in the refinement of the retino-collicular or cortico-collicular projections. This is because the developmental timing of the NADPH-diaphorase activity correlates with the functional development of the retino-collicular pathway that begins at the end of the first postnatal week and occurs primarily during the second week after birth. A role for NOS in retinogeniculate development is provided by the studies of Cramer et al. (1995) which indicate that there are high levels of NADPH-diaphorase in the lateral geniculate nucleus during the period of the ON/OFF sublamination phase of retinogeniculate development. A subsequent study (Cramer et al., 1996) finds that treatment with a NOS inhibitor during the third and fourth postnatal weeks reduces the formation of these ON/OFF sublaminae in the geniculate. They conclude that NOS acts together with postsynaptic NMDA receptors in the activity-dependent refinement of the size, position and synaptic connectivity of the presynaptic ganglion cell arbors. They suggest that NO may work through guanylate cyclase and cGMP-gated channels to increase intracellular calcium and axonal structure, and/or through adenosine diphosphate ribosyl transferase to act on growth-associated protein 43 (GAP-43) in remodeling these synaptic connections. They speculate that NO may inhibit growth cone elongation or that it may regulate local blood flow and growth factors. The idea that NO can influence growth cone elongation during development receives strong support from studies by Renterfa and Constantine-Paton (1996). These workers find that NO donors induce the collapse of ganglion cell axonal growth cones that are extending from explants of tadpole retina in culture. They conclude that this effect is not mediated by cGMP and that it is not due to the production of NO within the growth cones, because neither cGMP or NOS inhibitors induce collapse. They suggest that perhaps NO may cause ADP-ribosylation of G-actin or that there may be NO-dependent modifications of growth cone proteins such as GAP-43 or synaptic associated protein 25 (SNAP-25). Finally, they suggest that the NO produced by the activated target dendrites could alter the growth patterns of axons by causing a local collapse of growth cones. This collapse may then increase arborizations within a target of glutamate-sensitive dendrites to help establish new synapses on the depolarized dendrite targets. In summary, in various species, NO seems to play several roles in synaptic development of the visual system. In the case of the retina, optic tectum, superior colliculus and lateral geniculate there is a close temporal correlation in the developmental appearance of NOS, and 124

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the development and refinement of synaptic connections. Furthermore, at least in the case of the optic tectum and lateral geniculate, NO is required for the correct establishment of these connections. It is particularly intriguing that NO may be involved in the maturation of synapses at all levels of the visual system from within the retina itself, to its central projections.

4. BIOCHEMISTRY AND MOLECULAR BIOLOGY OF NOS IN THE RETINA 4.1. NOS A number of detailed studies have been made on NOS in the retina, many of which extend the knowledge obtained in the brain, and indicate that the roles of different NOS isotbrms may be much more complicated than previously realized. It is generally accepted that both nNOS and eNOS are calcium-dependent while iNOS is calcium-independent, and that more than 85% of the NOS in brain and macrophages is soluble, while 95% of eNOS is membrane-bound (Schmidt et al., 1993). However, several studies indicate that subcellular localization and calcium dependence of NOS in the retina may be more complicated. At least two studies use molecular methods to analyze NOS in the retina. As described above, Park et al. (1994) use primers of a conserved cofactor region of nNOS and iNOS for RT-PCR and find that both retinal nNOS and retinal iNOS have very close sequence homologies to brain NOS and human chondrocyte iNOS, respectively. The cDNA was subsequently cloned using RT-PCR and human retinal mRNA (Park et al., 1996), and the sequence shows a possible genetic polymorphism in comparison to brain NOS that is due to single-base substitutions and not due to frame shift by addition or deletions of bases. Northern blot analysis indicates that although the retinal nNOS mRNA is identical in size to brain and muscle types, there are lower levels of nNOS mRNA expression in the retina than in either human brain or skeletal muscle. These data could suggest that nNOS protein has a high level of stability in the retina. Other studies examine the regulation of NOS activity in the retina. Sheng and Ignarro (1996) measure NOS activity by citrulline formation in the rat retina and find that 70% of the activity is soluble in the supernatant, with the remaining being particulate. Inclusion of the calmodulin inhibitor, calmidazolium, reduces soluble NOS activity by only 75%, while 40% of the soluble NOS activity is retained under calcium-free conditions. This is in contrast to the total loss of activity in soluble cerebellar NOS activity under these same conditions. Although these authors find a soluble 160 kDa protein that cross-reacts to specific antibody to nNOS, they conclude that the retina contains more that one type of NOS. They speculate that some of the particulate NOS may be eNOS, but that the calcium-insensitive NOS is probably not iNOS in healthy eyes and may represent a new isoform of constitutively expressed NOS with tightly bound calcium and calmodulin. The idea that there may be a calcium-independent NOS present in retina is also supported by studies in both bird and bovine retina. De Carvalho et al. (1996) use a quantitative NADPH-diaphorase assay as a measure of NOS enzyme activity in the chick retina. They find that arginine-analogue NOS inhibitors inhibit the diaphorase activity by 65% in embryonic retinas and by 50% in post-hatch retinas, which may indicate the presence of other NADPH-dependent metabolic enzymes. Their developmental studies indicate that, during early embryonic development, NADPH-diaphorase is stimulated 50% by calcium, and that this stimulation decreases during development and is gone in post-hatched retinas. They 125

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conclude that there are two populations of NOS, a calcium-dependent diaphorase that is predominant in early embryonic development and is inhibited by low concentrations of NOS inhibitor, and a calcium-independent NOS that predominates in mature retinas and is only inhibited by high concentrations of NOS inhibitor. Although these studies also argue for the presence of calcium-independent NOS in retina, the interpretation of these results is hampered by the potential influence of other NADPH-dependent metabolic enzymes. A recent study uses the conversion of [3H]arginine to [3H]citrulline to study the regulation of retinal NOS activity by calcium (Margulis et al., 1998). They find that the total NOS activity is higher in synaptosomal fractions made from the inner retina than in synaptosomal fractions made from photoreceptors. In addition, 20-25% of the NOS activity in synaptosomal fractions is soluble and in all cases it is calcium-dependent, while 75-80% of NOS activity in synaptosomal fractions is membrane-bound and 50-60% is calcium-dependent. These investigators conclude that the calcium-independent NOS may be iNOS, which is consistent with the previous retinal localizations of iNOS by Park et al. (1994), L6pez-Costa et al. (1997) and Fujii et al. (1998). Margulis et al. (1998) calculate that synaptosomal fractions from the inner retina can produce 4 nM/s of NO under optimal conditions, which suggests that NO may reach the concentration of 250 nM NO needed for the half-maximal activation of soluble guanylate cyclase (sGC) (Stone and Marietta, 1995). Finally, Wellard et al. (1995) investigate the effects of NOS inhibitors on arginine uptake by examining the conversion of [3H]arginine to [3H]citrulline in intact chicken retinas. They find that several NOS inhibitors, like the arginine analogues L-NMMA, L-NIO, L-NA and L-NAME, all have IC50s in the micromolar range for retinal homogenates. In contrast, these same inhibitors have 20-3000 times higher ICsos in intact retina. These inhibitors also produce the following varying effects on arginine uptake in intact retina: L-NA causes a 31% reduction in arginine uptake, L-NMMA and L-NIO reduce arginine uptake by about 75%, and L-NAME has no obvious effect on arginine uptake. The authors conclude that because many NOS inhibitors may block arginine uptake, these inhibitors cannot distinguish arginine uptake from the effects of NOS inhibition, and thus citrulline production cannot be used as an accurate measure of NOS activity. Another implication of their results is that it will be very difficult to use isoform-selective NOS inhibitors to examine the function of specific NOS isoforms in intact retina. This is because most of these inhibitors show only a relative specificity for a certain isoform, and that any isoform selectivity may be obscured by the high dosages needed to inhibit NOS in intact tissue. 4.2. PHOTORECEPTORS cGMP plays a crucial role in phototransduction, and photoreceptors have many biochemical pathways to regulate cGMP levels, some of which involve NO. As discussed in one of the following sections, NO also plays distinct roles in modulating synaptic release from the terminals of photoreceptors. The majority of studies of the particulate guanylate cyclases (pGCs) in photoreceptor outer segments have concentrated on the two pGCs, retGC (Dizhoor et al., 1994) and retGC2 (Lowe et al., 1995), which are modulated by intracellular calcium and a soluble factor (Koch, 1992). However, one early study also indicates a potential role for NO in modulating cGMP in bovine photoreceptor outer segments. Horio and Murad (1991) examine a pGC that is solubilized from bovine outer segment membranes and find that it is activated by low calcium and inhibited by higher calcium. They state that they carefully washed their samples to eliminate potential sGC contamination and that antibodies against sGC did not recognize this pGC. Interestingly, they find that the NO donor sodium nitroprusside (SNP), 126

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and NO gas also activate this pGC up to 20 fold, although higher concentrations of NO inhibit this pGC. The potential source of NO in activating this outer segment GC is examined by Venturini et al. (1991). These workers use a crude preparation of rod and cone outer segments and find NOS activity that is similar to nNOS, in that it is NADPH- and calcium-dependent and that it is blocked by NOS inhibitor. They suggest that this NOS may provide the NO to activate the NO-sensitive pGC described by Horio and Murad ( 1991 ). The activation of guanylate cyclase (GC) in photoreceptor outer segments by NO has also been investigated by Margulis et al. (1992). In contrast to the results of Horio and Murad (1991), Margulis et al. (1992) find that although NO can produce a 1300-1400% activation of the sGC in photoreceptor outer segments, they find no NO activation of the pGC that they isolated from outer segments. They could not explain the difference between their results and those of Horio and Murad (1991). Margulis et al. (1992) conclude that the high levels of free Ca 2+ in dark-adapted outer segments can activate NOS to make NO and subsequently activate sGC. The characterization of NOS in photoreceptors and its interactions with GC are also examined by Koch et al. (1994). They report the existence of a calcium/calmodulin-regulated NOS in biochemical fractions made from outer segments and the adjacent portions of the photoreceptor inner segments from bovine retina. The biochemical properties of photoreceptor NOS resemble that of mammalian brain nNOS in terms of its cofactor requirements and its 160 kDa molecular weight on Western blots using antibodies against nNOS. They conclude that photoreceptor inner segments, but not the outer segments, contain a NOS that is functionally coupled to sGC, and that the pGC in the outer segments is not sensitive to NO. They suggest that the NOS in the outer segments is involved in the ADP-ribosylation of outer segment proteins to modulate phototransduction, as described below (Pozdnyakov et al., 1993). In contrast, the recent studies of Yoshida et al. (1995) report the existence of NOS in bovine rod outer segments. They find that both cytosolic and membrane fractions of rod outer segments have NOS activity. Adding calcium increases the activity of both the membrane and cytosolic NOS. However, in the absence of calcium and calmodulin, the cytosolic NOS is inactive, while the membrane NOS still retains basal activity. They speculate that the calcium-free basal activity of rod outer segment membrane NOS may be due to a tightly bound calmodulin like that found in iNOS, and that the non-calcium-dependent NOS could influence the rate of recovery of the light response when calcium is low. Again, they speculate that the NO may relate to ADP-ribosylation of G proteins in rod outer segments (Pozdnyakov et al., 1993). One mechanism of action of NO is the ADP-ribosylation of proteins (Bredt and Snyder, 1994), and the studies of Pozdnyakov et al. (1993) indicate that this occurs in retina as well. Pozdnyakov et al. (1993) find that both the membrane and cytosolic fraction of rod outer segments contain ADP-ribosyl transferases that can be regulated by NO. A NO donor or endogenously produced NO inhibits ribosylation of 116 kDa, 66 kDa and 46 kDa membrane proteins, while NO increases the ribosylation of 38 kDa cytosolic and 39 kDa membrane proteins. The 39 kDa protein is the ~ subunit of the G protein transducin and thus NO may modulate visual transduction through ADP-ribosylation. They conclude that rod outer segments may have at least two membrane ribosyl transferases, one that is inhibited by NO while the other is stimulated, and that both membrane and cytosolic NOS each have their own associated ADP-ribosyl transferases. In summary, the biochemical results indicate the presence of several distinct isoforms of NOS. There is good evidence for both soluble and particulate NOS in the retina, with more 127

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NOS activity being concentrated in the inner retina than in the outer retina. The most recent studies indicate that, in the retina as a whole, all of the soluble NOS is calcium-dependent, while only ~60% of the particulate NOS is calcium-dependent. In light of both biochemical and immunocytochemical studies, it is possible that at least some of the calcium-independent NOS is iNOS, although the possibility of other NOS isoforms cannot be eliminated. A similar situation occurs in photoreceptors where all of the soluble NOS is calcium-dependent, while the particulate NOS retains some basal activity in calcium-free solutions. Finally, there is good evidence that although NO can activate sGC or ADP-ribosyl transferases in photoreceptor outer segments, it probably has no effect on the pGCs in outer segments.

5. FUNCTION OF NO IN SPECIFIC RETINAL CELL TYPES 5.1. PHOTORECEPTORS In comparison to other retinal cell types, the role of NO in photoreceptors has been the most thoroughly investigated and has proven to be complex. In their investigation of tiger salamander rods, Kurenni et al. (1994) find that NO increases the Ica current in barium-conducting calcium channels by 183%. They postulate that an activation of ADP-ribosyl transferase or direct nitrosylation of cellular protein thiols is responsible for the shift in the activation curve of these channels, because cGMP analogs have no effect on this current. The effects of NO are specific for these channels, in that the nonselective cation current Ih and the non-inactivating IK~ currents remain unaffected. The activation of these calcium channels by NO could modulate the photoreceptor output, because the postsynaptic response in the horizontal cells is a third-power relation of the calcium influx at the photoreceptor synapse (Attwell et al., 1987). Kurenni et al. (1994) postulate that calcium influx through cGMP-gated channels in the outer segment, or through voltage-gated calcium channels in the inner segment, activates the NOS in the ellipsoids to produce NO that depolarizes the rod. This NO may then diffuse to neighboring photoreceptors to make a smooth gradient of channel modulation that spreads light adaptation through the photoreceptor mosaic. Rieke and Schwartz (1994) also examine effects of NO donors on synaptic exocytosis in salamander cones. They conclude that the cone soma and synaptic processes contain a sGC that can modulate the cGMP-gated current, Ca 2+ entry and thus, exocytosis. Interestingly, they find that this mechanism continues when the cones are hyperpolarized to more than - 4 0 mV, and thus serves to extend the voltage range of exocytosis in the cones. In contrast, the rods are driven primarily by voltage-gated Ca 2+ channels and their exocytosis is terminated by such hyperpolarizations. These results indicate that NO can serve to extend the range of the light response in cone photoreceptors. N611 et al. (1994) also speculate that the same mechanisms involved in the recovery of photoreceptor responses are involved in light adaptation and that NO is implicated in these processes. They find that, while recording from photoreceptors using whole-cell patch clamp electrodes, there is a reduction in the flash response due to a spontaneous hyperpolarization caused by the loss of intracellular cGMP. They find that addition of a NO donor in the electrode prevents this hyperpolarization and accelerates the speed of flash recovery, presumably by increasing both the concentration and turnover of cGMP. Lowering of intracellular calcium with EGTA blocks the effects of the NO donor, and inhibition of NOS produces the same retardation of the recovery of the light response. They suggest that the 128

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sGC is regulated by calcium, and that calcium and NO form two different feedback loops that regulate each other to modulate the light response. In a related study, Tsuyama et al. (1993) also examine the relationship between NO, the dark voltage and the recovery of light responses in isolated frog rods. They too find that during whole-cell patch recording there appears to be a loss of components that sustain the normal dark voltage, and that the cell hyperpolarizes. This hyperpolarization is slowed by including either GTP or L-arginine in the pipette, while including NADPH in the pipette depolarizes the rod photoreceptors and accelerates the repolarization of the light responses. They came to a similar conclusion as N611 et al. (1994), in that increases in NO activate sGC to increase cGMP which could depolarize the cells and accelerate repolarization to counterbalance the increased PDE activity in the light. The results of all these studies strongly support a role for NO in light adaptation in photoreceptors. The role of NO and cGMP-gated channels in photoreceptor synaptic exocytosis is also investigated by Savchenko et al. (1997). They do whole-cell patch clamp recordings of lizard cones that lack outer segments and find that the cGMP, analog pCPT-cGMP, produces an inward current in the synaptic terminals that is distinct from the voltage-gated calcium current. They show that this current is through channels directly activated by cGMP and not through protein kinase G (PKG). Interestingly, they find that only 31% of the patches made on these terminals have channels, but when present they have a high density of 13-34 channels in comparison to outer segment patches that have only 9-19 channels. This indicates that the cGMP-gated channels are clustered on the cone terminals. These workers also find that NOS inhibitors depress the feedback inhibition onto cone terminals by the horizontal cells, and conclude that the endogenous production of NO modulates communication between cones and horizontal cells. Finally, using pairs of horizontal cells and cone terminals, they find that NO, by working through cGMP to open cGMP-gated channels, increases glutamate release onto the horizontal cells. From this they conclude that the cGMP-gated channels clustered on the photoreceptor terminals may be tonically activated to allow calcium influx at hyperpolarized potentials and thus extend the photoresponse during light stimulus. Finally, Greenstreet and Djamgoz (1994) investigate the role of NO in light adaptation and retinomotor movements in cyprinid fish retinas. They find that NO donors produce two significant aspects of light adaptation in fish. Treatment of isolated dark-adapted cyprinid retinas with NO donors causes the inner segments of cone photoreceptors to contract, and the dendrites of horizontal cells to form finger-like projections called spinules that invaginate into cone pedicles. These authors speculate that these effects could originate directly in the photoreceptors and horizontal cells, because NADPH-diaphorase staining is demonstrated in both cell types (Weiler and Kewitz, 1993). These results indicate that NO may be a signal regulating light adaptation in photoreceptors. 5.2. HORIZONTAL CELLS A number of studies have also focused on the role of NO in horizontal cells. One of the first of these studies examines the effects of NO on the gap junction coupling between horizontal cells. De Vries and Schwartz (1988) find that when cGMP is introduced directly into solitary pairs of catfish horizontal cells in culture, there is a 40% decrease in the junctional conductance between the cells. They also obtain similar uncoupling by perfusing the cultures with the NO donor SNP. They conclude that horizontal cells can be uncoupled by concentrations of cGMP which can be produced by NO activation of the endogenous GC present in the horizontal cells. 129

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Miyachi et al. (1990) confirm these earlier results by finding that injection of either cGMP or L-arginine into H1 luminosity-type horizontal cells in turtle retina reduces their light responses. They also find a dramatic increase in the cells' input resistance, and that there is a decrease in the horizontal cells' response to a surround and an increase in their response to stimulation of their receptive field center. They conclude that L-arginine is blocking the gap junctions between horizontal cells, and because a NOS inhibitor blocks the effects of L-arginine, they conclude that the arginine effect is not a direct effect and works through NOS. These physiological results support the presence of a NOS pathway in these horizontal cells. The immunocytochemical studies of Haverkamp and Eldred (1998b) confirm the existence of NOS-LI in turtle horizontal cells. Taken with the results by De Vries and Schwartz (1988), it is apparent that the entire N O S / N O / s G C / c G M P signal transduction system is present in horizontal cells. As mentioned above, Greenstreet and Djamgoz (1994) find that NO donors stimulate the formation of spinules by fish horizontal cells. A study by Pottek et al. (1997) also addresses the formation of spinules and several other effects of NO on fish horizontal cells. They find that the NO donor SNP increases the full-field light response of HI, H2 and H3 cone horizontal cell types. Stimulation with cGMP produces similar increases in H1 cell responses, which suggests that NO is working through cGMP. Moreover, because SNP decreases the 455-nm response in H3 horizontal cells, but increases its depolarization to green and red stimuli, they suggest that red and green cones are involved but not blue cones. SNP also blocks spread of Lucifer Yellow to one-half of the adjacent H1 cells, although dopamine uncouples them all. Finally, in contrast to the results of Greenstreet and Djamgoz (1994), Pottek et al. (1997) find that the NOS inhibitor, nitroarginine, has no effect in vivo on spinules. They attribute the difference in spinule formation to the loss of efferents to the retina in vitro. These results indicate that NO can influence the physiology of multiple horizontal cell types, both in terms of their chromatic responses and their network coupling. A second study of fish horizontal cells also supports a role for NO in chromatic processing. Furukawa et al. (1997) report that H1 cells in carp have different postsynaptic mechanisms in response to short- and long-wavelength stimulation, and that short-wavelength stimulation is only active during the light-adapted state. They find that the spectral balance in H1 horizontal cells is modulated by NO and that NOS inhibitor eliminates the light-adapted changes. In contrast, a NO donor produces the same effect as light adaptation. They conclude that the short-wavelength transmission to H1 horizontal cells uses an APB-sensitive glutamate receptor coupled through NO as a light-adaptive messenger, much like the photoreceptors and ON-bipolar cells. Finally, McMahon and Ponomareva (1996) investigate the modulation of glutamate receptors on fish horizontal cells by NO. Fish horizontal cells mainly have AMPA receptors and dopamine/cAMP/protein kinase A (PKA) increases the sensitivity of these AMPA receptors. These workers report that NO donors reduce the response of these glutamate receptors to kainate by -~54%. These effects of NO are reported to work through cGMP/PKG because cGMP also reduces the kainate response by 34%, and intracellular injection of the PKG inhibitor peptide, RKRARKE, also blocks the NO effects. Bath application of L-arginine also decreases the kainate responses like cGMP and NO in H1 cells, and thus it is likely that the H1 cells can serve as their own source and target of NO. These authors conclude that the glutamate receptors on horizontal cells can be positively modulated by dopamine or cAMP to increase the gain at photoreceptor-horizontal cell synapses, or they can be negatively modulated by NO to decrease this gain. Although this decrease in gain is too slow to play a role in visual processing, it may protect the horizontal cells from the high rates of release 130

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of glutamate from photoreceptors during darkness or from excitotoxicity (McMahon and Ponomareva, 1996). As shown by Djamgoz et al. ( 1996) above. there are a number of possible sources of NO in the outer retina of the fish. 5.3. BIPOLAR CELLS The light response in ON-bipolar cells is produced by the decrease in glutamate release from photoreceptors in the light. This decreased transmitter release reduces the activation of an APB-sensitive glutamate receptor on the ON-bipolar cells, which is in turn linked through a G protein to PDE. The decrease in receptor activation, decreases the activity of the G protein and the PDE, which allows an increase in cGMP levels to open cGMP-gated channels and depolarize the bipolar cell (Shiells and Falk, 1990). Shiells and Falk (1992) have investigated the effects of NO on the light responses of these ON-bipolar cells by doing whole-cell voltage clamp recordings using a pipette filled with the NO donor SNP. They find that, as the NO donor diffuses into the bipolar cell, it produces an inward current accompanied by a rise in dim and bright flash response amplitudes and an increase in membrane conductance. Inclusion of sGC inhibitors in the pipette causes outward currents, and a suppression of the light responses and membrane conductances. They conclude that there is a NO-sensitive GC in bipolar cells and that the G protein probably acts by activation of a phosphodiesterase. Interestingly, by measuring the light response in the presence of different intracellular levels of cGMP, they conclude that small cGMP levels increase the light response until saturation, but that higher levels then decrease the light response. The results of Haverkamp and Eldred (1998b) indicate that nNOS-LI is present in ON-bipolar cells and may provide a cell autonomous endogenous source of NO in ON-bipolar cells. 5.4. AMACRINE CELLS In the mammalian retina, rod signals are integrated into cone pathways through A11 amacrine cells that are in turn connected through gap junctions to other A11 amacrines and cone bipolar cells. Just as NO modulates the coupling between horizontal cells (De Vries and Schwartz, 1988; Miyachi et al., 1990). Mills and Massey (1995) show that NO donors can also selectively modulate gap junction coupling between the A11 amacrine cells and cone bipolar cells. By using specialized biotin tracers, Mills and Massey (1995) show that dopamine and CAMP decrease the gap junction coupling between adjacent A11 amacrine cells, but not between the A11 amacrine cells and the cone bipolar cells. In contrast, although NO, the PDE inhibitor Zaprinast and cGMP all decrease the coupling between the A11 amacrine cells and the cone bipolar cells, they do not decrease the gap junction coupling between adjacent A11 amacrine cells. They conclude that by working through cGMP, the NO released by light stimulation (Koistinaho et al., 1993) decreases the rod input and increases the cone input during light adaptation by uncoupling the A11 amacrine cells from the cone bipolars. As described above, NO modulates glutamate receptors on fish horizontal cells (Mc'Mahon and Ponomareva, 1996). Recently, Wexler et al. (1998) extended this result by demonstrating that NO can depress the GABAAreceptor function on arnacrine cells. They tind that both a NO donor or cGMP decrease GABA responses by 34-4 1 %. In contrast. inhibiting sGC enhances the GABA response and blocks 45% of the NO-stimulated depression of the GABA current. Inhibition of PKG prevents the depression of GABA currents in the basal condition. but not when stimulated with cGMP, which suggests that cGMP may have other targets in addition to PKG. Because activation of the cGMP-activated CAMP-specific type 11 PDE also depresses

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GABA currents, the authors believe that the amacrine cells contain a PDE that depresses GABA currents. Wexler et al. (1998) conclude that NO stimulates sGC to increase cGMP levels which then increase phosphorylation by PKG. The increased cGMP also stimulates the cGMP-activated PDE to decrease cAMP levels and PKA phosphorylation. Thus, activators of adenylate cyclase, like dopamine, enhance GABAA currents, while activators of guanylate cyclase, like NO, do the opposite, and therefore cAMP and NO/cGMP function in a push-pull mechanism in GABAergic transmission. 5.5. GANGLION CELLS One of the clearest examples of the function of cGMP in the inner retina is the modulation of cGMP-gated channels. Ahmad et al. (1994) use in situ hybridization to localize rat rod photoreceptor cGMP-gated cation channels in the inner segments of rod photoreceptors, throughout the INL and in a subpopulation of cells in the GCL. They find that patch electrodes filled with cGMP are able to activate these nonspecific cation channels on ganglion cells and that stimulation with the NO donor SNP also activates these channels after a short delay. Ahmad et al. (1994) conclude that the cGMP-gated channels are regulated by a NO-sensitive sGC in these ganglion cells. In summary, NO plays a role in the physiology of every nerve cell type in the retina. In photoreceptors, NO can modulate transmitter release and may serve to spread this modulation to adjacent photoreceptors. NO also signals light adaptation and both speeds and extends the range of the light response in photoreceptors, and modulates the communication between photoreceptors and horizontal cells. The entire signal transduction cascade from NOS through NO and sGC to cGMP has been demonstrated in horizontal cells. In addition to NOs reducing the response of their glutamate receptors, NO and cGMP can influence both the chromatic responses and network coupling in horizontal cells. In bipolar cells, there is a NO-sensitive sGC that can increase cGMP levels and the gain of the light response. In amacrine cells, in addition to depressing responses of GABAA receptors, NO can also influence the network coupling of All amacrine cells from rods to cones during light adaptation. Finally, ganglion cells have been shown to have cGMP-gated channels that are regulated by a NO-sensitive sGC.

6. RELEASE OF NO IN THE RETINA

To date, only the study of Neal et al. (1998) has directly examined the release of NO in the retina. The difficulty in directly measuring NO release is that it is quickly broken down into its stable breakdown products nitrite and nitrate before it can be reliably measured. To overcome this limitation, Neal et al. (1998) cleverly took advantage of this fact and converted all of the nitrate to nitrite using nitrate reductase and then converted all of the resultant nitrite to NO by using KI/HzSO4. Thus, they could use nitrate-nitrite formation to measure NO release from the retina. They find that there is a spontaneous release of NO from unstimulated dark-adapted retinas of 1.2 nmol/min and that treatment with arginine doubles this resting release. Note that this is substantially below the 4 nM/s of NO that Margulis et al. (1998) calculate that synaptosomal fractions from the inner retina can produce under optimal conditions. Stimulations with flickering or continuous light increases the NO release and NOS inhibitor blocks the light-stimulated release of NO. They find that the photoreceptors are not releasing NO, because the blocking of the ON-bipolar cells with APB also blocks the light-stimulated release. In contrast, the blocking of glutamate receptors with the nonselective 132

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glutamate receptor antagonist PDA, blocks the NO release stimulated by continuous light, but has no effect on the release stimulated by flickering light. From these results, Neal et al. (1998) conclude that because PDA blocks all glutamate synapses except from the photoreceptors to the ON-bipolar cells, the NO release stimulated by flickering light must be from ON-bipolars or Mtiller cells. In contrast, because both APB and PDA block the NO release stimulated by continuous light, it must be from amacrine cells. They speculate that in most cases there is a diffuse release of NO in the IPL that may be related to the reduction of the coupling from cone bipolars to AII amacrines during light adaptation, or to regulation of the gain in ON-bipolar cells. In contrast, flickering light causes NO release from specific ON-bipolar cells.

7. MODULATION OF TRANSMITTER RELEASE BY NO NO has also been shown to increase and decrease release of specific neurotransmitters. Ientile et al. (1996) find that stimulation with NMDA releases GABA, glutamine and glutamate in chick embryo retina, and that this effect is blocked by the NOS inhibitor, nitro-t-arginine. Stimulation with the NO donors, SNP or (+)-S-nitroso-N-acetylpenicillamine (SNAP), also increases the release of GABA and glutamine, but stimulation with cGMP has no effect. These results indicate that NO can stimulate transmitter release through a mechanism that does not involve cGMP. A direct.role for NO in stimulating synaptic release is provided by the studies of Meffert et al. (1994), that report that NO directly stimulates synaptic vesicle release in a calcium-independent manner, and by the studies of Meffert et al. (1996), that indicate that NO can directly interact with proteins involved in synaptic vesicle docking fusion reactions. In addition to increasing transmitter release, Bugnon et al. (1994) report that NO exerts a negative feedback effect on the basal and potassium-induced release of dopamine in bovine retina. They find that the NO donor, hydroxylamine, decreases release similar to the effects of potassium stimulation under calcium-free conditions, and that the NOS inhibitor, t-nitroarginine, increases basal dopamine release with no effect on potassium-stimulated release. They conclude that NO does not work through cGMP because cGMP itself has no effect on basal or potassium-stimulated release, and they suggest that possibly NO reduces release by decreasing intracellular calcium levels. The inhibition of dopamine release by NO is also demonstrated in rabbit retina by Djamgoz et al. (1995), who report that the NO donors SNP and SNAP also cause a dramatic decrease in potassium-stimulated dopamine release in rabbit retina. Their results indicate that GABAA receptors are not involved, and suggest that NO could be activating calcium-dependent potassium channels as in smooth muscle. Further experiments will be required to determine the mechanism of action of NO, and whether NO modulates the release of other transmitters as well.

8. THE cGMP SIGNAL TRANSDUCTION PATHWAY IN RETINA AND ITS MODULATION BY NO Since perhaps the best established function of NO is the activation of sGC to increase levels of cGMP, a discussion of retinal cGMP is necessary to put the role of NO in a functional perspective. Many aspects of the NOS/NO/cGMP signal transduction pathways are summarized in Fig. 1. Clearly, there are many interrelated systems regulating the cGMP levels in the retina. Several earlier studies carefully biochemically quantify and localize cGMP in the retina (Berger et al., 1980; review by Ferrendelli and De Vries, 1983). These 133

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W.D. Eldred Fia. 1. This tieurc illustrates some o f the hiochcmical pathways involved in the nitric oxidc/cyclic GMP s i m a l transduction bystem, inany of' which have hccn demonstrated in retina. It is designed to he a diagrammatic summary of the results From numerous studies and is not intended to indicate the precise cellular location or coIocitli/:iltion of specitic hiochernical pathways in a known cell type.

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studies report that cGMP levels in the retina are 10-100 times higher than in any other part of the nervous system, ~5 gM in whole mouse retina. The concentration of cGMP in the photoreceptors is 10-20 times higher than in the inner retina, with the photoreceptor outer segments having 2-3 times the cGMP found in the inner segments. In contrast, the levels of cGMP in the inner portions of control retinas are the same as in the brain. Cyclic GMP is synthesized by either pGC or sGC and there are several isoforms of each type of GC (Fig. 1; Wong and Garbers, 1992: Murad et al., 1993). Microdissection studies find high levels of GC in photoreceptor outer segments, where they are 30-40 times higher than in the brain and 10-50 times higher than in other retinal layers (Berger et al., 1980; review by Ferrendelli and De Vries, 1983). Outside of the photoreceptors, the IPL and OPL have the highest levels of GC. Soluble GC mRNA is expressed at high levels in the INL of rat retina, while pGC mRNA is primarily localized in the outer retina (Ahmad and Barnstable, 1993). However, Ahmad and Barnstable (1993) report that both sGC and pGC mRNA are in all layers of the retina and that 25% of the pGC is in the inner retina. In the rabbit retina, sGC has been localized immunocytochemically in many photoreceptor somas in the distal layer of the ONL, and in amacrine, ganglion and bipolar cells (Haberecht et al., 1998). Soluble GC has also been functionally localized in many amacrine, bipolar and ganglion cells in turtle retina (Blute et al., 1998). There are several forms of pGC in the retina. In photoreceptors, there are two pGCs, retGC (Dizhoor et al., 1994) and retGC2 (Lowe et al., 1995), which are both modulated by intracellular calcium (Koch, 1992). In addition to the retGCs, there is biochemical or molecular evidence in the retina for (1) A-type particulate GC (pGC-A) that can be activated by atrial natriuretic factor (ANF) or brain natriuretic factor (BNF) (Kutty et al., 1992), (2) B-type particulate GC (pGC-B), which is activated by C-type natriuretic factor (CNF) (Chrisman et al., 1992), and (3) C-type particulate GC (pGC-C) (Duda et al., 1993) that can be activated by heat-stable enterotoxin or guanylin (Chrisman et al., 1992). The ligand for pGC-A, ANF, has been detected in retina using biochemical techniques (Stone and Glembotski, 1986), and it is found diffusely in both the IPL and OPL of rat and rabbit retinas using immunocytochemistry (Palm et al., 1989). ANF receptors are found biochemically in rat and human retinas (Fernandez-Durango et al., 1989; Pardhasaradhi et al., 1994). mRNAs encoding pGC-A and pGC-B, as well as mRNAs encoding BNF and CNF are found in rat retina (Fernandez-Durango et al., 1995). Two other pGCs, E-type (pGC-E) and F-type (pGC-F), are in the eye, but their cellular location and precise function are unclear. It is thought that pGC-E and pGC-F, along with D-type particulate GC (pGC-D) in olfactory epithelia, represent a subfamily of pGCs restricted to sensory systems (Yang et al., 1995). With few exceptions, although molecular or biochemical studies indicate their presence in retina, the precise cellular location of specific pGCs and their activating ligands, and their function in regulating retinal cGMP, is not known. In addition to their biochemical studies of retinal NOS described above, Margulis et al. (1998) also examine retinal sGC in bovine eyes. They find that conventional synaptosomes from the inner retina have a 4-5 fold higher sGC activity than the sGC in the cytosol or in photoreceptor synaptosomes, and that 80% of the sGC activity is in the synaptosomes from inner retina. None of this sGC activity is influenced by calcium. They use their data to calculate that the measured sGC activity in inner retina could produce 2.3 nM of cGMP per second, and that, when stimulated with NO, the inner synaptic layers could produce 76-934 nM of cGMP per second. Given the fact that the volume of the band with increased cGMP in response to NO stimulation (Koistinaho et al., 1993) represents only about 10-15% of the IPL or 3% of the total retinal volume, they conclude that cGMP production could reach 135

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0.6-7.2 ~M/s in synaptic terminals. This value for the stimulated cGMP level fits well with the fact that 10-40 laM cGMP produces half saturation of cGMP-gated channels in rod outer segments (Fesenko et al., 1985; Koutalos et al., 1995). Such cGMP concentrations also fit well with the immunocytochemical demonstrations of NO-stimulated increases in cGMP in specific retinal neurons described below (Koistinaho et al., 1993; Blute et al., 1998). Extensive studies of the sensitivity of the cGMP antibody employed by Koistinaho et al. (1993) and Blute et al. (1998) indicate that it is able to detect as little as 10-5 M cGMP (De Vente et al., 1987, 1989). The early biochemical studies (Berger et al., 1980; review by Ferrendelli and De Vries, 1983) find a very high PDE activity in photoreceptor outer segments that is 15-300 times the activity in other retinal layers and 40 times that in the brain. Outside of the photoreceptors, the PDE activity is slightly higher in the plexiform layers. In both bipolar and ganglion cells, PDEs reduce the levels of cGMP, in that PDE inhibitors raise intracellular levels of cGMP and open cGMP-gated channels (Nawy and Jahr, 1991; Abroad et al., 1994). Another well-established function of cGMP is the activation of PKG, and the PKG activity in retina is three times that in whole rabbit brain (Berger et al., 1980; review by Ferrendelli and De Vries, 1983). These studies report that there is little PKG in the photoreceptor outer segments and in the inner retina, but that the levels of PKG activity peak in the photoreceptor inner segments and in the OPL. PKG phosphorylates several proteins in photoreceptor outer segments that are dephosphorylated by light. These PKGs are regulated by changes in cGMP concentration that are in the same range as those found in rod outer segments. Other studies use an antibody against cGMP itself to investigate the modulation of cGMP in specific cell types. Using this method, Berkelmans et al. (1989) find some cGMP-LI in photoreceptors and ganglion cells in control rat retinas. However, they find that stimulation with SNP increases cGMP-LI in the OPL and IPL, in cell bodies in the INL, and in photoreceptors and ganglion cells. They conclude that sGC is present in the INL and OPL, in amacrine cells and in fibrous structures in the IPL. Koistinaho et al. (1993) use the same cGMP antibody (De Vente et al., 1987, 1989) to examine control rabbit retinas and find cGMP-LI in photoreceptor outer segments and in some faint neurons in the GCL. Like Berkelmans et al. (1989), Koistinaho et al. (1993) report that stimulation with a NO donor increases cGMP-LI in photoreceptor inner segments and sparse ganglion cells. However, in rabbit the increased cGMP-LI is most apparent in two populations of bipolar cells, the majority being ON cone bipolars and the minority being OFF-midget bipolar cells. None of these bipolar cells with increased cGMP have protein kinase C and therefore rod bipolar cells are not involved (Greferath et al., 1990). Koistinaho et al. (1993) postulate that the amacrine cells with NOS could regulate their synaptic input. They speculate that flashing light could stimulate glutamate release from ON-bipolars that could cause the type I NADPH-diaphorase-positive amacrine cells to release NO, and in turn this NO could increase the cGMP levels in these same ON-cone bipolar cells and enhance their light response and transmitter release. Some of the most detailed studies of the activation of sGC by NO are in the turtle retina. Blute et al. (1998) functionally localize sGC within cells by using cGMP immunocytochemistry to detect the cell autonomous increases in cGMP-LI in response to the activation of sGC by NO. In turtle, NO-stimulated increases in cGMP-LI occur in at least eight different amacrine cell types, three bipolar cell types, some somas in the GCL and in discrete bands of processes in the IPL. These results are consistent with the high levels of sGC mRNA expressed in the inner retina (Ahmad and Barnstable, 1993). ELISA measurements indicate that stimulation with NO increases total retinal levels of cGMP by ~1500%. The control baseline levels of cGMP primarily represent the cGMP in approximately one hundred million 136

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photoreceptors (Ferrendelli and De Vries, 1983). Thus, the magnitude of this increase in cGMP in less than one hundred thousand amacrine, bipolar and ganglion cells suggests that the NO-stimulated levels of cGMP are remarkably high in these neurons. The broad neuronal distribution of NO-stimulated cGMP-LI suggests that the NO/sGC/cGMP signal transduction cascade is involved at several levels of visual processing in the inner retina. The study by Blute et al. (1998) indicates that exogenous NO activates sGC in specific neurons in turtle retina. A subsequent study by B lute et al. (1999), shows that activation of either kainate or NMDA-sensitive glutamate receptors also produces sufficient increases in endogenous NO to activate sGC and increase levels of cGMP. In these studies (Blute et al., 1999), stimulation with NMDA or kainate increases cGMP in overlapping subsets of the previously demonstrated sGC containing neurons, including 4-5 amacrine cell types, some bipolar cells and some somas in the GCL. These increases in cGMP are blocked by NOS inhibitors, which confirms the involvement of NOS and NO. ELISA measurements indicate that stimulation with either NMDA or kainate approximately doubles the total retinal levels of cGMP, which is consistent with the number of cells seen with increased cGMP-LI, in comparison to the number seen with NO stimulation. These results also indicate that by working through NO, activation of glutamate receptors can increase levels of cGMP in cells that may or may not even have glutamate receptors. A recent study by Gotzes et al. (1998) also examines NO stimulated increases in cGMP, but goes on to suggest that there are interrelationships of NO and cGMP between the inner and outer retina. They find that, in control bovine retinas treated with IBMX, there is some immunocytochemically detectable cGMP in mostly OFF- and a few ON-bipolar cells, and that there is weak cGMP in rod photoreceptor spherules, somas and inner segments, but not in photoreceptor outer segments. If these retinas are treated with NOS inhibitors, there is a total loss of cGMP-LI in the inner retina and an increase in cGMP-LI in the rod photoreceptor somas, inner segments and spherules. Treatment with SNP decreases the cGMP-LI in the outer retina, and increases cGMP-LI in both OFF- and ON-bipolar cells, in the nerve fiber layer and in amacrine and ganglion cells. They suggest that the endogenous NO released from the inner retina may inhibit cGMP synthesis in the outer retina. They conclude that the NO is probably released from amacrine cells, and that this NO may inhibit cGMP synthesis or enhance the degradation of cGMP in the outer retina. Finally, they propose that all of this relates to light and dark adaptation; just as dopamine is released during the light from amacrine cells and influences photoreceptor responses, NO is released in the light and increases levels of cGMP in the inner retina. Given that NO can influence dopamine release (Djamgoz et al., 1995), they speculate that NO and dopamine might interact to regulate light and dark adaptation in the retina. As mentioned above, Wexler et al. (1998) also propose that dopamine and NO/cGMP may function in a push-pull mechanism in GABAergic transmission. In summary, many aspects of the cGMP signal transduction system are extremely well developed in the retina. The retina has very high overall levels of cGME particularly in photoreceptors. This cGMP is produced by many different GCs, including the sGC found in high concentrations in inner retina, and by the multiple types of pGC, some of which are localized in specific cell types like photoreceptors. There are also clearly established roles for both PKG and PDEs in the retina, although few details are available. NO can activate sGC in large numbers of retinal neurons, and functionally active levels of NO are produced endogenously by the activation of glutamate receptors. Finally, there is a complex interactive regulation of NO and cGMP between the inner and outer retina. Clearly, NO and cGMP are important for many diverse aspects of retinal function, but many critical details remain to be clarified. 137

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9. F U T U R E AREAS OF INVESTIGATION It should be apparent from this review that NO plays many important roles in the function of the retina. The last ten years have provided many interesting details, but many questions remain. The existence and subcellular localizations of all the potential isoforms of NOS give rise to many questions. Although the existence of nNOS seems a certainty, recent studies describe more than one isoform of nNOS. Eliasson et al. (1997) report the existence of nNOS(x, nNOS[3 and nNOSu in brain, of which both nNOS6 and nNOSu are cytoplasmic because they are missing the postsynaptic density 95 (PSD-95) binding domain that could anchor them to membranes. It will be important to determine if these isoforms also exist in retina, and if the nNOS6 and nNOSy are responsible for the NOS-LI seen in the cytoplasm of the strongly stained amacrine cells. It is possible that nNOS(x is the isoform found at synapses while nNOSI3 or nNOSy are found in the cytoplasm, and nothing is known about the differential regulation of these enzymes. The details concerning the existence of both soluble and membrane-bound NOS, some of which is calcium-sensitive and some of which is not, also calls for further investigation. The application of new methods for the direct imaging of NO (Blute et al., 2000) will greatly facilitate the exploration of the activation of NOS isoforms and the release of NO from specific cell types. For instance, in many ganglion cells there is NOS in the soma but not in the processes of these cells. Is NO released from these somas, or does it just represent NOS that may be transported to the axon terminals of these ganglion cells? The existence, localization and function of both eNOS and iNOS need to be clarified. There is evidence both for and against the existence of eNOS in the retina, while the presence of iNOS in the retina is more conclusive. The existence of iNOS and the continuous basal levels of NO that it might produce raise the possibility that in the retina, NO may not just function as a neurotransmitter/neuromodulator. Physiologically, the retina is an extremely complex tissue with a very high rate of metabolism. It is entirely possible, that in some cases, the NO is regulating metabolism and not functioning in synaptic processing. This has already been suggested by the studies that localize nNOS in cells with a close relationship to the retinal vasculature. Such a metabolic function is also supported by McKee et al. (1994), who report that NO can modulate the activity of Na+/K+-ATPase, and by Clementi et al. (1998), who describe interactions of NO with mitochondrial metabolism. It is also very likely that NO will modulate levels of intracellular calcium, as has been clearly demonstrated in the brain (Clementi and Meldolesi. 1997). The regulation of calcium by NO will be extremely interesting in that both eNOS and nNOS are activated by calcium, which raises the possibility for some complex biochemical feedback relationships. Many studies of NO in retina have focused on the function of NO in specific cell types or transmitter systems, such as NO modulation of dopamine release or NO regulation of the junctional coupling between horizontal cells. However, in retina, gap junction coupling has been demonstrated between large numbers of diverse cell types (Vaney, 1994), which raises the possibility that NO may modulate coupling between many different cells in the retina. A similar enlargement of the role of NO may occur in the relationship of NO to neurotransmitter release. In brain, NO increases or decreases the release of many neuropeptides or neurotransmitters (Zhu and Luo, 1992; Bugnon et al., 1994; Garry et al., 1994; Jones et al., 1995), NO inhibits glutamate or dopamine (Pogun and Kuhar, 1994) transport into synaptosomes, and NO can evoke GABA release through either calcium-dependent release or through reversal of the calcium-independent and Na+-dependent transporter-mediated GABA uptake system (Ohkuma et al., 1996). Similar mechanisms may occur in the retina, where all 138

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of these transmitter systems are well developed. Therefore, it will be important to determine if the functions of NO that have been demonstrated in specific retinal cell types occur in other retinal cell types, and which of the diverse functions of NO in the brain also occur in retina. In summary, although other second-messenger pathways, such as the dopamine/cAMP system, have been shown to function in select cells in the retina, it is becoming clear that the NO/cGMP signal transduction system may play some role in every cell type in the retina. Although the biochemistry and pharmacology of these NO/cGMP pathways is extremely complicated by the numerous interrelated control mechanisms, it will be vital for future studies to characterize these pathways in order to truly understand retinal function.

10. ABBREVIATIONS

ADP ANF BNF BSPT cAMP cGMP CNF ELISA eNOS GABA GC GCL IBMX INL iNOS IPL kDa LI mRNA NADPH NMDA NO NOS OPL PDE pGC PKA PKG RT-PCR S sGC SNAP SNP

adenine dinucleotide phosphate atrial natriuretic factor brain natriuretic factor 2-(2'-benzothiazolyl)-5-styryl-3(4'-phthalhydrazidyl)-tetrazolium chloride cyclic adenosine monophosphate cyclic guanosine monophosphate C-type natriuretic factor enzyme linked immunos0rbant assay endothelial nitric oxide synthase gamma amino butyric acid guanylate cyclase ganglion cell layer isobutyl methyl xanthine inner nuclear layer inducible nitric oxide synthase inner plexiform layer kilodalton like immunoreactivity messenger ribonucleic acid nicotinamide adenine dinucleotide phosphate N-methyl-D-aspartic acid nitric oxide nitric oxide synthase outer plexiform layer phosphodiesterase particulate guanylate cyclase protein kinase A protein kinase G reverse transcriptase polymerase chain reaction strata soluble guanylate cyclase (+)-S-nitroso-N-acetylpenicillamine sodium nitroprusside 139

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ACKNOWLEDGEMENTS

I w i s h to t h a n k Dr. T o d d A. B l u t e a n d M i c h a e l L e e for c r i t i c a l r e a d i n g o f the m a n u s c r i p t , a n d I e s p e c i a l l y t h a n k Dr. B l u t e for m a n y useful d i s c u s s i o n s .

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De Vries SH, Schwartz EA (1988): Modulation of an electrical synapse between solitary pairs of catfish horizontal cells by dopamine and second messengers. J Phvsiol Land 414:351-375. Dizhoor AM, Lowe DG, Olshevskaya EV, Laura RE Hurley JB (1994): The human photoreceptor membrane guanylyl cyclase retGC is present in outer segments and is regulated by calcium and a soluble activator. Neuron 12:1345-1352. Djamgoz MBA, Cunningham JR, Davenport SL. Neal MJ (1995): Nitric oxide inhibits depolarization-induced release of endogenous dopamine in the rabbit retina. Neulvsci Lett 198:33-36. Djamgoz MBA, Aguilo R, Greenstreet EH, Reynolds R, Wilkin GP {1996): Histochemistry of NADPH-diaphorase a marker for neuronal nitric oxide synthase - - in the carp retina. Neurochem lnt 28:283-291. Duda T, Goraczniak RM, Sitaramayya A. Sharma RK (1993): Cloning and expression of an ATP-regulated human retina C-type natriuretic factor receptor guanylate cyclase. BiochemistpT 32:1391-1395. Eliasson MJL, Blackshaw S, Schell MJ. Snyder SH (1997): Neuronal nitric oxide synthase alternatively spliced forms: prominent functional localizations in the brain. Proc Natl Acad Sci USA 94:3396-3401. Fernandez-Durango R, Sanchez D, Gutkowska J. Carrier F, Fernandez-Cruz A (1989): Identification of atrial natriuretic factor receptors in the rat retina. Life Sci 44:1837-1846. Fernandez-Durango R, Nunez DJR, Brown MJ (1995) Messenger RNAs encoding the natriuretic peptides and their receptors are expressed in the eye. Exp Eve Res 61"723-729. Ferrendelli JA, De Vries GW (1983): Cyclic GMP systems in the retina. Fed Ppvc 42:3103-3106. Fesenko EE, Kolesnikov SS, Lyubarsky AL (1985): Induction of cyclic GMP of cationic conductance in plasma membrane of retinal rod outer segments. Nature 313:310-313. Fujii S, Honda S, Sekiya Y, Yamasaki M, Yamamoto M, Saijoh K (1998): Differential expression of nitric oxide synthase isoforms in form-deprived chick eyes. Curr Eve Res 17:586-593. Furukawa T, Yamada M, Petruv R, Djamgoz MBA, Yasui S {1997): Nitric oxide 2-amino-4-phosphonobutyric acid and light/dark adaptation modulate short-wavelength-sensitive synaptic transmission to retinal horizontal cells. Neurosci Res 27:65-74. Garry MG, Richardson JD, Hargreaves KM (1994): Sodium nitroprusside evokes the release of immunoreactive calcitonin gene-related peptide and substance P from dorsal horn slices via nitric oxide-dependent and nitric oxide- independent mechanisms. J Neurosci 14:4329-4337. Goldstein IM, Ostwald P, Roth S (1996): Nitric oxide: a review of its role in retinal function and disease. Vision Res 36:2979-2994. Gotzes S, De Vente J, Mtiiler F (1998): Nitric oxide modulates cGMP levels in neurons in the inner and outer retina in opposite ways. Vis Neurosci 15:945-955. Goureau O, Hicks D, Courtois Y (1994a): Human retinal pigmented epithelial cells produce nitric oxide in response to cytokines. Biochem Biophys Res Commun 198:120-126. Goureau O, Hicks D, Courtois Y, De Kozak Y (1994b): Induction and regulation of nitric oxide synthase in retinal Mtiller glial cells. J Neurochem 63:310-317. Goureau O, R6gnier-Ricard E Jonet L, Jeanny JC, Courtois Y, Chany-Fournier F (1997): Developmental expression of nitric oxide synthase isoform I and III in chick retina. J Neurosci Res 50:104-113. Greenstreet EH, Djamgoz MBA (1994): Nitric oxide induces light-adaptive morphological changes in retinal neurones. Neuroreport 6:109-112. Greferath U, Grtinert U, W~issle H (1990): Rod bipolar cells in the mammalian retina show protein kinase C-like immunoreactivity. J Comp Neurol 301:433-442. Haberecht MF, Schmidt HHHW, Mills S, Massey SC, Nakane M, Redburn-Johnson DA (1998): Localization of nitric oxide synthase NADPH diaphorase and soluble guanylyl cyclase in adult rabbit retina. Vis Neurosci 15:881-890. Haverkamp SH, Eldred WD (1998a): Localization of the origin of retinal efferents in the turtle brain and the involvement of nitric oxide synthase. J Comp Neurol 393:185-195. Haverkamp SH, Eldred WD (1998b): Localization of nNOS in photoreceptor bipolar and horizontal cells in turtle and rat retinas. Neuroreport 10:2231-2235. Hering H, Kr6ger S (1996): Formation of synaptic specializations in the inner plexiform layer of the developing chick retina. J Comp Neurol 375:393-405. Horio Y, Murad F (1991): Solubilization of guanylyl cyclase from bovine rod outer segments and effects of lowering Ca 2+ and nitro compounds. J Biol Chem 266:3411-3415. Huxlin KR (1995): NADPH-diaphorase expression in neurons and glia of the normal adult rat retina. Brain Res 692:195-206. Huxlin KR, Bennett MR (1995): NADPH diaphorase expression in the rat retina after axotomy ~ a supportive role for nitric oxide. Eur J Neurosci 7:2226-2239. 141

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Ientile R, Picciurro V, Pedale S, Nucci C, Malecka B, Nistic6 G. Macaione S (1996): Nitric oxide enhances amino acid release from immature chick embryo retina. Neurosci Lett 219:79-82. Jones NM, Loiacono RE, Beart PM (1995): Roles for nitric oxide as an intra- and interneuronal messenger at NMDA release-regulating receptors: evidence from studies of the NMDA-evoked release of [3H]noradrenaline and O-[3H]aspartate from rat hippocampal slices. J Neutw'hem 64:2057-2063. Koch KW (1992): Biochemical mechanism of light adaptation in vertebrate photoreceptors. Trends Biochem 17:307-311. Koch KW, Lambrecht HG, Haberecht M, Redburn D. Schmidt HHHW (1994): Functional coupling of a CaZ+/calmodulin-dependent nitric oxide synthase and a soluble guanylyl cyclase in vertebrate photoreceptor cells. EMBO J 13:3312-3320. Koistinaho J, Swanson RA, De Vente J, Sagar SM I1993): NADPH-diaphorase (nitric oxide synthase)-reactive amacrine cells of rabbit retina: putative target cells and stimulation by light. Neuroscience 57:587-597. Koutalos Y, Nakatani K, Tamura T, Yau KW (1995): Characterization of guanylate cyclase activity in single retinal rod outer segments. J Gen Phvsiol 106:863-890. Kurenni DE, Moroz LL, Turner RW, Sharkey KA, Barnes S (1994): Modulation of ion channels in rod photoreceptots by nitric oxide. Neuron 13:315-324. Kurenni DE, Thurlow GA, Turner RW, Moroz LL, Sharkey KA, Barnes S (1995): Nitric oxide synthase in tiger salamander retina. J Comp Neurol 361:525-526. Kutty RK, Fletcher RT, Chader GJ, Krishna G (1992): Expression of guanylate cyclase-A mRNA in the rat retina: detection using polymerase chain reaction. Biochem Biophys Res Commun 182:851-857. Lau KC, So KF, Tay D, Leung MCP (1994): NADPH-diaphorase neurons in the retina of the hamster. J Comp Neurol 350:550-558. Liepe BA, Stone C, Koistinaho J, Copenhagen DR (1994): Nitric oxide synthase in Mtiller cells and neurons of salamander and fish retina. J Neurosci 14:7641-7654. L6pez-Costa JJ, Goldstein J, Saavedra JP (1997): Neuronal and macrophagic nitric oxide synthase isoforms distribution in normal rat retina. Neurosci Lett 232:155-158. Lowe DG, Dizhoor AM, Liu K, Gu Q, Spencer M, Laura R, Lu L, Hurley JB (1995): Cloning and expression of a second photoreceptor-specific membrane retina guanylyl cyclase (RetGC) RetGC-2. Proc Natl Acad Sci USA 92:5535-5539. Margulis A, Sharma RK, Sitaramayya A (1992): Nitroprusside-sensitive and insensitive guanylate cyclases in retinal rod outer segments. Biochem Biophys Res Commun 185:909-914. Margulis A, Pozdnyakov N, Dang L, Sitaramayya A (1998): Soluble guanylate cyclase and nitric oxide synthase in synaptosomal fractions of bovine retina. Vis Neurosci 15:867-873. Matsumoto T, Nakane M, Pollock JS, Kuk JE. F6rstermann U (1993): A correlation between soluble brain nitric oxide synthase and NADPH-diaphorase activity is only seen after exposure of the tissue to fixative. Neurosci Lett 155:61-64. McKee M, Scavone C, Nathanson JA (1994): Nitric oxide cGMP and hormone regulation of active sodium transport. Proc Natl Acad Sci USA 91:12056-12060. McMahon DG, Ponomareva LV (1996): Nitric oxide and cGMP modulate retinal glutamate receptors. J Neutvphysiol 76:2307-2315. Meffert MK, Premack BA, Schulman H (1994): Nitric oxide stimulates Ca2+-independent synaptic vesicle release. Neuron 12:1235-1244. Meffert MK, Calakos NC, Scheller RH, Schulman H (1996): Nitric oxide modulates synaptic vesicle docking fusion reactions. Neuron 16:1229-1236. Mills SL, Massey SC (1995): Differential properties of two gap junctional pathways made by All amacrine cells. Nature 377:734-737. Mitrofanis J (1989): Development of NADPH-diaphorase cells in the rat's retina. Neurosci Lett 102:165-172. Mitrofanis J, Robinson SR, Ashwell K (1992): Development of catecholaminergic indoleamine-accumulating and NADPH-diaphorase amacrine cells in rabbit retinae. J Comp Neurol 319:560-585. Miyachi E, Murakami M, Nakaki T (1990): Arginine blocks gap junctions between retinal horizontal cells. Neuroreport 1:107-110. Morgan IG, Miethke R Li ZK (1994): Is nitric oxide a transmitter of the centrifugal projection to the avian retina. Neurosci Lett 168:5-7. Mtiller FH, Wiissle H, Brecha NC (1988): NADPH-diaphorase positive amacrine cells show GABA-Iike immunoreactivity in cat retina. EurJ Neurosci [Suppl] 1:153. Murad F, Fostermann U, Nakane M, Pollock J, Tracey R, Matsumoto T, Buechler W (1993): The nitric oxide142

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Shiells RA, Falk G (1990): Glutamate receptors of rod bipolar cells are liked to a cyclic GMP cascade via a G-protein. Proc R Soc Lond 242:91-94. Shiells RA, Falk G (1992): Retinal on-bipolar cells contain a nitric oxide-sensitive guanylate cyclase. Neuroreport 3:845-848. Stanarius A, T6pel I, Schulz S, Noack H, Wolf G (1997): Immunocytochemistry of endothelial nitric oxide synthase in the rat brain: a light and electron microscopical study using the tyramide signal amplification technique. Acta Histochem 99:411-429. Stone JR, Marletta MA (1995): Heme stoichiometry of heterodimeric soluble guanylate cyclase. Biochemistry 34:14668-14674. Stone RA, Glembotski CC (1986): Immunoactive atrial natriuretic peptide in the rat eye: molecular torms in anterior uvea and retina. Biochem Biophys Res Commtm 134:1022-1029. Straznicky C, Gabriel R (1991): NADPH-diaphorase positive neurons in the retina of Bufo marinus: selective staining of bipolar and amacrine cells. Arch Histol Cvtol 54:213-220. Ten6rio F, Giraldi-Guimaraes A, Mendez-Otero R 11996): Morphology of NADPH-diaphorase-positive cells in the retinoceptive layers of the developing rat superior colliculus, lnt J Dev Neurosci 14:1-10. Tracey WR, Nakane M, Pollock JS, FOrstermann U (1993): Nitric oxide synthases in neuronal cells macrophages and endothelium are NADPH diaphorases but represent only a fraction of total cellular NADPH diaphorase activity. Biochem Biophys Res Commtm 195:1035-1040. Tsuyama 1", N611 GN, Schmidt KF (1993): L-Arginine and nicotinamide adenine dinucleotide phosphate alter dark voltage and accelerate light response recovery in isolated retinal rods of the lYog (Rana temporaria). Neurosci Lett 149:95-98. Vaccaro TM, Cobcroft MD, Provis JM, Mitrofanis J (1991): NADPH-diaphorase reactivity in adult and developing cat retina. Cell Tissue Res 265:371-379. Vaney DI (1994): Patterns of neuronal coupling in the retina. Pro~ Retin Eve Res 13:302-355. Vaney DI, Young HM (1988): GABA-Iike immunoreactivitv in NADPH-diaphorase amacrine cells of the rabbit retina. Brain Res 474:380-385. Venturini CM, Knowles RG, Palmer RMJ. Moncada S (1991): Synthesis of nitric oxide in the bovine retina. Biochem Biophys Res Commun 180:920-925. Villani L, Guarnieri T (1996): Localization of nitric oxide synthase in the goldfish retina. Brain Res 743:353-356. Vincent SR (1994): Nitric oxide: a radical neurotransmitter in the central nervous svstem. Prog Neutvbiol 42:129160. Vincent SR, Hope BT (1992): Neurons that say NO. Tremts Neurosci 15:108-113. W~issle H, Hoon Chun M, Mtiller F (1987)" Amacrine cells in the ganglion cell layer of the cat retina. J Comp Neurol 265:391-408. Weiler R, Kewitz B (1993): The marker for nitric oxide synthase NADPH-diaphorase colocalizes with GABA in horizontal cells and cells of the inner retina in the carp retina. Neu~vsci Lett 158:151-154. Wellard JW, Miethke R Morgan IG (1995): Neural barriers affect the action of nitric oxide synthase inhibitors in the intact chicken retina. Neurosci Lett 201:17-20. Wexler EM, Stanton PK, Nawy S (1998): Nitric oxide depresses GABA,x receptor function via coactivation of cGMP-dependent kinase and phosphodiesterase. J Neurosci 18:2342-2349. Williams CV, Nordquist D, McLoon SC (1994): Correlation of nitric oxide synthase expression with changing patterns of axonal projections in the developing visual system. J Neurosci 14:1746-1755. Wislocki GB, Sidman RL (1954): The chemical morphology of the retina. J Contp Neurol 101:53-91. Wong SKF, Garbers DL (1992): Receptor guanylyl cyclases. J Clin hn'est 90:299-305. Yamamoto R, Bredt DS, Snyder SH, Stone RA I1993): The localization of nitric oxide synthase in the rat eye and related cranial structures. Neutvscience 54:189-200. Yang RB, Foster DC, Garbers DL, Ftille HJ (1995): Two membrane forms of guanylyl cyclase found in the eye. Ppvc Natl Acad Sci USA 92:602-606. Ye X, Laties AM, Stone RA (1990): Peptidergic innervation of the retinal vasculature and optic nerve head. hn'est Ophthalmol Vis Sci 31:1731-1737. Yoshida A, Pozdnyakov N, Dang L, Orselli SM, Reddy VN, Sitaramayya A (1995): Nitric oxide synthesis in retinal photoreceptor cells. Vis Neurosci 12:493-500. Young HM, Vaney DI (1989): GABA-Iike immunoreactivity in NADPH-diaphorase amacrine cells of rabbit and rat retinae. Neurosci Lett [Suppl] 27:S114. Young HM, O'Brien AJ, Furness JB, Ciampoli D, Hardwick JR McCabe TJ, Narayanasami R, Masters BSS, Tracey WR (1997): Relationships between NADPH diaphorase staining and neuronal endothelial and inducible nitric 144

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oxide synthase and cytochrome P450 reductase immunoreactivities in guinea-pig tissues. Histochem Cell Biol 107:19-29. Zhu XZ, Luo LG (1992)" Effects of nitroprusside (nitric oxide) on endogenous dopamine release from rat striatal slices. J Neurochem 59:932-935.

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CHAPTER V

Nitric oxide signaling in the hypothalamus B. WOODSIDE AND S. AMIR

1. INTRODUCTION The mammalian hypothalamus plays an important role in the control of autonomic and neuroendocrine functions such as cardiovascular activity and secretion of stress hormones; it is also implicated in diverse physiological and behavioral processes such as temperature regulation, energy balance, circadian rhythmicity, and reproductive behavior. Comprised of a network of anatomically discrete nuclei, the hypothalamus serves an integrative function by altering metabolic, endocrine, and autonomic activities in anticipation of, or in response to, specific organismic or environmental challenges. It is now well recognized that an important mechanism underlying the functional integrity of the hypothalamic surveillance system is neurochemical diversity. Transduction of signals to and from various hypothalamic regulatory systems involves disparate classes of excitatory and inhibitory transmitters, hormones, peptides, and second messengers. An important new addition to this neurochemical diversity is nitric oxide (NO). NO is a highly reactive free radical gas which, in neurons, is produced from L-arginine by a calcium- and calmodulin-dependent and NADPH-requiring neuronal form of the enzyme, nitric oxide synthase (NOS) (Garthwaite, 1991; Bredt and Snyder, 1992). Neuronal NOS is a constitutive protein that can rapidly and transiently synthesize and release small amounts of NO in response to stimuli that increase intracellular calcium concentration (Bredt and Snyder, 1990; Alagarsamy et al., 1994). Once released, NO can interact with various target proteins within the generating cell or in adjacent cells to influence ion channels, transmitter receptors, enzymes, and second-messenger systems (Bredt and Snyder, 1989; Knowles et al., 1989; Schmidt et al., 1993; Schuman and Madison, 1994; Vincent, 1994; Bhat et al., 1996). In this chapter studies of NO as a messenger molecule within the hypothalamus are reviewed. The focus of this chapter is the involvement of this messenger in the regulation of hypothalamic systems involved in neuroendocrine and autonomic control.

2. NOS IN THE HYPOTHALAMUS

Studies of the distribution of NOS mRNA and neuronal NOS protein within the mammalian central nervous system have been conducted using in situ hybridization as well as immunohistochemical and NADPH-diaphorase (NADPH-d) histochemical methods (Bredt et al:, 1990, 1991; Dawson et al., 1991; Hope et al., 1991). These studies have revealed the presence of a large number of NOS-containing neurons in the magnocellular and parvocellular subdivisions of the hypothalamic paraventricular nucleus (PVN), supraoptic nucleus (SON), nucleus circularis, and lateral hypothalamus. Additional groups of hypothalamic neurons that express Handbook o f Chemical Neun~anatom ~; Vol. 17: Functional Neunmnatom v o f the Nit ric Oxide Svstem

H.W.M. Steinbusch, J. De Vente and S.R. Vincent, editors @ 2000 Elsevier Science B.V. All rights reserved.

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NOS have been demonstrated in the preoptic nucleus, periventricular nucleus and portions of the ventromedial nucleus (VMH), lateral anterior nucleus, bed nucleus of the stria terminalis, magnocellular neurons in the horizontal limb of the diagonal band, supraoptic decussation, tuber cinereum, the medial and lateral tuberal nuclei, the organ vasculosum of the lamina terminalis (OVLT) (Vincent and Kimura, 1992: Vincent, 1994) and the suprachiasmatic nucleus (SCN) (Decker and Reuss, 1994; Lupi et al., 1996; Wang and Morris, 1996b: Chen et al., 1997). NO-producing neurons within the PVN project to the posterior pituitary and the brain stem and nerve endings containing NOS have been demonstrated in the internal portion of the median eminence (ME) (Arevalo et al., 1992: Vincent and Kimura, 1992; Vanhatalo and Soinila, 1995; Kawakami et al., 1998). As well as being a source of NO neurons the hypothalamus is also the recipient of NOS-positive inputs. For example, a nitrergic input from the amygdala to the hypothalamus has been identified (Tanaka et al., 1997). Illustrations of major NOS-containing hypothalamic nuclei as revealed by NADPH-d histochemistry are shown in Figs. 1-7.

3. CO-LOCALIZATION Hypothalamic NOS has .been co-localized with several neuropeptides, classical neurotransmitters, and receptor-types. Within the PVN, NOS-containing cells have been shown to contain corticotropin-releasing hormone (CRH) (Siaud et al., 1994), vasopressin (AVP) (Calka and Block, 1993b; Sanchez et al., 1996), oxytocin (OT) (Miyagawa et al., 1994), angiotensin (Calka and Block, 1993a), pituitary adenylate cyclase-activating polypeptide (PACAP) (Okamura et al., 1994a), calbindin (Alonso et al., 1992a), and enkephalin (Yamada et al., 1996). Similar patterns of co-localization have been observed in the SON where cholecystokinin (CCK) immunoreactivity as well as galanin message-associated peptide-like immunoreactivity have also been observed in some NOS-stained neurons (Yamada et al., 1996). In both the medial preoptic area (MPOA) and the VMH, NOS has been co-localized with substance P-like (Yamada et al., 1996) and with estrogen-receptor immunoreactivity (Okamura et al., 1994b,c). Some NOS-stained cells in the VMH also show enkephalin- and somatostatin-like immunoreactivity (Yamada et al., 1996). Within the SCN, NOS neurons have been shown to stain for vasoactive-intestinal polypeptide (VIP) (Reuss et al., 1995). In the dorsomedial nucleus enkephalin-, galanin message-associated peptide- and substance P-like immunoreactivity have all been shown to co-localize with NOS, and enkephalin-like immunoreactivity has also been found in NOS-containing neurons in the arcuate nucleus (Yamada et al., 1996). Finally, CCK, substance P, and enkephalin-like immunoreactivity have all been co-localized with NOS in the mammillary region (Yamada et al., 1996). With respect to the classical neurotransmitter systems, co-localization of NOS and acetylcholine has been reported in the PVN, SON, and nucleus circularis and co-localization of NOS and tyrosine hydroxylase has been observed in some neurons in the SON (Johnson and Ma, 1993; Blanco et al., 1997; Crespo et al., 1998). Consistent with the close association

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Fig. 1. Examples of NADPH-d staining in three anterior-posterior levels of the PVN. Line illustrations (derived from Swanson, 1992) on the right side of the figure indicate the approximate level of the PVN, and the area within the black box the location of the photomicrographs. 149

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between NMDA receptor activation and the induction of NO, the mRNA for NMDA-R1 receptors has been shown in NOS-containing neurons in the OVLT, MPOA, VMH, PVN, and SON (Bhat et al., 1995).

4. REGULATION Levels of NOS as reflected in the density of staining for NADPH-d, NOS immunoreactivity or mRNA hybridization signal within specific nuclei of the hypothalamus are modulated both by naturally occurring changes in the state of the organism and by a variety of exogenous stimuli. One potent class of stimuli associated with endogenous changes in state are the gonadal hormones. In male rats, castration has been shown to decrease levels of NOS in the MPOA (Hull et al., 1997; Shi et al., 1998; Du and Hull, 1999) and ovariectomized females show lower levels of NOS in both the MPOA (Okamura et al., 1994b) and the VMH (Ceccatelli et al., 1996" Rachman et al., 1998) than intact animals. In ovariectomized females levels of the enzyme in these areas increases following estrogen administration when this hormone is given acutely so that it mimics changes that occur across the estrous cycle (Okamura et al., 1994b; Ceccatelli et al., 1996; Rachman et al., 1998). The co-localization of NOS with estrogen-receptor immunoreactivity in the VMH and MPOA suggests that this is a direct effect of estrogen. NOS levels in the PVN and SON are also modulated by reproductive state and are increased during late pregnancy and lactation (Ceccatelli and Eriksson, 1993; Woodside and Amir, 1996; Xu et al., 1996; Popeski et al., 1999) as is the mRNA for NOS in the OVLT, and lamina terminalis (Luckman et al., 1997). NOS staining in the PVN and SON is not affected by administration of estrogen alone but does increase in response to a exogenous hormone administration that mimics the changes in ovarian hormone levels seen in late pregnancy (Popeski et al., 1999). In contrast, 4 days of estrogen treatment alone may be sufficient to increase NADPH-d staining in the pituitary (Wang and Morris, 1999). Together these data demonstrate the importance of gonadal hormones in the regulation of NOS and emphasize the fact that their effects are both site-specific and dependent on the pattern of hormone exposure. It is interesting to note that in contrast to the powerful effects of gonadal steroids on NOS levels, adrenal steroids do not seem to play a major role in regulating NOS levels in the hypothalamus. Bilateral adrenalectomy, for example, produces only a small and non-significant increase in NADPH-d staining in magnocellular neurons of the PVN (Sanchez et al., 1996). Thyroid hormone status does have a marked effect on mRNA for NOS, however. Hypothyroidism induced by propylthiouracil administration reduces mRNA for NOS and this effect can be reversed by adding T3 to the food (Ueta et al., 1995a). Finally, there are diurnal influences on NOS. Neuronal NOS immunoreactivity in the MPOA increases in the afternoon in female rats and increases in MPOA cGMP efftux in the afternoon, which are assumed to be NO-dependent, have also been observed in both male and female rats (Pu et al., 1998a,b). NOS levels in the SCN also fluctuate with the light/dark cycle with high levels being associated with the dark phase of the cycle. These fluctuations

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Fig. 2. Examples of NADPH-d staining in the posterior PVN. The line illustration (derived from Swanson, 1992) indicates the approximate level of the PVN, and the area within the black box the location of the photomicrographs. The bottom panel represents the same area at a higher magnification. 151

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disappear when the animal is placed in constant darkness (Ferreyra et al., 1998). Further, it has been shown that the pineal hormone melatonin, the release of which is restricted to the dark phase of the cycle inhibits NOS activity in the hypothalamus in rats (Bettahi et al., 1996). NO has also been implicated as a mediator of melatonin's effects on the suppression of norepinephrine-induced prostaglandin E2 and cAMP production (Bettahi et al., 1998). Diverse types of manipulations that activate the stress axis have been shown to upregulate NOS levels in the PVN and/or SON. These include challenges to the immune system such as injection of lipopolysaccharide (Lee et al., 1995" Jacobs et al., 1997; Harada et al., 1999) or interleukin-1 (Brunetti et al., 1996) as well as psychogenic stressors such as forced swim 152

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and restraint (Calza et al., 1993" Kishimoto et al., 1996 • Sanchez et al., 1999). In addition, a variety of homeostatic stressors such as salt loading (Pow, 1994: Villar et al., 1994b), water deprivation (Calka et al., 1994" Ueta et al., 1995c" O'Shea and Gundlach, 1996) and 153

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hypovolemia induction following propylene glycol administration increase NOS levels in these structures (Ueta et al., 1998). Conversely, food deprivation decreases NOS staining in the PVN (Ueta et al., 1995b; O'Shea and Gundlach, 1996). Decreased levels of NOS have also been observed in cholestatic rats (Swain et al., 1997). The sensitivity of this system to homeostatic challenge raises the issue of how it is changed in mutant strains that show 154

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disturbances of homeostatic systems. Studies of ob/ob mutant mice have found increased levels of mRNA for NOS whereas Zucker fa/fa rats show a decrease in mRNA for NOS in the hypothalamus (Morley et al., 1995; Morley and Mattammal, 1996). 155

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Diabetes either inherited or induced by streptozotocin administration is associated with an increase in NOS in the PVN and SON (Wang and Morris, 1996a" Yamamoto et al., 1997; Serino et al., 1998) as is hypertension (Alaghband-Zadeh et al., 1996; Plochocka-Zulinska 156

Nitric oxide signaling in the hypothalamus

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and Krukoff, 1997; Takeda et al., 1997). On the other hand, decreases in levels of NOS in PVN have been observed after experimentally induced heart failure (Patel et al., 1996; Zhang et al., 1998). In humans, reduced levels of NOS have also been observed in the PVN of schizophrenic patients (Bernstein et al., 1998). In addition to changes in endogenous state a number of external stimuli have been shown to modulate NOS levels in the PVN and SON. Consistent with the upregulation of NOS following water deprivation and salt loading, the administration of oxytocin (Melis et al., 1997b; Popeski et al., 1999) or angiotensin II (Dawson et al., 1998), both of which are implicated in the control of drinking and water balance, increases NOS levels in the PVN and/or SON. In addition, a series of studies have demonstrated that NO is increased in the PVN in response to administration of dopamine and of the glutamate receptor agonist N-methyl-D-aspartate (NMDA) (Melis et al., 1996, 1997a). As in other systems axonal damage induced by pituitary stalk transection also increases NOS levels in neurosecretory cells of the PVN and SON (Wu and Scott, 1993: Villar et al., 1994a; Lumme et al., 1997). Finally, recent evidence suggests that treatment with NO donors decreases NOS pointing to a possible negative feedback effect of NO on its own production (Canteros et al., 1996).

5. FUNCTIONAL CONSIDERATIONS 5.1. STRESS AXIS The synthesis and release of glucocorticoids from the adrenal cortex is regulated by the anterior pituitary hormone adrenocorticotropin (ACTH), whose secretion is controlled by the hypothalamic corticotropin-releasing hormone (CRH) neurosecretory system. Cell bodies containing CRH as well as other ACTH-releasing factors such as vasopressin are located in the parvocellular subdivision of the PVN: these cells project axons to the portal capillary plexus in the external zone of the median eminence. A number of findings prompted investigation of the role of NO in the modulation of the hypothalamic-pituitary-adrenal (HPA) axis. These include: (a)co-localization of NOS and CRH in neurons of the PVN (Siaud et al., 1994): (b) upregulation of NOS following both psychogenic and immune stress (Calza et al., 1993; Lee et al., 1995: Kishimoto et al., 1996; Harada et al., 1999; Sanchez et al., 1999): and (c) induction of immediate early gene expression in NOS-containing neurons following a variety of stressors including restraint, glucoprivation, hypotension, and morphine withdrawal (Petrov et al., 1995; Hatakeyama et al., 1996; Jhamandas et al., 1996; Amir et al., 1997; Krukoff and Khalili, 1997; Dawson et al., 1998; Briski and Sylvester, 1999; Yang et al., 1999). The results of the research on this topic have pointed to a complex relationship between NO and the stress axis. Studies using the hypothalamic slice preparation have shown that treatment with either the NO precursor L-arginine, or the NO donors molsidomine and sodium nitroprusside, did not affect basal release of CRH but did block the stimulated release of CRH induced by KC1 or interleukin-1. The inhibitory effect of L-arginine could be reversed by treatment with the competitive NOS blocker NG-monomethyl-L-arginine (L-NMMA) and the inhibitory effect of the NO donors on CRH release was blocked by treatment with hemoglobin, a scavenger of NO. On the basis of these findings it has been suggested that NO exerts an inhibitory effect on stimulated CRH release by hypothalamic neurons (Costa et al., 1993). Contrary to these findings, evidence has been reported suggesting that NO exerts a permissive or facilitatory 157

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effect on CRH release (Karanth et al., 1993). In these experiments, prolonged treatment with L-arginine enhanced both the basal release of CRH as well as interleukin-2-induced stimulation of CRH release from hypothalamic slices. Treatment with the NOS blocker L-NMMA completely suppressed the interleukin-2-induced release of CRH but had no effect on norepinephrine-induced CRH release. Additional data supporting a permissive role of NO on CRH release has been reported (Brunetti et al., 1993). These experiments showed that treatment with the competitive NOS blocker NG-nitro-L-arginine (L-NOArg) inhibits interleukin-l-induced release of CRH and ACTH from cultured hypothalamic neurons in vitro. Similarly, Sandi et al. (1995) using a rat hypothalamus perifusion system showed that pre- and co-incubation with a NOS inhibitor or a NO scavenger produced a strong attenuation of interleukin-1-stimulated CRH secretion. More recently, NO has been identified as a mediator of IL-10 stimulation of CRF release from median eminence fragments (Stefano et al., 1998). Both inhibitory and permissive effects of NO have also been reported from in vivo studies. The effects of manipulating NO on the stress axis depend on the type of stimulus administered and the parameters measured. In general, NO appears to inhibit the HPA response to immune stressors but facilitate that to psychogenic stressors. Using the c-fos gene product Fos as a marker of neuronal activation, it was found that immobilization stress activates PVN cells that express the NOS histochemical marker, NADPH-diaphorase (Hatakeyama et al., 1996; Amir et al., 1997). Further, systemic treatment with NOS blockers, either NG-nitro-L-arginine methyl ester (L-NAME) or 7-nitro indazole, significantly attenuates immobilization stress-induced Fos expression in the PVN (Amir et al., 1997) and the release of adrenal corticosterone in rats (Rackover et al., 1994). Pretreatment with D-NAME, an inactive isomer of L-NAME had no effect. Interestingly, a low dose of L-NAME that was sufficient to block the immobilization-induced release of corticosterone did not reduce the stress-induced expression of Fos in the NOS-containing PVN neurons although a higher dose attenuated both the corticosterone and Fos response to restraint stress. Although NO has been implicated in basal steroidogenesis (Adams et al., 1992) as well as in the control of basal adrenal vascular tone in the rat isolated adrenal gland preparation, it was found not to be involved in either the vascular or steroidogenic response to ACTH stimulation (Cameron and Hinson, 1993). These data provide further support for the conclusion that the inhibitory effect of L-NAME on stress-induced corticosterone release is mediated centrally and may not involve peripheral actions at the level of the adrenal gland. Contrary to the finding suggesting that NO has a permissive role in the control of stress-induced corticosterone release, other authors have obtained data suggesting an inhibitory effect of NO on acute inflammation-, interleukin-1-, vasopressin-, and oxytocin-induced ACTH release in vivo (Rivier and Shen, 1994; Turnbull and Rivier, 1996). Specifically, Rivier and Shen (1994) found that peripheral administration of the NOS blocker L-NAME enhances the stimulatory effect of peripherally administered interleukin-1, vasopressin and oxytocin on ACTH release, whereas treatment with L-arginine blunts the effect of interleukin-1 on ACTH release. Furthermore, systemic treatment with L-NAME enhanced lipopolysaccharide-induced ACTH and corticosterone release. Interestingly, systemic L-NAME had no effect on centrally administered interleukin-1-induced release of ACTH, suggesting that the facilitatory effects of L-NAME on ACTH and corticosterone release following systemic treatment with the different secretagogues was mediated downstream from the hypothalamus (Rivier and Shen, 1994). Additional evidence in support of the latter view is that although administration of NOS blockers reduced NGFlt3 expression in the PVN following footshock it had no effect on that seen following IL-1 administration (Lee and Rivier, 1998). 158

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While the studies conducted by Costa et al. (1993) and Rivier and Shen (1994) suggest an inhibitory influence of NO on cytokine-induced release of CRH, ACTH, and corticosterone, the other studies cited above are consistent with the view that NO exerts a facilitatory effect on the release of these stress hormones in response to restraint stress and footshock. Specifically, they suggest that NOS-containing neurons in the PVN are activated in response to stress, and that upon activation these neurons are induced to synthesize and release NO, which has a permissive effect on CRH secretion. This ultimately leads to enhanced release of ACTH from the anterior pituitary and, consequently, to enhanced synthesis and secretion of corticosterone. It remains to be determined precisely how NO influences CRH secretion from PVN neurosecretory neurons, and what are the neurochemical signals that trigger NO production in NOS-containing PVN cells during stress. One hypothesis, advanced by Karanth et al. (1993), suggests that production of NO in NOS-containing PVN neurons is coupled to the activation of muscarinic receptors by the neurotransmitter acetylcholine. Such activation would trigger calcium influx into the NOS-containing cell, leading to the rapid activation of the enzyme and release of NO. NO diffuses to the CRH neuron where it stimulates E2-prostaglandin production by activating the cyclooxygenase enzyme. E2-prostaglandin, through activation of adenylate cyclase, stimulates the generation of cAME which induces exocytosis of CRH secretory granules (Karanth et al., 1993). In general, the mechanisms by which NO facilitates CRH release are better understood than those by which NO suppresses CRH release from hypothalamic neurons or inhibits secretagogue-stimulated ACTH release from the pituitary (Costa et al., 1993; Rivier and Shen, 1994). It has recently been suggested that whereas stressors such as footshock stimulate ACTH release through actions at CRH cell bodies, the effects of interleukin-1 on ACTH release are mediated primarily through effects in the median eminence (Rivier, 1998). Thus the differential effects of NO on HPA activation following these two types of stress might reflect its opposing actions at different levels of the stress axis. 5.2. MAGNOCELLULAR NEUROSECRETORY SYSTEM

5.2.1. Modulation of vasopressin and oxytocin release Co-localization mapping studies have shown that a proportion of NOS-containing magnocellular neurons in the PVN and SON also express the peptides vasopressin (Calka and Block, 1993b) and oxytocin (Miyagawa et al., 1994). These neuropeptides play an important role in the regulation of water balance: both vasopressin and oxytocin are released from the posterior pituitary in response to osmotic and hypovolemic challenges such as water deprivation, salt loading, and hemorrhage. In addition, vasopressin stimulates the secretion of ACTH from the pituitary and oxytocin plays a critical role in reproduction via its peripheral effects on uterine contraction and milk ejection. Further, recent studies have implicated the central release of oxytocin not only in modulation of activity patterns in oxytocinergic neurons but also in the control of maternal behavior, and parasympathetically mediated responses such as penile erection and yawning. Interestingly, in spite of the wealth of evidence suggesting that NOS levels are upregulated at times when magnocellular neurosecretory cells are most active as for example following salt-loading and during lactation (Ceccatelli and Eriksson, 1993; Pow, 1994; Luckman et al., 1997; Popeski et al., 1999), the preponderance of evidence indicates that NO has an inhibitory effect on the release of vasopressin and oxytocin. Inconsistencies in the data, however, suggest 159

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that the effects of NO on the release of vasopressin, at least, may vary as a function of the state of the organism. Both electrophysiological and neurochemical studies have been carried out on hypothalamic tissue in vitro to evaluate the effects of NO in the PVN and SON. Single cell recordings from hypothalamus slice preparations show that bath administration of NO either directly or through treatment with precursors or donors depolarized parvocellular but not magnocellular cells of the PVN (Bains and Ferguson. 1997a). Similar treatments were found to increase GABA-dependent inhibitory postsynaptic potentials in magnocellular-type neurons (Bains and Ferguson, 1997b). Consistent with this inhibitory effect of NO on magnocellular cells, Liu et al. made electrophysiological recordings of the activity of SON neurons in vitro and showed that NO donors inhibited cell discharges whereas treatment with NOS blockers enhanced activity (Liu et al., 1997). Studies of the involvement of NO in the control of vasopressin release from incubated rat hypothalami in vitro have shown that treatment with the NO precursor L-arginine reduces KC1- and interleukin-l-evoked vasopressin release (Yasin et al., 1993). Similar inhibitory effects on KCl-evoked vasopressin release were noted after treatment with the NO donor sodium nitroprusside. Further, studies using in vivo preparations have shown that intravenous infusion of the NOS blocker L-NAME increases vasopressin concentration in conscious rabbits (Goyer et al., 1994). L-NAME treatment had no effect on the increase in vasopressin induced by hypertonic saline or sodium nitroprusside infusion, however, indicating that NO is involved in the control of basal vasopressin secretion, but not in vasopressin responses to hyperosmolality and hypotension. The same pattern of results was reported in conscious rats where central injection of L-NAME was shown to increase basal secretion of vasopressin but had no effects on the increased plasma levels seen following subcutaneous injection of hypertonic saline (Kadekaro et al., 1994). Similarly, Summy-Long et al. (1993) reported no increase in plasma vasopressin levels in response to central administration of NOS inhibitors in dehydrated rats although this manipulation did increase oxytocin release. More recently, however, Liu et al. (1997) have reported a stimulatory effect of L-NAME administration on vasopressin release in dehydrated rats but found that this effect occurred after a much longer latency than that at which an increase in oxytocin secretion was observed. In contrast to these reports of inhibitory effects of NO on vasopressin release, Ota et al. (1993) demonstrated that central injection of the NO donor S-nitroso-N-acetylpenicillamine or the NO biosynthetic precursor L-arginine stimulate vasopressin release in vivo in rats and it has been shown that NO stimulates both basal and reflex release of vasopressin in anesthetized rats (Cao et al., 1996). As is the case with vasopressin, studies of the involvement of NO in oxytocin release from the posterior pituitary have shown that under most circumstances NO plays an inhibitory role but in contrast to vasopressin, NO apparently suppresses both basal and stimulated OT levels. Administration of NOS blockers increases basal oxytocin secretion, enhances the oxytocinergic response to dehydration and hemorrhage, and increases OT depletion from the pituitary in salt-loaded rats. These data are consistent with the suggestion that in this system NO plays a role in negative feedback (Luckman et al., 1997). Interestingly, Summy-Long et al. (1998) have reported that central administration of t,-arginine, the substrate for NOS, also enhances oxytocin release in dehydrated rats which would point to an excitatory role for NO in oxytocin release. These authors, however, offer the intriguing notion that kyotorphin, a dipeptide that is formed from L-arginine in the brain may be producing the effects attributed to increases in NO production. Consistent with this hypothesis they show that kyotorphin administration does, indeed, produce effects on oxytocin secretion similar to those seen following L-arginine administration. 160

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5.2.2. Drinking Stimuli such as osmotic challenge and hypovolemia induce drinking as well as the release of vasopressin and oxytocin. The results of a number of studies suggest that the suppression of NO production attenuates drinking in response to these challenges (Calapai et al., 1992, 1994; Liu et al., 1998). Angiotensin-II-stimulated drinking is also reduced by the central injection of a NOS inhibitor as well as by the central administration of a guanylate cyclase inhibitor, methylene blue (Zhu and Herbert, 1997).

5.2.3. Reproductive behavior In rats, administration of NO donors both disrupts parturition (Okere et al., 1996a) and inhibits milk ejection (Okere et al., 1996b, 1999) both of which are dependent on the synchronous bursting of oxytocin neurons in the SON and PVN that results in bolus release of oxytocin from the posterior pituitary. Inhibitors of NO production had no effect on either of these processes. This pattern of results was attributed to the disruptive effects of NO donors on the pattern of neuronal discharge of magnocellular neurons (Okere et al., 1996a,b, 1999). It has also been reported that NO inhibits maternal behavior (Okere et al., 1996a), the onset of which can be stimulated by exogenous oxytocin administration. These effects, however, were found in animals in which parturition was prolonged by the administration of NO donors; thus the extent to which the changes in maternal behavior reflect abnormalities in delivery of young rather than a direct effect on maternal behavior itself is unclear. Oxytocin injection into the PVN of male rats induces both penile erection and yawning (Argiolas, 1994; Argiolas and Melis, 1995, 1998; Melis and Argiolas, 1997). Oxytocin treatment also increases NO production within this nucleus (Melis et al., 1997b) and the behavioral effects of oxytocin administration can be prevented by co-administration of NOS blockers (Melis et al., 1997b). The injection of NO donors into the PVN stimulates penile erection and yawning (Melis and Argiolas, 1995), and treatment with oxytocin receptor blockers into the ventricles but not into the PVN can eliminate the effects of NO donors on penile erection and yawning (Melis et al., 1999). These data are consistent with the notion that oxytocin stimulates these behaviors through an NO-dependent pathway involving cell bodies in the PVN. The activation of NO, in turn stimulates increased activity in an oxytocinergic pathway, the terminals of which are outside the PVN (Melis et al., 1999). Both dopamine and NMDA administration into the PVN induce yawning and penile erection; these actions are also suppressed by inhibition of NO production within the PVN (Melis et al., 1994, 1996). 5.3. THE REPRODUCTIVE AXIS

5.3.1. Modulation of luteinizing hormone-releasing hormone secretion Pulsatile release of both luteinizing hormone (LH) and pulsatile follicle stimulating hormone release (FSH) as well as the ovulatory surge of LH are controlled by the hypothalamic releasing peptide luteinizing hormone-releasing hormone (LHRH). LHRH is synthesized by neurosecretory cells in the anterior hypothalamus and preoptic area and released from terminals in the median eminence into the hypophysial portal system from where it is carried to the anterior pituitary to stimulate FSH and LH release into the general circulation. LH and FSH then act at the gonads to stimulate follicular development in females, spermatogenesis in 161

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males and gonadal steroid release in both sexes. The possibility that NO might be involved in the regulation of the hypothalamic LHRH neurosecretory system is consistent with a number of lines of evidence. NOS-containing neurons are located in close proximity to LHRH neurons in the OVLT and MPOA (Grossman et al., 1994; Bhat et al., 1995) although co-localization of NOS with LHRH neurons has not been reported. NOS staining is also seen in the ME where NO production could modulate LHRH release from terminals of neurosecretory neurons (Rettori et al., 1993). In addition, studies carried out using in vitro preparations have implicated NO in both stimulated and basal LHRH release. Basal release of LHRH from hypothalamic explants was enhanced dose dependently by treatment with the NO donor sodium nitroprusside (Moretto et al., 1993). Similar results were obtained using the GT-1 immortalized cell line (Moretto et al., 1993; Jung et al., 1998) which has been shown to express NOS (Mahachoklertwattana et al., 1994; Lopez et al., 1997). Treatment with a NOS blocker was also able to suppress pulsatile GnRH release from this cell line (Lopez et al., 1997). Norepinephrine (NE), glutamate and NMDA all stimulate LHRH release and treatment of medial basal hypothalamic fragments with the NOS blocker L-NMMA has been shown to block the effect of these manipulations while leaving basal levels of secretion unaffected (Rettori et al., 1994). In addition, the stimulatory effect of leptin, the protein product of the obese gene, on LHRH release has been shown to be antagonized by a NOS blocker (Yu et al., 1997). Most recently, evidence has been obtained suggesting that the inhibitory effects of [3-endorphin and granulocyte-macrophage colony stimulating factor on LHRH release are mediated by the ability of these substances to suppress a NO pathway (Kimura et al., 1997; Faletti et al., 1999). In contrast to the relationships observed in hypothalamic explants, in the GT-1 cell line NMDA suppresses GnRH gene expression, an effect that is also mediated by NO (Belsham et al., 1996). Other studies have also demonstrated an inhibitory action of NO on basal and stimulated LHRH release from GTI-1 neuronal cell line (Sortino et al., 1994) and on LHRH-induced release of pituitary LH (Ceccatelli et al., 1993). In vivo studies have demonstrated a role for NO in both pulsatile and surge release of LH. For example, injection of L-NMMA into the third cerebral ventricle blocks pulsatile LH release in rats (Rettori et al., 1993, 1994). Recent evidence also suggests that the tonic restraint exerted on LHRH release by opiates is mediated through NO (Bhat et al., 1998). Systemic injection of L-NMMA was found to block progesterone-induced augmentation of LH surges in estrogen-primed ovariectomized rats (Bonavera et al., 1993). Similarly, treatment with antisense oligonucleotides to both neuronal and endothelial NOS attenuated the steroid-induced LH surge in steroid-primed female rats (Aguan et al., 1996). It has also been shown that the ability of neuropeptide Y to facilitate the proestrus LH surge is dependent on NO (Bonavera et al., 1996) and interesting recent data suggest that the diminution of the LH surge in middle-aged rats reflects, in part, the absence of an upregulation of NOS in the hypothalamus on the afternoon of proestrus (Sahu, 1998). A number of mechanisms have been proposed to underlie the effects of NO on LHRH release. For example, it has been suggested that the effect of NO on LHRH release is linked to its stimulatory action on E2-prostaglandin synthesis within the LHRH-secreting neurons (Rettori et al., 1992, 1993, 1994). Others have suggested that the effect of NO on LHRH secretion is mediated by cGMP (Moretto et al., 1993). In these experiments, treatment with sodium nitroprusside stimulated LHRH release as well as induced an elevation in both extra- and intracellular cGMP levels in hypothalamic explants in vitro. The effect of sodium nitroprusside on LHRH release was blocked by Rp-8-Br-cGMP, a cGMP analog which blocks cGMP-dependent protein kinase, implicating cGMP as a mediator of the stimulatory effect of NO on LHRH release. 162

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5.3.2. Modulation of prolactin release As well as its effects on reproductive function through the modulation of LHRH release NO has also been shown in in vitro studies to influence prolactin release. The administration of NOS blockers suppresses both the changes in activity of tuberoinfundibular dopamine neurons and the associated prolactin surge seen following estrogen priming in ovariectomized females. Intracerebroventricular administration of an antisense oligonucleotide to the mRNA for NOS had similar effects (Yen and Pan, 1999). Similarly, the administration of NO donors decreases tyrosine hydroxylase activity in the dopamine tuberoinfundibular neurons of male rats and results in a dose-dependent increase in circulating prolactin release (Gonzalez et al., 1998).

5.3.3. Sexual behavior In addition to its role in modulating the release of sex hormones NO has also been implicated in the control of sexual behavior. For example, systemic administration of L-NAME has been shown to prevent ejaculation in naive rats although it was ineffective in experienced animals (Benelli et al., 1995). Although some of the effects that NO has on sexual function may be mediated through modulation of peripheral blood flow, there is growing evidence that NO may also act centrally to influence sexual behavior. In behavioral tests in male rats, L-arginine administration into the MPOA has been shown to increase mount rate, which is an index of sexual motivation, whereas administration of NOS blocker reduces mounting (Sato et al., 1998). Given that castration, in males, downregulates NOS in the MPOA it has been suggested that one pathway through which testosterone maintains copulatory behavior in the male rat is by upregulating NOS in this structure (Hull et al., 1997). In support of this hypothesis, it has been shown that NO is an important regulator of dopamine and serotonin release in the MPOA of male rats and facilitates dopamine release during copulation (Lorrain and Hull, 1993; Lorrain et al., 1996). Although penile erection is mediated by the parasympathetic system, ejaculation depends on sympathetic activation and NO apparently acts centrally to inhibit sympathetic outflow to the penile gland because L-NAME administration into the MPOA has been found to increase seminal emissions in ex-copulo tests of genital reflexes (Moses and Hull, 1999). NO has also been implicated in female sexual behavior. Administration of a NOS blocker into the third ventricle prevents progesterone-induced facilitation of lordosis behavior (Mani et al., 1994). This effect of NO is thought to be mediated by cGMP production because blockers of cGMP also suppress lordosis behavior (Chu and Etgen, 1996, 1997). It has also been suggested that NO regulates female sexual behavior, indirectly, through modulation of LHRH (Mani et al., 1994). As noted above, NOS is present in lordosis-relevant areas of the VMH and is modulated by circulating levels of ovarian steroids (Rachman et al., 1996, 1998) suggesting that the effect of NO on female sexual behavior is physiologically relevant. 5.4. SOMATOSTATIN RELEASE Coexistence of NOS and somatostatin (SRIF) has been demonstrated in some PVN neurons (Alonso et al., 1992b). The possibility that NO is involved in the control of SRIF synthesis and release has been assessed using periventricular nuclei of male rats in vitro. Treatment with the NO donor sodium nitroprusside stimulated SRIF mRNA levels and SRIF release; similar effect was observed after treatment with the cGMP analog, dibutyryl cGMP (Aguila, 163

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1994). Treatment with growth hormone-releasing factor (GRF) increased cGMP formation and SRIF mRNA level and stimulated SRIF release. These effects were blocked by L-NMMA but not D-NMMA, implicating the NO/cGMP pathway in the control of SRIF synthesis and release (Aguila, 1994). Recent data suggests that NO may also play a role in AMPA-mediated stimulation of growth hormone release (Gonzalez et al., 1999). 5.5. CIRCADIAN REGULATION Neuronal NOS assessed either with immunocytochemistry or NADPH-d histochemistry has been shown in the SCN, site of the circadian clock in mammals. Photic information necessary for the daily resetting of the clock is communicated to the SCN via a glutamatergic projection from the retina. Glutamate, acting through NMDA receptors to increase intracellular calcium, is a primary stimulus for the activation of NOS in central neurons (Southam et al., 1991; Luo et al., 1993; Luo and Vincent, 1994). NMDA receptors have also been implicated in the transmission of photic signals from the retina to the SCN (Abe et al., 1991; Colwell et al., 1991; Castel et al., 1993; Gannon and Rea, 1993; Reghunandanan et al., 1993). These observations provided the impetus for the investigation of the role of NO as a mediator of light-induced resetting of the circadian clock. In vitro studies using hypothalamic slice preparations containing the SCN have shown that treatment with NO donors mimics the effect of glutamate on cell firing rhythm in the SCN whereas treatment with blockers of NOS inhibits glutamate-induced and serotonin-induced phase shifts in cell firing rhythm (Ding et al., 1994, 1997; Starkey, 1996). In vivo, photic stimulation in the subjective night induces both expression of Fos protein in the SCN and phase shifts in free running activity rhythms. Some of the Fos expression observed was found to be localized to NOS-positive neurons (Amir et al., 1995; Castel et al., 1997). Similarly, light-induced phosphorylation of the transcription factor cyclic AMP response element binding protein (CREB) is also exhibited in NOS-containing neurons in the SCN (Ding et al., 1997). Furthermore, treatment with NOS blockers can inhibit light-induced Fos expression in the SCN (Amir and Edelstein, 1997) as well as light-induced phase shifts in free running rhythms (Watanabe et al., 1994, 1995; Weber et al., 1995a). Finally, it is known that NO exerts its effects through activation of cGMP signaling cascade and treatment with a blocker of cGMP-dependent protein kinase has been shown to inhibit light-induced phase shifts in free running circadian rhythms (Weber et al., 1995b). Together, these data are consistent with the idea that the signaling mechanism involved in light-induced circadian clock resetting uses NO as a mediator (Rea, 1998). 5.6. AUTONOMIC REGULATION Diverse hypothalamic nuclei have been implicated in the modulation of the autonomic nervous system. The localization of NOS within the hypothalamic nuclei involved in autonomic control suggests a potential role for NO as a modulator of neuronal activity within these centers. To study the direct effect of NO on the sympathetic outflow from the PVN, Horn et al. (1994) injected the NO donor sodium nitroprusside into the PVN of urethane-anesthetized rats and evaluated the effect on blood pressure and release of amino acid using microdialysis. Intra-PVN injection of sodium nitroprusside caused a decrease in blood pressure and a rise in extracellular glutamate within the PVN. In another study, Bains and Ferguson (1994) have shown that treatment with the NOS blocker L-NAME enhances the pressor response elicited by electrical stimulation of the subfornical organ, an effect which is mediated by angiotensin 164

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II release in the PVN. These studies suggest that NO acts in the PVN to decrease sympathetic tone and blood pressure. In the SON, activation of NMDA receptors leads to a rapid increase in sympathetic activity and heart rate (Amir, 1994). Studies of the role of the NO/cGMP signaling system in this effect have revealed that infusion into the SON of compounds that either block NO synthesis by NOS (L-NAME) or inhibit NO-induced stimulation of cGMP production by soluble guanylate cyclase (methylene blue) attenuate the increase in heart rate induced by intra-SON injection of NMDA. Furthermore, intra-SON injection of the calcium ionophore A23187, which stimulates NOS by increasing intracellular calcium directly, or injection of a membrane permeating analog of cGME 8-Br-cGMP, mimicked the effect of NMDA injection on cardiac acceleration. Thus, activation of NMDA receptors in the SON leads to a sympathetically mediated increase in heart rate; the signal from NMDA receptor stimulation to cardiac acceleration involves NO production by NOS and consequent increase in cGMP, which is mediated by NO-induced activation of soluble guanylate cyclase. Photic stimulation leads to increases in sympathetic outflow, a process mediated by NMDA receptor activation in the SCN (Amir, 1989, 1992; Amir et al., 1989; Niijima et al., 1992, 1993). To test the possibility that the NMDA-mediated link between photic stimulation and sympathetic nervous system activation involves release of NO in the SCN region, Amir (1992) examined the effect of infusing L-NAME into the SCN on light-induced cardiac acceleration in urethane.-anesthetized rats. L-NAME infusion into the SCN region attenuated the light-induced increase in heart rate; the inactive analog D-NAME had no effect. Similarly, infusion of methylene blue, a blocker of the cGMP synthesizing enzyme soluble guanylate cyclase which is activated by NO, attenuated the light-induced increase in heart rate. Infusion of L-NAME or methylene blue into the SCN region had no effect on cardiac acceleration induced by a non-photic stimulus, tail pinch, pointing to a selective role of this NO/cGMP signaling mechanism in the neural system that transduces photic stimulation to cardiac acceleration (Amir, 1992). Neurons containing NOS have been demonstrated in the preoptic area of the anterior hypothalamus (POAH), a structure involved in diverse physiological functions, including temperature regulation and cold-induced thermogenesis. Prostaglandin E2 injection into the POAH increases body temperature in urethane-anesthetized rats by stimulating heat production in brown adipose tissue (Amir et al., 1991). This effect is blocked by co-injection of the NOS blocker L-NMMA, suggesting that the elaboration or transmission of the thermogenic signal involves NO production (Amir et al., 1991). In rabbits, injection of NO donors into the OVLT induces fever and the antipyretic effects of dexamethasone have been attributed to suppression of the inducible form of NOS (Lin and Lin, 1996: Lin et al., 1997). Conversely, treatments that increase NO have been shown to restrain the rise in core temperature associated with lateral hypothalamic lesions (Monda et al., 1994). 5.7. PLASTICITY The demonstrated role of NO in various forms of neural plasticity in other parts of the nervous system suggest that NO might also be implicated in the processes underlying neural plasticity within the hypothalamus. For example, it has been suggested that the release of NO is a necessary step in the expression of some forms of activity-dependent synaptic plasticity, such as hippocampal long-term potentiation (Zorumski and Izumi, 1993). In this context, it has been proposed that NO serves as a retrograde messenger, allowing transmission of signals from the postsynaptic membrane to the presynaptic membrane, signals that ultimately 165

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bring about increases in neurotransmitter release. In addition, NO has been implicated in neuron-glial interactions (Do et al., 1996) and a major role has been attributed to NO in the formation of gap junctions (O'Donnell et al., 1999). There is ample evidence that neurosecretory cells and glia in the SON do, indeed, undergo structural remodelling, including changes in glial conformation, neuronal cell size and synaptic rearrangement as well as an increase in dye coupling following exposure to stimuli such as suckling and osmotic stress (Hatton et al., 1987; Theodosis and Poulain, 1993). As mentioned earlier, these stimuli also increase NOS mRNA and protein in PVN and SON neurosecretory cells (Sagar and Ferriero, 1987; Kadowaki et al., 1994; Pow, 1994) as does axonal injury induced by hypophysectomy (Wu and Scott, 1993; Scott et al., 1995). Within the magnocellular neurosecretory system itself a number of studies have demonstrated a role for NO in both neural reorganization in response to increases in activity and neural regeneration following injury. Scott et al. (1995) have shown that treatment with a competitive antagonist of NO inhibits the regrowth of neurosecretory axons into the neurohypophysis. Further, although effects of NO on glial reorganization within the SON itself have not been demonstrated, there is evidence that NO can change pituicyte morphology in culture (Ramsell and Cobbett, 1996) and that in vivo administration of L-NAME reverses the ultrastructural changes seen in the pituitary of dehydrated rats (Beagley and Cobbett, 1997). Finally, Yang and Hatton (1999) have recently demonstrated that dye coupling in the SON of virgin rats was increased almost four-fold by treatment with an NO precursor or NO donor. Interestingly, these treatments were less effective in lactating rats which have high endogenous levels of NOS but in the latter group treatment with a NOS blocker reduced baseline levels of dye coupling. Further experiments demonstrated that the effect of NO on dye coupling in this area was cGMP-dependent. Together, these data suggest that NO may play a potent role in the structural reorganization of magnocellular neurons.

6. CONCLUSIONS The studies reviewed above demonstrate that NO is involved in the regulation of specific hypothalamic neurosecretory and autonomic neuronal systems. There is compelling evidence that neurons within functionally and anatomically defined hypothalamic circuits involved in neuroendocrine and autonomic regulation express NOS. Further, stimuli that are known to activate these circuits can influence the expression of NOS mRNA as well as bring about an increase in the number of neurons expressing NOS. In addition, pharmacological interventions that influence NO production, that interfere with the action of NO on target proteins, or that mimic the action of endogenous NO on these proteins, influence the function of these pathways. Notwithstanding, some key questions about the function of the NO signaling system within the hypothalamus remain to be answered. In many instances, although the effectiveness of a particular stimulus or state in upregulating NOS is well established, the precise relationship to increased NO production is less clear. As to the signal transduction mechanisms that might trigger NO synthesis by NOS, excitatory amino acids are perhaps the most likely candidates. Indeed, glutamate terminals and receptors are widely distributed in the hypothalamus and have been shown to play a key role in the regulation of neurosecretory neurons and autonomic neurons. Moreover, NMDA receptors have been shown to be critically involved in stimulation of NO production throughout the nervous system (Garthwaite et al., 1988, 1991). Mediation by cholinergic neurons via muscarinic cholinergic receptors is also possible (Hu and E1-Fakahany, 1993), and has been discussed in relation to the mechanism by 166

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which interleukins influence CRH neurosecretion from hypothalamic slices in vitro (Karanth et al., 1993). The role of other transmitter systems, such as the serotonergic system, is less well defined. Another question concerns the mechanism by which NO influences the activity of hypothalamic neurons. The best known target of NO is soluble guanylate cyclase (Knowles et al., 1989; Southam and Garthwaite, 1993; Luo and Vincent, 1994), and it is most likely that some of the NO-mediated changes in hypothalamic neuronal function involve an increase in intracellular cGMP in the cells targeted by NO. NO may also influence neuronal function by stimulating the ADP-ribosylation of proteins (Brune and Lapetina, 1989). These NO-mediated cellular actions are likely to influence transmitter release and/or neurotransmitter receptor function, as has been reported in other brain regions (Lonart et al., 1992; Manzoni et al., 1992; Prast and Philippu, 1992; Zhu and Luo, 1992: Prast et al., 1996, 1997). For example, NO donors have been shown to decrease histamine release from superfused hypothalami (Prast et al., 1996), to mediate NE stimulated PGE2 release (Rettori et al., 1992) and may modulate dopamine release from the medial basal hypothalamus (Seilicovich et al., 1995). Finally, NO may influence neuronal activity by inducing long-term changes via effects on gene expression (Peunova and Enikolopov, 1993). Finally, the idea that NO functions as an intercellular messenger molecule in hypothalamic systems involved in neuroendocrine and autonomic regulation is now well established. The studies reviewed here are consistent with a general view that NO participates in the control of many different neurosecretory and autonomic processes. There is less agreement, however, as to the direction in which NO influences these processes. These apparent contradictions may be due merely to differences in experimental models and procedures used across different studies. It may, however, also represent the true nature of this elusive messenger molecule, namely an ability to exert diametrically opposed effects within discrete systems depending on the state of the organism. It is possible, for example, that the effect exerted by NO within a given system will change from one which is inhibitory to one which is facilitatory on the system, or vice versa, depending on the level of basal activity within the system, the type of stimulus presented to the system, or both.

7. A C K N O W L E D G E M E N T S

This work was supported by grants from the 'Fonds pour la Formation de Chercheurs et l'Aide ~t la Recherche du Qu6bec', the 'Natural Sciences and Engineering Research Council of Canada', and the 'Medical Research Council of Canada' to both B.W. and S.A.

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CHAPTER VI

Nitric oxide systems in the medulla oblongata and their involvement in autonomic control T.F.C. BATTEN, L. ATKINSON AND J. DEUCHARS

1. INTRODUCTION Since the introduction of the NADPH-diaphorase (NADPH-d) histochemical staining method for nervous tissues (Thomas and Pearse, 1961), its modification for use on aldehyde-fixed material (Scherer-Singler et al., 1983), and the confirmation that this is a reliable method for visualising nitric oxide synthase (NOS) in nitric oxide (NO) producing neurones throughout the nervous system (Dawson et al., 1991" Hope et al., 1991), many authors have reported widely distributed systems of NO neurones in many vertebrate species. These extensive and prominent neuronal systems throughout the brain were first described using the NADPH-d technique (Vincent and Kimura, 1992), subsequently using immunohistochemical detection of NOS (Dun et al., 1994; Rodrigo et al., 1994), and most recently by the detection of NOS mRNA using in situ hybridisation (Iwase et al., 1998). These authors have all briefly described the system of NO neurones in the medulla oblongata, while others (ladecola et al., 1993: Ohta et al., 1993; Simonian and Herbison, 1996" Krowicki et al., 1997: Lawrence et al., 1998) have studied these neurones in relation to specific areas of the medulla concerned with the control of particular autonomic functions. In this chapter, we will describe in detail the location of NOS-immunoreactive (NOS-IR) neurones and NADPH-d-positive neurones (which we will collectively refer to as 'nitrergic neurones') in the medulla of the rat, and provide anatomical evidence for the functional role of nitrergic neurones in the medulla by (1) examining the extent of the co-localisation of NOS-IR with immunoreactivities for other neuronal markers, (2) presenting data concerning the presence of NOS in vagal afferent and efferent neurones, and (3) analysing the synaptic relationships between vagal afferents and NOS-IR neurones in the dorsal vagal complex. We will also discuss our results in relation to those from other anatomical, pharmacological and physiological studies that have strongly implicated the involvement of neuronal NO as a potential transmitter in autonomic, somatosensory and viscerosensory pathways at the level of the medulla oblongata, in addition to its more well-established functions in neuronal development and neurodegeneration (Schuman and Madison, 1994; Anderson, 1998).

2. M E T H O D S

(1) Preparation of tissue. Male rats (150-200 g) were transcardially perfused with fixatives containing 0.1-0.5% glutaraldehyde and 4% paraformaldehyde, in 0.1 M phosphate Handbook of Chenffcal Neuroanatom ~; Vol. 17." Functional Neuroanatomy of the ,Vttri~ Oxide System H.WM. Steinbusch, J. De Vente and S.R. Vincent, editors @~ 2000 Elsevier Science B.V. All rights reserved.

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TABLE 1. Priman" antibodies used, their dilutions and their sottlz'es Primary antibody to

Abbreviation

Raised in

Dilution

Source

Nitric oxide synthase

NOS

Sheep Rabbit

1: 15000 1:1000

E Emson Afliniti

Tyrosine hydroxylase

TH

Mouse Rabbit

1:2000 1:2000

lncstar Eugene Tech

Glutamate

Glu

Mouse

1:4000

Incstar

y-Aminobutyric acid

GABA

Rabbit Mouse

1:5000 1:1000

Sigma Sigma

5-Hydroxytryptamine

5-HT

Rabbit Rat

1:3000 1 : 1000

.H. Steinbusch Sera Lab

Somatostatin

SS

Rabbit

1:2000

Peninsula

Neuropeptide Y

NPY

Rabbit

1:40(/0

Amersham

Cholecystokinin-8

CCK

Rabbit

I :5000

J.-J. Vanderhaeghen

CalbindinD28K

Calb

Mouse

1:1000

Sigma

Calretinin

Calret

Goat

1:10(10

Sigma

Parvalbumin

Parv

Mouse

1: 1000

Sigma

buffer (PB), pH 7.4. Brains were removed and post-fixed in the same solution for 4-12 h at 4~ Coronal sections of the medulla were cut at 30-60 g m using a vibrating microtome, and collected in 0.1 M PB. Subsequent procedures varied depending on whether the tissue was required for fluorescence microscopy or for light and electron microscopy. (2) Light microscopy. Sections were preincubated with 5% normal donkey serum (NDS) diluted in phosphate-buffered saline (PBS) containing 0.1% Triton X-100 for 30 min. They were then incubated in primary antiserum raised in sheep against recombinant rat neuronal NOS (nNOS) kindly provided by Dr. 19. Emson (University of Cambridge, UK) diluted 1:15,000 in PBS-Triton for 12-24 h. The specificity of this antiserum for nNOS has previously been established (Herbison et al., 1996). Following 3 x 10 rain washes in PBS, the sections were placed into biotinylated secondary antibody to sheep lgG (Amersham Life Science) for 2-4 h, and then into Vectastain Elite ABC reagent (Vector Labs) for 2 - 4 h. The bound antibodies were visualised by immersing in VIP substrate (Vector Labs) for 2-3 min. The sections were rinsed in PBS and distilled water, then mounted onto gelatinised slides and dried at 60~ overnight. After dehydration in graded alcohols and clearing in Histoclear, the sections were mounted under coverslips in Hystomount. (3) Fluorescence microscopy. The sections were processed as above as far as the incubation in primary antibody. For dual- or triple-labelled preparations, primary antibodies raised in different species at the appropriate dilutions (Table 1) were mixed in PBS-Triton buffer, and the incubations performed simultaneously. Species-specific secondary antibodies (Jackson Immunoresearch) were conjugated to Cy2 (green), Cy3 (red) or AMCA (blue). These were used at dilutions of 1:200-1500 depending on the primary antibodies to be detected, and for multiple labelling they were mixed together. Following washing, the sections were dried onto gelatin-coated slides at 4~ for 4-12 h. Coverslips were mounted with VectaMount reagent (Vector Labs), or with Hystomount, following a brief immersion in absolute ethanol and 178

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Histoclear. Slides were examined using a Zeiss Axioskop fluorescent microscope fitted with appropriate filter sets. (4) Correlated light and electron microscopy. The sections were either washed in 50% ethanol for 1 h or freeze-thawed in liquid nitrogen to permeabilise the tissue. They were then placed in primary antibody diluted in PBS for 12-48 h. Following 3 x 10 rain washes in PBS the sections were placed into secondary antibody conjugated to either 1 nm colloidal gold particles or biotin (Amersham Life Sciences) for 8-24 h. Biotinylated secondary antibodies were localised using the Vectastain ABC Elite kit (Vector Labs) and visualised using diaminobenzidine (DAB) as the chromogen. Gold particles were silver-enhanced for 5-10 min using an IntenSE kit (Amersham Life Sciences). Sections were then post-fixed in 0.5% osmium tetroxide for 1 h and dehydrated through a series of alcohols before immersing in Durcupan resin. The following day the sections were mounted on glass slides on a warm hotplate, a coverslip placed over them and placed in an oven at 60~ for 48 h. Slides were subsequently examined at the light microscopic level. When areas with suitable staining for electron microscopy were selected, the coverslip was removed, the relevant area cut out and glued to the flat surface of a resin block. Following trimming of the block, serial ultrathin sections were cut using a Leica UltraCut S ultramicrotome, collected on Formvar-coated 1 mm slot grids and examined on a Phillips CM10 transmission electron microscope. (5) Labelling of vagal afferent and efferent neurones. Vagal afferent fibres were anterogradely labelled in 150-200 g rats under halothane anaesthesia (5% in 02) by the injection of 5-10 ~1 of 10% biotinylated dextran amine (BDA, MW 10,000 kDa, Molecular Probes) into the nodose ganglion. Vagal efferent neurones were retrogradely labelled by injecting 5-10 [xl of 1% cholera toxin B-subunit (CTb, List Biologicals) into the nodose ganglion or the cervical vagus nerve. Following 7-10 days recovery the rats were reanaesthetised and perfused with fixative as described above. Sections of the medulla were cut on the vibrating microtome at 40-70 rtm thickness and permeabilised by freeze-thawing in liquid nitrogen or by immersing in 50% ethanol. BDA was then detected by incubation with ABC reagent, and CTb was detected with antisera generated in either rabbit (Sigma) or goat (List Biologicals), using biotinylated secondary antibodies. For light and electron microscopy the bound peroxidase was visualised with DAB. Sections were then subsequently transferred to the primary antibody for immunohistochemical detection of transmitters as described above, using 1 nm gold-conjugated secondary antibodies. Tissue was then processed for electron microscopy as described above. For fluorescence microscopy, BDA was visualised by using ABC reagent with a fluorescein-labelled tyramide signal amplification reagent (TSA-direct kit, NEN Life Science). CTb was detected using Cy2-conjugated anti-rabbit or anti-goat secondary antibodies. In both cases, subsequent dual labelling for other neuronal markers was performed with Cy3-conjugated secondary antibodies as described above. (6) Labelling of neurones projecting to the NTS. Male Wistar rats (280-310 g) were anaesthetised with halothane (5% in oxygen) and placed in a stereotaxic frame (SAS 4100, Harvard Apparatus). Two injections (1-2 l,tl each) of a 1.5% solution of CTb in sterile saline were made into the left commissural NTS. The co-ordinates of the injection sites were AP -13.8, DV -8.0, ML 0.5 and AP -14.3, DV -8.2, ML 0.3 measured in milimetres from bregma (Paxinos and Watson, 1997). Following 3 days recovery the rats were reanaesthetised and perfused with fixative as described above. CTb and NOS immunoreactivities were simultaneously visualised in vibratome sections using the rabbit antiserum to CTb and the sheep antiserum to NOS, with Cy2- and Cy3-conjugated secondary antibodies as described above. 179

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(7) Image manipulation. For light microscopy images were captured directly from the slide using a Hamamatsu Argus 20 Image Processor attached to a monochrome video camera. Images were manipulated in CorelDraw 8 to adjust gamma, brightness and contrast to the desired levels. For electron microscopy, the negatives were digitised using an Agfa Duoscan scanner and once again manipulated in CorelDraw 8. Final images were printed on photo-quality glossy paper using a 720 dpi Epson Stylus colour printer.

3. RESULTS AND DISCUSSION 3.1. ANATOMY OF NOS IN THE MEDULLA OBLONGATA 3.1.1. Description of the location and morphological characteristics of NOS-IR neurones in the rat medulla oblongata ~ The distribution of NOS-immunoreactive fibres and terminals in four different rostrocaudal regions of the medulla oblongata of the rat is illustrated in Figs. 1-4. We have also illustrated the pattern of staining in the NTS at different rostrocaudal levels since it varies considerably (Figs. 5 and 6). Viscerosensorv areas. Area postrema (AP) - - many very small scattered neurones and fibres. Nucleus tractus solitarii (NTS) - - characterised by many fibres, varicosities and groups of small- to medium-sized neurones throughout the nucleus, the numbers and packing density of the cells increasing towards rostral levels. Subnucleus gelatinosus (sg) - - relatively few intensely labelled scattered neurones and many fibres" commissural (co) and medial (me) subnuclei - - small triangular cells with variable labelling intensity, forming clusters at some levels; central (ce) subnucleus - - a large cluster of small, mostly ovoid neurones, some rather weakly labelled, with a high packing density and surrounded by a dense aggregation of fibres; dorsal (d) and interstitial (i) s u b n u c l e i - many small elongated bipolar neurones and fibres in a band extending over tractus solitarius (TS); ventral (v) and ventrolateral (vl) subnuclei medium-sized, intensely labelled, triangular and ovoid neurones with 2-4 prominent dendrites, forming clusters and extending in a band into the dorsal reticular area. At rostral levels these are densely packed with prominent, thick, contorted and varicose dendrites. Somatosensoo' areas. Gracile nucleus (Gr) small or very small, widely scattered neurones most numerous towards the periphery of the nucleus, mostly ovoid and bipolar, surrounded by a dense network of fibres. Cuneate nucleus (Cu) a group of small, strongly labelled triangular and ovoid neurones at caudal levels, replaced by only weakly labelled, isolated cells at more rostral levels. Relatively few varicosities and fibres. External cuneate nucleus (ECu) virtually devoid of any immunoreactive neurones or fibres. Spinal trigeminal nuclei b caudal division (Sp5C) - - patchily scattered cells and varicose fibres radiating in the interior (medial) part, many small triangular neurones and fibres in superficial layers, especially prominent clusters of bipolar cells in the dorsomedial and ventral areas of the nucleus at more rostral levels. A few larger cells sited at the boundary of the 1 N e u r o n a l s o m a t a < 10 ~ m d i a m e t e r described as "ver~ ,,mall'. 10-15 It m d i a m e t e r as "small'. 1 5 - 2 0 it m d i a m e t e r as " m e d i u m ' . 2 0 - 2 5 btm d i a m e t e r as ' l a r g e ' . > 2 5 t.tm d i a m e t e r as "very large'.

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Figs. 1-4. Atlas of NOS-IR neurones in the rat medulla oblongata visualised with the recombinant nNOS antibody and Cy3 Iabelling. Four coronal levels of sectioning are shown corresponding to bregma -14.60, -13.80, -13.30 and -12.80 of the Paxinos and Watson (1997) rat brain atlas. The boxed areas marked on the low power photomontage indicate areas containing NOS-IR neurones that are shown at higher magnification in panels A-J.

Fig. 1. Caudal level of the medulla (-14.60). Nuclei illustrated in boxed areas: (A) Gr, (B) Cu, (C) dorsal part of MdD (or DRt), (D) dorsomedial part of Sp5C, (E) neurones close to the cc, encapsulating the XII, in the DVN and coNTS, (F) dorsal part of MdV, (G) IRt in the region of the NA, (H) superficial Sp5C.

S p 5 C and the r e t i c u l a r f o r m a t i o n ; i n t e r p o l a r division ( S p 5 C I ) - - m a n y f e w e r cells and fibres, r e s t r i c t e d to the m o s t superficial layers; d o r s o m e d i a l division ( D M S p 5 ) - - a p r o m i n e n t c l u s t e r o f t r i a n g u l a r and ovoid n e u r o n e s , with a high d e n s i t y o f i m m u n o r e a c t i v e fibres. 181

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Fig. 2. Intermediate level of the medulla (-13.80). Nuclei illustrated in boxed areas: (A) AR coNTS and meNTS, (B) ts, dNTS and Gr, (C) ventral part of Cu, dorsal part of MdD (or DRt) and dorsomedial pole of Sp5C, (D) dorsal part of IRt, (E) ventral part of Sp5I, (F) dorsal MdV (nucleus of Probst's bundle) and Ro, (G) ROb and caudal part of PMn, (H) ventromedial part of MdV, (I) ventral part of MdV between LRt and IO, (J) ventral part of IRt surrounding the NA.

Paratrigeminal nucleus (Pa5) very dense network of varicose fibres and small ovoid intensely immunoreactive cells. Vestibular/auditoi3'. Medial vestibular nucleus (MVe) rather evenly distributed, well labelled, small triangular and ovoid neurones, with many dendritic and fibre profiles. Spinal vestibular nucleus (SpVe) very few isolated, weakly labelled, medium-sized neurones and few fibres. Nucleus prepositus hypoglossi (Pr) and nucleus intercalatus (In) clusters of small ovoid 182

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Fig. 3. Central level of the medulla (-13.30). Nuclei illustrated in boxed areas: (A) Ro, (B) dNTS, sgNTS and ceNTS, (C) vlNTS, (D) IRt, (E) dorsal part of PCRt (probably forming the caudal pole of DMSp5), (F) PMn, (G) ROb, (H) Gi, overlying the LRt, (I) C1 cell area, (J) central part of PCRt. neurones and varicose fibres, most numerous towards the ventricular surface. Visceromotor areas. Dorsal vagal nucleus (DVN) - - some weakly positive, medium-sized ovoid neurones and a few more intense elongated neurones, mostly situated in the lateral part. Nucleus ambiguus ( N A ) - isolated bipolar neurones in this region, but only very weak cellular labelling and varicose fibres occur within the compact zone. Somatomotor areas. Hypoglossal motor nucleus (XII) - - contains fibres and varicosities, but very few isolated neurones occur within the confines of the nucleus itself. A few large intensely labelled multipolar cells located medially to the XII bordering the central canal (cc) at caudal levels, and at the lateral margin of the nucleus more rostrally, with dendrites and 183

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Fig. 4. Rostral level of the medulla (-12.80). Nuclei illustrated in boxed areas: (A) Ro, In and medial tip of MVe, (B) ventral part of NTS and dorsal part of PCRt, (C) MVe and dorsal part of meNTS, (D) ts and vlNTS, (E) DMSp5 and dorsal part of Sp5I, (F) ROb, (G) GiV between the component nuclei of the IO, (H) LPGi, (I) IRt, (J) compact formation of NA.

varicose fibres crossing the XII towards the midline (but see also Ro and Prb below). Facial motor nucleus (VII) - - no labelled neurones, very few fibres or varicosities. Reticular formation. Nucleus of Roller (Ro), nucleus of Probst's bundle (Prb), and nucleus intermedius ( I n M ) several prominent groups of medium-sized, mainly ovoid, bipolar or triangular neurones occur in the most dorsomedial part of the reticular formation near the intersection of XII and DVN. Dorsal reticular area (MdD) - - many large mostly triangular multipolar neurones with prominent branching dendrites in the dorsal part, stretching down below vlNTS and DVN (but 184

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Fig. 5. Photomontages of the NTS at central level taken from adjacent 30 lxm sections stained by NADPH-d histochemistry and NOS immunohistochemistry (ABC-VIP method). Note that the relative numbers and distributions of the labelled cell bodies and fibres in the various subnuclei of the NTS are almost identical with both methods, with densely packed aggregations of small neurones and fibres in the central subnucleus (ceNTS), and more loosely packed groups of larger neurones in the medial (raNTS). ventral I vNTS) and ventrolateral (vlNTS) subnuclei.

most of these lie within the IRt). Similar, isolated neurones in the lateral part adjoining the Sp5C, but only a few isolated cells in the more ventrolateral parts. Intermediate reticular area (IRt) - - an almost continuous band of large triangular and ovoid neurones with prominent dendrites radiating dorsomedially and ventrolaterally into the reticular formation. Neurones appear to avoid the NA and A1 cell areas, but dendrites and fibres extend into these areas. Cells most numerous in this area at intermediate and rostral levels of the medulla. 185

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Parvocellular reticular field (PCRt) - - dense networks of fibres and varicosities, with widely scattered ovoid and triangular multipolar neurones, many rather weakly labelled. Some larger clusters occur in the dorsal part bordering the IRt, DMSp5 and Sp51. Lateral reticular nucleus (LRt) - - very few neurones within this nucleus, although many are scattered along the medial and dorsal aspects in the reticular formation, with dendrites penetrating into the LRt. Scattered fibres and varicosities. Ventral reticular area (MdV) - - large triangular and ovoid neurones with 2-5 main dendrites radiating in all directions are scattered throughout this area. At caudal levels these are aggregated towards the dorsal edge of the pyramidal tract (py) and medial to the NA and A1 cell areas but mainly aggregated in the ventral zone over the inferior olive, between the ROb and the LRt. Paramedian reticular area (PMn) - - small clusters of mainly bipolar and triangular neurones, some within the medial longitudinal fasciculus (mlf). Gigantocellular field (Gi) - - rather evenly scattered medium-sized neurones, mostly triangular with branching dendrites. Ventral gigantocellular field (GiV) - - as Gi, but a higher density of labelled neurones, many bipolar with long prominent dendrites, extending over and between the IO nuclei. Lateral paragigantocellular field (LPGi) - - similar large neurones to those of the Gi, but forming large clusters. Those towards the ventral surface of the medulla are very large in size with extensive dendritic arborisations. Raphe obscurus (ROb) ~ varicose fibres and isolated bipolar and multipolar neurones throughout, extending into the mlf. Inferior olive (IO) ~ scattered beaded fibres, but extremely few NOS-IR neurones within olivary complex, although cells similar to those seen in ventral reticular field do penetrate between the olivary subnuclei.

3.1.2. Relative distribution of NOS-IR and NADPH-diaphorase activity Two different methods have been used to map the distribution of nitric oxide-producing neurones in the central nervous system: the NADPH-diaphorase histochemical method (Scherer-Singler et al., 1983; Bredt et al., 1990: Leigh et al., 1990; Vincent and Kimura, 1992), and the later immunohistochemical method using antisera raised against nitric oxide synthase (NOS) (Dun et al., 1994; Egberongbe et al., 1994; Rodrigo et al., 1994). Evidence has been presented for several areas of the nervous system that in all likelihood the two techniques result in the visualisation of the same neuronal population (Bredt et al., 1991a; Dawson et al., 1991; Hope et al., 1991; Schmidt et al., 1992). The distributions of NOS-IR and NADPH-diaphorase staining throughout the medulla

Fig. 6. NOS-IR neurones are present at various levels of the NTS of the rat medulla./A. B) At caudal levels, NOS immunoreactivity can be identified by the presence of gold particles in cells in the dorsal part of the commissural NTS (A). The neurones in the boxed area can be seen at higher magnification in B. (C, D) At a caudal level similar to that shown in A and B, a discrete group of NOS-IR neurones can be observed ventral to the central canal. D is a higher magnification of the boxed area in C and illustrates that the labelling is clearly in neuronal somata and dendrites. (E, F) In areas of the NTS below the rostral pole of the area postrema a densely packed group of immunoreactive neurones can be observed in the central subnucleus (encircled in E). At higher magnification it is clear that the labelled neurones are small, with rounded somata IF). (G. H) In the rostral pole of the NTS, NOS-IR neurones are numerous (G), loosely packed (H) and vary in size and shape (H). 187

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of the rat and the cat have been compared by double-labelling sections and by examining series of adjacent sections. In dual-labelled 50 ~m vibratome sections, in which the NADPH reaction was performed prior to NOS immunolabelling, virtually all NADPH-positive cells displayed blue and brown reaction product within the cytoplasm of their soma and proximal dendrites. Approximately 80% of NADPH-positive cells in the rat medulla showed NOS-IR, and approximately 95% of NOS-IR cells showed NADPH staining. In adjacent 30 ~tm sections at all transverse levels, there was a close correspondence between the positions of NOS-IR cells or fibres and NADPH-diaphorase-positive cells or fibres in most anatomical divisions of both cat and rat medulla (see Fig. 5). Groups of cells intensely or weakly stained by one method showed a similar staining intensity with the other method. A few regions showed slight discrepancies, i.e. external cuneate nucleus, hypoglossal nucleus, inferior olive, nucleus ambiguus, where a large number of weakly NADPH-diaphorase-positive cells were sometimes found, but few NOS-IR cells were seen. It can be concluded, therefore, that identical sets of neurones in the medulla are labelled by the two different techniques. The small discrepancies noticed in the above studies are most likely explained by technical reasons inherent in the two methods. Firstly, additional neurones might be stained by the NADPH method, due to its relative lesser specificity for the nNOS enzyme than the well-characterised NOS antisera used. It is possible that structurally related enzymes such as endothelial NOS and cytochrome P450 oxyreductase (Bredt et al., 1991b; Norris et al., 1994) could be responsible for additional weakly reactive neurones not labelled by the NOS antisera. Secondly, discrepancies could be due to differential penetration of reagents into vibratome sections, with the smaller molecular size of the reagents for the NADPH reaction better able to penetrate into the interior of the sections than the larger antibodies used in the immunolabelling. Finally, and probably most important, are the different sensitivities of detection of the methods for neurones with low levels of NOS expression, allied with the effects of fixation on the reactive sites in the enzyme. This is demonstrated by the numbers of cells detected on adjacent sections using different antisera to NOS, and by the fact that adding higher concentrations of glutaraldehyde (>0.2%) to the fixative dramatically reduces the numbers of neurones detected by the NADPH method, whereas immunostaining with some NOS antibodies can tolerate glutaraldehyde concentrations up to 2% without any noticeable reduction in labelling.

3.1.3. Co-localisation of NOS-IR or NADPH-diaphorase activity with other neuronal phenotypes in the medulla oblongata Many authors have described NADPH-d activity co-localised with neuroactive substances within specific regions of the central nervous system. The presence of the enzyme has been reported in serotonergic cells in the mesopontine areas of the brain (Johnson and Ma, 1993). Other studies have demonstrated co-localisation with calcitonin gene-related peptide (CGRP)-IR and substance P-IR in neurones within cranial and dorsal root sensory neurones (Aimi et al., 1991; Ichikawa and Helke, 1996), and in cholinergic, glycinergic and GABAergic neurones of the spinal dorsal horn (Spike et al., 1993). NOS-IR has been found in a subpopulation of GABAergic local circuit neurones in the ventrobasal complex of the cat (Meng et al., 1996), and of GABAergic amacrine cells in the retina (Oh et al., 1998). Both GABA and/or the calcium-binding protein calbindin D28K have been detected in NOS-IR cortical neurones of several species (Bertini et al., 1996; Yan and Garey, 1997), while calbindin has also been detected in nitrergic neurones of the intermediolateral spinal cord (Grkovic and Anderson, 1997), lateral septum (Doutrelant-Viltart and Poulain, 1996) and in the petrosal and 188

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nodose ganglia (Ichikawa and Helke, 1996). A further calcium-binding protein, calretinin, was reported in significant proportions of NADPH-d-reactive neurones of the hippocampus and dentate gyrus (Megias et al., 1997). In hypothalamic neurosecretory neurones, co-localisation of NADPH-d with somatostatin, vasopressin, angiotensin and calretinin has been reported (Alonso et al., 1992; Ar6valo et al., 1993; Calka and Block, 1993a,b), and in other areas of the hypothalamus immunoreactivities for enkephalin and substance-P were frequently found in NOS-IR neurones (Yamada et al., 1996), with other peptides such as galanin, cholecystokinin (CCK) and somatostatin (SS) also occurring in minor populations of nitrergic cells in some areas. SS and avian pancreatic polypeptide-like (probably NPY) IR were also found to coexist with NADPH-d activity in neurones of the striatum and neocortex (Vincent et al., 1983a). NADPH-d activity has also been found to co-exist with choline acetyl-transferase (CHAT) in a number of regions including the NTS, the compact zone of the NA, raphe nuclei, PGLi, lateral tegmental field, dorsal horn and sympathetic columns of the spinal cord (Ruggiero et al., 1993; Spike et al., 1993), and in neurones of the pontine reticular formation (Vincent et al., 1983b). Both galanin and 5-HT immunoreactivities were found to coexist with NOS-IR in dorsal raphe neurones (Xu and H6kfelt, 1997). Neurones of the areas of the medulla that control autonomic functions contain a wide range of neuropeptides and neurotransmitters (Leibstein et al., 1985; Ruggiero et al., 1989; Batten, 1995). Two of the neuronal groups that have been most extensively studied are the vagal parasympathetic preganglionic neurones, and the vasomotor neurones of the RVLM that project onto the sympathetic preganglionic motorneurones located in the thoracic spinal cord. The latter group of neurones have been identified as the C 1 group of adrenergic neurones (Ross et al., 1984), and these occur in a very similar area to a large group of NADPH-d-reactive neurones (Iadecola et al., 1993). It has also been suggested that the sympathoexcitatory vasomotor neurones of the RVLM may be glutamatergic rather than catecholaminergic (Sun et al., 1988a,b). Therefore, it was of great interest to examine the possible co-existence of NADPH-d or NOS activities with immunoreactivities for catecholamine synthesising enzymes and glutamate in this region, and furthermore with immunoreactivities for other neuropeptides and neurotransmitters (e.g. SS, NPY, CCK) which have been localised to RVLM neurones projecting to the area containing the sympathetic preganglionic neurones of the intermediolateral columns of the spinal cord (Mantyh and Hunt, 1984; Blessing et al., 1986, 1987; Millhorn et al., 1987). The following sections therefore discuss the co-localisation of NOS with vagal afferent and efferent structures, bioamines, neuropeptides, amino acids and calcium-binding proteins. (a) Presence of NOS in vagal afferent and efferent structures. One possible source of NO in the NTS is the vagal afferent fibres. Studies to support this include: (i) neurones in the nodose ganglia stain positive for NADPH-d (Aimi et al., 1991; Morris et al., 1993; Ruggerio et al., 1996); (ii) in situ hybridisation indicates that cell bodies in the nodose ganglia express mRNA encoding NOS (Lawrence et al., 1996). NOS could therefore be transported from these cell ~odies to vagal afferent terminals in the NTS. The presence of NOS in central vagal terminals has also recently been inferred by a decrease in NOS-IR in a restricted portion of the ipsilateral medial NTS following nodose ganglionectomy (Lawrence et al., 1998), and this was illustrated by the presence of NOS immunoreactivity in 67% of degenerating terminals in the NTS following a nodose ganglionectomy (Lin et al., 1998). However, we have been unable to confirm the presence of NOS in vagal afferents despite combination of anterograde tracing from the nodose ganglion with NOS immunohistochemistry (Fig. 7). There are many other possible sources of NOS-IR fibres and terminals in the NTS as shown by studies injecting tracers into this nucleus. These include the dorsal horn of the spinal cord (Esteves 189

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Fig. 7. Relationship between NOS-IR neurones (B, D, F) with vagal preganglionic efferent neurones (A) and vagal afferent fibres (C, E) shown by double labelling. (A) Cell bodies of vagal efferent neurones in the DVN labelled by CTb injection into the vagus nerve, visualised with Cv2. (B) Corresponding area, showing NOS-IR neurones and fibres labelled with Cy3. Some cells are strongly labelled for both CTb and NOS (longer arrows in A and B), some CTb-containing cells are weakly immunoreactive for NOS (open-headed arrows), and some cells labelled lot CTb are not NOS-IR (double-headed arrow in A). A strongly NOS-IR neurone that is not labelled for CTb is indicated by a larger arrowhead in B. (C) Vagal afferent fibres in the TS and central NTS at an intermediate level, labelled by injection of BDA into the nodose ganglion, and visualised by the TSA-fluorescein method. (D) Same area, showing NOS-IR neurones and fibres (Cy3 labelling). Although NOS-IR fibres are concentrated in the same areas as the vagal afferents, there is no clear evidence of dual-labelled fibres or terminals. (E) Vagal afferent fibres in the TS and central NTS at a more rostral level (TSA-fluorescein). (F) Same area showing NOS immunoreactivity (Cy3). Again, there is little evidence for dual labelling. 190

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Fig. 8. NOS-IR neurones are present in the dorsal vagal nucleus (DVN) and the nucleus ambiguus (NA). (A) Photomontage of the dorsomedial area of a coronal section of the rat medulla showing both dorsal vagal nuclei. NOS-IR neurones are indicated by the arrows. The DVN on the right-hand side contains more labelled neurones than the left-hand side, and the labelling of the cells is noticeably more intense. This increased NOS expression may be a result of damage to the axons of the vagal neurones on the right-hand side. due to injections of tracer into the vagus nerve. (B) NOS immunoreactivity in the compact region of the NA can be detected in a dense network of fibres as well as some somata. (C) A NOS-IR neurone in the compact NA is apposed by two NOS-IR terminals (arrows).

et al., 1999), paratrigeminal nucleus (Armstrong and Hopkins, 1998) and raphe obscurus (Fig. 14). NOS and NADPH-d have also been detected in vagal efferent neurones (Mizukawa et al., 1989; Krowicki et al., 1997; Zheng et al., 1999: Figs. 7 and 8). Several anatomical studies have localised NOS in DVN neurones (e.g. cat: Mizukawa et al., 1989; Figs. 7 and 8). These nitrergic neurones are likely to be vagal neurones, since intra-peritoneal injection of Fluorogold, which labels all preganglionic neurones, combined with NADPH-d histochemistry revealed double-labelled neurones in the caudal and rostral, but not intermediate, zones of the DVN (Krowicki et al., 1997; see also Figs. 7 and 8). Retrograde tracing from the gastric fundus labelled a discrete population of NOS-immunoreactive neurones in the medial portion of the DVN (Zheng et al., 1999). However, it can be difficult to interpret this kind of double labelling since neurones in the DVN may upregulate NOS expression in response to 191

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axonal damage, such as that which might occur during a retrograde tracing experiment as a result of nerve section or merely the injection of a tracer (Fig. 8; Jia et al., 1994; Yu, 1994). While it is therefore difficult to determine if NOS immunoreactivity would have been detectable in the double-labelled cells without the experimental intervention, it is clear that weak staining, probably representing low levels of NOS, occurs within the DVN in a few neurones with the morphology of parasympathetic efferent neurones even in intact animals (Figs. 8 and 9). A few moderately stained NOS-immunoreactive neurones have been localised within or near to the nucleus ambiguus (Dun et al., 1994; Fig. 8). However, these neurones are not immunoreactive for cholinergic markers (Dun et al., 1994) and are therefore unlikely to be vagal efferent neurones. This is consistent with lack of double-labelled neurones in the NA following combination of retrograde tracing from the cervical vagus or the heart with NOS immunohistochemistry (E. Corbett, unpubl, observations). (b) Bioamines. Our double-labelling studies using either enzymatic or fluorescence methods on cat and rat brain stem sections demonstrated that very few NOS-containing neurones in the medulla were immunoreactive for the catecholamine synthesising enzymes tyrosine hydroxylase (TH), dopamine-[3-hydroxylase (DBH) or phenylethanolamine N-methyl transferase (PNMT). In both the caudal and rostral ventrolateral medulla the NOS cells were generally located more dorsal and more medial than the TH-IR cells (of the A1 and C1 cell groups, respectively). This distinction was most striking at the more rostral levels of the brain stem, and we have found no dual-labelled cells in either area (Fig. 10). In the dorsomedial medulla, the two groups of neurones were more intimately mingled, but catecholamine cells of the DMV and NTS (A2 and C2 groups) were only rarely (~ 10% of cases) found to be NOS positive (Fig. 10). NOS-IR terminals were found apposing catecholamine neurones in both the NTS and ventrolateral medulla, and TH-IR or DBH-IR terminals were observed on NOS/NADPH-reactive neurones in both these areas. These results are in general agreement with the earlier findings of Iadecola et al. (1993), Ohta et al. (1993), Dun et al. (1994) and Simonian and Herbison (1996), although the latter authors reported a small degree of TH-NOS coexistence in both A 1 and C 1 regions of the ventrolateral medulla. NOS-IR neurones in the nucleus raphe obscurus and ventral gigantocellular reticular nucleus have a similar morphology and distribution to the 5-HT-IR neurones in these nuclei. Nevertheless, in agreement with Dun et al. (1994) dual-labelling studies in the rat have demonstrated that the two immunoreactivities are generally localised to separate neurones, with only occasional examples of dual-labelled cells being found (Fig. 11). These studies revealed, however, evidence for the existence of an intimate reciprocal innervation of NOS-IR cells by 5-HT-IR fibre varicosities and vice versa. (c) Neuropeptides. Neurones staining for both NOS and SS immunoreactivity were found scattered within the paramedian and lateral tegmental field of the cat medulla. These dual-labelled cells accounted for 6% of the total NOS cells in both these areas and comprised 15% of the paragigantocellular NOS cells (Maqbool et al., 1995). In the most rostral medulla oblongata a few double-labelled cells were observed in the NTS (12%) and in the Pr (9%). In the rat medulla, dual-labelled neurones could be found scattered throughout several areas of the reticular formation, particularly in the dorsal (Fig. 12) and intermediate medullary reticular nuclei, in the InM, Ro and Prb. SS-IR was co-localised to very few NOS-IR cells throughout the NTS (Fig. 12) and spinal trigeminal nuclei, and to a small percentage of neurones in the DMSp5. Similar double-labelling studies demonstrated that NOS-reactive neurones were not immunoreactive for CCK although they were co-distributed with CCK-IR cells within the 192

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Fig. 9. Vagal afferent fibres labelled from the nodose ganglion closely appose NOS-immunoreactive neurones in the dorsal vagal complex. (A) A low power view of the dorsal vagal nucleus tDVN). The tractus solitarius (TS) can be seen at the top of the micrograph. The boxed region indicates the area shown at higher magnification in B. (B) The DVN contains many neurones immunoreactive for NOS. The neurone enclosed by the box is magnified in C. (C) A NOS-IR neurone in the DVN, identified by the presence of silver intensified gold particles, is closely apposed by a vagal afferent terminal (arrow). (D) Another NOS-IR neurone in the DVN is also closely apposed by two vagal afferent fibres (arrows).

lateral reticular field, within the paramedian field and dorsal to the inferior olive. Neither was NOS-IR found to be present in neurones immunoreactive for NPY, even though the two types of cells were often co-distributed within the same brain stem section. However, numerous NPY-IR terminals were found making close appositions with NOS neurones in the ventrolateral medulla. (d) Amino acids and acetylcholine. Neurones double-labelled for both NOS activity and glutamate-immunoreactivity were observed in the ventrolateral region throughout the extent of the cat medulla oblongata (Maqbool et al., 1995). These cells were scattered in the lateral tegmental field (9% of glutamate neurones), nucleus ambiguus (7%), lateral reticular nucleus (49%) and gigantocellular tegmental field (10%). Double-labelled cells were also seen in the paramedian (23%), the external cuneate nucleus (62%) and within the infratrigeminal nucleus (30%). At the most rostral levels examined double-labelled cells were also found in the nucleus prepositus hypoglossi (12%) and occasionally in the NTS (1%). A slightly 193

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Fig. 10. Relationship between NOS-IR neurones and catecholaminergic (TH-IR) neurones in the dorsomedial medulla (A2 cell group) and RVLM (C1 group) shown by double labelling. (A) NOS-IR neurones in the medial NTS at an intermediate level of the nucleus (Cy3 labelling). (B) Same area. showing TH-IR neurones labelled by Cy2, several of which (longer arrows) are dual-labelled. Other neurones in this cell group are single-labelled for either TH or NOS (larger arrowhead). (C. D) As A and B. but at a central level of the NTS. (C) Clearly dual-labelled neurones (arrows) are far less numerous here. (D) NOS-IR neurones in the RVLM (Cy3). (E) Corresponding area, showing TH-IR neurones of the C I group labelled with Cy2. These are clearly separate from the NOS-IR neurones, which lie in a more ventromedial position than the TH-IR neurones.

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Fig. 11. Relationship between NOS-IR neurones and 5-HT-IR neurones shown by double labelling. (A) Group of 5-HT-IR neurones and terminals in the ROb (Cy3 labelling), tB) Same area labelled for NOS-IR with (~y2. One neurone (longer arrow) is clearly dual-labelled, while others are only immunopositive for 5-HT or NOS (larger arrowheads). (C) Group of 5-HT-IR neurones in the superticial ventral medulla, immediately medial to a rootlet of the XII nerve (Cy3). (D) Corresponding area labelled for NOS-IR with Cy2. Large multipolar cells with extensive dendritic arborisations are located both medially and laterally to the nerve rootlet. Note that the two groups of labelled cells shown in panels C and D are clearly separate populations.

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different pattern of co-existence was observed in the rat (Fig. 12). In the NTS, particularly in the commissural, medial, ventral and ventrolateral subnuclei (Fig. 12), NOS-IR cells were also found to be moderately glutamate-IR. In certain reticular areas, for example the PMn, IRt, dorsal PCRt, Prb and Ro (Fig. 12), clearly dual-labelled cells were quite numerous. Occasional examples of coexistence were also seen in the DMV, DMSp5, MeV, Pr and Sp5C. Despite the evidence for the coexistence of GABA in nitrergic neurones of the dorsal horn and forebrain, and for the coexistence of acetylcholine in nitrergic cells of the spinal cord and pontine reticular nuclei (see above), we have not been able to gather any conclusive proof from dual-labelling studies of such a coexistence in any nuclei of the medulla. (e) Calcium-binding proteins. Neurones immunoreactive for NOS and for calbindin were found to have a broadly similar distribution throughout the rat medulla, and were also similar in their morphology (Fig. 13). However, coexistence of these two markers was found to be restricted to certain areas (Fig. 13). In the sensory nuclei of the medulla calbindin-IR neurones far outnumbered NOS-IR neurones, and coexistence was either absent or rarely observed. The two populations of neurones were entirely separate in the dorsal column nuclei (Gr, Cu and ECu) and in the AP. Few cells were dual-labelled in the NTS, even in the central and rostral zones, where numerous small ovoid or triangular cells immunoreactive for the two markers were coextensive and intermingled (Fig. 13). In the spinal trigeminal nuclei, coexistence was restricted to a few cells in the dorsal superficial zone of the pars caudalis, and to a larger population of cells in the DMSp5 at rostral levels. In the reticular formation no coexistence was seen in the ventral, parvocellular, paramedian or lateral paragigantocellular fields, or in the NA or the LRt. Only a few scattered neurones displayed coexistence in the MdD or Gi. However, many of the large neurones forming the prominent group in the IRt were immunoreactive for both NOS and calbindin at levels both caudal and rostral to obex. Labelling for calretinin or parvalbumin immunoreactivities was found overlapping populations of small, densely packed ovoid neurones throughout the NTS, and were particularly numerous at rostral levels (Fig. 13). Dual labelling demonstrated that the cells containing these calcium-binding proteins were usually different from the nitrergic neurones, with which they were intimately intermingled within the NTS. A few examples of weakly dual-labelled calretinin-NOS neurones were, however, observed in some brains (Fig. 13). From the above evidence, it appears, therefore, that nitric oxide production in the medulla is not confined exclusively to neurones of one particular transmitter phenotype or function. Although high levels of coexistence of NOS-IR occur in certain neurone populations that are presumed excitatory by virtue of their labelling for glutamate, groups of nitrergic neurones in other areas label for SS and calbindin, more usually associated with inhibitory neurones.

Fig. 12. Relationships between NOS-IR neurones and neurones containing somatostatin (SS) and glutamate (Glu) immunoreactivities in different regions of the medulla shown by double labelling. (A) Somatostatin-IR neurones in the dorsomedial area of the reticular formation (Cy2 labelling). (B) Same area showing NOS-IR neurones (Cy3). Two neurones are clearly dual-labelled (arrows), while at least one other cell is labelled for NOS but not somatostatin (larger arrowhead). (C) Somatostatin-IR neurones and fibres in the medial NTS (Cy2). (D) Same area, showing that there is no coexistence of somatostatin-IR and NOS-IR in this area of the NTS. The larger, multipolar NOS-IR neurones are more ventrally located than the smaller, ovoid somatostatin-IR cells. (E) Glutamate-IR neurones lying immediately ventral to the XII motor nucleus, within the nucleus of Roller (Ro), labelled with Cy2. (F) Same area, showing that these neurones also contain NOS immunoreactivity (arrows), visualised by Cy3 labelling. However, other neurones in this nucleus are NOS-IR (larger arrowhead), but have undetectable levels of glutamate-IR. (G) Clearly glutamate-IR neurones (arrows) in the ventral INTS at intermediate level of this nucleus (Cy2). (H) The same neurones (arrow) also show intense NOS-IR (Cy3). 197

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Fig. 13. Relationship between NOS-IR neurones and neurones containing calcium-binding proteins shown by double labelling. (A) Group of NOS-IR neurones in the dorsal NTS at an intermediate level (Cy3 labelling). (B) Same area labelled for calbindin-IR with Cy2. Only a few dual-labelled neurones can be recognised. (C) NOS-IR neurones and fibres in the DMSp5 (Cy3). (DI Same area showing labelling for calbindin-IR with Cy2. Many neurones are clearly dual-labelled (longer arrow), but others are only labelled for calbindin (larger arrowhead). (E) Large group of weakly NOS-IR neurones in the medial NTS at a rostral level (AMCA labelling). (F) Same area labelled for calretinin-IR with Cy3. Only a few neurones are clearly identified as being dual-labelled for both markers (arrow).

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It is clear, however, that the primary neurotransmitters co-existing with NOS in many of the nitrergic cell groups of the medulla (most notably the NTS) remain to be established. These neurones may contain peptides such as enkephalins that are difficult to visualise in somata and dendrites by immunohistochemistry, and it may be that methods aimed at defining the transmitters present in the terminals of these neurones after intracellular filling with markers such as biocytin may prove to be a more fruitful approach to resolving this question. 3.2. ROLE OF NITRIC OXIDE IN MEDULLARY PATHWAYS INVOLVED IN AUTONOMIC FUNCTIONS Considerable evidence exists implicating NO as a modulator of various central control pathways in the medulla, including those regulating the cardiovascular, oesophageal, gastrointestinal, respiratory and (anti)nociceptive systems. 3.2.1. NO in the control of the cardiovascular system The centrally mediated baroreceptor reflex provides a rapid negative feedback mechanism that dampens fluctuations in cardiovascular parameters. A rise in blood pressure increases the firing of baroreceptors in the carotid sinus and aortic arch. Sensory information from these receptors is carried to the NTS via the aortic and carotid sinus nerves (components of the vagus and glossopharyngeal nerves, respectively). This rise in baroreceptor firing causes an increase in the discharge of first-order neurones that receive synaptic inputs from the baroreceptor nerve fibres. Outputs from the NTS, via neuronal pathways that are still only partly resolved, modulate the activity of neurones in other medullary nuclei that results in compensatory adjustments of parasympathetic and sympathetic outflow, from efferent neurones located in the vagal motor nuclei and the RVLM, respectively. Ultimately, an increase in blood pressure results in an increased vagal tone and a decrease in sympathetic outflow to the heart and blood vessels, resulting in a decrease in blood pressure. Anatomical studies, using either NOS immunohistochemistry or NADPH-d histochemistry, have demonstrated nitrergic neurones in regions of the medulla implicated in central cardiovascular control, e.g. the NTS (Figs. 1-5), the RVLM (see Figs. 1-4); the raphe nuclei (Fig. 14), the DVN (Figs. 7 and 8) and the nucleus ambiguus (Fig. 8), which contains the majority of the vagal preganglionic neurones projecting to the heart. Of these regions, the NTS and the RVLM are those most likely to contain nitrergic neurones involved in cardiovascular control, since we have been unable to demonstrate the presence of NOS within cardiac vagal preganglionic neurones retrogradely labelled by application of tracers to the heart (E. Corbett et al., unpubl, results). (a) The nucleus tractus solitarius. Nitrergic neurones have been localised in several subnuclei of the NTS which may have roles in central cardiovascular control NTS (Ruggerio et al., 1996; Lawrence et al., 1998; Fig. 6). At least some of these neurones may be baroreceptive, since a substantial proportion of neurones in the dorsal NTS that expressed c-fos in response to phenylephrine-induced hypertension also stained positive for NADPH-d and NOS mRNA (Chan and Sawchenko, 1998). This region of the NTS is known to contain some terminals from aortic depressor nerve afferent fibres (Wallach and Loewy, 1980; Ciriello and Calaresu, 1981; Youfsi-Malki and Puizillout, 1995). However, since it is impossible to determine if these c-fos immunoreactive cells are mono- or polysynaptically activated by baroreceptor stimulation, their exact role in cardiovascular control remains uncertain. Nevertheless, we 199

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provide some support for this notion by illustrating that NOS-immunoreactive neurones in the NTS receive synaptic input from vagal afferents (Figs. 6, 15 and 16). Experiments in vitro indicate clearly that NO can have cellular actions in the NTS. Single-unit extracellular recordings from rat brain stem slices have shown that perfusion of the slices with L-arginine increases the neuronal activity of NTS neurones (Tagawa et al., 1994). Perfusion of the same slices with L-NMMA (a NOS inhibitor) or haemoglobin (a NO trapper) blocks the increase in neuronal activity evoked by L-arginine, as does the guanylate cyclase inhibitor methylene blue (Tagawa et al., 1994). Since haemoglobin does not penetrate into neurones (Stewart et al., 1987; Ignarro, 1991), its action suggests that NO diffuses out of the neurones in which it is formed and penetrates the adjacent neurones to increase their activity through an elevation in cGMP levels. This is consistent with good correlation between cGMP and NOS immunostaining in many areas of the brain including the NTS (De Vente et al., 1998). Evidence for a physiological function of NO in the baroreceptor reflex pathway has come from administering NO-like substances and antagonists to the NTS in vivo. Microinjection of S-nitrocysteine (a NO-like substance) into the NTS of anaesthetised (Lewis et al., 1991; Tseng et al., 1996) and conscious (Machado and Bongama, 1992) rats resulted in an immediate bradycardia and hypotension. Conversely, L-NMMA (a NOS inhibitor) microinjected into the NTS of rats has been shown to attenuate the depressor effect evoked with glutamate microinjection into the NTS (Paola et al., 1991), and to elicit an increase in arterial pressure and heart rate (Harada et al., 1993). The bradycardia and hypotension caused by injection of NO-like substances into the NTS can be blocked by inhibition of soluble guanylate cyclase (methylene blue), suggesting that NO mediates the control of blood pressure and heart rate via its actions on guanylate cyclase (Lewis et al., 1991). (b) The ventrolateral medulla. The rostral portion of the ventrolateral medulla (RVLM) contains sympathoexcitatory neurones (Barman and Gebber, 1985; Dampney et al., 1985; Ciriello et al., 1986) which provide a tonic sympathetic drive to blood vessels. Injection of L-arginine (a precursor of NO) or NO donors into the RVLM of anaesthetised rats (Tseng et al., 1996; Kagiyama et al., 1997), cats (Shapoval et al., 1991; Zanzinger et al., 1995) and rabbits (Kagiyama et al., 1998) elicits a depressor effect. This action is blocked by inhibiting soluble guanylate cyclase with methylene blue (Shapoval et al., 1991; Kagiyama et al., 1998). Similar injections of NOS inhibitors (L-NMMA; L-NAME) into the RVLM elicit a pressor response and an increase in blood pressure (in rat: Tseng et al., 1996; Kagiyama et al., 1997; in cat: Shapoval et al., 1991; in rabbit: Kagiyama et al., 1998). Injection of the NOS inhibitor L-NMMA into the RVLM prior to injection of L-arginine also attenuates the cardiovascular effects of L-arginine (Tseng et al., 1996). In contrast, other studies have shown that microinjection of NO donors into the pressor region of the RVLM of anaesthetised rabbits (Hirooka et al., 1996) or the RVLM of freely moving rats (Martins-Pinge et al., 1997) elicits a pressor effect. In both of these studies this effect was attenuated by prior injection of methylene blue (a guanylate cyclase inhibitor) into the RVLM (Hirooka et al., 1996; Martins-Pinge et al., 1997). Similar injections of the NOS inhibitor (L-NAME) in anaesthetised rabbits caused a depressor effect (Hirooka et al., 1996). These studies suggest that NO has a pressor and sympathoexcitatory action in the RVLM. These contrasting results may be due to variations in the methods used. All of the studies used different volumes and doses of NO donors and three different animal species were used (cat: Shapoval et al., 1991; Zanzinger et al., 1995; rat: Liu et al., 1996; Tseng et al., 1996; Kagiyama et al., 1997; Martins-Pinge et al., 1997; rabbit: Hirooka et al., 1996). Studies were 200

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Fig. 14. Raphe obscurus nucleus (ROb) at an intermediate level of the medulla showing the presence of NOS immunoreactivity in neurones projecting to NTS. (A) Scattered cell bodies in the ROb labelled by injection of CTb into the commissural NTS (Cy2 labelling). (B) Same area showing labelling for NOS-IR with Cy3. Some neurones are clearly dual-labelled (longer arrow), whereas other cells containing CTb are not NOS-IR (larger arrowhead). (C, D) Higher magnifications of areas boxed in A and B, respectively. Again, two cells are clearly dual-labelled (longer arrows), whereas other cells are only single-labelled for CTb or NOS-IR (larger arrowheads).

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Fig. 15. Vagal afferent fibres closely appose NOS-IR neurones in the NTS. (A) Low power view of the NTS on one side of the medulla oblongata. A dense network of vagal afferent fibres, labelled by the injection of BDA into the nodose ganglion, can be detected in the NTS. Some retrogradely labelled neurones are visible in the dorsal vagal nucleus (DVN). The boxed area lateral to the capillary (Cap) is illustrated at higher magnification in B. (B) The capillary (Cap) is the same as that indicated in A. The boxed area contains a NOS-IR neurone and is shown at higher magnification in C. ((7) The NOS-IR neurone, identified by silver intensified gold particles, is closely apposed by DAB-labelled vagal afferent tei-minals.

p e r f o r m e d on both a n a e s t h e t i s e d and c o n s c i o u s a n i m a l s , and differences m a y be due to the level and type of a n a e s t h e s i a . T h e r e is also the possibility that the injections w e r e m a d e into different areas o f the R V L M . O n l y one study injected N O d o n o r s into a region f u n c t i o n a l l y identified as the p r e s s o r region of the R V L M ( H i r o o k a et al., 1996); no others m a d e this distinction. For d i s c u s s i o n see H i r o o k a et al. (1996). 202

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Fig. 16. Electron microscopic verification that vagal afferent synaptic terminals form synaptic contacts with somata and dendrites of NOS-IR neurones in the NTS. IA) A NOS-IR neurone in the NTS identified by the silver intensified gold particles (broken arrows) is apposed by a vagal afferent terminal containing electron dense DAB reaction product (boxed area). (B) Higher magnification of the boxed area in A. A synaptic specialisation (two arrows) occurs between the labelled vagal afferent bouton and the NOS-IR cell (silver indicated by the broken arrow). (C) A large-calibre dendrite of a silver containing NOS-IR neurone is apposed by two electron dense vagal afferent terminals (arrows), illustrated at higher magnification in D and E. (D) The labelled vagal afferent terminal forms a synaptic contact (arrows) with the silver (broken arrow) containing NOS-IR dendrite. (E) The second terminal also forms a synaptic contact (arrows) with the NOS-IR dendrite (broken arrowl.

N O released in the R V L M m a y arise from nitrergic cells located in this region (Figs. 1 - 4 and 10), or from the axon terminals (or possibly dendrites) of neurones located in other areas such as the raphe nuclei or NTS. One possible source is the group of N O S m R N A and N A D P H - d - p o s i t i v e neurones in the d N T S , which were shown to express c-fos in response to p h e n y l e p h r i n e - i n d u c e d hypertension, and which were also retrogradely labelled by tracers injected into the R V L M (Chart and S a w c h e n k o , 1998). This suggests that there is a direct route involving N O neurones, by which the b a r o r e c e p t o r inhibition of s y m p a t h e t i c outflow m a y be affected. 203

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3.2.2. NO in neuronal circuitry underlying control of the oesophagus Reflex central control of the oesophagus may be conducted via a disynaptic pathway. Sensory vagal neurones carry information from the oesophagus to a discrete part of the NTS, the central subnucleus (ceNTS; Fryscak et al., 1984; Altschuler et al., 1989). Vagal afferents form synaptic contacts with NOS-immunoreactive NTS neurones near this region (Figs. 15 and 16). Neurones in the ceNTS emit a dense and topographically discrete projection to the compact formation of the nucleus ambiguus (NAc), where terminals surround the oesophageal motorneurones that project back to the oesophagus (Cunningham and Sawchenko, 1989). NO may be involved at every step in this pathway. The ceNTS is also rich in nitrergic neurones. Histochemical localisation of NADPH-d has revealed a cluster of positive neurones in the central subnucleus in a number of species (rat, dog, guinea pig, ferret and cynomolgus monkey: Bieger and Sharkey, 1993; humans: Gai and Blessing, 1996; rat: Vincent and Kimura, 1992: cat: Maqbool et al., 1995). NOS-IR has also been found in the neurones of the ceNTS/see Figs. 5 and 6: Gai et al., 1995; Maqbool et al., 1995). These cells appear to receive information from oesophageal afferents since nitrergic cells in the ceNTS are innervated by afferents labelled from the oesophagus (rat: Wiedner et al., 1995; rabbit: Gai et al., 1995). The NOS neurones in the ceNTS act as oesophageal premotor neurones and project to oesophageal motor neurones in the NAc (Cunningham and Sawchenko, 1989). NOS terminals in the NAc were found to be derived from ceNTS neurones as NTS lesions caused a decrease in the density of the NOS terminals (Gai et al., 1995). Furthermore, terminals in the NAc labelled anterogradely from the ceNTS surround motorneurones and show NADPH and NOS-IR (rat: Wiedner et al., 1995; rabbit: Gai et al., 1995). The nitrergic neurones in the ceNTS therefore act as interneurones in a disynaptic pathway connecting afferent and efferent neurones controlling oesophageal peristaltic activity. By releasing NO they are thought to excite the oesophageal motorneurones of the NAc, the axons of which terminate in nicotinic cholinergic neuromuscular junctions on the oesophageal muscle fibres, and may thus potentiate nicotinic synaptic transmission (O'Sullivan and Burgoyne, 1990; Anderson et al., 1993). This prevalence of NOS in the neuronal circuitry involved with control of the oesophagus is consistent with a prominent role for NO as a neurotransmitter involved in oesophageal peristalsis.

3.2.3. NO in CNS control of the stomach and large intestine It is well known that NO is a nonadrenergic noncholinergic (NANC) neurotransmitter in the vagal nerve fibres that mediates the relaxation of the GI tract (Desai et al., 1991: Lefebvre et al., 1992). However, the role of NO in the central control of this system is less well described, although the location of NOS and NADPH-d-positive cells suggests a physiological role for NO in the central regulation of gastrointestinal activity. Vagal afferent fibres from the stomach enter the NTS at intermediate levels and project sparsely to the medial and commissural nuclei, and densely to the subnucleus gelatinosus (sgNTS; Shapiro and Miselis, 1985). Afferents terminating in the sgNTS have been shown to synapse directly onto the dendrites of vagal gastric efferent neurones situated in the nearby DVN and projecting back to the stomach (Rinaman et al., 1989). NO may play a role here, as NADPH-d histochemistry has visualised a relatively dense network of fibres and terminal in the sgNTS (e.g. Krowicki et al., 1997). In addition, vagal afferents closely appose the somata of NOS-immunoreactive neurones in the dorsal vagal nucleus (Fig. 9). 204

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As discussed earlier (Section 3.1.3), NOS has been localised in vagal preganglionic neurones in the dorsal vagal nucleus (see also Figs. 7 and 8). Retrograde labelling studies have shown that the DVN provides the major source of efferent innervation to the stomach (Norman et al., 1985; Shapiro and Miselis, 1985), and it has been estimated that 95% of preganglionic DVN neurones innervate the stomach specifically (Leslie et al., 1982). There is also a lesser innervation by neurones in the DVN of the small intestine (Shapiro and Miselis, 1985; Zhang et al., 1991), large intestine (Berthoud et al., 1991) and pancreas (Rinaman and Miselis, 1987). Recently, Zheng et al. (1999) studied specific areas of the DVN thought to mediate particular reflexes such as the adaptive response of the stomach after ingesting a meal (i.e. the capacity of the stomach to receive large volumes with only a slight increase in pressure). They combined injection of retrograde tracers into the gastric fundus of rats and labelled a discrete neuronal population of NOS-immunoreactive neurones in the medial portion. At a cellular level it is clear that NO can excite DVN neurones (Travagli and Gillis, 1994). Perfusion of the rat DVN in vitro with L-arginine (the NO precursor) or S-nitrosoN-acetylpenicillamine (SNAP, a NO donor) has been shown to increase the spontaneous firing rate of DVN motoneurones, an excitatory effect that can be counteracted by the NOS antagonist N-nitro-L-arginine (L-NNA). Perfusion of the cGMP analogue dibutyryl cGMP onto the DVN also increased the spontaneous firing rate of the DVN neurones, and perfusion with LY88583 (an inhibitor of guanylate cyclase) counteracted the excitatory effect of L-arginine. These results suggest that the excitatory effect of NO on DVN neurones is due to the accumulation of cGMP (Travagli and Gillis, 1994). As DVN cells are known to mediate gastrointestinal activity and be excited by NO, much research has been carried out on the effect of NO agonists and antagonists on the GI system. In vivo there is a rostrocaudal separation of function in the DVN. The excitation of rostral DVN neurones by L-glutamate increased gastric motility and lower oesophageal pressure, whereas a similar excitation of caudal DVN neurones resulted in gastric relaxation and a decrease in lower oesophageal pressure (Rossiter et al., 1990). In keeping with an excitatory effect of NO on DVN neurones, in vivo microinjections of L-arginine into the rostral DVN of cats increased gastric motility, and injection of SNAP increased antral activity (Panico et al., 1995). This effect was mediated by NOS since the NOS inhibitor N-nitro-L-arginine-methyl-ester (L-NAME) prevented L-arginine from exerting an effect (Panico et al., 1995). In contrast, microinjection of NO-liberating compounds into the region of the sgNTS decreased intragastric pressure and NOS inhibitors increased intragastric pressure (Krowicki et al., 1997). These latter authors attribute the discrepancy between the effect of NTS and DVN injections to the different locations of the injection sites. They suggested that the functionally more relevant changes are as a result of microinjections into the sgNTS since this is the area where vagal afferents terminate on the dendrites of DVN neurones. Since the NOS antagonist L-NAME inhibits NTS neuronal excitability (Ma et al., 1995), this suggests that the NTS neurones activated by NOS may either be inhibitory neurones, or may project to the rostral DVN, where stimulation evokes relaxation effects. However, since vagal afferents closely appose the soma of neurones in the DVN (Fig. 9), this raises some questions about where the functionally most relevant sites of action might be, as synapses onto somata may have a greater influence on neurones than those on distal dendrites.

3.2.4. NO in central respiratory control To date the most likely region of the medulla oblongata in which NO appears to be involved in central control of respiration is the NTS. The caudal NTS is involved in the control 205

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of respiration since it receives input from the carotid sinus nerve (Housley et al., 1987), which carries afferent information from peripheral chemoreceptors (Donoghue et al., 1984; Housley and Sinclair, 1988). Stimulation of the peripheral chemoreceptors by hypoxia releases glutamate from the central synaptic terminals of the carotid sinus nerve and a reflex increase in ventilation occurs to correct the hypoxia and maintain body homeostasis (Neubauer et al., 1990). Injection of a NO donor into the NTS enhances the ventilatory response to hypoxia by increasing glutamate release in the NTS, probably from the presynaptic terminals of the chemoreceptor afferents (Ogawa et al., 1995). The formation of NO in response to hypoxia is blocked by an NMDA antagonist, suggesting that NO is produced in NTS neurones. However, it cannot be excluded that NO is released from the chemoreceptor terminals themselves since vagal afferent fibres have recently been shown to contain NMDA receptor immunoreactivity (Aicher et al., 1999), although it is uncertain that NOS is in vagal afferent fibres (see earlier). A role for the nitrergic neurones of the ventral medulla in chemoreception and in the ventilatory responses to cerebral hypoxia has been proposed (Iadecola et al., 1993). The NOS-IR neurones in the RVLM are completely separate from the catecholaminergic C1 cell group (Iadecola et al., 1993; Ohta et al., 1993; Fig. 10 of Maqbool et al., 1995), being located more medially and ventrally, approaching the surface of the medulla. Thus they are ideally positioned to act as central chemoreceptors, although firm evidence in support of this hypothesis is still lacking.

3.2.5. NO in pathways coordinating autonomic and nociceptive responses It has been known for many years that there are mutual interactions between the central mechanisms controlling autonomic, particularly cardiovascular responses and those promoting nociception or anti-nociception, which may involve reciprocal neural connections between the spinal cord and the medulla (Randich and Maixner, 1986; Randich and Gebhart, 1992; Wilson and Hand, 1997). Thus, peripheral somatosensory and viscerosensory inputs can modify autonomic reflex responses, and vagally mediated afferent inputs can disengage nociceptive responses. These interactions might occur both at the level of the spinal cord and within the brain, either in the medulla or in higher centres, as the effects of stimulating several brain sites, including the RVLM and raphe nuclei, suggest that they have a role in coordinating cardiovascular and nociceptive responses (Lovick, 1991). Inhibitory responses elicited in RVLM neurones appeared to be mediated through a GABAergic input, and might involve a more indirect spinal projection via another area of the medulla such as the NTS (Sun and Spyer, 1991). Electrical stimulation of vagal afferent fibres, particularly those of the cardiopulmonary branches, inhibited nociceptive responses, and the prime relay site in the medulla oblongata concerned with this inhibition was the NTS. Application of local anaesthetics to the NTS abolished the anti-nociceptive effects of vagal stimulation, whereas glutamate injection or electrical stimulation in the NTS strongly inhibited spinal nociceptive responses (Randich and Maixner, 1986; Randich and Gebhart, 1992), although other brain stem areas, including the locus coeruleus, raphe magnus, RVLM and adjacent reticular formation were also shown to be involved. NTS neurones mediating nociceptive inhibition may do so through indirect descending connections to the spinal cord via one or all of these three noradrenergic or serotoninergic brain stem nuclei which are also known to contain opiate neurones (Basbaum and Fields, 1984). However, direct connections from the NTS to the dorsal horn have been reported (Kuypers and Maisky, 1975; Loewy and Burton, 1978; Basbaum and Fields, 1979), and the participation of such projections in vagal afferent modulation of nociception warrants further investigation. It is interesting that nitrergic neurones are present 206

NO systems in the medulla oblongata and in atttonomic control

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in all the areas of the medulla that have been shown to participate in these responses (Sun and Spyer, 1991), and it has been proposed that NO plays an important role in central nociceptive function (Haley et al., 1992; Schuman and Madison, 1994; Anderson, 1998). In this regard, it has recently been demonstrated that visceral hyperalgaesia is associated with an increase in the numbers of cells in the RVLM labelled tbr NOS or NADPH-d, and furthermore, that injection of L-NAME into this region attenuated the hyperalgaesic response (Coutinho et al., 1998). Although the exact nature of the pathways within the medulla involved in these responses are still poorly understood, there is now evidence that at least two areas of the CNS known to integrate and relay peripheral inputs, i.e. the paratrigeminal nucleus (Armstrong and Hopkins, 1998) and lamina I of the dorsal horn (Esteves et al., 1999), contain NOS-IR neurones projecting to NTS.

4. Abbreviations AP A1 CC Cu DVN DMSp5 ECu Gi GiV Gr IO In InM IRt k-NAME L-NMMA LPGi LRt MdD MdV mlf MVe NA NADPH-d NAc NANC nNOs NO NOS NTS

Pa5

area postrema catecholamine cell area of the caudal ventrolateral medulla central canal cuneate nucleus dorsal motor vagal nucleus spinal trigeminal nucleus, dorsomedial division external cuneate nucleus gigantocellular field medial gigantocellular field gracile nucleus inferior olive nucleus intercalatus nucleus intermedius intermediate reticular nucleus NW-nitro-L-arginine methyl ester (a NOS inhibitor) N~-monomethyl-L-arginine (a NOS inhibitor) lateral paragigantocellular field lateral reticular nucleus dorsal medullary reticular nucleus ventral medullary reticular nucleus medial longitudinal fasciculus medial vestibular nucleus nucleus ambiguus nicotinamide adenine dinucleotide phosphate (reduced) diaphorase compact region of the nucleus ambiguus nonadrenergic nonchloinergic neuronal nitric oxide synthase nitric oxide nitric oxide synthase nucleus tractus solitarii (subnuclei: ce = central; co = commissural; d = dorsal; i = interstitial; me = medial; ts = tractus solitarius; v = ventral; vl --- ventrolateral) paratrigeminal nucleus 207

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PCRt PMn Pr Prb PY Ro ROb RVLM SNAP Sp5C Sp5CI SpVe TS VII XII

T.F.C. Batten et al.

parvocellular reticular field paramedian reticular area nucleus prepositus hypoglossi nucleus of Probst's bundle pyramidal tract nucleus of Roller raphe obscurus nucleus rostral ventrolateral medulla S-nitroso-N-acetylpenicillamine (a NO donor) spinal trigeminal nucleus, caudal division spinal trigeminal nucleus, interpolar division spinal vestibular nucleus tractus solitarius facial motor nucleus hypoglossal motor nucleus

5. ACKNOWLEDGEMENTS We are especially grateful to Dr. Piers Emson (Molecular Neuroscience Group, University of Cambridge) for providing the antiserum to recombinant rat nNOS. We wish to thank Brenda Frater and Jean Kaye for their invaluable technical assistance, and Drs. Azhar Maqbool, Filomena Esteves and Eric Corbett who contributed greatly to many of the experiments we report here. We are also grateful to the Medical Research Council for Project Grant and JREI Equipment Award funding to TFCB, and to the Wellcome Trust for a project grant to JD, as well as an Equipment Grant awarded to JD, TFCB and others.

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Vincent SR, Kimura H (1992): Histochemical mapping of nitric oxide synthase in the rat brain. Neuroscience 46:755-784. Vincent SR, Johansson O, H6kfelt T, Skirboll L, Elde RE Terenius L, Kimmel J, Goldstein M (1983a): NADPH-diaphorase: a selective histochemical marker for striatal neurones containing both somatostatin- and avian pancreatic polypeptide (AAP)-like immunoreactivities. J Comp Neuro! 217:252-263. Vincent SR, Satoh K, Armstrong DM, Fibiger HC (1983b): NADPH-diaphorase: a selective histochemical marker for the cholinergic neurons of the pontine reticular formation. Neurosci Lett 43:31-36. Wallach JH, Loewy AD (1980): Projections of the aortic nerve to the nucleus tractus solitarius in the rabbit. Brain Res I88:247-251. Wiedner EB, Bao X, Altschuler SM (1995): Localization of nitric oxide synthase in the brain stem neural circuit controlling esophageal peristalsis in rats. Gastroenterology 108:367-375. Wilson LB, Hand GA (1997): The pressor reflex evoked by static contraction: neurochemistry at the site of the first synapse. Brain Res Rev 23:196-209. Xu ZQ, H6kfelt T (1997): Expression of galanin and nitric oxide synthase in subpopulations of serotonin neurons of the rat dorsal raphe nucleus. J Chem Neuroanat 13:169-187. Yamada K, Emson P, H0kfelt T (1996): Immunohistochemical mapping of nitric oxide synthase in the rat hypothalamus and colocalization with neuropeptides. J Chem Neuroanat 10:295-316. Yan XX, Garey LJ (1997): Morphological diversity of nitric oxide synthesising neurons in mammalian cerebral cortex. Hirnforschung 38:165-172. Youfsi-Malki M, Puizillout JJ (1995): Study of brainstem projections of aortic baroreceptor afferents in the rabbit using transganglionic anterograde transport of choleragenoid-HRP. Prim Sens Neuron 1:143-155. Yu WH (1994): Nitric oxide synthase in motor neurons after axotomy. J Histochem Cvtochem 42:451-457. Zanzinger J, Czachurski J, Seller H (1995): Inhibition of basal and reflex-mediated sympathetic activity in the RVLM by nitric oxide. Am.J Phvsiol 268:R958-R962. Zhang X, Fogel R, Simpson P, Renehan WE (1991): The target specificity of the extrinsic innervation of the rat small intestine. J Auton Nerv Svst 32:53-62. Zheng ZL, Rogers RC, Travagli RA (1999): Selective gastric projections of nitric oxide synthase-containing vagal brainstem neurons. Neuroscience 90:685-694.

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CHAPTER VII

Nitric oxide in the peripheral autonomic nervous system H.M. YOUNG, C.R. ANDERSON AND J.B. FURNESS

1. INTRODUCTION 1.1. BRIEF HISTORY OF THE IDENTIFICATION OF NO AS A PERIPHERAL NEUROT RA N S M ITTER In the mid-1960s, following the availability of specific drugs to block adrenergic receptors or to block noradrenaline release, it was realised that some responses in peripheral tissues were not mediated by either of the then-known neurotransmitters of peripheral neurones, noradrenaline or acetylcholine (Burnstock et al., 1966). Shortly afterwards, Burnstock et al. (1970) proposed that ATP is an inhibitory neurotransmitter to gastrointestinal smooth muscle. Subsequently, the discovery of peptides with potent pharmacological activities in many peripheral nerves led to the demonstration that substances such as vasoactive intestinal peptide (VIP), neuropeptide Y and substance P can also act as neurotransmitters (see H6kfelt et al., 1980; Lundberg and H6kfelt, 1986: Fumess et al., 1989). However, there were a number of locations where inhibitory neurotransmission to vascular or non-vascular smooth muscle did not appear to be mediated by noradrenaline, acetylcholine, ATP or any known peptide. Furchgott and Zawadzki (1980) reported that the relaxant action of acetylcholine on in vitro preparations of rabbit aorta was mediated by a substance released from endothelial cells, which they termed EDRF (endothelium-derived relaxing factor). In 1987, using chemical assay and bioassay, it was shown by two different groups that EDRF is nitric oxide (NO) (Ignarro et al., 1987; Palmer et al., 1987). The similarities between EDRF and the unidentified inhibitory neurotransmitter in the rat anococcygeus muscle and the bovine retractor penis muscle was pointed out by Gillespie (1987) and Furchgott (1988). Using drugs that selectively block nitric oxide synthase (NOS), the enzyme responsible for the synthesis of NO, it was shown by a number of groups that NO is a major neurotransmitter mediating inhibitory transmission in the anococcygeus muscle of the rat (Gillespie et al., 1989; Li and Rand, 1989; Ramagopal and Leighton, 1989) and in the bovine retractor penis muscle (Liu et al., 1991). In fact, these experiments performed on peripheral neurones innervating the anococcygeus muscle were the first demonstration that NO acts as a neurotransmitter, in addition to its role as an endothelium-derived relaxing factor. Shortly after it was shown that NO acts as a neurotransmitter in the anococcygeus muscle, Bredt and Snyder (1990) demonstrated, using an antibody raised to NOS, that many neurones in the central and peripheral nervous systems are capable of producing NO.

Handbook of Chemical Neuroanatomv, Vol. 17." Functional Neuroanatomv of the Nitric Oxide System H.W.M. Steinbusch, J. De Vente and S.R. Vincent. editors (~) 2000 Elsevier Science B.V. All rights reserved.

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1.2. GENERAL PROPERTIES OF NO-MEDIATED NEUROTRANSMISSION The general properties of NO as a neurotransmitter, that have been reviewed in detail elsewhere (Bredt and Snyder, 1992, 1994; Rand, 1992; Sanders and Ward, 1992; Snyder, 1992; Stark and Szurszewski, 1992; Rand and Li, 1995), are summarised below. NO (and L-citrulline) are synthesised from L-arginine by NOS. There are three main isoforms of NOS neuronal, endothelial and macrophage NOS - - which are encoded by different genes and have different molecular weights. Both endothelial and neuronal NOS are dependent on Ca 2+, calmodulin and NADPH. Neuronal NOS is a 160 kDa cytoplasmic protein that is found as a homodimer (see Marietta, 1993). There is no evidence (or expectation) that stores of pre-formed NO exist in neurones, and it appears that NO is formed, by NOS, on demand and immediately released. The synthesis of NO can be inhibited by a number of arginine analogues, which are taken up by cells and competitively inhibit the synthesis of NO (see Moncada et al., 1991); these NOS inhibitors have been critical for determining whether NO plays a physiological role in different tissues. Since the formation of NO is Ca2+-dependent, it is likely that the increase in intracellular free Ca 2+ that occurs following the arrival of an action potential in a nerve terminal results in the activation of NOS and the consequent release of NO. Although other possible molecular targets are known, the major receptor for NO is soluble guanylyl cyclase, which unlike receptors for other neurotransmitters, is present within the cytoplasm, not at the cell membrane of the effector cell. Activation of soluble guanylyl cyclase results in an increase in the production of cGME which in turn activates cGMP-dependent kinases; increases in cGMP might also have other effects such as direct actions on ion channels. Cells containing NOS can be localised using immunohistochemistry, or using a histochemical reaction called the NADPH diaphorase reaction (Hope et al., 1991; Dawson et al., 1991; see Vincent, Chapter II of this volume). In the peripheral nervous system, there have been relatively few studies that have examined the equivalence of NOS immunoreactivity and NADPH diaphorase staining. However, those studies that have been performed have all shown that NOS immunoreactivity and NADPH diaphorase staining show identical locations in peripheral neurones. Consequently, throughout this review we have often not specified whether the presence of NOS was demonstrated immunohistochemically or histochemically. The only ambiguity that can arise is in the staining of nerve cell bodies. All cells contain NADPH-dependent enzymes, other than NOS, and thus if the NADPH diaphorase reaction is left to run for a long time, all cells will show some staining. It is therefore possible that there could be some false-positive reports of NOS in nerve cell bodies, based only on the presence of NADPH diaphorase staining. 1.3. SCOPE OF THIS REVIEW This review is not an exhaustive survey of every report of NOS in different regions of the peripheral autonomic nervous system of different mammalian species. We have concentrated on locations where anatomical studies showing the presence of NOS neurones have been backed up by physiological and pharmacological studies that have examined the function of neurally released NO. Moreover, the presence and role of NO neurones innervating the pelvic organs is only covered briefly (in Section 3.1.1 (c)), but is dealt with in detail in Chapter VIII of this volume.

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2. NO IN A U T O N O M I C G A N G L I A 2.1. NITRIC OXIDE AND SYMPATHETIC PRE- AND P O S T G A N G L I O N I C NEURONES

2.1.1. Presence of NOS in sympathetic pre- and postganglionic neurones The cell bodies of sympathetic preganglionic neurones are found in the spinal cord, usually between the first thoracic segment (T1) and, depending on the species, the upper to mid-lumbar segments. The axons of preganglionic neurones project in the ventral roots to innervate the cell bodies of postganglionic neurones in the paravertebral and prevertebral ganglia, and chromaffin cells in the adrenal medulla. Most sympathetic preganglionic neurones are located in nuclei on the lateral margins of the intermediate grey matter of the spinal cord, called the intermediolateral nuclei (Petras and Cummings, 1972). Synaptic transmission from sympathetic preganglionic neurones to postganglionic neurones is mediated primarily by acetylcholine, and although sympathetic preganglionic neurones contain many other putative neurotransmitters (Morris and Gibbins, 1992), non-cholinergic components of transmission are not easily demonstrated. NOS is present in many sympathetic preganglionic neurones (Fig. 1). Using NADPH diaphorase staining, NOS was first identified in rat sympathetic preganglionic neurones in 1992 (Anderson, 1992; Blottner and Baumgarten, 1992; Dun et al., 1992; Valtschanoff et al., 1992). An earlier study by Thomas and Pearse (1961) had reported intense NADPH diaphorase activity in the intermediolateral column of the spinal cord, but since the enzyme underlying the reaction was unidentified at the time, the significance of the staining was not recognised. Later, NADPH diaphorase-stained axons were found in the ventral roots of rats, but the axons were not definitely identified as arising from preganglionic neurones (Aimi et al., 1991). Since 1992, many studies have confirmed the presence of NOS in sympathetic preganglionic neurones of a wide range of species using immunohistochemistry and/or NADPH diaphorase staining (see Table 1), and the two techniques appear to show the same population of fibres and cell bodies (Anderson et al., 1993; Furness and Anderson, 1994). NOS is present in many, but not all, sympathetic preganglionic neurones in the species that have been examined in detail. Most is known about the distribution of NOS in rat sympathetic preganglionic neurones. In the upper thoracic segments in the rat, NOS is present in around

TABLE 1. Species in which NOS-containing syml~athetic preganglionic neurones have been demonstrated Species

References

Rat

Anderson (1992); Blottner and Baumgarten (1992): Dun et al. (1992. 1993a,b). Valtschanoff et al. (1992); Morris et al. (1993): Ceccatelli et al. (1994): Saito et al. (1994); Domoto et al. (1995); Okamura et al. (1995): Ando et al. (1996): Soinila et al. (1996); Chiba and Tanaka (1998); Tang et al. (1998) Brtining (1992); Dun et al. (1993b) Anderson et al. (1994); Furness et al. (1994) Hakim et al. (1995) Dun et al. (1993b) Hisa et al. (1995); Vizzard et al. (1997) Xu et al. (1996) Marley et al. (1995) Dun et al. (1993b) Smithson and Benarroch (1996)

Mouse Guinea pig Rabbit Cat Dog Sheep Cow Monkey Human

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Fig. 1. (A) Horizontal (longitudinal) section through the T8 thoracic segment of a rat stained for the NADPH diaphorase reaction. A compact mass of NADPH diaphorase-stained sympathetic preganglionic neurones is present in the intermediolateral nucleus (IML) on the margin between the grey matter (G) and white matter (W). The neurones extend dendrites along the IML as well as across the grey matter towards the intermediate grey above the central canal. The positively stained neurones in the midline are not sympathetic preganglionic neurones. Scale bar -- 100 Ixm. (B) Sympathetic preganglionic neurones in the intermediolateral nucleus of the TI2 spinal cord segment. The reaction product is homogeneous, not granular and is found throughout the neurone, including into the finest dendrites, but is not present within the nucleus. Scale bar = 25 l.tm. (C) Fluorescence micrograph of rat sympathetic preganglionic neurones in the intermediolateral nucleus tlML)of the TI3 spinal cord segment stained with an antiserum to NOS. The appearance is identical to that seen with the NADPH diaphorase reaction. W = white matter; G = grey matter. Scale bar = 100 It m.

85% o f all p r e g a n g l i o n i c n e u r o n e s . In m i d d l e to low thoracic s e g m e n t s , N O S is f o u n d in 6 5 - 7 5 % o f all p r e g a n g l i o n i c n e u r o n e s , and in a slightly h i g h e r p r o p o r t i o n ( 7 0 - 8 5 % ) in low thoracic and u p p e r l u m b a r levels ( A n d e r s o n ,

1992; G r k o v i c and A n d e r s o n ,

1997). T h e r e

are d i f f e r e n c e s in the p r o p o r t i o n s o f p r e g a n g l i o n i c n e u r o n e s c o n t a i n i n g N O S m e d i o l a t e r a l l y across the i n t e r m e d i o l a t e r a l c o l u m n . N O S - c o n t a i n i n g n e u r o n e s are m o r e c o m m o n laterally in the i n t e r m e d i o l a t e r a l c o l u m n than medially, in both the rat ( A n d e r s o n ,

1992) and the

g u i n e a pig ( F u r n e s s and A n d e r s o n , 1994). S y m p a t h e t i c p r e g a n g l i o n i c n e u r o n e s in the spinal 218

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autonomic nuclei innervate postganglionic neurones in sympathetic ganglia throughout the body. Successively more caudal sympathetic ganglia receive preganglionic inputs from successively more caudal spinal segments (see Strack et al., 1988). Experiments in which retrograde tracers were injected into different ganglia have revealed that there are small differences in the proportions of all preganglionic neurones that contain NOS that project to different paravertebral sympathetic ganglia; these differences largely parallel the differences in the occurrence of NOS-containing preganglionic neurones at different levels of the spinal cord (the proportion of total preganglionic neurones that project to the superior cervical, stellate and L5 lumbar sympathetic chain ganglia that contain NOS are 84%, 77% and 89%, respectively; all figures calculated from data in Grkovic and Anderson, 1996). In contrast, different prevertebral ganglia of the rat differ markedly in the proportion of their preganglionic neurones that contain NOS (superior mesenteric ganglion 43%, inferior mesenteric ganglion 85%; Grkovic and Anderson, 1996). There are similar differences between prevertebral ganglia in guinea pigs, with a larger percentage of preganglionic neurones projecting to the inferior mesenteric ganglion containing NOS (69%, Anderson et al., 1994) than to the coeliac ganglion (42%, Furness and Anderson, 1994). Sympathetic postganglionic neurones innervate a range of target tissues. Most of them use noradrenaline, ATP and a variety of peptides, particularly neuropeptide Y, as neurotransmitters. The overwhelming majority of studies have found very few NOS-containing neurones in the sympathetic paravertebral ganglia of the mouse and rat (Fig. 2A; Grozdanovic et al., 1992; Anderson et al., 1993; Dun et al., 1993a; Santer and Symons, 1993; Ceccatelli et al., 1994; Klimaschewski et al., 1994; Vanhatalo and Soinila, 1994: Morales et al., 1995; Handa et al., 1996; Lumme et al., 1996; Soinila et al., 1996), and they are only slightly more common in the guinea pig (Kummer et al., 1992; Fischer et al., 1993, 1996a). However, NOS neurones are common in sympathetic paravertebral ganglia of larger species including the cat (Fig. 2C; Anderson et al., 1995), dog (Hisa et al., 1995, 1997), pig (Modin et al., 1994), humans (Klimaschewski et al., 1996) and cattle (Sheng et al., 1993; Majewski et al., 1995). Little is known about the targets of the sympathetic NOS neurones in these larger animals except in the cat, where their targets include hindlimb vasculature, and sweat glands and blood vessels in paw pads (Fig. 2D; Anderson et al., 1995). In prevertebral ganglia, NOS-containing neurones are also present in low numbers in the coeliaco-mesenteric complex and in the inferior mesenteric ganglion of rats (Santer and Symons, 1993), guinea pigs (Furness and Anderson, 1994) and pigs (Kaleczyc et al., 1994). Sympathetic postganglionic neurones that contain NOS are all reported to lack noradrenergic markers (e.g. Kummer, 1992; Kummer et al., 1992; Modin et al., 1994; Majewski et al., 1995), with the exception of the bovine superior cervical ganglion, where a proportion of the NOS-containing neurones also contain dopamine [3-hydroxylase (Sheng et al., 1993).

2.1.2. Functionally identified subclasses of sympathetic preganglionic NOS neurones The targets of sympathetic preganglionic neurones include postganglionic neurones of different function and the chromaffin cells of the adrenal medulla. The presence of NOS in preganglionic neurones is likely to correlate with functionally distinct pathways. For instance, when a retrograde tracer is injected into the adrenal medulla of the rat, all labelled preganglionic neurones are NOS-immunoreactive (Grkovic and Anderson, 1997), even though at middle to low thoracic levels of the spinal cord, only around 70% of the preganglionic neutones contain NOS (see above). Determining the presence or absence of NOS in preganglionic neurones projecting to functionally identified subpopulations of postganglionic neurones relies 219

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Fig. 2. (A) NOS-immunoreactive nerve terminals outline the nerve cell bodies (asterisks) of postganglionic neurones in the rat superior cervical ganglion. The NOS nerve terminals arise from sympathetic preganglionic neurones. Note that there is no sign of immunoreactivity in the postganglionic cell bodies. Scale bar = 50 lxm. (B) Horizontal section through the sacral parasympathetic nucleus (IML) in the S1 spinal cord segment of a rat, stained for the NADPH diaphorase reaction. Stained parasympathetic preganglionic neurones are present oriented transversely across the lateral portion of the grey matter (G), close to the white matter (W). Scale bar -- 100 Ixm. (C) Section through the stellate ganglion of the cat stained for the NADPH diaphorase reaction. NADPH diaphorase-stained postganglionic neurones are present and they represent sympathetic cholinergic neurones that project to either blood vessels or to sweat glands (see Anderson et al.. 1995). Scale bar = 50 ~m. (D) Micrograph of a blood vessel from the paw of a cat, stained for the NADPH diaphorase reaction. The stained nerve terminals are likely to be collaterals from sympathetic cholinergic neurones that also innervate sweat glands in the paw pad. Scale bar = 50 lxm.

on the identification o f s u b p o p u l a t i o n s of p o s t g a n g l i o n i c n e u r o n e s b a s e d on the p r e s e n c e o f u n i q u e c o m b i n a t i o n s o f i m m u n o h i s t o c h e m i c a l l y d e t e c t a b l e s u b s t a n c e s a n d / o r the injection o f r e t r o g r a d e t r a c e r into p a r t i c u l a r p e r i p h e r a l targets. T h e f u n c t i o n a l l y identified p o s t g a n g l i o n i c n e u r o n e s are then e x a m i n e d to d e t e r m i n e if they are s u r r o u n d e d by N O S - i m m u n o r e a c t i v e n e r v e t e r m i n a l s . In the rat, N O S is p r e s e n t in p r e g a n g l i o n i c n e u r o n e s i n n e r v a t i n g p o s t g a n glionic n e u r o n e s p r o j e c t i n g to s w e a t glands, s u b m a n d i b u l a r salivary gland, heart, skeletal m u s c l e , b l o o d v e s s e l s and b r o w n fat ( G r k o v i c and A n d e r s o n , 1995, 1997; H a n d a et al., 1996; 220

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Anderson, 1998; Chiba and Tanaka, 1998). NOS is absent from the preganglionic neurones innervating postganglionic neurones projecting to the iris (CRA, unpubl.). Other functional groups of preganglionic neurones almost certainly contain NOS.

2.1.3. Presence of NOS in parasympathetic pre- and postganglionic neurones NOS is also present in many parasympathetic preganglionic (Fig. 2B) and postganglionic (Figs. 3B,C, 5A and 7A) neurones. Parasympathetic preganglionic neurones are located in the brain and lumbosacral spinal cord. Cranial parasympathetic preganglionic neurones are found in distinct nuclei throughout the brain stem and their axons run in the cranial nerves to innervate postganglionic neurones in the ciliary, sphenopalatine, submandibular and otic ganglia, as well as many neurones within organs in the head and within the thoracic and abdominal cavities. Lumbosacral parasympathetic preganglionic neurones are present in the sacral intermediolateral nuclei, two parallel longitudinal nuclei in the lateral aspects of the intermediate grey matter analogous to the intermediolateral nuclei of the thoracic spinal cord (Petras and Cummings, 1972), and they project via ventral roots to innervate postganglionic neurones in ganglia associated with the pelvic viscera. Transmission from all parasympathetic preganglionic neurones to postganglionic neurones is cholinergic, and NO is not a primary neurotransmitter. The presence of NOS in pelvic parasympathetic postganglionic neurones is discussed in Chapter VIII of this volume. Cranial preganglionic NOS neurones. The presence of NOS-containing nerve terminals in the major parasympathetic ganglia has been reasonably well studied, but the presence of NOS terminals in ganglia within structures in the head, thorax and abdomen has not been systematically examined. If NOS expression is confined to specific pathways, as it is in the sympathetic nervous system, then NOS-containing preganglionic nerve terminals may not necessarily be present in all parasympathetic ganglia. As most parasympathetic ganglia contain nerve cell bodies and dendrites strongly immunoreactive for NOS or reactive for NADPH diaphorase (see Section 3.1.1 (a,c), Sections 5.2 and 6.1), NOS-containing preganglionic inputs to these cells may be obscured. Notwithstanding, in the rat and mouse, the terminals of parasympathetic preganglionic neurones projecting to cranial ganglia rarely seem to contain NOS. Postganglionic neurones in the sphenopalatine ganglion of the rat are not innervated by preganglionic neurones that contain NOS (Ceccatelli et al., 1994). Similarly, the nerve cell bodies in the submandibular ganglia of mice and rats are not innervated by NOS-containing preganglionic neurones (Grozdanovic et al., 1992). In contrast, following injection of a retrograde tracer into the rabbit sphenopalatine ganglion, 75% of the labelled preganglionic neurones in the brain stem are immunoreactive for NOS (Zhu et al., 1997). A similar study in which retrograde tracer was injected into the submandibular ganglion of the rabbit reported that 100% of the retrogradely labelled preganglionic neurones as NOS-immunoreactive (Zhu et al., 1996). A subpopulation of axons in the vagus nerve contain NOS; some of these are the peripheral extensions of vagal sensory neurones (Fischer et al., 1993, 1996a), and some are efferent fibres that originate in the dorsal motor nucleus and project to the gastrointestinal tract and the heart. Efferent vagal NOS fibres are also very likely to project to other organs, but this is yet to be demonstrated. Following intraperitoneal injections of fluorogold, which will label all preganglionic neurones (Anderson and Edwards, 1994), only about 10% of the labelled nerve cell bodies in the dorsal motor nucleus of the vagus of the rat are stained for NADPH diaphorase (Krowicki et al., 1997). In the guinea pig, at least some of the vagal preganglionic neurones containing NOS innervate intrinsic cardiac ganglia, because there are 221

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Fig. 3. (A) Whole-mount preparation of a pial vessel from a guinea pig stained for the NADPH diaphorase reaction. Stained nerve terminals are abundant in the perivascular plexus of this vessel. The background staining is due to the presence NOS in endothelial cells of this vessel. Scale bar = 25 [tm. (B) Section through the sphenopalatine (pterygopalatine) ganglion of a rat after processing for neuronal NOS histochemistry. There are many NOS-immunoreactive nerve cell bodies in the ganglion and some sparse NOS-containing nerve fibres. Scale bar = 50 gm. (C, D) Paired micrographs of a section through the sphenopalatine ganglion of a rat that had been processed for both NOS (C) and choline acetyltransferase (D, CHAT) immunohistochemistry. Most of the NOS-immunoreactive nerve cell bodies (arrows indicate two examples) are also ChAT-positive. There are a small number of cells that are ChAT-immunoreactive (asterisk). but not NOS-positive (the location of the cell in C appears as a black hole). Note the high density of ChAT-positive nerve terminals, which presumably arise from parasympathetic preganglionic neurones, surrounding the nerve cell bodies. Scale bar = 25 Itm.

NOS nerve fibres in the heart that degenerate following vagotomies (Tanaka and Chiba, 1998; see Section 3.2). In the mouse, some of the vagal NOS-containing nerve fibres originating in the dorsal motor nucleus project to the myenteric plexus of the oesophagus, but none of the neurones in the nucleus ambiguus, which innervate the striated muscle of the oesophagus, contain NOS (Sang and Young, 1998b; Sang et al., 1999). However, the NOS neurones comprise only about one third of the neurones projecting from the dorsal motor nucleus to the mouse oesophagus (Sang and Young, 1998b). 222

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Sacral preganglionic NOS neurones. NOS-containing neurones in the sacral spinal cord have been identified as preganglionic neurones, based on their location in the intermediolateral nucleus or by the use of retrograde tracer, in the mouse (Brtining, 1992; Briining and Mayer, 1996), rat (Anderson, 1992; Valtschanoff et al., 1992; Vizzard et al., 1993; Saito et al., 1994; Burnett et al., 1995), cat (Dun et al., 1993b) and dog (Vizzard et al., 1997). The sacral parasympathetic preganglionic outflow innervates postganglionic neurones that control a range of pelvic organs. There is little systematic knowledge of the functional pelvic pathways that may include NOS-containing parasympathetic preganglionic neurones. Virus-tracing experiments have shown that at least some preganglionic neurones projecting to the parasympathetic ganglia of the bladder in the rat contain NOS (Papka et al., 1995). Parasympathetic postganglionic NOS neurones. Many cranial and sacral parasympathetic postganglionic neurones contain NOS. The presence of NOS in cranial postganglionic neurones is discussed in Section 3.1.1 (a) and Section 6.1, and the presence of NOS in pelvic ganglia is discussed briefly in Section 3.1.1 (c) and in Chapter VIII of this volume.

2.2. NITRIC OXIDE AND GANGLIONIC TRANSMISSION Given the widespread occurrence of NOS in sympathetic preganglionic neurones, it is reasonable to expect that NO might influence transmission from the preganglionic terminals to postganglionic nerve cells. The primary transmitter at these synapses is acetylcholine which acts through nicotinic receptors to generate fast excitatory postsynaptic potentials (EPSPs). In the isolated rat superior cervical ganglion, NO donors increase the size of postganglionic compound action potentials when the preganglionic nerve supplying the ganglion is electrically stimulated (Briggs, 1992). The effect is likely to be mediated by increases in cGMP, as application of 8-bromo cGMP, the membrane permeant analogue of cGMP, has identical effects to nerve stimulation (Briggs, 1992). Ganglionic cGMP levels are elevated following either electrical stimulation of the preganglionic nerves or exposure of the ganglion to high potassium concentrations (Quenzer et al., 1980; Volle et al., 1981, 1982; Ando et al., 1983; Volle and Quenzer, 1983; De Vente et al., 1987a,b; Sheng et al., 1992) and this effect can be blocked by NOS inhibitors (Sheng et al., 1992), or by prior preganglionic denervation (Volle et al., 1982). Other studies, using patch-clamping techniques, have demonstrated that NO donors enhanced Ca 2+ currents in postganglionic neurones in the rat superior cervical ganglion (Chen and Schofield, 1993, 1995). The effect is again mediated by cGMP, as it is mimicked by direct application of cGMP analogues and inhibited by methylene blue. Thus, NO is concluded to augment cholinergic neurotransmission in the superior cervical ganglion. In contrast to paravertebral ganglia, the effect of NO release in prevertebral ganglia is to diminish the effectiveness of transmission. In the mouse superior mesenteric ganglion, NO donors directly hyperpolarised the membrane of two thirds of the neurones impaled (Mazet et al., 1996). In this study, stimulation of the colonic nerves in the presence of a NOS inhibitor led to an increase in the size of the slow EPSP, suggesting that under normal circumstances, NO released by nerve stimulation inhibited the slow EPSP. The fact that this effect was present when the colonic nerves were stimulated suggests that the origin of the NOS nerve fibres involved was in the gut (intestinofugal neurones, see Section 4.8), or perhaps collaterals of dorsal root ganglion neurones (Domoto et al., 1995), rather than from preganglionic neurones. In the rabbit coeliac ganglion, prolonged stimulation of the splanchnic nerves led to a decreased probability of a presynaptic impulse firing an action potential in the coeliac ganglion neurones (Quinson et al., 1998). The progressive decrease in the probability of firing an action potential was due to the release of NO, as it was prevented by NOS inhibitors 223

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and NOS scavengers. The NO was presumably acting via increases in cGMP levels to diminish nicotinic transmission, as the probability of firing an action potential was increased by inhibitors of cGMP production and decreased by inhibitors of phosphodiesterases.

3. ROLE OF NO IN THE NEURAL CONTROL OF THE VASCULATURE AND THE HEART

3.1. ROLE OF NEURALLY DERIVED NO IN THE CONTROL OF THE VASCULATURE NO is a potent vasodilator. NOS is present in vascular endothelial cells, and NO is produced by these cells under basal conditions and in response to a variety of stimuli, including acetylcholine released from autonomic nerve terminals (Furchgott, 1996). NO derived from endothelial cells is thus an essential component of many of the signalling pathways mediating relaxation of blood vessels. However, some blood vessels are also innervated by nerve fibres containing NOS, and neurally released NO can also mediate vasodilation. It can be difficult to determine the relative physiological importance of the two sources of NO, and prior to the recent development of drugs that selectively block endothelial NOS or neuronal NOS, the only way that the contribution of neurally derived NO could be distinguished from that of endothelial-derived NO was by pharmacological examination of the relaxations with and without the endothelium. In this section, we will only consider the role of neurally derived NO in the regulation of vascular tone. Although neural control of blood flow through most blood vessels is mediated primarily by sympathetic vasoconstrictor neurones, blood vessels in some tissues can be innervated by three types of neurones: sympathetic vasoconstrictor neurones, parasympathetic, enteric or sympathetic vasodilator neurones, and the peripheral processes of some dorsal root ganglion neurones which mediate vasodilation (see Morris et al., 1995). All vasoconstrictor neurones are thought to contain noradrenaline, and to use various combinations of noradrenaline, ATP and neuropeptide Y as neurotransmitters. Since NO is a potent vasodilator, not surprisingly, NOS does not appear to be present in any vasoconstrictor neurones (Kummer et al., 1992; Hohler et al., 1995). 3.1.1. Autonomic vasodilator neurones

Populations of non-noradrenergic vasodilator neurones are present in cranial parasympathetic ganglia, some sympathetic ganglia, pelvic ganglia and enteric ganglia. (a) Vasodilator neurones in cranial parasympathetic ganglia. Vasodilator neurones play a prominent role in the regulation of blood flow through many blood vessels in the head (see Lee et al., 1975; Faraci and Brian, 1994; Toda and Okamura, 1996). Thus, in addition to sympathetic nerve fibres, the arteries supplying the brain are innervated by parasympathetic vasodilator nerve fibres arising from cranial parasympathetic ganglia, primarily the sphenopalatine (pterygopalatine) ganglion (Chorobski and Penfield, 1932; Hara and Weir, 1986; Goadsby, 1990). In addition, blood vessels supplying other structures in the head such as the salivary glands, the tongue, and the nasal mucosa are innervated by parasympathetic neurones that are located either in intrinsic ganglia or in major cranial parasympathetic ganglia. Recent studies indicate that NOS is present in the terminals of vasodilator neurones in the perivascular plexuses of arteries supplying the brain and other structures in the head, including the eye, salivary glands, tongue and nasal mucosa (Fig. 3A; 224

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Bredt et al., 1990; Bredt and Snyder, 1992; Ceccatelli et al., 1992; Grozdanovic et al., 1992; Iadecola et al., 1993; Suzuki et al., 1993; Yamamoto et al., 1993; Minami et al., 1994; Saito and Goto, 1994; Yoshida et al., 1994; Lohinai et al., 1995; Toda et al., 1995: Goadsby et al., 1996; Toda and Okamura, 1996; Young et al., 1997; Zhu et al., 1997). The role of NO in axons innervating blood vessels supplying the salivary glands and nasal mucosa is discussed in Sections 6.1 and 6.3. Using NOS immunohistochemistry and/or NADPH diaphorase histochemistry, NOS-containing nerve cell bodies have been found to comprise up to 90% of neurones in cranial parasympathetic ganglia, including the sphenopalatine and ciliary ganglia, and intrinsic ganglia within the salivary glands and the tongue of a variety of species (Fig. 3B; Grozdanovic et al., 1992; Yamamoto et al., 1993; Yoshida et al., 1993; Ceccatelli et al., 1994; Minami et al., 1994; Sun et al., 1994; Lee et al., 1995; Goadsby et al., 1996; Hu et al., 1996; Hanazawa et al., 1997; Jeon et al., 1997; Zhu et al.. 1997; Lacroix et al., 1998). Studies in rats and dogs that have examined the origin of the NOS-containing nerve fibres innervating cerebral arteries have shown that many of them arise from the sphenopalatine ganglion. In the rat, sectioning of fibres leaving both sphenopalatine ganglia (Nozaki et al., 1993) or unilateral removal of a sphenopalatine ganglion (Minami et al., 1994) result in a marked decrease in the density of NOS-immunoreactive nerve fibres associated with vessels forming the rostral circle of Willis. Moreover, most of the labelled neurones in the sphenopalatine ganglion labelled by application of retrograde tracer to the middle cerebral artery are NADPH diaphorase-positive (Minami et al., 1994). In the dog, damaging the sphenopalatine ganglion, or sectioning of the nerve fibres leaving it, results in a dramatic reduction or total loss of NOS-immunoreactive nerve fibres in the ipsilateral middle and posterior cerebral arteries, demonstrating that the sphenopalatine ganglion is the major source of NOS-containing nerve fibres innervating these vessels (Nozaki et al., 1993; Yoshida et al., 1993). Cholinergic nerve terminals, revealed by choline acetyltransferase (CHAT) immunohistochemistry, form a perivascular plexus around the cerebral and basilar arteries of a range of animals (Saito et al., 1985). These cholinergic nerve terminals also contain VIP (Hara et al., 1985). Many of the NOS-containing neurones innervating cerebral blood vessels also contain VIP (Ceccatelli et al., 1992; Minami et al., 1994; Goadsby et al., 1996), and thus the cerebral vasodilator neurones are cholinergic/NOS/VIP neurones (Fig. 3C,D). Although acetylcholine appears to be the main mediator of the nerve-mediated vasodilation of cerebral vessels (Forbes and Cobb, 1938), there is also an endothelium-independent dilatory response to nerve stimulation that is not mediated by acetylcholine (Lee, 1980; Lee et al., 1984). There is convincing pharmacological evidence from organ-bath experiments and from whole-animal studies that NO is an important mediator of vasodilation in cerebral arteries (for reviews see: Iadecola, 1993; Faraci and Brian, 1994), but most studies have not distinguished between endothelial-derived and neurally derived NO. However, experiments on cerebral arteries from which the endothelium has been removed show that neurally released NO contributes to the vasodilator responses. For example, segments of pig basilar artery (Lee and Sarwinski, 1991), monkey cerebral and temporal arteries (Toda and Okamura, 1990; Yoshida et al., 1994), guinea-pig middle cerebral artery (Saito and Goto, 1994) and dog cerebral artery (Toda and Okamura, 1990, 1991) from which the endothelium has been removed respond to transmural electrical stimulation with relaxations that are abolished by tetrodotoxin, and abolished or dramatically reduced by NOS inhibitors. Thus, although parasympathetic vasodilator nerves innervating the cerebral vessels contain at least three potential neurotransmitters, NO appears to the be dominant neurotransmitter that is responsible for nerve-mediated endothelium-independent vasodilation, and acetylcholine is the predominant neurotransmitter responsible for the nerve-mediated endothelium-dependent vasodilation. 225

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(b) Vasodilator neurones in sympathetic ganglia. In many species, a small population of sympathetic vasodilator neurones innervate the arteries supplying skeletal muscle, and they contain acetylcholine-related markers and a variety of peptides (Lindh et al., 1989; Gibbins, 1992; Morris et al., 1998). The presence of NOS and other peptides in sympathetic vasodilator neurones varies between species. In the guinea pig, sympathetic vasodilator neurones contain VIP, but not NOS (Morris et al., 1998), and thus any involvement of NO in vasodilation must be exclusively that derived from the endothelium. Similarly, in the pig, there are no NOS-immunoreactive nerve fibres associated with blood vessels supplying skeletal muscle (Modin et al., 1994). In the rat the situation is unclear. Following exposure of the rat to air-jet stress (a noxious stimulus), there is a marked vasodilation of the hindquarter vasculature, which is thought to be part of a 'defence reaction' (Folkow, 1982). Davisson et al. (1994, 1997) have reported that neurally released NO is responsible for the air-jet stress-induced vasodilation in the rat. However, a previous report (Anderson et al., 1993) had shown that there are no NADPH diaphorase-positive or NOS-immunoreactive nerve cell bodies in lumbar sympathetic ganglia of the rat, and so the source of the NOS nerve fibres that were concluded to innervate the rat hindlimb vasculature is unclear. In the cat, vasodilator neurones innervating the hindlimb vasculature contain both NOS and VIP, but not CGRP (Anderson et al., 1995). Arterioles in the cat paw pad are also innervated by NOS nerve fibres (Fig. 2D), but the contribution of neurally released NO to vasodilatory responses in the hindlimb and paw pad arteries of the cat has yet to be examined. (c) Vasodilator neurones in peh,ic ganglia. Like cerebral arteries, arteries supplying erectile tissue and the uterus are densely innervated by vasodilator neurones. Vasodilation of pelvic arteries is important during the later stages of pregnancy and for sexual function (Bell, 1972). Within the penis of many mammalian species, the penile arteries, veins and cavernous tissue are surrounded by a plexus of nerve fibres showing NOS immunoreactivity and NADPH diaphorase staining (Burnett et al., 1992, 1993; Keast, 1992; Alto et al., 1993; Ding et al., 1993; Schirar et al., 1994; Dail et al., 1995: Hedlund et al., 1995; Tamura et al., 1995). Retrograde tracing studies have shown that the vast majority of these NOS-containing nerve fibres originate from pelvic ganglia (Keast, 1992; Ding et al., 1993, 1995; Domoto and Tsumori, 1994; Schirar et al., 1994, 1997; Vizzard et al., 1994; Vanhatalo and Soinila, 1995). A retrograde tracing study in which tracer was injected into the penile shaft revealed some labelled, NOS-immunoreactive neurones in the intermediolateral column of the upper lumbar levels of the spinal cord, suggesting that some NOS preganglionic neurones may project directly to the penis (Vanhatalo and Soinila, 1995). However, it is also possible that the tracer was taken up by the cell bodies of neurones along the pelvic nerves, which are part of the pelvic plexus. Lumbar level dorsal root ganglia may also be sources of some of the NOS-containing nerve terminals within the penis (McNeill et al., 1992). A large proportion of the NOS neurones in the pelvic plexus that innervate the penile vasculature also contain acetylcholine-related molecules and VIE but none contain tyrosine hydroxylase (Ding et al., 1995; Hedlund et al., 1995; Ehmke et al., 1995; Tamura et al., 1995; Dail, 1996; Vanhatalo et al., 1996). There is substantial evidence showing that both endothelial-derived and neurally derived NO contribute to relaxation of vascular smooth muscle necessary for penile erection (see Burnett, 1995, 1997). Electrical field stimulation induces relaxations of the smooth muscle of the corpus cavernosum that are abolished by tetrodotoxin and markedly reduced by NOS inhibitors (Ignarro et al., 1990; Kim et al., 1991; Knispel et al., 1992a,b; Rajfer et al., 1992; Wang et al., 1994; Hedlund et al., 1995; Hayashida et al., 1996). Relaxations can be elicited from both endothelium-denuded and endothelium-intact preparations, and in both types of preparations, the relaxations induced by low-frequency stimulation are abolished by NOS 226

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inhibitors, whereas higher-frequency stimulation induces relaxations that are partly resistant to NOS inhibition; thus neurally derived NO plus another unidentified molecule appear to contribute to penile vasodilation (Kim et al., 1991" Simonsen et al., 1995, 1997). Although VIP is co-localised with NOS in many of the axons innervating penile blood vessels, no role for VIP in the regulation of muscle tone has been detected (Hayashida et al., 1996). Mice with a targetted deletion of the neuronal NOS gene (Huang et al., 1993" Burnett et al., 1996) or the endothelial NOS gene (Huang et al., 1995) breed normally, suggesting that neither endothelial NOS nor neuronal NOS are, by themselves, essential for penile function. However, in the neuronal NOS knockout mice, penile erections can be blocked by non-isoform-specific NOS inhibitors such as L-NAME, and since the levels of endothelial NOS present within the endothelium of these mice are significantly higher than in wild-type mice, it is believed that NO released from the endothelium mediates vasodilation during erections in mice lacking neuronal NOS (Burnett et al., 1996). Hence, compensatory mechanisms may prevent the normal physiological role of each isoform of NOS from being revealed in the knockout mice. Little is known about the role in NO in erectile tissue (clitoris and bulbs) of females. Neuronal NOS immunoreactivity is present in nerve fibres, and endothelial NOS immunoreactivity is present in vascular and sinusoidal endothelial cells of the human clitoris (Burnett et al., 1997b). However, to our knowledge there have been no functional studies examining the role of NO in this tissue. The uterine artery is innervated by vasodilator neurones, and a significant component of the vasodilator response in this vessel is non-cholinergic (Bell, 1969) and endothelium-independent (Morris, 1993" Nelson et al., 1995). NOS-immunoreactive nerve fibres form a plexus around the uterine arteries of humans and guinea pigs (Kummer et al., 1992; Toda et al., 1994; Anderson et al., 1997). In the guinea pig, the vasodilator neurones innervating the uterine artery arise from pelvic (paracervical) ganglia and they contain five potential inhibitory neurotransmitters or their synthetic enzymes: acetylcholine, VIP, CGRP, neuropeptide Y and NOS (Morris et al., 1985" Morris and Gibbins, 1987" Anderson et al., 1997). Pharmacological studies have revealed endothelial-independent vasodilations of the uterine artery in guinea pigs and humans that are blocked by tetrodotoxin and inhibited, but not abolished, by NOS inhibitors (Morris, 1993" Toda et al., 1994; Nelson et al., 1995). Thus both neurally released NO plus another neurotransmitter mediate endothelium-independent vasodilation in the uterine artery. In the guinea pig, the other neurotransmitter is likely to be a peptide such as VIP or CGRP, because the non-NO component of the response is trypsin-sensitive (Morris, 1993). Both the endothelium-dependent and endothelium-independent vasodilatory responses observed in uterine arteries become more pronounced during pregnancy (Nelson et al., 1995, 1998; Jovanovic et al., 1997). (d) Vasodilator neurones in the enteric nervous system. Submucosal arterioles are densely innervated by axons of extrinsic vasoconstrictor neurones that arise exclusively from sympathetic ganglia, axons arising from extrinsic sensory (dorsal root ganglia) neurones that cause vasodilation, and intrinsic vasodilator neurones with cell bodies in enteric ganglia (see Vanner and Surprenant, 1996). The vasodilatory responses that occur following the ingestion of food appear to be mediated largely by intrinsic neurones, whereas extrinsic sensory neurones mediate vasodilation principally during inflammatory states. Although endothelial NOS immunoreactivity and NADPH diaphorase activity are present in the endothelial cells of submucosal arterioles (Nichols et al., 1992, 1993" O'Brien et al., 1995), there have been no reports of NOS-containing nerve fibres associated with these vessels, and neurogenic vasodilation in submucosal arterioles appears to be mediated by acetylcholine, VIP and possibly other peptides (Vanner and Surprenant, 1996). 227

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(e) Blood vessels innervated by NOS nerve fibres of unknown origin and role. In the human, guinea pig and ferret, the blood vessels supplying the airways are surrounded by a plexus of NOS-immunoreactive nerve fibres (Dey et al., 1993" Fischer et al., 1993; Fischer and Hoffmann, 1996). However, the source and the role of these perivascular NOS fibres in the airways have yet to be examined. Since there are no intrinsic NOS-immunoreactive nerve cell bodies in the guinea-pig trachea, the perivascular NOS fibres in the guinea-pig airways are likely to arise from nerve cell bodies in the oesophagus, non-noradrenergic sympathetic neurones in the stellate ganglion (Section 5) or extrinsic sensory neurones (Fischer et al., 1993, 1996a). In humans and ferrets, NOS-immunoreactive nerve cell bodies are present in intrinsic ganglia of the trachea, and hence at least some of the perivascular NOS fibres could arise from this source (Section 5). There do not appear to have been any pharmacological studies which have examined the role of NO in the neural control of the airway vasculature. 3.1.2. Sensory vasodilation

Perivascular sensory nerve fibres (arising from dorsal root ganglia and vagal sensory ganglia) are associated with many blood vessels (Furness et al., 1982; Gibbins et al., 1985, 1987), and release a variety of neurotransmitters that can regulate vascular function (see Holzer and Maggi, 1998). The actions of sensory neurones can be distinguished from autonomic vasodilator neurones by their sensitivity to capsaicin, which acutely causes excitation, and in the long-term, degeneration of their nerve endings (Holzer, 1991). When stimulated, perivascular sensory fibres cause a vasodilation that is mediated by a number of different neurotransmitters, primarily CGRP, although ATP and tachykinins can also be involved (for reviews see: Holzer et al., 1995a; Maggi, 1995). NOS is present in subpopulations of sensory neurones (Fig. 4), and the role of sensory neurone-derived NO in vasodilation varies between vascular beds and species (Holzer et al., 1995b). For example, in mesenteric arterioles of the rabbit, neurally released ATP and CGRP fully account for the capsaicin-sensitive relaxation, although endothelial-derived NO is involved in the response (Kakuyama et al., 1998). In the cerebral cortex of the cat, the increase in blood flow induced by stimulation of the nasociliary nerve (which stimulates the fibres of trigeminal neurones) is unaffected by NOS inhibition, and the cerebral vasodilatation appears to be mediated largely by CGRP (Edvinsson et al., 1998). However, NO derived from primary sensory neurones does appear to mediate vasodilation in the mesenteric artery of the guinea pig. In segments of mesenteric artery of the guinea pig from which the endothelium has been removed and which have been pre-treated with guanethidine (to block neurotransmitter release from sympathetic nerve terminals), vasodilatory responses are reduced by over 50% by the NOS inhibitor, L-NNA, and nerve terminals containing both NOS and substance P are found in the perivascular plexus of this artery (Zheng et al., 1997). In addition to acting directly on the vascular smooth muscle, there is also evidence in some vascular beds that NO can facilitate the release of CGRP from afferent nerve fibres (Holzer et al., 1995b). 3.1.3. Role of neurally released NO in regulation of blood vessels --- summary

(i) NO released from cranial parasympathetic neurones innervating cerebral blood vessels, and other blood vessels supplying structures in the head, is an important mediator of nerve-stimulated, endothelial-independent vasodilation. (ii) NO released from the terminals of pelvic parasympathetic neurones innervating the uterine artery and blood vessels supplying the penis is an important mediator of vasodilation. 228

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Fig. 4. (A) Frozen section through the nodose ganglion of a rat, showing scattered, NOS-immunoreactive neurones in the ganglion. Scale bar -- 100 ~tm. (B) Scattered NOS-immunoreactive neurones are also present in the dorsal root ganglia (DRG) of the rat. The ganglion illustrated is from T12 (thoracic level). Scale bar = 25 ~tm. (C, D) Paired, high-power micrograph showing co-localisation between NOS (C) and calcitonin gene-related peptide (D, CGRP) in a T12 dorsal root ganglion of the rat. Some CGRP-positive cells (asterisks) do not show NOS immunoreactivity. Scale bar -- 25 ~tm. (iii) In some species only, sympathetic vasodilator neurones innervating skeletal muscle m a y use N O as a neurotransmitter. (iv) Within the gastrointestinal tract, neurally derived NO plays no role in the vasodilation of submucosal arterioles mediated by enteric vasodilator neurones. (v) Neurally mediated vasodilation in some vascular beds is mediated by NO released from subpopulations of perivascular sensory nerve fibres arising from dorsal root ganglia. 3.2. R O L E OF N E U R A L L Y D E R I V E D NO IN T H E C O N T R O L OF T H E H E A R T The heart is innervated by sympathetic (excitatory) neurones, vagal parasympathetic (inhibitory) pathways and extrinsic sensory neurones. The sympathetic nerve fibres arise from extrinsic neurones located in the cervical and upper thoracic sympathetic ganglia, and the sen229

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sory innervation arises from dorsal root ganglia and vagal sensory ganglia. An early study by Schmiedeberg and Koppe (1869) showed that the effects of muscarine and vagal stimulation on the heart are similar, and both are antagonised by atropine. Loewi (1921) demonstrated that the slowing of the heart in response to vagal activation is due to the release of a substance (which he called 'Vagusstoff'), whose action was blocked by atropine. Although the vagal parasympathetic preganglionic neurones that innervate the heart are cholinergic, the acetylcholine that inhibits the cardiac muscle is actually released from intrinsic neurones that are located in small ganglia within the heart and its surrounding tissues (King and Coakley, 1958), and which receive synaptic input from vagal nerve terminals. Consistent with the pharmacological data, all of the intrinsic cardiac neurones in the guinea-pig heart are ChAT-immunoreactive, and thus cholinergic (Mawe et al., 1996). However, the cardiac neurones show considerable electrophysiological, morphological and neurochemical diversity (Baluk and Gabella, 1989; Hassall et al., 1992; Steele et al., 1994; Edwards et al., 1995). Subpopulations of the cardiac neurones contain a range of neuropeptides including somatostatin, tachykinins, neuropeptide Y and pituitary adenylate cyclase-activating polypeptide (PACAP) (Baluk and Gabella, 1989; Steele et al., 1994; Braas et al., 1998). In addition, a subpopulation of the neurones in the heart of a range of mammalian species, including the rat, guinea pig, human, monkey and dog, contains NOS, as demonstrated by immunohistochemistry or NADPH diaphorase staining (Fig. 5A; Hassall et al., 1992; Klimaschewski et al., 1992; Tanaka et al., 1993; Steele et al., 1994; Armour et al., 1995; Mawe et al., 1996; Sosunov et al., 1996; Yoshida and Toda, 1996). In the guinea pig and rat, less than 10% of the cardiac neurones contain NOS (Hassall et al., 1992; Klimaschewski et al., 1992; Steele et al., 1994; Mawe et al., 1996), but in the dog, 30-40% of the neurones are NADPH diaphorase-positive (Armour et al., 1995). In close apposition with nerve cell bodies within the cardiac ganglia are nerve terminals that are immunoreactive for ChAT (Mawe et al., 1996), NOS (Fig. 5B; Sosunov et al., 1996), tyrosine hydroxylase or substance P plus CGRP (Steele et al., 1994). Most of the cholinergic nerve terminals probably arise from vagal preganglionic neurones but some are also likely to arise from intrinsic cholinergic neurones, the tyrosine hydroxylase-

Fig. 5. (A, B) Whole-mount preparation from the guinea-pig atrium showing two cardiac ganglia which are adjacent to nerve bundles. The ganglion shown in A has many NOS-immunoreactive nerve cell bodies, whereas the ganglion shown in B has no NOS-positive nerve cell bodies, but varicose, NOS-containing nerve terminals are present in the ganglion. Scale bar = 50 ~m. Preparation kindly dissected by David Hirst.

230

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immunoreactive nerve terminals arise from sympathetic neurones and the substance P/CGRP nerve terminals arise from extrinsic sensory neurones (Steele et al., 1994; Mawe et al., 1996). The source of NOS-immunoreactive nerve terminals within the heart has only recently been examined. Tanaka and Chiba (1998) examined the origin of NOS-immunoreactive nerve terminals in different parts of the guinea-pig heart by performing immunohistochemistry following unilateral cervical vagotomies. They found that the baskets of NOS nerve terminals that are present around intracardiac nerve cell bodies in ganglia associated with the left atrium and the interatrial septum arise from neurones that project to the heart via the left vagus nerve, since they completely disappear following a left-side vagotomy. The authors concluded that these vagal NOS fibres must be parasympathetic preganglionic nerve fibres, although it is also possible that some were extrinsic sensory fibres, since there are many NOS neurones in the nodose ganglion. The NOS-immunoreactive nerve terminals that form baskets around ganglion cells in the atrioventricular nodal region, or that form a sparse plexus covering the fight atrium, are unaffected by vagotomies of the right or left side, and thus must arise from intrinsic, NOS-containing neurones. The physiological role of NO in the neural control of heart rate has been examined in a number of studies (Han et al., 1994; Conlon et al., 1996, 1997; Elvan et al., 1997; Sears et al., 1998a,b). All of the studies have reported some effect of NOS inhibitors and NO donors on the sympatho-vagal control of heart rate. However, most of the studies did not use isoform-specific NOS inhibitors, and since non-isoform-specific NOS inhibitors, such as L-NA, affect baseline heart rate and mean arterial pressure, the results can be difficult to interpret. However, a study was recently published which used the specific neuronal NOS inhibitors, 1-(2-trifluoromethylphenyl) imidazole (TRIM) and 7-nitroindazole (7-NiNa), to examine the effect of neuronal NOS inhibition on sympathetic and vagal control of heart rate (Sears et al., 1998b). Unlike non-isoform-specific NOS inhibitors, TRIM was found to have no significant effect on mean arterial blood pressure or baseline heart rate in cardiac sympathectomised and vagotomized anaesthetised rabbits. Following unilateral sympathetic stellate ganglion stimulation, TRIM caused a significant increase in the cardioacceleration, which was reversed by L-arginine, indicating that neurally released NO inhibits the heart rate response to sympathetic nerve stimulation. Since TRIM had no effect on the change of heart rate observed following intravenous infusion of isoprenaline, the neurally released NO appears to inhibit the heart rate response to sympathetic nerve stimulation by acting presynaptically on sympathetic nerve terminals, rather than directly on the cardiac muscle (Sears et al., 1998b). In contrast, TRIM, 7-NiNa or even the non-isoform-specific inhibitor L-NA, had no effect on the size of the heart rate response to vagal nerve stimulation in either the anaesthetised rabbit or in the isolated guinea-pig atria (Sears et al., 1998b). However, in the ferret, both TRIM (Conlon et al., 1997) and the non-isoform-specific NOS inhibitors, L-NAME and L-NOARG (Conlon et al., 1996, 1998), reduce the heart rate response to vagal nerve stimulation, demonstrating that in some species, neurally released NO can facilitate the vagal transmission to the heart. To summarise, in the rat and rabbit, neurally released NO acts presynaptically on sympathetic nerve terminals to inhibit the heart rate response to sympathetic nerve stimulation (Sears et al., 1998b). In the ferret (Conlon et al., 1996, 1997, 1998), but not in the rat and rabbit (Sears et al., 1998b), neurally released NO also facilitates the vagal effects on the heart. Thus, overall, neurally released NO has cardiodepressor actions by decreasing sympathetic transmission and, in some species, enhancing vagal transmission. It remains unclear whether the neuronal NO that modulates the sympathetic and vagal control of heart rate is released predominantly from the terminals of vagal preganglionic neurones or from intrinsic cardiac neurones. 231

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4. ROLE OF NO IN THE NEURAL CONTROL OF THE GASTROINTESTINAL TRACT 4.1. INTRODUCTION The gastrointestinal tract is innervated by intrinsic (enteric) neurones, and extrinsic neurones with cell bodies in sympathetic, parasympathetic and sensory ganglia. Within the enteric nervous system are different functional types of neurones, including sensory neurones, interneurones, secretomotor neurones, vasodilator neurones (see Section 3.1.1 (d)) and excitatory and inhibitory muscle motor neurones (Fumess et al., 1999). The evidence that NO is synthesised by subpopulations of enteric neurones, and has roles in enteric transmission, is supported by abundant data (Rand, 1992; Sanders and Ward, 1992; Stark and Szurszewski, 1992; Brookes, 1993). These data indicate that NOS is contained in inhibitory enteric muscle motor neurones of all mammals that have been investigated, and that transmission from enteric inhibitory neurones to gastrointestinal muscle is antagonised when the production or action of NO is compromised. Interneurones that form synapses in the ganglia of the gastrointestinal tract also contain NOS, but NO seems not to have a substantial role in enteric neuro-neuronal transmission. NOS is also in intrinsic neurones that supply one component of the innervation of motor endplates in the oesophagus. In addition, NOS is in neurones supplying the gastric mucosa, in enteric neurones that project from the gut to sympathetic ganglia (intestinofugal neurones), and in neurones of the gallbladder and pancreas. Each of these is discussed below, but greatest emphasis is given to the role of NO in neuromuscular transmission. Although a role of NO in neuromuscular transmission is not in dispute, there are several theories to suggest how this transmission is mediated. 4.2. NO IS A NEUROTRANSMITTER OF ENTERIC INHIBITORY MOTOR NEURONES The enteric inhibitory neurones relax gut muscle. They innervate both the longitudinal and circular layers of the external musculature, and the muscularis mucosae. In functional terms, it is the innervation of the circular muscle that seems the most important because mixing and propulsion of the gut contents depends on contractile activity in this muscle coat. The neurones relax the circular muscle in order to aid the passage of food along the gut and through its sphincters. Although enteric inhibitory neurones were known to physiologists from the beginning of the century, identification of their transmitter(s) has proved difficult, even controversial. In the early 1970s it was proposed that ATP is the transmitter, but soon after evidence for VIP was advanced, then came the evidence that NO participates, followed by evidence for PACAP and carbon monoxide (Burnstock, 1972; Fahrenkrug, 1979; Rand, 1992; Furness et al., 1995). The proposition that NO is a transmitter of enteric inhibitory neurones arose almost simultaneously from six laboratories, all of whom took advantage of the advent of NOS inhibitors and the simplicity of isolated gut muscles for studies of the pharmacology of transmission from these neurones (Gillespie et al., 1989; Li and Rand, 1989; Ramagopal and Leighton, 1989; Bult et al., 1990; Gibson et al., 1990; Hata et al., 1990). In each case, it was found that inhibition of NOS substantially reduced the amplitude of the relaxation of the muscle caused by stimulation of the inhibitory neurones. In the next two to three years, more than 20 papers were published, each showing, in circular and longitudinal muscle, including sphincter muscle, from several mammalian species, that inhibition of NOS or inactivation of NO by oxyhaemoglobin attenuated transmission from the enteric inhibitory neurones 232

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(Sanders and Ward, 1992; Stark and Szurszewski, 1992). The hypothesis that NO is an enteric neurotransmitter was consolidated by the histological demonstration of NOS in enteric neurones, either by NOS immunohistochemistry or by the NADPH diaphorase reaction, which label identical neurones in the gut (Fig. 6A,B; Branchek and Gershon, 1989; Bredt et al., 1990; Belai et al., 1992; Costa et al., 1992; Furness et al., 1992; Grozdanovic et al., 1992; Nichols et al., 1992; Ward et al., 1992; Young et al., 1992). Many more papers describing the distribution of enteric nerve cells and nerve fibres containing NOS were published in subsequent years. Consistently in all species, NOS nerve cells are in the myenteric plexus of the small and large intestines, and nerve fibres are found in the fibre bundles innervating the circular and longitudinal muscle (Fig. 6C). The projections and chemistries of the NOS neurones that innervate the muscle indicate them to be enteric inhibitory motor neurones (Costa et al., 1992; Furness et al., 1992; McConalogue and Furness, 1993; Ekblad et al., 1994a,b; Timmermans et al., 1994; Costa et al., 1996; Porter et al., 1997; Sang et al., 1997; Pfannkuche et al., 1998). The NOS nerve fibres in the circular muscle arise from myenteric nerve cells that send their axons to the underlying and more anally located muscle. VIP immunoreactivity was previously identified as a marker of enteric inhibitory motor neurones (see Furness et al., 1995) and is co-localised with NOS in the neurones that innervate the muscle (Costa et al., 1992, 1996; Furness et al., 1992; Ekblad et al., 1994a,b; Keef et al., 1994; Ward et al., 1994b; Sang and Young, 1996). NO is released from enteric neurones when they are stimulated (Boeckxstaens et al., 1991; Wiklund et al., 1993a; Shuttleworth et al., 1995; Chakder and Rattan, 1996), although this release cannot be attributed specifically to the inhibitory motor neurones; it could be from NOS interneurones. In summary, there is little reason to doubt that NO is a transmitter of the enteric inhibitory motor neurones. However, it is not the only transmitter of these neurones, and the mechanism by which information is transmitted from the neurones to the muscle has been the subject of several theories. 4.3. ROLE OF NO IN CO-TRANSMISSION FROM ENTERIC INHIBITORY MOTOR NEURONES It became apparent in the 1980s that there are separable components of transmission from enteric inhibitory neurones. Niel et al. (1983) reported that the inhibitory junction potential (IJP), elicited by stimulation of these neurones, consisted of two phases of hyperpolarisation in the muscle of the guinea-pig intestine, a fast component blocked by apamin (a blocker of small-conductance potassium channels), and a slow component resistant to this drug. Crist et al. (1992) showed that the fast component of the IJP was blocked by ~,13-methylene ATE thereby consolidating the earlier proposal that ATP is a transmitter. A survey of inhibitory transmission throughout the guinea-pig intestine, using mechanical rather than electrical recording, indicated that the relative contribution of an apamin-sensitive component varied considerably between gut regions (Costa et al., 1986). For example, apamin almost completely abolished inhibitory transmission in the longitudinal muscle of the ileum, but was without effect in the circular muscle of the fundus. Other comparisons of transmission that confirmed the involvement of several transmitters have been reviewed (Furness et al., 1995). In addition to NO and ATP, substances that possibly contribute to muscle relaxation include VIP (Fahrenkrug, 1979; Grider et al., 1985a,b), PACAP (Schworer et al., 1992; Gilder et al., 1994; Jin et al., 1994; McConalogue and Furness, 1993; Katsoulis et al., 1996) and carbon monoxide (Rattan and Chakder, 1993; Ny et al., 1995; Zakhary et al., 1997). 233

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Following the initial identification of NO as an inhibitory transmitter, studies were undertaken to determine which component(s) NO contributed to the transmission process, and how the actions of NO were related to those of the other transmitters. It was found

; i i.-

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that NOS inhibition blocked the slow component of transmission in the guinea-pig ileum (Lyster et al., 1992; He and Goyal, 1993). This seemed to lead to a simple interpretation, the existence of a fast apamin-sensitive component of transmission mediated by ATP, and a slow component mediated by NO. However, it was also reported that the slow component was blocked by the VIP antagonist, VIP 10-28 (Crist et al., 1992). This led to a cascade hypothesis of transmission, in which inhibitory neurones were proposed to release VIP and PACAP (which act on the common VPAC receptor; Harmar et al., 1998), and these transmitters in turn release NO from muscle, which acts back on the muscle, causing relaxation (Grider et al., 1992; Murthy and Makhlouf, 1994). This hypothesis itself is confounded to some extent by the observation that NO-mediated inhibitory transmission, at least in some regions, is mediated via the interstitial cells of Cajal (ICC). The ICC are spindle-shaped cells that are at the surfaces of, and within, the muscle, and are closely apposed by the axons of excitatory and inhibitory motor neurones (Sanders, 1996). Stimulation of inhibitory motor neurones elicited IJPs of reduced amplitude when the ICC were dissected away (Huizinga et al., 1990). A similar result was obtained by comparison of inhibitory transmission in the lower oesophageal sphincter, stomach and pyloric sphincter of normal mice and mutants ( W / W ~') lacking ICC in the circular muscle of these regions (Burns et al., 1996: Ward et al., 1998). The component of transmission that was mediated via NO was absent in the mutants. In a study in which the ICC were disrupted by treating neonatal mice with antibodies to Kit (this specifically damages ICC), excitatory and inhibitory transmission to the circular muscle of the ileum were impaired, and neural transmission to colonic circular muscle was abolished (Torihashi et al., 1995). Data published by Publicover et al. (1993) indicate that the ICC can produce NO. This could possibly amplify transmission to the muscle. Thus, NO might be involved in transmission at three levels: (1) NO can be produced by inhibitory motor neurones and act both directly on muscle and on ICC. (2) The ICC can in turn inhibit the muscle through electrical connections, and perhaps through the facilitated release of NO. (3) NO derived from muscle may amplify inhibitory transmission. The hypothesis that NO is produced by ICC is supported by the immunohistochemical localisation of one of the non-neuronal isoforms of NOS, endothelial NOS (eNOS) in ICC (Xue et al., 1994). Similarly, RT-PCR and Northern analysis indicates that intestinal muscle cells express eNOS (Teng et al., 1998). There is something unusual about the eNOS in intestinal ICC and muscle, in that they are not revealed by N A D P H diaphorase staining (which should reveal all isoforms of NOS), and not by all antibodies against eNOS (Young et al., 1997). There are few experiments which bear on the question of whether NO acts in a primarily serial fashion, via ICC, or mainly in a parallel fashion, both via ICC and directly on the

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Fig. 6. (A) Whole-mount preparation of a myenteric ganglion from the guinea-pig ileum stained for the NADPH diaphorase reaction. There are many stained nerve cell bodies within the ganglion and faintly stained nerve terminals are also present. Scale bar = 50 p.m. (B) Myenteric ganglion from the guinea-pig proximal colon stained for NADPH diaphorase histochemistry. Stained nerve terminals form basket-like arrays around unstained neurones (asterisks). Scale bar -- 25 Ixm. (C) Frozen transverse section through the sphincter of Oddi in the duodenum following processing for NOS immunohistochemistry. There is a high density of nerve terminals within the sphincter (SO), and nerve terminals are also present in the circular muscle layer (CM). LM = longitudinal muscle layer; MP = myenteric plexus. Scale bar = 50 ~m. (Reproduced from Furness et al. (1994), by courtesy of the editors of Cell and Tissue Reseapz'h.) (D, E) Paired micrographs of a myenteric ganglion in the guinea-pig colon, 10 days after the injection of the retrograde tracer, Fast Blue, into the inferior mesenteric ganglion. The retrogradely labelled neurone (arrow) is NOS-immunoreactive (D), indicating that this NOS neurone projected to the inferior mesenteric ganglion. Scale bar -- 25 ~tm. (D and E kindly provided by Alan Lomax.)

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muscle. A serial mechanism is dominant in the mouse stomach and lower oesophageal and pyloric sphincters (Burns et al., 1996; Ward et al., 1998). In these gut regions of normal animals, a substantial component of inhibitory transmission is blocked by NOS inhibitors, but in mutant animals lacking ICC, the inhibitory responses, which are of lesser amplitude, are unaffected by NOS inhibition. It can be concluded that the NO component is via ICC and the non-NO component is mediated by another neurotransmitter acting directly on the muscle. Because NO can act directly on the muscle, for example in the canine colon (Koh et al., 1995), studies of a broader range of nerve-muscle preparations from the gastrointestinal tract may reveal different mixes of in-parallel and in-series roles of NO. The interplay between VIP and NO has been investigated in several organs. In some instances, the two substances seem to act in parallel, whereas in others a VIP-stimulated NO release may also occur. In the rat fundus, the responses to inhibitory nerve stimulation are reduced by VIP immunoneutralisation and reduced further by the combined presence of a NOS inhibitor (Li and Rand, 1990). The relaxation caused by VIP was slightly reduced by NOS inhibition. These results suggest that there may be both VIP-induced NO release and independent actions of VIP and NO. In the ferret stomach, vagal stimulation caused an initial rapid relaxation which was blocked by NOS inhibition, and more slowly developing relaxation that was blocked by VIP immunoneutralisation, suggesting that the actions of NO and VIP are parallel and independent (Grundy et al., 1993). VIP has been reported to stimulate NO production by muscle cells of the rat colon (Gilder, 1993), but not in the dog colon (Keef et al., 1994). In the rat colon, but not in the dog colon, NOS inhibitors reduced the relaxation caused by VIE and in both preparations NOS inhibitors reduced the response to nerve stimulation. Thus in dog colon there appear to be parallel actions of VIP and NO, whereas in the rat, both direct effects of VIP and NO on the muscle, and VIP-stimulated NO release appears to occur. In mice, the ICC that are involved in NO-mediated neurotransmission are only a subpopulation of ICC; a separate subpopulation of ICC is essential for the generation of rhythmic contractions of the circular muscle (Ward et al., 1994a; Huizinga et al., 1995). All ICC depend on the receptor tyrosine kinase, Kit, for survival (Torihashi et al., 1995). W mice, in which there is no expression of Kit, die around birth from anaemia. However, W~ W v mice, which have a point mutation that reduces but does not abolish the tyrosine kinase activity of Kit, survive until adulthood. W~ W ~' mice lack the ICC in the small intestine that generate rhythmic activity, and they also lack the subpopulation of ICC essential for NO-mediated transmission in the stomach and lower oesophageal and pyloric sphincters (see above; Maeda et al., 1992; Ward et al., 1994a, 1998; Burns et al., 1996). W / W v mice show delayed gastric emptying and abnormal motility patterns (Huizinga et al., 1997; Der-Silaphet et al., 1998), and humans suffering from hypertrophic pyloric stenosis lack ICC within the hypertrophic circular muscle (Vanderwinden et al., 1996). Moreover, mice in which the gene encoding neuronal NOS has been inactivated have a dilated stomach (Huang et al., 1993). Hence, the absence of either ICC or NOS neurones in the stomach appears to be correlated with gastric motor disorders. However, apart from a dilated stomach, neuronal NOS knockout mice show no other gastrointestinal disorders, suggesting that (a) cotransmitters of enteric neurones can fulfil the role of inhibition in the absence of NO, probably because the component of relaxation mediated by NO is not major, except in the stomach, and/or (b) other molecules, such as endothelial NOS or other inhibitory transmitters, have been up-regulated in the neuronal NOS knockout mice to compensate for the absence of neuronal NOS.

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4.4. AXO-AXONAL INTERACTIONS INVOLVING NO The terminals of NOS inhibitory motor neurones and of excitatory motor neurones are in the same axon bundles within the muscle layers. Thus, the nerve terminals of excitatory motor neurones will be exposed to NO. Pharmacological data indicate that NO released from the nerve terminals of inhibitory motor neurones reduces the release of the excitatory co-transmitters, acetylcholine and tachykinins. Thus, inhibition of NOS enhances cholinergic and tachykinin transmission, without increasing responses to exogenous acetylcholine and tachykinins (Knudsen and Tottrup, 1992; Lefebvre et al., 1992; Wiklund et al., 1993b; Hryhorenko et al., 1994; Holzer-Petsche and Moser, 1996; Hebeiss and Kilbinger, 1998). Furthermore, when ODQ was used to prevent the activation of guanylyl cyclase (the target enzyme for NO) cholinergic transmission, tachykinin transmission and acetylcholine release were all significantly increased (Hebeiss and Kilbinger, 1998). Conversely, NO donors inhibit excitatory transmission to the muscle (Baccari et al., 1994; Yunker and Galligan, 1996). Part of this inhibitory effect may be post-junctional, as NO reduces the response of the muscle to tachykinins (Yunker and Galligan, 1996). 4.5. NOS INTERNEURONES WITHIN THE INTESTINE NADPH diaphorase staining and NOS immunohistochemistry have revealed NOS in nerve terminals within enteric ganglia. In the guinea pig, there are few NOS-immunoreactive nerve terminals in ganglia of the oesophagus, stomach and duodenum, but many in the ileum and large intestine (Fig. 6B; Furness et al., 1994). The terminals in the myenteric ganglia arise from myenteric nerve cells whose axons project anally (Costa et al., 1992; McConalogue et al., 1993; Ekblad et al., 1994a; Timmermans et al., 1994; Sang et al., 1997). The majority of these descending interneurones are also immunoreactive for VIP (Costa et al., 1992; Ekblad et al., 1994b; Timmermans et al., 1994; Uemura et al., 1995; Sang et al., 1997). In guinea-pig and mouse intestines, where this has been evaluated, many of the NOS interneurones are also immunoreactive for CHAT, the synthesising enzyme for acetylcholine, and many of their terminals contain the vesicular acetylcholine transporter (Li and Furness, 1998; Sang and Young, 1998a). Evidence in human intestine also suggests that the NOS interneurones are cholinergic (Porter et al., 1997). Application of NO, indirectly from the NO donor nitroprusside, has no effect on the membrane potential of myenteric neurones of the guinea-pig small intestine, and does not affect fast excitatory transmission (Tamura et al., 1993). Furthermore, neither membrane potential, nor fast and slow excitatory transmission are affected by NOS inhibitors. As cholinergic interneurones, which are responsible for some of the fast transmission, contain NOS, these results suggest that NO has a minimal role in modulating neuro-neuronal transmission. However, exogenously applied NO does reduce the amplitudes of slow EPSPs. Pharmacological analyses of enteric reflexes in the guinea-pig small intestine are consistent with NO having little or no role in fast excitatory, neuro-neuronal transmission. In isolated intestine set up in a three-chambered organ bath, so drugs could be separately applied to a region containing synapses of interneurones, transmission from the descending interneurones was unaffected by blocking NOS (Yuan et al., 1995). On the other hand, reflexes were enhanced by inhibiting NOS in the chamber in which intrinsic primary afferent neurones transmit to interneurones (Yuan et al., 1995). The intrinsic primary afferent neurones do not contain NOS, so the authors concluded that NO was released from the cell bodies of the interneurones, and acted retrogradely to reduce the effectiveness of transmission. 237

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Transmission from the intrinsic primary afferent neurones is through slow EPSPs (Furness et al., 1998), so this conclusion is consistent with the observation of Tamura et al. (1993), referred to above, that NO reduces the amplitudes of slow EPSPs. In contrast, endogenous NO seems to have a substantial role in modulating neuro-neuronal transmission in the guinea-pig colon (Smith and McCarron, 1998). In divided organ-bath experiments, these authors found that inhibition of NOS enhanced transmission in descending nerve pathways and increased reflex responses more than 6-fold. 4.6. NOS IN NERVE FIBRES INNERVATING OESOPHAGEAL MOTOR ENDPLATES Motor endplates in the striated muscle of the oesophagus receive a dual innervation, a cholinergic innervation that derives from nerve cells in the nucleus ambiguus in the brain stem, and a local (enteric) innervation from neurones with cell bodies in the myenteric ganglia of the oesophagus (Neuhuber et al., 1994; W6rl et al., 1994; Kuramoto et al., 1996; Sang and Young, 1998b). The motor programs that direct peristalsis during swallowing are mediated entirely through the vagal neurones, and if the vagal innervation is removed, then peristalsis is paralysed (Biancani and Behar, 1995). The enteric neurones that innervate the oesophageal motor endplates contain NOS (Neuhuber et al., 1994; Sang and Young, 1997; Rodrigo et al., 1998), and in the rat oesophagus, they also contain VIP and galanin immunoreactivity (W6rl et al., 1998; Kuramoto et al., 1999). In the mouse, where they are also VIP-immunoreactive, it has been shown that the enteric axons at the motor endplates are not immunoreactive for the vesicular acetylcholine transporter, and therefore not cholinergic (Sang and Young, 1997). This has not been tested in other species. The roles of the NOS fibres at the motor endplates have not been determined, although it has been speculated that they may affect transmission from the vagal fibres both pre- and postsynaptically (W6rl et al., 1998). 4.7. NOS INNERVATION OF THE MUCOSA IN THE STOMACH The numerous publications dealing with the distribution of NOS fibres in the small and large intestine all indicate that the mucosa is not innervated, although in the mouse small intestine there is a transient innervation of the mucosa by NOS nerve fibres during late embryonic and early postnatal stages (Young and Ciampoli, 1998). However, NOS-immunoreactive nerve fibres have been detected in the lamina propria of the rat and human stomach (Ekblad et al., 1994a; Manneschi et al., 1998), and NOS-immunoreactive nerve cell bodies in the myenteric plexus of the guinea-pig stomach are labelled by retrograde transport from the mucosa (Pfannkuche et al., 1998). The innervation of the mucosa is sparse, and at this time there is no data that indicate the role of NO released from nerve fibres in the gastric mucosa. There is evidence that NO released from an endogenous mucosal source has a role in vasodilation and gastric mucosal protection, but it is thought that the source of this NO is the vascular endothelium (Stark and Szurszewski, 1992). 4.8. NOS IN INTESTINOFUGAL NEURONES Intestinofugal neurones have cell bodies in the myenteric ganglia of the small and large intestine, and provide terminals in sympathetic prevertebral ganglia (Szurszewski and Miller, 1994). They are part of the afferent limb of intestino-intestinal inhibitory reflexes. They transmit to cell bodies of sympathetic neurones in prevertebral ganglia via cholinergic fast EPSPs, and immunohistochemical studies have confirmed their cholinergic nature (Mann et al., 1995). 238

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The pharmacological data clearly identify acetylcholine as the primary transmitter of intestinofugal neurones (Crowcroft et al., 1971; Kreulen and Szurszewski, 1979). Intestinofugal neurones also contain several neuropeptides including gastrin releasing peptide, enkephalin, dynorphin and VIP, which probably contribute to slow EPSPs that can be evoked by stimulation of the neurones (Szurszewski and Miller, 1994). In the guinea pig, although none of the intestinofugal neurones that project from the small intestine to the coeliac ganglion contain NOS, over 20-50% of the intestinofugal neurones that project from the large intestine to the coeliac ganglion or the inferior mesenteric ganglion are NOS-immunoreactive (Fig. 6D,E; Domoto et al., 1995; Mann et al., 1995). Hence, in addition to NOS preganglionic neurones (see Section 2), NOS intestinofugal neurones are a major source of NOS nerve terminals in prevertebral ganglia. 4.9. NOS IN THE BILIARY SYSTEM AND PANCREAS NOS nerve cells and fibres are both found in the gallbladder (Talmage and Mawe, 1993; De Giorgio et al., 1994; Grozdanovic et al., 1994: Uemura et al., 1997) and the sphincter of Oddi (Furness et al., 1994; Wells et al., 1995). Many of the NOS cell bodies also express VIE and NOS/VIP nerve terminals innervate the gallbladder muscle. Because both NO and VIP relax the muscle, it seems reasonable to conclude that the NOS neurones supplying the muscle are inhibitory motor neurones, equivalent to those in the stomach and intestine (Mawe, 1998). However, co-localisation studies present a number of puzzles that require resolution. In guinea-pig, dog and human gallbladder, all nerve cells (including the NOS/VIP nerve cells) are ChAT-immunoreactive, and are therefore likely to be cholinergic. If this is so, it is necessary to determine how inhibition of the gallbladder muscle can be mediated by neurones that contain both inhibitory and excitatory transmitters. It is presumed that the NOS innervation of the mucosa is involved in the control of water and electrolyte transport, but whether NO or another substance is the primary transmitter remains unresolved. NOS has also been detected in nerve cell bodies and fibres in the pancreas of cat, dog, guinea pig, human and rat (Shimosegawa et al., 1993; Kirchgessner et al., 1994; Sha et al., 1995; Umehara et al., 1997). A high proportion of nerve cells in pancreatic ganglia contain NOS, and NOS nerve fibres are found in the ganglia, and around pancreatic acini and ducts. There are also a small number of NOS nerve fibres in the islets, but islet cells are not reactive for NOS. Exogenously applied NO hyperpolarises pancreatic neurones and elicits fast EPSPs, and inhibition of NOS increases the amplitudes of slow EPSPs (Sha et al., 1995). The pancreatic ganglia receive innervation from the vagus nerve, and when the vagus is stimulated there is pancreatic exocrine secretion that has a prominent non-cholinergic component, that is thought to be mediated by VIP (Holst et al., 1984). Inhibition of NOS by L-NAME reduced pancreatic secretion caused by vagus nerve stimulation by 20% and that evoked by VIP was reduced by 50% (Holst et al., 1994). The NO donor, sodium nitroprusside, increased secretion, but did not increase VIP release. These data suggest that NO is a transmitter of pancreatic secretomotor neurones, and that there is co-operativity between NO and VIE NO may also influence the endocrine pancreas as NOS inhibitors have been shown to stimulate insulin secretion from the pancreas in vitro (Jansson and Sandler, 1991).

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5. ROLE OF NO IN THE NEURAL CONTROL OF THE TRACHEA AND LOWER AIRWAYS

5.1. TYPES OF NEURONES INNERVATING THE TRACHEA AND LOWER AIRWAYS The trachea and bronchi are innervated by extrinsic sensory nerve fibres that arise primarily in vagal sensory (jugular and nodose) ganglia and upper thoracic spinal ganglia, sympathetic nerve fibres that arise from postganglionic neurones in the stellate and superior cervical ganglia, and parasympathetic preganglionic nerve fibres that arise from the dorsal motor nucleus of the vagus nerve and synapse on postganglionic neurones located within the airways or within the oesophagus (see below). Within the airways, the respiratory epithelium, vasculature, exocrine glands and ducts and the smooth muscle are innervated by autonomic and extrinsic sensory nerve fibres, but the density of sympathetic axons (identified by formaldehyde-induced fluorescence or tyrosine hydroxylase immunoreactivity) within the smooth muscle varies enormously between species (see Lundberg et al., 1988). In humans and guinea pigs, ChAT-immunoreactive nerve terminals are present in the airway smooth muscle and associated with the exocrine glands (Fischer et al., 1996b; Canning and Fischer, 1997). The cholinergic nerve terminals appear to arise from neurones located in small ganglia within the trachea and bronchi, since acetylcholine-mediated responses to nerve stimulation are observed in guinea-pig tracheas grown in organotypic culture for 2 days, when nerve fibres of extrinsic origin would have degenerated (Canning et al., 1996). Moreover, in the guinea pig, all of the intrinsic neurones within the trachea and bronchi are ChAT-immunoreactive (Fischer et al., 1996b; Canning and Fischer, 1997), although in humans, there is a small proportion of intrinsic neurones that is ChAT-negative (Fischer et al., 1996b). Consistent with the anatomical studies, a large number of physiological and pharmacological studies have shown that acetylcholine is the predominant neurotransmitter mediating contraction of airway smooth muscle and glandular secretion, and it also plays a role in the regulation of the airway vasculature (Widdicombe, 1963, 1993; Richardson and Beland, 1976; Laitinen and Laitinen, 1987; Tokuyama et al., 1990; Davis and Tseng, 1991; Matran, 1991; Fung et al., 1992; Coleridge and Coleridge, 1994; Reinheimer et al., 1996). The presence of VIP in nerve fibres innervating the airways has been known for many years (Matsuzaki et al., 1980; Dey et al., 1981; Laitinen et al., 1985). In the guinea pig, the VIP-immunoreactive nerve terminals within the smooth muscle also contain neuropeptide Y (Bowden and Gibbins, 1992). NOS (localised by immunohistochemistry and NADPH diaphorase histochemistry) is also present in nerve fibres innervating the smooth muscle and the airway vasculature of guinea pigs, humans and ferrets, but the presence of NOS in nerve fibres associated with the submucosal glands and the lamina propria varies between species (Dey et al., 1993; Fischer et al., 1993; Fischer and Hoffmann, 1996). NOS and VIP co-exist in the same nerve fibres in the airway smooth muscle of humans, guinea pigs and ferrets (Dey et al., 1993; Shimosegawa and Toyota, 1994; Fischer et al., 1996b). In the guinea pig, the NOS/VIP/neuropeptide Y nerve fibres are a completely separate population of nerve fibres from those containing CHAT, and in humans, only a small proportion of the ChAT-immunoreactive nerve fibres is also immunoreactive for NOS and VIP (Fischer et al., 1996b). The presence of NOS in nerve cell bodies in ganglia within the airways varies between species. In the guinea pig, none of the intrinsic neurones, which are all ChAT-positive (see above), contain NOS, VIP or neuropeptide Y (Bowden and Gibbins, 1992; Kummer et al., 1992; Fischer et al., 1993, 1996b; Shimosegawa and Toyota, 1994; Canning et al., 1996; Canning and Fischer, 1997). However, in humans, 60-80% (depending on the 240

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location) of the intrinsic neurones in the lower airways show NADPH diaphorase staining and NOS immunoreactivity (Fischer and Hoffmann, 1996), and some of the NOS-immunoreactive neurones also contain ChAT (Fischer et al., 1996b). In the ferret and pig, nerve cell bodies showing both NOS and VIP immunoreactivity are also found in ganglia within the trachea (Dey et al., 1993; De Rada et al., 1993). 5.2. PHARMACOLOGY OF TRANSMISSION AND SOURCE OF NOS NEURONES IN THE TRACHEA An inhibitory innervation of the airway smooth muscle, that is not mediated by either noradrenaline or acetylcholine, has been demonstrated in many species (e.g. Campbell, 1971; Coburn and Tomita, 1973; Richardson and Beland, 1976; Cameron et al., 1983), but for many years the identity of the neurotransmitter(s) was unclear. Pharmacological studies initially identified a role for VIP in the vagally mediated relaxations, because the nerve-mediated relaxations are reduced after the tissue had been incubated with a VIP antibody or by immunisation to VIP (Matsuzaki et al., 1980; Cameron et al., 1983; Ellis and Farmer, 1989). Following the identification of NO as a neurotransmitter, it was very soon shown that NO also mediates a component of the relaxations, because the relaxations induced by electrical field stimulation are significantly reduced by NOS inhibitors in human, pig, guinea-pig and bovine airways (Tucker et al., 1990; Li and Rand, 1991; Belvisi et al., 1992; Ellis and Undem, 1992; Kannan and Johnson, 1992; Bai and Bramley, 1993; Takahashi et al., 1995). Hence in most species in which it has been investigated, the non-noradrenergic relaxations of the airway smooth muscle appear to be mediated by a combination of NO and VIP, although in the cat, VIP may have a minimal role (Fisher et al., 1993). In the guinea pig, the NOS/VIP neurones mediating the relaxation of the tracheal muscle do not arise from neurones within the trachea (see above), and it appears that they arise from the myenteric plexus of the oesophagus, the stellate ganglion, and possibly also from vagal sensory ganglia. The first evidence that neurones innervating the trachea might arise from the oesophagus was the demonstration that vagally mediated relaxations of isolated preparations of guinea-pig trachea cannot be elicited when the trachea is separated from the adjoining oesophagus, whereas vagally mediated contractions (mediated by acetylcholine) are unaffected by removal of the oesophagus (Canning and Undem, 1993a,b). It was later shown that tracheas maintained in organ culture, with the adjacent oesophagus intact, display relaxations that are mediated by NO and VIP (see above), and possess NOSand VIP-immunoreactive nerve fibres at a similar density to that seen in control animals (Canning et al., 1996). However, when tracheas are grown in organ culture without the oesophagus, NO- and VIP-mediated relaxations are not observed, and the density of NOSand VIP-immunoreactive nerve fibres is less than 20% that observed in control animals (Canning et al., 1996). In addition, following application of DiI to the trachealis muscle and growth in organotypic culture for 2 days, labelled nerve cell bodies showing NOS, VIP and neuropeptide Y immunoreactivity are observed in the myenteric plexus of the oesophagus (Fischer et al., 1998; Moffatt et al., 1998). Some of the NOS/VIP/neuropeptide Y nerve fibres within the smooth muscle of the guinea-pig trachea also arise from non-noradrenergic sympathetic neurones, since following application of DiI to the tracheal smooth muscle, labelled nerve cell bodies showing immunoreactivity to neuropeptide Y, VIP and NOS, but not tyrosine hydroxylase, are observed in the stellate ganglion (Bowden and Gibbins, 1992; Fischer et al., 1996a). NOS-immunoreactive cell bodies are common in the vagal sensory ganglia supplying the airways of guinea pigs, and a retrograde tracing study has shown that 241

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5-20% of neurones in the nodose and jugular ganglia that project to the trachea contain NOS (Fischer et al., 1993, 1996a). Thus, some of the NOS terminals within the trachea may also originate from sensory ganglia. The origin of the NOS/VIP nerve fibres in the airways has not been examined experimentally in species other than the guinea pig. However, in humans, ferrets and pigs, there are many intrinsic NOS-containing nerve cell bodies in ganglia within the airways (Dey et al., 1993: De Rada et al., 1993: Fischer and Hoffmann, 1996), and therefore it is almost certain that, unlike guinea pigs, many of the NOS/VIP nerve fibres arise from intrinsic neurones. In summary, NO together with VIP are released from the same nerve fibres to mediate relaxation of the airway smooth muscle. The origin of the NOS/VIP nerve fibres within the airways varies between species. In the guinea-pig airways, unlike the cholinergic parasympathetic nerve fibres which arise from intrinsic neurones, most of the NOS/VIP nerve fibres arise from nerve cells in the myenteric plexus of the oesophagus, but some also arise from non-noradrenergic sympathetic neurones in the stellate ganglion: none of the NOS/VIP nerve fibres arise from intrinsic neurones. However in humans, pigs and ferrets, some of the NOS/VIP nerve fibres are likely to arise from intrinsic neurones, since there are many NOS neurones within intrinsic ganglia of the airways of these species, but other sources have yet to be examined.

6. ROLE OF NO IN THE NEURAL CONTROL OF SALIVARY GLANDS AND OTHER SECRETORY TISSUES 6.1. SALIVARY GLANDS

6.1.1. Types of neurones innervating the salivary glands The salivary glands are innervated by nerve fibres arising from extrinsic sensory, parasympathetic and sympathetic neurones. The parasympathetic and sympathetic nerve fibres innervate the acini, ducts and blood vessels, but the sensory fibres innervate only the blood vessels and ducts (see Lundberg et al., 1988). VIP is present in parasympathetic, cholinergic nerve fibres that innervate the salivary glands (Wharton et al., 1979; Lundberg et al., 1980; Uddman et al., 1980). In the parotid, submandibular and sublingual glands of the ferret, cat, pig and rat, NOS-immunoreactive nerve terminals are associated with both the secretory (Fig. 7D) and vascular components, and NOS nerve cell bodies are present in the hilar regions of the submandibular (Fig. 7A) and sublingual, but not the parotid glands (Grozdanovic et al., 1992; Alto et al., 1993, 1997; Ceccatelli et al., 1994: Modin et al., 1994; Lohinai et al., 1995). Many of the NOS neurones in the submandibular gland also contain VIP (Fig. 7A,B; Ceccatelli et al., 1994; Modin et al., 1994), and thus these parasympathetic neurones are cholinergic/NOS/VIP neurones (Fig. 7C,D). The NOS nerve fibres in the submandibular and sublingual glands probably arise from local neurones, whereas those in the parotid gland arise from the otic ganglion because lesioning the auriculo-temporal nerve (the pathway by which the neurones in the otic ganglion project to the parotid gland) causes an almost total disappearance of NOS nerve fibres from this gland (Alm et al., 1997).

6.1.2. Role of NO in vasodilation and secretion in the salivary glands Stimulation of parasympathetic nerves induces secretion of saliva and vasodilation. In 1872, Heidenhain demonstrated that the two responses have different pharmacological properties 242

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A

Fig. 7. (A, B) Paired micrographs of a frozen section through a small ganglion associated with the hilus of the submandibular salivary gland of a rat. Almost all of the neurones are both NOS- (A) and VIP-immunoreactive (B). Scale bar = 50 p.m. (C, D) The local, parasympathetic neurones are also known to be cholinergic, and the NOS-positive nerve terminals (arrows) in the sublinguai glands are also vesicular acetylcholine transporter (VAChT)-immunoreactive. VAChT is exclusively found in cholinergic nerve terminals. Scale bar = 25 ~tm.

because atropine blocks or substantially reduces (depending on the species) the secretory response induced by stimulation of preganglionic parasympathetic nerve fibres, but the vasodilation is largely unaffected (see Emmelin, 1967; Lundberg et al., 1988). Hence the secretory response is primarily mediated by acetylcholine and the vasodilatory response by another neurotransmitter. In the cat, VIP is co-released along with acetylcholine from parasympathetic nerve terminals (Bloom and Edwards, 1980). Although acetylcholine is the primary neurotransmitter mediating salivary secretion, VIP enhances the secretion of saliva induced by acetylcholine (Lundberg et al., 1980). However, because the flow of saliva is correlated with blood flow, and because substances secreted from epithelial cells can cause vasodilation, it can be difficult to distinguish the role of NO in salivary secretion from that in vasodilation. The effect of NOS inhibitors on the acetylcholine-induced secretory response appears to differ in different 243

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species. In the ferret, NOS inhibitors reduce the acetylcholine-mediated secretory response, and it was suggested that the VIP-mediated enhancement of the fluid response to acetylcholine is dependent on NO (Tobin et al., 1997). However, in the rat, the NOS inhibitor, NOLA, enhances the salivary fluid secretory response induced by carbachol, suggesting that NO acts postsynaptically on the secretory cells to reduce the secretion of saliva induced by acetylcholine (Lohinai et al., 1997). In the cat, L-NAME does not affect the secretory response induced by acetylcholine (Edwards and Garrett, 1993). The non-cholinergic vasodilation observed in the salivary glands in response to parasympathetic nerve stimulation appears to be mediated mainly by VIP and PACAP (Lundberg et al., 1980; Lundberg et al., 1982; Tobin et al., 1995). Several studies have shown that NO also plays a role in the vasodilation of the submandibular gland, and although studies using isoform-specific NOS inhibitors have not yet been performed, endothelial-derived NO appears to be the main source of NO. There is, however, some evidence that neurally derived NO can facilitate the release of VIP (see below). In the cat, the vasodilator response to exogenous VIP is abolished by large doses of L-NAME whereas that to acetylcholine is unaffected; the most likely explanation is that VIP induces release of NO from the endothelium (Edwards and Garrett, 1993). However, because NOS inhibitors reduce the amount of VIP released by parasympathetic nerve stimulation, it appears that NO can also act presynaptically to facilitate VIP release; in contrast to VIE the effects of acetylcholine are independent of NO (Buckle et al., 1995). In the ferret, the vasodilation is mediated by acetylcholine, VIP and possibly PACAP. The actions of acetylcholine are independent of NO, but the action of VIP depends on the postsynaptic (presumably endothelial) release of NO (Tobin et al., 1997). In the pig, both the acetylcholine- and VIP-evoked vasodilation are mediated by NO (Modin et al., 1994), although it is unclear whether neurally or endothelial-derived NO is involved. 6.2. SWEAT GLANDS The sweat glands of rats and cats are concentrated in the front and hind paw pads and do not receive an innervation by sympathetic noradrenergic nerve fibres (see Landis, 1988). The glands are, however, innervated by sympathetic cholinergic nerve fibres that arise from the stellate ganglia (front paw pads) and lower lumbar paravertebral sympathetic ganglia (hind paw pads). These sympathetic cholinergic nerve fibres also contain VIP and CGRP (Lundberg et al., 1979; Landis and Fredieu, 1986; Lindh et al., 1988), but the role of the peptides is unclear, because sweat secretion can be blocked by atropine (Randall and Kumura, 1955; Weiner and Hellmann, 1959; Stevens and Landis, 1987), as in humans. In the rat, the cholinergic/VIP/CGRP fibres innervating sweat glands do not contain NOS, and none of the neurones in the stellate ganglion retrogradely labelled from the front paw pad contain NOS (S.M. Murphy, pers. commun.). However, the VIP-containing sympathetic nerve fibres innervating the sweat glands of cats do contain NOS (Anderson et al., 1995), and like the neuropeptides, the role of NO, which is presumably released from these fibres, is unclear. Nerve fibres associated with the sweat glands of humans also show NADPH diaphorase staining (Schulze et al., 1997). 6.3. THE NASAL MUCOSA In addition to sympathetic and extrinsic sensory neurones (originating from the superior cervical and trigeminal ganglia, respectively), the glands and blood vessels in the nasal mu244

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cosa are innervated by parasympathetic neurones that arise from the sphenopalatine ganglion (see Lundberg et al., 1988). A very early study showed that electrical stimulation of the sphenopalatine ganglion causes nasal secretion (by Prevost in 1868; see Lundberg et al., 1988). Subsequent studies showed that stimulation of the vidian nerve (which contains both parasympathetic preganglionic and sympathetic postganglionic nerve fibres) elicits secretion which can be blocked by atropine (Eccles and Wilson, 1973). Low-threshold stimulation of the vidian nerve, which avoids activating sympathetic postganglionic fibres, also evokes a vasodilation that is atropine-resistant (Malcolmson, 1959). The postganglionic parasympathetic nerves innervating the nasal mucosa contain acetylcholine and VIP (Lundberg et al., 1980), and, as in the submandibular glands, acetylcholine appears to be the main neurotransmitter mediating secretion, and VIP the major transmitter mediating vasodilation (Lundberg et al., 1981). NOS-immunoreactive and NADPH diaphorase-positive nerve fibres have been reported in the submucosal glands, the arterioles and the sub-epithelial layer of the nasal mucosa of a range of species including rat, human and dog (Hanazawa et al., 1993, 1994; Ceccatelli et al., 1994; Kulkarni et al., 1994; Lee et al., 1995; Jeon et al., 1997; Lacroix et al., 1998). Many of the NOS nerve fibres also contain VIP (Ceccatelli et al., 1994; Lee et al., 1995; Lacroix et al., 1998), and in the human, there is occasionally overlap with substance P and tyrosine hydroxylase (Tasman et al., 1998). In the rat, neurones in the sphenopalatine ganglion that are retrogradely labelled from the nasal mucosa are NADPH diaphorase-positive, whereas those in the trigeminal and superior cervical ganglia are NADPH diaphorase-negative (Jeon et al., 1997). Hence, in the nasal mucosa, parasympathetic nerve fibres contain acetylcholine, VIP and NOS and they arise from the sphenopalatine ganglion. Isoform-specific NOS inhibitors have not yet been used in pharmacological studies of the parasympathetic vasodilation in the nasal mucosa. NOS inhibitors have been shown to reduce (Lacroix et al., 1998) or abolish (Watanabe et al., 1995) the atropine-resistant vasodilation in the nasal mucosa of the dog. However, it is possible that at least part of the vasodilation is dependent on NO derived from the endothelium. Although acetylcholine is the primary mediator of secretion, in some species there is an atropine-resistant component. In the dog, the NOS inhibitor, L-NNA, reduces the atropine-resistant secretory response (Lacroix et al., 1998), but this may be a secondary response to the effects of NOS inhibition on the vasculature. Therefore, despite the presence of NOS in parasympathetic nerve fibres in the nasal mucosa, there is still no conclusive evidence that neurally derived NO is a major neurotransmitter in this tissue.

7. ROLE OF NO IN THE INNERVATION OF THE ADRENAL MEDULLA

7.1. PRESENCE OF NOS IN NEURONES IN THE ADRENAL MEDULLA The adrenal medulla is composed of clumps of noradrenaline- and adrenaline-containing chromaffin cells which are directly innervated by cholinergic sympathetic preganglionic neurones. In the rat, studies that have investigated the presence of NOS in the preganglionic cell bodies, (Grkovic and Anderson, 1997) and nerve terminals (Holgert et al., 1995), have shown that all of the preganglionic neurones innervating the adrenal medulla also contain NOS. In contrast, enkephalin is present in the cholinergic/NOS nerve terminals that innervate the adrenaline-containing medullary cells, but not in the cholinergic/NOS terminals that innervate the noradrenaline-containing medullary cells (Pelto-Huikko et al., 1985; Holgert et 245

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al., 1994). The proportion of sympathetic preganglionic neurones that project to the adrenal and that contain NOS has not been examined in species other than the rat. The adrenal is also innervated by a small number of intrinsic neurones, and in many species, NOS is present in a subpopulation of intrinsic neurones (Bredt et al., 1990; Dawson et al., 1991; Afework et al., 1992; Alm et al., 1993; Brtining, 1994; Ceccatelli et al., 1994; Afework and Burnstock, 1995; Marley et al., 1995; Cracco et al., 1996). In the rat, many of the intrinsic NOS neurones also contain VIE but not CHAT, and many of the NOS/VIP cell bodies and fibres are associated with blood vessels within the adrenal medulla (Afework and Burnstock, 1995; Holgert et al., 1995). In addition to preganglionic and intrinsic neurones, some of the NOS fibres in the adrenal also arise from dorsal root ganglia (Heym et al., 1995). As well as several possible neuronal sources of NO, NOS is also found in endothelial cells and there have been some reports of NOS immunoreactivity in the catecholamine-containing chromaffin cells within the adrenal medulla (Dun et al., 1993a; Heym et al., 1994; Schwarz et al., 1998). 7.2. ROLE OF NEURALLY DERIVED NO IN THE ADRENAL MEDULLA At the ultrastructural level, nerve terminals containing NOS have been found in close contact with both types of chromaffin cells and with vascular smooth muscle (Tanaka and Chiba, 1996), and thus neurally released NO could potentially affect both catecholamine secretion and adrenal blood flow. Putative targets of NO in the adrenal glands of mice, rats and guinea pigs have also been examined by localising cGMP immunoreactivity following stimulation with a NO donor (Holmberg et al., 1998). Complementing the ultrastructural location of NOS-containing nerve terminals, the NO donor, sodium nitroprusside, induced increases in cGMP immunoreactivity in both noradrenaline- and adrenaline-containing chromaffin cells and in the walls of blood vessels. However, the adrenaline cells required higher concentrations of sodium nitroprusside than the noradrenaline cells before increases in the levels of cGMP immunoreactivity could be detected. The nerve-stimulated release of catecholamines is mediated by acetylcholine acting on nicotinic receptors on the chromaffin cells. Studies that have examined whether NO has a major role in nerve-stimulated catecholamine secretion are inconsistent. In the anaesthetised dog, inhibition of NOS was reported to have no effect on the release of adrenaline caused by nerve stimulation (Breslow et al., 1993). Similarly, in the isolated bovine adrenal, nerve-stimulated release of adrenaline and noradrenaline was reported to be unaffected by NOS inhibitors and NO donors (Marley et al., 1995). However, later studies, again in anaesthetised dogs, reported that both the nerve stimulation- and acetylcholine-induced increases in catecholamine secre-

Fig. 8. Summary diagram showing the main locations in which NOS neurones are found in the peripheral autonomic nervous system of small laboratory mammals, such as the rat and guinea pig. Note that in other mammalian species, the distribution of NOS neurones may be different. The major differences are as follows. (i) NOS-containing nerve terminals are not common in the sphenopalatine and submandibular ganglia in rats {see Fig. 3C, and Ceccatelli et al., 1994), and thus there appear to be very few NOS parasympathetic preganglionic neurones projecting to these ganglia in the rat. However, NOS is present in the vast majority of cranial parasympathetic preganglionic neurones projecting to the sphenopalatine and submandibular ganglia in the rabbit (see Zhu et al., 1996, 1997). (ii) NOS neurones are extremely rare in the paravertebral and prevertebral sympathetic ganglia of rats, mice and guinea pigs, but are common in larger mammals such as cats tsee Fig. 2C) and humans. (iii) In the guinea pig, none of the NOS neurones innervating the trachea are found within the trachea, and most arise from the oesophagus. However in humans, ferrets and pigs, NOS neurones are present within the airways.

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tion are inhibited by NO donors (Nagayama et al., 1998a,b), suggesting that NO may play an inhibitory role in the regulation of acetylcholine-mediated catecholamine secretion. Consistent with these data, NOS inhibitors were found to enhance the catecholamine secretion induced by acetylcholine (Nagayama et al., 1998b). However, NOS inhibitors reduced the secretion in-

.~essels

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duced by nerve stimulation, and it was suggested that NO could also be acting presynaptically to facilitate the release of acetylcholine from preganglionic nerve terminals, and that when catecholamine secretion is evoked by nerve stimulation, the presynaptic excitatory action of NO is dominant over the inhibitory action on secretion (Nagayama et al., 1998b). It is possible that the discrepancies between the results obtained from different laboratories may be due to different actions of NO (presynaptic excitation and postsynaptic inhibition) being dominant under different experimental conditions. Catecholamines spontaneously efflux from chromaffin cells under basal conditions, and studies that have examined whether NO plays a role in regulating this efflux are also not in agreement. In the dog, one study found that NOS inhibitors increase, and NO donors decrease, the basal efflux of catecholamines, suggesting that endogenous NO inhibits basal catecholamine secretion (Ward et al., 1996), whereas using the same species, another group found that basal catecholamine secretion was unaffected by NO donors and NOS inhibitors (Nagayama et al., 1998b). The effect of NO donors and NOS inhibitors on the acetylcholine-evoked catecholamine secretion from cultured bovine adrenal medullary cells has been examined in a number of studies, and again, the results differ markedly. For example, one study reported that NOS inhibitors inhibit acetylcholine-induced catecholamine secretion (Uchiyama et al., 1994), suggesting that NO may facilitate the acetylcholine-induced secretion, whereas other studies have found that inhibition of NOS enhances K +- and acetylcholine-stimulated catecholamine secretion, and NO donors inhibit acetylcholine-induced secretion, indicating that NO may inhibit acetylcholine-induced catecholamine secretion (Torres et al., 1994; Rodriguez-Pascual et al., 1996). NO, of unknown origin, does appear to have a role in the regulation of medullary blood flow. In the dog, inhibition of NOS reduces resting medullary blood flow and blocks the nerve-stimulated increase in blood flow (Breslow et al., 1993), and in the rat, NOS inhibitors decrease the rate of perfusion through the adrenal (Hinson et al., 1996). However, in both studies, the source of NO mediating the vasodilation was not determined; it could arise from the endothelium or from one of several neuronal sources.

8. OVERVIEW OF PERIPHERAL AUTONOMIC NO NEURONES Neurones capable of synthesising NO are widespread in the peripheral autonomic nervous system (Fig. 8). They form significant subpopulations of sympathetic preganglionic, parasympathetic preganglionic, parasympathetic postganglionic, vagal and spinal sensory, and enteric neurones. NOS is always found co-localised with other potential neurotransmitters: NOS neurones innervating non-vascular smooth muscle usually also contain VIE but not ChAT (for example, enteric inhibitory neurones and non-noradrenergic airway dilator neurones), and NOS neurones innervating other neurones and vascular smooth muscle are often cholinergic (for example, sympathetic and parasympathetic preganglionic neurones, enteric interneurones, cerebral vasodilator neurones). There is excellent evidence that NO acts as neurotransmitter to relax both vascular and non-vascular smooth muscle in a variety of locations. However, NO usually only mediates a component of the inhibitory transmission to the muscle, and rarely is NO the sole neurotransmitter. The role of NO at peripheral neuro-neuronal synapses is unclear, because most of the NOS-containing neurones that innervate other neurones are cholinergic neurones, and acetylcholine is the primary neurotransmitter. Although, NOS neurones are widespread and the participation of NO in transmission is common in the 248

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p e r i p h e r a l n e r v o u s s y s t e m , n e u r o n a l N O S k n o c k o u t m i c e not only survive, but r e p r o d u c e ( H u a n g et al., 1993). T h e only a b n o r m a l i t i e s related to the p e r i p h e r a l a u t o n o m i c n e r v o u s s y s t e m are a dilated s t o m a c h and a v o i d i n g d i s o r d e r due to urinary b l a d d e r - u r e t h r a l s p h i n c t e r d y s f u n c t i o n ( H u a n g et al., 1993; B u r n e t t et al., 1997a). The small n u m b e r of a b n o r m a l i ties o b s e r v e d in m i c e l a c k i n g n e u r o n a l N O S p r o b a b l y reflect the facts that (i) N O rarely acts as the sole n e u r o t r a n s m i t t e r , and (ii) o t h e r m o l e c u l e s (e.g. e n d o t h e l i a l N O S ) are likely to have b e e n u p - r e g u l a t e d in the k n o c k o u t m i c e to c o m p e n s a t e for the lack of n e u r o n a l NOS.

9.

ACKNOWLEDGEMENTS

T h e a u t h o rs ' w o r k is s u p p o r t e d by the N at i onal H e a l t h and M e d i c a l R e s e a r c h C o u n c i l of Australia (NHMRC).

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The nitric oxide system in the urogenital tract M.S. DAVIDOFF AND R. MIDDENDORFF

1. THE URINARY TRACT 1.1. THE UPPER URINARY TRACT

1.1.1. The kidney Nitric oxide plays a crucial role in the regulation of kidney function and metabolism. Nitric oxide has been shown to be important for the regulation of the basal renal, glomerular and medullar hemodynamics, the myogenic and tubuloglomerular feedback response as well as the pressure natriuresis and extracellular fluid volume. Nitric oxide is involved in the pathogenesis of numerous experimental and pathological renal malfunctions and diseases (Kone, 1997). Some more or less detailed reviews on this topic have already been published (Navar et al., 1996; Ito, 1997; Kone, 1997; Kone and Baylis, 1997; Mattson et al., 1997; Star, 1997; Navar, 1998). In the kidney, as in other organs, NO, which in turn activates soluble guanylyl cyclase (sGC), is produced from L-arginine by the nitric oxide synthase (NOS) enzyme family. Early studies have shown the expression in kidney of the three well-known NOS isoforms (Hecker et al., 1991; Mohaupt et al., 1994a), namely the constitutive neuronal and endothelial NOS forms as well as the inducible or macrophageal NOS. In the present review the constitutive neuronal form is referred to as NOS-I, the constitutive endothelial form as NOS-III and the inducible macrophageal isoenzyme as NOS-II. According to Star (1997) of the five NOS isoforms four exist in the kidney (two NOS-II isoenzymes). Moreover, all four sGC subunits, the intracellular NO receptor, have been established in the structural components of the kidney of different species (Star, 1997). Recent findings provide evidence for the existence of additional neuronal and inducible NOS isoforms in the kidney (see Kone and Baylis, 1997 for review). Interesting in this respect is a protein that exhibits similar catalytic properties as the NOS-I, termed nNOS-II or nNOS-Ix (Magee et al., 1996; Silvagno et al., 1996). Also two iNOS isoforms that are expressed differentially along the rat (but not human) nephron and termed VSM-NOS (Vascular Smooth Muscle cells NOS) and MAC-NOS (MACrophage NOS) were described (Mohaupt et al., 1994b). Experimental studies showed that VSM-NOS plays a protective role, and MAC-NOS plays a destructive role in the pathogenesis of postischemic renal failure in rats (Kone and Baylis, 1997). Recently, an unusual NO- and citrulline-producing NOS isoform, that possesses different properties in comparison with the well-known NOS isoforms, was found in the rat kidney by Singh et al. (1997). In addition, a new NOS-I mRNA, product of alternative splicing, that lacks exon 2 was established in the kidney by Oberbaumer et al. (1998). Thus, the different structural components of the kidney Handbook of Chemical Neuroanatom.~; Vol. ! 7: Functional Neuroamm,m v of the Nitric Oxide Sw~tern H.W.M. Steinbusch. J. De Vente and S.R. Vincent. editor,, 2000 Elsevier Science B.V. All rights reserved.

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contain specific NOS isoforms which may exert variable functional effects. Biochemical studies showed that the renal medulla produces the greatest amount of NO (Nava et al., 1996; Zou and Cowley, 1997). Correspondingly, the activity of the NOS in the kidney medulla was three times higher than the NOS activity of the renal cortex (McKee et al., 1994). Distribution of NADPH-diaphorase activity in the kidney. Numerous studies using the histochemical visualization of NADPH-d reported about the existence of good accordance between the distribution of this enzyme activity and the immunocytochemical location of the NOS proteins (Dawson et al., 1991" Schmidt et al., 1992). However, for some structures in the kidney probably not all NADPH-d enzyme activity is related to the NOS isoforms. This is evident, at least for the mouse kidney, where discrepancy between the biochemical results mentioned above and the intensity of NADPH-d enzyme activity exists. We found stronger NADPH-d staining intensity in the cortex (especially in the labyrinth) whereas biochemical measurements have found higher NOS activity in the medulla. The NADPH-d enzyme activity showed a characteristic distribution pattern in the kidney of male and female rats, hen, mice, hamsters, gerbils, guinea pigs, marmosets and humans. A moderate to strong NADPH-d enzyme activity was found in the components of the renal corpuscle such as the epithelial cells of the glomerulus and the endothelial cells of the glomerular capillaries, the endothelium and smooth muscle cells of the afferent arteriole and the endothelium of the efferent arteriole (Nakos and Gossrau, 1994; Grozdanovic and Gossrau, 1995; Grozdanovic et al., 1995). In addition, NADPH-d enzyme activity was found in structures of the juxtaglomerular apparatus (JGA). The juxtaglomerular apparatus comprises cells of the afferent arteriole, the macula densa cells of the distal tubule and the extraglomerular mesangium cells situated at the vascular pole of the renal glomerulus. The juxtaglomerular cells are myoepithelial cells which are located in the wall of the terminal part of interlobar arteries and the juxtaglomerular part of the afferent arteriole (Davidoff and Schiebler, 1981). In addition to renin, these cells in some species showed distinct NADPH-d enzyme activity (Grozdanovic et al., 1995; Yanagisawa et al., 1998). Also cells of the macula densa (MD), a row of specialized cells located at the junction of the ascending limb of the Henle loop and the distal convoluted tubule, exhibited moderate to strong NADPH-d enzyme activity (Mundel et al., 1992" Wilcox et al., 1992; Grozdanovic and Gossrau, 1995; Nadaud et al., 1998). Both, the proximal and the distal parts of the nephron tubules (Fig. 1) showed distinct NADPH-d activity, which was qualified as especially strong in the medullar thick ascending limb segments of the rat kidney (McKee et al., 1994). Similarly, in the intercalated cells (not in the principal cells) of the cortical (Fig. 2) and medullar collecting ducts a strong to moderate enzyme activity was established in the rat (Nakos and Gossrau, 1994; Bachmann et al., 1995; Grozdanovic and Gossrau, 1995; Grozdanovic et al., 1995). As mentioned, the visualization of NADPH-d enzyme activity provides only a general information on the existence of NOS in an organ. It must be emphasized that existence, location and intensity of the NADPH-d enzyme activity depend on the applied technique and the investigated species (Grozdanovic and Gossrau, 1995" Grozdanovic et al., 1995). In addition, sex-related differences must be taken into consideration (Neugarten et al., 1997). The exact localization of the different NOS isoforms was achieved by immunocytochemistry using specific monoclonal and polyclonal antibodies. However, some of the results reported must be accepted with caution because of the possibility of false positive results. Immunohistochemical detection of the reactivity of diverse NOS isoforms has shown characteristic distribution patterns in the kidney of different species. 268

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Fig. 1. Mouse kidney. NADPH-d enzyme activity in renal cortex. The proximal convoluted tubules show a stronger enzyme activity in comparison with the epithelial cells of the distal tubules. Bar -- 20 ~tm. Fig. 2. Mouse kidney. NADPH-d enzyme activity in renal cortex. The proximal convoluted tubules show a stronger enzyme activity in comparison with the epithelial cells of the distal tubules. The arrow points to positive intercalated cells of a cortical collecting duct. Bar -- 20 I_tm. Fig. 3. Immunoreactivity for the NOS-I isoform. Cells of the macula densa (arrow) of a rat kidney. G = glomerulus. Bar = 20 gm. Fig. 4. Immunoreactivity for the NOS-I isoform. Human ureter. Immunoreactivity in the superficial cells of the urothelium. L = lumen. Bar = 20 ~tm.

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Localization of the neuronal NOS-I isoform in the kidner. Immunoreactivity for the neuronal NOS-I isoform was found in some structural components of the nephron in rat, rabbit, mice, gerbil, marmoset and human kidney" cells of the glomerular visceral epithelium, endothelium of the afferent and efferent arterioles, cells of the parietal layer of Bowman's capsule, epithelial cells in the proximal and distal tubules and the collecting ducts (Nakos and Gossrau, 1994; Bachmann et al., 1995" Grozdanovic and Gossrau, 1995; Grozdanovic et al., 1995). Immunoreactivity for NOS-I was also shown in a combined light- and electron-microscopic study being localized within the cytoplasm and the nuclei of cells of the inner medullar collecting duct (Roczniak et al., 1999). Recent studies provide RT-PCR evidence for the existence in the principal cells of the cortical collecting ducts of NOS-I in rats on a high K + diet. However, only some principal cells showed immunoreactivity for the neuronal isoform" the remaining principal cells were negative (Wang et al., 1998). Of all these locations, the NOS-I immunoreactivity in the cells of the juxtaglomerular apparatus is especially interesting. The predominant distribution of NOS-I in the rat macula densa cells was shown by Mundel et al. (1992). Both NADPH-d enzyme activity and NOS-I immunoreactivity (Schmidt et al., 1992" Wilcox et al., 1992; Tojo et al., 1994a; Bachmann et al., 1995" Grozdanovic and Gossrau, 1995" Pollock et al., 1995" Roczniak et al., 1999) are strongly positive in the macula densa cells (Fig. 3). The expression of this NOS isoform is also confirmed by in situ hybridization labeling of the macula densa cells with a probe detecting the NOS-I mRNA (Mundel et al., 1992). Using electron microscopic immunocytochemistry Tojo et al. (1994a) found a reaction precipitate within the cytoplasm of the macula densa cells (Roczniak et al., 1999), mainly associated with small vesicles. Recent findings suggest the occurrence of NOS-I immunoreactivity in the narrow portion of the juxtaglomerular outflow segment of the efferent arteriole that is characterized by endothelial cells which are protruding into the vessel lumen. It is presumed that this segment represents a specific shear stress receptor (Elger et al., 1998). The only immunocytochemical study on the localization of the subunits of sGC in the kidney was performed by Mundel et al. (1995). According to these authors the alpha-1 subunit was located in the glomerular podocytes, whereas the beta-2 subunit was located in principal cells of the cortical collecting duct. These results show only some sites of distribution of sGC in the kidney (Star, 1997), probably due to a lower sensitivity of the applied immunocytochemical technique or the quality of the antibodies used. Localization of the endothelial NOS-III isoform in the kidney. Using RT-PCR the NOS-III mRNA was detected in microdissected segments along the nephron system (Terada et al., 1992a,b). The strongest signal was detected in inner medullar collecting duct whereas a lower signal was displayed in the inner medullar limb, cortical collecting duct, outer medullar collecting duct, glomerulus, vasa recta, and arcuate artery. Endothelin-1 mRNA signal was established in glomerulus and inner medullar collecting ducts, but endothelin A-receptor mRNA only in glomerulus, vasa recta bundle and arcuate artery. Endothelin B-receptor mRNA was located mainly in glomerulus and collecting ducts, sGC mRNA could be found largely in glomerulus, proximal convolute tubule, proximal straight tubule, and cortical collecting duct as well as in small amounts in the medullar thick ascending limb, inner medullar limb, outer medullar collecting duct, inner medullar collecting duct, and the vascular system (Terada et al., 1992a,b). In addition, both endothelin-1 and NOS-III protein expression were found in the juxtaglomerular cells of the afferent arterioles (Yanagisawa et al., 1998). Similarly, in microdissected nephron segments and by RT-PCR, NOS-III mRNA was found at high levels in renal glomeruli, interlobar artery/afferent arterioles and less in arcuate vessels. NOS-III mRNA was found by these authors inconstantly in proximal tubules, thick ascending 270

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limbs and cortical intermediary collecting ducts (Ujiie et al., 1994). Immunoreactivity for the NOS-III in the kidney was studied by Bachmann et al. (1995) and found to be located within the endothelial cells of the glomerular capillaries, both the afferent and efferent arterioles, as well as in the medullar vasa recta. In addition, a faint immunostaining in the entire distal tubule was reported by Tojo et al. (1994a). In contrast to the mRNA results these authors were not able to establish NOS-III immunoreactivity in the arcuate or interlobular arteries. NOS-III immunoreactivity was established also in the juxtaglomerular cells of the afferent arteriole by Grozdanovic et al. (1995) in marmoset kidney. In a recent study Heeringa et al. (1998) reported that a strong NOS-III immunoreactivity possessed the glomerular capillaries as well as interstitial tubular capillaries and larger interstitial vessels in Brown Norway rats. Also in the human kidney NOS-III was present in glomerular endothelial cells and the endothelium of cortical vessels (Furusu et al., 1998). Direct evidence for the production of NO by isolated rat glomeruli was recently provided by Reverte and Lopez-Novoa (1998). Localization of the inducible, macrophageal NOS-II isoform ill the kidney. The results concerning the location of NOS-II in the renal structures are controversial. In the normal rat kidney NOS-II mRNA was found primarily in the outer medulla (Morrissey et al., 1994). Structures with significant amounts of NOS-II mRNA established in this study were the proximal straight tubule, the medullar thick ascending limb, medullar collecting duct, vasa recta bundle and the glomeruli. Lesser amounts were found in the cortical tubules and in the inner medulla. An important finding was the fact that in the kidney NOS-II may be expressed to some extent constitutively (Kone and Baylis, 1997). In situ hybridization studies and studies by RT-PCR and immunocytochemistry have shown that VSM-NOS-II was expressed in the glomerulus as well as in the interlobular and arcuate arteries of normal rats. The second isoform, the MAC-NOS-II was found within the basal compartments of different renal tubule segments, being abundant in the $3 proximal tubule segment, the cortical and medullar thick ascending limb, the distal convolute tubule, cortical convolute tubule, cortical collecting duct and inner medullar collecting duct (Ahn et al., 1994; Mohaupt et al., 1994b; Morrissey et al., 1994; Tojo et al., 1994a). Lipopolysaccharide (LPS)-treated rats exhibited a much greater proportion of VSM mRNA and a higher level of total NOS-II mRNA in the cortex and in the outer and inner medulla of rat kidney (Mohaupt et al., 1994b). MAC-NOS-II was also found in cultured mesangial cells and the cytokines TNF-alpha and IF-gamma preferentially induced the expression of VSM mRNA in these cells (Mohaupt et al., 1994b). The NOS-III gene in mIMCD-3 cells derived from murine medullar collecting duct seems to be under basal conditions transcriptionally inactive, but could be markedly induced by TNF-alpha and IF-gamma (Mohaupt et al., 1995). The NOS-II gene transcription in the inner medullar collecting duct cells is activated by cytokines (Mohaupt et al., 1995). Also immunohistochemical studies of control and stimulated rats resulted in different localizations of the NOS-II (Jansen et al., 1994; Tojo et al., 1994b, 1995). The immunocytochemical localization of NOS-II in the afferent arteriole could not be confirmed by NOS-II mRNA visualization (Ahn et al., 1994). Immunocytochemistry, using antibodies against the NOS-II showed strong labeling of the intercalated cells of the cortical collecting duct (Tojo et al., 1994b). Interestingly, in rat kidney immunoreactivity for the inducible NOS-II occurs after LPS treatment of the animals in histiocytes and endothelial cells (Bandaletova et al., 1993). In non-stimulated kidneys these cells do not possess any NOS-II immunoreactivity. Mesangial cells of the kidney produce and release the NO degradation products NO 3 / N O 2 following stimulation by LPS or cytokines (Shulz et al., 1991). This indicates the presence of the macrophage NOS-II form (Buttery et al., 1994). There is strong evidence that mesangial 271

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cells are able to produce reactive oxygen species such as superoxide anion, hydrogen peroxide, hydroxyl radical and hydrochlorous acid. It is suggested that these autacoids may be involved in processes accompanying some forms of glomerular injury (Baud et al., 1992). Endogenously produced NO by mesangial cells in the rat kidney may upregulate the copper/zinc superoxide dismutase in these cells (Frank et al., 1999). This may be a protective mechanism for the prevention of formation of high quantities of peroxynitrite during endotoxic shock induction of iNOS. In control human kidney mesangial cells NOS-II immunoreactivity was not found; however, in case of glomerulonephritis immunoreactivity was found not only in mesangial cells but also in glomerular epithelial cells and infiltrating cells (Furusu et al., 1998). As already mentioned, there are species-related differences concerning the expression of NOS-I. In marmoset kidney Grozdanovic et al. (1995) were not able to establish any NOS-I immunoreactivity and NADPH-d activity in cells of the macula densa. Also in human macula densa cells with normal or nephritic glomeruli no immunoreactivity for NOS-I was found (Furusu et al., 1998). In these cases it is difficult to decide whether these are true species-related differences or that the discrepancies are due to inappropriate processing of the material or to the qualities of the antibodies used. In addition to the species-related differences, sex-dependent differences in the expression of the NOS-III and NOS-II isoforms were also reported. In this respect, the levels (established by Western blot analysis) of NOS-II and NOS-III were found to be significantly higher in the inner medulla of female rats as compared with male rats (Neugarten et al., 1997)" for the NOS-II this was true even after stimulation with LPS. These results were confirmed by establishing the steady-state levels of mRNA for NOS-II that were higher in the inner medulla of female rats. These relations seem to be specific since sex hormones failed to influence NO production or NOS-II levels in mesangial cells in culture that were stimulated by LPS (Neugarten et al., 1997). NOS in neural structures of the kidney. In the rat kidney two populations of nerve cell bodies exist. The large population of these ganglionic cells exhibit NADPH-d activity that co-localize with immunoreactivity for the catecholamine synthesizing enzyme dopamine beta-hydroxylase (Liu et al., 1995). The second neuronal population shows only NADPH-d activity. The probably parasympathetic nerve cell bodies are located at the hilum of the kidney associated with nerve bundles, in proximity to the lower or middle portion of the interlobar arteries, and on the wall of the renal pelvis (Liu and Barajas, 1993). NOS-I immunoreactivity was also established in perivascular nerve fibers surrounding the arcuate and interlobular arteries as well as beneath the pelvic epithelium. Functional significance of NO in the kidney. Numerous experimental studies revealed that in the kidney NO generated by NOS-I and NOS-III is involved in the regulation of glomerular hemodynamics. There is evidence that the NO generated from these enzymes fulfils different functional roles. The NO produced by NOS-I in the macula densa is considered an integral modulator of the tubuloglomerular feedback response, whereas the endothelium-derived NO acts as a major vasorelaxant factor (Bosse and Bachmann, 1997). Disjunction of the enzymes may cause glomerular hypertension and increased intraglomerular platelet aggregation (King and Brenner, 1991; Kettler et al., 1998). One of the most intriguing events in the regulation of kidney homeostasis is the autoregulatory response connected with the autoregulatory behavior of the renal microvasculature in response to changes in the general and renal arterial perfusion pressure (Ichihara et al., 1998b; Navar, 1998). This response ensures that the renal blood flow, the glomerular filtration rate, the glomerular pressure, the proximal tubule pressure, and the peritubular capillary pressure remain relatively constant over a wide range of the renal perfusion pressure (from as low as 70 272

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mm Hg to over 180 mm Hg). The autoregulatory response comprises two mechanisms which interact with each other in a complex manner (Navar, 1998): the myogenic response and the tubuloglomerular feedback response. The myogenic mechanism is intrinsic to arterial smooth muscle cells and regulates blood flow by influencing transmural wall tension. An increase in the arterial pressure results in vasoconstriction of the renal vascular musculature. The vasoconstriction leads to maintenance of a higher vascular resistance. The tubuloglomerular feedback mechanism depends on the special features of the macula densa cells and is primarily associated with the regulation of tubular fluid composition and regulation of the afferent arteriolar tone (Kone and Baylis, 1997; Navar, 1998). Vascular endothelial cells are influenced by two distinct hemodynamic forces, namely cyclical strain and shear stress. In the kidney, the vascular smooth muscle cells of afferent and efferent arterioles show selective responsiveness to various stimuli. The complex autoregulatory responses of afferent arterioles comprise the myogenic mechanism in response to increases in renal perfusion pressure through 'stretch-activated' cation channels that lead to depolarization, calcium entry, and vascular contraction. The efferent arterioles utilize other possibilities for calcium entry as well as intracellular calcium mobilization and do not appear to have a myogenic mechanism (Navar, 1998). As the last author postulates, there is a close coupling of renal blood flow and glomerular filtration rate autoregulation over the major portion of the autoregulatory range and this seems consistent with the view that most or all of the autoregulatory-mediated changes in renal vascular resistance are localized at preglomerular (afferent) arterioles. Acute shear stress in vitro activates signaling cascades in endothelial cells with acute release of NO and prostacyclin, the activation of transcription factors such as the nuclear factor (NF) kappaB, c-fos, c-jun and SP-1 as well as transcriptional activation of genes such as IVAM-1, MCP-1, tissue factor, platelet-derived growth factor-B, TGF-beta-1, cyclooxygenase-II and NOS-III (Ballermann et al., 1998). Not only endothelial cells of the vessels but also glomerular epithelial cells produce paracrine factors that are able to influence the basal tone and reactivity of afferent and efferent arterioles (Navar et al., 1996; Navar, 1998). It becomes evident that one of the factors involved in the autoregulation of renal homeostasis is NO. The localization of NOS-I in the macula densa cells shows the possibility for a role of NO in mediating signal transfer from distal tubular fluid to glomerular arterioles. The cells of the macula densa produce and release substances involved in vasoconstriction of the adjacent afferent arteriole; thus, being a part of the tubuloglomerular feedback response they are involved in its negative-feedback mode of action to associate glomerular capillary pressure and tubular-fluid delivery and reabsorption (Thorup et al., 1996). The tubuloglomerular feedback response which responds mainly to flow-dependent changes in tubular fluid composition (NaC1 and total solute concentration) in the distal tubules at the level of macula densa and transmits signals to the afferent arterioles to alter the activation state of voltage-dependent calcium signals leading to changes in afferent arteriolar tone (arteriolar resistance), enhances the autoregulatory mechanism of the kidney glomeruli. Experiments, performed in the stenotic Goldblatt kidney, suggest that the stimulation of the NOS-I of the macula densa cells induces release of NO into the extraglomerular mesangium. From there an NO-dependent intermediate stimulus may reach the glomerular vasculature (Bosse and Bachmann, 1997). NO of macula densa cells seems to have an important role in counteracting tubuloglomerular feedback response-mediated afferent arteriolar constriction and NO in general exerts vasodilatory effects on both afferent and efferent arterioles probably caused by enhanced shear stress and/or angiotensin II levels (Bosse and Bachmann, 1997). There are numerous candidates for tubu273

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loglomerular feedback response mediators between the macula densa and the afferent arteriole acting at the juxtaglomerular apparatus of the kidney. In addition to NO generated by NOS-I in macula densa cells, other paracrine factors can mediate the tubuloglomerular feedback response and autoregulatory related signals to the afferent arterioles. Among these factors, extracellular ATE adenosine, intrarenal angiotensin II (exerts vasoconstrictor effects on both afferent and efferent arterioles), cytochrome P450 vasoconstrictor metabolites, and arachidonic acid metabolites (eicosanoids), are able to modulate vascular responsiveness in order to maintain an optimal balance between the metabolically determined reabsorptive capabilities of the tubules and the hemodynamic-dependent filtered load (Navar et al., 1996; Navar, 1998). It is important to note that the tubuloglomerular feedback mechanism, depending on increases or decreases in distal tubule flow, may cause either vasoconstriction or vasodilatation. Under normal conditions, the renal vasculature is subject to a strong continuous influence by endogenous NO. Endogenous endothelium-derived NO is involved in the regulation of renal cortical blood flow (Luscher and Bock, 1991; Walder et al., 1991). Tonic basal release of NO by vascular endothelial cells controls blood pressure (vascular tone) in the basal state (Baylis et al., 1992). However, although the tubuloglomerular feedback and myogenic response may be modulated by NO, the accuracy of the renal hemodynamic autoregulation as well as the mean glomerular filtration rate seem to be kept independent of the influence of mean renal blood flow. The autoregulatory capacity for both renal blood flow and glomerular filtration rate in response to static or dynamic pressure changes is not affected (Just, 1997). This underlines once again the relative regulatory autonomy of the blood pressure and the glomerular filtration rate in the kidney. The preferential expression of NO by NOS-III in the renal microvasculature and glomeruli is functionally connected with regulation of glomerular filtration rate by vasodilating the afferent and to a lesser extent the efferent arterioles and by increasing the glomerular capillary ultrafiltration coefficient (Kf) (Dussaule and Chatziantoniou, 1996). As an endogenous vasodilator NO contributes to renal arteriolar tone (arteriolar resistance) and modulates relaxation of the mesangium cells (Raij et al., 1996). Numerous studies concern the significance of NOS isoforms and NO on renal afferent and efferent arteriolar resistances (see Kone and Baylis, 1997 for review). NO, tonically generated by NOS-III, exerts a direct vasodilator effect on the renal microcirculation. This local regulation is different from the action of NO on the general circulation which involves the decreased activity of such vasoconstrictors as angiotensin II (Ang II), the sympathetic nervous system (Granger et al., 1997) and endothelin (ET) (Kone and Baylis, 1997). These events are very complex and variable. Apparently, exogenous NO may cause renal arteriolar vasodilatation through both cGMP-dependent and cGMP-independent mechanisms, and the cGMP-independent action did not require K + channel activation. The endogenous NOS-III action appears to act exclusively through cGMP (Trottier et al., 1998). However, there are also results showing an action of NO on cAMP PDE-3 (phosphodiesterase-3) which also may account for the vasodilator effects of NO on the renal vasculature, thus the PDE-3 activity appears to be an important determinant of renal vascular resistance (Sandner et al., 1999). It must be noted that NO is not the only vasodilator of the renal vessels. There is evidence that the natriuretic peptides CNP and ANP act as vasodilators on the juxtaglomerular afferent arteriole when it is preconstricted with norepinephrine. In this condition CNP dilates the afferent arterioles via the prostaglandin/NO pathway, whereas ANP dilates them directly (Amin et al., 1996). There is also strong evidence that the ANP-dependent system and the particulate guanylyl cyclase (pGC) may be compensatory to altered activity of NO and sGC (Lewko et al., 1997). Also prostaglandins may maintain renal vasodilation and hyperfiltration during chronic NO synthase blockage in 274

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conscious pregnant rats (Danielson and Conrad, 1996). Endothelin-3 (ET-3) was found to stimulate release of prostaglandins and NO which cause dilatation of renal vasculature. Under these circumstances ET-3 is acting as a vasodilator and diuretic peptide (Yamashita et al., 1991). There is also evidence that NO derived from macula densa cells is involved in the control of renin synthesis (Bosse et al., 1995). Possible involvement of NOS-I at the macula densa in the increased renin production was recently recognized in the angiotensin type l a receptor gene-knockout mice (Kihara et al., 1998). Inhibition of NOS decreases renin levels in the juxtaglomerular apparatus. Moreover, NOS activity and gene expression are inversely related to chronic changes in renal perfusion, salt balance, and salt transport at the distal tubule in parallel with the response of renin in these changes (Bosse et al., 1995). This participation of NO in the tubuloglomerular feedback mechanism (Ito et al., 1997) implies a role for NO in body fluid-volume and blood-pressure homeostasis (Wilcox et al., 1992; Wilcox, 1998). Study on the importance of changes in the systemic blood pressure and the renin-angiotensin system in the mechanism of regulation of NOS-I in the macula densa of the rat, provides evidence that the macula densa NOS-I changes in parallel with plasma Ang II, but not renin or systemic blood pressure (Muracami et al., 1997). In addition to NO, other local factors such as prostaglandins and endothelins can exert significant effects on renin secretion and renin gene expression. Both prostaglandin E2 (the main renotubular prostaglandin) and prostacyclin (PGI2, the main endothelial prostanoid) stimulate renin gene expression and renin secretion through a cAMP mechanism (Wagner et al., 1998). In contrast, endothelins seem to inhibit the renin system. These studies show that NO and prostaglandins may be involved in the regulatory mechanisms by which salt intake affects the renin-angiotensin system (Wagner et al., 1998). From these studies it is evident that the juxtaglomerular apparatus plays a central role in the regulation of glomerular hemodynamics and renin release. It is important to emphasize that NO has differential action in the afferent arteriole, the efferent arteriole and the macula densa which may have important significance in the control of glomerular hemodynamics under various physiological and pathological situations (Ito et al., 1997). Glomerular hemodynamics may be essentially involved in the pathogenesis of hypertension and in the mode of progression of renal dysfunction (for review see Ito, 1997). NO also plays a role in the regulation of renal sodium excretion of proximal tubules and collecting ducts. NO is involved in the relationship between renal perfusion pressure and natriuresis and influences the tubuloglomerular feedback control. NO is an important autocrine or paracrine factor directly regulating Na + transport in the proximal tubule (Roczniak and Bums, 1996). NO causes the induction of natriuresis which occurs independently of changes in renal perfusion pressure (Majid and Navar, 1997). It was shown that NO inhibits both Na + (H +) exchange and Na+/K+-ATPase activity in the proximal tubule (Stoos and Garvin, 1997). Recent findings suggest that NO is able to inhibit transcription of the Na+/K+-ATPase alpha-1-subunit gene in an medullar thick ascending limb cell line (Kone and Higham, 1999). In addition, NO is also involved in regulating the activity of the basolateral low-condhctance 30-pS K + channel of the cortical collecting duct as well as the apical 70-pS K + channel in the thick ascending limb of the rat kidney (Lu and Wang, 1966; Lu et al., 1998). As a result, NO inhibits sodium reabsorption in proximal tubules and collecting ducts. However, in the cortical collecting ducts of rat it was shown that the transport was not regulated via the NO pathway and that the applied NO donor sodium nitroprusside acts as a cGMP-independent activator of K + channels in the basolateral membrane of these cells (Hirsch et al., 1997). 275

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NO plays a role also under conditions of increased or decreased dietary salt intake (Singh et al., 1966). Intake of sodium and extracellular fluid volume regulate levels of mRNA of the three NOS isoforms in the renal cortex differentially, suggesting that each of them plays a specific role (Nadaud et al., 1998). High salt intake increases NO production and renal action that depend on NOS-I and macula densa solute reabsorption, providing evidence that changes in NO generation via NOS-I in the macula densa contribute to changes in whole kidney NO generation and vasodilator and possibly natriuretic action (Wilcox et al., 1998). NO plays also a role during increased sodium intake in healthy humans and represents an important part of adaptation with respect to renal hemodynamics, sodium excretion, and secretion of renin (Bech et al., 1998). However, it is likely that additional mechanisms are involved in the NO generation in salt intake, because studies performed by Ying and Sanders (1998) showed that dietary salt increased glomerular expression of TGF-beta-1, which in turn enhanced NO production through NOS-III. On the other hand, dietary sodium restriction leads to increased content of renin in the kidney. This appears to require the induction of cyclooxygenase-2, while neuronal NOS appears to affect basal (not stimulated) renin content (Harding et al., 1997). Concerning the distribution of NOSs in the structures of kidney medulla it was shown that NO selectively increased (following stimulation with bradykinin or acetylcholine) or decreased (after L-NAME application) renal medullary blood flow with parallel changes in sodium and water excretion (Mattson and Higgins, 1996; Mattson et al., 1997). The selective decrease in medullar blood flow is associated with retention of sodium and water and leads to the development of hypertension (Mattson et al., 1997). Moreover, these events were independent of alterations in renal cortical blood flow or glomerular filtration rate. Thus, NO in the renal medulla exerts a strong influence on fluid and electrolyte homeostasis and the control of blood pressure. There is evidence that NO plays an important role in mediating the renal vasodilatation during chronic aldosterone excess (Granger et al., 1999). Persistent hypertension occurs in case of an abnormal relationship between renal perfusion pressure and renal sodium excretion. Recent studies have shown that some animal models of genetic hypertension and human hypertension forms are associated with a decrease of NO synthesis. This reduction of NO synthesis leads to reduction of the renal sodium excretory function, not only through direct action on renal vasculature, but through modulation of other vasoconstriction processes and through direct and indirect alterations in tubular transport (Schnackenberg et al., 1997). In this respect it was shown that acute changes in arterial pressure induce changes in intrarenal NO production, which may directly alter the tubular reabsorptive function to manifest the phenomenon of pressure natriuresis (Salom et al., 1992; Kone and Baylis, 1997; Majid and Naval, 1997). Recent results provide evidence that angiotensin II stimulates synthesis of NOS-III and enhances NO production. Previous studies have suggested that NO may play an important role in protecting the renal vessels from Ang II-mediated vasoconstriction (Henington et al., 1998). In case of angiotensin II-induced hypertension the maintained NO production in afferent arterioles counteracts the concomitant enhanced afferent arteriolar reactivity (Chin et al., 1998; Ichihara et al., 1998a). Results concerning the importance of the sympathetic nervous system in the vasoconstrictor response to acute NOS inhibition are controversial (see Kone and Baylis, 1997). In contrast to results of Granger et al. (1997), studies performed by Baylis et al. (1997) showed that in the normal, conscious, chronically catheterized rat in which the sympathetic nervous system is working at basal level, renal nerve activity does not contribute to the pressor or renal vasoconstrictor response to NO inhibition or the renal vasodilator response to NO stimulation. On the other hand, an interaction between NO (suppressive effects) and 276

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endothelin (stimulatory effect) in modulating the sympathetic nervous system in conscious spontaneously hypertensive (not normotensive) rats was established by Kumagai et al. (1997). There is also evidence that under elevated peripheral and renal sympathetic nerve activity, NO has a role in modulating renal hemodynamics, but not sodium excretion. This action does not appear to involve Ang II (Vogel and Zambraski, 1996). There is also evidence that endothelin acts as an inhibitory modulator of renal noradrenergic neurotransmission through ETB-receptor mechanisms (Matsuo et al., 1997). These last authors presume that this effect may be caused by stimulation of NO production by ET. There is also evidence that NO in addition to its ability to modulate vascular resistance by a peripheral action may also play a role in the central regulation of sympathetic renal nerve activity (Sakuma et al., 1992). However, as emphasized by Kone and Baylis (1997) the exact mechanisms concerning the role of NOS and Ang II in the central and peripheral regulation of vascular tone by the sympathetic nervous system are not definitely elucidated. It seems likely that in the kidney tonically released NO, probably by NOS-III, has a direct vasodilatory effect on renal microcirculation, whereas in the systemic circulation the action of NO requires attenuation of Ang II/sympathetic nervous system/endothelin activity (Kone and Baylis, 1997). 1.1.2. The renal pelvis .

There is strong evidence for the existence of NOS-immunoreactive nerve cell somata and postganglionic nerve fibers in the wall of the rat renal pelvis (Liu and Barajas, 1993; Liu et al., 1996a,b). All somata showing NOS immunoreactivity also possessed NADPH-d enzyme activity (Liu et al., 1996b). Exogenously applied NO elicited relaxation in pre-contracted human renal pelvis, where a calcium-dependent NOS activity was detected by Iversen et al. (1995). 1.1.3. The ureter By means of double label immunofluorescence the distribution pattern and co-localization of NOS-I, vasoactive intestinal peptide (VIP), neuropeptide Y (NPY) and the catecholamine-synthesizing enzyme tyrosine hydroxylase (TH) was studied in nerve fibers supplying the human lower ureter (Smet et al., 1994). It was found that some non-adrenergic VIP-containing nerve fibers have the capacity to synthesize NO, and that NO may be involved in the regulation of ureteric motility. Neurons in the ureterovesical ganglion complex provide autonomic innervation to the pelvic ureter, the ureterovesical junction and the bladder trigone. Using NOS-I immunocytochemistry and NADPH-d enzyme histochemistry, Goessl et al. (1995) studied their distribution in neuronal perikarya and processes in the human ureterovesical junction. Both immunoreactivity and enzyme activity were found in nerve cells of extramural situated ureterovesical ganglia. Positive nerve fibers were seen running within large extramural nerve trunks and between intramural smooth muscle bundles. Positive varicose nerve fibers were found surrounding extramural and intramural blood vessels. A discrepancy between the two methodical approaches is also reported, namely that the urothelium showed a strong NADPH-d enzyme activity but showed no NOS immunolabeling. The conclusion was made that NO may have a physiologic role in opening of the human ureterovesical junction. These results were confirmed and extended by Grozdanovic and Baumgarten (1996) which showed that NOS-positive perikarya in the ureterovesical ganglia co-locates VIP and NPY 277

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immunoreactivities. Also studies of the human ureter have shown that NOS-positive neuronal axons and nerve-ending-like structures are distributed in the muscular layers and possibly play a vasorelaxation role (Stief et al., 1996). NOS-, CGRP-immunoreactive and cholinergic nerves were observed in the longitudinal musculature of the distal ureter and the ureteral orifices into the bladder of female rats where no adrenergic, NPY- and VIP-immunoreactive nerves existed (Alto et al., 1995). It seems likely that species-related differences exist in the chemical composition of the autonomic ganglia along the chicken ureter and the ganglion cells found in this region in mammals (Sann, 1998). This is probably true also for the functional role of NO in the ureter while in the sheep ureter electrical field stimulation shows no NO-mediated relaxation in comparison with the urethra (Garcia-Pascual et al., 1996). Immunohistochemical monitoring of NOS activity and functional inhibitory effects mediated by NO and CO were studied in human ureter (Iselin et al., 1998). The authors observed moderate NOS innervation of the ureter, that was more prominent in the distal segments. Few NOS-immunoreactive nerve fibers were found in the wall of the organ, whereas varicose nerve fibers containing also calcitonin gene-related peptide immunoreactivity were situated in the submucosa. In the distal parts of the ureter nerve trunks were observed which exhibited immunoreactivity for hemoxygenase-2 (HO-2). In strips exposed to NO, an increase of the cGMP levels could be established. The authors emphasized that the richer innervation of the distal ureter may serve to the co-ordination of ureteral peristalsis and the motility of the ureterovesical junction. Studies of the same group (Iselin et al., 1997) on the pig and human intravesical ureter provide evidence that the L-arginine-NO-cGMP pathway may play a role in the regulation of the valve function in the ureterovesical junction. There is evidence that NO acts as an inhibitory neurotransmitter in the pig intravesical ureter and leads to relaxation of the smooth muscle through a guanylyl cyclase-dependent mechanism and opening of glibenclamide-sensitive K + channels (Hemandez et al., 1997). Using a conventional protocol for the visualization of NADPH-d enzyme activity Grozdanovic and Gossrau (1995) and Grozdanovic et al. (1995) revealed a positive reaction product deposition in the epithelial cells of the ureter and the urinary bladder of rats, hens, hamsters, gerbils, guinea pigs and marmosets. However, incubation solutions with added formaldehyde or permanganate failed to detect NADPH-d enzyme activity in the urothelium and showed positivity only in nerve fibers in the marmoset ureter. The nerve fiber localization was also the only NOS-I immunoreactivity seen in the ureter of the marmoset (Grozdanovic et al., 1995). In contrast to these findings and those of Goessl et al. (1995), immunoreactivity for NOS-I (Fig. 4) was shown in the superficial cells of the urothelium of the human ureter by Schindelmeiser et al. (1997). In the same cells immunoreactivity for the Cu/Zn superoxide dismutase and the tartrate-resistant acid phosphatase were found. All these enzymes are involved in the production and metabolism of free oxygen radicals (e.g. peroxynitrite). Thus, the urothelium possess a cytotoxic system which may be directed against pathogenic micro-organisms and foreign cell material in the urine and at the same time a SOD system which may protect the superficial cells from their own cytotoxicity. In accordance with a localization of NOS-I immunoreactivity in the urothelium are results from direct measurements of endogenous NO production on the surface of urinary bladder strips. It was shown that norepinephrine releases NO from urinary bladder epithelium, and capsaicin releases NO both from the epithelium and the nervous structures of the urinary bladder (Birder et al., 1998). These results support the view for a role of NO in the modulation of bladder and urothelial function. 278

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1.2. THE LOWER URINARY TRACT

1.2.1. The urinary bladder The major representative NOS form in the lower urinary tract is the NOS-I and the mean structures in which NOS immunoreactivity was found were components of the peripheral sensory and vegetative nervous system. In different species NADPH-d enzyme activity was detected in blood vessels (Fig. 5), the urothelium and neural structures of the bladder, including extramural (Fig. 6) and intramural sensory and parasympathetic cells and nerve fibers (Grozdanovic and Gossrau, 1995; Grozdanovic et al., 1995). However, parallel immunocytochemical studies failed to find NOS-I immunoreactivity in the urothelium of rat bladder (Alm et al., 1993). By means of cGMP and NOS-I immunohistochemistry Smet et al. (1996) revealed the potential target structures in the lower urinary tract (bladder and proximal urethra) of guinea pig and human. In both species a large number of the intramural nerve cells of the bladder contained NOS-I immunoreactivity. Also a moderate innervation to the detrusor muscle by varicose nitrergic nerve fibers was established. In contrast, the innervation of the urethral muscle was more dense. NOS-positive nerve fibers were observed in the submucosa and around blood vessels of the bladder. Distinct regional differences were found after stimulation with sodium nitroprusside: the smooth muscle cells of the urethra exhibited strong cGMP immunoreactivity which was not seen in the detrusor muscle. Further target cells that showed cGMP immunoreactivity were interstitial cells with mesenchymal features, the uroepithelial cells, vascular smooth muscle cells and pericytes. All these cells could potentially mediate the effects of endogenously released NO. Small clusters of NOS-immunoreactive paraganglion cells and branching varicose nerves in their vicinity were observed in the muscle layer of the urinary bladder during the 13 and 20 weeks of gestation of human fetuses (Dixon et al., 1998). The nerve fibers entering sensory corpuscles (Timofeew's sensory corpuscles) also exhibited NOS-I immunoreactivity, indicating possible involvement of NO in the sensory transmission from the urinary bladder. In the guinea pig urinary bladder NADPH-d-positive and NOS-I-immunoreactive intramural ganglion cells were observed (Zhou et al., 1997; Zhou and Ling, 1998). The nitrergic neurons were most numerous in the bladder base. In addition, distinct differences between the number of NADPH-d-positive and NOS-positive nerve cells were established providing an additional evidence that not all NADPH-d enzyme activity accounts for NOS. Recent findings provide evidence for estrogen effects on NOS in the lower urinary tract. In contrast to the upper urinary tract (kidney, urinary pelvis, ureter), estrogen treatment of ovariectomized rabbits led to a significant reduction of cytosolic (not particulate) NOS in the bladder, trigonum and urethra (A1-Hijji and Batra, 1999). In the perfusion (blood flow) of dog bladder mucosa sex-related differences were established by Pontari and Ruggieri (1999), the perfusion of male dog mucosa being significantly greater than in females. Moreover, the perfusion of the bladder mucosa was regulated differently than the bladder musculature. Intravesical instillation of bacterial LPS led to a localized inflammatory response and induction of NOS-II in the rat urinary bladder (Olsson et al., 1998). Sensory neuron in rat dorsal root ganglia and efferent neurons in the major pelvic ganglia responded to acute experimental cystitis with increased NOS synthesis (Callsen-Cencic and Mense, 1997). A role of NO in neuronal cell death in the urinary bladder following acute urinary retention was presumed by Zhou and Ling (1997) who established an increase of NADPH-d enzyme activity in the intramural ganglion cells of the urinary bladder of the guinea pig. Complete transection of the spinal cord increases the number of NOS-I-immunoreactive dorsal root ganglion 279

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neurons as well as the number of bladder afferent neurons exhibiting NOS-I immunoreactivity (Vizzard, 1997). Functionally these changes may be related with the emergence of the spinal micturition reflex.

1.2.2. The urethra Numerous studies on the distribution of the components of the NO/cGMP pathway in the lower urinary tract have shown that they are present only in its proximal part, the distal urethra being devoid of NOS. This indicates that NO acts predominantly within the outflow regions of the lower urinary tract (Andersson, 1996; Garcia-Pascual et al., 1996). However, nitrergic nerve fibers represent only one type among the nerves supplying the lower urinary tract (Radzisewski et al., 1996). Early studies showed that NO may play a major role in the non-adrenergic, non-cholinergic (NANC) inhibitory response in the urethra, bladder trigone and bladder neck in different species (for literature see Persson et al., 1993). In the pig lower urinary tract NADPH-d enzyme activity and NOS-I immunoreactivity were found in nerve trunks and fine nerve fibers in and/or around muscular bundles in the detrusor and the trigone of the bladder as well as in the urethra. Studies performed by Werkstrom et al. (1998) on the guinea pig urethra showed a dense innervation of the proximal urethral musculature by nerve fibers and terminals containing NOS-I, VIP and tyrosine hydroxylase (TH). Also a small number of cGMP-immunoreactive cell bodies could be seen in the urethra. Experiments with electrical field stimulation suggested that in the urethra a non-NO/cGMP mediated relaxation may be predominant (see also Lundberg, 1996). In the proximal part of the hamster urethra it was established that the urothelium may contribute to relaxations of the smooth muscle after nerve stimulation. In addition, a role for NO in inhibition of noradrenaline release from adrenergic nerves in the rabbit urethra was recently reported by Yoshida et al. (1998). There is strong experimental evidence against the view that nitrergic nerve fibers in the rat lower urinary tract and especially in the urethra belong to sympathetic or sensory nerves (Persson et al., 1997). Recent results obtained after pelvic ganglionectomy suggest that NOS-immunoreactive nerves, that mediate inhibition of smooth muscle tone in the rat urethra, originate from the major pelvic ganglion and contain choline acetyltransferase, indicating that these nerves may be cholinergic (Persson et al., 1998). Although positive for NADPH-d, no NOS-I immunoreactivity could be observed in the pig and rabbit urothelium (Persson et al., 1993; Zygmunt et al., 1993). In contrast, recent Western blot results provide evidence for the existence of a constitutive NOS in the urothelium of hamster proximal urethra and that the urothelium may release nitric oxide in response to electrical field stimulation (Pinna et al., 1999). Similar results and the possibility for a role in

<_....__

Fig. 5. NADPH-d enzyme activity in mouse tissues. Urinary bladder. Distinct staining is seen in the endothelial and smooth muscle cells of this small vessel. Bar -- 20 Ism. Fig. 6. NADPH-d enzyme activity in mouse tissues. Part of a perivesical ganglion. Strong enzyme activity is seen within the cytoplasm of the nerve cell perikarya. Bar = 20 g m. Fig. 7. Human testis. Groups of intertubular Leydig cells. Immunoreactivity for NOS-I isoform with distinct differences in the staining intensity between the individual cells. Bar -- 20 Ixm. Fig. 8. Human testis. Groups of intertubular Leydig cells (LCs). NOS-III immunoreactivityin a small LC-group. T -- tubule. Bar = 20 Ixm.

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relaxation of NO released by nerve fibers in the proximal (not distal) female dog urethra were reported by Takeda and Lepor (1995). This finding was confirmed by Nishizawa et al. (1997) who found that in the isolated dog urethra capsaicin produced an endogenous NO-dependent relaxation. NADPH-d-positive nerve fibers were also found within the lamina propria of the female rabbit urethra around smooth muscle bundles and arteries, where NANC-mediated relaxations, sensitive to NOS inhibition, were produced by electrical stimulation and by acetylcholine (Zygmunt et al., 1993). These results suggest that the urethral lamina propria can be relaxed independently by NO derived from NANC nerves and the endothelium of the blood vessels. In a recent study the presence of nitrergic nerves was found in the striated muscle of the human membranous urethral sphincter, suggesting a role for NO in the innervation of striated muscle (Ho et al., 1998). Moreover, in this study NOS immunoreactivity was found in the sarcolemma of the striate muscle cells, which seems to be an additional indication for a role for NO in regulation of metabolism and contraction of the urethral striate muscle. It seems likely that NO is not the only relaxant in the urothelium. As Ohnishi et al. (1997) showed P2Y-purinoceptors are distributed in the male rabbit urethra, and ATP and related purine compounds may play a role in NANC neurotransmission. Respectively, the pathways mediating urethral relaxation by ATE NO or VIP may be different. A relaxant effect of ATP that was NO- and urothelium-independent was also confirmed by Pinna et al. (1998). These compensatory mechanisms are probably the reason for the relatively normal appearance and behavior of NOS-I knockout mice (Sutherland et al., 1997). As mentioned above, the NOS activity is dependent on the hormonal status. Treatment of rabbits with estradiol dipropionate resulted in reduction of NOS activity and inhibition of the relaxation caused by nitrergic nerve stimulation of smooth muscle. Similarly, A1-Hijji and Batra (1999) found a significant down-regulation of the cytosolic (not particulate) NOS activity of rabbit female urethra following ovariectomy and subsequent treatment with polyestradiol phosphate. The existence of a new isoform of the NOS-I in the urethra, that is expressed also in the rat penis, prostate, skeletal muscle and cerebellum, was evidenced by Magee et al. (1996) but little is known about its exact localization and functional significance.

2. THE GENITAL TRACT

2.1. THE MALE REPRODUCTIVE ORGANS 2.1.1. The testis

NOS formation was described in the testis of man (Ehr6n et al., 1994) and rat (Burnett et al., 1995). Compared to other male reproductive organs testicular NOS activity was low. A series of reports, however, showed production and activity of NO in testicular structures, suggesting a special importance of the messenger nitric oxide. The first study localizing NO-synthesizing enzymes in testis-specific structures was published by our group. The neuronal subtype of NOS (NOS-I) was found in Leydig cells and Sertoli cells as well as in endothelial cells of the human testis (Davidoff et al., 1995). NOS-I was found to be also present in human Leydig cell tumors (Middendorff et al., 1995) as well as in rat Leydig cells (Lissbrant et al., 1997). In a former study, however, Burnett et al. (1995), were not successful to demonstrate this enzyme 282

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in rat Leydig cells. Subsequently, Leydig cells were shown to express also the isoforms NOS-II (Davidoff et al., 1996; Tatsumi et al., 1997) and NOS-III (Davidoff et al., 1996; Zini et al., 1996) (Figs. 7-9). Consistent with the immunohistochemical and Western blot data (Figs. 13 and 14), histochemical demonstration of NADPH-d enzyme activity in Leydig cells (Fig. 10) was also successful (Davidoff et al., 1995; Gianessi et al., 1998; Sjostrand et al., 1998). Furthermore, novel testis-specific NOS-I mRNA variants have been detected (Brenman et al., 1997; Wang et al., 1997). The mRNA transcript identified by Wang et al. also seems to be present in Leydig cells (our unpublished results). Until now, however, the function of these new transcripts is unclear. It was described that in most of the Leydig cells displaying NOS-I immunoreactivity, the NObinding enzyme sGC and its product cGMP (Fig. 11) were also detectable, whereas some cells were only sGC- and cGMP-immunopositive (Davidoff et al., 1997), suggesting autocrine as well as paracrine NO effects in Leydig cells. In accordance with the presence of sGC, treatment of isolated cells with NO donors resulted in elevated levels of cGMP (Davidoff et al., 1997). Since testosterone production constitutes the primary function of Leydig cells, a possible physiological involvement of NO in androgen regulation has to be considered. In this context, an inhibitory effect of NO on testosterone secretion by rat and mouse Leydig cells has been described (Adams et al., 1994; Welch et al., 1995; DelPunta et al., 1996; Gaytan et al., 1997; Pomerantz and Pitelka, 1998). This inhibition is due to direct NO action on at least one of the P450 steroidogenic enzymes and may be mediated in part by macrophage-derived NO (DelPunta et al., 1996; Pomerantz and Pitelka, 1998). An involvement of NO was also suggested in the context of stress- and shock-impaired steroidogenesis (Kostic et al., 1998; Sharma et al., 1998). However, the inhibitory effect on production of androgen is independent of cGMP (DelPunta et al., 1996). Therefore, findings suggesting a reduced expression of functionally active NO receptors both in mouse (MA10) and human Leydig tumor cells (Davidoff et al., 1995; DelPunta et al., 1996; Middendorff et al., 1997a) may support the existence of testosterone-independent physiological functions. For example, down-regulation of receptors for NO might prevent autocrine actions possibly necessary for the maintenance of normal Leydig cell phenotype. Previous studies have shown that NO influences tumor growth (Jenkins et al., 1995) and triggers a switch to growth arrest during differentiation of neuronal cells (Peunova and Enikolopov, 1995). Contractile peritubular cells and blood vessels were also found to be sites of NO production (Holstein et al., 1996; Middendorff et al., 1997b; Ruffoli et al., 1997; Bauche et al., 1998) and activity (Middendorff et al., 1997b). In peritubular myofibroblasts and testicular blood vessels NOS-I, NOS-III, sGC and cGMP (Fig. 12) were shown to be present (Holstein et al., 1996; Middendorff et al., 1997b), and NO donors increased tubular and vascular cGMP production, whereas NOS inhibitors displayed the opposite effect (Middendorff et al., 1997b) (Fig. 15). In these structures NO might be involved in relaxation of seminiferous tubules and blood vessels to modulate sperm transport and testicular blood flow, respectively. In this context, the cGMP-dependent protein kinase I, known to mediate relaxation was also found in contractile cells of the human testis (Middendorff et al., 1997a), and an influence of NO on testicular blood flow was found by in vivo experiments in rat (Lissbrant et al., 1997). In Sertoli cells, moreover, all isoforms of NOS were detected (Davidoff et al., 1995; Strphan et al., 1995; Zini et al., 1996; Middendorff et al., 1997b). The physiological significance, however, of NO-dependent cGMP accumulation in these cells (Norton et al., 1994) remains to be elucidated. Recently, we were able to describe a further functionally active NO system in the 283

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contractile cells of the testicular capsule of man (Davidoff and Middendorff, in prep.). In the rat, an involvement of NO in oxytocin-induced contractions of the capsule has already been described (S~inchez et al., 1996). In germ cells of the human testis, however, NO might not play a crucial role. NO synthases were exclusively found in apoptotic germ cells (Zini et al., 1996; E1 Gohary et al., 1999),

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Fig. 13. Western blot analysis for NOS-I (bands at approximately 160 kDa) and the alpha-1 subunit of the soluble guanylyl cyclase (sGC) (bands at approximately 82 kDa) in extracts of whole human testis and isolated Leydig cells. Signals representing the antigens are marked by arrows. Control -- rat brain extracts.

but not in intact g e r m cells ( D a v i d o f f et al., 1995, 1997; Zini et al., 1996; M i d d e n d o r f f et al., 1997b) and the N O - b i n d i n g e n z y m e s G C was barely d e t e c t a b l e in tubular g e r m cells ( D a v i d o f f et al., 1997; M i d d e n d o r f f et al., 1997a). Different to a variety of other organs, it is still unclear, w h e t h e r N O s y n t h a s e s are p r e s e n t in testicular nerve fibers. The only study s u g g e s t i n g the p r e s e n c e of N O S within intertubular structures was p u b l i s h e d by Zini et al. (1996). T h e o t h e r reports did not find any N O S - s p e c i f i c staining of interstitial nerve fibers. Since L e y d i g cells have b e e n s h o w n

Fig. 9. Human testis. Groups of intertubular Leydig cells (LCs). NOS-II immunoreactivity with differences between the individual LCs. Differential contrast. Bar = 20 g m.

Fig. 10. Mouse testis. A Leydig cell group with strong NADPH-d enzyme activity situated between two seminiferous tubule cross-sections. Bar --- 20 g m.

Fig. 11. Human testis. Immunocytochemical visualization of cGME Two adjacent Leydig cell groups showing some differences in the immunoreactivity of the individual cells. Bar -- 20 I.tm.

Fig. 12. Human testis. Immunocytochemical visualization of cGME A small intertubular vessel with stronger immunoreactivity in the cytoplasm of the smooth muscle cells (arrow) and a lower staining intensity in some endothelial cells. Bar = 20 txm.

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Fig. 14. Western blot analysis for the alpha-I subunit of the soluble guanylyl cyclase (sGC)(bands at approximately 82 kDa) in cytosolic fractions of seminiferous tubules and testicular blood vessels. Signals representing the antigen are marked by arrow. Control = rat brain extracts.

to possess morphological and functional similarities with neuroendocrine and nerve cells (Davidoff et al., 1993, 1996), it is thinkable that, with respect to the NO system, Leydig cells may fulfil nerve cell-specific functions. The mechanisms by which testicular NO synthases are activated in vivo, are still unclear. An autocrine activation of NOS-I by glutamate was suggested in human Leydig cells (Davidoff et al., 1995) and an induction of NOS-II by cytokines and LPS was shown in Leydig (Tatsumi et al., 1997), Sertoli (St6phan et al., 1995) and peritubular cells (Bauche et al., 1998) of the rat. Moreover, photoperiod (light-darkness) dependence of NOS activity in hamster Leydig cells was described (Angelova et al., 1996).

2.1.2. The spermatozoa A series of studies provided evidence for an influence of NO on sperm motility. High concentrations of NO were shown to inhibit motility (Rosselli et al., 1995; Weinberg et al., 1995; Zini et al., 1995; Nobunaga et al., 1996; Perera et al., 1996; Zhang and Zheng, 1996), whereas low concentrations resulted in opposite effects (Hellstrom et al., 1994; Zhang and Zheng, 1996). It was speculated (Rosselli et al., 1998) that low concentrations of NO ('physiological conditions') might protect against superoxide-mediated reduction of sperm motility, whereas high concentrations might inhibit sperm motility by toxic reaction products 286

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Fig. 15. Schematic presentation of the presumed NO systems in Leydig cells, testicular blood vessels and seminiferous tubules. Possible sites of NO production (occurrence of NOS) and activity (occurrence of sGC) are indicated. Physiological effects may include an influence on testosterone production by Leydig cells, dilatation of testicular blood vessels and relaxation of peritubular myofibroblasts. Within seminiferous tubules, a further system of NO production and activity, whose function is yet unknown, has been described in Sertoli cells (not shown). Since NO can freely diffuse across membranes (indicated bv + signs), the different systems may influence one another.

of NO primarily occurring under 'pathological conditions" (e.g. infections). However, NO concentrations in most of the studies (exception is Rosselli et al., 1995, up to 0.25 mM sodium nitroprusside) were chosen in analogy to previous experimental NO studies. Alternatively, an involvement of the cGMP pathway in the NO-induced inhibition of sperm motility is thinkable. In this context, a cyclic nucleotide-gated cation (CNG) channel was identified in bovine spermatozoa (Weyand et al., 1994; Wiesner et al., 1998), which has been shown to serve as a Ca 2+ entry pathway and to be opened by cGMP in vivo (Wiesner et al., 1998). Weinberg et al. (1995), however, were not able to inhibit sperm motility by cGMP analogues, though NO donors were effective. Moreover, convincing results demonstrating cGMP production by a sGC in spermatozoa are missing. In addition, NO was suggested to increase sperm capacitation (Zini et al., 1995), acrosome reaction (Viggiano et al., 1996; Revelli et al., 1999) and the zona pellucida binding ability of human spermatozoa (Sengoku et al., 1998). At the moment it is unclear, which function of NO predominates in vivo under physiological conditions. Furthermore, it is still a matter of debate, whether (in vivo) NO effects on sperm function may be due to NO produced by the upper female genital tract interacting with the male gametes during their way toward the oocyte or whether NO is synthesized by spermatozoa. Zini et al. (1995) and Schaad et al. (1996) did not find any NOS activity in human sperm. Since the latter authors exerted that human native semen and seminal plasma inhibit NOS activity as assayed on rat brain cytosolic fractions, it is thinkable that sperm proteases, released by sperm preparation, may inhibit both endogenous and exogenous NOS activity. A series of other studies suggested presence and activity of NO synthases within spermatozoa (Rosselli 287

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et al., 1995; Herrero et al., 1996; Lewis et al., 1996: Perera et al., 1996; O'Bryan et al., 1998; Revelli et al., 1999). Both, NOS-I (Herrero et al., 1996; Lewis et al., 1996) and NOS-III (Lewis et al., 1996; O'Bryan et al., 1998) have been detected by immunohistochemistry. But in a recent Western blot investigation it was convincingly shown that only the endothelial NOS-III is present in spermatozoa (Revelli et al.+ 1999). These authors also demonstrated that endogenous NOS activity may be increased by a protein-enriched chromatographic fraction of human follicular fluid. Others suggested progesterone-induced stimulation of sperm NOS (Herrero et al.+ 1998). Nevertheless, it remains obscure that within the testis this enzyme was not detectable in normal germ cells by former immunohistochemical studies (Zini et al., 1995). But recently we were able to demonstrate NADPH-d activity in spermatids of the mouse testis (Fig. 16).

2.1.3. The epididymis NOS activity was described in the epididymis of different species (Ehr6n et al., 1994; Burnett et al., 1995). Substantial levels of NOS activity were measured in all three regions of the rat epididymis. The cauda of this organ was found to be particularly NOS-rich, containing more than seven times the amount of NOS activity in testis (Burnett et al., 1995). After castration of rats, NOS activity was shown to be significantly reduced by 88%, 73% and 54%, respectively in the caput, corpus and cauda epididymidis, indicating an involvement of androgen on epididymal NOS activity (Chamness et al., 1995). All isoforms of NOS were described. NOS-I (Burnett et al., 1995), NOS-II (Wiszniewska et al., 1997) and NOS-III (Zini et al., 1996) were found to be present in epithelial cells of the epididymal duct. Epithelial NOS-II mRNA was inducible by LPS in vivo and in vitro (Wiszniewska et al., 1997). Whereas NOS-I-positive cells were predominantly seen in the cauda (Burnett et al., 1995), NOS-III staining was more intense in the proximal epididymis (Zini et al., 1996). Our own studies also suggest the presence of NOS in the ductuli efferentes. Both NOS-I immunoreactivity and NADPH-d activity were found in ciliated epithelial cells (Figs. 17 and 18). The physiological relevance of NOS in epithelial cells of the epididymis is still unknown. On the one hand, NO may diffuse out into the tubule lumen affecting nearby spermatozoa in a paracrine function, on the other hand, NO may influence the function of epithelial cells themselves or of surrounding smooth muscle cells in context of sperm transport. In this context, it is of interest that we were able to demonstrate the NO-binding enzyme sGC, cGMP

Fig. 16. Mouse testis. NADPH-d enzyme activity in the middle piece of the tail of elongated spermatids. The arrows point to negative heads of the spermatids. Bar = 20 p.m. Fig. 17. Mouse epididymis. NADPH-d enzyme activity in the efferent ductules. The strongest staining intensity is seen in the basal and the apical part of the ciliated cells. Also the middle piece of the spermatozoa within the lumen of the tubule is positive. Bar = 20 lain. Fig. 18. Human epididymis. NOS-I immunoreactivity in ciliated cells of the enlarged site branches of the efferent ductules. Bar = 20 Ism. Fig. 19. Rat penis. NADPH-d enzyme activity in thick nerve fiber bundles in the vicinity of blood vessels and in the submucosa beneath the epithelial lining of the urethra. Bar = 20 l~tm. 288

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and cGMP-binding protein kinase I both in epithelial cells and in smooth muscle cells (our unpublished results). Besides epithelial cells, NO synthases were also described in nerve fibers and vascular endothelial cells of the epididymis (Burnett et al., 1995: Dun et al., 1996; Zini et al., 1996). The supply of NOS-I-immunoreactive nerve fibers around the epithelium was found to be

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highest in the cauda (Dun et al., 1996). NOS-I-positive fibers were shown to arise mainly from neurons in the major pelvic ganglia and to co-express VIP but not tyrosine hydroxylase (Dun et al., 1996). 2.1.4. The vas deferens

Compared to the other male reproductive organs, the levels of NOS activity in the vas deferens are very high (Ehren et al., 1994; Burnett et al., 1995). NOS-I was described to occur in fine nerve fibers distributed throughout the smooth musculature of the rat vas deferens and in discrete plexuses coursing throughout the lamina propria (Burnett et al., 1995). NOS-I was also found in the vascular endothelium, whereas smooth muscle cells were devoid of staining. In the opposite, NOS-III was described in the epithelium along the length of the vas deferens of man (Zini et al., 1996). Interestingly, with respect to the histochemical demonstration of NADPH-d activity, Burnett et al. (1995) found a staining pattern similar to that detectable after use of anti-NOS-I antibodies in rat, whereas the pattern described by Zini et al. (1996) in man was comparable to that obtained by immunohistochemical analyses of NOS-III. With respect to NOS-containing nerve fibers, NOS was suggested to occur in cholinergic nerve fibers (Kalczyc, 1998) and to be co-localized with VIP in such postganglionic parasympathetic fibers (Ventura et al., 1998a). Moreover, in tissue segments taken from aged rats, the number of NOS-immunoreactive nerve fibers was suggested to be increased, whereas the vas deferens from immature rats showed no NOS immunostaining (Ventura and Burnstock, 1996). In man, NOS was suggested to be present in a number of both noradrenergic and non-noradrenergic (Jen et al., 1997) as well as in cholinergic (Sjostrand et aI., 1998) nerve fibers. A series of studies investigated the physiological role of NO in the vas deferens of different species. Ventura and colleagues found that the NO donor sodium nitroprusside may enhance electrically evoked contractions of the guinea pig vas deferens (Ventura et al., 1998b) and Postorino et al. (1998) discussed NO as a modulator of sympathetic transmission. Other investigations provided evidence against nitrergic neuromodulation of the ductus deferens in rat (Ventura and Burnstock, 1997) and mouse (Souilem et al., 1998). Taken together, a great number of studies dealing with the NO system in the vas deferens have been performed. However, neither the exact localization of this enzyme, nor its physiological relevance have been described convincingly. Most of the studies, even by the same group, remain contradictory. 2.1.5. The seminal vesicle

NOS activity within the seminal vesicle seems to be characterized by great species-specific differences. Ehr6n et al. (1994) described very high activity in man, whereas in rat only little NOS activity was found (Burnett et al., 1995; Chamness et al., 1995). After castration, NOS activity was shown to increase significantly (Chamness et al., 1995). NOS was localized to the epithelial layer (Burnett et al., 1995; Sjostrand et al., 1998) and to different types of nerve fibers (Dail, 1996; Jen et al., 1997; Sjostrand et al., 1998). NO synthases were found within cholinergic fibers (Dail, 1996; Sjostrand et al., 1998) as well as in noradrenergic and non-noradrenergic fibers at different stages of postnatal development (Jen et al., 1997). It was suggested that glandular NO production might be a prerequisite for muscarinic fructose secretion in the seminal vesicle via a cGMP-independent pathway as studied in guinea pig (Ehr6n et al., 1997). 290

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2.1.6. The prostate High levels of NOS activity were described in the human prostate (Ehr6n et al., 1994) as well as in the dorsal prostatic lobe of the rat (Chamness et al., 1995), whereas in the ventral and lateral lobes no significant NOS activity was found in rat (Chamness et al., 1995). In the lateral lobe NOS formation seems to be inhibited by androgen, since castration of rats resulted in a significant increase of NOS activity. In the ventral and dorsal lobe, however, the same procedure was without any changes in NOS activity (Chamness et al., 1995). In the human prostate, histochemical NADPH-diaphorase staining and immunohistochemical analyses using anti-NOS-I antibodies revealed the existence of a dense nitrinergic innervation of glandular epithelium, fibromuscular stroma, and blood vessels (Bloch et al., 1997). NOS seems to be partly co-localized with acetylcholinesterase (Sjostrand et al., 1998). In benign prostatic hyperplasia the nitrinergic innervation was shown to be reduced (Bloch et al., 1997). Expression of NOS-III in the endothelium of prostatic vessels was described to change depending on the region of the vascular bed and the diameter of vessels, displaying a pronounced staining in most of the arteries (Bloch et al., 1997). Magee et al. (1996) have cloned from rat penile corpora cavernosa a cDNA coding for a novel NOS-I different from the cerebellar NOS-I (see below). They suggested that this new species might be the only NOS-I mRNA expressed in the prostate. Further studies, however, are necessary to elucidate the complex role of NO in this organ.

2.1.7. The penis In the penis, the NO/cGMP system has gained particular interest, since sildenafil (Viagra), an inhibitor of the cGMP-specific phosphodiesterase type 5, has been established recently for the treatment of erectile dysfunction. First evidence for an involvement of nitric oxide in penile erection was provided by Ignarro et al. (1990). They demonstrated endogenous production and release of NO within strips of rabbit corpus cavernosum in vitro after electric field stimulation. In vivo data indicating that the arginine/NO pathway mediates rabbit penile erection were subsequently published (Holmquist et al., 1991). Consistently, NO synthesizing enzymes were demonstrated both in rat (Burnett et al., 1992) and in human (Burnett et al., 1993) penile nerve fibers innervating the corpus cavernosum and plexuses found in the adventitia of penile arteries (Fig. 19). Most of the NO-synthesizing neurons in the rat were shown to originate in the major pelvic ganglion, but also sensory NOS neurons occurred, whereas sympathetic postganglionic neurons were missing (Vanhatalo et al., 1996). The NO synthase described in neuronal penile structures is not completely identical to cerebellar NOS-I. Magee et al. (1996) cloned from the rat penile corpora cavernosa a cDNA coding for a novel NOS-I differing from the cerebellar NOS-I by the presence of a 102 nucleotides stretch and other features. This NOS-I was shown to be the only neuronal NOS expressed in the rat penis (Magee et al., 1996). However, mating b~havior and erectile function remained unchanged in NOS-I-knockout mice, but were blocked by the NOS inhibitor L-NAME (Burnett et al., 1996). Simultaneously, an up-regulation of NOS-III in these animals was observed, indicating that both NOS-I and NOS-III are capable of regulating NO-dependent penile erection (Burnett et al., 1996). NOS-III is present at high levels in endothelial cells of penile vasculature and especially of helicine arteries (Burnett et al., 1996; Bloch et al., 1998) as well as in cavernosal smooth muscle cells (Bloch et al., 1998). The significance of this enzyme for erectile function under physiological conditions, however, 291

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remains obscure. In addition to these isoforms, NOS-II is also expressed in cavernosal smooth muscle cells (Garban et al., 1997). Cloning of rat and human penile NOS-II indicated that the coding regions showed several amino acid differences from their analogous isoforms in non-penile tissues (Garban et al., 1997). Subsequently the authors found that local injection of NOS-II inducers corrected the erectile dysfunction observed in old rats. Though erectile dysfunction is associated with reduction in NOS-I activity, the other NOS isoforms are capable of replacing this enzyme in experimental studies, suggesting interesting therapeutical perspectives. Meanwhile, it is established that the influence of androgens on erectile function might be mediated by the NO-cGMP pathway (Lugg et al., 1996; Penson et al., 1996; Zvara et al., 1996; Schirar et al., 1997). Castration of rats and the anti-androgen flutamide (Chamness et al., 1995; Lugg et al., 1996: Penson et al., 1996) resulted in reduced NOS-I and NOS-III activity. It is now established that penile erection is completely dependent on androgens, most likely because of their role in the maintenance of penile NOS activity (Penson et al., 1996). Moreover, the described effects of N-methyl-D-aspartic acid (Melis et al., 1994a), apomorphine (Melis et al., 1994b), oxytocine (Melis et al., 1994b), acetylcholine (Champion et al., 1997) and dopamine (Chang et al., 1996) on the erectile function are NO-dependent. Adrenalectomy was also shown to affect penile erection via NO (Penson et al., 1997). As compared to young rats, NOS activity and NOS-containing nerve fibers decreased in old rats (Garban et al., 1995; Carrier et al., 1997). Erectile dysfunction occurring in diseases such as diabetes was shown to be accompanied by a decrease in NOS-I content and activity (Vernet et al., 1995; Rehman et al., 1997). The significance of NO for penile erection is doubtless (Burnett, 1995), and NO donors have already been used in the context of erection disorders (Stief et al., 1992). However, the clinical success in the treatment of erectile dysfunction is due to mechanisms down-stream the production and binding of NO. NO activates guanylyl cyclase and induces intracellular cGMP synthesis in erectile trabecular smooth muscle cells. Increased cGMP levels reduce intracellular Ca 2+ concentrations, inhibiting smooth muscle contractility and thereby initiating the erectile response. The phosphodiesterase-5 (PDE-5) is the predominant enzyme responsible for cGMP hydrolysis in cavernosal smooth muscle cells. Activation of PDE-5 terminates NO-induced cGMP-mediated smooth muscle relaxation, resulting ultimately in restoration of basal smooth muscle contractility and penile flaccidity. Sildenafil is a reversible and potent PDE-5 inhibitor that blocks cGMP hydrolysis effectively. Under conditions of excessive adrenergic tone or impaired neurovascular status, following sexual stimulation, sildenafil acts to enhance and prolong NO-mediated smooth muscle relaxation, resulting in improved penile erection in men with erectile dysfunction (for review see Goldstein et al., 1998, and Moreland et al., 1999). 2.2. THE FEMALE GENITAL TRACT 2.2.1. The ovary

Evidence that NO participates in ovulatory processes has been provided by Shukovski and Tsafriri (1994), who showed an inhibition of ovulation in the rat following administration of NOS inhibitors in vivo. These results were confirmed repeatedly (Bonello et al., 1996). In rabbit ovaries, L-NAME significantly reduced the percentage of large follicles that ovulated in response to hCG, and NO donors were found to induce follicle rupture (Hesla et al., 1997: 292

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Yamauchi et al., 1997). In agreement, NOS-III-knockout mice showed a significant reduction in ovulatory efficiency compared with wild-type females (Jablonka-Sharif and Olson, 1998). These data suggest that the ovarian NO system is necessary for follicle rupture during ovulation. Increases in NO synthesis with follicular maturation during the menstrual cycle were described both in normal subjects (Rosselli et al., 1994) and in female rats (Chatterjee et al., 1996). In addition, investigation of ovaries from NOS-III-knockout mice showed that fewer oocytes had entered metaphase II of meiosis, and that a greater percentage remained in metaphase I or were atypical (Jablonka-Sharif and Olson, 1998). These results suggest that NO might also be a modulator of oocyte meiotic maturation. In the last few years it was repeatedly shown that within the rat ovary NOS-II and NOS-III, but not the neuronal NOS-I, are expressed (Van Voorhis et al., 1995; Zackrisson et al., 1996; Jablonka-Shariff and Olson, 1997). Van Voorhis et al. (1995) localized NOS-II in the granulosa cells of primary, secondary and tertiary follicles of rat ovaries, whereas NOS-III was demonstrated in endothelial cells. In a subsequent report (Jablonka-Shariff and Olson, 1997) it was shown that NOS-III is not only present in endothelial cells, but also in the theca cell layer, ovarian stroma, the surface of oocytes, during follicular development additionally in mural granulosa cells and after ovulation within cells of the corpus luteum. These authors localized NOS-II to the. theca cell layer, stroma, the external layers of the developing corpus luteum as well as in the entire degenerating corpus luteum. In contrast, NOS-II was barely detectable by Zackrisson et al. (1996) both in follicles and in the corpus luteum. Whereas NOS-III levels were shown to increase during follicle growth, no changes in levels of ovarian NOS-II were visible (Jablonka-Shariff and Olson, 1997). Despite the discrepancies among the single morphological studies, it is evident that within the ovary NO is produced, at least in part, in the organ-specific hormone-producing cells. In agreement, expression of NOS-II and NOS-III was demonstrated in isolated cells obtained from rat ovarian follicles and corpora lutea at all stages of development (Van Voorhis et al., 1994; Olson et al., 1996). Endothelial and inducible NOS isoforms were observed in separate cells (Olson et al., 1996). In these isolated ovarian cells, NO was shown to inhibit estradiol synthesis in an autocrine manner independent of cGMP by directly inhibiting aromatase (Van Voorhis et al., 1994; Olson et al., 1996). Consistently, a higher concentration of estradiol was observed in NOS-III-knockout mice compared with wild-type mice (Jablonka-Sharif and Olson, 1998). The plasma progesterone levels, however, remained unchanged. In addition, there are also several lines of evidence for an involvement of NO on ovarian regulation of prostaglandin, phospholipase A2 and plasminogen activator levels, respectively (Bonello et al., 1996; Ahsan et al., 1997; Hurwitz et al., 1997: Kol et al., 1997a; Motta et al., 1997; Yamauchi et al., 1997). In this context, induction of NOS-II by interleukin-ll3 and TNF (both in vivo and in isolated ovarian structures) has repeatedly been reported (Bonello et al., 1996; Ahsan et al., 1997; Brunswig-Spickenheier and Mukhopadhyay, 1997; Hurwitz et al., 1997). Furthermore, NO mediates IL-113-induced expression of glucose transporter 3 (Kol et al., 1997b). Glucose increases during midcycle. Therefore, it is thinkable that NO and glucose facilitate follicle development in a complementary fashion. Since folliculogenesis involves the participation of both growth and programmed cell death, it was suggested that NO, known to regulate both of these processes, might also be involved in such processes (see Rosselli et al., 1998). 293

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2.2.2. The Fallopian tube There are only few reports dealing with the function of NO in oviducts. Grozdanovic et al. (1994) were not able to detect NOS-positive nerve fibers in the Fallopian tube. However, NO may be involved in a specific relaxation system within this organ, since basal (Ekerhovd et al., 1997) and endothelin-induced (Rosselli et al., 1994) contraction increased in the presence of the NOS inhibitor L-NAME. NOS-II and NOS-III were described in this organ (Bryant et al., 1995; Chatterjee et al., 1996; Rosselli et al., 1996, 1998). NOS-III was found in mucosal epithelium and muscular wall (Chatterjee et al., 1996; Rosselli et al., 1998). NOS-III immunostaining was described to be maximal during pro-estrus and estrus; NOS activity, however, was found to be relatively low during pro-estrus (Rosselli et al., 1998). NOS-II was induced by interleukin-l[5 and the staining pattern of NOS-II-positive cells differed dependent on certain pathological conditions (Rosselli et al., 1998). In general, NOS-II was found among epithelial and smooth muscle cells.

2.2.3. The uterus Nitric oxide seems to play crucial functional roles in the nonpregnant and pregnant uterus. The dynamical structural and functional changes of the uterus during menstruation, pregnancy and pathology stimulated the interest of numerous scientists on this topic (Dong et al., 1996; Norman and Cameron, 1996; Sladek et al., 1997: Garfield et al., 1998; Rosselli et al., 1998; Yallampalli et al., 1998, among others). Variable aspects of NOS isoform distribution and NO production and function were recognized and discussed. In general, the following functional aspects have been considered in the numerous studies: the significance of NO for the biology of the endometrium; the myometrial NOS and the NO contribution to the relaxation of the uterus musculature during pregnancy, before term and at labor; the NO function during implantation of the embryo as well as the role of NO in the regulation of the uterine vascular tone, uteroplacental blood flow and vessel function; the pathogenesis and pathology of some abnormal conditions and diseases. In the uterus of different species all three main isoforms of NOS seem to be constitutively expressed. RT-PCR studies have shown that in the rat uterus the NOS-II mRNA was predominant (Ali et al., 1997). These authors established that during pregnancy the uterine NOS-II mRNA increased, whereas on day 22 (before labor) it decreased and this tendency was prolonged during labor and term. In contrast, cervical NOS-II mRNA was low until delivery and started to increase dramatically during labor. Immunocytochemical detection of the three NOS isoforms confirm their existence in the structures of the uterus (Dong et al., 1996) and show that NOS-II immunoreactivity increased during pregnancy and decreased during delivery (Garfield et al., 1998). In contrast, the NOS-III levels do not change during pregnancy and the NOS-I isoform was found only during the nonpregnant stage of the rat uterus. Presumably, the increase of the NOS-II expression during pregnancy may be important in maintaining uterine quiescence. Expression of NOS-II mRNA in rat uterus after in vivo stimulation by LPS showed a significant increase in contrast to the low level in the control animals (Nakaya et al., 1996). The authors presume a prgtective function of NO against micro-organisms in the nonpregnant uterus. Immunohistochemical studies have localized NOS-III in the endothelium of uterine vessels and the NOS-III in myometrial and vascular smooth muscle and in decidual epithelium (Riemer et al., 1997). Immunoblotting studies, performed by the last authors, showed that expression of NOS-III decreased nearly fivefold, whereas NOS-III declined twofold in laboring rats at term (Bansal et al., 1997). 294

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Early studies found NOS-I-immunoreactive nerve fibers and ganglion cells in the reproductive tract of female mouse and established a substantial difference between the NO innervation of the mouse uterus horns and the uterus corpus and cervix (Grozdanovic et al., 1994). The most abundant nitrergic innervation was seen by these authors in the uterine cervix and vagina. In a parallel study NADPH-d-positive epithelial cells, luteal and interstitial cells of the ovary and extravascular granulocytes were devoid of NOS immunoreactivity (see Matani et al., 1997). NOS-I-exhibiting nerve fibers in the rat uterus were specified as parasympathetic with origin from neurons in the pelvic paracervical ganglia and some sensory fibers originating from neurons in different parts of the spinal cord and sacral dorsal root ganglia (Papka et al., 1995). Furthermore, the parasympathetic NOS-I-positive neurons co-expressed VIP, NPY and acetylcholinesterase, whereas in the sensory nerves calcitonin gene-related peptide and substance P were found. These results were extended with the finding that inferior mesenteric ganglia also contained uterine-related neurons (Papka et al., 1996). In the human and rat uterus NOS-III and NOS-II mRNA and NOS mmunoreactivity have been found in endometrial glandular epithelium and to a lower degree in decidual stromal cells (Chatterjee et al., 1996; Telfer et al., 1997; Cameron and Campbell, 1998). Although the functional significance of these localizations is not clear, the authors hypothesize a role of NO in initiation and control of menstrual bleeding, the inhibition of platelet aggregation, myometrial relaxation and maintaining of myometrial quiescence as well as relaxation of the nonpregnant myometrium (Telfer et al., 1995). Recent findings suggest that NO is a potent inhibitor of spontaneous contractile activity in the nonpregnant uterus. Moreover, for this relaxation an increase in the cGMP concentration was not required (Bradley et al., 1998). These results are in agreement with data showing cGMP-independent NO-induced inhibition of contractile activity of the nonpregnant human myometrium through involvement of K + ATP channels (Modzelewska et al., 1998). The vast majority of the studies on the NO activities in the uterus is devoted to the events accompanying pregnancy, preterm, term and labor events. The main significance of NO during pregnancy seems to be its ability to relax the myometrium to maintain the uterine quiescence (Buhimschi et al., 1995a; Kuenzli et al., 1998; Rosselli et al., 1998; Yallampalli et al., 1998). NOS-II mRNA in the rat uterus is enhanced during pregnancy and decreased during labor and postpartum (Ali et al., 1997). Furthermore NOS-II is up-regulated by progesterone and down-regulated by estrogens and prostaglandins (Dong et al., 1998a). However, it seems that NO can also increase uterine contractions of pregnant rats by enhancing uterine prostaglandin (PGE) synthesis (Motta et al., 1998). The next potent myorelaxant is the calcitonin gene-related peptide (CGRP) which inhibits uterine spontaneous contraction in rats during pregnancy but not during labor and postpartum. This effect of CGRP appears to be modulated by NO in the rat uterus (Dong et al., 1998b). Also adrenomedullin, a peptide produced by placental trophoblast cells, may directly or indirectly affect contractility of myometrium during pregnancy (Di Iorio et al., 1998a). An important event at the end of pregnancy is the decrease of the NO concentration in the uterine corpus and the significant increase in the cervix. A special feature of the uterine cervix is the expression of all three NOS isoforms (Buhimschi et al., 1996). There is strong evidence that in contrast to the uterine corpus the NOS-II mRNA in rat cervix was low until delivery (day 22) when it increased and was dramatically enhanced during parturition (Buhimschi et al., 1996; Ali et al., 1997). Apparently, NO is involved in the control of the conditioning of uterus and the cervix for labor. As already discussed, in the corpus, NO in concert with progesterone, inhibits uterine contractility. At term, a decrease in the production of NO by the 295

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uterus and the placenta was established which allows the initiation and progress of labor. In contrast, NO in the cervix increases at the end of pregnancy and is most probably involved in the cervical ripening by activation of matrix rnetalloproteases (Garfield et al., 1998; Rosselli et al., 1998). However, studies in humans showed significant differences. In contrast to animal studies, Thomson et al. (1997) showed a decrease in NOS activity and conclude that NO may not be involved in the onset of parturition at term. An explanation of this discrepancy may be provided by experiments performed with guinea pigs. In the guinea pig, in which cervical ripening resembles the inflammatory reaction in humans, local application of the NO donor SNP results in similar changes, but does not induce labor in pregnant animals (Chwalisz et al., 1997). In the final phase of labor, uterine contractions caused by oxytocin and probably by prostaglandin release may be also regulated by NO (Chaud et al., 1997). Further evidence for the importance of NO in gravity is provided by experimental studies with inhibition of NO synthesis in combination with onapristone on pregnant rats, resulting in preterm labor (Yallampalli et al., 1996). Moreover, chronic inhibition of NO in pregnant rats simulated preeclampsia-like conditions of high blood pressure (Buhimschi et al., 1995b), providing indirect evidence for a role of NO in the regulation of blood pressure during pregnancy. Another aspect of possible NO activity in the uterus is its role in inducing endometrial receptivity and the process of implantation (Biswas et al., 1998). NOS activity increases before embryonic implantation in rats. To achieve a successful embryo implantation an enhanced NO production is needed. The NO production appears to be controlled by estradiol which modulates the NOS activity during nidation (Novaro et al., 1997). Interestingly, the constitutive NOS-III isoform was found to increase gradually in the preimplantation phase, whereas the NOS-II isoform increased just at the beginning of implantation (day 5); after that the activity of both NOS isoforms decreased (Novaro et al., 1997). However, higher NO concentrations inhibit both development of the mouse embryo in vitro and embryo implantation in vivo (Barroso et al., 1998). Other factors, such as the platelet-activating factor (PAF) may also have a role in implantation by stimulating prostaglandin production. As evidenced by Chaud et al. (1998), NO represents an important mediator of the interaction between PAF and the prostaglandins. Another target of NO action is the uteroplacental vasculature. Numerous studies concern the regulation of the uterine artery but the results are to some extent controversial. In the mechanism of vasorelaxation also other factors are involved. Jain et al. (1999) provided evidence for the vasorelaxant role of the corticotropin-releasing factor (CRF), released mainly from the placenta, on the rat uterine artery. CRF is increased during human pregnancy and causes vasorelaxation by receptor-operated, endothelium-dependent and endothelium-independent pathways. In the uterine artery the CRF-endothelium-dependent relaxation is mediated by the NO/cGMP pathway and one endothelium-derived hyperpolarizing factor (Jain et al., 1999). On the other hand, although increased NO production exists in uterine arteries during pregnancy, experimental studies failed to show any contribution of endothelin-1 and NO to the augmented tension to depolarization and receptor-operated stimulation of vascular smooth muscle cells in rat uterine arteries during pregnancy (Wight et al., 1998). Uterine arteries, but not systemic vessels, respond to ovarian or exogenous estrogens with vasodilatation and elevated production of NO from NOS-III, even after LPS treatment (Vagnoni and Magness, 1998; Vagnoni et al., 1998), thus providing evidence that estrogen plays a role in regulating uterine artery responses to LPS. In the human uterine artery, pregnancy enhances acetylcholine-induced nitric oxide synthesis and release (Nelson et al., 1998). Estrogen may be also important in the hyperemic 296

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response of the uterus during estrus in nonpregnant sheep (Veille et al., 1996). The action of VIP on the human uterine artery is apparently also mediated by NO generated by the endothelial NOS-III (Jovanovic et al., 1998). NO participates also in the regulation of the tone of uterine microvessels of rats. As Saha et al. (1998) established, NO and dilator prostaglandins are involved in these regulations. In fetal membranes, Dennes et al. (1997) found expression of both NOS-III mRNA and NOS-II mRNA in human amnion, chorion-decidua, and placenta by means of semiquantitative RT-PCR. The authors presume that NO in fetal membranes may act directly or indirectly to inhibit myometrial contractility.

2.2.4. The vagina As already mentioned, the uterine cervix and the vagina are the structures with the most abundant nitrergic innervation in the female mouse (Grozdanovic et al., 1994). The nerve fibers were found to run parallel to the smooth muscle bundles and beneath the epithelium. In addition, NOS-I-immunoreactive nerve cell bodies were also found in the vaginal muscular wall. Also clusters of NOS-positive neurons were seen in Frankenh~iuser's paracervical ganglion at the cervico-vaginal junction. In all these structures NOS-I immunoreactivity and NADPH-d enzyme activity co-localize. Numerous NOS-I-immunoreactive nerves and nerve fibers, partially co-stained with VIP or SP, were found in the vagina of cow and pig (Majewski et al., 1995). A dense nitrergic innervation and VIP-positive nerve fibers and perikarya were observed in the musculature of the vagina of the hen (Costagliola et al., 1997). Also the human vagina displays NOS-immunoreactive nerves densely distributed around the deep arteries and veins and the blood vessels in the propria as well as in form of a subepithelial plexus (Hoyle et al., 1996). Vaginal NOS-III immunoreactivity was found in the epithelium and the smooth muscle cells and showed maximal staining intensity during estrus and pro-estrus (Chatterjee et al., 1996). The epithelial occurrence of the enzyme is probably related to the NO stimulation of vaginal secretion. Biochemical comparative studies on the NOS activity in cerebellum, uterus and vagina have shown that NOS in both cytosolic and particulate fractions of these organs was highly calcium-dependent and the activity in the cytosolic fraction was approximately four times higher (Batra and A1 Hijji, 1998). Furthermore, the concentration of NOS was highest in the cerebellum followed by vagina and uterus which is in accordance with the morphological observations of Grozdanovic et al. (1994). The vaginal cytosolic NOS activity responds with down-regulation after estrogen treatment (Batra and A1 Hijji, 1998). Estrogen appears to play a critical role also in concomitantly regulating vaginal NOS expression and apoptosis in nerves, smooth muscle cells, vascular endothelium and epithelium of the rat vagina, which may be of importance for the changes in the vagina during the menopause (Berman et al., 1998).

2.2.5. The placenta and umbilical artery Expression, localization and function of NOS isoforms and NO in the placenta were very intensely studied in the past decade (Poston, 1997). Most attention was paid on the placental vascular bed as a very important system ensuring a low-resistance circulation in which blood flow is determined by the fetal cardiac output (Wieczorek et al., 1995). A striking feature of the placenta is the lack of nerve supply. Therefore. the regulation of the vascular resistance is under the control of different vasoactive humoral factors. As in other organs NO is considered 297

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to exert tonic reduction in vascular resistance (Poston, 1997). As mentioned above, the pathogenesis of preeclampsia and eclampsia reflects changes in the regulatory mechanisms of placental vasculature (Myatt et al., 1996, 1997; Lee et al., 1997). Using semiquantitative RT-PCR, expression of NOS-III mRNA and NOS-II mRNA were revealed in human chorion, chorion-decidua and placenta by Dennes et al. (1997). In the rat placenta NOS-III expression seems to be constitutive (Casado et al., 1997). However, in contrast to NOS-III, RT-PCR studies on syncytiotrophoblast of human term placenta in culture failed to show NOS-II transcripts neither basal, nor following induction by interleukin-1, tumor necrosis factor-alpha, interferon-gamma and LPS (Lyal et al., 1998). Probably, the syncytiotrophoblast in culture conditions loses its ability for expression of NOS-II. Studies performed by Myatt et al. (1991) suggested that the human placental vascular tree is able to generate and to respond to NO which seems to be important in maintenance of basal vascular tone and attenuation of the action of different vasoconstrictors in placental circulation (Myatt et al., 1992). Biochemical measurements and Western blot analysis provide evidence that in the villous vasculature predominantly the NOS-III isoform exists (Myatt et al., 1993a). Enzyme histochemistry of NADPH-d and immunohistochemistry of NOS-III showed that the endothelial NOS isoform was located in the umbilical cord artery and vein endothelium and the endothelium of the stem villous vessels. In the terminal villi immunostaining was seen within the syncytiotrophoblast (Myatt et al., 1996; Sladek et al., 1997). This study shows that NOS-III appears to be present in the vasculature which possess smooth muscle cells (Myatt et al., 1993b). However, in the first trimester of development NOS-III immunoreactivity was found in syncytiotrophoblast and fibroblast layers of the amnion (Eis et al., 1995). Moreover, the study of differentiation of early placenta in culture showed that the transformation of cytotrophoblast into syncytiotrophoblast parallels with NOS-III expression. These results are in agreement with studies of Sahin-Toth et al. (1997) who showed that NOS-III is responsible for approximately 90% of NO synthesizing activity in first trimester placenta. As evidenced, tetrahydrobiopterin activates the microsomal (not cytosolic) NOS-III in the primordial placenta by promoting its subunit assembly in the membrane. The activity of NOS has been established to be higher in normal mid and late pregnancy and to decrease with the advancement of pregnancy (Chen and Zheng, 1996). In agreement with these results are the findings of Ariel et al. (1998) who found in the human placenta during the first trimester of pregnancy NOS-III immunoreactivity in the intermediate cells of the cell columns of anchoring villi, in trophoblastic cells at the implantation site and in the apical border of the syncytiotrophoblast, which may be related to a possible role in implantation and vascular invasion. In accordance, measurements of NOS-III activity and the cGMP concentration showed highest levels in the early gestation (Lees et al., 1998). However, this seems to be somewhat paradoxical, because the high NOS-III activity and cGMP levels coincide when umbilical artery pulsatility index is also high, and decrease as umbilical pulsatility decreases. Thus, the question arises whether NO effects during early pregnancy are vasodilatory or contribute to formation of new villous vessels. In women with complicated pregnancies with preeclampsia and/or intrauterine growth restriction, NOS-III mRNA was significantly higher (Nasiell et al., 1998) and high staining intensity in immunocytochemical visualization for NOS-III protein was observed in placenta accreta, partial and complete hydatiform moles, and choriocarcinoma (Ariel et al., 1998). It was presumed that these changes reflect some compensatory mechanisms in the disturbed uterine circulation. On the other hand, studies performed by Faxen et al. (1997) showed that the mRNA expression of endothelin-1, endothelin-A receptor and NOS-III in hydatiform moles did not vary from 298

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the control placentas and did not explain the increased frequency of preeclampsia in molar pregnancy. In a comparative study of the placenta of humans, rhesus monkeys, baboons and sheep expression of NOS-III was established only in the human syncytiotrophoblast (Zarlingo et al., 1997). The production of NO by syncytiotrophoblast of the human placenta was related to prevention of platelet and leukocyte aggregation in the intervillous space and adhesion to the trophoblast surface as well as with the possibility to mediate peptide hormone release from trophoblast. However, the exact functional significance of the syncytiotrophoblast NO remains to be elucidated. In human normotensive, preeclamptic placenta and placenta with intrauterine growth restrictions, NOS-II isoform was localized predominantly in the villous Hofbauer cells and in a part of the cases in syncytiotrophoblast and vascular endothelium (Myatt et al., 1997). The possible production of NO by NOS-II in the Hofbauer cells reflects their cytostatic or cytotoxic capabilities. In contrast, in the rat placenta NOS-II immunoreactivity was found in the trophospongial cell layer at the fetal-maternal interface, mostly within islands of vacuolated 'glycogen cells' and this immunoreactivity strongly decreased during labor (Purcell et al., 1997). Either species-related or antibody differences may account for the discrepancy in the localization of NOS-II in these species. NOS-I immunoreactivity and corresponding NADPH-d enzyme activity was observed only in the epithelial cells of the intraplacental yolk sac of mice by Matejvic et al. (1996). The authors speculate that this location with the narrow spatial relationship to large fetal blood vessels that enter the placental labyrinth from the chorionic plate and the possibility that NO, produced by the yolk sac, may affect the contractile state of the smooth muscle cells of these fetal vessels. Another topic of interest is the role of NO in the control of expression of the corticotropinreleasing factor (CRF) by the placenta. There is strong evidence that CRF plays an important role in human parturition. Studies by Ni et al. (1997) showed that exogenous NO inhibits CRF exocytosis by human trophoblast without affecting its biosynthesis. There is also evidence that NO is involved in the regulation of release of placental growth hormone during pregnancy (Roe et al., 1996). There are conflicting results concerning the effect of hypoxia on the placental NOS expression and hemodynamics. Using dually perfused human placental cotyledons Byrne et al. (1997) found that the reduction in basal NO release mediates hypoxic fetoplacental vasoconstriction. The opposite effects were established in differentiating human placenta. Exposure of differentiated human syncytiotrophoblast to hypoxia for 24 h significantly stimulates expression of NOS-III mRNA in comparison with trophoblast maintained in normoxia (Seligman et al., 1997). Thus, depending on the differentiation state a differing response to hypoxia exists in the human placenta. Studies on the importance of NO in the pathogenesis of preeclampsia are controversial (Buhimschi et al., 1995b, 1998; Ghabour et al.. 1995: Myatt et al., 1996, 1997; Lee et al., 1997; Palma-Gamiz, 1998). It became evident that some of the maternal symptoms of preeclampsia can be produced by uterine ischemia. Chronic nitric oxide synthase inhibition in rats also produces changes that resemble preeclampsia symptoms (Podjarny et al., 1999). Biochemical studies showed a significant reduction of human placental NOS activity in preeclampsia (Brennecke et al., 1997). In accordance, treatment of pregnant rats with the NOS inhibitor L-NAME resulted in light-microscopic and electron-microscopic structural changes in the placenta similar to those found in human preeclampsia (Osawa, 1996). However, plasma cGMP levels, NOS-III gene expression as well as NO synthesis were identical 299

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in human umbilical vein endothelial cells and placenta from normal and preeclamptic women (Boccardo et al., 1996). Also the NADPH-d enzyme activity was not changed in preeclamptic human placenta in comparison with normal ones and shear stress did not affect the production of NO metabolites, suggesting that trophoblast NOS-III may not contribute to the development of preeclampsia (Poranen et al., 1998). In addition, placental vessels obtained from normotensive and preeclamptic patients possessed the same sensitivity toward the relaxant effects of different nitric oxide donors (Gonzalez et al., 1997). Studies by Di Iorio et al. (1998b) in the human placenta reveal increased concentration of NO metabolites in amniotic fluid and positive NOS-III immunostaining in the placental villi that suggest an up-regulation rather than down-regulation of the L-arginine-NO system. An enhanced immunoreactivity for nitrotyrosine residues in endothelium of villous vessels, the perivascular muscle cells and villous stroma was observed in human placenta by Myatt et al. (1996) and considered as an indication for vascular damage, caused by peroxynitrite, that contributes to the increased vascular resistance. Decreased NO synthesis and activity was established in diabetic pregnancies that were accompanied with reduced blood flow and increased vascular resistance (Dollberg et al., 1997). A disturbed NO/endothelin-1 equilibrium may be also involved in the pathogenesis of preeclampsia because endothelin-I was found to be elevated in the serum of preeclamptic patients. In agreement, Leszczynska-Gorzelak et al. (1998) established that increased ET-I concentration resulted in a dysfunctional mechanism of negative feedback regulation of endothelin-1 production by cGMP or NO release in endothelial cells of arterial blood vessels of human placenta in culture. However, an involvement of endothelin-1 in the preeclampsia-like syndrome during pregnancy seems to exist only in the early phase of experimentally L-NAME-induced preeclampsia in rats (Wight et al., 1998). The umbilical artery is also the object of intensive studies in relation to NO and reactive oxygen radicals. On the one hand NO seems to influence the contractile status of the umbilical artery, and on the other, the contractile status of the umbilical artery seems to influence the expression of NOS and the production of NO from the placenta. NO may exert a strong relaxant effect on serotonin-induced vasoconstriction in the human umbilical artery (Okatani et al., 1996). Recent results suggest that superoxide potentiates vascular tension in the human umbilical artery, in part, by suppressing NO synthesis in the endothelial cells (Okatani et al., 1998). Vascular tension of the human umbilical artery was also stimulated by hydrogen peroxide and hydrogen peroxide potentiated the umbilical artery contraction induced by serotonin. This effect of hydrogen peroxide may be mediated by a suppression of NO and prostacyclin activity (Watanabe et al., 1996: Okatani et al., 1997a). This presumption is at odds with results reported by Izumi et al. (1996) who showed that NO and prostacyclin released from endothelium exert a vasorelaxant effect on the musculature of umbilical artery. The vasoconstriction action of hydrogen peroxide may be suppressed by melatonin, probably due to the ability of melatonin to scavenge the hydroxyl radical (Okatani et al., 1997b). Inhibition of NO activity in lamb fetoplacental circulation resulted in an increase in the umbilical artery flow velocity waveform systolic to diastolic ratio (Giles et al., 1997a) and placenta from women with abnormal umbilical artery flow velocity waveforms showed decreased mean NOS activity compared to normal ones (Giles et al., 1997b). The resistance index of the umbilical artery was found to be higher in preeclamptic compared to normotensive patients (Makino et al., 1997).

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3. ACKNOWLEDGEMENTS T h i s w o r k w a s s u p p o r t e d by g r a n t s f r o m the D e u t s c h e F o r s c h u n g s g e m e i n s c h a f t ( D a 4 5 9 / 1 - 1 " Mi 637/1-1).

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Rosselli M, Imthurn B, Macas E, Keller PJ, Dubely RK (1994): Endogenous NO modulates endothelin-I induced contraction in bovine oviduct. Biochem Biophys Res Commun 201:!43-148. Rosselli M, Dubey RK, Imthurn B, Macas E, Keller PJ (1995): Effects of nitric oxide on human spermatozoa: evidence that nitric oxide decreases sperm motility and induces sperm toxicity. Hum Reprod 10:1786-1790. Rosselli M, Dubey RK, Rosselli MA, Macas E, Fink D, Lauper U, Keller PJ, Imthurn B (1996): Identification of nitric oxide synthase in human and bovine oviduct. Moi Hum Reprod 2:607-612. Rosselli M, Keller PJ, Dubey RK (1998): Role of nitric oxide in the biology, physiology and pathophysiology of reproduction. Hum Reprod Update 4:3-24. Ruffoli R, Fabris FM, Giambelluca MA, Giannessi F (1997): Ultrastructural demonstration of the NADPHdiaphorase activity in peritubular myoid cells and fibroblasts of the lamina propria of the mouse seminiferous tubules, hz Vivo (hit J hi Vivo Res) 11:319-324. Saha PR, Alsip NL, Henzel MK, Asher EF (1998): Role of nitric oxide and cyclooxygenase products in controlling vascular tone in uterine microvessels of rats. J Reprod Fertil 112:211-216. Sahin-Toth M, Kukor Z, Toth M (1997): Tetrahydrobiopterin preferentially stimulates activity and promotes subunit aggregation of membrane-bound calcium-dependent nitric oxide synthase in human placenta. Mol Hum Reprod 3:293-298. Sakuma I, Togashi H, Yoshioka M, Saito H, Yanagida M. Tamura M. Kobayashi T. Yasuda H. Gross SS, Levi R (1992): NG-methyl-arginine, an inhibitor of L-arginine-derived nitric oxide synthesis, stimulates renal sympathetic tone?. Circ Res 70:607-611. Salom MG, Lahera V, Miranda Guardiola F. Romero JC (1992): Blockade of pressure natriuresis induced by inhibition of renal synthesis of nitric oxide in dogs. Am J Phvsiol 262:F718-F722. Sfinchez M, Menendes L, De Boto MJG, Hidalgo A (1996) Role of cyclic nucleotides in contraction induced by oxytocin in the testicular capsule of the rat in vitro. Pharmacology 53:296-301. Sandner P, Kornfeld M, Ruan X, Arendshorst WJ, Kurtz A (1999): Nitric oxide/cAMP interactions in the control of rat renal vascular resistance. Citz" Res 84:186-192. Sann H (1998): Neuronal subpopulations in autonomic ganglia associated with the chicken ureter: an immunohistochemical study. Cell Tissue Res 292:477-485. Schaad NC, Zhang XQ, Campana A. Schorderet-Slatkine S (1996): Human seminal plasma inhibits brain nitric oxide synthase activity. Hum Reprod I 1:561-565. Schindelmeiser J, MiJnstermann D, Mayer B, Holstein AF, Davidoff MS (1997): Occurrence of enzymes of free radical metabolism suggests the possible cytotoxic capacity of the transitional epithelium of the human ureter. Cell Tissue Res 287:351-356. Schirar A, Chang C, Rousseau JP (1997): Localization of androgen receptor in nitric oxide synthase- and vasoactive intestinal peptide-containing neurons of the major pelvic ganglion innervating the rat penis. J Neuroendocrinoi 9:141-150. Schmidt HHHW, Gagne GD, Nakane M, Pollock JS, Miller ME Murad F (1992): Mapping of neural nitric oxide synthase in the rat suggests frequent co-localization with NADPH diaphorase but not with soluble guanylyl cyclase, and novel paraneural functions lbr nitrinergic signal transduction. J Histochem Cvtochem 40:1439-1456. Schnackenberg C, Patel AR, Kirchner KA, Granger JP (1997): Nitric oxide, the kidney and hypertension. Clin Exp Pharmacol Phvsiol 24:600-606. Seligman SE Nishiwaki T, Kadner SS, Dancis J. Finlay TH (1997): Hypoxia stimulates ecNOS mRNA expression by differentiated human trophoblasts. Ann NY Acad Sci 828:180-187. Sengoku K, Tamate K, Yoshida T, Takaoka Y, Miyamoto T. Ishikawa M (1998): Effects of low concentrations of nitric oxide on the zona pellucida binding ability of human spermatozoa. Fertii Steril 69:522-527. Sharma AC, Sam AD 2rid, Lee LY, Hales DB, Law WR, Ferguson JL, Bosmann HB (1998): Effect of NGnitro-L-arginine methyl ester on testicular blood flow and serum steroid hormones during sepsis. Shock 9:416421. Shukovski L, Tsafriri A (1994): The involvement of nitric oxide in the ovulatory process in the rat. Endocrinology 135:2287-2290. Shulz PJ, Tayeh MA, Marletta MA, Raij L ( 1991 ): Synthasis of nitric oxide in rat glomerular mesangial cells. Am J Phvsiol 261 :F600-F606. Silvagno F, Xia H, Bredt DS (1996): Neuronal nitric oxide synthase-tJ., an alternatively spliced isoform expressed in differentiated skeletal muscle. J Biol Chem 271:11204-11208. Singh I, Grams M, Wang WH, Yang T, Killen R Smart A. Schnermann J, Briggs JP (1966): Coordinate regulation of renal expression of nitric oxide synthase, renin, and angiotensinogen mRNA by dietary salt. Am J Phvsiol 270:FlO27-F1037. 311

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Response of nitric oxide synthase to neuronal injury W. WU

1. I N T R O D U C T I O N Investigations leading to the identification of roles of nitric oxide (NO) in the body can be traced back to a study by Furchgott and Zawadzki (1980) who found that there is a diffusible molecule derived from the endothelial cells of blood vessels which mediates smooth muscle relaxation. Murad et al. (1978) first reported properties of NO which were later shown to be very much like those of the endothelium-derived relaxing factor (EDRF), the diffusible molecule that causes relaxation of smooth muscles of the blood vessels. The EDRF was later identified as NO (Palmer et al., 1987; Garthwaite et al., 1988; Ignarro et al., 1990), which is synthesized by NO synthase (NOS) (Bredt et al., 1991: Hevel et al., 1991; Lamas et al., 1992). In the past decade, three major isoforms of NOS have been found and purified: neuronal NOS (nNOS) (Bredt et al., 1991; Bredt and Snyder, 1990), inducible NOS (iNOS) (Hevel et al., 1991; Lowenstein et al., 1992; Xie et al., 1992), and endothelial NOS (eNOS) (F6rstermann et al., 1991; Lamas et al., 1992). Roles of NO/NOS have been extensively studied in three major systems: the immune system, vascular system, and nervous system under normal and pathological conditions (Bredt and Snyder, 1992; Moncada, 1992; F6rstermann and Kleinert, 1995; Garthwaite, 1995; Zhang and Snyder, 1995). Under normal conditions, NO plays an important role in many physiological functions of the nervous system, such as regulation of cerebral blood flow, memory formation, host defence against infections and tumors, modulation of neurotransmission and neuroendocrine function, etc. (Moncada, 1992; Szabo, 1996; F6rstermann et al., 1998). On the other hand, improper production of NO/NOS in the nervous system has been shown to be neurotoxic and involved in some pathological processes, such as ischemia, stroke, neuronal injury, and neurodegenerative diseases (Bredt and Snyder, 1992; Moncada, 1992; Szabo, 1996: Yun et al., 1997). It has been reported that NOS can be induced in many pathological conditions and excess NO may be, at least in part, responsible for the neurotoxicity in those conditions. Since NO has a very short life after it is synthesized, major insights into NO disposition have come from studies of NOS. Expression of NOS in the nervous system can be regulated under pathological conditions. In most cases, up-regulation of NOS is found. Increased expression of NOS has been observed in neurons following different types of injuries. Neuronal injuries that cause increased expression of NOS include the following. (1) Axonal injuries. NOS can be induced following a crush, ligation, axotomy, or root avulsion of the peripheral nerve, or after other traumatic neuronal injuries which cause damage to the axons (Wu, 1993; Wu et al., 1994c: Yu, 1994). Handbook of Chemical Neuroanatom~; Vol. ! 7: Functional NeuroanatonO ~,/the Nitric Oxide System H.W.M. Steinbusch. j. De Vente and S.R. Vincent. editors @ 2000 Elsevier Science B.V. All rights reserved.

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(2) Ischemic-hypoxic injury. Involvement of NOS is found in hypoxic neuronal injury of cultured neurons (Cazevieille et al., 1993), in brain slices (Wallis et al., 1992), in the injured rat striatum after middle cerebral artery (MCA) occlusion (Nakashima et al., 1995), and in the ischemia-reperfusion-injured retina (Hangai et al., 1996). NO plays an important role in the development of neuronal injury after global ischemia (Nakagomi et al., 1997; Samdani et al., 1997a). NOS knockout mice show smaller motor deficits than wild-type mice after MCA occlusion (Iadecola et al., 1997). (3) Excitotoxic neuronal injury. Stimulation of glutamate receptors causes an up-regulation of NOS in the affected neurons (O'Hearn et al., 1995). Both in vitro and in vivo studies have shown that NO/NOS directly mediates glutamate neurotoxicity (Dawson et al., 1991, 1994; Schulz et al., 1995). (4) Chemical injury to the nervous system. Up-regulation of NOS is observed in affected neurons following subcutaneous injection of methylmercury chloride (Himi et al., 1996), intrathecal administration of dynorphin (Hu et al., 1996), or systemic administration of capsaicin, a chemical which can deplete neuropeptides and cause degeneration of C-fiber afferent pathways (Vizzard et al., 1995a). (5) Neural degenerative diseases. NO may be involved in some human neuronal degenerative diseases (Sherman et al., 1992). It has been reported that NO promotes neuronal and oligodendrocyte damage in patients with multiple sclerosis (Xiao et al., 1996). Unexpectedly high levels of NOS have also been found in CNS tumor tissues in human (Cobbs et al., 1995). (6) Other injuries. Up-regulation of NOS is found in neurons when the normal axonal transport is interrupted by colchicine (Lumme et al., 1997; Vanhatalo et al., 1998). Temperature changes in the nervous system also causes induction of NOS (Saxon and Beitz, 1994). The present review summarizes the changes of NOS expression in response to neuronal injury. Up-regulation and de novo expressions of NOS in different regions of the nervous system from different species of animals of various ages are reviewed. Regulation and potential roles of injury-induced NOS in neuronal degeneration and regeneration are also discussed. Emphasis is focused on neuronal NOS and its response to neuronal injuries in in vivo models.

2. INJURY-INDUCED EXPRESSION OF NOS Distribution of NOS-containing (expressing) neurons in the nervous system has been widely studied under normal conditions (Vincent and Kimura, 1992; Blottner et al., 1995; Jaffrey and Snyder, 1995; Szabo, 1996). NOS is present in many locations throughout the central and peripheral nervous system. In adult animal, NOS-containing neurons are normally found in the cerebral cortex, striatum, hypothalamus, cerebellum, brain stem, spinal cord, and peripheral ganglia, etc. However, not all neurons in these nuclei or regions express NOS. In the cerebral cortex, for example, only about 1-2% of cortical neurons are NOS-positive. Pyramidal neurons in layer V of the neocortex normally do not express the enzyme. Purkinje cells in the cerebellar cortex, spinal motoneurons, and many other types of neurons do not normally express NOS either. Following neuronal injuries, however, expression of NOS is observed in many of those neurons that normally do not express this enzyme. The first study that dealt with the injury-induced expression of NADPH-diaphorase activity was reported by Gonzalez et al. (1987). Their study demonstrated that cervical vagotomy caused increased expression of NADPH-diaphorase (NADPH-d; years later shown to be identical to NOS; Hope et al., 1991) staining in the ipsilateral dorsal motor nucleus of the 316

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vagus nerve. Since then, injury-induced expression of NOS has been found in many other nuclei and regions throughout the nervous system, including the cerebral cortex (Kitchener et al., 1993), cerebellar cortex (Chen and Aston-Jones, 1994), hypothalamic nuclei (Wu and Scott, 1993), cranial nerve nuclei (Yu, 1994), nuclei forming the descending and ascending pathways (Wu, 1993; Wu et al., 1994a,b,c; Jin et al., 1997), spinal cord (Wu, 1993), dorsal root ganglion (H6kfelt et al., 1994), and autonomic nervous system (Vizzard et al., 1993, 1995b; Vizzard, 1997). In the present review, expression of NOS due to neuronal injuries is termed 'injury-induced' expression of NOS. It is important to note that injury-induced NOS in many neurons is the neuronal isoform of NOS, which is normally constitutively expressed within the nervous system. Injury-induced expression of NOS is observed in two major forms, the up-regulation of NOS expression and the de novo expression of NOS. Up-regulation of NOS expression means that expression of NOS increases after injury, either .in the numbers or staining intensity of NOS-positive neurons, in a given region where NOS is normally present. De novo expression of NOS means that NOS is induced after injury in a given region where NOS is not normally present. 2.1. UP-REGULATION OF NOS EXPRESSION IN INJURED NEURONS

2.1.1. Up-regulation of NOS expression in the neurohypophyseal system Although some neurons in the supraoptic and paraventricular nuclei of the hypothalamus normally express NOS (Fig. 1A,B), NOS activity increases significantly in these nuclei following hypophysectomy (see Fig. 1D-L; Wu and Scott, 1993; H6kfelt et al., 1994). In these studies, both the enzyme and the mRNA of the neuronal NOS increase dramatically (see Fig. 2; Scott et al., 1995; Lumme et al., 1997). Increased NOS staining is also observed in regenerating axons that are re-growing into the adjacent median eminence and the surface of the third cerebral ventricle (Wu and Scott, 1993; also see Fig. 1F,I). Similarly, NOS up-regulation in the paraventricular and supraoptic nuclei is also seen after salt loading (H6kfelt et al., 1994) or colchicine treatment (Lumme et al., 1997). In the anterior pituitary, NOS expression is increased after gonadectomy (Yamada et al., 1997).

2.1.2. Up-regulation of NOS expression in the nuclei of cranial nerves Increased expression of NOS is reported in the ambiguus nucleus, the solitary tract and nucleus, and the dorsal motor nucleus of the vagus nerve following vagotomy (Jia et al., 1994; Kristensson et al., 1994; Yu, 1994, 1997; Zhao et al., 1996).

2.1.3. Up-regulation of NOS expression in the spinal cord Some spinal interneurons and neurons in the dorsal horn are normally NOS-positive. NOS expression is increased in the dorsal horn (laminae I-III) following peripheral nerve injury (Hama and Sagen, 1994) or after transient spinal cord ischemia (Marsala et al., 1997). Traumatic injury in the rat spinal cord also up-regulates NOS expression in the dorsal horn of the lesion area (Wu, 1992; Sharma et al., 1996). Following hemisection of the spinal cord, numbers of NOS-positive spinal interneurons are significantly increased in the lesion area, especially on the lesioned side (see Fig. 3A; Wu, 1993). 317

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Response of nitric oxide synthase to neuronal injury

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Fig. 2. In situ hybridization staining showing expression of neuronal NOS in the supraoptic (arrows) and paraventricular nuclei (PVN) in normal control rats (A, B), and in experimental rats 2 weeks following hypophysectomy (C, D). The levels of NOS mRNA in the experimental animals are significantly greater than the control. OC = optic chiasm. Bar = I00 ~m.

2.1.4. Up-regulation of NOS expression in the peripheral nerve and ganglia There is a dramatic increase of neuronal NOS in many neurons in the dorsal root ganglion (DGR) following peripheral nerve axotomy as compared with the normal controls (see Fig. 3B,C; Verge et al., 1992; Zhang et al., 1993; H6kfelt et al., 1994). Systemic administration of capsaicin, a chemical which can deplete neuropeptide and cause degeneration of C-fibers, also significantly up-regulates NOS expression in DRG neurons (Vizzard et al., 1995a). Besides the DRG, up-regulation of NOS expression due to injury is also found 319

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Fig. 3. (A) Horizontal section of spinal cord from a rat 2 weeks following hemisection of the cord. The section was stained with NADPH-d histochemistry. Increased NOS-positive neurons are found in the lesioned (left) side surrounding the lesion area (asterisks). (B, C) In situ hybridization staining showing expression of neuronal NOS in dorsal root ganglia (DRG): B is from experimental rat that underwent peripheral nerve transection; C is from a normal rat. Significant up-regulation of NOS mRNA is observed in DRG neurons after peripheral nerve axotomy. Bar = 100 ~m.

in the nodose ganglion (H6kfelt et al., 1994; Jia et al., 1994; Zhang et al., 1996), the trigeminal ganglion (H6kfelt et al., 1994; Zhang et al., 1996), and the autonomic nervous system (Vizzard et al., 1993, 1995b; Vizzard, 1997). Accumulation of NOS is found both 320

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caudal and cranial to the lesion following a crush of the hypogastric nerve (Elfvin et al., 1997). As reviewed above, neuronal injury causes up-regulation of NOS expression in many regions of the nervous system. However, it is interesting to note that injury to some normal NOS-expressing neurons causes down-regulation of NOS. Axotomy of the splanchnic nerve or removal of peripheral target (adrenal medullectomy) in rat results in quantifiable loss of NOS-positive preganglionic neurons on the lesioned side (Blottner and Baumgarten, 1992). Thus, regulation of injury-induced NOS seems to be system-specific. 2.2. DE NOVO EXPRESSION OF NOS IN INJURED NEURONS 2.2.1. De novo expression of NOS in the cerebral cortex

In normal adult rats, NOS-positive neurons are observed to be scattered throughout the neocortex in layers II, III, IV, and VI. As noted earlier, neocortical pyramidal neurons in layer V do not express NOS (Vincent and Kimura, 1992; Blottner et al., 1995). However, de novo expression of NOS in these neurons is observed following traumatic injuries of the cortex as shown by NADPH-d staining (Kitchener et al., 1993). NOS reactivity in neocortical pyramidal neurons is transiently expressed (Kitchener et al., 1993), and is specific to lesioned neurons (Fig. 4A). 2.2.2. De novo expression of NOS in the cerebellar cortex

The cerebellar cortex is the major source of neuronal NOS in adult rats. The strongest NOS staining is detected in the granule and basket cells of the granular and deep molecular layers, respectively (Vincent and Kimura, 1992). Purkinje cells do not normally exhibit NOS or NADPH-d activity (Bredt and Snyder, 1989; Vincent and Kimura, 1992). In adult rats, expression of NOS in Purkinje cells can be induced by either traumatic or chemical injuries. Following a knife cut in adult rat cerebellum, NOS-positive Purkinje cells are found in the lesioned area (Chen and Aston-Jones, 1994; Saxon and Beitz, 1994). Following traumatic injury, induction of NOS in Purkinje cells seems to be specifically related to axonal injury. NOS-positive Purkinje cells are found only in a limited area in the lesion site (Fig. 4B,C). De novo expression of NOS in Purkinje cells is also observed in adult animals following subcutaneous injection of methylmercury chloride (Himi et al., 1996), or after excitotoxic neuronal injury caused by ibogaine administration (O'Hearn et al., 1995). It is important to note that the onset of NOS expression in these Purkinje cells is delayed for a few days after injury. The late induction of NOS may indicate that NO is unlikely to participate in the initial phase of Purkinje cell damage, but is more likely involved in delayed cell death or other neuronal recovery processes (O'Hearn et al., 1995). The late expression of NOS in injured neurons seems to be a common phenomenon in many in vivo neuronal injury models (Section 2.3). 2.2.3. De novo expression of NOS in the nuclei of cranial nerves

NOS is highly and transiently expressed in neurons of the developing olfactory epithelium during migration and establishment of primary synapses in the olfactory bulb. Although the granule cell layer of the olfactory bulb in the adult animals is one of the areas in the nervous system where the strongest NOS expression can be detected (Vincent and Kimura, 321

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Fig. 4. (A) Adult rat neocortical pyramidal neurons express NOS as shown by NADPH-d staining following traumatic injury in the cortex by a needle. Asterisks indicate the needle trace. Only these neocortical pyramidal neurons whose axons are damaged by the needle express NOS. tB. C) Sections of cerebellar cortex from rats following traumatic injury. The injury was made by punching a micropipette into the cerebellar cortex which creates only minimal damage. A single punch of the micropipette results in damage on one Purkinje cell that expresses NOS (arrow in B). Punching the micropipette into two different points results in damage to two Purkinje cells, which also express NOS (arrows in C). Expression of NOS in this model seems to be specifically related to axonal damage. Bar = 50 I~m.

1992), expression of N O S in olfactory receptor neurons rapidly declines after birth and is undetectable by adulthood (Roskams et al., 1994). Following bulbectomy in the adult rat, expression of neuronal N O S (both protein and m R N A ) are rapidly induced in regenerating olfactory receptor neurons, being particularly enriched in outgrowing axons (Roskams et al., 1994). Motor nuclei of cranial nerves do not normally express N O S in the adult animals. De novo expression of N O S is observed in motoneurons of the facial nucleus (Yu, 1994; Ruan et al., 1995) and hypoglossal nucleus (Yu, 1997) after peripheral nerve axotomy. Ligation of cranial nerve also induces de novo expression of N O S (Fig. 5A,B).

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Fig. 5. (A, B) Cross-sections through the nucleus of facial nerve from a normal control rat (A), or an experimental rat 3 weeks following facial nerve ligation (B). Sections were stained with NADPH-d histochemistry and counterstained with neutral red. Motoneurons in the facial nucleus normally do not express NOS (red staining cells in A). Many motoneurons in the nucleus express NOS following facial nerve ligation (dark blue cells in B). (C, E) Cross-section of the lesioned side of the spinal cord from a rat 2 weeks following root avulsion. The section was first stained by fluorescent immunocytochemistry with the primary antibody against neuronal NOS (C). The same section was then stained with NADPH-d histochemistry (E). Both NOS immunocytochemistry and NADPH-d histochemistry stains the same population of neurons. The staining intensity of NADPH-d is consistent with that of NOS immunocytochemistry in each individual neuron (arrows in C and E indicate the heavily stained neurons and the arrowheads indicate the lightly stained one). (D, F) Cross-section of a lesioned spinal cord from a rat 2 weeks following root avulsion. The section was first stained with NADPH-d histochemistry (D). The same section was then further stained for neuronal NOS mRNA by in situ hybridization (F). Both NADPH-d and NOS in situ hybridization stain the same population of neurons (arrows in D and F). Bar = 150 ~m (A. B), bar -- 50 Ism (C-F). 323

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Fig. 6. Cross-sections of spinal cord from a normal control rat (A) and a rat 2 weeks after spinal root avulsion (B, C, D). Sections were immunocytochemically stained with antibodies against neuronal NOS (A, B), inducible NOS (C), or endothelial NOS (D). Normally, neurons surrounding the central canal e×press neuronal NOS (arrowhead in A and B). Spinal motoneurons do not e×press the enzyme under normal conditions. Neuronal NOS (arrows in B), but not inducible NOS (C) or endothelial NOS (D), is de novo induced in motoneurons following root avulsion. nNOS, neuronal NOS; iNOS, inducible NOS: eNOS. endothelial NOS. Bar = 200 l~m.

2.2.4. De novo expression of NOS in spinal motoneurons De novo expression of NOS in spinal motoneurons was first demonstrated by NADPH-d staining in an adult rat avulsion model (Wu, 1993). Further studies have shown that the induced NOS is of the neuronal isoform which is expressed specifically in response to the root avulsion injury (see Fig. 6; Wu et al., 1994c). Following spinal root avulsion, expression of NOS is observed in motoneurons on the lesion side of the spinal cord. Both NOS protein and mRNA are induced in the injured motoneurons (Fig. 5). It is interesting to note that both NADPH-d histochemistry and NOS immunocytochemistry (with the application of a specific antibody against neuronal NOS) stain the same population of neurons (Fig. 5C,E). In addition, NADPH-d histochemistry can also identify injured motoneurons which express neuronal NOS mRNA as shown by in situ hybridization (Fig. 5D,F). Besides axonal injury, expression of NOS in spinal motoneurons is also found after intrathecal administration of dynorphin (Hu et al., 1996).

2.2.5. De novo expression of NOS in nuclei associated with the long descending and ascending pathways Neurons that form the major descending or ascending tracts normally do not express NOS in adult animals. Following lesions of these tracts, such as spinal cord injury, some of the neurons express NOS. For example, neurons of the spinocerebellar tract in Clarke's nucleus are completely NOS-negative under normal conditions. Following a traumatic injury of the 324

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Fig. 7. (A, B) Cross-sections of the L1 spinal cord from rats following hemisection (A) or transection (B) of the cord at the thoracic segment. Sections were stained with NADPH-d histochemistry. NOS-positive neurons are found in Clarke's nucleus on the lesioned side following hemisection of the spinal cord (arrow in A) and on both sides of the nuclei following transection of the cord (arrows in B). (C, D) Cross-sections of the L1 spinal cord from a rat following hemisection of the cord at the thoracic segment. Sections were stained with NADPH-d histochemistry (C) or NOS in situ hybridization (D). Expression of NOS in both enzyme (arrow in C) and mRNA (arrow in D) are detected in Clarke's nucleus. Bar = 100 Ism.

spinal cord, N O S is dramatically induced in very high concentration in the injured Clarke's neurons (Wu, 1993; Wu et al., 1994a,b,c; Yick et al., 1998). The induction of NOS in these neurons is of neuronal isoform. Both e n z y m e and m R N A are detectable (see Fig. 7; Wu et al., 1994a,b,c). D a m a g e in the spinal cord also causes interruption of descending pathways. Neurons in the lateral vestibular nucleus which form the vestibulospinal tract also express neuronal N O S after spinal cord injury (Jin et al., 1997). However, induction of NOS is not found in other ascending and descending pathways, such as the corticospinal tract, following the same injury (Wu, unpubl, data). 2.3. T I M E C O U R S E OF NOS E X P R E S S I O N IN I N J U R E D N E U R O N S

2.3.1. Time course of NOS expression is different in different populations of neurons Knife lesion in the cerebellar cortex causes expression of NOS in Purkinje cells from 3 to 42 days post injury. NOS-positive Purkinje cells first appear 3 days post injury and increase in 325

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number and staining intensity from 7 to 14 days post injury (Chen and Aston-Jones, 1994). The same reaction is seen in neurons of the inferior olive nucleus after cerebellar lesions (Chen and Aston-Jones, 1994). Onset of NOS expression in chemically injured Purkinje cells is also 3 days after ibogaine treatment (O'Hearn et al., 1995). The number of NOS-positive Purkinje cells increases over 7 days and reaches a maximum at 12-15 days. Subsequently, NOS-positive Purkinje cells decrease in number and no positive cells are found 2 months after injury (O'Hearn et al., 1995). In a facial nerve lesion model, NOS appears 2 days following axotomy, increases between 5 and 20 days, and decreases after 30 days (Ruan et al., 1995). Expression of NOS in pyramidal cells of the neocortex is observed 3 to 21 days after a traumatic injury with maximal expression 7 to 14 days post injury (Kitchener et al., 1993). In a spinal root avulsion model, NOS-positive motoneurons begin to appear 7 days following root avulsion. The number of NOS-positive motoneurons increases rapidly between 2 and 4 weeks, and reaches a maximum of 80% by 4 weeks post injury. Thereafter, the number of NOSpositive neurons decreases and disappears by 9 weeks after the avulsion (Wu, 1993). In a spinal cord injury model, NOS-positive neurons are first observed in Clarke's nucleus 3 days following hemisection of the spinal cord. The number of NOS-positive neurons rapidly increases between 5 and 10 days post injury and reaches a maximum of 50% by 10-20 days post injury. Following this the number of NOS-positive neurons in Clarke's nucleus subsequently decreases to about 10% at 40 days post injury and remains unchanged at 60 days (Yick et al., 1998). 2.3.2. Time course of NOS expression can be different in the same population of neurons following different types of injuries

In a study using the hypoglossal nerve injury model, the time course of NOS expression in the hypoglossal nucleus are compared following different types of injuries including axonal crush, transection, transection plus ligation, and avulsion (Yu, 1997). In the crushed animals, the number of NOS-positive neurons reaches its maximum of 24% at 3 days post injury, decreases to about 5% at 28 days post injury, and finally falls to 0% at 70 days after the crush. In both transected and ligated/transected animals, the number of NOS-positive neurons reaches its maximum of 60% at 14 days post injury. Thereafter, it drops precipitously in the transected group to about 15% at 42 days post injury and remains unchanged 70 days after the injury. In the ligated/transected group, the number of NOS-positive neurons remains at maximal levels up to 42 days post injury, and then decreases to 25% at 70 days after the injury. In avulsed animals, the number of NOS-positive neurons rises rapidly from 46% at 3 days to 75% at 7 days, and reaches its maximum of 92% at 14 days post injury. The number of NOS-positive neurons is then reduced to about 46% at 42 days and remains unchanged at 70% post injury (Yu, 1997). These results seem to indicate that the time course of NOS expression in response to axonal injury is related to the nature of the injury. In a facial nerve injury model, however, both nerve crush and transection cause the same time course of NOS expression in the facial nucleus (Yu, 1997). Together, the above data indicate that both the type of neurons and the nature of the injury can affect the time course of injury-induced NOS expression.

3. CO-EXPRESSION OF NOS WITH OTHER INJURY-RELEVANT COMPONENTS

In the normal nervous system, NOS is co-localized with many other neurotransmitters and neuropeptides, including choline acetyltransferase (CHAT), vasoactive intestinal peptide 326

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(VIP), substance P (SP), somatostatin (SOM), calcitonin gene-related peptide (CGRP), 5-hydroxytryptamine (5-HT), galanin, etc. (Elfvin et al., 1997; Xu and H6kfelt, 1997). Following neuronal injury, expression of NOS is found to coincide with other injury-relevant gene expression. Such co-expression of NOS with other injury-relevant gene varies in different population of neurons. For instance, neuronal NOS is found to be co-expressed with c-jun, an immediate early gene (IEG), in the same population of neurons in the lateral vestibular nucleus following spinal cord injury (Jin et al., 1998). In contrast, following axonal injury of spinal motoneurons, neuronal NOS is expressed in different populations of motoneurons from those expressing c-jun (see Fig. 8; Wu et al., 1994b). Besides the IEG, NOS has been reported to be co-expressed with many other injury-relevant genes in injured neurons. In cortical pyramidal neurons, expression of neuronal NOS is confined to a subset of neurons which also express high levels of beta-amyloid precursor protein (APP) after cerebral cortex lesion (Luth and Arendt, 1997). Sciatic nerve transection induces an up-regulation of calcitonin gene-related peptide (CGRP), low-affinity nerve growth factor receptor (p75), and growth-associated protein 43 (GAP-43), but not NOS in rat spinal motoneurons (Piehl et al., 1998). In contrast, galanin message-associated peptide (GMAP) becomes up-regulated in motoneurons following root avulsion and is coincident with the up-regulation of NOS, while expression of CGRP and p75 is normal (Piehl et al., 1998). Co-localization of NOS mRNA and GMAP mRNA is also found in hypothalamic magnocellular neurosecretory neurons following hypophysectomy (Villar et al., 1994). Compression/ligation or cut of the hypogastric nerve causes accumulation of NOS, which is often coexistent with VIP, SP, or CGRP (Elfvin et al.. 1997). Long-lasting 327

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co-localization of galanin and NOS is also observed in neurons of the mammillary nucleus or DRG following injuries in the mammillo-thalamic tract or sciatic nerve, respectively (Brecht et al., 1997). In dorsal motor vagal and hypoglossal neurons, co-expression of NOS with an interferon-gamma-like molecule (N-IFN-y) is found, following transection of the vagal or hypoglossal nerve, respectively (Kristensson et al., 1994). Taken together, injury-induced NOS co-expresses with many other genes. The precise mechanisms of such co-expression are still unknown.

4. AGE-RELATED EXPRESSION OF NOS IN INJURED NEURONS

It is well established that immature neurons are more vulnerable to axonal injury than the adult ones. The age of the animals plays an important role on neuronal survival after axonal injury. For example, sciatic nerve cut causes a dramatic motoneuron death in developing animals, but the same injury does not cause any detectable cell death in adults. Recent studies have also shown that there are gene expression changes in injured neurons following axonal injury (Wu et al., 1994c; Wu, 1996), which is influenced by the age of the animals (Li et al., 1993; Y. Wu et al., 1995; Piehl et al., 1998). For instance, sciatic nerve transection induces expression of NOS in spinal motoneurons in the developing rat but not in adult (Li et al., 1993). Studies on the change of age-related gene expression may be helpful in understanding the different responses of neurons to axonal injury in the developing and adult animals. It has been shown that axotomy in adult animals induces a significant up-regulation of NOS in the DRG neurons (Zhang et al., 1993; H6kfelt et al., 1994), while axotomy in earlier postnatal animals does not cause similar changes (Rydh-Rinder et al., 1996). This finding is different from that of spinal motoneurons, since sciatic nerve cut induces expression of NOS in early postnatal but not in adult rats (Clowry, 1993; Li et al., 1993). The fact that expression of NOS coincides with the death of immature motoneurons following peripheral nerve injury (Clowry, 1993; Li et al., 1993) indicates that NO/NOS may play a role in the target-deprived degeneration of motoneurons in early postnatal development. Expression of NOS and its roles in neuronal injury may be varied in different populations of neurons. Transection of the facial nerve in new born or earlier postnatal (within the first postnatal week) rats does not induce NOS expression while extensive motoneuron death is detected in the facial nucleus. In contrast, the same injury causes expression of NOS in the facial motor nucleus in the adult rats while nerve cell loss is less severe (Mariotti et al., 1997). This finding seems to indicate that NOS is not involved in neuronal death in this particular injury model. In contrast, in spinal motoneuron injury models, distal nerve axotomy in developing rats causes induction of NOS which coincides with severe motoneuron death (Li et al., 1993), while the same injury does not cause NOS expression, nor motoneuron death in adult rats (Wu, 1993; Wu et al., 1994c). In addition, spinal root avulsion induces a more rapid and intensive expression of NOS in spinal motoneurons of early postnatal rats than that of adults (Fig. 9). Such rapid and intensive expression of NOS in spinal motoneurons is coincident with the rapid and severe loss of motoneuron in the developing rats (Y. Wu et al., 1995). Taken together, the expression of NOS in axonal injured neurons is influenced to a large degree by the age of the animals and the type of neurons. Hence, injury-induced NOS may play different roles in different ages and different populations of neurons. 328

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Fig. 9. (A) Cross-section of spinal cord from a rat 5 days following root avulsion which was performed at 8 days of age. The section was stained with NADPH-d histochemistry. Motoneurons in the lesioned side are heavily stained with NADPH-d (open arrow). Arrowheads indicate blood vessels which are also NADPH-d-positive. (B, C, D) Horizontal sections of the lesioned side of the spinal cord from rats 10 days following root avulsion performed at 15 days of age (B), 15 days following root avulsion performed at 30 days of age (C), and 15 days following root avulsion in adult (D). Sections were stained with NADPH-d histochemistry. The staining intensity of NADPH-d in younger animals is more intense (B, C) than in adult ones (D). Bar = 100 Ltm (A), bar = 50 Ltm (B, C. D).

5. SPECIES-RELATED EXPRESSION OF NOS IN INJURED NEURONS It has been shown that response of animals to injury, in terms of NOS expression and NO production, may be different from one species to another. Mouse microglia generate large 329

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quantities of NO in response to injury or inflammatory stimulation, but human and hamster microglia do not produce measurable amounts of NO under the same conditions (Colton et al., 1996). Activity of injury-induced NOS and the roles of NOS in ischemic and infarct brain injury following middle cerebral artery (MCA) occlusion are different between the rats and mice (Yoshida et al., 1995). More interestingly, although spinal root avulsion leads to dramatic loss of spinal motoneurons in both rats and mice, the injury causes de novo expression of NOS in injured motoneurons only in rats (Wu, 1993: Wu et al., 1994c), but not in mice (Li et al., 1995). Similar phenomena are observed in other animal species. For example, peripheral nerve axotomy in rats causes up-regulation of NOS in DRG neurons, but the same lesion does not cause an increase of NOS expression in guinea pigs (Rydh-Rinder et al., 1996). In the cerebellar cortex, however, de novo expression of NOS in Purkinje cells is observed in both mice (Himi et al., 1996) and rats (Chert and Aston-Jones, 1994) following neuronal injury. We have investigated the expression of NOS in spinal motoneurons after spinal root avulsion in different animal species from mice to monkeys. Motoneurons in mice do not express NOS following root avulsion. All other species studied express NOS after the avulsion. Smaller animals, for example rats and guinea pigs, exhibit a quicker response to injury in terms of NOS expression than do larger animals, such as cats and monkeys. In general, with the exception of the mouse, more NOS-positive motoneurons and higher staining intensity are found in smaller animals within the same post-injury time period (Wu et al., 1997). Interestingly, morphological characteristics of injury-induced NOS-positive neurons are different in different species after the same avulsion injury. For example, by 2 weeks following root avulsion, NOS-positive motoneurons in rats exhibit characteristics which are similar to apoptotic cell death (Figs. 10B and 13), while most motoneurons of guinea pigs show degenerative characteristics of necrosis (Fig. 10C). Studies of NOS expression patterns and different morphological changes in different animal species may help to understand the roles of NOS in neuronal injury. These species differences are also important to consider when modeling human disease processes from rodent studies.

6. ULTRASTRUCTURE OF INJURY-INDUCED NOS-POSITIVE NEURONS Distribution of NOS in the subcellular structures has been studied using both NADPH-d histochemistry and immunocytochemistry. Under normal conditions, NOS-positive neurons show a large amount of the electron-dense reaction product, formazan, which is not confined

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Fig. 10. (A) Cross-section of the ventral horn of the spinal cord from a normal rat. (B, C) Cross-sections of the ventral horn of the spinal cord from experimental rat (B) or guinea pig (C) 2 weeks following root avulsion. Sections were stained with NADPH-d histochemistry and counterstained with neutral red. Normal spinal motoneurons do not express NOS and are stained by neutral red into red in color (A). By 2 weeks alter the injury, the perikarya of NOS-positive motoneurons is shrunken and the normal multipolar cell body transforms and becomes round as compared to the normal motoneurons (arrows in B). Compared with the normal motoneurons in A, the cell body of these NOS-positive motoneurons is smaller (B). These changes more likely resemble apoptosis. In contrast, injured motoneurons in the guinea pig show the different morphological changes. Many of these NOS-positive motoneurons show characteristics of necrosis which include the swelling of the cell body (open arrows in C), breakdown of the cell membranes (many cells in C without a clear nuclear membrane, asterisk in C indicates a motoneuron with disrupted membrane). Still, some motoneurons in guinea pig is shrunken and the cell body of these cells becomes round and smaller compared with the normal motoneurons (arrows in C). Bar = 20 gm.

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to any particular subcellular organelle. The formazan, or NOS immunoreactivity, is scattered throughout the whole cytoplasm. Although there is more labeling associated with smooth and rough endoplasmic reticulum and near the Golgi apparatus, it is absent from the mitochondria and cell nuclei (Mizukawa et al.. 1988; Llewellyn-Smith et al., 1992). However, other investigators have reported that NOS reactivity is found in the nuclear envelope, inner and outer mitochondria membranes, Golgi apparatus, and endoplasmic reticulum (Calka et al., 1994). Different distribution of NOS in the subcellular organelles reported in various studies may be due to different staining methods used. With either NADPH-d or NOS immunocytochemistry, we have found that NOS reactivity is not specifically associated with any subcellular organelle or plasma membrane if Triton X-100 is added to the reactive solution. Without Triton X-100 in the reactive solution, NOS reactivity can be observed in some membrane structures, such as endoplasmic reticulum (Wu, unpubl, data). The subcellular distribution of injury-induced NOS in injured neurons is similar to that of normal NOS-containing neurons. Injury-induced NOS is found throughout the cytoplasm but not the nucleus, and is not specifically associated with any subcellular organelles (Fig. 11A,B). Morphologically, these injury-induced NOS-positive neurons seem normal in some areas of the nervous system (Fig. 11A), but abnormal in others (Fig. 11B). Neuronal death following injury is often assumed under two major forms, i.e. necrosis or apoptosis (Ellis et al., 1991; Oppenheim, 1991 ; Cohen and Duke, 1992). Whether expression of NOS plays any role in either necrosis or apoptosis is not clear and may vary in different neuronal populations, developmental ages, and animal species. The expression of NOS in the rat spinal motoneurons may be more likely associated with necrosis, especially during the early stage of the injury when NOS concentration is in a higher level. This is based on the following observations. (1) Injury of the peripheral nerve in the developing animals causes apoptosis in spinal motoneurons without NOS expression (Li et al., 1998). (2) Expression of NOS in adult spinal motoneurons following root avulsion is associated with a necrotic type of cell death as characterized by the disintegration of membrane (Fig. l lC) and the activation of macrophages in the ventral horn of the spinal cord (Fig. 12). However, injury-induced NOS-positive motoneurons have mixed morphology, showing characteristics of both apoptosis and necrosis. As shown in the earlier section of this review, different animal species may show different characteristics of degeneration, either apoptosis or necrosis, following a same injury. Even in the same animal species, morphological characteristics of NOS-positive neurons could be different at different post-injury stages. For instant, injured rat motoneurons show necrotic damage during the early stage of root avulsion lesion (as shown earlier in this section). However, at the late stage after injury, these motoneurons exhibit features of apoptosis, including increased electron density in both the cytoplasm and nucleus with intact cell and nuclear membranes, the shrunken nucleus, and the reduced soma size of the cell (Fig. 13A). Occasionally, apoptotic bodies are found in these cells (Fig. 13B). In the cerebellar cortex, the size of injury-induced NOS-positive Purkinje cells appears to decrease gradually after lesion. The average diameter of reactive Purkinje cells 6 weeks post injury is about 38% smaller than that observed 3 days post injury (Chen and Aston-Jones, 1994). These changes seem to resemble apoptosis. In contrast, subcellular structures of injury-induced NOS-positive neurons in the hypothalamus following hypophysectomy seem normal, morphologically and functionally (Wu and Scott, 1993; Scott et al., 1995). No evidence of necrosis or apoptosis is observed in those hypothalamic neurons (Scott et al., 1995). These data indicate that the role of NOS in injured neurons is likely complex and expression of injury-induced NOS in different populations of neurons may play different roles in neuronal injury. 332

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Fig. 12. Cross-sections of the spinal cord from a rat 2 weeks following root avulsion. The sections were

immunocytochemically stained with antibody against OX-42 (A. C)or ED-I (B. D). Activated macrophages labeled by ED-1 are found in both dorsal and ventral horns of the lesioned side of the spinal cord lB. D). Bar = 500 I~m (A, B), bar = 100 I~m (C, D).

7. R E G U L A T I O N O F N O S E X P R E S S I O N IN I N J U R E D N E U R O N S As reviewed above, up-regulation or de novo expression of NOS is observed in some populations of neurons after neuronal injury. Expression of NOS in injured neurons is influenced by the age of animals, type of injury, and location of neurons. In addition, the length of the remaining proximal axons after axotomy and other experimental manipulations also have great influence on the regulation of injury-induced NOS expression. 7.1. REGULATION OF INJURY-INDUCED NOS BY THE LENGTH OF THE R E M A I N I N G PROXIMAL A X O N S FOLLOWING A X O T O M Y The length of the proximal axons plays a crucial role on injury-induced NOS expression in spinal motoneurons of adult rats (Wu, 1993; Gu et al., 1997). Spinal root avulsion causes de novo expression of NOS in injured motoneurons, but distal axotomy of the spinal nerve, 10 mm from the cord, does not induce NOS expression (Wu, 1993). It is important to note that expression of NOS coincides with the death of injured motoneurons in avulsion injury. No motoneuron loss is observed when axotomy is performed 10 mm from the spinal cord. When the axotomy is made closer to the spinal cord, motoneurons begin to express NOS and, at the same time, motoneuron loss is observed (Gu et al.,

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Fig. 13.

Electron photomicrographs of spinal motoneurons from a rat during the later stage (about 2 weeks) following root avulsion. (A) Shrunken motoneuron (MN) surrounded by several glial cells (G). (B) The nucleus of an injured motoneuron (MN) is shrunken, but the cell and nuclear membranes are intact (arrows). An apoptotic body is found near the nuclear membrane (arrowheads). Bar - 2 ll.m.

.

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Cross-sections of ventral horns from rats which underwent peripheral nerve axotomy at 8, 6, 4, 2, or 0 mm from the spinal cord. Sections were stained with NADPH-d histochemistry and counterstained with neutral red. NOS-positive motoneurons (arrows in 2 mm and 0 mm sections) are observed when axotomy is performed at the peripheral nerve less than 4 mm from the spinal cord. 335

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1997). The minimal length of proximal axon needed to prevent expression of NOS and motoneuron loss in adult rats following axotomy is about 4 mm (see Fig. 14; Gu et al., 1997). On the other hand, some populations of neurons in the central nervous system express NOS following axonal injury regardless of the length of the proximal axon. For example, neurons in Clarke's nucleus express NOS following either proximal or distal (Yick et al., 1998). In the hypothalamo-pituitary pathway, axonal transection, i.e. pituitary stalk section, results in a significant increase of NOS expression in the supraoptic and paraventricular nuclei but a strong decrease in periventricular nucleus (Lumme et al., 1997). These results illustrate that groups of neurons that are located in the same brain area can respond very differently to neuronal injury. 7.2. R E G U L A T I O N OF INJURY-INDUCED NOS BY P E R I P H E R A L NERVE (PN) GRAFT TRANSPLANTATION The fact that the length of remaining axons play a crucial role on NOS expression in spinal motoneurons, but not in Clarke's neurons after axotomy suggests that certain molecules in the remaining axons of motoneurons may inhibit the expression of NOS. Axons of spinal motoneurons are myelinated by Schwann cells and considered as a peripheral type of nerve, while axons of Clarke's neurons are myelinated by oligodendrocytes and considered as a central type of nerve. It is logical to believe that a NOS-inhibiting molecule is produced by the peripheral type of nerve. Results from recent studies seem to support this hypothesis. Transplantation of a PN graft after root avulsion significantly enhances motoneuron survival and reduces NOS expression in injured motoneurons (Wu et al., 1994a; Wu, 1996). More importantly, expression of NOS is completely inhibited in all injured motoneurons that have re-grown into the transplanted PN graft, while all non-regenerating ones express NOS (Fig. 15). It is also interesting to note that the regenerating motoneurons express p75 and c-jun, while non-regenerating ones do not (see Fig. 15; Wu, 1996). These results indicate that the expression of NOS in spinal motoneurons is related to neuronal death, whilst the expression of p75 and c-jun is related to neuronal survival and regeneration. Transplantation of PN graft in the CNS, however, has a different effect on injury-induced NOS expression. Following axotomy of Clarke's neurons, PN graft transplantation increases the number of injury-induced NOS-positive neurons on the lesioned side (Yick et al., 1999a). More interestingly, up-regulation of NOS expression coincides with an increase in survival of injured Clarke's neurons (Yick et al., 1999a). In a lesion model of the lateral vestibular

Fig. 15. Horizontal sections through the ventral horn of the spinal cord from a rat after implantation of a peripheral nerve (PN) graft. Motoneurons were pre-labeled with Fast Blue (FB) 2 days before C7 root avulsion. A PN graft was then implanted into the C7 segment immediately follov,ing the lesion." Three weeks after implantation, diamidino yellow (DY) was applied into the PN graft. Two days following the injection of DY, the animal was perfused and prepared for fluorescent microscopy. Following fluorescent microscopy, sections were immunocytochemically stained with either p75 (A. B. C) or c-jun (D, E. F). Both sections were then further stained with NADPH-d histochemistry. Motoneurons that expressed NOS (NADPH-d-positive. small arrows in C and F) represent those that did not regenerate into the PN graft after the lesion (labeled by FB but not by DY, small arrows in A, B and D, E). Motoneurons that regenerated into the PN graft following the lesion (labeled by both FB and DY, large arrows in A, B and D, E) expressed p75 (large arrows in C) and c-jun (large arrows in F) but not NOS (large arrows in C and F). Bar = 100 jim.

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nucleus, PN graft transplantation following axotomy is also found to increase injury-induced NOS expression (Jin et al., 1998). Moreover, co-expression of NOS and c-jun is observed in regenerating vestibular neurons on the lesioned side following PN graft implantation (Jin et al., 1998).

8-*

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These results demonstrate that the same experimental manipulation has different effects on various types of neurons. Expression of injury-induced NOS may also play a different roles from one type of neurons to another. 7.3. REGULATION OF INJURY-INDUCED NOS BY NEUROTROPHIC FACTORS Regulation of NOS expression by neurotrophic factors has been reported in an in vitro study (Samdani et al., 1997b) and in in vivo studies in both the developing (Huber et al., 1995) and adult animals (Holtzman et al., 1994: Novikov et al., 1995, W. Wu et al., 1995). Whether neurotrophic factors up-regulate or down-regulate NOS expression is dependent upon the type of injury made, source of trophic factor applied, and the population of neurons studied. When NOS expression in adult spinal motoneurons was first reported a few years ago (Wu, 1993), we speculated that expression of injury-induced NOS may be due to a lack of neurotrophic factors. Recent studies have provided evidence to support the hypothesis that expression of injury-induced NOS is regulated by certain neurotrophic factors. Among the known trophic factors, BDNF, GDNF and IGF-I have been reported to down-regulate NOS expression following neuronal injury. An in vitro study has demonstrated that BNDF protects cultured cortical neurons from NMDA receptor-mediated glutamate neurotoxicity by reducing cytotoxic action of NO (Kume et al., 1997). Expression of injury-induced NOS in spinal motoneurons is completely inhibited by continuous infusion of BDNF or GDNF (Fig. 16), which coincides with the increase of motoneuron survival after spinal root avulsion (Novikov et al., 1995, 1997; W. Wu et al., 1995). Inhibition of injury-induced NOS is trophic-factor-dependent, and only certain trophic factors have such effect. For instance, infusion of CNTF or IGF-I following root avulsion does not inhibit expression of injury-induced NOS, nor increase the survival of injured motoneurons (W. Wu et al., 1995). If the right source of trophic factor is used, expression of injury-induced NOS can be inhibited long-term by just a single application of that factor. For example, a single application of BDNF (20 gg/rat) at the time of avulsion inhibits NOS expression in injured motoneurons up to 6 weeks post avulsion (Chai et al., 1998). The effect of neurotrophic factors on the expression of injury-induced NOS is also neuron-specific, i.e. a given trophic factor may have a regulating effect on injury-induced NOS on one population of neurons but not on other populations. For example, IGF-1 has been reported to show neuroprotective effects by down-regulating NOS expression in spinal interneurons after traumatic injury of the spinal cord (Sharma et al., 1997), while IGF-I does not show any effect on regulating NOS expression in spinal motoneurons following avulsion (W. Wu et al., 1995). Furthermore, injury-induced NOS in axotomized Clarke's neurons increases concomitantly with the survival of injured neurons after continuous treatment of BDNF, NT-3, or CNTF (Yick et al., 1999a). Taken together, these results indicate that injury-induced NOS can be regulated by neurotrophic factors.

8. MECHANISMS OF NOS EXPRESSION IN INJURED NEURONS

As reviewed above there is no doubt that NOS can be up-regulated or induced in some populations of neurons following injury. However, with regard to how and why NOS is induced in neurons after injury, little is known. Mechanisms underlying the expression of injury-induced NOS are not understood and may be different from one population of neurons 338

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Fig. 16. Cross-sections of the spinal cord from rats 2 weeks following root avuision. The sections were immunocytochemically stained with antibody against neuronal NOS (,4, D, G), c-jtm (B. E, H), or p75 (C, F, /). Sections A, B, and C were from a control rat that only underwent root avulsion. Sections D, E, and F were from an experimental rat that received BDNF treatment after the root avulsion. Sections G. H, and 1 were from an experimental rat that received GDNF treatment after the root avulsion. In the control animal, NOS is induced in spinal motoneurons following root avulsion (A), while c-jun and LNGFR are only weakly labeled (B and C, respectively). BDNF or GDNF inhibits completely the expression of NOS due to root avulsion (D and G, respectively) and, at the same time, significantly up-regulates the expression of c-jun and LNGFR (E, H, F and I, respectively). Bar = 200 p.m.

to another. Several hypotheses have been proposed to interpret the induction of injury-induced NOS in neurons. 8.1. EXPRESSION OF INJURY-INDUCED NOS IN NEURONS IS A GENERAL RESPONSE TO INJURY When up-regulation of NOS as shown by NADPH-d histochemistry in the dorsal motor nucleus of the vagus nerve after axotomy was first reported a decade ago, it was thought 339

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that the increased NADPH-d staining might result from the accumulation of the enzyme in the cell body due to a decrease of transport to the terminals after axonal injury (Gonzalez et al., 1987). It was also speculated that increased NADPH-d staining might be involved in the regenerative response following axonal injury since axotomy increased the metabolic rate of glucose in injured neurons (Gonzalez et al., 1987). More than ten years since then, it is now clear that these neurons stained by NADPH-d are NOS-containing neurons, and their functions in the normal nervous system have been extensively studied. Distributions and functions of NO/NOS have been reported to involve in many physiological processes in the nervous system. However, mechanisms of injury-induced NOS expression and the role of this expression is not fully understood. Therefore, it is still possible that increased expression of NOS in injured neurons is just a general response of neurons to injury and may simply represent an increased metabolic rate. 8.2. EXPRESSION OF INJURY-INDUCED NOS RESULTS FROM THE INTERRUPTION OF AXONAL TRANSPORT Axonal injury not only damages the integrity of neurons structurally, but also interrupts functional axonal transport of the injured neuron. It is not clear whether injury-induced NOS expression results from damage of the structural integrity of the neuron or from the interruption of axonal transport. A recent study in the neurosecretory hypothalamo-pituitary system seems to support the idea that up-regulation of NOS in injured neurons is a result of interruption of axonal transport (Lumme et al., 1997). Lumme's study has shown that intracerebroventricular injection of colchicine, a compound that stops axonal transport, increases the number of NOS-positive neurons in the paraventricular nucleus. This experiment may provide evidence that up-regulation of the expression of NOS results from the interruption of axonal transport. A similar result is observed in postganglionic sympathetic neurons of the superior cervical ganglion after intraganglionic colchicine injection (Vanhatalo et al., 1998). It is believed that up-regulation of NOS in these colchicine-treated neurons is simply due to the accumulation of NOS in the cell body. Neurons in the paraventricular nucleus and the superior cervical ganglion normally express NOS. Accumulation of NOS in the cell body seems to be a logical outcome following interruption of axonal transport. On the other hand, it is interesting to know whether treatment of colchicine could induce NOS expression in neurons that normally do not express NOS. Although there are no data available to answer this question, it is unlikely that there is a de novo induction of NOS by treatment with colchicine. 8.3. EXPRESSION OF INJURY-INDUCED NOS RESULTS FROM STIMULATION OF NMDA RECEPTORS Activation of glutamate receptors is thought to be a major mechanism for the generation of NO (Szabo, 1996). Recent in vitro studies have shown that stimulation of NMDA receptors causes the release of NO (Garthwaite et al., 1989; Snyder and Bredt, 1992). It is proposed that stimulation of NMDA receptors opens channels for calcium to enter the cell and bind to calmodulin, thereby activating NOS (Snyder, 1991). In primary cortical neuronal cultures, when NMDA is added to cultures briefly, 80% of the neurons die within 24 h. In this system, nitroarginine, an inhibitor of NOS prevents the neurotoxicity elicited by NMDA and related excitatory amino acids (Dawson et al., 1991). Depletion of arginine (the precursor of NO) in the culture medium also prevents NMDA toxicity (Dawson et al., 1991). These findings 340

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suggest that NOS is activated by NMDA receptors and is involved in the neurotoxicity of excitatory amino acids in vitro. Study in an in vivo model, however, does not support the idea that injury-induced expression of NOS results from the stimulation of NMDA receptors. Blockade of NMDA receptors with MK801 does not prevent the induction of injury-induced NOS in spinal motoneurons following root avulsion, nor does it prevent the death of the injured motoneurons (Wu, unpubl, data). This indicates that other mechanisms must be involved in the injury-induced expression of NOS. 8.4. EXPRESSION OF INJURY-INDUCED NOS RESULTS FROM DEPRIVATION OF NEUROTROPHIC FACTORS As discussed in the previous section of this review, expression of injury-induced NOS can be regulated by neurotrophic factors. Mechanisms by which neurotrophic factors regulate the induction of injury-induced NOS are not clear. In adult rat spinal motoneurons, however, it has been shown that induction of injury-induced NOS results from the deprivation of certain neurotrophic factors. It has been hypothesized that neurons synthesize certain endogenous toxic molecules when deprived of neurotrophic factors and such endogenous toxic molecules are responsible for the death of neurons (Oppenheim et al., 1988; Johnson et al., 1989). We have previously hypothesized that the expression of NOS in avulsed spinal motoneurons results from deprivation of trophic factors, normally produced by the PN component (Wu, 1993). This hypothesis is supported by the following observations. (1) Expression of NOS is inhibited in all motoneurons that regenerate into the PN graft (Wu et al., 1994a; Wu, 1996). (2) All regenerated motoneurons re-expressed p75 (Wu, 1996). Since up-regulation of p75 has been suggested to reflect an increase in retrograde transport of neurotrophic factors (Lindsay et al., 1994), re-expression of this receptor after PN graft implantation may indicate that avulsed motoneurons have received atrophic factor from the PN graft. This factor is needed for the survival of the cells. (3) Local infusion of trophic factors, such as BDNF or GDNF, following root avulsion completely inhibits the expression of injury-induced NOS in injured motoneurons (W. Wu et al., 1995; Novikov et al., 1995). Therefore, NOS may serve as an endogenous toxic molecule and be responsible for the death of injured motoneurons.

9. POTENTIAL ROLES OF NOS EXPRESSION IN NEURONAL DEGENERATION AND REGENERATION As mentioned in the introduction section, NO/NOS play roles in a wide range of physiological functions in the nervous system under normal conditions. However, the role of injury-induced NO/NOS in neuronal injury is not fully understood. Up-regulation of NOS in injured neurons has been reported to be involved in both neuronal degenerative and regenerative processes, depending on the population of neurons studied and the type of injury performed. In general, it seems that injury-induced NOS plays a negative role in neurons that do not normally express the enzyme. In these cases, injury-induced NOS seems to be neurotoxic and may be involved in neural degenerative processes. In contrast, NOS may not cause degeneration in neurons that normally express the enzyme. The potential mechanisms of different responses to injury-induced NOS between these two populations of neurons will be discussed later.

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9.1. POTENTIAL ROLE OF NOS IN NEURONAL DEGENERATION Since many different injury models and neuron populations have been used to study the roles of NOS in neuronal injury, there are contradictory findings regarding this issue. Some studies have shown that NOS may not play any role in neuronal injury (Cohen et al., 1996; Yezierski et al., 1996; Van Muiswinkel et al., 1998) or may have beneficial effects for the injured neurons (Uemura et al., 1990; Weissman et al., 1992), while others have demonstrated that expression of NOS is involved in neuronal degeneration.

9.1.1. Involvement of NOS in neural degeneration Although NO/NOS is thought to contribute to the pathological pathway of injury in a wide variety of acute and chronic neurological disorders, including focal ischemia, trauma, epilepsy, Huntington disease, Alzheimer disease, amyotrophic lateral sclerosis, AIDS dementia, and other neurodegenerative diseases (Bonfoco et al., 1995), most recent studies on the response of NOS to injury have been focused on traumatic, ischemic, and chemical injuries. Traumatic injur3". Following traumatic injury of the nervous system, involvement of NOS in neural degenerative processes has been demonstrated in several injury models. Either a nonspecific inhibitor of NOS, N~-nitro-L-arginine methyl ester (L-NAME), or a specific neuronal NOS inhibitor, 7-nitroindazole (7-NI), have shown to have neuronal protective effects after traumatic brain injury (Mesenge et al., 1996). It has been demonstrated that inhibition of NOS increases neuronal survival of spinal motoneurons (Wu and Li, 1993), motoneurons of the facial nerve (Ruan et al., 1995), and neurons in Clarke's nucleus (Yick et al., 1999b) following root avulsion, facial nerve axotomy, or hemisection of the spinal cord, respectively. It is important to note that neurons in these nuclei do not normally express NOS and the lesion causes de novo expression of NOS. These studies suggest that injury-induced NOS may be associated with neural degeneration. In other traumatic injury models, topical application of neuronal NOS antiserum has a neuro-protective effect after acute traumatic spinal cord injury and also down-regulates the expression of NOS in the lesioned areas (Sharma et al., 1996). These observations support the idea that NOS is involved in neuronal degeneration. In addition to studies using NOS inhibitors, evidence supporting the involvement of NOS in neuronal injury has also been obtained from other studies. It has been shown that insulin-like growth factor-1 (IGF-1) protects against degenerative change by attenuating edema, and down-regulating neuronal NOS expression in the spinal cord after traumatic injury (Sharma et al., 1997). Local infusion of trophic factors, such as BDNF or GDNF, following root avulsion not only inhibits the expression of injury-induced NOS but also prevents death of injured motoneurons (W. Wu et al., 1995; Novikov et al., 1995). Use of methylprednisolone (MP) after severe spinal cord compression can reduce the extracellular level of arginine (the substrate for NO production) (Farooque et al., 1996). This change may counteract free radical formation and may be an important mechanism by which MP exerts its beneficial actions. On the other hand, when substrate levels for NOS become too low, NOS activity leads to increased production of free radicals (Mayer et al., 1991; Klatt et al., 1993) Additionally, inhibition of NOS has been shown to protect against acute traumatic neuronal injury in hippocampal slices in vitro (Wallis et al., 1996). Taken together, injury-induced NOS is likely to be neurotoxic and may be involved in the degenerative process after traumatic injury. Hence, inhibition of NOS can protect against such injury. Ischemic injuo'. Ischemic injury is another model commonly used to study the possible role of NO/NOS in neuronal injury. Recent studies have shown that nonspecific inhibition of NOS 342

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protects neurons from ischemic damage in the retina (Roth, 1997), in the cortex and striatum (Hamada et al., 1994), and in the hippocampus (Wallis et al., 1992; Nagafuji et al., 1993" Nakagomi et al., 1997). This suggests an important role of NO/NOS in the pathogenesis of ischemic injury. Interestingly, among the three isoforms of NOS, the neuronal NOS seems to be involved in neuronal injury since 7-NI (the specific inhibitor of neuronal NOS) significantly inhibits apoptosis in hippocampus and neocortex after prolonged hypothermic circulatory arrest (Tseng et al., 1997). Moreover, neonatal mice lacking neuronal NOS are less vulnerable to hypoxic injury, which also supports the role of NOS in ischemic injury (Ferriero et al., 1996). The neuroprotective agent lubeluzole, which is efficacious in both clinical and experimental models of cerebral ischemia, is found to protect cultured hippocampal neurons in a dose-dependent manner against the toxic effects of NO (Maiese et al., 1997), which further supports the idea that NO/NOS is involved in ischemic neuronal injury. Chemical injury. Recent studies have suggested that NO may be a key mediator of excitotoxic neuronal injury in the CNS. For example, NO directly mediates glutamate neurotoxicity in primary cortical cultures (Dawson et al., 1991). In vivo studies also show that 7-NI significantly attenuated lesions produced by intrastriatal injection of NMDA, which implicated that neuronal NOS might be involved in the pathogenesis of excitotoxic neuronal injury (Schulz et al., 1995). Another neuronal NOS inhibitor, S-methylthiocitrulline, also shows neuroprotective effects against excitotoxic neuronal injury following intrastriatal administration of quinolinic acid (Bazzett et al., 1997) or methylphenyl tetrahydropyridine (MPTP) and malonate (Matthews et al., 1997). In addition to studies using NOS inhibitors, evidence for the involvement of NOS in excitotoxic injury has also been obtained from other studies. Gangliosides, the polar sugar-containing lipids, are neuroprotective in animal models of neurotoxicity. It is reported that the neuroprotective effects of gangliosides may arise from a blockade of nitric oxide formation since gangliosides prevent glutamate neurotoxicity in cortical cultures which closely parallel their potencies in binding calmodulin and inhibiting NOS (Dawson et al., 1995). Among the subunit isoforms of glutamate receptors, the NMDA R1 receptor is most likely to contribute to NO-mediated excitotoxic injury (Weiss et al., 1998). In contrast, metabotropic glutamate receptor (mGluR) may prevent NO-mediated cell death since activation of mGluR subtypes with 1S, 3R-ACPD and L-AP4, agents which are neuroprotective against NO, significantly limit the progression of programmed cell death (PCD) (Vincent et al., 1997).

9.1.2. Potential mechanisms of NO/NOS-mediated neurotoxicity The potential mechanisms of NO production in neuronal injury include direct cytotoxicity, peroxynitrite formation by NO and superoxide anion, and NO-mediated elevation of cellular cGMP that enhance tumor necrosis factor-alpha toxicity. NO is a free radical that can directly cause DNA damage. NO appears to elicit neurotoxicity by activating poly-adenosine 5'-diphosphoribose synthetase (PARS). Activation of PARS can lead to cell death through depletion of the energy stores, [3-nicotinamide adenine dinucleotide (NAD) and adenosine triphosphate (ATP) (Snyder et al., 1998). Inhibition of PARS with benzamide can block NMDA- and NO-mediated neurotoxicity (J. Zhang et al., 1994). NO-induced injury can be prevented in CA1 neurons from a hippocampal slice by inhibition of poly-ADP-ribosylation, suggesting that poly-ADP-ribosylation may play an important role in NO-mediated neuronal injury (Wallis et al., 1993). NO may react with superoxide anion (O;-) to form peroxynitrite (ONOO-). Since NO is a free radical gas with a very short lifetime after being synthesized, peroxynitrite (OONO-) 343

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is a key component responsible for the cytotoxic effects of NO. Peroxynitrite is a highly reactive compound with harmful effects and may mediate cell damage (Muijsers et al., 1997). It has been suggested that most of the neurotoxic actions of NO are mediated by peroxynitrite, which damages cellular constituents including proteins, DNA and lipids (Beckman, 1991; Althaus et al., 1994; Dawson and Dawson, 1994, 1996; Bonfoco et al., 1995; Dawson, 1995). It is interesting to note that exposure of cortical neurons to relatively short durations or low concentrations of NMDA, S-nitrosocysteine, or 3-morpholinosydnonimine (which generate low levels of peroxynitrite) induces a delayed form of neurotoxicity predominated by apoptotic features. In contrast, intense exposure to high concentrations of NMDA or peroxynitrite induces necrotic cell damage (Bonfoco et al., 1995). This may explain why spinal root avulsion induces necrotic damage in motoneurons when NOS is maximally expressed, and apoptotic damage when injury-induced NOS levels are lower (see Section 6). NO may mediate and elevate cellular cGMP which in turn enhances tumor necrosis factor-alpha toxicity (Sherman et al., 1992). NO and cGMP are likely to be functional partners throughout the nervous system (Garthwaite. 1995). The coupling of the NOcGMP pathway to activation of NMDA receptors appears to be widely applicable in both physiological and pathological conditions. In addition, NO has also been found to be involved in cytokine-mediated neuronal apoptosis (Hu et al., 1997).

9.1.3. Evidence suggesting that NO/NOS is not involved in degenerative processes after neuronal injury Evidence that NO/NOS is not critically involved in neural degenerative processes after injury is obtained in some recent studies. It has been suggested that NO produced by phagocytes, neurons, or other cells may not play a significant role in secondary pathology after spinal cord injury (Cohen et al., 1996). Microinjection of NOS inhibitor, L-NAME, into the spinal cord reveals a positive dose-response relationship between L-NAME and neuronal loss, which indicates that the basal level of NOS activity is important for the survival of spinal neurons (Yezierski et al., 1996). Van Muiswinkel et al. (1998) have recently found that, in the development of fetal rat mesencephalic neurons grafted into a 6-hydroxydopamine (6-OHDA) lesioned rat, the maturation of grafted dopaminergic neurons coincides with a gradual increase in the expression of neuronal NOS within the graft. Dopaminergic cell numbers are not changed upon administration of L-NAME, which indicates that endogenously produced and potentially toxic NO does not affect the survival of grafted fetal dopaminergic neurons (Van Muiswinkel et al., 1998). Moreover, although it is hypothesized that NMDA toxicity is mediated by NO, systemic pretreatment of rats with 7-NI has no effect on lesion volumes after intrastriatal injection of NMDA (Loschmann et al., 1995). Pretreatment with dizocilipine maleate (MK801), but not L-NAME, reduces lesion size after intrastriatal injection of NMDA (Taylor et al., 1995). Studies using brain and hippocampal slices of both developing and more mature animals also show that the acute neurodegeneration mediated by NMDA or non-NMDA receptors in slice preparations is not mediated by NO (Garthwaite and Garthwaite, 1994). These studies indicate that formation of NO subsequent to NMDA receptor stimulation is not critically involved in excitotoxicity of NMDA. As reviewed above, NO seems to play a double-edged role in neuronal injury. Whether NO is protective or destructive may depend on the type of injury, population of injured neurons, stage of evolution of the injury, and the cellular source of NO. Different isoforms of NOS may play different roles in neurons in response to injury. In addition, the mechanisms of NO 344

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toxicity are dynamic and reversible processes that, if left unaltered, will lead to neuronal injury (Maiese et al., 1997). In the cerebral ischemia model, the role of NO in neuronal damage is dependent on the stage of evolution of the ischemic process and on the cellular source of NO (Iadecola, 1997). NO production by inducible NOS does not contribute to ischemic injury within 24 h after MCA occlusion, but may contribute to infarct maturation 2-4 days after ischemia (Yoshida et al., 1995). It is interesting to note that neuronal NOS has been shown to be neurotoxic while endothelial NOS appears to be neuroprotective (Huang et al., 1994). Using pharmacological and genetic approaches to study the roles of different isoforms in focal cerebral ischemia, Samdani et al. (1997a) proposed that nNOS and iNOS play key roles in neurodegeneration, while eNOS plays a prominent role in maintaining cerebral blood flow and preventing neuronal injury. Results of experiments in a thromboembolic stroke model also support this idea (Stagliano et al., 1997). However, inappropriate activation of NOS in endothelial cells and microglia, in response to MCA occlusion/reperfusion, has also been found to be closely associated with the initiation and progression of ischemic neuronal injury in the rat striatum (Nakashima et al., 1995). As discussed above, injury-induced nNOS is thought to be neurotoxic in ischemic injury and may cause neuronal death in the lesioned area. However, the nNOS neurons themselves retain their morphological structure in the area of the infarct (Z.G. Zhang et al., 1994), suggesting that nNOS-containing neurons are more resistant to ischemic insult. Indeed, it has also been reported that normal nNOS-containing neurons are more resistant to toxic effects of excitatory amino acids (Koh et al., 1986) and other neuronal degenerative processes (Ferrante et al., 1985). It is important to point out that NO/NOS may play different roles in neurons that normally contain the enzyme from those that do not normally express the enzyme. Normal NOS-containing neurons kill adjacent neurons through the action of NMDA receptor activation, though they remain relatively resistant to the toxic effects of NMDA and NO (Dawson et al., 1991). The precise mechanism of how these NOS-containing neurons resist NO toxicity is unknown. However, a recent study has shown that manganese superoxide dismutase (MnSOD) is enriched in normal NOS-containing neurons and MnSOD provides dramatic protection against NMDA and NO toxicity in neurons (Gonzalez-Zulueta et al., 1998). In contrast, neurons that normally do not express NOS may not contain any protective component against NO toxicity. It is possible that, when NOS is induced by injury in neurons that do not normally express NOS, these injury-induced NOS-containing neurons kill themselves due to the lack of protective mechanism against NO toxicity. Indeed, in spinal motoneurons, MnSOD is not detected when injury-induced NOS is over-expressed in the avulsed spinal segment (Wu, unpubl, data). The lack of protective mechanisms against the toxic effects of NO may result in the death of motoneurons following root avulsion. 9.2. POTENTIAL ROLE OF NOS IN NEURONAL REGENERATION There are relatively few data on the role of NOS in regeneration after neuronal injury. In the normal nervous system, NOS is highly expressed in neurons of the developing olfactory epithelium during migration and establishment of primary synapses in the olfactory bulb. This expression rapidly declines after birth. Seven days after birth NOS is undetectable in the olfactory receptor neurons (ORNs). However, following bulbectomy, NOS expression is rapidly induced in the regenerating ORNs and is particularly enriched in their outgrowing axons (Roskams et al., 1994). These findings suggest a prominent function for NO in neuronal regeneration. 345

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In an experiment in which fetal mesencephalic neurons are grafted into a 6-hydroxydopamine lesioned rat, increased NOS expression is found to coincide with the maturation of dopaminergic neurons in neural graft. This observation indicates a role of NOS in neuronal growth and regeneration (Van Muiswinkel et al., 1998). In optic nerve axotomy, the up-regulation of NOS in the retina seems to be important for the survival of injured retina ganglion cells since inhibition of NOS results in increased loss of ganglion cells (Huxlin and Bennett, 1995). Apart from its role in development, NOS has also been found to be important for the regeneration of some CNS neurons after injury. In the neurohypophyseal system, neurons in the supraoptic and paraventricular nuclei can regenerate their axons into the median eminence and onto the surface of the third cerebral ventricle following hypophysectomy (Wu et al., 1989). Regeneration of these neurons occurs between 1 and 2 weeks post axotomy, which is exactly coincident with the time course of up-regulation of NOS in the injured neurons (Wu and Scott, 1993). Up-regulation of NOS appears to correlate well with successful regeneration in the neurohypophyseal system. In fact, axonal regeneration in the neurohypophyseal system can be prevented by the use of nitroarginine, a competitive antagonist of NOS (Scott et al., 1995). On the other hand, however, it has been reported that NO/NOS is not associated with neural regeneration. Expression of NOS in spinal motoneurons after root avulsion is completely inhibited when the injured motoneurons regenerate their axons into an implanted PN graft (Wu et al., 1994a). Moreover, inhibition of NOS with L-NAME enhances regeneration of spinal motoneurons following sciatic nerve transection (Zochodne et al., 1997). These findings strongly suggest that NOS does not support regeneration in this population of neurons and may have a negative effect on neuronal regeneration.

10. SUMMARY

In conclusion, in response to neuronal injury, nNOS (the constitutive isoform of NOS) can be either up-regulated in neurons that normally express the enzyme or de novo induced in neurons that normally do not express the enzyme. Injury-induced expression of NOS is observed in many populations of neurons from the central nervous system to the peripheral nervous system following different types of neuronal injuries including axonal injury, ischemichypoxic injury, excitotoxic neuronal injury, chemical injury, etc. Expression of injury-induced NOS has been shown to be co-expressed with some other injury-related genes such as IEG, APE and neurotrophin receptor genes. Expression of injury-induced NOS can be regulated by experimental manipulations, which in turn affects the survival and regeneration of the injured neurons. Injury-induced NOS appears to play a double-edged role in neuronal injury. Whether injury-induced NO/NOS is beneficial or not for the injured neurons depends on types of NOS isoform, groups of neurons, age artd species of animal, and stage of the injury. It also depends on whether the injured neurons normally express NOS or not. Further study is needed to understand the precise mechanisms of injury-induced NOS in neuronal injury.

11. A C K N O W L E D G E M E N T S The author's research is supported by CRCG grants from the University of Hong Kong and a research grant from the Hong Kong Research Grant Council. I would like to thank Drs. K.F. 346

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So, H.K.F. Yip, M . M . L . Tan and K.L. S p a l d i n g for their critical r e v i e w s o f the m a n u s c r i p t . I w o u l d also like to t h a n k Mr. K.Y. L e u n g for d e v e l o p i n g the p h o t o g r a p h s and Ms. W.M. W o n g for t y p i n g part o f the m a n u s c r i p t .

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Wallis RA, Panizzon KL, Girard JM (1996): Traumatic neuroprotection with inhibitors of nitric oxide and ADP-ribosylation. Brain Res 710:169-177. Weiss SW, Albers DS, Iadarola MJ, Dawson TM, Dawson VL, Standaert DG (1998): NMDAR1 glutamate receptor subunit isoforms in neostriatal, neocortical, and hippocampal nitric oxide synthase neurons. J Neurosci 18:17251734. Weissman BA, Kadar T, Brandeis R, Shapira S (1992): NG-nitro-L-arginine enhances neuronal death following transient forebrain ischemia in gerbils. Neurosci Lett 146:139-142. Wu W (1992): Neuronal NADPH-diaphorase are related to survival and regeneration after severe neuronal damage. Soc Neurosci Abstr •8:860. Wu W (1993): Expression of nitric-oxide synthase (NOS)in injured CNS neurons as shown by NADPH diaphorase histochemistry. Exp Neurol 120:153-159. Wu W (1996): Potential roles of gene expression changes in adult rat spinal motoneurons following axonal injury: a comparison among c-jun, low affinity nerve growth factor receptor (LNGFR). and nitric oxide synthase (NOS). Exp Neurol 141:190-200. Wu W, Li L (1993): Inhibition of nitric oxide synthase reduces motoneuron death due to spinal root avulsion. Neurosci Lett 153:121 - 124. Wu W, Scott DE (1993): Increased expression of nitric oxide synthase in hypothalamic neuronal regeneration. Exp Neurol 121:279-283. Wu W, Scott DE, Gilman AM (1989): Correlative scanning-immunoelectron microscopic analysis of neuropeptide localization and neuronal plasticity in the endocrine hypothalamus. Brain Res Bull 22:399-410. Wu W, Han K, Li L, Schinco FP (1994a): Implantation of PNS graft inhibits the induction of neuronal nitric oxide synthase and enhances the survival of spinal motoneurons following root avulsion. E.W Neurol 121:335-339. Wu W, Li Y, Schinco FP (1994b): Expression of c-jun and neuronal nitric oxide synthase in rat spinal motoneurons following axonal injury. Neurosci Lett 179:157-161. Wu W, Liuzzi FJ, Schinco FP, Depto AS, Li Y, Mong JA. Dawson TM, Snyder SH (1994c): Neuronal nitric oxide synthase is induced in spinal neurons by traumatic injury. Neuroscience 61:719-726. Wu W, Li L, Penix JO, Gu Y, Liu H, Prevette DM, Oppenheim RW I1995): GDNF and BDNF inhibit the induction of NOS and prevent the death of adult rat spinal motoneurons Iollowing root avulsion. Soc Neurosci Abstr 2•:279. Wu W, Ju G, Yick LW, Liu HL, Lin YH, Wang HJ. Wang J (1997): Expression of nitric oxide synthase in axonal injured spinal motoneurons: a comparative study in various mammalian species from mouse to monkey. Soc Neurosci Abstr 23:1156. Wu Y, Li Y, Liu H, Wu W (1995): Induction of nitric oxide synthase and motoneuron death in newborn and early postnatal rats following spinal root avulsion. Neurosci Lett 194:109-112. Xiao BG, Zhang GX, Ma CG, Link H (1996): The cerebrospinal fluid from patients with multiple sclerosis promotes neuronal and oligodendrocyte damage by delayed production of nitric oxide in vitro. J Neurol Sci 142:114-120. Xie QW, Cho HJ, Calaycay J, Mumford RA, Swiderek KM. Lee TD, Ding A, Troso T, Nathan C (1992): Cloning and characterization of inducible nitric oxide synthase from mouse macrophages. Science 256:225-228. Xu ZQ, H6kfelt T (1997): Expression of galanin and nitric oxide synthase in subpopulations of serotonin neurons of the rat dorsal raphe nucleus. J Chem Neuroanat 13:169-187. Yamada K, Xu ZQ, Zhang X, Gustafsson L, Hulting AL, De Vente J, Steinbusch HW, H6kfelt T (1997): Nitric oxide synthase and cGMP in the anterior pituitary gland: effect of a GnRH antagonist and nitric oxide donors. Neuroendocrinology 65:147-156. Yezierski RP, Liu S, Ruenes GL, Busto R, Dietrich WD (1996): Neuronal damage tbllowing intraspinal injection of a nitric oxide synthase inhibitor in the rat. J Cereb Blood Flow Metab •6:996-1004. Yick LW, Wu W, So KF, Wong SY (1998): Time course of NOS expression and neuronal death in Clarke's nucleus following traumatic injury in adult rat spinal cord. Neurosci Lett 241:155-158. Yick LW, Wu W, So KF, Yip HK (1999a): Peripheral nerve graft and neurotrophic factors enhance neuronal survival and expression of nitric oxide synthase in the Clarke's nucleus after hemisection of spinal cord in adult rat. Exp Neurol 159:131-138. Yick LW, Wu W, So KF (1999b): Additive effect of NOS inhibitor and neurotrophic factors on the survival of injured Clarke's neurons. Neuroreport •0:2569-2573. Yoshida T, Waeber C, Huang Z, Moskowitz MA (1995): Induction of nitric oxide synthase activity in rodent brain following middle cerebral artery occlusion. Neurosci Lett 194:214-218. Yu WH (1994): Nitric oxide synthase in motor neurons after axotomy. J Histochem Cvtochem 42:451-457. 352

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Yu WH (1997): Regulation of nitric oxide synthase expression in motoneurons following nerve injury. Dev Neurosci •9:247-254. Yun HY, Dawson VL, Dawson TM (1997): Nitric oxide in health and disease of the nervous system. Mol Psvchiatn' 2:300-310. Zhang J, Snyder SH (1995): Nitric oxide in the nervous system. Annu Rev Pharmacol Toxicol 35:213-233. Zhang J, Dawson VL, Dawson TM, Snyder SH (1994): Nitric oxide activation of poly (ADP-ribose) synthetase in neurotoxicity. Science 263:687-689. Zhang X, Verge V, Wiesenfeld Hallin Z, Ju G, Bredt DS, Snyder SH, H6kfelt T (1993): Nitric oxide synthase-like immunoreactivity in lumbar dorsal root ganglia and spinal cord of rat and monkey and effect of peripheral axotomy. J Comp Neurol 335:563-575. Zhang X, Ji RR, Arvidsson J, Lundberg JM, Bartfai "I". Bedecs K, H6kfelt T (1996): Expression of peptides, nitric oxide synthase and NPY receptor in trigeminal and nodose ganglia after nerve lesions. Etp Brain Res 111:393-404. Zhang ZG, Chopp M, Gautam S, Zaloga C, Zhang RL. Schmidt HH, Pollock JS, F6rstermann U (1994): Upregulation of neuronal oxide synthase and mRNA, and selective sparing of nitric oxide synthase-containing neurons after focal cerebral ischemia in rat. Brain Res 654:85-95. Zhao XL, Yanai K, Hashimoto Y, Steinbusch HWM. Watanabe T (1996): Effects of unilateral vagotomy on nitric oxide synthase and histamine H3 receptors in the rat dorsal vagal complex. J Chem Nemvanat 11:221-229. Zochodne DW, Misra M, Cheng C, Sun H (1997): Inhibition of nitric oxide synthase enhances peripheral nerve regeneration in mice. Neurosci Lett 228:71-74.

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CHAPTER X

Nitric oxide-cGMP signaling in the rat brain J. DE VENTE AND H.W.M. STEINBUSCH

1. I N T R O D U C T I O N In this chapter we present an overview of the neuroanatomical relationship between constitutive nitric oxide synthase (NOS) and nitric oxide-mediated cGMP synthesis in the rat brain. The presence and properties of NOS in the brain have been dealt with in detail in the other chapters of this volume and only those aspects relevant for the discussion of NO-cGMP signal transductions will be touched upon in this chapter. There is compelling evidence that a major target for NO in the brain is the soluble isoform of guanylyl cyclase (sGNC). Although the second isoform, the particulate guanylyl cyclase (Garbers and Lowe, 1994; Wedel and Garbers, 1997) is also present abundantly in the rat brain, there is as yet no evidence that NO is involved, directly or indirectly, in the activation of this enzyme. Therefore we will not discuss cGMP synthesis in the rat brain in relation to this particulate isoform. In recent years a number of target structures for cGMP have been found to be present in the nervous system. It has been known for a considerable period of time that cGMP can regulate phosphodiesterase activity, i.e. a stimulation of cAMP hydrolysis by phosphodiesterase-2 and an inhibition of cAMP hydrolysis by phosphodiesterase-3 (Beavo and Reifsnyder, 1990). Another class of target molecules for cGMP are the cGMP-dependent protein kinases (e.g. Smolenski et al., 1998). High concentrations of cGMP-protein kinase I are present in the Purkinje cells of the cerebellum (Lohmann et al., 1981), and the cGMP-protein kinase II has a widespread distribution in the mammalian CNS (Uhler, 1993; E1-Husseini et al., 1995a). Target proteins for the cGMP-dependent protein kinases have been demonstrated in the CNS, i.e. DARPP-32 (Tsou et al., 1993), the inositol 1,3,4-triphosphate receptor (Koga et al., 1994; Komalavilas and Lincoln, 1996), G-substrate (Detre et al., 1984), NOS (Bredt et al., 1992), and the GABAA receptor (Leidenheimer, 1996). Probably much more of these will be identified in the near future (Wang and Robinson, 1997). Nuclear localization of cGMP-PK (Gudi et al., 1997) and of cGMP (Truman et al., 1996) indicate an involvement of cGMP in gene transcription. Recent reports have shown that cyclic nucleotide-regulated ion channels are widely distributed in the brain also (E1-Husseini et al., 1995b: Bradley et al., 1997; Zufall et al., 1997). The present data indicate that these channels are involved in translocating calcium into the cell (Kaupp, 1995: Biel et al., 1998). In addition, binding studies using 3H-cGMP showed that cGMP-binding proteins are present in all brain areas with large regional differences in the capacity to bind cGMP (Bladen et al., 1996; Bonkale et al., 1997). The presence of these target molecules for cGMP throughout the nervous system suggests important, although still largely undefined, roles for cGMP.

H a n d b o o k o f Chemical Neur~mnatom ~; ~1. 17: Functional ,Veuroamaom v ~tl~the Nitric Oxide System

H.W.M. Steinbusch, J. De Vente and S.R. Vincent, editors @, 2000 Elsevier Science B.V. All rights reserved.

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2. S O L U B L E GUANYLYL C Y C L A S E sGNC is a heterodimeric heme containing protein (Ignarro, 1989, 1994; Koesling et al., 1990; Giuli et al., 1992; Murad, 1994). Presently, tour subunits for sGNC are known, i.e. two large o~ subunits and two smaller [3 subunits (Zabel et al., 1998). The enzyme is composed of one o~ (o~l or 0~2) and one 13 (131 or [32) subunit. The coexpression of o~1 or 0~2 with 61 results in an active enzyme which can be activated by NO (Harteneck et al., 1990). Homodimer formation has been demonstrated; however, this results in an inactive enzyme (Zabel et al., 1999). The function of the 132 subunit is not clear. The expression of the 132 subunit may have a regulatory function as the combination ~1 or 0~2 with 132 results in an inactive enzyme. Little is known about the regulation of sGNC in the brain. It was demonstrated that cAMP decreased mRNA levels for the [31 subunit in fibroblasts (Shimouchi et al., 1993) and the o~l subunit in vascular smooth muscle (Papapetropoulos et al., 1995). Similarly, cAMP was found to mediate the decrease in 0~1 and 131 subunit in rat PC12 cells induced by NGF (Liu et al., 1997). Another line of evidence also indicates that the expressions of both 0~1 and [31 subunits are regulated in a coordinated fashion, as a tandem organization of both subunits was demonstrated in the Medaka fish sGNC (Mikami et al., 1999). In order to become activated, the sGNC has a heme group complexed with the heterodimer (Ignarro, 1989, 1994). The heine group has a very high affinity for NO (Ignarro, 1994, 1989; Traylor and Sharma, 1992; Tsai, 1994) and functions as the NO receptor. Formation of the NO-heine complex results in activation of the sGNC (Murad et al., 1978; Ignarro, 1989, 1994; Stone and Marietta, 1995; Dierks et al., 1997). There are several excellent reviews available on the mechanism of activation of sGNC (Waldman and Murad, 1987; Ignarro, 1989, 1994: Schmidt, 1992; Murad, 1994: Nakane and Murad, 1994: Koesling, 1998; Denninger and Marietta, 1999). sGNC has been found in the nervous system of all animal species studied so far (see also Chapter XI by Scholz and Truman). Early studies on the presence of sGNC activity in the rat brain revealed that this enzyme was present in every brain region, although there were large differences in activity between the regions (Ferrendelli, 1978). The localization of the enzyme has been studied using a number of different histochemical techniques (Zwiller et al., 1981; Ariano et al., 1982; Nakane et al., 1983: Matsuoka et al., 1992; Furuyama et al., 1993; Burgunder and Cheung, 1994; Smigrodzki and Levitt, 1996). The application of antibodies against the different subunits of sGNC should provide the most detailed information about the localization of this enzyme. Unfortunately, there are presently only publications describing the immunocytochemical application of subunit-specific antibodies to tissues outside the central nervous system (Mundel et al., 1995: Kummer et al., 1996: Davidoff et al., 1997). The presence of o~l and 0~2 subunits was demonstrated immunocytochemically in rat and guinea-pig dorsal root ganglia (Kummer et al., 1996). In this study the presence of the [31 subunit was demonstrated using RT-PCR analysis in this tissue but could not be visualized using a [31 subunit antiserum, whereas the reverse situation was found for the 132 subunit. In addition, polyclonal antibodies against the isolated enzyme have been used in a number of studies and showed a widespread distribution of the enzyme throughout the rat brain in neurons and astroglial cells (Zwiller et al., 1981; Ariano et al., 1982; Nakane et al., 1983). A number of discrepancies between these studies were attributed to specificity differences of the antibodies, or the source of sGNC (Nakane et al., 1983). However, there are several arguments which necessitate that the results, and especially the interpretation of the results as cited in the literature, are evaluated with care. Therefore, these studies will be reviewed in detail in the section on the localization of the NO-mediated cGMP synthesis in the cerebellum (Section 6). 356

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3. PHOSPHODIESTERASE ACTIVITY AND THE TERMINATION OF THE N O - c G M P SIGNAL

The 3',5'-cyclic nucleotide phosphodiesterases represent a family of enzymes of bewildering complexity (Beavo, 1995). Presently, ten subfamilies have been identified based on structural data and biochemical properties (Soderling et al., 1999). Each subfamily might consist of many different variants (Beavo, 1995; Houslay, 1998). Phosphodiesterases have a high affinity for either cAMP or cGME or both nucleotides. In addition, there is a large number of inhibitors of phosphodiesterase activity available with a selective profile ranging from nonselective (e.g. isobutyl-methylxanthine, IBMX) to highly selective (e.g. sildenafil and zaprinast as inhibitors of cGMP-selective phosphodiesterase-5). It has been demonstrated that some phosphodiesterases are highly localized in a particular tissue or specific cell type. Calcium/calmodulin-dependent phosphodiesterase activity (phosphodiesterase-1) is highly expressed in different areas of the rat and mouse brain and was found to be present especially at postsynaptic sites (Kincaid et al., 1987; Ludvig et al., 1991; Sonneburg et al., 1993; Furuyama et al., 1994). Subsequently it was shown that there are different forms of phosphodiesterase-1 present in the brain, with a different expression pattern (Polli and Kincaid, 1994; C. Yan et al.. 1994). Thus, phosphodiesterase- 1B 1 (63 kDa isoform) was present at high levels in the mouse or rat caudate putamen, whereas phosphodiesterase-lA2 .(61 kDa isoform) was present in this area in only a subset of small interneurons (Polli and Kincaid, 1994; C. Yan et al., 1994). Similarly, different forms of phosphodiesterase-2 (cGMP-stimulated cAMP-/cGMP-hydrolyzing phosphodiesterase) have been characterized and the results presently available indicate differences between the localization of these isoforms (Sonnenburg et al., 1991; Repaske et al., 1993; Yang et al., 1994). It was shown that a subpopulation of olfactory receptor cells expressed a phosphodiesterase-2, but not all of them (Juilfs et al., 1997). Two isoforms of the cGMP-inhibited cAMP-hydrolyzing phosphodiesterase-3 have been characterized and were found to have a different expression pattern in the rat brain (Reinhardt and Bondy, 1996). A large number of different isoforms of the cAMP-specific phosphodiesterase-4 have been described (e.g. Houslay, 1998) and it was recently shown that there are important differences in the expression of these subclasses (Cherry and Davis, 1999). Of the cGMP-specific phosphodiesterases, the phosphodiesterase-5 is present in blood vessel walls; however, a neuronal localization of this isoform has not been reported yet. Phosphodiesterase-6, which is uniquely regulated through G-proteins, has so far only been described in the retina (e.g. Baylor, 1996). Recently, a new cGMP-specific phosphodiesterase has been described, the phosphodiesterase-9, which is present in the brain of mice and human (Fisher et al., 1998; Guipponi et al., 1998; Soderling et al., 1998). This phosphodiesterase-9 was not inhibited by IBMX and only at a relatively high concentration (30 btM) by zaprinast (Fisher et al., 1998; Soderling et al., 1998). Very recently, a new phosphodiesterase has been reported which occurs in high concentrations in the mouse or human brain. This enzyme has a very high affinity for cAMP and a somewhat lower affinity for cGME and is potently inhibited by dipyridamole (Kotera et al., 1999; Loughney et al., 1999; Soderling et al., 1999). Two isoforms of the phosphodiesterase-10 have been cloned, one of these is expressed in the brain especially during development (Kotera et al., 1999). In view of the kinetics of this phosphodiesterase-10, this enzyme might be considered as a cAMP-inhibited cGMP-phosphodiesterase (Soderling et al., 1999). The finding that a specific form of a phosphodiesterase is only expressed during development (Kotera et al., 1999) is not unique. Observations on the developmental regulation of 357

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phosphodiesterase activity have been made for the 63 kDa calmodulin-dependent phosphodiesterase-1 (Billingsley et al., 1990: Sakagami et al., 1995) and for phosphodiesterase-3B (Reinhardt and Bondy, 1996). Phosphodiesterase activity is regulated by a number of endogenous factors. As already indicated above, calcium/calmodulin are activators of the phosphodiesterase-1 isoforms (Beavo and Reifsnyder, 1990; Beavo, 1995) and cyclic nucleotides are regulators of several subclasses of the phosphodiesterases (see above). There are a number of reports on endogenous activators or inhibitors of phosphodiesterase activity which have not been identified yet (Bums et al., 1992; Lochhead et al., 1997). For several of these factors it still has to be investigated how specific these are for the respective subfamilies or isoforms. Only in a few reports a detailed analysis on the specificity of activation or inhibition has been presented. Activation or inhibition of phosphodiesterases by endogenous compounds present potentially important regulatory mechanisms in potentiating or terminating the messenger function of cGMR Phosphodiesterase activity may also be triggered by an external signal. Placed in a historical context, the activation of phosphodiesterase-6 in the retina is a prototypical example of phosphodiesterase activity activated by a exogenous signal. We presented evidence that in the hippocampus slice preparation, a mild extracellular stimulus like 10-20 mM K+-activated cGMP-hydrolyzing phosphodiesterase activity within seconds, and this activation seemed to be directly related to the increased potassium concentration, as the increase in cGMP per se did not activate this phosphodiesterase activity (De Vente and Steinbusch, 1992). This phosphodiesterase activity could not be inhibited by 1 mM IBMX, the compound which is often inhibitor of phosphodiesterase activity of choice as IBMX shows a broad range of nonspecific activity. However, there are other reports in the literature which showed that IBMX does not inhibit all phosphodiesterase activity (Mayer et al., 1992; Fisher et al., 1998; Soderling et al., 1999).

4. cGMP IMMUNOCYTOCHEMISTRY When antibodies against cyclic nucleotides were developed early in the 70s (Steiner et al., 1972) they were immediately introduced as tools for immunocytochemistry (Wedner et al., 1972). These antibodies were raised against the 2'O-succinyl-derivative of the cyclic nucleotides, which had been coupled to human serum albumin or keyhole limpet hemocyanin with carbodiimide (Steiner et al., 1972). The first study detected cAMP in mice parotid glands stimulated with isoproterenol (Wedner et al., 1972). In neural tissue, the first immunocytochemical study localized cAMP in specific cerebellar neurons, i.e. Purkinje neurons and granule cells (Bloom et al., 1972). cGMP immunocytochemistry using the Steiner-type of antibodies was successful in demonstrating cholinergic elevation of cGMP levels in the superior cervical ganglion (Kebabian et al., 1975). This study showed that it was possible to demonstrate selectively increased levels of cAMP or cGMP in one tissue using the Steinertype of antibodies. Notwithstanding this initial success, it had become apparent that during the subsequent steps of the immunocytochemical procedure there was an almost complete loss of nucleotides due to the necessity of using unfixed tissue sections (Wedner et al., 1972; Cumming et al., 1980; Ortez et al., 1980; Cumming, 1981). Fixation of tissue sections with routine fixatives resulted in decrease or loss of immunostaining (Wedner et al., 1972; Cumming et al., 1980; Ortez et al., 1980), although this problem appeared to be less serious for cGMP (Chan-Palay and Palay, 1979). It was proposed that these antibodies probably 358

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visualized cyclic nucleotides bound to proteins like protein kinases and/or phosphodiesterases (Steiner et al., 1976), although this has never been actually shown. In order to overcome the above-mentioned drawbacks in the use of cyclic nucleotide antisera, we decided to raise new antisera based on the criteria that (a) small water-soluble molecules need to be fixed to the tissue matrix in order to allow a reproducible visualization, and (b) the crosslinker used to prepare the immunogen should be identical to the fixative used in the immunocytochemical procedure. This approach was previously successfully applied in raising antibodies to various monoamines, i.e. serotonin and noradrenaline (Steinbusch et al., 1978, 1981, 1983). For cGMR freshly depolymerized formaldehyde, or formaldehyde vapor, proved to be the fixative of choice (De Vente et al., 1987b, 1989c). It was shown that using this approach the loss of cyclic nucleotides was about 70% in a gelatin model system using a similar procedure as used for tissue processing (De Vente et al., 1987b). This represents a significant improvement with respect to the method described by Wedner et al. (1972). Preliminary results of our group indicate that the loss of cGMP from tissue sections after formaldehyde fixation is even considerably less than 70% (De Vente, unpubl, results). Although some loss of cGMP from the tissue compartment still occurs, and the kinetics of fixating cGMP in the tissue are not known, application of the cGMP antiserum proved reproducible and the immunocytochemical results correlated very well with the biochemical assay of cGMP (De Vente et al., 1987a. 1989a.b, 1990; De Vente and Steinbusch, 1992; Hopkins et al., 1996; W. Van Staveren et al. in prep.).

5. I M M U N O C Y T O C H E M I C A L LOCALIZATION OF NOS AND NO-MEDIATED cGMP SYNTHESIS 5.1. A NOTE ON THE USE OF BRAIN SLICES For the first attempts to visualize cGMP using the newly developed antibodies against formaldehyde-fixed cGMP there seemed to be no obvious reason to use brain slices for the identification of brain structures capable of synthesizing cGMR However, early studies showed that a routine perfusion fixation with formaldehyde-based fixatives were disappointing in this respect (De Vente et al., 1987b). One causative factor might be the cGMP-lowering effects of anesthetics (e.g. Kant et al., 1980; see also De Vente et al., 1990). Preperfusing animals with sodium nitroprusside (SNP) resulted in cGMP-IS in some brain areas (Berkelmans et al., 1989); however, this appeared to be restricted to areas in which the blood-brain barrier was not complete. Later Southam and Garthwaite (1993) demonstrated that preperfusion of the animal with a high dose of SNP in conjunction with a phosphodiesterase inhibitor resulted in an enhanced cGMP accumulation in virtually all regions of the rat brain. As we were interested in the pharmacology of sGNC activation and breakdown of cGMP, we adopted the brain slice procedure as our method of choice. This method offers the possibility to construct dose-response curves and to test different compounds in one single experiment. One inherent problem with this approach is reproducibility as each brain slice is unique by itself. This may give problems in evaluating results obtained in consecutive slices (even from the same region of the same animal). The slicing method proved to be a reliable approach in studying NO-cGMP signaling in the CNS. Although many modifications of the basic methods to prepare and incubate brain slices in vitro can be found in the literature (for a review see Aitken et al., 1995, and subsequent articles in the same volume), we experienced that a speedy procedure from the moment of decapitation of the animal in combination with cooling of the 359

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tissue during the slicing procedure was the most critical step (Garthwaite et al., 1979; De Vente and Steinbusch, 1997). 5.2. A NOTE ON THE USE OF PHOSPHODIESTERASE INHIBITORS IN IN VITRO STUDIES In vitro incubation of slices from an adult rat brain for 45 rain in the absence of a phosphodiesterase inhibitor, followed by formaldehyde fixation, sectioning and processing for cGMP immunocytochemistry results in an almost negligible amount of cGMP-IR. Under such circumstances cGMP-IR might be visualized weakly in some blood vessel walls and occasionally an astrocyte might be observed. Incubation of slices in the presence of an NO-donor but without phosphodiesterase inhibition resulted, in addition, in the appearance of very few isolated fibers. So far, most studies have used IBMX as a nonselective phosphodiesterase inhibitor. Inclusion of this phosphodiesterase inhibitor during the incubation also results in little cGMP-IS in the unstimulated slices (e.g. Fig. l a), cGMP-IS might be increased in blood vessel walls and in some brain areas a more intense cGMP-IR is observed (see Section 5.3). Comparison of these results with the visualization of abundant cGMP-IR when SNP and IBMX were both present during the in vitro incubation (Fig. lb), immediately suggests that phosphodiesterase activity must be high in the brain. Excluding the retina, four phosphodiesterases are presently known which have a sufficiently low Km for cGMP to be considered as cGMP-selective enzymes (Beavo, 1995; Soderling et al., 1999), i.e. a cGMP-selective calcium-/calmodulin-dependent phosphodiesterase-1, -5, -9, and -10. Little is known about the localization of these enzymes in the CNS (see also Section 3). However, these enzymes may be of decisive importance when attempting to localize cGMP-synthesizing structures using immunocytochemistry. The selectivity of the available phosphodiesterase inhibitors might in principle be large enough to be of use in an attempt to localize cells which express specific phosphodiesterase isoforms. Although some attempts have been made (Shuttleworth et al., 1993: De Vente et al., 1996), the results have been disappointing so far. The concentration of zaprinast, a selective inhibitor of the cGMP-specific phosphodiesterase-5, which resulted in a detectable increase in cGMP-IR in the hippocampus,

Fig. 1. cGMP-IS in the rat frontal cortex: (a) slice incubated in the presence of 1 mM IBMX but without NO-donor; (b) slice incubated in the presence of 1 mM IBMX and 0.1 mM SNR Exposure time of the film was determined using b and kept the same when taking a. Further processing of the pictures was the same for a and b. For further experimental details consult the legend of Fig. 2. Bar represents 50 gin.

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was too high to be classified as a selective effect (De Vente et al.. 1996). Nevertheless, inhibition of phosphodiesterase-2 by eo'thro-9-(2-hydroxy-3-nonyl)adenine (EHNA) or inhibition of, possibly, phosphodiesterase-I0 by dipyridamole both in concentrations ranging from 1 to 10 ~tM, in combination with an NO-donor resulted in an abundant increase in cGMP-IS in the hippocampus as well as in other brain areas (W. Van Staveren et al. in prep.). As it is not known how many more members of the phosphodiesterase enzyme family will be discovered in the (near) future, it becomes a precarious endeavor to try to inhibit all phosphodiesterase activity in brain slice preparations at present. Nevertheless, IBMX has been shown to be a potent inhibitor of general phosphodiesterase activity, and, as long as no better tool is available, it remains the drug of choice. 5.3. NOS ACTIVITY IN BRAIN SLICES To assess ongoing NOS activity in brain slices, the L-arginine-L-citrulline conversion method (Bredt and Snyder, 1989) is difficult and complicated to interpret. NO has a very high affinity for sGNC and in principle cGMP immunocytochemistry can be used as a read-out parameter of NOS activity. This can be tested in various ways. Current concepts state that NOS is activated through a mechanism which is triggered by activation of glutamate receptors residing on NOS-containing postsynaptic elements. The NOS might even interact directly with the NMDA-type glutamate receptor through PDZ-domains (Kornau et al., 1995: Brenman et al., 1996; Niethammer et al., 1996). Glutamate-induced calcium influx results in activation of calmodulin and subsequently NOS (see Ignarro and Jacobs, Chapter I). NO diffuses away from the site of synthesis. The physicochemical properties of NO allow rapid diffusion through biological membranes (Lancaster, 1994: Wood and Garthwaite, 1994), making NO an ideal candidate for an intercellular, or retrograde messenger molecule (Garthwaite, 1991: O'Dell et al., 1991; Garthwaite and Boulton, 1995). The maximal diffusional spread of NO has been calculated to be around 300 ~tm (Lancaster, 1994: Wood and Garthwaite, 1994). This distance is probably too large as a number of tissue factors, e.g. iron-heine or the hydrophobic interior of biological membranes, decrease the life time of NO dramatically (Ignarro et al., 1993; Kharitonov et al., 1994: Hakim et al., 1996: Liu et al., 1998). Furthermore, studies by Lev-Ram et al. (1995) indicate that the diffusion distance of NO is close to 10 ~tm. Also, we showed that in the islands of Calleja the diffusion distance of NO is very limited compared to the reported value of 300 ~tm (De Vente et al., 1998b), the maximal distance being probably close to 20 ~tm. As stated before, incubation of brain slices in the presence of 1 mM IBMX but in the absence of an NO-donor shows only little cGMP-IS in blood vessel walls (Fig. l a). However, under these conditions there is intense cGMP-IR in some rat brain areas, i.e. olfactory bulb (Hopkins et al., 1996), islands of Calleja (De Vente et al.. 1989b, 1998b) (see also Fig. 24b), the paraventricular thalamic nucleus (Blokland et al., 1999), the superior colliculi, the cerebellum (De Vente et al., 1989b, 1990: De Vente and Steinbusch, 1992) (see also Fig. 22b), the medial vestibular nuclei, the nucleus of the solitary tract, and the nucleus ambiguus (De Vente et al., 1998a). This "basal" immunostaining can be completely abolished by incubating the slices in the presence of 0.1 mM of the NOS inhibitor NG-nitro-L-arginine methylester (L-NAME). This is evidence for the presence of ongoing endogenous NOS activity during incubation of the slice. In addition, we were able to show in slices from the olfactory bulb, using both cGMP immunocytochemistry and biochemical determination of cGMP, that NMDA-type glutamate receptors were involved in NO-cGMP signal transduction (Hopkins et al., 1996). 361

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J. De Vente and H.W.M. Steinbusch

5.4. LOCALIZATION OF NO-MEDIATED cGMP ACCUMULATION IN BRAIN SLICES

5.4.1. Telencephalon The localization of cGMP-IS in relation to NOS-IS in the olfactory bulb has been reported in detail before (Hopkins et al., 1996). In Fig. 2a,b we show both cGMP- and NOS-IS in the glomerular layer although not in the same structures, cGMP-IS is also prominent in the olfactory nerve layer, a structure which has been reported to be devoid of NOS. Similarly, we did not observe any colocalization between cGMP- and NOS-IS in the other layers of the olfactory bulb, although cGMP-IR was present in granule cells. cGMP- and NOS-IS were prominently present in a network of varicose fibers in the cingulate (Fig. 2c,d), the frontal, the parietal, and the gpiriform cortex. The density of the neuronal network appeared to be slightly greater in the more superficial layers (layers I-III) of the cortex (see also Figs. lb, 30a,c and 31a). In all cell layers, we observed occasionally glial and neuronal cell somata. In these cortical areas, NOS-IS was found in a varicose fiber network with few somata. NO activated the sGNC in a network which, like the NOS-containing network, stretches throughout the cortical areas; however, we did not observe any colocalization between cGMP- and NOS-IS. In the corpus callosum NOS-IS somata or fibers were observed only occasionally (Fig. 3a,b). In agreement with this observation, there were relatively few fibers which showed cGMP-IS in this area. In addition, many astrocytes were observed in the region of the genu of the corpus callosum, whereas cGMP-IR astrocytes were scarce in other parts of the corpus callosum. Also, abundant cGMP-IS fibers were observed in the bundle of Probst. cGMP-IS was not observed in NOS-positive structures in the corpus callosum. cGMP- and NOS-IS were found in a dense network of varicose fibers in the caudate putamen complex (Fig. 3a,b). Only a small number of NOS-IS cell somata were found in this region. The NO-induced cGMP accumulation in the varicose fiber network appeared to be uniformly distributed throughout the caudate putamen complex. Similar observations were made in the nucleus accumbens, the tubercle olfactorium, the ventral pallidum, and the septal region. The lateral globus pallidus presented an exception in the sense that a relative absence of cGMP- and NOS-IS was found: both markers were observed in fibers encircling the myelinated fiber bundles which often contained cGMP-IR astrocytes. The islands of Calleja, including the major island located in the septum, were stained intensely for both cGMP and NOS (see also Fig. 24b). As has been reported in detail (De Vente et al., 1998b), we observed extensive colocalization between cGMP-IR and NOS-IS in the islands. In the hippocampus a dense varicose, apparently neuronal fiber network was visualized which was cGMP-IR (Fig. 3c). Occasionally, cGMP-IS in neuronal cell somata and astrocytes was observed. The intense cGMP-IS contrasted with the relatively sparingly distributed NOS-IS. We observed some (inter)neurons which were NOS-IS (Fig. 3d). Colocalization between cGMP- and NOS-IS was observed in a small number of cells (not shown), probably interneurons. No NOS-IS could be demonstrated convincingly in the pyramidal and granule cells in the hippocampus. This finding contrasts with reports describing intense labeling for NOS in these cells (Egberongbe et al., 1994; Endoh et al., 1994; Rodrigo et al.. 1994). However, notwithstanding the fact that we used two different polyclonal nNOS antisera, raised in rabbits (Schmidt et al., 1992) or in sheep (Herbison et al., 1996), which showed identical staining for NOS in other brain areas, and in addition, applying NADPH-diaphorase histochemical staining, using different fixation protocols (e.g. Dinerman et al., 1994), staining of pyramidal 362

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and granule cells in the hippocampus was at best doubtful. A similar result has been described by Vincent and Kimura (1992). It has been described repeatedly that NADPH-diaphorase staining is to some extent dependent upon the degree of fixation (Matsumoto et al., 1993a; Dinerman et al., 1994; W6rl et al., 1994; Buwalda et al., 1995: Vaid et al., 1996: see also Chapter II by Vincent). Therefore, although the NADPH-diaphorase staining is reproducible, controversies in this field need to be resolved by comparing the results of application of NOS antibodies with the histochemical approach. Strong evidence for the presence of NOS in pyramidal neurons was presented by Chiang et al. (1994) using a single-cell PCR approach. The constitutively expressed NOS detected in the cytoplasm of cultured primary hippocampal neurons was the neuronal isoform. This contrasts with the endothelial NOS isoform described by Dinerman et al. (1994). In view of these confusing results, it is important that the presence of neuronal NOS or endothelial NOS is confirmed in pyramidal cells from the adult rat or mouse. The central amygdala was relatively devoid of NOS-IS, whereas cGMP-IS was strong in this region (Fig. 4). The piriform cortex was heavily stained for cGMP-IR which was observed in very thin ramifications. No evidence for colocalization was found in this region. In the anterior commissure we frequently observed glial cells which were cGMP-IR.

5.4.2. Diencephalon The bed nucleus stria terminalis showed intense cGMP-IS and in the same area NOS-IS was prominent in a large number of cells and fibers. Some basal cGMP-IS was found in the thalamic paraventricular nucleus (Blokland et al., 1999). Partial colocalization between cGMP- and NOS-IS was found in the medial habenula (Fig. 5a,b). The medial aspect of the lateral habenula showed a dense immunostaining for both cGMP and NOS in a multitude of fibers and punctate staining structures, demonstrating extensive colocalization in this area. In the subthalamic nucleus a number of NOS-IS cells were observed; in this region intense colocalization of cGMP-IS was also observed in a number of NOS-IS cells (Fig. 5c,d). In the dorsomedial and the dorsoventral thalamic nuclei we found a dense network of cGMP-IR fibers and a large number of cGMP-IS neurons, whereas NOS-IS was relatively scarce in varicose fibers. In the pretectal area and the lateral geniculate nucleus we found intense cGMP-IS, sometimes colocalizing with NOS-IS (Fig. 6). Intense cGMP-IS was observed in the fibers running along the dorsolateral side of the dorsal geniculate nucleus (Fig. 6c). Optical tract and internal capsule contained fibers which stained strongly for cGMP-IR (Fig. 7a,b). Some NOS-IS somata were also found in this region (Fig. 7c) although no colocalization was observed. The thalamic reticular formation showed strong cGMP-IS (Fig. 34a), similar to that found earlier in perfusion-fixed whole animals (De Vente et al., 1987b). cGMP-IS in the hypothalamic area was very dense in thin varicose fibers (Fig. 7d) often encircling NOS-IS cell somata (Fig. 7e). The density of the cGMP-IS fibers varied somewhat between different areas of the hypothalamus, with the preoptic area showing intense cGMP-IS and the ventromedial hypothalamic areas showing relatively little cGMP-IS. In the ventral part of the suprachiasmatic nucleus we found strong cGMP-IR, whereas the dorsal part was

Fig. 3. cGMP-IS and NOS-IS in the same sections from the areas of the caudate putamen (a, b), and the hippocampus (c, d). Antibodies used sheep anti-cGMP and rabbit anti-NOS. Abbreviation: cc = corpus callosum. Bar represents 50 lam.

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almost devoid of cGMP-IS. We could not demonstrate NOS or NADPH-diaphorase staining in the suprachiasmatic nucleus. Nevertheless, others have presented evidence for the presence of NOS in the suprachiasmatic nucleus (Wang and Morris, 1996; Chen et al., 1997). Clear colocalization between cGMP- and NOS-IS was observed in the median eminence. Intensity of the cGMP-IS was clearly distinct between the supramammillary nucleus and the medial mammillary region (Fig. 8c,d) and a similar situation was observed using NOS-IS (Fig. 8a,b). The intensity of the staining patterns, both for NOS and cGMI:', made it impossible to make a clear statement about colocalization between the two markers in this area.

5.4.3. Mesencephalon In the mesencephalon we observed dense cGMP-IS, generally in thin, ramifying varicose fibers. This staining pattern was observed in virtually all regions, cGMP-IS and NOS-IS overlapped in all regions. Colocalization between NOS and cGMP-IR was observed in thin fibers in the red nucleus (Fig. 9a,b). No colocalization was observed in the region of the substantia nigra (Fig. 10) although cGMP- and NOS-positive fibers, strikingly, often followed the same course. In the interpeduncular region cGMP-IS and NOS-IS both showed the same characteristic staining pattern, and the intensity and density of the immunostainings suggest that both markers were present in the same fibers (Fig. 11). A similar dense labeling was observed in both the colliculi (Fig. 12), the central gray region (Fig. 13a-d), the dorsal raphe nucleus (Fig. 13e-h), and the commissure of the inferior colliculus (Fig. 14).

5.4.4. Cerebellum cGMP-IS was identified in parallel fibers in the molecular layer when a 5-~m-thick section from an unstimulated slice was immunostained, next to Bergmann glial fibers and cell somata (Fig. 15a). NOS-IS in this section was also visible in parallel fibers and granule cells (Fig. 15b), and was absent in Purkinje cells. Although an occasional presence of NOS in Purkinje cells has been reported (Rodrigo et al., 1994: Buwalda et al., 1995). our finding is in general agreement with the literature (e.g. Schmidt et al., 1992: Vincent and Kimura, 1992). In the cerebellar slices stimulated with SNP, cGMP-IS in the molecular layer was too intense to observe cGMP-IR in individual parallel fibers (e.g. Figs. 17a, 22d and 35); however, cGMP-IR was observed in granule cells, glomeruli, and glial cells. As described before (De Vente et al., 1989b), adult Purkinje cells were devoid of cGMP-IS, or cGMP-IS was very weak (see also Fig. 22b,d). The question whether Purkinje cells express sGNC has a history of about twenty years. This question can be taken as a test-case for cGMP immunocytochemistry and therefore will be discussed in a separate section.

Fig. 4. cGMP-IS and NOS-IS in the same section from the area of the amygdala. In c and d an enlargement is shown from the area marked "A'. Abbreviations: A = central amygdala: BLA = basolateral amygdala: Pir = piriform cortex. Antibodies used: rabbit anti-cGMP and sheep anti-NOS. Bars are 100 ltm (a, b) and 50 Itm (c, d).

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5.4.5. Pons and medulla oblongata In all regions of the pons and the medulla oblongata we observed cGMP-IS in close proximity to NOS-IS without actual colocalization. Only in certain areas some colocalization between the markers was found. Especially in the pontine reticular formation and the intermediate reticular formation we observed frequent colocalization between cGMP- and NOS-IS (Fig. 18). Very dense immunostaining for cGMP and NOS was observed in the posterodorsal tegmental nucleus (Fig. 16). An intense cGMP-IS was observed in what appeared to be the plasma membrane of large cells in the region of the locus coeruleus (Fig. 16b,d) and this obviously colocalized with NOS-IS. Using SIN-1 as a NO-donor, Xu et al. (1998) reported cGMP-IS in the cell somata of the locus coeruleus, colocalizing with TH. A role for cGMP in signaling in the locus coeruleus is supported by the finding of the presence of cGMP-PKII in this area and the effects of activators and inhibitors of cGMP-PKII on neuronal firing (Pineda et al., 1996). The medial vestibular nucleus contained strong cGMP- and NOS-IS; however, no colocalization was observed (Fig. 15c,d). The prepositus of the hypoglossal nerve was densely immunostained for both cGMP and NOS, suggestive of a colocalization (Fig. 17). The adjacent genu of the facial nerve contained little cGMP-IS and NOS-IS was absent. Some colocalization was observed in the region of the lateral cuneate nucleus and the inferior olive (Fig. 19). However, colocalization was more the exception than the rule. All parts of the inferior olive showed an intensely cGMP-IS in fibers and no cGMP-IS was observed in cell somata. In the complex area of the nucleus of the solitary tract we found cGMP-IS in cell somata in the caudal part of the nucleus (Fig. 20a) even in the unstimulated slice. Although this area also showed an intense NOS-IS, we did not clearly observe a colocalization between cGMP- and NOS-IS (Fig. 20b). In the spinal cord, cGMP-IR was found to be abundantly present in the gray matter, especially in layers I-III in varicose fibers and astrocyte-like cells. No colocalization between cGMP and NOS was observed in the adult spinal cord (Fig. 21c,d) (see also Vles et al., 2000). Although the function of cGMP in the spinal cord is still unknown, is already evident that cGMP has different electrical effects on specific neurons in the spinal cord, either being inhibitory or excitatory (Schmidt and Pehl, 1996). 372

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5.5. COLOCALIZATION OF cGMP AND NOS Our data have shown convincingly, and extended observations by others (Southam and Garthwaite, 1993) that NOS- and cGMP-IS structures are generally situated to each other within close proximity. This is in line with a rather short effective diffusion distance of NO. This means that both fiber systems overlap each other in a complementary way (Southam and Garthwaite, 1993). At the same time, we presented evidence that colocalization between NOS-IS and cGMP-IS in one cellular compartment is possible. In the brain stem, cGMP and NOS were frequently observed in the same fibers, and, sometimes, in cell somata. Especially in the cerebellum extensive colocalization between the two markers was observed in granule cells. This finding confirmed one of the conclusions of the group of Garthwaite on NO-signaling in the cerebellum (Garthwaite, 1991). Nevertheless, colocalization between 378

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NOS and cGMP is only minor compared to the abundance of independent staining throughout the brain. The observation that sGNC and NOS can be both present in a particular structure raises interesting questions about the regulation of both enzymes. In previous studies we showed that incubation of cerebellar slices in the presence of oxyhemoglobin abolished cGMP-IS (De Vente et al., 1990). This implies that the NOS present in the granule cells is not likely to be active when sGNC is activated. Similar observations were made in the islands of 380

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Calleja (De Vente et al., 1998b). As constitutive NOS requires a raised Ca 2+ concentration in order to be activated, and sGNC is inhibited by Ca :+, this has been suggested as a regulatory mechanism (Bredt and Snyder, 1989: Knowles et al., 1989). However, it was shown that sGNC is not inhibited by calcium concentration in the physiological range which is required for NOS activation (0.3 I.tM) (Mayer et al., 1992). This was supported by a recent report which showed that sGNC was inhibited by micromolar concentrations of free intracellular calcium (Parkinson et al., 1999). Therefore, it was argued that the activation of a calcium-dependent phosphodiesterase activity, not inhibited by IBMX, is responsible for cGMP breakdown in NOS-containing cells (Mayer et al., 1992). Recently, the presence of such phosphodiesterase activity in cerebellar cell cultures was reported (Agullo and Garcia, 1997); however, we have not been able to find any effect of phosphodiesterase-1 inhibitors like vinpocetine and calmidazolium, on cGMP accumulation in brain slices (unpublished results). In view of the extensive ramifications of NOS-containing cells, and taking into account the physicochemical 382

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Fig. 20. cGMP-IS and NOS-IS in the same section from the area of the nucleus of the solitary tract. Shown in a is the cGMP-IS in the basal, unstimulated situation (no NO-donor compound present during the in vitro incubation of the slices). Antibodies used: rabbit anti-cGMP and sheep anti-NOS. Abbreviation: cc = central canal. Bar represents 50 lt m.

Fig. 21. cGMP-IS in the cervical spinal cord: (a) basal cGMP-IS in a section from a slice at age PNI4: (b) cGMP-IS at P N I 4 in a slice stimulated with SNP: c and d shov,, respectively, cGMP-IS and NOS-IS in the same section from an adult slice incubated in the presence of SNR Bars represent 50 Itm (a and b: c and el). 383

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properties of NO, it was proposed by Bicker et al. (1997) that NO might be synthesized in one part of a NOS-containing fiber, and, after diffusion, might activate sGNC in the same fiber, but at a more distant locus.

6. LOCALIZATION OF cGMP IN THE CEREBELLUM

The cellular localization of cGMP in the cerebellum has been controversial for many years. In view of the importance of NO-cGMP signaling for LTD (Shibuki and Okada, 1991) this is not a trivial item. Different approaches have been used in the past, centering on the Purkinje cells as a source of cGMP synthesis. Therefore, we review the available literature on this particular topic. The observation that cGMP levels in the CNS are highest in the cerebellum is almost 30 years old (Ferrendelli et al., 1970, 1973). By extrapolating the knowledge at that time of the working mechanism of a series of drugs to the changes in cGMP levels in the cerebellum caused by these drugs, it was postulated that cGMP served as a second messenger in Purkinje cells (Biggio and Guidotti, 1976; Biggio et al., 1977). This hypothesis was, apparently, supported by the finding that cGMP levels were reduced in mutant mice lacking Purkinje cells (Mao et al., 1975). The elegant studies of Rubin and Ferrendelli (1977) on the content of cGMP in the different layers of the cerebellum were interpreted as demonstrating the presence of cGMP in Purkinje cells. The first immunocytochemical studies on the localization of cyclic nucleotides in the cerebellum demonstrated cAMP localized in Purkinje cells and other cellular structures (Bloom et al., 1972). In this study the application of cGMP antisera to the cerebellum was not mentioned. However, a few years later, Cumming reported the absence of cGMP in Purkinje cells, although the presence of cGMP in astroglial cells and fibers, and in granule cells could be demonstrated (Cumming et al., 1977, 1979). Similarly, another group (Chan-Palay and Palay, 1979) readily demonstrated cGMP in neuroglial cells in the cerebellum; however, the presence of cGMP in Purkinje cells could not be demonstrated convincingly. Nevertheless, again the presence of cAMP in Purkinje cells was readily demonstrated, using a similar Steiner-type antibody (Chan-Palay and Palay, 1979). It had been proposed that the Steiner-type antibodies preferably demonstrate cyclic nucleotides bound to target proteins (Steiner et al., 1976; see also p. 359). Thus, when a high concentration of cGMP-protein kinase I was found to be present in Purkinje cells in the cerebellum (Lohmann et al., 1981), this latter hypothesis appeared to be contrary to the finding that especially in the Purkinje cells no cGMP could be demonstrated. This discrepancy has been discussed (Cumming, 1981) in terms of differences in the loss of cyclic nucleotides from different types of cells, the characteristics of individual antibodies and the epitope organization in tissue sections, in this case especially the Purkinje cells. The detailed biochemical studies of Garthwaite could not provide evidence for cGMP synthesis in Purkinje cells. Working with dissociated cell suspensions enriched or depleted in Purkinje cells it was found that cGMP synthesis was especially strong in glial cells; however, no increase in cGMP accumulation or guanylyl cyclase activity could be detected in Purkinje cell-enriched suspensions (Bunn et al., 1986; Garthwaite and Garthwaite, 1987). These studies already suggested that the neuronal stimulus leading to guanylyl cyclase activation and the sites of cGMP synthesis were located in different compartments (Bunn et al., 1986; Garthwaite and Garthwaite, 1987), a prelude to NO as interneuronal messenger molecule (Garthwaite et al., 1988). Studies on the effects of climbing fiber lesions in vivo showed that cGMP synthesis in vivo occurred also in other cells than Purkinje cells (if at all) (Oltmans et al., 1987). Using 384

Nitric oxide-cGMP signaling in the rat brain

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a similar model mutant mouse deficient in Purkinje cells as used previously by Mao et al. (1975), Wood et al. (1994) found that NO-mediated cGMP synthesis did not take place in cerebellar Purkinje cells. Summarizing biochemical/pharmacological data and the immunocytochemistry experiments using the Steiner-type of antibodies, it can be stated that there is no clear evidence that the Purkinje cells are an important source of cGMP in the cerebellum. One publication described results of cGMP immunocytochemistry using a monoclonal antibody raised against a cGMP-glutaraldehyde conjugate (Sakaue et al., 1988). This publication has been cited a number of times as demonstrating the presence of cGMP-IR in Purkinje cells (Luo et al., 1994; Linden et al., 1995: Fedele et al., 1997). However, it has been demonstrated that Purkinje cells are surrounded by baskets which can stain strongly for cGMP (Figs. 22 and 35) (De Vente et al., 1987b; Southam et al., 1992) and, in addition, no Purkinje cells dendritic trees were visualized using the glutaraldehyde-cGMP antibody (Sakaue et al., 1988). These aspects have not been taken into account when evaluating the data of Sakaue et al. and till today there has been no follow up on this antibody. Immunocytochemical experiments using the antisera raised against formaldehyde-fixed cGMP (De Vente et al., 1987a) provided evidence for the localization of cGMP in Purkinje cells in the immature rat cerebellum at PN10 (Fig. 22a,c). In the adult rat cerebellum, cGMP-IS was demonstrated in Bergmann glia and basket cells in the Purkinje cell layer, astrocytes and granule cells in the granule cell layer, and in stellate cells and especially parallel fibers in the molecular layer (Fig. 15a)(De Vente et al., 1989b, 1990, 1998a: De Vente and Steinbusch, 1992; Southam et al., 1992; Southam and Garthwaite, 1993). Using this preparation, no, or very low levels of cGMP-IR were found in Purkinje cells in presence or absence of NO-donors, irrespective of the type of phosphodiesterase inhibitor used (nonselective like IBMX, or phosphodiesterase isoform-selective inhibitors like zaprinast, rolipram, calmidazolium, or EHNA) (De Vente, unpubl, results) (Figs. 22b,d and 35b,f). In addition, application of the atypical sGNC activator YC-1 (no NO-donor) did not result in cGMP-IR in Purkinje cells, either in the presence or absence of a NO-donor compound (Fig. 35c,d) (De Vente, in prep.). The sensitivity of the immunocytochemical method might be too low to detect cGMP-IR in the Purkinje cells. However, this seems unlikely, as calculations on the content of cGMP in the respective layers indicate that, if cGMP is to be present in Purkinje cells, the concentration is comparable to cGMP levels in the other cerebellar cortex layers (Rubin and Ferrendelli, 1977). Evidence in favor for cGMP synthesis in Purkinje cells seems to come from (immuno)histochemical studies. These studies can be divided in: (a) immunohistochemical studies using antibodies against sGNC or the respective subunits; (b) in situ hybridization studies. Application of the first antibody raised against sGNC (Lewicki et al., 1980) by Ariano et al. (1982) demonstrated neurons and astrocytic fibers to be immunoreactive. However, the claimed demignstration of cerebellar Purkinje cells containing sGNC is not corroborated by the published photographs (Ariano et al., 1982), as the sGNC-IR cells claimed to be Purkinje cells might be better characterized as Bergmann glial cells. The antibody raised against sGNC described by Nakane et al. (1983) immunostained cells in the cerebellum which have some characteristics of Purkinje cells somata. However, no Purkinje cell dendrites were visualized and the size of the depicted cells is more in line with Bergmann glial cells than with Purkinje cells (Nakane et al., 1983). Immunohistochemical results with a third antibody raised against the isolated sGNC was described by Zwiller et al. (1981). Application of this antibody to cerebellar sections indeed resulted in immunostaining of sGNC in cells which have characteristics of Purkinje cells, and preabsorption experiments resulted 385

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J. De Vente and H.W.M. Steinbusch

in loss of immunoreactivity in Purkinje cells. It is unfortunate that no other studies using this antibody are available. Recently, another antibody raised against sGNC isolated from rat brain was presented (Weinberg et al., 1998). Interestingly, application of this antibody did not result in the detection of sGNC in Purkinje cells, whereas other immunostaining characteristics of this antibody were very much in line with previous results and the results on the localization of NO-mediated cGMR Presently, antibodies against the subunits of sGNC have been developed (Koesling et al., 1990: Zabel et al., 1998); however, apparently these antibodies have not yet been applied for immunocytochemistry on cerebellar tissue sections. In situ hybridization studies of sGNC demonstrated the 131 subunit of sGNC mRNA in the Purkinje cell layer (Matsuoka et al., 1992: Furuyama et al., 1993; Giuili et al., 1994). Similarly, in situ hybridization studies on the localization of the c~l subunit ascribed a hybridization signal to either the Purkinje cell layer (Furuyama et al., 1993) or to the granule

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386

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cell layer (Burgunder and Cheung, 1994). Thus, taking into account the presence of other cells in the Purkinje cell layer, like basket cells and Bergmann glia cells which can accumulate large quantities of cGMP, and the uncertainties in the boundary of the hybridization signal, the resolution of the method prevented the unambiguous localization of the sGNC subunits in the Purkinje cells. Electrophysiological studies on the induction of long-term depression (LTD) all take the line of thought in which excitatory input to Purkinje cells activates NOS in parallel fibers and/or basket cells. Although there are some exceptions in the literature (see p. 367), it is generally agreed that there is no NOS in Purkinje neurons, convincingly demonstrated by Crepel et al. (1994). It is a common line of thought that NO serves as the interneuronal messenger and activates sGNC in Purkinje cells (Daniel et al., 1993; Lev-Ram et al., 1995; Boxall and Garthwaite, 1996). Nevertheless, the observations that (a) bath application of the selective inhibitor of sGNC, ODQ (Garthwaite et al., 1995), or (b) a selective inhibitor of the cGMP-specific PDE-5 isoform, are interventions for inducing LTD cannot be taken as evidence for the localization of sGNC in Purkinje cells (Hartell, 1994; Linden et al., 1995; Lev-Ram et al., 1997). Even the different time requirements for inducing LTD using the technique of uncaging NO or cGMP in the presence of Ca -,+ can be explained by assuming a relatively slow intracellular diffusion of cGMP from the soma to the dendrites, versus a fast diffusion of NO to parallel fibers followed by extrusion of cGMP at the synapse into the Purkinje cell dendrite (Lev-Ram et al., 1997). Similarly, the application of inhibitors or activators of cGMP-dependent protein kinases (Daniel et al., 1993; Hartell, 1994, 1996) is no evidence for the presence of sGNC in Purkinje cells, but can at best be taken as corroborating evidence for the presence of the two cGMP-protein kinases in Purkinje cells (Lohmann et al., 1981; J. De Vente and S.M. Lohmann, unpubl, results) and a role for cGMP in LTD induction. The effect of intracellular application of ODQ resulted in a blockade of LTD (Boxall and Garthwaite, 1996). However, recently it became apparent that ODQ is less specific than hitherto assumed (Feelisch et al., 1999). Experiments using in vivo microdialysis have shown efflux of cGMP from the cellular constituents of the cerebellum (Vallebuona and Raiteri, 1993; Laitinen et al., 1994; Luo et al., 1994; Fedele et al., 1997; Hermenegildo et al., 1998). The efflux of cGMP closely follows the effects of NO-mediated activation of sGNC, inhibition of phosphodiesterase activity, and manipulation of glutamatergic receptors. Recent schemes (Luo et al., 1994; Fedele and Raiteri, 1999) on the organization of the NO-cGMP signaling in the cerebellum do not take into account that NO-cGMP synthesis in the molecular layer also takes place inside parallel fibers (De Vente et al., 1998a). In conclusion, the available evidence that NO-mediated cGMP synthesis takes place in adult cerebellar Purkinje cells is not particularly overwhelming. Nevertheless, it cannot be excluded that some cGMP synthesis takes place in Purkinje cells and that the cGMP is bound to the protein kinases, which leaves levels of unbound cGMP below the detection limit of cGMP immunocytochemistry. The target-protein-bound cGMP is no longer available for binding the cGMP antibody against formaldehyde-fixed cGMP. There is convincing e~,idence that in the cerebellum there exists a large efflux of cGMP from a cellular compartment. This cGMP may have a function in intercellular communication, as has been suggested previously (Laitinen et al., 1994; Luo et al., 1994). The reports on the electrophysiological effects of bath-applied cGMP (Linden et al., 1995) can be taken as corroborating evidence. In our opinion, the available evidence points to the parallel fibers as an important source for cGMP and NO synthesis (Lev-Ram et al., 1995; Shibuki and Kimura, 1997) and cGMP as well (De Vente et al., 1998a), although it still has to be established that in parallel fibers NO and cGMP 387

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synthesis take place in the same compartment of the same fiber. The cGMP might diffuse or is transported out of the fibers and taken up by Purkinje cells.

7. N O - c G M P SIGNALING IN OTHER VERTEBRATES Using cGMP-IR as a tool to visualize the target structures for NO, N O - c G M P signaling has been investigated in the central nervous system of only a limited number of vertebrates other than rats. Using the slice technique, the relationship between NOS and NO-mediated cGMP synthesis was investigated in the mouse (Mulder et al. in prep.; Fig. 23), the cat (Moreno-Lopez et al., 1998) and in the frog Xenopus laevis (Allaerts et al., 1997, 1998). In all three species it was found that there is a complementary overlap between NOS-IS and cGMP-IR after stimulation with a NO-donor. Nevertheless, there are considerable differences between the localization of NO-mediated cGMP synthesis in the mouse compared to the rat (Fig. 23). In the basal, unstimulated slices, cGMP-IR was found to be present in astrocytes in several brain areas studied, e.g. the cortex, the hippocampus, the hypothalamus, and colliculi. After stimulation of sGNC with SNP, cGMP-IS was intensified in the areas that already showed cGMP-IS in the unstimulated slice, and in addition was found in astrocytes in all cortical areas (Fig. 22a,d), the hippocampus (Fig. 22b,e), the cerebellum (Fig. 22c), and to a minor degree in the caudate putamen complex (Fig. 22f). Comparing these results with cGMP-IS in the rat brain, the neuronal component in NO-mediated cGMP synthesis is, surprisingly, almost absent, whereas mouse astrocytes seemed to contain sGNC more frequently than rat astrocytes, or, as an alternative, appear to be more sensitive to NO. On the other hand it might be that mouse astrocytes express a different number of phosphodiesterases, which might be less sensitive to inhibition by IBMX than the rat astrocyte. This finding may have important implications for the interpretation of the functional involvement of N O - c G M P signal transduction in LTP, LTD, and behavioral paradigms in mice.

Fig. 24. cGMP-IS in islands of Calleja of the rat at PNI0 (a) and in the adult brain (b). Slice was incubated in the presence of 1 mM IBMX only. Bar represents 100 ~tm.

389

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J. De Vente and H.W.M. Steinbusch

Fig. 25. NOS-immunostainingin section from a sagittal slice of a rat brain at PNI0.

8. N O - c G M P SIGNALING IN THE RAT BRAIN DURING DEVELOPMENT During development, from PN0 onwards, the cGMP content in brain regions was found to decline and also the degree of stimulation of cGMP synthesis declined during maturation and aging (Schmidt and Thornberry, 1978: Puri and Volicer, 1981; Burgal et al., 1982; De Vente et al., 1990; De Vente and Steinbusch, 1992; Wei et al., 1993: Vallebuona and Raiteri, 1995; Markerink-Van Ittersum et al., 1997; Prickaerts et al., 1998). This decline may be caused by either a decrease in GNC or an increase in phosphodiesterase activity. An increase during development and aging in cGMP-hydrolyzing phosphodiesterase activity of the guinea-pig brain was noted in the early literature (Smoake et al., 1974; Davis and Kuo, 19761). In prostate and heart a decrease in cAMP phosphodiesterase activity upon aging was noted (Moses et al., 1987). Thus, not surprisingly, there exists a highly tissue-specific regulation of phosphodiesterase activities. This is also more in line with the present knowledge about the different classes of isoenzymes within this family (Beavo. 1995: Houslay, 1998). 390

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Fig. 25 (continued).

There is a considerable number of publications describing increased expression of NOS during embryonal and postnatal development compared to adult, e.g. in the superior colliculus and the ventral lateral geniculate nucleus of the rat (Gonzalez-Hemandez et al., 1993; Tenorio et al., 1995), the rat neocortex (Matsumoto et al., 1993b; Tomic et al., 1994; X.X. Yah et al., 1994), the rat septum (Holzman et al., 1996), rat cerebellum (Matsumoto et al., 1993a; Li et al., 1996), the chick tectum (Williams et al., 1994), and mouse forebrain (Deter and Derer, 1993). There are several reports which described NOS expression during development in a large number of brain tissues from the rat (Bredt and Snyder, 1994; Samama et al., 1995; Keilhoff et al., 1996) and the human (Ohyu and Takashima, 1998). These studies show that an increase as well as a transient increase during embryonic life in NOS expression can be found, corroborating the detailed report on transient NOS expression in the facial and hypoglossal nucleus, and the nucleus ambiguus (Gonzalez-Hernandez et al., 1994). There are data which indicate that upon aging NOS expression declines in all tissues as demonstrated in a large number of brain tissues from rat and mouse (Brien et al., 1995; Mollace et al., 391

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1995; Tsukada et al., 1995; Vaid et al., 1998). We also noted high levels of NOS-IS in the rat brain at PN10 compared to the adult situation (Figs. 25 and 26). An increased number of varicosities of the NOS axonal or dendritic network is a characteristic feature of the NOS-IS in the adult rat, as exemplified in Fig. 26. A parallelism between neurogenetic sequences and the (transient) expression of NOS has been noted (Bredt and Snyder, 1994; Samama et al., 1995). NOS expression coincides with synaptogenesis in a particular brain area (Bredt and Snyder, 1994; Roskams et al., 1994; Keilhoff et al., 1996), either by forming or eliminating synaptic connections (Wu et al., 1994). Similar observations have been made in invertebrates (see Scholz and Truman, Chapter XI). However, this does not imply that NOS is always involved in patterning connections in such an area, as demonstrated by the absence of a role of NOS in the formation of thalamocortical connections, despite the presence of NOS (Finney and Shatz, 1998). On the other hand NO may be involved in the transition from proliferation to differentiation by inducing growth arrest as was demonstrated in PC12 cells (Peunova and Enikolopov, 1995). NO-mediated cGMP synthesis is intense and widespread throughout the rat brain during postnatal development (Fig. 27). NOS activity in the islands of Calleja is higher in the unstimulated slice at PN10 compared to the adult situation (Fig. 24a,b). Very dense NO-mediated cGMP-IS is found in all cortical areas in a dense network of fibers, astrocytes and neurons (Fig. 28a,b). Oligodendrocytes have been found to express an active sGNC transiently during development (Tanaka et al., 1997). Also in the hippocampus a dense network of astrocytic and neuronal fibers of NO-responsive elements is found at PN10, and, although pyramidal cells are often found encircled by cGMP-IR fibers, these cells do not contain cGMP-IR (Fig. 28c). A similar finding is shown in Fig. 21a,b for the spinal cord. All the data suggest that NO-cGMP signaling is important during development in vertebrates. Nevertheless, the finding that in neuronal NOS knock-out mice development is normal (Huang et al., 1993; Burnett et al., 1996; Klein et al., 1998) makes the above statement almost untenable. It has been suggested that a residual amount of NOS, various isoforms of neuronal NOS or endothelial NOS (Olgilvie et al., 1995; Eliasson et al., 1997; Koleshnikov et al., 1997) are sufficient to keep NO-cGMP signaling at a sufficient level (Huang et al., 1993). Nevertheless, prenatal inhibition of NOS by administering L-NAME to pregnant rats at E4, E7, or El4 results in severe hindlimb defects at birth (Diket et al., 1994; Pierce et al., 1995) and similar defects were recently described in endothelial NOS-deficient mice (Gregg et al., 1998). On the other hand it has been shown that postnatal treatment of neonatal pups can reduce NOS activity to 98% or more in the ferret (Finney and Shatz, 1998) and cGMP synthesis can be reduced to low levels, although not to such low levels as NOS activity (Prickaerts et al., 1998), without an appreciable effect at the cellular level or in several behavioral paradigms in the rat (W6rtwein et al., 1997; Prickaerts et al., 1998). Similarly, systemic treatment with L-NAME of rats receiving a fetal mesencephalic transplant in the 6-OH-dopamine lesioned caudate putamen did not affect the survival and outgrowth of fetal dopaminergic cells (Van Muiswinkel et al., 1998), although it could be shown that an active NO and cGMP synthesis was present in the implant. These findings raise the important question at what level of activity NO-cGMP signaling is still functioning optimally, and whether another messenger for sGNC

Fig. 26. Comparison of NOS-immunostaining in rat brain areas at PN 10 (a, c. e) versus adult (b, d,./3. Areas in a, c, and e are enlargements of areas shown in Fig. 23. (a, b) Parietal cortex. (c. d) Colliculus superior. (e, f) Caudate putamen complex. Bar represents 50 Ism for all pictures.

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may be involved that takes over the role of NO, or that there is a second-messenger system that can take over completely the role of N O - c G M P signaling. There is as yet surprisingly little information about the expression of the subunits of sGNC during development. In the rat brain, the c~l subunit was absent at El2, but was however strongly expressed at E14-E15 (Smigrodzki and Levitt, 1996). It has been demonstrated that in the mouse the [31 subunit was present at E19 in the lung and the intestine and was absent or weakly expressed in the brain (Giuili et al., 1994), but strongly expressed at PN3. The presence of 0~1 at E14-E15, coincides with the appearance of NOS expression at that stage (Bredt and Snyder, 1994). At PN4 we found the 131 subunit to be present throughout the rat brain in cell somata and in the neuropil (Fig. 29b,d,f). In addition, we found areas in the brain that contain high levels of NO-mediated cGMP-IR whereas the immunostaining for the 131 subunit is absent or very weak, e.g. layers 1 and 2 of the cortex (Fig. 29a,b), the islands of Calleja (Fig. 27c,d), and the septal area (Fig. 29e,f). In the caudate putamen at PN8 we 394

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Fig. 27 (continued).

found cells which were cGMP-IR but did not immunostain for the [31 subunit, whereas other cells were immunopositive for [31 but not for cGMR in addition to cells which showed both markers as expected (Fig. 29e,f). These experiments show that the presence of the [31 subunit in a cell does not imply that this cell will synthesize cGMP in response to NO and, taking the absence of staining for the [31 subunit as evidence that the subunit is not expressed, that other subunits of the sGNC might exist.

9. COLOCALIZATION OF cGMP WITH NEUROTRANSMITTER SYSTEMS IN THE RAT BRAIN Until recently there was little information on the localization of cGMP in relation to neurotransmitters in the CNS like acetylcholine, dopamine, serotonin, GABA, noradrenaline, 395

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Fig. 28. cGMP-IS in rat brain areas at PNI0. Depicted are enlargements from the section shown in Fig. 25. (a) Parietal cortex. (b) Orbital cortex and anterior olfactory nucleus. (c) Hippocampus. Abbreviations: Pa = parietal cortex; VO = ventral orbital cortex; AOM = medial anterior olfactory nucleus: slm = stratum lacunosum moleculare. Bar represents 50 g m for all pictures.

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and glutamate. Using Steiner-type cGMP antisera, Ariano et al. described the localization of cGMP in the caudate putamen and substantia nigra of the rat. cGMP was found in glial cells, neurons and a punctate-stained neuropil (Ariano et al., 1980; Ariano and Matus, 1981) in the caudate putamen, whereas a large proportion of the striatonigral neurons were immunopositive for cGMP (Ariano and Ufkes, 1983). It was subsequently suggested that cGMP synthesis in the rat caudate putamen occurred in medium spiny neurons, probably enkephalinergic or GABAergic (Ariano, 1983). In addition, it was shown that cGMP-containing neurons in the caudate putamen contained muscarinic cholinergic receptors (Ariano, 1985). However, we have shown that cGMP accumulation takes place in varicose fibers and a conspicuous absence of staining was noted in cell somata (Fig. 3a). Recently, we were able to show that this cGMP-IS in the caudate putamen colocalizes to a large extent with the neuronal acetylcholine transporter (De Vente et al., 2000). No colocalization was observed with dopamine, serotonin, GAD, parvalbumin, or the neuronal glutamate transporter. Another striking observation is the absence of cGMP-IS in glial cells in the caudate putamen, which is contrary to the findings of Ariano et al. (see above). A significant difficulty in comparing these different results is the use of different types of antisera, i.e. Steiner-type antibodies versus antisera against formaldehyde-fixed cGMP. As discussed above (see Section 4), different pools of cGMP may be visualized. In addition, extensive colocalization was observed between cGMP-IR and VAch in all cortical areas (e.g. the frontal cortex, see Fig. 30a,b). In other rat brain areas, little colocalization was observed between cGMP-IR and VAch, e.g. in the hippocampus (Fig. 30c,d). Thus, cholinergic fibers in the cortex and caudate putamen present an important target for NO, although it is not yet clear what the function of NO-mediated cGMP synthesis is in these cholinergic fibers. A partial colocalization was noted between cGMP-IS and the acetylcholine transporter in the rat spinal cord (Vles et al., 2000). Taking parvalbumin as a partial marker for GABAergic neurons, partial colocalization of cGMP-IS with parvalbumin was demonstrated in the rat cortex (De Vente et al., 2000). Thus, little parvalbumin was found in layers I and II of the cingulate cortex; however, in the deeper cortical layers abundant parvalbumin was visualized with little or no colocalization with cGMP-IR (Fig. 31a,b). An exception was the indiseum griseum, where an extensive colocalization between these two markers was observed (Fig. 3 l a,b). Partial colocalization was observed between parvalbumin in a larger number of other brain areas, e.g. hippocampus (Fig. 3 l c,d), reticular thalamic nucleus, colliculi, the cerebellum (Figs. 34a,b and 35a-d), and the spinal cord (Vles et al., 2000). The still existing uncertainties associated with the cGMP-immunocytochemistry, with respect to detection limit and the role of phosphodiesterases, became all the more evident when rat brain slices were incubated in the presence of 10 ~tM YC-1 in combination with 10 ~M SNP. YC-1 is a novel stimulator of sGNC (Ko et al., 1994; Friebe et al., 1996, 1998; Stone and Marietta, 1998). YC-1 sensitizes sGNC through a not yet completely understood mechanism to become highly responsive to CO and NO. In addition, YC-1 has been shown to exhibit phosphodiesterase inhibiting properties (Friebe et al., 1998; Galle et al., 1999). Although YC-1 strongly potentiated the effect of NO on cGMP synthesis in brain tissue, the phosphodiesterase inhibiting potency of YC-1 was found to be weak in this tissue (De Vente et al., 1999; W. Van Staveren et al. in prep.). The intriguing properties of YC-1 on sGNC raise the question whether endogenous compounds exist which regulate sGNC in a similar fashion. cGMP synthesis was potentiated considerably in the presence of 10 ~tM YC-1 compared to the effect of 10 ~M SNP only, and cGMP-IR was observed in a fiber network which was clearly more dense than that observed after stimulation with 100 ~tM SNP only. In 400

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addition, more cell somata were observed which showed cGMP-IS. In cortical areas the colocalization between cGMP-IR and parvalbumin was observed more frequently when slices were stimulated in the presence of YC-1 (Fig. 32a,b), and in layers 1 and 2 of the cortex the number of the cGMP-IR fibers appeared to be increased (compare Fig. lb or Fig. 3 la with

Fig. 33. Double immunostaining of cGMP (a) and parvalbumnin (h) in the same area of the same section of the hippocampus. Arrowheads point to astrocytes v
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Fig. 32a). We observed colocalization between cGMP-IS and parvalbumin in the hippocampus in slices stimulated with the combination of YC-1 and SNP, especially in the fibers running through and surrounding the pyramidal and granule cells (Fig. 33). Extensive colocalization between parvalbumin and cGMP-IR was observed in cell somata in the reticular thalamic nucleus (Fig. 34) after stimulation of sGNC with SNP in the presence of YC-1. It has been proposed frequently that NO acts as a retrograde messenger after activation of NOS through excitatory amino acid receptors (e.g. Garthwaite, 1991; O'Dell et al., 1991; Hawkins et al., 1994; Garthwaite and Boulton, 1995; Aoki et al., 1997; Zhuo et al., 1998). Therefore, it was expected to find a colocalization between cGMP-IR and the neuronal glutamate transporter molecule (EAAT1) (which may also be found on astrocytes). However, only in a few instances colocalization between cGMP-IR and EAAT1 has been found, e.g. in astrocytes in the corpus callosum (not shown) and in granule cells in the cerebellum (Fig. 35e,f). Especially in the hippocampus and the molecular layer of the cerebellum, a colocalization between cGMP-IR and EAAT1 could not be observed. In view of the uncertainties associated with cGMP immunocytochemistry in relation to activation of sGNC and the role of phosphodiesterases, the present data on colocalization between cGMP-IR and other neurotransmitter systems can only be viewed as a first approximation.

10. Abbreviations cGMP-IS sGNC pGNC IBMX L-NAME eNOS nNOS NOS-IS SNP

cGMP-immunostaining soluble guanylyl cyclase particulate guanylyl cyclase isobutylmethylxanthine N°~-nitro-L-arginine methyl ester endothelial nitric oxide synthase neuronal nitric oxide synthase nitric oxide synthase immunostaining sodium nitroprusside

11. A C K N O W L E D G E M E N T S We thank Prof. David Hopkins (Halifax) for discussions. We are obligated to Jeffrey Rothstein (Bethesda), Piers Emson (Babraham), Prof. H.H.H.W. Schmidt (Wtirzburg), Prof. D. Koesling (Berlin), and the late Dr. Ichikawa for the gift of antibodies. Hans Vles is acknowledged for Fig. 20. Our special thanks are for Marjanne Markerink-Van Ittersum for her continuous enthusiastic support.

Fig. 35. cGMP-IS in the cerebellum of the adult rat: (a, b) double immunostaining of parvalbumin (a) and cGMP-IR (b) in the same area of the same section of a slice incubated in the presence of 10 IxM SNP and 1 mM IBMX; (c, d) double immunostaining of parvalbumin (c) and cGMP-IR (d) in the same area of the same section of a slice incubated in the presence of 100 ~M YC-1, 10 IsM SNP, and 1 mM IBMX; (e, f) double immunostaining of neuronal glutamate transporter (EAAT) (e) and cGMP-IR ~ in the same area of the same section of a slice incubated in the presence of 10 gM SNP. Bar represents 50 ~m for all pictures.

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CHAPTER XI

Invertebrate models for studying NO-mediated signaling N.L. SCHOLZ AND J.W. TRUMAN

1. INTRODUCTION For more than half a century, research on invertebrate nervous systems has made important contributions to our general understanding of how individual neurons and neural networks give rise to animal behavior. The large number of invertebrate phyla represent a rich diversity of body forms and behavior. Correspondingly, invertebrate nervous systems range from the simple nerve nets of cnidarian jellyfish to the elaborate brains of cephalopod molluscs (squid and octopus), which rival their fish competitors in terms of total numbers of neurons and complex network organization. The nervous systems of invertebrates offer the twin advantages of numerical simplicity and the presence of comparatively large and identifiable nerve cells. These 'identified neurons' can be reliably located in different animals, which makes it possible to analyze the specific function of a given neuron in differing developmental and physiological contexts. This approach has been a valuable tool in the study of the cellular aspects of nervous system function and development, and it will be a common theme in the discussion of NO-mediated signaling in invertebrates that follows. NO is an ancient signaling molecule, presumably evolving before the radiation of the metazoans (Feelish and Martin, 1995). There is remarkable conservation of function for NO-mediated signaling, even among organisms that belong to different kingdoms. For example, NO plays a key role in disease resistance in both plants (Delledonne et al.. 1998) and animals (Nathan, 1995). Comparative studies have found NO-producing neurons in animals from all the major phyla, suggesting that cellular signaling using NO is an ancestral character of both vertebrate and invertebrate nervous systems. For example. the NO/cGMP pathway has been identified in the coelenterate HTdru i~ilgaris,the most primitive organism known to possess a nervous system (Colasanti et al., 1995). In mammals, the NO signaling pathway was first identified in the search for the agent that caused the relaxation of vascular smooth muscle (Palmer et al., 1987). It was subsequently discovered that NO is also an important anterograde as well as retrograde intercellular signaling molecule in the mammalian brain (Bredt and Snyder, 1992; Garthwaite and Boulton, 1995). Comparative studies soon found nitric oxide synthase (NOS) enzymatic activity in molluscs (Elofsson et al., 1993; Gelperin, 1994) and arthropods (Radomski et al., 1991: Elphick et al., 1993; Ribeiro et al., 1993; Johansson and Carlberg. 1994). In parallel with mammalian studies, initial research in invertebrates focused on roles for NO in olfaction (Gelperin, 1994; Muller and Bicker, 1994) and learning and memory (Robertson et a].. 1994).

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Research has expanded in recent years and widespread roles for NO in cell-cell signaling are now established for most of the major phyla. Moreover, specific roles lbr NO have been extended into other aspects of nervous system function and into development. Most of the work continues to focus on molluscs and arthropods, and comprehensive reviews by Jacklet (1997), MOiler (1997), and Bicker (1998) have recently surveyed the function of NO in these animals. We will not try to repeat these reviews here. Rather, we will consider emerging research that is providing mechanistic insights into the roles of NO in the physiology and development of the nervous system.

2. COMPONENTS OF THE NO SIGNALING PATHWAY IN INVERTEBRATES

Nitric oxide is produced when NOS converts arginine into citrulline and NO. A widely used tool in assessing the presence of NOS in invertebrates has been the distribution of fixation-insensitive, NADPH-diaphorase staining. This histochemical procedure identified NOS activity in vertebrate systems (Bredt et al.. 1991" Hope et al., 1991). In invertebrate systems diaphorase activity was subsequently shown to copurify with NOS in both bees and Drosophila (Mtiller, 1994). However, considering the great diversity in the invertebrates, patterns of diaphorase staining should be taken as an indication, not a proof, of the presence of NOS. There may be forms of NOS that do not show up by diaphorase staining, and likewise, some diaphorase staining may not be due to NOS. Despite these cautions, however, diaphorase staining has been a valuable tool in establishing the range of NOS distributions in animals. The biochemical studies on the brains of locusts (Elphick et al., 1993) and of the marine molluscs Pleurobranchia and Aplysia (Moroz et al., 1996) demonstrate NOS enzymatic activity that shows the requirements for 6-NADPH and calmodulin that characterize some mammalian NOSs. The large size of invertebrate neurons, especially those from molluscs, coupled with capillary electrophoresis, has subsequently allowed the detailed examination of NO metabolism from single, identified neurons (Moroz et al., 1999). A gene for NOS was isolated from Drosophila (Regulski and Tully, 1995). It has 43% sequence identity with rat neuronal NOS and contains the putative FMN, FAD, NADPH, heine and calmodulin binding sites that are general features of constitutive NOS isoforms. NOS has since been cloned from other insects (the moth Mand, ca sexta, Nighorn et al., 1998; the grasshopper Locusta, Ogunshola et al., 1995) and from the snail Lym,aea stagnalis (Korneev et al., 1998). In vertebrates, NO acts on target cells via multiple pathways. The best characterized pathway is through activation of and the production of cGMP (Schuman and Madison, 1994; Garthwaite and Boulton, 1995). More recently, production of poly-ADP-ribose has been shown to mediate some NO actions (Zhang et al., 1994) as have redox pathways involving nitrosylation of cellular proteins (Stamler et al., 1997). Actions through the latter two pathways are still largely unexplored in invertebrates, and the only target of NO signaling that has been well characterized are the soluble guanylate cyclases (sGCs). Both Drosophila and the moth Mand, ca have homologs of sGCal and sGC[31. These function as an obligate heterodimer that is stimulated by NO (Shah and Hyde, 1995; Nighorn et al., 1998). In addition to the NO stimulatable sGCs, both the lobster (Prabhakar et al., 1997) and Mand,ca (Nighorn et al., 1999) have an additional cytoplasmic cyclase that is not NO-sensitive. In Mand, ca this is a novel 6 subunit, MsGCI33, that has little or no NO sensitivity and which needs not form a heterodimer for activity. The MsGC[33 lacks some of the residues thought to be needed for heme binding and it is insensitive to the sGC inhibitor 418

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1H-[1,2,4]oxadiazolo[4,3-0~]quinoxalin-l-one (ODQ; Garthwaite et al., 1995). The function of this novel cyclase has not been established but it may be involved in a peptide signaling pathway that involves the peptide eclosion hormone (Morton, 1997).

3. DEVELOPMENTAL ROLES FOR NO/cGMP SIGNALING IN INVERTEBRATES The developmental role of the NO/cGMP signaling pathway has been studied primarily in molluscs and arthropods. Developmental studies in molluscs are in their early phases and represented by reports of changes in NADPH-diaphorase histochemistry during embryonic and metamorphic development of the gastropods llvanassa obsoleta (Lin and Leise, 1996) and Lvmnaea stagnalis (Serfozo et al., 1998). More extensive data are available for the arthropods, and these animals are the main focus of this section. 3.1. NO SIGNALING AND PROLIFERATION CONTROL The role of NO in the control of proliferation is an emerging area of developmental research in invertebrates. The first evidence that NO regulates cell division came from the study of the growth of imaginal discs in Dpvsophila (Kuzin et al., 1996). These structures grow in the larva to give rise to adult structures such as the legs, wings, genitalia, etc. NADPH-diaphorase staining indicated that NOS activity appears in imaginal discs early in the last larval stage, as these structures are entering their rapid phase of growth. Pharmacological inhibition of NOS in larvae resulted in excessive growth of the discs and hypertrophied structures in the adult. By contrast, the ectopic expression of NOS resulted in undergrown structures. These findings suggest that NO acts as an antiproliferative agent during Drosophila development. A role for NO in proliferation control is also evident during neurogenesis in the brain outer proliferation zone (OPZ) in the tobacco hornworm moth Manduca sexta. This zone is a band of proliferating stem cells that produce the interneurons of the outer two layers of the optic lobe (the lamina and medulla). Early in metamorphosis, proliferation in the OPZ requires the presence of the steroid hormone, 20 hydroxyecdysone (20E) (Champlin and Truman, 1998). Importantly, cells in the zone respond to steroid in concert, such that all of the cells are either cycling or quiescent depending on steroid concentration. This coordinated proliferation suggests that there must be a non-cell-autonomous aspect to the ecdysteroid response that allows the system to respond as a unit. This coordination appears to be established via NO signaling (D. Champlin and J. Truman, unpubl, data). The OPZ of Manduca shows intense NADPH-diaphorase staining at the beginning of metamorphosis. In addition, cells in the proliferation zone stain with an antibody directed against a conserved region of NOS. The possibility that NO coordinates mitotic activity within the proliferation zone was tested in vitro using brains cultured in subthreshold levels of 20E (20 ng/ml), which arrested their proliferation. Brains w~'re then shifted to threshold levels of 20E (60 ng 20E/ml) with or without the NOS inhibitor, L-NAME. In the absence of L-NAME, the proliferation zone showed the expected all-or-none response to the steroid. In the presence of L-NAME, by contrast, the proliferation zone responded asynchronously, with a scattering of neuroblasts cycling independently within each optic lobe (D. Champlin and J. Truman, unpubl, data). Importantly, proliferation was still steroid-dependent in the presence of L-NAME, but L-NAME reduced the threshold concentration at which the steroid could initiate scattered proliferation. These results argue 419

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that the NO diffusion serves to coordinate cell division within the proliferation zone and that it does so through a suppression of steroid-induced proliferation. Interestingly, in both Drosophila and Manduca NO may not act via the production of cGMP to regulate cell proliferation. For the imaginal discs of Drosophila, the cells do not produce detectable cGMP in response to NO donors as measured using cGMP immunocytochemistry (Wildemann and Bicker, 1999a). The same is true for the neuroblasts in the OPZ in Manduca (D. Champlin and J. Truman, unpubl, data). Moreover, in the latter, the inhibition of sGC activity using 1H-[1,2,4]oxadiazolo[4,3-~]quinoxalin-l-one (ODQ; Garthwaite et al., 1995) had no effect on coordinating proliferation within the zone. Therefore, NO may be acting through another pathway, such as protein nitrosylation (Stamler et al., 1997). 3.2. NO SENSITIVITY DURING NEURONAL DEVELOPMENT After their birth, neurons progress through a number of discrete developmental phases, starting with axonal outgrowth and growth cone navigation, progressing through target recognition and branching over its target, and ending with synaptogenesis. Studies on a number of developing arthropods, including embryonic grasshoppers (Truman et al., 1996; Ball and Truman, 1998), Drosophila (Wildemann and Bicker, 1999a), and Manduca (Grueber and Truman, 1999) as well as metamorphic lobsters (Scholz et al., 1998), moths (Schachtner et al., 1998), and flies (Gibbs et al., 2000) have shown that many neurons exhibit a transient phase of NO sensitivity during their development. This sensitivity is defined by the ability of the neuron to produce levels of cGMP, that can be detected by immunocytochemistry, in response to a challenge with an NO donor in the presence of IBMX. The relationship between the appearance of NO sensitivity and the progression of neuronal development has been precisely established for two systems: developing motor neurons in the embryonic nervous system of grasshoppers (Truman et al., 1996; Ball and Truman, 1998) and the developing photoreceptors in the visual system of Drosophila (Gibbs and Truman, 1998). Grasshopper embryos have been a model system for studying neurodevelopment for almost 25 years, and the early development of many identified neurons is known in great detail (e.g., Goodman et al., 1984). As seen in Fig. 1, filleted embryos exposed to an NO donor and IBMX show a dramatic production of cGMP throughout their peripheral and central nervous systems (Truman et al., 1996; Ball and Truman, 1998). These embryos are especially convenient for studying early aspects of neuronal development because abdominal segments are produced sequentially, with about a 6-h delay between successive segments. Consequently, in a single embryo one can examine the properties of a neuron at progressively earlier times in development by examining cells in more posterior segments. As seen in Fig. 2, for particular identified motor neurons, such as RP2 and aCC, the onset of NO responsiveness is relatively abrupt, appearing over the space of a few hours. At the time the cell first becomes

Fig. 1. Photomicrograph of a filleted grasshopper embryo at 57c~ of embryonic development, showing cGMP immunoreactivity that was induced by incubation with a NO donor and an inhibitor of phosphodiesterase (IBMX). The embryo was cut down the dorsal midline, gutted, and the body wall pinned out to provide maximal access to the nervous system by bathing chemicals, cGMP production was confined to neurons in the central and peripheral nervous systems. Lines and letters down the right of the figure indicate the boundaries between the subesophageal (S), thoracic (T), and abdominal (A) regions of the embryo. Inset shows a higher magnification view of the cell bodies and processes of neurons that are responding in the fourth abdominal ganglion. Scale bar = 300 mm for main figure. Modified from Ball and Truman. 1998. 420

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NO-sensitive, it has the form of a 'balloon on a string', having a cell body and long axon but with little or no central or peripheral branches. Dendritic branches begin to be elaborated soon after the cell becomes NO-sensitive (Fig. 2).

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Fig. 2. Drawings showing the changes in the central arbor of grasshopper motor neurons RP2 and aCC after they become NO-responsive. The abdominal segments (A I-A6) form sequentially with an approximate 6 h delay between segments. Therefore, the most posterior neurons are developmentally the youngest. When the cells first respond to NO, they already have an axon that extends to their target muscle but only a very weak cGMP response is observed in the cell body. Cells that are 6 h older in development show complete staining of their axon and initial process but these are relatively unadorned. Branches and filopodia begin to extend soon thereafter. The drawing of the axon is truncated in all cases. From Ball and Truman (1998).

The relationship between pre- and post-synaptic cells during the time of NO sensitivity can be best appreciated in the case of motor neurons at the developing neuromuscular junction. This is illustrated for neuron VML1 and its target, muscle 187 (Fig. 3). Detectable NO sensitivity first appears when the motor neuron's peripheral axonal growth cone is on its target muscle but still has a compact shape and lacks branches (Fig. 3A). Shortly thereafter, the growth cone begins to branch and spreads over the developing muscle target (Fig. 3 B - E ) . The photoreceptors in the developing eye of Drosophila also show an abrupt onset of NO sensitivity. The birth of photoreceptor neurons starts midway through the last larval stage as a morphogenetic furrow moves across the developing eye imaginal disc (Wolff and Ready, 1993). The photoreceptor axons grow into the optic lobe towards targets in the forming lamina or medulla. When they reach these targets, they then arrest their development until the onset of metamorphosis. Soon after the start of metamorphosis the photoreceptors terminate their arrest and their growth cones begin to move laterally within their neuropilar layer to make contact with the interneurons that will make up the lamina 422

Invertebrate models for studying NO-mediated signaling

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Fig. 3. Photomicrographs of the right ventral body wall in segment A4 from progressively older grasshopper embryos showing the spread of the axonal arbor of neuron VML1 over its target muscle (number 187). The arbor was revealed by its cGMP production after the filleted embryo was exposed to a NO donor and IBMX. The period spans the interval from the beginning of the peripheral cGMP response in segment A4 at about 50c~ of embryonic development (% E) (A) to the fadeout of cGMP expression at approximately 90c~ E (I). A-E span from 50 to 60% of embryonic development, whereas F-I span from about 65c~ to 90c~ E. Two prominent branch points are indicated in successive frames by the asterisk and the arrow. The body wall and muscle grow through this period so the branch point indicated by the asterisk is out of the image alter part (E). Scale bar = 25 mm in A, 50 mm in 1. From Ball and Truman (1998).

c a r t r i d g e s a n d the m e d u l l a c o l u m n s ( M e i n e r t z h a g e n a n d H a n s o n ,

1993). It is at this p o i n t

that the a x o n s o f the p h o t o r e c e p t o r s b e c o m e r e s p o n s i v e to N O (Fig. 4; G i b b s a n d T r u m a n , 1998). 423

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Fig. 4. Photomicrographs of whole mounts of Drosophila brains that have been immunostained for cyclic GMP after treatment with an NO donor in the presence of IBMX. The axons of the developing photoreceptors are not yet responsive to NO donors at A 10 h after puparium formation (APF) but are so by B 16 h APF (arrows). Many neurons in the medial region of the brain are already NO-responsive by 10 h APE E -- photoreceptors clusters in the developing compound eye; OL = optic lobes. Modified from Gibbs and Truman, 1998.

A number of points can be made by a comparison of the grasshopper and fly examples. NO sensitivity is not evident in the neurons as they are undergoing axonal extension and navigation. It appears abruptly, though, when they begin to interact with their synaptic targets, a situation that might require an active dialog between pre- and post-synaptic cells. Although the data are incomplete, a similar relationship appears to hold for a variety of neuronal types (e.g., sensory neurons and interneurons) in a number of organisms (Truman et al., 1996; Schachtner et al., 1998; Scholz et al., 1998). Interestingly, for interneurons that do not have an obvious axon, the cell becomes NO-sensitive after it produces a 'basic skeleton' of its arbor but before it makes extensive branches and elaborations (Ball and Truman, 1998). In parallel with the situation found in motor neurons and sensory neurons, this initial 'skeleton' may be a product of pathfinding mechanisms while the higher-order branching would then be established through an interactive dialog with synaptic partners. The latter would involve NO signaling. The timing of appearance of NO sensitivity in a circuit may provide an anatomical marker for the onset of the interactions between per- and postsynaptic targets. For example, in the developing visual system of Drosophila, the first cells to become NO-responsive are the photoreceptors and the interneurons in the last synaptic neuropil, the lobula. Interneurons in the intermediate visual layers, the lamina and medulla, showed NO sensitivity about 20 to 30 h later. This pattern suggests that wiring first starts from the input and output portions of this brain region and the intermediate area is filled in later (Gibbs and Truman, 1998). A similar relationship is evident in the developing antennal system in the moth, Manduca sexta. A subset of antennal lobe interneurons become NO-responsive at the time of major synapse formation within the antennal lobe (Schachtner et al., 1998, 1999). In the larval lobster, one finds that NOS is upregulated in the accessory lobes of the brain just before the animal undergoes metamorphosis (Fig. 5; Scholz et al., 1998). NOS staining is most intense in the developing cortex and glomeruli, which presumably reflects a role for NO in the progressive wiring of synaptic contacts between deuterocerebral interneurons and projection neurons. The signals that cause a developing neuron to become NO-responsive are poorly understood. Target contact may be necessary for sensitivity to occur but this alone appears not to be sufficient (Ball and Truman, 1998: Gibbs and Truman, 1998). For example, in the fly visual system the first-born photoreceptors grow into the lamina and medulla regions but then wait for about 24 h before the appropriate hormonal signals, provided by 20E, cause them 424

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Fig. 5. Developmental upregulation of NOS in the brain of a lobster just before metamorphosis. Panels Al-A3 represent single confocal sections, dorsal to ventral, through the deuterocerebrum. Panels Bl and B2 are highermagnification images corresponding to A2 and A~, respectively. Anti-NOS immunoreactivity becomes intense in the developing glomeruli of the accessory lobes arrowhead in A l and Bi at this stage, a time at which synaptic connections are presumably forming between midbrain interneurons and projection neurons. Note that NOS is expressed in both the cortex (arrowheads in Bi) and the glomeruli (B2). NOS label is also present in the deuterocerebral commissure neuropil (arrows in Al). Scale bar = 100 mm in A. 25 mm in B. From Schoiz et al., 1998.

to become NO-sensitive (Gibbs and Truman, 1998). For the developing antennal system in

Manduca, the steroid hormone 20E is also important for shifting developing neurons into an NO-sensitive state. The premature exposure of neurons to high levels of 20E results in the early appearance of NO sensitivity (Schachtner et al., 1999). For both the eye and antennal systems, though, it is not known if steroid exposure alone is sufficient or if the cells need both steroid and contact with potential targets. Although NO sensitivity is a dominant theme for developing neurons in the insect CNS, it is not a feature of every neuron. In the developing antennal lobe of Manduca, for example, only a subset of the antennal interneurons show NO sensitivity during metamorphosis (Schachtner et al., 1999). Also, in grasshopper embryos, many of the body wall muscles receive multiple innervation, but only one or two of the neurons projecting to a given muscle become NO-sensitive (Ball and Truman, 1998). The order by which axons arrive at their targets does not appear to determine NO sensitivity. It may relate instead to the types of synapses that are to be made. For example, among the motor neurons that innervate the extensor tibiae muscle in grasshoppers, the slow extensor tibiae motoneuron (SETi) becomes NO-responsive during its development whereas the fast extensor tibiae motoneuron (FETi) does not (Ball and Truman, 1998). Likewise, in Drosophila very few of the motor neurons are NO-responsive during embryonic development but they become so later when they innervate muscles in the adult (Wildemann and Bicker, 1999a; Gibbs et al., 2000). 425

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Fig. 6. Nuclear localization of anti-cGMP immunoreactivity in two different neurons in the ventral nerve cord of a lobster at metamorphosis. Scale bar = 25 mm in B, 10 mm in A. From Scholz et al., 1998.

For many neurons, NO sensitivity is a transient feature of their development. It occurs while synaptic sites are being selected but then wanes as synapses mature (Fig. 3; Truman et al., 1996; Ball and Truman, 1998: Gibbs and Truman, 1998). For many of these developing neurons, their cGMP distribution after a challenge with NO is unusual in that it includes a prominent accumulation of cGMP within the nucleus (e.g., Fig. 6) (Truman et al., 1996; Scholz et al., 1998). This nuclear targeting of cGMP may be a feature of a developmental, rather than a physiological function of NO/cGMP signaling. This conclusion comes from the examination in grasshopper embryos of neurons, like the 'H' cell, that are NO-sensitive during their early development but then continues to be NO-responsive as a mature neuron. Treatment of the immature cell results in accumulation of cGMP in the nucleus whereas in the mature cell cGMP is excluded from the nucleus and is associated with cytoplasmic or cell membrane sites (Truman et al., 1996). In invertebrates, it is not known which potential cGMP target is responsible for the nuclear localization of cGMP. One of the mammalian cGMP-dependent protein kinases translocates to the nucleus after binding cGMP (Gudi et al., 1997). A similar kinase may play a role in developing invertebrate neurons and be responsible for the nuclear cGMP that is observed. This possibility is exciting because it provides a direct intercellular signaling pathway between a neuron and the nucleus of its target cell. As described below, some neurons may continue to be NO-sensitive during post-embryonic life because NO signaling constitutes part of their mature physiology. In other cases, though, it may reflect a continuing developmental plasticity. The latter is thought to be the case for bristle sensory neurons in the grasshopper. During embryogenesis, these sensory neurons show a window of NO sensitivity after their axons enter the CNS. This sensitivity transiently reoccurs, though, as the insect subsequently undergoes each of its nymphal molts (Truman et al., 1996). A possible reason for this reoccurrence comes from studies on the related cricket, in which bristle afferents are known to shift their post-synaptic targets as the animal grows and this changing of targets involves retrograde signaling (Murphey and Davis, 1994). Similar shifts probably also occur in growing grasshoppers and these periods of NO sensitivity may show when the retrograde signaling is taking place to allow these changes. In grasshoppers, NO sensitivity of motor neurons is essentially lost at hatching. However, in larval stages of both Manduca (Truman, unpubl, data) and Drosophila (Wildemann and

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Bicker, 1999a) motor neurons continue to show persistent NO sensitivity as the larva grows. The larvae of both species grow rapidly, a process accompanied by the rapid enlargement of body wall muscles. Post-embryonic NO sensitivity in these animals may be involved with a need to adjust synaptic strengths as the target muscle grows. The proposal of NO signaling at the larval neuromuscular junction is supported by the experiments in Drosophila, which show that both NO and membrane-permeant cGMP can induce vesicle release at the neuromuscular junction (Wildemann and Bicker, 1999b). This section has focussed primarily on neurons that can show cGMP production in response to NO donors. In many of the systems, though, the nature of the normal signaling molecule is not known. It is likely NO in the case of the Drosophila visual system, since the optic neuropils exhibit both NADPH-diaphorase staining and NOS-like immunoreactivity (Gibbs and Truman, 1998). The diaphorase activity in the optic lobes appears to be dependent on the presence of the retinal axons (Atkinson and Panni, 1999). Also, as described below, the inhibition of NO synthesis, or the presence of NO scavengers, disrupts neuronal projections (Gibbs and Truman, 1998). For the developing neuromuscular system, though, the nature of the signaling molecule is unclear. There is no diaphorase activity associated with developing or growing muscle in either grasshopper or Dtvsophila (Truman et al., 1996; Wildemann and Bicker, 1999a). Possibly, this tissue may contain a form of NOS that lacks fixation-insensitive diaphorase activity. Alternatively, another signaling molecule, such as CO, may be involved. 3.3. FUNCTIONAL SIGNIFICANCE OF NO/cGMP SIGNALING The correlation of NO sensitivity with a particular phase of neuronal development suggests that NO/cGMP signaling is required for some aspect of synapse formation. The first experimental test of such a function in invertebrates was for the development of the visual system in Drosophila. As described above, the photoreceptor axons become NO-responsive at about 10 h after puparium formation (APF), when they begin to seek out contact with the visual interneurons in the lamina and medulla (Meinertzhagen and Hanson, 1993). Also, NOS immunoreactivity is highest in the developing optic ganglia from this time through to the midpoint of metamorphosis (Gibbs and Truman, 1998). The brain/eye imaginal disc complexes from Dtvsophila larvae can be cultured and respond to ecdysteroids by the production of an appropriately layered optic lobe. Pharmacological manipulations of the NO/cGMP pathway including inhibition of NOS activity (L-NAME treatment), the scavenging of NO (using PTIO), and the inhibition of sGC (with ODQ) all caused a disorganization of the retinal projections into the medulla (Gibbs and Truman, 1998). Photoreceptor axons often moved out of their normal synaptic layers and extended into deeper layers of the optic lobe or even into the central brain (Fig. 7). The disruptive effects of L-NAME were not observed when we used the inactive form, D-NAME. Also, the disorganizing effects of k-NAME were rescued by adding a 10-fold excess of arginine. Most interestingly, though, the abnormal development provoked by 1 mM L-NAME was prevented by the addition of 8-bromo-cGMP to the cultures. Thus, the developmental effects of blocking NO production can be rescued by a proposed, down-stream effector. This strongly supports the interaction of NO and cGMP in visual system development. The effects of the NOS inhibitors provide some insight into a role for NO/cGMP signaling in neuronal development. NO sensitivity appears at the time when the growth cone is switching its behavior from responding to cues important for pathfinding to interactive associations with potential synaptic partners. When NO signaling is blocked, the growth cone 427

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Fig. 7. Photomicrographs of the left optic lobe from brain-eye complexes that were cultured in various concentrations of the NOS inhibitor, L-NAME, and then immunostained with an antibody against chaoptin, which is an antigen expressed on photoreceptor axons. The staining shows the projection of photoreceptor axons into the lamina (1) and the medulla (m). Numbers refer to control complexes (0). and complexes that show progressively greater disruption of the projection pattern due to the presence of the NOS inhibitor (1 to 4). From Gibbs and Truman, 1998.

resumes extension into deeper brain layers rather than branching in its appropriate synaptic layer. Hence, the local action of NO at this time may serve to maintain the growth cone in the appropriate neighborhood as it selects synaptic partners. After these contacts are made and synaptogenesis has begun, such an arresting agent would no longer be necessary. This relationship is especially interesting considering recent work with cultured neurons showing that cyclic nucleotides change the manner by which growth cones respond to signaling molecules (Song et al., 1998).

4. THE ROLE OF NO IN FEEDING BEHAVIOR

Recent mechanistic studies have found that NO has certain conserved functions, even among animals that are only distantly related. A prominent example is the selective role of NO in feeding. This topic has been reviewed by Jacklet (1997), and we will only briefly consider recent studies linking NO to feeding-related behaviors in animals from different phyla. Nitric oxide mediates the chemosensory-induced activation of feeding behavior in 14ydra (Colasanti et al., 1997), the most primitive animal known to possess a nervous system. H~'dra express a calcium-dependent but calmodulin-independent NOS. When a feeding tentacle comes in contact with a prey item, transmitter-evoked calcium influx is thought to trigger local NO synthesis (Colasanti et al., 1997). NO is apparently involved in triggering the discharge of some of the nematocytes (Salleo et al., 1996). It also rapidly diffuses to neighboring tentacles thereby recruiting these appendages into the feeding response. An NO-sensitive guanylate cyclase is also expressed in this primitive organism (Colasanti et al., 1995) and NO-activated cGMP resets the system by inhibiting the feeding response on longer timescales. Thus, the selective association between NO/cGMP signaling and chemosensory function (e.g., Breer and Shepherd, 1993) appears to have been conserved throughout nervous system evolution. NO has also been shown to trigger feeding behaviors in molluscs and crustaceans. In the gastropod mollusc Lymnaea, NO is an anterograde transmitter between chemoreceptor neurons in the lip and the feeding motor network in the central nervous system (Elphick et al., 1995). NO release within the nervous system triggers feeding movements of the buccal mass (Moroz et al., 1993; Elphick et al., 1995). Similarly, in the crab Cancer productus the release of NO into the stomatogastric nervous system underlies the dynamic assembly of a feeding 428

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motor network (Scholz et al., 1997). As in Hydra, NO acts at least in part via the stimulation of a sGC in the feeding networks of molluscs and crustaceans alike. The role of NO in the feeding behaviors of other invertebrates is less well known. Among echinoderms, NOS-like immunoreactivity has been detected in the neurons that innervate the cardiac stomach of the starfish Marthasterias glacialis (Martinez et al., 1994). Starfish feed by everting their stomach, thereby encapsulating and digesting the soft tissues of their prey. Recently, Elphick and Melarange (1998) found that NO relaxes the smooth muscle of the starfish cardiac stomach, indicating that NO may play a role in feeding in these animals as well. Interestingly, the NO-induced relaxation of muscle tone in starfish is similar to NO-mediated non-adrenergic, non-cholinergic (NANC) neurotransmission in the mammalian gut (reviewed by Boeckxstaens and Pelckmans, 1997). Anatomical studies of the feeding systems of other invertebrates have generally revealed an enrichment of NO-synthesizing neurons in the networks associated with food intake or the gut. For example, in the planarian Dugesia tigrina NOS is almost exclusively expressed in the pharynx (Eriksson, 1996). A similar profile was observed for the nematode Ascaris suum (Bascal et al., 1995). Curiously, the leech Hirudo medicinalis seems to be an exception to this rule. Leake and Moroz (1996) found little or no NOS expression in the leech gut, although it was found in other annelids. The authors speculate that this may be an adaptation to episodic feeding on hemoglobin-rich blood. Since hemoglobin is an NO scavenger, the perfusion of the gut with blood would be expected to limit or entirely block NO-mediated neurotransmission.

5. THE ROLE OF NO IN REGULATING RHYTHMIC MOTOR NETWORKS In molluscs and crustaceans, the neurons which drive rhythmic feeding behaviors (e.g., rasping, chewing, and swallowing) are often assembled into central pattern-generating (CPG) networks. The NO/cGMP pathway has been identified in a few of the more extensively studied model systems, and electrophysiological studies have provided considerable insights into the role of NO in the organization of motor behaviors. This section will compare NO-mediated signaling in the feeding circuit in gastropod molluscs to its function in the stomatogastric ganglion of crustaceans. Subsequently, these networks will be contrasted to a non-feeding CPG, the cardiac ganglion which drives the neurogenic heart of crabs. 5.1. THE GASTROPOD FEEDING CIRCUIT NO activates a feeding motor program in the gastropod mollusc Lymnaea stagnalis (Moroz et al., 1993; Elphick et al., 1995), a freshwater pond snail that feeds via the scraping movements of a toothed radula. The behavioral elements involve a simple three-phase activity cycle that includes an extension of the radula, followed by a rasp, and then swallowing (Rose and Benjamin, 1979). These movements are driven by a CPG network, the components of which have been well-studied (reviewed by Benjamin and Elliott, 1989). The CPG consists of a network of interneurons that coordinate the activity of feeding motor neurons in the buccal ganglia. The CPG, in turn, is modulated by central interneurons in the buccal and cerebral ganglia. The feeding motor program can be directly activated by chemosensory inputs from the lips or by stimulating the modulatory interneurons. Many neurons in the buccal ganglion contain NOS, as shown in studies using NADPHdiaphorase histochemistry (Elofsson et al., 1993; Moroz et al., 1993) and anti-NOS immunocytochemistry (Moroz et al., 1994). Moreover, exogenous applications of the NO donor 429

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S-nitrosocysteine or L-arginine elicit movements of the buccal mass, which led to the early conclusion that the NO-producing neurons in the buccal ganglion activate the CPG (Moroz et al., 1993). It was subsequently shown that lip chemosensory afferents are the primary source of NO in the feeding motor system (Elphick et al., 1995). Bipolar neurons in the lip stain intensely with NADPH-diaphorase histochemistry. These cells project to the cerebral ganglia and arborize in close proximity to modulatory interneurons in the feeding network. Chemosensory stimulation of the lips evokes a fictive feeding motor pattern in in vitro preparations of the Lymnaea nervous system. NO donors and a cGMP analogue mimic this effect when applied directly to the isolated central nervous system (Elphick et al., 1995). In addition, chemosensory-induced activation of the CPG is blocked when extracellular NO is scavenged with hemoglobin. NO thus appears to act as an anterograde neurotransmitter between sensory neurons in the lip and the CPG. The central release of NO presumably activates a guanylate cyclase in an as yet unidentified group of target neurons within the CNS. While central NO diffusion is required to initiate motor output from the CPG, it does not modulate synaptic interactions within the motor network itself (Elphick et al., 1995). 5.2. THE CRUSTACEAN STOMATOGASTRIC GANGLION The crustacean stomatogastric ganglion (STG) is a model system for studying how neuromodulatory transmitters act on a common pool of neurons to specify different functional networks. There are only ---30 neurons in the STG, and the physiological and synaptic properties of these cells have been extensively studied for more than 30 years (Harris-Warrick et al., 1992). STG neurons are arrayed into two functionally distinct motor networks which direct the movements of feeding structures in the foregut. These two networks, the gastric mill and pyioric, drive the grinding teeth of the gastric mill and the contractions of the pylorus, respectively. Rhythmic motor output from the STG is not fixed. Instead, the functional configuration of each circuit depends upon neuromodulatory and hormonal inputs to the system. At present, more than 20 different biogenic amines and neuropeptides are thought to be released into the STG neuropil (Harris-Warrick et al., 1992). Each transmitter evokes complex changes in the cellular and synaptic properties of individual neurons within the ganglion. These changes range from subtle modifications of an individual cell's activity pattern to the complete reorganization of an entire circuit. It has only recently become possible to study the distribution of NOS in the crustacean nervous system. The localization of crustacean NOS proteins has lagged behind parallel studies in other invertebrates, in part because conventional NADPH-d histochemistry does not label the majority of NOS-containing cells in the crustacean nervous system (Scholz et al., 1998). This may reflect an unusual sensitivity of the crustacean enzyme to paraformaldehyde fixation, as has been shown for the endothelial form of vertebrate NOS (Dinerman et al., 1994). However, the introduction of specific antibodies to universal NOS (uNOS) and citrulline (a product of NO biosynthesis) has now made it possible to detect NO-producing neurons in crustaceans. The anatomical distribution of key enzymes in the NO/cGMP signaling pathway suggests that NO acts as an anterograde transmitter in the STG. NOS-like proteins have been identified in the STG of the crab Cancer productus using an antibody to uNOS (Scholz et al., 1997). Citrulline accumulates in the terminals of neurons that project to the STG (Scholz et al., 1997), suggesting that modulatory inputs release NO directly into the synaptic neuropil of the ganglion. NO donors activate cGMP synthesis (Scholz et al., 1996) and cGMP immunocytochemistry reveals a distinct subset of sGC-expressing cells in the ganglion 430

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l Fig. 8. NO-induced sGC activation in the crab stomatogastric ganglion. The surrounding glial sheath was removed and the ganglion was treated with an NO donor (SNP) in the presence of IBMX. Following NO stimulation, the ganglion was fixed, processed for anti-cGMP immunocytochemistry, and imaged on a confocal microscope. As previously reported (Scholz et al., 1996), NO activates cGMP synthesis in --~13 of the 30 neurons in the ganglion. Note that cGMP staining intensity varies among individual neurons. Within a neuron, staining is diffusely distributed in somata, the central neuropil (the principal site of synaptic contacts in the ganglion), and in the axons that project out to the muscles that line the foregut. Scale bar = 50 ram. (N.L. Scholz, unpublished results.)

(Fig. 8). These NO-sensitive neurons (~ 13) innervate both gastric and pyloric muscles (Scholz et al., 1997), providing little clue as to how NO might modulate STG networks. Importantly, NO/cGMP signaling specifies the temporary assembly of a complete functional network within the adult STG. Electrophysiological studies from in vitro preparations of the STG have shown that a tonic release of NO within the ganglion is required for the production of gastric mill behavior (Scholz et al., 1997). Blocking anterograde NO diffusion 431

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Fig. 9. Inhibition of an NO-sensitive sGC results in the functional reorganization of the gastric mill motor circuit in the crab stomatogastric ganglion. Under basal conditions, the DG motor neuron (as recorded extracellularly from the dorsal gastric nerve, or dgn) and the GM motor neurons (as recorded from the anterior ventricular nerve, or avn) exhibit rhythmic bursts of coordinated activity. The motor pattern rpidly breaks down when the ganglion is bathed in ODQ, a specific inhibitor of NO-sensitive sGC. The DG neuron falls silent, and the GM cells switch pattern and begin firing in time with a separate network (the pyloric circuit). The effect is reversible, and the gastric motor pattern reforms when the ganglion is returned to normal saline. (N.L. Scholz, unpublished results.)

within the synaptic neuropil results in the rapid and selective disassembly of the gastric mill CPG. Some gastric mill neurons fall silent while others switch pattern and begin firing in phase with the pyloric CPG. The STG is similarly reconfigured when the NO-sensitive sGC is blocked by the inhibitor ODQ (Fig. 9). When PTIO or ODQ treatments are alternated with normal saline, the gastric mill network is selectively dismantled and reassembled over the span of a few seconds. This can be repeated multiple times during the course of a single experiment. Therefore, NO-releasing inputs can functionally rewire an active neural circuit by selectively modulating neurons that share a common diffusional space. NO-mediating signaling in the crab STG differs in some respects from the situation in the gastropod feeding network. In both systems NO originates from external inputs and serves to activate a feeding motor rhythm. However, in molluscs NO simply initiates the feeding motor program and is not involved in the central organization of the behavior (Elphick et al., 1995). By contrast, NO triggers an extensive reorganization of central circuitry in the STG, as manifest by pattern-switching in the gastric mill neurons (Scholz et al., 1997). Although specific mechanisms are unknown, network assembly is likely to involve complex changes in the cellular and synaptic properties of each of the 13 NO-sensitive neurons in the ganglion. In both molluscs and crustaceans, NO subserves specific functions that can be readily traced to the behavior of the intact animal. In crustaceans, NO-induced network partitioning in vitro closely resembles dynamic changes in motor output that have been recorded from freely behaving animals. Chronic recordings from intact lobsters have shown that while the pyloric rhythm is produced continuously, the gastric mill rhythm rapidly switches on and off (Clemens et al., 1998). Similarly, neurons that switch pattern in response to NO/cGMP neuromodulation are the same ones that switch between networks in vivo (Heinzel et al., 1993). We propose that the central release of NO triggers the dynamic assembly of the gastric mill motor network, which in turn drives the teeth of the gastric mill during bouts of feeding. This arrangement is likely to remain in place only as long as there is food in the gut. Upon the cessation of feeding, NO/cGMP signaling would be expected to terminate, returning the STG to a unitary network configuration. There is an intriguing relationship between NO-mediated signaling and the development of STG networks. In the lobster (genus Homarus) STG, the temporal appearance of the NO-sensitive sGC coincides with the first production of gastric mill motor behavior (Scholz et al., 1998). Both appear relatively late during post-embryonic development. At earlier 432

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stages (e.g., in newly hatched larvae) the foregut lacks a gastric mill. This condition persists throughout the larval stages of development, a period when the animal feeds on zooplankton in the water column. During this time the planktonic larva shreds prey tissues using its mandibles (Factor, 1981). The teeth of the gastric mill first appear at metamorphosis when the animal begins its transition down to the seafloor. However, the complete formation and calcification of the teeth is a gradual process that spans several developmental stages. While STG neurons are born and innervate their respective muscles in the embryo (Casasnovas and Meyrand, 1995), motor output from the ganglion is limited to a simple, unique rhythm throughout embryonic and larval stages of development. Adult STG networks begin to elaborate from this larval network at metamorphosis, in parallel with the gradual formation of the gastric mill. Interestingly, NO-responsiveness in the STG first appears in a single neuron at this time (Scholz et al., 1998). Additional neurons become NO-responsive as the teeth form and the gastric motor pattern takes shape (Fig. 10). The close developmental association between NO/cGMP signaling, the formation of the gastric mill, and the elaboration of adult STG networks further supports a key role for NO in the specification of gastric mill motor behavior. 5.3. THE CRUSTACEAN CARDIAC NETWORK The crustacean heart contains a very high level of NOS enzymatic activity relative to other tissues, including both skeletal muscles and the nervous system (Labenia et al., 1998). This raises the possibility that NO regulates cardiac rhythmicity in certain invertebrates, perhaps by mechanisms that are similar to those identified in vertebrates. NO plays an extensive and complex role in the regulation of vertebrate cardiac function, where it acts as a second messenger in the cholinergic control of pacemaking activity in mammalian sinoatrial myocytes (Han et al., 1994, 1995). Unlike mammals, the crustacean heart is neurogenic. The underlying cardiac nervous system consists of only nine neurons: five large motor neurons and four small pacemaking cells contained within the cardiac ganglion (Alexandrowicz, 1932). Together these cells generate stable bursts of activity which drive the rhythmic contractions of the cardiac muscles. Basic mechanisms of pattern generation have been studied in the crustacean cardiac ganglion for more than 50 years (reviewed by Cooke, 1988). Inhibitory and excitatory synaptic inputs originate from a pair of nerves which represent the only connection between the ganglion and the rest of the nervous system. The cardiac network is arrayed along the dorsal lumen of the heart and therefore also receives extensive modulatory input from blood-borne neurohormones. Although the oscillatory properties of crustacean cardiac and stomatogastric neurons are similar (Benson and Cooke, 1984), NO appears to play a very different role in the two nervous systems. Whereas NO acts as an anterograde transmitter in the STG, it appears to serve as an intracellular messenger within the cardiac ganglion. All nine cardiac cells label with an antibody to uNOS (Labenia et al., 1998). Citrulline accumulates in the somata and axons alike (Fig. 11), indicating a distributed release of NO throughout the entire "cardiac network. Moreover, all neurons show detectable cGMP accumulation under basal conditions, and the total cGMP content of the ganglion can be elevated by treatment with NO donors. Surprisingly, basal cGMP levels are --~10 times higher in the cardiac ganglion than in the STG. While overall cyclase activity is due in part to the NO-sensitive sGC, there is also a significant contribution from an NO-insensitive enzyme, the one that has been previously identified in the crustacean nervous system (Prabhakar et al., 1997). Taken together, the anatomical and biochemical data suggest that cardiac neurons both synthesize and respond to NO. 433

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Preliminary electrophysiological studies support a role for NO as an intracellular cardioinhibitor (Labenia et al., 1998). PTIO, which scavenges extracellular NO and was very effective on the crab STG, has no effect on the cardiac motor rhythm. However, blocking sGC activation with O D Q accelerates the firing frequency of the large motor neurons. Intrinsic NO production appears to be sufficient to saturate the physiological system since NO donors and c G M P analogs have no effect on firing activity. NO presumably modulates the oscillatory properties of individual cardiac neurons or synaptic transmission within the ganglion, and may be intermediate to excitatory or inhibitory synaptic inputs from the central nervous system. However, specific mechanisms have yet to be determined. Based on anatomical evidence, NO rarely acts as an intracellular messenger in the crustacean nervous system (Scholz et al., 1998). In this respect cardiac neurons are more similar to mammalian sinoatrial myocytes than to stomatogastric neurons. This raises the interesting possibility that NO may play an evolutionarily conserved functional role in cardiac pacemaking. In both mammals (Han et al., 1994) and crabs, NO acts as an inhibitory messenger. In mammalian primary pacemaker cells, acetylcholine activates N O / c G M P signaling, which in turn inhibits an I_-type calcium current via crosstalk with the c A M P pathway (Han et al., 1995). While a similar mechanism may be in place in the cardiac ganglion, the upstream activator is probably not acetylcholine since this transmitter increases rhythmic burst frequency in the network (Sullivan and Miller, 1990). In summary, N O / c G M P signaling serves different functions in the three model systems surveyed here. The specific role for NO ranges from an anterograde network activator (Lymnaea feeding circuit) to a network reorganizer (crab STG) to an intracellular messenger (carb cardiac ganglion). In all three cases, NO is likely to be a cotransmitter or a parallel modulator in a simple circuit that receives multiple chemical inputs. Therefore, the variation may arise both from subtle differences in circuitry and the presence of other transmitters. Together these models provide an opportunity to compare different mechanisms of N O / c G M P neuromodulation in networks that give rise to discrete behaviors.

6. NO AND NOCICEPTION N O / c G M P signaling is involved in the nociceptive pathway in the rat somatosensory system (e.g., Salter et al., 1996). This pathway may also be involved in nociception in invertebrates. In the moth Manduca sexta, early in metamorphosis the transection of peripheral nerves results in a dramatic production of c G M P in a small set of central neurons, many of which are serotonergic (Schachtner et al., 1998). This c G M P increase in response to injury is suppressed by the presence of L-NAME (J. Schachtner and J. Truman, unpubl, data), indicating the involvement of NO in mediating this response.

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Fig. 10. Post-embryonic development of NO-sensitive sGC activity in the lobster stomatogastric ganglion. The input nerve (stomatogastric nerve, labeled stn in the figure) is to the left and the dorsal ventricular nerve (dvn) is to the right. Each panel shows NO-induced cGMP immunoreactivity at successive developmental stages from hatching (stage I at top) to metamorphosis (stages III and IV) to adulthood IA at bottom). NO-sensitivity first appears in a single neuron at stage III. Additional cells begin to respond to NO as the animal undergoes metamorphosis and settles from the plankton to a life on the seafloor. The gradual onset of sGC activity coincides with a developmental reorganization of the ganglion from a simple larval circuit to a network which will eventually generate adult-specific motor behaviors (Casasnovas and Meyrand, 1995). Scale bar = 100 mm in A, 25 mm for I-V. From Scholz et al., 1998. 435

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Potential nociceptive sensory neurons were identified in larval Manduca using anti-cGMP immunocytochemistry (Fig. 12). Most sensory neurons show NO sensitivity during their development but become unresponsive as mature cells. These sensory neurons, by contrast, continue to be NO-sensitive after their development is complete (Grueber and Truman, 1999). These cells have large branching dendrites that form a network of processes that cover the surface of the animal and the dendrites terminate in naked endings in the epidermis. Electrophysiological recordings show that the cells respond to a range of noxious stimuli, including high heat and severe deformation of the cuticle (W. Grueber, unpubl, data). These cells are likely to be involved in nociception and their NO sensitivity may be related to this function. This is the first time that nociception in insects has been attributed to a particular type of sensory neuron. Because of their accessibility, these cells may provide a valuable model for examining the role of NO in nociception.

7. NO AS A POTENTIAL BLOOD-BORNE NEUROHORMONE The vasoactive properties of NO have been extensively studied in vertebrates. The first evidence that NO could serve as a cell-cell signaling molecule came with the discovery that NO was in fact an 'endothelial derived relaxing factor', which triggers vasorelaxation in mammalian smooth muscle (Palmer et al., 1987). NO is a reactive and relatively short-lived gas and was thought until recently to have a limited range of action in the mammalian vascular system. NO also binds to metal-containing proteins like hemoglobin, and the abundance of hemoglobin in blood was thought to further limit NO's sphere of diffusion. However, it was subsequently discovered that NO groups can also attach to hemoglobin by S-nitrosylation (Jai et al., 1996). Hemoglobin therefore acts as a 'carrier protein'. Since NO-binding proteins can transport and deliver NO throughout the body, NO can also function as a circulating neurohormone. One of the first functional demonstrations that invertebrates make use of NO-binding

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Fig. 12. Development of a set of putative nociceptive sensory neurons in the body wall of the second abdominal segment of the developing caterpillar of Manduca sexta. Embryos were pinned open, exposed to an NO donor and IBMX for 15 min and then fixed and stained for cyclic GME (A) At approximately 50% of embryogenesis (% E) the neurons become NO-sensitive after their axons have projected into the CNS through the dorsal (DN) or ventral (VN) nerves. (B) By 59% E, the neurons show extensive fine branches and filopodia extending from the cell bodies. (C) By 67% E, dendritic branches are more varicose and fine filopodial extensions have been lost. GN, ganglion A2. For each hemisegment, the dorsal midline is up and anterior is to the right. From Grueber and Truman (1999).

proteins comes from the bloodsucking bug Rhodnius prolixus. In the salivary glands of Rhodnius, a constitutive NOS produces NO (Ribeiro and Nussenzveig, 1993). NO is bound to and stored by a hemoprotein (Ribeiro et al., 1993) which is released into a host's bloodstream when the animal feeds. The subsequent liberation of NO relaxes the host's vasculature, enhancing the supply of food to the insect. NO-binding nitrosylheme proteins appear to have originated very early in eukaryotic evolution. Several invertebrates use hemoglobin as a respiratory protein. Others, including some molluscs and crustaceans, use hemocyanin. The NO-binding properties of hemoglobin and hemocyanin are similar. Like hemoglobin, hemocyanin binds NO (Solomon, 1981; Schoot Uiterkamp and Mason, 1973). Since S-nitrosylheme proteins can donate NO under the appropriate redox conditions (Pan et al., 1996), hemocyanin, like hemoglobin (cf. Stamler et al., 1997), could potentially deliver NO to a range of invertebrate tissues, especially in animals like crabs that have a semi-open circulatory system. If NO is released into the blood, what are the sources? There is some preliminary evidence from crustaceans that NO originates within the heart itself. In the crab Cancer productus, the heart has the highest levels of NOS enzymatic activity when compared with nervous system, muscle, and hepatopancreas (Labenia et al., 1998). The crab heart is neurogenic and, as described above, every neuron in the cardiac nervous system contains NOS (as detected by universal NOS and L- citrulline immunostaining). This nervous system branches extensively along the dorsal lumen of the heart, in direct contact with the circulating hemolymph. NO generated in the cardiac nervous system could thus be released directly into the blood in 437

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addition to acting on local targets within the heart. NO in the blood might target the bicuspid m u s c u l a r valves that guard the entrance to each artery, thereby controlling patterns of blood flow throughout the body. A similar function has been found for other neurotransmitters and n e u r o h o r m o n e s (Wilkens, 1997). Alternatively, blood-borne NO may act on e l e m e n t s within the nervous system such as the stomatogastric ganglion, which resides in the lumen of the ophthalmic artery. However, these inferences are only speculative and additional work on the physiological significance of N O - b i n d i n g proteins and blood-borne NO is needed.

8. ACKNOWLEDGEMENTS We would like to thank Jan de Vente, University of Maastricht, for the gift of the a n t i - c G M P antibody and S o l o m o n Snyder and Mikael Eliasson, Johns Hopkins University, for the gift of the anti-citrulline antibody. We are also grateful to Jana Labenia for technical assistance, and to Kathy G r a u b a r d and Michael G o y for their critical discussions. This work was supported by NIH Grant 15697 and an NIH predoctoral traineeship to NLS.

9. REFERENCES Alexandrowicz JS (1932): Innervation of the heart of Crustacea, I. Decapoda. Q J Microsc Sci 75:182-249. Atkinson J, Panni MK (1999): Optic target regulation of NADPH-diaphorase by larval retinal axons in Drosophila. Neurosci Lett 262:21-24.

Ball EE, Truman JW (1998): Developing grasshopper neurons show variable levels of guanylyl cyclase activity on arrival at their targets. J Comp Neurol 394:1-13. Bascal ZA, Montgomery A, Holden-Dye L, Williams RG, Walker RJ (1995): Histochemical mapping of NADPHdiaphorase in the nervous system of the parasitic nematode Ascaris suum. Parasitology 110:625-637. Benjamin PR, Elliott CJH (1989): Snail feeding oscillator: the central pattern generator and its control by modulatory interneurons. In: Jacklett JW led). Neuronal and Celhdar Oscillators. New York, NY: Dekker, pp 173-174. Benson JA, Cooke IM (1984): Driver potentials and the organization of rhythmic bursting in crustacean ganglia. Trends Neurosci 7:85-91. Bicker G (1998): NO news from insect brains. Trends Nemvsci 21:349-355. Boeckxstaens GE, Pelckmans PA (1997): Nitric oxide and the non-adrenergic non-cholinergic neurotransmission. Comp Biochem Phvsiol 118A:925-937.

Bredt DS, Snyder SH (1992): Nitric oxide, a novel neuronal messenger. Neuron 8:3-11. Bredt DS, Glatt CE, Hwang PM, Fotuhi M, Dawson TM. Snyder SH (1991): Nitric oxide synthase protein and mRNA are discretely localized in neuronal populations of the mammalian CNS together with NADPH diaphorase. Neuron 7:615-624. Breer H, Shepherd GM (1993): Implications of the NO/cGMP system in olfaction. Trends Neurosci 16:5-8. Casasnovas B, Meyrand P (1995): Functional differentiation of adult neural circuits from a single embryonic network. J Neurosci 15:5703-5718. Champlin DT, Truman JW (1998): Ecdysteroid control of cell proliferation during optic lobe neurogenesis in the moth Manduca sexta. Development 125:269-277. Clemens S, Meyrand P, Simmers J (1998): Feeding-induced changes in temporal patterning of muscle activity in the lobster stomatogastric system. Neurosci Lett 254:65-68. Colasanti M, Lauro GM, Venturini G (1995): NO in hydra feeding response. Nature 374:505. Colasanti M, Venturini G, Merante A, Musci G, Lauro GM t1997): Nitric oxide involvement in Hydra vulgaris very primitive olfactory-like system. J Nel,vsci 17:493-499. Cooke IM (1988): Studies on the crustacean cardiac ganglion. Comp Biochem Phvsiol 91C:205-218. Delledonne M, Xia Y, Dixon RA, Lamb C (1998): Nitric oxide functions as a signal in plant disease resistance. Nature 394:585-588.

Dinerman JL, Dawson TM, Schell MJ, Snowman A. Snyder SH (1994): Endothelial nitric oxide synthase localized to hippocampal pyramidal cells: implications for synaptic plasticity. Plvc Natl Acad Sci USA 91:4214-4218. 438

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Elofsson R, Carlberg M, Moroz L, Nezlin L, Sakharov D (1993): Is nitric oxide (NO) produced by invertebrate neurons?. Neuroreport 4:279-282. Elphick MR, Melarange R (1998): Nitric oxide function in an echinoderm. Biol Bull 194:260-266. Elphick M, Green I, O'Shea M (1993): Nitric oxide synthesis and action in an invertebrate brain. Brain Res 619:344-346. Elphick MR, Kemenes G, Staras K, O'Shea M (1995): Behavioral role for nitric oxide in chemosensory activation of feeding in a mollusc. J Neulvsci 15:7653-7664. Eriksson KS (1996): Nitric oxide synthase in the pharynx of the planarian Du,~esia ti~rina. Cell ~'ssue Res 286:407-410. Factor JR ( 1981): Development and metamorphosis of the digestive system of larval lobsters, Homarus americamls (Decapoda, Nephropidae). J Motphol 169:225-242. Feelish M, Martin JF (1995): The early role of nitric oxide in evolution. Tremts Ecol Evol 10:496-499. Garthwaite J, Boulton CL (1995): Nitric oxide signaling in the central nervous system. Annu Rev Phvsiol 57:683706. Garthwaite J, Southam J, Boulton CL, Nielsen EB, Schmidt K, Mayer B (1995): Potent and selective inhibition of nitric oxide-sensitive guanylyl cyclase by 1H-[ 1,2,4]oxadiazolo[4.3-c~]quinoxalin-l-one. Mol Pharmacol 48:184188. Gelperin A (1994): Nitric oxide mediates network oscillations of olI:actory interneurons in a terrestrial mollusc. Nature 369:61-63. Gibbs SM, Truman JW (1998): Nitric oxide and cyclic GMP regulate retinal patterning in the optic lobe of Drosophila melanogaster. Neuron 20:83-93. Gibbs SM, Currie D, Truman JW (2000): Nitric oxide synthase expression and cyclic GMP production in the Drosophila nervous system during larval life and metamorphosis (submitted). Goodman CS, Bastiani MJ, Doe CQ, Du Lac S, Helfand SL, Kawada JY, Thomas JB (1984): Cell recognition during neuronal development. Science 225:1271-1279. Grueber WB, Truman JW (1999): Identification and development of a nitric oxide-sensitive peripheral plexus in larvae of the moth, Manduca sexta. J Comp Neutvl 404:127-141. Gudi T, Lohmann SM, Pilz RB (1997): Regulation of gene expression by cyclic GMP-dependent protein kinase requires nuclear translocation of the kinase: identification of a nuclear localization signal. Mol Cell Biol 17:52445254. Hart X, Shimoni Y, Giles WR (1994): An obligatory role for nitric oxide in autonomic control of mammalian heart rate. J Phvsiol 476:309-314. Hart X, Shimoni Y, Giles WR (1995): A cellular mechanism for nitric oxide-mediated cholinergic control of mammalian heart rate. J Gen Phvsiol 106:45-65. Harris-Warrick RM, Marder E, Selverston AI, Moulins M (1992): Dynamic Biological Networks: The Stomatogastric Nen'ous S~'stem. Boston, MA: MIT Press. Heinzel HG, Weimann JM, Marder E (1993): The behavioral repertoire of the gastric mill in the crab. Cancer pagurus: an in situ endoscopic and electrophysiological examination. J Neurosci 13:1793-1803. Hope BT, Michael GJ, Knigge KM, Vincent SR (1991): Neuronal NADPH diaphorase is a nitric oxide synthase. Proc Natl Acad Sci USA 88:2811-2814. Jacklet JW (1997): Nitric oxide signaling in invertebrates, lm'ert Neu~vsci 3:1-14. Jai L, Bonaventura C, Bonaventura J, Stamler J (1996): S-nitrosohaemoglobin: a dynamic activity of blood involved in vascular control. Nature 380:221-226. Johansson KUI, Carlberg M (1994): NADPH-diaphorase histochemistry and nitric oxide synthase activity in the deutocerebrum of the crayfish, Pacifilsmcus ieniuscuhls (Crustacea, Decapoda). Brain Res 649:36-42. Korneev SA, Piper MR, Picot J, Phillips R, Korneeva E, O'Shea M (1998): Molecular characterization of NOS in a mollusc: expression in a giant modulatory neuron. J Nemvbiol 35:65-76. Kuzin B, Roberts I, Peunova N, Enikolopov G (1996): Nitric oxide regulates cell proliferation during Drosophila development. Cell 87:639-649. Labenia J, Scholz NL, Goy ME Graubard K (1998): NO/cGMP modulates the crustacean cardiac ganglion. Soc Neurosci Abstr 24:360. Leake LD, Moroz LL (1996): Putative nitric oxide synthase (NOS)-containing cells in the central nervous system of the leech, Hirudo medicinalis: NADPH-diaphorase histochemistry. Brain Res 723:115-124. Lin MF, Leise EM (1996): NADPH-diaphorase activity changes during gangliogenesis and metamorphosis in the gastropod mollusc llvanassa obsoleta. J Comp Neurol 374:194-203. Martinez A, Riveros-Moreno V, Polak JM, Moncada S, Seesma P (1994): Nitric oxide (NO) synthase immunoreactivity in the starfish Marthasterias glacialis. Cell Tissue Res 275:599-603. 439

Ch. XI

N.L. Scholz and J.W. Truman

Meinertzhagen IA, Hanson TE (1993): The development of the optic lobe. In: Bate M, Martinez Arias A (Eds), The Development of Drosophila melanogaster. Cold Spring Harbor: Cold Spring Harbor Press, pp 1363-1491. Moroz LL, Park J-H, Winlow W (1993): Nitric oxide activates buccal motor patterns in Lvmnaea stagnalis. Neuroreport 4:646-646. Moroz LL, Winlow W, Turner RW, Bulloch AGM, Lukowiak K, Syed NI (1994): Nitric oxide synthase-immunoreactive cells in the CNS and periphery of Lvmnaea. Neuroreport 5:1277-1280. Moroz LL, Chen D, Gillette MU, Gillette R (1996): Nitric oxide synthase activity in the molluscan CNS. J Neurochem 66:873-876. Moroz LL, Gillette R, Sweedler JV (1999): Single-cell analysis of nitrergic neurons in simple nervous systems. J Exp Biol 202:333-341. Morton DB (1997): Eclosion hormone action on the nervous system. Intracellular messengers and sites of action. Ann NY Acad Sci 814:40-50. Miiller U (1994): Ca 2+/calmodulin-dependent nitric oxide synthase in Apis mellifera and Drosophila melanogaster. Eur J Neurosci 6:1362-1370. Mtiller U (1997): The nitric oxide system in insects. Prog Neurobiol 51:363-381. Miiller U, Bicker G (1994): Calcium-activated release of nitric oxide and cellular distribution of nitric oxide-synthesizing neurons in the nervous system of the locust. J Neurosci 14:7521-7528. Murphey RK, Davis GW (1994): Retrograde signalling at the synapse. J Neurobiol 25:595-598. Nathan C (1995): Natural resistance and nitric oxide. Cell 82:873-876. Nighorn A, Gibson NJ, Rivers DM, Hildebrand JG, Morton DB (1998): The nitric oxide-cGMP pathway may mediate communication between sensory afferents and projection neurons in the antennal lobe of Manduca sexta. J Neurosci 18:7244-7255. Nighorn A, Byrnes KA, Morton DB (1999): Identification and characterization of a novel 13 subunit of soluble guanylyl cyclase that is active in the absence of a second subunit and is relatively insensitive to nitric oxide. J Biol Chem 274:2525-2531. Ogunshola O, Picot I, Piper M, Korneev S, O'Shea MR (1995): Molecular analysis of the NO-cGMP signalling pathway in insect and molluscan CNS. Soc Neurosci Abstr 21:631. Palmer RMJ, Ferridge AG, Moncada S (1987): Nitric oxide release accounts for the biological activity of endothelium-derived relaxing factor. Nature 327:524-526. Pan Z, Segal M, Lipton S (1996): Nitric oxide-related species inhibit evoked neurotransmission but enhance spontaneous miniature synaptic currents in central neuronal cultures. Proc Natl Acad Sci USA 93:15423-15428. Prabhakar S, Short DB, Scholz NL, Goy MF (1997): Identification of nitric oxide-sensitive and -insensitive forms of cytoplasmic guanylate cyclase. J Neurochem 69:1650-1660. Radomski MW, Martin JF, Moncada S (1991): Synthesis of nitric oxide by the haemocytes of the American horseshoe crab (Limutus polyphemus). Philos Trans R Soc Lond B Biol Sci 33:129-133. Regulski M, Tully T (1995): Molecular and biochemical characterization of dNOS, a Drosophila Ca 2+/calmodulin-dependent nitric oxide synthase. Proc Natl Acad Sci USA 92:9072-9076. Ribeiro JMC, Nussenzveig RH (1993): Nitric oxide synthase activity from a hematophagous insect salivary gland. FEBS Lett 330:165-168. Ribeiro JM, Hazzard JM, Nussenzveig RH, Champagne DE, Walker FA (1993): Reversible binding of nitric oxide by a salivary heme protein from a bloodsucking insect. Science 260:539-541. Robertson JD, Bonaventura J, Kohm AP (1994): Nitric oxide is required for tactile learning in Octopus vulgaris. Proc R Soc Lond B Biol Sci 256:269-273. Rose RM, Benjamin PR (1979): The relationship of the central motor pattern to the feeding cycle of Lvmnaea stagnalis. J Exp BioI 80:137-163. Salleo A, Musci G, Barra PFA, Calabrese L (1996): The discharge mechanism of aconital nematocytes involves the release of nitric oxide. J Exp Biol 199:1261-1267. Salter M, Strijbos PJ, Neale S, Duffy C, Follenfant RL, Garthwaite J (1996): The nitric oxide-cGMP pathway is required for nociceptive signalling at specific loci within the somatosensory pathway. Neuroscience 73:649-655. Schachtner J, Klassen L, Truman JW (1998): Metamorphic control of cyclic GMP expression in the nervous system of the tobacco hornworm, Manduca sexta. J Comp Neurol 396:238-252. Schachtner J, Homberg U, Truman JW (1999): Regulation of cyclic GMP elevation in the developing antennal lobe of the sphinx moth, Manduca sexta. J Neurobio141:359-375. Scholz NL, Goy MF, Truman JW, Graubard K (1996): Nitric oxide and peptide neurohormones activate cGMP synthesis in the crab stomatogastric nervous system. J Neurosci 16:1614-1622. Scholz NL, Truman JW, Graubard K (1997): Modulation of flexible motor circuits by nitric oxide and cGMP. Soc Neurosci Abstr 23:1787.

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Invertebrate models for studying NO-mediated signaling

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Scholz NL, Chang ES, Graubard K, Truman J (1998): The NO/cGMP pathway and the development of neural networks in postembryonic lobsters. J Neurobiol 34:208-226. Schoot Uiterkamp AJM, Mason HS (1973): Magnetic dipole-dipole coupled Cu(II) pairs in nitric oxide-treated tyrosinase: a structural relationship between the active sites of tyrosinase and hemocyanin. Proc Natl Acad Sci USA 70:993-996. Schuman EM, Madison DV ( 1994): Nitric oxide and synaptic function. Annu Rev Neurosci 17:153-183. Serfozo Z, Elekes K, Varga V (1998): NADPH-diaphorase activity in the nervous system of the embryonic and juvenile pond snail, Lvmnaea stagnalis. Cell Tissue Res 292:579-586. Shah S, Hyde DR (1995): Two Drosophila genes that encode the alpha and beta subunits of the brain soluble guanylyl cyclase. J Biol Chem 270:15368-15376. Solomon El (1981): Binuclear copper active site. In: Spiro G (Ed), Copper Proteins. New York, NY: Wiley-Interscience, pp 41-108. Song H-j, Ming G-I, He Z, Lehmann M, McKerracher L, Tessier-Lavigne M, Poo M-m (1998): Conversion of neuronal growth cone responses from repulsion to attraction by cyclic nucleotides. Science 281:1515-1518. Stamler J, Toone EJ, Lipton SA, Sucher NJ (1997): (S)NO signals: translocation, regulation, and consensus motif. Neuron 18:691-696. Sullivan RE, Miller MW (1990): Cholinergic activation of the lobster cardiac ganglion. J Neurobiol 21:639-650. Truman JW, De Vente J, Ball EE (1996): Nitric oxide-sensitive guanylate cyclase activity is associated with the maturational phase of neuronal development in insects. Development 122:3949-3958. Wildemann B, Bicker G (1999a): Developmental expression of nitric oxide/cyclic GMP synthesizing cells in the nervous system of Drosophila melanogaster. J Neurobiol 38:1 - 15. Wildemann B, Bicker G (1999b): Nitric oxide and cyclic GMP induce vesicle release at Drosophila neuromuscular junction. J Neurobiol 39:337-346. Wilkens JL (1997): Possible mechanisms of control of vascular resistance in the lobster Homarus americanus. J Exp Bio1200:487-493. Wolff T, Ready D (1993): Pattern formation in the Drosophila retina. In: Bate M, Martinez Arias A (Eds), The Development of Drosophila melanogaster. Cold Spring Harbor: Cold Spring Harbor Press, pp 1277-1325. Zhang J, Dawson VL, Dawson TM, Snyder SH (1994): Nitric oxide activation of poly (ADP-ribose) synthetase in neurotoxicity. Science 263:687-689.

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Subject Index

A1 cell group 37, 185, 187, 192 A2 cell group 192, 194 accessory olfactory bulb 21, 59, 63, 72 acetylcholine 159, 215, 223225, 227, 230, 237, 241, 243, 244, 435 nNOS colocalization 26, 28, 35, 36, 38, 149, 188, 193, 197, 243, 245 acetyicholinesterase 28 nNOS colocalization 91 ACTH 157, 158 ACTH release 158, 159 ADP-ribosylation 128 phototransduction 127 adrenal gland 38, 88, 158 cGMP 246 adrenal medulla 245, 246 chromaffin cells 217, 219 adrenal medullectomy 321 adrenal steroids 151 adrenalectomy 151 adrenocorticotropin see ACTH amacrine cells (see also retina) dopamine 132, 137 GABA 131, 132 NO release 137 periphery 121 subclasses 115-118, 120, 131, 136, 137 regional distribution 120, 121 amygdala 26, 59, 63, 67, 75, 82, 149, 365 13-amyloid percursor protein nNOS colocalization 327 angiotensin II 149, 157, 165, 189 anococcygeus muscle 215

anterior commissure 27, 65, 82, 365 anterior hippocampal rudiment 21 anterior hypothalamus 161 anterior intralaminar nucleus 28 anterior olfactory nucleus 21, 22, 63, 73, 82. 396 anterior pituitary 317 anterograde tracing 179, 189 anteromediai nucleus 28 anteroventral nucleus 28 aorta 215 Aplysia 418 apoptosis 92, 330, 332, 343 root avulsion 335 spinal motoneurons 335 Apteronotus leptorhynchus 55,57 aqueduct 30. 33 aqueductai-isthmic region 55 area postrema 33, 87, 180. 182, 187, 197 L-arginine 9 astrocytes 38 availability 126, 342 conversion of 5 gap junctions 130 immunoreactivity 38 metabolism 38 nitrate/nitrite precursor 3.4 storage 38 vasopressin release 160 arginine deiminase 3 arginine vasopressin 33 nNOS colocalization 3 l argininosuccinate lyase 38 argininosuccinate synthase 38 arthropods 419 Ascaris suum 429 astrocytes 166, 400

ATP 215, 219, 224, 228, 232, 233, 235 auditory area 182 auriculo-temporal nerve 242 autonomic ganglia 217 autonomic nervous system 320 coordination 206 peripheral overview 246 autonomic regulation ! 64, 177 hypothalamus 164 subfornical organ 164 suprachiasmatic nucleus 165 supraoptic nucleus 165 autonomic vasodilator neuron 224 axo-axonal interactions 237 axonal transport interruption 340 axotomy 321,326, 328. 334 barrel fields 24 basilar artery 225 BDNF 338 bed nucleus of the stria terminalis 149, 365 biliary system 239 biocytin 199 biotinylated dextran amine 179, 202 bipolar cells (see also retina) glutamate receptors 131 bladder 223 brain slices 359 brain stem 28, 149 9 cyclostomes 57 fish 57 brown fat 220 buccal ganglion 429, 430 bulbectomy 322, 345

c-fos 157. 158, 164, 199, 203 c-jun 327, 339 443

Subject Index

nNOS colocalization 327, 336 c-jun regenerating motoneurons 336 C1 cell group 37, 183, 189, 192, 194, 206 C2 cell group 192 calbindin nNOS colocalization 21, 27. 28, 31, 36, 91, 149, 188, 197, 198 calcineurin 28 calcitonin gene-related peptide see CGRP cairetinin nNOS colocalization 21, 28, 31, 91, 189, 197, 198 Cancer productus 428, 430, 437 capsaicin 228, 319 Carassius auratus 55 carbon monoxide 39, 232, 233,427 cardiac ganglion 221, 230, 435,436 pattern generation 433 cardiac network invertebrates 433, 435 cardiovascular system 199, 200 carotid sinus nerve 206 castration 151, 163 catecholamine secretion 247, 248 caudal ventrolateral medulla 192 caudate putamen 75, 78, 82, 362, 388, 389, 393, 394, 397, 400 celiac ganglion 219, 223, 239 ceUaco-mesenteric complex 219 central canal 181, 183, 187 central gray 70, 77, 85,376 cerebellar nuclei 65, 70, 86 cerebellum 37, 57, 63, 65, 70, 86, 321,322, 325, 332, 361, 367, 378, 388, 389, 400, 405 Bergmann glia 367, 385 cGMP 387 444

cGMP immunocytochemistry 384, 385 cGMP-dependent protein kinase I 384 development cGMP 386 granule cells 36 parallel fibers 367 Purkinje cells 36, 367. 378, 385 cGMP 384. 386, 387 soluble guanylate cyclase localization 384-387 cerebral artery 225 cerebral blood flow 92 cerebral cortex 22. 23.63.73, 88, 189, 321,326. 327, 360. 389, 394. 398. dendritic branching 92 layer V 321 cerebral cortical blood flow 24 cerebral ganglia 429 cerebrospinai fluid 33 cervical sympathetic ganglia 229 cGMP circadian rhythm 164 LHRH release 162 cGMP immunocytochemistry 358. 359 colocalization 395 acetylcholine transporter 4O0 enkaphalin 400 GABA 400 muscarinic receptors 400 nNOS 36, 362, 365, 372, 378.381-383 parvalbumin 398, 400, 401. 403. 405 soluble guanylate cyclase 397 vesicular acetylcholine transport 398 phosphodiesterase inhibitors selectivity 360. 361 retina 136 sensitivity 136, 359 species differences 389 cGMP-dependent protein

kinase 92, 130-132, 136, 162, 164, 426 type I 36 type II 29, 33, 372, 384 cGMP-gated channels 129, 131, 132, 136 retina 128 CGRP 188. 226-230, 231, 244, 327 nNOS colocalization 188, 226, 327 chaoptin 428 ChAT see choline acetyltransferase chemical injury 343 cholecystokinin nNOS colocalization 33, 149, 189, 192 cholera toxin B subunit 122, 179, 190, 201 choline acetyltransferase 27, 222, 225, 230, 240 nNOS colocalization 91 choroid 114 choroid plexus 33 chromaffin cells 245, 246, 248 ciliary ganglion 221,225 cingulate cortex 362, 363, 398, 400 circadian rhythm 24, 151, 152, 164 circle of Willis 225 circular nucleus 75 citrulline immunoreactivity 38 Clarke's nucleus 324-326, 336, 338 claustrum 26, 73 clitoris 227 cochlea 31 cochlear nucleus 76 colchicine 317. 340 colon 236, 238 commissure.of the inferior colliculus 377 comparative aspects adrenal glands mammals 246 airways nerve fibers mammals 240 basal forebrain mammal 27

Subject Index central nervous system 55 amphibians 59, 61,389 birds 67, 70, 71 fish 59 mammals 71-76, 389 reptiles 63, 65, 67 diencephalon amphibians 59 birds 70 cyclostomes 57 fish 57 mammals 75, 83 reptiles 65 enteric inhibitory motor neurons mammals 236 fish 55 gallbladder mammals 239 invertebrates 418, 419 mesencephalon amphibians 61 birds 70 cyclostomes 57 fish 57 mammals 75, 85 reptiles 65 nerve injury mammals 330 nucleus of the solitary tract mammals 204 retina amphibians 119-122, 124, 128, 129 birds 117, 121, 123, 124, 133 fish 119, 129, 131 human 114 mammals 114-117, 120, 121, 123, 127, 131, 133, 135-137 reptiles 118, 119, 129, 135-137 rhombencephalon amphibians 61 birds 71, 70 mammals 76, 86 reptiles 65 species differences amphibians 89-91 birds 89-91 fish 89-91

mammals 89-91 reptiles 89-91 spinal cord mammals 38 sweat glands mammals 244 telencephalon amphibians 59 birds 67 cyclostomes 55 fish 55 mammals 71, 77. 78.82 reptiles 63, 65 copulatory behavior 163 corpus callosum 24, 73, 362, 405 corpus cavernosum 3, 226 cortico-collicular projections 124 corticospinal tract 325 corticosterone 158 corticosterone release 158, 159 corticotropin-releasing hormone see CRH

Coturnix japonica 67-69 C-PON 21 nNOS colocalization 27 cranial nerve ligation 322 cranial nerve nuclei 317, 321 cranial parasympathetic neuron 228 cranial parasympathetic preganglionic neuron 221 CRH 149, 157 nNOS colocalization 36. 149, 157 CRH release 158, 159 cuneate nucleus 37. 76, 87, 180-182, 188. 193, 197. 372, 382 dark adaptation 129, 137 dark-adapted outer segments 127 dentate gyrus 25, 26, 73. 189 development 76-86, 328, 390-395, 419-428 retina 122 eNOS 123 nNOS 124 dexamethasone 165

diabetes 155 diagonal band of Broca 2 I, 27, 59, 63, 65, 73, 149 differentiation 92 5,6-dihydroxytryptamine 116 DiI 241 dopamine 157, 161 amacrine cells 132, 137 prolactin release 163 dopamine ~-hydroxylase 219 nNOS colocalization 192, 219 dorsal geniculate nucleus 365 dorsal medullary nucleus 181, 182 dorsal motor vagal nucleus 36, 71, 76, 181, 182, 183, 185, 187, 190, 191, 192, 193, 197, 199, 202, 221, 222, 240, 316, 317, 328, 340 esophagus 205 intestine 205 pancreas 205 stomach 204, 205 dorsal raphe nucleus 34, 35, 87, 189, 367, 376 dorsal reticular area 184 dorsal root ganglion 223, 226-230, 240, 246, 319, 320, 328 dorsolateral periaqueductal gray 30 dorsomedial hypothalamic nucleus 149 dorsomedial medulla 192 dorsomedial reticular field 197 dorsomediai thalamic nucleus 365 dorsoventral thalamic nucleus 365 drinking 161 Drosophila 418-420, 424-426 Dugesia tigrina 429 dye coupling 166 dynorphin 239, 324 ecdysteroids 419, 425,427 ectostriatum 67 Edinger-Westphai nucleus 57, 76 445

Subject Index

EDRF 1,215 cerebellum 2 ejaculation 163 embryogenesis 426 cGMP 420, 422, 423, 425-427 fl-endorphin LHRH release 162 endothelial cells L-arginine storage 38 endothelial NOS see eNOS endothelium-derived relaxing factor see EDRF enkephalin 239 nNOS colocalization 33, 38, 149, 189, 245 eNOS 4 caveolae 8 cerebellar granule cells 24 hippocampus 88 interstitial cells of Cajal 235 intestinal muscle cells 235 olfactory bulb 24 palmitoylation 9 retina 113, 115, 125, 138 development 123 spinal cord root avulsion 324 striatum 24 subcellular localization 8 enteric nervous system 227, 232-239 enteric reflex 237 entopeduncular nucleus 57, 73, 82 esophageal motor endplates 238 esophagus 204, 222, 228, 238, 240, 241 estrogen 33, 151 LH release 162 prolactin release 163 estrogen receptor 151 nNOS colocalization 32, 149, 151 estrous cycle 151 eukaryotic evolution 437 excitotoxicity 131 nNOS expression 343 extensor tibiae NO-responsive 425 446

external cuneate nucleus 180 extrinsic sensory neurons 229 eye (see also retina) autonomic vasodilator neurons 224 facial nerve 372. 380 facial nerve lesion 326. 328 facial nucleus 76. 184. 322. 323.326. 328. 391 fasciculus retroflexus 75 Fast Blue 235 feeding behavior invertebrates 428 feeding motor neurons 429. 430 fetal mesencephalic neurons 346 flickering light NO release 133 Fluorogold ! 91.221 follicle stimulating hormone see FSH food deprivation 154 footshock 158. 159 fornix 32, 73, 75 fovea 12 l foveola 121 frontal cortex 362, 388. 397 FSH 161 fundus 236 GABA 27, 28, 160 nNOS colocalization 22, 38. 91. 188. 197 retina 114. ll5 galanin 27 nNOS colocalization 149. 189 galanin message-associated peptide 327 gallbladder 239 Gallus domesticus 67 ganglion cell degeneration 115 ganglion cells (see also retina) cGMP-gated channels 132 gap junction amacrine cells 131 cone bipolar cells 131 horizontal cells 129. 130

NO 138, 166 gastric fundus 191 gastric mill 430-433 gastric motility 205 gastric relaxation 205 gastrin releasing peptide 239 gastrointestinal tract 3, 204, 205. 232 GDNF 338 Gekko gecko 63-65, 67 geniculotectai pathway 28 gigantocellular field 183, 187, 197 gigantocellular reticular nucleus 192 gigantocellular tegmental field 193 globus pallidus 73 glucocorticoid 157 glucoprivation 157 glutamate 157 circadian rhythm 164 LHRH release 162 glutamate receptors nNOS colocalization 37. 189. 193. 197 glutamate release cGMP 93 ON cone bipolar cells 136 glutamic acid decarboxylase 27 glycine nNOS colocalization 38. 188 gonadal steroids 162 gonadectomy 317 gracile nucleus 36, 76, 87, 180-182, 197 granuiocyte-macrophage colony stimulating factor LHRH release 162 growth arrest 92 growth cone 124, 427, 428 growth hormone related peptide nNOS colocalization 36 growth-associated protein 43 327 habenula 55, 57, 65, 67, 70, 75.365.368 heart 220, 224, 229, 231

Subject Index

heart failure 157 heme oxygenase 39 heme proteins NO transport 437 hemorrhage 159 hindlimb vasculature 226 hippocampus 24, 27, 70, 73, 81, 82, 189, 362, 388, 389, 396, 398, 400, 403, 405 development 393 NADPH-diaphorase 25, 88, 89 Hirudo medicinalis 429 histamine release 93, 167 Homarus 432 horizontal cells (see also retina) 119, 129-131 1H-[1,2,4]oxadiazolo[4,3-c~]quinoxaUn-l-one see ODQ human retinal pigment epithelial cells 114 huntingtin 28 Huntington's disease 28 Hydra vulgaris 417, 428, 429 hyperosmolality 160, 161 hyperstriatum 67 hypertension 156 hypertrophic pyloric stenosis 236 hypogastric nerve crush 321 hypoglossal nerve 326, 328, 372, 380 hypoglossal nucleus 65, 76, 77, 86, 87, 183, 188, 193, 322, 326, 328, 391 hypophysectomy 166. 317319, 327, 332, 333 hypotension 157, 160 hypothalamic paraventricular nucleus 75 hypothalamic slice preparation 164 hypothalamus 28, 32, 34, 38, 55, 57, 59, 61, 70, 75, 147, 154-157, 159, 189, 317, 332, 365, 389 hypothalamus anterior 161 hypothyroidism 151 hypovolemia 161 hypoxia 206 ibogaine 321,326

ICC see interstitial cells of Cajal IGF-I 338 ileum 235 llyanassa obsoleta 419 immediate early gene expression see c-fos c-jun immobilization stress 158 immune stress 157 indiseum griseum 400 inducible isoform of NOS see iNOS

inferior colliculus 30, 31.75. 367 inferior mesenteric ganglion 219, 235, 239 inferior olive 182. 184, 187. 188, 193. 326. 372. 382 infratrigeminal nucleus 37 infundibulum 61 iNOS 3 hypothalamus 165 macrophages 4 retina 113. 114. 125. 126. 138 spinal cord root avuision 324 insulin secretion 239 intercollicular commissure 31 interfascicular nucleus 76 interleukin-1 160 intermedial reticular nucleus 197 intermediate reticular formation 372 intermediate reticular nucleus 182-185, 187. 192. 381 intermediolateral cell column (see also spinal cord) 38, 217. 218, 221,223, 226 internal capsule 365. 368, 370 interpeduncular nucleus 33. 34, 76, 367. 374 interspecies differences see comparative aspects interstitial cells of Cajal 235. 236 intestinofugal neurons 223. 232. 238 invertebrate

development 419, 420, 422-427 nervous systems 417 iris 221 ischemic injury 342 islands of Calleja 23, 26, 82, 361. 362. 382. 389. 394, 397 development 393 isthmo-optic nucleus 122 isthmus 55, 57, 67, 70 jugular ganglion 240, 242 KC! 160 kyotorphin 160 lactation 151, 159 Lampetra planeri 55 Landolt clubs 119 lateral anterior hypothalamic nucleus 149 lateral geniculate nucleus 28, 65, 70, 75, 83. 365, 369, 391 development 92 NMDA receptors 124 lateral hypothalamus 147 lateral medial mammillary nucleus 32 lateral paragigantocellular field 187. 189, 197 lateral posterior nucleus 28 lateral reticular field 193 lateral reticular nucleus 37, 182. 187, 193. 197 lateral septum 188 lateral tegmental field 189. 192, 193 lateral tegmental nucleus 28 lateral vestibular nucleus 325, 327, 337 laterodorsal tegmentai nucleus 35, 81, 85 lemniscal nucleus 87 lentiform nucleus 65 leptin 162 LH 93. 161, 162 LHRH 161-163 LHRH release 93, 162 light adaptation 128-132, 137 lip chemosensory afferents 430 447

Subject hzdex Locusta 418

locus ceruleus 34, 65, 70, 122, 206, 372, 379 longitudinal fasciculus 57 lordosis behavior 163 lower airways 240 lower esophageal pressure 205 lower esophageal sphincter 235, 236 Lucifer Yellow 130 lumbar sympathetic ganglion 226 lumbosacrai parasympathetic preganglionic neuron 221 luteinizing hormone see LH luteinizing hormone-releasing hormone see LHRH Lymnaea stagnalis 418, 419, 428-430, 435 Macaca fascicularis

retina 116 macrophages 334 magnocellular preoptic nucleus 61 mammillary body 33, 57, 75, 149 mammillary nucleus 328, 371 mammilio-thalamic tract 83, 328 Manduca sexta 418-420, 424-426, 435-437 Marthasterias glacialis 429 maternal behavior 161 medial geniculate nucleus 28, 31 medial gigantocellular field 184 medial habenula 33 medial longitudinal fasciculus 187 medial mammillary nucleus 32 medial parabrachial nucleus 379 medial preoptic area 162 gonadal hormone 151 medial vestibular nucleus 36, 76, 182, 184, 197, 361,372, 378 448

median eminence 149, 156159. 161. 162. 317. 318, 367 median raphe nucleus 76 mediodorsal thalamic nucleus 28 medulla obiongata 177, 180, 181 melatonin 152 Melopsittacus undulatus 67 mesencephalic nucleus profundus 61 mesencephalic reticular formation 57.70, 379 mesencephalic trigeminal formation 57 mesencephalic trigeminal nucleus 61.65.70 metamorphosis 419. 422. 424--427. 433. 435 methylmercury chloride 321 middle cerebral artery 225 midline thalamic nucleus 28 milk ejection 161 mitochondrial metabolism 138 molluscs 419 motor networks invertebrates 429 morphine withdrawal 157 Mus musculus 75 myenteric ganglion 235, 237 esophagus 238 ileum 235 proximal colon 235 myenteric plexus 233. 235 esophagus 241 myoids 117 Na'-/K+-ATPase 138 NADPH-diaphorase astrocytes 89 BSPT as a substrate 113 oligodendrocytes 89 validation 20, 25. 53, 54, 87, 89, 112, 188. 216 nasal mucosa 21. 224, 225. 244, 245 nasociliary nerve 228 natriuretic factors nNOS colocalization 36 retina 135

necrosis 330, 332 nerve growth factor 158 nervus abducens 57 nervus vagus see vagus nerve neural regeneration 341,345 bulbectomy 345 nNOS expression 345, 346 neurodegeneration 342 excitotoxicity 344, 345 nNOS expression 341,344, 345 neuronal NO synthase see nNOS neuropeptide Y see NPY neuroprotection nNOS expression 345 neurotrophic factors 338, 341 deprivation 341 nNOS expression 341 spinal motoneurons survival 338 N~-hydroxyarginine 4, 5 nitric oxide synthase see NOS nitrosylation 128 nitrovasodilators clinical background 1 nitroglycerin 2 S-nitrosothiols 2 nitroxyl 6 protonated 5 NMDA 151, 157, 161 LHRH release 162 NMDA-R1 receptor 151 nNOS colocalization 31 nNOS (see also NOS) 10 aging 328 antisense probe 162, 163 chromosomal localization human 10 development 21, 78-83, 85, 86, 329, 390, 391,393 mossy fibers 91 nigro-striatal connections 92 retina 122-124 thalamo-cortical connections 92 gene regulation 10, 345 negative feedback 11 retina regional distribution 120 retrograde tracing

Subject Index locus ceruleus 122 retina I 15, 117 spinal cord 329 subcellular distribution 10, 332, 333 superoxide production 10 upregulation 163 proestrus 162 nNOS colocalization 21 A2 cell group 194 adrenal gland 246 after nerve injury 327 airways nerve fibers 240 basal ganglia 27, 28 cerebellum 36 cerebral cortex 22, 24 comparison 188, 189 cuneate nucleus 382 diagonal band of Broca 26, 27 dorsal root ganglion 229 enteric inhibitory motor neurons 233 esophagus 238 gallbladder 239 granule cells 378 habenula 365 hypothalamus 149 inferior olive 382 intermediate reticular nucleus 381 intestinofugal neurons 239 islands of Calleja 362 lateral geniculate nucleus 365 laterodorsal tegmental nucleus 35 mammillary body 33 medial preoptic area 151 medulla oblongata 188, 192, 193, 197 myenteric ganglia 237 nasal mucosa 245 nucleus of the solitary tract 198 olfactory bulb 21 paraventricular hypothalamic nucleus 31 pedunculopontine tegmental nucleus 35 perivascular plexus 225 raphe nuclei 34

raphe obscurus nucleus 195 regenerating vestibular neurons 336 reticular formation 372.381 rostral ventral medulla 37 rostral ventrolateral medulla 189 salivary gland 242 species differences 91 sphenopalatine ganglion 222 spinal cord 38 spinal trigeminal nucleus 198 stellate ganglion 244 sublingual gland 243 submandibular ganglion 243 suprachiasmal nucleus 31 supraoptic nucleus 159 tegmental nuclei 36 vasodilator neurons 226. 227 ventromedial hypothalamic nucleus 32. 151 nNOS de-novo expression apoptosis 344 axotomy 326. 328 axon length 334-336 bulbectomy 322 c-tim 327. 339 cerebellar cortex 321,322, 325.332 cerebral cortex, layer V 321 Clarke's nucleus 324-326. 336, 338 neurodegeneration 342 corticospinal tract 325 cranial nerve ligation 322 cranial nerve nuclei 321 development 328 dorsal motor vagal nucleus 328 dorsal root ganglion 328 dynorphin 324 facial nerve lesion 326. 328 facial nucleus 322. 323. 326. 328 neurodegeneration 342 hypoglossal nerve 326. 328 hypoglossal nucleus 322. 326, 328 hypophysectomy 327. 332. 333

hypothalamus 332 ibogaine 32 I. 326 inferior olive 326 lateral vestibular nucleus 325. 327. 337 mammillary nucleus 328 mammillo-thalamic tract 328 methylmercury chloride 321 necrosis 344 neocortex 321. 326, 327 neurodegeneration 342, 343 neurotrophic factors 339 NMDA receptor stimulation 340 olfactory epithelium 321 oxidative stress 344 p75 339 paraventricular hypothalamic nucleus 333 peripheral nerve graft 336 Purkinje cells 321,322, 325, 326, 332 root avulsion 323, 324, 327, 328, 330, 332, 334, 336, 339 sciatic nerve transection 327, 328 species differences 330 spinal cord 323, 326. 327 activated macrophages 334 spinal cord hemisection 325 spinal motoneurons 324, 326-328. 330, 332-334, 336 neurodegeneration 342 neurotrophic factors 338, 341 spinal root avulsion 326 spinocerebellar tract 324 traumatic injury 321. 322, 324, 326 vagus nerve 328 vestibulospinal tract 325 nNOS downregulation 163 adrenal medullectomy 321 axotomy 321 spinal motoneurons neurotrophic factors 338 survival 338 splanchnic nerve 321 449

Subject hldex nNOS knockout mice 25, 236 neurodegeneration 343 nNOS retrograde tracing 38, 179 adrenal medulla 219 cardiac vagal preganglionic neurons 199 dorsal motor vagal nucleus 192, 202, 205 gastric fundus 191 inferior mesenteric ganglion 235 intermediolateral cell column 223 middle cerebral artery 225 myenteric plexus 244 nucleus of the solitary tract 203 pelvic ganglia penile shaft 226 raphe obscurus nucleus 201 sphenopalatine ganglion 245 stellate ganglion 244 submandibular ganglion 221 trachealis muscle 241 nNOS upregulation anterior pituitary 317 autonomic nervous system 320 axonal transport interruption 340 capsaicin 319 colchicine 317 cranial nerve nuclei 317 dorsal motor vagal nucleus 316, 317, 340 dorsal root ganglion 319, 320 gonadectomy 317 hypogastric nerve crush 321 hypophysectomy 317-319 hypothalamus 317 median eminence 317, 318 neurodegeneration 343 nodose ganglion 320 nucleus ambiguus 317 nucleus of the solitary tract 317 paraventricular hypothalamic nucleus 317-319 peripheral nerve 319 450

peripheral nerve axotomy 319 peripheral nerve injury 317 peripheral nerve transection 320 salt loading 317 solitary tract 317 spinal cord 317 spinal cord ischemia 317 spinal cord hemisection 317. 320 supraoptic nucleus 317-319 hypophysectomy 166 osmotic stress 166 suckling 166 trigeminal ganglion 320 vagotomy 316. 317 vagus nerve 340 NO diffusion 361,378 identification 4 ion channel 128 neurotoxicity 3 retrograde messenger 3 NO-binding proteins 436. 437 nociception 37. 206. 435 development cGMP 437 invertebrates 435-437 nodose ganglion 36. 189. 193. 202. 229, 240, 242, 320 nonadrenergic-noncholinergic neurotransmission 3, 215, 232 noradrenaline 215.219, 224. 241 LHRH release 162 NOS activity 54 calmodulin 8.9 chemistry 4. 8.9 crystal structure 9. 10 expression estradiol 33 heme 4, 7.8 history 1 human 7 identification 2 inhibitors 8 isoforms 3.6. 7. 10. 93. 138 negative feedback 157 nomenclature 4

polypeptide insert 8 protein structure 6, 7 retrograde tracing 38 tetrahydrobiopterin 9 functions 5 NOS development invertebrates 419, 420, 425, 427 cGMP 420 NOS interneurons intestine 237 NOS knock-out mice GABA release 92 glutamate release 92 NPY 219, 224, 227, 230, 240 estrogen release 162 nNOS colocalization 21, 24, 27, 38, 189, 193, 240, 241 nucleus accumbens 27, 59, 63, 65, 73,362 nucleus ambiguus 37. 182185. 187-189, 191-193, 197. 204, 222, 238, 317, 361,391 esophageal afferents/efferents 36 nucleus arcuatis 149 nucleus basalis magnocellularis 27 nucleus circularis 33, 147, 149, 154 nucleus fasciculi longitudinalis 61 nucleus intercalatus 182 nucleus intermedius 184, 192 nucleus of Probst's bundle 182. 184, 192, 197, 362 nucleus of Roller 182-184, 192, 197 nucleus of the olfactory tract 82 nucleus of the optic tract 29 nucleus of the solitary tract 37, 63, 65, 71, 179, 189, 190, 192-194, 198, 199, 201-203, 317, 361, 372, 383 blood pressure 200 carotid sinus nerve 206 esophagus 204 hypoxia 206

Subject Index intestine 204, 205 nociception 206 nodose ganglionectomy 189 respiratory control 205, 206 stomach 204, 205 subnuclei 36, 180-182, 184, 185, 187, 197, 204, 205 nucleus prepositus hypoglossus 36, 182 oculomotor nucleus 57 ODQ 237, 387, 419, 432, 435 olfactory bulb 21, 22, 34, 55, 59, 63, 67, 71, 75, 363 development 21 olfactory epithelium 21,321 olfactory placode 21 olfactory tubercle 63, 67, 73, 80, 362 Oncorhynchus mykiss 55 optic chiasma 65 optic disk 121 optic nerve cGMP 137 optic nerve head 118, 122 optic nerve transection 117 optic tectum 124 optic tract 32, 57, 65 optic tract lesions 114, 365, 370 orbital cortex 396 organum vasculosum 149. 151, 162 fever 165 organum vasculosum of the lamina terminalis 32, 33 osmotic stress 166 otic ganglion 221,242 ovariectomy 151, 162, 163 oxytocin 157, 159, 161 nNOS colocalization 31, 91, 149, 159 oxytocin release 93, 159-161 L-arginine 160 kyotorphin 160 NO 160 p75 27, 327, 339, 341 regenerating motoneurons 336 spinal motoneurons 341 PACAP 149, 230, 232, 233, 235, 244

paleostriatal-paraolfactory lobe complex 67 paleostriatum 69 pallidum 26. 55.59. 63.77 pancreas 239 pancreatic ganglia 239 parabrachial nucleus 76. 87 paragigantocellular nucleus 37.71 paralemniscal nucleus 76 paramedian paragigantocellular field 197 paramedian reticular field 183, 187, 193. 197 paramedian reticular nucleus 182 paramedian tegmental field 192, 193 paranigral nucleus 76 parapineal organ 55 parateniai nucleus 28.83 77 paratrigeminal nucleus 76, 87. 181. 191 paraventricular hypothalamic nucleus 75. 83. 147, 149.151. 152. 154. 156-161. 166. 317-319. 333. 340 blood pressure 164 colchicine 340 paraventricular thalamic nucleus 28.29. 75.361. 365 paravertebral ganglion 217. 219. 223, 244 parietal cortex 362. 396 parotid gland 242 particulate guanylate cyclase NO-sensitivity 127 retina 126, 127 colocalization 135 parturition 161 parvaibumin 398, 400 nNOS colocalization 2 I. 91. 197 parvocellular paragigantocellular field 197 parvoceilular reticular field 183-185. 197 pectin 122 pedunculopontine nucleus 69 pedunculopontine tegmentai nucleus 28, 70, 85 pelvic ganglion 226. 227

pelvic plexus 226 penile erection 161 penis 226, 227 periaqueductai gray 31.33 pericytes retina 117 perilemniscic nucleus 57 periolivary nuclei 31 peripheral autonomic nervous system 215, 248 peripheral nerve 319 peripheral nerve axotomy 319 peripheral nerve graft 336 nNOS expression 341 peripheral nerve injury 317 peripheral nerve transection 320 peripherin nNOS colocalization 24 peristalsis 238 periventricular hypothalamic nucleus 57, 65, 149, 153. 163 periventricular thalamic nucleus 70

Petromyzon marinus 55 petrosai ganglion 188 phenylethanolamine N-methyl transferase nNOS colocalization 192 phosphodiesterase 129. 131, 132. 136. 137. 224. 357, 382, 389, 390, 400. 405, 420 cGMP immunocytochemistry 360. 361 hippocampus 358 localization 357 regulation 358 phosphodiesterase-2 131 photoreceptor axons Drosophila 428 NO-responsive Drosophila 427 photoreceptors (see also retina) cGMP-gated channels 128, 129 contraction NO 129 dark adaptation 129 451

Subject Index Drosophila 420, 422, 424 light adaptation 129 cGMP 128 phototransduction 126 phylogeny 90 pineal 33, 55, 59, 152 piriform cortex 21, 67, 73, 77. 362, 365 pituicytes 166 pituitary 31, 59, 149, 159-161 gonadal hormones 151 pituitary adenylate cyclaseactivating polypeptide see PACAP pituitary stalk transection 157 Pleurobranchia 418 Pleurodeles waltl 59 polymorphism 125 pons 372 pontine nuclei 71, 87 pontine reticular formation 76, 189, 372 pontine reticular nucleus 381 posterior central thalamic nucleus 57 posterior cerebral artery 225 posterior commissure 33, 34, 57, 61, 83 posterior commissure nucleus 61 posterodorsal tegmental nucleus 372, 379 postsynaptic density 95, 138 pregnancy 151,227 preoptic area 61, 65, 67, 75, 83, 161, 165, 365 preoptic magnoceilular nucleus 57 preoptic nucleus 149 preoptic region 57 pretectal nucleus 65, 70, 75, 369 prevertebral ganglion 217, 219, 223, 238, 239 progesterone LH release 162 prolactin release 163 proliferation zone Manduca sexta 419 prostaglandin E2 159, 165, 167

452

LHRH release 162 protein kinase A 130, 132 protein kinase C 136 proximal colon 235 Pseudemys scripta elegans 63. 65 psychogenic stress 157, 158 pterygopalatine ganglion 224 Purkinje cells 37, 57, 70, 86, 321. 322, 325, 326, 332 cGMP 384-387 pyloric sphincter 235, 236, 430-432 pyramidal decussation 86 pyramidal tract 187 Rana esculenta 61 Rana perezi 59-61.63 raphe magnus 206 raphe nuclei 34. 37, 63, 67, 86, 87, 189, 203, 206 raphe obscurus 86, 183, 184, 187, 191, 192, 195, 201 red nucleus 57, 367,372 reproductive behavior 161 respiratory control 205.206 restraint stress 159 reticular formation 55. 59. 65, 71.86. 184. 381 reticular nucleus 86 reticular thalamic nucleus 365,400, 403. 405 retina 55, 59. 61, 111, 115, 117, 125, 164. 188 afferents 122 blood flow 115 cGMP content 135, 136 cGMP signal transduction pathway 133-137 degeneration 123 development 124 preprogramming 123 dopamine 130. 131, 137 dopamine release GABAA receptors 133 electron microscopy 115, 118 eNOS 125 development 122-124 GAB A re lease 133 GABAergic transmission 137

ganglion cell death 123 glutamate receptors 132 glutamate release 129, 133 photoreceptors 131 glutamine release 133 iNOS 125 natriuretic factors 135 nNOS regional distribution 120 NO donor release 133 NO release 132 NOS isoforms 113, 138 NO synthesis 126 projections Drosophila 427 vasculature 115, 117, 123, 138 retino-collicular projections 124 retinogeniculate development 124 retinorecipient cells 31 retractor penis muscle 215 retrograde tracing (see also nNOS retrograde tracing) technique 179 Rhodnius prolixus 437 rhomboid nucleus 28 ribbon synapse 115, 118 root avuision 323, 324, 327-330, 332, 334-336, 339 rostral medulla 192 rostral ventrolateral medulla 189, 192, 194, 199, 200, 202, 203, 206 GABA 206 nociception 206 sacral preganglionic NOS neuron 223 salivary gland 220, 224, 225, 242-244 Salmo salar 55 salt loading 157, 159, 317 schizophrenia 157 sciatic nerve 328 sciatic nerve transection 327, 328 sensory vasodilation 228 septohippocampal pathway 27 species differences 90

Subject Index

septum 27, 59, 65, 67, 70, 73, 75, 82, 391 development 394 serotonin 34 circadian rhythm 164 nNOS colocalization 34, 188, 192, 195 retina 116 sexual behavior 163 skeletal muscle 220, 226 solitary tract 190, 193, 317 soluble guanylate cyclase 131-133 carbon monoxide 39 cerebellum 384-387 YC- 1 385 development 389, 390, 393, 394, 396, 397, 432 glutamate receptors 137 immunocytochemistry 356, 386 in situ hybridization 386 invertebrates 418 localization development 394, 397 NO insensitive 433 retina 126-129, 135-137 somatomotor areas 183 somatosensory areas 180 somatostatin 230 nNOS colocalization 21, 24, 27, 31, 37, 91, 149, 163, 189, 192, 197 somatostatin release 163 cGMP 163 growth hormone-releasing factor 164 species differences see comparative aspects sphenopalatine ganglion 21, 33, 221,222, 224, 225, 245 sphincter muscle 232 sphincter of Oddi 235 spinal cord (see also intermediolateral cell column) 37, 38, 88, 188, 189, 206, 317, 323, 326, 327, 329, 330, 334, 336, 339, 372, 383, 400 amphibians 63, 67 cervical segments 63 cyclostomes 59

development 393 lumbar segments 38. 226 mammals 76 sacral segments 38, 76. 220, 223 teleosts 59 thoracic segments 38. 76. 189. 217-219. 221. 240 spinal cord hemisection 317, 320, 325 spinal cord ischemia 317 spinal motoneurons 324, 326-328, 330, 333-336, 341 spinal root avuision 326 spinal trigeminal nucleus 36. 37, 57. 180-185. 187, 192, 193. 197. 198 spinal vestibular nucleus 182 spinocerebellar tract 324 splanchnic nerve 223, 321 stellate ganglion 219. 220. 228, 240, 241,244 steroidogenesis 158 stomach 204, 235, 236 stomach mucosa 238 stomatogastric ganglion 428. 430-433 development cGMP, 435 stratum album profundum 31 streptozotocin 156 stress 152, 153. 157-159 stria terminalis 29 striate cortex 27 striatum 23, 26. 27. 28, 59. 63, 65, 67.73, 77, 82, 189 subcommissural organ 33, 55 subcortical white matter see corpus callosum subfornical organ 32, 33 subiculum 24, 25, 73 sublingual gland 242. 243 submandibular ganglion 221, 243 submandibular gland 242 substance P 215, 230, 231 nNOS colocalization 28, 33, 36, 149, 188, 189, 245, 327 substantia innominata 73

substantia nigra 34, 367, 373, 400 substantia nigra pars compacta 65, 76 substantia nigra pars reticulata 65 subthalamic nucleus 29, 83, 365, 368 suckling 166 superficial ventral medulla 195 superior cervical ganglion 219, 220, 240, 244, 245, 340 cGMP 223 colchicine 340 superior colliculus 29, 30, 75, 77, 85, 124, 361,367, 375, 389, 391,400 superior mesenteric ganglion 219, 223 superior olive 63 superoxide dismutase 28 suprachiasmatic nucleus 70, 149, 151,365, 367 c-fos 164 cGMP 165, 365 heart rate 165 suprachiasmatic process 61 suprageniculate nucleus 31 supramammillary nucleus 32, 367 supraoculomotor cap 34 supraoptic decussation 149 supraoptic nucleus 32, 75, 83, 147, 151, 152, 156, 157, 160, 161, 166, 317-319 A23187 165 cGMP 165 dye coupling cGMP 166 heart rate 165 NMDA receptors 165 sweat glands 220, 244 sympathetic excitatory neurons 229 sympathetic neurons pre- and postganglionic 217-219 species differences 217 synaptic vesicle 133 453

Subject Index synaptogenesis 122, 123, 393, 420, 424, 425,427 tachykinins 228, 230, 237 taenia tecta 67 Taeniopygia guttata 67 tectum 30, 57, 61, 63, 65, 67, 69, 70 tegmentai nucleus 76 tegmentum 33, 35, 61, 63, 65. 69, 85 teleosts 89 temperature body 165 temporal artery 225 temporal cortex 31 testosterone 163 tetrahydrobiopterin 10 function in NOS catalysis 10 thalamus 28, 29, 38, 55, 57, 61, 65, 75, 83 thoracic sympathetic ganglia 229 thyroid hormones 151 Tinca tinca 55 tongue 224, 225 trachea 228, 240, 241 transplantation 336 trapezoid 31 traumatic injury 321, 322, 324, 326, 342 trigeminal ganglion 244, 245. 320 trigeminal nerve 59, 63 trigeminal nucleus 61, 65, 69. 71, 76, 86, 87 Trimeresurus flavoviridis 63

454

trochlear nucleus 57 tuber cinereum 149 tyrosine hydroxylase 230 nNOS colocalization 21.3 I. 34. 149. 192. 194. 226. 245 prolactin release 163 uNOS 430 uterine artery 227 uterus 226 uveitis 114 vagai afferents 189 vagal efferents 191 vagal parasympathetic pathways 229 vagal preganglionic neurons 230 vagal sensory ganglion 228 vagotomy 222. 231, 316. 317 vagus nerve 36. 179. 191.221. 230. 231. 236. 239. 240. 328, 340 vasculature neural control 224-231 vasoactive intestinal peptide see VIP vasopressin 157. 159 nNOS colocalization 91, 149. 159. 189. 237, 238. 239. 242-244 vasopressin release 159-161 ventral gigantocellular field 187 ventral medullary nucleus 181 ventral pallidum 362

ventral paragigantocellular field 197 ventral reticular area 187 ventrolateral geniculate 29 ventrolateral medulla 192, 193 blood pressure 200, 202, 203 ventromedial hypothalamic area 149, 151, 155, 163 ventroposterior thalamic nucleus 38 vestibular nuclei 65, 71, 87, 187 vestibulospinal tract 325 vidian nerve 245 VIP 31,215. 225-227, 232. 233, 235-237. 239-244, 327 nNOS colocalization 31, 149. 226, 227, 23, 245. 246 VIP release submandibular gland 244 visceromotor areas 183 viscerosensory areas 180 visual cortex 24 visual streak 120, 121, 123 vomeronasal organ 21, 59, 72, 88 water deprivation 157, 159

Xenopus laevis 59 Xiphophorus helleri 55 yawning 16 l YC-1 400. 401. 405 Zaprinast 131